The Rat Nervous System, Third Edition (Paxinos, The Rat Nervous System)

THE RAT NERVOUS SYSTEM THIRD EDITION

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THE RAT NERVOUS SYSTEM THIRD EDITION Edited by

GEORGE PAXINOS Prince of Wales Medical Research Institute The University of New South Wales Sydney, Australia

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Elsevier Academic Press 525 B Street, Suite 1900, San Diego, California 92101-4495, USA 84 Theobald’s Road, London WC1X 8RR, UK This book is printed on acid-free paper.

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Cover image: Figure 17.12, Panel A: illustrates the mixing of neurons that stain with antiserum against ORX (brown) and with a digoxygenin-labeled probe for MCH mRNA (blue) in the perifornical region of a rat. Although the two types of neurons cluster closely with one another around the edge of the fornix, there is virtually no colocalization within individual neurons. Modified from Elias, C.F., Saper, C.B., Maratos-Flier, E., Tritos, N.A., Lee, C., Kelly, J., Tatro, J.B., Hoffman, G.E., Ollmann, M.M., Barsh, G.S., Sakurai, T., Yanagisawa, M., and Elmquist, J.K. (1998b). Chemically defined projections linking the mediobasal hypothalamus and the lateral hypothalamic area. J. Comp. Neurol. 402, 442–459.

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Contents

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Contributors xiii Foreword xvii

PERIPHERAL NERVOUS SYSTEM AND SPINAL CORD 3. Autonomic Nervous System

I

GIORGIO GABELLA

DEVELOPMENT

Localization of Autonomic Ganglia 77 Structure of Autonomic Ganglia and Nerves 84

1. Gene Maps and Related Histogenetic Domains in the Forebrain and Midbrain

4. Primary Afferent Projections to the Spinal Cord

LUIS PUELLES, SALVADOR MARTÍNEZ, MARGARET MARTÍNEZ-DE-LA-TORRE, AND JOHN L. R. RUBENSTEIN

GUNNAR GRANT AND BRITA ROBERTSON

Projection of Primary Afferent Fibers to Different Laminae and Some Spinal Cord Nuclei 112 Somatotopic Organization of Primary Afferent Projections 114

Molecular Versus Anatomical Distinction of Brain Subdivisions: The Specification State 3 Sharing of Molecularly Distinct Brain Domains Among Vertebrates 5 Differential Aspects of Histogenesis 6 The Bauplan of the Brain 7 The Neural Plate Subdivisions 11 The Closed Neural Tube 13 Basal Plate Regions 14 Alar Plate Regions 16 Telencephalic Patterns 17 About Mechanisms 20 Relevant Genetic Mechanisms 20

5. Spinal Cord Cytoarchitecture GUNNAR GRANT AND H. RICHARD KOERBER

Lamina I 121 Lamina II 122 Lamina III 122 Lamina IV 122 Lamina V 123 Lamina VI 123 Lamina VII 123 Lamina VIII 124 Lamina IX 124 Area X 125 Lateral Spinal Nucleus 126 Lateral Cervical Nucleus 126

2. Development of the Telencephalon: Neural Stem Cells, Neurogenesis, and Neuronal Migration SHIRLEY A. BAYER AND JOSEPH ALTMAN

Neurogenetic Timetables in the Telencephalon 28 Maps of Stem Cell Mosaics in the Telencephalic Neuroepithelium 36 Development of the Lateral, Rostral, and Dentate Migratory Streams 66 Stem Cell Dynamics in Cortical Germinal Zones 68

6. Substantia Gelatinosa of the Spinal Cord ALFREDO RIBEIRO-DA-SILVA

Definition 129 Characteristics of Neurons of the Superficial Laminae of the Spinal Cord 130

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Ultrastructure of the Spinal Dorsal Horn 133 Neurochemistry of the Dorsal Horn 136 Final Remarks 143

Efferent Projections of LC Neurons 277 Other Metencephalic Noradrenergic Neurons (A5 and A7 Cell Groups) 284 Conclusions 284

7. Ascending and Descending Pathways in the Spinal Cord

12. Oromotor Nuclei

DAVID TRACEY

JOSEPH B. TRAVERS

Ascending Pathways 149 Descending Pathways 154

III BRAINSTEM AND CEREBELLUM 8. Precerebellar Nuclei and Red Nucleus TOM J. H. RUIGROK

Pontine Nuclei 167 Lateral Reticular Nucleus 174 Inferior Olivary Nucleus 180 Red Nucleus 187

9. Cerebellum JAN VOOGD

The Gross Anatomy of the Cerebellum 205 The Cerebellar Nuclei and Their Efferent Pathways 208 Longitudinal, Zonal Organization of Purkinje Cells in the Cerebellar Cortex: Chemoarchitecture and Connections 216 Afferent Mossy Fiber Systems 229 Terminations of Mossy Fiber Systems in Different Regions of the Cerebellum 231

Motor Trigeminal Nucleus 295 Facial Nucleus 301 Hypoglossal Nucleus 305 Summary and Conclusions 311

13. Central Nervous System Control of Micturition GERT HOLSTEGE

Motoneurons Innervating Bladder and Urethral Sphincter 321 Sacral Cord Micturition Reflexes 323 Brain Stem–Spinal Cord Pathways Coordinate Bladder and Sphincter Motoneurons 323 Afferent Systems 324 Forebrain Involvement in the Control of Micturition 325 Micturition Control in Humans 326 Conclusions 327

IV DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

10. Periaqueductal Gray

14. Anatomical Substrates of Hypothalamic Integration

KEVIN A. KEAY AND RICHARD BANDLER

RICHARD B. SIMERLY

PAG Columnar Organization 244 Anatomical Studies 247 The PAG And Parallel Circuits for Emotional Coping 249 Conclusions 253

11. Locus Coeruleus, A5 and A7 Noradrenergic Cell Groups GARY ASTON-JONES

Cytoarchitecture 259 Afferents to the Nucleus Locus Coeruleus 263 The Pericoerulear Region: The “Extranuclear LC” 274

Morphological Organization of the Hypothalamus 336 Hypothalamic Integration 352

15. Hypothalamic Supraoptic and Paraventricular Nuclei WILLIAM E. ARMSTRONG

Pituitary Gland 369 Supraoptic Nucleus 370 Paraventricular Nucleus 375 Accessory Magnocellular Neurosecretory Neurons 382 Conclusion 382

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16. Circumventricular Organs BRIAN J. OLDFIELD AND MICHAEL J. MCKINLEY

General Features 389 Subfornical Organ 390 Vascular Organ of the Lamina Terminalis 394 Area Postrema 397 Median Eminence and Neurohypophysis 398 Subcommissural Organ 399 Pineal Gland 400 Choroid Plexus 400

17. Thalamus HENK J. GROENEWEGEN AND MENNO P. WITTER

Some General Aspects of Thalamic Organization 408 Principal Thalamic “Relay” Nuclei 411 Association Thalamic Nuclei 425 Midline and Intralaminar Thalamic Nucle 433 Reticular Nucleus 441

18. Basal Ganglia CHARLES R. GERFEN

Cortical Input to the Striatum 458 Striatum 464 Indirect Pathway 478 Basal Ganglia Outputs 482 Dual Output Systems of Striatal Output Pathways 484 Nigrostriatal Dopamine System 488 Striatal Patch/Matrix Compartments 490 Summary 497

19. Amygdala and Extended Amygdala of the Rat: A Cytoarchitectonical, Fibroarchitectonical, and Chemoarchitectonical Survey JOSE S. DE OLMOS, CARLOS A. BELTRAMINO, AND GEORGE ALHEID

General Topography and Terminology 510 Description: Observation Procedures 514 Organization of the Rat Amygdaloid Complex and Extended Amygdala 514 The Extended Amygdala (EXA) 547 The Laterobasal Nuclear Complex (LBNC) 572 Unclassified Cell Groups in the Amygdala and Bed Nucleus of the Stria Terminalis 588

20. The Septal Region P. Y. RISOLD

Development of the Septal Region 602 Morphological Overviews and Cytoarchitecture of the Septal Nuclei 603

The Chemoarchitecture of the Septal Region 606 Connections of the Septal Region 615 Functional Organization of the Septal Region 621

V CORTEX 21. Hippocampal Formation MENNO P. WITTER AND DAVID G. AMARAL

Dentate Gyrus 637 Hippocampus 647 Overview of the Subiculum, Presubiculum, and Parasubiculum 658 Subiculum 660 Presubiculum 666 Parasubiculum 669 Entorhinal Cortex 670 Perirhinal and Postrhinal Cortices 684 Conclusions: The Organization of Hippocampal Circuitry and the Flow of Information Processing 687

22. Cingulate Cortex and Disease Models BRENT A. VOGT, LESLIE VOGT, AND NURI B. FARBER

Regional Organization 704 Is “Infra” Limbic Area IL Ventral to Limbic Cortex? 705 Cytology of Limbic Area 25 705 Modified Brodmann Nomenclature 705 Cytology of the Perigenual Anterior and Midcingulate Regions 707 Cytology of Retrosplenial Cortex 708 Opioid Architecture: Regional Differences and Neuronal Expression Patterns 709 Area 24B: Movement, Vision, and Pain Behaviors 710 Cortical Connections of Retrosplenial Cortex and Role in Visuospatial Function 711 Thalamic Afferents 712 NMDA Receptor Antagonist-Induced Neurotoxicity in Retrosplenial Cortex 714 Polysynaptic Circuit Disinhibition Underlies NRHypo Neurotoxicity 717 NRHypo-Induced Psychosis 719 NRHypo and Neurodegeneration in Alzheimer’s Disease 719 Comparison of Medial Cortex in Rat and Monkey 720 Rodent Models of Disease 721

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23. Isocortex

28. Gustatory System

NICOLA PALOMERO-GALLAGHER AND KARL ZILLES

ROBERT F. LUNDY, JR. AND RALPH NORGREN

Isocortex 728 Transition Regions Between Isocortex and Allocortex 747

VI SYSTEMS 24. Central Autonomic System CLIFFORD B. SAPER

Medullospinal Level: Reflex Control 761 Mesopontine Level: Modulation and Integration of Reflex Control and Arousal 766 Forebrain Level: Behavioral and Metabolic Integration of Autonomic Control and Arousal 774 Summary and Conclusion 784

25. Somatosensory System DAVID TRACEY

Somatosensory Receptors 795 Cell Bodies and Central Processes of Somatosensory Receptors 797 Ascending Spinal Pathways 798 Medullary Relay Nuclei 799 Somatosensory Thalamus 802 Somatosensory Cortex 803

26. Trigeminal Sensory System P. M. E. WAITE

Adult Sensory Trigeminal System 815 Development of the Trigeminal System 834

27. Pain System WILLIAM D. WILLIS, KARIN N. WESTLUND, AND SUSAN M. CARLTON

Nociceptors 853 Dorsal Horn Interneurons 856 Ascending Nociceptive Pathways 862 Thalamus and Cortex 866 Descending Control Systems 866 Plastic Changes in Pathological Conditions 868

Peripheral Anatomy 890 Central Organization 891 Cytoarchitecture 903 Neurochemistry 905 Functional Considerations 908 Conclusion 911

29. Olfactory System MICHAEL T. SHIPLEY, MATTHEW ENNIS, AND ADAM PUCHE

The Olfactory Epithelium 922 The Main Olfactory Bulb 925 Primary Olfactory Cortex 935 The Accessory Olfactory System 946 “Nonolfactory” Modulatory Inputs to the Olfactory System 950

30. Vestibular System PIERRE-PAUL VIDAL AND ALAIN SANS

The Vestibular Message: From the Periphery to the Center 964 The Vestibular Nuclear Complex: Morphofunctional Properties 966 Neurotransmitters and Neuromodulators of Central Vestibular Neurons 975 Conclusion 985

31. Auditory System MANUEL S. MALMIERCA AND MIGUEL A. MERCHÁN

The Organ of Corti 996 The Cochlear Nuclear Complex 1001 The Superior Olivary Complex 1011 The Nuclei of the Lateral Lemniscus 1018 The Inferior Colliculus 1027 He Medial Geniculate Body 1039 The Auditory Cortex 1049 The Descending Auditory Pathway 1056

32. Visual System ANN JERVIE SEFTON, BOGDAN DREHER, AND ALAN HARVEY

Visual Pathways 1082 Retinal Output 1084 Retino-Recipient Nuclei 1087 Associated Visual Nuclei 1119 Visual Cortex 1121

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33. Cerebral Vascular System OSCAR U. SCREMIN

Methodology 1165 Cerebral Blood Vessels 1166 Spinal Cord Blood Vessels 1190 Vascular Innervation 1192 Functional Localization With Blood Flow 1193

VII NEUROTRANSMITTERS 34. The Serotonin and Tachykinin Systems ANTONY HARDING, GEORGE PAXINOS, AND GLENDA HALLIDAY

Serotonin System 1203 Tachykinin System 1212

Coexistence of Serotonin and Tachykinins 1244 Functional Interaction Between Serotonin and Tachykinins 1245

35. Cholinergic Neurons and Networks Revisited LARRY L. BUTCHER AND NANCY J. WOOLF

Cholinergic Neuroanatomy in the Context of Function 1257 Central Cholinergic Neurons: Modes of Operation 1262 Genesis of Alzheimer’s Disease: A Hypothesis 1263

36. Glutamate JONAS BROMAN, ERIC RINVIK, MARCO SASSOE-POGNETTO, HOSSEIN KHALKHALI SHANDIZ, AND OLE PETTER OTTERSEN

Anatomical Systems 1269 Conclusion 1280

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Contributors

Numbers in parentheses indicate the pages on which the authors’ contributions begin. Alheid, George, (509) Department of Physiology and Institute for Neuroscience, Feinberg School of Medicine, Northwestern University, Chicago, Illinois, USA Altman, Joseph, (27), Department of Biology, Purdue University, Indianapolis, Indiana, USA Amaral, David G., (635), Department of Psychiatry, California National Primate Research Center and The M.I.N.D. Institute, University of California, Davis, California, USA Armstrong, Willaim E., (369), Department of Anatomy and Neurobiology, University of Tennessee, Memphis, Tennessee, USA Aston-Jones, Gary, (259), Laboratory of Neuromodulation and Behavior, Department of Psychiatry University of Pennsylvania School of Medicine, Philadelphia, USA Bandler, Richard, (243), Department of Anatomy and Histology, University of Sydney, Sydney, Australia Bayer, Shirley A., (27), Department of Biology, Purdue University, Indianapolis, Indiana, USA Beltramino, Carlos A., (509) Department de Neurofisiologia y Psicofisiologia, Facultad de Psicologia, Universidad Nacional De Cordoba, Cordoba, Argentina Broman, Jonas, (1269), Dept of Physiological Sciences, Lund University, Lund, Sweden Butcher, Larry L., (1257), Department of Psychology UCLA, Los Angeles, California, USA Carlton, Susan M., (853), Department of Anatomy, Neuroscience, Marine Biomedicine Institute, University of Texas Medical Branch, Galveston, Texas, USA

DeOlmos, Jose, S., (509), Instituo de Investigacion Medica, Mercedes y Martin Ferreyra, Cordoba, Argentina Dreher, Bogdan, (1083), Department of Physiology, University of Sydney, Sydney, Australia Ennis, Matthew, (923), Dept Anatomy and Neurobiology, University Maryland School of Medicine Baltimore, Maryland, USA Farber, Nuri B., (705), Department of Psychiatry, Washington University, St. Louis, Missouri Gabella, Giorgio, (77), Department of Anatomy and Developmental Biology, University College London, London, UK Gerfen, Charles, R., (455), Laboratory of Systems Neuroscience, Bethesda, Maryland, USA Grant, Gunnar, (111, 121), Department of Neuroscience, Karolinska Institutet, Stockholm, Sweden Groenewegen, Henk J., (407), Department of Anatomy, Vrije Universiteit, Amsterdam, The Netherlands Halliday, Glenda, (1205), Prince of Wales Medical Research Institute, Randwick, Australia Harding, Antony, (1205), Prince of Wales Medical Research Institute, Randwick, Australia Harvey, Alan, (1083), School of Anatomy and Human Biology, The University of Western, Crawley, Australia Holstege, Gert, (321, 1269), Department of Anatomy and Embryology, University Groningen Oostersingel, Groningen, The Netherlands Keay, Kevin A., (243), Department of Anatomy and Histology, University of Sydney, Sydney, Australia

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CONTRIBUTORS

Koerber, Richard, (121), Deparment of Neuroscience, Karolinska Institute, Stockholm, Sweden Lundy, Robert F., Jr. (891), Department of Behavioral Science, Pennsylvania State University, College of Medicine, Hershey, Pennsylvania, USA Malmierca, Manuel S., (997), Laboratory for the Neurobiology of Hearing, The Institute of Neuroscience of Castilla y Leon Faculty of Medicine, University of Salamanca, Salamanca, Spain Martinez, Salvador, (3), Department of Anatomy, University of Murcia, Murcia, Spain Martínez-de-la-Torre, Margaret, (3), Department of Morphological Sciences, University of Murcia, Murcia, Spain McKinley, Michael J., (389), Howard Florey Institute, University of Melbourne, Parkville, Australia Merchán, Miguel A., (997), Laboratory for the Neurobiology of Hearing, The Institute of Neuroscience of Castilla y Leon Faculty of Medicine, University of Salamanca, Salamanca, Spain Norgren, Ralph, (891), Department of Behavioral Science, Pennsylvania State University, College of Medicine, Hershey, Pennsylvania, USA Oldfield, Brian J., (389), Howard Florey Institute, University of Melbourne, Parkville, Australia Ottersen, Ole Petter, (1269), Centre for Molecular Biology and Neuroscience, Institute of Basic Medical Sciences, University of Oslo, Oslo, Norway Palomero-Gallagher, Nicola, (729), Institute of Medicine, Research Center, Julich, Germany Paxinos, George, (1205), School of Psychology, The University of New South Wales, Sydney, Australia Puche, Adam, (923), Dept Anatomy and Neurobiology, University Maryland School of Medicine Baltimore, Maryland, USA Puelles, Luis, (3), Department of Morphological Sciences, University of Murcia, Murcia, Spain Ribeiro-da-Silva, Alfredo, (129), Department of Pharmacology and Therapeutics, McGill University Montreal, Quebec, Canada Rinvik, Eric, (1269), Centre for Molecular Biology and Neuroscience, Institute of Basic Medical Sciences, University of Oslo, Oslo, Norway Risold, P. Y., (605), Laboratoire d’Histologie, Fac. Med. Université Franche-Comte, Besancon, France Robertson, Brita, (111), Department of Neuroscience, Karolinska Institutet, Stockholm, Sweden

Rubenstein, John L. R., (3), University of California, San Francisco, California Ruigrok, Tom J. H., (167), Department of Anatomy, Erasmus University, Rotterdam, The Netherlands Sans, Alain, (965), INSERM Unite 432, Neurobiologie et Developpement du Systeme Vestibulaire, Universite Montpellier II, Place Eugene Bataillon F34095 Montpellier, France Saper, Clifford B., (761), Department of Neurology, Beth Israel Hospital, Boston, Massachusetts, USA Sassoe-Pognetto, Marco, (1269), Dipartimento di Anatomia, Farmacologia e Medicina Legale, University of Turin, Italy Scremin, Oscar U., (1167), Veterans Affairs Greater Los Angeles Healthcare System, and Department of Physiology, UCLA School of Medicine, Los Angeles, California, USA Sefton, Ann Jervie, (1083), Department of Physiology, University of Sydney, Sydney, Australia Shandiz, Hossein Khalkhali, (1269), Centre for Molecular Biology and Neuroscience, Institute of Basic Medical Sciences, University of Oslo, Oslo, Norway Shipley, Michael T., (923), Dept Anatomy and Neurobiology, University Maryland School of Medicine Baltimore, Maryland, USA Simerly, Richard B., (335), Division of Neuroscience, Oregon National Primate Research Center, Oregon Health and Sciences University, Beaverton, Oregon Tracey, David, (149, 797), School of Medical Sciences, University of New South Wales, Sydney, Australia Travers, Joseph B., (259), Section of Oral Biology, OSU College of Dentistry, Columbus, Ohio, USA Vidal, Pierre Paul, (965), Laboratoire de Neurobiologie des Reseaux Sensorimoteurs, Universiteˇ Paris V, Paris, France Vogt, Brent A., (705), Cingulum Neurosciences Institute, Manlius, New York, USA Vogt, Leslie, (705), Cingulum Neurosciences Institute, Manlius, New York, USA Voogd, Jan, (205), Department of Anatomy, Erasmus University, Rotterdam, The Netherlands Waite, P. M. E, (817), School of Anatomy, The University of New South Wales, Sydney, Australia Westlund, Karin N., (853), Department of Anatomy, Neuroscience, Marine Biomedicine Institute, University of Texas Medical Branch, Galveston, Texas, USA

CONTRIBUTORS

Willis, William D., (853), Department of Anatomy, Neuroscience, Marine Biomedicine Institute, University of Texas Medical Branch, Galveston, Texas, USA

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Witter, Menno P, (407, 635), Department of Anatomy, Vrije Universiteit, Amsterdam, The Netherlands Zilles, Karl, (729), Vogt Brain Research Institute, University of Dusseldorf, Dusseldorf, Germany

1. SECTION TITLE

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Foreword

The structure of the nervous is the backbone of neuroscience. The aim of this book is to describe all parts of the rat nervous system in the context of modern hypotheses of structural and functional organization. No individual scientist could write with ultimate authority on every part of the brain, spinal cord and peripheral nervous system. It is for this reason that experts on different regions were asked to contribute to this book. The reader will notice that many of these

experts generated major hypotheses and original observations that today guide research in their field. It is hoped that the combined effort of contributors to the third edition of The Rat Nervous System will make this one an even friendlier and helpful companion to the graduate student or scientist wanting to learn the highlights and fundamentals of the structure of the nervous system. George Paxinos, Sydney January 2004

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SECTION

I

DEVELOPMENT

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C H A P T E R

1 Gene Maps and Related Histogenetic Domains in the Forebrain and Midbrain LUIS PUELLES1, SALVADOR MARTÍNEZ2, MARGARET MARTÍNEZ-DE-LA-TORRE1 and JOHN L. R. RUBENSTEIN3 1

Department of Human Anatomy and Psychobiology University of Murcia, Murcia, Spain

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Institute of Neuroscience/CSIC, University Miguel Hernandez San Juan de Alicante, Spain 3

Nina Ireland Laboratory of Developmental Neurobiology Langley Porter Psychiatric Institute, UCSF, San Francisco, USA

brain and midbrain patterns. To keep this essay within manageable size, we refer mainly to selected transcription factor genes, i.e., those whose coded proteins enter the cell nucleus and interact with DNA to regulate further genetic transcription into RNA. They belong to the class of “developmental genes,” influential in the generation of embryonic pattern, and whose lack of function can lead to profound alterations in the development of specific brain regions. Secreted morphogens and structural products, such as cell-adhesion proteins, are mentioned occasionally. We do not address “housekeeping” genes involved in metabolism, or genes that are generally related to neuronal and glial differentiation, as they tend to be broadly expressed and therefore provide little or no regional morphological information.

The developing neural tube shows over time an increasing number of regional subdivisions. These are known to us primarily as morphological entities (i.e., vesicles, neuromeres, lobes, gyri, eminences, recesses), but also can be characterized by their patterns of gene expression. We allude to the latter properties by the term “molecular brain subdivision,” referring the reader to a rapidly growing new field of neuromorphology. In recent years many observations have accumulated showing that such combined molecular topographies frequently show reproducible boundaries and are topologically invariant during ontogenesis (though some genes do show changing expression patterns over time), and many early patterns are remarkably resistant to evolutionary change. The largescale comparability of mouse brain gene patterns with counterparts in human, avian, amphibian, teleost, and agnathan species is providing a substantial new impulse to comparative neuroanatomy (i.e., SmithFernandez et al., 1998; Puelles et al., 2000; Bachy et al., 2001; Murakami et al., 2001; Hauptmann et al., 2002). This also accounts for the singularity of this mousebased chapter in a rat brain book. Space and time limitations have led us to concentrate upon the fore-

The Rat Nervous System, Third Edition

MOLECULAR VERSUS ANATOMICAL DISTINCTION OF BRAIN SUBDIVISIONS: THE SPECIFICATION STATE Molecular brain subdivisions start to appear at neural plate stages, earlier than morphological parts

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LUIS PUELLES ET AL.

can be distinguished (Fig. 1) (Shimamura et al., 1995; Rubenstein et al., 1998). Various transcription factors and morphogen-coding genes are expressed either homogeneously or gradientally within distinct anteroposterior (AP) and dorsoventral (DV) neuroectoderm regions, usually symmetrically relative to the midline. At each stage of development, the genes that are active in a locus (known genes plus unknown ones) are jointly held to represent the molecular specification state of the local tissue. The specification state at any stage may be more or less reversible, being sensitive to various microenvironmental aspects, particularly at early stages (activities of surrounding cells; composition of the intercellular matrix and fluid). Each state evolves with time as a result of ongoing intrinsic genetic regulations (genes up- or downregulated) and of epigenetic shortor long-range intercellular signaling effects; these phenomena are all encompassed in the concept of “patterning.” So-called planar neural patterning occurs by signal communication spreading within the ectoderm itself (the neuroepithelium and adjacent nonneural ectoderm, normally within a limited spatial range), whereas vertical patterning is imposed by signals emitted by tissues adjacent to the neural primordium, i.e., derivatives of mesoderm or endoderm. Fate-map analysis suggests that early differential specification of various regions and subregions in the neural plate and neural tube often correlates with prospective fates, or at least with specified levels of developmental “competence.” A state of competence implies a partial advance into a developmental route that still allows multiple outcomes (thus implying a partially unstable specification state), but some alternative developmental routes are now largely excluded. Prospective fate, referred usually to a grossly defined later morphological entity, like the “eye field” or the “telencephalon” (see Fig. 2), simply suggests that under normal circumstances the early region is on its way to produce the corresponding later brain part, without implications as regards competence or determination (the same cells might be able to do something completely different if grafted elsewhere). Fate maps (i.e., Inoue et al., 2000; Cobos et al., 2001; Fernández-Garre et al., 2002) thus apparently locate topologically invariant causal environments where capable cells usually achieve a particular developmental result, whereas specification maps identify the cells actually progressing along a definite developmental route, as they change their competence. Prospective fate can be determined before much molecular specification has taken place (i.e., at blastula stages). Therefore, correspondence of a prospectivefate region with a particular gene expression, or a

molecular constellation, is merely indicative of what may be the first steps in the specification of the correlative later brain part. As the neural primordium progresses through successive stages, following ever more diverse specification routes (reminiscent of the epigenetic landscape of Waddington, 1957), new differentially specified regional subdivisions are introduced. In contrast to the very dynamic changes at the earlier stages, more stable specification states tend to characterize the “definitive” determined fate of each particular subregion. This genetic identity of each locus, articulated with local epigenetic constraints, influences all aspects of histogenesis (proliferation properties, types of neurons and glia cells differentiated, cell adhesivity, and intercellular matrix composition—with consequences in cell migration and axonal navigation—and even differential synaptogenetic and trophic interaction capacities). A fully determined state cannot be proven in principle, since only limited testing conditions can be employed to show that the tissue cannot change its fate. The primary patterning mechanisms, by causing diverse genetic programs to be followed at different loci of the neural wall, lead to secondary histogenetic processes (proliferation, migration, differentiation, establishment of connections) characteristic for each genetic program and locus. Neurohistogenesis produces the characteristic neuronal and glial populations of each brain part and thereby mediates in a complex way the differential growth in surface and thickness of the neural wall. As a consequence of these varied developmental processes, where literally thousands of molecules and millions of cells participate, the anatomical landmarks that were studied in classical neuroembryology and neuroanatomy appear and are further transformed into adult shape. Brain morphogenesis is accordingly a tertiary process, consequent to fine-grain histogenetic and molecular phenomena in the brain wall. Brain wall shape is not directly related to (is not used for) functional fitness of the animal, whereas the local microscopic cellular structure—brain texture—clearly is strongly relevant for fitness. Thus, during evolution, patterning, histogenesis, and functional capacities are the essential aspects of brain development that are fixed in the genes by natural selection, whereas morphogenesis is an epiphenomenon. In some cases it is possible to directly correlate given anatomical landmarks with underlying details of molecular specification (Puelles and Rubenstein, 1993; Puelles, 1995, 2001a). However, there are also cases where anatomical landmarks widely taken as boundaries do not correlate precisely with molecular bound-

I. DEVELOPMENT

1. GENE MAPS AND RELATED HISTOGENETIC DOMAINS IN THE FOREBRAIN AND MIDBRAIN

aries. This is often the case with brain ventricular sulci, i.e., the sulcus limitans of His, or Herrick’s thalamic sulci (Puelles and Rubenstein, 1993). Another clear example is the “pons,” whose apparent upper and lower “limits,” at least as described in classical anatomy, lack a precise correlation with rhombomeric differential molecular identities (Rubenstein and Puelles, 1994; Marin and Puelles, 1995) (see Fig. 2). It is remarkable that pontine nuclei originate widely in the medullary rhombic lip and reach via rostralward tangential migration the pontine region in rhombomeres 2–3, whereas the cerebellum largely forms in the isthmus and rhombomere 1. This knowledge clearly disrupts the traditional concept of the metencephalon or pontocerebellum as a fundamental morphological unit of the hindbrain. Moreover, molecular boundaries sometimes cross a seemingly undivided anatomical region (i.e., no corresponding anatomical boundary was described there before) (Rubenstein and Puelles, 1994). In these comparisons, it seems reasonable to think that the molecular boundaries, which likely represent fate-determining primary causal mechanisms, weigh more as regards defining significant subdivisions than the tertiary anatomical landmarks. After all, differentially specified adjacent brain regions with independent histogenesis do not need to develop morphologically visible boundaries (i.e., the case of individual cortical areas or pallial and subpallial parts of the septum). Conversely, morphologic bulges or sulci tend to be rather variable ontoand phylogenetically, since they are not themselves fixed by genetic information and may depend heavily on epigenesis. Such landmarks often are subjects of vague and preconceived definition and thus can be traced somewhat tendentiously (it is surprising to see the variety of ventricular relief that has been taken as a “sulcus” in the literature; Kuhlenbeck once mentioned a “ridge-like” sulcus). When ventricular or surface sulci are mapped accurately, they tend to show a changing relationship relative to the topologically static molecular boundaries. Exceptions to this rule result when there is direct causal interaction of landmark-forming structures with the molecular boundaries, as occurs in the growth of some axonal tracts parallel to a molecular boundary. The axonal growth cones often are influenced by the underlying molecular discontinuity (i.e., growth-permissive versus nonpermissive substrates, as apparently occurs with the posterior commissure and the retroflex tract; see also Marin et al., 2002). Sometimes the boundary itself, by virtue of specific differentiation of its cells, acquires a mature ridge-like or sulcus-like appearance (i.e., the zona limitans intrathalamica or the hindbrain median raphe).

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SHARING OF MOLECULARLY DISTINCT BRAIN DOMAINS AMONG VERTEBRATES Accumulating comparative results over the last decade strongly indicate that in large measure there is a common pattern of differentially specified neural regions among all vertebrates (Smith-Fernandez et al., 1998; Puelles et al., 2000; Hauptmann and Gerster, 2000; Hauptmann et al., 2002; Bachy et al., 2001; Murakami et al., 2001). This shared pattern, where the relative topology of neighboring differentially specified radial histogenetic fields remains constant despite substantial quantitative differences in field size or in growth, cell migration, and differentiation properties, may be conceived as representing a topological neural tube molecular Bauplan common to all vertebrates. Its existence implies a developmentally constrained process of primary anatomical regionalization of the brain that underlies the overt morphological Bauplan, irrespective of its various modulations in different vertebrate lineages. It is possible to understand how this occurs. In different vertebrates, embryonic neuroepithelial domains with comparable topological positions express spatially and temporally characteristic sets of genes, thus acquiring sequentially the same (causally comparable) specification states. As a result, the neuroepithelial precursors and the neuronal or glial derivatives of these domains become similarly patterned as regards their fundamental proliferative and differentiative properties. A given shared gene combination thus instructs the relevant set of neuroepithelial cells to form a distinct histogenetic subdivision of the neural wall (a morphogenetic field) which is “the same” from diverse points of view: relative position (topology), significant causal mechanisms, overall field fate, and field competence. This situation is known as a field homology (see Puelles and Medina, 2002). Such widely shared molecular regionalization entails first large primary expression domains at neural plate and early neural tube stages, which secondarily become subdivided by intersection with other expression domains, as new genes are upregulated or downregulated selectively via cell-to-cell interactions along the anteroposterior (AP) and dorsoventral (DV) dimensions of the neural primordium or via cellautonomous genetic regulation of transcription. This accounts for implementation of a common Bauplan and for the existence of structural and functional homologies across vertebrates and particularly for the existence of developmental and anatomic similarity across those species that are closely related evolutionarily (i.e., among rodents; see Fig. 3 for the brain territory covered in this chapter).

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DIFFERENTIAL ASPECTS OF HISTOGENESIS The subsequent histogenetic development of each distinct field in the Bauplan under species-specific parameter and variable values—i.e., variant patterns of proliferation, genesis, differentiation, migration, neurite growth, synaptogenesis, and cell death of daughter cells—has important morphogenetic consequences which frequently are species-specific, both intrinsically and considered in the wider context of the other fields in the immediate neighborhood (Puelles and Medina, 2002). The more expansive fields of the neural wall will tend to compress or deform morphogenetically the less expansive adjacent ones, with various shaping results, depending on quantitative differences in the mechanical forces generated and the particular equilibrium states reached typically in each species (i.e., tangential expansion or compression, increase or decrease in thickness, protrusion at the surface or at the ventricle, bending of the radial dimension). This probably accounts for most species-specific aspects of brain structure and form. Normally the histogenesis and morphogenesis of each histogenetic field leads, via the predominant radial migration of derivatives into the local mantle layer, to a characteristic, more or less deformed radial block of the mature brain wall. Each “bidimensional” field of the undifferentiated neural tube wall (if we abstract the thickness of the neuroepithelium) transforms into a tridimensional block, or radial domain, of the mature brain wall (concept comparable to the migration areas of Berquist and Källén, 1954). The radial domains logically extend from the ventricular lining to the pial surface (along lines conceptually radiating from the central axis in the cavity of the neural tube). Frequently, we can distinguish superficial (subpial), intermediate, and periventricular strata of the derived mantle structure, though in some cases most neurons accumulate in only one or two of these positions, allowing space in the rest for fasciculated fiber tracts (i.e., deeply in the cortex or superficially in the spinal cord). Radial layering of neurons relative to their birthdates varies in different radial domains (inside-out, outside-in, or mixed patterns). There is also the heterochronic property of some differentially specified histogenetic fields that nevertheless share some properties. In such cases neighboring radial domains may implement similar programs with different relative timing, i.e., heterochrony of even and odd rhombomeres collaborating in plurisegmental longitudinal structures such as the sensory or motor columns of the cranial nerves (even segments are precocious relative to the adjacent odd rhombomeres). In contrast, different neocortical

areas tend to form continuous gradients of maturation across their boundaries (boundaries clearly may have various sorts of molecular and histological characteristics). Gradiental distribution of differentiation timetables across the brain surface is nevertheless regionspecific in orientation and magnitude. The boundaries of such heterochronic phenomena do correlate easily with underlying molecular boundaries (i.e., the isthmomesencephalic boundary separates mirror-image neurogenetic gradients in the AP dimension of the midbrain and the rostral hindbrain, and many segmental boundaries are transverse loci of minimal proliferation). There are special cases, nevertheless, where derivatives from a given histogenetic field migrate tangentially to a different location in the brain wall, colonizing, so to speak, one or several ectopic histogenetic domains, and thus also depleting variably the population of their original radial domain (such tangentially migrating neurons often keep at least partially their original molecular identity). One recently investigated example is the migration of populations of inhibitory interneurons from the telencephalic ganglionic eminences into the overlying pallium (Anderson et al., 1997, 2001; Marin and Rubenstein, 2001). Extreme cases are known where all the derivatives of one region seem to move away tangentially, as occurs for instance at the rhombic lip in relation to the formation of the inferior olive, the pons, and the external granular layer of the cerebellum. The olfactory bulb is a contrary extreme case in that most of its cells—except the mitral and tufted projection neurons—come from outside its histogenetic field proper (the interneurons come from the ganglionic eminences via the rostral migration stream). A special case of differences obtained among vertebrates, notably among mammals, relates to the number of areal fields one can distinguish within the cerebral cortex. Comparative data suggest that the areas increase in number in proportion to the overall size of the cortex. This apparently bespeaks of developmental constraints limiting the maximal size of a cortical area. It is unclear at the moment whether such constraints relate to regulation of proliferation and fate determination (patterning) or to histogenesis, or both. Protracted patterning might cause more regionalization when the morphogenetic fields overstep a given size limit—note that the intercellular signaling mechanisms involved in patterning have limited effective ranges and probably are sensitive to primordium size. As regards the histogenetic mechanisms, at the moment neither glia-guided radial cell migration or layer-specific neuronal differentiation seem candidates for such effects, since it is unclear how they would be affected by overall size. Possibly there may be size-related limits to the establishment of

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THE BAUPLAN OF THE BRAIN

ordered sets of projections—note that cortical areas thought to have subdivided evolutionarily typically show mirror-image topographic connection patterns relative to their mutual boundaries (Krubitzer, 2000). This interesting arealization phenomenon is a mechanism that converts quantitative changes in cell population into qualitative changes in emergent functional properties. It may operate not only in cortical areas but also in complex nuclear regions (i.e., nuclear subdivisions in the thalamus, the amygdala, or the pons). Understanding of the peculiar histogenetic and morphogenetic properties of the diverse anatomical regions of the brain is thus promoted by correlating the local histogenetic processes both with the underlying molecular specification history and the resulting formation of anatomical landmarks and functionally distinct structures. Comparative analysis of these issues across a range of animals wider than the usual rat– monkey–human spectrum frequently results revealing (Puelles, 2001a).

RP

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There are clear reasons for conceiving of the brain as an elongated tubular formation with DV and AP subdivisions. The precursor of the neural tube is the neural plate (here the prospective DV dimension appears flattened into symmetric mediolateral dimension; Fig. 1). The neural plate starts as a radially symmetrical area around the primary organizer (the planar radial dimension corresponds to the future DV axis of the neural tube) and then elongates dramatically, leading to secondary appearance of the AP axis (see review of clonal data in Rubenstein et al., 1998). Subsequent neurulation rolls up the edges of the neural plate and closes the tube, building the roof plate (Fig. 1). The most anterior end of the roof plate overlies the anterior commissure in the mature brain (Rubenstein et al., 1998; Inoue et al., 2000; Cobos et al., 2001) (see Fig. 2), whereas the anterior end of the floor plate corresponds to the locus of the neurohypophysis

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eye AP BP FP

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FIGURE 1 Graphic schematic representation of the neurulation process viewed from the rostrolateral end of the brain. Nonneural structures of the head primordium are not shown. As the borders of the neural plate approach each other and fuse at the dorsal midline, the eye vesicles evaginate and the forebrain and midbrain (delimited by a transverse dash-line) gradually acquire their characteristic shape. The schema ends when the telencephalic vesicles (Tel) start to evaginate and the rostral neuropore closes. The fundamental four-tiered longitudinal structure of the CNS has its simplest form at initial neural plate stages. The drawings display the topology of the prospective floor plate (FP), basal plate (BP), alar plate (AP), and roof plate (RP) longitudinal domains, which are continuous from left to right across the midline at the front of the neural plate. The initial eye field, which is conceived to lie within the alar plate, also bridges the rostral midline. The anterior neural ridge (ANR) is postulated as a secondary organizer for the forebrain. We base the represented size of this hypotetic organizer area on the early expression of Hesx1 and Six3 genes. As neurulation proceeds, the mutual topologic relationships of these longitudinal domains are not altered. The rostral end of the roof plate will form choroidal tissue at the roof of the third ventricle and at the caudomedial wall of the lateral ventricles (Tel). The ANR persists as the telencephalic commissural plate (median septum), continuing into the lamina terminalis and the prospective optic chiasm. The rostral-most floor plate area is the site where the neurohypophysis will develop, surrounded by the median eminence (compare with Fig. 2).

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TEL lt

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iorg

IV r1 r2 r3 r4 r5 r6 r7

Ped.pon. Pons Pontobulb. Medulla Raphe

Isthmus Cb Rhomb

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FIGURE 2 Fate map of the mouse neural plate (right side; based on data from Inoue et al., 2000, and extrapolating some chicken data from Fernández-Garre et al., 2002; Cobos-Sillero et al., 2001; and Marín and Puelles, 1995).. The fate map appears divided in longitudinal domains, corresponding to the early floor, basal, alar, and roof plates (choroidal roof plate in dark gray; see Fig. 1), as well as in transversal domains (rhombomeres, isthmus, midbrain, prosomeres, secondary prosencephalon). The floor region is divided into a prechordal part (rostral and caudal hypothalamus, RH, CH, plus Mam), and an epichordal part (note distinct AP parts for diencephalon proper and midbrain, and for hindbrain). These epichordal floor subdivisions correspond to two plurisegmental domains: the ventral tegmental area (VTA, underlying the nigral tegmentum in the mes–diencephalic basal plate) and the hindbrain raphe domain. These two domains are differentially characterized by the formation of dopaminergic and serotoninergic neurons, respectively. The midbrain and isthmic basal plate areas contain the oculomotor and trochlear nuclei (III, IV). The main prospective areas derived from the alar plate are indicated. The alar hypothalamus (AH) surrounds the eye domain and includes caudally the peduncular area (which is also known as the supraoptoparaventricular area, because it contains the paraventricular and supraoptic nuclei; the name used here refers to its nature as bed of the telencephalic peduncle; Marín et al., 2002). Rostral to the midbrain, a series of dorsal specialized territories like the subcommissural organ (Sco; under the posterior commissure) and the epithalamus and pineal gland (Eth, Pin) are continuous rostrally with the eminentia thalami (Emt) and the anterior entopeduncular area (AEP); the last two jointly form the so-called hemispheric stalk domain (telencephalic border). The Emt borders upon the pallial telencephalon (which includes the olfactory bulb, OB), whereas AEP borders upon the subpallial telencephalon. The neural plate rim rostral to the forebrain choroidal roof contains the prospective septal domain. The topologic arrangement of pallium and subpallium in the neural plate shows that the pallium is primarily caudal to the subpallium. Abbreviations used: Acust, cochlear column; AEP, anterior entopeduncular area; Cb, cerebellum; CH, caudal hypothalamus; DT, dorsal thalamus; Emt, eminentia thalami; Eth, epithalamus; III, oculomotor nucleus; iorg, isthmic organizer; IV, trochlear nucleus; lt, lamina terminalis; Mammammillary area; Mes, mesencephalon; NH, neurohypophysis; Nigral Tegm, nigral tegmentum; OB, olfactory bulb; och, optic chiasma; p1–p3, prosomeres 1–3; Ped. area, peduncular (supraoptoparaventricular) area; Ped. pon., pedunculopontine region; Pin, pineal gland (epiphysis); Pontobulb, pontobulbar region; r1–r6, rhombomeres; r7, pseudorhombomere 7; rch, retrochiasmatic area; RH, rostral hypothalamus; Rhomb, rhombencephalon; RM, retromammillary area; Sco, subcommissural organ; TEL, telencephalon; Trig, trigeminal column; Vest, vestibular column; VT, ventral thalamus; VTA, ventral tegmental area; ZLI, zona limitans intrathalamica.

(Rubenstein et al., 1998) (see Fig. 2). The neural tube wall placed between the floor and the roof plates is divided into basal and alar primary longitudinal zones, distinguished initially by a characteristic neurogenetic heterochrony already recognized by His (1904), such that the basal plate differentiates precociously (Fig. 1; see also Puelles et al., 1987). Molecular specification data

and experimental causal analysis have consistently corroborated this basic DV distinction (an equilibrium state between opposed “ventralized” and “dorsalized” patterning effects), incidentally showing the inconstant sulcus limitans of His to be, even in favorable cases, just a mere approximation of the molecular boundary (see below). However, we still lack a precise and

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complete molecular definition of the alar–basal boundary at different stages, or of the secondary subdivisions formed within the primary longitudinal zones, in correlation with locally emergent anatomical structure. Nevertheless, this issue recently has seen considerable progress in the spinal cord (see review in Muhr et al., 2001) and is conventionally assisted by some degree of differential histological appearance (Fig. 3; see also Shimamura et al., 1995; Puelles, 1995). At the rostral end of the neural tube, the bilateral basal and alar plates converge respectively together at the midline, in the space existing between the neurohypophysis and the anterior commissure. The alar–basal boundary crosses the rostral midline between the optic chiasm (an alar structure, like the evaginated eyes) and the anterobasal (retrochiasmatic) nucleus (Figs. 1 and 2) (Shimamura et al., 1995, Puelles, 1995, 2001a; Marcus et al., 1999). The anlage of the anterobasal (retrochiasmatic) nucleus builds with the hypothalamic tuberal area the rostral end of the basal plate, whereas the suprachiasmatic, supraoptic, and preoptic areas represent the rostral end of the alar plate and converge rostromedially upon the optic chiasm and the lamina terminalis (Figs. 1 and 2). Further DV patterning within the basal and alar plates shows differential characteristics at different AP levels. In general, each part of the neural tube develops its own pattern of DV subdivisions. These are best understood in the hindbrain, where they form longitudinal sensory and motor columns (and other associated formations) related to cranial nerve nuclei

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(Fig. 2). DV patterning becomes quite complex in the forebrain, where the evaginated eyes and telencephalon, as well as various diencephalic centers, develop (Figs. 2 and 3). AP divisions of the neural tube are bounded early on by molecular limits orthogonal to the longitudinal DV zones; such boundaries already start to become established at neural plate stages (Figs. 4A–4C), and their number subsequently increases by gradual AP subdivision of the early primary regions, partly under the control of so-called “secondary organizers” or via retinoic acid-mediated caudalization and further neighborhood interactions. Differential growth generated by the region-characteristic molecular specification states soon leads to “vesiculation” and “segmentation” of the neural tube primordium. Vesicles are localized outpouchings created by proliferative singularities largely restricted to one of the longitudinal zones; they typically have a boundary (stalk) that is not orthogonal to the brain length axis—consider, i.e., the eye vesicles, the midbrain tectum, the cerebellum, or the telencephalon. Segments (neuromeres), in contrast, are serial transverse outpouchings whose limits are orthogonal to the complete set of DV neural zones; the transverse intersegmental boundaries generally become quiescent proliferatively, though there are exceptions (i.e., at the isthmus). Note that the term “vesicle” in the past was applied indiscriminately to vesicles and segments. Early segments are also known as proneuromeres, following a usage that restricts the term “neuromere” to the definitive set of smaller transverse subdivisions

FIGURE 3 Sagittal section through the brain of an E14.5 mouse embryo, immunoreacted for calbindin to illustrate some of the major anatomic subdivisions in the forebrain. Abbreviations used: cb, cerebellum; cth, caudal (dorsal) thalamus; emt, eminentia thalami; hb, hindbrain; ist, isthmus; mam mammillary area; mes, mesencephalon; mteg, midbrain tegmentum; ob, olfactory bulb; oem, ootoeminential domain; pall, pallium; poa, preoptic area; pt, pretectum; rm, retromammillary area; rth, rostral (ventral) thalamus; se, septum; spv, supraoptoparaventricular (peduncular) area; tm, tuberomammillary area; tu, tuberal area; ZLI, zona limitans intrathalamica.

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FIGURE 4 In situ hibridized mice embryos at late neural plate stages (A–C; whole mounts) and early neural tube stages (D–F; halved neural tubes flat-mounted for organotypic culture) (single- or double-color reactions). The characteristic expression domains of diverse genes are displayed; their names are color-coded depending on the respective arbitrary red versus blue reaction product. Approximate position of the postulated prosomeric domains is indicated in D relative to the Irx3 expression pattern. Note that the eye vesicle lies in a different plane than the rest of the forebrain and is seen superposed.

found in the closed neural tube. In our concept of neuromeres, the serially iterated fundamental morphologic constitution that warrants treating neuromeres as serial homologs or “metameres” is provided by their sharing a complete set of fundamental DV zones and consequently a generally comparable causal history of DV patterning. Note we do not expect that interneuromeric boundary properties, such as clonal restriction, which are themselves secondary to AP patterning, and may vary along the length axis (see Wilkinson, 2001), define the metamery of the neural segments, as is currently postulated by other authors. Our conception has the advantage that the full extent of the neural tube is “segmented,” providing an all-encompassing morphological framework for causal understanding and anatomical reference. In contrast, definition of neural metamery on the basis of boundary properties—i.e., clonal restriction—actually portrays neural segmentation as a transient oddity of a minor part of the neural tube (Larsen et al., 2001). A related concept is that of “tagma,” which refers to a series of adjacent segments sharing special regional characteristics. The caudal epichordal diencephalic segments (prosomeres 1 and 2) possibly build, jointly with the midbrain, a caudal forebrain tagma (i.e., as suggested by common early expression of the Irx3 gene; Bosse et al., 1997; see our Fig. 4D). The midbrain by itself seems segmentally undivided, in contradiction to various classical or recent accounts describing two mesomeres, though it certainly differentiates differentially along

the AP axis; some earlier authors were unaware of the differential growth process caused at the isthmomesencephalic junction by the isthmic organizer (reviews in Puelles et al., 1996; Martínez, 2001; Figs. 4A–4E). Some aspects of midbrain molecular specification (Figs. 3–6), structure and connectivity (i.e., development of dopaminergic cell populations), strongly support its inclusion in the caudal forebrain tagma, though in other aspects the midbrain resembles the hindbrain (i.e., having a motor nucleus). Prosomere 3 and the secondary prosencephalon (SP; we have come recently to consider this a rostral unsegmented proneuromere; Puelles and Rubenstein, 2003) can be grouped in a rostral forebrain tagma encompassing all forebrain elements rostral to the zona limitans intrathalamica (i.e., the domain unified by alar expression of Dlx and Arx genes and basal expression of Nkx2.1; Bulfone et al., 1993; Kitamura et al., 1997). These rostral and caudal forebrain tagmata might as well be joined into a forebrain protagma—SP, p1–p3, plus midbrain— defined by the early overall domain of Otx2 expression (Simeone et al., 1992b). The hindbrain tagma is represented by a set of 12 overt or hidden transverse units (rhombomere 0—the isthmus, classic rhombomeres 1–7, and pseudorhombomeres 8–11; see Cambronero and Puelles, 2000, on the concept of pseudorhombomeres). The spinal cord may be considered a spinal tagma as well, encompassing the full set of cervical, thoracic, lumbar, and sacral myelomeres (spinal neuromeres or pseudoneuromeres).

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FIGURE 5 In situ hibridized mice embryos at late neural tube stages (whole mounts; eye vesicles cut out, leaving eye stalk openings). The characteristic expression domains of diverse genes are displayed; their names are color-coded depending on the respective arbitrary red (in red) versus blue reaction product (in white). Description in the text. Note the outgrowth of the telencephalic vesicles inside the alar plate. Compare in D, E, and F the increasing size of the ZLI core, labeled with Shh.

The fundamental longitudinal, transverse, and vesicular subregions of the neural tube may become variously deformed or hidden during subsequent morphogenesis, but there clearly remain significant molecular traces of them, manifested in differential cytoarchitectony, chemoarchitectony, and adult gene expression. As mentioned above, the ventricular sulci abundantly used in earlier periods of neuroanatomy are tertiary results of brain development and therefore represent imprecise approximations to the relevant causal molecular boundaries. Patterns of neural differentiation, including cyto-, myelo-, and chemoarchitecture, result directly from the diversified histogenetic patterns in the neural wall, secondary to the early molecular specification states. They therefore represent the most reliable guides to the functionally relevant divisions in the mature brain wall.

THE NEURAL PLATE SUBDIVISIONS Molecular regions traceable already at neural plate stages underline the concept sketched above of DV longitudinal zones curving rostrally around a fixed floor plate endpoint, as opposed to wedge-shaped AP transverse regions (Figs. 1, 2, and 4). A good number of mouse genes are expressed early on along restricted DV longitudinal zones, either across the whole or a

large AP extent of the neural plate (i.e., SCF, Matsui et al., 1990; HNF3␣/␤, Sasaki and Hogan, 1993; BF-1, Hatini et al., 1994; Nkx2.2, Shimamura et al., 1995; PLZF, Avantaggiato et al., 1995; Grg4/3, Koop et al., 1996; Zic1, Nagai et al., 1997; ENC-1, Hernandez et al., 1997; Nkx2.9, Pabst et al., 1998; sFRP1, Leimeister et al., 1998). Other genes also are expressed longitudinally, but in shorter AP portions of the neural plate (Evx1, Bastian and Gruss, 1990; Wnt7b/3a/1, Parr et al., 1993; Nkx2.1, Shimamura et al., 1995; Ebk, Ellis et al., 1995; Sim2, Fan et al., 1996; Otlx2, Mucchieli et al., 1996; Fgf3, Mahmood et al., 1996; AP2/AP2.2, Chazaud et al., 1996; Nkx2.1, Pax6, BF-1, Fgf8, BMP7, Shimamura and Rubenstein, 1997; BMPs, Furuta et al., 1997; TCF4, Cho and Dressler, 1998; Dac, Caubit et al., 1999; Six6, Jean et al., 1999; Emx2, Suda et al., 2001). In contrast, various genes appear expressed in wedge-shaped transverse domains of the neural plate, encompassing all DV zones at different AP locations (i.e., Boc, Mulieri et al., 2002; Irx3, Bosse et al., 1997; Six3, Oliver et al., 1995; Six6, Toy and Sundin, 1999; Jean et al., 1999; Lhx5, Sheng et al., 1996; BF-1, Lai et al., 1990, 1991; Shimamura and Rubenstein, 1997; Krox20, Lobe, 1997; Hes1/3, Lobe 1997; En1, Lobe, 1997; Mf3, Labosky et al., 1997; Sax1, Schubert et al., 1995; Gsh1, Valerius et al., 1995; Gbx2, Wassarman et al., 1997; Hesx1, Dattani et al., 1998; Martinez-Barbera et al., 2000; Martinez-Barbera and Beddington, 2001; Ras/Rx,

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FIGURE 6 Microphotographic and schematic illustration of gene expression patterns characteristic of either the alar or the basal plate domains of the neural tube. (A,B) In situ hibridized mice embryos at E11.5 (A; single ISH) and E12.5 (B; double ISH) (whole mounts). The characteristic expression domains of three genes are displayed; their names are color-coded depending on the respective arbitrary red versus blue reaction product. Note in A the rostral thalamic Dlx2 expression stopping at the ZLI core, whereas Nkx2.2 labels the ZLI shell area around the core. Abbreviations are as in Fig. 3. (C,D) Schematic representation of the prosomeric model, showing the main postulated longitudinal and transversal subdivisions, superposed with some characteristic molecular genetic markers. See text for description. The mode in which extratelencephalic boundaries might continue into the telencephalon, with or without connection with the palliosubpallial boundary, remains tentative.

Dattani et al., 1998; Pax2/5/6, Schwarz et al., 1999; Otx2, Simeone et al., 1992a; Shimamura et al., 1995; sFRP2, Leimeister et al., 1998; Ptc, Gli, and Shh, Platt et al., 1997). This nonexhaustive list of examples reveals that a remarkable degree of differential molecular specification and regionalization at neural plate stages is already known, displaying patterns consistent with the proposed model of DV and AP axial dimensions and available fate maps (Figs. 1, 2, and 4) (Rubenstein et al., 1998; Inoue et al., 2000; Cobos et al., 2001). It should be noted that there arises a singularity, due to the radial symmetry found at the rostral midline of the neural primordium: the apparent AP dimension along the median neuroepithelium extending from the anterior commissure locus (rostral neuropore or end of

the roof plate) to the locus of the prospective neurohypophysis (rostral end of floor plate) is coded genetically consistently with its content of a full set of DV longitudinal zones (compare with Fig. 1). More laterally, this topological singularity partially affects as well the prospective telencephalon and eye fields (Figs. 1 and 2). Patterning of this median and paramedian rostral domain of the forebrain notably depends on blockage of BMP and Wnt signaling mechanisms, both of which represent “dorsalizing” signals (Wilson and Rubenstein, 2000). Thus, paradoxically, the topographically “anterior” end of the neural plate shows “dorsal” molecular characteristics. This part of the neural tube is also peculiar in being prechordal (overlying the prechordal plate and rostral mesendoderm) while the

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rest of the neuraxis is epichordal (overlies the notochord). This implies vertical inducing effects different from those of the local axial mesoderm and exclusive rostral patterning influences from the anterior visceral endoderm (Rubenstein et al., 1998; Wilson and Rubenstein, 2000). These important differences probably underlie the morphological singularities of the rostral forebrain.

THE CLOSED NEURAL TUBE The incipient bending of the length axis already observed at late neural plate stages increases considerably after neurulation, leading to the well-known cephalic and cervical (ventral) flexures of the neural tube, as well as to the oppositely oriented pontine flexure (Figs. 3–5). Observations of gene marker expression and fate mapping in different vertebrates consistently support the notion that molecularly distinct regions of the early neural tube bend coherently with it, implying that relative specification of DV and AP position along the neural wall is not affected by this macroscopic morphogenetic process. Precise reference to brain AP and DV “relative molecular” positions thus will henceforth need topological terms referred to the bent length axis (Fig. 3) (Puelles and Rubenstein, 1993; Puelles, 1995, 2001b), against current usage (compare, e.g., the BMPs expressed dorsally all along the neural tube roof in Furuta et al., 1997). It is important to realize that during the rebirth of neuroanatomy after the 2nd World War, topographic references for the brain became founded upon the now obsolete convention of a brain length axis lacking a cephalic flexure and ending rostrally inside the telencephalic vesicle—the bent axis was erroneously supposed to be transient during development (see recent use of this convention in Swanson, 1992, his Figs. 3–5). That assumption is clearly irreconcilable with the concept presented here and poses various terminological sources of confusion, due to its widespread usage. The possible confusion resulting from partial or complete adherence to such a brain model affects notably two brain areas: (a) the isthmic and pedunculopontine hindbrain regions frequently are misinterpreted as midbrain derivatives (the midbrain entirely lies rostral to them), and (b) some diencephalic areas are also wrongly attributed to the midbrain (i.e., the pretectum and the prerubral tegmentum extending into retromammillary areas). The classical rendering of the main diencephalon (in Herrick’s model) as consisting of superposed longitudinal areas (i.e., dorsal and ventral thalami, hypothalamus) is exactly 90º wrong when we refer to the observed bent axis of the forebrain (Figs. 3–5), since these diencephalic areas actually are transverse domains orthogonal to the causally

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determinant axial elements in the floor of the neural tube and in the underlying mesoderm (Puelles and Rubenstein, 1993; Rubenstein et al., 1994, 1998; Shimamura et al., 1995; Puelles, 1995, 2001b). Note these two problematic brain regions caudal and rostral to the midbrain precisely encompass the stretch of neural tube bent at the cephalic flexure. A further common cause of morphologic confusion lies in regarding the telencephalon, an entirely dorsal derivative of the rostral forebrain, as an area where palliosubpallial regionalization can be described as “dorsoventral” patterning (see pallium and subpallium in Figs. 2–6), implicitly assuming a phantom length-axis bifurcated inside each of these paired vesicles; see Puelles and Rubenstein, 1993). Fate mapping at neural plate stages has shown conclusively that the subpallium is rostral relative to the pallium (Inoue et al., 2000; Cobos et al., 2001) (see Fig. 2). A number of genes show domains of expression that cover the whole DV extent of the neural tube wall, with transverse boundaries at diverse AP locations. The prosomeric model identifies a number of morphologic boundaries that correlate with these molecular subdivisions (Figs. 4D, 5C–5E, 6C, and 6D) (Puelles and Rubenstein, 1993; Puelles, 1995, 2001a; Rubenstein et al., 1994, 1998; see Echevarria et al., 2001, for neural tube flat-mount culture). This model initially postulated six prosomeres in the forebrain (counted caudorostrally from the mes–diencephalic border in the order of appearance) and one mesomere. In a recent update of the model, we acknowledged that no strong evidence has accumulated for the postulated prosomeres 4–6 within the secondary prosencephalon, and we are now proposing that the secondary prosencephalon as a whole should be regarded as an unsegmented proneuromere, which is further patterned by singular mechanisms into specific telencephalic and hypothalamic subdivisions nonanalogous to either DV or AP divisions found more caudally in the neural tube (Puelles and Rubenstein, 2003). The rest of the forebrain, the caudal diencephalon or diencephalon proper, subdivides into prosomeres 1–3, for which there is strong evidence. Note we regard the prosomeric model as an advanced and empirically potent construct, which nevertheless remains essentially nonfinished. It already has been modified in recent years in light of accruing evidence (tracing of some boundaries or identity of neural derivatives of given areas) and will probably continue to evolve in the quest of greater consistency and usefulness. At closed neural tube stages, Six3 first appears restricted to the entire rostromedian DV domain originally postulated by us as p6 (Oliver et al., 1995) (compare Figs. 4D and 4E); this region later gives rise to the

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rostralmost forebrain, including the telencephalic septum, the optic vesicles, and the hypothalamus (some secondary domains appear elsewhere later). Interestingly, elimination of Six3 activity leads to severe stunting of the forebrain, amounting to complete loss of the entire secondary prosencephalon (Lagutin et al., 2003). Otx2 expression first appears in the whole forebrain tagma, down to the caudal limit of the midbrain (Fig. 4A). This boundary remains constant, while some rostral forebrain areas secondarily downregulate Otx2 (Simeone et al., 1992b). The Irx1/2/3 genes, present in the brainstem and midbrain, jointly show a rostral transverse boundary coinciding with the zona limitans intrathalamica in the diencephalon (ZLI ; p2/p3 limit; Bosse et al., 1997) (Figs. 3, 4D, and 5E). The genes En1 and En2 define overlapping transverse domains across the isthmic boundary, but their signal reaches, at least initially, the rostral midbrain limit. Gbx2 is widely expressed in the spinal cord and hindbrain and stops rostrally at the isthmomesencephalic boundary, abutting the caudal boundary of Otx2 and a transverse ring of Wnt1 expression in the caudal midbrain (Figs. 4A and 4B).

BASAL PLATE REGIONS Gene expression patterns illustrating longitudinal zones in the closed neural tube are well known. For instance, the basal plate domain in the extended forebrain (including midbrain) is defined at the ventricular lining by continued expression of the Shh gene, coding for a secreted protein largely responsible for early ventralizing patterning effects (Figs. 4F, 5A, 5D–5F). Though initially also expressed in the forebrain floor plate, Shh later becomes downregulated in the hypothalamic floor region. This forebrain–midbrain pattern contrasts with the hindbrain and spinal cord expression of Shh, where it remains restricted to the floor plate. Other genes contribute to the regional specification of the basal plate, with various AP restrictions. In the secondary prosencephalon, the basal plate zone of the hypothalamus characteristically expresses the transcription factors Nkx2.1 and Nkx5.1/Nkx5.2, overlapping the local Shh signal (Figs. 5B and 6B) (Shimamura et al., 1995; Rinkwitz-Brandt et al., 1995). Fkh-4 and Fkh-5/Mf3 signals are restricted to the mammillary region (Kaestner et al., 1996; Wehr et al., 1997). Hypocretin (orexin) also seems restricted to the posterior hypothalamic basal plate, above and behind the mammillary complex (Peyron et al., 1996). Dbx seems to be expressed selectively in the basal plate of p3 (Fujii et al., 1994). Emx1 characterizes the basal plate of p2 (Simeone et al., 1991; Puelles, unpublished observations), whereas Mf2

appears restricted to the basal domain of p1 and, separately, to the secondary prosencephalon (Sareina et al., 1998). The basal domain of Lim1 initially coincides just with p1 and the midbrain, though it later expands rostrally (Mastick et al., 1997). In the caudal forebrain (p1–p3, mes), as well as in the hindbrain and spinal cord, the medial part of the basal plate (where motoneurons are produced) expresses the Sax1 and Nkx6.1 genes (Schubert et al., 1995; Qiu et al., 1998). The Nkx6.1-positive column of basal neurons unifies the hindbrain, midbrain, and forebrain medial tegmentum up to the retromammillary area, irrespective of the change in expression pattern of Shh and several other genes at the isthmus (Fig. 6D) (Qiu et al., 1998). Rostral to the midbrain oculomotor nucleus, this neuronal band forms a recently characterized compact “periventricular tegmental area” where LHRH neurons are formed (Puelles et al., 2001); classical literature refers somewhat vaguely to this formation as the “nucleus of Darkschewitsch.” Isl-1 appears in all postmitotic motoneurons, as well as in other basal and alar derivatives in the forebrain (Ericson et al., 1995). The lateral part of the forebrain and midbrain basal plate, adjacent to the alar–basal boundary, expresses the genes Nkx2.2, Nkx2.9, and Ptc (Fig. 6B; all three continue in the hindbrain and spinal cord in a thin band, found adjacent to the floor plate expression of Shh; Shimamura et al., 1995; Platt et al., 1997; Pabst et al., 1998). Wnt7a and Wnt5a are expressed along much—if not all—of the epichordal basal plate (Parr et al., 1993). A band of Gli-1 and Ptc expression apparently parallels the alar–basal boundary at the alar side in the midbrain and diencephalon, apparently just outside the Shh expression in the basal plate (Hynes et al., 1997; Platt et al., 1997). It has not been determined yet whether the neighboring band of Nkx2.2 and Nkx2.9 expression, which seems to abut on this boundary from the basal side, overlaps partially with the Gli-1 and Ptx bands. Most genes expressed in the forebrain basal plate— i.e., Shh, Sim1, Sim2, Sax-1, Six3, Otlx2, Brx1, Nkx2.2, Nkx2.9, Ptc, Plp, DM20, Emx1, and PLZF—are deflected transversally into the core (i.e., Shh in Fig. 5F) or shell regions (i.e., Nkx2.2 in Fig. 6B) of the ZLI (this encloses the p2/p3 limit) at the transverse boundary between the classical ventral and dorsal thalamic regions (thalamus and prethalamus in our present terminology; TH and PTh in Figs. 3–6). Comparative data in zebrafish and chick embryos suggest that this paradoxical transverse ZLI spike of the longitudinal basal markers does not exist at early stages, when the longitudinal zones are first established, but is formed secondarily (Barth and Wilson, 1995; Hauptmann and Gerster, 2000; Larsen et al., 2001). Formation of the ZLI spike in zebrafish accompanies dorsal expansion of the rostral-most

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expression of the gene axial, the homolog of the mammalian HNF-3␤ gene, another basal plate marker (Sasaki and Hogan, 1993; Barth and Wilson, 1995). It seems that the transverse dorsalward deflection of longitudinal ventral markers at the ZLI (followed by other genes controlled by them, such as Nkx2.2; Shimamura et al., 1995; see also Platt et al., 1997) results from homeogenetic induction of “basal” genes in primitively alar cells, followed by consequent changes in the neighboring cell populations (S. Martínez, unpublished observations). Curiously enough, some roof plate gene expressions also become deflected ventrally into the ZLI during early development (e.g., Fgf8, Bmp4, and Wnt3a; see Crossley et al., 2001). The expression of diverse genes coding for diffusible morphogens at the ZLI (SHH, WNTs) underlies the current hypothesis that the ZLI may represent a diencephalic (thalamic) organizer. Rostrally to their deflection into the ZLI, most basal plate gene domains in the diencephalon curve into the floor at retromammillary or mammillary levels, after passing longitudinally ventral to the prethalamus (p3). It is not yet clear why this occurs. It is attractive to speculate that this phenomenon relates to the transition between epichordal and prechordal parts of the neural tube and portrays the end of epichordal induction effects. Genes such as HNF3␤ might depend directly or indirectly on signals diffusing exclusively from the notochord (whose rostral tip lies close behind the mammillary pouch) and thus are not present more rostrally, next to the prechordal plate. Several classical neuroanatomists postulated the mammillary or retromammillary regions to be at the rostral end of the basal plate (see reviews in Shimamura et al., 1995, and Puelles, 1995). Floor plate markers and local differentiation patterns comparable to those in the midbrain tegmentum extend for a distance rostral to the midbrain. This is consistent with the idea that the ventral neural tube directly exposed to chordal influence ends at the retromammillary region (the latter actually lies in the secondary prosencephalon, closely behind the mammillary complex; note that literature often refers to this area as the “supramammillary region,” a less apt term that implicitly assumes a non-bent longitudinal axis at the cephalic flexure). Due to the dogmatic perdurance of His’ (1904) tentative definition of the meso–diencephalic border (extending from the posterior commissure to the mammillary pouch), the retromammillary and prerubral tegmentum have traditionally been assigned to the midbrain tegmentum. This has obscured our understanding of this entire brain region, which turns out to contain distinct parts belonging to mes, p1–p3, and the caudal-most part of the secondary prosencephalon (hypothalamus);

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molecular specification patterns described above reveal both shared gene expressions (reflecting the general character of the epichordal forebrain basal plate) and differentially expressed genes, which provide specific identities to the diverse AP sectors. For instance, as a reflection of a shared causal pattern, dopaminergic neuronal populations of the mammalian ventral tegmental area have their origin throughout the paramedian floor continuum extending from the isthmus to the retromammillary region (Verney et al., 2001). The mammillary region itself apparently lies under the crossed inductive influences of the tip of the notochord and the caudal end of the prechordal plate, which possibly leads to its molecular, anatomical, and functional singularities. The rest of the ventral forebrain (tuberomammillary and tuberal hypothalamus) would be selectively influenced by prechordal mesodermal signals. Prechordal Nodal signaling upon the secondary prosencephalon is needed to impede cyclopia and holoprosencephaly; accordingly, a particular mechanism mediates the bilateral formation of the eyes and the telencephalic vesicles. Thus, while Shh and Isl-1 are expressed along the whole forebrain and midbrain basal plate, defining the generic ventralized character of this longitudinal zone, other basal plate genes show either prechordal or epichordal expression domains, apparently divided at the mammilloretromammillary boundary. The literature is extremely confusing about whether or not the rostrally convex curve of the epichordal basal genes encloses the mammillary bodies. Recently we have discovered this ambiguity to be related to the fact that the mammillary primordium is morphologically inconspicuous at early stages, and only becomes clearly identifiable as a distinct bulging pouch approximately after E14–15 in the mouse. At earlier stages there exists in this area of the forebrain floor a similar ventral bulge that actually corresponds to the prospective retromammillary area; this is later overshadowed by the more prominent mammillary protrusion. The precocious retromammillary pouch apparently has been frequently misinterpreted in developmental and gene mapping studies (including our own ones; i.e., Bulfone et al., 1993) as the mammillary pouch. The mouse gene Otlx2, for instance, described as mapping to the mammillary and retromammillary basal areas, clearly ends restrictedly at the retromammillary area (and also labels its laterally displaced derivative, the subthalamic nucleus), while the incipient mammillary pouch in front of it is distinctly devoid of this signal (Mucchielli et al., 1996; their Fig. 5B showing an E13.5 mouse embryo). A similar reinterpretation may be needed for Brx1 (Kitamura et al., 1997) and Ebf1/Ebf2/Ebf3 (Garel et al., 1997), among other genes. On the other hand, the Sim1

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and Sim2 genes, depicted early on as expressed along a basal band that surrounds the mammillary pouch, actually map precisely upon the mammillary primordium in the terminal arc where their respective bands approach the forebrain midline floor (Fan et al., 1996; their Fig. 6). Moreover, the final mammillary curve of the Sim1/2 basal domains coincides with expression of the Otx1, Otp, and PLZF genes (Avantaggiato et al., 1995), as well as with the genes Mf3 (Labosky et al., 1997) and Fkh-4 (Kaestner et al., 1996), and is delineated as well by the caudal boundary of some genes expressed in the tuberal or tuberomammillary hypothalamus, including the Dlx family genes (Avantaggiato et al., 1995; Puelles, unpublished observations) and Arx (Miura et al., 1997). Finally, the similarly arc-shaped posterior boundary of the Nkx2.1 expression domain in the basal hypothalamus, which initially one tends to believe respects the mammillary pouch (see Fig. 5B), actually contours the retromammillary region, as becomes clear later on (see Figs. 6B and 6D), when the mammillary pouch proper and all its derivatives stand out as strongly Nkx2.1-positive. Realization of this cause of confusion has led us in recent publications to, first, “move” the mammillary primordium from p4 to p5 (Puelles, 2001b), and, most recently, to simply consider it a specialized part of the unsegmented hypothalamic basal plate (Puelles and Rubenstein, 2003). The retromammillary tegmental region, jointly with the so-called “posterior hypothalamus,” represents the basal plate and floor domains at the epichordal–prechordal transition (Figs. 2, 3, and 6). There occurs also a more detailed, microzonal DV patterning inside the basal plate itself, driven by signals from the midline cells (Ruiz I Altaba and Jessell, 1993) and leading to a detailed neuroepithelial DV zonal specification where each microzone produces different subtypes of neurons. This process has been best studied in the mouse spinal cord and hindbrain (Osumi et al., 1997; Ericson et al., 1997; Briscoe et al., 1999; Sander et al., 2000; McMahon, 2000; Stone and Rosenthal, 2000).

ALAR PLATE REGIONS There are genes expressed in the closed neural tube whose signal extends selectively throughout the alar plate from end to end. Such is the case of the Zic genes (Zic1-3; Nagai et al., 1997). Other genes, like Pax7 and Pax3 (Stoykova and Gruss, 1994), are found in the alar plate of the spinal cord, the hindbrain, the midbrain, and part of the caudal diencephalon (p1; Fig. 6C). Pax2 and Pax8 also have extensive alar domains of expression, stopping rostrally at the isthmus; Pax5 is expressed only in a smaller domain centered upon the isthmo-

mesencephalic boundary (Stoykova and Gruss, 1994). Interestingly, another Pax gene—Pax6—is expressed initially throughout all alar forebrain domains, stopping caudally at the p1–mes boundary (Figs. 5C and 6C), though it is secondarily partly downregulated both rostral and caudal to the ZLI (incipient at the stage illustrated in Fig. 5C). At later stages this gene becomes largely restricted to the rostral half of the prethalamus in p3, the epithalamus in p2, and the caudal (commissural) pretectum in p1 (Fig. 8D) (Stoykova and Gruss, 1994; Mastick et al., 1997). The definitive Pax6 alar domain also extends through the eminentia thalami into the adjacent pallial part of the telencephalon (Figs. 5C, 6D, and 8B–8H; the eminentia thalami is a topologically dorsal area of p3 (see Figs. 2, 3, and 6), whose ventricular surface bulges at the back of the interventricular foramen, forming sulcus terminalis with the basal ganglia at the bottom of the caudal lateral ventricle—see EMT in Fig. 8; it represents the bed of the stria medullaris tract, rostral to the epithalamus). Wnt7b, similar to Pax6, is expressed in the eminentia thalami and expands rostrally into the telencephalic pallium in p5 (Parr et al., 1993). Other genes showing a connection of eminentia thalami expression with caudal (amygdaloid) telencephalic pallium signal are Dbx (Lu et al., 1994, 1996) and Otp (Wang and Lufkin, 2000). Finally, both the gene Lhx5 and the gene coding for R-cadherin appear expressed in the whole extratelencephalic alar plate rostral to the ZLI (Sheng et al., 1997; Redies and Takeichi, 1996). Other gene patterns in the alar plate are less extensive along the AP axis, but still respect some postulated interneuromeric boundaries. The forkhead gene BF-2 appears in the forebrain alar plate rostral to the ZLI (p2/p3) limit, but is excluded from the telencephalon, where a complementary pattern of BF-1 is found (Tao and Lai, 1992; Hatini et al., 1994; Xuan et al., 1995). The gene Arx (Miura et al., 1997) appears in the entire alar extent of p3 (prethalamus and the overlying eminentia thalami); this pattern displays most clearly the p3–SP boundary across the alar plate (for prethalamic molecular subdivisions, see Kitamura et al., 1997, and Nakagawa and O’Leary, 2001). Arx also shows a longitudinal thin alar band of expression along the alar–basal boundary of the secondary prosencephalon (SP), which ends in the suprachiasmatic area of the hypothalamus. It may be useful to call this the “chiasmatic band” (Figs. 6C and 6D). The Dlx family genes overlap the Arx-positive domain in the prethalamus, excluding the eminentia thalami, and in the chiasmatic band (Fig. 6), but their signal also appear additionally in the tuberomammillary and arquate hypothalamic areas (Bulfone et al., 1993). The Arx- and Dlx-negative alar region immediately dorsal to the

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chiasmatic band is known as the “optoeminential domain” (due to its end at the optic stalk; see oem in Fig. 3). Here typically the genes Brn2, Otp, and Sim1, are expressed, though there are suggestions that they do not completely overlap, possibly defining subdivisions within the optoeminential domain (Fig. 6C) (Simeone et al., 1994; Nakai et al., 1995; Fan et al., 1996; Michaud et al., 1998; Acampora et al., 1999). This area is the locus where the magnocellular and some parvocellular hypophysiotropic neuronal populations of the paraventricular, anterior periventricular, and supraoptic nuclei are formed, jointly with part of the entopeduncular nucleus (another part of the latter arises in the caudal part of the chiasmatic band). The gene Six6 is present early on, jointly with other genes, in the optic field and optic vesicles (Toy and Sundin, 1999; Crossley et al., 2001). Wnt3 defines the entire alar plate of p2 (thalamus plus epithalamus), whereas Gbx2 appears restricted therein to the dorsal thalamus area (Fig. 6C) (Bulfone et al., 1993; see later thalamic subdivisions in Nakagawa and O’Leary, 2001, and Gonzalez et al., 2001). The gene TCF-4 labels jointly the alar plate of p1 (pretectum) and p2 (thalamus and epithalamus; Fig. 5D) (Cho and Dressler, 1998), whereas the entire alar p1 expresses the gene Lim1 (Fujii et al., 1994). The Ebf family genes characterize the rostral (precommissural) pretectum (p1), as well the optoeminential and eminentia thalami domains mentioned above (Garel et al., 1997). On the other hand, Sax1 is expressed in caudal alar p1 (Schubert et al., 1995), a domain similarly marked by the gene AP-2 (Chazaud et al., 1996) and the gene Lmbx1 (Gogoi et al., 2002; Broccoli et al., 2002), all of which also extend into the adjoining midbrain. The rostral pretectum shows signals of the genes Dbx1 and Dbx2 (extending into p2; Shoji et al., 1996), as well as of PLZF (the latter also present in the epithalamus; Avantaggiato et al., 1995). As an example of a more heterogeneous pattern, Gsh-1 is expressed distinctly not only in alar p1, but also in the alar midbrain, the caudal bank of the ZLI in p2, alar p3, alar SP, and the subpallium (Valerius et al., 1995). Some gene expression patterns subdivide the alar midbrain into anterior and posterior parts corresponding to the superior and inferior colliculi. For instance, Hes3 appears in the caudal midbrain (inferior colliculus; Lobe, 1997), whereas Otlx2 appears in the alar rostral midbrain (superior colliculus; Mucchielli et al., 1996).

TELENCEPHALIC PATTERNS The evaginated telencephalic vesicle is divided basically into subpallial and pallial molecular domains

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(Figs. 6A–6D and 9) (Puelles et al., 2000). We presently conceive these two telencephalic primary domains as differential dorsal alar derivatives of the secondary prosencephalon, though they possibly do not represent straightforwardly either DV or AP subdivisions (Puelles and Rubenstein, 2003). It is doubtful that the palliosubpallial boundary, as defined for instance by abutting domains of Dlx and Tbr1 expression in all vertebrates examined so far (Figs. 6A, 6C, 7, 8, and 9), is strictly continuous with an extratelencephalic transversal boundary in the neural tube wall. It forms a curve that encloses the basal ganglia and the subpallial amygdala, a large part of the septum, the anterior entopeduncular area (at the telencephalic stalk), and the preoptic area (Figs. 6C–6D). The prospective pallidal subdomain marked by expression of Nkx2.1 is limited by another curve, enclosed by the larger one (Figs. 5B and 6A–6D); moreover, the smaller anterior entopeduncular area selectively expressing Shh also appears enclosed within the Dlx+Nkx2.1-positive domain, in a Russian-doll-like arrangement. None of these domains reach caudally the rostral border of p3 (represented here by the eminentia thalami, Emt), from which they are separated by intratelencephalic spikes of the gene expression territories of the optoeminential domain, which extend in front of the rostral border of p3 into the caudal telencephalic pole (amygdala; Figs. 6C and 6D). These spikes form part of the stria terminalis band, while other parts of this complex, representing as well the extended amygdala, develop within the striatal and pallidal subdomains (Bst). All this complexity suggests that several special patterning effects are superposed upon any fundamental prosomeric topology of the SP, possibly initiated by the action of a rostral organizer at the anterior neural ridge of the neural plate (ANR in Fig. 1; see Crossley et al., 2001), which acts upon nearby areas of the SP. The optoeminential domain at the transition between the telencephalic subpallium and the hypothalamus seems to be an alar pattern that just escapes the rostral organizer influence and probably relates causally to the eye field patterning mechanisms (Wilson and Rubenstein, 2000). The entire telencephalon falls within the expression domain of BF-1, coinciding already at neural plate stages with fate-map data (TEL; Fig. 2) (Shimamura et al., 1995) and bulging out as distinct paired vesicles shortly after neural tube closure (Tao and Lai, 1992; Hatini et al., 1994; Xuan et al., 1995). Curiously, the BF-1 expression domain extends outside the evaginated vesicles into the preoptic area and nasal portion of the optic vesicles (this, together with the preoptic sharing of some subpallial markers—Fig. 6—gives some molecular support to the old morphological conception of an

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FIGURE 7 Autoradiographic in situ hibridizations of parallel section series through an E13.5 mouse brain, cut in a plane horizontal to the forebrain axis. The upper range of photographs illustrates subpallial and prethalamic expression of Dlx2. The range at the bottom shows pallial expression of Emx1 in adjacent sections. Note the facing boundaries of these two patterns do not touch, leaving a thin negative space, conceived to represent the ventral pallium region (Puelles et al., 2000; see Fig. 9).

FIGURE 8 Autoradiographic in situ hibridizations of parallel section series through an E13.5 mouse brain, cut in a plane horizontal to the forebrain axis (same brain as in Fig. 7). The upper range of photographs (a,c,e,g) illustrates expression of Tbr1 throughout the pallial mantle zone and in derivatives of the eminentia thalami (EMT). The range at the bottom (b,d,f,h) shows in adjacent sections expression of Pax-6 in the pallial ventricular zone, extending into the EMT ventricular zone, and in parts of the prethalamus and commissural pretectum. Note in panels (d) and (f) a stream of Pax-6-positive neurons migrating radially inside the outer striatal subpallium and accumulating in the olfactory tuberculum and anterior amygdala (ms; TO; AA; compare with Fig. 9).

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impar telencephalic portion, associated essentially with the preoptic area. Attribution of the latter to the hypothalamus never was more than a convention, which seems increasingly obsolete; see Keyser, 1972). Shortly after the evagination of the telencephalic vesicle, the molecular palliosubpallial boundary becomes established (it coincides with the highest expression of tenascin-C in the neuroepithelium; Stoykova et al., 1997). The subpallium sensu lato (subpallium proper plus telencephalon impar) starts to express the Dlx genes (Dlx1, Dlx2, Dlx5, Dlx6; Bulfone et al., 1993; Liu et al., 1997) (Fig. 7), as well as Mash1 (Porteus et al., 1994; Tuttle et al., 1999), Isl-1 (Bulchand et al., 2001), netrin (Métin et al, 1997; Tuttle et al., 1999), Olig2 (Takebayashi et al., 2000), and Gsh2 (Yun et al., 2001), among other general subpallial markers. The basal ganglia primordia and centromedial part of the amygdala formed within this field soon bulge into the ventricular cavity. Both the striatal and pallidal domains of the basal ganglia extend rostromedially into the septum (Figs. 3 and 9) (Puelles et al., 2000). First there appears the medial ganglionic eminence (MGE), the pallidal primordium, which expresses, in addition to the Dlx genes, the Nkx2.1 and Lhx6 genes (Grigoriou et al., 1998; Sussel et al., 1999; Lavdas et al., 1999; Tuttle et al., 1999; Figs. 5B and 6B). The Nkx2.1 domain also covers the anterior entopeduncular area and the anterior preoptic area (AEP, POA). Coexpression of other Nkx genes—Nkx5.1 and Nkx5.2—distinguishes selectively the POA (Rinkwitz-Brandt et al., 1995),

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while the AEP selectively combines the expression of Shh in its ventricular zone (Figs. 5A and 5D–5F) (Sussel et al., 1999; Bulchand et al., 2001) and displays also selectively DM20 signal, associated with the local appearance of a source of oligodendrocyte precursors (Timsit et al., 1995). The telencephalic magnocellular cholinergic neurons possibly originate also from the AEP, colonizing thereafter the basal nucleus of Meynert, the diagonal band, and the septum; smaller cholinergic neurons invade the striatum (Marín and Rubenstein, 2001). Note that the boundaries between diverse subpallial regions in the mantle zone are less clear than those in the ventricular zone, because of the complex tangential cell migrations into adjacent histogenetic domains. The septum, on its part, strongly coexpresses steel as a distinctive molecular character (Bulchand et al., 2001). The lateral ganglionic eminence (LGE), representing the striatal primordium, bulges slightly later into the lateral vetricle, lateral and rostral to the MGE; both eminences fuse caudally at the amygdaloid caudal ganglionic eminence (Figs. 6B–6D). Apart from Dlx and other general subpallial marker genes, the striatal domain also displays some selective markers, such as Ebf1, SCIP, RAR␣, CRABP-1, and mCad8 (Garel et al., 1999) or Pax6 and Six3 (Puelles et al., 2000; Yun et al., 2001). Selective expression of striatin was described in the rat striatum (Fig. 6C) (Salin et al., 1998). On the other hand, the pallial domain of the telencephalon is generally delineated by a number of marker genes, showing strong neuroepithelial expression of

FIGURE 9 Comparison of characteristic molecular subdivisions in sauropsidian and mammalian telencephali to illustrate conservation of topology of molecular domains and differential morphogenesis/histogenesis, consistent with the respective postulated “field homologies” for both the pallium and the subpallium. Septum is shown unlabeled. DP, dorsal pallium; LP, lateral pallium; MP, medial pallium; mz, mantle zone; PA, pallidum; ST, striatum; VP, ventral pallium; vz, ventricular zone. Asterisk, radial migration of Pax6 cells; double asterisk, tangential migration of pallial cells. Note that the massive tangential migration of GABAergic interneurons from the subpallium into the pallial mantle is not represented.

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Pax6 (Figs. 5C and 8b–8h), R-cadherin, Ngn1/2 (Stoykova and Gruss, 1994; Stoykova et al., 1997, 2000; Wilson and Rubenstein, 2000; Yun et al., 2001), and many other genes, e.g., AP2.2 (Chazaud et al., 1996) and vzg-1 (Hecht et al., 1996). Curiously, a lower signal of Pax6 also appears in the striatal ventricular zone, accompanied by migration of Pax6-positive neurons in the local mantle, but the pallium itself is completely devoid of Pax6-expressing mantle cells (Figs. 5C, 8d, 8f, 8h, and 9) (Puelles et al., 2000). The pallial mantle shows initially general expression of Tbr1 (Figs. 8a, 8c, 8e, 8g, and 9) (Bulfone et al., 1995). We proposed a subdivision of the pallium into four molecularly distinct parts, named medial, dorsal, lateral, and ventral pallial portions (MP, DP,LP, VP; Fig. 9) (Puelles et al., 2000; see also Stoykova et al., 2000; Yun et al., 2001; Puelles, 2001a,b). The medial pallium largely includes hippocampal and parahippocampal cortex, while the dorsal pallium corresponds to the isocortex. On the other hand, the lateral and ventral pallial portions separately contribute to olfactory piriform cortex and the underlying claustral, endopiriform, and basolateral amygdaloid nuclei (Puelles et al., 2000). The Wnt7a gene characterizes the MP and DP. Emx1 is expressed in MP, DP, and LP, but is excluded from VP (Fig. 7; note VP does show a stream of possibly tangentially migrated superficial cells). Lhx2 is more strongly expressed in MP and DP than in LP and VP, though one of the derivatives of these, the claustrum, selectively expresses Lhx2 (Rétaux et al., 1999;Yun et al., 2001; Bulchand et al., 2001). Claustrum and endopiriform nuclei are also selectively positive for latexin (Arimatsu and Ishida, 1998). A good number of genes have been reported recently that are expressed regionally or gradientally in various ways in the mouse, rat, or human cortex. The reader is referred here to some relevant sources (Bulfone et al., 1995; Suzuki et al., 1997; Inoue et al., 1998; Grove et al., 1998; Rubenstein et al., 1999; Miyashita-Lin et al., 1999; Rétaux et al., 1999; Donoghue and Rakic, 1999; Bertuzzi et al., 1999; Nakagawa et al., 1999; Wilson and Rubenstein, 2000; Yun et al., 2001, 2003).

ABOUT MECHANISMS One may ask what is the rationale for the consistent formation of topologically conserved gene expression domains and resulting distinct brain wall regions. The conservativeness of the invariant Bauplan topology against random mutational effects that hypothetically should promote its variation (and consequent evolution) bespeaks of strong internal constraints, which maintain constant the number and relative spatial arrangement

of the primary histogenetic units, irrespective of their individual variations in secondary histogenetic aspects and tertiary morphogenesis and function. Present-day thoughts about how morphogenies achieve partial morphostasis (lack of change) during millions of years of evolution underline the role of causal nets of changebuffering activities of many transcription factor genes. These are arranged in retroactively and horizontally interconnected causal cascades, which build superposed and multiply redundant layers of regulated regulators within causal “attractor fields,” also known as “morphogenetic fields” (Thomson, 1988; Arthur, 1998; Striedter, 1998; Puelles and Medina, 2002). Such complex multi-stable causal entities represent basins in the organismic epigenetic landscape (Waddington, 1957), which tend to produce particular developmental outcomes despite considerable variations in the individual molecules or in the number and type of cellular elements enacting the “field effect.” Such collections of regulatory genetic loops and the resulting morphogenetic fields are held to be of ancient evolutionary origin, possibly coincident with the establishment of the fundamental animal morphotypes. Subsequent evolution indeed seems to have deepened considerably, rather than diminished, the regulatory capacity of the “attractor” mechanisms (their developmental “buffering” potential, or the depth of the morphogenetic basin). Evolution obviously diversified simultaneously the levels and types of emergent novelty that can be viably made consistent with these attractors. This may explain the coexistence of evolving phenotypes, eventually with emergent morphological and functional novelty, and a conserved Bauplan.

RELEVANT GENETIC MECHANISMS In the DNA sequence, specific “enhancer” or “repressor” motifs within gene promoters regulate the expression of each gene in particular brain (or body) locations and eventually also under particular functional conditions. These regulatory sites are the binding targets of particular transcription factors (frequently with associated regulating proteins), which thus combinatorially direct gene expression. As development proceeds, the region-specific constellations of expressed transcription factors interact in the nucleoplasm with the gene enhancers and suppressors. This process translates the positional aspects of the previous specification state (the spatially nonhomogeneous distribution of transcription factors and other molecules) into the next stage of regionalization, which results from the new combination of genes activated and resulting specification changes. Some genes reactivate themselves,

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or pairs (sets) of genes activate each other mutually, leading to extended temporal maintenance of some fundamental molecular aspects of the specification states. This phenomenon leads to progressive recruitment of a defined regional identity, as more such “permanent” genes are upregulated in the course of development. During evolution, some enhancer or suppressor sequences of one gene may have been duplicated and/or translocated by recombination into other genes. This phenomenon can explain the typical situation where different genes (notably redundant genes of the same family, formed by partial or complete chromosomal duplication) show identical, or very similar, expression domains (i.e., the neurally expressed Dlx1,2,5,6 or Irx1,2,3 genes). The resulting congruent expression control via comparable enhancers contributes to the developmental fixation of Bauplan elements. Viable promoter configurations are highly resistant to change and contribute by potential interaction of multiple and redundant transcription factors to the buffering of aleatory changes introduced by mutation in a single gene of the constellation. Simultaneously, the nature of these mechanisms allows for novel genes to be added to preexisting constellations (a relatively rare event) and, more importantly, allows deviant genes no longer performing an essential role within the constellation to vary further (a more frequent event), exploring novel functional possibilities, so to speak, and potentially leading to phenotypic variation and brain evolutionary change.

Acknowledgments We thank Oxford University Press, Wiley–Liss, Inc., and Christoph Redies for permission to use previously printed microphotographic or graphic materials.

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1. GENE MAPS AND RELATED HISTOGENETIC DOMAINS IN THE FOREBRAIN AND MIDBRAIN

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C H A P T E R

2 Development of the Telencephalon: Neural Stem Cells, Neurogenesis, and Neuronal Migration SHIRLEY A. BAYER and JOSEPH ALTMAN Laboratory of Developmental Neurobiology, Indiana University–Purdue University Indianapolis, Indiana, USA

Neural stem cell research is an intensely studied topic today. Most of the work is either on gene expression studies in vivo or biochemical characterizations in vitro. The research that is presented here is histological and cytological. Spatiotemporal changes in developing brain morphology (normal histology) and cell proliferation dynamics ([3H]thymidine autoradiography) are used to map stem cell heterogeneity in the neuroepithelium of the telencephalon and in the secondary germinal matrices derived from the telencephalic neuroepithelium from embryonic day (E) 11 to E22. The maps shown here often correlate with recently published gene expression maps of the telencephalon (Stoykova et al., 2000; Zappone et al., 2000; Bulchand et al., 2001; Schuurmans and Guillemot, 2002). Migration patterns throughout the telencephalon are briefly reviewed and the development of three migratory streams, the lateral, rostral, and dentate, is documented in greater detail. Because both the rostral and dentate migratory streams contain neural stem cells that persist in adult rodents, a brief review of adult neurogenesis is also included. Finally, we summarize the population dynamics of neural stem cells during normal development. The most important data used to generate the mosaic maps of the neuroepithelium and secondary germinal matrices are the timetables of neurogenesis of telencephalic neurons. These timetables were quantitatively determined in adult rats that were exposed on 2 to 4 consecutive days to [3H]thymidine during development (long-survival autoradiography). Figures 1

The Rat Nervous System, Third Edition

through 6 summarize telencephalic neurogenetic timetables. These data reveal when a group of neural stem cells is actively producing neurons, either in the expanding and receding parts of the primary neuroepithelium or in the expanding and receding secondary germinal matrices derived from the primary neuroepithelium. The actual location of a germinal zone and proliferation dynamics are based on observations in embryonic and fetal rats that survived 2 h after single [3H]thymidine injections (short-survival autoradiography). Sojourn and migration patterns are based on observations in embryonic and fetal rats that survived for successive 24-h intervals after single [3H]thymidine injections (sequential-survival autoradiography). When all of these methods are used together, and the observations are correlated between specimens, the fundamental processes of brain morphogenesis can be understood. For a detailed description of all three methods of [3H]thymidine autoradiography and their uses, see Bayer and Altman (1974, 1987, 1991a). The maps themselves are presented in two sets of sections from normal rat embryos and fetuses that were not exposed to [3H]thymidine. These are plates chosen from the Atlas of Prenatal Rat Brain Development (Altman and Bayer, 1995). The first set (Figs. 7 through 14) shows coronal (frontal) sections through the telencephalon that are slightly anterior to or just grazing the most anterior extent of the preoptic area of the diencephalon. The second set (Figs. 15 through 24) shows sagittal sections through the part of the anterior telencephalon that evaginates into the olfactory bulb.

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Copyright 2004, Elsevier (USA). All rights reserved.

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These two cutting planes contain the neuroepithelia that will generate most telencephalic structures, including the hippocampus, the medial limbic cortex, the neocortex, the lateral limbic cortex, the primary olfactory cortex, the striatum, portions of the basal telencephalon, portions of the globus pallidus, the nucleus accumbens, the septum, portions of the amygdala, and the olfactory bulb.

NEUROGENETIC TIMETABLES IN THE TELENCEPHALON Pallidum The pallidum contains a diffuse collection of large neurons, many of which are cholinergic, scattered throughout the basal telencephalon: the entopeduncular nucleus, the globus pallidus, the substantia innominata, the horizontal limb of the diagonal band of Broca, and the large polymorph neurons in the olfactory tubercle. The magnocellular neurons get their major input from

either the caudate/putamen complex (striatum) or the ventral striatum (olfactory tubercle and the nucleus accumbens), and they project topographically to the cerebral cortex. The degeneration of pallidal axons may be associated with senile dementia of Alzheimer’s type (reviewed in Bayer, 1985b). In rats, the entopeduncular nucleus is embedded in the posteroventral part of the internal capsule and is the homolog of the internal segment of the primate globus pallidus. It contains the oldest neurons in the pallidum (graph 1, Fig. 1) that originate mainly between E12 and E14 in a sandwich gradient where neurons in the core are older than those in either the anterior or the posterior poles (Bayer, 1985b). The globus pallidus is a body of large neurons sandwiched between the lateral border of the internal capsule and the ventromedial border of the striatum. Most neurons originate between E13 and E16 (graph 2, Fig. 1) in a complex three-way neurogenetic gradient: posterior, ventral, and lateral (older) to anterior, dorsal, and medial (younger). The oldest neurons (birthdays on E13) are located in the posteroventral

FIGURE 1 Timetables of neurogenesis for major neuronal populations in dorsal and ventral pallidum (graphs 1–5) and the striatum (graphs 6–9). From Bayer and Altman (1995).

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globus pallidus; the youngest neurons (birthdays on E16) are located anteromedially (Bayer, 1985b). The substantia innominata contains medium- to largesized neurons lying beneath the striatum (see below) and the globus pallidus. These neurons are generated from E13 to E17 (graph 3, Fig. 1) in a two-way posterior and lateral (older) to anterior and medial (younger) neurogenetic gradient (Bayer, 1985b). The magnocellular preoptic nucleus or the nucleus of the horizontal limb of the diagonal band of Broca (MPO-HDB) in rats is identical to the basal nucleus of Meynert in primates (reviewed in Bayer, 1985b). In rats, this nucleus contains large neurons that sweep forward through the anteromedial and anterolateral basal telencephalon to blend in with the large polymorph neurons scattered in layer III of the olfactory tubercle. Anteromedially, the horizontal limb of the diagonal band is continuous with the vertical limb of the diagonal band of Broca. Neurons in the MPO-HDB are generated mainly from E13 to E16 (graph 4, Fig. 1) in a two-way posterior and lateral (older) to anterior and medial (younger) neurogenetic gradient (Bayer, 1985b). The large polymorph neurons in the olfactory tubercle are the most anterior pallidal neurons and resemble those in the globus pallidus, the substantia innominata, and the horizontal limb of the diagonal band. These neurons are generated mainly from E14 to E16 (graph 5, Fig. 1) in a lateral (older) to medial (younger) neurogenetic gradient (Bayer, 1985a).

1981). The fiber bundles of the internal capsule do not traverse the nucleus accumbens; rather, the anterior commissure passes through its medial part. Nucleus accumbens neurons are generated over a protracted period, from E15 through P3 (graph 7, Fig. 1). Neurogenetic gradients within the nucleus are similar to those in the caudoputamen complex; lateral and ventral neurons are older than medial and dorsal neurons. The small neurons in the olfactory tubercle are similar in size and dendritic structure to the medium spiny neurons in the caudoputamen complex and have a similarly long period of neurogenesis, from E14 to E20 (graph 8, Fig. 1). A few (3%) are generated from postnatal day (P) 0 to P3. These neurons settle in a lateral (older) to medial (younger) gradient (Bayer, 1985a), in a similar pattern but at an earlier time than those in the caudoputamen complex (compare graphs 6 and 8 in Fig. 1). The islands of Calleja are dense clusters of small granule cells in the olfactory tubercle and bordering the nucleus accumbens. Neurons are generated in the islands from E16 to E22 (graph 9, Fig. 1) in a two-way neurogenetic gradient (Bayer, 1985a); ventral and lateral neurons are older than dorsal and medial neurons. In the large island on the medial border of the nucleus accumbens, anterior neurons are older than posterior neurons.

Striatum

The anterior amygdaloid area is a diffuse collection of variably sized neurons lying lateral to the preoptic area and deep to the nucleus of the lateral olfactory tract and the anterior cortical nucleus. Anteromedially, there are scattered large cells resembling those in the horizontal limb of the diagonal band. Neurogenesis occurs mainly from E13 to E15 (graph 1, Fig. 2), but a few neurons originate as early as E12, and some are not generated until E21 (Bayer, 1980c). The long time span is due to the overlapping production of several distinct neuronal populations. Medium-sized neurons are generated mainly between E13 and E15, with a peak on E14; large neurons are generated on E14 and E15, with a peak on E14; small neurons are generated from E18 to E21. The nucleus of the lateral olfactory tract is a distinct spherical cluster of densely packed medium-sized neurons in the anteromedial amygdala. Its neurons are generated mainly on E14 and E15 (graph 2, Fig. 2) and settle in a medial (older) to lateral (younger) order. The nucleus of the accessory olfactory tract contains diffusely packed smaller cells posteromedial to the nucleus of the lateral olfactory tract. Its neurons are generated in a biphasic pattern, most on E12 and E13, and a few on E15 (graph 3, Fig. 2).

The medium-spiny neurons in the caudoputamen complex are generated mainly from E16 to E21–E22 (graph 6, Fig. 1). There are several neurogenetic gradients between these neurons (Bayer, 1984). The most prominent gradient is that ventrolateral neurons are older than dorsomedial neurons. But there are divergent neurogenetic gradients between anterior and posterior parts of the striatum. In the anterior part, older neurons are in superficial and posterior positions; younger neurons are deep and anterior. In the posterior part, the reverse is true; older neurons are in deep and anterior positions, and younger neurons are superficial and posterior. There is also a neurogenetic gradient between patch neurons (older) and matrix neurons (younger) throughout the striatum (Bayer, unpublished observations). The nucleus accumbens surrounds the inferior horn of the lateral ventricle and extends forward to the anterior olfactory nucleus; it blends posteriorly with the anterior part of the bed nucleus of the stria terminalis. The nucleus accumbens has been included in the septal region, but the structure of its neurons and its developmental patterns place it within the striatum (Bayer,

Amygdala

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FIGURE 2 Timetables of neurogenesis for major neuronal populations in the amygdala (graphs 1–14). From Bayer and Altman (1995).

The central nucleus lies in the dorsal part of the amygdala, just beneath the striatum and medial to nuclei in the basolateral group. Its neurons are generated from E13 to E18 (graph 4, Fig. 2) and settle in an anteromedial (older) to posterolateral (younger) order (Bayer, 1980c). The amygdalo–hippocampal area in rats is a small region in the posteromedial amygdala where the medial nucleus blends with the ventral hippocampus. Some of the youngest neurons in the amygdala are located here; neurogenesis occurs from E16 through

E19 (graph 5, Fig. 2) and the cells settle in a superficial (older) to deep (younger) gradient (Bayer, 1980c). The intercalated masses are clumps of densely packed small cells interspersed between other nuclei in the amygdala. Anteriorly, they are clustered around the temporal limb of the anterior commissure; posteriorly, they are clustered among the fibers in the core of the amygdala between nuclei in the corticomedial and basolateral groups. These neurons are generated late (E15 to E19; graph 6, Fig. 2) and settle in an anterior (older) to posterior (younger) order (Bayer, 1980c).

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The medial nucleus forms the medial wall of the amygdala from the anterior amygdaloid area to the ventral tip of the hippocampus. These neurons are generated from E13 to E16 (graph 7, Fig. 2) in an anteroventral (older) to posterodorsal (younger) neurogenetic gradient (Bayer, 1980c). The cortical nuclei are subdivided into anterior and posterior parts that have a strong anterior (older) to posterior (younger) neurogenetic gradient between them (graphs 8 and 9, Fig. 2). Both nuclei have superficial (older) to deep (younger) neurogenetic gradients, and the posterior nucleus has a medial (older) to lateral (younger) gradient (Bayer, 1980c). The basomedial nucleus lies in the core of the amygdala throughout much of its rostrocaudal extent, while the basolateral and lateral nuclei form a pyramidlike structure apposed to the white matter that borders the piriform cortex. Neurons in the basomedial and basolateral nuclei are generated from E14 through E17, while the lateral nucleus has younger neurons, with some arising as late as E19 and E20 (graphs 10–12, Fig. 2). All nuclei have anterior (older) to posterior (younger) neurogenetic gradients (Bayer, 1980c). In addition, the basolateral nucleus has a lateral (older) to medial (younger) gradient, while the lateral nucleus has a dorsal (older) to ventral (younger) gradient. The stria terminalis is a major fiber tract that leaves the amygdala and reaches various targets in the basal forebrain. Neurons that are scattered within this fiber tract collectively form the bed nucleus of the stria termi-

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nalis. The anterior part of the nucleus encircles the interbulbar part of the anterior commissure, extending from the posterior nucleus accumbens to the decussation of the anterior commissure. Neurons in the anterior part are generated from E13 to E20 with an overall peak on E15–E16 (graph 13, Fig. 2) and settle in a posterior (older) to anterior (younger) gradient (Bayer, 1979a, 1987). In addition, ventromedial neurons are generated earlier than dorsolateral ones. The preoptic continuation extends ventromedially into the posterior preoptic area. These neurons are generated mainly from E13 through E16, also with an overall peak on E15–E16 (graph 14, Fig. 2) and settle in a ventrolateral (older) to dorsomedial (younger) order (Bayer, 1987).

Septum The septum forms the subcallosal anteromedial wall of the telencephalon and is a prominent structure in the rat basal forebrain. The triangular and medial septal nuclei are in the midline. The anterior medial septal nucleus blends in with the vertical limb of the diagonal band of Broca just lateral to the midline. A large lateral septal nucleus, situated on either side of the midline nuclei, extends throughout the entire anteroposterior extent. The tiny bed nucleus of the anterior commissure resembles cells in the triangular nucleus and forms a compact cluster of small neurons where the columns of the fornix descend behind the decussation of the anterior commissure (Fig. 24).

FIGURE 3 Timetables of neurogenesis for major neuronal populations in the septum (graphs 1–5). From Bayer and Altman (1995).

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Beginning with the midline septal nuclei, neurons in the triangular nucleus (graph 1, Fig. 3) are generated from E13 to E17, with a peak on E15; those in the lateral part are generated slightly earlier than those in the midline (Bayer, 1979a). Neurons in the medial septal nucleus are generated from E13 to E17 (graph 2, Fig. 3) and settle in a posterior (older) to anterior (younger) order. Indeed, the oldest neurons in the septal complex lie in the medial septal nucleus above the decussation of the anterior commissure (Bayer, 1979a). Neurons in the nucleus of the diagonal band of Broca (vertical limb) are also generated from E13–E17 (graph 3, Fig. 3) and settle in a posterior (older) to anterior (younger) order, continuing the gradient in the medial septal nucleus. Neurons in the lateral septal nucleus are generated from E14 to E19 (graph 4, Fig. 3) and settle in a medial (older) to lateral (younger) order. The youngest neurons in the septal complex lie in this nucleus adjacent to the ependymal lining of the lateral ventricle. Neurons in the bed nucleus of the anterior commissure are generated from E14 to E17 and settle mainly on E15 and E16 (graph 5, Fig. 3) following a timetable similar to that of neurons in the triangular septal nucleus (Bayer, 1979a).

Cerebral Cortex The cerebral cortex is the largest structure in the rat telencephalon. It can be subdivided into the neocortex, the limbic cortex, the piriform (primary olfactory) cortex, and the hippocampal region. Neocortex and Limbic Cortex The neocortex has five cell-dense layers (II–VI), large Cajal–Retzius neurons sparsely distributed in layer I, and scattered neurons in the deep white matter (layer VII or the subplate). In rats, the superficial layers (II–IV) are thinner than those in the human neocortex but layers VI and V are thick (Bayer and Altman, 1991a). In the limbic neocortex, five cell-dense layers are also present. Generally, layers V and VI are easily distinguished from each other, but they tend to be thinner than the same layers in the neocortex. The superficial layers (IV–II) are quite reduced and are often grouped together in a single layer. Most neocortical and limbic cortical neurons are generated between E14 and E20 (graphs 1–5, Fig. 4) and settle in strict gradients to form the layers of the mature cortex. That settling can be separated into three major epochs (Bayer and Altman, 1991a). During the first epoch, the sequentially generated Cajal–Retzius neurons in layer I and subplate neurons (layer VII) settle in a superficial (older) to deep (younger) gradient. The peak time of origin of Cajal–Retzius neurons is on E14 (graph 1, Fig. 4), while subplate neurons are generated

on E14 and E15 (graphs 5, Fig. 4). Taken together, these two populations form the top and bottom “crust” of older neurons around a “sandwich” of younger neurons in layers VI–II. Neurons in layers VI–II are generated during the second and third epochs and settle in a deep (older) to superficial (younger) radial gradient, one of the most prominent neurogenetic gradients in the entire brain (reviewed in Bayer and Altman, 1991a). Layers VI–V are sequentially generated from E15 to E17 (second epoch, graphs 3 and 4, Fig. 4); layers IV–II are sequentially generated from E17 to E20 (third epoch graph 2, Fig. 4). Only a few of the most superficial neurons in layer II are generated on E21, the last day of cortical neurogenesis. Besides the radial gradient between layers VI and II, there are two additional neurogenetic gradients within these layers. First, a transverse gradient that runs parallel to the plane of coronal sections: neurons situated ventrolaterally (those closer to the rhinal sulcus) tend to be older than neurons situated dorsomedially (those closer to the cingulate cortex). Second, a longitudinal gradient that runs parallel to sagittal sections near the midline: neurons situated anteriorly (those closer to the frontal pole) tend to be older than neurons situated posteriorly (those closer to the occipital pole). The deep layers (VI–V) have strong transverse and longitudinal gradients, irrespective of any boundaries between cortical areas. However, the primary sensory areas of the rat cortex always contain neurons in layers II and III younger than the neurons in the same layers of the secondary sensory areas, even if the primary area is lateral to a secondary area. For example, the primary somatosensory and the primary visual areas contain superficial neurons younger than those in their medially situated secondary areas. The limbic cortex surrounds the neocortex. Although there is the same stacking of older to younger cells in the radial dimension, the medial limbic cortex (cingulate and retrosplenial areas) reverse the neocortical transverse gradient and the lateral limbic cortex (insular area) has a longitudinal gradient different than the one in the neocortex (Bayer and Altman, 1991a). These findings lend support to the argument that the limbic and neocortical parts of the cerebrum have different phylogenetic roots; the limbic neocortex may be partially linked to neurogenetic gradients in the paleocortex (Bayer and Altman, 1991a). Piriform (Primary Olfactory) Cortex The piriform cortex is located below the rhinal sulcus and forms the ventrolateral portion of the cerebral hemispheres. It extends nearly 7.4 mm in the rostrocaudal direction, from the anterior olfactory nucleus

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FIGURE 4 Timetables of neurogenesis for major neuronal populations in the neocortex and limbic cortex (graphs 1–5) and in the piriform or primary olfactory cortex (graphs 6–9). From Bayer and Altman (1995).

to the lateral entorhinal cortex. Comparative neuroanatomists consider the piriform cortex to be “paleocortex” because, unlike the neocortex, it is represented in the forebrains of fish, amphibians, and reptiles. Another characteristic that distinguishes the piriform cortex from the neocortex is that it receives monosynaptic input from the main olfactory bulb throughout its entire length and breadth (Bayer, 1986b), while all sensory input to the neocortex is through a relay in the thalamus. Most anatomists consider the piriform cortex to have three cell layers in addition to the cell-sparse external plexiform layer (I). Layer II contains densely packed small pyramidal cells, layer III sparsely packed medium-sized pyramidal cells, and layer IV very sparse large-sized pyramidal cells and polymorph cells. Neurogenesis in the piriform cortex proceeds in two gradients: deep neurons are older than superficial neurons, and posterior neurons are older than anterior neurons (graphs 6–9, Fig. 4). Deep neurons in layers III and IV are generated mainly from E13 to E16, 80% of

the posterior deep neurons on or before E15, and over 30% of the anterior deep neurons on or after E16. The superficial neurons are generated mainly from E15 through E18, 75% of the posterior ones on or before E16, and 37% of the anterior ones on or after E17. In the anterior piriform cortex, layer II is quite thick, and the small pyramidal cells are stacked two to three deep rather than in a monolayer. Here, the superficial neurons are generated slightly earlier than the deep ones (see Fig. 6 in Bayer, 1986b). These data indicate that the piriform cortex does not have the same type of radial gradient as the neocortex where younger neurons are always superficial to older deep neurons. The Hippocampal Region The hippocampal region is a prominent component of the rat cerebral cortex, containing five contiguous structures. The entorhinal cortex, the presubiculum, the parasubiculum, and the subiculum take up most of the ventroposterior cortical wall; the hippocampus proper extends forward beneath the corpus callosum and the

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deep white matter of the neocortex and the lateral limbic cortex. The hippocampus proper is one of the best known neuroanatomical structures due to intense study with a variety of descriptive and experimental anatomical methods (reviewed in Bayer, 1980a). Our studies in the rat include quantitative determinations of the timetables of neurogenesis (Altman, 1966; Bayer and Altman, 1974; Bayer, 1980a, 1982; Bayer et al., 1982), cell migration and settling (Bayer, 1980b; Altman and Bayer, 1990a,b,c), and vulnerability to X-irradiation (Bayer and Altman, 1975a,b).

sized pyramidal cells. Layer IV is a cell-sparse zone (lamina dessicans) with a few scattered large pyramidal cells. Layer V–VI contains relatively densely packed medium- and small-sized neurons. The neurogenetic timetables (graphs 1–4, Fig. 5) indicate a modified deep (older) to superficial (younger) neurogenetic gradient. Neurons in layer V–VI are generated mainly on E15, those in layers II and IV mainly on E15 and E16, and the youngest neurons in layer III mainly on E17. All layers have lateral (older) to medial (younger) neurogenetic gradients (Bayer, 1980a).

Entorhinal cortex The entorhinal cortex contains five layers that, with the exception of layer I, are substantially different from those found in the neocortex. Layer II contains the cell bodies of large stellate cells, grouped into islands laterally and separated from layer III by a cell-sparse zone. Layer III contains medium-

Subicular region The structures between the medial edge of the entorhinal cortex and the hippocampus are the parasubiculum, the presubiculum, and the subiculum proper. The parasubiculum and the presubiculum form a wedge in the posteromedial angle of the cortical wall; these parts of the subiculum are best seen

FIGURE 5 Timetables of neurogenesis for major neuronal populations in the entorhinal cortex (graphs 1–4), the subiculum (graphs 5–9), and the hippocampus (graphs 10–13). From Bayer and Altman (1995).

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in horizontal sections and are therefore not labeled in any of the photographs in Figs. 7–24. They share a triangular-shaped core of deep neurons that are generated from E15 to E17 (graph 7, Fig. 5), later than the deep neurons in the laterally situated entorhinal cortex. There is a lateral (older) to medial (younger) neurogenetic gradient in the superficial cells of the parasubiculum and the presubiculum. Those in the parasubiculum are generated earlier (mainly on E17 and E18) than the small superficial neurons in the presubiculum (compare graphs 5 and 6, Fig. 5). The pyramidal layer in the subiculum proper appears to be an extension of the deep neurons of the parasubiculum and the presubiculum. Within the layer, deep neurons are generated earlier (peak on E16) than superficial neurons (peak on E17, compare graphs 8 and 9, Fig. 5). When taken as a whole, neurons in the subiculum proper are generated later than the deep neurons in the parasubiculum and the presubiculum. These patterns indicate that the lateral (older) to medial (younger) neurogenetic gradient that begins in the entorhinal cortex continues throughout the subiculum (Bayer, 1980a). Hippocampus The hippocampus contains two interlocked C-shaped layers of cortex, dominated by the pyramidal cells of Ammon’s horn (fields CA1–3) and the granule cells in the dentate gyrus. In Ammon’s horn of the rat, the oldest pyramidal cells are in field CA3ab (peak on E17, graph 11, Fig. 5) and younger pyramidal cells flank them in fields CA1 (closer to the subiculum, graph 10, Fig. 5) and CA3c (in the hilus of the dentate gyrus, graph 12, Fig. 5). It is remarkable that the lateral (older) to medial (younger) neurogenetic gradient seen throughout the entorhinal cortex and the subiculum is broken by the sandwich gradient seen in the pyramidal cells of Ammon’s horn (Bayer, 1980a). The granule cells in the dentate gyrus are noted for their exceptionally late time of origin (reviewed in Bayer and Altman, 1974; Bayer, 1980a). Approximately 85% of these neurons are generated after birth in rats, mainly during the first postnatal week (graph 13, Fig. 5). Neurogenesis gradually tapers off during the second and third postnatal weeks, so that the dentate gyrus appears mature by the time of weaning (21 days). Most of the dentate granular neurons settle in a superficial (older) to deep (younger) gradient, opposite to the gradients between and within the layers of the neocortex (Bayer, 1980a). The dentate granular layer is also unusual because there are always a few neurons that can be labeled after [3H]thymidine injections are given to juvenile and adult rats (Altman, 1963; Altman and Das, 1965; Bayer, 1982; Bayer et al., 1982). It has

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been established that the neurons produced in adults add to the total population of dentate granule cells. One-year-old rats have significantly more neurons than 1-month-old rats (Bayer, 1982; Bayer et al., 1982).

Olfactory Bulb and Peduncle The Olfactory Bulb The olfactory bulb is a prominent component of the rat forebrain. Olfactory nerve fibers that originate from the olfactory epithelium in the upper part of the nasal cavity penetrate small foramina in the cribriform plate and terminate in the superficial glomerular layer of the olfactory bulb. In the glomerular layer, olfactory nerve axons synapse with the primary dendritic branches of mitral and tufted neurons. Axons of the mitral and tufted cells leave the olfactory bulb to terminate in various parts of the olfactory peduncle and in the primary olfactory cortex. There are three populations of short-axon interneurons: a large population of granule cells that forms a thick layer beneath the layer of mitral cells, small neurons scattered diffusely in the external plexiform layer, and small neurons dispersed between the glomeruli. The external plexiform layer lies between the glomerular and the mitral cell layers; it is a region where the secondary branches of mitral cell and tufted cell dendrites interact with input from granule cells and the external plexiform interneurons; it also contains the scattered cell bodies of the tufted output neurons (for details of olfactory bulb anatomy, see Bayer, 1983). Neurogenetic timetables in the olfactory bulb (graphs 1–7, Fig. 6) show a highly sequential pattern of generation between different neuronal populations (Bayer, 1983). The oldest neurons are the mitral cells that originate mainly on E14–E16. The internal, external, and interstitial tufted cells follow, with peaks on E16–E17, E18–E19, and E20–E22, respectively. The interneurons in the glomerular, external plexiform, and granular layers are generated mainly after birth (graphs 5–7, Fig. 6). Nearly all of the external plexiform cells and the periglomerular cells are generated by the end of the first postnatal week, but the large population of granule cells continues to be generated up to and beyond P19 (graph 7, Fig. 6). Granule cells in the olfactory bulb continue to be generated during the adult period since a few are always labeled within a few weeks after [3H]thymidine injections are given to adult rats (reviewed in Bayer, 1983). The Olfactory Peduncle The anterior olfactory nucleus is located posterior to the olfactory bulb in the olfactory peduncle. It is one of the major olfactory processing centers; the olfactory

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bulb is its major afferent input and is also the principal target of its axons. The anterior olfactory nucleus can be divided into a pars externa, an ectopic group of neurons in the anterior dorsolateral part of the peduncle, and the anterior olfactory nucleus proper, which contains the pars dorsalis, pars lateralis, pars ventralis, and pars medialis. The posterior parts of the anterior olfactory nucleus proper form transition areas to the primary olfactory cortex in the piriform lobe. A posterior (older) to anterior (younger) neurogenetic gradient is found both within and between components of the olfactory peduncle (Bayer, 1986a). Neurons in the pars externa are generated mainly between E16 and E19 (graph 8, Fig. 6), those in the anterior olfactory nucleus proper from E15 to E20 (graph 9, data in Fig. 6 are combined for all subdivisions), and those in the posterior transition areas from E14 to E19 (data are not shown in Fig. 6, see Bayer, 1986a). Only 3–4% of the neurons in the most anterior pars lateralis and pars dorsalis originate after birth. All parts of the anterior olfactory nucleus proper have a strong superficial (older) to deep (younger) neurogenetic gradient, while many of the transitional areas

have a gradient in the opposite direction, deep (older) to superficial (younger). These data suggest that characteristic patterns of neurogenesis, namely, the “insideout” versus the “outside-in” gradients, distinguish nuclear and cortical components of the olfactory brain. There is evidence that the pattern in which the anterior olfactory nucleus sends axons into the olfactory bulb is related to time of neuron origin: posterior parts project to the bulb first, anterior parts project later (reviewed in Bayer, 1986a).

MAPS OF STEM CELL MOSAICS IN THE TELENCEPHALIC NEUROEPITHELIUM The maps are presented in Figs. 7 through 24 from E11 to E22 in both the coronal (Figs. 7–14) and the sagittal planes (Figs. 15–24). In both sets of figures, the maturing components of the brain contained within the sections are identified as soon as they become recognizable. The maps are best viewed together, so they are placed near the end of Part IV of this chapter.

FIGURE 6 Timetables of neurogenesis for major neuronal populations in the olfactory bulb (graphs 1–7) and the anterior olfactory nucleus (graphs 8–9). From Bayer and Altman (1995).

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The Two Sources of Neurons: The Neuroepithelium and the Secondary Germinal Matrices The Neuroepithelium The primary source of cells specific to the nervous system (neurons, glia, choroid plexus, and ependyma) is a proliferative germinal matrix that is variably referred to as (1) the primitive ependymal layer, (2) the ventricular zone, or (3) the neuroepithelium (Langman et al., 1966). The latter term is now widely used. During gastrulation, the neuroepithelium differentiates from a sheet of columnar proliferative cells on the dorsal surface of the embryo, the neural plate. The neural plate subsequently forms a neural tube with a central fluidfilled lumen. The neural tube extends throughout the entire neuraxis. Its caudal part becomes the spinal cord, which retains its tubular appearance throughout life. Its rostral parts form three primary brain vesicles, the rhombencephalon (hindbrain), the mesencephalon (midbrain), and the prosencephalon (forebrain). Figures 7, 15, and 16 show high-magnification photographs of the prosencephalic neuroepithelium at an early stage after neural tube closure. Continuing cell proliferation in specific loci greatly expands the primary brain vesicles. The rhombencephalon subdivides into the myelencephalon (medulla) and the metencephalon (cerebellum and pons). The mesencephalon subdivides into the tectum (superior and inferior colliculi) and the tegmentum. The prosencephalon subdivides into the diencephalon (thalamus and hypothalamus) and the paired telencephalic vesicles (cerebral cortex and basal ganglia). Midbrain and hindbrain structures are shown in the thumbnail photographs of sagittal sections in Figs. 15–24. There is growing evidence that, notwithstanding its apparently homogeneous cellular composition, the neuroepithelium contains a heterogeneous population of neural stem cells that constitute the blueprint (Bauplan) of the mature central nervous system. In the following discussion we concentrate on the spatiotemporal neuroepithelial mosaic in the telencephalon where committed neural stem cells generate specific neurons according to strict timetables. The most probable germinal sources of telencephalic neurons are presented with comments about the initial radial migratory paths many of these neurons take as they settle in the maturing brain. Complex and stepwise migration patterns in the neocortex and major migratory streams in the telencephalon are summarized in Part IV. The Subventricular Zone Cells leaving the neuroepithelium have three fates: (1) they become postmitotic and differentiate as neurons, (2) they become glia or glial precursors, and

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(3) they retain their proliferative capacity and form secondary germinal matrices. The best-known secondary germinal matrix is the external germinal layer (egl) of the cerebellum. The egl forms a discreet layer over the surface of the developing cerebellum and produces granule cells, basket cells, and stellate cells in the cerebellar cortex (Altman and Bayer, 1996). The secondary germinal matrices of the telencephalon form other types of aggregates. In the developing cerebral cortex and basal ganglia, the secondary germinal matrix forms a subventricular zone adjacent to the primary neuroepithelium. In the developing olfactory bulb, proliferating cells migrate in a rostral migratory stream from the subventricular zones of the septum, accumbens, and cortex (Altman, 1969). In the developing hippocampus, the dentate migratory stream (Altman and Bayer, 1990a,b,c) contains the progenitors of granule cells in the dentate gyrus.

Germinal Sources of Telencephalic Neurons Neuroepithelium of the Ventral Telencephalon This neuroepithelium generates the pallidum, striatum, olfactory tubercle, nucleus accumbens, amygdala, septum, and part of the primary olfactory cortex. Its medial border extends to the point above the septal neuroepithelium (region 5 in the maps) where the neuroepithelium thins in the medial telencephalic wall (region 1B in the maps) and invaginates into the lateral ventricle as the choroid plexus. There are two prominent swellings of this neuroepithelium, a lateral ganglionic eminence (region 4A in the maps) and a medial ganglionic eminence (region 4B in the maps). The neuroepithelium in the corticopallidal angle is designated as region 3/4 in the maps because it generates neurons in cortical as well as ganglionic structures. Genetic expression studies (Stoykova et al., 2000; reviewed in Schuurmans and Guillemot, 2002) indicate that the neuroepithelium in regions 5, 4A, and medial 4B express genes Nkx2.1, several Dlx genes, Vax1, and Mash1. The neuroepithelium in lateral region 4B expresses Pax6, a gene expressed in the cortical neuroepithelium. Region 3/4 neuroepithelium expresses many of the same genes expressed in the cortical neuroepithelium including Pax6, Ngn1/2, Emx1/2 (Stoykova et al., 2000; Schuurmans and Guillemot, 2002), and Sox2 (Zappone et al., 2000). Pallidum The entopeduncular nucleus neurons originate in a strip of neuroepithelium that bridges the ventral telencephalic/diencephalic border (Altman and Bayer, 1986). That germinal source is part of the neuroepithelium that generates the zona incerta and (Text continues on p. 62)

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FIGURE 7 Nissl-stained 10-μm parrafin sections of the rat anterior coronal forebrain on E11 (A) and E12 (B). A is modified from E11 coronal plate 2; B is modified from E12 coronal plate 3 in Altman and Bayer (1995).

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FIGURE 8 Nissl-stained 10-μm parrafin section of the rat anterior coronal forebrain on E13; modified from E13 coronal plate 3 in Altman and Bayer (1995). The right half of the photograph is at low contrast so that labels are more visible. Arows indicate cells exiting the ganglionic and septal parts of the neuroepithelium. The incorporated table lists the compartments in the neuroepithelium and the major events taking place, based on timetables of neurogenesis in Figs. 1–6.

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FIGURE 9 As in Fig. 8 for the rat anterior coronal forebrain on E14; modified from E14 coronal plate 3 in Altman and Bayer (1995).

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FIGURE 10 As in Fig. 8 for the rat anterior coronal forebrain on E15; modified from E15 coronal plate 5 in Altman and Bayer (1995).

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FIGURE 11 As in Fig. 8 for the rat anterior coronal forebrain on E16; modified from E16 coronal plate 6 in Altman and Bayer (1995).

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FIGURE 12 As in Fig. 8 for the rat anterior coronal forebrain on E17; modified from E17 coronal plate 7 in Altman and Bayer (1995).

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FIGURE 13 As in Fig. 8 for the rat anterior coronal forebrain on E18; modified from E18 coronal plate 5 in Altman and Bayer (1995).

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FIGURE 14 As in Fig. 8 for the rat anterior coronal forebrain on E20; modified from E20 coronal plate 7 in Altman and Bayer (1995).

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FIGURE 15 Nissl-stained 10-μm parrafin sagittal section of the rat brain on E11; modified from E11 sagittal plate 2 in Altman and Bayer (1995). A shows the entire section with a box indicating the limits of the enlarged photograph in B.

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FIGURE 16 Nissl-stained 10-μm parrafin sagittal section of the rat brain on E12; modified from E12 sagittal plate 4 in Altman and Bayer (1995). A shows the entire section with a box indicating the limits of the enlarged photograph in B.

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FIGURE 17 Nissl-stained 10-μm parrafin sagittal section of the rat brain on E13; modified from E13 sagittal plate 3 in Altman and Bayer (1995). A shows the entire section with a box indicating the limits of the enlarged photograph in B.

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FIGURE 18 Nissl-stained 10-μm parrafin sagittal section of the rat brain on E14; modified from E14 sagittal plate 3 in Altman and Bayer (1995). A shows the entire section with a box indicating the limits of the enlarged photograph in B.

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FIGURE 19 Nissl-stained 10-μm paraffin sagittal section of the rat brain on E15; modified from E15 sagittal plate 3 in Altman and Bayer (1995). A shows the entire section with a box indicating the limits of the enlarged full-contrast photograph in B. C on the facing page shows a low-contrast copy of B with labels and an explanatory table.

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FIGURE 19, cont’d

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FIGURE 20 As in Fig. 19 for the sagittally sectioned rat brain on E16; modified from E16 sagittal plate 4 in Altman and Bayer (1995).

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FIGURE 20, cont’d

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FIGURE 21 As in Fig. 19 for the sagittally sectioned rat brain on E17; modified from E17 sagittal plate 3 in Altman and Bayer (1995).

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FIGURE 21, cont’d

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FIGURE 22 As in Fig. 19 for the sagittally sectioned rat brain on E18; modified from E18 sagittal plate 4 in Altman and Bayer (1995).

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FIGURE 22, cont’d

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FIGURE 23 As in Fig. 19 for the sagittally sectioned rat brain on E20; modified from E20 sagittal plate 3 in Altman and Bayer (1995).

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FIGURE 23, cont’d

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FIGURE 24 As in Fig. 19 for the sagittally sectioned rat brain on E22; modified from E22 sagittal plate 3 in Altman and Bayer (1995).

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FIGURE 24, cont’d

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the lateral hypothalamus, and it is not represented in any of the sections shown in the maps. Neurons migrate rapidly away from this site soon after their generation, and the entopeduncular nucleus is recognizable as early as E13, presumably containing neurons that were generated on E12 (Altman and Bayer, 1986). With the exception of the entopeduncular nucleus, all of the other pallidal neuronal populations have linked neurogenetic gradients that suggests these nuclei represent a single large system of magnocellular neurons populating the basal telencephalon (graphs 2–5, Fig. 1; Bayer, 1985b). It further implies that these neurons are generated sequentially in an anterior– posterior (longitudinal) ridge of neuroepithelium. The most likely source is the medial eminence of basal ganglia neuroepithelium (region 4B, Figs. 8–11 and 16–20). A few pallidal neurons migrate out of the basal ganglia neuroepithelium on E13 (Fig. 8) and many more migrate between E14 and E16 (Figs. 9–11). Posterior and ventral globus pallidus neurons settle within 2 days after their generation (see Figs. 29 and 30 in Altman and Bayer, 1986), and other pallidal neurons probably migrate predominantly radially from the medial eminence to settle throughout the ventral telencephalon. Striatum and small olfactory tubercle neurons Striatal medium spiny neurons and small neurons in the olfactory tubercle are probably generated in two steps by the basal ganglia neuroepithelium. (1) The primary neuroepithelium produces secondary neural stem cells that move into the subventricular zone and continue to proliferate; this takes place from E16 to E18 (Figs. 11–13). (2) Secondary neural stem cells in the subventricular zone produce medium spiny striatal neurons. Some of the oldest striatal neurons in the ventrolateral striatum are generated directly by primary neural stem cells in the lateral ganglionic eminence neuroepithelium (region 4A in the maps), because presumptive striatal neurons are already outside of the striatal subventricular zone on E16 (Fig. 11). At the time when striatal neurons start to be produced in large numbers (E18–19, graph 6, Fig. 1) there is only one eminence in the basal ganglia neuroepithelium and a single large subventricular zone (region 4, Fig. 13). Prior to that both eminences are present, each with a large subventricular zone. Nucleus accumbens The nucleus accumbens is another structure where most neurons are generated in a two-step process. Secondary neural stem cells committed to produce nucleus accumbens neurons move into the subventricular zone immediately surrounding the neuroepithelium in the inferior horn of the lateral ventricle (region 6 in the maps). As early as E17 (Figs. 12

and 21), a band of differentiating cells, presumably the oldest nucleus accumbens neurons, surrounds the subventricular zone in the inferior horn. Younger neurons settle consecutively inside surrounding rings of older neurons. Peak proliferative activity in the nucleus accumbens subventricular zone coincides with the peak period of neurogenesis. Low-level exposures to X-irradiation massively kill cells in the subventricular zone during the peak time of neurogenesis, while the neuroepithelium itself is less affected, indicating that the neuroepithelium is probably producing the ependyma (Bayer, 1979b). It is important to note that the subventricular zone producing the nucleus accumbens is also one source of secondary neural stem cells in the rostral migratory stream. Amygdala Early generated medium-sized and large-sized neurons in the anterior amygdaloid area are probably produced by primary neural stem cells in both eminences of the basal ganglia neuroepithelium (regions 4A and 4B in the maps); later generated small neurons may originate from secondary neural stem cells in the large basal ganglia subventricular zone (outside region 4 in the maps). Large- to medium-sized anterior amygdaloid neurons settle among neurons of the ventral pallidum just outside the basal ganglia neuroepithelium as early as E14. The anterior parts of the central nucleus and the basolateral group most likely are generated in the lateral eminence of the basal ganglia neuroepithelium (region 4A). Some modified pyramidal-like cells in the basolateral amygdala may be generated either in the cortical neuroepithelium (region 3) or in the corticopallidal angle (region 3/4) and may be the migrating neurons expressing Pax6 shown by Stoykova et al. (2000). Younger neurons in the posterior central nucleus appear to be part of the striatum and may be generated in the striatal subventricular zone. Neurons in the intercalated masses may be generated in the cortical neuroepithelium (region 3), migrate in the lateral migratory stream to the reservoir of the stream, and then leave to disperse between various amygdaloid nuclei. Neurons in the nuclei of the lateral and accessory olfactory tracts are not visible in the amygdala until relatively late. Possibly, these neurons are generated in a neuroepithelium that produces olfactory bulb mitral cells in the anterior basomedial floor of the ventral telencephalon (anterior region 6/posterior region 7 in the E15 map, Fig. 19). The late appearance of these nuclei suggests that they are generated in a distant source, then migrate posteriorly to settle in the amygdala. Morphogenetic studies of the bed nucleus of the stria terminalis indicate two distinct neuroepithelial sources (Bayer, 1987). The anterior bed nucleus is generated by

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the primary neuroepithelium in the inferior horn of the lateral ventricle posterior to the nucleus accumbens neuroepithelium (region 6A in the maps). Neurons migrate radially from that source so that older neurons settle ventromedially, and younger neurons settle dorsolaterally, closest to the inferior horn. The preoptic continuation of the strial bed nucleus is generated in the neuroepithelium at the base of the posterior part of the medial horn of the lateral ventricle, lateral to the area of fusion with the anterior thalamic neuroepithelium (region 4C in the maps). This is also part of the basal ganglia neuroepithelium (Figs. 19–21). Neurons migrate radially from that source and settle in a downward medial-curving pattern that extends toward the sexually dimorphic nucleus in the medial preoptic area. Younger neurons accumulate adjacent to the older lateral neurons. Throughout the anterior and posterior bed nucleus, neurons begin to migrate within 1 day after their generation and usually settle 1 or 2 days later (Bayer, 1987). Septum The source of most neurons in the septal complex is the neuroepithelium lining the ventromedial wall of the lateral ventricle (region 5, Figs. 7–14 and 19–24; Bayer, 1979b). Already by E15 in rat embryos, a thick band of young neurons has accumulated outside of that neuroepithelium (see it best in the coronal section, Fig. 10). The young neurons are presumably those that will form the medial septal and diagonal band nuclei. By E17, the neuroepithelium is less prominent, and more young neurons, presumably those of the lateral septal nucleus, accumulate just outside it, and the vertical limb of the diagonal band is recognizable (Fig. 12). More neurons accumulate on E18 (Fig 13) and E19. The septal neuroepithelium is thin by E20 (Fig. 14) and is changing to the primitive ependyma, while the zone of differentiating cells continues to enlarge. Neuroepithelium of the Dorsal Telencephalon This part of the neuroepithelium contains two major parts, the cortical (region 3) and the hippocampal (regions 2A, 2B, and 2C). The cortical neuroepithelium forms the roof of the telencephalon and is the source of Cajal–Retzius neurons, subplate neurons, and pyramidal cells throughout all layers in the limbic cortex and neocortex. Along with the neuroepithelium in the corticopallidal angle (region 3/4), the cortical neuroepithelium probably generates layer II neurons in the primary olfactory cortex and some neurons in the amygdala. The hippocampal neuroepithelium forms the medial hem of the cortex and is the source of neurons in the subiculum, Ammon’s horn of the hippocampus, and the dentate gyrus; it also contains a

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glial matrix for the fimbria and contributes cells to the invaginating choroid plexus. Neuroepithelial cells in regions 3 and 2A express Pax6, Ngn1/2, Emx1/2 (Stoykova et al., 2000; Schuurmans and Guillemot, 2002), Otx1 (Tarabykin et al., 2001), Sox2 (Zappone et al., 2000), and Lhx2 (Bulchand et al., 2001). The Bulchand et al. (2001) study provides a detailed map of additional genes expressed in the hippocampal neuroepithelium that correlate with its unique progeny in the cortex. Region 2A expresses EphB1; regions 2B and 2C have highly segregated expression of several Wnt genes, including Wnt2b, Wnt3a, and Wnt5a; regions 2A/B and the choroid plexus (region 1B) express Bmp4 and Bmp7; and the choroid plexus expresses Msx1. Neocortex and limbic cortex Using short- and sequential-survival [3H]thymidine autoradiography and exposures to low-level X-irradiation, timedependent successive transformations in the cortical neuroepithelium can be detected and linked to the three epochs of cortical neurogenesis (see Chapters 4 and 10 in Bayer and Altman, 1991a). During the first epoch (E13–E15), the cortical neuroepithelium is composed mainly of primary neural stem cells and radial glial cells. There are sparse founder cells of secondary neural stem cells that will move into the subventricular zone, glial stem cells, and ependymal stem cells. A single exposure to 200-R X-rays kills most cells in the neuroepithelium and the dying cells drop down into the ventricular lumen. [3H]Thymidine autoradiography indicates that most neuroepithelial cells undergo interkinetic neuronal migration. In that process the nucleus migrates within the cytoplasm during every cell cycle. DNA is duplicated when the nucleus is in the basal part of the cell, and mitosis occurs at the cell apex near the ventricular lumen. Two-hour-survival [3H]thymidine autoradiography shows a band containing many heavily labeled nuclei (the synthetic zone) at the base of the neuroepithelium and few labeled nuclei near the ventricular lumen (the mitotic zone). Cajal–Retzius and subplate neurons are the chief progeny of the primary neural stem cells during the first epoch. These neurons form clumps of heavily labeled cells in the basal part of the neuroepithelium 24 h after a single [3H]thymidine injection; 48 h after the injection, heavily labeled Cajal–Retzius and subplate neurons are outside all parts of the cortical neuroepithelium and may represent the Tbr1expressing cells shown by Stoykova et al. (2000). Also during this time, the cortical neuroepithelium is increasing in size (compare the size changes in Figs. 10–12) because it contains a large population of selfreplicating primary neural stem cells that will produce neurons later.

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During the second epoch (E16–E17), primary neuronal precursors undergoing interkinetic nuclear migration still predominate in the cortical neuroepithelium. Two hours after a [3H]thymidine exposure, nuclei are heavily labeled in a distinct synthetic zone but the number of heavily labeled nuclei in the mitotic zone is also increasing. Although low-level X-ray still kills most cells in the neuroepithelium, many cells survive. The shedding of dead cells into the ventricle declines on E16 and nearly stops on E17 (see Chapter 10 in Bayer and Altman, 1991a). The surviving cells are probably ependymal stem cells at the ventricular lumen and glial precursors; both cells are more radioresistant than neural stem cells. Despite the fact that primary neural stem cells are declining on a relative basis, the cortical neuroepithelium is still expanding on an absolute basis, and it reaches its greatest volume on E17. The chief progeny of the primary neural stem cells during this epoch are pyramidal cells that will settle in layers VI and V. These neurons rapidly move out of the neuroepithelium after they are generated and sojourn in layer-specific bands in the intermediate zone. Secondary neural stem cells also leave the neuroepithelium during this time to continue proliferating in the cortical subventricular zone, where they will generate some of the upper layer neurons in layers IV–II (Smart and McSherry, 1982; Tarabykin et al., 2001). During the third epoch (E18–E21), primary neuronal precursors undergoing interkinetic nuclear migration decline and disappear in the cortical neuroepithelium. Two hours after a [3H]thymidine exposure, heavily labeled nuclei are randomly distributed in the neuroepithelium; the distinction between the synthetic and the mitotic zones disappears. Low-level X-ray kills fewer cells in the neuroepithelium, indicating that ependymal stem cells form an intact layer at the ventricular lumen. It is during this third epoch that the volume of the cortical neuroepithelium shrinks both relatively and absolutely. By E20, the cortical neuroepithelium is already a primitive ependyma. The chief progeny of the neural stem cells during this epoch are in neurons in layers IV–II (Bayer and Altman, 1991a). The heterogeneous array of neurons in these layers probably originate from primary neural stem cells in the cortical neuroepithelium (regions 3 and 3/4) and secondary neural stem cells in the cortical subventricular zone. The neurons generated in the cortical neuroepithelium sojourn in a prominent band in the lower subventricular zone before migrating to the cortical plate (see Chapter 7, Figs. 7–3 to 7–8 in Bayer and Altman, 1991a). Secondary neural stem cells and their progeny in the subventricular zone express Svet1 during the peak times of layer IV–II neurogenesis.

Svet1 cells migrate through the intermediate zone, through the lower cortical plate, and settle in the upper cortical plate (Tarabykin et al., 2001). It is possible that the layer IV–II neurons generated in the cortical neuroepithelium also begin to express Svet1 during their sojourn in the lower subventricular zone, making that zone critical in determining an upper layer fate (Tarabykin et al., 2001) A new hypothesis is that most neocortical interneurons are generated in the medial eminence of the basal ganglia neuroepithelium (region 4B) and migrate into the cortex (Anderson et al., 1997, 2001; Denaxa et al., 2001). The dorsal and ventral parts of the telencephalic neuroepithelium and subventricular zone are continuous at the cortical–pallidal angle (region 3/4, Figs. 9–14), and cells could migrate through that region from the basal ganglia to the cortex (De Carlos et al., 1996). Piriform (primary olfactory) cortex Neurons in layers III–IV are generated in the corticopallidal angle (region 3/4) and in the lateral part of region 4A (Figs. 8–10; Bayer and Altman, 1991b; De Carlos et al., 1996). Neurons in layer II are probably generated in the cortical neuroepithelium (regions 3 and 3/4) and migrate laterally and ventrally for several days before settling above the layer III–IV neurons (Figs. 10–12; Bayer and Altman, 1991a). The layer II neurons in the piriform cortex appear late as a thin ventrolateral extension of the lateral limbic cortical plate. The posterior part of the piriform cortex matures earlier than the anterior part, in accordance with the posterior to anterior neurogenetic gradient (reviewed in Bayer, 1986b). Hippocampal region The neuroepithelium in the posterolateral cortical primordium is the presumed source of neurons in the entorhinal cortex (Bayer, 1980b). That neuroepithelial area is not in any of the sections shown, so a brief verbal summary of events follows. In rat embryos on E16, the entorhinal neuroepithelium is thick and a zone of young neurons migrate outward from its lateral part. The entorhinal neuroepithelium is still prominent on E17, and there are layers resembling the cortical intermediate zone and a cortical plate; both of these are thicker laterally than medially, reflecting the prominent lateral (older) to medial (younger) neurogenetic gradient that is found throughout the entorhinal cortex. By E18, the entorhinal neuroepithelium thins, coinciding with reduced neurogenesis, while the cortical plate becomes thicker, again more so laterally than medially. Also on E18, a cell-sparse region develops beneath the cortical plate in the entorhinal region that is similar to the upper intermediate zone in the neocortex. On E19, a

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split appears in the entorhinal cortical plate, possibly representing the lamina dessicans in layer IV. That morphological feature distinguishes the entorhinal area from the limbic and neocortical areas, where the cortical plate has no cell-sparse zones. The splitting of the cortical plate proceeds from a lateral to a medial direction up to E20. Although the lamination of the entorhinal cortex is obvious by E22, there are still a few spindle-shaped radially oriented cells in the intermediate zone, indicating that entorhinal neurons are still migrating. The presumed source of the subiculum (Bayer, 1980b) is the part of the hippocampal neuroepithelium (region 2A, Figs. 8–11 and 19–22) that is continuous with the cortical neuroepithelium. Between E18 (Fig 22) and E20 (Fig 23) the cortical plate extends into the differentiating zones outside the subicular/ammonic part of the hippocampal neuroepithelium. In horizontal sections of rat embryos, the bifurcation of the entorhinal cortical plate ceases to progress medially by E20, and the adjacent wedge-shaped nonbifurcated cortex can be more accurately delineated as the parasubiculum and the presubiculum. The neurons in this part of the cortical plate arrive there from E18 through E20 and probably represent deep neurons in the parasubiculum, the presubiculum, and the subiculum proper. The parasubiculum cannot be distinguished from the presubiculum until E22 because the migration of small neurons to the superficial layers of the presubiculum is exceptionally late (Bayer, 1980b). [3H]Thymidine autoradiographic studies indicate that the “bulge” in the neuroepithelium of the hippocampus has three components (Altman and Bayer, 1990a,b,c). One (region 2A) gives rise to the pyramidal cells of Ammon’s horn, a second (region 2B) gives rise to the granule cells in the dentate gyrus, and a third (region 2C) gives rise to glia that will populate the fimbria and probably part of the choroid plexus. Region 2A appears on E14 in rat embryos (Figs. 9 and 18). That neuroepithelium shows a high level of proliferative activity up to E18 (Figs. 9–12 and 18–22); relatively few pyramidal neurons are generated on E20 and the neuroepithelium declines (not shown). After a single [3H]thymidine injection on E18, the migratory routes of the pyramidal cells were tracked by killing animals at daily intervals after the injection (Altman and Bayer, 1990b). All of the pyramidal cells move out of the neuroepithelium 1 day after their generation and form a band of heavily labeled cells just outside it. On subsequent days, the pyramidal cells leave this band and migrate into the pyramidal layer. CA1 neurons migrate radially and take 4 days to reach their destina-

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tions. Although the CA3 neurons are generated earlier than the CA1 neurons, they take longer to migrate to the pyramidal layer because part of their migratory trajectory is a curved path around the edge of the accumulating CA1 neurons. Possibly the earlier time of origin of CA3 neurons is related to their longer pathway of migration. At the time of birth in rats, many CA3 pyramidal cells are still migrating into the lengthening pyramidal layer. The granule cells of the dentate gyrus are ultimately derived from the dentate neuroepithelium that indents slightly at the edge of the Ammonic neuroepithelium, a region called the “dentate notch” (region 2C, Figs. 11, 12, and 19–21; Altman and Bayer, 1990a,b,c), but the secondary neural stem cells that give rise to granule cells migrate from the notch in the dentate migratory stream before producing granule cells (see Part IV, Figs. 21–24). Olfactory Bulb and Peduncle Olfactory nerve fibers reach the telencephalon on E14 (Fig. 18) and define the part of the telencephalic neuroepithelium that begins to evaginate into an olfactory bulb on E15 (region 7, Fig. 19). However, [3H]thymidine autoradiography indicates that the mitral neurons of the olfactory bulb are generated in the septal/accumbal neuroepithelium (regions 5 and 6) and migrate into the olfactory bulb (Figs. 19–21; Bayer, unpublished observations). On E15 (Fig. 19), mitral neurons accumulate at the base of the brain behind the olfactory nerve fibers and their axons start to form the lateral olfactory tract. On E16 (Fig. 20), some earlygenerated mitral cells reach the evaginating olfactory bulb. On E17 (Fig. 21) and E18 (Fig. 22), more mitral neurons migrate into the bulb from a ventral direction and curve around it dorsally. The mitral cell layer appears on E17 (Fig. 21) and becomes more definite from E18 to E22 (Figs. 22–24). The neuroepithelium in the olfactory bulb itself may give rise to accessory olfactory bulb mitral cells and tufted cells that migrate radially to settle in the external plexiform layer outside the mitral cell layer (Figs. 23 and 24). Interneurons and granule cells are produced by secondary neural stem cells that migrate in the rostral migratory stream from the subventricular zones of the cortex and ventral telencephalon (Figs. 21–24). The anterior olfactory nucleus in the olfactory peduncle is generated just after the mitral neurons in the main olfactory bulb. It is likely that these neurons are produced by either the same neuroepithelium as mitral neurons (regions 5 and 6) or by the most posterior and ventral part of the evaginating olfactory neuroepithelium (region 7, Figs. 21–23).

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DEVELOPMENT OF THE LATERAL, ROSTRAL, AND DENTATE MIGRATORY STREAMS Lateral Migratory Stream and Cell Migration in the Cerebral Cortex The lateral migratory stream is a prominent band of horizontally oriented cells adjacent to the cortical subventricular zone in the lower intermediate zone that extends from the dorsomedial to the ventrolateral extent of the cortex. It is best seen in coronal sections (Figs. 12–14). In sagittal sections, only a thin part of the stream is visible (Figs. 21–24). This is the most complex migratory stream in the telencephalon for three reasons. First, only some neocortical neurons migrate in the stream. The Cajal–Retzius and subplate neurons migrate and settle without using the lateral migratory stream. Neurons settling in the dorsal neocortex probably do not use the stream. Second, the neocortical neurons in the stream are in the second stage of their migration after they have sojourned for approximately 24 h in layer-specific bands either in the intermediate zone or in the subventricular zone (Bayer and Altman, 1991a). Third, the cortical neuroepithelium produces neurons that migrate in the stream to destinations outside the neocortex before, during, and after neocortical neurons migrate in the stream. Migration of Cajal–Retzius and Subplate Neurons The Cajal–Retzius and subplate neurons are sequestered in the neuroepithelium after they are generated and migrate out radially within 1 day (Bayer and Altman, 1991a) to immediately settle in the primordial plexiform layer without entering the lateral migratory stream. The Cajal–Retzius cells assume a horizontal orientation, settle beneath the pial membrane among a subpial system of extracellular channels, and rapidly begin to differentiate. These channels are soon filled with early-arriving axons that synapse with Cajal– Retzius dendrites. The subplate neurons take several days before settling permanently in a morphologically distinct subplate. First, they accumulate beneath the Cajal–Retzius neurons in the primordial plexiform layer. Second, the subplate neurons become radially oriented and form a cortical plate (see it ventrolaterally on E16, Fig. 11). The thin cortical plate that extends to the dorsomedial cortex on E17 is composed mainly of subplate neurons (Fig. 12). As the cortical plate expands from ventrolateral to dorsomedial on E16 and E17, a deep set of extracellular channels appears (prominent white bands beneath the ventrolateral cortical plate on the left half of the photographs in Figs. 11 and 12). Third, subplate neurons delaminate from the cortical plate in a

ventrolateral to dorsomedial direction to settle permanently among the deep extracellular channels on E18 (Fig. 13). By E20 (Fig. 14) the subplate is a distinct layer below the cortical plate. The subplate neurons rapidly differentiate and form synaptic contacts with axons that grow into the deep channel system. It is postulated that these early migratory patterns of the Cajal–Retzius and subplate neurons are necessary prerequisites for the normal development of layers VI–II (reviewed in Bayer and Altman, 1991a). Migration of Layer VI–II Neurons Stage 1: Sojourn in layer-specific bands The neurons destined to settle in layers VI–V and many neurons in layers IV–II move out of the cortical neuroepithelium within 1 day after their generation and first sojourn in narrow layer-specific bands in the subventricular and intermediate zones (Bayer and Altman, 1991a). These bands can only be seen in autoradiographic sections 24 h after an exposure to [3H]thymidine. To save space, the pictures are not reproduced here, but see Chapter 7 in Bayer and Altman (1991a) for a detailed description of the layer-specific sojourn hypothesis. By correlating the timetables of neurogenesis with their sequential appearance, each band can be linked to a specific population of cortical neurons. The neurons destined to settle in layer VI sojourn in the upper intermediate zone very close to the base of the cortical plate; this is called the first superior band (sb1). The sb1 is in the rat cortex on E16 and E17, 1 day after [3H]thymidine injections on E15 and E16, and during the time that maximum numbers of layer VI neurons should be located outside of the neuroepithelium. Neurons bound for layer V sojourn in the second superior band (sb2) at a lower level in the intermediate zone. The sb2 is in the rat cortex on E17 and E18, just at the time when large numbers of layer V neurons should be outside of the neuroepithelium. Neurons bound for layers IV–II sojourn in the first inferior band (ib1) in the lower subventricular zone. The ib1 appears on E18, correlating with the first peak of neurogenesis of layer IV neurons, and remains until E21 when the youngest neurons bound for layers III and II are generated in the cortical subventricular zone. Many neurons in the ib1 are postulated to express Svet1 during their sojourn (Tarabykin et al., 2001) Stage 2: Predominantly radial migration to the cortical plate Layer VI–II cortical neurons continue their migration to the cortical plate after their sojourn. Those bound for layers VI–II in the dorsal neocortex migrate in the radial direction and settle on the next day; the entire sequence from birth to settling takes only 2 days. Layer VI and V neurons that settle in the

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dorsolateral neocortex (those generated on E15) also migrate predominantly radially and settle in 2–3 days because the cortical neuroepithelium is either immediately below or only slightly medial to the lateral cortical plate when they are migrating on E16 (Fig. 11) and E17 (Fig. 12). Stage 2: Predominantly lateral migration Layer V–II neurons generated on or after E17 that settle in the ventrolateral cortical plate cannot follow a radial migratory path. These neurons are generated approximately 0.5 to 1.0 mm medial to the points where they will penetrate the cortical plate because growth of the basal ganglia displaces the cortical neuroepithelium and the subventricular zone. After sojourning in the intermediate and the subventricular zones for 1 day, these neurons orient tangentially and enter the lateral migratory stream 2 days after their generation and migrate along the outside edge of the external capsule. The ventrolateral extension of the lateral migratory stream is first seen on E18 (Fig 13). One day later neurons leave the stream, turn, migrate radially through the upper intermediate zone, and finally enter the ventrolateral cortical plate around E20 4 days after their generation. Neurons bound for the far ventrolateral neocortex, such as the insular area, migrate laterally for 3 days and take 5 or more days to reach their destinations in the cortical plate. The later time of arrival of neurons in the ventrolateral and farventrolateral parts of the cortical plate is documented with [3H]thymidine autoradiograms in Chapter 9 in Bayer and Altman (1991a). Primary Olfactory Cortex Neurons and Others in the Basal Telencephalon The cortical neuroepithelium also gives rise to neurons that are destined to settle in the primary olfactory cortex and other sites in the basal telencephalon, such as the basolateral complex of the amygdala and the intercalated masses of the amygdala. Sequentialsurvival [3H]thymidine autoradiography after an injection on E15 can track heavily labeled cells moving from the neocortical lower intermediate zone into layer II of the primary olfactory cortex (Altman, unpublished observations). Layer II does not appear until E17 in the primary olfactory cortex as a thin ventrolateral extension of the neocortical plate (Fig. 12). In accordance with a ventral (older) to dorsal (younger) neurogenetic gradient, layer II continues to lengthen in the ventral direction as younger neurons are added dorsally on E18 (Fig. 13), E20 (Fig. 14), and beyond. Contrary to the neocortex, the layer III–IV neurons in the primary olfactory cortex do not form part of the cortical plate,

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possibly because they are generated by part of the basal ganglia neuroepithelium. Sequential-survival [3H]thymidine autoradiography after an injection on E15 can also track heavily labeled cells moving into deep parts of the ventral telencephalon, wrapping around and under the external capsule (Altman, unpublished observations). These neurons may be settling in the basolateral group of amygdaloid nuclei where some neurons resemble pyramidal cells in the cortex. Observations after later injections indicate that labeled cells migrate to and accumulate in a reservoir at the base of the lateral migratory stream (see Fig. 14). Cells move out of the reservoir and appear to migrate into the intercalated masses of the amygdala (Bayer, unpublished observations).

Rostral Migratory Stream and Adult Neurogenesis in the Olfactory Bulb The rostral migratory stream (Altman, 1969) is a large subventricular zone in the anterior forebrain that starts in the region of the septum and nucleus accumbens, curves around the olfactory neuroepithelium itself, and ends at the anterior border of the cortical subventricular zone. It appears on E17 both above and below the olfactory bulb neuroepithelium (rms, Fig. 21). From E18 on, the rostral migratory stream is continuous around the most anterior extent of the olfactory bulb neuroepithelium (Figs. 22–24). It is not known which part of the primary neuroepithelium contributes to the secondary neural stem cells that move into the rostral migratory stream, but it is likely that contributions come from cortical, septal, and nucleus accumbens neuroepithelia. Once inside the stream, secondary neural stem cells proliferate as well as migrate forward. Some secondary neural stem cells produce postmitotic neurons (olfactory granule cells, periglomerular granule cells, and other olfactory interneurons) far back in the stream and their progeny continues to migrate into the olfactory bulb; others may go all the way into the bulb before producing neurons (Altman, 1969; Kaplan and Hinds, 1977; Peretto et al., 1999). Once the cells reach the olfactory bulb, postmitotic neurons leave the stream and move into the granular layer. The outpouring of young neurons is especially prominent in the E22 brain (Fig. 24). The rostral migratory stream persists into adulthood and is a source of new olfactory bulb interneurons throughout adult life (Fig. 25A). Morphologic studies reveal that the adult rostral migratory stream contains bipolar undifferentiated cells (presumably neural stem cells) surrounded by astrocytes that form elongated tubes extending toward the olfactory bulb (Peretto et al., 1999). Quantitative studies (Roselli-Austin and Altman, 1979; Kaplan et al.,

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1985) have shown that the number of granule cells in the olfactory bulb increases in adults, a feature they share with granule cells in the dentate gyrus. [3H]Thymidine autoradiographic data indicate that many adult-generated neurons die and are replaced by the constant supply of new adult-generated neurons (Bayer, 1983; Kaplan et al., 1985); the subset of new adult neurons that show turnover may be related to turnover of the primary olfactory sensory neurons in the olfactory epithelium (Graziadei and Graziadei, 1979).

increases during adult life (Bayer et al., 1982; Bayer, 1982). In contrast to the granule cells of the olfactory bulb, there is little evidence of cell death in the granule cells of the dentate gyrus (Bayer, 1985c), and the functional importance of granule cell neurons may increase in the adult hippocampus.

Dentate Migratory Stream and Adult Neurogenesis in the Dentate Gyrus

Figure 26 summarizes the hypothetical changes that take place in neural and glial stem cell lines between E15 and E20 in the dorsomedial cortex. The diagram distinguishes two stages in cortical germinal zones. On E15 (Fig. 26A) and E16 (Fig. 26B) there is still only one germinal zone, the cortical neuroepithelium, which may be specified as early as E11 (Figs. 7A and 15). From E17 on (Figs. 26C–26F) there are two germinal zones, the primary cortical neuroepithelium and the secondary subventricular zone created by glial stem cells and neural stem cells exiting from the neuroepithelium. As cortical development proceeds, the primary germinal zone shrinks while the secondary germinal zone expands to reflect the continually changing number and types of neural, glial, and ependymal stem cell populations. The diagram in Fig. 26 shows neural stem cells and glial stem cells existing as separate populations in the early neuroepithelium (Figs. 26A and 26B), mainly for simplicity because the emphasis of our analysis is on the proliferation dynamics of stem cells, not on the time of commitment. Probably unique populations of stem cells arise sequentially in the cortical neuroepithelium through restriction of pluripotent stem cell lines; similar to “model B” in the Kalyani and Rao (1998) review of stem cell lineages in the spinal cord and peripheral nervous system. Retroviral cell lineage studies in the cortex (Grove et al., 1993; Krushel et al., 1993; McCarthy et al., 2001) support the initial heterogeneity of some stem cells (clones generate neurons and glia) and the sequential commitment and heterogeneity of others (clones restricted to specific types of neurons or specific types of glia).

The dentate/fimbrial neuroepithelium can be distinguished as a slight medial curve or “notch” at the base of the Ammon’s horn neuroepithelium on E15 (region 2B/C, Figs. 10 and 19) and E16 (Figs. 11 and 20). Region 2C of that neuroepithelium is the source of secondary neural stem cells that migrate into the dentate primordium and establish a secondary germinal matrix in the subgranular layer of the dentate gyrus. Beginning on E17, cells move out of the dentate neuroepithelium along the pia at the very edge of the cortex (dms, Fig. 21) and accumulate in a rounded area, the dentate primordium. The migrating cells on E17 are mostly postmitotic neurons (basket cells and large neurons in the molecular layer of the dentate gyrus) because short-survival [3H]thymidine autoradiography indicates no label uptake (Altman and Bayer, 1990c). By E18 (Fig. 22) many of the migrating cells are proliferating, indicating that secondary neural stem cells are in the stream (Altman and Bayer, 1990c). Up to E20 (Fig. 23), secondary neural stem cells continue to migrate into the dentate gyrus, following a curved path across the fimbria and around the expanding edge of Ammon’s horn. By E22 (Fig. 24), the migratory path is less distinct, and the subgranular layer of the dentate gyrus contains many secondary neural stem cells generating granule cell neurons. The external (or ectal) limb of the granular layer is already visible. During the first postnatal week, most of the granule neurons are generated and accumulate in a thick layer with older neurons stacked above younger neurons. Secondary neural stem cells thin out during the second and third postnatal weeks but many remain in a distinct subgranular layer (Fig. 25B). Secondary neural stem cells in the subgranular layer continue to produce granule neurons during juvenile and adult life. These new neurons move into the lower third of the granule cell layer (Altman, 1962; Altman and Das, 1965; Kaplan and Hinds, 1977; Bayer et al., 1982; Bayer, 1982; Kaplan and Bell, 1984). Quantitative studies indicate that the number of granule cells significantly

STEM CELL DYNAMICS IN CORTICAL GERMINAL ZONES

Neural Stem Cells Two different types of neural stem cells populate the cortical germinal zones. Primary neural stem cells (PNS, red and yellow ellipses, Fig. 26) are presumed to be columnar cells that go through the mitotic phase of the cell cycle at the ventricular lumen and generate neurons in the neuroepithelium. Secondary neural stem cells (SNS, orange ovals, Fig. 26) are initially in the neuroepithelium, move out, and generate neurons in the sub-

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FIGURE 25 A diagram summarizing adult neurogenesis in the olfactory bulb (A) and in the dentate gyrus of the hippocampus (B) based on the findings of Altman and Bayer and their coworkers in publications that appeared between 20 and 40 years ago (Altman, 1962, 1969; Altman and Das, 1965; Roselli-Austin and Altman, 1979; Bayer et al., 1982; Bayer, 1982, 1985c). Neural stem cells are represented as green circles in A and B; adult generated neurons are yellow-green ellipses in A and yellow-green ovals in B; neurons generated during the late-fetal, early postnatal period are magenta ellipses in A and magenta ovals in B. The large arrow in A indicates the direction taken by migrating secondary neural stem cells. The small arrows in A and B indicate the directions taken by migrating new adult neurons. The fact that olfactory sensory neurons are continually being renewed in the mammalian olfactory sensory epithelium has been extensively studied by Graziadei and Graziadei (1979).

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FIGURE 26 Summary diagram of the postulated changes in stem cell type and density from E15 (A) through E20 (F) in the germinal zones of the dorsomedial neocortex that produce motor cortex and somatosensory cortex. Primary neural stem cells (yellow and red ellipses) generate neurons in the neuroepithelium, secondary neural stem cells (orange ovals) generate neurons in the subventricular zone, glial stem cells (blue ovals) generate glia in both germinal zones, ependymal stem cells (violet squares) proliferate in the neuroepithelium, and ependymal cells completely replace the primary germinal matrix by E20 (F). The density changes and proportions of self-replicating and final neurogenetic divisions in both types of neural stem cells are based on timetables of neurogenesis in the motor cortex and the somatosensory cortex (Bayer and Altman, 1991a).

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ventricular zone. While in the neuroepithelium, SNS are presumed to be randomly distributed and go through the mitotic phase of the cell cycle at various distances from the ventricular lumen. Mitotic figures at some distance from the ventricular lumen are in the cortical neuroepithelium before the subventricular zone emerges, continually increase as the neuroepithelium declines, and are randomly distributed throughout the subventricular zone (Bayer and Altman, 1991a). One major contribution of comprehensive [3H]thymidine labeling is finding that throughout the central nervous system neurons in specific nuclei and laminae are generated over short time periods. The timetables of neurogenesis for layer VI–II cortical neurons (graphs 2–4, Fig. 4) show broader time spans because neurogenetic timetables are combined for all cortical areas, including the medial and lateral limbic cortices. But in the dorsomedial motor neocortex for example, peak neurogenesis of specific layers occurs on 2 days: 80% of layer VI neurons originate on E15 and E16, 78% of layer V neurons on E16 and E17, 88% of layer IV and lower layer III neurons on E17 and E18, and 77% of upper layer III and layer II neurons on E18 and E19 (see Fig. 14–4 in Bayer and Altman, 1991a). How can so many neurons be generated in such a short time? It is postulated that layer-specific neural stem cells take several days to prepare for a short burst of neurogenesis by continually renewing themselves at an exponential growth rate; all progeny remain in the cortical germinal zones and increase the number of layer-specific neural stem cells (self-replicating division). As neurogenesis begins, a few stem cell divisions produce either one or two postmitotic neurons (neurogenetic division). During peak neurogenesis, nearly all layer-specific neural stem cells produce only postmitotic neurons. Neurogenesis ends when the last layerspecific stem cells disappear from the neuroepithelium or the subventricular zone as they go through their final neurogenetic divisions to generate the youngest neurons in a given layer. The density and type of neural stem cells change hourly in the cortical germinal zones as neurogenesis takes place; some hypothetical changes are illustrated in Fig. 26 for the neuroepithelium and subventricular zone generating neurons in the dorsomedial neocortex. The proportions listed in Fig. 26 of neural stem cells in self-replicating and final neurogenetic divisions are based on the actual data of neuron origin using longsurvival [3H]thymidine autoradiography. Layer VI–V PNS predominate on E15 and reach peak density on E16; numbers decline on E17, and the last day any layer VI–V PNS exist in the neuroepithelium is E18 (red ellipses, Figs. 26A–26D). Stem cells generating neurons in layers IV–II have more complex changes because

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some are in the neuroepithelium (PNS, yellow ellipses, Fig. 26) and others are in the subventricular zone (SNS, orange ovals, Fig. 26). Another complication is that dorsomedial germinal zones in the neocortex are generating layer IV–II neurons not only for the motor cortex directly above but also for the ventrolaterally placed somatosensory cortex, where there are robust numbers of layer IV–II neurons. It is postulated that layer IV–II PNS generate a subset of older neurons in the superficial layers that have peak time of origin on E17 when the subventricular zone is still small (Fig. 26C); these stem cells have self-renewal exponential growth on E15 and E16, reach peak density on E17, decline sharply on E18, and generate their last neurons on E19. We also postulate that layer IV–II SNS (the Svet1 proliferating cells discovered by Tarabykin et al., 2001) generate a subset of younger neurons that has a peak time of origin on E19, when the subventricular zone is larger than the neuroepithelium (Fig. 26E); these stem cells show mainly self-duplicating division up to E18, reach peak density on E19 when most are either in neurogenetic division or in final neurogenetic division, and start to decline on E20. A few layer IV–II PNS generate the youngest neurons in layer II on E21 (not shown). It is possible that a very small fraction of layer IV–II PNS are retained in adults because layer IV neurogenesis can take place in the mature rat cortex (Kaplan, 1981).

Glial and Ependymal Stem Cells Glial and ependymal stem cells are different from neural stem cells because they are present in the developing brain and are still common in adult brains. The density of glial (blue ovals) and ependymal stem cells (violet squares) is lower than that of neural stem cells in cortical germinal zones up through E17 (Figs. 26A– 26C); by E19 and E20, they are the predominant stem cells (Figs. 26E and 26F). Ependymal stem cells always multiply in the neuroepithelium at the ventricular lumen. They are sparse on E15 and E16 and increase considerably on E17, but still do not form a continuous layer (Figs. 26A–26C). X-irradiation exposures result in dead cell debris shedding into the ventricular lumen. A continuous layer exists on E18 (Fig. 26D) because dead cell debris no longer sheds into the ventricular lumen after X-irradiation (see Chapter 10 in Bayer and Altman, 1991a). The ependymal layer continues to become more dense throughout the rest of development; neural stem cells are absent by E20, when the proliferating cells at the ventricle form the primitive ependyma (Figs. 26E and 26F). Glial stem cells multiply at random locations within the neuroepithelium on E15 and E16 (Figs. 26A and 26B). On E17 and E18 many glial stem cells move into

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the adjacent subventricular zone and they continue to proliferate in both germinal zones (Figs. 26C and 26D). Glial stem cells are mainly in the subventricular zone on E19 and are located outside of the primitive ependyma on E20 (Figs. 26E and 26F). The density of glial precursors continually increases in the subventricular zone; it is still large on E22 (Fig. 24) and populated with glial precursors because few to no neurons are being generated in the cortex (Bayer and Altman, 1991a). These stem cells generate both astrocytes and oligodendrocytes. The astrocyte precursors disperse out of the subventricular zone on E20 and E21 to locally multiply in the intermediate zone and cortical plate (Bayer and Altman, 1991a). Tekki-Kessaris et al. (2001) postulate that, similar to the spinal cord, the ventral forebrain has a focus of oligodendroglial stem cells in the neuroepithelium that eventually populate the dorsal forebrain, including the cortex. They show indirect evidence that oligodendroglial precursors migrate into the ventrolateral rat cortical subventricular zone from a source in the ventromedial neuroepithelium of the medial ganglionic eminence. The subventricular zones of the cortex and basal ganglia are continuous around the corticopallidal angle, and cells could migrate between telencephalic regions. Possibly, these pioneer oligodendroglial stem cells function to restrict uncommitted glial precursors to an oligodendroglial fate (Tekki-Kessaris et al., 2001).

References Altman, J. (1962). Are neurons formed in the brains of adult mammals? Science 135, 1127–1128. Altman, J. (1963). Autoradiographic study of cell proliferation in the brains of rats and cats. Anat. Rec. 145, 573–591. Altman, J. (1966). Autoradiographic and histological studies of postnatal neurogenesis. II. A longitudinal investigation of the kinetics, migration and transformation of cells incorporating tritiated thymidine in infant rats, with special reference to postnatal neurogenesis in some brain regions. J. Comp. Neurol. 128, 431–474. Altman, J. (1969). Autoradiographic and histological studies of postnatal neurogenesis. IV. Cell proliferation and migration in the anterior forebrain, with special reference to persisting neurogenesis in the olfactory bulb. J. Comp. Neurol. 137, 433–457. Altman, J., and Bayer, S. A. (1986). The development of the rat hypothalamus. Adv. Anat. Embryol Cell Biol. 100. Altman, J., and Bayer, S. A. (1990a). Mosaic organization of the hippocampal neuroepithelium and the multiple germinal sources of dentate granule cells. J. Comp. Neurol. 301, 325–342. Altman, J., and Bayer, S. A. (1990b). Prolonged sojourn of developing pyramidal cells in the intermediate zone of the hippocampus and their settling in the stratum pyramidale. J. Comp. Neurol. 301, 343–364. Altman, J., and Bayer, S. A. (1990c). Migration and distribution of two populations of hippocampal granule cell precursors during the perinatal and postnatal periods. J. Comp. Neurol. 301, 365–381. Altman, J., and Bayer, S. A. (1995). “Atlas of Prenatal Rat Brain Development.” CRC Press, Boca Raton, FL.

Altman, J., and Bayer, S. A. (1996). “Development of the Cerebellar System in Relation to Its Evolution, Structure, and Functions.” CRC Press, Boca Raton, FL. Altman, J., and Das, G. D. (1965). Autoradiographic and histologic evidence of postnatal neurogenesis in rats. J. Comp. Neurol. 124, 319–335. Anderson, S. A., Eisenstat, D. D., Shi, L., and Rubenstein, J. L. (1997). Interneuron migration from basal forebrain to neocortex: Dependence on Dlx genes. Science 278, 474–476. Anderson, S. A., Marin, O., Horn, C., Jennings, K., and Rubenstein, J. L. (2001). Distinct cortical migrations from the medial and lateral ganglionic eminences. Development 128, 353–363. Bayer, S. A. (1979a). The development of the septal region in the rat. I. Neurogenesis examined with [3H]thymidine autoradiography. J. Comp. Neurol. 183, 89–106. Bayer, S. A. (1979b). The development of the septal region in the rat. II. Morphogenesis in normal and X-irradiated embryos. J. Comp. Neurol. 183, 107–120. Bayer, S. A. (1980a). Development of the hippocampal region in the rat. I. Neurogenesis examined with [3H]thymidine autoradiography. J. Comp. Neurol. 190, 87–114. Bayer, S. A. (1980b). Development of the hippocampal region in the rat. II. Morphogenesis during embryonic and early postnatal life. J. Comp. Neurol. 190, 115–134. Bayer, S. A. (1980c). Quantitative [3H]thymidine radiographic analyses of neurogenesis in the rat amygdala. J. Comp. Neurol. 194, 845–875. Bayer, S. A. (1981). A correlated study of neurogenesis, morphogenesis and cytodifferentiation in the rat nucleus accumbens. In “The Neurobiology of the Nucleus Accumbens” (Chronister, R. B., and De France, J. F., Eds.), pp. 173–197. Haer Institute, Brunswick, ME. Bayer, S. A. (1982). Changes in the total number of dentate granule cells in juvenile and adult rats: A correlated volumetric and [3H]thymidine autoradiographic study. Exp. Brain Res. 46, 315–323. Bayer, S. A. (1983). [3H]Thymidine-radiographic studies of neurogenesis in the rat olfactory bulb. Exp. Brain Res. 50, 329–340. Bayer, S. A. (1984). Neurogenesis in the rat neostriatum. Int. J. Dev. Neurosci. 2, 163–175. Bayer, S. A. (1985a). Neurogenesis in the olfactory tubercle and islands of Calleja in the rat. Int. J. Dev. Neurosci. 3, 135–147. Bayer, S. A. (1985b). Neurogenesis of the magnocellular basal telencephalic nuclei in the rat. Int. J. Dev. Neurosci. 3, 229–243. Bayer, S. A. (1985c). Neuron production in the hippocampus and olfactory bulb of the adult rat brain: Addition or replacement? Ann. N. Y. Acad. Sci. 457, 163–172. Bayer, S. A. (1986a). Neurogenesis in the anterior olfactory nucleus and its associated transition areas in the rat brain. Int. J. Dev. Neurosci. 4, 225–249. Bayer, S. A. (1986b). Neurogenesis in the rat primary olfactory cortex. Int. J. Dev. Neurosci. 4, 251–271. Bayer, S. A. (1987). Neurogenetic and morphogenetic heterogeneity in the bed nucleus of the stria terminalis. J. Comp. Neurol. 265, 47–64. Bayer, S. A., and Altman, J. (1974). Hippocampal development in the rat: Cytogenesis and morphogenesis examined with autoradiography and low-level x-irradiation. J. Comp. Neurol. 158, 55–80. Bayer, S. A., and Altman, J. (1975a). Radiation-induced interference with postnatal hippocampal cytogenesis in rats and its long-term effects on the acquisition of neurons and glia. J. Comp. Neurol. 163, 1–20. Bayer, S. A., and Altman, J. (1975b). The effects of x-irradiation on the postnatally forming granule cell populations in the olfactory bulb, hippocampus, and cerebellum of the rat. Exp. Neurol. 48, 167–174.

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Bayer, S. A., and Altman, J. (1987). Directions in neurogenetic gradients and patterns of anatomical connections in the telencephalon. Prog. Neurobiol. 29, 57–106. Bayer, S. A., and Altman, J. (1991a). “Neocortical Development.” Raven Press, New York. Bayer, S. A., and Altman, J. (1991b). Development of the endopiriform nucleus and the claustrum in the rat brain. Neuroscience 45, 391–412. Bayer, S. A., and Altman, J. (1995). Neurogenesis and neuronal migration. In “The Rat Nervous System.” (Paxinos, G., Ed.), 2nd ed., pp. 1041–1077. Academic Press, San Diego. Bayer, S. A., Yackel, J. W., and Puri, P. S. (1982). Neurons in the rat dentate gyrus granular layer substantially increase during juvenile and adult life. Science 216, 890–892. Bulchand, S., Grove, E. A., Porter, F. D., and Tole, S. (2001). LIMhomeodomain gene Lhx2 regulates the formation of the cortical hem. Mech. Dev. 100, 165–175. De Carlos, J. A., López-Mascaraque, L., and Valverde, F. (1996). Dynamics of cell migration from the lateral ganglionic eminence in the rat. J. Neurosci. 16, 6146–6156. Denaxa, M., Chan,C.-H., Schachner, M., Parnevelas, J. G., and Karagogeos, D. (2001). The adhesion molecule TAG-1 mediates the migration of cortical interneurons from the ganglionic eminence along the corticofugal fiber system. Development 128, 4635–4644. Graziadei, P. P., and Graziadei, G. A. (1979). Neurogenesis and neuron regeneration in the olfactory system of mammals. I. Morphological aspects of differentiation and structural organization of the olfactory sensory neurons. J. Neurocytol. 8, 1–18. Grove, E. A., Williams, B. P., Li, D-Q.,Hajihosseini, M., Friedrich, A., and Price, J. (1993). Multiple restricted lineages in the embryonic rat cerebral cortex. Development 117, 553–561. Kalyani, A. J., and Rao, M. S. (1998). Cell lineage in the developing neural tube. Biochem. Cell Biol. 76, 1051–1068. Kaplan, M. S. (1981). Neurogenesis in the 3-month-old rat visual cortex. J. Comp. Neurol. 195, 323–338. Kaplan, M. S., and Bell, D. H. (1984). Mitotic neuroblasts in the 9-day-old and 11-month-old rodent hippocampus. J. Neurosci. 4, 1429–1441. Kaplan, M. S., and Hinds, J. W. (1977). Neurogenesis in the adult rat: Electron microscopic analysis of light radioautographs. Science 197, 1092–1094.

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Kaplan, M. S., McNelly, N. A., and Hinds, J. W. (1985). Population dynamics of adult-formed granule neurons of the rat olfactory bulb. J. Comp. Neurol. 239, 117–125. Krushel, L. A., Johnston, J. G., Fishell, G., Tibshirani, R., and van der Kooy, D. (1993). Spatially localized neuronal cell lineages in the developing mammalian forebrain. Neuroscience 53, 1035–1047. Langman, J., Guerrant, R. L., and Freeman, B. G. (1966). Behavior of neuroepithelial cells during closure of the neural tube. J. Comp. Neurol. 131, 15–26. McCarthy, M., Turnbull, D. H., Walsh, C. A., and Fishell, G. (2001). Telencephalic neural progenitors appear to be restricted to regional and glial fates before the onset of neurogenesis. J. Neurosci. 21, 6772–6781. Peretto, P., Merighi, A., Fasolo, A., and Bonfanti, L. (1999). The subependymal layer in rodents: A site of structural plasticity and cell migration in the adult mammalian brain. Brain Res. Bull. 49, 221–243. Roselli-Austin, L., and Altman, J. (1979). The postnatal development of the main olfactory bulb of the rat. J. Dev. Physiol. 1, 295–313. Schuurmans, C., and Guillemot, F. (2002). Molecular mechanisms underlying cell fate specification in the developing telencephalon. Curr. Opin. Neurobiol. 12, 26–34. Smart, I. H., and McSherry, G. M. (1982). Growth patterns in the lateral wall of the mouse telencephalon. II. Histological changes during and subsequent to the period of isocortical neuron production. J. Anat. 134, 415–442. Stoykova, A., Treichel, D., Hallonet, M., and Gruss, P. (2000). Pax6 modulates the dorsoventral patterning of the mammalian telencephalon. J. Neurosci. 20, 8042–8050. Tarabykin, V., Stoykova, A., Usman, N., and Gruss, P. (2001). Cortical upper layer neurons derive from the subventricular zone as indicated by Svet1 gene expression. Development 128, 1983–1993. Tekki-Kessaris, N., Woodruff, R., Hall, A. C., Gaffield, W., Kimura, S., Stiles, C. D., Rowitch, D. H., and Richardson, W. E. (2001). Hedgehog-dependent oligodendrocyte lineage specification in the telencephalon. Development 128, 2545–2554. Zappone, M. V., Galli, R., Catena, R., Meani, N., De Biasi, S., Mattei, E., Tiveron, C., Vescovi, A. L., Lovell-Badge, R., and Ottolenghi, S. (2000). Sox2 regulatory sequences direct expression of a β-geo transgene to telencephalic neural stem cells and precursors of the mouse embryo, revealing regionalization of gene expression in CNS stem cells. Development 127, 2367–2382.

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3 Autonomic Nervous System GIORGIO GABELLA Affiliation

body and the glands and the afferent (sensory) neurons that support the reflexes and the sensations from visceral organs. Many autonomic ganglia can be recognized with the naked eye as swelling or protrusions along nerve trunks or as knots within a mesh of nerves. Certain ganglia are connected in a sequence or ganglionated chain (the paravertebral sympathetic chain), while others are connected by a mesh of nerve trunks with which they form a plexus (the abdominal plexus or the pelvic plexus). Countless other microscopic ganglia are buried within a nerve or in the wall of viscera, and there are many single neurons along nerve trunks. In contrast. the sensory component consists mainly of neurons located in dorsal root ganglia (and in the nodose ganglion of the vagus nerve); in addition, afferent neurons are present within enteric ganglia. The autonomic nervous system is organized in groups of ganglia, which can be schematically subdivided into four groups, namely, paravertebral, prevertebral, paravisceral, and intramural (Fig. 1). Other ganglia are located in the head and provide motor innervation to salivary glands, cranial blood vessels, and the eye. The paravertebral ganglia are connected to each other and form two chains or ganglionated nerves, the sympathetic chains, which lie on either side of the vertebral column and are connected to the spinal nerves (hence to the spinal cord) by short nerve trunks, the “white” rami communicantes. The prevertebral ganglia are connected to each other and form a plexus (the abdominal plexus, which includes the celiac and the superior mesenteric ganglia) by the abdominal

LOCALIZATION OF AUTONOMIC GANGLIA General Organization The nerve cells and the nerves of the autonomic nervous system supply heart and blood vessels and intestinal, airway, urinary, and genital organs. The nerves regulate and coordinate bodily functions based on secretory activity of glands, on contraction and relaxation of smooth muscles and cardiac muscle, and on sensation arising from deep viscera. Muscles in the eye and skin and parts of the striated musculature of the esophagus and urethra are also innervated by autonomic nerves. While the central component of the autonomic nervous system consists of a few neuronal columns in the spinal cord and nuclei in the brain stem and the hypothalamus (see the relevant sections in other chapters), the peripheral part is scattered throughout the body. The peripheral autonomic nervous system of the rat, like that of other mammals, is of an extensive array of nerves and ganglia, connected to the CNS (spinal cord and brain stem) on one side and the viscera on the other. Viscera include the organs of the thoracic, abdominal, and pelvic cavities, the blood vessels, the organs of the head and neck, and the components of the skin. The autonomic nervous system has a common structural plan in all mammals, the rat being a species that has been studied extensively in this respect. The system comprises the efferent (motor) neurons that innervate the entire smooth musculature of the

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FIGURE 1 Highly schematic representation of the main groups of autonomic ganglia. The numbers indicate the topographical positions of the main autonomic nerves and of some individual ganglia: 1, superior cervical ganglion; 2, stellate ganglion; 3, lumber sympathetic ganglia; 4, intermediolateral column in thoracolumbar spinal cord; 5, rami communicantes; 6, thoracic splanchnic nerves; 7, lumbar splanchnic nerves; 8, mesenteric nerves; 9, hypogastric nerve; 10, perivascular nerves to blood vessels; 11, ciliary, otic, and sphenopalatine ganglia; 12, vagus nerve; 13, pelvic nerves; 14, cardiac ganglia; 15, pelvic ganglion; 16, prevertebral ganglia.

aorta. The paravisceral ganglia lie in the proximity of some viscera; the main groups are in the cardiac plexus and in the pelvic plexus, and other smaller ganglia are in a plexus close to the trachea and bronchi. Last, the intramural ganglia, which are too small to be seen with the naked eye, are situated within the wall of the gastrointestinal tract and biliary pathways.

Paravertebral and prevertebral ganglia are the main elements of the sympathetic outflow or the sympathetic pathway (referred to by many authors as the “sympathetic system”), which originates in the thoracic and lumbar segments of the spinal cord. The autonomic ganglia of the head are the cranial component of the parasympathetic pathway, which includes also the

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vagus nerve, all rooted in nuclei of the brain stem. The sacral component of the parasympathetic system originates from sacral segments of the spinal cord and consists chiefly of the main pelvic ganglion and a web of nerves known as the pelvic plexus. Tracheal, cardiac, and pelvic ganglia are situated along parasympathetic pathways; the pelvic ganglia, however, receive also a large input of sympathetic nerves. The intramural ganglia of the gut are neither sympathetic nor parasympathetic, although they are connected to both pathways; they constitute the enteric nervous system (Langley, 1921), a relatively autonomous component of the autonomic nervous system. The viscera have an abundant afferent (sensory) innervation, and if these fibers are regarded as part of the autonomic nervous system, as it seems they should be, they cannot be included in either the sympathetic or the parasympathetic pathway.

Sympathetic Chains The sympathetic chain is a bilaterally symmetric structure extending from the base of the skull to the sacrum. In the neck, the cervical sympathetic chain lies dorsal to the vagus nerve and the common carotid artery and ventral to the transverse processes of vertebrae and the prevertebral muscles. The chain has at this level two prominent ganglia, the superior cervical ganglion, and the inferior cervical ganglion (or stellate ganglion, which includes the uppermost thoracic sympathetic ganglia); a small intermediate cervical ganglion is sometimes found (Baljet and Drukker, 1979; Hedger and Webber, 1976) (Fig. 2). The superior cervical ganglion is spindle shaped, some 5 mm in length (often with a constriction in the middle), and lies dorsal to the bifurcation of the carotid artery. Among its (postganglionic) nerves, a carotid branch leaves the cranial pole of the ganglion and follows the internal carotid artery, and other smaller branches form a plexus around the external carotid artery. Other constant branches (rami) can be traced to the carotid body, to cranial nerves 9–12, and to cervical nerves 1–4 (Hedger and Webber, 1976). The stellate ganglion, consisting of the inferior cervical ganglion and the first two or three thoracic ganglia fused together, is located at the level of the first two thoracic vertebrae, on the right side being medial to the innominate artery (Hedger and Webber, 1976) (Fig. 2). Branches from the stellate ganglion join the lowermost cranial and the uppermost thoracic spinal nerves. Other branches connect with a plexus around the vertebral artery and with a plexus on the ventral side of the arch of the aorta (Hedger and Webber, 1976). In the thorax, the sympathetic chains lie ventral to the head of the ribs and dorsal to the parietal pleura

FIGURE 2 The cervical sympathetic trunk and its branches in the rat. A, aorta; B, brachiocephalic trunk; C, left common carotid artery; CN, carotid nerve; CT, costocervical trunk; C1, first cervical spinal nerve; I, internal carotid artery; LS, left subclavian artery; S, stellate ganglion; SC, superior cervical ganglion; ST, sympathetic trunk; V, vertebral artery; VN, vertebral nerve; E, external carotid artery; M, middle (intermediate) cervical ganglion; RS, right subclavian artery; T1, first thoracic spinal nerve. It should be noted that the cervical sympathetic trunk and superior cervical ganglion are located dorsal to the carotid artery (reproduced with permission from Hedger and Webber, 1976).

(Fig. 3). Each chain is made up of some 10 ganglia (including those usually fused with one another and with the inferior cervical ganglion). The lowermost ganglion lies opposite the 10th intercostal space. Small horizontal nerve trunks connect the two chains across the midline (Baljet and Drukker, 1979). In addition to the branches to the spinal nerves (rami communicantes) and small branches to the blood vessels, the thoracic sympathetic chains issue the splanchnic nerves (see below). In the abdomen the sympathetic chain is retroperitoneal and is embedded in the psoas muscle. There are five or six pairs of ganglia with rami communicantes to the spinal nerves, branches to blood vessels, and

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FIGURE 3 Lower cervical and thoracic sympathetic trunks of the rat (reproduced with permission from De Lemos and Pick, 1966).

small branches to the abdominal plexus (lumbar splanchnic nerves).

Rami Communicantes Rami communicantes are short nerve trunks connecting the ganglia of the sympathetic chain to the spinal nerves. The rami are particularly short in the rat

(De Lemos and Pick, 1966), and even when they are multiple and separated into two or more bundles they cannot be distinguished as white and gray rami. Preganglionic and postganglionic fibers are, therefore, mixed within each ramus, and because they have no distinctive structural features to indicate their orientation and nature, their identification can be made only after experiments of selective nerve sections.

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Splanchnic Nerves

Prevertebral Ganglia

Descending branches from the thoracic sympathetic ganglia give rise to the greater (major) splanchnic nerve, and occasionally to a lesser splanchnic nerve. Its apparent origin is usually from the 9th and the 10th ganglia (Baljet and Drukker, 1979). Both right and left greater splanchnic nerves enter the abdominal cavity, by piercing the diaphragm, between the medial and the lateral crus, and end in the abdominal plexus. The nerve, about 25 mm long, contains 190,000 nerve fibers, both afferent and efferent, the latter type pre- and postganglionic, and approximately 25,000 neurons (Isomura et al., 1985). There is often a paraaortic nerve, which originates in the lowermost thoracic sympathetic ganglia, enters the abdominal cavity, along the aorta, and terminates in the abdominal plexus. The lumbar splanchnic nerves are variable in number (usually four or five), size, and origin, and they extend from the lumbar sympathetic ganglia to the prevertebral plexus (Baljet and Drukker, 1979).

The prevertebral ganglia constitute the abdominal plexus, a large assembly of ganglia and nerve trunks lying close to the abdominal aorta and its main branches (Fig. 4). Blood vessels are the chief guide to the identification of prevertebral ganglia. Two sets of vessels stem from the abdominal aorta (Greene, 1935): parietal arteries (inferior phrenic arteries, lumbar arteries, ileolumbar arteries, middle caudal artery and terminal trunk, and common iliac artery) and visceral arteries (celiac artery, superior mesenteric artery, inferior mesenteric artery, renal arteries, and ovarian/ testicular arteries). The inferior mesenteric artery is often a branch of the right common iliac artery (Baljet and Drukker, 1979). Two major components can be distinguished in the abdominal plexus, the ciliac plexus [Paxinos et al., 1991 (Fig.4)], and the inferior mesenteric plexus, an intermesenteric plexus being interposed between them. The celiac plexus is situated around and between the

FIGURE 4 A left lateral representation (A) and a right lateral representation (B) of the celiac–superior mesenteric ganglion complex of the rat, as derived from serial sections. aa, abdominal aorta; ag, aorticorenal ganglion; ca, celiac artery; cg, celiac ganglion; cn, celiac nerve; ima, inferior mesenteric artery; ipa, inferior phrenic artery; ipv, inferior phrenic vein; isv, inferior suprarenal vein; k, kidney; msn, major splanchnic nerve; oa, ovarian artery; ra, renal artery; sg, suprarenal ganglion; sma, superior mesenteric artery; smg, superior mesenteric ganglion; srg, suprarenal gland; st, splanchnic trunk (reproduced by permission from Hammer and Santer, 1981).

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celiac artery and the superior mesenteric artery and extends dorsally between the adrenal glands and the cranial half of the kidneys. The left celiac ganglion is crescent shaped and lies on the lateral side of the celiac and superior mesenteric artery; the right celiac ganglion is triangular, smaller than the left one, and lies on the opposite side of the same arteries, dorsal to the inferior vena cava. Both celiac ganglia extend caudally, without distinct boundaries, into the superior mesenteric ganglia. A dorsal extension of the celiac ganglion forms the aorticorenal ganglion (Hammer and Santer, 1981) or ganglia, also more developed on the left than on the right side (Baljet and Drukker, 1979). Along the terminal part of the major splanchnic nerve, shortly before the celiac ganglion, there is a small ganglion called the suprarenal ganglion (Baljet and Drukker, 1979). Innumerable nerve trunks contribute to, and issue from, the celiac plexus. In addition to the thoracic and abdominal splanchnic nerves, there are nerve trunks (which include smaller or microscopic ganglia) within the plexus itself, including many nerves lying across the midline both ventral and dorsal to the aorta. Nerve trunks emerging from the celiac plexus and directed to abdominal organs reach the suprarenal arteries, the celiac artery, the superior mesenteric artery, the renal arteries, and the inferior phrenic arteries. Caudally, the celiac plexus continues into the intermesenteric plexus, an array of nerve trunks and very small ganglia lying on the ventral and lateral aspects of the aorta. The inferior mesenteric plexus of the two sides are extensively interconnected and lie around the initial segment of the inferior mesenteric artery. The inferior mesenteric ganglion, also called the hypogastric ganglion (Langworthy, 1965), is a spindle-shaped expansion along the main nerve trunk of the plexus. Caudally, the continuation of the inferior mesenteric ganglion is the hypogastric nerve, a bilaterally symmetric nerve that reaches the pelvic plexus. Nerve branches from the intermesenteric plexus can be followed to the kidneys, the ovaries, and the uterus or the testis. The main branches from the inferior mesenteric plexus, apart from the hypogastric nerve, are directed to the periphery along the inferior mesenteric artery.

Pelvic Plexus The pelvic plexus is a large and elaborate crossroads of nerves and ganglia supplying the rectum, the lower urinary tract, and the genital tract. The anatomy of this plexus, which is somewhat less complex in the rat than in other species, was investigated by Langworthy (1965), who used microdissection after vital staining with methylene blue and by Purinton et al. (1973) and Hulsebosch and Coggeshall (1982).

In the male rat, the main component of the plexus is a single, large, bilaterally symmetric ganglion, the right and left pelvic ganglia, sometimes referred as the hypogastric ganglion (Bentley, 1972; Sjöstrand, 1965) (Fig. 5). The ganglion is diamond-shaped, measuring about 2×4 mm, and lies on the side of the prostate, closely apposed to its fascia, ventral to the rectum, and caudal to the ureter and vas deferens. It shows lobulations that protrude in the direction of the surrounding organs. The ganglion is accompanied by a few, small, accessory ganglia, mainly related to the seminal vesicles and the vas deferens.

FIGURE 5 Pelvic ganglion stained in situ and in toto for acetylcholinesterase in an adult male rat. The right major pelvic ganglion (the dark triangular mass immediately left of center, with a perforation near its top which in vivo gives passage to a large artery for the bladder) is connected to the pelvic nerves (to the left), to the genital nerve (caudalward or downward, and to the left), to the hypogastric nerve (cranialward or upward), and to numerous nerves for the pelvic organs (to the right). Part of the bladder and its neck are visible near the right edge and in the right bottom corner. The right seminal vesicle, with a bulbous profile, appears vertically through the middle of the field; the left seminal vesicle is next to it and the ductus deferens (dark and cylindrical) is further to the right; all three are crossed by the right ureter (light, cylindrical, and widening near the point of entry into the bladder). To the far left is part of the rectum, displaying its myenteric plexus. Accessory ganglia are seen along the nerves to the ductus deferens and the urethra and along the hypogastric nerve.

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The main incoming nerve trunks to the ganglion are the hypogastric and the pelvic nerves. The hypogastric nerve, carrying the bulk of the sympathetic input, originates as the caudal continuation of the inferior mesenteric ganglion and reaches the cranial pole of the pelvic ganglion. It is retroperitoneal, medial to the external iliac artery, and passes behind the ureter, where it branches into the main and accessory hypogastric nerves. The terminal portion of the latter is slightly expanded as it contains the minute hypogastric ganglion (the hypogastric ganglion proper). The nerve contains about 1600 fibers, including sympathetic pre- and postganglionic fibers and sensory fibers (Hulsebosch and Coggeshall, 1982). The pelvic nerve, carrying the parasympathetic input to the pelvic ganglion, originates from the last lumbar (L6) and first sacral (S1) spinal nerves (Purinton et al., 1973). It consists of five to seven fascicles (Hulsebosch and Coggeshall, 1982), traveling mainly ventrally and reaching the dorsolateral aspect of the ganglion. The nerve contains about 5000 axons, mainly the preganglionic parasympathetic fibers for the pelvic neurons, but also afferent fibers and sympathetic postganglionic fibers (from paravertebral ganglion neurons). Numerous small efferent nerve trunks arise from the ganglion and reach the rectum, the ureter the vas deferens, the seminal vesicles, the prostate, the bladder, and the urethra. The largest trunk is the one that supplies the urethra and then proceeds to innervate the penis; this main penile nerve, or genital nerve, also contains hundreds of ganglion neurons (Dail et al., 1989). Eight or more small nerves are directed to the bladder; they divide into two groups, passing in front and behind the ureter and reaching the ventral and dorsal surface of the bladder. Minute ganglia can be seen along these nerves and those supplying the vas deferens and seminal vesicles. The largest of these accessory ganglia contains about 400 neurons, and the hypogastric ganglion has about 250 (Hondeau et al., 1995). In the female rat, the pelvic ganglion, also referred to as the paracervical or Frankenhauser ganglion (Kanerva, 1972; Marshall, 1970), is smaller and more difficult to expose than that in the male (Purinton et al., 1973). It is flat and lies firmly adherent on the side of the uterine cervix. Its largest branch, arising from the caudal portion of the ganglion, innervates the clitoris after giving branches to the urethra, vagina, and rectum. Other fine nerves issuing from the ganglion run to the bladder, cervix of the uterus, and upper portions of the vagina. A few small accessory ganglia can be found, usually on the ventral wall of the vagina near the bladder neck (Purinton et al., 1973). Extensive decussation of fibers on the midline occurs over the ventral surface of the cervix.

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In addition to topographical differences, the pelvic ganglion shows structural differences between male and female rats. The most prominent of them is that there are more than twice as many neurons in the male (about 14,000) than in the female (about 6000) (Greenwood et al., 1985), a difference in neuron numbers that becomes established postnatally. In the rat bladder intramural nerve ganglia are absent or very few in number.

Parasympathetic Nerves and Ganglia of the Head These ganglia are very small or microscopic, and they are associated with branches of certain cranial nerves. The ciliary ganglion lies lateral to the optic nerve and is attached to the initial part of the branch of the oculomotor nerve for the medial rectus muscle. Malmfors and Nilsson (1966) have described the exact position of the ganglion and the surgical approach for its excision [Paxinos et al., 1991 (Figs. 26–28)]. The ganglion is made up of about 200 ganglion neurons (Wigston, 1983). The optic ganglion of the rat [Paxinos et al., 1991 (Figs. 17, 43–45, and 103–106)] has no direct connections with the glossopharyngeal nerve, as is the case in man and other mammals (Al-Hadhithi and Mitchell, 1987), and the preganglionic fibers reach the ganglion via a connection with the facial nerve. The sphenopalatine ganglion is associated with the facial nerve [Paxinos et al., 1991 (Figs. 5, 6, 25–39, 75–80, and 82–87)]; its postganglionic nerves reach the lacrimal gland and the nasal mucosa. Otic and sphenopalatine ganglia are also major contributors of vasomotor fibers to cerebral vessels (Suzuki et al., 1988; Suzuki and Hardebo, 1991). The submandibular ganglion is a collection of minute ganglia located around the excretory ducts of the submandibular and the sublingual glands, in the connective tissue between these ducts and the lingual nerve, and within the submandibular gland itself (Ng et al., 1992). Two large aggregates of ganglion neurons, close to each other and located at the confluence of the internal carotid nerve and the greater superficial petrosal nerve, are known as the internal carotid ganglion (Mitchell, 1953; Suzuki et al., 1988). On the basis of the histochemical features of its neurons, the ganglion is considered an aberrant sympathetic ganglion or a rostral expansion of the superior cervical ganglion (Hardebo et al., 1992). The vagus nerve emerges from the cranial cavity through the jugular foramen. A prominent swelling of the nerve immediately after its emergence is known as the nodose ganglion [Paxinos et al., 1991 (Fig. 6, 59–61, 63, 67, 137–139, and 141)]; the ganglion issues two branches, the cranial, or pharyngeal, branch, forming a plexus with branches from the glossopharyngeal

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nerve, and the caudal branch, giving rise to the superior laryngeal nerve (Greene, 1935). There are about 6000 neurons in the nodose ganglion of the rat, and the full complement of ganglion neurons is already present at birth (Cooper, 1984). Neuronal mitoses in this ganglion are completed by the end of the second week in utero (Altman and Bayer, 1982). From the nodose ganglion, the trunk of the vagus nerve runs caudally into the neck. The main cervical branches are the cardiac branches and the recurrent nerve, which terminates as the inferior laryngeal nerve; from the thoracic vagus originate esophageal and pulmonary branches. The nerve reaches the abdominal cavity and terminates, spreading over the anterior (left vagus) and the posterior (right vagus) surface of the stomach. Details of the fine composition and distribution of the abdominal part of the vagis nerve are given by Prechtl and Powley (1990) and Berthoud et al. (1991, 1992).

STRUCTURE OF AUTONOMIC GANGLIA AND NERVES Preganglionic Neurons The parasympathetic preganglionic neurons are located in the spinal cord between the cervical 8 and the lumbar 2 levels (C8 and L2) [Paxinos and Watson, 1986 (Fig. 116, 118, and 119)]. The distribution and the morphology of preganglionic neurons have been studied to great advantage with the horseradish peroxidase (HRP) technique (retrograde filling). This has produced results that are more reliable than those obtained by degeneration methods and silver impregnation methods. The majority of the preganglionic neurons are in a column called the intermediolateral nucleus. A similar column in two or three sacral levels of the spinal cord contains the preganglionic neurons of the sacral parasympathetic outflow. The preganglionic neurons innervating the superior cervical ganglion of the rat are located in segments C8–T5 (90% of them in thoracic segments T1–T3). Seventy-five percent of the total of 1600 neurons retrogradely filled from the cervical sympathetic trunk of one side of the animal are in the intermediolateral nucleus, 23% are in the lateral funiculus, and the remainder is in the other central autonomic area and in the intercalated region (Rando et al., 1981). Injected neurons are exclusively homolateral to the injected cervical trunk (Rando et al., 1981). In contrast, Navaratnam and Lewis (1970) found chromatolytic neurons on both sides of the spinal cord after unilateral section of the pelvic nerves. An additional column of preganglionic neurons, termed the dorsal commissural nucleus, projecting into

the hypogastric nerve has been identified in the spinal cord levels L1-L2 (Hancock and Peveto, 1979). Schramm and collaborators (1975) reported labelling of preganglionic neurons at all the levels between T1 and L1 after injection of horseradish peroxidase into the adrenal medulla. Approximately 1000 neurons were labeled after injection into one gland (Schramm et al., 1976). They also reported that the neurons have a marked longitudinal orientation, as has been observed in other species, with the dendrites grouped into two bundles directed cranially and caudally. This high polarization of the dendrites arises rather late in development; it is absent in 3-week-old rats and is still far from complete in 7-week-old rats (Schramm et al., 1976). Preganglionic neurons in the rat spinal cord display an intense acetylcholinesterase activity [Navaratnam and Lewis, 1970; Paxinos and Watson, 1986 (Fig. 116)]. Nerve endings containing noradrenaline or serotonin are present around the cell bodies (Dahström and Fuxe, 1965).

Preganglionic Fibers Sympathetic preganglionic axons issue from neurons located mainly in the intermediolateral column of the thoracic and lumbar levels of the spinal cord. The fibers emerge from the cord within the ventral roots (T1–L2) bundled with the somatic motor fibers. From the ventral nerves the preganglionic fibers pass to the sympathetic chain via very short connections (rami communicantes). The latter also contain postganglionic fibers that travel to the periphery within somatic nerves. Depending on the level of origin, preganglionic fibers travel some distance up or down the sympathetic chain, forming synaptic contacts with neurons in more than one ganglion. In the lumbar segment of the chain, the preganglionic fibers are mainly descending (caudally directed). The length of the preganglionic fibers, therefore, can be considerable. In the upper thoracic segment they are mainly ascending (cranially directed) and in the cervical sympathetic trunk all the preganglionic fibers are directed cranially. In this trunk, however, there are also caudally directed postganglionic fibers, originating in the superior cervical ganglion, and cranially directed postganglionic fibers, originating in the middle and lower cervical ganglia (Bowers and Zigmond, 1981). Other sympathetic preganglionic fibers, having reached the paravertebral chain, pass into a splanchnic nerve and travel to prevertebral ganglia in the abdominal cavity and, in a smaller number, as far as the pelvic ganglion. In the rat, unlike other species, such as humans and cats, the great majority of preganglionic fibers are unmyelinated. For example, less than 1% of the axons

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in the cervical sympathetic trunk are myelinated (Brooks-Fournier and Coggeshall, 1981; Dyck and Hopkins, 1972; Hedger and Webber, 1976). The terminal branches of preganglionic fibers have varicosities and terminal boutons that synapse on ganglion neurons. In the ganglionic relay there is divergence; that is, each preganglionic fiber innervates several ganglion neurons. The numeric ratio between preganglionic and postganglionic neurons, as calculated by different investigators, varies from 1:5 to 1:20. The total number of neurons in the spinal cord that are retrogradely filled with horseradish peroxidase from the cervical sympathetic trunk is about 1600 (Rando et al., 1981), a value that should be compared with the number of neurons they innervate, that is, the neurons in the superior cervical ganglion (see Table 1). The divergence accounts for the fact that all ganglion neurons are innervated by preganglionic fibers, despite the smaller number of these fibers. In addition, each ganglion neuron receives synapses from more than one preganglionic neuron; that is, there is convergence of synaptic inputs onto each neuron. The latter process is ideally analyzed with electrophysiological techniques. Typical values obtained in rabbits (Wallis and North, 1978), hamsters (Lichtman and Purves, 1980), and guinea pigs (Njå and Purves, 1977) range between 7 and 11 preganglionic fibers per ganglion neuron; moreover, each ganglion neuron receives an input from several levels of the spinal cord (Njå and Purves, 1977). With the same technique, Purves et al. (1986) have obtained a figure of about 1000 preganglionic neurons for the superior cervical ganglion of the rat (against about 26,000 ganglion neurons), each preganglionic neuron innervating on average 240 ganglion neurons

TABLE 1

Number of Ganglion Neurons in the Superior Cervical Ganglion of the Rat

Mean number of neurons ± standard deviation (number of cases)

Study

45,000 ± 600

Davies, 1978

35,000 ± 600

Davies, 1978

39,000 ± 500 (4)

Klingman, 1972

37,000–38,000 (2)

Hedger and Webber, 1976

36,000

Ostberg et al., 1976

32,000 (1)

Levi-Montalcini and Booker, 1960

26,000–32,000 (6)

Eränkö and Soinila, 1981

25,000 ± 1940 (3)

Johnson et al., 1980

21,500 ± 3400 (3)

Santer, 1991

15,600 ± 6100 (17)

Brooks-Fournier and Coggeshall, 1981

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and each ganglion neuron being innervated by 9 preganglionic neurons, on average (Purves et al., 1986). Parasympathetic preganglionic fibers of the cervical region originate in nuclei of brain stem (nucleus of Edinger-Westphal, salivatory nuclei, dorsal vagal nucleus) and project onto the autonomic ganglia of the head. Sacral parasympathetic fibers originate from neurons in a short column at levels L6–S1 of the spinal cord (Tanaka and Zukeran, 1981) and project to the pelvic main and accessory ganglia. These preganglionic fibers are unmyelinated (Mallory et al., 1989), but their conduction velocity is faster than that of the corresponding unmyelinated postganglionic fibers. The degree of divergence and convergence in parasympathetic ganglia is considerably less than that in the sympathetic chain, as discussed below.

Sympathetic Ganglia The sympathetic ganglia are scattered along the sympathetic chain (paravertebral ganglia) and in the abdominal plexus (prevertebral ganglia). The best known of them is the superior cervical ganglion (Fig. 2). Because of its large size, its accessibility, its vascular supply, and the layout of its preganglionic and postganglionic nerves, it has been investigated more extensively than any other ganglion, the rat being one of the species of choice. Many of the structural features of the superior cervical ganglion are reproduced in the other sympathetic ganglia, although important differences are being found with more detailed studies, especially between prevertebral and paravertebral ganglia. In addition to ganglion nerve cells (principal ganglion neurons) (Fig. 6), sympathetic ganglia contain several other cell types. These include small granular cells (or small intensely fluorescent cells), cells of the glial type (Schwann cells and satellite cells), vascular cells (mainly endothelial cells), mast cells, and fibroblasts (in thin septa of connective tissue and in the capsule). The capsule, which is in continuation with the sheath of incoming and outgoing nerves, is relatively thick and offers a strong barrier to the diffusion of substances (Arvidson, 1979). However, substances (for example, horseradish peroxidase) injected systemically can diffuse into the ganglion from fenestrated capillaries (Jacobs, 1977) (see below). Injected tracers diffuse around the cells but do not penetrate the narrow space between a neuron and its satellite glial cells (Ten Tuscher et al., 1989). The satellite cells form a tight, continuous sheath around each ganglion neuron, blocking any diffusion of extracellular fluids, in contrast to the situation in sensory ganglia, in which satellite cells are loosely arranged and a tracer such as

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horseradish peroxidase freely reaches the cell membrane of the ganglion neurons (Ten Tuscher et al., 1989). Tight junctions are only occasionally found between satellite cells in rat sympathetic ganglia (Ten Tuscher et al., 1989), whereas there are numerous gap junctions [as Elfvin and Forsman (1978) have documented in the guinea pig]. By contrast, there is no extravasation of a systemically injected tracer in sympathetic nerve trunks, and it has been suggested that at this level (but not within ganglia) there is a blood– brain barrier to proteins (Jacobs, 1977). While the capillaries running between ganglion neurons are of the continuous type, have tight junctions, and are impermeable to injected tracers, those running close to the small intensely fluorescent cells are fenestrated and let tracers diffuse in the interstitial space of the ganglion (Chau and Lu, 1996). Most of the incoming fibers in paravertebral sympathetic ganglia, and virtually all the synaptic endings, originate from preganglionic neurons. Postganglionic fibers and fibers en passage, either pre- or postganglionic, are present within ganglia. Last, there are afferent fibers, probably en passage. For example, tracers injected in the superior cervical ganglion label a small number of dorsal root ganglion neurons in T1–T3 (Yamamoto et al., 1989).

Principal Ganglion Neurons The total population of sympathetic neurons in the rat probably numbers a few hundred thousand, but accurate counts are available only for the superior cervical ganglion (Table 1). The wide range in the values published by different authors is probably accounted for by a certain amount of experimental error (Hendry, 1976), but also by some variability between individual animals and possibly also between strains of rats. The variability in the number of neurons in the superior cervical ganglion in mammalian species in general is wide, ranging from about 4200 in a bat (Webber and Kallen, 1968) to nearly a million in a human (Ebbeson, 1968). The final number of neurons is established during fetal life, with few mitoses occurring in ganglion neurons of the rat after birth (Eränkö, 1972). From the first week after birth the number of principal cells remains unchanged (Davies, 1978) or shows a slight decrease (Eränkö and Soinila, 1981; Henry and Campbell, 1976). Brooks-Fournier and Coggeshall (1981) have obtained the lowest counts (15,000 on average, in 17 ganglia) and have also reported large differences in neuron number (up to 80%) between right and left ganglions. Santer (1991) has shown that there is no loss of neurons in the superior cervical ganglion of senescent rats (24 months old).

Principal ganglion neurons of the rat are multipolar neurons measuring up to 50 μm in diameter, mostly 25–40 μm (Tamarind and Quilliam, 1971). By comparison with other animal species, especially those of large body size, the dendritic arborization of sympathetic neurons in the rat is not extensive and the relative volume of neuropil is small (Fig. 6). There is little or no evidence of substantial structural differences within the population of ganglion neurons. Cajal (1911) suggested that the number and pattern of dendrites characterize neuronal ganglionic subpopulations; however, it has proven difficult to demonstrate, by silver methods, the dendrites in sympathetic ganglia, especially in the rat. Recently, a combination of electrophysiology and intracellular injection of HRP has allowed Kiraly et al. (1989) to see, in considerable detail, the dendritic trees in the superior cervical ganglion of the rat. On the basis of the dendritic patterns, neurons were classified as radiate (poorly branching dendrites arising all around the cell), tufted (extensively branching dendrites arising, clustered, from one area of the cell or from two opposite areas and running in opposite directions), and intermediate (Kiraly et al., 1989). The size and extension of dendrites in the superior cervical ganglion are correlated during development with the size of the animal (Voyvodic, 1987); experimentally induced changes in the size of the target (e.g., the submandibular gland) influence the size of dendritic trees (Voyvodic, 1989). There is marked expansion of the dendritic trees during postnatal development (Purves et al., 1986). In aged rats this process is reversed (at least in the superior cervical ganglion) and there is reduction in soma size, total dendritic length, number of branch points, and total area of dendritic arborization (Andrews et al., 1994). Ganglion neurons are individually ensheathed by satellite cells. This glial sheath is continuous over the neuronal soma; in some areas it is reduced to a very thin cytoplasmic process interposed between the neuron on one side and the basal lamina and connective tissue on the other. A glial sheath extends over the dendrites. Here, however, there are areas where the neuronal cell membrane is uncovered and lies directly apposed to the basal lamina and connective tissue (Fig. 7B). In these dendritic regions, there are large clusters of vesicles and, occasionally, membrane specializations similar to dense projections. Similar clusters of vesicles can also be found in the more superficial parts of the cytoplasm of the cell body and along the dendrites (Fig. 7A). Each neuron has an axon, which can be clearly recognized in silver-impregnated preparations; the axon travels within the ganglion, often along a tortuous path but without dividing or giving off branches. Under the electron microscope, the axon can be recognized only at some distance from the cell body, and

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FIGURE 6 Superior cervical ganglion of a rat. Araldite section stained with toluidine blue, showing ganglion neuron profiles (some nucleated) and the surrounding neuropil. The latter include satellite cells, neurites, blood vessels, and connective tissue (from Gabella, 1976). (800×)

its exact point of origin is therefore not usually seen. Dendrites from different cells are often in membraneto-membrane contact with each other (dendrodendritic contacts), or they form dendrosomatic contacts (Kiraly et al., 1989). These commonly occurring contacts, which do not have synaptic features, have been interpreted as evidence of intraganglionic connectivity, enabling the neurons to modulate or regulate each other’s activity (Kiraly et al., 1989). Only a limited topographic subdivision of ganglion neurons into groups is apparent in sympathetic ganglia. The question of the localization of these neurons in relation to the organ they innervate has been investigated through the retrograde reaction of axotomized neurons (Matthews and Raisman, 1972), the retrograde transport of nerve growth factor (Hendry et al., 1974) and HRP (Bowers and Zigmond, 1979), and the increased utilization of glucose in stimulated neurons (Yarowski et al., 1979). Neurons tend to be located in the part of the ganglion near the site of emergence of their (postganglionic) fibers (Matthews and Raisman, 1972), but this localization is ill defined, and in practice neurons projecting to a particular organ may be found in any part of the ganglion (Hendry et al., 1974). However, the neurons whose axons extend in the

internal carotid artery are located mainly in the cranial part of the superior cervical ganglion; those projecting in the external carotid nerve are mainly located in the caudal portion of the ganglion, where there are also neurons sending their axons in the cervical trunk (Bowers and Zigmond, 1979).

Small Intensely Fluorescent Cells Small intensely fluorescent (SIF) cells are identified by their small size, their distribution in small clusters, and their very intense formaldehyde-induced fluorescence (Eränkö and Harkonen, 1965; Norberg et al., 1966). Under the electron microscope their cytoplasm appears rich in large, dense-cored vesicles, hence the term small-granule-containing cells (Matthews and Raisman, 1972) (Fig. 8). These cells are strikingly heterogeneous in distribution, morphology, and chemical composition and are part of a large group of chromaffin and chromaffin-like cells (see review in Taxi, 1979), which includes cells of the paraganglia and the adrenal medulla and cells scattered in many tissues outside the nervous system. They all originate from the neural crest, probably from a common progenitor (Anderson, 1989). In the rat embryo, SIF cells appear

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FIGURE 7 Superior cervical ganglion of rat. (A) Electron micrograph of a ganglion fixed by immersion in glutaraldehyde. A preganglionic nerve ending (center) synapses on a cell process (dendrite) containing a mitochondrion, ribosomes, neurofilaments, and a large cluster of electronlucent vesicles. An ill-defined band of electron-dense material lies beneath the postsynaptic membrane (56,000×). (B) Electron micrograph of a dendrite with a large number of vesicles lying directly underneath its surface. The cell membrane has some dense projections attached to it and is here devoid of a satellite cell sheath (38,000×). (C) Fluorescence micrograph (by Dr. Lars Olson) (Falck–Hillarp method for catecholamines). The nerve cell bodies and some of their processes show specific fluorescence of varying intensity (270×).

later than principal ganglion neurons, and, therefore, at least in the superior cervical ganglion, they cannot be regarded as precursors of ganglion neurons (Hall and Landis, 1991). In vitro, however, SIF cells from the rat superior cervical ganglion develop long processes and convert into nerve cells, if the corticosteroids

necessary for their differentiation are withdrawn from the incubation medium and replaced with nerve growth factor (NGF) (Doupe et al., 1985). The number of SIF cells is variable even in the same ganglion. In the rat superior cervical ganglion, up to 1000 (Santer et al., 1975) or 370 (Williams et al., 1977)

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FIGURE 8 Rat pelvic ganglion. The electron micrograph shows a small granular cell (SIF cell) packed with dense-cored vesicles. The cytoplasm also displays mitochondria, microtubules, and the endoplasmic reticulum; at the right is part of the nucleus. The cell is surrounded by a continuous capsule of satellite cells (14,000×).

SIF cells have been counted. Eränkö and Soinila (1981) found an average of about 450 SIF cells per ganglion and observed a twofold increase in the number of SIF cells during the first 3 weeks of postnatal life. However, there seems to be a subsequent decrease in the number of SIF cells during development (Lempinen, 1964). The trend can be counteracted by glucocorticoid injection: treatment of rats with hydrocortisone at birth induces an increase of up to 10 times the number of SIF cells (Eränkö and Eränkö, 1972). The SIF cells of rat sympathetic ganglia measure 10–15 μm in diameter, and many of them are grouped into tight clusters, sheathed by satellite glial cells, and have extensive membrane-to-membrane contact between adjacent SIF cells. There are occasional discontinuities in the thin glial sheath and at these points the surface of a SIF cell lies bare and is usually directly opposite a fenestrated capillary. The cytoplasm contains a large number of dense-cored vesicles, rather variable in size, electron density, and shape; on the basis of these features of the vesicles, two or three types of SIF cells have been identified (Taxi, 1979): Type I SIF cells have granular vesicles measuring 80–100 nm (versus

40–50 nm for synaptic vesicles), whereas type II SIF cells have vesicles of 150–300 nm in diameter, similar to those of adrenal medullary cells. The SIF cells contain and probably release a biogenic amine; the type of amine varies from species to species and even between different ganglia of the same species. In the rat superior cervical ganglion the SIF cells, which are predominantly of type I, contain mainly dopamine (Björklund et al., 1970), but it has subsequently been reported that in this ganglion there are separate groups of SIF cells, containing dopamine or containing noradrenaline or serotonin (5-HT) (Konig, 1979; Verhofstad et al., 1981). Some SIF cells display two or more processes; a few are tens of micrometers in length, are ultrastructurally similar to the cell body, and have a varicose outline. Other cells, especially those in large clusters, have no processes. The SIF cells receive synapses from preganglioninc fibers, predominantly on the cell body. A few SIF cells (and notably some in the superior cervical ganglion) form specialized contacts with principal ganglion neurons, which are described as efferent synapses (Matthews and Raisman, 1969); these are usually somadendritic,

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and large granular vesicles are grouped around dense projections on the presynaptic membrane. Those SIF cells that have afferent and efferent synapses can be regarded as interneurons (Williams, 1967). Many, and possibly the majority of SIF cells, have no efferent synapses, and these, if not all SIF cells, are considered to be endowed with an endocrine role, releasing substances (amines) in the interstitial space or in the local circulation of a ganglion. The stimuli can be of central nervous system origin (via preganglionic fibers) or local (via chemoreceptors) A chemoreceptive role is suggested by the structural similarity of certain SIF cells to cells of the carotid body and the aortic glomus (Kondo, 1977). It is noteworthy that, according to Grillo (1978), about 10% of the synapses onto SIF cells in the superior cervical ganglion of the rat are not affected by preganglionic denervation; moreover, some efferent synapses of the SIF cells are not on ganglion neurons, but on large axons originating from the glossopharyngeal nerve, and are probably afferent.

Nerve endings, Synapses, and Other Cell Junctions There are abundant nerve endings synapsing onto principal neurons of sympathetic ganglia (Fig. 7A). The synapses are usually on dendrites or on dendritic spine-like processes and only rarely on the cell soma; in contrast, axosomatic synapses are common in ganglia of immature rats (Smolen and Raisman, 1980). The ultrastructural features of ganglionic synapses in rat sympathetic ganglia have been thoroughly investigated (see review by Matthews, 1983). Most of the intraganglionic endings are packed with small agranular vesicles (49–60 nm in diameter), in addition to a few mitochondria, endoplasmic reticulum, and microtubules; large granular vesicles, although representing no more than a small proportion of the vesicle population, are usually well in evidence. In the superior cervical ganglion, virtually all synaptic endings disappear after decentralization of the ganglion, and this confirms that they are of preganglionic origin. The synaptic cleft is rich in acetylcholinesterase (Somogyi and Chubb, 1976), acetylcholine is released (in quantal form) upon stimulation, and decentralization reduces acetylcholine levels by nearly 60% (Klingman and Klingman, 1969). Transmission across the ganglion is achieved by cholinergic synapses mainly operating through nicotinic receptors (see review by Skok, 1983). There are also synapses from preganglionic fibers onto SIF cells and occasionally from SIF cells onto a ganglion neuron. Numerous junctions of the adherens type (presumably of mechanical significance) occur between

neurons and satellite cells and between neuronal elements. Dendrodendritic contacts are numerous (Kiraly et al., 1989), but the suggestion of Kondo et al. (1980) that they are synaptic has not been confirmed.

Neurotransmitters and Related Substances The great majority of neurons in sympathetic ganglia are adrenergic (Fig. 7C). Catecholamines are stored in the cell body, in dendrites, in the axon, and, in a much higher concentration, in the varicosities of the terminal portion of the axon. The catecholamine content of the cell bodies, as detected histochemically by fluorescence microscopy, is variable from neuron to neuron and tends to decrease with age (Santer, 1979). Ultrastructurally, the biogenic amines are localized in large dense-cored vesicles, in small dense-cored vesicles clustered beneath the cell membrane, and in tubules of endoplasmic reticulum (Richards and Tranzer, 1975). A small percentage of sympathetic ganglion neurons (about 4% in the superior cervical ganglion; Yamauchi and Lever, 1971) are intensely positive for acetylcholinesterase and are negative for monoaminoxidase and catecholamines. These neurons, which also contain vasointestinal peptide (Landis and Fredieu, 1986), supply vasodilator cholinergic fibers to some blood vessels and secretomotor fibers to the eccrine sweat glands (Langley, 1922; Wechsler and Fisher, 1968). However, there is firm evidence that in the rat there is no cholinergic sympathetic innervation to the limb muscle blood vessels (Guidry and Landis, 2000). Sympathetic neurons (from the superior cervical ganglion) obtained from newborn rats and grown in vitro under certain conditions (which include the presence of nonneuronal cells; Patterson and Chung, 1977) undergo a transition from adrenergic to cholinergic (Furshpan et al., 1976, 1982; Johnson et al., 1976). A similar transition seems to occur in vivo: the ganglion neurons that innervate sweat glands are adrenergic in very young rats, and only adrenergic fibers are found around the developing glands. By the end of the third week of age, however, the same neurons have become cholinergic and only cholinergic fibers are found within the glands (Landis and Keefe, 1983). However, these nerve fibers, which are, both functionally and histochemically, cholinergic, maintain a limited ability to take up and store catecholamines, and some of them remain able to synthesize small amounts of catecholamines (Landis and Keefe, 1983). Neuropeptides are found within the rat sympathetic ganglia, although their amounts, as assessed by immunofluorescence, are lower than those in other species, for example, the guinea pig. A few single fibers immunoreactive for substance P are found in the stellate

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and the superior cervical ganglion (Hökfelt et al., 1977a), and a small number of fibers weakly immunoreactive for vasoactive intestinal polypeptide (VIP) occur in the superior cervical ganglion (Hökfelt et al., 1977b). There are no positive cell bodies for either peptide. There are also a few fibers immunoreactive for enkephalin and somatostatin (Hökfelt et al., 1977c); after colchicine treatment a positive reaction can be detected in some cell bodies. Many neurons in rat sympathetic ganglia (paravertebral, prevertebral, and pelvic) have intense NADPH-diaphorase activity (Santer and Symons, 1993): this enzyme is a cofactor of nitric oxide synthase (Hope et al., 1991), and these neurons are therefore capable of producing and presumably releasing nitric oxide, as suggested by pharmacological studies (Gillespie et al., 1989).

Prevertebral Ganglia The prevertebral ganglia are in many respects structurally similar to the ganglia of the sympathetic chain. An important difference, established mainly in the guinea pig (Crowcroft and Szurszewski, 1971), is that the ganglion neurons receive inputs not only from the spinal cord via the splanchnic nerves but also from neurons located in the wall of the gut and from neurons located in adjacent ganglia of the abdominal plexus. Several neuropeptides are localized in nerve fibers in prevertebral ganglia, including substance P, VIP, and enkephalin, although their occurrence is sparser in the rat than in the guinea pig (Schultzberg, 1983). Of particular interest is the localization of substance P fibers in some nerve endings abutting on ganglion neurons in the guinea pig. These are afferent fibers from dorsal root ganglion cells and they innervate the viscera; however, while in transit through the prevertebral ganglia, they issue collateral branches that synapse on ganglion neurons (Matthews and Cuello, 1982). This arrangement allows a reflex involving direct spread of stimuli (for example, nociceptive stimuli) from afferent axons to efferent neurons. Leranth and Ungvary (1980) have described the presence of several ultrastructural types of axons and have commented on the complexity of the synaptic connections in rat prevertebral ganglia.

Pelvic Ganglia The main components of the ganglion are the principal neurons, measuring 20–40 μm in diameter, sheathed by satellite cells (Fig. 9). Both cell types are similar in appearance to those found in the abdominal ganglia (Dail et al., 1975; Kanerva and Teräväinen, 1972). The neurons, however, visualized by intracellular injection of Lucifer yellow, have only one to four processes,

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one of which is the axon, and are therefore in this respect simpler than paravertebral and prevertebral ganglion neurons (Tabatabai et al., 1986). The dendrites are not only few in number but are also short, thin, and unbranched. Several neurons have no dendrites. The number of preganglionic inputs per neuron is on average only two, indicating that only two preganglionic fibers converge on each ganglion neuron. In addition there are ganglion neurons with large vacuoles (vacuolated neurons) (Fig. 9B). The vacuoles, measuring up to 20 μm in diameter, greatly enlarge the cell and displace the other components of the neuron, which is otherwise similar in structure and synaptic connections to the principal ganglion neurons. The significance of the vacuoles is obscure. The vacuolated neurons are about 0.8% of the neuronal population in the pelvic ganglion of pregnant rats and less than 0.2% in rats that are not pregnant (Lehmann and Stange, 1953). Vacuolated neurons are found also in the pelvic ganglion of male rats (Dail et al., 1975) in the range of 0.8–1.2% (Partanen et al., 1979); they make their appearance around the 7th week of life, but they disappear in castrated animals (Partanen et al., 1979). Small intensely fluorescent cells form a complete wrapping around ganglion neurons; the glial capsule of some neurons is formed by concentric layers of glial cell processes, surrounded by a basal lamina, and layers of collagen fibrils, so that a characteristic onion-like appearance that is not found in other ganglia is generated (Fig. 10). Small intensely fluorescent cells are also consistently found, in large numbers, usually in clusters. Under the electron microscope, they are recognized by their size and by the large dense-cored vesicles (Fig. 8). Most or all of them are of type II (Dail et al., 1975). Cholinergic and adrenergic neurons are both found in the pelvic ganglion. The adrenergic neurons (identified by formaldehyde-induced fluorescence) are about one-third of the neuronal population in the pelvic ganglion of the female rat (Kanerva et al., 1972) whereas they constitute the majority of neurons in the male (Dail et al., 1975). Cholinergic neurons (whose identification, based on an intense reaction for acetylcholinesterase, is less certain) are about one-fifth of all neurons. In the female rat, cholinergic neurons are among the largest in the ganglion (Kanerva, 1972); in the male they are small (15–25 μm in diameter) and are mainly found near the entrance of the pelvic nerve into the ganglion (Dail et al., 1975). A large proportion of the small neurons contain VIP; the axons of these neurons (VIP-ergic fibers) are plentiful in the smooth musculature of the penis, in the helicine arteries (Dail et al., 1983), and in the myometrium (Gu et al., 1984). It has been shown, in some autonomic neurons of the cat, that VIP neurons are often acetylcholinesterase positive (Lundberg, 1981).

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FIGURE 9 Pelvic ganglion of adult female rats (from Gabella et al., 1992). (A) The micrograph illustrates various types of cells: e, endothelial cell; f, fibroblast; g, nucleus of satellite glial cell; h, Schwann cell nuclei; m, mast cell; n, principal ganglion neurons; p, pericyte; and s, small granule cells or small intensely fluorescent (SIF) cells. (B) Vacuolated neuron. (C) Binucleate neuron. (D) The micrograph illustrates the laminar appearance of the glial capsule around some neurons (arrows). The scale bar represents 30 μm.

Most of the ganglion neurons (including the vacuolated cells) receive synapses, of the cholinergic type, from preganglionic fibers: unlike the situation in paravertebral ganglia, the nerve endings mainly abut the soma or somatic spines (Kanerva and Teräväinen, 1972a, 1972b). Some endings are found tunneling deep inside a perikaryon. Other synapsing nerve endings are not readily identified as cholinergic in that they contain a vast number of larger granular vesicles (Kanerva and Teräväinen, 1972a, 1972b). Some of the cholinergic neurons are surrounded by adrenergic terminals (as seen in fluorescence microscopy), which are regarded as collaterals of adrenergic neurons in the same ganglion (Dail et al., 1975). Adrenergic varicosities abutting the adrenergic neurons

have also been observed by means of fluorescence microscopy (Dail et al., 1975). The origin of these structures remains uncertain: they could be collaterals from other ganglion neurons, processes of SIF cells, or short dendritic processes from the same ganglion cell. Ultrastructural evidence of adrenergic endings synapsing on pelvic ganglion neurons has been found in the guinea pig (Watanabe, 1971); it is possible that the same situation occurs in the rat. Substance P-containing fibers form baskets around 10–20% of rat pelvic ganglion neurons. Most of the fibers disappear after section of the pelvic nerve and probably originate from dorsal root ganglia; the rest originate from SIF cells, which also stain for substance P (Dail and Dziurzynsky, 1985). Several peptides are found in pelvic ganglion

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FIGURE 10 Pelvic ganglion of an adult female rat. The electron micrograph shows a neuron, with nucleus and nucleolus. Its glial capsule is made of several layers of satellite processes and collagen (6400×).

neurons (neuropeptide Y and VIP), in nerve terminals (somatostatin, substance P, and enkephalin), or in both (Keast, 1991). There is good physiological evidence of reflex activity mediated through the pelvic ganglion in rats of either sex, without relay in the dorsal root ganglia (Purinton et al., 1971). This has led to the suggestion that there are peripheral afferent neurons (“sensory perikarya”), and selective denervation experiments have shown that these neurons are located distal to the pelvic and the hypogastric nerves and are probably situated in the pelvic ganglion itself (Purinton et al., 1971). The pelvic ganglion has two separate preganglionic inputs: sympathetic fibers originating from the lumbar levels of the spinal cord (and reaching the ganglion via rami communicantes, lumbar splanchnic nerves, and the hypogastric nerve) and parasympathetic cholinergic fibers originating in the spinal cord at L6 and S1 levels (and reaching the ganglion via the pelvic nerve). This traditional notion is confirmed by the electron microscopy study of Hulsebosch and Coggeshall (1982),

which, however, has shown an unexpected complexity in the nerve pathways connected to the pelvic ganglion. Thus, of the 1600 axons in the hypogastric nerve 58% are sympathetic postganglionic, 34% are sympathetic preganglionic, and 8% are sensory. Of the nearly 5000 axons of the pelvic nerve 34% are sensory and 49% are parasympathetic preganglionic; the remaining 17% are sympathetic postganglionic axons. Sympathetic fibers (preganglionic and postganglionic) are present also in the pudendal nerve, a nerve that is mainly a somatic sensory nerve (Hulsebosch and Coggeshall, 1982). Only 12% of all the preganglionic fibers to the pelvic ganglion are myelinated. A small proportion of postganglionic fibers are also myelinated. The two types of preganglionic fibers do not mix or converge, but they instead project onto separate neurons (Tabatabai et al., 1986). This property, together with the paucity of dendrites (Tabatabai et al., 1986), suggests that the integrative role of the pelvic ganglion is markedly smaller than that in other autonomic ganglia, for example, the superior cervical ganglion (Brown and McAfee, 1981).

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The cells of origin of the preganglionic fibers, identified with HRP after retrograde transport from the hypogastric nerve, are localized in the spinal segments L1 and L2, but not in the intermediolateral nucleus. Most of the neurons form a column along the midline in the dorsal gray commissure [dorsal commisural nucleus (DNC); Hancock and Peveto (1979)].

Cardiac and Tracheal Ganglia The cardiac plexus is made by many minute ganglia located beneath the epicardium and at the base of the aorta and pulmonary artery [King and Coakley, 1958; Paxinos et al., 1991 (Fig. 2), 1994 (Fig. 8); Pauza et al., 2002]. There is no bilateral symmetry, of course, and the total number of neurons is about 4000 (Pardini et al., 1987). About half the neurons are located near the superior border of the interatrial septum; other groups lie posterior to the right atrium (23.5%), and between superior vena cava and aorta (2.5%) (Pardini et al., 1987). The neuronal population consists of cholinergic neurons; these also contain various peptides and some of them display aspects of the adrenergic phenotype, such as immunoreactivity for dopamine β-hydroxylase. There are also a few SIF cells, some adrenergic fibers in transit, probably originating from the stellate ganglion and sensory fibers. In the rat the adrenergic fibers are only in transit and do not appear to form pericellular synaptic nests around ganglion neurons. The ultrastructure of the rat cardiac ganglia, in many respects similar to that of other autonomic ganglia, is described by Ellison and Hibbs (1976). The incoming synapses are mainly axosomatic and most of the synapsing nerve endings appear to be cholinergic; other endings contain mainly flat and lucent vesicles, whereas axons with dense-cored vesicles do not make contacts with ganglion neurons in this species. On the dorsal aspect of the trachea, overlying the tracheal muscle, a few scores of small nerve ganglia and connecting strands constitute the tracheal plexus, linked mainly with nerve trunks from the vagus nerve and made of postganglionic parasympathetic neurons. A few ganglia of this plexus are found near the primary bronchi and nerves extend to reach glands and smooth musculature in the lung.

Parasympathetic Ganglia of the Head Three quarters of the 250 neurons of the rat submandibular ganglion are each one innervated by a single preganglionic fiber, the remaining ones being innervated by two or three fibers (Lichtman, 1977). The arrangement whereby most ganglion neurons are driven by a single preganglionic neuron arises during

postnatal development. At birth most ganglion neurons are innervated by four to six preganglionic fibers. During the first 6 to 7 weeks of postnatal life, there is a progressive reduction in the number of preganglionic fibers converging on each neuron until the majority of neurons have a single input. At the same time, however, the total number of synaptic endings on each neuron increases (Lichtman, 1977). In the adult rat the neurons are usually devoid of large dendrites, but they have numerous minute cytoplasmic projections from the cell body and from the initial portion of the axon. Synaptic boutons are mainly associated with these projections. In preparations stained with zinc-iodide osmium, an average of 44 boutons per neuron was counted (Lichtman, 1977). Two distinct populations of neurons are recognized electrophysiologically (Kawa and Roper, 1984): the neurons innervating the submandibular gland and those innervating the sublingual gland. About one-third of the former (and none of the latter) are electrically coupled (however, gap junctions have not been found; Lichtman, 1977). After decentralization, intrinsic synapses (that is, synapses arising from other ganglion neurons) are found in 72% of the submandibular neurons and in only 12% of the sublingual neurons (Kawa and Roper, 1984). The extent to which interneuronal connections among submandibular neurons are present in the absence of decentralization remains to be established. In the submandibular ganglion of another species, the mouse, all synapses disappear after decentralization, a clear sign of the absence of interneurons or interneuronal synaptic connections (Yamakado and Yohro, 1977). The general ultrastructure of the submandibular ganglion of the rat is described by Ng et al. (1992) Of the 200 or so ganglion neurons in the rat ciliary ganglion, some are without dendrites, some have few long dendrites, and some have several long dendrites (Wigston, 1983). Each neuron is innervated by one to four preganglionic axons (on average 2.2), which are cholinergic and form synapses on perikarya and dendrites (Wigston, 1983).

Intramural Ganglia of the Gut A myriad of small intramural ganglia, joined by connecting strands, are gathered into two ganglionated plexuses, the myenteric and the submucosal plexus. The myenteric plexus is intramuscular, being located between the circular and longitudinal muscle layers; it extends without interruption from the esophagus (including the portion where the musculature is striated rather than smooth), through the stomach and small and large intestine, to the anal canal. In the stomach, myenteric ganglia are larger and more numerous near

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the lesser curvature. In the small intestine of the rat (unlike other mammalian species) myenteric neurons are distributed in cords parallel to the circular musculature, rather than discrete ganglia. By contrast, the myenteric ganglia of the large intestine are larger and have a better defined outline. Myenteric ganglia are found throughout the full length of the anal canal. In the small intestine there are about 9400 myenteric neurons per square centimeter of serosal surface. The submucosal plexus is found in the submucosa of the small and large intestines, usually close to the inner aspect of the circular muscle layer. Its neurons are about half as numerous as those in the myenteric plexus, and they are also, on average, smaller in size. Submucosal neurons are not found in the stomach. The intrinsic neurons of the gut form a complex and varied population (Fig. 11). Several types of neurons have been distinguished on the basis of the number of cell processes (as visualized by intracellular injection or by methylene blue staining, affinity for silver salts, and cell size (see a review in Gabella, 1979). Most of the more recent studies have been carried out on the guinea pig: fewer data are available for the rat. For the guinea pig myenteric plexus several investigators have put forward classifications based on ultrastructural features (Cook and Burnstock, 1976), on distribution of

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neuropeptides (Furness and Costa, 1980), and on electrophysiological properties (Wood, 1981). The myenteric ganglia (but not the submucosal ones) also contain neurons, well documented in the rat, projecting centripetally to the celiac and superior mesenteric ganglia (Furness et al., 2000). In the rat the terminal portion of the rectum contains a substantial number of neurons projecting to the spinal cord via the dorsal roots (Dörffler-Melly and Neuhuber, 1988). The enteric ganglia have a compact structure with tightly packed cells and cell processes (Fig. 12). The shape and thickness of myenteric ganglia change greatly with the contraction of the adjacent muscle layers. Collagen fibrils, fibroblasts, interstitial cells, and capillaries do not penetrate the ganglia but lie around them without forming a proper capsule. A single basal lamina is spread over the surface of the whole ganglion. Tracers injected intravenously in high concentrations diffuse from perigangliar capillaries and penetrate the interstices of the enteric ganglia of the rat (Jacobs, 1977). The cell types found within the ganglia are neurons and glial cells, the latter outnumbering the former by about three to one. The processes of glial cells and the neuronal processes (partly of intrinsic and partly of extrinsic origin) constitute the neuropil. The ganglion neurons, when examined under the electron microscope,

FIGURE 11 Whole-mount preparation of the muscle coat of the rat cecum, showing some neurons of the myenteric plexus. The faint staining in the background is due to the muscle cells of the circular layer (160×).

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FIGURE 12 Electron micrograph (montage) of a ganglion of the myenteric plexus of the rat small intestine. To the right is the longitudinal musculature; to the left is the circular musculature (in transverse section). The ganglion displays a neuronal cell body of complex shape, with its nucleus, and a large number of neuronal and glial processes (13,000×).

have a complex and irregular shape. Characteristically, parts of the neuronal perikarya reach the surface of the ganglion, and their membrane is in direct contact with the basal lamina and the connective tissue surrounding

the ganglion. Axosomatic and axodendritic synapses are numerous. The great majority of them survive an extrinsic denervation of the gut and are therefore of intrinsic origin. Tentative classifications of the vesicle-containing

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nerve endings of the enteric ganglia have been proposed for species other than the rat. The distribution of neuropeptides in the enteric ganglia of the rat has been studied in detail by Schultzberg and collaborators (1980). More than 50% of the neurons in the submucosal plexus of the ileum display immunofluorescence for VIP, about 20% for substance P, and 19% for somatostatin neurons (Schultzberg et al., 1980). There are no adrenergic ganglion neurons. Adrenergic fibers are seen within the muscle layers, around the intramural blood vessels, and within the ganglia (Van Driel and Drukker, 1973). The enteric glial cells pervade all the spaces within the ganglia, lying over parts of the surface of neurons and between nerve processes. Glial processes reach the surface of the ganglia and are the more conspicuous features of these glial cells. The gliofilaments (intermediate filaments) are inserted in dense plaques anchored to the cell membrane at the surface of the ganglion (Gabella, 1981). They are immunologically identical to the gliofilaments found in astrocytes (Jessen and Mirsky, 1980). There are many specialized contacts between vesicle-containing nerve endings and enteric glial cells (Gabella, 1981).

Neuromuscular Junctions Most of the axons issued by ganglion neurons, the so-called postganglionic fibers, terminate on muscle cells or on gland cells. The terminal portion of each branch of a fiber has a beaded structure, made of expansions, or varicosities, and intervaricose segments. The branching of preterminal autonomic axons and the presence of varicosities allow an extremely large number of endings (the term is used here to include varicosities) to be deployed by each axon. The number of varicosities per neuron can be two orders of magnitude larger than the number of endings made by a somatic motoneuron. Because of the sequential distribution of varicosities, each varicosity is both a point of transmission of an action potential along the axon and a potential point of transmitter release. In some smooth muscles, notably those of the intestine, axons are grouped in progressively smaller bundles and they tend to remain grouped together even in their terminal or varicose portions. Other muscles, such as those of the iris, bladder, and ductus deferens, in contrast, contain arborizations of nerve bundles that lead to individual axons with little or no Schwann cell wrapping in their terminal portions. Varicosities are very variable in length, diameter, and spatial frequency even within the same fiber. The separation between nerve varicosity and muscle cell, as well as the extent of the glial wrapping, is also

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variable. Varicosites are usually packed with axonal vesicles, but they also contain mitochondria, microtubules, and a few elements of smooth endoplasmic reticulum. Axonal vesicles may be clustered against the membrane of the varicosity. Structural specializations that characterize interneuronal synapses or neuromuscular junctions of skeletal muscles are absent or very faint at the nerve endings in smooth and cardiac muscles (Canale et al., 1986). As regards the innervation of blood vessels, detailed studies have shown that sympathetic nerve endings have structural relations with muscle cells that range from close contacts, with less than 20 nm of separation and fusion of the two basal laminae, to loose contacts, with a separation of 100 nm or more; the range is continuous, although at least two types of varicosities are distinguished (Luff and McLachlan, 1989). In small arterioles the majority of the endings have features indicating that they form neuromuscular junctions (Luff et al., 1991). In visceral muscles, various patterns of termination of the efferent fibers are observed. It may be useful to present two different arrangements of the intramuscular termination of autonomic motor nerves—as exemplified in the intestine and the bladder, although it is not clear whether they represent two exclusive patterns, or whether they are two patterns over a continuum of varying structural arrangements. In the intestine the intramuscular terminal nerves form a plexus rather than a tree-like terminal distribution. Isolated axons are rare and short. Most of the axons, including the majority of varicosities, are found within nerve bundles, and the varicose portion of an axon is usually located near the surface of the bundle, beneath its basal lamina. All the axons are tightly packed together with extensive membrane-to-membrane contact, even in large nerve trunks, and the Schwann cell component of the nerves is relatively small. The vesicle-containing varicosities thus face the extracellular space at large: the nearest cellular structures, apart from other axons and Schwann cells, are muscle cells and interstitial cells of Cajal. There is anatomical evidence of juxtaposition of vesicle-containing varicosities and muscle cells, strongly suggesting sites of transmission from nerve to muscle. Whether these juxtapositions represent true neuromuscular junctions is a mute point, because the structural configurations are very variable and often indistinct and rarely intimate. The possibility of nerve transmission to interstitial cells is much discussed (Sanders, 1996). In contrast, the intramuscular nerves of the bladder have a tree-like terminal pattern and the branching of the nerve bundles continues until each axon runs singly between muscle cells (Fig. 13). The varicosities are large and become progressively larger along the

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FIGURE 13 Muscle layer of the rat bladder. The electron micrograph shows three smooth muscle cells, approximately in transverse section. Between them is a nerve ending (varicosity) packed with axonal vesicles. The neuromuscular gap is approximately 40 nm and is occupied by an amorphous material that probably corresponds to two fused basal laminae. The ending is completely bare; that is, there is no Schwann cell sheath (35,000×)

terminal segment of the axon; they are variable in size but markedly larger than the varicosities of the intramuscular nerves of the gut. The intramuscular axons are individually wrapped by a Schwann cell (unlike those of intestinal nerves) and make no contact with each other, both in the nerves proper (Fig. 14) and in the nerve trunks that penetrate the wall of the organ; early varicosities have part of their surface uncovered by Schwann cell and abutting on the basal lamina (an arrangement refered to as a “window”); further down the axon the varicosities, fully loaded with vesicles, lose progressively their Schwann cell sheath and the last two or three of them are devoid of Schwann cell sheath, the axon extending some micrometers further than the Schwann cell (Fig. 15). The structure of terminal varicosities and the individual point of termination of an axon can be identified by serial sections (Gabella, 1995) (Fig. 16). The varicosities are packed with vesicles and lie mostly close to the surface of a muscle cell; sometimes they abut on two or three muscle cells, or lie in a groove of a muscle cell. The distance between the two membranes is often reduced to 30–50 nm. It seems correct in these situations to talk of autonomic neuromuscular junctions and to assume that these are

the discrete points of transmission from nerves to muscle. Although the structural configurations remain very variable, these neuromuscular junctions are characterized by the axon expanding into a varicosity and losing (completely or over a window) its Schwann cell cover, the separation from the muscle cells being reduced to a few tens of nanometers (the gap, or junctional cleft, being occupied or not by a basal lamina, but not by collagen or other structures extracellular materials) (Fig. 17). Subjunctional specializations are not observed on the muscle cell membranes lying beneath autonomic varicosities, even when the latter form identified neuromusuclar junctions; there are no postjunctional membrane densities or folds or specific subjunctional structures. However, clustering of postjunctional ATPreceptors has been documented in the muscle cells of the rat detrusor; the receptors are gathered into patches, about 1 μm across, that lie beneath the nerve varicosities (Hansen et al., 1998). The vesicles packing the varicosities in the bladder detrusor muscle are of uniform type although both acetylcholine and ATP are released from these nerve terminals. The high density of ATP-receptors (P2X-receptors) on muscle cells of the

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FIGURE 14 Electron micrograph of an intramural nerve of a rat bladder, showing several axon–glial bundles with unmyelinated axons. Nuclei of Schwann cells are visible. Many collagen fibrils, here mainly in transverse section, occupy the spaces between the axon–glial bundles. The axons display mitochondria, neurofilaments, some microtubules, and, occasionally, a cluster of small lucent vesicles.

bladder detrusor and the intensity of the atropineresistant excitatory input are characteristic of the rat.

Sensory fibers The sensory component of the autonomic nervous system consists mainly of neurons in dorsal root ganglia. Their central projections to the dorsal horn of the

spinal cord intermingle in part with somatic afferent projections, thus providing the anatomical basis of referred pain. The central projection synapses on interneurons that project onto preganglionic autonomic neurons, providing the basis for polysynaptic reflexes, or directly onto the preganglionic neurons (monosynaptic reflexes). The peripheral projections of these sensory neurons reach the adventitia of blood vessels

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FIGURE 15 Electron micrographs from serial sections of a rat detrusor muscle showing an axon coursing between muscle cells, all in transverse section. The axon is devoid of Schwann cells over the entire length studied (10 μm). The varicosity at the start of the series contracts into an intervaricose segment of 0.15 μm in diameter, then expands into a varicosity of about 0.7 μm in diameter, then contracts again into an intervaricose segment, and then expands again into a varicosity. The intervaricose segment in (xv) measures 0.05 μm and is occupied by a single microtubule. There are four microtubules in (i–v), six in (vi–xi), two in (xii), and one in the remaining ones. The roughly triangular space outlined by the three muscle cells and occupied by the axon is larger at the level of the varicosity, e.g., in (vi), than at the level of the intervaricose portion (xiv–xviii) (from Gabella, 1995).

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FIGURE 16 Electron micrographs from serial sections of a rat detrusor muscle showing an axon coursing between muscle cells, all in transverse section. In this series, an axon, accompanied by a slender Schwann cell process, terminating in (vi), expands into a varicosity, which is the terminal one since the axon ends in (x). This terminal varicosity is not lying particularly close to the surrounding muscle cells (from Gabella, 1995).

and the wall of most viscera. These afferent fibers reach the viscera traveling within splanchnic nerves and in postganglionic nerves; afferent and efferent fibers are thoroughly mixed within these nerves, and without seeing their origin or the terminal portion they cannot be distinguished anatomically from one another (Fig. 14). Afferent fibers for blood vessels in the body wall and limbs travel initially within somatic nerves.

The density of distribution of sensory endings is very high in certain structures, for example, the mucosa of the bladder, while it is sparse in others, for example, the intestinal mucosa. It has been calculated that the rat bladder is innervated by about 16,000 ganglion neurons approximately half of which are efferent (motor) and half sensory (Gabella, 1999). The terminal portions of the sensory fibers in the bladder are found in the mucosa,

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FIGURE 17 Electron micrographs from serial sections of a rat detrusor muscle showing an axon coursing between muscle cells, all in transverse section. This series illustrates the formation of an autonomic neuromuscular junction in the bladder musculature The 18 micrographs are taken from a set of serial sections covering a thickness of 10 μm. In the top left micrograph an intervaricose portion of the axon is fully wrapped by a Schwann cell process. The Schwann cell wrapping then retracts and a “window” appears where the axonal membrane is in contact with the basal lamina. The axon grows, increasing over 35-fold in the cross-sectional area, the bundle of microtubules is displaced to one side, and many axonal vesicles appear and pack the varicosity. The “window” expands and the distance between axolemma and smooth muscle cell membrane is reduced to about 20 nm over a wide area. The postjunctional membrane shows no typical structural specializations (from Gabella, 1995).

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mostly lying very close to the deep aspect of the urothelium. These fibers (readily identified by histochemical methods because of their high content of the neuropeptides substance P and, especially in the rat, CGRP) form a tight mesh which is particularly well developed in the caudal part of the bladder near the neck (Figure 18, top) and is sparse in the equatorial region and almost absent in the cranial region of the bladder. The last few hundreds of micrometers of a sensory fiber are distinctly varicose; these varicosities are smaller in size than those of efferent axons, they contain some vesicles but not as densely packed as in efferent axons, and they do not appear to be associated with specialized cellular structures or special components of the extracellular materials. Some fibers extend into the epithelium reaching very close to the luminal surface (Fig. 19, bottom). The mechanism of transduction at these sensory endings (which are regarded as “free” endings in that they are not associated with a corpuscular receptor) is not clear. Release of neurochemicals from the afferent nerve ending itself is probably part of the process. Recent evidence from the bladder (of mice and rabbits) shows that ATP is produced by the epithelial cells and is released upon mechanical stimulation Ferguson et al., 1997), and it can then act on the P2X-receptors present in subepithelial afferent nerve fibers (Evans and Surprenant, 1996) and trigger an afferent impulse (Cockayne et al., 2000). The release of neurochemical from afferent nerve endings (which can occur upon chemical or mechanical stimulation or by antidromic nerve impulses) is a crucial mechanism in the process of neurogenic inflammation which is known to occur, in rodents, in some viscera, including bladder and trachea.

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FIGURE 18 Whole-mount preparation of the bladder mucosa stained with CGRP antibody. A dense plexus of immunoreactive fibers lies parallel to the lumenal surface and very close to the epithelium. These fibers originate from dorsal root ganglia and provide the main sensory innervation to the bladder (from Gabella and Davis, 1998).

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FIGURE 19 (Top) Whole-mount preparation of the bladder mucosa of a rat, stained for CGRP, showing terminal and branching points of sensory axons, sequence and range of varicosities, and an axonal loop from a single axon, all spreading in a very flat plane immediately below the epithelium. (Bottom) Varicose terminal portion of sensory fiber, stained for CGRP, situated within the epithelium of the rat bladder mucosa (from Gabella and Davis, 1998, modified).

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C H A P T E R

4 Primary Afferent Projections to the Spinal Cord GUNNAR GRANT and BRITA ROBERTSON Department of Neuroscience, Karolinska Institutet Stockholm, Sweden

The primary afferent fibers projecting to the rat spinal cord enter via the dorsal roots. It has been suggested, however, that also ventral roots contribute afferents. It appears from studies conducted on the cat that the ventral roots may contain afferents at sacral levels and that these fibers terminate in the spinal gray matter (Mawe et al., 1984). Unmyelinated afferent fibers have been demonstrated in the L7 ventral root in the cat. Instead of penetrating into the central nervous system, these fibers either make U-turns or enter the pia mater (Risling and Hildebrand, 1982; Risling et al., 1984). Physiological studies suggest the occurrence of an additional category of sensory axons in feline sacral ventral roots. Blindly ending sensory axons with small, circumscribed spot-like receptive fields in ventral roots can be activated by chemical or mechanical stimuli such as a slight stretch of the roots (Jänig and Koltzenberg, 1991). Such “nervi nervorum” seem indeed to represent the dominating sensory component in the S2 ventral roots of the cat (Häbler et al., 1990). Electrophysiological data suggest that the ventral root afferents in the cat finally enter the central nervous system via the dorsal root rather than directly through the ventral root. Hence, dorsal root axons and neurons in the spinal gray matter can be activated from the distal but not the proximal stump of a divided ventral root (Chung et al., 1983, 1985; Kim et al., 1988; Shin et al., 1985, 1986). Unmyelinated ventral root afferents have also been found in the rat, but they appear to be significantly less numerous than their feline and human counterparts (Coggeshall et al., 1977). Further, there is evidence for the existence of myelinated

The Rat Nervous System, Third Edition

sensory axons making U-turns in rat ventral roots (Baik-Han et al., 1989; Bostock, 1981). The dorsal roots in the rat are grouped in pairs of 8 cervical, 13 thoracic, 6 lumbar, 4 sacral, and 3 caudal (coccygeal; for example, Waibl, 1973). The number of dorsal root ganglion cells in single pairs may vary considerably between the two sides (Ygge et al., 1981). A similar variation can therefore be expected to exist also in the number of dorsal root axons. Their actual number, however, may be larger than the number of ganglion cells (Chung and Coggeshall, 1984; Langford and Coggeshall, 1979). In the monkey, small caliber dorsal root axons are segregated into a lateral bundle as the dorsal rootlets enter the spinal cord (Snyder, 1977). This is not the case in the cat and does not appear to be prominent in the rat either (Willis and Coggeshall, 1991). After entering the spinal cord, the dorsal root afferents are distributed differently depending upon size. Coarse calibered fibers run medially into the dorsal funiculus, whereas fine fibers approach the dorsal horn via the dorsolateral fasciculus (Lissauer’s tract). More than two-thirds of the axons in the dorsolateral fasciculus at lumbosacral and midthoracic levels in the rat have been demonstrated to be of primary afferent origin (Chung et al., 1979). Anatomical studies on primary afferent projections to the spinal cord have been conducted mainly in cat and rat. Data from these studies, taken together, suggest that the primary afferent terminations in the spinal gray matter largely follow two principles of organization. First, fine calibered fibers are distributed preferentially

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in superficial laminae of the dorsal horn, whereas coarse fibers project more ventrally (for references see below). Second, somatic primary afferents terminate somatotopically along a mediolateral axis in the dorsal horn (Smith, 1983; Ygge and Grant, 1983; Cervero and Connell, 1984b; Molander and Grant, 1985, 1986; Swett and Woolf, 1985; Woolf and Fitzgerald, 1986; Ueyama et al., 1987; Shortland et al., 1989; Rivero-Melián and Grant, 1990, 1991; Shortland and Woolf, 1993; RiveroMelián, 1996; see also Willis et al., Chapter 27). With regard to visceral primary afferents, such afferents do not seem to have been studied specifically with regard to somatotopic organization. If present, such an organization may be difficult to reveal, due to the more widespread distribution of the afferents (e.g. Morgan et al., 1981; Kuo et al., 1983; Nadelhaft and Booth, 1984; Neuhuber et al., 1986; Sugiura et al., 1989; Sugiura and Tonosaki, 1995; Wang et al., 1998). In the sections below, we first describe the distribution of primary afferents to different laminae and some specific nuclei. Thereafter, their somatotopic arrangement is considered.1

PROJECTION OF PRIMARY AFFERENT FIBERS TO DIFFERENT LAMINAE AND SOME SPINAL CORD NUCLEI As in the cat and monkey (LaMotte, 1977; Réthelyi, 1977; Light and Perl, 1979a; Beal and Bicknell, 1981; Gobel et al., 1981), both lamina I (the marginal layer) and lamina II (substantia gelatinosa) of the rat spinal cord are reported to receive unmyelinated as well as fine myelinated primary afferent fibers (e.g., Light and Perl, 1979a; Janscó and Király, 1980; Nagy and Hunt, 1983; McMahon and Wall, 1985; Cruz et al., 1987; Fitzgerald, 1989; LaMotte et al., 1991). In the guinea pig, physiologically characterized, horseradish peroxidase (HRP)-labeled cutaneous and visceral C fiber afferents have been shown to terminate in both lamina I and lamina II (Sugiura et al., 1986, 1989, 1993; see also Willis et al., Chapter 27).

Lamina I The major primary afferent input appears to be provided by Aδ fibers, although there is also a cutaneous unmyelinated C fiber input (Sugiura et al., 1986; see 1 The distribution of different kinds of neurotransmitters/ neuromodulators and their receptors is not dealt with here. The reader is referred to the chapters dealing specifically with neurotransmitters; see also Ribeiro-da-Silva, Chapter 6; Weihe, 1990; Willis and Coggeshall, 1991; Hunt et al., 1992; Lawson, 1992.

also Gobel et al., 1981). In the primate, LaMotte (1977) found that afferent endings in lamina I degenerated slower than those in lamina II and suggested that Aδ primary afferents gave rise to a lamina I input. This was supported by evidence from physiological studies on cat and monkey (Kumazawa and Perl, 1978; Mense and Prabhakar, 1986). Direct evidence for a termination of Aδ fibers in lamina I in these two species was shown by Light and Perl (1979b) and further supported by morphological work on the monkey (Ralston and Ralston, 1979). With respect to muscle afferents, the number of afferents projecting to lamina I seems sparse and appears to vary not only between different muscle groups but also between species. Some studies on the cat show a clear terminal projection to lamina I (Craig and Mense, 1983; Nyberg and Blomqvist, 1985; Mense and Prabhakar, 1986; Mense and Craig, 1988), whereas others report essentially no or only sparse labeling following application of tracer in cats and rats (Mysicka and Zenker, 1981; Ammann et al., 1983; Abrahams et al., 1984; Bakker et al., 1984; Molander and Grant, 1987; Rivero-Melián, 1996). Projections of articular afferents have been the subject of a study in the cat (Craig et al., 1988). In addition to a projection to lamina I, the deep dorsal horn was found to receive afferents, similar to the situation for muscle (see below). As the authors point out, their data therefore support the existence of a common pattern for the central distribution of deep somatic afferents. They also found it reasonable to suggest that the articular afferent input to lamina I would comprise small diameter myelinated and unmyelinated (Group III and IV) fibers and may be primarily nociceptive. Apart from a somatic primary afferent input to lamina I, projections of visceral afferents have also been reported in monkey, cat, guinea pig, and rat (DeGroat et al., 1978; Morgan et al., 1981, 1986; Neuhuber, 1982; Ciriello and Calaresu, 1983; Kuo et al., 1983; Nadelhaft et al., 1983; Cervero and Connell, 1984a, 1984b; Neuhuber et al., 1986; Sugiura et al., 1989; Wang et al., 1998). In a quantitative analysis of unmyelinated C fiber afferents in guinea pig, more than 60% of the central synaptic enlargements of the visceral afferents were found to be localized superficially in lamina I, and the adjacent area (Sugiura et al., 1993). This seemed to be the main region of termination. Ten to twenty percent of the boutons appeared in deeper layers.

Lamina II Unmyelinated C fiber afferents have been found to provide the main primary afferent input in monkey, cat, guinea pig, and rat (Light and Perl, 1979a; Sugiura et al.,

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1986, 1989, 1993; Cruz et al., 1991). Such fibers are reported to have their main termination site in the same transverse band where fluoride-resistant acid phosphatase (FRAP)-positive terminals have been found (Coimbra et al., 1974; see also Ribeiro-da-Silva and Coimbra, 1982; Nagy and Hunt, 1983; and Ribeiro-daSilva, Chapter 6). Some fine myelinated afferent fibers, supposedly Aδ fibers subserving high-threshold mechanoreceptors, appear to terminate preferentially in the superficial parts (lamina IIo; Nagy and Hunt, 1983; lamina IIA of Ribeiro-da-Silva, Chapter 6). This has also been reported in the cat and monkey (Beal and Bicknell, 1981; LaMotte, 1977). Additionally, some fine myelinated fibers, presumably Aδ D-hair, and cutaneous mechanoreceptive Aβ afferents reach the deepest part of lamina II from the more ventrally situated lamina III in the rat (Cruz et al., 1987, 1991; Woolf, 1987; Beal et al., 1988; Shortland et al., 1989; Shortland and Woolf, 1993). Although lamina II appears to receive mainly unmyelinated cutaneous afferents, a small number of unmyelinated visceral afferents have been shown to terminate in this lamina in the guinea pig (Sugiura et al., 1989, 1993). Furthermore, presumed preterminal axons and/or terminals of visceral afferents have been found in the superficial part of lamina II in the rat, both from the inferior mesenteric plexus and hypogastric nerve (Neuhuber, 1982) and from the greater splanchnic nerve (Neuhuber et al., 1986). In the cat, the arborizations of unmyelinated afferents entering from the superficial part of the dorsal horn have been found distributed in narrow (150 μm wide) zones, and their terminals in still narrower (16–28 μm thick) sagittal sheets (Réthelyi, 1977). A zonal organization can also be seen for primary afferents in lamina II in the rat (Ygge and Grant, 1983; Molander and Grant, 1985, 1986; Tong et al., 1999). Recently, a group of itch-specific, mechanically insensitive dorsal horn neurons connected to very slowly conducting primary afferents (C fibers) were identified in monkey (Andrew and Craig, 2001). Primary afferents with similar properties have been found in humans (Schmelz et al., 1997). In the rat, histamine-responsive dorsal horn neurons have been described, but these respond also to mechanical stimuli and, thus, do not appear to be itch-specific (Jinks and Carstens, 2000).

The Lateral Spinal Nucleus In the dorsolateral funiculus, just lateral to the superficial dorsal horn, the rat spinal cord contains an aggregation of neurons called the lateral spinal nucleus (LSp); (see Grant and Koerber, Chapter 5). This has been described to receive visceral primary afferent fibers (Neuhuber, 1982; Neuhuber et al., 1986) and to respond

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to subcutaneous and/or deep structures (Menétrey et al., 1980). Willis et al. (Chapter 28) consider a possible role of the lateral spinal nucleus in nociception.

Laminae III–VI The primary afferent fibers projecting to the deep parts of the dorsal horn, including laminae III–V, and in the enlargements also lamina VI, are in general of a caliber coarser than those projecting to the superficial laminae. One exception to this is certain types of muscle and visceral afferents, terminating in lamina V, which have been identified in the cat (Light and Perl, 1979b; Craig and Mense, 1983; Kuo et al., 1983; Cervero and Connell, 1984a, 1984b; Morgan et al., 1986). Studies in the rat suggest that fine fibers, derived from muscle and viscera, also project to lamina V in this species (Ciriello and Calaresu, 1983; Neuhuber et al., 1986; Molander and Grant, 1987; Wang et al., 1998). The large-diameter afferent fibers enter the dorsal horn from the dorsal funiculus. For the cat, the course and termination of these fibers have been studied extensively, using different methods, including intraaxonal application of HRP to physiologically identified units. The results of these studies have been reviewed in a monograph on organization in the spinal cord (A. G. Brown, 1981; see also Maxwell and Bannatyne, 1983; Semba et al., 1983; Fyffe, 1984; Ralston et al., 1984; Willis and Coggeshall, 1991). With regard to largediameter primary afferent fibers in the rat, only a few studies have been published. The most extensive of these are the studies by Woolf (1987), Shortland et al. (1989), and Shortland and Woolf (1993), in which the method of intraaxonal application of HRP was used for an analysis of three types of low-threshold cutaneous mechanoreceptors. The general pattern of the terminal arborizations was one of mediolaterally compressed, rostrocaudally oriented sheets. The terminal arborizations of the hair follicle afferents had a distinctive morphology, identical to the “flame-shaped arbors” of Scheibel and Scheibel (1968). They had a recurrent course, distributing arborizations with synaptic boutons from lamina IV ventrally to inner lamina II dorsally. The two other types of units arborized within laminae III–V: rapidly adapting glabrous skin mechanoreceptors, which also may have some arbors projecting dorsally into lamina II, and slowly adapting type I afferent fibers, which in general have their terminal arbors somewhat deeper, in laminae IV, V, and even VI. Cutaneous afferent projections to the deep dorsal horn have also been demonstrated by transganglionic tracing in the rat (LaMotte et al., 1991; Maslany et al., 1992; RiveroMelián and Grant, 1991; Woolf and Fitzgerald, 1986). This method has also revealed muscle and visceral

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afferent projections, presumably of fine calibered fibers, to lamina V, as was commented on above. In addition, muscle afferent fiber projections to lamina VI, as well as to more ventral laminae, have been found (Molander and Grant, 1987; Rivero-Melián, 1996).

Lamina VII The intermediate gray, lamina VII, has been shown to receive a projection from muscle nerves in the rat (Mesulam and Brushart, 1979; Smith, 1983; Molander and Grant, 1987; Rivero-Melián, 1996). Muscle nerve projections to the dorsal nucleus (D; Clarke’s column) have also been demonstrated (Brushart and Mesulam, 1980; Molander and Grant, 1987; Rivero-Melián, 1996). In addition, cutaneous primary afferents project to part of this nucleus (Rivero-Melián and Grant, 1991). In the cat, articular nerves also contribute afferents to the dorsal nucleus (Craig et al., 1988). In the rat, neck muscle afferents project to the central cervical nucleus (CeC) which is located in the upper cervical segments of the spinal cord. This has been demonstrated both by tracing (Mysicka and Zenker, 1981; Ammann et al., 1983; Örnung et al., 1995) and in electrophysiological studies (Popova et al., 1995). Furthermore, there is evidence that the sacral parasympathetic nucleus, which is found at the dorsolateral border of lamina VII at L6 and S1 levels in the rat, also receives a primary afferent projection, although of visceral origin (Nadelhaft and Booth, 1984; Wang et al., 1998).

Area X Area X and the dorsal gray commissure have been shown in several studies to receive visceral afferents in the rat. These have been derived both from renal (Ciriello and Calaresu, 1983) and pelvic (Nadelhaft and Booth, 1984; Neuhuber, 1982; Wang et al., 1998) nerves, as well as from the greater splanchnic nerve (Neuhuber et al., 1986). The area has also been shown to receive somatic afferents, both in rat (Neuhuber et al., 1986) and cat (Cervero and Connell, 1984b; Honda, 1985). Furthermore, Honda (1985) found that cells in this area in the sacral spinal cord in the cat received primary afferent inputs converging from a wide range of receptor types in somatic and visceral structures. A possible involvement of cells in this area in the transmission of visceral nociception has recently been discussed (Wang et al., 1999; see also Willis et al., Chapter 27).

The Ventral Horn The organization of primary afferent projections to the ventral horn has been studied in detail by tracing

in the rat. Smith (1983) investigated the development and postnatal organization of primary afferents to the thoracic cord, using unconjugated HRP. She found significant labeling, however, only in animals less than 17 days of age. Other investigators have studied the projections of muscle afferents in the adult animal at lumbar levels (Mesulam and Brushart, 1979; RiveroMelián and Grant, 1990; Rivero-Melián, 1996) and in the upper cervical cord (Mysicka and Zenker, 1981; see also Örnung et al., 1995). Very prominent labeling was achieved by using HRP conjugated to the B-fragment of cholera toxin (Rivero-Melián and Grant, 1990; RiveroMelián, 1996). This seems superior for the labeling of somatic myelinated afferents. The results achieved by Smith confirmed that there are connections between afferents and motoneurons with axons in the same nerve and showed that the afferent boutons were distributed widely across the dendritic arbors of the motoneurons. At lumbar levels the densest primary afferent projection from each injected muscle nerve was found in the homonymous group of motoneurons (Rivero-Melián, 1996). It is obvious that the principle of organization of spinal cord primary afferent fibers, that fine-calibered fibers terminate in superficial laminae of the dorsal horn and coarse-calibered fibers more ventrally in the gray matter, is not an absolute one. Fine-calibered primary afferents are found in lamina V and visceral, presumed fine fibers terminate both in lamina VII, at the sacral level, and in area X and the dorsal gray commissural region. Indeed, a highly specialized central projection of primary afferent endings related to sensory function and not to fiber diameter was proposed by Light and Perl (1979b). Furthermore, afferent fibers from different types of peripheral targets, such as skin, muscle, and viscera have different, characteristic termination sites (Fig. 1). The finding of a partial overlap of some types of afferent fibers, such as visceral and cutaneous fibers, as in lamina I, and somatic and visceral in the area around the central canal, would have to be expected if convergence of different types of afferent fibers is to be made possible.

SOMATOTOPIC ORGANIZATION OF PRIMARY AFFERENT PROJECTIONS Early physiological studies on the cat demonstrated a somatotopic organization of cells in the dorsal horn activated by low-threshold cutaneous afferent fibers, suggesting a similar organization for the incoming afferents (Koerber and Brown, 1982). Physiological studies have confirmed such an organization (see A.G. Brown, 1981; P.B. Brown et al., 1991, 1992). There was

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FIGURE 1 Schematic drawings of transverse sections of the spinal cord gray matter on one side of one lower lumbar (left) segment and one lower thoracic (right) segment in the rat, showing principal termination sites for afferent fibers from skin and muscle (left) and viscera (right).

also anatomical support for a somatotopic arrangement of the incoming afferents (see Grant and Ygge, 1981; Ygge and Grant, 1983). As will be described below, the HRP tracing method has confirmed such an arrangement and extended our knowledge as to the intricacy of the organization. The thoracic level, where the segmental pattern of the body is clearly preserved, should be well suited for studies of the general principles of organization of primary afferent projections in the spinal gray matter. In such studies conducted on the rat, it was found that the dorsal and ventral rami, the two main components of the spinal nerve, have their projections restricted to the lateral and medial parts of the dorsal horn, respectively (Smith, 1983; Ygge and Grant, 1983). Furthermore, the projections of the two main branches of the ventral ramus were found to be somatotopically organized, within the projection compartment of their parent nerve (Ygge and Grant, 1983) (Fig. 2). This principle of organization was confirmed in a later study of projections of cutaneous afferent fibers from different dorsoventral sectors of the cat’s tail, using single-unit labeling of physiologically identified primary afferent fibers (Ritz et al., 1989). The analysis of the rostrocaudal extension of the projections showed that the mediolateral compartments extended into neighboring segments, resulting in an overlap between compartments from corresponding rami of adjacent spinal nerves (Ygge and Grant, 1983). The central branches of hindlimb and forelimb nerves and lumbar dorsal root ganglia, as well as afferents

from cutaneous regions of the paws (see Fig. 2), have also no been shown to be organized somatotopically in mediolateral compartments in the dorsal horn similar to those of the thoracic spinal nerve (Molander and Grant, 1985, 1986; Nyberg and Blomqvist, 1985; Swett and Woolf, 1985; Woolf and Fitzgerald, 1986; Shortland et al., 1989; Rivero-Melián and Grant, 1990, 1991; Maslany et al., 1992; Shortland and Woolf, 1993; Rivero-Melián, 1996). Furthermore, central branches of the pudendal nerve in rat are similarly organized (Ueyama et al., 1987). Studies by Mirnics and Koerber (1995) indicate that the somatotopic organization of the incoming afferents is established very early in development and requires little refinement to match that seen in the adult. The somatotopic arrangement does not exclude the possibility that afferent fibers also project outside their predicted termination sites. An example of this, although not from the dorsal horn, is the projection from the rat sciatic nerve not only to the gracile but also to the cuneate nucleus in the rat (Grant et al., 1979). Such “exterior” projections might conceivably serve interactions between different peripheral sources, necessary for proper sensory function.

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FIGURE 2 (Left) Schematic drawing of transverse section of the spinal cord gray matter on one side of the 10th thoracic segment in the rat, showing somatotopically arranged termination of nerves from dorsal, lateral, and ventral sectors of the trunk. (Right) Schematic dorsal view map of lamina II, depicting density centers of projections from various parts of the rat’s hindpaw skin (bottom) and the somatotopic termination of nerve branches from different sectors of the trunk (top) (compare with the drawing on the left, which was modified from Fig. 5 in Molander and Grant, 1990). Ammann, B. M., Gottschall, J., and Zenker, W. (1983). Afferent projections from the rat longus capitis muscle studied by transganglionic transport of HRP. Anat. Embryol. 166, 275–289. Andrew, D., and Craig, A. D. (2001). Spinothalamic lamina I neurons selectively sensitive to histamine: A central neural pathway for itch. Nat. Neurosci. 4, 72–77. Baik-Han, E. J., Kim, K. J., and Chung, J. M. (1989). Electrophysiological evidence for the presence of looping myelinated afferent fibers in the rat ventral root. Neurosci. Lett. 104, 65–70.

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Schmelz, M., Schmidt, R., Bickel, A., Handwerker, H. O., and Torebjörk, H. E. (1997). Specific C-receptors for itch in human skin. J. Neurosci. 15, 8003–8008. Semba, K., Masarachia, P., Malamed, S., Jacquin, M., Harris, S., Yang, G., and Egger, M. D. (1983). An electron microscopic study of primary afferent terminals from slowly adapting Type I receptors in the cat. J. Comp. Neurol. 221, 466–481. Shin, H. K., Kim, J., and Chung, J. M. (1985). Flexion reflex elicited by ventral root afferents in the cat. Neurosci. Lett. 62, 353–358. Shin, H. K., Kim, J., Nam, S. C., Paik, K. S., and Chung, J. M. (1986). Spinal entry route for ventral root afferent fibers in the cat. Exp. Neurol. 94, 714–725. Shortland, P., and Woolf, C. J. (1993). Morphology and somatotopy of the central arborizations of rapidly adapting glabrous skin afferents in the rat lumbar spinal cord. J. Comp. Neurol. 329, 491–511. Shortland, P., Woolf, C. J., and Fitzgerald, M. (1989). Morphology and somatotopic organization of the central terminals of hindlimb hair follicle afferents in the rat lumbar spinal cord. J. Comp. Neurol. 289, 416–433. Smith, C. L. (1983). The development and postnatal organization of primary afferent projections to the rat thoracic spinal cord. J. Comp. Neurol. 220, 29–43. Snyder, R. (1977). The organization of the dorsal root entry zone in cats and monkeys. J. Comp. Neurol. 174, 47–70. Sugiura, Y., Lee, C. L., and Perl, E. R. (1986). Central projections of identified, unmyelinated (C) afferent fibers innervating mammalian skin. Science 234, 358–361. Sugiura, Y., Terui, N., and Hosoya, Y. (1989). Differences in distribution of central terminals between visceral and somatic unmyelinated (C) primary afferent fibers. J. Neurophysiol. 62, 834–840. Sugiura, Y., Terui, N., Hosoya, Y., Tonosaki, Y., Nishiyama, K., and Honda, T. (1993). Quantitative analysis of central terminal projections of visceral and somatic unmyelinated (C) primary afferent fibers in the guinea pig. J. Comp. Neurol. 15, 315–325. Sugiura, Y., and Tonosaki, Y. (1995). Spinal organization of unmyelinated visceral afferent fibers in comparison with somatic afferent fibers. In “Visceral Pain” (Gebhart, D. F., Ed.), pp. 41–59. IASP Press, Seattle. Swett, J. E., and Woolf, C. J. (1985). The somatotopic organization of primary afferent terminals in the superficial laminae of the dorsal horn of the rat spinal cord. J. Comp. Neurol. 231, 66–77.

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Tong, Y.-G., Wang, H. F., Ju, G., Grant, G., Hökfelt, T., and Zhang, X. (1999). Increased uptake and transport of cholera toxin B-subunit in dorsal root ganglion neurons after peripheral axotomy: Possible implication for sensory sprouting. J. Comp. Neurol. 404, 143–158. Ueyama, T., Arakawa, H., and Mizuno, N. (1987). Central distribution of efferent and afferent components of the pudental nerve in the rat. Anat. Embryol. 177, 37–49. Waibl, H. (1973). Zur Topographie der Medulla Spinalis der Albinoratte (Rattus norvegicus). Adv. Anat. Embryol. Cell Biol. 47(fasc. 6), 1–42. Wang, C.-C., Willis, W. D., and Westlund, K. N. (1999). Ascending projections from the area around the spinal cord central canal: A Phaseolus vulgaris leucoagglutinin study in rats. J. Comp. Neurol. 415, 341–367. Wang, H. F., Shortland, P., Park, M. J., and Grant, G. (1998). Retrograde and transganglionic transport of horseradish peroxidase conjugated cholera toxin B subunit, wheat germ agglutinin and isolectin B4 from Griffonia simplicifolia I in primary afferent neurons innervating the rat urinary bladder. Neuroscience 87, 275–288. Weihe, E. (1990). Neuropeptides in primary afferent neurons. In “The Primary Afferent Neuron: A Survey of Recent MorphoFunctional Aspects” (W. Zenker and W. L. Neuhuber, Eds.), pp. 161–172. Plenum, New York. Willis, W. D., and Coggeshall, R. E. (1991). “Sensory Mechanisms of the Spinal Cord,” 2nd ed. Plenum, New York. Woolf, C. J. (1987). Central termination of cutaneous mechanoreceptive afferents in the rat lumbar spinal cord. J. Comp. Neurol. 261, 105–119. Woolf, C. J., and Fitzgerald, M. (1986). Somatotopic organization of cutaneous afferent terminals and dorsal horn neuronal receptive fields in the superficial and deep laminae of the rat lumbar spinal cord. J. Comp. Neurol. 251, 517–531. Ygge, J., Aldskogius, H., and Grant, G. (1981). Asymmetries and symmetries in the number of thoracic dorsal root ganglion cells. J. Comp. Neurol. 202, 365–372. Ygge, J., and Grant, G. (1983). The organization of the thoracic spinal nerve projection in the rat dorsal horn demonstrated with transganglionic transport of horseradish peroxidase. J. Comp. Neurol. 216, 1–9.

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C H A P T E R

5 Spinal Cord Cytoarchitecture GUNNAR GRANT Department of Neuroscience, Karolinska Institutet Stockholm, Sweden

H. RICHARD KOERBER Department of Neurobiology, The University of Pittsburgh School of Medicine Pittsburgh, USA

The cytoarchitectonic scheme presented for the cat spinal cord by Rexed (1952, 1954) has gained wide acceptance in the neuroscience literature. It is based on the morphology and arrangement of Nissl-stained cell bodies in transverse sections, providing a framework to which localized features, anatomical, physiological, or histochemical, may be related. The gray matter is divided into 10 cytoarchitectonic regions, laminae I–IX and an area around the central canal (area X). As emphasized Rexed (1952), the borders between the different laminae may be indistinct and should be recognized as zones of transition rather than as strict borderlines. Although originally presented for the cat, Rexed’s scheme has, with some alterations, been found to be applicable also to the rat spinal cord (Fukuyama, 1955; Steiner and Turner, 1972; McClung and Castro, 1976; Molander et al., 1984, 1989; Paxinos and Watson, 1986). The following description of the rat spinal cord cytoarchitecture represents an updated version of that of the chapter by Molander and Grant in the 2nd edition of The Rat Nervous System, which was based primarily on previously published observations by Molander et al. (1984, 1989).

Lima and Coimbra (1986) described fusiform, multipolar, flattened, and pyramidal cell types, most of which have their major dendrites within lamina I or in the adjacent white matter. Some send dendrites ventrally as far as lamina V (Beal et al., 1989). The flattened and pyramidal neurons correspond to the marginal cells described in the classical literature (Rexed, 1952; Lima and Coimbra, 1986). A large proportion of the cells with perikarya in lamina I or the adjacent white matter seem to be wide dynamic range neurons activated by both low- and high-intensity stimulation (Menétrey et al., 1977; McMahon and Wall, 1983; Woolf and Fitzgerald, 1983). Other neurons seem to respond primarily to either low- or high-threshold stimulation (McMahon and Wall, 1983; Woolf and Fitzgerald, 1983). Furthermore, there is evidence that neurons in lamina I, as well as in lamina II, respond to both pruritic and algesic chemical stimuli and thus might participate in transmitting sensations of itch and/or chemogenic pain (Jinks and Carstens, 2000; cf. Grant and Robertson, Chapter 4). Subpopulations of neurons in lamina I project to the brain stem, including among other structures the nucleus of the solitary tract, the parabrachial nucleus, and the periaqueductal gray (Menétrey et al., 1982; Chaouch et al., 1983; Cechetto et al., 1985; Lima and Coimbra, 1989, 1990; Esteves et al. 1993; Tavares et al., 1993; Feil and Herbert, 1995; Kayalioglu et al., 1999; Bester et al., 2000), to hypothalamus (Burstein et al., 1987, 1990a; Kayalioglu et al., 1999), and to thalamus (Granum, 1986; Kemplay and Webster, 1986; Lima and Coimbra, 1988; Hylden et al., 1989; Burstein et al., 1990b; Kobayashi, 1998; Kayalioglu et al., 1999). The type of

LAMINA I Lamina I (marginal zone) forms a thin rim along the dorsal and dorsolateral edges of the dorsal horn. Most of the cells are small but a few mediolaterally elongated large cells can usually be seen in each section. The neuropil is oriented tangentially to the lamina. Using three-dimensional reconstructions from Golgi sections,

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dendritic pattern seems to be correlated to projection target (Beal et al., 1989; Lima and Coimbra, 1990). Earlier studies in rat suggested a lack of correlation between cell morphology and functional properties of lamina I cells (Woolf and Fitzgerald, 1983). More recently, however, Han et al. (1998) showed a strong correlation between morphological type and functional characteristics in cat. They report that cells responding specifically to nociceptive stimuli were identified as different varieties of fusiform cells. Those responding to innocuous cold were pyramidal and those classified as polymodal nociceptive were morphologically identified as multipolar (also see Light and Willcockson, 1999).

neurons in lamina II are relatively few, but some have been found to project to the caudal ventrolateral reticular formation of the medulla oblongata (Lima and Coimbra, 1991) and a few cells in a region approximately corresponding to lamina II have been found to project to the lateral cervical nucleus and the brain stem (Giesler et al., 1978) and thalamus (Burstein et al., 1990b). There appear to be morphological differences in dendritic tree shape between projection neurons and locally projecting neurons (Beal et al., 1989), but no clear correlation has been found between morphology and functional types [Light et al., 1979 (cat); Woolf and Fitzgerald, 1983; Light and Willcockson, 1999 (rat)].

LAMINA II

LAMINA III

Lamina II (substantia gelatinosa) is subjacent and parallel to lamina I. It is wider than lamina I and is characterized in Nissl-stained material by its dominance of small round cell bodies with sparse Nissl substance. At the levels of the enlargement lamina II diverges from the usual parallel arrangement with lamina I. At these level lamina II is depressed ventrally away from lamina I (Woodbury et al., 2000). Interspersed between lamina I and lamina II is a distinct hemi-lamina that differs from lamina I as it lacks myelinated fiber inputs and large marginal layer neurons. This area does not receive input from fibers that bind the IB4 isolectin (Wang et al., 1994; Woodbury et al., 2000) but does receive a rich peptidergic innervation (e.g., Silverman and Kruger, 1988, 1990). At the lumbar level, the location of this dip in lamina II somatotopically corresponds to the glabrous skin of the feet (Woodbury et al., 2000). Lamina II has an intensely stained outer zone (IIo; named IIA by Ribeiro-da-Silva, Chapter 6) with densely packed cells and a less compact inner zone (IIi; named IIB by Ribeiro-da-Silva, Chapter 6). Myelinated fibers are spars except for bundles of myelinated fibers that cross the lamina, particularly in its medial part. Neurons in lamina IIo are most often characterized as nociceptive, responding best to high-intensity stimulation. Most neurons in IIi respond maximally to brush stimuli and are classified as non-nociceptive (Woolf and Fitzgerald, 1983; Light, 1992; Light and Willcockson, 1999). These results are consistent with the demonstration of extensive input from low-threshold mechanoreceptors in IIi (Woodbury et al., 2000). A description of rat lamina II neurons, which was based on Golgi and retrograde tract tracing techniques, was published by Beal et al. (1989). On the basis of dendritic morphology, the cells were classified into limiting, central, islet, stalked, inverted stalk, arboreal, spiny, vertical, and star-shaped cells. Tract

Lamina III runs just ventral and parallel to lamina II. It has a cytoarchitectonic appearance similar to that of lamina II but shows a slightly wider range of cell sizes and is less compact. The border between lamina II and lamina III is difficult to recognize from cell morphology, but it can often be distinguished by a clearly visible transition from the homogeneous neuropil characteristic of IIi to a more heterogeneous neuropil in lamina III. If a myelin stain is used, the almost myelin-free lamina IIi stands out clearly against lamina III, which contains numerous fine myelinated fibers. The dendritic fields of many lamina III neurons are oriented rostrocaudally [Scheibel and Scheibel, 1968 (cat and rat); Beal et al., 1988 (rat)]. Many cells in lamina III respond only to weak mechanical stimuli (Cervero et al., 1988). Cells in a region corresponding approximately to lamina III have been shown to project to other regions within the same segment of the spinal cord [Light and Kavookjian, 1988 (cat and monkey)], the dorsal column nuclei [Giesler et al., 1984 (rat)], the lateral cervical nucleus [Baker and Giesler, 1984 (rat)], and the thalamus [Burstein et al., 1990b (rat)]. Brown (1981) noted that postsynaptic dorsal column neurons in the cat have wide-spread dendrites, some reaching laminae I and IIo, whereas the dendrites of spinocervical tract cells are oriented more rostrocaudally and do not reach laminae I and IIo. Whether this is true also in the rat is as yet unknown.

LAMINA IV Lamina IV forms the base of the head of the dorsal horn and curves ventrally along its medial border. It becomes continuous with the contralateral lamina IV in the dorsal commissure at lumbar and sacral levels and ends at area X (see below) at thoracic and cervical levels.

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The cells in lamina IV appear more loosely arranged than those in lamina III, and some of them are multipolar and considerably larger. In the thoracic and upper lumbar region, lamina IV becomes interrupted by the elongated cells in Clarke’s column at the base of the dorsal horn. Cells in this nucleus have their dendrites oriented rostrocaudally [Scheibel and Scheibel, 1968 (cat and rat); Loewy, 1970 (cat)], respond to proprioceptive and cutaneous stimuli [Oscarsson, 1973 (cat)], and project to the cerebellum (Matsushita and Hosoya, 1979; RiveroMelián and Grant, 1990). A small group of spinocerebellar neurons have also been found in the dorsal column white matter just dorsal to Clarke’s column at levels T13 through L2 (Beal et al., 1990; Rivero-Melián and Grant, 1990). This group of neurons is, however, not the same as the network of neurons located along the midline of the dorsal column white matter described by Abbadie et al. (1999). The cells in lamina IV have widespread dendritic fields [Scheibel and Scheibel, 1968 (cat and rat)]. Many of the neurons seem to have their major dendritic orientation toward superficial laminae, so called “antenna-like neurons” [Szentágothai, 1964 (cat); Scheibel and Scheibel, 1968 (cat and rat); Schoenen, 1982 (man)]. As in lamina III, many cells in lamina IV respond only to light mechanical stimuli, although nociceptivespecific and wide dynamic range neurons are present as well (cf. Cervero et al., 1988). Furthermore, as in lamina III, subpopulations of cells in a region corresponding approximately to lamina IV project locally within the spinal cord [Mannen, 1975 (cat)] to the lateral cervical nucleus (Baker and Giesler, 1984), to the dorsal column nuclei (Giesler et al., 1984), and to the thalamus (Burstein et al., 1990b; Kayalioglu et al., 1999).

tract neurons in the lamina V region are the lateral cervical nucleus (Baker and Giesler, 1984), the dorsal column nuclei (Giesler et al., 1984), the brain stem reticular formation (Chaouch et al., 1983), the midbrain (Menétrey et al., 1982), the cerebellum (Matsushita and Hosoya, 1979; Rivero-Melián and Grant, 1990), the thalamus (Burstein et al., 1990b; Kayalioglu et al., 1999), and the amygdala (Burstein and Potrebic, 1993). Some neurons project to other parts of the spinal cord [Mannen, 1975 (cat)].

LAMINA VI Lamina VI forms the base of the dorsal horn. It consists of a narrow band of darkly stained compactly arranged neurons and is present mainly in the enlargements. The borders with the neighboring laminae V and VII are ambiguous. The dendritic fields of the cells in this layer are similar to those described above for lamina V, although they may be more extensive [Brown, 1981 (cat)]. Cells in this layer have been described to respond to cutaneous and proprioceptive inputs (Wall, 1967). Some respond primarily to noxious stimuli, others are of the wide dynamic range type (Cervero et al., 1988). A subpopulation of the cells in this layer seems to project to ventral horn motoneurons [Hongo et al., 1989 (cat)]. Cells in the medial part of lamina VI in the upper cervical segments give rise to axons projecting to the cerebellum (Matsushita and Xiong, 2001).

LAMINA VII LAMINA V Lamina V forms the neck of the dorsal horn and is the widest layer situated here. The wide lateral part of this layer can easily be recognized by its reticulated appearance; the medial nonreticulated part narrows as it approaches the midline dorsal to the central canal. The neurons in lamina V appear more heterogeneous in shape and size than those in lamina IV, but the border between lamina IV and V is difficult to distinguish, particularly medially. The dendritic fields of the cells in this layer radiate primarily in the transverse plane and to a lesser extent rostrocaudally [Scheibel and Scheibel, 1968 (cat and rat); Mannen, 1975 (cat); Brown, 1981 (cat)]. Correlating morphology with function, Ritz and Greenspan [1985 (cat)]. noted that cells responding to both noxious and light mechanical stimuli were larger than those responding only to noxious or only to light mechanical stimuli. The projection targets of

Lamina VII corresponds to the intermediate zone of the gray matter and to parts of the ventral horn not occupied by laminae VIII and IX. It has a lighter and more homogeneous appearance in Nissl-stained sections than the adjacent laminae. Lamina VII contains the intermediolateral nucleus in segments T1–L3 (preganglionic sympathetic neurons) and L6–S1 (preganglionic parasympathetic neurons) and the intermediomedial nucleus at all levels. The central cervical nucleus can be seen in segments C1–3. Cells in this nucleus, as well as some of the other cells in lamina VII, are known to project to the cerebellum (Matsushita and Hosoya, 1979; Rivero-Melián and Grant, 1990; Matsushita, 1991; Matsushita et al., 1991; Matsushita and Yaginuma, 1995), as well as to the vestibular nuclei (Matsushita et al., 1995; Sato et al.,1997). There are also cells in lamina VII that project to other parts of the spinal cord [Mannen, 1975 (cat)], to the brain stem reticular formation (Chaouch et al., 1983; Shokunbi

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et al., 1985), and to the thalamus (Chaouch et al., 1983; Burstein et al., 1990b).

LAMINA VIII Lamina VIII, located in the ventral or ventromedial part of the ventral horn, has a more heterogeneous appearance and generally slightly larger cells than lamina VII. This lamina contains commissural cells. As with lamina VII, some cells in this region project to the brain stem reticular formation (Chaouch et al., 1983; Schokunbi et al., 1985) and to the thalamus (Burstein et al., 1990b).

LAMINA IX Lamina IX consists of collections of cell groups bordering the lateral and ventral edges of the ventral horn. Many of the darkly stained large cells in these groups are motoneurons projecting through the ventral roots. The groups can often be ascribed to particular muscles or groups of muscles (cf. Swett et al., 1986; RiveroMelián, 1996). The range of soma sizes shows a bimodal pattern, presumably representing larger α-motoneurons and smaller γ-motoneurons (Swett et al., 1986). The dendrites of these neurons have a wide distribution, occasionally extending as far dorsally as lamina III (Cook and Woolf, 1985).

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FIGURE 1, cont’d Schematic drawings of transverse sections from different levels of the spinal cord. Outlines from Figs. 77 and 78 in The Rat Brain in Stereotaxic Coordinates (George Paxinos and Charles Watson, Eds.), Academic Press, San Diego. I–IX, cytoarchitectonic laminae; X, area X; dl, dorsolateral fasciculus; CeCv, central cervical nucleus; D, dorsal nucleus (Clarke); IML, intermediolateral nucleus; IMM, intermediomedial nucleus; LSp, lateral spinal nucleus; LatC, lateral cervical nucleus; py, pyramidal tract.

AREA X Area X is the area surrounding the central canal. It borders the white matter ventrally and dorsally, except for the lumbosacral levels, where it borders dorsally dorsal horn layers crossing the midline. The cells are generally smaller and more densely packed than those

in the adjacent lamina VII. They are pyramidal, stellate, and fusiform, and many of the cells respond to noxious stimuli (Nahin et al., 1983). Cells in the area have been found to project to the brain stem (Menétrey et al., 1982; Nahin et al., 1983; Wang et al., 1999), amygdala (Burstein and Potrebic, 1993), the hypothalamus (Burstein et al., 1987), and the thalamus (Burstein et al.,

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1990b). At lumbosacral levels, a rostrocaudally arranged column of cells has been found to be immunoreactive to different opioids (Nicholas et al., 1999). Subpopulations of these contribute to ascending pathways to the reticular formation of the medulla oblongata (Nahin and Micevych, 1986) and to the thalamus (Ju et al., 1987). A possible involvement of cells in area X in the transmission of visceral nociception has recently been discussed (Wang et al., 1999; see also Willis et al., Chapter 27).

LATERAL SPINAL NUCLEUS The lateral spinal nucleus consists of multipolar neurons in the white matter ventrolateral to the lateral edge of the dorsal horn and is present at all levels of the spinal cord. Cells in this nucleus generally respond only to stimulation of subcutaneous and/or deep structures (Menétrey et al., 1980). They have been shown to project bilaterally to the midbrain (Menétrey et al., 1982; Kayalioglu et al., 1999), hypothalamus (Burstein et al., 1987; Kayalioglu et al., 1999) and the thalamus (Burstein et al., 1990b; Kayalioglu et al., 1999). In addition, neurons in the lateral spinal nucleus as well as the lateral funiculus in the upper cervical spinal cord have been shown to project directly to the sympathetic preganglionic neurons (Jansen and Loewy, 1997).

LATERAL CERVICAL NUCLEUS The lateral cervical nucleus consists of mainly rounded neurons and is found just lateral to the lateral spinal nucleus in the C1–3 segments. All cells in this nucleus respond to hair movement and some to noxious stimulation (Giesler et al., 1979). They project mainly contralaterally to the midbrain (Giesler et al., 1988) and the thalamus (Granum, 1986; Kemplay and Webster, 1986; Burstein et al., 1990b).

References Abbadie, C., Skinner, K., Mitrovic, I., and Basbaum, A. I. (1999). Neurons in the dorsal column white matter of the spinal cord: Complex neuropil in an unexpected location. Proc. Natl. Acad. Sci. USA 98, 9836–9841. Baker, M. L., and Giesler, G. J. (1984). Anatomical studies of the spinocervical tract of the rat. Somatosens. Mot. Res. 2, 1–18. Beal, J. A., Knight, D. S., and Nandi, K. N. (1990). Nerve cell bodies in the dorsal funiculus of the rat spinal cord. Exp. Brain Res. 81, 372–376. Beal, J. A., Nandi, K. N., and Knight, D. S. (1989). Characterization of long ascending tract projection neurons and nontract neurons in the superficial dorsal horn (SDH). In “Processing of Sensory Information in the Spinal Cord” (Cervero, F., Bennett, G. J., and Headly, H. M., Eds.), pp. 181–197. Plenum, New York.

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Jansen, A. S., and Loewy, A. D. (1997). Neurons lying in the white matter of the upper cervical spinal cord project to the intermediolateral cell column. Neuroscience 77, 889–898. Jinks, S. L., and Carstens, E. (2000). Superficial dorsal horn neurons identified by intracutaneous histamine: Chemonociceptive responses and modulation by morphine. J. Neurophysiol. 84, 616–627. Ju, G., Melander, T., Ceccatelli, S., Hokfelt, T., and Frey, P. (1987). Immunohistochemical evidence for a spinothalamic pathway cocontaining cholecystokinin- and galanin-like immunoreactivities in the rat. Neuroscience 20, 439–456. Kayalioglu, G., Robertson, B., Kristensson, K., and Grant, G. (1999). Nitric oxide synthase and interferon-γ receptor immunoreactivities in relation to ascending spinal pathways to thalamus, hypothalamus and the periaqueductal grey in the rat. Somatosens. Mot. Res. 16, 280–290. Kemplay, S. K., and Webster, K. E. (1986). A qualitative and quantitative analysis of the distributions of cells in the spinal cord and spinomedullary junction projecting to the thalamus of the rat. Neuroscience 17, 769–789. Kobayashi, Y. (1998). Distribution and morphology of spinothalamic tract neurons in the rat. Anat. Embryol. 197, 51–67. Light, A. R. (1992). The initial processing of pain and its descending control: Spinal and trigeminal systems. In “Pain and headache” (Gildenberg, P. L., Ed.), Vol. 12. Karger, Basel. Light, A. R., and Kavookjian, A. M. (1988). Morphology and ultrastructure of physiologically identified substantia gelatinosa (lamina II) neurons with axons that terminate in deeper dorsal horn laminae (III–V). J. Comp. Neurol. 267, 172–189. Light, A. R., Trevino, D. L., and Perl, E. R. (1979). Morphological features of functionally defined neurons in the marginal zone and substantial gelatinosa of the spinal dorsal horn. J. Comp. Neurol. 186, 151–172. Light, A. R., and Willcockson, H. H. (1999). Spinal laminae I–II neurons in rat recorded in vivo in whole cell, tight seal configuration: Properties and opioid responses. J. Neurophysiol. 82, 3316–3326. Lima, D., and Coimbra, A. (1986). A Golgi study of the neuronal population of the marginal zone (lamina I) of the rat spinal cord. J. Comp. Neurol. 244, 53–71. Lima, D., and Coimbra, A. (1988). The spinothalamic system of the rat: Structural types of retrogradely labelled neurons in the marginal zone (lamina I). Neurosci. Lett. 27, 215–230. Lima, D., and Coimbra, A. (1989). Morphological types of spinomesencephalic neurons in the marginal zone (lamina I) of the rat spinal cord, as shown after retrograde labeling with cholera toxin subunit B. J. Comp. Neurol. 279, 327–339. [Erratum: J. Comp. Neurol. (1989) 286, 542] Lima, D., and Coimbra, A. (1990). Structural types of marginal (lamina I) neurons projecting to the dorsal reticular nucleus of the medulla oblongata. Neuroscience 34, 591–606. Lima, D., and Coimbra, A. (1991). Neurons in the substantia gelatinosa Rolandi (lamina II) project to the caudal ventrolateral reticular formation of the medulla oblongata in the rat. Neurosci. Lett. 132, 16–18. Loewy, A. D. (1970). A study of neuronal types in Clarke’s column in the adult cat. J. Comp. Neurol. 139, 53–80. Mannen, H. (1975). Reconstruction of axonal trajectory of individual neurons in the spinal cord using Golgi-stained serial sections. J. Comp. Neurol. 159, 357–374. Matsushita, M. (1991). Cerebellar projections of the central cervical nucleus in the rat: An anterograde tracing study. Neurosci. Res. 12, 201–216. Matsushita, M., Gao, X., and Yaginuma, H. (1995). Spinovestibular projections in the rat, with particular reference to projections from the central cervical nucleus to the lateral vesticular nucleus. J. Comp. Nuerol. 361, 334–344.

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Matsushita, M., and Hosoya, Y. (1979). Cells or origin of the spinocerebellar tract in the rat, studied with the method of retrograde transport of horseradish peroxidase. Brain Res. 173, 185–200. Matsushita, M., Ragnarson, B., and Grant, G. (1991). Topographic relationship between sagittal Purkinje cell bands revealed by a monoclonal antibody to zebrin I and spinocerebellar projections arising from the central cervical nucleus in the rat. Exp. Brain Res. 84, 133–141. Matsushita, M., and Xiong, G. (2001). Uncrossed and crossed projections from the upper cervical spinal cord to the cerebellar nuclei in the rat, studied by anterograde tracing. J. Comp. Neurol. 432, 101–118. Matsushita, M., and Yaginuma, H. (1995). Projections from the central cervical nucleus to the cerebellar nuclei in the rat, studied by anterograde axonal tracing. J. Comp. Neurol. 353, 234–246. McClung, J. R., and Castro, A. J. (1976). Neuronal organization in the spinal cord of the rat: An analysis of the nine lamina scheme of Rexed. Anat. Rec. 184, p. 474. McMahon, S. B., and Wall, P. D. (1983). A system of rat spinal cord lamina I cells projecting through the contralateral dorsolateral funiculus. J. Comp. Neurol. 214, 217–223. Menétrey, D., Chaouch, A., and Besson, J. M. (1980). Location and properties of dorsal horn neurons at origin of spinoreticular tract in the lumbar enlargement of the rat. J. Neurophysiol. 44, 862–877. Menétrey, D., Chaouch, A., Binder, D., and Besson, J. M. (1982). The origin of the spinomesencephalic tract in the rat: An anatomical study using the retrograde transport of horseradish peroxidase. J. Comp. Neurol. 206, 862–867. Menétrey, D., Giesler, G. J., and Besson, J. M. (1977). An analysis of response properties of spinal cord dorsal horn neurones to nonnoxious and noxious stimuli in the spinal rat. Exp. Brain Res. 27, 15–33. Molander, C., Xu, Q., and Grant, G. (1984). The cytoarchitectonic organization of the spinal cord in the rat. I. The lower thoracic and lumbosacral cord. J. Comp. Neurol. 230, 133–141. Molander, C., Xu, Q., Rivero-Melián, C., and Grant, G. (1989). Cytoarchitectonic organization of the spinal cord in the rat. II. The cervical and upper thoracic cord. J. Comp. Neurol. 289, 375–385. Nahin, R. L., Madsen, A. M., and Giesler, G. J. (1983). Anatomical and physiological studies of the gray matter surrounding the central canal. J. Comp. Neurol. 220, 321–335. Nahin, R. L., and Micevych, P. E. (1986). A long ascending pathway of enkephalin-like immunoreactive spinoreticular neurons in the rat. Neurosci. Lett. 65, 271–276. Nicholas, A. P., Zhang, X., and Hökfelt, T. (1999). An histochemical investigation of the opioid cell column in lamina X of the male rat lumbosacral spinal cord. Neurosci. Lett. 270, 9–12. Oscarsson, O. (1973). Functional organization spinocerebellar paths. In “Handbook of Sensory Physiology” (Iggo, A., Ed.), Vol. 2, pp. 340–380. Springer-Verlag, Berlin. Paxinos, G., and Watson, C. (1986). “The Rat Brain in Stereotaxic Coordinates.” Academic Press, Sydney. Rexed, B. (1952). The cytoarchitectonic organization of the spinal cord in the cat. J. Comp. Neurol. 96, 415–496. Rexed, B. (1954). A cytoarchitectonic atlas of the spinal cord in the cat. J. Comp. Neurol. 100, 297–379. Ritz, L. A., and Greenspan, J. D. (1985). Morphological features of lamina V neurons receiving nociceptive input in cat sacrocaudal spinal cord. J. Comp. Neurol. 238, 440–452. Rivero-Melián, C. (1996). Organization of hindlimb nerve projections to the rat spinal cord: A choleragenoid horseradish peroxidase study. J. Comp. Neurol. 364, 651–663. Rivero-Melián, C., and Grant, G. (1990). Lumbar dorsal root projections to spinocerebellar cell groups in the rat spinal cord: A double labeling study. Exp. Brain Res. 81, 85–94.

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Sato, H., Ohkawa, T., Uchino, Y., and Wilson, V. (1997). Excitatory connections between neurons of the central cervical nucleus and vesticular neurons in the cat. Exp. Brain Res. 115, 381–386. Scheibel, M. E., and Scheibel, A. B. (1968). Terminal axon patterns in cat spinal cord. II. The dorsal horn. Brain Res. 9, 32–58. Schoenen, J. (1982). The dendritic organization of the human spinal cord: The dorsal horn. Neuroscience 7, 2057–2087. Schokunbi, M. T., Hrycyshyn, A. W., and Flumerfelt, B. A. (1985). Spinal projections to the lateral reticular nucleus in the rat: A retrograde labelling study using horseradish peroxidase. J. Comp. Neurol. 239, 216–226. Silverman, J. D., and Kruger, L. (1988). Lectin and neuropeptide labeling of separate populations of dorsal root ganglion neurons and associated “nociceptor” thin axons in rat testis and cornea whole-mount preparations. Somatosens. Res. 5, 259–267. Silverman, J. D., and Kruger, L. (1990). Selective neuronal glycoconjugate expression in sensory and autonomic ganglia: Relation of lectin reactivity to peptide and enzyme markers. J. Neurocytol. 19, 789–801. Steiner, T. J., and Turner, L. M. (1972). Cytoarchitecture of the rat spinal cord. J. Physiol. (London) 222, 123–125. Swett, J. E., Wikholm, R. P., Blanks, R. H. I., Swett, A. L., and Conley, L. C. (1986). Motoneurons of the rat sciatic nerve. Exp. Neurol. 93, 227–252.

Szentágothai, J. (1964). Neuronal and synaptic arrangement in the substantia gelatinosa Rolandi. J. Comp. Neurol. 122, 219–239. Tavares, I., Lima, D., and Coimbra, A. (1993). Neurons in the superficial dorsal horn of the rat spinal cord projecting to the medullary ventrolateral reticular formation express c-fos after noxious stimulation of the skin. Brain Res. 623, 278–286. Wall, P. D. (1967). The laminar organization of dorsal horn and effects of descending impulses. Physiol. (London) 1888, 403–423. Wang, H., Rivero-Melián, C., Robertson, B., and Grant, G. (1994). Transganglionic transport and binding of the isolectin B4 from Griffonia simplicifolia I in rat primary sensory neurons. Neuroscience 62, 539–551. Wang, C.-C., Willis, W. D., and Westlund, K. N. (1999). Ascending projections from the area around the spinal cord central canal: A Phaseolus vulgaris leucoagglutinin study in rats. J. Comp. Neurol. 415, 341–367. Woodbury, C. J., Ritter, A. M., and Koerber, H. R. (2000). On the problem of lamination in the superficial dorsal horn of mammals: A reappraisal of the substantia gelatinosa in postnatal life. J. Comp. Neurol. 417, 88–102. Woolf, C. J., and Fitzgerald, M. (1983). The properties of neurones recorded in the superficial dorsal horn of the rat spinal cord. J. Comp. Neurol. 221, 313–328.

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C H A P T E R

6 Substantia Gelatinosa of the Spinal Cord ALFREDO RIBEIRO-DA-SILVA Departments of Pharmacology & Therapeutics and Anatomy & Cell Biology McGill University, Montréal, Québec, Canada

The superficial layers of the dorsal horn of the spinal cord and of the trigeminal subnucleus caudalis, particularly the substantia gelatinosa (or lamina II of Rexed), are the areas that have been traditionally associated with the modulation of nociceptive information since the classical clinical studies of Ranson (1914). Considerable attention to this area was triggered in the 1960s and 1970s by the publication of physiologically based pain theories (Cervero and Iggo, 1978; Melzack and Wall, 1965), which postulated the existence of synaptic circuits involving interneurons and afferent fibers conveying distinct inputs. These theories introduced the basic concept that nociceptive transmission can be modified by the concomitant activation of other fiber systems. In recent years, there has been considerable progress in understanding the anatomical and neurochemical characteristics of the relevant cells and systems. However, our knowledge of this area is still far from complete, and, unfortunately, too many oversimplified schemes can be found in textbooks and even in reviews. In this chapter, I provide an overview of the substantia gelatinosa of the spinal cord in the rat, with emphasis on its anatomical, ultrastructural, and immunocytochemical aspects. Although I will focus on lamina II, or the substantia gelatinosa proper, I will also describe briefly lamina I (marginal layer) and lamina III (superficial part of the nucleus proprius) because of their close interrelations and physiological relevance. Certain issues are discussed elsewhere in this volume and will be described here only very briefly. For an overview of the spinal cord cytoarchitecture see Grant and Koerber, Chapter 5. Readers interested in primary

The Rat Nervous System, Third Edition

sensory fibers should consult Grant and Robertson, Chapter. The ascending projections of the spinal cord are discussed briefly here. Readers interested in the ascending and descending pathways in the spinal cord are advised to consult the chapter by Tracey (see Chapter 7). An integrated view of pain mechanisms is presented in the chapter by Willis and collaborators (see Chapter 27).

DEFINITION The substantia gelatinosa of the spinal cord was given its name by Rolando in 1824 (quoted by Ramón y Cajal, 1909), based on the translucent and gelatinous appearance it possesses when examined in fresh tissue. In the cat, Rexed (1952) utilized 100-μm-thick freezing microtome sections stained for Nissl substance to subdivide the spinal cord into several horizontal laminae, based on cell density and size and the morphology of the Nissl bodies. Lamina II had a particularly high cellular density, because of the occurrence of many small neurons. Rexed made lamina II correspond to the substantia gelatinosa, which he divided into dorsal (more cellular) and ventral (less cellular and thicker) parts. More recent studies usually subdivide lamina II into outer lamina II (or lamina IIo) and inner lamina II (or lamina IIi). Rexed’s cytoarchitectonic classification has been adapted to the rat (see Grant and Koerber, Chapter 5) and to other species, such as the monkey (Ralston, III, 1979). The laminar pattern can also be recognized in samples examined with dark

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field (e.g., cryostat sections processed for receptor binding studies), stained with fiber stains (e.g., the Mahon method), or osmicated and Epon-embedded (see Light, 1992, for details). Using Epon-embedded 1or 2-μm-thick semi-thin sections, lamina I can be separated from outer lamina II by the abundant small myelinated fibers in the former; these are less abundant in outer lamina II and are almost entirely absent from inner lamina II. The lamina II–III border is easy to identify on semi-thin sections because of the numerous small myelinated fibers that occur in lamina III. At the light microscopic level, the subdivisions of the superficial laminae in the rat are therefore very similar to those observed in other mammals such as the cat and monkey. Unfortunately, the ultrastructural observation of the rat dorsal horn creates unexpected problems. In fact, in contrast with the dorsal horn of the monkey and cat (Ralston, III, 1968, 1979; Ribeiro-da-Silva and Coimbra, 1982), outer lamina II in the rat is virtually devoid of synaptic glomeruli (see below for details) (Ribeiro-da-Silva and Coimbra, 1982). Inner lamina II has a narrow dorsal band which is rich in synaptic glomeruli of type I (with an electron-dense central varicosity) and has very few of type II (which possess a light and large central varicosity) (Ribeiro-da-Silva and Coimbra, 1982). The more extensive ventralmost part of lamina II is rich in synaptic glomeruli of type II and has very few glomeruli of type I (Ribeiro-da-Silva and Coimbra, 1982). In contrast, in the cat and monkey, the glomeruli of the dense type prevail in lamina IIo (Knyihár-Csillik et al., 1982b; Maxwell et al., 1990). This difference in the distribution of synaptic glomeruli is very likely the result of interspecies differences in the distribution of primary sensory fibers. Therefore, while the cytoarchitectonically defined outer lamina II looks similar in rat, cat, and monkey, there are probably differences among species in primary afferent input. It is as if outer lamina II in the rat had certain features of lamina I. As a result of these interspecies differences in the distribution of synaptic glomeruli, I usually prefer to use an alternative nomenclature when dealing with rat dorsal horn: lamina IIA (instead of lamina IIo) and lamina IIB (instead of lamina IIi). Further, I subdivide lamina IIB into two sublaminae: sublamina IIBd (corresponding to the dorsalmost part of inner lamina II) and sublamina IIBv (corresponding to most of inner lamina II). In cross sections of the cervical dorsal horn (C4–C5 level) of young adult rats (200–250 g in weight), lamina I is approximately 20 μm thick, lamina IIA and sublamina IIBd are 20 μm each, and Sublamina IIBv is 40 to 60 μm-thick (Ribeiro-da-Silva and Coimbra, 1982). A diagram of these laminar subdivisions is shown in Fig. 1. At lumbar levels, which are frequently used for studies on animal models of chronic pain, the major

difference is that the thickness of the most superficial laminae is less in the lateral than in the intermediate and medial parts. However, at midlumbar levels, Todd et al. (1998) propose a lamina I in the middle much thicker than that in the lateral and medial parts of the dorsal horn. This view is based on the distribution of projection neurons and pattern of immunostaining for substance P receptors and does not follow the standard cytoarchitectonic criteria. Todd et al. (1998) compare this region of possibly thicker lamina I to the “dorsal cap” described in the cat by Snyder (1982). In my opinion, when defining the limits of the main laminae, it is important to stick to the parameters defined in studies using classical cytoarchitectonic methods, as outlined in the chapter by Grant and Koerber (see Chapter 5). Unfortunately, this is not often followed. As a consequence of the use of poorly defined criteria when delimiting dorsal horn laminae, many published micrographs and diagrams show a lamina I that is too thick and includes part of lamina II. One approach to define the laminae on sections processed for immunocytochemistry is to stain an adjacent section using a Nissl method. The Rexed lamination can be easily marked on a micrograph of the Nisslstained section and a transparency with the lamination overlaid on images from the sections that were immunostained. An example of the use of this approach is shown in Fig. 2.

CHARACTERISTICS OF NEURONS OF THE SUPERFICIAL LAMINAE OF THE SPINAL CORD Lamina I Lima and Coimbra (1986) have described four morphological types of neurons in the rat using the Golgi method, a classification still followed by most researchers. Fusiform neurons are elongated rostrocaudally and are more abundant in the lateral part of the lamina. Multipolar neurons have characteristically radiating dendritic trees and prevail in the medial part of the lamina. Pyramidal neurons have cell bodies of triangular shape and occur throughout the entire mediolateral extension of lamina I, always at the edge of the white matter. Flattened cells have dendritic trees that spread in the mediolateral and rostrocaudal axes. Cells of each of the four types are occasionally (6% of the total) two to three times the regular size (Lima and Coimbra, 1986). The larger versions of the pyramidal and flattened cells probably represent the classical Waldeyer cells (Lima and Coimbra, 1988; Puskar et al., 2001). Lamina I is considered an important

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FIGURE 1 Rexed’s laminae of the rat spinal cord at C4 level and details of subdivisions of lamina II. (A) On top, drawing of the limits of Rexed’s laminae. The framed area (enlarged underneath) represents the part that is equidistant from both the medial and the lateral edges of the dorsal horn. The equivalence of the two possible ways of subdividing lamina II is shown in the enlarged area. I have suggested a different nomenclature for the rat from that used for the cat, on the basis of ultrastructure and termination of sensory fibers. Panels B and C are shown at the same magnification as that of the lower part of panel A to facilitate the correlation of the lamination in Epon-embedded semi-thin section (B) and Nissl preparation (C). (B) Micrograph obtained from a 5-μm-thick plastic section of material incubated for the demonstration of acid phosphatase (FRAP) activity (arrows). Note that the FRAP-reactive band corresponds very closely to the limits of sublamina IIBd. (C) Micrograph originated from a 50-μm-thick section stained for Nissl. It shows the characteristic aggregation of neurons in lamina IIA. DC, dorsal columns. Scale bar (for both micrographs) = 20 μm.

projection area to higher structures. Main projection sites from lamina I are the thalamus (for reviews see Lima and Coimbra, 1988; Todd et al., 2000) and certain areas of the brain stem, particularly the lateral reticular nucleus, the parabrachial nucleus, and the periaqueductal gray matter (for reviews see Lima, 1997; Todd et al., 2000). Although it has been proposed, based on some experimental evidence, that the morphological types of lamina I neuron differ in their neurotransmitter/modulator content and supraspinal projection pattern (Lima, 1997), this issue remains controversial. In fact, evidence is accumulating that favors a correlation between the morphological and physiological properties of neurons in lamina I (Prescott and De Koninck, 2002).

Lamina II Despite several studies, our understanding of lamina II neurons in the rat is less than it is in the cat. In the latter species, lamina II cells have been extensively studied using anatomical and physiological approaches. Cells of lamina II have been classified since the work of Ramón y Cajal (1909) into two main morphological types: the central cell, which is widespread throughout the lamina, and the limiting cell, which occurs in an outer band close to the laminae I–II border. These types were identified by Gobel (1975, 1978) in the cat and named islet cells and stalked cells, respectively. In the rat, Todd and Lewis (1986) using the Golgi method confirmed the occurrence of

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FIGURE 2 Laminar distribution of CGRP and IB4 in the rat lumbar dorsal horn, as an example of the application of Rexed’s lamination to confocal microscope images. A section, adjacent to the one that was processed for immunocytochemistry, was stained with a Nissl method and the laminar limits were marked on a transparent sheet. This was placed over the images at the same magnification as that obtained with the confocal microscope. The limit of lamina IIBd was defined based on our previous work that shows that its thickness is approximately the same as that of lamina IIA. Both panels A and B were derived from the same confocal micrograph, corresponding to an optical section about 1 μm in thickness, after elimination in the confocal microscope software of the display corresponding to the other channel. In A, corresponding to immunostaining for CGRP, note that the fiber terminal density is high in laminae I–IIBd and low in lamina IIBv. In B, corresponding to lection IB4 binding, note that the highest density of fiber terminals is in the middle third of lamina II.

both stalked and islet cells in lamina II. The stalked cells corresponded to half of the stained cells in the outer part of lamina II, whereas islet cells were found throughout the entire lamina and corresponded to about one third of the entire stained neuronal population. However, Todd and Lewis (1986) also reported that about half of the cells in lamina IIBv could not be classified as either stalked or islet cells, although they could be subdivided into groups based on their dendritic arborization. The axons of these cells either passed to lamina III or remained in lamina II. Some of these cells may correspond to the stellate and LII–III border cells previously described in other species (Todd and Lewis, 1986). In the cat, both islet and stalked cells were electrophysiologically characterized, filled with horseradish peroxidase (HRP), and studied at the light and electron microscopic levels by Bennett, Gobel, and collaborators (Bennett et al., 1980; Gobel et al., 1980). At least some stalked cells, with an axonal arborization in lamina I, seem to relay excitatory impulses to lamina I cells and, therefore, represent feed-forward excitatory interneurons. The main electrophysiological findings have been that the physiological properties of islet cells differed according to their localization, as those situated in deep lamina II did not respond to noxious stimuli while those in outer lamina II responded specifically to these stimuli. This agrees with previous studies in the cat by Light and collaborators (1979) who found that the cells in the outer half of lamina II responded to noxious cutaneous stimuli while those in the inner half

of lamina II only responded to innocuous stimuli. Also in the cat, when examining lamina II cells, the type of response elicited seemed to have little correlation with their morphology (light et al., 1979) but depended more on the localization of the dendritic arborization. As dendrites receive most of the information from incoming fibers, the fibers which terminate deeply in lamina II seemed not to transmit nociceptive information in the cat (Light and Perl, 1979). However, this is not likely the case in the rat because of the differences in the termination pattern of sensory fibers. In fact, as explained below, the nonpeptidergic subpopulation of small-diameter sensory fibers in rodents terminates mostly in the outermost part of inner lamina II (sublamina IIBd), and the available evidence indicates that these fibers are nociceptive (Alvarez and Fyffe, 2000; Snider and McMahon, 1998). In agreement with this, studies in C fibers which combine intracellular recording with intracellular injection with a marker revealed a considerable termination of unmyelinated polymodal nociceptive fibers in a certain region of ventral lamina II in the guinea pig (Sugiura et al., 1986). The available evidence indicates that in animals such as the guinea pig and the rat, unmyelinated fibers terminate deeper in lamina II than they do in cats and monkeys. Originally, lamina II was considered to be a closed system (Szentágothai, 1964), receiving afferents but not projecting to any area of the brain. However, there is now evidence that a small number of lamina II neurons project to the brain (thalamus, lateral cervical

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nucleus, or pontine–medullary junction) (for review see Willis and Coggeshall, 1991). One study claims that a considerable number of islet cells project to the reticular formation of the medulla (Lima and Coimbra, 1991).

Lamina III In the cat, neurons of lamina III have been described as a heterogeneous population of nonnociceptive cells (Maxwell et al., 1983), based on intracellular injections of physiologically characterized neurons. However, the concept that the cells are all nonnociceptive needs revision at least for the rat, based on the detection in laminae III and IV of neurons that express the substance P receptor and possess dorsally oriented dendrites that branch in laminae I and II (Brown et al., 1995; Littlewood et al., 1995; Liu et al., 1994a; Naim et al., 1997). Most of these neurons project to supraspinal levels (Todd et al., 2000). Little is known concerning the other lamina III neuronal populations in rat.

ULTRASTRUCTURE OF THE SPINAL DORSAL HORN The features that allow the characterization of each lamina and sublamina under the electron microscope are the density of small myelinated fibers and the distribution of synaptic glomeruli (see above). Readers interested in a detailed description of the general ultrastructural characteristics of each lamina and sublamina should consult Ribeiro-da-Silva and Coimbra (1982). In this chapter, we focus on synaptic glomeruli.

Synaptic Glomeruli The ultrastructure of the spinal and medullary dorsal horn has been studied in detail in the rat (Coimbra et al., 1974; Ribeiro-da-Silva et al., 1985; Ribeiro-da-Silva and Coimbra, 1982). The most striking ultrastructural feature of the dorsal horn is the presence of synaptic glomeruli, which are complex synaptic arrangements in which a “central” (core) axonal bouton is surrounded by several dendrites and axonal boutons (surrounding boutons). The core (C) axonal bouton is of primary sensory origin, as demonstrated by studies showing their degeneration after multiple dorsal root transections or labeling after the injection of tracers (Coimbra et al., 1984; Cruz et al., 1987). The C bouton interacts with the dendrites of spinal cord interneurons (Gobel et al., 1980) or projection neurons (Maxwell et al., 1985). Some of these dendrites contain synaptic vesicles (presynaptic dendrites) and are presynaptic to the C bouton and/or to other dendrites (Gobel, 1976;

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Ribeiro-da-Silva et al., 1985). Surrounding glomerular axonal boutons, most likely originating from neurons intrinsic to the dorsal horn (see below under Neurochemistry of the Dorsal Horn), are presynaptic to the glomerular C boutons and to glomerular dendrites. Synaptic glomeruli are thought to play an important role in sensory mechanisms because they constitute a significant part of the synaptic population of the superficial dorsal horn (Duncan and Morales, 1978; Murray and Goldberger, 1986) and display complex synaptic arrangements (Gobel, 1976; Knyihár-Csillik et al., 1982a; Ribeiro-da-Silva et al., 1985). See Fig. 3 for details of the ultrastructure of a synaptic glomerulus. The literature does not have a consistent definition of what should be considered a synaptic glomerulus. However, a proper definition is essential, as glomeruli can be excellent markers to identify the termination of sensory fibers at the ultrastructural level. Terminals of fibers originating from the brain stem or from neurons intrinsic to the spinal cord (identified by antigenic markers such as serotonin and GABA) are sometimes the core element of synaptic arrangements that are simpler than synaptic glomeruli. On an isolated electron micrograph, a complex synaptic arrangement can be classified as a synaptic glomerulus if it meets all of the following criteria: (a) it must have a C bouton possessing agranular round synaptic vesicles, (b) the C bouton must be in apposition to at least four “surrounding” dendritic profiles (one or more can be replaced by axonal boutons and presynaptic dendrites), and (c) two or more synaptic specializations must be found between C and surrounding profiles. Types of surrounding profiles are: (1) dendrites devoid of synaptic vesicles (“plain” or “common” dendrites – D), (2) vesicle-containing or presynaptic dendrites (V1), and (3) surrounding axonal boutons (V2). In the rat (but not in the cat or monkey), lamina I has very few synaptic glomeruli. Glomeruli become abundant only in lamina IIB, particularly in sublamina IIBd. Glomeruli are rather frequently encountered in lamina III.

Types of Synaptic Glomeruli Two main types of synaptic glomeruli have been described in the rat (Ribeiro-da-Silva and Coimbra, 1982). Type I glomeruli possess a relatively small C bouton of scalloped contour, with closely packed synaptic vesicles and very few mitochondria (Fig. 3). Two varieties can be described. Glomeruli of type Ia (or type I “nonpeptidergic”) have a particularly electrondense C bouton, with vesicles displaying a very wide variation in diameters, and have on average one V 1 and one V2 terminal per glomerulus. Glomeruli of type Ib (or type I “peptidergic”) possess more than three

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FIGURE 3 Diagrammatic representation of a synaptic glomerulus. The drawing was based on an electron micrograph of a type Ia glomerulus and shows the morphological features and the synaptic circuits involving the core and surrounding profiles. C, central or core bouton; D, “regular” dendrite; V1, presynaptic dendrite; V2, peripheral axonal bouton; G, glial profile.

dense-core vesicles in the C bouton, are immunoreactive for sensory peptides, and have simplified synaptic architecture (virtually all surrounding profiles are dendrites postsynaptic to the central bouton). Type II glomeruli have a larger C bouton, of less scalloped contour, which is lighter and richer in mitochondria than their type I counterparts. Furthermore, type II glomeruli are richer in surrounding axonal boutons (V2) than type I glomeruli. Two varieties of type II glomeruli can be recognized (Fig. 4): type IIa (devoid of neurofilament bundles in the C bouton) and type IIb (with neurofilament bundles in the C bouton). Type IIb glomeruli are particularly rich in V2 boutons. For a more detailed description of glomerular types the reader should consult Ribeiro-da-Silva et al. (1985, 1989). Figure 4 displays diagrammatically the glomerular types in lamina II and gives the relative frequency in which they occur at cervical levels C4–C5 in rat.

Functional Role of Synaptic Glomeruli The functional role of glomeruli is far from known. Most varicosities in primary sensory fibers are unrelated to glomeruli (see e.g., Coimbra et al., 1984). However, the available evidence strongly indicates that glomeruli are “multiplier systems,” i.e., devices via which primary

sensory information is transmitted to several dorsal horn neurons by means of a single axonal bouton. In turn, synaptic glomeruli are important integrators, being often postsynaptic to other neuronal profiles. Therefore, synaptic glomeruli are very important elements in sensory transmission. Most likely, the C boutons of type I glomeruli (CI) represent unmyelinated nociceptive fibers, because they correspond to the termination of fibers that are capsaicin-sensitive (Ribeiroda-Silva and Coimbra, 1984). However, capsaicin also damages the smaller Aδ fibers; therefore some CI boutons may represent the termination of Aδ fibers. It is tempting to state that all C boutons of type I glomeruli represent the termination of nociceptive sensory fibers. If this is the case, then type I glomeruli are of the utmost importance for the transmission of pain-related information. Most peptidergic small-diameter primary afferents are nonglomerular. However, about 20% of type I glomeruli are of the peptidergic type. These peptidergic (or Type Ib) glomeruli are most likely only multiplier systems, because their peptide-containing core boutons share an important characteristic with nonglomerular endings of the same fiber population: the fact that they are virtually never postsynaptic to other neuronal profiles. This is in complete contrast with the arrangement of type I glomeruli of the

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FIGURE 4 Diagrammatic representation of the morphological properties and relative incidence of the several types of synaptic glomeruli in lamina II.

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Results of the colocalization of substance P and CGRP were based on a double-labeling study involving a combination of an anti-substance P internally radiolabeled monoclonal antibody with anti-CGRP antibody (Ribeiro-da-Silva, 1994); results of the colocalization of CGRP and somatostatin were based on a study combining preembedding (somatostatin) and postembedding (CGRP) immunocytochemistry (Ribeiro-da-Silva, 1994). As recent studies have shown that boutons colocalizing SOM and CGRP also bind IB4 (see, e.g., Alvarez and Fyffe, 2000), a dashed arrow was added to the figure to illustrate that very likely the subdivision between peptidergic and nonpeptidergic type I glomeruli is not absolute. D, “regular” dendrite; V1, presynaptic dendrite; V2, peripheral axonal bouton; G, glial profile; neurofil., neurofilaments; glom., glomerular; FRAP, fluoride-resistant acid phosphatase; SOM, somatostatin; CGRP, calcitonin gene-related peptide; SP, substance P.

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nonpeptidergic type (type Ia) in which both presynaptic dendrites and peripheral axons are presynaptic to the core bouton and therefore are likely also very important integrating devices. These nonpeptidergic terminals correspond to the fibers that express the P2X3 receptor and bind the lectin IB4. In relation to type II glomeruli, we can extrapolate from ultrastructural studies of physiologically characterized fibers in the cat which show glomerular morphology very similar to that of the C boutons of Type II glomeruli (CII) (for review see Maxwell and Réthelyi, 1987). Therefore, the C boutons of type IIa glomeruli probably represent the termination of Aδ D-hair fibers, and those of type IIb (with neurofilaments) the termination of thicker fibers. In the rat, both varieties of type II glomeruli display rather complex synaptic arrangements, involving both presynaptic dendrites (V1) and surrounding axonal boutons (V2 in type IIa or V2 in type IIb).

Electron Microscopic Properties of Lamina II Neurons Stalked and islet cells were studied at the ultrastructural level by Todd (1988), using the Golgi method. As previously demonstrated in the cat, stalked cells did not give origin to presynaptic dendrites, in contrast with islet cells. Both cell types participated in synaptic glomeruli through their dendritic processes.

NEUROCHEMISTRY OF THE DORSAL HORN Since the publication of the previous edition of this book, significant advances have been made in our understanding of the chemical substances that occur in the superficial laminae of the dorsal horn. Unfortunately, there are no recent reviews on the topic. What follows is a brief integrated overview of the most studied neurochemicals in the region and, when applicable, of their respective receptors.

nated or thinly myelinated sensory fibers that terminate mainly in laminae I and II (Cuello and Kanazawa, 1978; Hökfelt et al., 1975). Immunoreactivity for substance P is particularly intense in lamina I and outer lamina II, but decreases substantially in inner lamina II (Ribeiro-daSilva et al., 1989). In lamina III, SP immunoreactivity is reduced even further and represents mostly fibers crossing toward deeper laminae. In laminae IV–V, there are clusters of substance P-immunoreactive (IR) fibers and boutons separated by areas with sparse immunoreactivity (Ruda et al., 1986). It should be clearly stated that, contrary to common belief, not all substance P immunoreactivity in the superficial dorsal horn is of primary sensory origin as multiple dorsal rhizotomies and capsaicin treatment are unable to fully deplete substance P immunoreactivity. Furthermore, substance P-containing cell bodies have been identified in spinal laminae I and II, both with immunocytochemistry (Ljungdahl et al., 1978; Ribeiro-da-Silva et al., 1991) and in situ hybridization (Warden and Young, 1988). Also, although most substance P-containing systems descending from the brain stem terminate in the ventral horn (Gilbert et al., 1982; Hökfelt et al., 1978), some may terminate in the superficial laminae of the dorsal horn. It is interesting to note that most, if not all, substance P-IR cell bodies in the spinal dorsal horn colocalize enkephalin immunoreactivity (Ribeiro-da-Silva et al., 1991). At the ultrastructural level, substance P immunoreactivity in the central boutons of synaptic glomeruli is particularly meaningful, because such profiles are of known sensory origin (Coimbra et al., 1984; Murray and Goldberger, 1986). Substance P immunoreactivity has also been detected in glomerular C boutons in several animal species, including the rat (Ribeiro-daSilva et al., 1989; Ribeiro-da-Silva and Cuello, 1987). In the rat, substance P immunoreactivity has been detected in 10% of the C boutons of synaptic glomeruli in lamina II (Figs. 4 and 5A). All these glomerular boutons had large dense-core vesicles (characteristic of glomeruli of the type Ib variety).

Other Neurokinins Neurokinins The three main mammalian neurokinins are substance P, neurokinin A, and neurokinin B. They all occur in the superficial laminae of the dorsal horn. Substance P There is now unquestionable evidence of the involvement of the neurokinin substance P in the processing of sensory information in the region of the first sensory synapse (for reviews see Cuello, 1987; Henry, 1982; Otsuka and Yanagisawa, 1990). Immunocytochemically, substance P has been shown to occur in either unmyeli-

Virtually all substance P-containing neurons in the rat express precursors that produce both substance P and neurokinin A (Carter and Krause, 1990) which means that their distribution is essentially the same. However, neurokinin B derives from a different precursor and, in contrast to substance P, it does not occur in primary sensory neurons (Ogawa et al., 1985). A recent light and electron microscopic immunocytochemical study of neurokinin B in the dorsal horn of the spinal cord revealed that, in the superficial laminae, its signal was detected in axon terminals in laminae I–II, with a peak in lamina IIB, as well as in cell bodies and dendrites

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mostly in lamina IIB (McLeod et al., 2000). Lamina III showed much less immunolabeling. So, in contrast to substance P, immunoreactivity for neurokinin B increased from lamina I to lamina IIB. Interestingly, neurokinin B immunoreactivity occurred in dendrites of type I glomeruli, suggesting a participation in the modulation of nociception (McLeod et al., 2000). Neurokinin Receptors The original descriptions of substance P receptor (neurokinin-1 receptor) immunoreactivity in the dorsal horn reported that it was present in neurons with cell bodies located in lamina I and in deeper layers (laminae III–IV) (Liu et al., 1994a; Nakaya et al., 1994). According to these reports, the substantia gelatinosa proper did not have any cell bodies immunoreactive for the neurokinin-1 receptor (NK-1r) and possessed much less immunoreactivity than lamina I, as this was restricted to cell processes mostly from neurons in deeper laminae. However, more recent studies reported immunoreactivity for the NK-1r in cell bodies in both lamina I and outer lamina II (LIIA) (McLeod et al., 1998; Ribeiro-daSilva et al., 2000). It should be pointed out that most of these NK-1r-IR neurons project to higher levels: the thalamus (Marshall et al., 1996), the parabrachial nucleus (Ding et al., 1995; Todd et al., 2000), the lateral reticular nucleus, the dorsal part of caudal medulla, and, to a minor extent, the periaqueductal gray (Todd et al., 2000). In contrast with the NK-1r, the receptor for neurokinin A (neurokinin-2 receptor) hardly occurs in the CNS (for review see Ribeiro-da-Silva et al., 2000), indicating that either neurokinin A acts through another receptor in the CNS or it acts mainly in the periphery. In the superficial laminae of the dorsal horn, some immunoreactivity for the neurokinin-2 receptor was detected in a narrow band in the lateral part of lamina I, but seemed to be located in glial cells (Zerari et al., 1998). Concerning the preferential receptor for neurokinin B, the neurokinin-3 receptor, it occurs in cell bodies located in laminae I and, mostly, in lamina II of the spinal cord (Ding et al., 1996; Mileusnic et al., 1999; Zerari et al., 1997).

Calcitonin Gene-Related Peptide (CGRP) ‘Immunoreactivity for CGRP has been shown to occur in dorsal root ganglia and in primary sensory fibers which project mainly to the superficial laminae of the spinal cord (Ju et al., 1987; Wiesenfeld-Hallin et al., 1984). In the dorsal horn, CGRP-IR boutons occur mostly in laminae I, IIA, and IIBd (Fig. 2), as well in patches in lamina V. One of the interesting features of CGRP immunoreactivity in sensory systems is its colocalization with substance P (Ju et al., 1987; Wiesenfeld-Hallin et al., 1984). In reality, substance P immunoreactivity is almost invariably colocalized with CGRP in dorsal root

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ganglion cells, although CGRP immunoreactivity occurs in a considerably higher percentage of these cells than substance P (Ju et al., 1987). Another interesting feature of CGRP immunoreactivity in the dorsal horn is its almost complete disappearance after dorsal rhizotomy (Chung et al., 1988; Traub et al., 1989). This finding suggests that all CGRP immunoreactivity in the dorsal horn originates from primary sensory fibers, a finding that is confirmed by in situ hybridization studies which do not reveal any dorsal horn neurons synthesizing the peptide (Réthelyi et al., 1989). Therefore, it seems legitimate to use the colocalization of CGRP and substance P in the same terminal as a marker of primary sensory origin. At the ultrastructural level, CGRP occurs mostly in nonglomerular varicosities in the dorsal horn of the rat spinal cord, although a few varicosities are of the glomerular type (Merighi et al., 1989, 1991; Ribeiro-daSilva and Cuello, 1991). Most CGRP immunoreactivity in the dorsal horn is colocalized with either substance P (Fig. 5A) or somatostatin immunoreactivities (Fig. 5B) (Ribeiro-da-Silva, 1994). The distribution of CGRP receptors in the spinal cord has been studied with ligand binding approaches. They occur in high densities in lamina I and in deeper laminae but occur in low densities in lamina II (Yashpal et al., 1992). However, following peripheral denervation, considerable CGRP binding was detected in lamina II, indicating that the neurons have the capacity to produce the receptor (Kar et al., 1994)

Somatostatin Somatostatin-like immunoreactivity occurs both in primary sensory fibers and in neurons of the spinal cord (Alvarez and Priestley, 1990a; Hökfelt et al., 1976; Ribeiro-da-Silva and Cuello, 1990b). In the superficial laminae, somatostatin-IR neurons occur mainly in lamina II (Alvarez and Priestley, 1990a; Ribeiro-daSilva and Cuello, 1990b). Somatostatin receptors form a family of five receptors (sst1 to sst5), all belonging to the G protein-coupled receptor superfamily (Dournaud et al., 2000). Immunoreactivity for receptor subtypes has been detected in cell bodies and processes in the superficial laminae of the dorsal horn (Schulz et al., 1998; Von Banchet et al., 1999).

Opioid Peptides Enkephalin Since their discovery, endogenous opioid peptides have been considered important candidates for presynaptic interactions in the dorsal horn of the spinal cord. The opioid peptides met- and leu-enkephalin occur in high concentrations in lamina I and II of the spinal

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FIGURE 5 Examples of neurochemicals and their colocalizations in synaptic glomeruli as detected by immunocytochemistry. A Core (C) bouton of a type Ib synaptic glomerulus colocalizing CGRP (immunogold particles) and substance P (dense precipitate). Note the immunogold-labeled dense-core vesicles (arrowheads) in the C bouton (CIb). The C bouton was never found postsynaptic to surrounding profiles in this peptidergic glomerulus. Sublamina IIBd. (B) Core (C) bouton of a type Ib synaptic glomerulus colocalizing CGRP (immunogold particles) and somatostatin (dense precipitate). Note the immunogold-labeled dense-core vesicles (arrowheads) in the C bouton (C Ib). As in A, the C bouton was never found postsynaptic to surrounding profiles in this peptidergic glomerulus. Sublamina IIBd. (C) Immunocytochemistry of GABA as demonstrated by means of an anti-GABA polyclonal antibody revealed by an anti-rabbit IgG conjugated to 10-nm gold particles (postembedding protocol). Two type I glomeruli of the nonpeptidergic subtype are shown. Note the intrinsic electron-dense core boutons (CIa) typical of this glomerular variety. Some of the glomerular surrounding profiles show GABA immunoreactivity (gold particles). D, “regular” dendrite; V1, presynaptic dendrite; V2, surrounding axonal bouton. GABA+, profiles possessing GABA immunoreactivity. Sublamina IIBd. Scale bars (in both micrographs) = 1 μm.

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cord (Höfelt et al., 1977; Hunt et al., 1981) and have been demonstrated in nerve cell bodies in laminae I–III (Del Fiacco and Cuello, 1980; Hunt et al., 1981; Miller and Seybold, 1989). Double-labeling studies combining radioimmunocytochemistry and DAB-based immunocytochemistry have demonstrated that sometimes enkephalin and substance P-IR varicosities establish separate synapses on a common dendrite and that substance P-IR C glomerular boutons are presynaptic to enkephalin-IR dendrites in the rat (Cuello, 1983; Ribeiro-da-Silva et al., 1991). Enkephalin-IR boutons were never presynaptic to substance P-IR boutons (Ribeiro-da-Silva et al., 1991). Such results demonstrate (together with data indicated below) that substance Pcontaining glomerular C boutons excite dendrites of enkephalinergic interneurons in the substantia gelatinosa and that the axons of such neurons inhibit the dendrites of neurons that have been excited by substance P. The discovery of substance P and enkephalin colocalization in a considerable number of neurons and axonal varicosities in both rat and cat (Ribeiro-daSilva et al., 1991; Senba et al., 1988; Tashiro et al., 1987) added a new dimension to the problem. In reality, almost all substance P-IR neurons in the rat dorsal horn colocalize enkephalin and approximately 50% of enkephalin-IR cells colocalize substance P (Ribeiro-daSilva et al., 1991; Senba et al., 1988). It seems likely that most of the enkephalin immunoreactivity comes from neurons intrinsic to the dorsal horn. Enkephalin has been localized in serotonergic neurons of the raphe nuclei that project to the spinal cord, but most such fibers terminate in the ventral horn (Menétrey and Basbaum, 1987; Tashiro et al., 1988). Also, some enkephalin immunoreactivity may originate from primary sensory fibers. However, enkephalin has never been detected in a significant number of neurons in the dorsal root ganglia. Based on the above, it is clear that the colocalization with enkephalin can be used as a marker for substance P immunoreactivity in nerve terminals of dorsal horn origin. In the cat, enkephalin-IR boutons have been shown to synapse on spinothalamic neurons (Ruda et al., 1984) and on neurons of the dorsal column postsynaptic pathway (Nishikawa et al., 1983). Dynorphins Dynorphin immunoreactivity has been detected in neurons of laminae I and II (Miller and Seybold, 1987). Endormorphins Of the two endomorphins, endomorphin-2 is the most abundant in the superficial laminae of the dorsal horn, where it occurs in laminae I and IIA with a distribution similar to that of substance P, with which it is colocalized in sensory fibers (Martin-Schild et al., 1997,

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1998, 1999). In contrast, endomorphin-1 is intrinsic to the CNS and occurs in fibers in laminae I and II (MartinSchild et al., 1999). Opioid Receptors As dorsal rhizotomy leads to a reduction of both μand δ-opioid receptor binding (Fields et al., 1980), it has been postulated that such receptors should be, at least in part, localized in primary sensory fibers. In situ hybridization cytochemistry has confirmed the occurrence of opioid receptors in dorsal root ganglia neurons and in dorsal horn neurons (Minami and Satoh, 1995). Combined light and electron microscopic studies utilizing antibodies against the δ-opioid receptor have shown conflicting results. One group claims that the receptors occur both on cell bodies and dendrites of dorsal horn neurons and in axon terminals (Cheng et al., 1995, 1997). Surprisingly, another group has found the δ-opioid receptor associated mainly with densecore vesicles on sensory fibers and not with the plasma membrane, as expected for a G-protein-coupled receptor (Zhang et al., 1998). The receptor has been shown to be colocalized in terminals with enkephalin (Cheng et al., 1995) and in sensory fibers with substance P (Zhang et al., 1998). In contrast, the κ receptor has been localized mostly postsynaptically (Arvidsson et al., 1995), whereas μ receptor immunoreactivity occurs mostly in lamina II (Honda and Arvidsson, 1995; Kemp et al., 1996), in axon terminals, in dendritic profiles, and in cell bodies of dorsal horn neurons (Cheng et al., 1996, 1997). Interestingly, the great majority of cell bodies immunoreactive for the μ-opioid receptor, which were mostly located in lamina II, did not contain either GABA or glycine immunoreactivities, suggesting that the neurons that express the μ receptor might be mostly excitatory interneurons (Kemp et al., 1996).

Glutamate It has been shown by immunocytochemistry that glutamate is localized in virtually all sensory fibers and that at the ultrastructural level it occurs in virtually all the central varicosities of glomeruli, supporting the hypothesis that glutamate is the fast excitatory transmitter of primary sensory fibers (Battaglia and Rustioni, 1988; De Biasi and Rustioni, 1988). Glutamate and substance P have been shown to be colocalized in a considerable number of dorsal root ganglia cells (Battaglia and Rustioni, 3 1988) and terminals in the dorsal horn (De Biasi and Rustioni, 1988). Aspartate is colocalized with glutamate in some of these sensory fibers, particularly in those of small diameter (Tracey et al., 1991). Glutamate receptors have been studied in the dorsal horn by applying receptor binding and in situ hybridization

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(Henley et al., 1993), as well as immunocytochemistry. The latter studies have shown that AMPA receptors have a widespread distribution in cells of the dorsal root ganglia and dorsal horn (Tachibana et al., 1994). One study has shown that Ca2+-permeable AMPA receptors were localized mostly in GABAergic inhibitory interneurons and NK-1r-IR neurons (Albuquerque et al., 1999). Interestingly, NMDA receptors have been shown to occur also in sensory fibers using light and electron microscopy (Liu et al., 1994b), a localization that has also been described for glutamate metabotropic receptors (Ohishi et al., 1995).

Inhibitory Amino Acids GABA and Glycine It has been shown that some neurons are immunoreactive for GABA or GAD or take up [3H]GABA and are assumed to be GABAergic (Barber et al., 1982; Hunt et al., 1981; Ribeiro-da-Silva and Coimbra, 1980; Todd and McKenzie, 1989), while others specifically incorporate [3H]glycine or are immunoreactive with antibodies against glycine and are considered to be glycinergic (Ribeiro-da-Silva and Coimbra, 1980; Todd and Sullivan, 1990). By combining immunocytochemistry with Wallerian degeneration in the rat (Barber et al., 1978) or with the intracellular filling of identified sensory fibers in the cat (Maxwell and Noble, 1987), GABAergic neurons have been shown to be presynaptic to primary sensory boutons (Fig. 5C). The available evidence indicates that GABA, like enkephalin, is present mostly in local circuit neurons, and a colocalization of both neurochemicals has been demonstrated at the light microscopic level in the superficial dorsal horn (Todd et al., 1992). We have confirmed this finding at the ultrastructural level (see Fig. 6). Furthermore, it has been suggested that virtually all glycinergic neurons in laminae I–III are also GABAergic (Todd, 1991; Todd and Sullivan, 1990). However, only about half of the GABAergic cells colocalize glycine (Todd, 1991; Todd and Sullivan, 1990). Recent evidence also indicates the costorage of GABA and glycine within the same vesicles at synapses in the superficial dorsal horn (Chéry and De Koninck, 1999). Glycinergic varicosities, like the GABAergic ones, can be presynaptic to primary sensory fibers in glomeruli (Todd, 1996). GABA and Glycine Receptors GABAA receptors have been described by in situ hybridization methods in cells of both the dorsal root ganglia and the spinal cord (Persohn et al., 1991), supporting the morphological finding that GABA-IR fibers are frequently presynaptic to sensory fibers (see above). Using immunocytochemistry, GABAA receptor sub-

units were shown to occur in laminae I–III (Alvarez et al., 1996; Bohlhalter et al., 1996). An ultrastructural study with an antibody generated against the GABAA receptor subunits β2–β3 revealed that most of the immunostaining was localized to dendrites and cell bodies, although some central elements of glomeruli were also labeled, confirming that the receptor also occurs in primary sensory neurons (Alvarez et al., 1996). The light microscopy distribution of GABAB receptor immunoreactivity has recently been described in the spinal cord and was highest in laminae I and II, where it occurred both in cell bodies and in the neuropil (Margeta-Mitrovic et al., 1999). In contrast to GABA receptors, glycine receptors are restricted to dorsal horn neurons. Gephyrin (a glycine receptor-associated protein) has been found postsynaptic to boutons immunopositive for GABA (Mitchell et al., 1993), a finding that was to be expected as many neurons colocalize GABA and glycine immunoreactivities (see above). The specificity of the mixed GABA/glycine synapses in the superficial dorsal horn appears to be determined by the expression, properties, and subsynaptic localization of the target GABAA, GABAB, and glycine receptors (Chéry and De Koninck, 1999, 2000) and to change during development (Keller et al., 2001).

Other Classical Transmitters and Other Neuropeptides Cell bodies immunoreactive for choline acetyltransferase (ChAT) have also been described in this area of the central nervous system (CNS) (Barber et al., 1984; Kimura et al., 1981; Todd, 1991). Such cholinergic neurons occur mainly in laminae III–IV and are presynaptic to primary sensory fibers in synaptic glomeruli and to cells of the dorsal horn (Ribeiro-da-Silva and Cuello, 1990a). A study from my laboratory has demonstrated that most of these ChAT-IR neuronal cell bodies (Fig. 7A) and boutons (Figs. 7B–7D) colocalize GABA immunoreactivity. Serotonin originates from cell bodies located in the brain stem (for review see Ruda et al., 1986). In the cat, retrograde tracing has shown that serotonin-IR profiles have direct contacts with projection neurons (Ruda, 1986). Despite two ultrastructural studies, very little is known concerning the synaptic contacts of noradrenergic fibers in the dorsal horn, except that they are presynaptic to dorsal horn neurons (Doyle and Maxwell, 1991; Hagihira et al., 1990). However, the light microscopic distribution of noradrenergic fibers in the dorsal horn and their origin in the brain stem are well known (Fritschy and Grzanna, 1990; Westlund et al., 1983). Neurotensin immunoreactivity occurs in neurons in laminae I and II (Hunt et al., 1981; Seybold and Elde, 1982).

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FIGURE 6 Examples of colocalization of GABA and enkephalin (ENK) in laminae II–III as detected by immunocytochemistry. (A) ENK/GABA colocalization in a presynaptic dendrite (V1) of a type Ia synaptic glomerulus; note another presynaptic dendrite possessing exclusively GABA immunoreactivity (V1 GABA+). (B) ENK/GABA colocalization in a peripheral axon (V2 profile) of a type Ia glomerulus; note a presynaptic dendrite (V1 GABA +) which is immunoreactive for GABA only. (C, D) Colocalization of ENK and GABA immunoreactivities in V2 profiles of type II glomeruli. D, “regular” dendrite; V1, glomerular presynaptic dendrite; V2, glomerular peripheral axon; CIa, central varicosity of type Ia glomerulus; CIIa, central varicosity of type IIa glomerulus; CIIb, central varicosity of type IIB glomerulus. Scale bars = 0.5 μm.

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FIGURE 7 Examples of colocalization of ChAT and GABA immunoreactivities in laminae II–III, as detected by immunocytochemistry. (A,B) In nonglomerular profiles, (C,D) in synaptic glomeruli. (A) Nerve cell body (CB) in lamina III colocalizing ChAT (immunoprecipitate) and GABA (gold particles) immunoreactivities. (B) ChAT/GABA colocalization in an axonal bouton (arrow) in lamina IIA. (C) GABA+ChAT colocalization in a peripheral axon (V2 profile) of a type I synaptic glomerulus; note a presynaptic dendrite (V1) which is immunoreactive exclusively for GABA. (D) Two V2 profiles in a type II glomerulus colocalizing ChAT/GABA immunoreactivities; a third V2 profile possesses exclusively GABA immunoreactivity. (D), dendrite; V1, glomerular presynaptic dendrite; V2, glomerular peripheral axon; CIa, central varicosity of type Ia glomerulus; CIIb, central varicosity of type IIb synaptic glomerulus. Scale bars = 0.5 μm.

Markers of Nonpeptidergic Primary Sensory Fibers Since a seminal article by Hunt and Rossi in 1985 (Hunt and Rossi, 1985), the concept of the occurrence of two populations of sensory fibers conveying nociceptive information, the peptidergic and the nonpeptidergic, has emerged. The first express sensory neuropeptides (in particular substance P), and the second display fluoride-resistant acid phosphatase (FRAP)

activity. This concept was largely neglected for a decade while investigators focused mostly on the terminations of fibers expressing sensory neuropeptides, in particular substance P. However, interest in the concept has been revived in recent years. It was clarified that the population that expressed FRAP activity, originally described a few years earlier by two groups independently (Coimbra et al., 1970, 1974; Knyihár, 1971; Knyihár and Gerebtzoff, 1973), could specifically bind the isolectin IB4 and be recognized by the monoclonal antibody

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LA4 (Alvarez et al., 1989a,b; Dodd and Jessell, 1985; Jessell and Dodd, 1985, 1989). However, the real interest in the nonpeptidergic population emerged following the discovery that the two populations differed in neurotrophic support in the adult. In fact, during development, both populations require nerve growth factor (NGF) for survival, but shortly after birth only the peptidergic type continue to respond to NGF, whereas the nonpeptidergic population starts to respond instead to glial cell line-derived neurotrophic factor (GDNF) (Bennett et al., 1998). Accordingly, the peptidergic population expresses the NGF high-affinity receptor trkA, whereas the nonpeptidergic expresses GDNF receptors. It was shown that the latter population also expressed the purinergic receptor P2X3 (Bradbury et al., 1998; Snider and McMahon, 1998) and the capsaicin VR1 receptor (Guo et al., 1999). Although the distinction between two populations of primary sensory fibers, peptidergic and nonpeptidergic, seems attractive, it is not completely accurate as a small proportion of sensory fibers that colocalize CGRP and somatostatin do not respond to NGF in the adult and bind the lectin IB4 (Alvarez and Fyffe, 2000). It should also be noted that, in all of the above putative nociceptive fibers, the “classical” synaptic transmitter is very likely glutamate (Battaglia and Rustioni, 1988; De Biasi and Rustioni, 1988; Merighi et al., 1991), or both glutamate and aspartate (Merighi et al., 1991; Tracey et al., 1991). Of the markers of nonpeptidergic nociceptive sensory fibers described above, the one that has been best studied is FRAP. At the light microscope level, FRAP activity is localized in a band in the middle third of lamina II, corresponding to sublamina IIBd (Ribeiroda-Silva et al., 1986) (see also Fig. 1). At the ultrastructural level, FRAP occurs in the C boutons of synaptic glomeruli of type I but not type II (Ribeiro-da-Silva et al., 1986). The physiological role of this enzymatic activity is still unknown, but it is useful as a marker of a subset of small-diameter sensory fibers. In most recent studies, the binding of IB4 has been used as the marker of the nonpeptidergic nociceptive fiber population (see Fig. 2).

Neurochemistry of Synaptic Glomeruli The C boutons of synaptic glomeruli are likely all immunoreactive for glutamate and possibly for aspartate (De Biasi and Rustioni, 1988; Tracey et al., 1991). Of the neuropeptides in the C boutons of glomeruli, CGRP is the most abundant, as it occurs in virtually all the C boutons of type Ib (i.e., with dense-core vesicles – see Figs. 4, and 5A, and 5B). Substance P immunoreactivity occurs in a subpopulation of those CGRP-IR C boutons of type I glomeruli (Ribeiro-da-Silva et al., 1989; Ribeiro-da-Silva and Cuello, 1991) (Figure 5A).

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Somatostatin also occurs in the C boutons of glomeruli (Alvarez and Priestley, 1990b; Ribeiro-da-Silva and Cuello, 1990b), where it is colocalized with CGRP (Ribeiro-da-Silva, 1994). Relative to surrounding glomerular profiles, several neurochemicals have been detected in “regular” glomerular dendrites (D): substance P (Ribeiro-da-Silva et al., 1989), neurokinin B (McLeod et al., 2000), enkephalin (Ribeiro-da-Silva et al., 1991), somatostatin (Ribeiro-da-Silva and Cuello, 1990b), GABA (Fig. 5C), glycine (Todd, 1990), and ChAT (Ribeiro-da-Silva and Cuello, 1990a). In presynaptic dendrites (V 1), the following antigenic sites have been detected, among others: somatostatin (Ribeiro-daSilva and Cuello, 1990b), enkephalin, GABA (Todd and Lochhead, 1990) (Fig. 5C), and glycine (Todd, 1990). In glomerular peripheral axons (V2), the following were detected: GABA (Barber et al., 1978; Todd and Lochhead, 1990), glycine (Todd, 1990), ChAT (Ribeiroda-Silva and Cuello, 1990a), and enkephalin (Ribeiroda-Silva et al., 1991). Certain colocalizations have been found in the surrounding profiles of glomeruli: GABA+ChAT (Figs. 7C and 7D) in V2 profiles and dendrites (D); GABA + enkephalin in V1, D, and V2 profiles; (Fig. 6) and enkephalin +substance P in D profiles (Ribeiro-da-Silva et al., 1991). GABA+glycine colocalization has been demonstrated in cell bodies (Todd and Sullivan, 1990) at the light microscopic level. Subsequently, Todd (1996) has provided evidence of GABA+glycine colocalization in V2 profiles and some V1 profiles in type II, but not in type I, glomeruli. The occurrence of GABA+glycine colocalization in presynaptic dendrites (V1 profiles) of type II glomeruli is not surprising as Spike and Todd (1992) had detected such colocalization in islet cells. However, it should be pointed out that GABA, and not glycine receptors, has been detected on primary sensory fibers. Therefore, it is likely only GABA that acts on the C boutons, whereas glycine targets other glomerular profiles.

FINAL REMARKS In conclusion, since the influential theoretical paper of Melzack and Wall (1965) introducing the spinal gate control theory, many complex synaptic arrangements have been postulated in the rat superficial dorsal horn, fitting or contradicting their main hypothesis. Despite the recent progress, the fact is that there is still insufficient direct evidence integrating the circuitry of the dorsal horn, the physiological characteristics of neurons, and the chemical nature and type of synapses involved. In the rat, it seems that the outer two-thirds of lamina II play a major role in the modulation of nociception. However, the details of modulatory mechanisms and neurochemicals involved are still not well known.

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Acknowledgments I dedicate this book chapter to Professor António Coimbra (Porto, Portugal), who was my thesis supervisor and to whom I owe a significant part of my scientific training. Many of the results reported here on the classification of synaptic glomeruli derive from my long-term collaboration with him. The author is particularly grateful to Drs. Andrew J. Todd (Glasgow, UK) and Yves De Koninck (U. Laval, Québec, Canada) for critical reading of the manuscript. I am also grateful to Mrs. Marie Ballak and Ms. Johanne Ouelette for expertise in electron microscopy, to Mr. Sid Parkinson for editorial assistance, to Mr. Alan Forster for photographic expertise, and to Mrs. Manon St. Louis for help with immunocytochemistry. The original data on transmitter colocalization given in this chapter are the result of research supported by the Canadian Institutes of Health Research (CIHR).

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C H A P T E R

7 Ascending and Descending Pathways in the Spinal Cord DAVID TRACEY School of Medical Sciences, University of New South Wales Sydney, New South Wales, Australia

by the axons of spinal neurons projecting to other segments of the spinal cord, to the dorsal column nuclei, the sensory trigeminal nuclei, and the reticular formation. The dorsal columns also contain axons descending from the dorsal column nuclei. In the rat, most of the fibers of the corticospinal tract descend in the basal part of the dorsal columns. The classic view of the dorsal columns is that they are composed of the myelinated axons of mechanoreceptors and transmit sensory information from these receptors in skin, muscles, and joints. Recent work shows that the dorsal columns also transmit information from visceral nociceptors (Feng et al., 1998; Willis et al., 1999).

Ascending pathways in the spinal cord conduct information from sensory receptors and interneurons to the brain, while descending pathways transmit motor commands as well as signals which modulate the transmission of sensory information from the spinal cord to supraspinal levels. This chapter provides an overview of the neuroanatomy of these pathways, with some reference to functional aspects. The chapter is arranged according to the level of origin or termination of the pathways (medulla, pons, midbrain, and telencephalon). Some attention is given to the neurochemistry and extent of collateralization of the neurons involved. References are to studies carried out on the rat unless another species is mentioned.

Direct dorsal column pathway The axons are ascending collaterals of sensory neurons with cell bodies in the dorsal root ganglia. These axons are somatotopically organized so that fibers from the tail run close to the midline, while fibers from the hindlimb, trunk, and forelimb are added to the lateral border of the column at progressively more rostral levels. Primary afferents entering the cervical and upper thoracic segments of the spinal cord have collaterals which terminate in the cuneate and external cuneate nuclei, while those entering the cord at lower thoracic and lumbosacral levels terminate in the gracile nucleus. These terminations have a complex somatotopic organization, which is discussed in the chapter on the somatosensory system (Tracey, Chapter 25). Lumbar afferents also have dense projections to spinocerebellar tract cells in the dorsal nucleus (Ganchrow and Bernstein, 1981; RiveroMelián and Grant, 1990) with minor projections to other

ASCENDING PATHWAYS Pathways from the Spinal Cord to the Medulla The best known of these is the dorsal column pathway, but there are also ascending pathways from the spinal cord to the sensory trigeminal nuclei, the lateral cervical nucleus, the vestibular nuclei, nuclei X and Z, the nucleus of the solitary tract, the reticular formation, and the inferior olivary nucleus. Dorsal Column Pathways The dorsal columns contain two groups of ascending fibers. The first group is made up of the ascending collaterals of primary afferents and constitutes the direct dorsal column pathway. The second group is formed

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brain stem nuclei (Ganchrow and Bernstein, 1981) including nucleus Z (Leong and Tan, 1987). However, few of the fibers which enter the dorsal columns reach the dorsal column nuclei. Only 15% of fibers entering the cord at lumbar levels reach the cervical dorsal columns, so that most leave the dorsal columns within two to three segments of their level of entry (Smith and Bennett, 1987). Some of these fibers terminate on neurons giving rise to other ascending tracts, such as the dorsal spinocerebellar tract (RiveroMelián and Grant, 1990), while about 25% of the axons in the dorsal column are propriospinal fibers, i.e., fibers originating and terminating in the spinal cord (Chung et al., 1987). Primary afferents also have collateral branches which descend in the dorsal columns, but only 3% reach as far as two segments from the site of entry (Smith and Bennett, 1987). The dorsal columns are often thought to be composed of the myelinated axons of mechanoreceptors. However, about 25% of primary afferents in the dorsal columns are unmyelinated (Chung et al., 1987). In the gracile fasciculus, many of these axons are collaterals of primary afferents which ascend at least as far as C3 and might reach the gracile nucleus (Patterson et al., 1992, 1990, 1989). However, injection of retrograde tracers into the dorsal column nuclei labeled only large cell bodies in the dorsal root ganglia. Small cell bodies, which give rise to unmyelinated axons, were not labeled (Giuffrida and Rustioni, 1992), suggesting that unmyelinated fibers in the dorsal columns do not terminate in the dorsal column nuclei. They may terminate instead on postsynaptic dorsal column neurons or on spinothalamic neurons—many of which are located at high cervical levels (see section on spinothalamic tract). Postsynaptic dorsal column pathway A substantial number of primary afferent fibers do not project directly to the dorsal column nuclei, but terminate on spinal neurons whose axons then project to the gracile and cuneate nuclei. In the rat, as in other animals, these postsynaptic dorsal column (PSDC) neurons are located primarily in lamina 4 (de Pommery et al., 1984) or central gray (Al-Chaer et al., 1996; Wang et al., 1999) and constitute about 30–40% of neurons projecting to the dorsal column nuclei (Giesler et al., 1984). There is a somatotopic arrangement of PSDC cells, such that those at lumbar levels of the cord project to the gracile nucleus, while those in the cervical enlargement project to the cuneate nucleus (Giesler et al., 1984). The axons of PSDC neurons terminate at all rostrocaudal levels of the gracile and cuneate nuclei and also terminate in the external cuneate nucleus; they make apparent synaptic contacts with lemniscal neurons projecting to the ventrobasal thalamus (Cliffer and Giesler, 1989). Post-

synaptic dorsal column neurons appear to provide the most important ascending pathway for nociceptive signals from the pelvic viscera (Al-Chaer et al., 1996; Willis et al., 1999). See also Willis et al., Chapter 27. Spinoreticular Tracts There are three main groups of spinoreticular neurons: (1) those projecting to the lateral reticular nucleus (LRt); (2) a group projecting to the medial nuclei of the pontomedullary reticular formation, including the gigantocellular reticular nucleus (Gi), the paragigantocellular nucleus (PGi), and the caudal part of the pontine reticular nucleus (PnC); and (3) neurons that innervate the dorsal reticular nucleus of the medulla (MdD). Each group receives a projection from the area around the central canal (Wang et al., 1999). Spinal neurons projecting to the LRt originate in the intermediate gray, ventral horn, and lateral spinal nucleus (Menétrey et al., 1983). The axons ascend in the ventrolateral funiculus (Zemlan et al., 1978) and terminate in a topographically organized manner in the caudal three-quarters of the nucleus (Rajakumar et al., 1992). However, these neurons appear to be heterogeneous in function. One subgroup responds to innocuous changes in joint position (Menétrey et al., 1984a) and may be implicated in motor control (Magnuson et al., 1998). This role is consistent with the cerebellar projection of LRt. A second subgroup of spinal neurons projecting to LRt responds only to noxious inputs (Menétrey et al., 1984a) and is more likely to be involved in nociception. This is consistent with the finding that a region of the LRt receives visceral and cutaneous nociceptive inputs (Ness et al., 1998) and is an important site of pain modulation (Janss and Gebhart, 1988). Neurons projecting to the medial pontomedullary reticular formation are located mainly in contralateral laminae 5, 7, and 8 (Chaouch et al., 1983; Van Bockstaele et al., 1989), where their distribution overlaps with that of spinothalamic neurons (q.v.). In fact about 8% of spinoreticular neurons send collaterals to the thalamus (Kevetter and Willis, 1983). Many spinoreticular axons terminating in the Gi ascend in the ventrolateral funiculus (Zemlan et al., 1978). Electrophysiological experiments in the cat and monkey found that a surprisingly large proportion of spinal neurons projecting to the medial pontomedullary reticular formation were not activated by peripheral stimuli, although a few were activated by low- and/or high-threshold cutaneous afferents (Haber et al., 1982; Sahara et al., 1990). It is possible that some of the unresponsive spinoreticular neurons are sensitized and activated under conditions of chronic pain (Pezet et al., 1999). A third group of spinoreticular neurons can be distinguished, which projects to the MdD. Neurons in the MdD are

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activated by noxious stimuli (Villanueva et al., 1989) and may form an important link in the transmission of signals from the spinal cord to the medial thalamic nuclei (Villanueva et al., 1998). The cells of origin of spinal neurons projecting to the MdD are mostly ipsilateral and located at all levels of the spinal cord (Lima, 1990; Villanueva et al., 1991); they are particularly dense in laminae 5–7 of the cervical enlargement (Raboisson et al., 1996). Physiological evidence suggests that the axons of these spinoreticular neurons (like those projecting to other parts of the reticular formation) run in the ventrolateral funiculus (Bing et al., 1990). Spinocervical Tract The spinocervical tract consists of axons ascending in the dorsolateral fasciculus to the lateral cervical nucleus in the upper cervical cord (see Grant and Koerber, Chapter 5). Cells of origin are concentrated in layers 3–5 at all levels of the cord, but are far fewer in number in the rat than in the cat (Baker and Giesler, 1984). The lateral cervical nucleus of the rat is small relative to that in the cat and shows no evidence of somatotopic organization (Giesler et al., 1988). In the cat, most spinocervical tract neurons are excited by hair deflection, although there is evidence for input from other modalities as well (Brown, 1981). Other Ascending Pathways to the Medulla In addition to the pathways described above, there are ascending spinal pathways to the nucleus of the solitary tract, trigeminal nuclei, vestibular nuclei, and nuclei, X and Z. The caudal part of the nucleus of the solitary tract receives afferent fibers from the spinal cord (Torvik, 1956) and from the spinal trigeminal nucleus. The cells of origin are located in laminae 1, 5, and 10 and in the lateral spinal nucleus (Guan et al., 1998; Menétrey and Basbaum, 1987; Wang et al., 1999) and overlap with neurons giving rise to the spinomesencephalic and spinoreticular tracts. Spinosolitary fibers transmit sensory data from the viscera (Hubscher and Berkley, 1995) and may be involved in integrating somatic and visceral information from the trunk (Guan et al., 1998; Menétrey and Basbaum, 1987). The sensory trigeminal complex receives afferent fibers from the spinal cord (Phelan and Falls, 1991; Torvik, 1956; Xiong and Matsushita, 2000). These axons ascend in the dorsal columns and the lateral funiculus (Phelan and Falls, 1991) and end in a narrow band just deep to the spinal trigeminal tract. They are likely to be important in reflex control of head and neck orientation. Projections from the spinal cord to the vestibular nuclei terminate primarily in the medial, spinal, and lateral vestibular nuclei (MVe, SpVe, and LVe), with particularly strong projections from the C2 and C3

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segments containing the central cervical nucleus (Matsushita et al., 1995). Retrograde labeling studies suggest that the superior vestibular nucleus also receives terminations from the spinal cord (Vincent and Rubertone, 1984). A significant part of the spinovestibular projection appears to relay sensory information from the neck. Sensory data from the neck need to be integrated with vestibular information to generate appropriate postural reflexes. See also Vidal and Sans, Chapter 30. Nuclei X, Y, and Z were originally described as accessory vestibular nuclei. Nucleus Z receives ascending collaterals of the dorsal spinocerebellar tract (Low et al., 1986) and projects in turn to the ventrobasal thalamus. This pathway transmits proprioceptive information from the hindlimb to the somatosensory cortex in the cat (McIntyre et al., 1985) and other mammals. Nucleus Z also receives sparse terminations from ascending collaterals of primary afferents from the hindlimb (Leong and Tan, 1987); fibers ascending in the lateral funiculus also project to nucleus X. Spinal inputs to the inferior olivary nucleus are discussed below with the spinocerebellar pathways.

Pathways from the Spinal Cord to the Pons These include projections to the parabrachial and Kölliker–Fuse nuclei, pontomedullary reticular formation, and basilar pontine nuclei. Parabrachial Nucleus The parabrachial nucleus is located around the superior cerebellar peduncle at the junction of the pons and midbrain. Parabrachial neurons are implicated in autonomic processing and nociception; they receive terminations from laminae 1 and 2 and the lateral spinal nucleus (Feil and Herbert, 1995), from laminae 5 and 7 (Kitamura et al., 1993), and from lamina 10 (Wang et al., 1999). The majority of spinoparabrachial neurons are excited only by noxious stimuli and the pathway is thought to be involved in autonomic and emotional or aversive reactions to painful stimuli (Bester et al., 2000). The Kölliker–Fuse nucleus is a subnucleus of the parabrachial nucleus. Its neurons are involved in respiratory and cardiac regulation and receive terminations from lamina 1 (Cechetto et al., 1985), from the lateral spinal nucleus of upper cervical segments (Feil and Herbert, 1995), and from lamina 10 (Wang et al., 1999). Other Projections from Spinal Cord to Pons A region of the dorsolateral pontine tegmentum corresponds to the micturition reflex center of Barrington. This region receives projections from the

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parasympathetic nuclei of the lumbosacral cord (Ding et al., 1997; Hamilton et al., 1995). Caudal regions of the basilar pontine nuclei receive a sparse projection from neurons located in the intermediate basilar nucleus of C1–C2 and in laminae 6–7 of the lumbar cord (Mihailoff et al., 1989; Swenson et al., 1984; Yamada et al., 1985). The locus coeruleus and the subcoerulear region receive terminals from neurons in lamina 10 (Wang et al., 1999). Spinoreticular projections to the pons are discussed previously together with those to the medulla under Spinoreticular Tracts.

Pathways from the Spinal Cord to the Cerebellum Pathways from the spinal cord to the cerebellum include two, the dorsal and the ventral spinocerebellar tracts, which transmit information from mechanoreceptors in the hindlimb, while several pathways transmit information from receptors in the forelimb and neck. Spinocerebellar axons terminate in lobules 1–5 of the anterior lobe and lobule 8 of the posterior lobe (Berretta et al., 1991; Tolbert et al., 1993) and in the deep cerebellar nuclei (Matsushita, 1999). In the granular layer of the cerebellar cortex, these axons terminate as mossy fiber terminals, arranged in longitudinal aggregates subjacent to sagittal bands of Purkinje cells (Ji and Hawkes, 1994). The general pattern of organization of the spinocerebellar tracts in the rat is the same as that in the cat and other animals. Dorsal and Ventral Spinocerebellar Tracts The cells of origin of the dorsal spinocerebellar tract are located in Clarke’s column (dorsal nucleus) from about T1 to L3 (Matsushita and Hosoya, 1979). The axons enter the lateral funiculus where they ascend (Zemlan et al., 1978). The axons overlap with those of the ventral spinocerebellar tract in the lateral funiculus and enter the cerebellum via the inferior cerebellar peduncle (Yamada et al., 1991). They signal information about position and movement of the hindlimb in the cat (Bosco et al., 2000). Neurons which give rise to the ventral spinocerebellar tract include groups of large cells in laminae 7 and 9 of the lumbar spinal cord, referred to as spinal border cells. Most of these cells have axons that cross the cord in the anterior commissure; they ascend in the lateral funiculus and enter the cerebellum via the superior cerebellar peduncle (Yamada et al., 1991) and then decussate a second time to terminate ipsilateral to their cells of origin.

lamina 6 and the other in central lamina 7 (Matsushita and Hosoya, 1979). These neurons project in the superior cerebellar peduncle to the ipsilateral cerebellum (Matsushita and Hosoya, 1979; Yamada et al., 1991). Neurons in the external cuneate nucleus and the rostral cuneate nucleus contribute to the cuneocerebellar tract (Tolbert and Gutting, 1997) and receive proprioceptive inputs from primary afferent fibers in the dorsal columns. This tract is in some ways a forelimb analog of the dorsal spinocerebellar tract and carries information about the position and movement of the forelimb. The central cervical nucleus is located just lateral to the central canal from C1 to C3, and projects to the cerebellar cortex and nuclei in the inferior and superior cerebellar peduncles (Matsushita and Yaginuma, 1995). It transmits information from receptors in the neck and labyrinths. Spinoolivary Tract and Other Indirect Spinocerebellar Pathways There is an indirect projection from the spinal cord to the cerebellum via the inferior olivary nucleus, whose climbing fibers terminate exclusively in the cerebellum (see Voogd, Chapter 9). Spinoolivary neurons terminate in the medial and dorsal accessory olivary nuclei (Azizi and Woodward, 1987; Swenson and Castro, 1983b); their cells of origin are located in the medial aspect of the nucleus proprius and in the central cervical nucleus (Swenson and Castro, 1983a). Other polysynaptic pathways from the spinal cord to the cerebellum include the projection from the spinal cord to the lateral reticular nucleus (see above) and the projection from the dorsal column and trigeminal nuclei to the inferior olivary nucleus (Molinari et al., 1996).

Pathways from the Spinal Cord to the Midbrain Spinomesencephalic neurons are located mainly in the cervical cord (Yezierski and Mendez, 1991) and project to three main regions of the midbrain: the superior colliculus, the central gray, and the midbrain reticular formation. All three regions receive nociceptive inputs and form part of the neural circuitry involved in the localization or descending control of pain (see Willis et al., Chapter 27). There are also terminations in the anterior and posterior pretectal nuclei, the red nucleus, the Edinger–Westphal nucleus, and the interstitial nucleus of Cajal (Menétrey et al., 1982; Willis and Westlund, 1997; Yezierski, 1988).

Other Spinocerebellar Pathways

Superior Colliculus

In the cervical enlargement, there are two distinct groups of spinocerebellar neurons—one in medial

Most spinotectal neurons are located contralaterally in laminae 3 to 5 and in laminae 7 to 8 (Morrell and Pfaff,

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1983). Spinotectal neurons are also found in lamina 1 and in the lateral spinal nucleus (Menétrey et al., 1982). The axons run in the lateral funiculus and terminate in the deep and intermediate layers of the superior colliculus (Antonetty and Webster, 1975) as well as the intercollicular nucleus (Zemlan et al., 1978) and pretectal nuclei (Yezierski, 1988). The tract provides somatic input to the superior colliculus, where it is integrated with visual and auditory information to play a role in head orientation. Central Gray Spinal neurons project to the caudal part of the central gray via the ventrolateral funiculus (Bernard et al., 1995; Bianchi et al., 1990). Functionally distinct columns have been recognized in the periaqueductal gray, with nociceptive input from deep somatic and visceral structures activating neurons in the ventrolateral column and nociceptive input from the skin activating the lateral column (Clement et al., 2000). Cells of origin are located mainly in the upper cervical and sacral spinal cord, and those projecting to the lateral column are organized in a manner somewhat similar to that of the spinoparabrachial projection (Keay et al., 1997). There is a strong projection from lamina 10 (Wang et al., 1999). See also Keay and Bandler, Chapter 10. Midbrain Reticular Formation Anterograde labeling of spinomesencephalic neurons in the lumbosacral cord showed that they have relatively dense terminations in the cuneiform nucleus in the caudal part of the midbrain and sparser terminations in the rostral part of the midbrain, including the deep mesencephalic nucleus (Veazey and Severin, 1982; Yezierski, 1988). The cells of origin are similar in location to those projecting to the superior colliculus and central gray (Menétrey et al., 1982). They include lamina 1 cells with axons in the contralateral dorsolateral fasciculus and collateral branches to the cuneiform nucleus (McMahon and Wall, 1985) and neurons in lamina 10 (Wang et al., 1999).

Pathways from the Spinal Cord to the Diencephalon The major projection from the spinal cord to the diencephalon is the spinothalamic tract (Fig. 1). Spinal neurons also project to the hypothalamus and to parts of the subthalamus and epithalamus (Cliffer et al., 1991). Spinothalamic Tract The spinothalamic tract (STT) is the main pathway for information from receptors signaling pain and temperature (Willis and Westlund, 1997). Several groups of STT neurons can be distinguished, including cells in

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medial laminae 4–6 and lateral laminae 7–9 (Granum, 1986; Kemplay and Webster, 1986; Kobayashi, 1998). In the rat, at least 50% of STT neurons are located in the first four cervical segments, and about 90% of STT neurons have axonal terminations contralateral to their cell bodies. The axons generally cross the midline within a few segments of the cells of origin (Granum, 1986; Kemplay and Webster, 1986) and ascend in the ventral or ventrolateral funiculus (Giesler et al., 1981). The spinothalamic tract terminates in three main regions—the ventroposterolateral nucleus (Peschanski et al., 1983); the intralaminar nuclei, primarily the central lateral nucleus (Ma et al., 1987); and the posterior complex (Dado et al., 1994a; Ledoux et al., 1987). STT axon terminals are also found in the nucleus gelatinosus (submedius) (Craig and Burton, 1981), but they are sparse (Cliffer et al., 1991). A study of STT neurons in the cervical enlargement of the rat found that about 50% responded to both noxious and innocuous stimuli (wide dynamic range) while 44% responded only to noxious stimuli (high threshold) and 6% responded preferentially to innocuous stimuli (Dado et al., 1994b). These data are consistent with those from STT neurons in the lumbar enlargement of monkeys (Willis and Coggeshall, 1991; Willis and Westlund, 1997). However, lumbar STT neurons of the rat differ from those of primates in that only a small percentage of the rat’s complement of STT neurons are located in the lumbar enlargement, and most of these respond primarily to innocuous stimuli (Dado et al., 1994b; Menétrey et al., 1984b). In the sacral spinal cord of the rat, a significant proportion of STT neurons responded to noxious visceral inputs such as distension of the colon, rectum, and vagina (Katter et al., 1996). Spinothalamic neurons send collaterals to several regions of the CNS (Lu and Willis, 1999), including the medullary reticular formation (Kevetter and Willis, 1983), the periaqueductal gray (Harmann et al., 1988; Liu, 1986), and the parabrachial area (Hylden et al., 1989). Spinal Projections to Other Parts of the Diencephalon There is a substantial projection from the spinal cord to the hypothalamus. Spinohypothalamic neurons are located bilaterally, mostly in the deeper laminae of the dorsal horn and the lateral spinal nucleus (Burstein et al., 1990). They terminate in several regions of the hypothalamus, including the lateral, posterior, and dorsal hypothalamic areas (Cliffer et al., 1991; Wang et al., 1999). Their response properties are similar to those of spinothalamic neurons (Burstein et al., 1991; Dado et al., 1994b; Katter et al., 1996). There is also a projection from the spinal cord to the zona incerta, mainly from the dorsal horn and intermediate gray of the cervical

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A

PO VPL CL

CM

B

gr cu dlf

from Lamina I

lfu Lateral STT

VMnF vfu

Medial STT 0.5 mm

C

1

gr cu dlf

4 5 6

lfu 10 8

7

sparse moderate

2

3

9 9

VMnF

9

9 vfu

dense

FIGURE 1 Spinothalamic tract (STT). (A) Sites of termination in the centrolateral thalamic nucleus (CL), the posterior thalamic nuclear group (PO), and the ventral posterolateral thalamic nucleus (VPL). Note the patchy distribution of spinothalamic terminals (Cliffer et al., 1991). (B) Course of spinothalamic axons (Giesler et al., 1981; Hylden et al., 1989). (C) Cells of origin (Kemplay and Webster, 1986).

and lumbar cord (Shaw and Mitrofanis, 2001; Wang et al., 1999).

Pathways from the Spinal Cord to the Telencephalon There is a direct projection from neurons in the spinal cord to some regions of the telencephalon, including the basal ganglia, amygdala, and infralimbic and medial orbital cortex (Cliffer et al., 1991; Newman et al., 1996; Wang et al., 1999). Spinal neurons projecting to these regions are found bilaterally in the deeper layers of the dorsal horn, lamina 10, and the lateral spinal nucleus

(Burstein and Potrebic, 1993; Wang et al., 1999). These projections provide a direct pathway for somatosensory information from the spinal cord to the limbic system and basal ganglia.

DESCENDING PATHWAYS Pathways from the Telencephalon to the Spinal Cord The vast majority of neurons in the telencephalon which project to the spinal cord are in the cerebral cortex.

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Corticospinal Tract

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et al., 1990). Corticospinal neurons are not restricted to primary motor and sensory cortex, but are found in a number of other cortical regions. In frontal cortex, corticospinal neurons are found in an area corresponding with the supplementary motor area (Fr2) and in the prefrontal cortex (Li et al., 1990; Miller, 1987). In parietal cortex they are found in the second somatosensory area (Par2) and in the posterior parietal cortex (part of Par1); corticospinal neurons are also located in the visual association cortex, the anterior cingulate cortex (Miller, 1987), and the infralimbic cortex (Hurley et al., 1991).

Most corticospinal neurons are located in the primary motor cortex, equivalent to Fr1 + Fr3, and in the forelimb and hindlimb parts of the primary sensory cortex, equivalent to FL and HL (Fig. 2). In primary motor cortex, corticospinal neurons are located throughout layer 5, whereas in primary sensory cortex they are restricted to layer 5b (Miller, 1987). Neurons in the forelimb area of motor and sensory cortex project to the cervical enlargement, while those in the hindlimb area project to the lumbar enlargement (Li

A RSG

Cg1 Fr2

Fr1

RSA

HL

FL

29D

Oc2M

Oc1M

Fr3 Oc1B

Par1 Par2 Par2(S2)

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Te2

gr cu

vfu 0.5 mm

C 2

dlf lfu

1 3 4 5 6

9 7 9 9

sparse moderate

gr cu

8

10

9

dense

FIGURE 2 Corticospinal tract. (A) Cells of origin (Miller, 1987). The dorsal aspect of the cortex is shown, with nomenclature after Zilles (Palomero-Gallagher and Zilles, Chapter 23). Par1+FL+HL together constitute the first somatosensory area, often referred to as SI; while Par2 is equivalent to the second somatosensory area, SII. Fr1+Fr3 together make up the motor cortex, while Fr2 corresponds to the supplementary motor area. (B) Course of corticospinal axons. Note the bundle of uncrossed axons in the ventral funiculus, vfu (Casale et al., 1988). (C) Terminations in the spinal cord (Casale et al., 1988).

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The corticospinal tract of the rat decussates in the caudal medulla and runs in the base of the dorsal columns (Armand, 1982) but there is also a ventral uncrossed tract in the ventral funiculus (Brösamle and Schwab, 1997), as well as other minor components (Liang et al., 1991). In the rat, corticospinal neurons send collaterals to the midbrain and trigeminal nuclei (Catsman-Berrevoets and Kuypers, 1981; Killackey et al., 1989). A few corticospinal axons send collaterals to the pontine nuclei and the red nucleus (Akintunde and Buxton, 1992). Corticospinal axons terminate in all spinal laminae contralateral to the cells of origin (Fig. 2), with dense terminations in laminae 3–7 of the dorsal horn and less dense terminations in laminae 1 and 2 and the ventral horn (Casale et al., 1988; Liang et al., 1991). Corticospinal axons make synaptic contacts with motoneurons in the rat (Liang et al., 1991), as they do in primates. The corticospinal tract plays a role in the control of movement through its terminations in the intermediate gray and ventral horn, which include direct terminations on motoneurons (Liang et al., 1991). However, the corticospinal tract is not essential for the control of skilled limb movements such as reaching for and grasping food (Whishaw et al., 1998). Diencephalic Projections to the Spinal Cord There is a major projection from the hypothalamus to the spinal cord, terminating primarily in lamina 1 of the dorsal horn and in the preganglionic sympathetic and parasympathetic cell columns. The cells of origin are located in several distinct groups—the paraventricular, arcuate, and perifornical nuclei and the lateral hypothalamus. These neurons use peptides such as oxytocin and vasopressin as neurotransmitters and terminate bilaterally in the spinal cord (Cechetto and Saper, 1988; Palkovits, 1999). There is also a dopaminergic projection from the All group, located in the posterior and dorsal hypothalamic areas and in the caudal thalamus (Skagerberg and Lindvall, 1985). Paraventricular neurons containing vasopressin terminate directly on preganglionic sympathetic neurons and are involved in the response to stressful stimuli (Motawei et al., 1999). The spinal terminations of neurons in the perifornical nuclei and lateral hypothalamic area contribute to the control of blood pressure (Allen and Cechetto, 1992), while the projections of arcuate and retrochiasmatic neurons to the thoracic cord are implicated in production of melatonin and regulation of energy balance (Elias et al., 1998; Ribeiro-Barbosa et al., 1999). Cells in the medial part of the zona incerta project to the cervical and lumbar regions of the cord, with most terminations in laminae 4, 5, and 10 (Schwanzel-Fukuda et al., 1984; Shaw and Mitrofanis, 2001; Watanabe and

Kawana, 1982). There is a sparse projection to the ventral horn of the cervical cord from the parafascicular nucleus of the thalamus (Marini et al., 1999).

Pathways from the Midbrain to the Spinal Cord There are projections to the spinal cord from the red nucleus and from those parts of the midbrain which receive spinal inputs, i.e., the superior colliculus, the central gray, and the midbrain reticular formation. Rubrospinal Tract Neurons in both the parvicellular and magnocellular parts of the red nucleus project to the spinal cord. There is a somatotopic organization of the nucleus such that the ventrolateral part projects to the lumbar cord, while dorsomedial parts project to the cervical cord (Daniel et al., 1987; Strominger et al., 1987). This organization reflects the sequence of descent of rubrospinal fibers during development (Lakke and Marani, 1991). Most rubrospinal axons terminate in contralateral laminae 5–7 where they establish contact with excitatory and inhibitory interneurons (Antal et al., 1992). Lesions of the red nucleus impair locomotion and skilled reaching movements in the rat (Muir and Whishaw, 2000; Whishaw et al., 1998). Other Projections from the Midbrain to the Spinal Cord Tectospinal axons originate in the deep and intermediate layers of the superior colliculus and project to the cervical cord, mainly to contralateral lamina 5, 7, and 8 (Murray and Coulter, 1982; Yasui et al., 1998). They are implicated in the control of head movements. There are also projections to the spinal cord from the periaqueductal gray (Masson et al., 1991), the midbrain reticular formation (Satoh, 1979; Veazey and Severin, 1980a, 1980b; Waldron and Gwyn, 1969), and the dorsal raphe (Bowker et al., 1981; Kazakov et al., 1993; Skagerberg and Björklund, 1985) and from accessory oculomotor nuclei such as the Edinger–Westphal nucleus, the nucleus of Darkschewitsch, and the nucleus of the posterior commissure (Leong et al., 1984).

Pathways from the Pons to the Spinal Cord The pons contains several groups of neurons that project to the spinal cord. These include neurons in the pedunculopontine tegmental nucleus and the pontine reticular formation (Rye et al., 1988; Sirkin and Feng, 1987; Spann and Grofova, 1989). The parabrachial nucleus (including the Kölliker– Fuse nucleus) is involved in the modulation of nocicep-

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tion (Blomqvist et al., 1994) and autonomic functions. Its neurons project to the dorsal horn and intermediolateral cell column (Fulwiler and Saper, 1984; Yoshida et al., 1997). Several groups of noradrenergic neurons project to the spinal cord. These are the A5 group in the ventrolateral brain stem, the locus coeruleus (A6), and the A7 group which overlaps with the Kölliker–Fuse nucleus and the subcoeruleus nucleus (Kwiat and Basbaum, 1992). Their descending axons have been implicated in the control of autonomic functions, modulating the perception of pain (Jones, 1991) and modifying motor behavior such as locomotion. The course and terminations of axons descending from these noradrenergic neurons depend on the substrain of rat. In Sprague– Dawley rats supplied by Harlan, most locus coeruleus neurons projected to the dorsal horn. In contrast, in Sprague–Dawley rats supplied by Sasco, most locus coeruleus neurons projected to the ventral horn (Sluka and Westlund, 1992). Such differences in spinal projections have also been shown for A5 and A7 groups, although the A5 group appears to have consistent terminations in the intermediolateral cell column (Clark and Proudfit, 1991, 1993; Fritschy and Grzanna, 1990; Sluka and Westlund, 1992). The intermediolateral cell column also receives axon terminals from enkephalincontaining neurons in these “noradrenergic” groups (Romagnano et al., 1991). In the dorsolateral pontine tegmental region there is a group of neurons which corresponds to the micturition reflex center of Barrington. Neurons in this region project to the spinal parasympathetic nucleus and to pudendal motoneurons (Ding et al., 1995). Barrington’s nucleus may also influence the activity of sympathetic neurons in the spinal cord (Cano et al., 2000).

Pathways from the Cerebellum to the Spinal Cord Neurons in the deep cerebellar nuclei project to the cervical spinal cord (Bentivoglio, 1982; Leong et al., 1984). Some of these neurons send collaterals to the diencephalon or superior colliculus (Bentivoglio and Kuypers, 1982).

Pathways from the Medulla to the Spinal Cord These include projections from the trigeminal and dorsal column nuclei, the medullary reticular formation, and the raphe nuclei, as well as the vestibular complex. Trigeminal and Dorsal Column Nuclei Neurons in all three subnuclei of the spinal trigeminal nucleus (Sp5C, Sp5I, and Sp5O) send axons as far as

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the thoracic cord (Lakke, 1997) or further (Ruggiero et al., 1981). Trigeminospinal neurons are also found in the principal sensory nucleus (Phelan and Falls, 1991) and the mesencephalic trigeminal nucleus (Leong et al., 1984). Descending projections from the gracile and cuneate nuclei are present in the rat (Burton and Loewy, 1977; Villanueva et al., 1995); there is also a spinal projection from the external cuneate nucleus (Leong et al., 1984; Zemlan et al., 1979). The gracile nucleus seems to project to the lumbar cord and sacral cord, while neurons in the cuneate nucleus project mainly to the cervical cord (laminae 1, 4 & 5). Cuneospinal neurons are concentrated in the ventral parts of the nucleus (Leong et al., 1984; Masson et al., 1991). Reticular Formation Thirteen groups of reticulospinal neurons have been described in the medulla (Newman, 1985), including the raphe nuclei (see below). The gigantocellular complex (Gi) has several components which project to the spinal cord. Their axons course bilaterally in the ventral and lateral funiculi and terminate in all laminae, including laminae 1 and 2, the intermediolateral cell column, and the sacral parasympathetic nucleus (Martin et al., 1985). Terminations of Gi reticulospinal axons have been implicated in the modulation of blood pressure (Aicher et al., 2000) and the control of axial musculature (Robbins et al., 1992; Sasaki, 1999). The ventrolateral part of the intermediate reticular nucleus contains a cell column which includes the ventral respiratory group and its rostral pole, the Bötzinger complex. These groups form part of the medullary respiratory network and project to the phrenic nucleus in the cervical cord (Ellenberger, 1999). Adjacent to the ventral respiratory group is the rostral ventrolateral medulla, a functionally distinct group whose descending axons modulate the activity of preganglionic sympathetic neurons (Aicher et al., 2000; Lipski et al., 1996). Reticulospinal neurons are also found in the dorsal and ventral medullary nuclei (MdD and MdV) (Tavares and Lima, 1994; Villanueva et al., 1995). Those in the dorsal reticular nucleus are involved in pain modulation (Villanueva et al., 1996). The retroambiguus nucleus projects to sympathetic and spinal motoneurons and is implicated in respiration, vocalization, and copulation (Hardy et al., 1998; Holstege et al., 1997). Raphe Nuclei The medullary raphe nuclei are the raphe magnus (RMg), the raphe obscurus (ROb), and the raphe pallidus (RPa); all of these have projections to the spinal cord (Bowker et al., 1982) and up to 85% of raphe–spinal neurons contain 5-HT (Bowker and Abbott, 1990).

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Neurons in the raphe magnus and the adjacent ventral part of the gigantocellular reticular formation project to the spinal cord in the dorsolateral funiculus (Fig. 3) (Bowker et al., 1982; Skagerberg and Björklund, 1985). Their axons terminate mainly in the dorsal horn, with sparse projections to the ventral horn as well (Jones

A

ROb GiV

RMg RPn

and Light, 1990). This pathway plays an important role in the descending control of pain (Mason, 1999; Wei et al., 1999). Neurons in the raphe obscurus and raphe pallidus nuclei project in the ventrolateral white matter (Fig. 3) (Skagerberg and Björklund, 1985) and many terminate on motoneurons in the ventral horn (Hermann et al., 1998; Holstege and Kuypers, 1987a, 1987b). Most of these raphe–spinal neurons have multiple neurotransmitters, including serotonin, glutamate, GABA, and neuropeptides such as substance P and thyrotropinreleasing hormone (Hökfelt et al., 2000). Sympathetic preganglionic neurons in the intermediolateral cell column receive synaptic contacts from axons originating in both the raphe pallidus and the raphe magnus (Bacon et al., 1990). The function of the medullary raphe nuclei has been described in general terms as a system for integration and gain control in autonomic and somatomotor systems (Lovick, 1997). Vestibular Nuclei

B

Vestibulospinal projections arise from all four divisions of the vestibular nuclear complex: the lateral, medial, spinal, and superior vestibular nuclei. Neurons in all four nuclei send axons as far as the lumbosacral cord, but most spinally projecting neurons are located in the lateral vestibular nucleus (Leong et al., 1984; Masson et al., 1991). The medial vestibular nucleus projects predominantly to the dorsal horn of the upper cervical spinal cord (Bankoul and Neuhuber, 1992). Some vestibulospinal neurons send collateral projections to the oculomotor nuclei (Tracey and Wenderoth, 1992).

dlfu

vlfu

C gr

2

Propriospinal Connections

1

3

10

CC 9

4 LSp 5 IML 7 8 9

VMnF

FIGURE 3 Serotoninergic pathways to the spinal cord. (A) Cells of origin. (B) Course of serotoninergic axons. (C) Terminations of serotoninergic axons. Neurons can be divided into two groups: A ventral group (cross hatch) includes the raphe magnus nucleus (RMg) and the ventral part of the gigantocellular reticular nucleus (GiV) and has axons which course mainly in the dorsal part of the lateral funiculus (dlfu) and terminate primarily in the dorsal horn. The second group (stipple) includes the raphe obscurus nucleus (ROb) and the raphe pallidus nucleus (RPa) and has axons which course mainly in the ventral part of the lateral funiculus (vlfu) and terminate primarily in the ventral horn. There is overlap between the projections of the two groups (Bowker et al., 1982; Jones and Light, 1990; Skagerberg and Björklund, 1985).

Propriospinal neurons connect one part of the spinal cord with another. They have been implicated in the control of movement (Cowley and Schmidt, 1997) and in nociception (Sandkuhler et al., 1993). Propriospinal axons constitute approximately 33% of axons in the white matter of the sacral cord (Chung and Coggeshall, 1983; Chung et al., 1987). Their cell bodies are located in all laminae except lamina 9 (Bice and Beal, 1997; Menétrey et al., 1985; Verburgh et al., 1990) and project to the ipsilateral dorsal and ventral horns and lamina 10, as well as to the contralateral cord (Matsushita, 1998; Petko and Antal, 2000). However, Petko and Antal argue that there are significant differences between the medial and lateral parts of the dorsal horn, with the lateral part of the superficial dorsal horn receiving most C-fiber terminations and giving rise to most of the ascending pathways dealing with nociception. Propriospinal neurons in the lateral dorsal horn differ from those in the medial dorsal horn in that they have reciprocal connections with the whole rostrocaudal extent of the

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cord, have commissural connections with the contralateral dorsal horn, and are the main source of projections to the ventral horn and supraspinal centers (Petko and Antal, 2000). These differences in connectivity suggest that it is the propriospinal neurons in the lateral part of the dorsal horn which play a significant role in nociception.

Acknowledgments I thank Professor M. Matsushita, Dr. J. Mitrofanis, and Dr. R.B Simerly for constructive comments and Ms. Alicia Fritchle for assistance with illustrations. I also thank Professor Peter Grafe for providing the facilities for the preparation of the manuscript.

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8 Precerebellar Nuclei and Red Nucleus TOM J. H. RUIGROK Department of Neuroscience, Erasmus MC Rotterdam The Netherlands

The cerebellum receives afferents from many sources in the brain stem and spinal cord that are collectively known as the precerebellar nuclei. Clearly, these precerebellar nuclei cannot be regarded as a single group, but should be differentiated into various functionally and morphologically distinct clusters. The largest group consists of neurons that terminate as the so-called mossy fibers in the granular cell layer of the cerebellar cortex (Ramón y Cajal, 1888, 1911). Well-known and quantitatively important sources of mossy fibers are the spinal cord (see Tracey, Chapter 7), the pontine nuclei, the vestibular nuclei (see Vidal and Sans, Chapter 31), the lateral reticular nucleus, the dorsal column nuclei, the trigeminal nuclei, and a score of reticular nuclei. A second group supplying afferents to the cerebellum consists of only a single nucleus. The inferior olivary complex gives rise to all climbing fibers, terminating upon the dendritic tree of the cerebellar Purkinje cells (Ramón y Cajal, 1888, 1911; Szentágothai and Rajkovits, 1959; Desclin, 1974). Finally, a third group of precerebellar nuclei provide monoaminergic inputs to the cerebellum and are most likely involved in modulatory functions (André et al., 1991, 1993; Strahlendorf et al., 1991). Serotonergic afferents are found in diffusely beaded terminals within the granular cell layer as well as in the cerebellar nuclei (Bishop and Ho, 1985). Noradrenergic terminals mainly originate within the locus coeruleus and are found within all layers of the cerebellar cortex and cerebellar nuclei (Hökfelt and Fuxe, 1969; Olson and Fuxe, 1971; Chan-Palay, 1977). This chapter deals with two of the main sources of mossy fibers, the basal pontine nuclei (including the

The Rat Nervous System, Third Edition

reticulotegmental nucleus) and the lateral reticular nucleus, as well as with the source of the climbing fibers, the inferior olivary complex. In addition, this chapter reviews and discusses the role of the red nucleus in cerebellar functioning. This prominent center in the midbrain, which gives rise to the rubrospinal tract, is intimately related to cerebellar function since it receives a major input from the cerebellar nuclei. It, furthermore, supplies afferents to some important precerebellar centers such as the lateral reticular nucleus and the inferior olive, but also sends input to the cerebellum directly.

PONTINE NUCLEI The basilar pontine nuclei (Pn) consist of a large cluster of rather densely grouped small to mediumsized neurons that are located near the ventral surface of the metencephalon. These nuclei receive a massive input from the cerebral cortex, but also from a number of subcortical brain areas. Their neurons project to the cerebellum by way of the middle cerebellar peduncle and terminate as mossy fiber terminals within the granular cell layer of large areas of the cerebellar cortex. The nucleus reticularis tegmenti pontis or reticulotegmental nucleus, located directly dorsal to the Pn and also providing a major mossy fiber projection to the cerebellum, has several features in common with the pontine nuclei (Schwarz and Thier, 1996) and both are often considered simultaneously. Other authors, however, regard the RtTg as a specialized nucleus of

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the pontine reticular formation (Newman and Ginsberg, 1992) since it differs both in ontogeny (Altman and Bayer, 1987) and in pattern of connectivity (see below) from the Pn. Therefore, it is described in a separate subsection.

Cytoarchitecture The rat Pn cover almost 2 mm in the rostrocaudal direction and are found between the level of the interpeduncular nucleus and that of the trapezoid body. They are surrounded and traversed by a number of large fiber bundles. Laterally, the middle cerebellar peduncle emerges from the Pn. It carries the pontine efferent fibers that course to the cerebellum. Most efferent fibers of the pontine neurons cross the midline at the ventral surface, before entering the middle cerebellar peduncle. Descending fibers of the cerebral peduncle take up a position as the longitudinal fibers of the pons (lfp) in the dorsal part of the Pn but also as scattered bundles throughout the Pn. Ascending fibers of the medial lemniscus are located dorsal to the lfp and the Pn. Four main subdivisions are generally recognized with respect to their position relative to the lfp (Mihailoff et al., 1981b) (Fig. 1). The medial, ventral, and lateral subdivisions consist of rather tightly packed and homogeneously distributed neurons, and the peduncular nuclei immediately surround the lfp. A number of smaller subnuclei may also be identified. However, borders are not easy to delineate between the various subdivisions.

FIGURE 1 Schematized diagram of the pontine nuclei (modified from Mihailoff et al., 1981b). Abbreviations used: dL, dorsolateral pontine area; dM, dorsomedial pontine area; dPd, dorsal peduncular area; L, Lateral nucleus; lfp, longitudinal fascicle of the pons; M, medial nucleus; mcp, medial cerebellar peduncle; ml, medial lemniscus; RtTgC, central part of the reticulotegmental nucleus of the pons; RtTgP, pericentral part of the reticulotegmental nucleus; V, ventral nucleus; vM, ventromedial area; vPd, ventral peduncular area.

Neurons in the Pn have been shown to demonstrate a rather variable morphology. The projection neurons in the dorsal and medial regions of the Pn are generally somewhat larger and possess more dendrites compared to the neurons in the ventral Pn areas. Cluster analysis indicated that these differences are due to a dorsoventral gradient rather than reflecting different cell types (Schwarz and Thier, 1996). Mihailoff and collaborators (1981b), studying Golgi material, suggested that a small population of small neurons displaying local axon collaterals, could indicate an inhibitory feedback mechanism. However, although some of these neurons indeed may be GABAergic, the impact of an intrinsic GABAergic source at best is very limited (Border and Mihailoff, 1985; Aas and Brodal, 1990), or may even be nonexistent (Mock et al., 1999).

Afferents to the Basilar Pontine Nuclei Cerebral Cortex In the rat, most afferents to the Pn arise from layer V neurons located throughout the entire ipsilateral cortex (Legg et al., 1989). However, there are clear regional differences in the relative contribution of each cortical area to the corticopontine system. Most fibers originate from the sensory motor and visual cortices. In addition, the primary auditory (rostral temporal cortex) as well the cingulate, the retrosplenial, and the agranular insular cortices provide an appreciable corticopontine projection also. Relatively small contributions are derived from the caudal temporal cortex and from the perirhinal cortex (Burne et al., 1978; Mihailoff et al., 1978; Wiesendanger and Wiesendanger, 1982a, 1982b; Mihailoff et al., 1985; Legg et al., 1989). The corticopontine fibers usually also send fibers to other subcortical structures (O’Leary and Stanfield, 1985; Ugolini and Kuypers, 1986; Leergaard et al., 1995). Within the corticopontine projections a topographical pattern has been frequently reported. Projections from the motor cortex predominantly terminate in the medial subdivision of the Pn, whereas the somatosensory and visual cerebral cortices project to more central and lateral areas, respectively (Wiesendanger and Wiesendanger, 1982b; Mihailoff et al., 1985). These earlier studies reported that cortical areas containing hindlimb and face representation terminate in one or several clusters that show an essentially longitudinal orientation (also see Panto et al., 1995). However, recent studies using three-dimensional analysis of serial sections indicated that these clusters are not distributed primarily in rostrocaudal columns but in curved and elongated lamellar clusters, displaying an internal-to-external somatotopic arrangement (Leergaard et al., 2000b). As such the

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projections from the peri-oral regions of the head are found in the center of the Pn whereas the projections from extremities and trunk are located in multiple shelllike and complementary regions more externally in the Pn (Figs. 2A and 2B). Hence, within the distribution of corticopontine terminals, the spatial relationships from the major cortical body representations appear to be preserved (Panto et al., 1995; Leergaard et al., 2000b). The establishment of this specific pattern of terminations may be related to the differences in maturation of the cerebral cortex in conjunction with the ontogeny within the Pn as has been hypothized by Leergaard and collaborators (Fig. 2A: Leergaard et al., 1995, 2000b). Also, at even more detailed levels, such as the pontine projection from individual cortical barrel fields, the shell-like, multiple, and essentially inside-out representation of the terminal fields holds true (Fig. 2C) (Leergaard et al., 2000a). The terminal fields of the cortical barrel fields representing whiskers of the same row were found to be located in different shells (anterior whisker represented more centrally, compared to the more posterior whisker), whereas barrels across rows (i.e., located in columns) resulted in terminal fields in the same shell. Despite the fact that the somatotopic pattern of the corticopontine projections is largely preserved in a complementary (i.e., nonoverlapping) way, some overlap (up to 20%) was reported in the pontine projections from the individual barrel fields (Panto et al., 1995; Leergaard et al., 2000a). As yet, the visual corticopontine input has not been investigated in similar detail but was earlier found to terminate predominantly within the lateral third of the basal Pn with exception of its lateral-most part. In addition, small patches of labeling are found rostromedially (Wiesendanger and Wiesendanger, 1982b). Auditory projections arising from the primary temporal cortex are rather weak and terminate ventrally within the lateral third of the nuclei. Anatomical and physiological studies (Ruegg et al., 1977; Potter et al., 1978; Wiesendanger and Wiesendanger, 1982b) have suggested that there are major zones of convergence from widely differing cortical regions. Mihailoff et al. (1981b) attributed this convergence not only to overlap of the corticopontine termination clusters, but also to the tendency of the dendritic trees of the pontine neurons to invade neighboring corticopontine termination zones. However, a detailed morphological study by Schwarz and Thier (1995) showed that the dendritic fields of most pontine projection neurons respect the borders of cortical afferent fields. Cerebellar Nuclei A second major input system to the Pn originates in the cerebellar nuclei (Watt and Mihailoff, 1983a, 1983b; Angaut et al., 1985b; Lee and Mihailoff, 1990; Teune et al.,

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2000; also see Voogd, Chapter 9). Although considerable overlap with the corticopontine projection is evident, the level of specificity between the corticopontine and the cerebellopontine projections remains to be investigated (Leergaard et al., 2000a). The cerebellopontine projection is predominantly contralateral and reaches the Pn via the crossed descending limb of the superior cerebellar peduncle. Fascicles of cerebellopontine fibers leave the main bundle of the descending limb throughout the rostrocaudal extent of the Pn and pass around or through the medial lemniscus and the cerebral peduncle to terminate in the pontine gray. The cerebellopontine system is composed of collateral branches of cerebellar efferent fibers that project to either the thalamus or the inferior olive (Lee et al., 1989) and was claimed to consist, at least in part, of glutamatergic and GABAergic fibers (Border et al., 1986; Border and Mihailoff, 1987). This finding is consistent with an electrophysiological study that reports both excitatory as well as inhibitory monosynaptic responses of Pn neurons as the result of electrical stimulation of the lateral cerebellar nucleus (Lat) (Berretta et al., 1991). However, an ultrastructrural double-labeling study has clearly concluded that the cerebellopontine projection is nonGABAergic (Schwarz and Schmitz, 1997). Also, it was recently shown that electrical stimulation of the Lat is capable of inducing enhanced expression of the immediate early gene c-Fos suggesting that the activity of the cerebellar nuclei may significantly influence activity patterns in the Pn presumably by way of the direct cerebellar nucleopontine projections (Bosco et al., 2000). Although all divisions of the cerebellar nuclei provide input to the Pn, the largest number of cerebellopontine fibers, as well as the largest Pn domain covered by terminal arborizations, are derived from the Lat. Basically, the Lat projections are distributed to three longitudinal columns, one in each major subdivision of the Pn. An inverted topographical pattern in this projection, so that caudal parts of the Lat tend to project to more rostral Pn regions than do the rostral Lat areas, has been described by Angaut and colleagues (1985b). Projections from the interposed nuclei to the Pn appear to be restricted to the ventral peduncular nucleus at more caudal levels of the pontine gray matter. However, according to Watt and Mihailoff (1983a), the interpositus projection may also extend into the ventral pontine subnucleus, where it overlaps with projections from the Lat. The medial cerebellar nucleus provides only few projections to the Pn that are mainly found in the dorsomedial area, where they may partly overlap with projections from the Lat, but also with input from the mammilary nucleus and the cingulate cortex (Wiesendanger and Wiesendanger, 1982b; Watt and Mihailoff, 1983b).

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FIGURE 2 Topographic organization of corticopontine projections in the rat. (A) Diagram of the hypothesis explaining the establishment of the general topographic organization in the rat corticopontine system. Temporal gradients of development, from early to later, are illustrated by dark, medium, and light shading. Early arriving corticopontine fibers (red) innervate the early established central core of the pontine nuclei, whereas later arriving fibers (yellow and blue) innervate progressively more external volumes. Modified from Leergaard et al. (2000b) with permission; see also Leergaard et al. (1995). (B) Three-dimensional surface model of the projections from major SI body representations to the pontine nuclei. (B1) Diagram of the right cerebral hemisphere indicating the position of the SI body map. The different areas of the body map are color coded. (B2) Three-dimensional reconstruction of the corresponding clustered pontine projection regions seen in a ventral view of the brain stem. Note that the clusters together form concentric layers with an overall inside-out organization. Modified from Leergaard et al. (2000a,2000b). (C) Reconstruction showing the topography of pontine terminal fields arising in the rat SI whisker barrel field (modified from Leergaard and Bjaalie, 2002, with permission; see also Leergaard, 2003). (C) (Left) The anterograde tracers biotinylated dextran amine (BDA, blue) and fluororuby (FR, red) were injected into electrophysiologically defined individual whisker representations in SI (upper left inset) and the distribution of labeling was computer reconstructed in three dimensions (lower left inset). (Middle) Computer-generated dot map showing the distribution of BDA-labeled (blue) and FR-labeled (red) fibers within the ipsilateral pontine nuclei. The clusters of red dots surround the clusters of blue dots externally. (Right) The outer boundaries of labeled clusters are demonstrated by solid surfaces. The labeled clusters arising from the same row of SI barrels are located in dual lamellae that are shifted from internal to external.

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Tectum Another important input station to the Pn is provided by the tectum. Especially the superior colliculus sends a major, ipsilateral projection to the peduncular and lateral regions of the caudal Pn. The latter projection appears to be topographically organized: the medial part of the superior colliculus projects primarily to peduncular regions whereas its lateral part rather terminates in the ventrolateral pontine gray. A contralateral projection to the dorsomedial and medial parts of the peduncular subnuclei has also been described (Burne et al., 1981). In the rabbit, it was established (Wells et al., 1989) that the projection is limited to the dorsolateral Pn when the injection was restricted to the superficial laminae of the superior colliculus. Pretectal areas also project to the lateral-most pontine areas but at somewhat more rostral levels. The inferior colliculus appears to provide a rather sparse projection to the lateral Pn areas caudal to the termination fields of the superior colliculus (Burne et al., 1981). Additional visually related input to the Pn is derived from the ventral lateral geniculate nucleus (Ribak and Peters, 1975; Wells et al., 1989). Hypothalamus The hypothalamus, in particular the mammilary nuclei, provides another well-known projection to, primarily, the medial and dorsomedial parts of the pontine gray (Cruce, 1977; Hosoya and Matsushita, 1981; Aas and Brodal, 1988; Mihailoff et al., 1989; Allen and Hopkins, 1990; Liu and Mihailoff, 1999). These projections show convergence with projections from the prefrontal cortex implying that some form of integration of limbic and/or autonomic processes may take place within the Pn (Allen and Hopkins, 1998). Other Sources Apart from the major projections mentioned above, a study by Mihailoff et al. (1989) revealed that the Pn receive afferents from a score of other spinal and brain stem centers, which are very briefly mentioned here. Most of these projections have been confirmed in the cat (Aas, 1989). A spinopontine projection arises from marginal regions of the dorsal nucleus (Clarke’s nucleus: see also Yamada et al., 1985) and terminates predominantly in the caudal Pn. A corresponding projection from the external cuneate nucleus to the caudal Pn has also been identified (Kosinski et al., 1986). In addition, a pontine projection also originates from the dorsal column nuclei and includes an area directly ventrolateral to the cuneate nucleus. This projection mainly arises as collaterals of medial lemniscus fibers that project to the ventral posterolateral nucleus of the

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thalamus (Kosinski et al., 1988b) and was already identified by Ramón y Cajal (1911). Additional somatosensory projections to the Pn arise from the interpolar subdivision of the spinal trigeminal nucleus (Swenson et al., 1984). Spinal, trigeminal, and dorsal column nuclear projections to the Pn provide a pathway for peripheral sensory information to be transmitted rather directly to the Pn, thus contrasting to incoming sensory information from the somatosensory cortex (Azizi et al., 1986; Kosinski et al., 1988a). A vestibulopontine projection originates specifically from the spinal vestibular nucleus and adjacent nucleus X. Reticulopontine projections are described as rather sparse and originate mainly from the ventral reticular, gigantocellular pars alpha and the paragigantocellular nuclei. At least some of these projections appear to be GABAergic (Border et al., 1986). In the cat, it has been shown that a cholinergic pontine input may originate from the dorsolateral pontine tegmentum (Aas et al., 1990). The accessory optic system, such as the medial and lateral terminal nuclei, but also certain accessory oculomotor nuclei such as the nucleus of Darkschewitsch and the Edinger–Westphal nucleus, also provide input to the Pn. Noteworthy, a relatively large, and partially GABAergic (Border et al., 1986), pontine projection arises from the region near the lateral and ventral borders of the red nucleus as well as from the adjacent deep mesencephalic nucleus. A scant projection to the Pn finds its origin from neurons located within the peripeduncular nucleus and adjacent portions of the substantia nigra and from ventral and lateral regions of the periaquaductal gray. Besides the already mentioned diencephalic projection from the ventral lateral geniculate nucleus to the Pn, a notable and predominantly ipsilateral projection arises from the zona incerta (Mihailoff, 1995). Finally, pontine input from the locus coeruleus and from the raphe nuclei suggests noradrenergic and serotonergic projections, respectively. When considering all the subcortical afferent sources to the Pn, it is noted that some of these areas (i.e., dorsal column nuclei, vestibular nuclei, reticular formation) also send mossy fibers to the cerebellum, and, thus, besides establishing a rather direct connection with the cerebellum also may provide an indirect cerebellar pathway via the Pn (Mihailoff et al., 1989). As yet, it is not known if these projections arise as collaterals of the projection to the Pn and so the functional significance of this particular circuitry must await further analysis.

Efferents of the Basilar Pontine Nuclei The Pn project to the cerebellum mostly via the contralateral middle cerebellar peduncle. A sparse to

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moderate projection to the cerebellar nuclei, in particular to the lateral cerebellar nucleus, has been demonstrated with physiological (Shinoda et al., 1992) and anatomical techniques (Eller and Chan-Palay, 1976; Mihailoff, 1993, 1994) but could not be substantiated anatomically in the cat (Dietrichs and Walberg, 1987). The relative sparsity of the, presumably collateral, projection to the cerebellar nuclei from the Pn contrasts the clear-cut and rather dense collateral projection which arises from the reticulotegmental nucleus of the pons (Gerrits and Voogd, 1987; Mihailoff, 1993). Pontocerebellar fibers terminate as mossy fibers in the granular cell layer where they are distributed widely in the hemispheres, including the paraflocculus, and to the vermal lobules VI to IX. The vestibulocerebellar parts (nodulus and flocculus) as well as the anterior lobe do not appear to receive a pontocerebellar innervation, although Gerrits (1985) demonstrated a major pontine projection to especially the intermediate and hemispheral parts of the contralateral anterior lobe in the cat. Some evidence suggests that pontocerebellar mossy fibers use glutamate as the major transmitter (Beitz et al., 1986; Border and Mihailoff, 1991). It is generally believed that the pontocerebellar projection is precisely organized and characterized by complex patterns of convergence as well as divergence; that is, it has been demonstrated that single pontine neurons can project to two different lobules within the same hemisphere or even to both hemispheres (Mihailoff, 1983; Rosina and Provini, 1984). Convergence is readily apparent in retrograde transport studies where relatively small cortical injections may label different areas within the Pn (Azizi et al., 1981; Eisenman, 1981; Mihailoff et al., 1981a). Nevertheless, in the cat, it has been suggested that the projections to paraflocculus are mostly governed by topographical patterns. Layers of pontine neurons project to different parafloccular folia as was demonstrated in this anatomical study employing multiple fluorescent tracers (Nikundiwe et al., 1994). The number of double labeled pontine neurons as well as the position of lamellar-like layers of labeled pontine neurons was strongly dependent on the interfoliar distance of the injection sites. In a recent anterograde study in the rat, it was shown that labeling of relatively small groups of pontine neurons results in mossy fiber projections to multiple, parasagittally organized, strips of cerebellar cortex, which were observed bilaterally but with a contralateral preponderance (Serapide et al., 2001). The strips with mossy fiber terminals were well-defined and frequently sharply bordered. Even the smallest injections still resulted in the labeling of multiple strips, whereas larger injections failed to show a zonal-like pattern of termination. The authors conclude that the pontocerebellar projection is

organized in multiple pathways that project to specific sets of strips of cerebellar cortex. Since the corticonuclear projection also follows a zonal pattern, with each output to a different part of the cerebellar nuclei, which in turn are suggested to subserve different functional purposes, it would be interesting to study the potential relation between the corticonuclear and pontocortical zonal organization. Mostly due to the converging and diverging termination characteristics, it is still difficult to describe the precise anatomical relations between the Pn and the cerebellum. Using retrograde tracer techniques, it has been possible to state that the posterior vermis mainly receives pontine-relayed afferences from visual, auditory, and somatosensory regions of the cerebral cortex. In addition, certain areas of lobules VI and VII also receive mossy fibers from pontine regions that are under tectal influence (Azizi et al., 1981). In line with the anterograde study of Serapide et al. (2001), a parasagittal organization of pontocerebellar projections was also established for lobule VIII (Eisenman, 1981). A medial zone receives mossy fibers from medial and ventrolateral regions of the caudal part of the pontine gray; an intermediate zone from the intermediate pontine region; and a lateral zone from medial, ventrolateral, and dorsal areas of the pons. Pontocerebellar projections to the hemispheres, originate from many regions in the Pn (Mihailoff, 1993). It was noted that the projections to the lobulus simplex are organized rather differently from those to the other hemispheral components because most of its mossy fibers originate from neurons located ipsilaterally within the ventral part of the rostal pontine gray. Crus I receives most mossy fibers from neurons distributed along the medial, ventral, and lateral regions of the contralateral pontine gray, whereas more central pontine areas project to the Crus II. The paramedian lobule is supplied by mossy fibers emanating from the central region of the contralateral pons. The peduncular subnuclei also supply mossy fibers to the cerebellar hemispheres. The paraflocculus receives a projection from a medial and a lateral column of cells as well as from scattered neurons within the peduncular subnuclei, which may convey both auditory and visual signals (Azizi et al., 1985; Glickstein and Stein, 1991). Different pontine projections are directed to the dorsal and ventral paraflocculus (Eisenman, 1980), which may well reflect the difference in function of these two parafloccular regions (Gerrits and Voogd, 1982; Gerrits et al., 1984; Azizi et al., 1985).

Functional Considerations The Pn tie the cerebral and the cerebellar cortices via two conspicuously large fiber bundles, i.e., the cerebral

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peduncle and the medial cerebellar peduncle. However, it has become evident that the Pn itself is not a mere link between both cortices but can also be seen as a major integrating center. Not only because different cerebral cortical areas may converge upon individual pontine neurons (Ruegg et al., 1977; Potter et al., 1978; Leergaard et al., 2000a, 2000b), but also since many diverse brain stem centers as well as the spinal cord also provide an input to the Pn (Mihailoff et al., 1989). It is interesting that many of these centers also provide a direct mossy fiber input to the cerebellum. Finally, cerebellar output feeds back into the Pn mostly by nonGABAergic pathways. As such, the Pn cannot be interpreted as a mere station of passage of the information flow from cerebrum to cerebellum, but should be regarded as a highly integrative center, capable of updating the cerebellum with integrated information on ongoing somatosensorimotor, visual, auditory, and autonomic and affective processes. Glickstein and Stein (1991), have suggested that the visual and somatosensory input to the Pn and their relay to the cerebellum may present an important pathway for the control of visually guided movements. Schwarz and Thier have recently argued that the Pn are specifically adapted to form a necessary interface enabling cerebellar processing of cerebral information. The necessity of such an interface would transpire from the greatly different computational principles that govern the cerebral cortex and the cerebellum (Schwarz and Thier, 1999).

Reticulotegmental Nucleus of the Pons The reticulotegmental nucleus of the pons (RtTg) is located dorsal to the medial lemniscus, along the midline. It appears to be continuous with the dorsomedial aspect of the pontine gray, especially at rostral levels. Whether or not the neurons located within the medial lemniscus should be regarded as part of the RtTg or of the Pn remains a matter of conjecture (Mihailoff et al., 1988; Schwarz and Thier, 1996). Torigoe et al. (1986b) describe two cytoarchitectonically distinct portions of the RtTg. A central part (RtTgC) consists of rather tightly packed cells, whereas a pericentral part (RtTgP) is composed of loosely packed small neurons. The RtTgC is located dorsal to the medial lemniscus over the caudal two thirds of the Pn. At some points it is continuous with the dorsomedial Pn area (Mihailoff et al., 1981b). Caudal to the Pn, the RtTg extends caudodorsally until it dissipates just rostral and ventral to the abducens nucleus. The pericentral part is found at the dorsolateral margins of the RtTgC rostrally, but is positioned ventral to its caudal part (Fig. 1). Schwarz and Thier (1996), based on a morphological study of intracellularly injected projection neurons in both Pn

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and RtTg, claim that differences in size and dendritic pattern of these cells can be explained by a dorsoventral gradient that basically ignores the border between both nuclei. RtTg afferents originate from the cerebellar nuclei, the cerebral cortex, and a number of brain stem centers. Excitatory cerebellar afferents are mainly derived from the Lat and, to a lesser degree, from the interposed nuclei (Angaut et al., 1985a; Torigoe et al., 1986a; Schwarz and Schmitz, 1997; Verveer et al., 1997; Buisseret-Delmas et al., 1998; Teune et al., 2000). Afferents from the cerebral cortex stem from layer V neurons. The cingular cortex, thought to be homologous with the frontal eye fields of the cat and primates, projects to the RtTgC, whereas the pericentral RtTg receives afferents from the somatomotor cortex, in particular (Torigoe et al., 1986b). Brain stem projections arise from various visuomotor areas such as the nucleus of the optic tract, the contralateral superior colliculus, the medial terminal nucleus, the ventral lateral geniculate nucleus, the ventral tegmental relay zone, the anterior and posterior pretectal nuclei, and the supraoculomotor periaquaductal gray. Moreover, a number of areas that may subserve a wider variety of motor behaviors also supply afferents to the RtTg, such as the zona incerta, the fields of Forel, the interstitial nucleus of the mlf, the vestibular nuclei, and the reticular formation (Burne et al., 1981; Torigoe et al., 1986a; Redgrave et al., 1987; Gayer and Faull, 1988; Matsuzaki and Kyuhou, 1997). A limbic input is furthermore supplied via afferents from the mammilary nuclei, the lateral hypothalamic area, the habenula, the preoptic nuclei and the diagonal band of Broca (Cruce, 1977; Terasawa et al., 1979; Hosoya and Matsushita, 1981; Torigoe et al., 1986a; Allen and Hopkins, 1990). The RtTg sends mossy fibers to most lobules of the cerebellar cortex with a slight contralateral preponderance. In the cat only the nodulus is reported to be devoid of RtTg afferents (Gerrits and Voogd, 1986). Strong RtTg input has been described to terminate within the flocculus and ventral paraflocculus and the oculomotor vermal areas of lobules VI, VII and VIII (Blanks et al., 1983; Cazin et al., 1984; Yamada and Noda, 1987; Gayer and Faull, 1988; Päällysaho et al., 1991). Mossy fiber input to the flocculus arises from caudal regions in the RtTg whereas projections to the paraflocculus originate from more rostral parts (Osanai et al., 1999). Collaterals of the RtTg mossy fibers terminate within the Lat and, to a lesser degree, in the interposed nuclei (Brodal et al., 1986; Gerrits and Voogd, 1987; Schwarz and Schmitz, 1997; Verveer et al., 1997). Furthermore, RtTG efferents have been described to terminate within the medial vestibular nucleus of the rabbit (Balaban, 1983), the nucleus prepositus hypoglossi of the rat (Cazin et al., 1984; Korp et al., 1989), and even the visual cortex of

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the mouse (Castaneyra-Perdomo, 1984). It is not known if the same neurons also provide input to the cerebellum. Due to the input of many visuomotor centers to the RtTg and its projection to the oculomotor vermis and flocculus, it is obvious that this precerebellar nucleus functions as an important relay center for optokinetic processes and various visuovestibular interactions (Averbuch-Heller and Leigh, 1996; Buttner-Ennever and Horn, 1997). However, lesion, behavioral, and electrophysiological studies have implied that many neurons of the RtTg may be involved in the execution of movements and locomotion as well (Chesire et al., 1983, 1984; Brudzynski and Mogenson, 1984; Matsunami, 1987; Hammer and Klingberg, 1991). Whether or not this behavioral involvement of the RtTg is in any way related to the visuomotor input and/or to the limbic inputs of the RtTg remains to be determined.

LATERAL RETICULAR NUCLEUS The lateral reticular nucleus (LRt) is situated ventrally in the medulla oblongata, dorsolateral to the inferior olivary complex and ventromedial to the spinal tract of the trigeminal nerve. Its caudal pole first appears slightly caudal to the inferior olive and it extends to the level of the rostral quarter of the inferior olive. As a major source of mossy fiber projections to the cerebellar cortex, the LRt plays an important role in the control of motor activity and coordination, especially when related to the realization of fine and target-reaching movements of the forelimb (Oscarsson, 1958; Clendenin et al., 1974b; Illert et al., 1977; Arshavsky et al., 1978; Alstermark et al., 1987b; Ekerot, 1990a). More recently, involvement of the LRt in nociceptive processes has received increasing attention (Murphy and Behbehani, 1993; Ness et al., 1998). In particular neurons with descending projections to the dorsal horn of the spinal cord are thought to play a role in these processes (Gebhart and Ossipov, 1986; Liu et al., 1989, 1990). However, although precerebellar neurons within the LRt have been implicated, especially in visceral nociceptive processing (Ness et al., 1998), as yet, it is not known to what extent precerebellar LRt neurons are involved in these processes (Lee and Mihailoff, 1999). Here, we discuss the LRt only as a structure that mainly consists of precerebellar neurons.

Cytoarchitecture As a “reticular” nucleus in the very general sense of the word, the boundaries of the LRt cannot always be precisely defined. Only at caudal levels can its contours be clearly delineated, while at rostral levels the nucleus becomes more diffuse and sometimes poorly discernible

from the surrounding reticular formation. The caudorostral extension of the LRt is about 2800 μm, while mediolaterally and dorsoventrally its greatest dimensions are about 1400 and 800 μm, respectively. Kapogianis et al. (1982a) estimated the caudorostral length of the LRt to be about 1000 μm shorter. This discrepancy is due to the fact that the rostral pole of the LRt has been redefined. Using immunohistochemical techniques, Kaneko et al. (1989) first suggested that Paxinos and Watson’s (1986) linear nucleus of the medulla should be included in the LRt, a hypothesis that was confirmed by a retrograde tracing study of Newman and Ginsberg (1992) and by cytologic, enzyme histochemical, and anterograde and retrograde transport studies of Cella et al. (1992). In most mammals studied, it is possible to recognize a parvicellular, a magnocellular, and a subtrigeminal part within the LRt (Brodal, 1943; Walberg, 1952; Valverde, 1961; Ramón-Moliner and Nauta, 1966; Kitai et al., 1972; Hrycyshyn and Flumerfelt, 1981a; Hrycyshyn et al., 1982; Kapogianis et al., 1982a). However, in each subdivision, cells of a predominant average diameter are intermingled with cells of other sizes (Kapogianis et al., 1982b). The subtrigeminal part is characterized by predominantly medium-sized cells, but comprises many large cells as well. Moreover, the only distinct subdivision is represented by the subtrigeminal part, while the magnocellular and the parvicellular subdivisions partially overlap, particularly at midrostral levels. Their boundaries, therefore, remain arbitrary (Menétrey et al., 1983; Shokunbi et al., 1986). Caudally, in transverse sections, the LRt first appears at about 200 μm below the caudal pole of the inferior olive as a cluster of cells which rapidly develops into a principal, dorsomedially oriented, ovoid part and a more superficially located cell strip which is connected to the principal division by thin bridges of neuropil. Through this caudal extension of the LRt, it is possible to approximately delineate its magnocellular part first dorsomedially (stippled area in Fig. 3, levels 1–3) and then dorsomedially and dorsolaterally (Fig. 3, levels 4 and 5), while its parvicellular part corresponds to the ventral region of the principal LRt and to the strip of cells running along its ventrolateral boundary. At about 800 μm of the LRt length, the medial portion of this strip begins to fuse with the principal division, while the thin connecting bridges become progressively larger until no separation is left between the medial twothirds of the strip and the principal LRt. The lateral third of the strip virtually disappears. The very lateral part of the LRt now begins to develop in what more rostrally will become the subtrigeminal division (Fig. 3, levels 6–8). At the same levels, the ventral displacement of the caudo- and rostroventrolateral reticular nucleus (CVL and RVL) (Paxinos and Watson, 1986), joined a

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FIGURE 3 Diagrammatic representation of the rat inferior olivary complex and lateral reticular nucleus (LRt), based on transverse sections, spaced 160 μm. Caudal is at the bottom left (level 1); rostral is at the top right (level 21). The various olivary subdivisions are indentified by different hatching patterns. Abbreviations used: Amb, nucleus ambiguus; IOD, dorsal accessory olive; dfIOD, dorsal fold of IOD; DC, dorsal cap of Kooy; DM, dorsomedial group; DMCC, dorsomedial cell column; CVL, caudoventrolateral reticular nucleus; β, nucleus β; Li, linear nucleus of the medulla; LRtm, magnocellular part of the LRt; LRtp, parvicellular part of the LRt; LRts5, subtrigeminal part of the LRt; PO, principal olive; RVL, rostroventrolateral reticular nucleus; VLO, ventrolateral outgrowth; IOM, medial accessory olive. Bar equals 1 mm.

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little more rostrally by the nucleus ambiguus, causes a flattening and indentation of the dorsal boundary of the LRt. Ultimately this leads to the division of the LRt into a medial and a lateral (subtrigeminal) subdivision (LRtS5, Fig. 3, levels 7–10). Along with these developments in the morphology of the LRt, a rearrangement of the internal cytoarchitecture also takes place: the magnocellular part now has shifted ventromedially, but many large cells are also located within the surrounding parvicellular areas. Of the two divisions in which the LRt has been split by the CVL/RVL, the medial one is rather well-defined in its ventral aspect, while its dorsal part is more diffuse with two cell-poor extensions toward the medial reticular formation (Fig. 3, levels 10–14). The lateral division, on the other hand, has wellcircumscribed boundaries throughout its extent and represents the most rostral subdivision of this nucleus. Roughly triangular in shape at more caudal levels, it becomes progressively narrower in its mediolateral diameter while extending dorsally to adapt the form of a diagonal bar (nucleus linearis, Li). This part of the LRt consists of a long arm, directed dorsomedially, and of a short arm, directed more medially. The medial extremity of this short arm shows a tendency to bend steeply ventrally, but the boundaries at this point are less distinct than those at the lateral side. Finally, the two arms of the nucleus progressively shorten until their joining point remains as the rostral-most part of the LRt (Fig. 3, level 18).

Afferents to the Lateral Reticular Nucleus Data available from investigations in different mammals indicate that the main afferent input to the LRt is a bilateral, topographically organized projection from the spinal cord (Brodal, 1943; Morin et al., 1966; Kunzle, 1973; Mizuno and Nakamura, 1973; Corvaja et al., 1977; Martin et al., 1977; Corvaja and d’Ascanio, 1981; Hrycyshyn and Flumerfelt, 1981b; Flumerfelt et al., 1982; Menétrey et al., 1983; Shokunbi et al., 1985; Westman et al., 1986; Cella et al., 1991; Rajakumar et al., 1992). Moreover, the LRt receives afferents from several supraspinal centers, such as the contralateral red nucleus, the medial cerebellar nucleus, the cerebral cortex, the vestibular nuclei, and, as demonstrated in the cat, the superior colliculi and the hypothalamus (Kuypers, 1958; Walberg, 1958a; Walberg, 1958b; Hinman and Carpenter, 1959; Walberg and Pompeiano, 1960; Kawamura et al., 1974; Künzle and Wiesendanger, 1974; Corvaja et al., 1977; Hrycyshyn and Flumerfelt, 1981b; Qvist et al., 1984; Dietrichs et al., 1985; Shokunbi et al., 1986; Rajakumar et al., 1992). Current knowledge about the organization of the massive spinal input to the LRt has been mainly derived

from electrophysiological studies in the cat, which have revealed the existence of at least three distinct afferent tracts, all ascending in the lateral funiculus of the spinal cord, namely, the bilateral ventral flexor reflex tract (bVFRT), the ipsilateral forelimb tract (IFT) and a group of propriospinal neurons located in the third and fourth cervical segments (Holmqvist et al., 1960; Lundberg and Oscarsson, 1962; Rosén and Scheid, 1973a, 1973b; Clendenin et al., 1974b; Clendenin et al., 1974c; Illert et al., 1978; Alstermark et al., 1981a, 1984; Ekerot, 1990a, 1990b, 1990c). Apart from the three tracts in the lateral funiculus, one ascending tract in the dorsal funiculi has been distinguished (Clendenin et al., 1975; Ekerot, 1990a), but not yet completely characterized. The spinal projections to the LRt have been shown to result in both monosynaptic excitatory and inhibitory responses. No comparable experiments, however, have been performed in the rat and no conclusive data yet exist about the anatomical equivalent of the physiologically identified spinal pathways. The following data on the spinal afferents to the LRt only refer to retrograde and anterograde tracer studies performed in the rat and demonstrate the projecting spinal cells and their termination area within the LRt. As a result of the retrograde labeling following HRP injections in the LRt, Menétrey et al. (1983) found that labeled neurons were present at all spinal levels and in particularly large numbers in the cervical and lumbar enlargements. Labeled cells were located, with contralateral predominance, in all segments of the spinal cord, within laminae VII, VIII, and X, in the reticular expansion of the dorsal horn, in the superficial layers of the dorsal horn, and in the nucleus of the dorsolateral funiculus. Next to this labeling pattern common to all spinal segments, a specific pattern was shown to be exclusively present in the cervical and lumbar enlargements, which contained additional labeled neurons in the ipsilateral lamina VII and in the contralateral laminae III and IV, respectively. These results were only partly confirmed by Shokunbi et al. (1985), who, using the same technique, showed that discrete HRP placements in the caudomedial part of the LRt resulted, in the cervical segments, in labeled cells mainly in laminae V and VII and, less heavily, in laminae III and IV ipsilaterally, in laminae VII and VIII contralaterally, and in lamina X. At thoracic levels, few cells were labeled bilaterally in lamina VII, while the lumbar segments showed a certain amount of labeled neurons in the contralateral lamina VIII and adjacent lamina VII. Injections of HRP in the caudolateral part of the LRt, instead, resulted in retrograde labeling of a few ipsilateral cervical neurons located in lamina VII and a prominent labeling of the contralateral lumbar segments, where the HRP-containing cells were mainly located in

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laminae IV and V. No labeled neurons were encountered within the superficial-most laminae of the dorsal horn. This last conclusion was in accordance with a retrograde and anterograde transport study by Cella et al. (1991) who noticed that small injections of WGA–HRP in the dorsal LRt regions mainly resulted in retrogradely labeled cells within the ipsilateral cervical cord. Injections placed in the ventral LRt predominantly labeled neurons located bilaterally within the lumbar spinal cord. Subtrigeminal injections are characterized by a combination of bilateral cervical and lumbar labeling. The anterograde labeling in the LRt subsequent to injections with Phaseolus vulgaris-Leucoagglutinin (PHA-L) in the spinal cord resulted in four different labeling patterns, depending on the segmental level and laminae where the injection had been placed. Injections in the cervical enlargement demonstrated, when placed in the dorsal horn, an essentially ipsilateral projection to the LRt, in the caudal dorsolateral magnocellular area of the nucleus and, when placed in ventral horn, a bilateral projection to the LRt, in a more dorsomedial magnocellular area. Instead, injections in the lumbar enlargement demonstrated a bilateral projection to the LRt in more ventral, parvicellular areas. Termination patterns varied only slightly with injections placed in either dorsal or ventral horn. Labeled terminals were occasionally found in the rostromedial and rostrolateral regions of the LRt, which fits the attribution of these two areas to the LRt. These results are somewhat at variance with those obtained by Rajakumar et al. (1992), who, using anterograde transport of WGA–HRP, failed to note an ipsilateral spinoreticular projection arising from lumbar levels. Furthermore, they were unable to differentiate between dorsal and ventral horn projections. Common to these different studies is the finding that the largest amount of spinal neurons projecting to the LRt is found within the cervical and lumbar enlargements and that a specific ipsilateral pathway takes its origin from the cervical cord. It is attractive to speculate that these two different projecting patterns to the LRt could represent anatomical equivalents of the physiologically demonstrated iFT and bVFRT of the cat, respectively. Apart from these considerations most retrograde and anterograde tracer studies basically resulted in bilateral labeling patterns in the spinal cord and LRt, respectively. However, a retrograde double labeling study by Koekkoek and Ruigrok (1995) demonstrated that individual spinoreticular neurons located throughout the spinal cord project to either the ipsilateral or the contralateral LRt, but only seldom to both regions simultaneously. In conclusion, it is obvious that the anatomical projections of the spinal cord to the LRt, related to the establishment of multiple, physiologically distinguishable, spinoreticular tracts, suggest a highly

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complex functional organization of the spinoreticulocerebellar pathway. In the cat, physiological evidence was found that part of the spinoreticular projection may arise as collaterals of the spinocerebellar pathway (Alstermark et al., 1990). However, for the rat no systematic study seems to be available on the degree of collateralization of the spinoreticular, spinocerebellar, and spinoolivary projections. With respect to the nonspinal afferents to the LRt the rubral projection appears to be the most extensive. It arises from neurons in the caudal two-thirds of the contralateral red nucleus. In a retrograde tracer study employing HRP, Shokunbi et al. (1986) showed that terminals of neurons located ventrally and ventrolaterally in the red nucleus distributed terminals to the rostrolateral part of the LRt, while neurons located dorsally and dorsomedially projected to the rostromedial LRt. In a subsequent anterograde study using WGA–HRP (Rajakumar et al., 1992), only a small projection from the contralateral red nucleus to the lateral part of the rostral half of the LRt was found. It is generally believed that the rubral projection to the LRt consists of collateralizing rubrospinal tract fibers. A cerebellar input to the LRt mostly arises from the medial cerebellar nucleus (Shokunbi et al., 1986; Rajakumar et al., 1992; Teune et al., 2000). According to Shokunbi and colleagues (1986), this projection is bilateral with ipsilateral preponderance and distributes diffusely throughout the LRt, particularly to its ventral and medial parts, whereas Rajakumar et al. (1992) found that medial afferents only terminated in the contralateral LRt, mainly in the dorsomedial aspect of the rostral twothirds of the magnocellular division. It was noticed by Rajakumar et al. (1992) that these different results might be due to the use of different tracers and the possible uptake of HRP by passing fibers. The input from the cerebral cortex is scant in the rat. Shokunbi et al. (1986) as well as Rajakumar et al. (1992) agree by reporting that it arises from neurons located in layer V of the contralateral frontoparietal sensorimotor cortex and which project to the rostromedial part of the LRt. Projections from the anterior pretectal nucleus to the ventrolateral regions of the caudal medulla oblongata also incorporate the LRt and have been suggested to be related to the nociceptive functions of this region (Zagon et al., 1995). It can be concluded that, while spinal projections to the LRt have already been rather extensively investigated, much less is known about supraspinal afferents to the LRt. Most authors agree in attributing to the caudal one-half of the nucleus the function of integrating the various spinal signals, while the middle third of the nucleus, where rubral, cerebellar, and spinal terminals appear to overlap, would subserve the function of integrating spinal and supraspinal impulses to the

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cerebellum. The rostral part of the LRt could take part in a separate corticocerebellar pathway. On the afferents to the newly defined LRt areas (i.e., the rostromedial and linear parts) no specific experimental data appear to be available.

Efferents of the Lateral Reticular Nucleus The LRt represents one of the most important sources of mossy fibers to the cerebellar cortex, also reaching, by way of collaterals, the cerebellar and lateral vestibular nuclei (Brodal, 1975; Künzle, 1975; Bishop et al., 1976; Matsushita and Ikeda, 1976; Chan-Palay, 1977; Martin et al., 1977; McCrea et al., 1977; Dietrichs and Walberg, 1979; Eisenman, 1982; Hrycyshyn et al., 1982; Dietrichs, 1983; Payne, 1987; van der Want et al., 1987; Qvist, 1989b; Päällysaho et al., 1991; Ruigrok et al., 1995; Parenti et al., 1996; Wu et al., 1999). A few authors also report a minor efferent path to the inferior olive, suggesting a possible role of the LRt within the spinoolivocerebellar system (Brown et al., 1977; Swenson and Castro, 1983a). The LRt projection to the cerebellar cortex passes through the ipsilateral inferior cerebellar peduncle and terminates as mossy fibers in the granular cell layer. Although the pathway is mainly uncrossed, a single nucleus projects to both sides of the cerebellum, except for the input to the lobulus paramedianus, which is exclusively ipsilateral. The mossy fiber terminal arborizations within the cortex are arranged, at least partly, in parasagittal bands (Eisenman, 1982; Hrycyshyn et al., 1982; Wu et al., 1999), possibly interdigitating with the mossy fibers arriving by way of the cuneocerebellar tract (Voogd et al., 1969). Studies using electrophysiologic techniques in the cat, retrograde fluorescent double-labeling techniques in the rat, and a single fiber anterograde study in the rat have demonstrated that individual LRt neurons may innervate both left and right sides of the cerebellar hemispheres, both within and between parasagittal zones (Clendenin et al., 1974a; Payne, 1983; Ghazi et al., 1987; Payne, 1987; Wu et al., 1999). In the rat as well as in the cat, the LRt mossy fibers project to lobules I through V of the anterior lobe, to the rostral part of lobule VI, to the most caudal part of lobule VII, to lobule VIII, to the medial part of the ansiforme lobule, and to the simple and paramedian lobules. Minor projections have also been demonstrated to the caudal part of lobule VI, to the rostral part of lobule VII, to the lateral part of the ansiforme lobule, to the paraflocculus, and to the flocculonodular lobe (Brodal, 1943; Clendenin et al., 1974a; Künzle, 1975; Kimoto et al., 1978; Dietrichs and Walberg, 1979; Eisenman, 1982; Hrycyshyn et al., 1982; Payne, 1987; Qvist, 1989a; Päällysaho et al., 1991).

There is agreement among different authors that within the anterior lobe, lobules II and III, the classic hindlimb-related area, receive input mainly from the ventral, parvicellular LRt, which in turn receives mainly terminals from lumbosacral neurons (see above). Lobules IV and V, which constitute the forelimb area, receive input mainly from the magnocellular LRt, which is the predominant relay station for the cervical cord. Moreover, the pyramis (lobule VIII) is supplied exclusively by the parvicellular LRt and the copula pyramidis by the subtrigeminal part. The simple, ansiform and paramedian lobules mainly receive projections from the magnocellular LRt (Eisenman, 1982; Hrycyshyn et al., 1982). However, a considerable overlap characterizes the projections from different LRt regions to the various subdivisions of the cerebellum, suggesting a rather diffuse organization of the LRt– cerebellar system. Both in the cat and the rat, the LRt projection to the vermis is much denser than the projections to the hemispheres, but in the rat, unlike in the cat, the projection to the anterior part of the cerebellar hemispheres notably exceeds the one to the lobulus paramedianus. In the rat, as well as in the opossum, it is suggested that the LRt projection also reaches nonspinal areas of of the cerebellum, like the simple and ansiforme lobules (Künzle, 1975; Martin et al., 1977). Moreover, the topographic arrangement in the projection of the LRt found in the cat, has been denied (Eisenman, 1982) or could be only partly confirmed (Hrycyshyn et al., 1982) for the rat. Very little is known about the cerebellar projections arising from the rostromedial and rostrolateral regions of the LRt, but there is agreement between the results of Newman and Ginsberg (1992) and of Cella et al. (1992) in showing that these two areas participate in the ascending pathway to the cerebellum. The cerebellar nuclei as well as the lateral vestibular nucleus also receive predominantly ipsilateral projections from the LRt (Ruigrok et al., 1995; Parenti et al., 1996; Wu et al., 1999). According to the detailed retrograde study by Parenti et al. (1996), the lateral cerebellar nucleus receives its terminals mostly from the rostral dorsomedial LRt, whereas the interposed nuclei are targeted by efferents from its dorsolateral and central parts. Finally, the caudal regions of the intermedioventral region would seem to prefer to project to the medial cerebellar nucleus. However, Wu et al. (1999) showed that individual neurons can provide collaterals to multiple nuclei simultaneously. Neurons within the boundaries of the LRt have been reported to project to the spinal cord (e.g., Janss and Gebhart, 1988; Liu et al., 1989). Some of these cells may possess collaterals to the periaquaductal central gray (Lee and Mihailoff, 1999). However, as yet it is not

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known if and, if so, to what extent these cells project to the cerebellum.

Functional Considerations From the afferent projections to the LRt it will be obvious that this center is predominantly concerned with the regulation of somatic sensorimotor events. Spinal and supraspinal impulses necessary for the control of movements of the ipsilateral forelimb are integrated and possibly related or coordinated with motor activity in other parts of the body. The outcome of LRt integrative properties is relayed to the cerebellum where additional integrative processes result in an adequate cerebellar output (Parenti et al., 1996; Ezure and Tanaka, 1997). Of the three characterized spinal pathways, the bVFRT, the iFT, and the C3–C4 propriospinal neurons, the first is thought to participate in a more global motor control while the latter two appear to be specifically related to the movements of the ipsilateral forelimb (Clendenin et al., 1974b, 1974c; Alstermark et al., 1981b; Ekerot, 1990c). These two general functions to which the LRt contributes very probably correspond to different regions of the nucleus, although some overlap may exist. Not only do they receive inputs from different levels of the spinal cord but they also send afferents to different regions of the cerebellum. In the regulation of sensorimotor control, the bVFRT has the specific characteristic of being activated by cutaneous afferents as well as by group II and III muscle afferents that participate in limb flexor reflexes (Eccles and Lundberg, 1959). Moreover, the integration and transfer of information concerning peripheral motor behavior is also influenced by descending motor pathways, since the bVFRT is monosynaptically activated by stimulation of the lateral vestibulospinal tract (Holmqvist et al., 1960; Clendenin et al., 1974b). This integration of input is indicative for the role of the bVFRT in postural control and coordination of movements and it has suggested that the bVFRT carries information about activity in spinal motor centers influenced by segmental afferents and descending motor paths (Clendenin et al., 1974b; Arshavsky et al., 1978). Besides the inhibitory and excitatory inputs mediated by the bVFRT, single LRt units responding to peripheral stimuli have also been shown to receive inhibitory or excitatory input from the cerebral cortex (Agree and Kitai, 1967). Indeed, a direct route from the cerebral cortex reaches the LRt through the pyramidal tract, while an indirect cortical influence is probably mediated by the red nucleus (Allen and Tsukahara, 1974). The combination of both excitatory and inhibitory cortical control of the LRt may serve as a mechanism

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for fine regulation of the tonic excitatory drive from the LRt to the cerebellar granule cells and cerebellar nuclei (Shokunbi et al., 1986). With regard to the motor control of the ipsilateral forelimb, the C3–C4 propriospinal neurons play an important role in target-reaching movements (Alstermark et al., 1981b). These neurons are activated by the corticospinal, rubrospinal, and tectospinal tracts but also, though weakly, by afferents of the ipsilateral forelimb (Illert et al., 1977; Alstermark et al., 1987a, 1987b, 1987c; Ekerot, 1990a, 1990b). In the rat, it is not known if the neurons in the central cervical nucleus (Verburgh et al., 1989) have similar characteristics. The iFT controls fine movements of the forelimb, such as grasping movements following the target-reaching movement. It is strongly activated from peripheral receptive fields of the ipsilateral forelimb. Both the bVFRT and the iFT consist of an inhibitory and an excitatory component (I-iF and E-iF) (Ekerot, 1990b). The large number of LRt neurons with a convergent input from the bVFRT and the I-iF tract suggests that the two functional parts of the LRt represent different aspects of a common physiologic system (Ekerot, 1990a, 1990b, 1990c). It has been suggested that the LRt is involved not only with sensorimotor activity but also with pain mechanisms, because electrical stimulation of brain stem sites, including the LRt, has been shown to modulate or suppress nociceptive reflexes (Sessle and Hu, 1981; Dostrovsky et al., 1982; Gebhart and Ossipov, 1986; Tanaka and Toda, 1986; Liu et al., 1990; Sotgiu and Bellinzona, 1991; Mineta et al., 1995). Moreover, a retrograde tracer technique study concerning spinal afferents to the LRt (Menétrey et al., 1983) has shown that the lateral portion of the nucleus may receive a projection from laminae I and II of the dorsal horn. These two laminae of the dorsal horn are known to receive terminals from thin afferent (Aδ and C) fibers. However, the location of these neurons within the boundaries of the LRt as well as their participation as a precerebellar source is doubted (e.g., see Shokunbi et al., 1986; Cella et al., 1991; Tavares and Lima, 1994; Lee and Mihailoff, 1999). Neuroanatomical and electrophysiologic studies have implicated the LRt in controlling autonomic cardiovascular activity. Some authors consider the LRt as a part of the system of neuron populations known in the cat as the ventrolateral medulla (VLM) (Henry and Calaresu, 1974; Loewy and McKellar, 1981; Spyer, 1981; Reis et al., 1984; Ciriello et al., 1986; Duffin and Aweida, 1990). However, as yet, the relation, if any, between the LRt and the VLM is far from clear. Furthermore, it must be noted that the involvement of the LRt in cardiorespiratory events may be also related to its functions in motor control, since the mechanic respiratory movements also depend on pure somatic motricity (Cella

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et al., 1991). Indeed, a convergence of information on respiratory and locomotor rhythms on the same neurons within the LRt has recently been demonstrated (Ezure and Tanaka, 1997). The role of the linear and rostromedial parts of the LRt also remains to be elucidated. Therefore, at the moment, it is premature to attempt a more global explanation of the way the various functional properties of the LRt complete or influence each other or to try to define which of its regions are specifically involved in a particular function.

INFERIOR OLIVARY NUCLEUS The inferior olivary nuclear complex is a prominent bilateral aggregation of neuronal cells located ventromedially in the caudal part of the medulla oblongata. It receives afferent connections from many different sources in the brain stem and spinal cord. Its efferent fibers are the sole source of cerebellar climbing fibers (Desclin, 1974) and, as such, this nucleus has a marked effect on cerebellar functioning.

Cytoarchitecture As in most mammals, the inferior olivary complex of the rat consists of three main subdivisions, supplemented by several smaller neuronal subnuclei (Fig. 3). In the rat, the medial accessory olive (IOM) is the largest of the three subdivisions. Its caudal-most border is found just rostral to the pyramidal decussation (approximately 1500 μm caudal to the obex) and extends over approximately 2600 μm to terminate about 200 μm caudal to the rostral-most pole of the olivary complex. The dorsal accessory olivary nucleus (IOD) is, at caudal levels, initially found directly dorsolateral to the IOM (Fig. 3) (levels 10 and 11). At more rostral levels, the principal olive (IOPr) becomes interpolated between IOD and IOM. Further rostrally, the IOD and IOPr merge and this aggregate forms the rostral pole of the olivary complex. Opinions differ somewhat as to the number and name of the various smaller subdivisions and cell groupings as is outlined below. The IOM is usually divided into a caudal and a rostral half or lamella. Azizi and Woodward (1987) further divided the caudal half into a horizontal and a vertical lamella. Within the caudal part of the IOM (levels 7–9, Fig. 3), three subgroupings are generally recognized and referred to as cell groups a, (IOA), b (IOB), and c (IOC) from lateral to medial (Gwyn et al., 1977). Dorsomedially, the nucleus β (IOBe) adjoins IOC. In agreement with Bernard (1987), Nelson and Mugnaini (1988), Ruigrok and Voogd (1990, 2000), and BuisseretDelmas and Angaut (1993), we include the caudal part

of IOC in IOBe (Fig. 3, levels 7 and 8; see also Ruigrok, 1997). IOBe gradually moves dorsally, thus making way for the more rostral part of IOC (Fig. 3, levels 8–10). It terminates about 1250 μm rostral to its caudal pole. A small cell cluster separates from the dorsomedial margin of the IOBe. This cell cluster enlarges somewhat and, after merging with its contralateral counterpart, terminates after approximatly 300 μm (level 16). In analogy with the situation in the cat, this cell group was identified as the dorsomedial cell column (IODMCC) by Azizi and Woodward (1987), Bernard (1987), Nelson and Mugnaini (1988) and Ruigrok and Voogd (1990, 2000). In the cat, however, the IODMCC is associated with the ventral lamella of the IOPr at more caudal levels and merges rostrally with the IOM, thus showing no relation to the IOBe (Brodal and Kawamura, 1980). Some authors, therefore, refer to a conspicuous enlargement on the medial end of the ventral lamella of the IOPr as the IODMCC (Gwyn et al., 1977; Furber and Watson, 1983; Sotelo et al., 1986; Apps, 1990). However, this olivary region has been shown to be linked with the dorsolateral hump (IntDL) of the rat cerebellar nuclei (Ruigrok and Voogd, 1990; Buisseret-Delmas and Angaut, 1993), and, most probably, is not homologous to the IODMCC of the cat. The term dorsomedial group (IODM), therefore, is applied to this specific IOPr region. In a study of the inferior olive using immunostaining for glutamic acid decarboxylase (Nelson and Mugnaini, 1988), the IODMCC and IODM were found to be linked at caudal levels. Since the IODMCC of both sides are intimately linked in the midline (Fig. 3, levels 15 and 16), the related regions of IODM and IODMCC have been referred to as the transition area (T-area, Fig. 4) Nelson and Mugnaini, 1988; De Zeeuw et al., 1996). Directly dorsal to the caudal aspect of IOBe (level 7 of Fig. 3) a small cell cluster appears which is generally referred to as the dorsal cap of Kooy (IOK). The IOK extends rostrally for about 800 μm. Approximately 600 μm rostral to its caudal tip a small cell grouping detaches itself ventrolaterally from the IOK and becomes adjacent to the lateral border of IOBe. This is the ventrolateral outgrowth (IOVL), which eventually merges rather suddenly with the ventral lamella of the IOPr (level 14, Fig. 3; see also Nelson and Mugnaini, 1988). The caudal pole of the IOPr is first recognized about 1300 μm rostral to caudal pole of the IOM. Its dorsolateral part is associated with the ventromedial part of the caudal IOD. As this point of fusion of the two subdivisions progressively moves dorsomedially in succesively more rostral areas, it becomes obvious that (1) the IOPr divides into a dorsal and a ventral lamella which are connected laterally and (2) the IOD also appears as a folded structure (Fig. 3, levels 12 and 13; Fig. 4). At more rostral

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levels the dorsal IOPr lamella fuses with its ventral counterpart and finally also with the IOD. The IOD is first recognized approximately 200 μm caudal to the caudal pole of the IOPr as a small cell cluster lateral to the IOBe at level 10 (Fig. 3). More rostrally, this so-called dorsal fold of the IOD (IODdf) (Azizi and Woodward, 1987) expands ventrolaterally, until it reaches the ventral surface, where it curves medialward. This medial extension becomes adjacent to the dorsal lamella of the IOPr and enlarges dorsomedially to become the ventral fold of the IOD or the IOD proper. For most of its rostrocaudal length the medial IOD is continuous with the dorsal fold of the IOPr, making the demarcation of both divisions difficult. Careful examination of sections incubated for acetylcholine esterase or cytochrome oxidase and of the afferent and efferent connections indicates that the medial tip of the dorsal lamella of the IOPr should be included with the IOD. As such the IOD becomes immediately adjacent to the IODM group of the ventral lamella of the IOPr (Fig. 3, level 15; Fig. 4) (Ruigrok and Voogd, 1990). Rat IO neurons are small (soma diameter range, 12–25 μm) (Gwyn et al., 1977) and possess four to seven primary dendrites. In many mammalian species olivary

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cells can be divided into a type with dendrites that recurve toward the soma, giving the dendritic tree a rather globular appearance. A second, simpler, type possesses dendrites that radiate away from the soma. In the rat, these two cell types are found throughout the olivary complex. Golgi impregnations of young animals sometimes show axons with recurrent collaterals (King, 1980) for review); however, these collaterals are not encountered in intracellularly HRP-injected olivary neurons in adult cats (Ruigrok et al., 1990). The dendritic tree of olivary cells is occupied by simple as well as rather complex dendritic appendages. In cat it has been demonstrated that appendages of a number of different cells are entwined and surrounded by GABAergic as well as nonGABAergic terminals with an excitatory appearance (De Zeeuw et al., 1989b, 1990a, 1990b). The whole, packed in a glial sheath, constitutes a glomerulus and is characteristic of the olivary neuropil (King, 1976). Another characteristic feature of olivary neurons is their electrotonic coupling by gap juntions. These gap junctions are frequently found between the spiny appendages (Llinás et al., 1974; Sotelo et al., 1974; De Zeeuw et al., 1989a, 1990a, 1990b). Within the rostralmost part of the IOM and within the IODdf, small

FIGURE 4 Reconstructed dorsal view of the inferior olivary complex of the rat. Different (sub-)nuclei are indicated with different shadings. Note the transition area (T-area) where the neuropil of the various regions seem to intertwine and are also in contact with the contralateral side (modified after De Zeeuw et al., 1996).

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neuronal somata that stain positive for glutamic acid decarboxylase have been noted, suggesting that in these areas GABAergic local interneurons may be present (Fredette et al., 1992).

Afferents to the Inferior Olivary Nucleus The inferior olive receives afferent connections from large parts of the central nervous system. However, the origin of the afferent projections to any particular subdivision appears to be rather restricted and coincides with the modular organization of the efferent climbing fiber projection. It follows that the various subdivisions of the inferior olive are incorporated in an impressive range of cerebellar functions. It has been convincingly demonstrated that the IOD is the recipient of pathways from somatosensory nuclei that relay cutaneous tactile and nociceptive information (Gellman et al., 1983; Molinari et al., 1990; Garwicz et al., 1992, 1996, 1997). These pathways arise from the spinal cord, the spinal trigeminal nucleus, and the dorsal column nuclei (Boesten and Voogd, 1975; Swenson and Castro, 1983a, 1983b; Apps, 1998). Lumbosacral fibers terminate laterally within the dorsal and ventral folds of the IOD, whereas cervical levels project to more medial areas. Projections from the interpolar part of the trigeminal nucleus are found in the rostromedial part of the IOD (Huerta et al., 1983, 1985; Van Ham and Yeo, 1992, 1996). Terminal arborizations from the gracile and cuneate nuclei tend to overlap with projections from the lumbosacral and cervical spinal cord, respectively (Boesten and Voogd, 1975; Berkley and Hand, 1978). Using electrophysiological techniques (mapping of responsive fields; Gellman et al., 1983), and with the aid of detailed anterograde tracer techniques (Molinari et al., 1991; Matsushita et al., 1992), it has been demonstrated in the cat that a detailed somatotopical map can be found within the rostral half of the IOD. An additional, but more crude, somatotopical representation appears to be present within the caudal half of the cat IOD, which most probably is homologous with the rat dorsal fold. The IOD, furthermore, receives input from the lateral mesencephalic nucleus, the pretectal area, and the reticular formation (Swenson and Castro, 1983a, 1983b; Bull et al., 1990). It is of interest that these areas are also the recipients of cerebellar input as has been demonstrated in the cat (Bull and Berkley, 1991). In the rat it has been shown that the anterior interposed nucleus (IntA) of the cerebellum provides a GABAergic projection to the caudolateral, hindlimb-related, and rostromedial, facerelated, parts of the IOD. A similar GABAergic projection from the lateral vestibular nucleus has been demonstrated to terminate in the IODdf (Ruigrok and Voogd,

1990; Fredette and Mugnaini, 1991). GABAergic projections to the intermediate, presumably forelimb-related, part of the IOD have been suggested to arise from the cuneate area (Nelson and Mugnaini, 1989). However, in the cat it has been reported that the forelimb recipient zones of the IOD also receive a GABAergic cerebellar projection (Molinari, 1992). The caudal part of the IOM (horizontal lamella, groups A and B) also processes somatosensory information since it also receives input from the spinal cord, dorsal column nuclei, and spinal part of the trigeminal nucleus. However, contrary to the situation in the IOD, a somatotopical representation is less obvious (Gellman et al., 1983; Huerta et al., 1985). Moreover, input from the vestibular nuclei may overlap with the somatosensory information (Swenson and Castro, 1983a, 1983b). Teleceptive information arising from the deep layers of the superior colliculus predominantly terminates within the C group (Fig. 5) (Hess, 1982; Kyuhou and Matsuzaki, 1991; Akaike, 1992). The rostral periaquaductal gray and the interstitial nucleus of the medial longitudinal fascicle (Fig. 5) also provide extensive projections to the caudal half of the IOM including IOBe. A prominent input to the IOBe is also derived from the vestibular nuclei (Brown et al., 1977; Swenson and Castro, 1983a, 1983b; Kaufman et al., 1991, 1996). Part of this projection, at least, has been shown to be GABAergic (Nelson and Mugnaini, 1989). A sparse, but definite, and presumably GABAergic projection to the lateral and medial parts of the caudal IOM, as well as to the IOBe and IODMCC, has been shown to arise from the medial cerebellar nucleus (Med) (Ruigrok and Voogd, 1990). However, these connections cannot account for the massive GABAergic projection also found in the caudal IOM. An additional GABAergic projection to the IOMC has been shown to arise from the ipsilateral parasolitary region (Nelson and Mugnaini, 1989; De Zeeuw et al., 1993). The origin of a major GABAergic pathway to the IOMA and IOMB groups has yet to be identified. The IOK and IOVL both play an important role in the control of compensatory eye movements as has been amply demonstrated in the rabbit (Leonard et al., 1988). It receives its main input from the nucleus of the optic tract (OT, to the IOK) and the accessory optic nuclei such as the medial terminal nucleus (MT, to the IOVL). Additional IOVL projections stem from the periaquaductal central gray and the visual tegmental relay zone (VTRZ) (Giolli et al., 1984, 1985). A dopaminergic projection to the IOVL has been shown to be derived from neurons in the medial mesodiencephalic junction (Toonen et al., 1998). A projection from the medial cerebellar nucleus to the IOVL, described by Angaut and Cicirata (1982) and Swenson and Castro (1983b), could not be verified by Ruigrok and Voogd (1990) who,

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FIGURE 5 Sources of inferior olivary afferents in the midbrain. (A–D) Microphotographs of consecutively more caudal levels of the midbrain. Some nuclei and tracts have been plotted as have been cells that were retrogradely labeled from the injection in the inferior olivary complex shown in panel E. Examples of the retrogradely labeled neurons are shown in the inset in panel E (arrows). Note that most labeled cells were found surrounding the fasciculus retroflexus and that virtually none were located within the confines of the red nucleus. Also see Fig. 6. Plots were constructed using Neurolucida (Microbrightfield, Inc.) software (Ruigrok, unpublished results).

instead, mention a IOVL projection from a small area in the parvicellular part of the lateral cerebellar nucleus (see also Nelson and Mugnaini, 1989; Ruigrok et al., 1992). The IOK and IOVL both receive a GABAergic as well as a nonGABAergic projection from the prepositus hypoglossal nucleus (De Zeeuw et al., 1993). The rostral lamella of the IOM and the IOPr receive afferents from areas located at the mesodiencephalic junction. A continuum of neurons surrounding the fasciculus retroflexus, defined as area parafascicularis prerubralis by Carlton et al. (1982), encompassing the rostral part of the nucleus of Darkschewitsch, the medial accessory oculomotor nucleus (MA3), the rostral interstitial nucleus of the mlf, and the prerubral field can be found to project to the inferior olive (Figs. 5, and 6) (Brown et al., 1977; Swenson and Castro, 1983a; Swenson and

Castro, 1983b; Bentivoglio and Molinari, 1984; De Zeeuw et al., 1990a). In the cat, it has been demonstrated that presumably homologous areas project in a topographical fashion to the rostral half of the IOM, to the ventral lamella of the IOPr, and to its dorsal lamella, respectively (Saint-Cyr and Courville, 1982; Onodera, 1984; Holstege and Tan, 1988). Moreover, specific projections from the sensorimotor and parietal association cortices have been attributed to these regions in various animal species. As to the much debated olivary projections arising from the red nucleus, the reader is referred to the section dealing with this nucleus (also see Figs. 5 and 6). Prominent GABAergic projections from the posterior interposed nucleus (IntP) of the cerebellar nuclei to the rostral lamella of the IOM and from the Lat to the IOPr have been amply demonstrated (Angaut

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et al., 1987; Angaut and Sotelo, 1989; Ruigrok and Voogd, 1990; Fredette and Mugnaini, 1991). The IODM group of the ventral lamella of the IOPr receives a specific innervation from the dorsolateral hump (IntDL) of the rat cerebellar nuclei, in addition to projections stemming from the spinal trigeminal nucleus and the adjacent reticular formation (Huerta et al., 1983; Swenson and Castro, 1983a; Ruigrok and Voogd, 1990). A schematic summary of the cerebellar nucleoolivary projection is given in Fig. 7. Here, the IO and the

cerebellar nuclei are represented as more or less continuous sheets of cells, folded in their subnuclei. In this scheme there is a rather simple topographical relation between both cell masses, especially when the IntP is regarded as a caudal projection originating from the Med and Lat, and its target nucleus, the rostral lamella of the IOM, as a rostral expansion of the vertical lamella of the IOM. Likewise, one can consider the IntDL as an enlargement intercalated between the dorsal Lat and the lateral IntA that projects to the IODM group of the inferior olive which can be interpreted as a medial bulge between the medial IOD and the dorsal lamella of the IOPr, which, thus, becomes located adjacent the ventral lamella of the IOPr (Ruigrok and Voogd, 1990). Indoleaminergic as well as catecholaminergic projections to specific subdivisions of the rat inferior olivary complex have been described (Wiklund et al., 1977; Bishop and Ho, 1984, 1986; Toonen et al., 1998). The neuropeptides enkephalin, substance P, cholecystokinin, and corticotropin-releasing factor have also been encountered in olivary afferent profiles (King et al., 1989). In summary, the specific subdivisions of the IO all receive a major GABAergic as well as a nonGABAergic projection. The GABAergic projection is derived from the cerebellar nuclei, but, for some subdivisions, may also originate from the vestibular nuclei, from the prepositus hypoglossal nucleus, or from parasolitary and cuneate regions. In certain regions GABAergic boutons may arise from locally found interneurons. The nonGABAergic afferent projections to the caudal IOM and IOD may be subdivided into those that relay, more or less directly, sensory information and those that relay more highly integrated information including cerebellar output. The rostral IOM and IOPr, on the other hand, appear to process integrated information from both the cerebellum and the cerebral cortex. Modulatory influences from various aminergic and peptidergic systems may influence this information processing (van der Steen and Tan, 1997).

Efferents of the Inferior Olive

FIGURE 6 Three-dimensional reconstruction of mesodiencephalic junction of the same experiment shown in Fig. 5. However, the junction is now shown as a three-dimensional reconstruction (Neurolucida, Microbrightfield, Inc.) based on serial plots of sixteen 40-μm sections (1 of 4 sections was plotted). (A) Caudal view, (B) rostral view. Note that within the confines of the red nucleus (R) as well as of the nucleus of Darkschewitsch (Dk) only a few labeled cells were found. Most labeled neurons were located surrounding the fasciculus retroflexus. Also, note the vast quantities of labeled cells in the deep mesencephalic nucleus, pretectum, central gray, and zona incerta (cf. Fig. 5; Ruigrok, unpublished results).

The inferior olive is the sole source of climbing fibers to the cerebellum (Desclin, 1974). They terminate extensively on the dendritic tree of Purkinje cells (Szentágothai and Rajkovits, 1959; Eccles et al., 1966b) and provide a collateral projection to the cerebellar nuclei (Kitai et al., 1977; van der Want et al., 1989; Ruigrok, 1997; Ruigrok and Voogd, 2000). The climbing fiber projections from the various subdivisions of the IO are topographically organized into a number of sagittally oriented strips that are characterized by the projection of the Purkinje cells in those strips to the cerebellar and vestibular nuclei. The correlation between

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FIGURE 7 Schematized cerebellar nucleoolivary relationships. The cerebellar nuclei and olivary nuclear complex are visualized as unfolded continuous sheets of cells. Projections from the olivary sheet to the cerebellar nuclei are excitatory (Audinat et al., 1992) and are indicated with arrowheads. The reciprocal projection from the cerebellar sheet to the inferior olive is GABAergic (Fredette and Mugnaini, 1991) and is indicated with filled circles. Folds in both sheets are indicated by open arrows (modified after Ruigrok and Cella, 1995, also see Ruigrok and Voogd, 2000). See text for further explanation.

the olivocerebellar and the corticonuclear projection bear similar characteristics in the rat, cat, and monkey. The collateral projection of the climbing fibers to the cerebellar nuclei as well as the GABAergic nucleoolivary projection appears to be aligned with the olivocorticonuclear projection. Indeed, the formation of a number of parallel circuits linking the inferior olive and the cerebellum appears to constitute the essence of cerebellar functioning. The reader is referred to Chapter 9 by Voogd for a comprehensive review on the olivocerebellar projection in the rat (see also Buisseret-Delmas and Angaut, 1993). Initial studies have suggested that the climbing fiber system may use aspartate as a neurotransmitter (Wiklund et al., 1982, 1984; Kimura et al., 1985), but more recent reports claim that glutamate (Vollenweider et al., 1990; Zhang and Ottersen, 1993; Grandes et al., 1994; Dzubay and Jahr, 1999; Laake et al., 1999; Wadiche and Jahr, 2001) is the most likely neurotransmitter candidate of the climbing fibers. N-Acetylaspartylglutamate, in addition, has also been suggested to be involved as a neurotransmitter/neuron modulator in at least a subset of climbing fibers (Sekiguchi et al., 1989; Renno

et al., 1997). Corticotropin-releasing factor, enkephalin, and cholecystokinin have also been reported as neuropeptides present in particular subsets of climbing fibers (King et al., 1986; Young et al., 1986; Palkovits et al., 1987; van den Dungen et al., 1988; Bishop, 1990; King and Bishop, 1990; King et al., 1992).

Functional Considerations The role of the inferior olive in the operation of the cerebellum remains rather enigmatic. Indeed, the function(s) of the cerebellum itself are not entirely understood. Although its role in motor functions has been amply documented (e.g., see Holmes, 1939; Ito, 1984; Arshavsky et al., 1986), later studies have suggested that the cerebellum, and the IO, also may be involved in visceral functions and affective behavior (Bradley et al., 1987; Nisimaru et al., 1991; Waldrop and Iwamoto, 1991), and even in certain mental activities (Ito, 1990; Schmahmann, 1991; Fiez et al., 1992, 1996; Leiner et al., 1993; Schmahmann and Sherman, 1998). Furthermore, it has been shown that the IO plays a role in classical conditioning (Voneida et al., 1990; Sears and Steinmetz,

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1991; Yeo and Hesslow, 1998). It is obvious that the function of the IO will be reflected in its sources of afferent information, in its integrating capabilities, and in the effect of its efferent impulses. IO afferents are known to transmit a wide variety of sensory modalities that may be integrated at various levels with cerebral input, cerebellar input, or both (e.g., Kistler et al., 2000; Schwarz and Welsh, 2001). These characteristics of the IO afferent systems have prompted the suggestion that the IO may function as a detector of events and/or errors of a variety of physiological processes (Oscarsson, 1980; Ito, 1984; De Zeeuw et al., 1998b; Ito, 1998). Indeed, climbing fiber responses mediated via a number of direct or indirect spinoolivocerebellar pathways have been implicated to signal specific somatosensory conditions (Garwicz et al., 1992; Ekerot et al., 1997). Recently, it has become clear that the excitability of these pathways may be subject to gating processes, purportedly in order to minimize or eliminate propagation of expected signals (Apps et al., 1997; Apps and Lee, 1999; Apps, 2000). When considering the integrative capabilities of olivary neurons, the rather peculiar conductances of olivary neurons should be taken into account together with the fact that olivary cells are electrotonically coupled to each other (Llinás and Yarom, 1981a, 1981b; 1986; Benardo and Foster, 1986; Yarom, 1991). These features, in conjunction with the intricate network of interwoven dendrites with their many long and frequently complex spiny appendages (Sotelo et al., 1974; Gwyn et al., 1977; Ruigrok et al., 1990), would appear to warrant rather complex input–output relationships for this nucleus (De Zeeuw et al., 1998b). The observation that these spiny appendages of olivary neurons form the core of the olivary glomeruli (King, 1976) and are surrounded by excitatory as well as GABAergic synapses can be interpretated as representing a structural correlate of a device that may be very well suited to detect, and respond to, changes and/or differences in the temporal resolution of the various inputs (Segev and Rall, 1988, 1998; De Zeeuw et al., 1990a; 1998b; Ruigrok et al., 1990; Segev and Rall, 1998; Lang, 2001). Climbing fiber impulses result in a powerful excitation of the innervated Purkinje cells, which respond with a characteristic “complex spike.” The complex spike consists of a series of characteristic dendritic calcium spikes, which may induce somatic firing (Llinás and Sugimori, 1980). As the result of a complex spike, up to six action potentials with a frequency of about 500 Hz may be conducted along the Purkinje cell axon (Eccles et al., 1966a; Ito and Simpson, 1971). Nevertheless, at a first inspection, the rather low firing frequency of olivary neurons (1–2/s) would appear to preclude an important contribution of the IO to the overall firing

rate of the Purkinje cells. However, cooling and lesioning of the IO has demonstrated quite the reverse. In effect, behavioral and physiological studies suggest that lesions of the IO, especially in its initial stages, resemble lesions of the whole cerebellum (Llinás et al., 1975; Colin et al., 1980; Batini and Billard, 1985; Demer et al., 1985). This effect is, most likely, due to the interaction of the IO climbing fiber and mossy fiber–parallel fiber input at the Purkinje cell level. The climbing fiberinduced complex spike (CS) gives rise to a temporal depression of the mossy fiber–parallel fiber activated simple spikes (SSs) of the Purkinje cells. Conversely, a reduction in frequency or a complete abolition of CSs enhances the frequency of SSs. Due to the GABAergic action of the Purkinje cell terminals on their target (Ito and Yoshida, 1966; Ito, 1984), this will result in a massive inhibition of the cerebellar nuclei. Apart from the tonic effects of CSs on the frequency of SS firing (Strata, 1984) more subtle and long-term effects between CS and SS responsiveness have also been demonstrated. Particularly, it has been noted that repeated stimulation of climbing fibers depresses the synaptic efficacy of near-simultaneously firing parallel fibers. This long-term depression (LTD) has been hypothesized to enable the climbing fibers, acting as an error signal, to adapt the output of the Purkinje cells semipermanently to new or specific requirements in motor function (Marr, 1969; Albus, 1971; Ito, 1984, 1994, 2001). Indeed, it was recently shown that genetically modified mice (in which protein kinase C had been blocked in Purkinje cells only) not only failed to show LTD but also failed to show adaptation of the vestibuloocular reflex (De Zeeuw et al., 1998a) Another school suggests that the IO acts as a real afferent system. As such it would not serve as a modulator of the SS responsiveness at the Purkinje cell level, but it is suggested that the climbing fiber-induced CS may bring about specific responses in Purkinje cell target areas that are necessary to adjust to a particular situation (Llinás and Mühlethaler, 1988; Llinás, 1991; Welsh et al., 1995). The tremorogenic action of the indolamine harmaline, which results in rhythmic firing of the inferior olive and in related bursting of neurons in the cerebellar nuclei and brain stem, has been put forward as evidence of direct impact of the inferior olive on motor programming (De Montigny and Lamarre, 1973; Llinás and Volkind, 1973; Llinás and Sasaki, 1989). Timing and synchrony of the olivary activations are key elements in this theory. Indeed, it has been demonstrated that the electrotonic coupling by gap junctions may be modulated by GABAergic synapses (De Zeeuw et al., 1989b, 1990a; Llinás and Sasaki, 1989; Lang et al., 1996). In theory, this would enable the IO to respond with different aggregates of

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coupled cells to a specific input in different situations. The relevance of the timing of events may be related to the intrinsic oscillation of the membrane potential found in the coupled olivary cells (Benardo and Foster, 1986; Llinás and Yarom, 1986; Yarom, 1991). Rhythmic firing of groups of olivary cells related to ongoing oscillatory movements such as locomotion have recently been noted in freely moving animals (Smith, 1998) and also resulted in increased Fos labeling in related regions of the inferior olive and the cerebellar nuclei (Ruigrok et al., 1996). Some evidence obtained in Lurcher mice that lack Purkinje cells would indeed suggest that the collaterals of climbing fibers may participate in inducing Fos in the cerebellar nuclei (Oldenbeuving et al., 1999). A somewhat undervalued aspect of cerebellar functioning in general and of olivary functioning in particular may be found in the likelihood that activity patterns may be reverberating and thus be maintained within the various circuits between brain stem and cerebellum. The interaction between intrinsic subthreshold oscillations of the membrane potential of inferior olivary neurons (Lampl and Yarom, 1993) and the activity pattern in olivocerebelloolivary and olivocerebello– midbrain–olivary circuits may be specifically revelant for cerebellar functioning (Ruigrok and Voogd, 1995; Kistler and van Hemmen, 1999; Kistler et al., 2000).

RED NUCLEUS The red nucleus, which gets its name thanks to the pinkish color of the large rounded structure found in fresh human tissue, is a conspicuous nucleus located on either side of the midbrain tegmentum of limb-using vertebrates (ten Donkelaar, 1988). It subserves a premotor function and is closely related to the cerebellum since it not only receives a prominent input from the cerebellar nuclei but also acts as a source of information for various precerebellar nuclei. In most mammals studied, a magnocellular part is distinguished from a more rostrally placed parvicellular part. In primates this subdivision is easy to recognize (Paxinos et al., 2000), although the magnocellular part is only poorly developed in anthropoids and, especially, in man. It follows that the human red nucleus essentially consists of the parvicellular division of the nucleus (Nathan and Smith, 1982; Paxinos and Huang, 1995). In most mammals, however, the parvicellular part is not so well defined. Caudally, it appears to overlap with the magnocellular part, whereas its rostral boundaries are difficult to establish. The two subdivisions are generally believed to serve a different function, but their connections, as reported in the literature are subject to many controversies. The next paragraph attempts to

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elucidate some of these issues in the cytoarchitecture and connectivity pattern of especially the rat red nucleus.

Cytoarchitecture The red nucleus of the rat is roughly ovoid in shape and located bilaterally in the midbrain tegmentum. Its caudal tip is found approximately 2.5 mm rostral to the interaural line (Paxinos and Watson, 1986), where it emerges as a prominent cell group within the crossed superior cerebellar peduncle. Its rostral boundary is more difficult to establish but, by convention (Reid et al., 1975b), is defined just caudal to the level of the fasciculus retroflexus, at approximately 3.7 mm rostral to the interaural plane. Rostral to this level the prerubral field is more or less continuous with the red nucleus. The superior cerebellar peduncle envelops and traverses the red nucleus at all rostrocaudal levels. Outgoing fibers of the oculomotor nucleus pass medially from and through its caudal aspect. The medial lemniscus (ml) is found adjacent to the ventrolateral border of the rostral half of the nucleus. The dorsal tegmental decussation, the medial longitudinal fasciculus, and the medial tegmental tract border the red nucleus on its dorsomedial side. Clearly, different cell types are found within the confines of the red nucleus of the rat. In a first, detailed, study by Reid et al. (1975a, 1975b), four types were recognized, primarily based on soma size. Giant (soma diameter, >40 μm) and large (26–40 μm) neurons, which possess similar structural characteristics, predominated in the caudal third of the nucleus. Medium neurons (20–25 μm) displayed a lower cytoplasmic to nuclear ratio and contained fewer and poorly organized Nissl bodies. Small neurons (<20 μm) were usually achromatic and were frequently seen in association with neuroglial cells. Medium and small cells made up most of the rostral two thirds (i.e., the parvicellular part). Strominger et al. (1987) only recognized three neuronal populations: neurons with coarse Nissl bodies, neurons with a fine granular Nissl substance, and achromatic neurons. The soma sizes of neurons with coarse and fine Nissl substance overlap considerably, whereas the achromatic neurons are clearly smaller than the other two cell types. “Coarse” neurons were found exclusively in the caudal tip, while the rostral tip harbored predominantly “fine” neurons. In the intermediate red nucleus both cell groups were found. The small neurons were found throughout the nucleus. Finally, studies by Tucker et al. (1989) differentiated between neurons with a relatively small, oval, and eccentrically placed nucleus and displaying a large cytoplasmic–nuclear ratio (magnocellular neurons) and neurons with a relatively large rounded and a centrally placed nucleus displaying a

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small cytoplasmic to nuclear ratio (parvicellular neurons). Approximately 3000 magnocellular neurons were counted which could be divided into a largesized (>25 μm) and a small-sized (<25 μm) population and which dominated the caudal and ventral part of the nucleus. Parvicellular neurons (±20,000) were classified as large-sized (30–45 μm), medium-sized (15–30 μm), or small-sized (>15 μm) cells and may all be found throughout the nucleus. Although there is no consensus between different authors, it is agreed that the caudal pole harbors the largest neurons. Between 250 and 400 μm from the caudal pole, this large-celled part can be subdivided into a rather compact ventrolateral part and a more loosely arranged dorsomedial part (Reid et al., 1975b; see also Figs. 8A and 8B). The ventrolateral part is continuous with the so-called lateral horn of the rat red nucleus (Reid et al., 1975b) and is somewhat separated from the dorsomedial part, which appears to divide into a dorsal and a dorsomedial grouping of neurons. Rostral to this level no subdivisions are apparent. At least a part of the small neuronal population may represent local GABAergic interneurons, which have been demonstrated in turtle, cat, and monkey (VuillonCacciuttolo et al., 1984; Keifer et al., 1992; Ralston, 1994). Large neurons in the caudal two-thirds of the rat red nucleus frequently stain positive for calbinding D28k, whereas medium to large neurons in the rostral two-thirds frequently contain parvalbumin. Both populations, however, were strongly intermingled and double-labeled cells were only seldomly encountered (Hontanilla et al., 1995). The nucleus minimus deserves some comment. This aggregation of small cells lateral to the parvicellular part of the red nucleus was first described by Von Monakov (1909) in the rabbit and later identified in the cat by others (Brodal and Gogstad, 1954; Taber, 1961). In the rat, data are controversial. Reid et al. (1975b) did not find a nucleus minimus. However, they described the lateral horn as a subdivision of the red nucleus and as consisting of predominantly small- and mediumsized neurons. Faull and Carman (1978), who based their opinion on the pattern of terminal degeneration after superior cerebellar peduncle lesions, concluded that a nucleus minimus complying to Von Monakov’s description may be found intercalated between fascicles of the medial lemniscus. However, the position of this nucleus is rather similar to that of the lateral horn of Reid. Finally, Paxinos and Watson’s atlas of the rat brain (1986), based on Faull and Carman’s paper, delineates a nucleus minimus lateral to the dorsal part of the red nucleus, where it appears to be dorsal or dorsolateral from the lateral horn of Reid et al. Based on our own material of cerebellar nuclear projections to this area

(see below), we propose that the term nucleus minimus should be abandoned. Rather, we would advocate the term pararubral area, which forms an integral part of the parvicellular part of the red nucleus rostrally but becomes discernible as a more or less separated cluster of neurons located dorsolateral to its magnocellular part (Figs. 8H and 9: also see Fig. 6 of Ruigrok and Cella, 1995). As such, the pararubral area is located within the medial part of the nucleus reticularis subcuneiformis of Newman (1985). As mentioned earlier, the rostral boundary of the parvicellular red nucleus is difficult to establish. Based on the efferent connections of this area, it has been suggested by Kennedy (1987; Tucker et al., 1989) that the area directly surrounding the fasciculus retroflexus should be considered a part of the red nucleus. In rat, the area dorsomedial to this fiber bundle is recognized as (the rostral part of) the nucleus of Darkschewitsch and its ventrocaudal continuation as the nucleus accessorius medialis of Bechterew (?edial accessory oculomotor nucleus, MA3, of Paxinos and Watson, 1986). The rostral interstitial nucleus of the medial longitudinal fascicle and the prerubral field are situated ventral and ventrolateral to the fasciculus retroflexus, respectively (Paxinos and Watson, 1986). Carlton et al. (1982) designated the whole area surrounding the retroflex bundle as the nucleus parafascicularis prerubralis on the basis of its efferent connections. In this respect, it is noteworthy that in primates the parvicellular red nucleus is indented rostromedially by the retroflex bundle and extends rostrally to it. In human, the fasciculus retroflexus even traverses the parvicellular red nucleus completely (Paxinos and Huang, 1995), thus separating a dorsomedial part, which may be homologous to (part of) the nucleus parafascicularis prerubralis of the rat. It will be evident that extensive and detailed studies on the afferent as well as the efferent connections of these areas in the various animals are needed in order to establish whether areas with similar names indeed subserve similar functions.

Afferents to the Red Nucleus The red nucleus receives input from the cerebellar nuclei as well as from the cerebral cortex. Projections arising from the posterior thalamic nucleus (Roger and Cadusseau, 1987), zona incerta (Ricardo, 1981), hypothalamic areas, central pontine gray, nuclei raphe dorsalis and magnus, gigantoreticular nucleus, parvicellular reticular nucleus, parabrachial nuclei, and locus coeruleus (Bernays et al., 1988) have also been described. In cat, projections from the dorsal column nuclei and the spinal cord have been demonstrated to reach and terminate within the red nucleus (Boivie, 1988).

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FIGURE 8 Relation between the origin of the rubrospinal tract and the cerebellar nucleorubral connections. (A) Retrograde labeling of rubral neurons after WGA–HRP injection into the contralateral cervical spinal cord. (B) Retrograde labeling after WGA–HRP injection into the contralateral lumbar spinal cord. Note that the labeled cells are positioned ventral to those labeled in panel A. (C,E,G) Injection sites of the anterograde tracer PHA-L into the lateral part of the IntA, the medial part of the IntA, and the ventral, parvicellular part of the Lat, respectively. (D,F,H) Corresponding terminal labeling in the contralateral red nucleus. Note the the resultant labeling after a medial IntA injection (D) overlaps with the location of rubrospinal neurons that project to the lumbar cord, whereas the lateral IntA projects to the dorsal red nucleus (F) where the rubrocervical neurons are found. The parvicellular Lat gives rise to a conspicuous terminal labeling in the pararubral area (small arrows in panel H, also noted labeled, nonterminal, fibers within confines of the red nucleus). Open arrow in panels A, B, D, F, and H, idicates the lateral horn. Bar equals 200 μm (Ruigrok, unpublished results).

Various studies employing degeneration (Caughell and Flumerfelt, 1977) and retrograde and/or anterograde techniques (Daniel et al., 1987; Angaut and Cicirata, 1988) have recognized that the projections from the contralateral cerebellar nuclei to the red nucleus in the rat are organized in a somatotopical fashion. Our

material of small injections into the cerebellar nuclei with the anterograde tracer PHA-L (Ruigrok and Voogd, 1990) confirms and extends these observations (Figs. 8C–8H). Massive projections arise from the anterior interposed nucleus (IntA); its lateral part projects to the dorsomedial part of the magnocellular

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red nucleus, whereas its medial part provides heavy terminal arborizations in its ventrolateral part and lateral horn. The projection from the IntA stems from coarse axons and terminates on somata and proximal dendrites of the magnocellular neurons (Flumerfelt, 1978, 1980; Dekker, 1981). The Lat provides a major input to especially the parvicellular part of the nucleus. Here, a topographical organization also can be recognized. The rostral part of the Lat projects rather ventromedially, whereas more dorsolateral areas of the parvicellular part are supplied by the caudal part of the Lat (Angaut and Cicirata, 1988). A conspicuous projection to the cell aggregate located lateral to the magnocellular red nucleus (pararubral area) was found to arise specifically from the parvicellular subdivision of the Lat (Ruigrok et al., 1993; Teune et al., 2000: see also Figs. 8G and 8H). Red nuclear projections from the posterior interposed nucleus are sparse and are found along its dorsomedial aspect. This cerebellar nucleus distributes terminals mainly to the medial accessory oculomotor nucleus and to the medial part of the nucleus parafascicularis prerubralis (Teune et al., 2000, however, cf. Daniel et al., 1987). A small but definite projection is found from the medial cerebellar nucleus to the medial part of the base of the lateral horn. It should be recognized that the projections from the medial, posterior interposed as well as from the lateral cerebellar nuclei consist of rather fine caliber fibers that terminate on intermediate and distal dendrites and are not as massive when compared to the projections from the anterior interposed nucleus (Caughell and Flumerfelt, 1977; see also Figs. 8C and 8F). It is suggested that all rubral projections from the cerebellum are collaterals from ongoing fibers that eventually terminate in the thalamus (Shinoda et al., 1988) and are excitatory (Toyama et al., 1970; Oka, 1988). Most likely they make use of an excitatory amino acid as neurotransmitter (Bernays et al., 1988; Giuffrida et al., 1993; Schwarz and Schmitz, 1997). In the cat, using physiological techniques, it was found that at least some of the nucleoolivary projection neurons (thought to be all GABAergic: De Zeeuw et al., 1989b, Fredette and Mugnaini, 1991) collateralize to the red nucleus and/or thalamus (McCrea et al., 1978; Andersson and Hesslow, 1987). However, so far this notion could not be corroborated by anatomical studies (Teune et al., 1994; Schwarz and Schmitz, 1997). The projections from the rat sensorimotor cortex are not as well documented as those arising from the cerebellar nuclei. Gwyn et al. (1974) using degeneration techniques, reported that the sensorimotor cortex provides projections to the parvicellular part of the red nucleus only. This was confirmed by Brown (1974a) who also noticed that the prerubral field and an area dorsolateral

to the red nucleus (i.e., the pararubral area) receive a particularly heavy cortical input (cf. projections from the sensorimotor cortex to the subcuneiform reticular nucleus, Newman et al., 1989). Physiological studies by Giuffrida et al. (1988a, 1988b), however, claim that both magno- and parvicellular parts of the red nucleus of the rat are controlled by cerebral as well as cerebellar influences. This agrees well with a study using retrograde transport of small injections of lectin coupled to colloidal gold in different parts of the red nucleus in the guinea pig (Giuffrida et al., 1991). Retrogradely labeled neurons in layer V of the agranular frontal cortex were found after injections into the parvi- and into the magnocellular part of the red nucleus. In a study employing the use of selective retrogradely transported tracers, Bernays et al. (1988) suggested that the cerebral afferents to the red nucleus make use of an excitatory amino acid as transmitter (also see Giuffrida et al., 1993). Cadusseau and Roger (1987, 1988) have described a conspicuous and essentially reciprocal projection from the posterior thalamic nucleus to the parvicellular part of the red nucleus in the rat. The posterior thalamic nucleus is positioned within the sub-pretectal area at the mesodiencephalic junction and appears to be involved in the assimilation of somatosensory and/or nociceptive information. In this regard it is interesting that the zona incerta, considered to be a highly integrative somatosensory center, also provides input to both the posterior thalamic nucleus and the parvicellular red nucleus (Ricardo, 1981; Watanabe and Kawana, 1982; Roger and Cadusseau, 1985; Roger and Cadusseau, 1987). A somatosensory input from the spinal cord to the red nucleus that bypasses the cerebellum and cerebral cortex has been reported in the cat (Wiberg and Blomqvist, 1984a, 1984b; Padel et al., 1986; Boivie, 1988; Rathelot and Padel, 1997; Steffens et al., 2000) and in the monkey (Wiberg et al., 1987; Kerr and Bishop, 1991). In the rat, evidence has been obtained that direct reciprocal connections between the red nucleus and the trigeminal complex may function in a potentially similar way (Godefroy et al., 1998). Finally, a serotonergic projection arising from the raphe magnus and the raphe dorsalis nuclei to the red nucleus as well as a noradrenergic projection has been demonstrated in rat and cat (Bosler et al., 1983; André et al., 1987; Bernays et al., 1988) and has been shown to exert a powerful modulatory action on the activity of rubral neurons (Schmied et al., 1991; Ciranna et al., 1996; Faherty et al., 1997; Licata et al., 1998, 2001).

Efferents of the Red Nucleus When examining the efferent connections of the red nucleus a distinction should be made in the efferents

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of its parvicellular part and in those of its magnocellular part. However, it has been pointed out that, contrary to the situation in primates, in the rat this distinction is not easily made. Various authors have pointed out that large neurons may be found well into the parvicellular region. Yet, in monkeys, available evidence indicates that a rather strict separation of the efferent connections of both subdivisions of the red nucleus exists. The magnocellular part gives rise to the crossed rubrospinal and rubrobulbar projection, whereas the parvicellular part gives rise to the central tegmental tract which descends to the ipsilateral inferior olivary complex. In the cat, a similar organization has been suggested, although here, as in the rat, a division of the red nucleus in a parvi- and magnocellular part is less clear. Obviously, the efferent connections of the rat red nucleus may give insight into its subdivisions. However, data on the projections arising from the red nucleus are confusing. Originally, only the caudal third of the nucleus with its giant- and large-sized neurons was thought to give rise to the rubrospinal tract (Gwyn, 1971; Flumerfelt and Gwyn, 1974; Murray and Gurule, 1979). Its dorsomedial part was found to project to the cervical enlargement, whereas its ventrolateral part connected with the lumbosacral cord (Figs. 8A and 8B). However, it has become clear that at least some parvicellular neurons also project to the spinal cord (Huisman et al., 1981, 1983; Shieh et al., 1983; Tucker et al., 1989; Kennedy, 1990). The rubrospinal tract descends dorsally within the contralateral lateral funiculus where it is separated from the substantia gelatinosa by a small spinocervical tract. It terminates at the base of the dorsal horn and intermediate regions of the ventral horn (Brown, 1974b). Rubrospinal terminals may synapse with both excitatory and inhibitory interneurons and a considerable number of fibers may send projections to the ipsilateral side of the cord (Antal et al., 1992). Rubrospinal projections are considered to be excitatory and most likely make use of either glutamate or aspartate as neurotransmitter (Benson et al., 1991) The parvicellular red nucleus is generally thought to provide a projection to the inferior olive. In the rat, using anterograde as well as retrograde techniques, such a projection has been described by Swenson and Castro (1983a, 1983b). However, Rutherford et al. (1984) and Carlton et al. (1982) specifically denied this projection and claimed that most olivary projections stem from the area surrounding the fasciculus retroflexus (i.e., their nucleus parafascicularis prerubralis: also see Figs. 5, 6, and and 9). Finally, Kennedy (1987), using a modified HRP visualization technique, claimed that many neurons not only in the parvicellular part but also in magnocellular part were (faintly) retrogradely

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labeled after inferior olivary injections. Initial studies by Kennedy (Tucker et al., 1989; Kennedy, 1990; Tucker and Kennedy, 1990), employing double retrograde labeling techniques with fluorescent tracers, suggested that the rubroolivary pathway, at least in part, may exist of collaterals of the rubrospinal pathway but these ideas could not be corroborated in more recent years. In analogy with the situation in monkey, Kennedy furthermore proposed that the nucleus parafascicularis prerubralis should be redefined as being a part of the parvicellular red nucleus. Although we can agree with Kennedy on this point, it should be appreciated that (A) the retrograde labeling as the result of inferior olive injections is considerably more abundant and intense in the area parafascicularis prerubralis as compared to the labeling of cells in the parvicellular part of the red nucleus, suggesting the origin of a dense projection to the inferior olive from the former area, and (B) the pararubral area, with its input from the ventral part of the Lat, should also be incorparated in the rubral complex (Figs. 8 and 9). No specific data on the efferent projections of this area have been published as yet. Although Newman (1985) claims that the subcuneiform reticular nucleus projects to the spinal cord, it can be seen from Fig. 8 that the large, retrogradely labeled cells located dorsolateral to the red nucleus do not seem to correspond to the area of anterogradely labeled fibers derived from the parvicellular part of the lateral cerebellar nucleus (compare Figs. 8A and 8B with Fig. 8H). In addition to the rubrospinal and rubroolivary pathways, the red nucleus also provides a crossed rubrobulbar projection, presumably via collaterals of the rubrospinal pathway, which terminate in various brain stem centers like the lateral part of the facial nuclei, the parvicellular reticular formation, the rostral part of the lateral reticular nucleus, the oral part of the spinal trigeminal nucleus, and the principal sensory trigeminal nucleus, the descending vestibular nucleus, and the dorsal column nuclei (Flumerfelt and Gwyn, 1974; Hinrichsen and Watson, 1983; Godefroy et al., 1998). As a fourth major terminal source, the red nucleus has been shown to give rise to a rubrocerebellar pathway (Huisman et al., 1983; Yarom et al., 1991). Presently no information is available on the special characteristics of the rubronuclear and/or rubrocortical projection patterns in the rat, but in the cat these projections have been reported to specifically target the IntA (Nakamura et al., 1987). Finally, rubrothalamic projections, in particular arising from the parvicellular part of the nucleus, have been described to terminate in the posterior thalamic nucleus (Roger and Cadusseau, 1987) and, in the cat, in the ventrolateral thalamic nucleus (Condé and Condé, 1980).

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Functional Considerations Obviously, many questions still remain unanswered concerning the function of the red nucleus. The, previously made, distinction between magno- and parvicellular subdivisions that serve different functions, may still be useful, but appears to be rather naive. Present knowledge suggests that the red nuclear area may comprise at least three populations of neurons. A caudally located, predominantly large-celled, group participates in the execution of motor behavior, it receives its main input from the anterior interposed nucleus of the cerebellum and gives rise to the rubrospinal tract. A more rostrally located cell group consisting of slightly smaller cells, possibly assisted by a group of small cells located lateral to the magnocellular subdivision, receives its main input from the cerebellar dentate, the sensorimotor cortex, and the posterior thalamic nucleus. In the rat, many of these parvicellular neurons appear to project to the spinal cord but may also project to the inferior olive. Finally, a rostrally located group of small neurons surrounding the fasciculus retroflexus (area

parafascicularis prerubralis) receives a massive input from the sensorimotor cortex, but also from the dentate and the posterior interposed nuclei and appears to project mainly to the inferior olive. Interestingly, this region has also been shown to project to the nucleus raphe magnus (Carlton et al., 1982) and has been implied to play a role in antinociceptive functions (Peschanski and Mantyh, 1983). In this respect it should be noted that electrical stimulation of the red nucleus area has been shown to inhibit the tail flick response to noxious heat (Prado and Roberts, 1985; Kumar et al., 1995). Some of the various (suggested) divisions and efferents of the whole rubral complex are schematized in Fig. 9. In monkey, the rubrospinal tract has been implicated to be involved in controlling limb movements and more specifically of that of the hand and fingers (Lawrence and Kuypers, 1968a, 1968b; Gibson et al., 1985a, 1985b; van Kan and McCurdy, 2001). Indeed, recently, it was noted that in rat also activity of the red nucleus could be clearly related to skilled movements of the forelimb (Whishaw et al., 1998; Jarratt and Hyland, 1999). However, a rubrospinal impact on more general limb actions, such as

FIGURE 9 Diagrammatic representation of some of the efferent connections of the rubral area. Hatched lines represent minor pathways. Rubroolivary fibers are found in the central (ctt) and in the medial (mtt) tegmental tracts and arise from the prerubral field (PR), from the accessory oculomotor nucleus (MA3), and, in particular, from the area surrounding the fasciculus retroflexus (fr). Rubrospinal fibers (rst) originate mainly from the caudal, magnocellular, part of the red nucleus (RMC) but also from more rostral parts (RPC). The projections specifically originating from the pararubral area (paraR) have not yet been investigated.

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scratching or locomotion, has also been firmly established (Arshavsky et al., 1986; Muir and Whishaw, 2000). Although it has been convincingly demonstrated that this rubral complex functions as an important premotor center, at present it is not clear exactly what signals are conveyed and how its function relates to the function of the corticospinal system. Many physiologic similarities appear to exist between both systems, although some differences have also been noticed (Massion, 1988; Kennedy, 1990; Cheney et al., 1991). Either system may, at least partly, compensate for lesions in the other system (Lawrence and Kuypers, 1968b; Whishaw et al., 1998). Also, it has been suggested that the red nucleus may provide a tonic framework against which the motor cortex can produce more precise movements (Whishaw and Gorny, 1996). The red nucleus has also been implicated in mediating conditioned responses (Chapman et al., 1990; Pananceau et al., 1996; Ryou et al., 1998; Voneida, 1999). On the role of the parvicellular part of the red nucleus, i.e., in the rat those parts that project to the inferior olive (prerubral and parafascicular parts), even less is known. The role of the dentatorubroolivocerebellar circuit would appear to gain in importance in higher evolved species as this coincides with a dramatic expansion of the lateral cerebellar nucleus, the parvicellular red nucleus, the central tegmental tract, the principal olive, and the cerebellar hemisphere. This circuit may reverberate signals and match them with either descending cerebral input or ascending spinal input or both and with the intrinsic oscillations of the inferior olivary neurons (Oscarsson, 1980; De Zeeuw et al., 1998b; Kistler and van Hemmen, 1999). Kennedy (1990) has proposed that the parvicellular part of the red nucleus may be functioning as a switching device designed for automatization of learned movements. Once movements are learned by the motorcortex, its connections to the parvicellular red nucleus serve, by way of the rubroolivocerebellar pathway, to automize or condition the rubrospinal pathway. Indeed, lesion studies seem to be in accordance with such a notion (Kennedy and Humphrey, 1987; Fanardjian et al., 1999). However, although this attractive hypothesis has its merits, clearly many questions remain unanswered, one of them being that it does not explain why the rubrospinal pathway is virtually nonexistent in humans (Nathan and Smith, 1982; Massion, 1988; Paxinos et al., 1990), whereas an impressive range of automated movements are an essential element of our everyday life.

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9 Cerebellum JAN VOOGD Department of Neuroscience, Erasmus MC Rotterdam The Netherlands

(Larsell, 1948) and the cerebellum of the rat. Consequently he subdivided the cerebellum of both species into 10 lobules, numbered 1 to 10 from rostral to caudal (Fig. 1e). The deep primary fissure separates the anterior lobe from the posterior lobe. The primary and preculminate fissures, which subdivide the anterior lobe in the lobules 1 to 3 and 4 to 5, are foliated on their anterior and posterior walls and reach the lateral margin of the cerebellum. The other interlobular fissures of the anterior lobe do not reach as far laterally (Fig. 1c). Shallow indentations in the surface of lobules 4 and 5 indicate the border between the vermis and the hemispheres. These indentations are more distinct in lobule 6 (Bolk’s [1906] lobulus simplex), caudal to the primary fissure. A deep paramedian sulcus is present lateral to lobule 7, but absent in lobule 8 (the pyramis). The cortex of the pyramis continues uninterruptedly into the hemisphere as the copula pyramidis. None of the interlobular fissures in the segment of the posterior lobe, located between the primary and prepyramidal fissures, is completely continuous between the vermis and the hemisphere (Fig. 1b). At the junction of lobules 6c, lobule 7, and the hemisphere, the cerebellar cortex is interrupted and the white matter comes to the surface (Figs. 1b, 6f, 6h, and 6j). Three fissures come together at this point: the vermal segment of the posterior superior fissure, located between lobules 6 and 7; the hemispheral segment of the posterior superior fissure, which separates the simple lobule from the crus 1 of the ansiform lobule; and the intercrural fissure of the ansiform lobule. The ansoparamedian fissure, located

The cerebellum of the rat is used extensively in neurobiologic research, but no systematic description of its morphology is available. This chapter deals with the anatomy of the lobes and lobules, their afferent and efferent connections, the zonal distribution of Purkinje cells, and the structure of the cerebellar nuclei of the rat. The precerebellar nuclei and their cerebellar projections are reviewed in this volume by Ruigrok (Chapter 8). This chapter does not include a review of the histology of the cerebellar cortex of the rodent. For information on this subject the reader is referred to Ramon y Cajal’s (1911) original studies, to the monograph of Palay and Chan-Palay (1974), and to the recent surveys of Dino et al. (1999, 2000). The chemical neuroanatomy of the cerebellum, including the “diffuse” catecholaminergic and cholinergic afferent systems, was reviewed by Voogd et al. (1996b). A short reviews of the structure and connections of the cerebellum was published by Voogd and Glickstin (1998).

THE GROSS ANATOMY OF THE CEREBELLUM Early references to the gross anatomy of the cerebellum of the rat can be found in the papers of Bradley (1904), Bolk (1906), Ingvar (1919), Riley (1928), and Açiron (1951). The development and the adult configuration of the lobes and lobules of the cerebellum of the rat were described by Larsell (1952) and Larsell and Jansen (1970). Larsell was struck by the close similarity between midsagittal sections of the avian cerebellum

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FIGURE 1 The cerebellum of the rat: graphic reconstructions from serial sections. The anterior lobe has been removed on the right side of the figures to show the simple lobule in the caudal bank of the primary fissure. Interruptions of the cortex are hatched. (a) Caudal aspect. (b) Dorsal aspect. (c) Rostral aspect. (d) Ventral aspect. (e) Midsagittal section. (f) Diagram of the cortical loop of the paraflocculus and the flocculus. Abbreviations for this and subsequent figures: 1–10, lobules of the cerebellum; A, cerebellar zone A; ANT, anterior lobe; ApmF, ansoparamedian fissure; B, cerebellar zone B; C1, crus 1 of the ansiform lobule; C1–C3, CX, cerebellar zone C1–C3 or CX; c2, cervical segment 2; C2, crus 2 of the ansiform lobule; CeC, cerebellar commissure; CL, central lateral nucleus; CO, cochlear nucleus; COP, copula pyramidis; D1, D2, D0, cerebellar zones D1, D2, D0; DA, dorsomedial subnucleus of the POv; DAOv, d, dorsal or ventral fold of the dorsal accessory olive; DCo, dorsal cochlear nucleus; DLH, dorsolateral hump (central cerebellar nuclei); DLP, dorsolateral protuberance; DMC, dorsomedial crest (central cerebellar nuclei); dmcc, dorsomedial cell column; FB, fast blue; FL, flocculus; FLped, floccular peduncle; GABA, γ-aminobutyric acid; GI, gigantocellular nucleus; Glu, glutamate; ic, internal capsule; IC, interstitial cell groups; IcF, intercrural fissure; icp, inferior cerebellar peduncle; Inf, infracerebellar nucleus; Int, interposed cerebellar nucleus; IntA, anterior interposed nucleus; IntP, posterior interposed nucleus; IntPpc, parvocellular part of the posterior interposed nucleus; IO, inferior olive; IPfls, intraparafloccular sulcus; jrb, juxtarestiform body; Lat, lateral cerebellar nucleus; Latpc, parvocellular part of the lateral cerebellar nucleus; LD, laterodorsal nucleus (thalamus); LVe, lateral vestibular nucleus; MAO, medial accessory olive; MAOc, r, caudal or rostral part of the medial accessiory olive; mcp, middle cerebellar peduncle; MD, mediodorsal nucleus of the thalamus; Med, medial cerebellar nucleus; MedCM, caudomedial subdivision of the medial cerebellar nucleus; MedDLP, dorsolateral protuberance of the medial cerebellar nucleus; MedM, middle subdivision of the medial cerebellar nucleus; MedMpc, parvocellular part of the middle subdivision of the medial cerebellar nucleus; MV, medial vestibular nucleus; NI, interposed cerebellar nucleus; NL, lateral cerebellar nucleus; NM, medial cerebellar nucleus; NY, nuclear yellow; ocf, olivocerebellar fibers; PCrt, parvocellular reticular formation; PFL, paraflocculus; PflS, parafloccular sulcus; PIF, posterolateral fissure; PM, paramedian lobule; PmS, paramedian sulcus; Pod, v, dorsal or ventral lamina of the principal olive; PpF, prepyramidal fissure; Pr, nucleus prepositus hypoglossi; PreculF, preculminate fissure; PrF, primary fissure; PsF, posterior superior fissure; SC, superior colliculus; scp, superior cerebellar peduncle; SecF, secondary fissure; Sim, simple lobule; smv, superior medullary velum; spS, spinal trigeminal nucleus; SpVE, spinal vestibular nucleus; SuVe, superior vestibular nucleus; unc, uncinate tract; Vco, ventral cochlear nucleus; VL, ventrolateral nucleus (thalamus); VM, ventromedial nucleus (thalamus); vsc, ventral spinocerebellar tract; X, zone X; Y, group Y of the vestibular nuclei.

between the caudal folium of crus 2 of the ansiform lobule and the rostral folium of the paramedian lobule, ends in the paramedian sulcus lateral to lobule 7. The relation between the vermis and the hemispheres clearly differs for different segments of the cerebellar cortex. Functionally, the mediolateral continuity of the cortex depends on the presence of parallel fibers, that is, of a molecular layer (Marani and Voogd, 1979; Voogd, 1975). In the anterior lobe, the simple lobule, and lobule 8, the cortex of the vermis continues uninterruptedly into the hemispheres. In between lobules 6b and 6c and 7, and the ansiform and paramedian lobules, however, the cortex bridging the paramedian sulcus is greatly constricted or even completely absent. In Bolk’s (1906) terms, the folial chains of vermis and hemisphere of this part of the posterior lobe are completely independent of each other. The cortex of lobules 9 (the uvula) and 10 (the nodule) and the secondary and posterolateral fissures ends in a deep paramedian sulcus which separates these lobules from the copula pyramidis (Fig. 1d). Laterally the copula continues into the paraflocculus. The cortex of the paraflocculus constitutes a laterally directed loop, which is continuous with the cortex of the flocculus at the bottom of the hemispheral segment of the posterolateral fissure (Fig. 1f). The cortex of the paraflocculus is interrupted in the center of the loop in the so-called intraparafloccular sulcus. These areas, where the central white matter comes to the surface,

are found at the caudoventral and rostral aspects of the paraflocculus. For descriptive purposes the dorsal and ventral limbs of the loop are distinguished as the dorsal and ventral paraflocculus, but this distinction is secondary to the essential continuity of the folial chain of the hemisphere. The paraflocculus of the rat is located in the fossa subarcuata, a bony cavity on the posterior surface of the petrosal bone. Larsell (1952) has stated that a lateral extension of the secondary fissure separates the dorsal from the ventral paraflocculus. No such continuity exists in the rat. The fissures of this part of the cerebellum develop independently in the cortex of the caudal vermis and in the hemisphere and end at the white matter in the paramedian and interparafloccular sulci, which separates the caudal vermis from the paraflocculus and the flocculus. The taenia of the roof of the fourth ventricle is attached to the margin of the nodule, the copula pyramidis, and the flocculus. A posterior medullary velum is not present, although the areas devoid of cortex, bordering the tenia in the paramedian sulcus and the ventral aspect of the paraflocculus, could be considered as such. The superior medullary velum is continuous with the cerebellar commissures in the central white matter of the cerebellum. The morphology of the cerebellum of the rat conforms to the general mammalian pattern as described by Bolk (1906), Riley (1928), and Voogd et al. (1998). The

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cortex of the vermis and hemispheres is continuous in the anterior lobe and the simple lobule and in a restricted portion of the posterior lobe between the prepyramidal and secondary fissures. In the intermediate and caudal segments of the posterior lobe, the vermis and hemispheres behave as independent folial chains, and mediolateral connections between them are absent or greatly attenuated. In most respects, this description of the cerebellum of the rat closely corresponds to the observations of Larsell (1952) and to the description of the mouse cerebellum by Marani and Voogd (1979), to which the reader is referred for further details. Interruptions of the cerebellar cortex between the ansiform lobule and the vermis, and in the center of the parafloccular loop, were also found to be present in the mouse and in most other mammalian species investigated (Voogd et al., 1998). Larsell’s account remains indispensible as the basis for the nomenclature of the rat cerebellum. It should be noted that the vermal segment of the lobulus simplex is denoted as lobule 6a, and the vermal segment of the crus I as the lobules 6b and c. This differs from the usage in other mammals, where the vermis of the lobulus simplex constitutes the entire lobule 6, and the vermal segment which is in continuity with the crus I corresponds to the rostral segment of lobule 7 (lobule 7A). Lobules 6 and 7, therefore, should not be considered as homologs of the lobules bearing the same numbers in other mammalian species. In recent years studies of deviations from the normal folial pattern in mice mutants or transgenic mice have appeared. Combined with an analysis of modifications of longitudinal zonal patterns in the expression of Purkinje cell-specific substances, it was proposed that the cerebellum is subdivided into five transverse zones characterized by independent variations in their transverse and longitudial patterns (Ozol et al., 1999).

THE CEREBELLAR NUCLEI AND THEIR EFFERENT PATHWAYS The Subdivision of the Cerebellar Nuclei The cerebellar nuclei usually are subdivided according to Weidenreich (1899). His scheme was applied to pinnipedia and cetacea by Ogawa (1935) and extended by Ohkawa (1957), whose comparative anatomic studies included rodents. These authors divided the cerebellar nuclei into two groups of interconnected nuclei. The caudal group consists of the medial cerebellar or fastigial nucleus and the posterior interposed nucleus; the rostral group consists of the anterior interposed nucleus and the lateral cerebellar or dentate nucleus (Fig. 2). Myelinated fibers occupy the space between the two nuclear groups;

the border between the two nuclei within a group often is more difficult to define. Korneliussen (1968) applied this subdivision to the cerebellar nuclei of the rat. His description takes account of the presence of certain subnuclei which are peculiar to the rat and which were first described by Goodman et al. (1963). His description was adopted in most experimental studies of the connections of the nuclei. It also served as the starting point for the detailed Golgi and morphometric studies of the dentate nucleus (Chan-Palay, 1977) and the medial cerebellar nucleus (Beitz and Chan-Palay, 1979a, 1979b) of the rat. Additional cerebellar nuclei which should be considered as separate structures are the “interstital cell groups,” located between the caudal medial and the posterior interposed nucleus (Buisseret-Delmas et al., 1993) and the basal interstitial nucleus (Langer, 1985). The latter is a group of small, acetylcholinesterase-positive neurons, which extends from the white matter of the flocculus, in the roof of the fourth ventricle, next to the cerebellar nuclei, to the white matter of the nodulus. It exists in the rat, but has not been studied in great detail (Komei et al., 1983). Neurons of the cerebellar nuclei constitute a mixed population of cells of all shapes and sizes. Several authors noticed a binominal distribution for cell size in the cerebellar nuclei (Courville and Cooper, 1970; ChanPalay, 1977, monkey; Palkovits et al., 1977, cat). This distribution can be explained by the presence of a population of small, inhibitory, GABAergic neurons and a population of excitatory, presumably glutaminergic neurons of different sizes, as reported by Batini et al. (1992) for the rat cerebellar nuclei (Fig. 3). The excitatory neurons give rise to highly branching axons, with collaterals which may descend to the spinal cord and ascend to the thalamus (Fig. 7) (Bentivoglio and Kuypers, 1982; Bentivoglio and Molinari, 1986; Gonzalo-Ruiz and Leichnetz, 1987; Lee et al., 1989; Teune et al., 1995; Teune, 1999). The small GABAergic neurons project preferentially to the inferior olive (see Ruigrok, this volume, Chapter 8, for particulars). GABAergic nuclear cells with projections to the cerebellar cortex have been found by several authors (Chan-Palay et al., 1979; Angaut et al., 1988; Batini et al., 1989). However, most nucleocortical fibers take their origin as collaterals from the putative glutaminergic relay cells of the cerebellar nuclei and terminate as mossy fibers in the cerebellar cortex (McCrea et al., 1978; Hámori and Takács, 1989; Hámori et al., 1990; see also Tolbert et al., 1980). Consequently the sizes of nucleocortical cells in the cat follow the same distribution as neurons which could be retrogradely labeled from the thalamus (Tolbert et al., 1978). The nucleocortical projection in the rat was studied by Buisseret-Delmas and Angaut (1988, 1989b).

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FIGURE 2 The cerebellar nuclei of the rat. Transverse section 1 is the most caudal one. The three-dimensional diagram is a dorsal view of the nuclei. Redrawn from Voogd et al. (1996a). Abbreviations as in Fig. 1

A population of small glycinergic interneurons of the cerebellar nuclei, some of which colocalize GABA, was found by Chen and Hillman (1993b) in the rat. Their content of glycine distinguishes these cells from the GABAergic neurons which project to the olive, because these cells never colocalize glycine (de Zeeuw, personal communicaton). According to Gruesser-Kornehls and Bäurle (2001), the appearance of parvalbuminexpressing neurons in the cerebellar nuclei of certain mouse mutants is due to the activity of glycinergic and GABAergic interneurons which exert an increased inhibition of the nuclear neurons and which compensate for the loss of inhibition by the Purkinje cells. The organization of the cerebellar nuclear efferents generally supports the distinction of the two groups of nuclei. Voogd (1964) and Verhaart (1970) described a

subdivision of the superior cerebellar peduncle, which contains the ascending fibers of several nuclei, into a smaller medial part and a larger lateral portion, in most mammals studied. The medial third of the superior cerebellar peduncle of the cat contains fibers from the medial cerebellar and posterior interposed nuclei. The lateral two-thirds of the peduncle contain efferents from the anterior interposed and lateral cerebellar nuclei. This localization was confirmed by Haroian et al. (1981) for the rat (Fig. 4). The small caliber, GABAergic nucleoolivary tract connects the lateral and interposed nuclei with the contralateral inferior olive. These fibers collect in the lateral angle of the fourth ventricle and ascend in a bundle located ventral to the superior cerebellar peduncle to their decussation (Chan-Palay, 1977, monkey; Legendre and Courville, 1987, cat; Cholley et al., 1989, rat).

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FIGURE 4 The fiber composition of the superior cerebellar peduncle in the rat. Relabeled and reproduced from Haroian et al. (1981). Abbreviations are as in Fig. 1.

FIGURE 3 Cell-diameter distributions in the nucleus medialis (Fsg), the nucleus interpositus (Int), and the nucleus lateralis (Lat) of GABAergic and glutaminergic neurons in rat cerebellar nuclei. The populations of GABA-immunoreactive neurons are tabulated under panel A, and glutamate-immunoreactive neurons under panel B. In panel 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 micrometers (class interval, 2.5 μm). Ordinates: percentage of neurons in each diameter class. From Batini et al. (1992).

The Medial (Fastigial) Cerebellar Nucleus The medial cerebellar nucleus of the rat is characterized by the prominent dorsolateral protuberance of Goodman et al. (1963), a group of large neurons extending far dorsally into the white matter of the posterior lobe (Figs. 2 and 6). Korneliussen (1968) further subdivided the medial nucleus into middle and caudomedial portions. The caudomedial subdivision is the most distinct one. Most of its cells are small (Beitz and Chan-Palay, 1979a; Beitz, 1982). The caudomedial subdivision of the medial nucleus is located at the base of the nodule and the uvula. Dorsally, it remains separated from the rest of the nucleus by myelinated fibers;

ventrally, where it lines the roof of the fourth ventricle, it merges with the middle portion of the medial nucleus. The middle subdivision is distinguished by its high content of myelinated fibers which belong to two groups. The uncinate tract emerges from and traverses the nucleus on its way to the cerebellar commissure. Smaller, so-called “perforating fibers,” traverse its caudal part, medial to the dorsolateral protuberance, on their way to the vestibular nuclei. These fibers originate from Purkinje cells of the anterior vermis (Voogd et al., 1991). The unique shape of the dorsolateral protuberance and its afferent corticonuclear connections from the hemisphere of the posterior lobe (Goodman et al., 1963; Buisseret-Delmas, 1988a; Buisseret-Delmas and Angaut, 1993; Armstrong and Schild, 1978a, 1978b) set it apart from the rest of the medial nucleus, which receives its corticonuclear projection from the vermis, and preclude its identification with any of the subdivisions of the medial cerebellar nucleus of other species such as the cat and monkey. The uncinate tract takes its origin from the entire medial nucleus, including the dorsolateral protuberance. It crosses the midline in the caudal part of the cerebellar commissure (Fig. 5), rostral to the gliotic, interfastigial area, in small bundles dorsal and caudal to this area, and in the superior medullary velum. Contralaterally, uncinate fibers pass rostral to and through the hilus of the medial nucleus (Fig. 6). The tract arches dorsal to the superior cerebellar peduncle, immediately rostral to the anterior interposed nucleus, to join the inferior cerebellar peduncle in its course lateral to the vestibular nuclei. Some of the fibers of the uncinate tract join the medial part of the superior peduncle as the crossed ascending limb of the uncinate tract (Haroian et al., 1981) (Fig. 4). Over most of their intracerebellar course, the efferent fibers of the uncinate and the superior cerebellar peduncle remain separated from the spino- and reticulocerebellar fibers

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or uncinate tract fibers passing through the nucleus, or did not pay attention to the intracerebellar course of the system. The main projections of the rostromedial portion of the nucleus are bilateral and include the vestibular nuclei (mainly the magnocellular portion and more caudal ventral and lateral regions of the medial vestibular nucleus, the spinal vestibular nucleus, and the parasolitary nucleus) and the bulbar reticular formation. Rostrally this projection extends into the ipsilateral pontine reticular formation, with minor targets in the mesencephalon and the diencephalon. The dorsolateral protuberance gives rise to a major projection to the contralateral bulbar medial reticular formation, extending into the pons in a region bordering on the trigeminal nuclei and including the parabrachial nuclei. In the mesesencephalon terminations include the deep mesencephalic nucleus and the adjoining central gray. Collateralization to the bulbar reticular formation and the thalamus has been observed (Fig. 7) (Bentivoglio and Kuypers, 1982). Efferents of the caudomedial medial nucleus focus on the contralateral paramedian pontine reticular formation, with strong projections to the pararubral area, the deep mesencephalic nucleus, the central gray, deep layers of the superior colliculus, and regions adjoining the oculomotor nuclei and the fasciculus retroflexus. Thalamic targets include the parafascicular, ventromedial and ventrolateral nuclei. Collaterals of the same neuron may terminate in the spinal cord, the bulbar reticular formation, the tectum, and the thalamus (Fig. 7). Nucleoolivary fibers from the medial nucleus are considered by Ruigrok (this volume, Chapter 8). FIGURE 5 (Top) Midsagittal section through the cerebellum of the rat. For symbols see Fig. 6. (Bottom) Drawing of Häggqvist-stained section through the cerebellar commissure, with contributions from the uncinate tract and the inferior and middle cerebellar peduncles. Abbreviations are as in Fig. 1.

of the inferior cerebellar peduncle by a layer of thin, olivocerebellar fibers. Uncrossed fastigiobulbar fibers take their origin from the middle and caudomedial subdivisions of the fastigial nucleus, but a contribution of the dorsolateral protuberance seems to be small or absent (Voogd et al., 1985). The few experimental studies on the efferents of the medial cerebellar nucleus of the rat (Achenbach and Goodman, 1968; Angaut and Cicirata, 1982; Ruigrok and Voogd, 1990; Watt and Mihailoff, 1983; Teune, 1999; Teune et al., 2000) either used silver impregnation methods for degenerated axons after lesions of the medial nucleus, which always interrupt the corticofugal

The Posterior Interposed Nucleus and the Interstitial Cell Groups The posterior interposed nucleus is the smallest of the central nuclei of the rat, but it has a very high cell density (Figs. 6c–6i). It contains rather large cells; small cells are more numerous ventrally. A cell group located between the posterior interposed nucleus and the fastigial nucleus, which formerly was included with the posterior interposed, was considered as an independant cerebellar nucleus by Buisseret-Delmas et al. (1993), because it it serves as the target nucleus for one of the corticonuclear projection zones of the anterior lobe (the X zone, see also Trott and Armstrong, 1987b). It is known as the interstitial cell group (Figs. 2 and 10C). The efferent connections of the posterior interposed nucleus of the rat have been studied by Haroian et al. (1981), Daniel et al. (1987), and Teune et al. (2000). They cross in the dorsal part of the decussation of the superior cerebellar peduncle and terminate along the

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FIGURE 6 (A–F) Sagittal sections through the cerebellum of the rat. (a, c, h, g, and i) Nissl-stained sections through the medial part of the cerebellar nuclei. (b, d, f, h, j, and k) Drawings of parallel Häggqvist-stained sections. Coarse fibers of the restiform body are indicated with open circles; olivocerebellar fibers are in black. Pontocerebellar fibers are stippled; the efferent fibers in the superior cerebellar peduncle are hatched. Calibration (bar, 1 mm) refers to Nissl-stained sections. Abbreviations are as in Fig. 1.

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FIGURE 6, cont’d (A–F) Sagittal sections through the cerebellum of the rat. (a, c, h, g, and i) Nissl-stained sections through the medial part of the cerebellar nuclei. (b, d, f, h, j, and k) Drawings of parallel Häggqvist-stained sections. Coarse fibers of the restiform body are indicated with open circles; olivocerebellar fibers are in black. Pontocerebellar fibers are stippled; the efferent fibers in the superior cerebellar peduncle are hatched. Calibration (bar, 1 mm) refers to Nissl-stained sections. Abbreviations are as in Fig. 1.

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FIGURE 7 (a–c) Double-labeling of cells of the cerebellar nuclei of the rat after combinations of injections of fluorescent tracers in the thalamus, the superior colliculus, the medial bulbar reticular formation, and the spinal cord. Relabeled and reproduced from Bentivoglio and Kuypers (1982). (d) Retrograde labeling of cells of the dorsolateral hump after injections of fast blue (FB) in the lateral, parvocellular reticular formation and double labeling of cells in more ventral parts of the interposed nucleus after combined injections of nuclear yellow (NY) in the thalamus. From Bentivoglio and Molinari (1986). Abbreviations are as in Fig. 1.

medial margin of the red nucleus, the central gray, the deep mesencephalic nucleus, the deep layers of the superior colliculus, the nucleus of Darkschewitsch, the subparafascicular nucleus, and the zona incerta. Their thalamic targets include the ventromedial, ventrolateral, and the intralaminar nuclei. The contribution of the posterior interposed nucleus to the crossed descending limb of the superior cerebellar peduncle is small. In the

rat, the posterior interposed nucleus does not contribute fibers to the pontine nuclei (Watt and Mihailoff, 1983). Small caliber fibers descend dorsolateral to the pyramidal tract to the level of the inferior olive, where they terminate on the rostral part of the medial accessory olive and the ventral lamella of the principal olive (Swenson and Castro, 1983a, 1983b; Daniel et al., 1987; Ruigrok and Voogd, 1990; Ruigrok, this volume,

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Chapter 8). The retrograde labeling studies of Bentivoglio and Kuypers (1982) in the rat did not distinguish between the anterior and posterior interposed nuclei. Cells projecting to the spinal cord appeared to be most numerous in the region of the interstitial cell group (Fig. 9C). They collateralize to the superior colliculus, the thalamus, and the medial reticular formation (Bentivoglio and Kuypers, 1982) (Fig. 7). Other connections of the interstitial cell groups were detailed by Buisseret-Delmas et al. (1998).

The Anterior Interposed (Interpositus) Cerebellar Nucleus, the Dorsomedial Crest, and the Dorsolateral Hump The dorsomedial crest and the dorsolateral hump were described by Goodman et al. (1963) as lateral and dorsal protrusions of the undivided interposed nucleus. The cells of the dorsomedial crest and the adjoining medial part of the anterior interposed nucleus are smaller than the cells of the lateral part of this nucleus. A distinct border is present between the small cells of the dorsomedial crest and the larger cells of the posterior interposed nucleus. The dorsolateral hump is a ridge of small cells on the rostrolateral and dorsal surface of the anterior interposed nucleus. Korneliussen (1968) included the lateral fourth of the anterior interposed nucleus in the hump. When the hump is defined

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in this way, it includes the large cells of the caudal pole of the anterior interposed nucleus, which lie intercalated between the posterior interposed and the lateral nucleus. Hump and caudal pole can also be distinguished as separate bulges in the surface relief of the nuclei. (Fig. 2). According to Woodson and Angaut (1984), the dorsolateral hump is the main origin of the uncrossed descending branch of the superior cerebellar peduncle. According to Ruigrok and Voogd (1990) it also includes the portion of the anterior interposed nucleus ventromedial to the hump. Originally, this system was described by Ramon y Cajal (1903, 1911) with the Golgi method (Fig. 8). Mehler (1967, 1969) retraced it with the Nauta method in rats and guinea pigs. Its fibers enter the brain stem between the motor and principal sensory nuclei of the trigeminal nerve (Fig. 10B) and descend in the lateral reticular formation to terminate here or in deep layers of the principal and spinal trigeminal nuclei. Some fibers descend as far as the spinal cord (Achenbach and Goodman, 1968; Faull, 1978; Woodson and Angaut, 1984). The origin of the uncrossed descending branch of the superior peduncle was illustrated by Bentivoglio and Molinari (1986) (Fig. 7). Efferents of the anterior interposed nucleus, including those of its dorsolateral hump, travel in the middle part of the superior cerebellar peduncle (Fig. 4). The anterior interposed nucleus contributes to the crossed ascending

FIGURE 8 Sagittal section showing the origin of the uncrossed descending branch of the superior cerebellar peduncle of the mouse using the Golgi method. Reproduced from Ramon Y Cajal (1911). Original labeling: A, rootfibers of the trigeminal nerve; B, bifurcation of the vestibular nerve; C, superior cerebellar peduncle; D, uncrossed descending branch of the superior cerebellar peduncle; E, inferior cerebellar peduncle; G, middle cerebellar peduncle; H, trapezoid body; O, lateral cerebellar nucleus; a, ascending branch of the trigeminal root; b and d, spinal tract of the trigeminal nerve.

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and descending branches of the superior cerebellar peduncle. The terminations of the ascending branch in the magnocellular part of the red nucleus and the ventrolateral complex of the thalamus, of the descending branch in the basilar pons and the reticulotegmental nucleus, and of the nucleoolivary fibers in the dorsal accessory olive are discussed by Ruigrok (this volume, Chapter 8). Cells projecting to the ventrolateral complex of the contralateral thalamus and the contralateral medial bulbar reticular formation, including the inferior olive, are found over the entire anterior interposed nucleus. Cells with collateral projections to the thalamus and superior colliculus are located in the lateral part of the nucleus; cells with double projections to the thalamus and the medial bulbar reticular formation extend more laterally into the dorsolateral hump. Fibers of the anterior interposed nucleus do not descend into the spinal cord. These features are illustrated in Fig. 7.

The Lateral (Dentate) Cerebellar Nucleus, Group Y, and the Basal Interstitial Nucleus of Langer The lateral cerebellar nucleus of the rat consists of a dorsolateral magnocellular portion and a ventromedial parvocellular portion (Korneliussen, 1968). The cytoarchitecture of the lateral nucleus was analyzed by ChanPalay (1977). Fusiform cells, belonging to the infracerebellar nucleus of Gacek (1977, 1979), corresponding to the dorsal group Y of Highstein and Reisine (1979) in the cat and/or the basal interstitial nucleus of Langer (1985), are located ventral to the lateral nucleus, within the floccular peduncle. Cells of the ventral group Y are located as a compact subnucleus ventromedial to the floccular peduncle, capping the inferior cerebellar peduncle (Fig. 6). The efferent connections of the lateral nucleus are contained in the ventral and lateral parts of the superior cerebellar peduncle (Haroian et al., 1981) (Fig. 4). Some of the afferents of the group Y probably take the same route. The ventral group Y neurons give rise to commissural and cerebellar connections; the infracerebellar nucleus (dorsal group Y) projects to the oculomotor complex (Highstein and Reisine, 1979). The lateral nucleus contributes to the crossed ascending and descending branches of the superior cerebellar peduncle. The terminations of the crossed ascending fibers in the parvocellular red nucleus and in the thalamus and of the crossed descending fibers in the pontine nuclei and the reticulotegmental nucleus and the nucleoolivary projection to the principal olive were reviewed by Teune et al. (2000) and Ruigrok (this volume, Chapter 8).

The double-labeling study of Bentivoglio and Kuypers (1982) confirmed the projection of the lateral, magnocellular part of the lateral nucleus to the thalamus. Collateral projections to the superior colliculus and the spinal cord and the medial bulbar reticular formation arise from different cell groups (Fig. 7).

LONGITUDINAL, ZONAL ORGANIZATION OF PURKINJE CELLS IN THE CEREBELLAR CORTEX: CHEMOARCHITECTURE AND CONNECTIONS Corticonuclear Projection Zones Although the paradigm of the essential similarity in the longitudinal organization of the corticonuclear and olivocerebellar projections was established in anatomical and electrophysiological studies in the cat (Voogd, 1964, 1969; Voogd and Bigaré, 1980; Armstrong et al., 1974; Trott and Armstrong, 1987a, 1987b; Oscarsson, 1969, 1973), the relation between this longitudinal pattern and the chemoarchitecture of the cerebellar cortex only can be studied in rodents, with a clear, zonally distributed chemical heterogeneity of the Purkinje cells (Scott, 1964; Hawkes et al., 1985). In this section I review studies on the corticonuclear and olivocerebellar projections in the rat, culminating in Buisseret-Delmas’ (1988a, 1988b) demonstration of their organization in similar zonal patterns, the evidence of the chemical heterogeneity of the Purkinje cells and their astroglial satellites, the Bergmann glia, and the still fragmentary evidence of the relationship of the connections of the Purkinje cells to their histochemical identity. It has been known since the Marchi studies of Klimoff (1899) in the rabbit that the corticonuclear projection is strictly uncrossed and that the cerebellar vermis is connected with the medial cerebellar nucleus and the hemisphere with the interposed and lateral cerebellar nuclei. Corticovestibular fibers originate from the vermis and the flocculus. Since Klimoff’s time, an impressive amount of detail has been assembled, mainly in the cat and the rabbit, on the projection of different lobules to different combinations of the cerebellar and vestibular nuclei. Most of these older studies, which were reviewed by Voogd (1964), Larsell and Jansen (1972), and Haines et al. (1982), used large lesions and employed arbitrary criteria to define the borders between the vermis and the hemisphere and between the different central cerebellar nuclei. Information on the lobular organization of the corticonuclear projection in the rat is fairly substantial, but, as pointed out by Armstrong and Schild (1978a), the localization in the corticonuclear

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projection is sharper in a mediolateral direction than in a rostrocaudal direction. Attention, therefore, must be focused on the existence in the cerebellar cortex of longitudinal zones with a specific projection to the central cerebellar or vestibular nuclei. A subdivision of the cerebellar cortex into medial (vermal), intermediate, and lateral zones projecting to the medial, interposed, and lateral cerebellar nuclei, respectively, was introduced by Jansen and Brodal (1940, 1942) and applied to the rat cerebellum by Goodman et al. (1963). The position of the borders between these three cortical zones depended on the position of the arbitrary borders between the medial, interposed, and lateral nuclei, and not on specific landmarks in the cortex itself. This also holds for the border between the vermis and the hemisphere in the rat, which is distinct only for the lobules 6b, 6c, 7, 9, and 10. An important deviation from Jansen and Brodal’s three-zone concept was the observation of Goodman et al. (1963) that the medial hemisphere of the rat cerebellum projects to the dorsolateral protuberance of the fastigial nucleus. This unique projection, which is not present in carnivores and primates, later was confirmed in the studies of Armstrong and Schild (1978a, 1978b) and Haines and Koletar (1979). Umetani et al. (1986) established that it originates from the region located between the primary and prepyramidal fissures. Historically ideas on the longitudinal organization in the corticonuclear projection are based on the observation that Purkinje cell axons use morphologically distinct, parasagittal compartments in the cerebellar white matter to reach their target nuclei. Myelins stains, like the Häggqvist method, which allow the distinction of the fairly coarse, myelinated axons of the Purkinje cells from other constituants of the white matter (Voogd, 1964, 1969), or acetylcholinesterase staining, which accentuates the borders between the white matter compartments (Hess and Voogd, 1986), has been used to define the architecture of the white matter compartments in carnivores, primates, and the rabbit. These methods have never been systematically and successfully applied to the cerebellum of the rat. In other species, knowledge of the compartmentalized architecture of the white matter, in combination with studies using antegrade or retrograde axonal tracing techniques, resulted in the designation of a fairly stereotyped zonal pattern in the corticonuclear projection. This pattern is characterized by the projection of one or more longitudinal Purkinje cell zones to a single cerebellar or vestibular target nucleus (Voogd and Bigaré, 1980). A major advance in our knowledge of the longitudinal organization of the cerebellar cortex was made when it became clear that the organization of the olivocerebellar and the corticonuclear projections is very similar (Voogd,

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1969; Courville et al., 1974; Groenewegen and Voogd, 1977; Groenewegen et al., 1979). More recently, BuisseretDelmas and her collaborators were able to confirm the principles and the longitudinal pattern in the corticonuclear and olivocerebellar projection in the rat, in a systematic analysis of small injections of WGA–HRP in the cerebellar cortex of this species (Buisseret-Delmas, 1988a, 1988b; Buisseret-Delmas and Angaut, 1989a, 1993; Buisseret-Delmas et al., 1993; Yatim et al., 1995). According to Buisseret-Delmas (Fig. 9) three corticonuclear projection zones—A, X, and B—can be distinguished in the vermis. The medial A zone extends over the entire vermis and projects to the middle and caudomedial subdivisions of the fastigial nucleus (see also Päällysaho et al., 1990; Umetani, 1989; Tabuchi et al., 1989; Voogd and Ruigrok, 1997). The X zone is present in the anterior lobe, probably extends into lobule 6a, and, according to Yatim et al. (1995), is also represented in lateral lobules 9 and 10 (Fig. 10). It projects to the interstitial cell groups. The B zone occupies the lateral vermis of the anterior lobe and lobule 6a and projects to the lateral vestibular nucleus (Voogd et al., 1991). In the cerebellar hemisphere the A2 zone, three C zones, and three D zones were distinguished. The A2 zone (the lateral extension of the A zone of BuisseretDelmas, 1988a) projects to the dorsolateral protuberance of the fastigial nucleus. A2 is represented in the lobulus simplex, the crura of the ansiform lobule, and the paramedian lobule (Fig. 11I). According to BuisseretDelmas (1988a) A2 is continuous with the vermal A zone; in our experience the B zone separates A from A2 in the lobulus simplex. Lateral to A2 the C2 zone, with the flanking C1 and C3 zones, is located. In carnivores C1 and C3 were found to project to the anterior interposed nucleus and C2 to the posterior interposed nucleus. A different projection was advocated by BuisseretDelmas (1988b) in the rat: C1 is connected with medial and C2 with more lateral portions of both interposed nuclei and C3 only projects to the anterior interposed nucleus (e.g., Pardoe and Apps, 2002). In a recent study we confirmed the original target nuclei, as established for carnivores, in the rat (Fig. 19) (Voogd et al., 2003). C1 and C3 are confined to the anterior lobe, the lobulus simplex, the crus II, and the paramedian lobule. C1 proceeds into the copula pyramidis. C1 and C3 are absent from the crus I, the paraflocculus, and the flocculus. C2 extends over the entire cerebellum, including the paraflocculus and the flocculus. Three D zones, projecting to the lateral cerebellar nucleus, were distinguished by Buisseret-Delmas and Angaut (1989a). D0 projects to the dorsolateral protuberance, D1 to the ventral and caudal portions of the lateral nucleus, and D2 to its dorsal and rostral portions. D0 is confined to the anterior lobe, the lobulus simplex, crus

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FIGURE 10 Diagrams of the connections of the X and Cx zones in the cerebellum of the rat. (A) Localization of the anterior and posterior X and the Cx zones. Compare with Fig. 9 for their position relative to the B and C1–C3 zones. (B) Horizontal projection of the medial accessory olive, indicating the origin of the projection to the rostral X and Cx zones as an oblique band at the border of the rostral and caudal subdivisions of the medial accessory olive (vertical hatching) and to the caudal X zone from the dorsomedial cell column (dots). Compare with Fig. 13 for the climbing fiber projection of the dorsomedial cell column to lobules 9 and 10. (c) Termination of Purkinje cell axons within the interstitial cell groups. Panels A and B were modified from Buisseret-Delmas et al. (1993), panel C was redrawn from Buisseret-Delmas and Angaut (1993). Abbreviations: I–X, lobules I–X; β, subnucleus beta; CX, CX zone; dmcc, dorsomedial cell column; IC, interstitial cell groups; MAO, medial accessory olive; NI, interposed nucleus; NM, medial cerebellar nucleus; X, X zone. FIGURE 9 Topographical arrangement of the olivo- and corticonuclear connections of the cerebellum in the albino rat. (A) The subdivisions of the inferior olive; (1) is the most caudal transverse section. (B) Diagram of the zonal organization of the rat cerebellar cortex. (C) The cerebellar nuclei; (1) is the most caudal transverse section. Redrawn, and modified by addition of the connections of the dorsolateral hump (DLH; Buisseret and Angaut, 1989a) from BuisseretDelmas and Angaut (1993). Compare with Fig. 1g, for differences in the projection of the C1, C2, and C3 zones to the anterior and posterior interposed nucleus, of the D1 and D2 zones in their connections with the lateral cerebellar nucleus and the inferior olive, and in the sequence of the D0, D1, and D2 zones. Abbreviations are as in Fig. 1.

II, and the paramedian lobule. D1 and D2 extend over all lobules. The conclusion that each cerebellar zone projects to a single cerebellar target nucleus was challenged by Panto et al. (2001), who found a systematic distribution of corticonuclear terminals in other cerebellar nuclei. It is argued in the last section of the chapter that these additional foci of terminal labeling may represent

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mossy fiber collaterals, labeled from the cortical injection sites.

Climbing Fibers: The Olivocerebellar Projection

FIGURE 11 (I) Lateral extension of zone A (or A2 zone) in the posterior lobe with its projection to the dorsolateral protuberance (dlp) of the fastigial nucleus in the rat. The cumulative results of eight injections of WGA–HRP in the posterior lobe are illustrated. The injection sites are represented as gray areas in panel B and the corresponding afferent and efferent connections are represented in black in panels A and C, respectively. Dots indicate single labeled neurons in panel A and few sparsely labeled terminals in panel C. The lateral extension of the A zone receives a projection from the medial subnucleus C of the caudal medial accessory olive (MAO) Buisseret-Delmas (1988a). (II) Tectorecipient zones of Akaike (1992) in the medial part of the simple lobule (SI) and the crus 2 (CII). This zone is interrupted in the crus 1 and it is absent from the anterior lobe and the copula. The tectorecipient zone in lobule 7 is separated from the corresponding zone in the crus 2 by a strip with other, nonspecified, olivocerebellar connections. The tectal response zone in the hemisphere differs from the lateral extension of the A zone of Buisseret-Delmas (1988a) because it is absent from the crus 1 and from the rostral folia of the paramedian lobule. From Akaike (1992). Abbreviations: ANT, anterior lobe; COP, copula pyramidis; CrI, II, crus 1,2 of the ansiform lobule; PMD, paramedian lobule; SI, simple lobule; IV–IX, lobules IV–IX of Larsell.

The olivocerebellar projection to the cerebellar cortex and the cerebellar nuclei is topically organized. This was already known (Brodal, 1940) before it was realized that the inferior olive is the main source of climbing fibers in the rat (Desclin, 1974). Brodal (1940) investigated the projection of the inferior olive with the retrograde cell degeneration method in young cats and rabbits and concluded that specific lobules received olivocerebellar fibers from specific subdivisions of the contralateral olivary nucleus. Voogd (1969) and Oscarsson (1969) showed that each subdivision of the inferior olive projects to a particular longitudinal strip of cortex, which can be traced through a number of successive lobules. According to Voogd (1969), the olivocerebellar fibers reach the Purkinje cells of these strips through the same myeloarchitectonic compartments which contain the projection of these Purkinje cells to the central cerebellar nuclei. He concluded that the organization of the olivocerebellar projection and organization of the corticonuclear projection are essentially similar. The concept of the longitudinal zonal organization of the olivocerebellar projection was further developed in the anterograde axonal transport studies of Courville et al. (1974), Groenewegen and Voogd (1977), Groenewegen et al. (1979), and Gerrits and Voogd (1982) in the cat and applied in the retrograde transport studies, summarized in the monograph of Brodal and Kawamura (1980). Direct evidence for a longitudinal organization of the olivocerebellar projection in the rat was provided by Chan-Palay et al. (1977) using autoradiography of 35 S-labeled methionine to demonstrate the sagittal organization of the olivocerebellar projection in the rat. According to their findings, the projection is bilateral, and banded areas that receive labeled climbing fibers alternate with areas that receive climbing fibers from extraolivary sources. Convincing evidence that the injections of the inferior olive in this study must have been incomplete and that, instead, the entire cortex of the cerebellum is provided with climbing fibers from the inferior olive was obtained by Armstrong et al. (1982) and Campbell and Armstrong (1983a). These authors were unable to confirm the presence of an uncrossed component in the olivocerebellar projection of the rat. A zonal arrangement was recognized by Sotelo et al. (1984) in their autoradiographic studies in neonatal rats. Sugihara et al. (1999, 2001) reconstructed the entire trajectory of individual olivocerebellar fibers, from small injections of biotinylated destran amine (BDH) in the

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inferior olive of the rat. Figure 14 from Sugihara et al. (2001), illustrates the two or three climbing wide strip, innervated from a small focus in the caudal medial acessory olive (MAO), which extends over more than 15 mm over the cerebellar surface. The subdivision and the afferent connections of the inferior olive of the rat were discussed by Ruigrok (this volume, Chapter 8). An important contribution to the morphology of the inferior olive of the rat was made by Azizi and Woodward (1987). They subdivided the MAO into a horizontal lamella (corresponding to the subnuclei A and B of the caudal MAO), a vertical lamella (including the subnucleus C, the group beta, the dorsal cap (DC), and the ventrolateral outgrowth (VLO) and a rostral lamella (corresponding to the rostral MAO and the dorsomedial cell column (DMCC). They distinguished a dorsal fold of the caudal dorsal accessory olive (DAO), which is joined laterally to the rest of the DAO, which they indicated as the ventral fold. They distinguished the enlarged, medial extension of the ventral lamina of the principal olive (PO) from the DMCC as the dorsomedial subnucleus (DM) (Fig. 12). The zonal projections of subnuclei of the inferior olive to the Purkinje cells of the cerebelllar cortex, their collateral projections to the cerebellar nuclei, the reciprocally organized nucleoolivary connections, and the corticonuclear projections all are in perfect register (Buisseret-Delmas and Angaut, 1993; Ruigrok and Voogd, 1990, 2000; Ruigrok, this volume, Chapter 8). The zonal pattern in the olivocerebellar projection, proposed by Buisseret-Delmas (Fig. 9) (BuisseretDelmas, 1988a, 1988b; Buisseret-Delmas and Angaut, 1989a, 1993; Buisseret-Delmas et al., 1993) differs in some respects from Azizi and Woodward’s (1987) scheme of the projection of the different lamellae and folds of the rat inferior olive (Fig. 12). Both recognized a medial vermal A zone, innervated by the caudal MAO (zones 3 and 4 of Azizi and Woodward, 1987), and a lateral B zone (zone 1), innervated by the dorsal fold of the DAO. The X zone, which receives climbing fibers from an oblique strip at the border of the caudal and rostral halves of the MAO (Fig. 10), was not recognized by Azizi and Woodward (1987). The C2 zone (zone 5), flanked by the C1 (zone 2) and C3 zones, innervated by the rostral MAO and the rostral DAO, respectively, was recognized by Buisseret-Delmas (1988b) in the hemisphere, but an equivalent of the C3 zone is lacking in Azizi and Woodward’s scheme. Two zones in the intermediate part of the hemisphere which were not accounted for by Azizi and Woodward (1987) are the A2 zone (the “lateral extension of the A zone” of Buisseret-Delmas, 1988a) located in the medial hemisphere of lobules 6 and 7 and the Cx zone (Fig. 11A). The A2 zone corresponds to the lateral “tectal response

FIGURE 12 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 (MAO, 3) appear to receive afferents mainly from the spinal cord and dorsal column nuclei while projecting to the anterior vermis and parts of the intermediate cerebellum. The medial MAO (vertical lamella, 4) receives from the vestibular and visual areas and projects to the posterior vermis as well as the flocculus. The rostral lamella of the MAO and both lamellae of the principal olive (PO) receive projections from higher centers and sends 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. From Azizi and Woodward (1987).

zone” of Akaike (1986a, 1986b, 1987, 1989, 1992). The medial “tectal response zone” is located in the medial half of lobule 7. They contain the climbing fiber-evoked potentials on stimulation of the ipsilateral superior colliculus (Fig. 11II). They receive their climbing fibers from two separate, but overlapping, populations of neurons in the rostral subnucleus C (the tectorecipient zone of the MAO. The CX zone, originally, was described in electrophysiological studies of climbing fiber branching in the cat (Ekerot and Larson, 1982). It was found to receive branches from the same climbing fibers terminating in the X zone. It is located in the hemisphere, immediately medial to C1, but its peripheral response properties were found to be indistinguishable from those of the C1 zone (Trott and Armstrong, 1987a).

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Contrary to C1, it receives its climbing fibers from the MAO (Campbell and Armstrong, 1985). In the rat a CX zone, receiving climbing fibers from the DMCC, was distinguished by Buisseret et al. (1993). Its presence in the posterior lobe also was advocated by Atkins and Apps (1997) and Pardoe and Apps (2002). In the lateral hemisphere both Buisseret-Delmas and Angaut (1989a) and Azizi and Woodward (1987) distinguished medial D1 (zone 6) and lateral D2 (zone 7) zones, which receive projections from the dorsal and the ventral lamina of the PO, respectively. An additional D0 zone, innervated by the dorsomedial subnucleus of Azizi and Woodward (1987), was distinguished by Buisseret-Delmas and Angaut (1989a). The projection from the caudal MAO to the vermis is not a uniform one. Subzones, receiving their climbing fibers from different subnuclei of the caudal MAO and the group beta, have been reported for several lobules. In the anterior lobe a lateral strip of Purkinje cells may receive its climbing fibers from the subnucleus B, which receives a projection from the vestibular nuclei. These Purkinje cells have been shown to project to both the fastigial and the vestibular nuclei (Voogd and Ruigrok, 1997). Two zones have been distinguished in lobule 7. The medial zone corresponds to the medial tectorecipient zone of Akaike (1992, cf. Hess, 1982; Sugita et al., 1989). The lateral zone may receive its climbing fibers from the group beta (Furber and Watson, 1983). The olivocerebellar projection to lobules 8 and 9 was defined by Eisenman (1981a, 1984) and Apps (1990) for the rat. Alternating strips, innervated by the caudal MAO and the group beta, were present in lobule 8 (Fig. 22). The complicated innervation pattern of lobules 9 and 10 is illustrated in Fig. 13 (Voogd and Ruigrok, 1997). Branching of olivocerebellar fibers, which terminate in different sites of the cortex, has been reported in many studies. Estimates of the number of climbing fiber collaterals issued by a single neuron, based upon a comparison of the total numbers of Purkinje cells and cells of the inferior olive (Schild, 1970, about 7) and on direct observations of individual olivocerebellar fibers (Sugihara et al., 1999, 8.5 ±3.7; Sugihara et al., 2001, 7 climbing fibers on average), both in the rat, are in good accordance. The presence and the typical distribution of collaterals of single olivocerebellar fibers in the cat was first described in the electrophysiological studies of Armstrong et al. (1973), Oscarsson and Sjölund (1977a, 1977b), and Ekerot and Larson (1982). Two types of collateralization were distinguished: sagittal and transverse. In sagittal collateralization the climbing fibers were distributed over different anterior and posterior segments of the same zone or set of zones. Transverse branching of olivocerebellar fibers occurred between zones innervated by the same olivary subnucleus, i.e.,

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FIGURE 13 Diagram comparing the olivocerebellar projection zones of the uvula and the nodulus of the rat (left side) to the zebrin pattern (right side). Zebrin-positive areas are shaded; zebrin positive bands P1+ – P4+ are indicated with the numbers 1–4. Olivary subnuclei in the horizontal projections of the principal olive (PO) and the medial accessory olive (MAO) and their projections zones are indicated in with the same symbols. Note that P2+ and P3+ are bisected by climbing fiber bands from the rostral and caudal group Beta and the dorsomedial cell column (DMCC). Zebrin-negative bands P2- and P3- are innervated by the caudal MAO and the dorsomedial group (DM). DC, dorsal cap; VLO, ventrolateral outgrowth. Reproduced from Voogd et al. (1996a).

between the X and the CX zones, and between C1 and C3 (Ekerot and Larson, 1982). Most of the anatomical studies on climbing fiber branching used double retrograde labeling methods and confirmed the presence of sagittal branching patterns in cat and rat (Brodal et al., 1980; Rosina and Provini, 1983; Hrycyshyn et al., 1989; Wharton and Payne, 1985; Eisenman, 1981a; Eisenman and Goracci, 1983; Lawes and Payne, 1986; Payne et al., 1985). Transverse branching between the X and lateral CX zones was studied by Apps et al. (1991) in the cat. Direct observations of climbing fiber branching have been made by Wiklund et al. (1984) with D-aspartate tracing, by Sugihara et al. (1999, 2001) with labeling of single olivocerebellar axons (Fig. 14), and by Chen and

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Aston-Jones (1998) and Voogd et al. (2003) using cortical injections of cholera toxin B unit (CTb). Voogd et al. (2003) found collateral labeling of climbing fibers after injections of the C1, Cx, and C2 zones of the copula in the anterior lobe and after injections of A2, C1, C2, C3, and D1 zones of the paramedian lobule in corresponding

zones of the lobulus simplex. Evidence of transverse branching was found for the C1 and C3 zones of the posterior lobe, which collateralize to both the anterior C1 and C3 zone, and for the CX zone of the copula which shares collaterals with the X and CX zones of the anterior lobe.

FIGURE 14 Climbing fibers originating from small areas in the inferior olive distribute within narrow longitudinal bands in the cerebellar cortex. (a) Forty-two climbing fibers arising from six axons. The color-coding of the individual axons in the original figure has been omitted. (Inset) Lateral view of the entire axonal trajectories from the biotinylated dextran amine (BDA) injection site in the centromedial portion of the medial accesory olive. (b) The distribution of the climbing fibers plotted on the unfolded vermal cortex from the midline to the left by 1.3 mm. Blank and dotted areas in the unfolded scheme represent the cerebellar cortex exposed in the cerebellar surface and hidden in the sulci, respectively. Dotted line indicates the contour of the distribution area. (Inset) The area for the unfolded display. Reproduced from Sugihara et al. (2001).

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Zonal Organization of the Vestibulocerebellum and the Paraflocculus The vestibulocerebellum includes the nodulus (lobule 10), the flocculus, and adjacent portions of lobule 9 and the paraflocculus. Lobule 10 receives climbing fibers from a combination of vestibular-innervated (group beta, DMCC) and optokinetic olivary subnuclei (DC and VLO; Fig. 13). Climbing fibers to the flocculus mainly stem from the optokinetic subnuclei of the olive (Blanks et al., 1983; Bernard, 1987). A large proportion of the climbing fibers from the the DC and the VLO are branches from climbing fibers which also terminate in the nodulus (Takeda et al., 1989a, 1989b; and b; Maekawa et al., 1989). The corticonuclear and vestibular projectons of lobules 9 and 10 of the rat have been reported by Bernard (1987), but a complete analysis of its zonal organization (compare Wylie et al., 1994, in the rabbit) is not yet available. Ruigrok et al. (1992) analyzed the zonal organization of the olivocerebellar projection to the flocculus and the adjacent paraflocculus in the rat. Two pairs of interdigitating zones, innervated by the DC and the VLO, respectively, and a lateral C2 zone could be distinguished in the rat (Fig. 15). The distal segments of these zones crossed the posterolateral fissure and were found to receive climbing fibers from more rostromedial levels of the DC (for the FE/FE´ zones) and the ventral leaf of the PO (for the FD/FD´ zones). The projections from the DC to lobule 10 and the flocculus earlier were traced with parvalbumin immunohistochemistry in rat pups by Wassef et al. (1992b). Corticonuclear projections of the flocculus of the rat include the caudoventral parts of the lateral and interposed nuclei, the group Y, and certain vestibular nuclei (Umetani, 1992). Balaban et al. (2000) used retrograde transport from the vestibular nuclei to define its zonal organization. The zonal pattern in the dorsal flocculus and the adjacent paraflocculus closely corresponds to the spatial organization of the olivocerebellar projection as published by Ruigrok et al. (1992). Zones FD/FD´ project to the superior vestibular nucleus, and zones FE/FE´ project to the medial vestibular and rostral lateral vestibular nuclei. In the ventral flocculus the pattern is less clear (Fig. 16). These observations are in accordance with the situation in other species (Voogd et al., 1996a). Nothing is known about a posssible zonal organization in the olivocerebellar projection to the paraflocculus of the rat. According to Furber and Watson (1983), Azizi and Woodward (1987), and Buisseret-Delmas and Angaut (1993) it receives climbing fibers from the rostral MAO and the PO. Its corticonuclear projection is directed at the ventral parvocellular part of the lateral

FIGURE 15 Diagram of the projection from the inferior olive to the flocculus and the ventral paraflocculus in the rat. The medial accessory olive (MAO) and the principal (PO) are drawn as diagrams of the unfolded inferior olive; the cortex of the flocculus and the ventral paraflocculus are unfolded. The symbols in the olive flocculus and the ventral paraflocculus correspond to each other. Compare with Fig. 16. Abbreviations: C2, C2 zone; caud, caudal; dc, dorsal cap; FD, FD´, FD and FD´ zones, (projections of vlo and PO); FE, FE´, FE and FE´ zones, (projections of dc); FLOd and v, dorsal and ventral surface of the flocculus; MAO, medial accessory olive; PO, principal olive; rost, rostral; vlo, ventrolateral outgrowth; VPFL, ventral paraflocculus. From Ruigrok et al. (1992).

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FIGURE 16 The corticovestibular projection of the flocculus and the adjacent paraflocculus in the rat. Summary diagram showing the localization of retrogradely labeled Purkinje cells on the ventral and dorsal surface of the flocculus and the adjacent ventral paraflocculus, based on a series of experiments with injections of the different vestibular target nuclei. The outlines of the olivocerebellar projection zones FE(´), FD(´), and C2 based on Ruigrok et al. (1992, Fig. 15) are indicated in each diagram. (A) Injections of the caudal medial vestibular nucleus (MVe), inferior vestibular nucleus (SpVe), and nucleus prepositus hypoglossi. (B) Injections of the superior vestibular nucleus (SuVe). (C) Injections of the rostral MVe. (D) Injections of the ventral lateral vestibular nucleus (SuVe). Caudal and rostral MVe and ventral LVe receive Purkinje cell axons from different strips within the projection of the FE and FE’zones. SVN receives its main projection from the FD and FD´ zones. Redrawn from Balaban et al. (2000).

cerebellar nucleus and the lateral pole of the posterior interposed nucleus (Armstrong and Schild, 1978a, 1978b; Buisseret-Delmas and Angaut, 1993).

Longitudinal Zones: Chemoarchitecture The relative lack of information on the organization of the corticonuclear and olivocerebellar projections in rats, in the period before Buisseret-Delmas started to contribute to this topic, constrasts strongly with the abundance of information on the histochemical differentiation of longitudinal zones in the cerebellar cortex of rodents. The enzyme histochemical studies of Scott (1964, 1965) and Marani (1982, 1986) already showed the presence of alternating longitudinal zones of high and low 5´-nucleotidase (5´-N) activity in the cerebellar cortex of the vermis and the hemispheres in mice and rats. The pattern of 5´-N-positive and -negative zones is complete in the sense that it is present in all lobules of the vermis and the hemisphere and unequivocal

because, in the mouse at least, the bands are clearly delineated. The pattern is less distinct in the rat because a high background activity is present all over the molecular layer. The cellular localization of this enzyme is still disputed. Marani (1982, 1986) favored a primarily neural localization in the Purkinje cells. Others advocate a Bergmann glial plasma membrane localization (Kreutzberg et al., 1978; Schoen et al., 1987, 1988). A Purkinje cell-dependent, presumed Bergmann glial localization of 5´-N was demonstrated by Hess and Hess (1986) in Purkinje cell-deficient mutant mice. A midline band, flanked by six, symmetrically disposed, 5´-N-positive bands can be recognized in mice. The bands in the anterior and posterior lobes are not necessarily continuous. The 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 in the rest of the posterior lobe, where the 5´-N-negative zones are reduced to narrow slits.

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An identical, “zebrin” pattern was discovered by Hawkes et al. (1985). The epitope recognized by the Hawkes family of monoclonal antibodies, known as the “anti-zebrins,” is exclusively localized in Purkinje cells. The first of the zebrins (mabQ113, zebrin I) was documented by Hawkes et al. (1985) and Hawkes and Leclerc (1987) as a monoclonal which recognized a 120-Kda protein in a subset of Purkinje cells of the rat. The epitope is present in dendrites, soma, axons, and axon terminals of these Purkinje cells and these cells are arranged in longitudinal zones alternating with strips of nonimmunoreactive Purkinje cells (Fig. 17). A second antibody (anti-zebrin II), which reacts with tissue of nonmammalian and mammalian species, was produced by Brochu et al. (1990). The epitope of zebrin II is associated with a 36-kDa polypeptide, identified as aldolase C (Hawkes, 1992; Ahn et al., 1992). The zonal distribution of zebrin I- and zebrin II-immunoreactive Purkinje cells is identical (Hawkes and Leclerc, 1987; Brochu et al., 1990). The congruence of the 5´nucleotidase and the zebrin pattern was shown in mice by Eisenman and Hawkes in 1989. The zonal pattern in the distribution of zebrinimmunoreactive and nonimmunoreactive Purkinje cells is identical or very similar to that of nerve growth factor receptor protein (Koh et al., 1989; Sotelo and Wassef, 1991; Dusart et al., 1994), to the monoclonal antibody B30 of Stainier and Gilbert (1989), which recognizes two minor gangliosides, to protein kinase C delta (Chen and Hilman, 1993a), to the glutamate transporter EAAT4 (Dehnes et al., 1998), and to the GABAB slice variant GB1b (Fritschy et al., 1999). A reversed pattern, i.e., colocalization with zebrin-negative Purkinje cells, has been found for P-path-immunoreactive

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Purkinje cells in the cerebellum of the mouse (Edwards et al., 1989) and for the anti-BW66 antibody, against the microtule-associated protein MAP 1a (Touri et al., 1996). Colocalization of zebrin II and P-path immunoreactivity only occurs in restricted regions (Leclerc et al., 1992). A distribution which partially complements the zebrin pattern was found for mouse Purkinje cells displaying HNK immunoreactivity (Hawkes, 1992; Eisenman and Hawkes, 1993) and for the constitutive expression of the 25-kDa heat-shock protein Hsp25 (Armstrong and Hawkes, 2000; Armstrong et al., 2001). Hsp25 is not expressed by Purkinje cells of rat cerebellum (Plumier et al., 1997; Armstrong et al., 2001). Bergmann glial fibers, immunoreactive for an antibody against FAL (3-fucosyl-N-acetyl-lactasamine), are arranged in a zonal pattern in adult mice. This pattern also appears to be complementary to the distribution of zebrin-immunoreactive Purkinje cells (Bartsch and Mai, 1991; Marani and Mai, 1992). Zebrin-positive and -negative Purkinje cells display a selective vulnerability to different noxes: in the Nervous mutation in mice the zebrin-negative Purkinje cells are most resistant (Wassef et al., 1987; Edwards et al., 1994), however, in a transgenic mouse model for Niemann– Pick’s disease the zebrin-negative Purkinje cells are the first to disappear (Sarna et al., 2001). A transient, zebrin-like pattern in the expression of L7 in mouse Purkinje cells was shown to be under genetic control (Oberdick et al., 1993; Oberdick, 1994). The zonal distributions of other Purkinje cell markers mentioned in the literature are not necessarily similar to the zebrin pattern (Ingram et al., 1985; Chan-Palay et al., 1981, 1982a, 1982b; Nilaver et al., 1982; Nunzi et al., 1999; Fusco et al., 2001).

FIGURE 17 The reconstruction of parasagittal bands of mabQ113+ (zebrin I-immunoreactive) Purkinje cells in the adult rat cerebellar cortex as seen anteriorly (a) and posteriorly (b). The band pattern is based upon the serial reconstruction of nine complete and five partial cerebellums from sections cut in the horizontal plane and four complete reconstructions from sections cut coronally. Bands P1+ through P7+ are labeled. From Hawkes and Leclerc (1987).

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The zebrin pattern, therefore, is the common denominator in the distribution of a number of different, seemingly unrelated substances and properties of Purkinje cells and their Bergmann glial covering. A unifying theory, explaining this distribution is still lacking, and no information is available on the possible differences in electrophysiological parameters of zebrin-positive and negative Purkinje cells. The nomenclature used for the bands of zebrinpositive and -negative Purkinje cells was introduced by Hawkes and Leclerc (1987). It is based on the staining of Purkinje cell dendrites in the molecular layer. The axons of zebrin-positive Purkinje cells can be traced into the white matter, where they appear more or less as distinct bundles. Hawkes and Leclerc (1987) grouped the zebrin I-positive Purkinje cells in a midline band (P1+) and seven, symmetrically disposed, parasagittal bands (P2+–P8+, Figs. 17 and 18). The midline band consists of the fused median zebrin I-immunoreactive Purkinje cells of both sides: the axons of these cells collect in distinct bundles on both sides of the midline. The intercalated zebrin-negative areas were indicated with the same number as the next medial zebrin Ipositive band (i.e., P1− is located between P1+ and P2+). The diagram of Hawkes and Leclerc (1987) (Fig. 17) suggests that all zebrin-positive bands, with the exception of P8+, continue uninterruptedly from the posterior lobe into the anterior lobe. In the anterior lobe six zebrin-positive bands, numbered P1+ to P6+ were distinguished. With the exception of the P3+ band, the zebrin-positive bands are distinct and clearly delineated. Staining of Purkinje cells in P3+ is less complete and its borders are fuzzy. The P1+ and P2+ bands increase in width in the dorsal part of the anterior lobe and in dorsal lobule V one or two satellite bands are usually present between them. Bands P3+, P4+, and P5+ cannot be precisely identified in the ventral lobules of the anterior lobe (lobules 1–3), while the lateral P6+ band is often divided into a narrow medial portion and a wide lateral portion. The pattern of zebrin banding in the lobulus simplex (the hemispheral expansion of lobule 6a) is continuous with and very similar to that of the anterior lobe. Apart from P3+, several narrow satellite bands are present in the region between P2+ and P4+. Bands P4+ and P5+ can be followed over the surface of the two sublobules of the lobulus simplex and continue on the rostral surface of crus I of the ansiform lobule. In the ansiform lobule (the hemispheral region lateral to lobules 6b and 6c) the bands coalesce into a uniform, zebrin-positive area. The band pattern reappears in crus II where the two narrow P4b+ and P5a+ bands and the more lateral and wider P5+, P6+, and P7+ bands can be distinguished. In the paramedian lobule the pattern shifts

laterally relative to crus II, and the P6+ and 7+ bands are often fused. Bands P4b+ and P5a+ generally fuse in the ventral part of the paramedian lobule. The pyramis (lobule 8) is characterized by the distinct zebrin-positive bands P1+ to P4+. Its hemisphere, the copula pyramidis, contains a zebrin-positive patch on the apex of the lobule. Laterally, the P5+/P7+ bands fuse into a zebrin-positive area, which is shifted further laterally with respect to the corresponding bands in the paramedian lobule. Laterally this zebrin-positive area continues into the paraflocculus and the flocculus. Wide, zebrin-positive separated by zebrin-negative slits are present in lobule 9. Most Purkinje cells of lobule 10 are zebrin-positive, although zonally distributed regions with higher and lower immunoreactivity can be recognized in the bottom of the posterolateral fissure and in lobule 10.

Correlations of the Corticonuclear and Olivocerebellar Projections with the Zebrin Pattern Gravel et al. (1987) and Wassef et al. (1992a) demonstrated that the position of antegradely labeled longitudinal strips of climbing fibers correlates with the zebrin pattern, but were unable to show a precise correspondance of specific zebrin-positive or -negative zones with the olivocerebellar projection of the individual subnuclei of the olive. For some regions of the cerebellum this object was achieved in our studies using antegrade axonal tracing from small injection sites restricted to individual olivary subnuclei, mapping of retrogradely labeled Purkinje cells from injections confined to particular cerebellar or vestibular nuclei, and/or the tracing of climbing fiber collaterals from injections of CTb of electrophysiologically and anatomically identified single zones in the posterior lobe of rat cerebellum (Fig. 19) (Voogd et al., 1993, 1996a; Voogd and Ruigrok, 1997; Voogd et al., 2003). The DAO, intermediate regions of the MAO, and and the DM were found to project to zebrin-negative territory. Rostral MAO with the DMCC, the PO with the VLO, and the DC and the group beta innervated zebrin-positive Purkinje cells. The projections from the caudal MAO include both zebrin-positive and zebrin-negative regions, but could not be studied in sufficient detail. The situation in the hemisphere of the rat cerebellum is fairly clear. The C2 zone, innervated by the rostral MAO, occupies the zebrin-positive P4+ band in the anterior lobe and the lobulus simplex and the P5+ band in the crus II, the paramedian lobule, and the copula pyramidis. The C1 and C3 zones, innervated by the rostral pole (ventral fold) of the DAO, correspond to the zebrin-negative bands, located at either side of

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FIGURE 18 Reconstruction of the localization of zebrin-immunoreactive Purkinje cell zones in the cerebellum of the rat. Numbers indicate the zebrin-positive Purkinje cell zones P1–P7 of Hawkes and Leclerc (1987). Abbreviations: 1–7. Zebrin-positive bands 1–7; COP, copula pyramidis; CrI and CrII, crus I and II of the ansiform lobule; PMD, paramedian lobule; SI, simple lobule; I–X, lobules I–X.

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FIGURE 19 Diagram comparing the pattern of zebrin-positive and-negative bands in the anterior cerebellum (anterior lobe and lobulus simplex) and the posterior cerebellum (lobules 7 and 8, crus II, paramedian lobule, and copula) with the olivocerebellar and corticonuclear projection zones in the rat. The zebrin-postive bands are numbered and hatched. For each projection zone the source of the climbing fiber projection in the inferior olive (stippled) and the cerebellar target nucleus is indicated. The topographical relations between connections and the P1+/P4a+ bands in the posterior cerebellum have not yet been verified. The projections of the C1–C3, D1, and D2 zones to single cerebellar target nuclei and the relative position of the D0 zone differ from those in the diagram of Buisseret-Delmas and Angaut (1993; Fig. 9). The A2 zone is not indicated in the anterior cerebellum because its relation to the zebrin pattern is incompletely known. Data are from Voogd et al. (1993; in preparation). Abbreviations: 1–7, zebrin-positive zones P1+ / P7+; A–D2, corticonuclear and olivocerebellar projection zones A–D2; Dd, dorsal fold of the dorsal accessory olive; Dh, dorsolateral hump; DM, dorsomedial group of the ventral lamina of the PO; DP, dorsolateral protuberance of the medial cerebellar nucleus; Dv, ventral fold of the dorsal accessory olive; IA, anterior interposed nuxleus; IC, interstitial cell groups; IP, posterior interposed nucleus; Lc, caudal part of the lateral cerebellar nucleus; Lr, rostral part of the lateral cerebellar nucleus; LV, lateral vestibular nucleus; Mc, caudal medial accessory olive; Mi, intermediate medial accessory olive; Mm, middle subnucleus of medial cerebellar nucleus; Mr, rostral medial accessory olive; Pd, dorsal lamina of the principal olive; Pv, ventral lamina of the principal olive.

P4+ in the anterior lobe and the lobulus simplex and of P5+ in the posterior lobe. The B zone, which receives its climbing fibers from the caudal pole (dorsal fold) of the DAO, similarly, occupies the medial part of the zebrin-negative P2− band in the anterior cerebellum. Climbing fibers from intermediate levels of the MAO were found to branch to zebrin-negative strips, medially to P4+ and laterally to P2+, corresponding to the X and CX zones, respectively. In the lateral hemisphere, the PO-innervated D1 and D2 zones correspond with the zebrin-positive P5+ and P6+ bands in the anterior lobe and with P6+ and P7+ in the posterior lobe, respectively. The DM-innervated D0 zone is located in the zebrin-negative strip, located between P5+ and P6+ in the anterior cerebellum and between P6+ and P7+ in the posterior lobe. This D1–D0–D2 sequence is at variance with the D0–D1–D2 sequence, as originally reported by Buisseret-Delmas and Angaut (1989a) (compare Figs. 9 and 19). The A2 zone corresponds with a medial region in the crus II and the paramedian lobule, containing the zebrin-positive P4b+ and P5a+ bands and with a similar region in the medial lobulus simplex. The uniform, zebrin-positive appearance of the crus I and the paraflocculus is in accordance with the lack of projections of the DAO and the DM to zebrin-negative territory in these lobules. Similarly, the fusion of the

P5+, P6+, and P7+ bands in the paramedian lobule and the copula pyramidis, corresponds with the absence of zebrin-negative C3 and D 0 zones in these lobules. The question as to whether the zebrin-negative P4b− and the zebrin-postive P4b+ and P5a+ bands of the A2 zone are innervated by different groups of olivary neurons in the caudal MAO cannot, as yet, be answered. The shift in the numbering of zebrin bands, corresponding to particular climbing fiber zones, between the anterior and posterior cerebellum (compare Figs. 17, 18, and 19), can be explained by the absence of zebrin-negative DAO and DM-innervated zones in crus I. The uniform zebrinpositive appearance of crus I prevented Hawkes and Leclerc (1987) from establishing their continuity. The correspondance of the climbing fiber zones with the zebrin-postive and negative bands in lobules 9 and 10 is illustrated in Fig. 13. It should be noted that the zebrin-positive bands P2+ and P3+ are bisected by branching projections from caudal group beta to P1+ and medial P2+, from rostral group beta to lateral P2+ and medial P3+, and from the DMCC to lateral P3+. Moreover, the DM and the caudal MAO project to the narrow, zebrin-negative strips P2− and P3−, respectively (Voogd et al., 1996a; Voogd and Ruigrok, 1997). Overall, the zebrin pattern of the rat cerebellar hemisphere can be considered, therefore, to be the result of zebrin-negative zones innervated by DAO, DM, and

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intermediate MAO, interdigitating with zebrin-positive zones innervated by rostral MAO and PO and the vestibular and optokinetic subnuclei of the olive (group beta, DMCC, DC, VLO). In crus I, the flocculus and the paraflocculus, where zebrin-negative zones are not represented, the Purkinje cells are uniformly zebrinpositive. It remains to be determined whether this distinction between zebrin-negative and -positive zones is linked to the fact that subnuclei of the olive innervating the former all receive a somatotopically organized input from the periphery (Molinari et al., 1996; Yatim et al., 1996), whereas olivary subnuclei innervating the latter are dominated by afferent connections from higher levels of the brain stem.

AFFERENT MOSSY FIBER SYSTEMS The origin and the termination of the main afferent mossy fiber systems of the cerebellum are reviewed by Ruigrok (this volume, Chapter 8). Here we address the question of the topographic distribution of some of these tracts. Mossy fiber systems enter the cerebellum through the inferior and middle cerebellar peduncles. The ventral spinocerebellar tract, which courses rostral to the entrance of the trigeminal nerve, reaches the cerebellum dorsal to the superior cerebellar peduncle, where it rejoins the fibers of the inferior cerebellar peduncle. Within the cerebellum they remain separated from the cerebellar nuclei by a layer of olivocerebellar fibers. Mossy fibers cross in a separate portion of the cerebellar commissure, rostral and dorsal to the fibers of the uncinate tract (Fig. 6). The common features in the distribution of mossy fibers are well-illustrated in the recent paper of Wu et al. (1999) on the course and termination of individual mossy fibers from the lateral reticular nucleus of the rat (Fig. 20). The parent fibers enter the cerebellum laterally and sweep medially, as semicircular fibers, located rostral and dorsal to the cerebellar nuclei. They usually cross in the rostral and dorsal portion of the cerebellar commissure and, therefore, distribute bilaterally. The parent fibers emit collaterals at certain preferential positions, which enter the white matter of the folia and terminate as ill-defined, longitudinal stripes of mossy fiber rosettes in the granular layer (see also Scheibel, 1977). Some mossy fiber systems also send collaterals to the cerebellar nuclei. The primary orientation of the “parent” mossy fibers, therefore, is a transverse one, contrasting with the strictly longitudinal (parasagittal) orientation of the olivocerebellar and corticonuclear projections. The termination of a mossy fiber system usually is restricted

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to a particular combination of lobules or folia. This type of distribution of mossy fiber systems has been used to subdivide the cerebellum into functional regions, such as its spinal, vestibular, visual, and cerebropontine territories. Within these territories mossy fibers terminate as ill-defined longitudinal stripes of mossy fiber rosettes, separated by empty areas, as shown for the mossy fiber system of the lateral reticular nucleus (Fig. 20) and several other mossy fiber pathways, including the pontocerebellar projection. The termination in multiple longitudinal aggregates raises important questions on (1) the spatial organization of overlap and segregation of different mossy fiber systems, (2) the paradox that possible effects of such a termination in segregated longitudinal aggregates of mossy fiber terminals would be erased by the transverse orientation in the next link of the pathway consisting of the axons of the granule cells, the parallel fibers, and (3) the relation of the longitudinal aggregates of mossy fiber terminals to the compartmental organization of the corticonuclear and olivocerebellar projections. The question of the overlap and the segregation of mossy fiber systems is considered in the next sections on specific mossy fiber pathways. The effect of the transverse orientation of the parallel fibers on the patterns in the termination of mossy fibers has been mitigated by the observation that granule cells preferentially influence Purkinje cells located immediately superficial to them, by multiple syanapses on their ascending branches (Llinas, 1982; Bower and Woolston, 1983). Any pattern in the termination of the mossy fibers, therefore, will be reproduced at the level of the Purkinje cells. The transverse branches of the parallel fibers extend over large distances; over the entire width of the lobules of the vermis and the hemisphere in the rat. As a consequence, mossy fibers exert a strong influence on certain arrays of Purkinje cells and a weaker influence on the Purkinje cells in between. A correspondance in the distribution of climbing fiber and mossy fiber evoked potentials has been shown by Eccles et al. (1968a, 1968b, 1971, 1972) for ill-defined patches of the cat cerebellar cortex, which received both mossy and climbing fiber input from the same receptive area on the limbs. Ekerot and Larson (1973, 1980) found a similar spatial correspondance for the effects of stimulation of different limb nerves mediated by climbing fibers terminating in the C1, C2, and C3 zones of the anterior lobe of the cat and the slightly wider zones of termination of the exteroceptive component of the cuneocerebellar tract. This correspondance was extended to the level of the microzones by Garwicz et al. (1998). Recent studies of tactile projections to crus II of the rat cerebellum (Brown and Bower, 2001) revealed a similarity in the peripheral receptive field organization

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FIGURE 20 Course and termination of single mossy fibers in the cerebellum of the rat. (A) Frontal view of a completely reconstructed, biotinylated dextran amine-labeled, single mossy fiber from the lateral reticular nucleus. The fiber entered the cerebellum through the ipsilateral restiform body, projected bilaterally to the cortex and the cerebellar nuclei, and formed a multiple, longitudinal zonal projection pattern by its cortical arborescent collaterals. Scale bars, 0.5 mm. (B) Mapping of mossy fiber rosettes in the cerebellar cortex after biotinylated dextran amine injection in the left lateral reticular nucleus, showing bilateral projction with ipsilateral predominance and zone-like terminal distribution in the cerebellar cortex (a–f, rostral to caudal). Scale bar, 1 mm. From Wu et al. (1999). Abbreviations: COPab, copula pyramidis, lobules a and b; D, dorsal; DN, lateral cerebellar nucleus; FN, fastigial nucleus; icp, inferior cerebellar peduncle; IP, posterior interposed nucleus; LRNm, magnocellular part of the lateral reticular nucleus; Lt, left; LVN, lateral vestibular nucleus; PM, paramedian lobule; Rt, right; Sima, Simb, lobulus simplex, lobules a and b; V, ventral.

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of Purkinje cell complex spikes (generated by activity in climbing fibers) and the immediately subjacent granule cells (which receive their principal input from mossy fibers). These observations are in accordance with our anatomical observations on the distribution of mossy fiber collaterals, namely, that collaterals of mossy fibers terminating in a particular locus always terminate subjacent to the strips of climbing fiber collaterals arising from olivocerebellar fibers terminating in the same cortical locus (Voogd et al., 2003). When the similarity in receptive field organization of the Purkinje cells and the subjacent granule cells applies to the entire cerebellum, this may have important consequences for our understanding of the organization of the Purkinje cells in longitudinal zones and microzones. Questions on overlap and segregation in the termination of different mossy fiber systems in a particular zone, on somatotopical localization and the relation of Welker’s (1987) maps of a discontinuous (fractured) localization to the multiple representations of the body in the microzones, and on the nature of the mossy fiber input of zones and regions that lack a somatopical organization now have become relevant issues. One feature of the termination of mossy fiber systems which has not been discussed is their termination in the cerebellar nuclei. It has been attempted to investigate this type of termination using injections of retrograde tracers into the cerebellar nuclei. However, the proximity of the afferent tracts to the central nuclei may easily lead to false positive results. The experiments of Eller and Chan-Palay (1976) with injections of HRP into the lateral cerebellar nucleus of the rat showed a multitude of extracerebellar afferents to this nucleus. Of these sources, the pontine nuclei and the nucleus reticularis tegmenti pontis (Gerrits and Voogd, 1987; Shinoda et al., 1995; Mihailoff, 1993), the lateral reticular nucleus (Ruigrok et al., 1995; Wu et al., 1999), and the raphe nucleus (Chan-Palay, 1977) were confirmed with anterograde axonal tracing methods. The question of the projection of the basal and reticular pontine nuclei to the cerebellar nuclei recently was studied by Mihailoff (1993) with antegrade axonal transport of the more sensitive tracer Phaseolus vulgaris leucoagglutinin. Both the pontine nuclei and the tegmental pontine nuclei were found to send fibers to the cerebellar nuclei, but the density of the projection was greater for the reticular nucleus. Pontine nuclei were found to project to dorsal regions of the fastigial (to the dorsolateral protuberance), interposed, and lateral nuclei, the nucleus reticularis tegmenti pontis to ventral and caudal parts of the nuclei. The rostral and medial parts of the anterior interposed nucleus are spared. The lateral reticular nucleus, similarly, projects to all cerebellar nuclei, but the projection focuses on more or

less complementary regions of the nuclei, including the ventral and ventrolateral fastigial nucleus, the interstitial cell groups, medial parts of the interposed nuclei, and the lateral vestibular nucleus (Ruigrok et al., 1995; Wu et al., 1999) (Fig. 20). Projections to the same regions of the fastigial and interposed nuclei were traced from different levels of the spinal cord (Matsushita, 1999a, 1999b; Matsushita and Yaginuma, 1995; Matsushita and Xiong, 2001; Matsushita and Gao, 1997). All the collateral projections of the reticular nuclei and the cord are bilateral, with a predominance of the lipsilateral side. The pertinent negative results of Eller and ChanPalay (1976) with respect to possible collateral projections to the nuclei of the dorsal column nuclei were confirmed in the anterograde transport study of Gerrits et al. (1985) in the cat. Similar data are not available for the other precerebellar nuclei of the rat. The tentative conclusion is that not all mossy fiber systems distribute collaterals to the central nuclei. The reticular nuclei, including the reticulotegmental and lateral reticular nuclei, the spinal cord, and the monoaminergic systems seem to be the main sources of the extracerebellar nuclear afferents. The rubrocerebellar pathway (Brodal and Gogstad, 1954), a collateral pathway of the rubrospinal tract, was investigated with double-labeling techniques and shown to terminate in the anterior interposed nucleus in the rat (Huisman et al., 1983). It is the only pathway known to terminate in the cerebellar nuclei and not in the cortex. In our experiments on the collateralization of mossy fibers from small injection sites in the cerebellar cortex of the rat (Voogd et al., 2003) we often observed multple, bilateral patches of thin varicose axons in the cerebellar nuclei, apart from the coarse terminals of the Purkinje cell axons. We considered these fine plexuses as collateral projections of the mossy fibers. Panto et al. (2001), similarly, observed multiple foci of termination in the cerebellar nuclei after injections of the cortex. These authors did not mention the bilaterality of these projections and considered all of them as terminals from Purkinje cell axons.

TERMINATIONS OF MOSSY FIBER SYSTEMS IN DIFFERENT REGIONS OF THE CEREBELLUM Projections from the Spinal Cord, the Dorsal Column Nuclei, the Trigeminal Nuclei, and the Lateral Reticular Nucleus Regions dominated by spinocerebellar, cuneocerebellar, and trigeminocerebellar input correspond to the

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anterior lobe and the lobulus simplex, lobule 8, the crus II, the paramedian lobule, and the copula pyramidis. As pointed out in the previous section, this part of the cerebellum is characterized by the presence of X, B, C1, C3, and D0 zones, which receive somatosensory climbing fiber input from the intermediate MAO, the DAO, and the dorsomedial subnucleus. The output of these zones is directed at the spinal cord (X zone, through the interstitial cells group, B zone through the lateral vestibular nucleus), the lateral reticular formation and the trigeminal nuclei (D0 zone through the dorsolateral hump and the uncrossed descending limb of the brachium conjunctivum), and the red nucleus (C1 and C3 zones, through the anterior interposed nucleus). The lateral reticular nucleus, which projects to the same lobules, relays information from the spinal cord and the red nucleus to the cerebellum (Ruigrok, this volume, Chapter 8). Spinocerebellar tracts in the rat were traced by Anderson (1943) with the Marchi method and illustrated by Voogd (1967). These tracts are distributed bilaterally to the anterior lobe, with a preference for its ventral part, to the bottom of the primary fissure, the pyramis, and the dorsal part of the uvula and to the ventral part of the paramedian lobule and the copula pyramidis. Fibers of

the ventral spinocerebellar tract terminate more medially and do not reach lobule 1, the uvula, or the paramedian lobule. The termination of the individual spinocerebellar tracts has not been studied in the rat. The origin of the spinocerebellar fibers from the cord in the rat was studied by Snyder et al. (1978), Matsushita and Hosoya (1979), and Beretta et al. (1991a). Branching of spino and cuneocerebellar fibers to the anterior lobe and the posterior lobes of the cerebellum of the rat was demonstrated by Beretta et al. (1991a, 1991b). A zonal pattern has been demonstrated in the termination of the spinocerebellar, cuneocerebellar, and lateral reticular–cerebellar tracts. The mediolateral periodicity in the termination of the spinocerebellar tracts is already present at birth (Arsenio-Nunes and Sotelo, 1985). The semiquantitative plotting study of Tolbert et al. (1993) used a semiquantitative method to plot the spinocerebellar terminals. They showed that spinocerebellar projections from the thoracic and lumbar cord are often restricted to certain transverse bands often centered at the bottom of the interlobular fissures. These bands are fractured in multiple longitudinal aggregates of mossy fiber terminals (Fig. 21). These longitudinal aggregates, therefore, are often discontinuous, with interruptions usually located at the apex of the lobules.

FIGURE 21 Surface reconstruction of the lobules I–V of the anterior lobe of the rat, showing the position of spinocerebellar terminals (SpCb) from a bilateral injection of the lumbar and thoracic cords (grey figures) and an injection of the right cuneate nucleus (ICu/ECu; black figures). The anterior–posterior (AP) dimension of the cerebellar surface has been reduced, relative to its width (ML). In lobule V, three concentrations of spinocerebellar terminals are indicated (Lat-SpCb, Me-SpCb, and Mi-SpCb), which altermate with a patches of cuneocerebellar terminals (CuCb). From Alisky and Tolbert (1997).

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The zonation in the spinocerebellar projection also was shown by Gravel and Hawkes (1990) for the projection of the lumbar and thoracic cord to the lobules of the anterior lobe of the rat. Their mossy fiber terminal aggregates usually were more discrete than those illustrated in the paper of Tolbert et al. (1993), and they were able to correlate the positions of these aggregates with the zebrin bands in the overlying molecular layer (Fig. 22). In subsequent papers Tolbert and Gutting (1997), Alisky and Tolbert (1997), and Ji and Hawkes (1984) showed that the multiple zonal termination of the cuneocerebellar and the spinocerebellar fibers from the lumbar and thoracic levels of the cord remains segregated in the cerebellar cortex. The cuneocerebellar fibers terminate in a lateral strip in the anterior vermis and in patches in the hemisphere of lobule 5 (Fig. 21). Spinocerebellar fibers from the cervical cord overlap with the cuneocerebellar projection (Matsushita et al., 1991; Ji and Hawkes, 1994). The relationship between the zebrin bands and the projections of the external cuneate nucleus and the spinocerebellar tracts in lobules 2 and 3 is illustrated in Fig. 22, from the paper of Ji and Hawkes (1993). In the case of the rat lateral reticular nucleus (Fig.20) (Wu et al., 1999) the striped termination pattern is bilaterally symmetrical and occupies the dorsal part of the anterior lobe (lobules 4 and 5), the lobulus simplex (6), and the pyramis with the copula pyramidis (8). The topographical relations of this pattern with the

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spinocerebellar and the cuneocerebellar projections and the zebrin pattern have not been studied. Trigeminocerebellar mossy fiber projections have been studied in the rat by Shambes et al. (1978a, 1978b), Woolston et al. (1981), Huerta et al. (1983), Akaike (1989), Arends et al. (1991), and Phelan and Falls (1991). Systematic studies of this system in the rat, using antegrade axonal transport techniques, have not been published thus far. However, the electrophysiological studies of Welker and Shambes on the spatial organization of somatosensory projections to the granule layer of the rat cerebellar cortex (reviewed by Welker, 1987) are mostly confined to receptive fields on the face and, therefore, concern the trigeminocerebellar mossy fiber projection. These studies were restricted to the posterior lobe of the cerebellum. Receptive fields were mostly cutaneous, varied widely in size, and had differentially enlarged representations on specific folia for certain body parts. Somatosensory short-latency projections to the granular layer occupy all hemispheric folia (except for the copula) as well as dorsal lobule 9 in the posterior vermis. They are mainly ipsilateral, with a few bilateral projections. They are shaped as irregular patches, which vary in size between 0.3 and 0.3 mm3. Arrays of juxtaposed patches form mosaics. Receptive fields often have multiple representations on different folia. The somatotopical localization in the receptive mosaic is “fractured”; i.e., the original continuity of the receptive fields has become lost. Multiple patches

FIGURE 22 Lobules II and III of the rat anterior vermis: a schematic representation of the relationships between Zebrin-positive and -negative bands of Purkinje cells and the mossy fiber projections from the external cuneate nucleus, and lumabr, thoracic and cervical levels of the cord. There is one zebrin (P1+) compartment at the midline and two others (P2+, P3+) positioned laterally. Terminals of the cuneocerebellar tract (Cu1) are clustered under P1+ and within P2- (Cu2) and P2- (Cu3). Only the medial edge of Cu3 clearly corresponds to a zebrin+/− boundary. The lumbar and thoracic spinocerebellar terminals (Gravel and Hawkes, 1990) distribute complementary to the cuneocerebellar fields. The lumar L2 fields fade toward their medial edge. The mossy fiber terminals from the cervical cord are shown as spread uniformly across the vermis. From Ji and Hawkes (1994).

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sharing the same receptive field are innervated by branches from the same trigeminocerebellar mossy fibers. Projections from the somatosensory cortex (Bower et al., 1981) and the superior colliculus (Kassel, 1980), similarly, are organized in patches which exhibit homotopy with the patches activated from the periphery. These projections are confined to the hemisphere and, presumably, are relayed by the pontine nuclei. Unfortunately, this fractured somatotopical pattern has not been extended to the anterior regions of the rat cerebellum and has not been correlated with the longitudinal pattern in the olivocerebellar projection. A partial correspondance of the tactile map with the zebrin pattern has been reported by Hallem et al. (1999) for the crus II of the rat.

Pontocerebellar Mossy Fibers The projection from the basal and reticular tegmental nuclei of the pons was charted by Burne et al. (1978b), Azizi et al. (1981), and Mihailoff et al. (1981) using retrograde transport of HRP in the rat. The retrogradely labeled cells formed multiple foci or discontinuous lamellae, in medial, lateral, dorsal, and central regions of the contralateral, and in some cases the ipsilateral, pontine nuclei and in the nucleus reticularis tegmenti pontis. Most neurons projecting to vermal lobules 6a, 8, and 9 were located in caudal regions of the pons, with most neurons projecting to 6a located within the reticular nucleus, cells projecting to lobule 8 in medial and lateral patches in the pontine nuclei (Eisenman, 1981b), and cells projecting to lobule 9 in dorsal and medial patches in the caudal pons. Most of the neurons projecting to the copula, the paramedian lobule, and the crus II are found in central and medial patches in the caudal pons and in the reticular nucleus. Rostral regions of the pons project to the crus I. The pontocerebellar projections as they appear from these studies generally are not sharply focused. The anterior lobe of the rat never was included in these or similar studies. The tecto– and pretecto–pontine cerebellar pathways were traced by Burne et al. (1981). They were found to influence extensive regions in the hemisphere and the posterior vermis. No equivalent appears to exist for the focused olivocerebellar projection to the tectorecipient zone in lobule 7, reported by Hess (1982) and Akaike (1992). A localized projection was described for the pathways from the visual cortex, via medial and lateral patches in the rostral pons, to the contralateral paraflocculus (Burne et al., 1978a; Eisenman, 1980). A similar pathway seems to exist for the projection of the auditory cortex to the paraflocculus. Auditory pontocerebellar projections to the caudal vermis are relayed by the inferior colliculus (Azizi et al., 1985).

Serapide et al. (2001) recently reported on an often bilateral pattern of longitudinal stripes separated by “interstripes” in the projection of the pontine nuclei to all lobules of the cerebellum, with the exception of the anterior lobe. Stripes were present after small injections of antegrade tracers in different parts of the pontine nuclei; larger injections resulted in diffuse labeling. The stripes and the interstripes, therefore, are innervated by different, but adjacent, parts of the pontine nuclei. The precise, topical relations between the pontine nuclei and the stripes and the interstripes in a particular lobule cannot be established from their data. The topographical relations of these projections from the pontine nuclei to the zonal patterns in the termination of other mossy fiber systems and to the zebrin pattern presently are unknown.

Vestibular Projections: Mossy Fibers Terminating in Flocculus, Nodulus, and Adjacent Areas Vestibular root fibers enter the cerebellum from the superior vestibular nucleus, as part of the ascending branch of the vestibular root (Lorente de No, 1933; 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 recent 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 rat by Rubertone et al. (1995). Both primary and secondary vestibulocerebellar fibers terminate in lobule 10 and ventral lobule 9, in lobules 1 and 2, 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 vestibulocerebellar projection to the hemisphere is restricted to the flocculus and the adjacent cortex of the ventral paraflocculus. A primary vestibulocerebellar projection to the flocculus is absent in the rabbit (Gerrits et al., 1989) and probably in most other mammals including the rat. 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. 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. Widespread projections of the prepositus hypoglossal

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nucleus and neighboring perihypoglossal nuclei terminate bilaterally with an ipsilateral predominance in the vermis, in the flocculus and the paraflocculus, and in the cerebellar nuclei (see McCrea and Baker, 1985, and Roste, 1989, for particulars). Some cerebellumprojecting neurons of the nucleus prepositus hypoglossi and the caudal parts of the vestibular nuclear complex contain acetylcholine and/or CRF. Choline acetyltransferase (ChAT)-immunoreactive mossy fibers recently were charted in lobules 9 and 10 of the caudal vermis and the flocculus in the rat and other mammalian species by Barmack et al. (1992a, 1992b). They were most numerous in the caudal vermis, the rosettes were large in lobule 10 and smaller in ventral lobule 10 and lobules 1–3 of the anterior lobe. The cholinergic mossy fiber innervation of the flocculus was restricted to the ventral folium and the ventral half of the dorsal folium of this lobule. A particularly dense plexus of thin, beaded fibers, which may detach from the mossy fibers, was present in the flocculus. The origin of these cholinergic mossy fibers was traced from the caudal medial vestibular nucleus, the vestibular efferent, and the nucleus prepositus hypoglossi, with double-labeling of retrogradely transported HRP from injections in the caudal vermis and the flocculus and ChAT immunohistochemistry. Single- and doublelabeled ChAT-immunoreactive neurons were absent from the superior nucleus. Single HRP-labeled cells were present in all vestibular nuclei with the exception of the lateral vestibular nucleus. In other brain stem nuclei only a few neurons in the lateral reticular nucleus were double-labeled for HRP and ChAT following injections in lobules 9 and 10 (Barmack et al., 1992b). The origin of mossy fiber projections to the nodulus, the uvula, and the flocculus of the cat were quantified by Akaogi et al. (1994). Major projections to the nodulus (>95%) and the ventral uvula (>70%) were derived from the vestibular nuclei. The dorsal uvula receives its main projection (>80%) from the pons, and the flocculus receives projections from the reticular formation and the raphe nuclei (>50%) and the nucleus prepositus hypoglossi (20%). A similar situation may be inferred from the studies of Blanks et al. (1983) and Osanai et al. (1999) for the flocculus of the rat.

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10 Periaqueductal Gray KEVIN A. KEAY and RICHARD BANDLER Department of Anatomy and Histology, and Pain Management Research Institute Royal North Shore Hospital, The University of Sydney Sydney, New South Wales, Australia

Darkschewitz, and the interstitial nucleus of Cajal), the dorsal raphe nucleus, the mesencephalic trigeminal nucleus, and the dorsal tegmental nuclei. Cytoarchitectural studies of the PAG (Beitz, 1985; Beitz and Shepard, 1985; Gioia et al., 1985; Hamilton, 1973a, 1973b; Liu and Hamilton, 1976, 1980; Meller and Dennis, 1990a, 1990b, 1993) have revealed that it contains predominantly small- to medium-sized (5–20 μm in diameter) fusiform-, triangular-, and stellate-shaped neurons, whose soma and axons are oriented usually in a rostral–caudal direction. It has not proved possible to subdivide the PAG into discrete subnuclei using cytoarchitectonic criteria such as soma size, pigmentation, or dendritic morphology and orientation. However, many investigators have commented on an increase in the size of neurons and their packing density, the greater the distance from the aqueduct (e.g., density: 10,780 mm3 (near aqueduct) to 21,950 mm3 (dorsolateral to aqueduct) (Beitz, 1985; Beitz and Shepard, 1985)). This has led to suggestions that the PAG is divisible radially into: (i) a relatively cell-sparse juxtaaqueductal zone, next to the aqueduct; (ii) an adjacent, inner or medial zone, containing a moderate density of small neurons; and (iii) an outer or lateral zone, with a higher density of small- to medium-sized neurons (Beitz, 1985; Gioia et al., 1985; Hamilton, 1973a, 1973b; Onstott et al., 1993; Ramon y Cajal, 1911). The conceptualization of the PAG as radially organized has yet to prove fruitful in the reinterpretation of existing anatomical and functional data or in the generation of new experimental approaches to the study of the PAG (although see (Ruiz-Torner et al., 2001; Vanderhorst et al., 2000)).

Early Cytoarchitectural Studies The periaqueductal gray (PAG), also known as the midbrain central gray, constitutes a cell-dense region, bordered laterally by the descending tectospinal fibers, which surrounds the midbrain aqueduct (cerebral aqueduct/aqueduct of Sylvius). In the rostral midbrain (at the level of posterior commissure) as the third ventricle narrows to become the midbrain aqueduct, the PAG forms an elongated, oval-shaped collection of neurons in continuity with the periventricular gray matter of the hypothalamus (Fig. 1A). More caudally (at the level of the superior colliculus) as the aqueduct shortens dorsoventrally, the dorsal two-thirds of the PAG expands laterally and the PAG becomes an inverted pear shape structure (Figs. 1B and 1C). More caudally still (at a midcollicular level) as the mediolateral diameter of the aqueduct increases and the dorsal raphe nucleus appears, the ventral third of the PAG expands and the PAG takes on a rhomboid shape (Fig. 1D). Finally at the caudal end of the midbrain, as the aqueduct expands to become the fourth ventricle, the dorsal two-thirds of the PAG progressively narrows in its mediolateral extent (Fig. 1E), eventually disappearing. The ventral third of the PAG becomes continuous, however, with the pontine gray matter which constitutes the floor of the fourth ventricle. The common usage of the term PAG usually excludes a number of nuclei which, although technically within its boundaries, are structurally and functionally distinct from the PAG. These include ocular-related nuclei (i.e., oculomotor and trochlear nuclei, the Edinger–Westphal nucleus, the nucleus of

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A

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FIGURE 1 Serial coronal sections of the periaqueductal gray region spaced 1 mm apart showing the radial and columnar organization described in the text. (Sections are modified from the Stereotaxic Atlas of Paxinos and Watson, 1982.) Abbreviations used: III, oculomotor nucleus; EW, Edinger–Westphal nucleus; Dk, nucleus of Darkschewitz; dm, dorsomedial column of the PAG; dl, dorsolateral column of the PAG; lat, lateral column of the PAG; vl, ventrolateral column of the PAG; m, medial PAG; ja, juxtaaqueductal PAG; DR, dorsal raphe nucleus; Me5, mesencephalic nucleus of the trigeminal; LDTg, lateral dorsal tegmental nucleus; DTgP, dorsal tegmental nucleus.

Overall, the cytoarchitectonic organization of the PAG of the rabbit, cat, monkey, and human appears identical to that of the rat (Beitz, 1985; Beitz and Shepard, 1985; Hamilton, 1973a, 1973b; Liu and Hamilton, 1976, 1980; Mantyh, 1982; Meller and Dennis, 1990a, 1990b, 1993; Paxinos and Huang 1995; Paxinos et al., 2000; Ramon y Cajal, 1911).

PAG COLUMNAR ORGANIZATION Early Functional Studies Early functional studies reported that aversive or defensive reactions, hypertension, sexual behavior (lordosis), and analgesia were readily evoked by electrical stimulation within or immediately adjacent to the PAG (Depaulis et al., 1988; Fardin et al., 1984a, 1984b; Liebeskind et al., 1973; Morgan and Franklin, 1988; Morgan et al., 1989; Sakuma and Pfaff, 1979; Sandner et al., 1987; Schmitt and Karli, 1980; Yeung et al., 1977). Although, there were suggestions from these studies that defensive and sexual reactions were most easily evoked from the dorsal PAG, and analgesia was more readily evoked from the ventral PAG, the appreciation that PAG organization took the form of a series of rostrocaudally oriented, longitudinal neuronal columns emerged only later, from studies that utilized the technique of microinjection of excitatory amino acids (EAA). As summarized in Fig. 2, these studies revealed: (i) that EAA microinjections made within the dorsal PAG evoked active defensive reactions and (ii) that the particular

defensive strategy expressed reflected the rostrocaudal position of the injection cannula. Thus, stimulation, within the dorsal PAG, at rostral sites triggered a “confrontational defense reaction,” the animal facing and threatening the stimulus (e.g., experimenter, another animal), often with vocalization; whereas stimulation, within the dorsal PAG, at caudal sites triggered an escape or flight response, the animal turning and running away from, rather than confronting, the stimulus (Bandler and Carrive, 1988; Bandler et al., 1985, 1991; Bandler and Depaulis, 1988; Bandler and Shipley, 1994; Carrive et al., 1987, 1989; Depaulis et al., 1989, 1992; Depaulis and Vergnes, 1986; Morgan and Franklin, 1988; Morgan et al., 1998; Yardley and Hilton, 1987; Zhang et al., 1990). The different defense reactions were accompanied always by hypertension and tachycardia, although the two strategies, confrontational defense/ threat (rostrally) versus escape/flight (caudally), were each characterized by a distinct set of regional blood flow changes (see Fig. 2). In contrast to the active defensive strategies that characterized the effects of dorsal PAG excitation, microinjections of EAA within the ventrolateral part of the caudal PAG (vlPAG) evoked a passive coping reaction/disengagement from the environment, characterized by quiescence and immobility, decreased vigilance, hyporeactivity, hypotension, and bradycardia. It was found also that (i) that the passive coping reaction evoked by vlPAG stimulation was associated with an opioid-mediated analgesia, whereas (ii) the active defense reactions evoked by dorsal PAG stimulation were associated usually with a non-opioidmediated analgesia (Bandler et al., 1985, 1991; Depaulis

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active coping strategies evoked from the "dorsal" PAG confrontational defense / threat non-opioid mediated analgesia hypertension and tachycardia extracranial vasodilation hindlimb & renal vasoconstriction dm

escape / flight non-opioid mediated analgesia hypertension and tachycardia hindlimb vasodilation extracranial & renal vasoconstriction

dorsal dm dorsal dm dorsal

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quiescence and hyporeactivity opioid mediated analgesia hypotension and bradycardia

dm dorsal

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passive coping strategies evoked from the vlPAG FIGURE 2 Schematic illustration of the dorsal and ventrolateral neuronal columns within (from left to right) the rostral PAG, the intermediate PAG (two sections), and the caudal PAG. Injections of excitatory amino acids (EAA) within the dorsal (dark shading) vs ventrolateral (vlPAG) (light shading) columns evoke fundamentally opposite, active vs passive emotional coping strategies. EAA injections made within the rostral portions of the dorsal PAG evoke a confrontational defense/threat reaction, tachycardia, and hypertension (associated with decreased blood flow to limbs and viscera and increased blood flow to extracranial vascular beds). EAA injections made within the caudal portions of the dorsal PAG evoke escape/flight, tachycardia, and hypertension (associated with decreased blood flow to visceral and extracranial vascular beds and increased blood flow to limbs). In contrast, EAA injections made within the vlPAG evoke cessation of all spontaneous activity (quiescence), a decreased responsiveness to the environment (hyporeactivity), hypotension, and bradycardia. Non-opioid-mediated and opioid-mediated analgesia are evoked, respectively, from the dorsal and the vlPAG. (Modified from Fig. 1 of Keay and Bandler, 2001).

et al., 1992, 1994; Krieger and Graeff, 1985; Lovick, 1992, 1993, 1996; Morgan et al., 1998; Yaksh et al., 1976). To summarize, then, excitation of neurons within two longitudinally oriented columns of PAG neurons evokes fundamentally distinct, integrated patterns of somatic, autonomic, and antinociceptive adjustments. Overall, the dorsal PAG-evoked, active coping reactions correspond remarkably well to the natural strategies employed by an animal when threatened or attacked, whereas the ventrolateral PAG-evoked, passive coping reactions are strikingly similar to the reaction of an animal to serious traumatic injury (i.e., blood loss, deep pain) or chronic stress (e.g., repeated defeat in social encounters) (Bandler and Keay, 1996; Bandler et al., 2000a, 2000b; Keay et al., 2000; Lovick, 1993).

Neurochemical Studies Although the functional studies provided a basis to subdivide the PAG into dorsal and ventrolateral

longitudinal columns, neurochemically there are good grounds for further subdivision of the dorsal PAG. As illustrated in Fig. 3, within the dorsal PAG there lies a wedge-shaped, dorsolateral (dl) zone which stains intensely for specific neurochemicals, e.g., NADPH– diaphorase (Fig. 3), also nitric oxide synthase (NOS), cholecystokinin, acetylcholinesterase, and met– enkephalin (not illustrated), but not for other neurochemicals, e.g., glycine-2 transporter (Fig. 3) and cytochrome oxidase (not illustrated). Conversely, adjacent to the wedge-shaped dorsolateral zone lie dorsomedial (dm) and lateral (l) zones, which: (i) stain intensely for substances for which there is absence of staining in the dorsolateral zone, e.g., cytochrome oxidase and glycine 2 transporter; but (ii) stain weakly or not at all for those substances that intensely label the dorsolateral zone (e.g., NADPH–diaphorase). The restricted distributions of these neurochemical markers led to the suggestion (Bandler et al., 1991; Bandler and Keay, 1996; Bandler and Shipley, 1994) that the dorsal PAG was broadly

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dm dl dm dl l

dm dl l dm dl l

FIGURE 3 (Top) Photomicrographs of the PAG stained for the presence NADPH–diaphorase-containing neurons. Note the dense labeling in the dorsolateral PAG column. (Middle) Photomicrographs of the PAG stained for the presence of a glycine 2 transporter (GLY2-T). Note the labeling predominantly in the dorsomedial, lateral, and ventrolateral PAG columns and the absence of label in the dorsolateral PAG column. (Bottom) Schematic subdivision of the dorsal PAG into three columns: dm, dorsomedial; dl, dorsolateral; l, lateral (also see Fig. 1). Note that active emotional coping reactions are evoked readily by EAA microinjections made within the dlPAG or the lPAG (see Fig. 2). Active emotional coping reactions have been reported also following electrical or chemical stimulation of the dmPAG/deep layers of the superior colliculus (see for example, Krieger and Graeff, 1985).

divisible into three longitudinal columns, usually designated dorsomedial (dmPAG), dorsolateral (dlPAG), and lateral (lPAG). The utility of this particular columnar scheme has been supported by anatomical and functional–anatomical studies that are considered below. Other patterns of distinct PAG immunohistochemical staining have been reported. For example, VIP and galanin staining is restricted to the juxtaaqueductal zone

(Melander et al., 1986a; 1986b; Moss and Basbaum, 1983; Moss et al., 1983); phenylethanolamine N-methyltransferase and calcitonin staining are particularly intense within parts of the vlPAG (Fabbri et al., 1985; Herbert and Saper, 1992); calbindin staining and to a large extent calretinin staining are most pronounced in the dlPAG, the vlPAG, and the ventral part of the lPAG, but are absent in the dorsal part of the lPAG and the dmPAG

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(Paxinos et al., 1999); neurofilament protein SMI-32 staining is present in the outer (lateral) zone of dmPAG, lPAG, and vlPAG, but is absent in the inner (medial) zone of these columns and in the dlPAG (Paxinos et al., 1999). In addition, the distribution within the PAG of low-abundance peptides, transmitter substances, and their receptors, have been studied using in situ hybridization (De Belleroche et al., 1990; Harlan et al., 1987; Jansen et al., 1998; Lanaud et al., 1989; Mansour et al., 1994, 1995; Rattray et al., 1992; Seroogy et al., 1989; Smith et al., 1994; Tolle et al., 1993) and receptor autoradiographic techniques (Gundlach, 1991; Williams and Beitz, 1989a, 1989b, 1990). These studies have revealed additional discrete inter- and intracolumnar patterns of substancespecific cellular label (mRNAs) and receptors (silver grain densities). These variations may well reflect important organizational properties of the PAG. However, before significance can be attached to any of these distinctive patterns, anatomical and functional studies focused on these more subtle differences are needed.

ANATOMICAL STUDIES Brain Stem Efferents The somatomotor, cardiovascular, and antinociceptive adjustments which characterize active and passive coping can be readily elicited naturally, or by PAG stimulation, in a precollicular decerebrate rat or cat (i.e., a preparation in which the brain stem has been disconnected from the forebrain, including the hypothalamus). Such findings suggest that projections to the lower brain stem (there is a relative paucity of direct PAG–spinal projections) provide the essential substrates for PAG-mediated active and passive coping. As illustrated schematically in Fig. 4 both the vlPAG and lPAG project extensively to ventromedial and ventrolateral medullary regions controlling cardiovascular, somatomotor, and antinociceptive functions (Bandler et al., 1991; Bernard and Bandler, 1998; Carrive et al., 1988; Ennis et al., 1991; Hamilton, 1973b; Hamilton and Skultety, 1970; Henderson et al., 1998; Lovick 1985, 1992b, 1996; Morgan et al., 1989; Van Bockstaele et al., 1991; Vanderhorst et al., 2000). Interestingly, the “functionally opposed” vlPAG and lPAG broadly target the same ventromedial and ventrolateral medullary regions, suggesting that the different sets of projections: (i) use different neurotransmitters or (ii) target different neural populations in each area. Consistent with either hypothesis there is good evidence that vlPAG and lPAG excitation have opposite physiological effects on RVLM and VMM neural activity (Lovick, 1985, 1992a, 1992b, 1993, 1996; Morgan, 1991; Morgan et al., 1998; Verberne

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and Guyenet, 1992; Verberne and Boudier, 1991). The dmPAG column, whose functions are less frequently studied, projects also to the ventromedial and ventrolateral medulla. The dlPAG column, however, has no direct projections to the medulla (Fig. 4). Instead, it strongly targets: (i) the cuneiform nucleus, a region which projects to the ventrolateral medulla and from which active defensive behaviors (freezing, flight, hypertension) are evoked (Mitchell et al., 1988a, 1988b; Redgrave et al., 1988), and (ii) the superolateral parabrachial nucleus, a region which strongly innervates retrochiasmatic and ventromedial (dorsomedial division) hypothalamic nuclei (Bernard and Bandler, 1998; Krout et al., 1998).

Diencephalic Efferents Distinct PAG columns project also to specific hypothalamic and midline and intralaminar thalamic regions (Floyd et al., 1996 ; Krout and Loewy, 2000). These PAG–diencephalic projections likely contribute to active and passive coping responses in the intact animal. The lateral hypothalamic area, a region from which hypotension and bradycardia are readily evoked by EAA microinjection (Allen and Cechetto, 1992; Gelsema et al., 1989; Spencer et al., 1990), is selectively targeted by the vlPAG, whereas dorsal and medial hypothalamic areas from which hypertension, tachycardia, and somatomotor activation are evoked by microinjection of EAAs or GABA antagonists (Allen and Cechetto, 1992; DeNovellis et al., 1995; Gelsema et al., 1989; Sun and Guyenet, 1986; Waldrop et al., 1988) receive inputs from the lPAG and the dlPAG (see Fig. 6). In addition, the vlPAG provides the heaviest input to the thalamus, specifically to centromedial, centrolateral, intermediodorsal, and paraventricular nuclei, and to a restricted, medial part of the caudal, ventromedial nucleus (VMc) (Floyd et al., 1996; Krout and Loewy, 2000). The lPAG provided a more moderate input to same regions, with the exception of the VMc. The dlPAG, in contrast, projects predominantly to the paraventricular thalamic nucleus (Krout and Loewy, 2000).

Spinal and Medullary Afferents In terms of “direct” sensory inputs, the PAG is dominated by afferents (somatic and visceral) arising from the spinal cord, the medullary dorsal horn (Sp5) and the nucleus of the solitary tract (Sol) (Bjorkeland and Boivie, 1984; Clement et al., 1998; Cowie and Holstege, 1992; Herbert and Saper, 1992; Keay and Bandler, 1992; Keay et al., 1997a, 1997b; Mantyh, 1982; Menetrey et al., 1982; Wiberg and Blomqvist, 1984; Yezierski, 1988; Yezierski and Mendez, 1991; Yezierski and Schwartz,

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Nucleus of the Solitary Tract Spinal Cord

Sol

Medullary Dorsal Horn dm

dl

Sp5 l dm

PAG

vl

dl

l dm vl

dl l

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CVLM FIGURE 4 Schematic illustration of somatic and visceral afferents to different PAG neuronal columns (top half of figure) and medullary projections arising from different PAG neuronal columns (bottom half of figure). It can be seen that the ventrolateral column of the PAG (vlPAG) receives afferents from the spinal cord and the nucleus of the solitary tract (Sol) and projects to both the rostral and caudal ventrolateral medulla (RVLM, CVLM) and the ventromedial medulla (VMM). The lateral column of the PAG (lPAG) receives afferents from spinal cord and the medullary dorsal horn (Sp5) and projects to the RVLM, CVLM, and VMM. The dorsolateral column of the PAG (dlPAG) has no significant somatic or visceral inputs arising from spinal cord, Sp5, or Sol. The dlPAG does not project directly to the medulla, but can influence the rostral ventrolateral and ventromedial medulla via a projection to the cuneiform nucleus (cnf). (Modified from Fig. 2 of Keay and Bandler, 2001).

1986; Yezierski et al., 1987). These inputs terminate almost exclusively in the vlPAG and lPAG columns; i.e., there is no anatomical evidence of significant, direct spinal, Sp5, or Sol projections to either the dlPAG or the dmPAG. The lPAG receives a somatotopically organized input (i.e., lumbar enlargement to caudal lPAG, cervical

enlargement and Sp5 to progressively more rostral parts of lPAG); whereas, afferents from Sol and spinal cord converge onto the vlPAG without any apparent topographical organisation (see Fig. 4). Interestingly, the Sol does not project directly to the lPAG, nor does the Sp5 directly innervate the vlPAG.

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The spinal projections to both lPAG and vlPAG arise predominantly from the superficial and deep dorsal horn, with two notable exceptions: (i) in the sacral cord, vlPAG- and lPAG-projecting neurons are located predominantly in the region of the sacral parasympathetic nucleus; and (ii) in the upper cervical cord (segments C1–C3), vlPAG projections arise predominantly from neurons in deep dorsal horn and the intermediate and ventral gray. It should be noted that approximately 30% of all spinal neurons that project to the vlPAG are located in the C1 spinal segment, with an additional 20% found in segments C2–C3. Approximately 50% of the spinal neurons that project to the lPAG are located also within the superficial and deep dorsal horn of the upper cervical cord (C1–C3). The functional significance of this extraordinarily large upper cervical input to the vlPAG and lPAG has yet to be investigated (although see Clement et al., 2000; Keay and Bandler, 2002; Keay et al., 1997b, 2000). In addition to Sol–vlPAG projections, there are extensive projections to the PAG arising from the ventrolateral, ventromedial, and dorsomedial medulla. The general principle underlying these projections is that ventromedial and dorsomedial medullary regions preferentially target the vlPAG (Herbert and Saper, 1992); whereas ventrolateral medullary neurons project to dmPAG, lPAG, and vlPAG (Beitz, 1982; Clement et al., 1998; Hamba et al., 1990; Herbert and Saper, 1992; Kwiat and Basbaum, 1990). The absence of any substantial medullary projection to the dlPAG is a particularly striking finding.

Forebrain Afferents The potential significance of prefrontal cortical (PFC) projections to specific PAG columns (although there were earlier reports of PFC–PAG projections (e.g., Hurley et al., 1991; Sesack et al., 1989) was appreciated only after the work of Shipley and colleagues (Shipley et al., 1991). Their study provided the first systematic evidence that projections arising from specific PFC fields had patterns of termination that respected PAG columnar boundaries. Recent studies in both rat and primate have confirmed and extended these observations. In the rat, projections to the PAG arise from the medial wall, dorsomedial convexity, and select orbital and insular PFC areas (Floyd et al., 2000). As summarized schematically in Fig. 5, discrete PFC subgroups can be identified on the basis of their projections to a specific PAG column(s). Thus, dorsolateral orbital (DLO), agranular insular (AId, AIp), and medial orbital (MO, VO, VLOm) cortical areas project almost exclusively to the vlPAG. As well, the rostroventral medial PFC wall (PL and IL, blue shaded) projects predominantly to the vlPAG. In contrast, strong projections to the dlPAG

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originate from the caudodorsal medial PFC wall (PL and IL, purple shaded) and the cortex of dorsomedial convexity (Acd and Acv, pink shaded) (Fig. 5). (Note: there are also projections to the lPAG (not illustrated in Fig. 5) that arise from PL, ACd, DLO, and AId; although each of these cortical areas has a stronger projection to either the vlPAG or the dlPAG (Floyd et al., 2000)). In the macaque, medial and select orbito-insular PFC areas are divisible also into subgroups which share projections to different PAG columns (see An et al., 1998). A striking feature of the primate brain is a dramatic growth in the size and complexity of the medial PFC (macaque PFC is predominantly dygranular and granular cortex, whereas rat PFC is exclusively agranular cortex) and an associated remarkable increase in the density of medial PFC projections to the dlPAG (An et al., 1998). The fact that the dlPAG receives no direct spinal or medullary afferents highlights the potential significance of its medial PFC input. Projections to the PAG from the amygdala and the hypothalamus also respect columnar boundaries. Amygdaloid projections to PAG arise principally from the central nucleus and terminate in all but the dlPAG column (Price and Amaral, 1981; Rizvi et al., 1991). With respect to the hypothalamus (see Fig. 6), the lateral hypothalamic area (LHA) projects selectively to the vlPAG. The vlPAG and lPAG also receive inputs from dorsal and medial hypothalamic regions (not illustrated). Hypothalamic projections to the dlPAG are more restricted, arising primarily from the dorsal (DHA), posterior (PHA), and anterior (AHA) hypothalamic areas, as well as from the dorsomedial subdivision of ventromedial hypothalamic nucleus (VMHdm) (Floyd et al., 2001; Veening et al., 1982, 1987, 1991).

THE PAG AND PARALLEL CIRCUITS FOR EMOTIONAL COPING Anatomical Overview In both the rat and the primate, it has been further revealed that the PFC subgroups which project preferentially to specific PAG columns project also to distinct regions of the hypothalamus (An et al., 1998; Bernard and Bandler, 1998; Floyd et al., 2000, 2001; Öngür et al., 1998). As illustrated schematically in Fig. 6: (i) the orbitoinsular and rostroventral medial PFC areas that project to the vlPAG project selectively to the LHA; (ii) the caudodorsal medial PFC areas that project to the dlPAG project selectively to DHA, PHA, and AHA; and (iii) the PAG columns and distinct hypothalamic regions that receive common PFC inputs, in turn, are reciprocally interconnected, i.e., vlPAG–LHA; dlPAG–DHA/

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AId

PRh AIp

MO

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VO/VLO m

MO

ACd ACv PL IL

dlPAG

vlPA G

FIGURE 5 Surface diagrams of lateral (left) and medial (right) views of the rat brain are drawn in the top row (modified from Krettek and Price, 1977) below which is a coronal drawing of the PAG with the location of dlPAG (pink) and vlPAG (blue) columns indicated. Blue shading (top panels) indicates the cortical regions that project to the vlPAG column. These include rostral PL, rostral IL, and MO on the medial wall (top row, right); and VO, VLOm, AIp, AId, and DLO on the orbital and lateral cortical surfaces (top row, left). Pink and purple shading (top panels) indicates the cortical regions that project exclusively (pink) or predominantly (purple) to the dlPAG column. These comprise caudal PL, caudal IL, ACd, and ACv on the medial wall and PRh cortical area on the lateral wall. Abbreviations used: ACd, dorsal anterior cingulate cortex; ACv, ventral anterior cingulate cortex; AId, dorsal agranular insular cortex; AIp, posterior agranular insular cortex; DLO, dorsolateral orbital cortex; IL, infralimbic cortex; MO, medial orbital cortex; PL, prelimbic cortex; PRh, perirhinal cortex; VLOm, ventrolateral orbital cortex, medial part; VO, ventral orbital cortex. (Modified from Fig. 17 of Flody et al. 2000)

PHA/AHA. These anatomical findings suggest that the dorsolateral and ventrolateral (to a lesser extent the lateral) PAG columns are embedded with distinct, but parallel, circuits that extend rostrally to include specific orbital and medial prefrontal cortical areas and specific hypothalamic regions (Fig. 6). To summarize, two PAG columns, the ventrolateral and lateral, are spinal-, Sp5-, and Sol-recipient (Fig. 4). That is, they are regions within which signals from physical stressors (e.g., cutaneous or deep pain, hemorrhage) gain access to neural substrates mediating distinct emotional coping reactions. In contrast, the dorsolateral PAG column (which mediates active coping) has neither direct spinal nor medullary afferents, but does receive strong medial PFC input (more so in the primate than in the rat). These anatomical connections suggest that active coping mediated by the dlPAG may be driven by stressors that have a predominantly psychological component (i.e., are dependent on PFC neural processing), whereas active coping mediated

by the lPAG is likely to driven by physical stressors. Passive emotional coping, whether the precipitating event is physical or psychological, should be mediated exclusively by the vlPAG.

Functional Studies (Immediate Early Gene Expression) Consistent with this anatomical analysis, studies which have utilized immediate early gene (c-fos) expression as a marker of neuronal activation indicate that PAG columns are selectively activated by specific classes of stress (Bellchambers et al., 1998; Canteras and Goto, 1999; Chieng et al., 1995; Clement et al., 1996; Keay and Bandler, 1993, 1998, 2002; Keay et al., 1994, 1997a, 2000, 2001; Tassorelli and Joseph, 1995). For example, three distinct “inescapable” physical stressors (muscle pain (intramuscular formalin), visceral pain (intravenous 5-HT), inescapable cutaneous pain (clip of neck)), each of which evoke passive coping as their

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Prefrontal Cortex DLO AId FPm

ACv

PL

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MO

Hypothalamus

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f LHA a

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f

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FIGURE 6 Blue shading indicates the orbital and medial prefrontal cortical (PFC) areas that project to both the vlPAG and the lateral hypothalamus (LHA). Yellow shading indicates the orbital and medial PFC areas that project to the dlPAG and the medial hypothalamic regions (AHA, anterior hypothalamic area; DHA, dorsal hypothalamic area; PHA, posterior hypothalamic area). Note also that hypothalamic areas and PAG columns projected upon by common PFC areas are also interconnected. See Fig. 5 for cortical abbreviations.

primary response, selectively activate neurons in the vlPAG column (Fig. 7A). Other manipulations (not illustrated) which also evoke passive coping as their primary response (e.g., hypotensive haemorrhage (Keay et al., 1997a); noxious stimulation of cerebral vessels

(Keay and Bandler, 1998); systemic injection of nitroglycerin (Tassorelli and Joseph, 1995)) similarly elicit selective vlPAG Fos expression. In contrast, stressors which as their primary response evoke an active coping reaction elicit Fos expression in the lPAG and/or the

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Intravenous 5-HT

Clip of the Neck (Keay et al., 2001b)

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B Opiate Withdrawal (modified from Chieng et al., 1998)

Radiant Heat (Keay & Bandler, 1993) -5.8

C

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Exposure to a Cat (modified from Canteras & Goto, 1999) -5.8

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FIGURE 7 Camera-lucida drawings of coronal sections through the PAG (5.8, 6.8, 7.8, and 8.8 mm caudal to bregma) showing patterns of neural activation following physical or psychological stressors which trigger as their primary reaction either active or passive coping. (A) Patterns of neural activation following three physical stressors which evoke passive coping as their primary response: (i) intramuscular injection of formalin (top row), (ii) intravenous injection of 5HT (middle row), and (iii) clip applied to the dorsal skin of the neck (bottom row). The shaded area indicates the ventrolateral PAG column. (B) Patterns of neural activation following two physical stressors which evoke active coping as their primary response: (i) opiate withdrawal (top row) and (ii) radiant heat applied to the skin of the neck (bottom row). The shaded area indicates the lateral PAG column. C Pattern of neural activation evoked by a “psychological stressor,” exposure of a rat to a cat (additional data generously provided by Professor Newton Canteras). The shaded area indicates the dorsolateral PAG column. (Modified from Fig. 5 of Keay and Bandler, 2001)

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dlPAG (Figs. 7B and 7C). Note, however, that when active coping is triggered by a physical stressor (e.g., opiate withdrawal, radiant heat) Fos expression is much stronger in the lPAG than in the dlPAG (Fig. 7B). In contrast, when a “psychological” stressor is used (e.g., the presence of a cat, but without physical contact) Fos expression in the dlPAG predominates (Fig. 7C). The presence of significant vlPAG Fos expression in Figs. 7B and 7C likely reflects the fact that a stress-triggered period of active coping may be followed by a delayed period of recovery and healing (i.e., passive coping/ recuperation), if the opportunity is available (Bandler et al., 2000a, 2000b).

CONCLUSIONS All mammals possess the capacity to affect appropriate responses to “escapable” or “inescapable” stressors and to facilitate recovery and healing once the stress passes. Different stressors possess, to varying degrees, physical and psychological components. A substantial body of evidence has been reviewed which supports the concept that the PAG is divisible into a number of distinct, longitudinal neuronal columns, each of which lies embedded within a circuit that extends rostrally to include specific PFC and hypothalamic areas. These PFC–PAG/hypothalamic circuits, in turn, project caudally to affect somatic and autonomic premotor neurons within the ventrolateral medulla and the ventromedial (raphe) and paramedian medullary neural pools. The evidence reviewed suggests that different PAG columns (and their distinctive forebrain and brain stem connections) play important roles in coordinating distinct emotional coping strategies for dealing with different classes of stress. Specifically, it has been proposed: (i) that the lPAG column (and its associated circuit) is activated preferentially by “escapable” physical stressors to which an active defensive reaction(s) is the primary response; (ii) that the dlPAG column (and its associated circuit) is activated preferentially by “escapable” psychological stressors to which an active defensive response is the primary response; and (iii) that the vlPAG column (and its associated circuit) is activated (a) by “inescapable” physical or psychological stressors for which passive coping behavior is the primary response or (b) as a delayed/secondary response, to promote recovery and healing following any stressor.

Acknowledgments The authors acknowledge the support of NHMRC (Australia) for their work described in this chapter.

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Functional, Anatomical and Neurochemical Organization.” (Depaulis, A., and Bandler, R., Eds.). pp. 417–448. Plenum Press, New York. Smith, G. S., Savery, D., Marsden, C., Lopez Costa, J. J., Averill, S., Priestley, J. V., and Rattray, M. (1994). Distribution of messenger RNAs encoding enkephalin, substance p, somatostatin, galanin, vasoactive intestinal polypeptide, neuropeptide Y, and calcitonin gene-related peptide in the midbrain periaqueductal grey in the rat. J. Comp. Neurol. 350, 23–40. Spencer, S. E., Sawyer, W. B., and Loewy, A. D. (1990). L-Glutamate mapping of cardioreactive areas in the rat posterior hypothalamus. Brain Res. 511, 149–157. Sun, M. K., and Guyenet, P. G. (1986). Hypothalamic glutamatergic input to medullary sympathoexcitatory neurons in rats. Am. J. Physiol. 251, R798–R810. Tassorelli, C., and Joseph, S. A. (1995). Systemic nitroglycerin induces fos immunoreactivity in brainstem and forebrain structures of the rat. Brain Res. 682, 167–181. Tolle, T. R., Berthele, A., Zieglgansberger, W., Seeburg, P. H., and Wisden, W. (1993). The differential expression of 16 NMDA and non-NMDA receptor subunits in the rat spinal cord and in periaqueductal gray. J. Neurosci. 13, 5009–5028. Van Bockstaele, E. J., Aston-Jones, G., Pieribone, V. A., Ennis, M., and Shipley, M. T. (1991). Subregions of the periaqueductal gray topographically innervate the rostral ventral medulla in the rat. J. Comp. Neurol. 309, 305–327. Vanderhorst, V. G., Terasawa, E., Ralston, H. J., III, and Holstege, G. (2000). Monosynaptic projections from the lateral periaqueductal gray to the nucleus retroambiguus in the rhesus monkey: Implications for vocalization and reproductive behavior. J. Comp. Neurol. 424, 251–268. Veening, J., Buma, P., Ter Horst, G. J., and Roeling, T. A. P. (1991). Hypothalamic projections to the pag in the rat: Topographical, immuno-electron microscopical and functional aspects. In “The Midbrain Periaqueductal Gray Matter: Functional, Anatomical, and Neurochemical Organization” (Depaulis, A., and Bandler, R., Eds.). Plenum Press, New York. Veening, J. G., Swanson, L. W., Cowan, W. M., Nieuwenhuys, R., and Geeraedts, L. M. (1982). The medial forebrain bundle of the rat. II. An autoradiographic study of the topography of the major descending and ascending components. J. Comp. Neurol. 206, 82–108. Veening, J. G., Te Lie, S., Posthuma, P., Geeraedts, L. M., and Nieuwenhuys, R. (1987). A topographical analysis of the origin of some efferent projections from the lateral hypothalamic area in the rat. Neuroscience 22, 537–551. Verberne, A., and Guyenet, P. (1992). Midbrain central gray: Influence on medullary sympathoexcitatory neurons and the baroreflex in rats. Am. J. Physiol. 263, R24–R33. Verberne, A. J., and Boudier, H. R. S. (1991). Midbrain central grey: Regional hemodynamic control and excitatory amino acidergic mechanisms. Brain Res. 550, 86–94. Waldrop, T. G., Bauer, R. M., and Iwamoto, G. A. (1988). Microinjection of GABA antagonists into the posterior hypothalamus elicits locomotor activity and a cardiorespiratory activation. Brain Res. 444, 84–94. Wiberg, M., and Blomqvist, A. (1984). The spinomesencephalic tract in the cat: Its cells of origin and termination pattern as demonstrated by the intraaxonal transport method. Brain Res. 291, 1–18. Williams, F. G., and Beitz, A. J. (1989a). Production and characterization of a novel monoclonal antibody against neurotensin: Immunohistochemical localization in the midbrain and hypothalamus. J. Histochem. Cytochem. 37, 831–841. Williams, F. G., and Beitz, A. J. (1989b). A quantitative ultrastructural analysis of neurotensin-like immunoreactive terminals in the

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C H A P T E R

11 Locus Coeruleus, A5 and A7 Noradrenergic Cell Groups GARY ASTON-JONES Laboratory of Neuromodulation and Behavior, Department of Psychiatry University of Pennsylvania School of Medicine, Philadelphia, USA

CYTOARCHITECTURE

Several significant advances in understanding the structure and function of the locus coeruleus (LC) system have taken place since the previous edition of this book. In particular, numerous studies have extended our understanding of neurotransmitter-defined inputs to the LC. Also, substantial progress has been made in identifying inputs to dendrites of LC neurons that lie outside of the LC nuclear core. The recent application of viral transsynaptic tract-tracing techniques has begun to extend our understanding of how these systems are regulated by circuits that feed into these norepinephrine (NE) cells, including second- and higher-order afferents. This chapter also extends the previous version by including advances in functional understanding of these NE systems, ranging from the role of the A5 and A7 systems in pain to the role of the LC system in attention and cognitive processing. The purpose of this chapter is twofold. One goal is to provide a succinct synopsis of the most salient anatomical features of the LC and other metencephalic NE neuronal systems in the rat. A second goal is to emphasize new anatomical results obtained since the previous editions of this book, so as to update readers with the most current information. Owing to length limitations, only a skeleton outline of the extensive anatomical knowledge that exists for these neurons can be provided. I have tried to include features most likely to be of interest to a large number of readers.

The Rat Nervous System, Third Edition

Cell Types and Subnuclei The A4 and subcoeruleus cell groups are considered here to be part of the LC. However, the A5 and A7 cell groups are considered to be separate from the subcoeruleus; the subcoeruleus is the ventral extension of the LC and does not encroach into the periolivary region of the A5 cells or into the periparabrachial area of the A7 cells. In the rat, the LC nucleus is readily recognized in Nissl-stained sections where it appears as a densely packed cluster of darkly stained cells in the rostral rhombencephalic tegmentum (Fig. 1). The nucleus stretches for 1.2 mm along the ventrolateral edge of the IVth ventricle, abutting the medial side of the mesencephalic nucleus of the trigeminal nerve. Caudally the nucleus extends to the genu of the facial nerve; rostrally it reaches into the periaqueductal gray (Grzanna and Molliver, 1980). The LC reaches its largest mediolateral extent at the level of the motor nucleus of the trigeminal nerve. Estimates of the number of neurons in the rat LC (unilaterally) range from about 1400 to 1800 (Descarries and Saucier, 1972; Goldman and Coleman, 1981; Loughlin et al., 1986; McBride et al., 1985; Swanson, 1976) depending on the staining method used to delineate the nucleus.

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FIGURE 1 Bright-field photomicrographs of two adjacent sections through the rat locus coeruleus. The section shown on the left was stained with cresyl violet. The section shown on the right was processed for dopamine β-hydroxylase immunohistochemistry to visualize noradrenergic neurons and extensive dendritic processes. The bar represents 100 μm. Dorsal is at the top and medial is at the left of each figure.

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Studies with the Golgi method have identified several different types of neurons in the rat LC. Swanson (1976) distinguished medium-sized fusiform cells and somewhat larger multipolar neurons in the rat LC. Other investigators distinguished a third class of ovoidshaped cells in the rat LC (Cintra et al., 1982; Pfister and Danner, 1980; Shimizu and Imamoto, 1970). The cell types differ not only in size and shape but also in the orientation of their dendrites. Small fusiform cells are obliquely positioned with the long axis extending mainly in the anterior–posterior directions and tilted in a dorsolateral to ventromedial orientation suggesting a dendritic arborization mainly in the sagittal and horizontal plane. In contrast, the dendritic arborization of multipolar LC neurons appears to be primarily in the frontal plane (Sievers et al., 1981). Loughlin et al. (1986) analyzed the appearance of retrogradely filled LC neurons following injections of horseradish peroxidase into selected terminal fields and were able to distinguish six different subpopulations of cells in the rat LC. A principal finding of this study was that morphologic subpopulations of LC neurons have different efferent targets. Despite differences in their morphology, nearly all neurons in the rat LC contain noradrenaline, and it has become customary to consider the presence of this transmitter and its biosynthetic enzymes the defining property of the rat LC. In addition to the morphological heterogeneity among its cells, the rat LC can be divided into subnuclei based upon the size of its cells and the orientation of its dendrites. Swanson (1976) distinguished a large dorsal portion of the LC comprised of small and tightly packed cells. This dorsal subdivision is readily distinguished from a somewhat smaller ventral subdivision in the caudal third of the nucleus consisting of larger, less densely packed cells. The number of cells in the ventral LC has been estimated to be 200 per nucleus. Extending laterally and dorsally is an outpost of subependymal NE neurons in the roof of the fourth ventricle referred to as the A4 group by Dahlstrom and Fuxe (1964). A distinct feature of these neurons is the set of processes that extend between ependymal cells toward the ventricular surface. A small subdivision of the LC is formed by a small group of large multipolar neurons scattered in the ventral portion of the periadqueductal gray (Grzanna and Molliver, 1980; Grzanna et al., 1980). These cells lie approximately 500 μm rostral to the compact portion of the LC and can only be distinguished from neighboring cells in the rostral periaqueductal gray (PAG) by their NE content and the presence of the corresponding biosynthetic enzymes. These cells have prominent dendrites that radiate in all directions within the ventrolateral portion of the periaqueductal gray.

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Delineation of the subcoeruleus in the rat has been inconsistent and no satisfactory nomenclature has yet emerged. Unlike the LC, the subcoeruleus is a biochemically heterogeneous cell group containing both NE and non-NE neurons. Following the introduction of the histofluorescence method there has been a tendency to focus on the NE cells of the subcoeruleus. Olson and Fuxe (1972) refer to a subcoeruleus area which contains NE cell bodies connecting the ventral part of the LC, the A5 and the A7 group. Amaral and Sinnamon (1977) suggested that the term subcoeruleus should apply only to the cluster of NE-containing cell bodies that lies immediately ventral to the LC. However, this cell cluster is continuous with a sheet of scattered NE neurons that extends ventrally toward the A5 group and rostrally toward the A7 group. Grzanna et al. (1987) delineated the NE-containing cells of the subcoeruleus and counted approximately 130 cells per nucleus. Based on retrograde transport studies, these scattered NE cells in the pontine tegmentum between the ventral portion of the LC proper and the A5 and A7 groups share projections with the ventral LC.

Neurotransmitters within Rat LC Neurons In the rat, the LC nucleus proper is a dense collection of noradrenergic neurons as evidenced by either catecholamine-induced fluorescence or immunohistochemistry for the synthetic enzymes tyrosine hydroxylase (TH) or dopamine β-hydroxylase (DBH; Fig. 1). Analyses with these stains indicated that all neurons within this nucleus are noradrenergic. However, more recent studies indicate the presence of a limited population of small, round neurons within the LC that stain with an antibody to GABA (Ijima et al., 1987; Ijima and Ohtomo, 1988). The functional significance of these small neurons is unknown. It is notable that the rostral pole of the LC in the rat is more neurochemically heterogeneous than the nucleus proper and contains many non-NE neurons intermixed with NE neurons. This characteristic of the rostral pole may yield properties not seen in more caudal LC areas (e.g., additional afferents, interneurons). Other putative neurotransmitter molecules in addition to NE can distinguish subsets of rat NE neurons. Most prominent among these is the neuropeptide galanin (Austin et al., 1990; Holets et al., 1988; Levin et al., 1987; Melander et al., 1986; Skofitsch and Jacobowitz, 1985; Sutin and Jacobowitz, 1988) which is present in most, though not all, LC NE neurons. Additional peptides present in a smaller percentage of noradrenergic LC neurons include neuropeptide Y (NPY) (Everitt et al., 1984; Sutin and Jacobowitz, 1988), vasopressin, neurophysin (Caffe et al., 1985, 1988), neurotensin, vasoactive

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intestinal peptide (VIP), and atrial natriuretic factor (Sutin and Jacobowitz, 1988). Coexistence of NE, vasopressin, and neurophysin has been reported in some subcoeruleus neurons (Caffe et al., 1985). The majority of the NPY-containing neurons were found in the dorsal LC, and very few noradrenergic neurons in the subcoeruleus were found to also contain either NPY or galanin (Holets et al., 1988). While enkephalin has been reported to exist within some NE neurons of cat LC (Charnay et al., 1982, 1984) as well as in some early reports in rat LC neurons (Finley et al., 1981; Khachaturian et al., 1983), more recent work by us and others has found no evidence for enkephalin–NE coexistence within the rat LC (Drolet et al., 1992; Fallon and Leslie, 1986; Murakami et al., 1987). Similarly, although it has been suggested that corticotropin-releasing hormone (CRH) may be contained within some LC NE neurons, we (Valentino et al., 1992) and others (Sutin and Jacobowitz, 1988) find no evidence for this in more recent studies. It has also been reported that a higher percentage of LC neurons that contain NPY or galanin project to the hypothalamus than to the spinal cord or cerebral cortex (Holets et al., 1988), indicating that LC neurons may have specialized targets corresponding to colocalized peptides. Non-peptide transmitter candidates have also been identified within a large percentage of LC NE neurons, including N-acetylaspartylglutamate (Forloni et al., 1987) and the putative marker of glutamatergic neurons glutaminase (Drolet and Aston-Jones, 1991; Kaneko et al., 1989, 1990). A recent study reported that some rat LC neurons express tryptophan hydroxylase mRNA as well as immunoreactivity for serotonin (Ijima and Sato, 1991). These results have yet to be confirmed by other studies. In addition, the functional significance of possible colocalization of other transmitter molecules within LC neurons is unclear. For example, it has been reported that neuropeptide Y and galanin (Moore and Gustafson, 1989), as well as vasopressin (Caffe et al., 1988), may not be transported to LC terminals in cortical areas, despite being located within LC somata that innervate these areas.

NE Transporters Recent studies have cloned the molecule responsible for reuptake of NE, termed the NE transporter (NET) (Barker and Blakely, 1995; Pacholczyk et al., 1991). Immunolocalization of NET in rat brain has shown that it is confined to NE neurons and processes and is not present within, e.g., DA or serotonin (5-HT) neurons (Schroeter et al., 2000). NET labeling exhibited a nonuniform pattern of expression along axons, reflecting a high degree of spatial organization of NE clearance. In

addition, ultrastructural analysis of NET in the prefrontal cortex revealed a prominent cytoplasmic localization in presynaptic terminals, indicating possible regulated trafficking of the transporter controlling NE clearance (Schroeter et al., 2000). There is evidence that NET may be phosphorylated in a protein kinase Cdependent manner, and it appears that transmitters, antagonists, and second messengers can modify the intrinsic activity and surface expression or protein levels of amine transporters including NET (Blakely and Bauman, 2000). Thus, it is likely that NET activity is a mechanism contributing to synaptic plasticity rather than a static determinant of transmitter clearance. NET has also been found in A5 and A7 NE neurons, as expected (Comer et al., 1998).

Electrotonic Coupling within the LC In neonatal rats Christie and colleagues have provided compelling evidence that LC neurons are electrotonically coupled (Christie and Jelinek, 1993; Christie et al., 1989). In rats less than about 1 week of age, these investigators found abundant dye coupling and electrotonic conduction between pairs of LC neurons. Although this evidence for electrotonic coupling among LC cells wanes as animals approach adulthood, recent evidence from Williams and colleagues reveals that coupling may persist into adulthood. They found that the reversal potential for opiates (Osborne and Williams, 1996; Travagli et al., 1995), the effects of locally applied neurotransmitters on LC neurons in the slice (Travagli et al., 1995), and synchronous activity among adult LC neurons (Ishimatsu and Williams, 1996; Travagli et al., 1995) could be explained by electrotonic coupling. The apparent coupling was reduced after application of the coupling blocker carbenoxolone. Coupling was also reduced by cutting slices of the LC in the coronal plane instead of the horizontal plane (Osborne and Williams, 1996; Travagli et al., 1996) or by isolating the cell body region of the LC in horizontal slices from the rostral and caudal dendritic zones (Ishimatsu and Williams, 1996). This indicates that coupling may predominantly occur between distal dendrites of LC neurons in the peri-LC region. More recent studies have also shown coupling between LC neurons and astrocytes in neonatal LC. This coupling was shown anatomically with dye coupling and with staining for connexin proteins and also found to be functional in that depolarization of astrocytes increased the firing rate of LC neurons (AlvarezMaubecin et al., 2000). Studies recording LC neurons in behaving monkeys have found that electrotonic coupling among LC neurons may have important behavioral consequences

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as well (Usher et al., 1999). Different modes of LC activity were simulated by changes in coupling among LC neurons, and these different modes were found to be capable of producing changes in attentiveness on a target detection task. Thus, coupling among cells in the LC could be an important mechanism regulating functions of this noradrenergic efferent network.

AFFERENTS TO THE NUCLEUS LOCUS COERULEUS Neurotransmitter Inputs to the LC and Peri-LC Area There are various lines of evidence that a wealth of neurotransmitters innervate the LC. Here, we briefly

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review neurochemically identified fibers that have been localized in the rat LC, as well as the receptor content and pharmacological sensitivity of these neurons. However, due to the very small size of the LC nucleus and the marked neurochemical topography in the LC region, studies employing punch or other tissue sampling of the LC for biochemical measures are unable to specifically ascribe results to the LC nucleus vs surrounding areas, and these studies are not included here. The recently discovered dense innervation of the LC nucleus by the novel neuropeptide hypocretin (also known as orexin) is particularly intriguing (Fig. 2) (Cutler et al., 1999; de Lecea et al., 1998; Hervieu et al., 2001; Horvath et al., 1999). This peptide is made only by neurons in the hypothalamus. Neuropharmacologic studies have shown that hypocretin strongly activates LC neurons by decreasing a resting potassium conductance

FIGURE 2 (A) Hypocretin (orexin) innervation of the LC. Bright-field photomicrograph of a frontal section through the rat LC stained with an antibody against hypocretin. Note dense innervation of the LC nucleus by fibers containing this peptide. Neutral red counterstain. Fourth ventricle is to the left (medial); dorsal is up. (B) Innervation of peri-LC zone from the medial prefrontal cortex. Dark-field photomicrograph showing fibers anterogradely labeled with Phaseolus vulgaris leucoagglutinin (PHA-L; gold color) after an injection in the medial prefrontal cortex of a rat. The LC nucleus appears as a dark cluster of neurons to the right of the gold-labeled PHA-L fibers. Note that the prefrontal fibers provide very little input to the LC nuclear core, but instead preferentially terminate in the ventromedial peri-LC area. This innervation is among LC extranuclear dendrites (compare to location of LC dendrites in Fig. 1). Stimulation of this area of the cortex activates LC neurons (see Fig. 5). The fourth ventricle is medial (at left); dorsal is up.

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FIGURE 3 Voltage-dependent depolarization of LC neurons by hypocretin (HCRT; orexin) in brain slice. (A1, A2) Two depolarizing responses to a puff application of HCRT. Both responses were recorded in the same cell, either at resting membrane potential (A1) or 10 min later when the cell was hyperpolarized to −85 mV (A2). Note a smaller depolarization in the latter. (B) Three puff applications of HCRT (as above, except 30-s application) to the same cell, at 10-min intervals and different membrane potentials as indicated. The first application evoked a clear depolarization at resting membrane potential (B1). The second application, after the membrane potential was shifted to 90 mV, produced no depolarization (B2). The third application again depolarized the cell when the membrane potential was returned to the previous resting level (B3). TTX (1 μM) and Co2+ (1 mM) were added to the bath 3 min before trace B1 was recorded and were continuously present during the following records. Note that Ca2+ spikes and associated membrane oscillations, but not the HCRT-induced depolarization, were blocked by Co2+. The dotted lines with the numbers below indicate the level of membrane potential in all records. (A, B) LC neurons from different slices. Taken from Ivanov and Aston-Jones, 2000.

(Fig. 3) (Hagan et al., 1999; Horvath et al., 1999; Ivanov and Aston-Jones, 2000). Hypocretin has attracted a great deal of attention due to its apparent role in feeding, as well as in regulation of sleep and waking (Willie et al., 2001). The former would be consistent with the recent demonstration of leptin receptors in the LC and A7 cell groups (Hay-Schmidt et al., 2001). However, the latter may also be particularly relevant for innervation of the LC, as the LC NE system has long been implicated in regulation of arousal (AstonJones and Bloom, 1981a).

As reviewed in Table 1, a host of immunohistochemically defined fibers have been localized within the LC or in the peri-LC region containing LC dendrites (discussed below) (Shipley et al., 1996). As seen in Table 1, most such observations have been consistently made by different investigators. Opiate receptors (Atweh and Kuhar, 1977; Van Bockstaele et al., 1996b, 1996c), α1 and α2 adrenoceptors (Jones et al., 1985; Lee et al., 1998a, 1998b; Young and Kuhar, 1980), calcitonin gene-related peptide binding sites (Skofitsch and Jacobowitz, 1985), and moderate levels of muscarinic cholinergic receptors (Rotter et al., 1979) have been reported in the LC. More recently, receptors for glutamate (Van Bockstaele and Colago, 1996), purines (Kanjhan et al., 1999; Yao et al., 2000), hypocretin/orexin (Sunter et al., 2001), neurokinins (Chen et al., 2000; Hahn and Bannon, 1999), and CRH (Morin et al., 1999) have been identified in the LC. There are limitations to the interpretation of these studies, however. For example, it is well documented (Herkenham, 1987) that receptor localization does not always correlate with anatomical inputs containing the appropriate corresponding neurotransmitters. In addition, receptor localization and immunohistochemical studies alone do not specify the sources or anatomic pathways responsible for the proposed chemically defined inputs. In neuropharmacologic studies, LC neurons are strongly influenced by a wide range of putative neurotransmitters. For example, LC cells are potently inhibited by α2-adrenergic agonists (Aghajanian et al., 1977; Cedarbaum and Aghajanian, 1976, 1977), GABA (Cedarbaum and Aghajanian, 1977; Guyenet and Aghajanian, 1979) and μ opiate agonists (Bird and Kuhar, 1977; Williams and North, 1984). These neurons are excited by substance P (Guyenet and Aghajanian, 1977), adrenocorticotropin hormone (Olpe and Jones, 1982; Valentino et al., 1983), CRH (CRH; Valentino et al., 1983), acetylcholine (ACh; Aston-Jones et al., 1991a, 1991b; Bird and Kuhar, 1977; Guyenet and Aghajanian, 1979), and glutamate (Aston-Jones et al., 1991a, 1991b; Cherubini et al., 1988); neurotensin and serotonin have yielded more complex results (Aston-Jones et al., 1991a, 1991b; Guyenet and Aghajanian, 1977; Young et al., 1978). It is difficult to draw conclusions about the sources of afferents to the LC from such studies, and it is possible these drugs could be acting on presynaptic terminals of afferents to the LC. In fact, a recent study found that 5-HT and opioid receptors are localized on non-NE terminals within the LC, as well as on LC NE processes (Van Bockstaele, 2000; van Bockstaele et al., 1997). This suggests that 5-HT and opioids may regulate other afferents to the LC presynaptically, as well as regulating LC neurons directly.

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TABLE 1 Fibers in peri-LC

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Transmitter

LC

References

ACh

−

+

Altschuler et al., 1984; Butcher and Woolf, 1984; Hartman et al., 1986; Kimura et al., 1984; Ruggiero et al., 1990; Sutin and Jacobowitz, 1988

Epinephrine

+

+

Astier et al., 1986, 1987; Berod et al., 1984; Haselton and Guyenet, 1987; Hokfelt et al., 1974, 1985; Pieribone and Aston-Jones, 1991; Pieribone et al., 1988

serotonin

+

+

Beitz, 1982; Bowker et al., 1981; Charnay et al., 1984; Hunt and Lovick, 1982; Pieribone et al., 1989; Steinbusch, 1984; Thor et al., 1988; Van Bockstaele, 2000

excitatory amino acids

+

+

Aston-Jones and Ennis, 1988; Ennis and Aston-Jones, 1986, 1988; Ottersen and StormMathisen, 1984a, 1984b

GABA

+

+

Aston-Jones et al., 1990, 1991c; Ennis and Aston-Jones, 1989a, 1989b; Mugnaini and Oertel, 1985; Ottersen and Storm-Mathisen, 1984a, 1984b; Shipley et al., 1988

Enkephalin

+

+

Cassini et al., 1989; Charnay et al., 1985; Conrath-Verrier et al., 1983; Drolet et al., 1992; Fallon and Leslie, 1986; Finley et al., 1981; Guthrie and Basbaum, 1984; Hokfelt et al., 1977, 1979; Hunt and Lovick, 1982; Khachaturian et al., 1983; Lynch et al., 1984; Miller and Pickel, 1980; Sar et al., 1978; Uhl et al., 1979; Van Bockstaele et al., 2000; Watson et al., 1980

Substance P

+

+

Cassini et al., 1989; Hokfelt et al., 1978; Ljungdahl et al., 1978; Nomura et al., 1982; Sutin and Jacobowitz, 1988; Triepel et al., 1985

Neurotensin

+

+

Beitz, 1982; Jennes et al., 1982; Minagawa et al., 1983; Papadopoulos et al., 1986; Triepel et al., 1984; Uhl et al., 1979

VIP

+

+

Eiden et al., 1982; Martin et al., 1987; Sutin and Jacobowitz, 1988; Wang and Aghajanian, 1989

Somatostatin

+

+

Johansson et al., 1984; Vincent et al., 1985

CRH

+

+

Bloom et al., 1982; Cummings et al., 1983; Merchenthaler, 1984; Merchenthaler et al., 1982; Olschowka et al., 1982; Sakanaka et al., 1987; Swanson et al., 1983; Valentino et al., 1992; Van Bockstaele et al., 1998a

Hypocretin/orexin

+

+

Cutler et al., 1999; de Lecea et al., 1998; Hervieu et al., 2001; Horvath et al., 1999

Galanin

+

+

Melander et al., 1986; Skofitsch and Jacobowitz, 1985; Sutin and Jacobowitz, 1988

Coexistence of Neurotransmitters in Afferents to LC Evidence indicates that the LC is heavily innervated by glutamate, GABA, and enkephalin inputs (see above). The high incidence of neurons in medullary afferents to the LC that stained for enkephalin (Drolet et al., 1992), combined with the strong GABA or glutamate-mediated effects of stimulating these same regions on LC activity (Ennis and Aston-Jones, 1988; Ennis and Aston-Jones, 1989a, 1989b), made it seem very likely that opiates would colocalize in GABAergic or glutamatergic LC inputs. Indeed, recent ultrastructural studies have revealed that enkephalin inputs to LC neurons can also contain either GABA (Van Bockstaele and Chan, 1997) or glutamate (Van Bockstaele et al., 2000). It seems likely that other examples of colocalized transmitters in afferents to the LC will be identified as well.

ventromedial and lateral hypothalamus (Saper et al., 1976, 1979), ventrolateral medulla (Loewy et al., 1981; McKellar and Loewy, 1982; Sawchenko and Swanson, 1982), and central nucleus of the amygdala (Cedarbaum and Aghajanian, 1978). Deutch et al. (1986) report anterograde transport of Phaseolus vulgaris leucoagglutinin (PHA-L) into LC from the ventral tegmental area in rat (unconfirmed in our studies, described below). There are some qualifications to these studies, however. First, amino acids may be transported transsynaptically, so that such labeling in LC must be viewed with caution. In addition, limited documentation in many previous amino acid studies leaves it unclear whether the labeling is actually within the LC (a relatively tiny nucleus) or in immediately adjacent structures. In other cases it is not clear whether the transported label is in terminals or in fibers passing through or nearby the LC.

Retrograde and Anterograde Tract Tracing Early Tract-Tracing Studies Using tritiated amino acids for anterograde transport, projections have been reported to the LC from the ventral tegmental area (Beckstead et al., 1979), median and dorsal raphe (Conrad et al., 1974; Pierce et al., 1976),

Major afferents Early experiments used retrograde transport of unconjugated horseradish peroxidase (HRP) revealed with diaminobenzidine (DAB) as the peroxidase substrate to examine inputs to the LC in rat (Cedarbaum and Aghajanian, 1978; Morgane and Jacobs, 1979; Clavier, 1979). All three of these studies reported

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similar findings: retrogradely labeled neurons in a complex array of structures in the brain and spinal cord. In the forebrain, major inputs were identified from the central nucleus of the amygdala, the bed nucleus of the stria terminalis, the medial preoptic area, and the dorsomedial and paraventricular nuclei of the hypothalamus. From the brain stem, prominent inputs were reported from the contralateral LC, midbrain central grey, vestibular nuclei, lateral reticular nucleus, and nucleus tractus solitarius (Sol). Substantial inputs were also reported from the fastigial nucleus of the cerebellum and from the marginal zone of the dorsal spinal horn. Although these studies revealed a number of possible regions that project to the LC, these early HRP techniques had several limitations. First, free HRP diffuses substantially, and it is possible that some labeled areas reflect inputs to peri-LC regions that may receive a different set of afferents than the LC nucleus proper (see below). In addition, HRP is avidly taken up by nonterminal fibers of passage, so that labeled neurons could project through, but not to, the LC. Finally, the HRP– DAB method is relatively insensitive, so that the full constellation of inputs to the LC may not have been detected. Others and we reinvestigated afferents to the LC using more sensitive and selective techniques. iontophoretic wheat germ–agglutinin-conjugated HRP (WGA–HRP), as revealed by the tetramethylbenzidine (TMB) reaction, produced injections restricted to the LC nucleus (AstonJones et al., 1986). Such deposits yielded retrogradely labeled neurons most consistently and strongly in two areas, both located in the rostral medulla: the nucleus paragigantocellularis lateralis (LPGi) and the area on the medial edge of the nucleus prepositus hypoglossi. This latter region, however, may not be a component of the prepositus hypoglossi but corresponds to an area subsequently defined in human tissue as a distinct structure, the epifasicular nucleus (EF) (Paxinos and Huang, 1995). The most prominent afferent to LC (in terms of number of densely labeled neurons) was the LPGi. This nucleus, described in rat by Andrezik et al. (1981), is located in the rostral ventrolateral medulla. Retrogradely labeled LPGi neurons were predominantly ipsilateral to the injection site. These cells were scattered in a region extending caudally from a zone just medial to the facial nucleus to the rostral pole of the lateral reticular nucleus; medially, these neurons extended to the inferior olive and, laterally, to the trigeminal sensory nuclei. The second major source of afferents to the LC was located in the dorsomedial rostral medulla, in the region corresponding to the EF. This group of neurons, located on the medial border of the prepositus hypoglossi and centered slightly rostrally to the LPGi-labeled cells, was densely aggregated along the dorsal-most border

of the medial longitudinal fasiculus where it meets the IVth ventricle; labeled cells were also scattered ventrally along the lateral aspects of, and occasionally within, this fasiculus. Retrograde labeling in EF was bilateral, but slightly greater contralaterally. Minor afferents In our initial studies with WGA– HRP (Aston-Jones et al., 1986), two additional areas consistently exhibited retrograde transport, but with only a few sparsely labeled cells. This labeling was found in the dorsal cap of the paraventricular hypothalamic nucleus (Pa) and in the intermediate zone of the spinal gray. For the Pa dorsal cap cells, labeling was bilateral but slightly greater ipsilaterally. Neurons in more central portions of the Pa were only labeled when injections substantially exceeded the boundaries of LC. Labeled neurons in the spinal cord were scattered in the intermediate zone near the central canal, predominantly contralaterally. Many of these cells were so weakly labeled as to be at the threshold of detection. The ventral tegmental area (VLTg), dorsal and median raphe nuclei, PAG, Sol, A5 area, and lateral as well rostral hypothalamus and preoptic area also contained a few labeled cells in some animals, but were unlabeled in other cases. Additional anterograde tracing studies indicated that the VLTg, dorsal spinal horn, and rostral Sol do not project to the core LC nucleus, but instead project to structures adjacent to the LC. Recent Tracing Studies Some studies employed focal LC injections of the more sensitive tracers WGA–HRP (inactivated) coupled to colloidal gold (WGA–apoHRP–Au) or cholera toxin b subunit (CTb). These experiments yielded more consistent retrograde labeling in the PAG (Ennis et al., 1991) and medial preoptic area (Rizvi et al., 1992), indicating that these areas send projections into the LC. My collaborators and I have confirmed with anterograde transport of PHA-L that the ventrolateral PAG and the lateral aspect of the medial preoptic area send sparse projections into the LC nucleus (Ennis et al., 1991). It is noteworthy that these areas send much more pronounced terminations into the immediately adjacent ventromedial and rostral peri-LC regions, areas that contain abundant LC dendrites (described below). The innervation of these dendrites by extrinsic inputs is being elucidated in part with ultrastructural analyses (see below). Retrograde labeling with WGA–apoHRP–Au or CTb from injections centered in the LC nucleus revealed labeled neurons in certain areas more consistently than seen with WGA–HRP tracing, including the Kölliker– Fuse nucleus (A7 area), A5 area, median raphe (B9 area), and caudal and lateral hypothalamus (Luppi et al., 1991). However, the extent of effective injection sites with

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these tracers has not been well studied, so that confirmation by anterograde tracing is needed to confirm such possible additional inputs to the LC. This is especially important as these nuclei send prominent projections to areas immediately surrounding the LC. Again, ultrastructural analysis is required to determine if any of these areas innervates extranuclear LC dendrites. Other conceivable sources of afferents to the LC were more difficult to examine with tracing methods because of their proximity to the LC. In particular, nearby areas including the LDT nucleus, Barrington’s nucleus, parts of the PAG, and other pericoerulear areas were usually contained within the halo of LC injections and were, therefore, very difficult to assess as possible LC afferents. However, recent anterograde studies confirmed inputs to the LC nucleus from Barrington’s nucleus (Valentino et al., 1996) and the ventrolateral PAG (Bajic and Proudfit, 1999). Cases with LC injections that were “off center” such that they impinged on neighboring structures such as the PB, vestibular, or LDT nuclei contained substantial retrograde labeling in areas (e.g., amygdala, spinal dorsal horn, Sol, insular cortex) previously reported to prominently project to the LC nucleus. These areas were unlabeled after injections restricted to the LC. Taken together, these results indicate that the LC receives major afferents from the rostral medulla, as well as from more limited sources of inputs. It appears that many of the previously inferred afferents to the LC terminate, in fact, in the PB and PAG nuclei which are directly adjacent to the LC; the extensive connections of these two areas were unknown when the earlier HRP–DAB studies of LC afferents were published. Recent Studies Using Anterograde Tracing Methods To confirm our retrograde labeling for specific afferentation of the LC, we examined anterograde labeling in the LC area from various nuclei previously reported to be prominent afferents to the LC. After WGA–HRP injections into the central nucleus of the amygdala, the principal LC nucleus was devoid of anterograde fiber labeling; only the rostral pole (where NE and non-NE neurons are interdigitated, as described previously) contained scattered terminals. Dense anterograde labeling was present in the adjacent PB nucleus. Recent studies employing PHA-L anterograde tracing have largely confirmed this result (Wallace et al., 1992). This region also contains LC extranuclear dendrites, and recent ultrastructural analyses indicates that amygdala projections may terminate in part on these LC processes (Van Bockstaele et al., 1996a, 1996e). Our injections of WGA–HRP or PHA-L into the dorsal horn of the thoracic spinal cord, VLTg, prefrontal cortex, or rostral Sol yielded similar results: no

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anterograde labeling in the core of the LC nucleus but robust labeling in the PB lateral to the LC (from Sol) or the PAG medial to the LC (from VLTg, cortex, and dorsal spinal horn) nuclei (Fig. 2B). In contrast to results in rat, recent results indicate that dorsal spinal horn neurons project into the LC in monkey (Westlund and Craig, 1996); this may represent an interesting species difference. Recent ultrastructural analysis reveals that the Sol innervates LC dendrites in the lateral peri-LC region (see below). Labeled fibers following PHA-L injections into the VLTg have been noted in the rostral pole (containing interdigitated NE and non-NE neurons), but not the main body, of the LC nucleus (Deutch et al., 1986). Injections into the caudal Sol (commissuralis region) yielded occasional scattered fibers in the LC (Blessing and Aston-Jones, unpublished observations); as LC injections did not consistently label neurons in the Sol, these fibers could be axons of passage projecting to the adjacent PB area. Overall, the results of these anterograde tracing experiments were consistent with our retrograde data indicating that many previously reported afferents to the LC actually terminate in neighboring structures. Earlier retrograde studies of afferents to the LC did not use anterograde tracing to confirm that candidate afferents actually terminated in the LC. Anterograde labeling was prominent in the LC following WGA–HRP injections into either the LPGi or the EF, confirming that these areas are major sources of inputs to the LC. Similar results have also been obtained using PHA-L (Guyenet and Young, 1987; Van Bockstaele and Aston-Jones, unpublished observations) or biotinylated dextran (BDA) anterograde tracing (Van Bockstaele et al., 1998). Our recent studies with PHA-L tracing have revealed three distinct pathways by which LPGi projections reach the LC (Van Bockstaele et al., 1989): (i) The medullary adrenergic bundle uses a dorsomedial pathway to reach the LC (reviewed below). (ii) PHA-L injections into the medial LPGi labeled fibers reaching the LC primarily by a ventromedial pathway. (iii) Finally, laterally placed injections of PHA-L in the LPGi revealed a lateral pathway which proceeds rostrally lateral to the superior olive and ascends through the lateral pontine tegmentum, through the ventral and dorsal parabrachial regions, to enter the LC from its lateral and rostral aspects. This pathway closely resembles that reported by others (Guyenet and Young, 1987). These results were recently confirmed with other tracers as well (Van Bockstaele et al., 1998b).

Microphysiology Studies of Afferents to the LC The above anatomic results indicate that major LC afferents arise from the LPGi and the EF. In additional experiments, single cell recordings and electrical stimu-

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lation substantiated these conclusions. Space constraints prevent a full description. In brief, no neurons were antidromically activated in the Sol or contralateral LC from LC stimulation, and only 2 of 44 neurons in the lateral reticular nucleus were driven antidromically from the LC (Ennis and Aston-Jones, 1989a, 1989b). One of these was located in the rostral pole of this nucleus, at the caudal border of the LPGi, and may be a member of this latter set of cells. By contrast, a high percentage of neurons in the LPGi (25%) and the EF (28%) were antidromically activated by LC stimulation. These studies revealed two physiologically distinct subpopulations of LPGi neurons projecting to LC. Other findings described below indicate that there are at least three different neurotransmitter systems in this pathway as well. Taken together, the results of these antidromic stimulation experiments are entirely consistent with our anatomic findings. As LC dendrites extend into a shell-like region surrounding the LC nucleus itself (reviewed below), it is possible that fibers innervating areas adjacent to the LC may contact dendrites of LC neurons. We (AstonJones et al., 1986) have investigated this possibility with physiological techniques. High intensity (2 mA) stimulation of the central nucleus of the amygdala elicited only weak and inconsistent synaptic activation in a few LC neurons. Five other LC neurons were antidromically activated, as expected from the fact that the LC projects to the central nucleus of the amygdala. In contrast to this lack of consistent response in the LC, central amygdala stimulation caused strong, short-latency synaptic activation in 19 of 30 neurons in the adjacent PB area. Similar results have been obtained for another area that heavily innervates the peri-LC region, the Sol (Ennis and Aston-Jones, 1989b). These electrophysiologic studies suggest that the central nucleus of the amygdala and Sol have strong inputs to the PB but not to the LC. In contrast, stimulation of a specific region of the medial prefrontal cortex that also sends projections to the peri-LC region (Zhu and Aston-Jones, 1996) produced consistent activation of LC neurons (Fig. 4) (Jodo and Aston-Jones, 1997; Jodo et al., 1998). It has also been reported that stimulation of a similar area in ketamine-anesthetized rats produced inhibition of LC neurons, indicating that excitatory and (presumably indirect) inhibitory influences from the medial prefrontal cortex on LC activity may exist (Sara and Herve-Minvielle, 1995).

Indirect Afferents to the LC—Defining Afferent Circuitry Knowledge of direct afferents is critical to understanding the function of a brain structure. However,

this is not sufficient, as each input could convey several different types of information depending upon the circuits that the input neurons are enmeshed within. To unravel this circuit-level question of afferent control of the LC, we have begun using the transynaptic retrograde tracer pseudorabies virus (PRV). PRV is a live organism that is transported selectively retrogradely and avidly crosses synaptic junctions to label second and higher order afferents to neurons at the site of the injection. We (Aston-Jones and Card, 2000; Chen et al., 1999) have characterized the use of this tracer with central injections and found that it is a reliable and powerful tool for circuit-level afferent analysis. Focal injection of PRV into the LC yielded labeling in the LPGi and the EF at short survival times, as expected given that these are major direct inputs. However, at longer survival periods (greater than 44 h), neurons were also labeled in (among other sites) the suprachiasmatic nucleus (SCh; Fig. 5) (Aston-Jones et al., 2001). This labeling was particularly intriguing because it suggested the first specific circuit for circadian regulation of sleep/waking and performance. We determined with double labeling that the dorsomedial nucleus of the hypothalamus (DM) is a likely relay from the SCh to the LC. As the DM does not strongly innervate the LC nucleus proper, these results imply that the DM may innervate LC processes in the peri-LC dendritic zone. The role of the DM as an SCh–LC relay was confirmed when lesions of the DM substantially reduced the PRV labeling in the SCh after injection in the LC (Aston-Jones et al., 2001). Additional studies showed that the LC exhibits a circadian rhythm in discharge activity and that this rhythm was eliminated with DM lesions (Aston-Jones et al., 2001). Therefore, the SCh– DM–LC circuit is functionally important for circadian regulation of LC activity and, by implication, also of arousal and cognitive performance. Additional studies are currently underway to test these ideas. These findings are but one result among many intriguing outcomes showing the importance of circuit-level afferent analysis and the utility of the PRV tool in carrying afferent anatomy to this next functionally important step.

Conclusions of Tract-Tracing and Microphysiology Studies Tract-tracing and microphysiology studies reveal that the LPGi and EF are major afferents to the LC. Minor afferents with consistent WGA–HRP retrograde labeling are the dorsal cap of the PA and the intermediate zone of the spinal cord. Additional afferents identified retrogradely with WGA–aopHRP–Au or CTB and confirmed with anterograde tracing include the medial preoptic area and the ventrolateral PAG. Additional areas have

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FIGURE 4 Activation of LC neurons by stimulation of the medial prefrontal cortex in rat. Electrical stimulation was given at three different depths below the surface of the brain (1, 2, and 3 mm) in the prefrontal cortex, corresponding approximately to superficial, middle, and deep locations in the dorsomedial prefrontal cortex along the same stimulation electrode tracks. In the left PSTHs single pulse stimulation is designated by arrows. In the right PSTHs train stimulation was given during the epoch designated by small dots. Bin width in each PSTH was 5 ms. This response may indicate the the prefrontal cortex innervates extranuclear LC dendrites in the peri-LC area (see Fig. 2B). Taken from Jodo et al., 1998.

been identified as afferents to the LC based upon the neurotransmitter contents of fibers, including the tuberomammillary nucleus and the posterior hypothalamus (described below). Many areas previously thought to project to the LC (including the central nucleus of the amygdala, Sol, prefrontal cortex, and dorsal horn of the cord among others) densely innervate pericoerulear regions, but not the LC nuclear core. Some of these (Ace, Sol) have been demonstrated to contact extranuclear LC dendrites. The prefrontal cortex may also innervate these processes as stimulation of this area causes consistent activation of LC neurons. Finally, tracing with transynaptic methods has revealed a circuit from the SCh to the LC. Additional functionally identified circuit

afferents will be defined with these techniques in the near future.

Neurochemical Identity of Afferents to the LC Table 1 lists immunocytochemical studies indicating that the LC is innervated by fibers that stain for a variety of neurochemical markers. It is noteworthy that many of these neurotransmitters proposed to innervate the LC are contained in LPGi and EF neurons. For example, immunocytochemical studies reveal that LPGi and EF neurons stain for markers of adrenaline (Hokfelt et al., 1974; Howe et al., 1980), acetylcholine (Butcher and Woolf, 1984; Kimura et al., 1984), excitatory amino

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FIGURE 5 (Left panels) Labeling of neurons in the suprachiasmatic nucleus (SCh) after injection of pseudorabies virus (PRV) in the LC. Brightfield photomicrographs of frontal sections taken through the SCh at different survival times after PRV injection in the LC, as indicated. The number of labeled neurons in the SCh increased from 44 h (A) to 66 h (C) of survival. Also, labeling was more prominent on the side ipsilateral to the LC injection site (right side in these photographs). Sections were counterstained with neutral red. 3V, third ventricle; ox, optic chiasm. Bar = 200 μm. (Right panels) Histograms showing the distributions of LC firing rates during different epochs of the circadian cycle. Three paired groups of rats were maintained in either 12-h/12-h dark/light (A and C) or 24-h darkness (B). One paired group received bilateral ibotenic acid lesions of the DM (C). Impulse activity was recorded in LC neurons during either the dark or light photoperiod (A and C) or the active or inactive epoch of the rat’s circadian cycle in 24-h darkness (B). There was a significant difference in LC firing rates in animals taken from their dark vs light periods (A) and from their active vs inactive periods when maintained in continuous darkness (B; P<0.0001, each). Lesions of the DM eliminated the difference in LC firing rates during the dark vs light photoperiods (C). The scale on the y axis only corresponds to the histogram bars. The solid lines on each of the graphs represent best-fit normal distribution curves for the histogram data. Taken from Aston-Jones et al., 2001.

acids (Forloni et al., 1987; Toomin et al., 1987), enkephalin (Hokfelt et al., 1977; Khachaturian et al., 1983), CRH (Swanson et al., 1983), substance P (Ljungdahl et al., 1978), 5HT (Steinbusch, 1984), and GABA (Mugnaini and Oertel, 1985). We have performed singlelabeling immunohistochemical screening studies for the locations of several of these neurotransmitter markers in the LPGi and the EF, and we find that neurons staining for phenylethanolamine N-methyltransferase (PNMT, a marker for adrenergic neurons), 5-HT, enkephalins, substance P, GABA/GAD, ChAT, and dynorphin are located in regions that also contain neurons labeled retrogradely from the LC in other studies. Thus, it is possible that multiple neurotransmitter systems arise from neurons in either or both of these areas. Indeed, studies discussed below provide direct evidence for adrenergic, glutamatergic, GABA, and CRH inputs from the LPGi. It should be borne in mind that at least some of the neurotransmitter diversity in the afferent innervation of the LC may stem from colo-

calization of neurotransmitters, e.g., neuropeptide Y is colocalized with adrenergic neurons of the C1 group in the LPGi (Blessing et al., 1986). Here we review evidence for neurotransmitter inputs to LC neurons based upon the neuropharmacology of responses of LC neurons to stimulation of afferent sources and upon double-labeling anatomical studies employing retrograde transport combined with immunohistochemistry of neurotransmitter markers. Pharmacology of Inputs to the LC Excitatory amino acid input from the LPGi Singlepulse electrical stimulation of the LPGi elicits predominantly short-latency phasic activation of LC neurons; a minority of LC neurons is inhibited by such stimulation (described below). Intracerebroventricular (icv) or local intracoerulear microinfusion of non-NMDA-type excitatory amino acid (EAA) antagonists strongly attenuated or completely blocked LPGi-induced excitation in all LC neurons tested (Ennis and Aston-Jones, 1986,

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1988). In addition to blocking LPGi responses, these EAA antagonists simultaneously attenuated footshock (sciatic nerve stimulation)-evoked excitation of LC neurons. Inhibitory adrenergic input from the LPGi As noted above, some LC neurons were inhibited by LPGi stimulation. To determine whether C1 adrenergic neurons located in the LPGi were involved in this inhibitory projection, we characterized the effects of the α2 adrenoceptor antagonist idazoxan (IDA) on inhibitory responses of LC neurons to LPGi stimulation (Aston-Jones et al., 1992). Intravenous administration of IDA attenuated LPGi-evoked inhibition and often revealed an underlying weak excitation. Intraventricular administration of EAA antagonists eliminated excitation from LPGi and disclosed an underlying inhibitory response in many LC neurons that were previously excited by LPGi stimulation. These results revealed that about 90% of LC neurons receive inhibition from the LPGi. Intravenous or intracoerulear IDA significantly reduced such LPGi-evoked inhibition, completely blocking this response in many LC cells tested. These results indicate that C1 adrenergic neurons in the LPGi provide a direct inhibitory input to the great majority of LC noradrenergic neurons. These findings are compatible with our anatomical results showing that the C1 neurons in the LPGi provide the bulk of adrenergic innervation of the LC (described below) and that most or perhaps all LC cells receive both EAA and adrenergic inputs from this area. Inhibitory GABA input from the EF In contrast to the predominant excitation of LC from LPGi, stimulation of the EF yielded nearly uniform inhibition of LC activity (Ennis and Aston-Jones, 1989a; Ennis and AstonJones, 1989b). The GABA antagonist picrotoxin significantly attenuated inhibition of the LC from the EF, completely blocking such inhibition in most cells. Iontophoretically applied or microinfused bicuculline (BIC) blocked EF-evoked inhibition in nearly all LC cells tested. Local application of this agent into the LC did not affect footshock-evoked excitation of LC neurons and had no overall effect on postactivation inhibition following such excitation. Interestingly, continuous iontophoretic application of BIC alone increased the mean spontaneous discharge rate of LC neurons, suggesting a tonic GABAergic input in vivo. In contrast to the effect of the GABA antagonists picrotoxin or BIC, iontophoretic application of the glycine antagonist strychnine hydrochloride did not effect EFevoked inhibition of LC discharge. Hypocretin/orexin input to the LC As noted above, recent anatomical studies revealed a dense innervation

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of LC by hypocretin/orexin fibers. The sole source of this peptide is in the lateral and dorsal hypothalamus (de Lecea et al., 1998; Sakurai et al., 1998). Our studies in brain slices have revealed that this peptide strongly excites LC neurons directly, so that excitation is observed in the absence of synaptic transmission in the slice preparation (Fig. 3) (Horvath et al., 1999). This response appears to be due to inhibition of a resting “leak” potassium conductance in LC cells (Ivanov and Aston-Jones, 2000). Double Labeling Anatomical Studies Glutaminase-positive afferents to the LC from the LPGi We (Drolet and Aston-Jones, 1991) combined WGA–apoHRP–Au retrograde tracing with immunohistochemistry for phosphate-activated glutaminase, a purported marker of glutamatergic neurons. Numerous glutaminase-immunoreactive cell bodies were observed in the LPGi. Double-labeling revealed that such glutaminase-positive neurons represent a substantial proportion of the LC-projecting neurons in the LPGi. This provides anatomical support of the above pharmacological evidence for a major EAA input to the LC from the LPGi. Adrenergic inputs to the LC from C1 cells in the LPGi The LC is densely innervated throughout with PNMT-immunoreactive fibers (Berod et al., 1984; Hokfelt et al., 1974; Milner et al., 1989; Pieribone and Aston-Jones, 1991; Pieribone et al., 1988). In fact, the LC appears to be the primary target in the pons for adrenergic fiber innervation. In ultrastructural studies, PNMT-immunoreactive varicosities are numerous and make conventional symmetric and asymmetric synapses onto dendrites of LC neurons (Milner et al., 1989). We combined retrograde transport with immunohistochemistry for PNMT to identify the source of adrenergic afferents to the LC (Pieribone and AstonJones, 1991; Pieribone et al., 1988). Double-labeled LC afferent neurons were located among both C1 and C3 adrenergic neurons in the LPGi and the EF, respectively. No afferent neurons were found in the vicinity of the C2 adrenergic cell group in the Sol or the dorsal motor nucleus of the vagus, confirming our previous results with WGA–HRP (Aston-Jones et al., 1986; described above) Overall, approximately 21% of LC afferent neurons in LPGi are PNMT-positive (C1 neurons), while only 4% of LC afferents in the EF (C3) are PNMT-positive (Pieribone and Aston-Jones, 1991). This study also identified two pathways for PNMTimmunoreactive fibers to innervate the LC. Fibers from neurons in both C1 and C3 were found to enter the medullary adrenergic bundle and to ascend within this bundle to innervate the LC. This is consistent with

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previous results that electrolytic lesions of this adrenergic bundle in the rostral medulla cause an almost total loss of adrenergic fiber innervation of the LC (Astier et al., 1987). Fibers from the C3 cell group were found to travel on the ventral surface of the IVth ventricle and then distribute laterally to innervate the LC and surrounding areas. Both in vivo iontophoretic studies and in vitro experiments (discussed above) conclude that adrenaline potently inhibits the spontaneous activity of LC neurons via α2 receptors (Cedarbaum and Aghajanian, 1976; Williams et al., 1985). Together with the above anatomic data, these studies indicate that adrenergic neurons, most prominently in the C1 cell group, exert an α2-mediated inhibitory effect on LC neurons. We confirmed that stimulation of the C1 area inhibits LC neurons via α2 receptor activation (Aston-Jones et al., 1992). Serotonergic inputs to the LC The LC receives a dense serotonin innervation as seen with autoradiography for serotonin uptake sites (Leger and Descarries, 1978), tryptophan hydroxylase immunoreactivity (Pickel et al., 1977), and serotonin immunoreactivity (Steinbusch, 1984). Ultrastructural examination of tryptophan hydroxylase immunoreactivity (Pickel et al., 1977), 5HT (Van Bockstaele, 2000) and serotonin uptake sites (Leger and Descarries, 1978), in LC indicated that many of these terminals may not form classical synapses within the LC. No obvious fiber pathway to the LC is evident in serotonin-stained material and the source of serotonin innervation to the LC has been controversial. Lesions of the dorsal raphe reduced biochemical markers and uptake sites for serotonin in micropunches containing the LC and surrounding areas, leading Pujol and coworkers to propose that such innervation derives from the dorsal raphe complex (MacRae-Degueure and Milon, 1983; Pujol et al., 1978). However, labeled cells were not consistently observed in the dorsal raphe or raphe magnus following WGA–HRP (Aston-Jones et al., 1986), Fluorogold (Pieribone and Aston-Jones, 1988), CTb (Luppi et al., 1991), or WGA–apoHRP–Au injections restricted to the nucleus LC (unpublished observations). Furthermore, in preliminary studies injections of WGA–HRP or PHA-L into the dorsal raphe have failed to produce anterograde transport into the LC. In addition, serotonergic fiber innervation of the LC remains apparently undiminished following electrolytic lesions of the dorsal raphe area (Pieribone et al., 1989). Thus, our results disclose no substantial serotonin input to the LC from the dorsal raphe. In other preliminary studies, serotonergic neurons in the lateral wings of the B9 area were retrogradely labeled with CTb from the LC (Luppi et al., 1991). However, as noted above, afferents revealed by this tracer require anterograde

labeling for confirmation, which has not yet been conducted for this possible pathway. Recent studies reveal that although serotonin has no consistent effect on spontaneous LC discharge, this agent dramatically reduces glutamatergic, but not cholinergic, activation of LC neurons (Aston-Jones et al., 1991a–1991c). GABAergic inputs to the LC Several studies (Berod et al., 1984; Ijima and Ohtomo, 1988; Ottersen and StormMathisen, 1984a, 1984b) have identified dense glutamic acid decarboxylase (GAD)- or GABA-immunoreactive fiber staining in the LC. We have confirmed such findings with both light and electron microscopy. There is strong physiologic and pharmacologic evidence that at least a part of the GABAergic innervation of the LC arises from GABA neurons in the EF (Ennis and AstonJones, 1989a, 1989b, reviewed above) and may be responsible for a short-latency synaptic potential recorded in vitro (Cherubini et al., 1988). Additionally, in unpublished anatomical studies we have found a restricted population of GABA-positive neurons in the EF region that is a major source of LC afferents. In studies using retrograde transport of WGA–apoHRP–Au or CTb from the LC combined with immunohistochemistry for GABA or GAD, we have found that many LC-projecting neurons in the EF are putatively GABAergic (unpublished observations). Paradoxically however, electrolytic lesions of the EF have failed to eliminate GAD-immunoreactive terminals in the LC (Shipley, Ennis, and Aston-Jones, unpublished). This raises the possibility that some GABA terminals present in the LC do not arise from the EF but rather from other sources. In this regard, it is noteworthy that GAD- and GABA-positive neurons are found in the LPGi area that also contains LC afferent neurons (Ruggiero et al., 1985; Mugnaini and Oertel, 1985; AstonJones, unpublished). In addition, there are numerous GABAergic neurons in the adjacent peri-LC region (Jones, 1991). We have recently determined that many such putatively GABAergic neurons become retrogradely labeled after injection of retrograde tracers into the LC (Aston-Jones et al., 2001). We have also confirmed previous reports (Ijima and Ohtomo, 1988) for a population of small GABA-staining neurons located within the LC nucleus proper. These results indicate that neurons within the LC or peri-LC region may also provide a substantial GABAergic innervation of the LC. As expected, GABA has a strong inhibitory effect on LC neurons (Shefner and Osmanovic, 1991; also see above). Luppi and colleagues recently found that bicuculline, an antagonist of GABA-A receptors, disinhibits LC neurons during sleep (Gervasoni et al., 1998). This confirms previous proposals that LC neurons are actively inhibited during sleep (Aston-Jones and Bloom, 1981a,

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1991b), and identifies at least one possible behavioral function of the strong GABA input to LC neurons. Enkephalin inputs to the LC There is abundant evidence for opioid inputs to the LC. Numerous fibers within the LC nucleus stain with an antibody raised against enkephalin. Enkephalinergic terminals are reported to make synapses upon neurons in the LC (Pickel et al., 1979), and opioid binding sites are dense in the LC (Pert et al., 1976). In addition, LC spontaneous activity is potently suppressed by directly applied opiates (Bird and Kuhar, 1977; Guyenet and Aghajanian, 1979; Williams and North, 1984); these effects are mediated primarily through μ receptors on LC neurons (Williams and North, 1984). Our studies using an antibody directed against the extended arg–gly–leu–met–enkephalin molecule (derived from the enkephalin precursor, proenkephalin A) have revealed that the enkephalin innervation of the LC is denser than previously appreciated (Drolet et al., 1992). Enkephalin-positive fibers are especially dense in the rostral and ventromedial pericoerulear regions containing extranuclear LC dendrites, raising the possibility that distal dendrites of LC neurons also receive enkephalin inputs. Our retrograde tracing with WGA–apoHRP–Au combined with enkephalin immunohistochemistry revealed that a substantial percentage (more than 50%) of LC-projecting neurons in the LPGi and EF contain enkephalin (Drolet et al., 1992). These findings have been confirmed and extended by recent work at the ultrastructural level. Van Bockstaele and colleagues found that met– or leu–enkephalinpositive terminals make synaptic contacts with THpositive dendrites in the LC (Van Bockstaele et al., 1995; Van Bockstaele and Chan, 1997); some of these LC neurons were identified as cortically projecting (van Bockstaele et al., 1996d). Additional studies have localized μ receptors on the dendrites of LC neurons, both outside of apparent synaptic contact (Van Bockstaele et al., 1996b), and in apposition to leu–enkephalinpositive terminals (Van Bockstaele et al., 1996c). This group also found δ opioid receptors on presynaptic axon terminals in the LC (van Bockstaele et al., 1997). Finally, there is evidence (as noted above) that enkephalin is colocalized in both GABAergic (Van Bockstaele and Chan, 1997) and glutamatergic inputs to LC neurons (Van Bockstaele et al., 2000). The significance of these opioid peptide projections to the LC is the subject of ongoing research. One recent study found that stress induces increased endogenous opioid inhibition of LC neurons (Curtis et al., 2001), consistent with previous work where endogenous opioid influences on LC neurons were only found under stressful conditions (Abercrombie and Jacobs, 1988).

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CRH inputs to the LC Using an antibody directed against rat/human CRH, Valentino et al. (1992) found numerous CRH-positive fibers in the LC and CRHpositive cells in areas that contain LC afferents. WGA– apoHRP–Gold retrograde tracing combined with immunohistochemistry revealed that a population of LC-projecting neurons in the LPGi stain for CRH. A few doubly labeled neurons were also found in the dorsal cap of the paraventricular hypothalamic nucleus. In addition, there are many CRH cells in the pericoerulear area, raising the possibility that local pericoerulear neurons contribute to the CRH innervation of the LC. Indeed, we found that Barrington’s nucleus, a peri-LC structure densely populated with CRH neurons, send many CRH inputs to the LC (Valentino et al., 1996). No LC NE neurons were observed to stain for CRH. Ultrastructural studies found that CRH fibers directly terminate on LC processes (Van Bockstaele et al., 1996a). Some of the CRH innervation of LC processes in the rostral peri-LC region originates from the central nucleus of the amygdala (Van Bockstaele et al., 1996a; Van Bockstaele et al., 1998a). Recent work has found direct evidence for CRH-1 receptor localization in NE neurons of the rodent LC (Sauvage and Steckler, 2001), confirming that CRH has direct effects on the activity of LC cells. Effects of CRH, and putative activation of CRH afferents, on LC activity are described elsewhere (Valentino, 1989; Valentino et al., 1991). Ultrastructural Studies of Inputs to LC Neurons Shimizu and Imamoto (Shimizu and Imamoto, 1970; Shimizu et al., 1979) and Groves and Wilson (Groves and Wilson, 1980a) provided the first systematic studies of the ultrastructure of the LC. LC neurons have abundant cytoplasmic organelles including a highly developed Golgi apparatus that extends into the dendrites. Most afferent terminations in the LC nucleus synapse on dendrites ranging between 0.5 and 2.5 μm in diameter and onto spine-like appendages on dendrites and cell bodies. The majority of these synapses fell into four categories: (i) synapses with small round, densely packed vesicles (41%); (ii) those with large rounded vesicles (20%); (iii) synapses with large flattened vesicles (23%); and (iv) those with numerous small flattened vesicles (11%). The remaining 5% had mixtures of these and/or contained dense-core vesicles. In a second study, Groves and Wilson (1980b) found that 5-hydroxydopamine labeled a small percentage of all these synaptic types; these results were taken as indirect evidence for monoaminergic synapses of various types. In early studies, Pickel and co-workers (Pickel et al., 1979) demonstrated substance P- and met–enkephalincontaining axons and terminals in the LC although they

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did not use double-labeling to confirm that these peptidergic inputs synapsed onto noradrenergic neurons. This could be important because numerous noncatecholaminergic dendrites are present within the LC nucleus; these dendrites presumably derive from pericoerulear neurons. Milner et al. (1989) demonstrated a high percentage of synapses in the LC that were immunoreactive for PNMT. This study supported previous light microscopic and neurophysiological evidence for an adrenergic projection to LC neurons from the ventrolateral medulla. More recent electron microscopic studies by Van Bockstaele and colleagues elucidate afferentation of LC neurons in greater detail. Several of these studies are reviewed previously under sections corresponding to specific transmitter inputs. In addition, analyses of anterograde transport of biotinylated dextran amine (BDA) from the LPGi found that this structure monosynaptically innervates NE processes in the LC and peri-LC region (Van Bockstaele et al., 1998), confirming previous tract-tracing studies (Aston-Jones et al., 1986). Ultrastructural studies have also revealed that the bed nucleus of the stria terminalis (BST) innervates extranuclear dendrites of LC neurons in the rostral peri-LC region (Van Bockstaele et al., 1999b). Similar experiments have also demonstrated projections from the NTS (Van Bockstaele et al., 1999a) and from the ventrolateral PAG to extranuclear LC dendrites in the periLC zone (Bajic et al., 2000). Other Putative Neurotransmitter Candidates Innervating the LC The inputs reviewed above represent the most thoroughly investigated neurochemical afferents to the LC. As indicated in Table 1, a variety of other neurotransmitter markers are found in fibers that innervate the LC, but the sources of these markers have not been identified.

Glia in the LC Recent studies have shown that the LC is richly endowed with glia and that astroglia in particular intimately comingle with NE neurons and processes within the LC (Alvarez-Maubecin et al., 2000). These studies also found that glia and neurons in the LC were functionally connected through electrotonic coupling. The additional finding that neurons and glia stained for the molecular substrate of electrotonic coupling, connexin proteins 26 and 32, supported this result. Although reported only for neonatal animals, these findings support prior observations for electrotonic coupling in the LC (described above) and extend those results to also include coupling between LC neurons and astroglia.

Conclusions The results concerning afferent regulation of the LC mandate the need to know more about the anatomy and physiology of the LPGi and EF, as well as other sources of inputs. The integrative functions of these areas are critical for understanding the functional role of the LC in brain and behavioral processes. The set of activities represented in the LPGi and EF are diverse, but cluster around functions involved in autonomic regulation, sensory processing, and behavioral orientation (reviewed in Aston-Jones et al., 1991c). This is particularly significant for understanding LC function since as reviewed above (Aston-Jones and Bloom, 1981a, 1981b) the most intense output of the LC reliably corresponds to sympathoexcitatory stimuli and behavioral orienting responses. In addition, it has become clear in the last few years that the extranuclear dendrites of LC neurons are also important to understand, as they receive afferents that do not directly innervate the LC nucleus proper. This “extranuclear LC” area is reviewed in more detail below.

THE PERICOERULEAR REGION: THE “EXTRANUCLEAR LC” Inputs to the Pericoerulear Region It has been commonly believed that the region surrounding the LC in the rat is largely composed of fiber tracts with few if any neurons. However, our work (Aston-Jones et al., 1986, 1991c; Shipley et al., 1996) along with that of other groups (e.g., Cechetto et al., 1985; Saper, 1982, 1987; Sesack et al., 1989; Wallace et al., 1989) has shown that several brain areas contribute dense terminal fields to the neuropil immediately adjacent to the cell bodies that make up the nucleus LC proper (see below). There are dense inputs to the PB area lateral to the LC from the Sol (Mantyh and Hunt, 1984; Ennis and Aston-Jones, 1989a, 1989b; Van Bockstaele et al., 1999a), the dorsal horn of the spinal cord (Cechetto et al., 1985; Standaert et al., 1986), and the central nucleus of the amygdala (Aston-Jones et al., 1986; Van Bockstaele et al., 1996a; Wallace et al., 1989) among others. The pontine gray rostral, medial, and ventral to the LC, on the other hand, receives inputs from the frontal cortex (Hurley et al., 1991; Sesack et al., 1989; Zhu and Aston-Jones, 1996), the dorsal raphe (Bobillier et al., 1979; Conrad et al., 1974; Pieribone et al., 1989), the central nucleus of the amygdala (Aston-Jones et al., 1986; Wallace et al., 1989), and the VLTg (Deutch et al., 1986). In addition, we have shown that there is a dense projection to the rostromedial peri-LC region from the medial preoptic area (MPA) (Rizvi et al., 1994). This

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MPA input also terminates heavily in the region between the medial edge of the LC and the ependymal layer of the IVth ventricle. A few fibers from the MPA enter the nuclear core of the LC but the preponderance of the fibers terminates among the extranuclear dendrites of LC neurons (see below). Our recent studies also demonstrate input to this rostromedial peri-LC region from the DM. Thus, injections of retrograde tracers into this area produce labeled neurons in the DM, whereas similar injections confined to the LC nuclear core produce little labeling in the DM. Our recent results indicate that this input is an important relay for circadian information reaching the LC (Aston-Jones et al., 2001) (summarized above). This same rostroventromedial pericoerulear region is heavily targeted by fibers containing several peptides and includes Barrington’s nucleus which is a dense cluster of CRH neurons. Many of these neurons project into the LC proper (Valentino et al., 1996).

Architecture of the Pericoerulear Region These findings cast doubt on the view that the pericoerulear area is a relatively cell-free area and prompted an analysis of the neuropil in the region surrounding the “nuclear core” of the LC. Several interesting observations have emerged. In addition to Barrington’s nucleus and the laterodorsal tegmental nucleus, already well documented by others, we have found that the pericoerulear area immediately rostral, ventral, and medial to the LC proper contains many neurons which vary in size, shape, and dendritic orientation; these neurons do not fall easily into the described boundaries of Barrington’s or the lateral dorsal tegmental nuclei. Thus, inputs to areas immediately adjacent to the LC, particularly those in this rostroventromedial pericoerulear area, could terminate on these unidentified neurons. On the other hand, additional studies of this pericoerulear region have revealed that it also contains a dense, focal plexus of “extranuclear” LC dendrites (Shipley et al., 1996). This extranuclear dendritic plexus, which may be targeted by some or all of the discrete terminal systems found in this rostroventromedial pericoerulear area, is discussed next.

Extranuclear Dendrites of LC Neurons Several reports have shown that processes of LC neurons extend for a few hundred micrometers outside of the nuclear core of the LC (Cintra et al., 1982; Grzanna and Molliver, 1980; Grzanna et al., 1980; Shimizu and Imamoto, 1970; Shimizu et al., 1978; Swanson, 1976). However, in most cases it has not been clear whether these extranuclear processes are dendrites, axons, or

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both. Moreover, it is not known whether these “extranuclear dendrites” arise mainly from those LC neurons situated at the periphery of the nucleus or if the majority of LC neurons have extranuclear dendrites. Using gold–silver intensification (Gallyas et al., 1982) of dopamine β-hydroxylase (DBH) immunohistochemically stained material, we found that extranuclear LC processes have a remarkable degree of spatial organization. The vast majority of these processes ramify in two distinct, focal pericoerulear zones—rostroventromedial to the LC and a narrow strip that extends from the caudal juxtaependymal boundary of the LC closely apposed to the ependymal cell layer of the IVth ventricle. At the light microscopic level, many of these processes appeared to be obvious dendrites; others, however, were very thin and beaded, and could be axons. To unambiguously identify the nature of these extranuclear processes, we analyzed DBH- and tyrosine hydroxylase (TH)-stained material at the EM level (Shipley et al., 1996). Of more than 500 labeled processes in the rostroventromedial and caudal juxtaependymal peri-LC zones, all but three labeled processes were dendrites. We also found that these extranuclear LC dendrites are heavily targeted by noncatecholaminergic afferent synapses. Thus, LC neurons have an appreciable postsynaptic surface that lies a considerable distance outside the confines of the nucleus proper. Moreover, this “receptive surface” is preferentially distributed in two discrete peri-LC zones. These studies could not resolve whether these extranuclear dendrites arise primarily from a subset of LC neurons (e.g., those located near the periphery of the nucleus) or whether extranuclear dendrites were a characteristic feature of most LC neurons. To address this issue, we studied the morphologies and dendritic domains of individual LC neurons filled with biocytin in in vitro tissue slices (Shipley et al., 1990). Filled LC neurons had remarkably similar morphologies, with four or five major dendrites extending from the soma in all directions and a single thin process (presumably the axon) often emanating from a proximal dendrite. As shown in Fig. 6, dendrites of individual LC neurons typically exited the nuclear core of LC at its medial, rostral, or caudal boundary and extended for up to 500 μm into the rostral, medial, or caudal pericoerulear region. Most commonly, dendrites that exited the lateral margin of the nucleus coursed for ≤100 μm and then made an abrupt turn to extend in a rostral or caudal direction. These extranuclear dendritic fields were characteristic of the majority of LC neurons independent of the location of their cell body in the nucleus. Thus, individual LC neurons typically have extranuclear dendrites that extend into one or both of the preferred extranuclear dendritic zones.

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FIGURE 6 Extranuclear dendrites of LC neurons. (A) Horizontal section through the LC, showing a reconstruction of a single biocytin-filled LC neuron. The majority of the dendrites of LC neurons extend outside the LC nucleus proper into two focal zones: the rostromedial and the caudomedial pericoerulear regions. The dendrites of the LC neuron shown here extend into the rostromedial pericoerulear area. Arrows indicate the axon of this cell. At the bottom right is a drawing of the horizontal section. Rostral is at the top; medial is at the left. (B) Electron micrograph of extranuclear LC dendrites (D) contacted by synapses containing presynaptic vesicles (pv). Arrowheads indicate symmetrical synapses; the arrow indicates an asymmetrical synapse. LC dendrites labeled here by antibodies to DBH are located in the rostromedial pericoerulear region.

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The fact that several brain areas (see above) project densely and selectively into the zones containing these extranuclear LC dendrites raises the interesting possibility that some of these inputs synapse with the extranuclear dendrites of LC neurons. Both the LPGi and the EF, the two major inputs to the nuclear core of the LC, also project to these two pericoerulear dendritic zones and, thus, may terminate both on proximal and more distal LC dendrites. However, the rostroventromedial peri-LC receives input from several brain areas (frontal, insular, and perirhinal cortices; amygdala; MPA; dorsal raphe; and VLTg) that do not appreciably innervate the nuclear core of the LC. Recent studies (reviewed above) reveal that some of these areas directly innervate extranuclear LC dendrites. Although inputs to the soma-rich LC nucleus might exert a greater influence on LC discharge than inputs to extranuclear dendrites, the later inputs could significantly regulate LC activity.

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grade tracer pseudorabies virus (PRV) into the LC proper (Aston-Jones et al., 2001). This tracer also has a very limited diffusion, due to the large size of the viral particles and the high affinity for extracellular neuronal membrane molecules, and produces no apparent halo around the LC injection site (Aston-Jones and Card, 2000; Chen et al., 1999). One important property of this tracer is that it is transported through the length of the dendrites of infected neurons and transsynaptically labels inputs to distal dendrites (Aston-Jones and Card, 2000; Card et al., 1993). Therefore, results with focal PRV injections in the LC nuclear core include neurons that innervate distal extranuclear LC dendrites. We hypothesize that at least some of the neurons labeled with PRV in these studies in the peri-LC region may serve as interneurons and innervate distal LC dendrites. Note that many of these peri-LC neurons that are labeled after LC injections of either WGA– apoHRP–gold or PRV stain for GABA or GAD (AstonJones et al., 2001).

Pericoerulear Inputs to LC? As noted above, there are many neurons in the pericoerulear area. However, it is not known if any of these neurons innervate the LC, i.e., if there are “local” afferents to the LC. Our previous studies using injections of retrograde tracers into the LC (Aston-Jones et al., 1986) could not adequately address this issue because the injected tracer, WGA–HRP, produced an injection “halo” in the pericoerulear area, particularly in the rostromedial pericoerulear zone. We were able to tentatively rule out inputs from the PB area, but possible inputs from rostromedial pericoerulear areas were not analyzed. Indeed, results using the tracers PHA-L and WGA–apoHRP–Gold suggest that pericoerulear neurons may innervate the LC. One region in which retrogradely labeled cells were often found within the injection halo of WGA–HRP cases was the caudal ventrolateral PAG, rostral to the LC. Anterograde transport of PHA-L from the caudal ventrolateral PAG revealed prominent innervation of the central gray ventromedial to LC and scattered light innervation of the LC as well (Ennis et al., 1991). Stimulation this PAG region strongly influences pericoerulear neurons adjacent to the LC and has a moderate influence on LC neurons (Ennis et al., 1991). Recently, we found that injections of WGA– apoHRP–Gold into the LC, which produce very focal injections lacking a “halo,” yielded retrograde labeling in this peri-LC dendritic zone, indicating that at least some cells in this area may project into the LC nucleus proper. In additional studies, we found that many small neurons in the peri-LC dendritic zone were labeled following injections of the transynaptic retro-

EFFERENT PROJECTIONS OF LC NEURONS Ascending Projections Olfactory Structures The olfactory bulb and olfactory cortex compose a major fraction of the telencephalon in rats and other macrosomatic animals. In a quantitative analysis of LC NE innervation of the olfactory bulb (Shipley et al., 1985), it was determined that at least 40% of all LC neurons project to the bulb. Actually, this number may be in error on the low side as counts of labeled LC neurons were limited to those cases where injections of the retrograde tracer WGA–HRP were restricted to the rostral half of the main olfactory bulb in order to rule out any cells labeled by tracer that might spread into the caudally adjacent accessory olfactory bulb or anterior olfactory nucleus. Retrograde labeling was not found in any other structures containing NE neurons, thus it was concluded that the LC is the sole source of NE innervation of the bulb. This conclusion was supported by a subsequent study using anterograde tracing methods and DBH immunocytochemistry which demonstrated that LC NE fibers provide a dense and remarkably specific innervation of both the main and accessory olfactory bulbs (McLean et al., 1989). LC NE axons and varicosities form a plexus that is especially dense in the granule cell and internal plexiform layers of the main olfactory bulb. A sparser plexus of fibers innervates the more superficial external plexiform layer but the most superficial cellular layer

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of the bulb—the glomerular layer—is essentially devoid of NE fibers. This highly selective innervation pattern is unusual for the LC NE system. However, as the identical LC NE innervation pattern was observed in the accessory olfactory bulb which, though architectonically similar to the main olfactory bulb, has entirely distinct afferent and efferent connections, it was concluded that the LC NE system provides a uniquely laminar-specific innervation of the main and accessory olfactory bulbs. As the laminar segregation of specific neuron types in the olfactory bulb is more specific than that in any other cortical structure in the rat, the observed selective termination of LC NE fibers in the granule cell, internal plexiform, and (to a lesser extent) external plexiform layers suggests that the most likely cellular target of LC NE fibers in both the main olfactory bulb and the accessory olfactory bulb is the granule cells. Recent neurophysiological evidence indicates that LC projections may also terminate on mitral cells in the bulb. These experiments in slices of the bulb revealed that NE fibers act directly on mitral cells at α1 adrenoceptors to increase an “up” state and enhance responses to weak inputs (Hayar et al., 2001). This suggests a mechanism whereby LC stimulation in vivo increases responses of mitral cells to low-level olfactory nerve inputs (Jiang et al., 1996). In contrast to the olfactory bulb, much less is known about LC NE innervation of the set of structures known collectively as the primary olfactory cortex, which is directly innervated by the main and accessory olfactory bulbs. While earlier studies using the histofluorescence method leave little doubt that the primary olfactory cortex is richly innervated by catecholaminergic fibers, far less is known about the specific contributions by LC and/or non-LC NE fibers or by DA inputs from the brain stem. NE fibers richly innervate the entire rostrocaudal extent of the primary olfactory cortex, which stretches from the caudal end of the olfactory bulb throughout all the piriform, periamygdaloid, and lateral entorhinal cortices. NE fibers show a moderate to strong degree of laminar selectivity in these temporal cortical fields, strongly innervating the superficial half of layer I (layer Ia) and the middle and deep parts of layer III. Layer Ib has the sparsest innervation and layer II and the superficial part of layer III are intermediate. This pattern of innervation gives fewer clues about the possible neuronal targets of LC NE targets in olfactory cortex than in the olfactory bulb. Neocortex and Hippocampus Neocortex Shortly following the demonstration of NE projections from the brain stem to the cerebral cortex (Fuxe et al., 1968), the LC was identified as the

sole source of this cortical afferent (Jones and Moore, 1977; Mason and Fibiger, 1979; Ungerstedt, 1971). Later studies revealed substantial projections of the rat LC to all areas and spanning all layers of the neocortex (Levitt and Moore, 1978; Morrison et al., 1978, 1979). In the rat the pattern of LC axon distribution in cortex varies little between different areas of cortex. The general features of the coeruleocortical projections in the rat are described by Morrison et al. (1978). Layer I is characterized by a dense grid-like pattern of axons running tangentially to the cortical surface. Layers II and III contain mostly radial fibers while layer IV contains short and somewhat tortuous axons. Layers V and VI contain numerous axons oriented predominantly in the anteroposterior direction. Retrograde transport studies established that the coeruleocortical projection in the rat is almost exclusively ipsilateral with only 5–10% of LC neurons projecting to the contralateral neocortex (Ader et al., 1980; Room et al., 1981; Simpson et al., 1997; Waterhouse et al., 1993). In agreement with the finding of Morrison et al. (1981) concerning the rostrocaudal course of LC axons within the cortex, Loughlin et al. (1982) showed that coeruleocortical axons collateralize most heavily in the anterior–posterior dimension within the cortex and only to a limited extent in the mediolateral dimension. A study by Sakaguchi and Nakamura (1990) found that over 61% of randomly sampled LC neurons were antidromically activated from neocortical sites. Intracortical trajectory of LC axons By using dopamine β-hydroxylase immunocytochemistry in combination with intracortical lesions, Morrison et al. (1981) analyzed the course of LC axons within rat cortex. Based upon their findings these authors proposed the existence of a tangential intracortical course of the coeruleocortical projection: LC axons that enter the cortex through the frontal pole travel caudally within the gray matter forming a sheet of fibers that extends tangentially across cytoarchitectural fields of cortex. Hippocampus LC projections account for the entire NE input to the hippocampal formation. LC neurons projecting to the hippocampus are found primarily in the dorsal one-third of the LC (Loughlin et al., 1986a, 1986b; Mason and Fibiger, 1979) and originate in fusiform cells. LC axons reach the hippocampal formation via three distinct routes: (i) the ventral amygdaloid pathway, (ii) the fornix, and (iii) the cingulum bundle. Some authors hold that the projection originates predominantly in the larger, multipolar LC neurons (Loughlin et al., 1986a, 1986b; Loy et al., 1980), although another study (Haring and Davis, 1983) found exclusively

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small fusiform neurons labeled in the dorsal LC after injections of retrograde tracers into the dorsal parts of the dentate gyrus. Fibers traveling in the ventral amygdalofugal bundle innervate the entire hippocampal gyrus and the midseptotemporal and ventral regions of the dentate gyrus (Haring and Davis, 1983); fibers entering via the fornix innervate the septal pole of the dentate gyrus; and fibers entering from the cingulum bundle innervate mainly the ventral dentate gyrus.

out as perhaps the only major brain region essentially devoid of NE fibers. However, recent studies reveal a considerable NE innervation in the ventral striatum, particularly in the caudal medial shell of the nucleus accumbens (Berridge et al., 1997; Delfs et al., 1998). It is notable that this NE innervation originates primarily from the A2 and A1 cell groups of the caudal medulla, with relatively little contribution from the LC or other NE cell groups (Delfs et al., 1998).

Thalamus

Most areas of the basal forebrain receive at least moderate NE innervation. For example, NE inputs ramify throughout the substantia inominata. These terminals are relatively large in diameter and make asymmetric endings primarily on dendrites (Chang, 1989). One basal forebrain system that has received considerable attention, particularly in conjunction with the increased interest in Alzheimer’s disease, is the magnocellular cholinergic system including the medial septum, the nucleus of the diagonal band and the nucleus basalis. All these structures receive a moderate to strong NE innervation (Zaborszky and Cullinan, 1996). Based on a variety of tract-tracing and immunocytochemical studies, it appears that the majority of these NE fibers may derive from the LC. This suggests that the LC NE system influences the magnocellular cholinergic basal forebrain system although it must be recognized that the cholinergic neurons of the basal forebrain system are embedded in structures that are complex and heterogeneous, containing multiple neurotransmitters/peptides other than ACh. Relatively few studies have examined specific LC NE influences on the cholinergic system. An exception to this is the work of Zaborszky and Cullinan (1996) who have provided biochemical and EM immunocytochemical evidence for a direct influence of NE terminals on identified basal forebrain cholinergic neurons. Specifically, 6-hydroxydopamine lesions of ascending catecholaminergic fibers cause a reduction in ChAT levels in basal forebrain areas containing cholinergic neurons. The LC NE system does not appear to selectively target cholinergic neurons, however, as the entire septal complex receives a moderate to strong NE innervation yet the cholinergic neurons are restricted to the medial septal nucleus. Indeed, the heaviest NE innervation of the septum appears to be in its ventral and ventrolateral parts of the septum, well outside of the known dendritic spread of the medial septal cholinergic neurons. There is also moderate to dense NE innervation of the BST. The BST has several distinct subdivisions, which receive distinct patterns of NE inputs. This basal telencephalic complex appears to be involved in a number of complex functions including sexual

There have been relatively few studies of the specific LC NE innervation of the thalamus with tract-tracing and/or immunocytochemical methods. The most comprehensive account of NE innervation of the thalamus is the glyoxylic fluorescence study of Lindvall et al. (1974). NE fibers, primarily from the LC, differentially innervate different thalamic nuclei. The most prominent innervation is to the anterior thalamic group; the anteroventral nucleus, particularly its lateral aspect, is one of the most heavily innervated structures in the brain. The dorsal lateral geniculate and the ventrobasal complex, sensory nuclei for the visual and somatosensory systems, respectively, receive moderate to dense innervation but the principal sensory relay in the auditory pathway, the medial geniculate, receives a relatively sparse innervation. The lateral dorsal and posterior nuclei also receive an innervation less robust than that of the dorsal lateral geniculate and ventrobasal complex. The ventroanterior and ventromedial nuclei receive moderate innervation. In general, the midline nuclei are sparsely innervated by NE fibers but the paraventricular nucleus receives a very dense innervation of presumed adrenergic fibers as this plexus is robustly stained by specific antibodies to PNMT. The lateral and medial habenular nuclei (epithalamus) have a moderate to dense innervation although the source of this input appears to derive mainly from noncoerulear fibers. The thalamic reticular nucleus, a key structure for coordinating thalamocortical activity, has a dense NE plexus that derives from the LC. A recent study by Simpson and colleagues (1997) found that LC neurons retrogradely labeled from ventrobasal or posterior thalamic nuclei are found throughout the rostrocaudal extent of the nucleus, but clustered somewhat caudally overall. In contrast to neurons that project to the cerebral cortex, they found that LC neurons projecting to these thalamic nuclei were located bilaterally, but with an ipsilateral predominance. Striatum NE innervation is very sparse or absent in most areas of the striatum. In fact, the dorsal striatum stands

Basal Forebrain

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behavior and autonomic function. Recent work has also implicated the BST in anxiety (Davis and Shi, 1999) and in the aversiveness of opiate withdrawal (Delfs et al., 2000). Overall, DBH fiber innervation in medial BST is greater than that in the lateral BST (Phelix et al., 1992). However, the ventral BST receives perhaps the densest NE innervation in the brain. In ultrastructural studies DBH terminals formed synapses with dendritic shafts and spines of BST neurons (Phelix et al., 1992). In addition, there are frequent contacts from DBH terminals onto CRH-positive neurons in the ventrolateral BST (Phelix et al., 1994). Recent tract-tracing studies in our lab (Delfs et al., 2000) demonstrated that the majority of NE in the ventral BST originates in the A2 and A1 neurons of the caudal medulla, and not in the LC as had been previously supposed. Amygdala The amygdala receives a topographically organized innervation by NE fibers throughout its various subnuclei (Fallon et al., 1978). There are several sources of this NE input, including projections from the LC, the A2 neurons in the Sol, and the A1 neurons in the caudal ventrolateral medulla (Clayton and Williams, 2000; Zardetto-Smith and Gray, 1990, 1995). We have observed that the LC preferentially innervates the medial subcomponent of the central nucleus of the amygdala, whereas the NE input to the lateral division of the central amygdala nucleus originates from nonLC sources (Aston-Jones et al., 1996). NE input to the amygdala has attracted considerable attention in studies of memory. McGaugh and colleagues, as well as others, have found that memory for emotionally salient events is enhanced as a consequence of elevated NE release in the basolateral amygdala (McGaugh, 2000). Interestingly, this memory-enhancing effect requires NE from the A2 group in the Sol (Williams and McGaugh, 1993), while other studies indicate that the LC is also involved (Clayton and Williams, 2000). Preoptic Area

innervated by NE fibers and receives direct input from the LC and the A1 and A2 NE cell groups in the caudal medulla (Chou et al., 2002). This has implications in regulation of sleep and waking, as the ventrolateral preoptic area has been found to critically regulate sleep (Lu et al., 2000), and the LC has long been implicated in arousal and waking (Aston-Jones et al., 1984). Hypothalamus and Midbrain The hypothalamus is densely innervated by NE axons. The majority of these axons originate in cells of the A1 and A2 cell groups of the medulla, with only a sparse contingent of LC axons. Mason and Fibiger (1979) observed retrogradely labeled neurons in the rostral pole of the LC following HRP injections into the hypothalamus, a finding later confirmed by Loughlin et al. (1986a, 1986b). The vast majority of LC axons traverse the lateral hypothalamus en route to telencephalic regions, and the possibility that some labeling after hypothalamic injections was due to uptake by passing fibers cannot be ruled out. Cunningham and Sawchenko (1988) traced fibers from the LC to the medial wall of the paraventricular nucleus of the hypothalamus, but did not find LC projections to the supraoptic nucleus. This is consistent with our observations for anterograde transport of WGA–HRP from the LC (unpublished observations). LC axon terminals in the midbrain are most prominent in the tectum. By using anterograde transport of the lectin PHA-L in combination with dopamine βhydroxylase immunocytochemistry, Fritschy and Grzanna (1990b) demonstrated a bilateral distribution of LC axons in the inferior and superior colliculus, the dorsal and lateral periaqueductal gray, and the interpeduncular nucleus. Numerous PHA-L-labeled LC axons were present in the dorsal bundle but only a few labeled fibers could be detected in the midbrain tegmentum.

Descending Projections

The medial preoptic area has moderate to strong catecholaminergic inputs that would appear to be composed primarily of NE- and epinephrine-containing fibers. There is retrograde labeling evidence for direct projections from the LC and from adrenergic neurons in the ventral medulla to the preoptic area, a sexually dimorphic structure that plays a pivotal role in sexual behaviors and the regulation of gonadal steroid function. The cells of origin of this complex catecholaminergic innervation and the relative importance of epinephrine and NE to these functions are largely unknown. However, recent studies by Saper and colleagues have found that the ventrolateral preoptic area is densely

Cerebellum The innervation of the cerebellum by LC NE fibers has been described in considerable detail. Early studies indicated that the NE innervation of the cerebellar cortex was derived entirely from the LC (Moore and Bloom, 1979; Olson and Fuxe, 1971). Light microscopic observations using formaldehyde- or glyoxylic acidinduced fluorescence in the cerebellum revealed widespread innervation of the cerebellar cortex by fluorescent (presumably noradrenergic) fibers. These fibers enter the cerebellum via the superior peduncle (Jacobowitz and Kostrezewa, 1971; Pickel et al., 1973)

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and pass from the underlying white matter through the granule cell layer without considerable branching or apparent terminations. NE fibers then ramify in the molecular layer with numerous branches and varicosities, particularly in the vicinity of the proximal dendrites of Purkinje cells (Bloom and Battenburg, 1976; Bloom et al., 1971, 1972; Moore and Bloom, 1979). In the outer molecular layer the fibers appear to bifurcate and run parallel to the folium surface. Detailed ultrastructural analyses of NE synapses in the cerebellar cortex were made by Bloom and colleagues using autoradiography of sites taking up 3NE or by localizing degenerating fibers induced by treatment with the selective catecholamine neurotoxin 6-hydroxydopamine. These studies revealed that the majority of the NE synapses were onto dendrites of Purkinje cells in the mid-to-outer molecular layer (Bloom et al., 1971, 1972). Occasionally presumed NE terminals were seen to contact Purkinje cell dendrites in the inner molecular layer. These fibers appeared to climb along Purkinje dendrites before giving off terminallike varicosities. The estimated percentage of total cerebellar terminals that are presumably noradrenergic was only about 1%, but it was concluded that sufficient arborization existed to allow NE innervation of each Purkinje dendritic field (Bloom et al., 1972). These anatomical studies formed the basis for a great deal of physiological and pharmacological work which provided the initial detailed characterization of NE functional effects on central neurons, using the LC– Purkinje cell synapse as a model system (Bloom et al., 1972). Pons and Medulla The entire brain stem is richly innervated by NE axons. Early studies of the projections of the LC to the pons and medulla concluded the existence of widespread distribution of the efferent network in the pons and medulla. However, studies employing bilateral LC lesions revealed a distinct pattern of organization for the NE innervation of the brain stem. According to the observations by Levitt and Moore (1979), the LC projects primarily to sensory and association nuclei of the brain stem with NE neurons from non-LC sources projecting to motor nuclei and autonomic centers. This latter view was supported by a study of LC axon distribution by Fritschy and Grzanna (1990b) who used anterograde transport of PHA-L to demonstrate a remarkably restricted distribution of LC axons primarily to the trigeminal complex, especially the trigeminal nucleus pars caudalis. Spinal Cord Despite the obvious conceptual interest in knowing the detailed distribution of LC axons in the spinal cord, this topic has remained a subject of considerable debate.

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Since the demonstration of LC projections to the spinal cord (Ader et al., 1980; Commissiong et al., 1978; Nygren and Olson, 1977; Westlund et al., 1983), several laboratories have attempted to characterize their distribution within the cord. Early studies suggested that the LC projects primarily to the ventral horn thus implicating this nucleus in the control of motor output of the spinal cord (Commissiong et al., 1978). Anterograde transport of PHA-L by Fritschy and Grzanna (1990a) traced LC axons to the superficial laminae of the dorsal horn, implicating the LC in the control of sensory inputs to the spinal cord. This distribution is in good agreement with the pattern of distribution described in the brain stem where LC axons are distributed primarily to sensory and association nuclei (Fritschy and Grzanna, 1990b; Levitt and Moore, 1979). Within the spinal cord LC axons appear to descend the length of the cord with the superficial laminae of the dorsal horn. However, an entirely different pattern of LC axon distribution was reported by Clark and Proudfit (1991). These authors traced LC axons to the ventral horn of the spinal cord and were unable to detect any LC axons in the superficial layers of the dorsal horn. However, these authors noted striking substrain differences when they compared the noradrenergic innervation of the spinal cord in Sprague–Dawley rats from Sasco, Inc., vs Harlan Sprague–Dawley, Inc. (Clark et al., 1991). Using discrete injections of the retrograde tracer fluoro-gold into the superficial dorsal horn of Harlan rats (the strain used by Fritschy and Grzanna, 1990a, 1990b), Clark et al. (1991) observed more retrogradely labeled neurons in the LC than in the A5 and A7 cell groups, a finding in agreement with the anterograde study of Fritschy and Grzanna (1990a).

General Properties of LC Projections The functional significance of the broad projections from the LC nucleus depends upon the degree to which individual LC neurons share in such a multitarget innervation strategy. It is possible that different LC neurons project to select target areas and convey functions specialized for their specific targets. On the other hand, it is possible that individual LC neurons innervate a number of different targets, so that their function is more general and integrative. Several recent studies have examined these issues and are reviewed below in two sections: (i) the collateralization of individual LC neurons, and (ii) the topography of neurons within the LC based upon the targets that they innervate. Efferent Collateralization of LC Neurons This issue has been addressed using both electrophysiological methods to antidromically activate LC

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neurons from multiple brain areas and anatomical methods to doubly retrogradely label LC neurons from multiple target regions. Together, these studies indicate that single LC neurons are highly branched with projections often leading to distant and functionally different target sites. Retrograde labeling of LC neurons from multiple sites Several studies have found that two retrograde tracers injected into different target areas doubly label individual LC neurons. Some LC neurons were doubly labeled after injections of fluorescent tracers into pairs of sites in different hemispheres, including cortex–cortex, hippocampus–cortex, thalamus–cortex, thalamus– hippocampus, and in a few cases hippocampus– hippocampus (Ader et al., 1980). These results are compatible with those of others (Room et al., 1981) that individual LC neurons could project to at least two of the following sites (either ipsilaterally or bilaterally): prefrontal cortex, thalamus, hippocampus, and spinal cord. Injections into the cortex, thalamus, and hippocampus revealed not only ipsilateral but also contralateral labeling of cells in the LC. In addition, injections of fluorescent tracers into any forebrain area combined with an injection into the spinal cord produced doubly labeled LC neurons. A study in mice found that over 50% of cerebellar cortical-projecting LC neurons were also retrogradely labeled from ipsilateral forebrain sites and that these doubly labeled neurons appeared to be scattered throughout the LC nucleus (Steindler, 1981). Similar results were found in rat where doubly labeled LC neurons occurred after injections of retrograde tracers into frontal cortex and either cerebellar cortex, occipital cortex, or spinal cord (Nagai et al., 1981). Single LC neurons are reported to innervate both dorsal and ventral aspects of the area dentata of the hippocampus (Haring and Davis, 1983). Branched projections from single LC and subcoeruleus neurons (presumed to be noradrenergic) to both amygdala and the cerebellum have also been reported using double retrograde tracing (Dietrichs, 1985). Detailed studies of the distribution of LC projections to rat cortical areas by Loughlin et al. (1982) revealed that while individual LC neurons often were labeled from two distinct cortical areas, double labeling was much more frequent following injections into cortical areas aligned in the rostrocaudal axis (e.g., frontal and parietal cortices) than with injections placed at different mediolateral positions in the cortex (e.g., cingulate and suprarhinal cortices). This result is consistent with results of Morrison et al. (1981) for rostrocaudally oriented NE fibers in rat cortex. Double labeling was especially frequent with injections placed in ventral and deep sites of the same cortical region.

Evidence of branched projections from subcoeruleus neurons to the medial preoptic area and cervical spinal cord (Leanza et al., 1989), or to the raphe pontis and hypothalamic zona incerta (Leanza et al., 1988), has also been reported using double-labeling techniques. However, it is unknown whether these branched neurons are noradrenergic. A recent study by Waterhouse and colleagues has revealed additional organization in LC projections of an intriguing nature (Simpson et al., 1997). These investigators used retrograde labeling to study the innervation of different nuclei within the somatosensory pathway by individual LC neurons. They found that LC neurons were more likely to be doubly retrogradely labeled in cases where the two injections were made into functionally related areas (e.g., barrel field cortex and the ipsilateral ventrobasal thalamus) than when cortical and thalamic injections were made into functionally unrelated areas (e.g., barrel field cortex and ipsilateral dorsolateral geniculate nucleus). These results present a novel and potentially functionally important topography and specificity in the anatomy of the “ubiquitous” set of LC efferents. As one possible functional upshot, given the proposed role of LC neurons in attention (Aston-Jones et al., 2000), such “functionally homologous” branching could support selective attention to one vs other modalities of sensory stimuli. This possibility remains to be tested. Antidromic activation of LC neurons from multiple sites The above results with double retrograde labeling are largely compatible with “electro-anatomy” experiments, where antidromic activation from electrical stimulation in different target sites has been used to chart branched LC projections. The earliest such study by Nakamura and Iwama (1975) revealed that individual LC neurons projected to frontal and visual cortices, hippocampus and frontal cortex, hippocampus and visual cortex, and from three sites in one cell, hippocampus and frontal and visual cortices. In addition, of 3 cells driven antidromically from the cerebellar cortex, 2 were similarly driven from the cerebral cortex. Faiers and Mogenson (1976) found that 7 of 21 LC neurons antidromically identified as projecting through the supracallosal bundle under the frontal cortex also projected to the olfactory bulb. More evidence of branched LC axons was obtained in a study by Takigawa and Mogenson (1977), who found individual LC neurons antidromically activated from olfactory bulb, supracollosal bundle, and bed nucleus of the stria terminalis. This same study revealed that many subcoeruleus neurons could also be antidromically activated from the supracollosal bundle. Single LC neurons were similarly antidromically activated

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from the olfactory bulb and the cingulate cortex (Aston-Jones et al., 1980). Finally, a study by Guyenet (1980) found that 4 of 8 spinally projecting LC neurons were also antidromically driven by stimulation of the midbrain dorsal noradrenergic bundle and that each of the 4 neurons driven antidromically from the cerebellum also projected rostrally through the dorsal noradrenergic bundle. Topography of LC Neurons with Specific Targets Possible topographical distinctions within the LC are relatively difficult to study due to the small volume and number of cells in the nucleus and the highly divergent nature of the overall projection. This situation seems compatible with the scenario that most cells project to more than one target area, so that strictly segregated subgroups of LC neurons distinguished by efferent targets may not be prevalent. Nonetheless, examples of topographic specificity for LC neurons with particular targets have been described. The clearest example of a subgroup of selectively projecting LC neurons is the set projecting to the spinal cord. Many studies have shown using a variety of techniques that spinally projecting LC neurons are highly preferentially located in the ventral caudal aspect of the nucleus (Loughlin et al., 1986a, 1986b; Room et al., 1981; Satoh et al., 1977; Westlund et al., 1981, 1983). Other studies indicate that most LC neurons identified by a particular target overlap spatially within the nucleus with other sets of LC neurons that project to different target areas. Nonetheless, there are subtle distinctions among such groupings, such that interdigitated cells defined by different targets exhibit topographically distinguishable gradients of localization within the nucleus. Loughlin et al. (1982, 1986a) found that LC cells retrogradely labeled from the hippocampus were clustered in the dorsal aspect of the nucleus, while those labeled from the hypothalamus clustered strongly in the rostral aspect of the LC. Cells projecting to the cerebral or cerebellar cortices, on the other hand, were more evenly distributed throughout the nucleus. Morphological distinctions were also found among groups of LC neurons with different targets, such that fusiform cells project to the cerebral cortex and hippocampus, large multipolar neurons in ventral LC innervate the spinal cord and cerebellum; and small round cells in the central and anterior LC, along with multipolar anterior cells, preferentially project to the hypothalamus (Loughlin et al., 1986b). Cells centrally located in the nucleus were numerously retrogradely labeled following injections into many different target areas. Room et al. (1981) found that medial cerebellar injections yielded retrogradely labeled neurons throughout the LC nucleus. Haring and Davis (1983)

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reported that LC neurons retrogradely labeled from the dorsal area dentata were preferentially located within the dorsal aspect of the LC nucleus, whereas cells labeled from ventral area dentata injections were more widely dispersed in the LC nucleus. Fusiform and multipolar cells were most commonly labeled in the LC from area dentata injections. These investigators also reported that the septal and ventral poles of the dentate gyrus are innervated by distinct fiber pathways from the LC, employing the fornix and cingulum, respectively (Haring and Davis, 1985). In a detailed study of cortically projecting LC neurons, Waterhouse et al. (1983) found that LC cells projecting to the cerebral cortex tended to be localized within the caudal three-fifths of the dorsal ipsilateral LC nucleus. Within this portion of the nucleus, groups of neurons identified as projecting to occipital, sensorimotor, or frontal cortices were observed to form a dorsal to ventral gradient, while occipital-projecting LC cells tended to be more caudally placed in the LC nucleus than those projecting to the sensorimotor or frontal cortices. Only the frontal cortex received innervation from LC neurons in both dorsal and ventral subdivisions of the nucleus. Overlapping sets of cells were labeled from cerebrocortex, caudate, hippocampus, and cerebellum. These findings differ in some regards with earlier conclusions that cortically projecting LC neurons in rat are randomly distributed throughout the nucleus except for its ventral division (Mason and Fibiger, 1979). It is noteworthy that the density of LC innervation varies considerably among different cortical or other target areas, as reviewed above. It is not known whether these distinct innervation patterns or densities are related to select populations of neurons that innervate different regions. Ultrastructural Characteristics of LC Terminals The terminals of LC neurons have been examined in EM studies in several brain areas. Early reports from Descarries and colleagues (Descarries and Beaudet, 1983; Descarries et al., 1977) using EM autoradiographic examination of NE terminals in the cerebral cortex concluded that most NE terminals did not make synaptic contacts but rather existed as nonsynaptically arranged terminals to provide NE in a paracrine fashion to a local cortical area. Similar results have been reported by this lab more recently using EM localization of cortical terminals stained with antibodies against NE (Seguela et al., 1990). However, quantitative studies by Koda et al. (1978) in the hippocampus found that NE terminals (identified by the presence of small granular vesicles following permanganate fixation) formed specialized synaptic junctions with other neurons as frequently as

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non-NE terminals. In addition, studies by Papadopoulos, Parnavelas and colleagues (Papadopoulos and Parnavelas, 1990; Papadopoulos et al., 1987) using serial EM reconstruction of noradrenergic synapses in cerebral cortex identified with DBH immunohistochemistry found that the great majority of NE terminals formed conventional synaptic contacts onto neuronal profiles. These results are consistent with those of similar studies by Olschowka et al. (1981) who found that most DBHpositive terminals in areas of the diencephalon, the cerebellum, and the limbic cortex formed axodendritic synapses characterized by specialized junctional appositions. Thus, although results from different labs vary as to the frequency with which LC terminals form synaptic contacts onto target neurons, considerable evidence exists to indicate that this system uses conventional synaptic transmission at many of its terminations. The possibility remains, of course, that both synaptic and paracrine modes of neurotransmission exist at the same or different NE terminals. More recent studies by Aoki (1992) have found that terminals in the cerebral cortex that stain for TH are often located in apposition to astrocytic processes that stain for an antibody raised against a β-adrenergic receptor. These observations are consistent with other studies of Stone and colleagues (Stone and John, 1991) for a glial localization of β-adrenergic receptors and open the possibility for additional modes of LC influence in target areas involving glial cell activities.

OTHER METENCEPHALIC NORADRENERGIC NEURONS (A5 AND A7 CELL GROUPS) The neuroanatomy of the A5 and A7 cell groups has been studied much less extensively than that of the LC. These two groups form what has been termed the pontine lateral tegmental noradrenergic cell groups (Moore and Bloom, 1979; Moore and Card, 1984). They are distinct from the more medially and dorsally placed LC neurons in cytology, location, and connections. As summarized below, the lateral tegmental noradrenergic neurons have much efferent connections much more restricted than those of the LC cells, innervating primarily brain stem and spinal structures. Noncoerulear NE neurons form a nearly continuous column of cells in the ventrolateral portion of the metencephalon extending from the rostral region of the motor nucleus of the facial nerve to the pontine– midbrain junction. Recognizing two distinct clusters of NE cells, Dahlstrom and Fuxe (1964) divided this cell column into a rostral and a caudal portion which they labeled the A7 and A5 groups, respectively. This sepa-

ration of NE cells in the ventrolateral metencephalon into two separate groups has been questioned. Other investigators (Moore and Bloom, 1979) consider these neurons the metencephalic portion of a column of NE neurons that is continuous with A1 cells in the ventrolateral medulla and refer to the entire cell complex as the lateral tegmental group. A5 and A7 cells differ from LC neurons in that they lack ascending projections to the forebrain. The significance of the A5 and A7 groups has remained controversial. Nevertheless, there is a considerable body of evidence that a distinction between A5 and A7 cells is justified on connectional grounds. Tracer studies indicate that the A5 and A7 groups have distinct projection fields. Grzanna et al. (1987) noticed different labeling patterns in the A5 and A7 groups following tracer injections into the motor nuclei of the trigeminal and facial nerves, respectively. In double-labeling experiments, Lyons and Grzanna (1988) demonstrated that A7 cells have divergent projections to the motor nucleus of the trigeminal nerve and the spinal cord, suggesting a role of A7 cells in the control of motor output. There is general agreement that A5 cells project to autonomic nuclei of the brain stem and spinal cord (Byrum and Guyenet, 1987). More recent studies have focused on A7 neurons and their role in nociception. Proudfit and colleagues have shown that spinally projecting A7 neurons, which are implicated in antinociception, receive a direct input from the ventrolateral PAG, indicating a possible route for PAG-induced NE-dependent antinociception (Bajic and Proudfit, 1999). In addition, these neurons contain receptors for substance P (Chen et al., 2000) and receive substance P inputs (Proudfit and Monsen, 1999). These investigators have also implicated GABA neurons that project to A7 cells in antinociception (Nuseir and Proudfit, 2000). Recent studies also found that A5 NE neurons serve as a relay between the caudal ventrolateral medulla and the spinal cord (Tavares et al., 1996). This may indicate a pathway by which the caudal ventrolateral medulla exerts effects on blood pressure and respiratory function (Koshiya and Guyenet, 1994).

CONCLUSIONS The LC NE system in rat remains the premier chemically characterized neural system in the brain. This system is notable because of it has very widespread and divergent projections, and it provides the bulk of the brain’s NE. Although extensive work has revealed that LC projections to many target areas form conventional synapses onto neurons, this does not rule out

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the possibility of additional, nonsynaptic release of the NE transmitter. LC neurons contain several peptides in addition to NE. This provides some of the clearest evidence for heterogeneity among LC neurons, which otherwise appear in many respects to resemble one another closely. At the level of the LC nucleus, detailed immunohistochemical studies have found that LC dendrites extend outside of the LC proper in a polarized fashion, creating dendritic zones surrounding the LC nucleus. Recent results show the potential importance of this peri-LC surround (dendritic zone) in the afferent regulation of LC neurons. Other recent findings show an important circuit from the SCh through the DM to the LC. This circuit may participate in the circadian regulation of arousal and cognitive performance. Major afferents to the LC nucleus proper include the rostral medullary nuclei LPGi and the EF. These findings reveal that the LC is closely linked with autonomic circuitry in the CNS, consistent with a host of previous results. These results are consistent with the view that the LC system serves as the cognitive limb of a global rapid response system, whereby LC and sympathetic systems are coactivated in parallel to yield rapid adaptive responses to urgent stimuli (AstonJones et al., 1991b; 1996). A broader view of LC function has recently emerged from recent studies of LC activity in behaving monkeys. This hypothesis states that the NE LC system regulates attention and behavioral flexibility, either promoting focused attention and responding to stimuli relevant to an existing task or facilitating the search for another task that is more adaptive in the current context (Aston-Jones et al., 2000). In addition to these results for the LC system, recent work has significantly extended our knowledge of the connections and possible functions of the A5 and A7 metencephalic cell groups. Examination of the anatomy of these metencephalic NE systems continues to reveal unexpected specificity with new and deeper functional implications. However, many additional studies at the circuit and ultrastructural levels are needed to fully benefit from the new findings.

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C H A P T E R

12 Oromotor Nuclei JOSEPH B. TRAVERS Department of Oral Biology,Ohio State University Columbus, Ohio, USA

This chapter will describe the intrinsic organization of the trigeminal (Mo5), facial (7), and hypoglossal (12) motor nuclei and identify the sources of central projections to them. Brain stem projections to these nuclei originate from both sensory and reticular formation structures and serve a wide range of oral behaviors that range from simple oral reflexes, e.g., the dysynaptic jaw-opening reflex, to complex oral functions such as swallowing, mastication, and respiration. Although swallowing, mastication, licking, and respiration are each thought to be organized by brain stem central pattern generators that provide a coordinated output to the appropriate oromotor pools (reviewed in Jean, 2001; Nakamura and Katakura, 1995; Rekling and Feldman, 1998; Travers et al., 1997), there is increasing evidence that oromotor nuclei are more than simple output neurons. In addition to receiving excitatory and inhibitory projections utilizing classic neurotransmitters, the oromotor nuclei also receive inputs that utilize a variety of neuromodulatory substances. The intrinsic membrane properties of motoneurons (Del Negro et al., 1999) as well as the colocalization of neuromodulators confer the potential to further shape patterned motor output. γ-Aminobutyric acid (Davidoff and Schulze, 1988), calcitonin gene-related peptide (Skofitsch and Jacobowitz, 1985), urotensin II (Dun et al., 2001), and urocortin (Bittencourt et al., 1999) have each been localized within oromotoneurons, and mRNA for cholecystokinin has been identified as well (Ingram et al., 1989; Sutin and Jacobowitz, 1990). Further adding to the complexity of the oromotor nuclei are interneurons within 12 (Boone and Aldes, 1984; Takasu and

The Rat Nervous System, Third Edition

Hashimoto, 1988), possible axon collaterals within Mo5 (Moore and Appenteng, 1989), and, in species other than rat, central projections originating from Mo5 and 7 (cat: Kotchabhakdi and Walberg, 1977; Roste, 1989; rabbit: Bukowska and Grottel, 1997). Although each of the oromotor nuclei have unique characteristics, the coordination between muscles innervated by Mo5, 7, and 12 during feeding, grooming, and respiration is perhaps the most compelling reason to discuss the neuroanatomy of these nuclei within one chapter.

MOTOR TRIGEMINAL NUCLEUS [REFER TO INT. 0.2 TO −0.8 MM] Intrinsic Organization Myotopic Organization The motor trigeminal nucleus of the rat is conventionally divided into a large dorsolateral division defining the rostrocaudal length of the nucleus and a smaller ventromedial division in the caudal two-thirds (Jacquin et al., 1983; Limwongse and DeSantis, 1977; Lynch, 1985; Mizuno et al., 1975; Rokx and van Willigen, 1985; Sasamoto, 1979). Although an accessory nucleus for Mo5 was designated using cobalt labeling of the peripheral nerve (Szekely and Matesz, 1982), these neurons almost certainly correspond to the ventromedial subdivision of the main nucleus rather than to motoneurons outside the main nucleus. Figure 1 depicts the myotopic organization of Mo5. The jaw closing muscles—the masseter (superficial,

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Ms; anterior deep, Ma), temporalis (T), and medial (external) pterygoid (Pm)—are innervated from the dorsolateral division (DL); the jaw opening muscles— the anterior digastric (AD) and mylohyoid (MY)—are innervated from the ventromedial division (VM). A third jaw opening muscle, the lateral (internal) pterygoid (Pl) is grouped with jaw closing motoneurons in the ventral aspect of the dorsolateral division. In the rat, the location of medial and lateral pterygoid motoneurons have not been located individually within the DL; however, medial pterygoid motoneurons are located either ventral (guinea pig: UemuraSumi et al., 1982) or lateral (rabbit: Matsuda et al., 1978) to lateral pterygoid motoneurons. A laminar organization further defines the distribution of motoneurons within the DL (Rokx et al., 1985). Motoneurons innervating the anterior deep masseter muscle are interposed between motoneurons innervating the superficial masseter muscles further laterally and the temporalis muscle dorsomedially. Different “compartments” (Weijs, 1996) or “anatomical partitions” (Saad et al., 1997) of the rabbit masseter muscle function with some degree of independence during mastication. The spatial separation of motoneurons innervating these different compartments potentially allows for the segregation of input as a source for differential activation. The degree of motoneuron separation, however, is unclear. Weijs (1996) found that superficial masseter motoneurons occupied a dorso-

lateral position within the dorsal subdivision compared to deep masseter motoneurons that were more medial. The overlap between these two populations of neurons was on the order of 50%. In contrast, after applying separate fluorescent tracers to different nerve branches, the overlap between deep and superficial masseter motoneurons was estimated at over 90% (Saad et al., 1997). Motoneurons innervating the transverse mandibular muscle have not been specifically located in the rat but are clustered together in the same region as motoneurons innervating the superficial masseter muscle in guinea pig (Segade, 1990). Within the ventromedial division of Mo5, anterior digastric motoneurons are dorsomedial to mylohyoid motoneurons. Innervation appears strictly ipsilateral. Evidence that a small number of Mo5 neurons innervate the contralateral AD (Kemplay and Cavanagh, 1983) was not confirmed (Rokx et al., 1985). The tensor palatini is also innervated by motoneurons in the ventromedial division (cat: Keller et al., 1983). In rat, the tensor tympani is innervated by trigeminal motoneurons located ventral to Mo5 and medial to exiting trigeminal rootlets and thus appears in approximately the same anatomical location as group k neurons (Spangler et al., 1982). Group k neurons are a cell column outside the cytoarchitectonic boundaries of Mo5 (rabbit: Meesen and Olszewski, 1949) and contains (additional) motoneurons innervating the masseter and anterior digastric muscles (rabbit: Donga

FIGURE 1 The myotopic organization of the motor trigeminal nucleus is depicted in three coronal sections. The larger dorsolateral (DL) subdivision extends over the rostrocaudal length of the nucleus, the smaller ventrolateral (VM) subdivision only over the caudal two-thirds (b & c). Motor pools innervating primarily jaw closure muscles in the DL are largely segregated: superficial masseter (Ms), anterior deep masseter (Ma), temporalis (T), medial pterygoid (Pm), and lateral pterygoid (Pl). Jaw openers in the VM include the anterior digastric (AD) and mylohyoid (MY).

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et al., 1992; Saad et al., 1997). Group k motoneurons are smaller than those in Mo5 and mirror the topology seen in the main nucleus, i.e., masseter motoneurons are dorsal to digastric motoneurons. However, there are few references to group k in rat (e.g., Li et al., 1995) and group k innervation of masseter and digastric muscles has not been reported. Cytoarchitectonics Each Mo5 contains approximately 2265 to 2979 cells (Limwongse and DeSantis, 1977), the majority of which innervate jaw closing muscles (Kemplay and Cavanagh, 1983; Szekely and Matesz, 1982). The dorsolateral division contains both small (8–14 μm) and large (28–42 μm) diameter multipolar cells, which correlates with a bimodal distribution of nerve fiber diameters (Limwongse and DeSantis, 1977). Muscle spindles in the jaw closing muscles of rats (Karlsen, 1965) may terminate on small (presumed) γ efferent motoneurons in dorsolateral Mo5. Two classes of multipolar cells in the ventromedial division are also evident, although the large cells (24–34 μm in diameter) are not as large as their dorsolateral counterparts and the small cells (16–20 μm in diameter) are not as small. An absence of very small neurons in the ventromedial division is consistent with the lack of muscle spindles in jaw opening muscles (Limwongse and DeSantis, 1977). Axosomatic synapses cover, on average, 78% of the soma of large multipolar Mo5 cells in contrast to the smaller multipolar or fusiform cells that have proportionally far fewer synapses (Card et al., 1986). In addition to somata size, intracranial axonal trajectory further differentiates between dorsolateral and ventromedial cells. Axons from the dorsolateral division exit the brain directly, forming a few large roots that pass just ventral to the principal trigeminal sensory nucleus (Pr5). Axons from the ventromedial division, however, first course dorsally to form a genu rostral to the more prominent facial nerve (7n) genu before turning ventrolaterally as a separate root, parallel to fibers from the dorsolateral division (Jacquin et al., 1983; Szekely and Matesz, 1982). Although classical descriptions of trigeminal motoneurons do not include axon collaterals (Ramon Y. Cajal, 1972), collaterals were observed for each of four Mo5 cells filled intracellularly with horseradish peroxidase (HRP) (Moore and Appenteng, 1989). For three of the four cells, the exiting motoneuron axon arborized within Mo5 and the fourth axon arborized in Pr5. Unfortunately, axon collaterals have not been reported in any other intracellular labeling studies (Lingenhohl and Friauf, 1991; cat: Shigenaga et al., 1988; Yabuta et al., 1996; Yoshida et al., 1987), leaving their existence in question.

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As with all the cranial motor nuclei, Mo5 is cholinergic. In addition, calcitonin gene-related peptide (CGRP) (Skofitsch and Jacobowitz, 1985), urotensin II (Dun et al., 2001), and urocortin (Bittencourt et al., 1999) have been localized in Mo5 cells. Messenger RNA for cholecystokinin (CCK) has also been widely reported (e.g., Ingram et al., 1989; Sutin and Jacobowitz, 1990); however, there is little evidence for immunohistochemical detection of the peptide. Dendritic Architecture Dendrites of neurons located centrally within Mo5 radiate symmetrically from the soma compared to more peripherally located cells with dendrites that radiate preferentially toward the interior of the nucleus or follow the contour of the nucleus (Fig. 2) (Card et al., 1986; Lingenhohl and Friauf, 1991). Dendrites are largely confined to the nucleus; however, a few dendrites of jaw closing motoneurons extend dorsally and laterally beyond the borders of Mo5 into the mesencephalic trigeminal nucleus (Me5), the principal trigeminal nucleus (Pr5), and the supra- and intertrigeminal regions or medially into the pontine reticular formation (Lingenhohl and Friauf, 1991; Mong et al., 1988), (cat: Yoshida et al., 1987). Dendrites of motoneurons that innervate non-oral musculature do not overlap the dendritic fields of orally related trigeminal motoneurons. Thus, dendrites of tensor tympani motoneurons do not overlap the dendritic or soma fields of nearby jaw closing or opening motoneurons but are in close proximity to the superior olive, providing a possible substrate for auditory reflex function (cat: Friauf and Baker, 1985). A quantitative ultrastructural analysis of synapses on masseter motoneurons indicated a differential distribution such that putative inhibitory synapses were preferentially distributed on proximal dendrites (cat: Bae et al., 1999). Interneurons A class of small fusiform-shaped neurons located primarily around the borders of Mo5 that receive few synaptic terminals may be interneurons (Card et al., 1986). Other, presumably nonmotoneuronal cells within Mo5 have central projections. Cells within Mo5 were retrogradely labeled from injections of HRP into the cerebellum (cat: Kotchabhakdi and Walberg, 1977; Roste, 1989) (rabbit: Bukowska and Grottel, 1997) and a projection from Mo5 to the hypoglossal nucleus has also been reported (Manaker et al., 1992). In addition, a small projection from neurons in the jaw closing region of Mo5 to the homologous region on the contralateral side could contribute to bilateral jaw closing (Juch et al., 1993; Ter Horst et al., 1990; cat: Mizuno et al., 1983; rabbit: Kolta et al., 2000).

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FIGURE 2 Intracellular fills of masseter motoneurons. (a) Motoneurons near the edge of the nucleus have dendrites that radiate toward the center of the nucleus and follow the nucleus contour. (b) Motoneurons in the center of the nucleus have dendrites that radiate symmetrically. In many cases, dendrites extend into the adjacent reticular formation. From Lingenhohl and Friauf, 1991.

Afferent Projections Forebrain Pathways Although there is little anatomical evidence for direct forebrain projections to Mo5 in rat based on retrograde (Travers and Norgren, 1983; Vornov and Sutin, 1983) or anterograde tracers (Zhang and Sasamoto, 1990), this view may be in need of modification. Recent work in cat (Fung et al., 2001) suggests direct hypothalamic projections of orexin-containing neurons to the motor trigeminal and hypoglossal nuclei, and mRNA for the receptor for melanin-concentrating hormone, another hypothalamic feeding-related peptide, is found in all of the rat oromotor nuclei including the motor trigeminal nucleus (Saito et al., 2001). Forebrain structures certainly have a direct influence on trigeminal motoneuron activity. Electrical stimulation of either the frontal cortex or the central nucleus of the amygdala can produce rhythmic jaw movements in a number of species including rat (Ohta, 1984; Sasamoto and Ohta, 1982). In addition, the relatively short-latency onset of EPSPs in jaw opening motoneurons and IPSPs in jaw closing motoneurons was suggestive of a monosynaptic pathway (Ohta, 1984). One brief report describes a few direct cortico-trigeminal fibers (Mishima et al., 1983). The predominant view, however, is that rhythmic masticatory movements are

organized in the brain stem reticular formation (reviewed in: Nakamura and Katakura, 1995; Travers et al., 1997). The short-latency responses observed in Mo5 neurons from cortical stimulation might result from descending cortical projections (Takeuchi et al., 1988) onto Mo5 dendrites that extend into the lateral tegmental field. Most direct projections originate from the medulla and pons with a small contribution from the midbrain; major direct projections are summarized in Fig. 3. Midbrain Pathways Midbrain projections to Mo5 consist almost entirely of cells from the rostral extension of the ipsilateral Me5. Axon terminals of individual Me5 cells are distributed preferentially in the dorsolateral region of Mo5 (Dessem et al., 1997; Lingenhohl and Friauf, 1991; Matesz, 1981; cat: Shigenaga et al., 1989) and monosynaptically excite masseteric neurons in response to peripheral stimulation of either muscle spindles or periodontal ligament (Jerge, 1963). Recent studies tracing the trajectory of intracellularly filled Me5 neurons indicate that muscle spindle afferents most sensitive to muscle stretch project ipsilaterally directly onto trigeminal motoneurons (Dessem et al., 1997). Muscle spindle afferents signaling muscle length can influence trigeminal motoneurons bilaterally via the supratrigeminal area. Glutamate

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Pons regio h

A7 Pr5 (SubC) NE

Medulla PCRt/IRt MdD/MdV Gi LPGi raphe

EAA IAA ACh M-ENK Midbrain Mo5 5-HT

EAA

MeV

SP

EW

Sp5 FIGURE 3 Afferent projections to the trigeminal motor nucleus. Abbreviations: ACh, acetylcholine; EAA, excitatory amino acid; EW, Edinger–Westphal nucleus; Gi, gigantocellular nucleus; IAA, inhibitory amino acids; IRt, intermediate subdivision of reticular formation; LPGi, lateral paragigantocellularis; MdD, dorsal subdivision of the reticular formation (MdD); MdV, ventral subdivision of the reticular formation; M-ENK, methionine–enkephalin; Me5, mesencephalic trigeminal nucleus; NE, noradrenaline; PCRt, parvicellular reticular formation; Pr5, principal trigeminal sensory nucleus; regio h, reticular formation surrounding the motor trigeminal nucleus; SP, substance P; Sp5, spinal trigeminal nucleus; SubC, nucleus subceruleus; 5-HT, serotonin.

acting on a non-N-methyl-D-aspartic (non-NMDA) receptor may mediate transmission between Me5 and Mo5 (guinea pig: Chandler, 1989). Other midbrain projections to Mo5 include a projection from the Edinger–Westphal nucleus, immunohistochemically identified as containing substance P (cat: Fort et al., 1990) and an indirect pathway from the red nucleus. The red nucleus may influence Mo5 via the juxtatrigeminal area, that part of the lateral reticular formation interposed between Pr5 and the ventrolateral aspect of Mo5 (Godefroy et al., 1998; cat: Mizuno et al., 1974). The transynaptic transfer of pseudorabies virus (PRV) further suggests that the red nucleus influences Mo5 motoneurons via multisynaptic pathways (Fay and Norgren, 1997a). Pontine Projections Pontine projections to Mo5 begin rostrally at the level of the decussation of the brachium conjunctivum, just medial to the lateral lemniscus in an area corresponding to the A7 group (Grzanna et al., 1987; Lyons and Grzanna, 1988; Vornov and Sutin, 1983; cat: Fort et al., 1990). This projection is the primary source of noradrenaline (NE) input to Mo5 (Fuxe, 1965; Levitt and Moore, 1979; Vornov and Sutin, 1983), although other noradrenergic sources to Mo5 include cells in the subceruleus nucleus (SubC) (Grzanna et al., 1987; cat: Fort et al., 1990). Noradrenergic pathways influence both reflex and rhythmic Mo5 activity. The masseteric reflex induced by electrical stimulation of Me5 is

enhanced either by systemic treatment with the catecholamine precursor L-dihydroxyphenylalanine (L-DOPA) (Morilak and Jacobs, 1985) or by direct electrical stimulation of the A7 area (Vornov and Sutin, 1986). As with excitatory amino acids and serotonin (5hydroxytryptamine, 5-HT), the iontophoresis of NE onto anterior digastric motoneurons potentiates rhythmic responses evoked by cortical stimulation (Katakura and Chandler, 1990). Pontine cell groups projecting bilaterally to Mo5 include cells surrounding Mo5 in the supratrigeminal and intertrigeminal nuclei (Appenteng et al., 1990; Rokx et al., 1986; Travers and Norgren, 1983; Vornov and Sutin, 1983; cat: Mizuno et al., 1983; rabbit: Inoue et al., 2002). This area (rabbit: Regio h, Meesen and Olszewski, 1949) receives numerous inputs that potentially influence Mo5 neurons including projections from the red nucleus (Godefroy et al., 1998; cat: Mizuno et al., 1974) and midline pontine and medullary reticular formation (RF) (rabbit: Kolta et al., 2000). These structures were also labeled transynaptically with PRV following injections into masticatory muscles and relatively long survival times, suggesting an indirect influence on Mo5 neurons (Fay and Norgren, 1997a). The principal trigeminal nucleus, primarily the dorsomedial region, also projects bilaterally to Mo5. This part of Pr5 receives cutaneous afferent projections from the oral region (Hamilton and Norgren, 1984) and is part of a jaw opening reflex pathway (reviewed in Nakamura, 1980). A few cells in Pr5 contain GABA and

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could be inhibitory to jaw closing motoneurons during jaw opening activity (Saha et al., 1991b). Inhibition of masseteric motoneurons during jaw opening is also mediated by cells in the supratrigeminal area (Kidokoro et al., 1968), which receives projections directly from primary trigeminal afferents and indirectly via Pr5 (Mizuno, 1970; Torvik, 1956). Medullary Projections Projections to Mo5 from the medulla are from distributions of cells continuous with those from the pons (Cunningham and Sawchenko, 2000; Fay and Norgren, 1997a; Li et al., 1995; Travers and Norgren, 1983; Vornov and Sutin, 1983; cat: Holstege et al., 1977; Mizuno et al., 1983). Clusters of small-sized neurons in the parvicellular (PCRt) and intermediate zone (IRt) of the reticular formation project, primarily ipsilaterally, to Mo5, whereas clusters of larger neurons, more medially situated in the reticular formation, project bilaterally. At the level of 12 the medial distribution is primarily contralateral in the reticular nucleus of the medulla pars dorsalis (MdD) and pars ventralis (MdV). Based on dual injections of different retrograde fluorescent tracers, a small number of lateral medullary RF cells project bilaterally to Mo5 (Li et al., 1993c). Double-label studies have also demonstrated that a small number of cells in the RF project to both Mo5 and 12 or Mo5 and 7 (Li et al., 1993b). Somewhat more double-labeled cells were obtained with the Mo5 and 7 combination compared to the number obtained with Mo5 and 12. Neurons in the rostral lateral medullary RF (PCRt and IRt) appear to play an important role in the generation and coordination of rhythmic oral behavior as muscimol infusions into this region suppress both anterior digastric and lingual EMG activity associated with licking (Chen et al., 2001). Although the majority of direct premotor neurons to Mo5 use excitatory and inhibitory amino acids (Li et al., 1996; Rampon et al., 1996; guinea pig: Turman and Chandler, 1994; rabbit: Kolta et al., 2000), some cells in PCRt and MdD that project to Mo5 are cholinergic, and a population of more medially situated cells in the gigantocellular nucleus (Gi) were immunohistochemically identified as containing methionine–enkephalin (cat: Fort et al., 1990). Ultrastructural studies indicate that PCRt neurons form both symmetrical (inhibitory) and asymmetrical (excitatory) synapses on dendrites and soma in Mo5 (Mogoseanu et al., 1993). In addition, the gasseous neuromodulator nitric oxide depolarized Mo5 neurons, and NADPH-containing neurons in the medullary RF may be the source of nitrergic input to Mo5 (Abudara et al., 2001). Fluorogold (Cunningham and Sawchenko, 2000; Li et al., 1995) or fluorescent dextran injections (rabbit:

Kolta et al., 2000) into Mo5 further labeled neurons in the ventral medulla including the caudal raphe and paragigantocellular nuclei. These neurons may control state-dependent atonia during sleep (Hajnik et al., 2000) or the recruitment of masticatory muscles when respiratory drive is high (see Jacquin et al., 1999). Cells from the spinal trigeminal complex (pars oralis, interpolaris, and caudalis) project to Mo5 (Cunningham and Sawchenko, 2000; Li et al., 1995; Travers and Norgren, 1983; Vornov and Sutin, 1983; cat: Yoshida et al., 2001). The projection is predominantly ipsilateral. Differential projections to Mo5 have been studied by making injections of PRV into jaw opener and jaw closer muscles (Fay and Norgren, 1997a) and by injecting small volumes of conventional retrograde tracers directly into specific motor pools (Li et al., 1995). Some differences were apparent. Mesencephalic 5, the medial parabrachial nucleus (MBN), the supratrigeminal nucleus, and the dorsal cap of Pr5, oralis, and interpolaris all had more extensive projections to closer motoneurons in the dorsal subdivision compared to opener motoneurons in the ventromedial subdivision. In contrast, the lateral parabrachial nuclei (PB), ventrally situated neurons in the trigeminal sensory complex, and neurons in the α subdivision of the Gi all preferentially projected to opener motoneurons in the ventromedial subdivision. Serotonergic projections Serotonergic projections to Mo5 (Cropper et al., 1984; Fritschy et al., 1988; Palkovits et al., 1974; Schaffar et al., 1982, 1984; Steinbusch, 1981) account for approximately 13% of all synapses within Mo5 and are primarily axodendritic (Saha et al., 1991a; Takeuchi et al., 1983). In the midbrain and pons, retrograde labeling combined with immunohistochemistry identified the raphe dorsalis as one source of 5-HT to Mo5 (Fritschy et al., 1988; Li et al., 1993a; cat: Fort et al., 1990); however, autoradiographic studies identified the raphe centralis superior (median raphe) and not the raphe dorsalis as the principal source of raphe projections to Mo5 (cat: Bobillier et al., 1976). Further caudal in the brain stem, serotonergic-containing neurons in the raphe magnus, pallidus, and obscurus project to Mo5 as do neurons from the paragigantocellular nucleus (Fort et al., 1990; Fritschy et al., 1988; Li et al., 1993a). Electrical stimulation of the caudal raphe depolarized Mo5 neurons, an action that is blocked with serotonin antagonists (cat: Nagase et al., 1997). The motor trigeminal nucleus, together with the facial and hypoglossal nuclei show particularly dense labeling for 5-HT2A and 5-HT3 receptor subtypes (Fay and Kubin, 2000; Morales et al., 1998). Microiontophoretic application of 5-HT onto either masseter or anterior digastric motoneurons did not produce a change in firing rate (Kurasawa et al., 1990).

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Serotonin did, however, potentiate an excitatory response from these motoneurons induced by iontophoresis of glutamate into Me5, activation of jawstretch afferent fibers, or electrical stimulation of Me5 (Kurasawa et al., 1990) (cat: Ribeiro-do-Valle et al., 1991). More recently, Trueblood and colleagues demonstrated that serotonin increased both NMDA and nonNMDA-mediated responses in Mo5 neurons that were induced by electrical stimulation of Me5 (guinea pig: Trueblood et al., 1996). Serotonergic inputs are also involved in rhythmic jaw movements. Iontophoresis of 5-HT onto trigeminal motoneurons increased the number of rhythmically occurring spikes evoked by cortical stimulation (Katakura and Chandler, 1990). Serotonin may also play an important role beyond simply altering excitability to neuronal inputs. In a tissue slice preparation, Mo5 neurons showed bursting patterns when serotonin was applied (Hsiao et al., 1998), suggesting that intrinsic motoneuron membrane properties contribute to shaping rhythmic masticatory movements (guinea pig: Del Negro et al., 1999; Hsiao et al., 1997, 1998). Amino acid neurotransmitters Ultrastructural studies combined with immunohistochemistry have identified GABAergic, glycinergic, and glutaminergic synapses in the motor trigeminal nucleus (Bae et al., 2002; Yang et al., 1997a, 1997b) All three neurotransmitters formed both axosomatic and axodendritic synaptic terminals. In addition to labeled boutons exclusive for GABA (22%) and glycine (32%), glycine and GABA coexisted in 46% of the labeled boutons (Yang et al., 1997b). Similar results were obtained in an ultrastructural analysis of intracellularly filled and identified masseter motoneurons. However the proportion of coextensive GABA/glycine terminals was somewhat lower (cat: Bae et al., 1999). Axoaxonic synapses were evident in Mo5 with GABA localized presynaptically to glutamate-labeled axons (Yang et al., 1997a). Glutamate receptors on trigeminal motoneurons have not been extensively characterized but include NMDA receptors (Turman et al., 1999; Rema and Ebner, 1996). GABAergic and glycinergic neurons with projections to Mo5 originate primarily from Regio h (mostly supratrigeminal and the region medial to Mo5) and from PCRt (Li et al., 1996; Rampon et al., 1996; rabbit: Kolta et al., 2000). Fewer such cells originate from the nucleus raphe magnus and the sensory trigeminal complex (Pr5 and interpolaris). There is extensive overlap in the regional distribution of glutamatergic projections to Mo5 with GABAergic and glycinergic projections originating from both Regio h and PCRt (guinea pig: Turman and Chandler, 1994; rabbit: Kolta et al., 2000). An extensive body of literature implicates excitatory and inhibitory

amino acid neurotransmitters in the generation of rhythmic masticatory-like activity elicited by cortical stimulation (reviewed in Nakamura and Katakura, 1995), as well as natural consummatory behavior (Chen, Z. and Travers, J.B. Inactivation of amino acid receptors in the medullary reticular formation modulate and suppress ingestion and rejection in the awake rat. American Journal of Physiology, 285:R68–R83, 2003).

FACIAL NUCLEUS [REFER TO INT. –1.3 TO –2.3 MM] Intrinsic Organization Myotopic Organization Cells within 7 are segregated into several subdivisions consisting largely of cell groupings that are separated by white matter, but not by obvious cytoarchitectonic differences (Friauf and Herbert, 1985; Klein and Rhoades, 1985; Martin and Lodge, 1977; Martin et al., 1977; Papez, 1927; Semba and Egger, 1986; Szekely and Matesz, 1982; Tsai et al., 1993; Watson et al., 1982). Lateral, dorsolateral, intermediate, and medial subdivisions are readily apparent in most Nissl-stained coronal sections. These subdivisions are apparent over the rostrocaudal length of the nucleus; however, the medial division does not extend as far caudally as the intermediate and lateral subdivisions and the dorsolateral subdivision does not extend as far rostrally (Martin and Lodge, 1977; Semba and Egger, 1986). In some sections, a ventromedial cluster of cells, distinct from both the intermediate and the medial divisions, is apparent (Klein and Rhoades, 1985; Martin and Lodge, 1977; Semba and Egger, 1986) and in other sections a dorsal “cap” over the intermediate and medial divisions can also be distinguished (Faulkner et al., 1997; Friauf and Herbert, 1985; Watson et al., 1982). The divisions designated in the atlas [Int. −2.3 mm] correspond both to aggregates of neurons evident in Nissl-stained sections and myotopic representations derived from experimental studies (vide infra). The organization of 7 has been mapped for the representation of facial nerve branches within the nucleus (Faulkner et al., 1997; Martin and Lodge, 1977; Papez, 1927; Semba and Egger, 1986; Szekely and Matesz, 1982; Tsai et al., 1993) and for the representation of individual muscles (Faulkner et al., 1997; Friauf and Herbert, 1985; Klein and Rhoades, 1985; Shohara and Sakai, 1983; Watson et al., 1982). Minor discrepancies among these studies may be accounted for by the multiple innervation of a single muscle by different nerve branches and the innervation of several muscles by one nerve branch (Watson et al., 1982). Nevertheless,

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a general pattern of organization appears across several species in which dorsal muscles are represented dorsally, ventral muscles ventrally, and the anterior– posterior axis lateromedially (Friauf and Herbert, 1985; Klein and Rhoades, 1985; Martin and Lodge, 1977; Papez, 1927; cat: Courville, 1966a; Kume et al., 1978; Papez, 1927; opossum: Dom, 1982; Papez, 1927; Provis, 1977). This topographical organization for the rat is shown in Fig. 4 in which it can be seen that motoneurons innervating the proboscis are lateral within 7, peri-oral and orbital motoneurons intermediate, and facial motoneurons innervating the ear and neck medial. Detailed descriptions of the myotopic organization of individual auricular muscles indicate that although there is considerable overlap within the medial subdivision, motoneurons innervating the interscutular and posterior levator muscles are sandwiched between motoneurons innervating the transverse auricular muscle ventromedially and motoneurons innervating the anterior auricular muscle dorsally (Friauf and Herbert, 1985). Within the lateral subdivision, motoneurons innervating the intrinsic muscles of dorsally situated vibrissae are located lateral to those innervating more ventrally situated vibrissae (Klein and Rhoades, 1985). The anterior–posterior position of individual vibrissae is not myotopically organized within 7. Obicularis oculi motoneurons have also been studied in some detail (Faulkner et al., 1997). Approximately

257 motoneurons per eyelid are distributed in a dorsal “cap” that spans the mediolateral extent of 7. As with other motor nuclei, 7 is cholinergic; however, CGRP (Skofitsch and Jacobowitz, 1985) and urotensin II (Dun et al., 2001) are also present in some 7 cells as is mRNA for CCK (Sutin and Jacobowitz, 1990) and urocortin (Bittencourt et al., 1999). Other muscles innervated by 7n include the posterior digastric, stylohoid, and stapedius and are innervated by motoneurons outside the main motor nucleus (Huber and Hughson, 1926–27; Provis, 1977). Motoneurons innervating both the posterior belly of the digastric and stylohyoid (active primarily during jaw opening) are found dorsal to the anterior pole of 7 along the exiting facial root and form the accessory facial nucleus (Acs7) (Hutson et al., 1979; Rokx and van Willigen, 1985; Semba and Egger, 1986; Shohara and Sakai, 1983; Szekely and Matesz, 1982; Watson et al., 1982). Within Acs7, motoneurons innervating the stylohyoid muscle are ventral to those innervating the posterior belly of the digastric (Shohara and Sakai, 1983). The size and shape of cells in Acs7 is similar to those in 7 although there is a tendency for the soma and dendrites to be elongated in the dorsoventral direction (Szekely and Matesz, 1982). The axonal trajectory of Acs7 neurons is similar to that of other facial motoneurons but the axons form a distinct bundle parallel to the main exiting root. The location of Acs7, dorsal to

FIGURE 4 The myotopic organization and afferent projections to the facial nucleus (7) are depicted in a coronal section. Subdivisions within 7 include lateral (L), dorsolateral (DL), intermediate (I), and medial (M). Some subdivisions receive differentially strong or specific projections from structures indicated. Numerous regions in the medulla, pons, and midbrain, however, project to all subdivisions. Abbreviations: CG, central gray; Gi, gigantocellular nucleus, IRt, intermediate subdivison of the reticular formation; KF, Kolliker–Fuse nucleus; MdD, dorsal subdivision of the reticular formation; MdV, ventral subdivision of the reticular formation; M-ENK, methionin – enkephalin; NST/RF, nucleus of the solitary tract and subjacent reticular formation; PB, parabrachial nucleus; PCRt, parvocellular reticular formation; R, red nucleus; SubC, nucleus subceruleus; SP, substance P; Sp5, spinal trigeminal nucleus. Adapted from Watson et al., 1982.

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the rostral end of 7, places these cells nearly continuous with trigeminal motoneurons innervating the anterior belly of the digastric with which they share both common morphological (Szekely and Matesz, 1982) and functional characteristics (Basmajian, 1979). Facial motoneurons innervating the stapedius muscle of the middle ear have not been studied in the rat but have been located anterior to 7, adjacent to the superior olive in the cat (Lyon, 1978).

cat: Courville, 1966b; Edwards, 1972; Henkel and Edwards, 1978; Holstege et al., 1977; Takeuchi et al., 1979; opossums: Dom et al., 1973; Panneton and Martin, 1978, 1979, 1983). Similar to Mo5, there appear to be no forebrain projections to 7 in rat. In several instances, welldefined central projections to 7 differentiate between those subdivisions that innervate muscles of the orofacial region and those that innervate the facial muscles of the ear and eye.

Cytoarchitectonics and Dendritic Architecture

Midbrain Projections

Neurons within 7 are stellate shaped, ranging from 15 to 55 μm in diameter (Friauf, 1986; Martin et al., 1977; Szekely and Matesz, 1982; Watson et al., 1982). Estimates of the number of cells in 7 range from 3400 to 5800 (Friauf and Herbert, 1985; Martin et al., 1977; Semba and Egger, 1986; Watson et al., 1982) with the lower estimates including a correction factor for split cells (Martin et al., 1977; Semba and Egger, 1986). The dendritic arborization of individual HRP-filled neurons located in the medial, intermediate, and lateral subdivisions has been examined in some detail (Friauf, 1986). Primary dendrites extend in all directions but the overall dendritic field remained within a subdivision by branching primarily in the rostrocaudal direction. Nevertheless, dendritic fields of neurons in the lateral and intermediate subdivisions overlapped somewhat in the mediolateral direction but dendrites of neurons in the intermediate and medial subdivisions did not. Thus, within 7 itself, dendritic fields were functionally segregated into those involved with orofacial function (intermediate and lateral subdivisions) and those innervating the auricular muscles (medial subdivision). The overlap of dendritic fields between facial motoneurons from all the subdivisions was more apparent in the RF dorsal to 7. Unlike Mo5, there is no evidence for either interneurons or axon collaterals in 7. Nevertheless, not all neurons in 7 innervate facial muscles. A small projection from 7 to the cerebellar flocculus (cat: Kotchabhakdi and Walberg, 1977) does not originate from the cells that project to the facial musculature (Roste, 1989) and a few cells in 7 travel with the hypoglossal nerve to innervate lingual musculature (O’Reilly and Fitzgerald, 1990) (dog: Chibuzo and Cummings, 1982).

In the midbrain, cells anterior to the contralateral nucleus of the lateral lemniscus, i.e., the paralemniscal zone (cat: Henkel and Edwards, 1978) or retrorubral nucleus (Isokawa-Akesson and Komisaruk, 1987), project specifically to the medial subdivision of 7 (Hinrichsen and Watson, 1983; Li et al., 1997; Travers and Norgren, 1983). Some of these projection neurons are either glycinergic or GABAergic (Li et al., 1997). In the cat, this region of the midbrain receives input from the superior colliculus and has been implicated in pinna orientation to sound (Henkel and Edwards, 1978). Cells in the central gray, periocular nuclei (Edinger– Westphal nucleus, interstitial nucleus of Cajal, and Darkschewitch nucleus), olivary pretectal nucleus, and midbrain reticular formation also project to 7 and provide a pathway for facial movements associated with emotional behaviors (rage), reflexes of the eye (blinking), and vocal–facial behavior (Morcuende et al., 2002; cat: Fort et al., 1989; opossum: Panneton and Martin, 1983). Using the rabies virus as a transneuronal tracer, Morcuende and colleagues (2002) elegantly demonstrated specific projections from the periolary nuclei, red nucleus, oculomotor nuclei, and deep cerebellar nuclei to the orbicularis oculi motoneurons in the dorsolateral subdivision of 7. Some of the midbrain pathways appear neurochemically specific. A subset of neurons in the central grey and Edinger–Westphal nucleus, retrogradely labeled from injections of cholera toxin into 7, was immunocytochemically labeled for substance P (cat: Fort et al., 1989). Consistent with this observation, the greatest density of substance P immunoreactivity in 7 was concentrated over motoneuron pools located in the intermediate and dorsal subdivisions that innervate the orbital facial muscles (Senba et al., 1985). Retrogradely labeled cells in the olivary pretectal nucleus were immunocytochemically labeled for methionine–enkephalin (cat: Fort et al., 1989), however, the greatest enkephalinergic (ENK) immunoreactivity in 7 was concentrated over facial motoneurons in the medial subdivision that innervate the ear (Senba et al., 1985). The lateral and intermediate subdivisions of 7 receive projections from the contralateral red nucleus and

Afferent Projections Different divisions of 7 receive substantial and differential input from nuclei of the midbrain, pons, and medulla (Erzurumlu and Killackey, 1979; Fay and Norgren, 1997b; Hinrichsen and Watson, 1983; Isokawa-Akesson and Komisaruk, 1987; Li et al., 1997; Mogoseanu et al., 1994; Pinganaud et al., 1999; Senba and Tohyama, 1983a, 1983b; Travers and Norgren, 1983;

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provide a relay for cortical and cerebellar input to 7 (Hattox et al., 2002, Isokawa-Akesson and Komisaruk, 1987; Hinrichsen and Watson, 1983; Travers and Norgren, 1983) (opossum: Panneton and Martin, 1983). Electrical stimulation of the red nucleus produced movement of both the vibrissae and the eyelid (Isokawa-Akesson and Komisaruk, 1987). Pontine Pathways Differential projections to the subdivisions of 7 are evident from the pons. More cells in the ipsilateral Kolliker–Fuse (KF), supratrigeminal, and parabrachial (PB) nuclei project to the lateral and intermediate divisions of 7 than project to the medial subdivision (Isokawa-Akesson and Komisaruk, 1987; Hinrichsen and Watson, 1983; Travers and Norgren, 1983; opossum: Panneton and Martin, 1983). The PB region receives second-order vagal input (Norgren, 1978; Ricardo and Koh, 1978) and may be involved in exploratory sniffing, nasal breathing, and vibrissae movements, activities requiring coordination between respiratory and facial motoneurons. Likewise, the PB receives second-order gustatory input (Norgren, 1978) and could mediate gustatory-evoked facial responses. Projections from Pr5 to 7 are relatively sparse and are limited mainly to the lateral divisions (Erzurumlu and Killackey, 1979; Hattox et al., 2002; Travers and Norgren, 1983). A greater number of cells in the reticular formation dorsal to the superior olive at the level of Mo5 project to the medial division of 7 than project to the lateral division. Projections from the supratrigeminal region (Rokx et al., 1986) may serve as interneurons to mediate trigeminal reflex inhibition of intermediate subdivision 7 motoneurons (Minkels et al., 1995). Likewise, neurons in both the supratrigeminal area and PB may serve as interneurons for basal ganglia control of orofacial movement via projections from the entopeduncular nucleus (Takada et al., 1994). Catecholaminergic projections to 7 from the KF, nucleus subcoeruleus, and A5 region (Grzanna et al., 1987; Levitt and Moore, 1979; cat: Fort et al., 1989) project to all of the subdivisions of 7 (Senba et al., 1985). Medullary Projections In the medulla, most of the projections to 7 originate from cells in the reticular formation, bilaterally distributed in PCRt and IRt at the level of 7, and MdD and MdV further caudally (Li et al., 1997; Ter Horst et al., 1991; Travers and Norgren, 1983). A recent study using the more sensitive retrograde tracer cholera toxin, however, labeled additional populations of prefacial neurons in more medial, e.g., gigantocellular nucleus, areas of the medullary RF (Hattox et al., 2002). Doublelabel experiments with injections into both 7 and 12 or

7 and Mo5 established that as many as 10% of the labeled cells projected to both motor nuclei (Dauvergne et al., 2001; Li et al., 1993b). A somewhat higher percentage of labeled neurons (15%) projected bilaterally to 7 (Li et al., 1993c). Many of these cells are either GABAergic or glycinergic (Li et al., 1997). Ultrastructural studies confirm a monosynaptic connection between the medullary RF and 7 and indicate the presence of both symmetrical and asymmetrical synapses (Mogoseanu et al., 1994). Different regions of the medullary RF project to different divisions of 7, although these differences are not as pronounced as projections from the midbrain and pons. At the level of 12 many reticular neurons, bilaterally distributed in MdD, MdV, and along the medial border of the ipsilateral Sp5C project to the intermediate and lateral divisions of 7 (Hinrichsen and Watson, 1983; Travers and Norgren, 1983). Cells in the caudal medulla that project to the medial subdivision of 7 tend to be ventrally located, i.e., dorsal to the lateral reticular nucleus and medial to Sp5C, and include cells in and around the nucleus ambiguus (Hinrichsen and Watson, 1983; Isokawa-Akesson and Komisaruk, 1987; Senba and Tohyama, 1983a; Travers and Norgren, 1983). Reticular neurons that project preferentially to the medial division of 7 extend into the cervical levels of the spinal cord, originating in the intermediolateral gray column and lateral part of the ventral horn (Hinrichsen and Watson, 1983; Senba and Tohyama, 1983a; cat: Takeuchi et al., 1987). Disynaptic spinofacial reflexes may be mediated in part by spinofacial projection neurons that receive primary afferent terminals via dendrites that extend into spinal lamina IV (cat: Takeuchi et al., 1987). Some projections reach the facial nucleus from the ventral medulla and may be involved with respiratory (Fay and Norgren, 1997b; Hwang et al., 1998) or gasping (Fung et al., 1997) activity. Other medullary projections to 7 originate from the sensory trigeminal complex (Erzurumlu and Killackey, 1979; Hattox et al., 2002, Hinrichsen and Watson, 1983; Pinganaud et al., 1999; Travers and Norgren, 1983). The most complete description of these pathways employed both anterograde and retrograde tracers and further tested for collateral projections to 12 using dual tracers (Pinganaud et al., 1999). All subdivisions of the sensory complex projected to 7 but there were substantive differences in the patterns of projections. Neurons from the spinal trigeminal complex were located throughout the nuclei but clustered preferentially within the dorsal part of 5Sp. Projections were densest to the lateral subdivisions of 7 and decreased systematically more medially. The projection from principal 5 was from neurons almost exclusively in the ventral part of the nucleus. These trigeminofacial and spinal dorsal horn

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projections appear topographically organized such that facial motoneurons receive cutaneous input from areas overlying the muscles they innervate (Panneton and Martin, 1983). Thus, projections from the mandibular branch of 5 to the dorsal region of 5Sp can influence vibrissae motoneurons located intermediate and lateral in 7. Likewise, periocular afferents that terminate in the ventral part of 5P project onto obicularis oculi motoneurons distributed dorsally within 7 to mediate a disynaptic blink reflex (Pellegrini et al., 1995; cat: Hiraoka and Shimamura, 1977). Minor projections to 7 include projections to the medial division of 7 from cells in the medial vestibular nuclei (Hinrichsen and Watson, 1983; cat: Shaw and Baker, 1983; opossum: Panneton and Martin, 1983) and projections from the nucleus of the solitary tract (sol) to the intermediate subdivision (Norgren, 1978; Travers and Norgren, 1983). Cholinergic and peptidergic pathways Cholinergic terminals in 7 have been ultrastructurally characterized (Davidoff and Irintchev, 1986). One source of this cholinergic input is from cells in the MdD, lateral to 12 and ventral to the sol (cat: Fort et al., 1989). Substance P projections to the intermediate and dorsolateral subdivisions of 7 also originate from this region of the reticular formation (Senba and Tohyama, 1985). Other sources of substance P projections to 7 include the ventrolateral medullary reticular formation and the nuclei raphe pallidus and obscurus. Neurons in these regions, immunohistochemically identified as containing both substance P and 5-HT, project primarily to the ventral regions of 7 (cat: Fort et al., 1989). The ventrolateral medullary reticular formation is also a source of ENK projections to 7, primarily to the medial subdivision (Finley et al., 1981; Senba and Tohyama, 1983a, 1983b; cat: Fort et al., 1989). As with substance P, enkephalinergic projection neurons to 7 from the nuclei raphe obscurus and pallidus also contained 5-HT (cat: Fort et al., 1989). Serotonergic pathways Serotonergic fibers have been identified in 7 (Aghajanian and McCall, 1980; Palkovits et al., 1974; Steinbusch, 1981) and appear either throughout 7 (Senba et al., 1985) or concentrated in the ventrolateral subdivision (cat: Fort et al., 1989). The sources of serotonergic projections appear to be raphe nuclei obscurus, magnus, and pallidus (Senba et al., 1985; cat: Fort et al., 1989) and form both axosomatic and axodendritic synapses (Aghajanian and McCall, 1980). The iontophoretic application of 5-HT does not directly activate facial motor neurons but facilitates excitatory responses induced by other means (McCall and Aghajanian, 1979). In vivo studies indicate that this facilitation is postsynaptic and

mediated by receptor subtypes 5-HT2 and 5-HT1C (Rasmussen and Aghajanian, 1990).

HYPOGLOSSAL NUCLEUS Intrinsic Organization Myotopic Organization The tongue consists of both extrinsic (genioglossus, GG; styloglossus, STY; hyoglossus, HY; palatoglossus, PG) and intrinsic (vertical, V; transverse, T; superior longitudinal, SL; inferior longitudinal, IL) musculature. Although the geniohyoid (GH) is not a true lingual muscle (Chibuzo and Cummings, 1982), it has a common embryological origin with lingual muscles (Edgeworth, 1907) and often functions together with the GG, especially during tongue protrusion (Cunningham and Basmajian, 1969; Travers and Jackson, 1992). The palatoglossus (poorly defined in rat) functions as a pharyngeal muscle (Lowe, 1981). Motoneurons innervating the lingual musculature originate from 12 and travel in the hypoglossal nerve (12n) (Fig. 5). However, a small population of motoneurons within 7 exit the brain stem via 12n to innervate lingual muscles (O’Reilly and Fitzgerald, 1990; dog: Chibuzo and Cummings, 1982). A few motoneurons from the ventral region of 12 travel in the ansa cervicalis to innervate motor fibers of the intrinsic and GH muscles (O’Reilly and Fitzgerald, 1990). The hypoglossal nucleus consists of a dorsal and ventral subdivision (Aldes, 1995; Barnard, 1940; Kitamura et al., 1983, 1985; Krammer et al., 1979; Lewis et al., 1971; McClung and Goldberg, 1999; Odutola, 1976; O’Reilly and Fitzgerald, 1990; Uemura-Sumi et al., 1988). Geniohyoid motoneurons form a distinct cluster of neurons ventrolateral to the main nucleus. Axons arising from the dorsal subdivision travel in the lateral branch of 12n to innervate muscles that retract the tongue; ventral subdivision axons, together with GH axons, travel in the medial branch to innervate muscles that protrude the tongue. In general, motoneurons innervating the intrinsic musculature are medial to those innervating the extrinsic musculature. The main cell groupings of 12 are schematically depicted in Fig. 6. Each styloglossus muscle is innervated by approximately 45 motoneurons located at the rostral one-third of the dorsal subdivision (Guo et al., 1996; McClung and Goldberg, 1999). The hyoglossus is innervated by approximately twice as many motoneurons in the dorsal subdivision, caudal to the STY motoneuron distribution. Intrinsic lingual motoneurons that retract the tongue are also located in the dorsal subdivision. The hydrostatic model of lingual function predicts that

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IL V/T

SL

HY

V/T

GG

STY

R

GG

GH

SL V/T

HY GG GH

C

FIGURE 5 Organization of the hypoglossal nerve. Abbreviations: AC, ansa cervicalis; C1, C2, C3, ventral roots of first three cervical nerves; DXII, distal hypoglossal nerve; GG genioglossus; GH, geniohyoid; HB, hyoid belly of geniohyoid; HG, hyoglossal; JNG, jugulonodose ganglion; LD, lateral division of hypoglossal nerve; MB, mandibular belly of geniohyoid; MD, medial division of hypoglossal nerve; PXII proximal trunk of hypoglossal nerve; S, sympathetic; Sal, salivatory nucleus; SCG, superior cervical ganglion; SG, styloglossus; 7, facial nucleus, 7n, facial nerve; 12d, dorsal subdivision of hypoglossal nucleus; 12v, ventral subdivision of hypoglossal nucleus. From O’Reilly and Fitzgerald, 1990.

contraction of (inferior and superior) longitudinal muscles shorten the tongue (Kier and Smith, 1985), and this prediction has been borne out directly by measuring retraction forces following microstimulation of individual longitudinal motor units (Sokoloff, 2000; Sutlive et al., 1999). Superior and inferior longitudinal motoneurons are located medial to STY and HY motoneurons with the inferior longitudinal motoneurons distributed rostral to superior longitudinal motoneurons. The number of intrinsic motoneurons greatly exceeds extrinsic motoneurons, with longitudinal motoneurons making up about 75% of neurons in the dorsal subdivision (McClung and Goldberg, 1999). Motoneurons that protrude the tongue are located in the ventral compartment with (extrinsic) genioglossus motoneurons located lateral to motoneurons innervating the intrinsic horizontal and vertical muscles (Aldes, 1995; Uemura-Sumi et al., 1988). Motoneurons in the ventral compartment innervating the intrinsic muscles are topographically represented such that the

FIGURE 6 Myotopic organization of 12 from rostral (R) to caudal (C). Muscle abbreviations: GG, genioglossus; GH, geniohyoid; HY, hyoglossus; IL, inferior longitudinal; SL, superior longitudinal; STY, styloglossus; T, transverse; V, vertical. Adapted from Aldes, 1995.

tip of the tongue is represented caudal within 12 and the base of the tongue rostral (Aldes, 1995). Immunohistological labeling for various neurotransmitters, their precursors, derivatives, or receptors frequently demarcate or fall within myotopic subdivisions of 12. The distribution of catecholaminergic fibers, for example, determined by immunohistochemical labeling of tyrosine hydroxylase (TH) was evident throughout 12 but particularly dense in the ventromedial quadrant of the caudal half of 12 (Aldes et al., 1988b). In contrast, 5-HT immunoreactivity was particularly dense in both the dorsal division of the caudal two-thirds of 12 and in the ventrolateral region, over GH motoneurons. Although receptor binding sites for 5-HT1A and 5-HT1B were particularly dense in the dorsal subdivision of 12 (Manaker and Zucchi, 1998), corresponding to the distribution of immunoreactive 5-HT terminals (Aldes et al., 1988b), similar binding sites were noticeably absent in the ventrolateral region of 12 (GH motoneurons) despite the presence of concentrations of immunoreactive terminals. Immunocytochemical labeling of the converting enzymes dopamine β-hydroxylase (DBH) and phenylethanolamine (PNMT) was greatest in the ventromedial region of 12. This pattern of label confirmed the distribution of catecholamine (CA) innervation observed previously and further specified

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NE as the major source of CA innervation of 12 (Aldes et al., 1988a). Ionotropic glutamate receptor subunits also show some myotopic variation within 12 (Garcia del Cano et al., 1999). In particular, GluR1 (AMPA) was particularly absent in 12 motoneurons except for caudally located cells. In contrast, other AMPA (GluR2/3, GluR4), kainate (GluR5/6/7), and NMDA (NR1) receptors were more evenly distributed on 12 motoneurons. Putative interneurons in the dorsolateral quadrant of 12 stained well for GluR1. Physiological studies indicate that NMDA and non-NMDA receptors are co- localized within 12 (O’Brien et al., 1997). Immunohistochemical labeling for adenosine deaminase was restricted to the dorsal division of 12 (Senba et al., 1987a). Adenosine may function in neurotransmission or other neuromodulatory actions (Cooper et al., 1996; Stone, 1981). Adenosine labeling was transient in both 12 and (oro)facial motoneurons during development and disappeared by postnatal day 25. A possible correlation between the transient expression of adenosine and suckling behavior was noted by Senba and colleagues (Senba et al., 1987b). Enkephalinergic input to 12 was also differentially distributed (Aldes, 1998). Immunohistochemically labeled ENK terminals were particularly dense over the ventrolateral subdivision, corresponding to the location of GG motoneurons. Cytoarchitectonics Each half of the hypoglossal nucleus contains approximately 3500 neurons (Lewis et al., 1971). Most of the cells within 12 are motoneurons but there is anatomical evidence (Boone and Aldes, 1984; Cooper, 1981; Takasu and Hashimoto, 1988; monkey: Takasu et al., 1987) for a small population of interneurons, confirming evidence obtained earlier from neurophysiological studies (cat: Green and Negishi, 1963; Sumi, 1969). Motoneurons Cytoarchitectonic characteristics of size, shape, and dendritic orientation further differentiate motoneurons in different regions of 12 (Aldes, 1995; Boone and Aldes, 1984; Cooper, 1981; Kitamura et al., 1983; McClung and Goldberg, 1999; Odutola, 1976). Cells in the dorsal division appear fusiform, oriented along the mediolateral axis and range from 17 to 40 μm in diameter. Dorsal subdivision motoneurons innervating intrinsic muscles were significantly smaller (X=25.7 μm) compared to extrinsic muscle motoneurons (X=30.25 μm) (derived from McClung and Goldberg, 1999). Likewise, cells in the central region of the ventral subdivision that innervate intrinsic muscles were smaller on average (X=23 μm) and more globular in shape than the larger more lateral multipolar motoneurons in the ventral division (X=29.5 μm) (Aldes, 1995). Cell diameters (average

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= 28.7 μm) were unimodally distributed in the ventrolateral division, suggesting to Kitamura and colleagues (Kitamura et al., 1983) a lack of γ efferents, but subsequent studies (O’Reilly and Fitzgerald, 1990; Smith, 1989) have clearly confirmed the earlier observation (Maier, 1979) that both the GH and longitudinal muscles possess a few muscle spindles. Hypoglossal motoneurons are cholinergic (e.g., Connaughton et al., 1986; Wamsley et al., 1981) and a subset colocalize GABA (Davidoff and Schulze, 1988). Peptides in 12 motoneurons include both galanin and CGRP (Moore, 1989) as well as urotensin II (Dun et al., 2001). Messenger RNA for both urocortin (Bittencourt et al., 1999) and CCK (Abelson and Micevych, 1991; Cortes et al., 1990) is also found in 12 motoneurons. Interneurons Interneurons in 12 were initially identified as cells not labeled following the application of HRP to lingual muscles (Boone and Aldes, 1984). These neurons were smaller (10–18 μm) than labeled motoneurons (25–50 μm), had fewer dendritic processes, and were confined to the ventrolateral or dorsolateral borders of the nucleus. Evidence obtained from Golgiimpregnated neurons localized to 12 and subsequently (re)sectioned for ultrastructural analysis confirmed that small neurons make synaptic contact within the borders of 12 (Takasu and Hashimoto, 1988). An example obtained from a juvenile rat demonstrated multiple axodendritic synapses between an axon collateral to a small interneuron and a hypoglossal motoneuron. The GABAergic nature of interneurons is suggested by both radioactive uptake studies and immunohistochemical staining for GABA and its synthetic enzyme, glutamic acid decarboxylase (GAD) (Aldes et al., 1988c; Simon et al., 1985; monkey: Takasu et al., 1987). Immunocytochemical labeling of GABA and GAD fibers were evident throughout 12, although staining appeared denser in ventral regions of the nucleus, particularly in the caudal half (Aldes et al., 1988c). A few positively stained GABAergic neurons were located laterally within 12 and along the dorsolateral border, i.e., in the same location as HRP-identified interneurons. Neurons staining positive for GABA were small, 12–18 μm, round, and few in number. GABA is inhibitory to 12 motoneurons (Altmann et al., 1972; Gregg and Carpenter, 1982). Interneurons in 12 may also have collaterals outside the nucleus (Popratiloff et al., 2001). Injections of Fluorogold into the facial nucleus consistently labeled a few small neurons laterally located within 12. Dendritic Architecture Neurons in 12 can be further classified based on dendritic morphology. Using Golgi-stained material, Ramon Y. Cajal (1972) designated hypoglossal cells as

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“external” when dendrites extended beyond the borders of the nucleus and “internal” when dendrites were confined to the nuclear region. Two particularly dense dendritic zones within 12 include the dorsalmost part of the nucleus, just ventral to the dorsal motor nucleus of the vagus (10) and the midline of the nucleus around the central canal (Lorente de No, 1947). Some motoneuron dendrites extend into the contralateral nucleus (Altschuler et al., 1994; Wan et al., 1982). Dendritic orientation of cells labeled from either the Golgi technique (Odutola, 1976) or cholera toxin HRP (Wan et al., 1982) further distinguish between cells in the various subdivisions. Dendrites of cells of the dorsal division can be either internal or external (Odutola, 1976). External dendrites may reach 1 mm in length, extending laterally into the adjacent reticular formation (MdD and MdV) and, in some cases, reach Sp5 (Wan et al., 1982). Dendrites from these cells also extend into the ipsilateral SOL and a very few traverse 10 to reach the contralateral nucleus of the solitary tract (SOL) (Wan et al., 1982). Dendritic fields from cells in the ventral division include an apical dendrite that ramifies within the dorsal divisions of 12 and basal dendrites that ramify laterally within 12 and the adjacent reticular formation (Odutola, 1976), extending ventrally into the medial longitudinal fasciculus (MLF) and nucleus raphe obscurus (Wan et al., 1982). Dendrites from multipolar cells in the center of 12 fan out to ramify throughout the nucleus (Odutola, 1976). Dendrites from motoneurons within a given compartment also form dendritic bundles in the rostral/caudal dimension (Fig.7) and may provide a substrate for simultaneous activation of synergistic motoneurons (Altschuler et al., 1994).

Afferent Projections Afferent projections reach 12 from cells in the midbrain, pons, and medulla (Aldes, 1980; Borke et al., 1983; Cooper and Fritz, 1981; Dobbins and Feldman, 1995; Fay and Norgren, 1997c; Travers and Norgren, 1983). Many of these projections are from specific RF regions organizing the complex oromotor functions of mastication, licking, swallowing, and respiration (Hwang and St. John, 1987; Nakamura and Katakura, 1995; Rekling and Feldman, 1998; Travers et al., 1997). Other RF projections control state-dependent muscle tone in the tongue (Hajnik et al., 2000) and projections from the nucleus of the solitary tract and sensory trigeminal complex mediate lingual reflexes (Lowe, 1981). Forebrain Pathways There appear to be no direct cortical projections to 12 in the rat (Travers and Norgren, 1983; Walberg, 1957).

FIGURE 7 Photomicrograph of cholera toxin-labeled geniohyoid motoneurons in horizontal plane. Arrowheads point to dendritic bundles running in rostrocaudal orientation. From Altschuler et al., 1994.

Descending pathways, however, activate oromotor responses of mastication and licking that are organized in the brain stem reticular formation (reviewed in Nakamura and Katakura, 1995; Travers et al., 1997). Damage to a small zone of anterior-lateral (orbital) cortex inhibits some tongue protrusion activity but not protrusions occurring during rhythmic licking (Whishaw and Kolb, 1989). Cortical sites that support microstimulation-elicited ororhythmic responses send projections to widespread regions of the lateral medullary and pontine reticular formation (Zhang and Sasamoto, 1990). These regions in the brain stem overlap the location of neurons that respond rhythmically during licking (Travers et al., 2000) or to cortical microstimulation (Moriyama, 1987; cat: Sahara et al., 1996) as well as the location of neurons that project to 12 (Travers and Norgren, 1983; cat: Sahara et al., 1996). Descending projections from limbic structures to the midbrain provide additional pathways through which forebrain structures involved in feeding can ultimately influence 12 motoneurons (White, 1986). Midbrain Projections In the midbrain, a few cells in the contralateral reticular formation project to 12 as demonstrated with HRP (Aldes, 1980; Aldes and Boone, 1984; Travers and Norgren, 1983). A similar study (opossum: Panneton and Martin, 1979), however, localized midbrain projections primarily from within the central gray. Some controversy exists as to whether Me5 projects directly to 12 via Probst’s tract (see Lowe, 1981). Although this projection was evident using degeneration techniques (cat: Mizuno and Sauerland, 1970; Szentagothai, 1948), relatively few Me5 cells were labeled in the retrograde

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HRP experiments. This projection, however, may be to dendrites of 12 motoneurons extending laterally into the adjacent RF (Matesz, 1981; Ruggiero et al., 1982) and thus provide a substrate for proprioceptive interactions between the muscles of mastication and lingual muscles (Lowe, 1981). Proprioceptive information from jaw muscles may also reach 12 through the supratrigeminal region in the pons (Mizuno, 1970). This region receives monosynaptic input from Me5 (Matesz, 1981) and projects to 12 (Travers and Norgren, 1983). Pontine Projections Several distributions of neurons in proximity to Mo5 project to 12. These include the ventral PB and intertrigeminal nuclei (bilaterally), the KF (ipsilaterally), and the supratrigeminal area (Aldes, 1980; Borke et al., 1983; Fay and Norgren, 1997c; Luo et al., 2001; Saper and Loewy, 1980; Travers and Norgren, 1983). The KF, part of the lateral tegmental group of catecholaminecontaining cells (Stevens et al., 1982), may be a source of NE projections to 12 (Levitt and Moore, 1979). Another source of pontine CA projections to 12 is from the SubC (Aldes, 1990). Injections of HRP into 12 labeled scattered cells ventral to locus coeruleus and medial to Mo5 in the SubC and injections of tritiated amino acids into SubC produced reciprocal anterograde label within 12. Projections from the supratrigeminal region were prominent to the dorsal subdivision of 12 (Luo et al., 2001). A subpopulation of supratrigeminal neurons stained for GABA or glycine as did other projection neurons to 12 from neuron groups near Mo5 (Li et al., 1997). A primarily ipsilateral projection from Pr5 reaches 12 from a distribution of relatively large cells (30–50 μm) located dorsomedially in the caudal two-thirds of the nucleus (Aldes, 1980; Aldes and Boone, 1985; Borke et al., 1983; Travers and Norgren, 1983). Pr5 is topographically organized such that somatosensory afferent axons from the tongue project to the dorsomedial region (Hamilton and Norgren, 1984). Thus, somatosensory input from the tongue can influence hypoglossal motoneurons through disynaptic pathways. Similarly, descending cortical fibers from the face region of somatosensory cortex can influence lingual motoneurons via Pr5 (discussed in Aldes and Boone, 1985). Medullary Projections Reticular formation The majority of cells projecting to 12 originate from the medullary RF (Aldes, 1980; Borke et al., 1983; Borke and Nau, 1985; Cooper and Fritz, 1981; Dobbins and Feldman, 1995; Fay and Norgren 1997c; Travers and Norgren, 1983; Zhang and Luo, 2003; cat: Basbaum et al., 1978; Holstege et al., 1977). The central core of the medullary brain stem, encompassing most

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of the Gi, is not involved in these direct projections; however, there are direct projections from the midline raphe and ventral subdivisions of the gigantocellular and paragigantocellular nuclei (Aldes et al., 1989; Henry and Manaker, 1998; Manaker and Tischler, 1993; Travers and Rinaman, 2002; cat: Bobillier et al., 1976). Dorsal medulla In the dorsal medulla, reticular neurons projecting to 12 form a continuous column along the rostrocaudal axis. At the spinomedullary junction, projection cells are bilaterally distributed lateral to 12 in the MdD and MdV (less). Rostral to 12, these cells are bilaterally distributed in the IRt and, to a lesser extent, in the PCRt. It is unclear if there is spatial segregation of reticular neuron projecting to different lingual motoneurons. Neurons projecting to lingual retractor motoneurons have been described as either dorsal to those projecting to protrudors (Dobbins and Feldman, 1995) or ventral (Fay and Norgren, 1997c). Some RF neurons project to more than one oromotor nucleus (Amri et al., 1990; Li et al., 1993b; Popratiloff et al., 2001), and some RF interneurons project to multiple pools of 12 motoneurons (Travers and Rinaman, 2002). Double-label studies with injections of retrograde fluorescent markers into combinations of 12, Mo5, and 7 double-labeled approximately 5–6% of labeled RF neurons, scattered among single projection cells. The medullary RF is a likely source of cholinergic input to 12 (Connaughton et al., 1986; Davidoff and Irintchev, 1986). Cells in the medullary RF projecting to 12 also stain positively for GABA and glycine (Li et al., 1997). The dorsal medullary RF projections to 12 are likely involved in the lingual component of consummatory behavior. Neurons lateral to 12 are active during both licking and swallowing (Gestreau et al., 1996; Ono et al., 1998; Travers et al., 2000), and the area subjacent to the caudal SOL is part of the central pattern generator for swallowing (Jean, 2001). Projection neurons more rostral in the medulla in the IRt are also active during licking (Travers et al., 2000) and muscimol infusions into this region suppress this behavior (Chen et al., 2001). Ventral medulla Neurons in the ventral RF form a critical substrate for respiratory rhythmogenesis (reviewed in Rekling and Feldman, 1998) and direct projections to 12 have been demonstrated in both rat (Lipski et al., 1994) (Fig. 8) and cat (Ono et al., 1994). In the rat, neurons active during both inspiration and expiration were found quite far rostral in the ventrolateral medulla, just caudal to the facial nucleus, and appeared to overlap with the rostral ventrolateral reticular formation (Ezure et al., 1988). In cat, some neurons in the ventral medulla project to both the phrenic nucleus and 12 (Ono et al., 1994).

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FIGURE 8 Reconstruction of an intracellularly filled respiratoryrelated neuron in the ventral medulla show axons (no dendrites) that ramify in both dorsal and ventral subdivisions of 12 and the nucleus of the solitary tract. Arrow indicates spinally projecting axonal branch. Abbreviations: PY, pyramidal tract; SOL, nucleus of the solitary tract; 12, hypoglossal nucleus. From Lipski et al., 1994.

Neurons in the paragigantocellular and ventral gigantocellular nuclei also project to 12. These neurons were not well-labeled using HRP (Borke et al., 1983; Travers and Norgren, 1983; Yang et al., 1995), but were somewhat more visible using Fluorogold (Manaker and Tischler, 1993), Fast blue (Yang et al., 1995), or cholera toxin (Travers and Rinaman, 2002). Neurons in this region control muscle atonia during sleep (Hajnik et al., 2000). Stimulation of ventral and medial RF sites including the ventral gigantocellular and paragigantocellular nuclei decreased muscle tone in neck muscles and it was recently reported that stimulation of this region released an inhibitory neurotransmitter, glycine, in 12 (Kodama et al., 2000). Other reports of electrical stimulation in gigantocellular nucleus, however, in sites that appear to overlap with the previous study, suggest a primarily excitatory effect on hypoglossal motoneurons (Yang et al., 1995). Raphe nuclei Neurons in the caudal raphe (obscurus, pallidus, and magnus) project to 12 (Henry and Manaker, 1998; Manaker and Tischler, 1993). Many projection neurons, mostly from obscurus and palidus stained for 5-HT and serotonergic terminals in 12 have been identified ultrastructurally (Aldes et al., 1989). A number of 5-HT projection neurons also contained substance P; a smaller number contained enkephalin (Henry and Manaker, 1998). The physiological effects of 5-HT on 12 motoneurons are complex. Microinjections of 5-HT directly onto 12 motoneurons increased activity levels (Kubin et al.,

1992) and microdialysis delivery of 5-HT raised GG activity in an awake rat across sleep–wake cycles (Jelev et al., 2001). On the other hand, hypoglossal motoneurons were inhibited by the infusion of 5-HT in a tissue slice preparation, and response decrements also occurred to electrical or chemical excitation of the raphe (Monteau et al., 1990; Morin et al., 1990). It has been proposed that serotonin may depolarize hypoglossal motoneurons postsynaptically, but inhibit excitatory glutamatergic input presynaptically (Singer and Berger, 1996). Because raphe neurons are more active during awake states, glutamatergic inputs, possibly respiratory-related could thus be selectively suppressed and allow raphe neurons to augment hypoglossus function associated with other oromotor functions such as feeding. Binding sites for 5-HT1A and 5-HT1B receptors are widely distributed across the hypoglossal nucleus but are sparse in the ventrolateral nucleus that contains geniohyoid motoneurons (Manaker and Zucchi, 1998). Messenger RNA for 5-HT1B, 5-HT2A, and 5-HT2B receptors were found in 12 but mRNA for 5-HT1A was not (Okabe et al., 1997) Trigeminal and solitary nucleus input A second bilateral distribution of medullary cells projecting to 12 originates from the spinal trigeminal complex (Aldes, 1980; Borke et al., 1983, 1988; Borke and Nau, 1987; Pinganaud et al., 1999; Travers and Norgren, 1983). Projection neurons are clustered primarily in the dorsal part of Sp5 in the area receiving the bulk of the trigeminal afferents from the tongue (Jacquin et al., 1983), although some projection neurons are scattered throughout the nuclei. Similarly, small injections of anterograde tracers into Sp5C labeled both dorsal and ventral subdivisions of 12 (Pinganad et al., 1999). In the cat these neurons are associated with polysynaptic trigeminolingual reflexes elicited from electrical stimulation of the trigeminal nerve (reviewed in Lowe, 1981). Multiple routes from Sp5 to 12 include projections via short axon interneurons in the adjacent RF (Scheibel, 1955a, 1955b; Valverde, 1961) and by the extension of 12 motoneuron dendrites into this area (Odutola, 1976; Wan et al., 1982). A more recent study using anterograde tracers injected into Sp5 demonstrated that most RF projections terminate in the PCRt and MdD with fewer in the IRt and MdV and the least in the Gi (Zerari-Mailly et al., 2001). Cortical projections to Sp5 have also been proposed as part of a corticohypoglossal pathway in the cat (Porter, 1967). Inhibitory interneurons adjacent to 12 have also been implicated in trigeminohypoglossal reflexes (Sumino and Nakamura, 1974). Relatively few cells from the SOL project to 12. Those that do are ipsilateral, located primarily in the caudal half of the nucleus (Borke et al., 1983; Norgren, 1978; Travers and Norgren, 1983) in an area that receives

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afferents via the glossopharyngeal and vagus nerves (Contreras et al., 1982; Hamilton and Norgren, 1984; Torvik, 1956). Hypoglossal reflexes induced by stimulation of these nerves (reviewed in Lowe, 1981) may be mediated by interneurons in SOL or by direct synaptic contact via hypoglossal dendrites that extend into SOL (Odutola, 1976; Wan et al., 1982). Neuropeptide input Synaptic terminals within 12 that contained substance P and enkephalins have been described in studies combining immunohistochemistry and electron microscopy (Connaughton et al., 1986; Gatti et al., 1996; Hinrichsen and Weston, 1999). Substance P terminals appeared distributed throughout 12 (Gatti et al., 1996); however, ENK terminals were densest in the ventral compartment of 12, particularly at rostral levels that include GG muscles (Aldes, 1998). ENK endings were predominantly axodendritic; however, substance P terminals can be either axosomatic or axodendritic. In rat, it appears that substance P terminals are on interneurons (Hinrichsen and Weston, 1999), but in cat there is clear anatomical and physiological data for substance P terminals directly onto motoneuron soma and dendrites (Gatti et al., 1996, 1999). A major source of substance P is probably from the caudal raphe (Henry and Manaker, 1998). Substance P and enkephalin colocalize in different populations of 5-HT-containing cells in the caudal raphe; however, this represents a relatively sparse ENK projection (Henry and Manaker, 1998). Other sources of ENK input to 12 could come from the medullary RF, the SOL, or the PB (discussed in Connaughton et al., 1986), the spinal trigeminal complex (Borke and Nau, 1987), or sites implicated in respiratory function with projections to 12 including the KF and ventrolateral medullary RF (discussed in Aldes, 1998). Another peptide present in 12 is thyrotropinreleasing hormone (TRH) (Hokfelt et al., 1975; Kubek et al., 1983). In a tissue slice preparation, TRH depolarized 12 neurons, increased spontaneous activity, and potentiated responses evoked by iontophoresis of NMDA (Rekling, 1990, 1992). Lingual proprioceptors Another set of afferent projections to 12 are proprioceptive (Lowe, 1981). Approximately six muscle spindles were consistently observed in the GH (O’Reilly and Fitzgerald, 1990; Smith, 1989) and a smaller, more variable number in the longitudinal, HY, and GG muscles. No muscle spindles were observed in the other extrinsic or intrinsic muscles. Although few in number, these muscle spindles are proportional to those found in primates when expressed as a ratio of spindle number/muscle volume (Smith, 1989). Intrinsic muscle proprioceptor afferent fibers with cell bodies in spinal C2 and C3 travel in the ansa

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cervicalis; those from extrinsic muscles (including the GH) have their cell bodies in the jugulonodose or trigeminal ganglia and travel with 12n (O’Reilly and Fitzgerald, 1990; Neuhuber and Mysicka, 1980; dog: Chibuzo and Cummings, 1981: cat: Nazruddin et al., 1989). Fibers traveling in 12n with cell bodies located in the trigeminal or jugulonodose ganglia terminated in Sp5i and Sp5c (Fig. 5). In addition, 12n axons with cell bodies in the trigeminal ganglion also terminated in Pr5 in contrast to the central end of jugulonodose ganglion cells that terminated in the caudal SOL. Spinal afferents from the tongue terminated in lamina I and V of the cervical dorsal horn. In all cases, terminal endings were not evident within 12, indicating a lack of monosynaptic contacts (see also Lodge et al., 1973). Lingual proprioceptive input also influences other related oral motor nuclei and electrical stimulation of 12n elicits responses in both facial (Tanaka, 1975) and trigeminal motoneurons (e.g., cat: Nakamura et al., 1978; Takata et al., 1979). These projections do not constitute a complete description of all the direct influences on hypoglossal motoneurons. Extensive dendritic arborizations of 12 motoneurons throughout the adjacent RF, nucleus raphe magnus, and MLF make further monosynaptic influence from other regions of the CNS a certainty. Cerebellar influences on 12, for example, may be mediated by 12 motoneuron dendrites in the MLF (Wan et al., 1982). Projections onto 12 dendrites extending into the RF may also mediate sensory input to 12 from visual, somatosensory, and vestibular sources (Elmund et al., 1983; Mameli and Melis, 1992; Mameli et al., 1988).

SUMMARY AND CONCLUSIONS Each of the orofacial motor nuclei is organized into multiple subdivisions defined both myotopically and cytoarchitectonically. In the case of Mo5 and 12, muscle antagonists are segregated in distinct subdivisions in which motor agonists have adjacent representation. In 7, a pronounced topography related to peripheral musculature is evident. Myotopic subdivisions are further defined by the specificity of the central projections they receive. Mesencephalic 5 and the supratrigeminal region project preferentially to jaw closer motoneurons in Mo5; lateral PB and Gi project preferentially to jaw-opener motoneurons. Within 7, aural musculature motoneurons in the medial subdivision receive highly specific projections from the paralemniscal zone in the midbrain and ENK projections from the olivary pretectal nucleus. These projections contrast with those to the orally related intermediate and lateral divisions of 7 from the KF and ventral PB in the pons and the MdD in the medulla.

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Although these central projections serve to differentiate between motor nuclei subdivisions, there is also overwhelming evidence of overlap in the central projections to the oromotor nuclei that unifies them functionally. Neurons in the PCRt and IRt rostrally, the MdD and MdV caudally, and the ventral medulla have extensive projections to the oromotor nuclei, and double-label studies reveal single neurons within these fields with multiple projections. Although such projections no doubt serve as outputs from central pattern generators for mastication, swallowing, and respiration, systems, driving the oromotor nuclei may more accurately be characterized as interacting multifunctional systems (Marder, 2000; Marder and Calabrese, 1996). Such systems must not only act as pattern generators to, for example coordinate between lingual and trigeminal motoneurons during mastication but must also coordinate muscle participation between competing functions, e.g., respiration and swallowing. The substrates appear multifunctional in that neurons involved in patterning one function, e.g., licking may also participate in another, e.g., swallowing (Ono et al., 1998; Travers et al., 2000). Following the progress made in invertebrate systems (Marder, 2000; Marder and Calabrese, 1996), the identification of neuromodulators within the afferent pathways driving oromotoneurons as well as the motoneurons themselves provides future direction for exploring complex oromotor function.

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13 Central Nervous System Control of Micturition GERT HOLSTEGE Department of Anatomy and Embryology Faculty of Medical Sciences, University of Groningen Groningen, The Netherlands

Micturition does not take place at random, but is directly related to the survival of the individual or species. An individual animal micturates only when the environment is relatively safe, because the animal is vulnerable during the release of urine. Moreover, many animals use urine as a marker for territorial demarcation or sexual attraction (by leaving a scent trace a female lets the males know that she is in estrus). These facts explain why micturition is under strong control of the emotional motor system (Blok and Holstege, 1996, Holstege, 1998). The present chapter discusses the bladder and bladder sphincter motoneurons, their regulation by the sacral cord, and how the sacral cord centers are regulated by distinct regions in brain stem, diencephalon, and cortex. Micturition depends on a coordinated action between the smooth (detrusor) muscle of the bladder and the striated external urethral sphincter (EUS), which closes the bladder outlet. During the storage of urine, the detrusor muscle remains relaxed and the EUS is tonically contracted. This activation pattern is reversed during micturition: the EUS relaxes and the bladder contracts, resulting in expulsion of urine. The motoneurons of both bladder and EUS are located in the sacral spinal cord. Their mutual coordination, however, takes place in the pontine micturition center (PMC) in the brain stem. For example, transection of the spinal cord in animals as well as in humans interrupts the descending fibers from the PMC to the sacral cord. The result is dyssynergic micturition, which means that contraction of the bladder is accompanied by simultaneous contraction of the sphincter. Consequently, in

The Rat Nervous System, Third Edition

order to expel urine through the tonically closed urethral sphincter, the bladder has to produce an extremely high intravesical pressure. The result is a thick bladder wall and a small bladder capacity, the so-called overflow bladder. Brain lesions rostral to the pons never result in bladder–sphincter dyssynergia, but in urge incontinence, i.e., hyperactivity of the bladder and inability to delay voiding. Apparently, the PMC coordinates micturition as such, but regions rostral to the pons determine the beginning of micturition.

MOTONEURONS INNERVATING BLADDER AND URETHRAL SPHINCTER Bladder Motoneurons The smooth musculature forming the detrusor muscle of the bladder and the smooth musculature of the descending colon is innervated, via the pelvic nerve, by parasympathetic preganglionic motoneurons in the sacral intermediolateral cell group or sacral parasympathetic nucleus (SPN). In rat the SPN is located in the L6–S1 spinal segments (Nadelhaft and Booth, 1984). In cat (Morgan et al., 1979) and human the SPN is located in the S2–S3 spinal cord, and in cat a viscerotopic organization has been demonstrated wherein the colon is innervated primarily by cells in a dorsal band and the urinary bladder by cells in a lateral band (Nadelhaft et al., 1980). There is also a spatial segregation within the SPN in rat, in which penis motoneurons

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are located rostral (rostral L6–rostral S1) to the bladder motoneurons (caudal L6-caudal S1; Banrezes et al., 2002). Sympathetic preganglionic motoneurons, located in the intermediolateral cell group of the upper lumbar cord (L1–L4), also innervate the bladder. Their fibers run via the pelvic and hypogastric nerves to have direct access to the bladder or to have indirect access via connections with the paravesical ganglia of the parasympathetic system. Sympathetic fibers have inhibitory effects on the detrusor muscle and excitatory effects on the smooth musculature of the urethra and the base of the bladder, perhaps via suppression of transmission between parasympathetic preganglionic and postganglionic neurons within the pelvic ganglia (see De Groat et al., 1999, for review).

1985), and human (Schrøder, 1981) pelvic floor motoneurons are located in one nucleus on the ventrolateral border of the ventral horn in the upper sacral cord. In these species the motoneurons innervating the anal sphincter are located dorsomedially, and those innervating the bladder sphincter are located ventrolaterally. In the pig (Blok et al., 1996) and Mongolian gerbil (Ulibarri et al., 1995), anal sphincter motoneurons are located in a completely different location, dorsal and dorsolateral to the central canal, in the area of the intermediomedial cell group (IMM) (Fig. 1). In nonexperimental sections, the ON can be recognized by its wealth of longitudinally oriented dendrites (Dekker et al., 1973), indicating that the motoneurons have many connections within the nucleus itself. In the rat, in the dorsolateral nucleus, as well as in the spinal nucleus of the bulbocavernosus, so-called gap junctions have been demonstrated, the number of which were influenced by testosterone levels (Matsumoto et al., 1989). Such gap junctions could not be detected in the ON of the cat (Takahashi and Yamamoto, 1979). ON motoneurons belong to a distinct class of motoneurons. They belong to neither somatic nor autonomic (sympathetic and parasympathetic) motoneurons. Similar to somatic motoneurons ON motoneurons innervate striated muscles of the pelvic floor, including the urethral and anal sphincters (cat: Sato et al., 1978; human: Schrøder, 1981), and are under voluntary control. Animals and humans can voluntarily contract the two sphincters, although not separately. Arguments against ON motoneurons to be somatic are: (1) in contrast to most other motoneuronal cell groups, they do not receive direct cortical afferents (Iwatsubo et al., 1990); (2) they play a role in “autonomic” motor activities such as micturition, ejaculation, defecation, and parturition; (3) they receive direct hypothalamic afferents (Holstege, 1987); (4) they have an appearance similar to that of autonomic motoneurons; (5) unlike all other somatic motoneurons, they are not affected in

Pelvic Floor Motoneurons The pudendal nerve innervates the intrinsic external urethral sphincter, which forms part of the pelvic floor musculature. In most vertebrates the pelvic floor motoneurons are located in the so-called nucleus of Onuf (ON), after Onufrowicz, who called himself Onuf, and who described this nucleus already in 1899. Onuf thought that the nucleus was involved in sexual activity and not in continence for feces and urine. For a long time ON was called group Y (Romanes, 1951), and only 25 years ago ON has been identified as containing pelvic floor motoneurons (Sato et al., 1978). In rat (Schrøder, 1980; McKenna and Nadelhaft, 1986), ON consists of two separate nuclei. The dorsolateral nucleus contains motoneurons innervating the ischiocavernosus muscle and the external urethral sphincter and differs between male and female rats (Jordan et al., 1982). The dorsomedial nucleus innervates the bulbospongiosus muscle and the anal sphincter (McKenna and Nadelhaft, 1986). In other species like hamster (Gerrits et al., 1997), cat (Sato et al., 1978; Kuzuhara et al., 1980; Ueyama et al., 1984), dog (Kuzuhara et al., 1980), monkey (Roppolo et al.,

A A A

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FIGURE 1 Schematic overview of the pelvic floor motoneurons in various mammalian species. A, external anal sphincter; U, external urethral sphincter.

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amyotrophic lateral sclerosis, (Mannen et al., 1977); and (6) they degenerate in Shy–Dräger syndrome (Mannen et al., 1982; Chalmers and Swash, 1987), which is characterized by degeneration of autonomic (sympathetic and parasympathetic) but not somatic motoneurons. In conclusion, ON motoneurons have properties of both somatic and autonomic motoneurons and occupy a position in between these two kinds of motoneurons.

SACRAL CORD MICTURITION REFLEXES Behavioral evidence for the existence of sacral micturition reflexes was given by De Groat et al. (1975, 1981), who observed that micturition as well as defecation are elicited in neonatal kittens when the mother licks the perineal region. This is the primary stimulus for micturition, because separation of the kittens from the mother results in urinary retention. The perineal-tobladder reflex is quite prominent during the first 4 postnatal weeks, after which it becomes less effective and usually disappears by the age of 7–8 weeks, the approximate age of weaning. Thoracic cord transections of the spinal cord did not abolish the perineal-tobladder reflex, indicating that it is a sacral cord reflex. After 7–8 weeks the bladder-to-bladder reflexes, which involve supraspinal structures, have replaced the perineal-to-bladder reflex. Transection of the spinal cord in older kittens or adult cats causes reemergence of perineally induced micturition within 1–2 weeks. The same is true in humans in which this spinal cord reflex system is functionally nonexistent, except in patients with spinal cord transection rostral to the sacral cord. Apparently, in adult animals and humans pathways exist within the sacral cord that can produce bladder and sphincter contractions, although these contractions are not necessarily well coordinated (i.e., they are dyssynergic).

BRAIN STEM–SPINAL CORD PATHWAYS COORDINATE BLADDER AND SPHINCTER MOTONEURONS Diffuse Descending Systems The SPN and the ON play a role not only in micturition but also in several other mechanisms. For example, level-setting mechanisms (see Holstege, 1991, for review) send fibers to almost all parts of the spinal cord including SPN and ON motoneurons. These serotonergic and other projections from the caudal raphe nuclei and the adjoining ventral tegmentum of the

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medulla to the SPN and the ON are sometimes mistaken for specific micturition control systems. Also the noradrenergic projections from the dorsolateral pontine tegmentum (locus coeruleus and nucleus subcoeruleus) and the dopamine projections from the A11 cell group in the rostral mesencephalon (Holstege et al., 1996) take part in very diffuse descending systems but seem not to play a specific role in micturition control. Nevertheless, studies of serotonin (5-HT) receptors and reuptake mechanisms might represent targets for the development of new drugs for treatment of patients suffering from bladder (detrusor) overactivity and urinary incontinence. One might expect, however, many side effects of these drugs, because of the diffuse nature of these projections (DeGroat, 2002).

Abdominal Pressure Control Systems The pelvic floor forms the bottom of the abdominal cavity. Increased abdominal pressure is needed for strong expiration, vocalization, vomiting, and parturition, and the pelvic floor plays an important role in these motor activities. It is not surprising, therefore, that the nucleus retroambiguus, which controls abdominal pressure by innervating abdominal wall muscle motoneurons, also innervates ON motoneurons (Holstege and Tan, 1987) (Fig. 2).

Micturition Control in Animals Since the work of Barrington (1925) it is known that a crucial structure of the micturition reflex is located in the dorsolateral pontine tegmentum, because bilateral lesions in this area in the cat result in urinary retention. In the cat two different pontine projection systems have been identified. One group of neurons in the medial part of the dorsolateral pons projects to the parasympathetic bladder motoneurons in the sacral intermediolateral cell column (Loewy et al., 1979, in rat; Holstege et al., 1979, 1986, in cat) (Fig. 3 left), whereas neurons in the lateral part of the dorsolateral pons specifically project to the nucleus of Onuf (Holstege et al., 1979, 1986; Fig. 3 right). The medial cell group was called M-region (M = medial) and the lateral cell group L-region (L = lateral). The M-region is also called the pontine micturition center (PMC) or Barrington’s nucleus. Ultrastructurally, it has been demonstrated that the PMC projection to the bladder parasympathetic motoneurons in the sacral cord is monosynaptic and excitatory (Blok and Holstege, 1997). In accordance with these findings, electrical and chemical stimulation in the PMC in the cat produces a steep rise in the intravesical pressure and an immediate and sharp decrease in the urethral pressure and pelvic floor electromyogram

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XII

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Phrenic motoneurons C5-C6 External intercostal motoneurons T1-T13 Abdominal muscle motoneurons T5-L3 Onuf (pelvic floor) motoneurons S1-S2 FIGURE 2 Schematic overview of the pathways controlling respiration and abdominal pressure. For the sake of clarity the few ipsilaterally descending pathways have not been indicated.

(Holstege et al., 1986, in cat; Mallory et al., 1989, in rat) (Fig. 4). In all likelihood, this decrease is the result of the PMC projection to GABA- and glycinergic interneurons in the sacral intermediomedial cell group (Blok et al., 1997a; Sie et al., 2001). Retrograde tracing studies with the pseudorabies virus have shown that this area contains interneurons projecting to the nucleus of Onuf (Nadelhaft and Vera, 1996), and electrical stimulation in the intermediomedial cell column results in an inhibition of the EUS via GABAergic interneurons in the intermediomedial cell group (Blok et al., 1998). Bilateral lesions in the PMC result in total urinary retention leading to depressed detrusor activity and increased bladder capacity (Griffiths et al., 1990; Holstege et al., 1986). In rats, unilateral chemical lesions of the PMC attenuate the bladder response (Mallory et al., 1989). Stimulation in the L-region results in strong excitation of the pelvic floor musculature and an increase in the urethral pressure (Holstege et al., 1986) (Fig. 5). Bilateral lesions in the L-region give rise to an inability to store urine; bladder capacity is reduced and urine is expelled prematurely by excessive detrusor activity, accompanied by urethral relaxation. Outside the episodes of detrusor activity, the urethral pressure is

not depressed below normal values (Mallory et al., 1989). These observations suggest that during the filling phase the L-region has a continuous excitatory effect on the nucleus of Onuf resulting in inhibition of urethral relaxation coupled with detrusor contraction. Perhaps the L-region should be considered as a “continence” center, especially since the PET scan results suggest the existence of such a center in humans (see below).

AFFERENT SYSTEMS Peripheral Afferent Nerves Most afferent fibers from the bladder enter the sacral cord via the pelvic nerve. The peripheral fibers of the dorsal root ganglia neurons of the pelvic nerve contact the bladder wall mechanoreceptors. The proximal fibers, many of which contain calcitonin gene-related peptide (CGRP) (Chung et al., 1993), enter Lissauer’s tract and terminate mainly in Rexed’s (1954) laminae I, V, VII, and X of the lumbosacral spinal cord at segments L4–S2 (Morgan et al., 1981). The majority of these afferents are thin myelinated and unmyelinated axons, and their

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Spinal Cord-Brain Stem Pathways Involved in the Micturition Reflex To empty the bladder at an appropriate time, bladder filling information from sensory neurons in the sacral cord must reach the PMC. To a limited extent in rat (Ding et al., 1997; Blok and Holstege, 2000), but not in cat (Blok et al., 1995), the lumbosacral cord projects directly to PMC neurons. This raises the question of what other area relays bladder filling information to the PMC. The micturition reflex is not abolished by precollicular decerebration (Tang and Ruch, 1956), which means that lumbosacral projections to forebrain areas, such as the thalamus and the hypothalamus, are not essential for this reflex. However, the very strong projections from the lumbosacral cord to the lateral and dorsal parts of the mesencephalic periaqueductal gray (PAG) probably play a much more important role (Blok et al., 1995; VanderHorst et al., 1996) (Fig. 6). The lateral PAG is the only brain stem structure known to project specifically to the PMC (Blok and Holstege, 1994). Moreover, stimulation of afferent bladder nerves in the rat evokes short latency potentials (13 ± 3 ms) in the most caudal PAG, while much longer latency potentials (42 ± 7 ms) are found in the PMC (Noto et al., 1991). In the cat electrical stimulation of the PAG has been shown to evoke complete micturition (Skultety, 1959), facilitate bladder reflexes, and reduce bladder capacity (Kruse et al., 1990). The PAG has also been implicated in the control of micturition in humans (see below).

FIGURE 3 Bright-field photomicrographs of autoradiographs showing the tritiated leucine injection areas and dark-field photomicrographs showing the spinal distributions of labeled fibers after an injection in the area of the M-region (on the left) and after an injection in the area of the L-region (on the right) in the cat. Note the dense distribution of labeled fibers to the sacral intermediolateral (parasympathetic motoneurons) and intermediomedial cell groups after an injection in the M-region (S2 segment on the left). Note also the pronounced projection to the nucleus of Onuf (arrows in the S1 segment) in the case with an injection in the L-region (right).

conduction velocities are in the A∂ and C fiber range, respectively (Hulsebosch and Coggeshall, 1982). Most A∂ fibers originate from slowly adapting mechanoreceptors in the bladder wall, because excitation of these fibers results in activation of the micturition reflex. In all likelihood, the A∂ fibers are the peripheral afferent fibers for this reflex (DeGroat et al., 1982; Mallory et al., 1989), because the unmyelinated C fibers in the pelvic nerve do not respond to distention and contraction of the urinary bladder (Jänig and Morrison, 1986).

FOREBRAIN INVOLVEMENT IN THE CONTROL OF MICTURITION The forebrain is not essential for the basic micturition reflex, but it is thought to determine the beginning of micturition (Blaivas, 1982). In the cat, stimulation of forebrain structures, such as the anterior cingulate gyrus, the preoptic area of the hypothalamus, the amygdala, the bed nucleus of the stria terminalis, and the septal nuclei, have been shown to elicit bladder contractions (Gjone and Setekleiv, 1963; Gjone, 1966). Although most of these regions send fibers to the PAG and other parts of the brain stem (see Holstege, 1991, for review), only the preoptic area has been shown to project directly to the PMC (Holstege, 1987, in cat, Rizvi et al., 1994, in the rat). One might speculate that the projections from the many limbic structures to the PAG are tools of the emotional motor system for giving a “safe signal” to the PMC to start micturition and that the direct projection of the preoptic area to the PMC controls micturition for sexual and territorial marking. Figure 7 shows a schematic overview of all the pathways that are thought to play a role

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FIGURE 4 Recordings of urethral pressure, pelvic floor EMG, intravesical pressure, and stimulus timing during M-region stimulation in the cat. Note the immediate fall in urethral pressure and pelvic floor EMG after the beginning of the stimulus and the steep rise in the intravesical pressure about 2 s after the beginning of the stimulus. This pattern mimics complete micturition (from Holstege et al., 1986).

FIGURE 5 Recordings of urethral pressure, pelvic floor EMG, intravesical pressure, and stimulus timing during L-region stimulation in the cat. Note the immediate increase in the urethral pressure and the pelvic floor EMG at the beginning of each period of stimulation (from Holstege et al., 1986).

in micturition control. The PET studies of Blok et al. (1997b, 1997c, 1998a, 1998b) suggest that this might also be true in humans (see below).

MICTURITION CONTROL IN HUMANS Blok and Holstege have demonstrated that with

respect to pelvic floor activities such as micturition in men and women (Blok et al., 1997a, 1998b), as well as in voluntary contraction (Blok et al., 1997c), the supraspinal control systems seem to be similarly organized in humans and cats. Of the volunteers that were asked to micturate, while lying with their heads in the PET scan, half of them could satisfactorily perform the task. In this group the same regions as those observed in the cat

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L7

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bladder, colon, vagina, vaginocervix, uterine cervix, pelvic floor, perineal skin

PAG-projecting neurons

penis and clitoris

parasympathetic preganglionic motoneurons

S1 caudal

S2

S3

FIGURE 6 Schematic overview of the location of the pelvic and pudendal primary afferents, the PAG projecting neurons, and the sacral parasympathetic preganglionic motoneurons (from VanderHorst et al., 1996).

were found to be active in humans, i.e., an area in the dorsal pontine tegmentum, probably representing the PMC (Fig. 8, left), an area in the PAG (Fig. 9, left), and one in the hypothalamus or the preoptic region (Fig. 9, right). The other group of volunteers, who were willing to micturate, but, for emotional reasons, could not perform, tightly contracted their pelvic floor. The PET scan results in these volunteers revealed an area in the ventrolateral pontine tegmentum, which might represent the L-region or continence center (Fig. 8, right). With respect to the voluntary contraction of the pelvic floor it was found that none of the micturition control regions were active, but only the motor cortical regions that were directly involved in voluntary pelvic floor contraction (Blok et al., 1997c). It also means that the motor cortical system is completely separate from the basic micturition control system that takes part in the emotional motor system (Holstege, 1998). The PET scan results in men and women (Blok et al., 1997b, 1997c, 1998a, 1998b) point to two cortical areas involved in micturition. The right dorsolateral prefrontal cortex is active when micturition takes place and also when the volunteers are asked to micturate, but cannot perform. Another part that might play a role in micturition is the rostral part of the right cingulate gyrus, which receives strong afferent connections from many parts of the limbic system. It is interesting that those parts of the rostral cingulate gyrus that are active during successful micturition differ from those that are activated during nonsuccessful micturition (Fig. 10). Conversely, activation in the right anterior cingulate gyrus was significantly decreased during voluntary withholding of urine (Blok et al., 1997b). Possibly, activation of the prefrontal cortex and the anterior cingulate gyrus are not specifically involved in micturition, but

in general mechanisms, such as attention and response selection (Pardo et al., 1990) or emotional distress and pain (Drossman et al., 2003). Forebrain lesions including the anterior cingulate gyrus are known to cause urge incontinence (Andrew and Nathan, 1966), which might be due to an attention deficit that interferes with the patient’s ability to recognize a full bladder. A striking observation of the PET studies of Blok et al. (1997b, 1997c, 1998b) was that the control areas were found predominantly on the right side of the brain and brain stem (cortex and pons), which corresponds with studies reporting that urge incontinence is correlated with lesions in the right hemisphere (Kuroiwa et al., 1987).

CONCLUSIONS The organization of the central nervous system control of the basic micturition reflex is probably the same in humans and cats, but differs from that in rat. Several levels of the central nervous system are involved in the control of micturition. Sensory information concerning bladder filling is relayed, via the pelvic nerve and dorsal root ganglion cells, to the sacral cord, where second-order sensory neurons are located. These neurons convey the information to the caudal PAG where it is processed and filtered. When the amount of urine in the bladder makes voiding desirable, PAG neurons activate the premotor interneurons in the Mregion, which, via long descending fibers to the bladder motoneurons in the sacral cord, let micturition take place. Projections from many limbic structures to the PAG, and the projections from the preoptic area to the M-region, may influence this spinal–brain stem–spinal reflex and may determine the beginning of micturition.

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L-region

GABA-ergic / glycinergic neurons

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Onuf's nucleus

)-(

Onuf's nucleus

)+(

S2

S2

Primary afferents

Bladder

Bladder )+(

External urethral sphincter

External urethral sphincter

)+(

)+(

FIGURE 7 On the left a schematic overview of the micturition pathways with the ascending and descending components in the cat. On the right an overview of the possible pathways involved in continence control. The question mark indicates that it is not yet known what controls the L-region, although the structures involved, in all likelihood, are located in the limbic system.

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FIGURE 8 PET scan results during normal micturition (left) and nonsuccessful micturition, leading to “emotional” contraction of the pelvic floor (right).

FIGURE 10 (Top) PET scan results showing activation in the anteFIGURE 9 PET scan results showing on the left activation in the area of the periaqueductal gray (arrow) and on the right in the hypothalamus.

References Andrew, J., and Nathan, P. W. (1964). Lesions of the anterior frontal lobes and disturbances of micturition and defecation. Brain 87, 233–262. Banrezes, B., Andrey, P., Maschino, E., Schirar, A., Peytevin, J., Rampin, O., and Maurin, Y. (2002). Spatial segregation within the sacral parasympathetic nucleus of neurons innervating the bladder or the penis of the rat as revealed by three-dimensional reconstruction. Neuroscience 115, 97–109. Barrington, F. J. F. (1925). The effect of lesions of the hind- and midbrain on micturition in the cat. Q. J. Exp. Physiol. Cogn. Med. 15, 81–102. Blaivas, J. G. (1982). The neurophysiology of micturition: A clinical study of 550 patients. J. Urol. 127, 958–963. Blok, B. F. M., DeWeerd, H., and Holstege, G. (1995). Ultrastructural evidence for the paucity of projections from the lumbosacral cord to the M-region in the cat: A new concept for the organization of the micturition reflex with the periaqueductal gray as central relay. J. Comp. Neurol. 359, 300–309. Blok, B. F. M., DeWeerd, H., and Holstege, G. (1997a). The pontine micturition center projects to sacral cord GABA immunoreactive neurons in the cat. Neurosci. Lett. 233, 109–112. Blok, B. F. M., and Holstege, G. (1994). Direct projections from the periaqueductal gray to the pontine micturition center (Mregion): An anterograde and retrograde tracing study in the cat: Neurosci. Lett. 166, 93–96.

rior cingulate gyrus, comparing successful micturition with voluntary withholding of urine. (Bottom) PET scan results comparing unsuccessful micturition with voluntary withholding of urine. Note that a different part of the cingulate gyrus is active during both activities.

Blok, B. F. M., and Holstege, G. (1996). Neuronal control of micturition and its relation to the emotional motor system. Prog. Brain Res. 107, 113–126. Blok, B. F. M., and Holstege, G. (1997). Ultrastructural evidence for a direct pathway from the pontine micturition center to the parasympathetic preganglionic motoneurons of the bladder of the cat. Neurosci. Lett. 222, 195–198. Blok, B. F. M., and Holstege, G. (2000). The pontine micturition center in rat receives direct lumbosacral input: An ultrastructural study. Neurosci. Lett. 282, 29–32. Blok, B. F. M., Maarseveen, J. T. P. W., and Holstege, G. (1998a). Electrical stimulation of the sacral dorsal gray commissure evokes relaxation of the external urethral sphincter in the cat. Neurosc. Lett. 249, 68–70. Blok, B. F. M., Roukema, G., Geerdes, B., and Holstege, G. (1996). Location of external anal sphincter motoneurons in the sacral cord of the female domestic pig. Neurosci. Lett. 216, 203–206. Blok, B. F. M., Sturms, L. M., and Holstege, G. (1997b). A PET study on cortical and subcortical control of pelvic floor musculature in humans. J. Comp. Neurol. 389, 535–544. Blok, B. F. M., Sturms, L. M., and Holstege, G. (1998b). Brain activation during micturition in women. Brain 121, 2033–2042. Blok, B. F. M., Willemsen, A. T. M., and Holstege, G. (1997c). A PET study on the brain control of micturition in humans. Brain 120, 111–121. Chalmers, D., and Swash, M. (1987). Selective vulnerability of urinary Onuf motoneurons in Shy–Dräger syndrome. J. Neurol. 234, 259–260.

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Holstege, J. C., Van Dijken, H., Buijs, R. M., Goedknegt, H., Gosens, T., and Bongers, C. M. (1996). Distribution of dopamine immunoreactivity in the rat, cat and monkey spinal cord. J. Comp. Neurol. 376, 631–652. Hulsebosch C. E., Coggeshall R. E. (1982). An analysis of the axon populations in the nerves to the pelvic viscera in the rat. J Comp Neurol. 211, 1–10. Iwatsubo, T., Kuzuhara, S., Kanemitsu, A., Shimada, H., and Toyokura, Y. (1990). Corticofugal projections to the motor nuclei of the brainstem and spinal cord in humans. Neurology 40, 309–312. Jänig, W., and Morrison, J. F. (1986). Functional properties of spinal visceral afferents supplying abdominal and pelvic organs, with special emphasis on visceral nociception. Prog. Brain Res. 67, 87–114. Jordan, C. L., Breedlove, S. M., and Arnold, A. P. (1982). Sexual dimorphism and the influence of neonatal androgen in the dorsolateral motor nucleus of the rat lumbar spinal cord. Brain Res. 249, 309–314. Kruse, M. N., Noto, H., Roppolo, J. R., and De Groat, W. C. (1990). Pontine control of the urinary bladder and external urethral sphincter in the rat. Brain Res. 532, 182–190. Kuroiwa, Y., Tohgi, H., Ono, S., and Itoh, M. (1987). Frequency and urgency of micturition in hemiplegic patients: Relationship to hemisphere laterality of lesions. J. Neurol. 234, 100–102. Kuzuhara, S., Kanazawa, I., and Nakanishi, T. (1980). Topographical localization of the Onuf’s nuclear neurons innervating the rectal and vesical striated sphincter muscles: A retrograde fluorescent double labeling in cat and dog. Neurosci. Lett. 16, 125–130. Loewy, A. D., Saper, C. B., and Baker, R. P. (1979). Descending projections from the pontine micturition center. Brain Res. 172, 533–539. Mallory, B., Steers, W. D., and De Groat, W. C. (1989). Electrophysiological study of micturition reflexes in rats. Am. J. Physiol. 257, R410–421. Mannen, T., Iwata, M., Toyokura, Y., and Nagashima, K. (1977). Preservation of a certain motoneurone group of the sacral cord in amyotrophic lateral sclerosis: Its clinical significance. J. Neurol. Neurosurg. Psychiatry 40, 464–469. Mannen, T., Iwata, M., Toyokura, Y., and Nagashima, K. (1982). The Onuf’s nucleus and the external anal sphincter muscles in amyotrophic lateral sclerosis and Shy–Drager syndrome. Acta Neurophathol. 58, 255–260. Matsumoto, A., Arnold, A. P., and Micevych, P. E. (1989). Gap junctions between lateral spinal motoneurons in the rat. Brain Res. 495, 362–365. McKenna, K. E., and Nadelhaft, I. (1986). The organization of the pudendal nerve in the male and female rat. J. Comp. Neurol. 248, 532–549. Merrill, E. G. (1970). The lateral respiratory neurones of the medulla: Their associations with nucleus ambiguus, nucleus retroambigualis, the spinal accessory nucleus and the spinal cord. Brain Res. 24, 11–28. Morgan, C., Nadelhaft, I., and De Groat, W. C. (1979). Location of bladder preganglionic neurons within the sacral parasympathetic nucleus of the cat. Neurosci. Lett. 14, 189–195. Morgan, C., Nadelhaft, I., and De Groat, W. C. (1981). The distribution of visceral primary afferents from the pelvic nerve to Lissauer’s tract and the spinal gray matter and its relationship to the sacral parasympathetic nucleus. J. Comp. Neurol. 201, 415–440. Nadelhaft, I., and Booth, A. M. (1984). The location and morphology of preganglionic neurons and the distribution of visceral afferents from the rat pelvic nerve: A horseradish peroxidase study. J. Comp. Neurol. 226, 238–245. Nadelhaft, I., DeGroat, W. C., and Morgan, C. (1980). Location and morphology of parasympathetic preganglionic neurons in the sacral

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C H A P T E R

14 Anatomical Substrates of Hypothalamic Integration RICHARD B. SIMERLY Division of Neuroscience, Oregon National Primate Research Center Oregon Health & Sciences University, Beaverton, Oregon, USA

The primary biological imperatives imposed on an organism are to first ensure its survival and second to propagate its species. To satisfy these imperatives an animal must coordinate complex physiological processes with diverse environmental conditions and elaborate appropriate adaptive responses to specific sensory cues. Thus, the maintenance of homeostasis represents a key component of biological adaptation and depends upon the successful integration of endocrine, visceral, and somatomotor control mechanisms. An extensive body of literature supports an important role for the hypothalamus in mediating this integration and the hypothalamus is generally viewed as an essential interface between the endocrine, autonomic, and somatomotor systems. In mammals the hypothalamus has been shown to regulate the cardiovascular system, the thermoregulatory responses, and the abdominal viscera, as well as defensive–aggressive behavior and ingestive behaviors that provide nutrients and water. In addition, the hypothalamus plays an essential role in assuring the survival of the species by controlling the expression of sexual and maternal behaviors. Of fundamental importance to the function of the hypothalamus is its intimate relationship with the pituitary gland and the now clearly established pathways for neural control of endocrine secretion patterns (Harris, 1948; Sawyer, 1978; Swanson, 1986). In addition, the hypothalamus has extensive connections with the major ascending and descending fiber systems that allow it to sample and influence activity in all major parts of the brain and spinal cord. Thus,

The Rat Nervous System, Third Edition

the unique relationship between the hypothalamus and the pituitary gland, together with its position in the neuraxis, make the hypothalamus the logical neurological substrate for the integration of diverse and dynamic environmental cues with the biological requirements of the individual. Although several interesting and insightful hypotheses have been advanced to explain hypothalamic function (Stellar, 1954; Szentágothai et al., 1962; Valenstein et al., 1969; Morgane, 1979), only recently has the relationship between form and function begun to be elucidated. This is due, at least in part, to the fact that cell groups in the hypothalamus are not clearly differentiated and the region is traversed by diffuse fiber systems that rank among the most complex fiber systems of the nervous system. Nevertheless, work carried out with modern neuroanatomical methods has yielded new insight into the organization of the hypothalamus and suggests that functionally distinct neural systems can be identified and studied in great detail (Swanson, 1999). This chapter presents an overview of hypothalamic anatomy and attempts to illustrate some of the ways in which functional integration is accomplished. Space does not allow for a complete description of the connections and neurochemistry of the hypothalamus, but interested readers may consult excellent reviews of these subjects by Nauta and Haymaker (1969), Palkovits and Zaborszky (1979), Zaborszky (1982), Hoffman et al. (1986), Silverman and Pickard (1983), Swanson (1986, 1987), Markakis and Swanson (1997).

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MORPHOLOGICAL ORGANIZATION OF THE HYPOTHALAMUS The hypothalamus occupies the ventral half of the diencephalon on both sides of the third ventricle and lies immediately above the pituitary gland. Dorsally, the hypothalamus is bounded along most of its length by the zona incerta, and the medial edge of the cerebral peduncle corresponds to its lateral border. Caudally, the hypothalamus merges with the periaqueductal gray and ventral tegmental area of the midbrain. The preoptic region represents the rostral-most part of the hypothalamus and is bordered dorsally by the anterior commissure and anteroventrally by the nucleus of the diagonal band of Broca. The medial preoptic area does not extend beyond the lamina terminalis, but the lateral preoptic area (see below) appears to extend medially, where it replaces the medial preoptic area at rostral levels (Simerly and Swanson, 1988). The hypothalamic parcellation scheme adopted for the third edition of The Rat Nervous System is that described in detail by Swanson (1987, 1998) and is based on the work of Gurdjian (1927) and Krieg (1932). This nomenclature and organizational scheme is the most commonly used and in general is supported by neurochemical and hodological studies. Reference to other interpretations of hypothalamic anatomy and nomenclature can be found in the work of Diepen (1962) and Bleier et al. (1979, 1985). Crosby and Woodburne (1940) recognized that the hypothalamus is best regarded as three distinct longitudinal zones (periventricular, medial, and lateral) and this perspective is supported by physiological and behavioral analyses (Swanson, 1987). The periventricular zone contains most of the neurons that project to the pituitary and is therefore primarily involved in regulating secretion of hormones from this gland. The medial zone of the hypothalamus consists of a rostrocaudally organized series of distinct cell groups that receive their major inputs from various parts of the limbic region of the telencephalon such as the septum and the amygdala. Neurons located in the lateral zone of the hypothalamus are scattered among the fibers of the medial forebrain bundle and can be viewed as a bed nucleus for this complex fiber system. Based on the organization of cell groups in the medial zone, Le Gros Clark (1938) divided the hypothalamus into four rostrocaudal levels designated the preoptic, supraoptic (replaced here by the more common term anterior), tuberal, and mammillary regions. The superposition of LeGros Clark’s regions onto Crosby and Woodburne’s zones divides the hypothalamus into 12 compartments that contain all of the recognized hypothalamic nuclei (Fig. 1). Although this

general organizational plan remains a useful framework for understanding the functional organization of the hypothalamus, certain exceptions and modifications are suggested by recent anatomical data (Swanson, 2000).

Periventricular Zone The hypothalamus represents the final common pathway for the neural control of the anterior, intermediate, and posterior lobes of the pituitary gland and most of the neurons that contain hormone-releasing hormones reside in the periventricular zone (Swanson, 1986; Markakis and Swanson, 1997). Two notable exceptions to this organizational plan are the supraoptic and accessory supraoptic nuclei, which consist of magnocellular cells that during development migrate away from the periventricular zone, and gonadotropinreleasing hormone neurons, which are derived from the olfactory epithelium and migrate to the septal and preoptic regions of the forebrain. By analogy with the motor neurons in the spinal cord, these neuroendocrine neurons have been referred to as “the motoneurons of the neuroendocrine system,” since these cells are the effector units of the neuroendocrine hypothalamus in a way similar (to the way that the) motor neurons are the effector units of neuromuscular pathways (Swanson, 1986; Swanson et al., 1987). In addition, the periventricular zone contains other cell groups that are intimately connected to the neuroendocrine cells and thereby play important roles in controlling neuroendocrine secretion. Cytoarchitectonically the periventricular zone is characterized by small, vertically oriented fusiform neurons and is traversed by a complex array of ascending and descending fibers that are part of the periventricular system connecting the periventricular zone of the hypothalamus with the midline thalamus and the midbrain periaqueductal gray (Sutin, 1966; Nauta and Haymaker, 1969). The periventricular zone is composed of the four periventricular nuclei (preoptic, anterior, intermediate (or tuberal), and posterior) and six other distinct cell groups that represent differentiated parts of this zone. Swanson (2000) has recently proposed that a hypothalamic periventricular region should also be recognized that consists of neurons in the periventricular zone that do not project to the median eminence (are not neuroendocrine) and the diffusely organized parts of the medial zone (those neurons not included in the large nuclei). Although such a region is hard to conceptualize in purely anatomical terms, the proposal that neurons in this region constitute a visceromotor pattern generator network represents a potentially important functional notion that deserves experimental attention.

IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

Regions Preoptic

Anterior

Tuberal

LPO

LH

LH

Mammillary MCLH

LH

Lateral

Zones

MCPO PS MPA Medial

PD MPO

PePO

SCh

DMH AH

Pa

SuM

TU PHA PMV

RCh

TA

PeA

PeI

VMH Arc

LM

T M V

PMD MM

TMD

PeP

Areas and Nuclei Periventricular zone Preoptic periventricular n. (PePO) Vascular organ of the lamina terminalis (vo) Median preoptic n. (MnPO) Anteroventral periventricular n. (AVPV) Suprachiasmatic preoptic n. (PSCh)

Medial zone Medial preoptic a. (MPA) Medial preoptic n. (MPO) Anterodorsal preoptic n. (AD) Anteroventral preoptic n. (AV) Parastrial n. (PS) Posterodorsal preoptic n. (PD)

Lateral zone Lateral preoptic a. (LPO) Magnocellular preoptic n. (MCPO)

Periventricular zone Anterior periventricular n. (PeA) Suprachiasmatic n. (SCh) Paraventricular n. (Pa)

Medial zone Anterior hypothalamic a. (AHA) Anterior hypothalamic n. (AH) Retrochiasmatic area (RCh) Nucleus circularis (NC)*

Lateral zone Lateral Hypothalamic a. (LH) Supraoptic n. (SO)*

Periventricular zone Tuberal periventricular n. (PeI) Arcuate n. (Arc)

Medial zone Tuberal a. (TA) Ventromedial n. (VMH) Dorsomedial n. (DMH)

Lateral zone Lateral hypothalamic a. (LH) Tuberal n. (TU)

Periventricular zone Posterior periventricular n. (PeP) Dorsal tuberomammillary n. (TMD)**

Medial zone Dorsal premammillary n. (PMD) Ventral premammillary n. (PMV) Mammillary complex medial mammillary n. (MM) lateral mammillary n. (LM) supramammillary n. (SuM) Posterior hypothalamic area

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IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

AD AV MnPO PSCh Periventricular vo AVPV

SO AHA NC

Lateral zone Lateral hypothalamic area (LH) Magnocellular n. of LHA Ventral tuberomammillary n. (TMV)

FIGURE 1 Organization of the hypothalamus. A highly schematic representation of the morphological organization of the hypothalamus to show a simplified view of its zones, regions, areas, and nuclei. Abbreviations used: a, area; n, nucleus. Notes: *The supraoptic n. and n. circularis are best considered a displaced part of the periventricular zone. **Despite the location of the TMD in the periventricular zone, the connections and presumed function of both parts of the tuberomammillary n. are more closely related to the lateral zone.

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Preoptic Region In addition to the periventricular preoptic nucleus (PePO) the preoptic part of the periventricular zone contains three identifiable cell groups. The median preoptic nucleus (MnPO) is a dense cluster of small cells in the lamina terminalis that extends along the anterior border of the third ventricle from the tip of the preoptic recess of the third ventricle dorsocaudally to a small triangular area between the descending columns of the fornix (Atlas plates 18–20). The MnPO plays a critical role in neural circuits controlling cardiovascular responses and fluid homeostasis (Saper and Levisohn, 1983; Saper, 2003). Consistent with this role it receives strong inputs from the subfornical organ and the parabrachial nucleus (Lind et al., 1982; Saper and Levisohn, 1983; Fulwiler and Saper, 1984; Lind, 1987; Bester et al., 1997) and sends projections to the paraventricular nucleus and magnocellular cell groups, as well as to the dorsomedial hypothalamic nucleus (Gu and Simerly, 1997; Thompson and Swanson, 1998). The anteroventral periventricular nucleus (AVPV) occupies a ventral position in the periventricular zone of the preoptic region and is located immediately caudal to the vascular organ of the lamina terminalis. This oval cluster of densely packed, darkly stained neurons is readily distinguishable from the periventricular preoptic nucleus, which lies immediately caudal and dorsal to the AVPV, and is limited laterally by a cell-sparse region that separates the AVPV from the anteroventral preoptic nucleus (Atlas plates 19 and 20). Ventrally the AVPV is separated from the optic chiasm by the suprachiasmatic preoptic nucleus (PSCh), which consists of obliquely oriented fusiform cells also known as the ventromedial preoptic nucleus (Paxinos and Watson, 1998). The morphology of the AVPV and PSCh were first defined by Bleier et al. (1979, 1985) who referred to them as dorsal and ventral subdivisions of the “medial preoptic nucleus,” a term that is generally used to describe the major preoptic nucleus of the medial zone (Gurdjian, 1927; Christ, 1969; Simerly et al., 1984b). This discrepancy in nomenclature derives from different interpretations of Gurdjian’s description of preoptic nuclei (Gurdjian, 1927; see Simerly, 1995a, for review). However, on the basis of cytoarchitectonic (Simerly et al., 1984b), neurochemical (Simerly et al., 1985; Simerly and Swanson, 1987; Simerly, 1989; Herbison, 1992), connectional (Simerly and Swanson, 1988; Gu and Simerly, 1997; Simerly, 1998), and functional grounds (Wiegand and Terasawa, 1982; Herbison, 1998; Le et al., 1999), the AVPV and PSCh are clearly part of the periventricular zone of the hypothalamus. The AVPV has also been included in a region called the “nucleus preopticus,

pars suprachiasmatica” (König and Klippel, 1963), and in a region termed the “AV3V” region, which is defined functionally on the basis of its involvement in the regulation of fluid and electrolyte metabolism (Saper and Levisohn, 1983; Lind and Ganten, 1990; Saper, 2003). The afferents of the AVPV have not been mapped in detail, but results of retrograde (Wiegand and Price, 1980; Wiegand, 1984; Simerly, 1998) and anterograde tracing studies (Simerly and Swanson, 1988; Gu and Simerly, 1997) support its proposed functional role as a nodal point in neural circuitry controlling gonadotropin secretion (Wiegand and Terasawa, 1982). Of primary importance are the strong inputs from the posterior and medial nuclei of the amygdala and from the principal nucleus of the bed nuclei of the stria terminalis (Simerly et al., 1989; Canteras et al., 1992a; Hutton et al., 1998), which presumably convey olfactory information (Halpern, 1987; Segovia and Guillamön, 1993; Guillamon and Segovia, 1996). The AVPV also receives a strong input from the ventral part of the lateral septal nucleus, which relays multimodal information from the hippocampal formation to periventricular and medial parts of the hypothalamus (Risold and Swanson, 1996, 1997b; Risold, 2003). The AVPV is heavily innervated by all parts of the periventricular zone of the hypothalamus, the medial preoptic nucleus, and the dorsomedial hypothalamic nucleus and receives a particularly dense input from the ventral premammillary nucleus as well (Wiegand, 1984; Ter Horst and Luiten, 1986; Simerly and Swanson, 1988; Canteras et al., 1992b; Thompson et al., 1996). Circadian regulation of AVPV activity may be conveyed by direct inputs from the suprachiasmatic nucleus, which appear stronger in females (Watts et al., 1987; Van der Beek et al., 1997). The AVPV also receives afferents from many of the same brain stem nuclei that provide inputs to the medial part of the medial preoptic nucleus (Simerly and Swanson, 1986), such as the nucleus of the solitary tract, ventrolateral tegmentum, and periaqueductal gray. Consistent with its proposed functional role, neurons in the AVPV appear to provide direct inputs to gonadotropin-releasing hormone containing neurons in the region adjacent to the vascular organ of the lamina terminalis, as well as to tuberoinfundibular dopaminergic neurons in the arcuate nucleus (Gu and Simerly, 1997). A broader role in neuroendocrine regulation is suggested by the observation that the heaviest projections from the AVPV are to nuclei within the periventricular zone of the hypothalamus, including the paraventricular nucleus, and these regions also regulate autonomic function (Westerhaus and Loewy, 1999). In addition, strong connections between the AVPV and the subfornical organ, the median preoptic nucleus, and the

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parastrial nucleus (see below) suggest a possible role for these neurons in the regulation of fluid homeostasis (Lind and Ganten, 1990; Simerly, 1995a; Saper, 2003). Although the AVPV has not been implicated in the neural control of behavior, and does not generally provide major inputs to medial zone nuclei, it does send strong projections to the medial part of the medial preoptic nucleus and dorsomedial hypothalamic nucleus. The only major telencephalic projection of the AVPV is to the ventral part of the lateral septal nucleus (Gu and Simerly, 1997) and it sends more diffuse projections to the bed nuclei of the stria terminalis (BST). Anterior Region The anterior periventricular nucleus (PeA) is a caudal extension of the PePO and has a similar morphological appearance, although the width of the PeA is somewhat greater. Two of the most recognizable and clearly differentiated of all hypothalamic nuclei reside in the anterior part of the periventricular zone. The first of these, the suprachiasmatic nucleus (SCh), consists of small, closely packed, darkly staining neurons. It is bordered anteroventrally by the optic chiasm and posteroventrally by the supraoptic commissure. The SCh is generally divided into dorsomedial and ventrolateral parts that can be seen readily in Nissl or histochemically stained frontal sections (Fig. 2) (van den Pol, 1980). The SCh receives a direct input from the retina and many studies have established its critical role in the control of rodent circadian and diurnal rhythms (Moore and Lenn, 1972; Rusak and Zucker, 1979; Moore, 1983). Because biological rhythmicity appears to be relatively free of feedback modulation (Zucker, 1983) it is not surprising that the SCh receives few other inputs, although the ventral part of the nucleus receives a strong serotonergic input from the midbrain raphe nuclei (Conrad et al., 1974; Moore et al., 1978; Moga and Moore, 1997) and a projection from the ventral

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lateral geniculate nucleus that contains neuropepitede (NPY) (Swanson et al., 1974; Moore et al., 1984; Watts and Swanson, 1987). The projections of the SCh have been analyzed in detail with the surprising finding that the strongest terminal field lies in a region designated the “subparaventricular zone,” which is bordered by the periventricular nucleus, the paraventricular hypothalamic nucleus, and the anterior hypothalamic nucleus (Swanson and Cowan, 1975a; Watts et al., 1987). Moreover, recent observations with retrograde tracers indicate that the subdivisions of the SCh have distinct projections (Leak and Moore, 2001). Another well-differentiated nucleus in the anterior part of the periventricular zone is the paraventricular nucleus, which perhaps best typifies the functional importance of the periventricular zone in that it contains neurons that express hypothalamic-releasing hormones (such as corticotropin-releasing hormone) and project to the median eminence. It also contains neurons that provide direct projections to regions containing preganglionic autonomic neurons, as well as to neurons that send projections to the posterior pituitary (Sawchenko and Swanson, 1983a, 1983b; Swanson and Sawchenko, 1983; Armstrong, 2003). Accordingly, the paraventricular nucleus is thought to play a central role in mediating hypothalamic responses to stress, feeding, and drinking behavior and participates in a variety of autonomic responses (Swanson and Mogenson, 1981; Loewy, 1991; Sawchenko, 1991, 1998; Sawchenko et al., 1996; Elmquist et al., 1999). The nucleus is composed of parvicellular and magnocellular divisions, and each division is composed of distinct subdivisions that have characteristic connections and distributions of neurotransmitter–specific populations of neurons (Armstrong et al., 1980; Swanson and Kuypers, 1980b). As a whole, the parvicellular parts of the paraventricular nucleus share strong bidirectional connections with other nuclei in the periventricular zone such as the AVPV and the

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FIGURE 2 The suprachiasmatic nucleus (SCh). (A,B) Fluorescence photomicrographs of the same field, taken with two different filter systems, to show the localization of vasopressin-immunoreactive neurons (A) to the dorsal part of the SCh. (C) Fluorescence photomicrograph of an adjacent section to show a dense plexus of serotonin (5-HT)-immunoreactive fibers and terminals in the ventral part of the SCh (original magnification, ×60).

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arcuate nucleus (Levin et al., 1987; Gu and Simerly, 1997; Li et al., 2000) as well as with the medial part of the medial preoptic nucleus, dorsomedial nucleus, and ventral premammillary nucleus (Sawchenko and Swanson, 1983b; Simerly and Swanson, 1986, 1988; Canteras et al., 1992b; Thompson et al., 1996). The paraventricular nucleus is considered in detail in Chapter 15 by Armstrong. The magnocellular neurons of the supraoptic nucleus have much in common with magnocellular neurons of the paraventricular nucleus and, as such, may be considered to be a part of the periventricular zone that was displaced by the optic chiasm (Cunningham and Sawchenko, 1988; Cunningham et al., 1990). Tuberal Region The majority of the tuberal part of the periventricular zone is occupied by the intermediate periventricular nucleus, which is continuous with the PeA and extends caudal to the arcuate hypothalamic nucleus (Arc) (van den Pol and Cassidy, 1982) along the walls of the mammillary recess. The Arc begins just rostral to the infundibular recess and lies along the walls of this recess throughout its length. Two distinct cytoarchitectonic subdivisions of the Arc are generally recognized: a small-celled dorsomedial part and a larger ventrolateral part that contains medium-sized neurons. These two subdivisions of the Arc are apparent in histochemically stained material (Meister and Hauokfelt, 1988; Simerly and Young, 1991) and many Arc neurons contain hypophysiotropic hormones that are released from the terminals located in the neurohemal zone of the median eminence into the hypophysial portal system which carries them to the anterior pituitary (Harris, 1948; Sawyer, 1975, 1978). The strongest inputs to the Arc are from other parts of the periventricular zone, including the paraventricular hypothalamic nucleus and the anteroventral periventricular nucleus, as well as from medial parts of the medial zone, such as the medial subdivision of the medial preoptic nucleus, the dorsomedial nucleus (Thompson and Swanson, 1998) and the ventral premammillary nucleus (Zaborzsky, 1982; Sawchenko and Swanson, 1983b; Simerly and Swanson, 1988; Canteras et al., 1992b). Similarly, the projections of the Arc are largely confined to the periventricular zone, but notably avoid the suprachiasmatic nucleus, with the densest terminal fields found in many of the regions that supply strong afferents to the Arc (Fig. 3), Extrahypothalamic connections are sparse, but the Arc receives strong inputs from the principal nucleus of the BST and the posterodorsal part of the medial nucleus of the amygdala (Simerly et al., 1989; Canteras et al., 1992a; Gu et al., 2003), as well as from the ventral part of the

lateral septal nucleus (Risold and Swanson, 1997b). It also receives substantial inputs from several brain stem sites thought to convey ascending visceral sensory information to the Arc (Jones and Moore, 1977; Azmitia and Segal, 1978; Ricardo and Koh, 1978; Swanson, 1987). Mammillary Region The mammillary part of the periventricular zone is occupied solely by the caudal part of the posterior periventricular nucleus (PeP) which surrounds the posterior end of the third ventricle. This diffuse cell mass is continuous dorsally with the dorsomedial hypothalamic nucleus and ventrally with the Arc. Because its cells resemble those of the Arc it is often included as part of this nucleus, but on neurochemical and connectional grounds it appears to more closely resemble the other periventricular nuclei (Everitt et al., 1986; Swanson, 1987) and contains few neurons that project to the median eminence (Wiegand and Price, 1980).

Medial Zone The medial zone of the hypothalamus contains a series of large nuclei that collectively play key roles in the initiation of motivated behaviors such as copulatory, aggressive, and appetitive behaviors (see Swanson, 1987, 2000). Accordingly, the connections of these nuclei are exceedingly complex with each nucleus possessing strong connections with widely distributed parts of the telencephalon, diencephalon, and brain stem that are thought to mediate the somatomotor integration necessary for the elaboration of appropriate adaptive responses to specific exteroceptive cues. In addition, many of the nuclei of the medial zone are in a position to be influenced by every major sensory modality. Much of this sensory information is relayed by nuclei in the limbic region of the telencephalon (Swanson, 1983a) and these limbic afferents may represent the key to understanding the functional anatomy of the medial zone of the hypothalamus (Risold and Swanson, 1996; Swanson and Petrovich, 1998; Swanson, 2000; Petrovich et al., 2001; Fig. 4). Superimposed on the limbic inputs are afferents from a well-defined set of brain stem nuclei, some of which relay visceral sensory information (see Saper, 2003). A notable feature of nuclei in the medial zone is that they share strong bidirectional connections with most limbic and brain stem regions that supply medial zone afferents. In addition, nuclei in the medial zone of the hypothalamus share strong intrahypothalamic connections with each other, and all project to cell groups in the periventricular zone, thereby providing a route for limbic modulation of neuroendocrine function. Medial nuclei also have connections with the lateral zone of the hypothalamus, which may be involved in

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FIGURE 3 Projections of the arcuate nucleus (Arc). (A) Bright-field photomicrograph to illustrate the appearance and location of a Phaseolus vulgaris leucoagglutinin (PHA-L) injection site centered in the Arc. (B–D) Dark-field photomicrographs that illustrate projection axons and terminals, emanating from the injection site shown in panel A, observed in the anteroventral periventricular nucleus (AVPV), the medial part of the medial preoptic nucleus (MPOm; c, central part of the MPO), and the medial parvocellular part of the paraventricular nucleus (PaMP; original magnification, ×50). This largely periventricular projection pattern was typical of nuclei located in the periventricular zone.

mediating generalized aspects of behavioral state and arousal (Wayner et al., 1981), and contain subpopulations of neurons that appear to participate in a variety of complex behaviors (Pfaff et al., 1994; Canteras et al., 1997; Siegel, 1999; Watts, 2000; Willie et al., 2001; Blaustein and Erskine, 2002; Hull et al., 2002; Schneider and Watts, 2002; Thompson and Swanson, 2002).

Like the periventricular zone, the medial zone of the hypothalamus can be divided into preoptic, anterior, tuberal, and mammillary levels. Thus, the medial zone is divided into discrete areas designated (from rostral to caudal) the medial preoptic, the anterior, the tuberal, and the posterior hypothalamic areas and the mammillary region. Each of these areas consists of a

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FIGURE 4 Limbic inputs to the hypthalamus Photomicrographs to illustrate the appearance of an injection of PHA-L centered in the nucleus of the BST (BSTp) of a male rat and the inputs to the AVPV, ventral premammillary nucleus (PMV), and medial preoptic nucleus (MPO). (Data from Hutton et al., 1998; Gu et al., 2003) (See Ju and Swanson, 1989, for nomenclature, but see also Alheid et al., 2003)

somewhat undifferentiated hypothalamic gray in which several cellular condensations, or nuclei, are embedded. As described in detail by Swanson (1987), axonal transport studies have revealed fundamental organizational principals regarding the functional role of medial zone nuclei. Briefly, the mammillary region appears to be primarily involved in the modulation of cortical information processing, whereas the other nuclei of the medial zone are more directly involved in the production of behavioral responses to visceral, gustatory, and olfactory stimuli. These generalizations are supported by a large body of evidence that defines critical roles for the major nuclei of the medial preoptic, anterior, and tuberal regions of the hypothalamus in the expression of homeostatic and reproductive behaviors,

and the anatomical relationships of the mammillary region suggest that it may indirectly participate in the modulation of auditory and visual influences on these essential behaviors. Preoptic Region The preoptic region is the most complex and differentiated part of the medial zone of the hypothalamus and, not surprisingly, causes the most confusion in the literature regarding its morphological limits, cytoarchitecture, and nomenclature. Because of this, the normal morphology of the medial preoptic area is addressed here in some detail. At least five distinct cell groups are embedded in the undifferentiated gray that makes up the medial preoptic area (MPA). The large oval-shaped

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medial preoptic nucleus (MPO) extends nearly the entire length of the preoptic region and is composed of three distinct subdivisions: a cell-sparse lateral part (MPOl), a cell-dense medial part (MPOm), and a compact, very cell-dense central part (MPOc) that is embedded in the medial subdivision (Simerly et al., 1984b). The MPO as described here corresponds to Gurdjian’s “medial preoptic nucleus” as depicted in Fig. 12, p. 23, of Gurdjian (1927), and to his “nucleus b” of the medial preoptic area (p. 79). Bleier et al. (1979, 1985) designate this nucleus as the “anterior hypothalamic nucleus,” and the term “medial preoptic nucleus” has also been

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used to refer to the medial preoptic area as a whole (Le Gros Clark, 1938; König and Klippel, 1963; Nauta and Haymaker, 1969; Palkovits and Zaborszky, 1979). The connections of the MPO are among the most complex of any cell group of the hypothalamus (Simerly, 1995b, 1998). In addition to extensive intrahypothalamic connections, the MPO receives strong inputs from the posterior and medial nuclei of the amygdala, the principal nucleus of the BST (Fig. 5), caudal and ventral parts of the lateral septal nucleus (Canteras et al., 1992a, 1995; Risold and Swanson, 1997b; Hutton et al., 1998; Risold, 2003), and distinct brain stem regions including the

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FIGURE 5 Projections of medial zone nuclei Dark-field photomicrographs that illustrate marked differences in the patterns of projections from the medial preoptic nucleus (MPO) and central part of the anterior nucleus (AHC; B) to the tuberal region of hypothalamus. Note the dense projections from the MPO to the arcuate nucleus and ventrolateral part of the VMH (A; original magnification, ×40). In contrast, projections from the AHC avoid nuclei in the periventricular zone such as the paraventricular nucleus (PaV; B; original magnification, ×50) and do not innervate the ventrolateral part of the VMH (C; original magnification, ×50).

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ventral tegmental area, nucleus of the solitary tract, and parabrachial nucleus (Berk and Finkelstein, 1981; Simerly and Swanson, 1986; Murphy et al., 1999). Each of the inputs to the MPO are distributed topographically within its three subdivisions, with projections from hypothalamic nuclei of the periventricular zone terminating primarily in the MPOm, and projections from the ventral subiculum, caudal part of the lateral septum, and brain stem serotonergic neurons ending in the MPOl (Simerly et al., 1984a; Simerly and Swanson, 1986; Canteras and Swanson, 1992b; Kishi et al., 2000). The projections of the MPO are equally extensive (Swanson, 1976; Conrad and Pfaff, 1977; Simerly and Swanson, 1988) and include regions thought to mediate the neuroendocrine, autonomic, and somatomotor responses that are important components of the many functions associated with this nucleus such as reproductive and maternal behavior (Numan, 1992; Meisel and Sachs, 1994; Pfaff et al., 1994). Like its inputs, the projections of each subdivision of the MPO are unique. The MPOc is the most sexually dimorphic part of the MPO and sends its strongest projections to other sexually dimorphic parts of the forebrain such as the ventral lateral septal nucleus, the ventrolateral part of the ventromedial nucleus of the hypothalamus, the ventral premammillary nucleus, the principal nucleus of the BST, and the posterodorsal part of the medial nucleus of the amygdala. The MPOm sends its strongest projections to the parts of the hypothalamus involved in the regulation of hormone secretion from the anterior pituitary such as the AVPV, the arcuate nucleus, and the paraventricular nucleus. The cellsparse lateral part of the MPO has the most diffuse projections with a projection to the lateral septum and perifornical region of the hypothalamus, as well as projections to several brain stem nuclei. In addition to the MPO, four smaller nuclei can be distinguished in the medial preoptic area, although little is known regarding the functional role of these nuclei. The first of these smaller cell groups is the anteroventral preoptic nucleus, which lies at the base of the medial preoptic area between the AVPV and the nucleus of the diagonal band of Broca and is separated from each nucleus by clearly demarcated cell-sparse zones. Cells located in this intermediate zone have been termed the ventrolateral preoptic nucleus (VLPO) by Saper and colleagues, and contain GABAergic and galanin containing neurons that provide direct projections to the tuberomammillary nucleus (Sherin et al., 1996, 1998). In addition, the VLPO appear to play an important role in production of sleep, whereas the anteroventral preoptic nucleus seems to participate in neural pathways related to temperature control (Lu et al., 2000). The anterodorsal preoptic nucleus (ADPO)

lies in the dorsal part of the medial preoptic area and appears roughly round in frontal sections through rostral levels of the preoptic region, but caudally it merges imperceptibly with undifferentiated parts of the medial preoptic area. In the nomenclature of Bleier et al. (1979, 1985) the ADPO together with the ventral part of the lateral septal nucleus make up the “septohypothalamic nucleus.” However, the ADPO and the lateral septum are quite different in terms of their cytoarchitecture (Swanson and Cowan, 1979; Simerly et al., 1984b), neurochemistry (Simerly et al., 1988), and connections (Swanson and Cowan, 1979; Simerly and Swanson, 1988). Immediately lateral to the ADPO is a triangular region of low cell density that contains the precommissural component of the stria terminalis. This region was studied in detail by Raisman and Field (1973), who referred to it as the “strial part of the preoptic area.” Raisman and Field also identified a distinct “round nucleus” just lateral to the strial area (StA). This round cluster of cells is replaced caudally by a lens-shaped group of fusiform cells and these two cell groups here are referred to collectively as the parastrial nucleus (PS) (Simerly et al., 1984b; Ju and Swanson, 1989). The PS receives a strong input from the AVPV (Gu and Simerly, 1997) and is notable for its direct projections to magnocellular neurons in the paraventricular nucleus (Simerly and Swanson, 1988). The posterodorsal preoptic nucleus (PDPO) is a small cluster of large, darkly staining neurons that lie at caudal levels of the preoptic region near the posterior tip of the PS, just ventrolateral to the anterior magnocellular part of the paraventricular nucleus (Swanson and Kuypers, 1980b) along the medial border of the BST. Although the PS and PDPO are often considered to be part of the BST, neither nucleus receives inputs from the amygdala and each appears to be more closely related to hypothalamic circuits. Due to its direct projections to magnocellular neurons the parastrial nucleus is likely involved in regulation of fluid homeostasis, whereas the posterodorsal nucleus has been implicated in the neural control of male sexual behavior (Coolen et al., 1998). Because the caudal border of the medial preoptic area is difficult to identify in Nissl-stained sections, it is often included erroneously within the anterior hypothalamic area. Axonal transport studies clearly indicate that the projections of the MPO and medial preoptic area are quite different from those of the cell groups in the anterior hypothalamic region (Fig. 5) (Saper et al., 1978; Simerly and Swanson, 1988; Risold et al., 1994) and therefore support the interpretation of Saper et al. (1978) regarding the caudal limits of the preoptic area. The caudal part of the medial preoptic area, including much of the caudal MPO and BST, has also been

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referred to as the “preoptic–anterior hypothalamic junctional area” (de Olmos and Ingram, 1972). In addition, the term “striohypothalamic nucleus” (Paxinos and Watson, 1998) has been used to define a region between the BST and the dorsolateral aspect of the MPO. However, the results of anterograde tract-tracing studies indicate that the area designated “striohypothalamic nucleus” is part of the terminal field formed by projections of neurons in the medial amygdala to the principal nucleus of the BST and to the MPO (Simerly et al., 1989) and that this region (see Ju and Swanson, 1989) contains numerous fibers en route to the MPO. Anterior Region Compared with the medial preoptic area, the anatomical organization of the anterior hypothalamic area (AHA) is rather simple. The bulk of the AHA is occupied by the oval-shaped anterior hypothalamic nucleus (AH), which is first recognizable near the rostral end of the suprachiasmatic nucleus and lies ventrolateral to the caudal pole of the medial preoptic nucleus. As it increases in size it replaces the medial preoptic area, and caudally the AH is displaced dorsally by the ventromedial hypothalamic nucleus before merging with the rostral end of the dorsomedial hypothalamic nucleus. On cytoarchitectural grounds the AH can be divided into anterior, central, and posterior components (Saper et al., 1978). In addition, a small discrete cluster of neurons (AHd; Atlas plates 26 and 27) in the dorsal aspect of the caudal part of the AH can be recognized and has been designated elsewhere as the stigmoid nucleus (Paxinos and Watson, 1998) or dorsal tuberal nucleus (Bleier et al., 1979; Bleier and Byne, 1985). In general, the connections of the AH appear to be a subset of the connections of the medial preoptic nucleus: complex connections with other parts of the medial zone, sparse inputs from the lateral zone, and only limited inputs from telencephalic regions such as the ventral subiculum, BST, and ventral part of the lateral septal nucleus (Swanson, 1987). The overall pattern of projections from each subdivision of the AH is similar, but there are distinct differences in the relative densities of inputs to specific regions. The major efferents of the AH are to those parts of the medial zone that provide AH afferents and to widespread areas of the lateral zone with a particularly dense terminal field in the perifornical region, as well as to the paraventricular nucleus of the thalamus (Conrad and Pfaff, 1976; Saper et al., 1978; Risold et al., 1994). Significant projections are also directed toward the anterior periventricular and paraventricular nuclei of the hypothalamus and a strong descending projection ends in the periaqueductal gray. The rest of the anterior hypothalamic area is occupied by an undifferentiated

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hypothalamic gray. Ventral to the third ventricle, and between the optic chiasm and the arcuate hypothalamic nucleus, lies a region designated the retrochiasmatic area (RCh) which contains fibers of the supraoptic commissure and neurons that project to the brain stem and spinal cord (Swanson and Kuypers, 1980a). Tuberal Region The tuberal area (TA) of the hypothalamus contains two large and well-differentiated nuclei: the ventromedial (VMH) and dorsomedial (DM) hypothalamic nuclei. The VMH is the largest cell group in the tuberal region and is composed of two distinct cell condensations designated the dorsomedial (VMHDM) and ventromedial (VMHVL) subdivisions (Gurdjian, 1927), which are separated from each other by a cell-sparse zone (VMHc). A fourth subdivision is apparent at anterior levels and is designated the anterior part (VMHA) (Canteras et al., 1993). The VMH is surrounded by a thick fiber capsule or “shell” (Millhouse, 1973) that separates it from the surrounding hypothalamic gray. The major telencephalic afferents to the VMH are from the amygdala and ventral subiculum, with inputs from the posterior nucleus of the amygdala (Canteras et al., 1992a) and ventral subiculum (Canteras and Swanson, 1992b; Kishi et al., 2000) innervating the ventrolateral part of the shell of the VMH. In contrast, the medial and basolateral nuclei of the amygdala send fibers to the cellular core of the nucleus (Krettek and Price, 1977, 1978). The VMH also receives inputs from all parts of the medial zone of the hypothalamus (except the medial and lateral mammillary nuclei) with the input from the DM being especially strong to the VMHDM (Ter Horst and Luiten, 1986). In addition, the VMH receives inputs from the lateral zone, the posterior hypothalamic area, and the suprachiasmatic nucleus (Saper et al., 1979; Watts et al., 1987; Fahrbach et al., 1989). Among the brain stem inputs to the VMH, afferents from the parabrachial nucleus have received the most attention due to the possible importance of this pathway in feeding behavior (Fulwiler and Saper, 1984, 1985; Bester et al., 1997). The VMH sends massive projections to other parts of the medial zone of the hypothalamus and tends to avoid periventricular regions and the lateral zone. The VMH also sends widespread projections to the amygdala and the septum, with the strongest projection to the bed nuclei of the stria terminalis, as well as providing descending projections to brain stem regions such as the periaqueductal gray (Saper et al., 1976b; Canteras and Simerly, 1994) that are known to project either to the spinal cord or to the basal ganglia. These descending projections are consistent with the proposed role of the VMH in mediating somatomotor aspects of

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complex motivated behaviors (Pfaff et al., 1994). The subdivisions of the VMH appear to have quantitatively different projections, but the overall pattern of projections is qualitatively similar. In general, the VMHVL shares connections with regions of the forebrain and brain stem that also contain high densities of neurons that express sex steroid receptors (Fig. 6) and are involved in mediating reproductive behavior, such as the medial amygdala and the medial preoptic area, while the VMHDM shares strong connections with regions involved with appetitive behaviors, such as the paraventricular nucleus and dorsomedial hypothalamic nuclei (Saper et al., 1976b; Canteras and Simerly, 1994; see Keay and Bandler, 2003, for review of PAG anatomy). In addition, the central, lateral, and medial nuclei of the amygdala receive substantial inputs from the VMH; other forebrain regions that receive inputs from the VMH include the BST, the nucleus accumbens, and the infralimbic cortex. Each part of the VMH sends a massive projection to the periaqueductal gray (PAG) that is topographically organized with the VMHDL preferentially innervating rostral and dorsal parts, and the VMHVL providing more widespread inputs to caudal parts of the PAG (Canteras and Simerly, 1994).

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Immediately lateral to the VMHVL lies the tuberal nucleus (TU), which consists of an oval cluster of smallto medium-sized neurons embedded in the tuberal part of the lateral hypothalamic area. The TU was originally described by Malone (1910) and is also known as the lateral tuberal nucleus (Diepen, 1962; Nauta and Haymaker, 1969). Bleier and colleagues (1979, 1985) named this cell group the medial tuberal nucleus and referred to what is described here as the magnocellular nucleus of the lateral hypothalamus (see below) as the lateral tuberal nucleus. Paxinos and Watson (1998) utilized the term medial tuberal nucleus and recognized a lateral condensation of neurons that is demarcated in Timm’s and acetylcholinesterase stained preparations, which they termed the nucleus terete (Te; “Atlas” plates 31 and 32). Many neurons in the TU express estrogen receptor mRNA (Simerly et al., 1990) and the connections of the TU appear to represent a subset of those described for the VMHVL. Thus, the TU provides dense inputs to other hypothalamic nuclei that contain high densities of neurons that express sex steroid receptors, including the medial preoptic nucleus, the VMHVL, and the ventral premammillary nucleus. However, the TU appears to provide stronger inputs

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FIGURE 6 Hormone–sensitive neurons in the hypothalamus Dark-field photomicrographs to show the distribution and density of neurons in the arcuate nucleus (Arc) and ventrolateral part of the ventromedial (VMHVL) nucleus that express estrogen receptor (ER) mRNA (A). Panel B illustrates the density of androgen receptor (AR) mRNA containing neurons in the ventral premammillary nucleus (PMV; X40). Note the relative absence of neurons that express ER or AR mRNA in adjacent cell groups. (C) Cluster of neurons in paraventricular nucleus (PaV) that express ERβ mRNA. (D) Expression of PR mRNA in the AVPV of a female rat (original magnification, ×50).

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than the VMHVL to regions that participate in regulating neurosecretion, such as the arcuate, anterior periventricular, and paraventricular nuclei of the hypothalamus, and its projections to the amygdala, midbrain, and caudal brain stem are weaker than those of the VMHVL (Canteras and Simerly, 1994). The DM occupies the dorsal half of the tuberal area between the anterior hypothalamic nucleus, rostrally, and the posterior hypothalamic area, caudally. It is generally considered to be composed of three parts which are best appreciated at mid levels through the nucleus where the cell-dense posterior part of the DM (which corresponds to the compact part of the DM of Paxinos and Watson (1998) separates the dorsally positioned anterior subdivision from the ventral part of the nucleus (Atlas plate 31). The DM receives afferents from the BST and the ventral part of the lateral septal nucleus, and all parts of the brain stem that provide inputs to the MPO except for the nuclei of the raphe (Thompson and Swanson, 1998). In addition, the DM receives inputs from most parts of the hypothalamus including the AVPV, MPO, AH, VMH, and SCh. Although descending fibers from the DM provide inputs to the periaqueductal gray and appear to innervate Barrington’s nucleus, the parabrachial nucleus, and the nucleus of the solitary tract, the projections of the DM are mostly intrahypothalamic (Ter Horst and Luiten, 1986; Thompson et al., 1992). Thus, the most densely innervated areas are the anterodorsal and suprachiasmatic preoptic nuclei, the parastrial nucleus, the preoptic and anterior periventricular nuclei, and the parvicellular parts of the paraventricular nucleus. The functional role of the DM remains a matter of discussion, but it has been implicated in the regulation of ingestive behavior, stress, reproduction, circadian rhythms, and thermogenesis (Bernardis and Bellinger, 1998; Sawchenko, 1998), (Gunnet and Freeman, 1983; Rothwell, 1994; Polston and Erskine, 1995; Coolen et al., 1996; Thompson and Swanson, 1998, 2002). Mammillary Region The mammillary part of the medial zone of the hypothalamus is occupied by the mammillary complex— which consists of the medial and lateral mammillary (mammillary body) and supramammillary nuclei— together with the premammillary nuclei (dorsal and ventral) and the posterior hypothalamic area. Although we also consider the tuberomammillary nucleus here, it should be considered as part of the lateral zone of the hypothalamus because of its widespread telencephalic projections and postulated role in modulating behavioral state (Köhler et al., 1985). The mammillary body is divided into a medial mammillary nucleus (MM) that occupies the majority of the mammillary

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body and a lateral mammillary nucleus (LM) that is easily distinguished by its large, darkly stained neurons. The MM is further subdivided into three parts: a small medial part (MMm) located at the midline, the lateral part (MMl) which is by far the largest, and the posterior part (MMp). Perhaps the most important input to the mammillary body is that from the subicular region of the hippocampal formation that travels along the postcommissural fornix (Swanson and Cowan, 1975b; Canteras and Swanson, 1992b; Kishi et al., 2000; Witter and Amaral, 2003). Other limbic inputs include afferents from the lateral septal nucleus and BST, which appear to preferentially terminate in the LM, while the medial septal–diagonal band complex preferentially innervates the medial nucleus (Swanson and Cowan, 1979). In contrast to the other nuclei of the medial zone, the mammillary body receives only modest inputs from the brain stem, although the dorsal tegmental nucleus of Gudden sends a projection to the LM and the ventral tegmental nucleus appears to preferentially innervate the MM. The projections of the mammillary body are equally unique among medial zone nuclei. Projection axons form two major fiber tracts: a descending mammillotegmental tract containing fibers from both the MM and the LM that terminate in the tegmental nuclei of Gudden, and the ascending mammillothalamic tract that terminates in the anterior complex of the thalamus. The anterior thalamic nuclei in turn send projections to a continuous strip of limbic cortex that includes the anterior limbic area, retrosplenial area, presubiculum, and parasubiculum and receives inputs from visual and auditory areas of the isocortex. The connections of the mammillary body serve to distinguish it from the other nuclei of the medial zone and separate this zone into distinct rostral and caudal (mammillary) components. In contrast to the strong olfactory, gustatory, and visceral sensory information that regulates the MPO, AH, DM, and VMH, the mammillary body appears to be most strongly influenced by visual and auditory information. In addition, the mammillary body lacks the extensive intrahypothalamic connections typical of the other medial zone cell groups. Finally, the MPO, AH, DM, and VMH all send projections to regions of the brain stem and spinal cord that control somatomotor and autonomic responses, whereas the mammillary body appears to lack such projections, although at least some intrahypothalamic connections have been reported (Gonzalo-Ruiz et al., 1992), and its cortical connections may provide an indirect pathway for inputs to the striatum (Swanson, 1987). The premammillary nuclei lie at the level of the mammillary recess of the third ventricle immediately caudal to the ventromedial hypothalamic nucleus and rostral to the mammillary body. The dorsal

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premammillary nucleus (PMD) appears as a trapezoidalor rectangular-shaped cluster of pale neurons in Nisslstained frontal sections and is bordered laterally by the fornix column and medially by a cell-poor zone that separates the two nuclei. The ventral premammillary nucleus (PMV) is somewhat easier to distinguish and was first recognized by Gurdjian (1927) as a compact cluster of darkly staining neurons. It is most clearly delimited by retrograde labeling, for example, after injections of tracers into the medial preoptic nucleus (Simerly and Swanson, 1986), or by labeling neurons in the PMV that express sex steroid receptors (Fig. 8) (Pfaff and Keiner, 1973; Simerly et al., 1990). The major inputs to the PMV arise in sexually dimorphic forebrain regions that compose a distinct circuit (see Simerly and Swanson, 1986; Simerly and Swanson, 1988; Canteras et al., 1992b) that includes the posterior nucleus of the amygdala, the medial preoptic nucleus, the principal nucleus of the BST (a.k.a. BSTMP) (Paxinos and Watson, 1998), and the posterodorsal part of the medial nucleus of the amygdala (see Simerly, 1995b, for review). Thus, olfactory information relayed by these nuclei appears to provide the major sensory input to the PMV. The PMV contributes strong inputs to most parts of the periventricular zone, including the anteroventral periventricular and arcuate nuclei, as well as sends projections to other sexually dimorphic regions that innervate it, which may represent feedback connections (Canteras et al., 1992b). The connections of the PMV are consistent with functional data that suggest a possible role in mediating certain aspects of reproductive behavior and physiology (Beltramino and Taleisnik, 1978, 1980; Akesson and Micevych, 1995), and perhaps aggressive behavior as well (Van den Berg et al., 1983). In contrast, the PMD has quite a different pattern of connections and Canteras and Swanson (1992a) have suggested that it may be viewed as a rostral component of the mammillary body that serves as an interface between the anterior nuclei of the medial zone and the mammillary body. Unlike the mammillary body, which lacks strong inputs from the hypothalamus, the PMD receives its major input from the anterior hypothalamic nucleus (Saper et al., 1978; Risold et al., 1994), which in turn receives inputs from the prefrontal cortex, the amygdala, and the hippocampus. The PMD is also innervated by the interfascicular nucleus of the BST and receives substantial input from the infralimbic and prelimbic areas, as well as from the lateral septal nucleus (Comoli et al., 2000). In addition to inputs from the anterior hypothalamus, the PMD is innervated by the perifornical region and the anterior and dorsomedial parts of the ventromedial hypothalamic nucleus. The projections of the PMD more closely resemble those of the mammillary body than those of the PMV. Like the

mammillary body the PMD sends a branched projection that ends in the anterior thalamus and brain stem, but it also provides afferents to the anterior hypothalamic nucleus and a descending projection to the periaqueductal gray, superior colliculus, and adjacent parts of the reticular formation (Canteras and Swanson, 1992a). Thus, the PMD represents an interface between medial zone nuclei of the rostral hypothalamus and mammillary nuclei. By virtue of its projections to the hippocampus (by way of the anteromedial nucleus of the thalamus and its projections) and to motor-related brain stem regions, it provides a means for communication between medial zone nuclei and brain regions involved in spatial memory and diverse behavioral responses including expression of fear and defensive responses to environmental stimuli (Canteras et al., 1997; Comoli et al., 2000). The supramammillary nucleus (SuM) lies between the mammillary body and the posterior hypothalamic area and is bordered dorsolaterally by the ventral tegmental area (VTA). It can be divided into a largecelled lateral part and small-celled medial part that contains many dopaminergic neurons (Swanson, 1982). Although its inputs have not been examined systematically, it appears to receive a massive input from the ventral part of the septal nucleus, the BST, the medial preoptic nucleus, and the lateral zone of the hypothalamus (Saper et al., 1979; Swanson and Cowan, 1979; Simerly and Swanson, 1988; Risold and Swanson, 1997b; Dong et al., 2000). The SuM projects to most major telencephalic regions, including the entire cortical mantle, including a prominent projection to the dentate gyrus and the entorhinal cortex. It provides weaker inputs to other parts of the hippoccampal formation and avoids the corpus striatum (Haglund et al., 1984; Vertes, 1992). The SuM also sends a projection to the central nucleus of the amygdala (Swanson, 1982). Descending projections of the SuM appear to be relatively minor, but a few fibers appear to end in the periaqueductal gray (Vertes, 1992). The posterior hypothalamic area (PHA), the most caudal and dorsal hypothalamic region, is limited laterally by the mammillothalamic tract and merges in the midline at the level of the mammillary recess. It begins just dorsal to the DM—a region often referred to as the dorsal hypothalamic area–which it replaces caudally until it merges with the periaqueductal gray of the midbrain. The connections of the PHA appear to have much in common with those of the periaqueductal gray. The PHA receives inputs from the amygdala, the septum, and the hippocampal formation, as well as from much of the hypothalamus, and many of these connections appear to be bidirectional (Abrahamson and Moore, 2001; Cavdar et al., 2001). The PHA sends the

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majority of its ascending projections via the medial forebrain bundle (mfb) and provides significant inputs to cortical regions related to limbic structures such as perirhinal, insular, limbic, and prelimbic regions of the cerebral cortex (Vertes et al., 1995). In addition, the PHA innervates subcortical regions in the hypothalamus, thalamus, amygdala, septum, and basal forebrain. Because of the close association between the PHA and the forebrain regions connected to the hippocampus it has been suggested that the PHA may play a role in various aspects of emotional behavior (Vertes et al., 1995). The majority of the tuberomammillary nucleus (TM) lies along the base of the caudal diencephalon and is best considered as a nuclear complex that forms a shell around ventral parts of the mammillary body. It can be divided into two magnocellular parts with scattered cells in between. Thus, the complex consists of a dorsomedial TM that appears as a dense cluster of large neurons along the dorsal aspect of the mammillary recess, just dorsal to the rostral part of the posterior periventricular nucleus (Atlas plate 34); a ventral nucleus (TMV) that is considerably larger than the TMD and lies along the base of the hypothalamus ventral and lateral to the PMV (Atlas plates 34 and 35); and a diffuse component (TMdif) that consists of scattered, darkly staining neurons between the TMD and the TMV within the posterior periventricular nucleus (Köhler et al., 1985). Various parcellation schemes have been used to classify these cell groups (for review see Ericson et al., 1987), but it was the advent of immunohistochemical methods that distinguished the TM from the more rostral supraoptic magnocellular neurons and clarified the organization of the complex. Most, if not all, neurons in the TM stain with antisera to histidine decarboxylase and amino decarboxylase, and all parts of the TM receive catecholaminergic afferents from the C1–C3 and A1–A2 cell groups in the brain stem, as well as serotonergic inputs from the B5–B9 cell groups (Ericson et al., 1989). The TM also receives afferents from the lateral septal nucleus and the bed nuclei of the stria terminalis, the medial preoptic nucleus, the lateral preoptic and hypothalamic areas, and the ventromedial nucleus (Ericson et al., 1991). The projections of the TM resemble those of the locus coeruleus and dorsal raphe in that they are distributed to widespread parts of the brain, including the entire cortical mantle, the basal ganglia, the septum and amygdala, parts of the thalamus, the cerebellum, and several brain stem nuclei. The extensive pattern of projections has led to the suggestion that the TM plays a role in the modulation of arousal and behavioral state and, therefore, conceptually belongs to the lateral zone of the hypothalamus. However, the strongest projections of TM neurons are to the hypothalamus, with particularly

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dense terminal fields in the paraventricular and supraoptic nuclei (Ericson et al., 1987), suggesting an important role for the TM in regulating neuroendocrine function as well.

Lateral Zone Although the functional importance of the lateral zone of the hypothalamus has been appreciated for several decades (Stevenson, 1969; Hitt et al., 1970; Siegel and Edinger, 1981), it is safe to say that no aspect of hypothalamic circuitry is less clear. Few distinct regions can be distinguished on cytoarchitectural or neurochemical grounds and attempts to functionally dissect this complex region have largely been impeded by technical limitations. The primary reason for this is that the lateral zone is traversed by the mfb, undoubtedly the most complex fiber system in the mammalian brain (Veening et al., 1982). The mfb contains ascending and descending fibers that arise from more than 50 different cell groups which extend from the prefrontal cortex, through the hypothalamus and ventral brain stem reticular formation, to sacral levels of the spinal cord, and many of these projections appear to contribute inputs to cells in the lateral zone. Thus, it has proven difficult to interpret experimental findings based on methods that fail to distinguish between specific manipulations of neurons and perturbations of fibers passing through the lateral zone. Nevertheless, the lateral zone of the hypothalamus has been implicated in the processing of sensory information and the expression of behaviors associated with hunger and thirst (Swanson and Mogenson, 1981; Swanson, 1987; Elmquist et al., 1999; Siegel, 1999; Watts, 2000), aggression (Kruk, 1991; Siegel et al., 1999), and reproduction (Pfaff et al., 1994). In more general terms, the functional evidence suggests that the lateral zone is involved in mediating general arousal and sensory sensitization associated with motivated behavior and may modulate spinal pathways and so affect the likelihood that specific behavioral patterns will be expressed (Swanson, 1987). However, by virtue of its strong connections with telencephalic regions such as the cerebral cortex, the amygdala, and the septum, and its connections with parts of the periventricular zone, it is in a good position to coordinate motivational aspects of behavior with visceromotor responses. Like the other zones of the hypothalamus the lateral zone can be divided into preoptic, anterior, tuberal, and mammillary regions. However, because clear criteria for subdivision have not emerged it is generally considered to have only two major divisions, the lateral preoptic area (preoptic level) and the lateral hypothalamic area (anterior, tuberal, and mammillary levels).

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Lateral Preoptic Area (LPA)

forebrain bundle, a definition that is supported by the results of anterograde tract-tracing studies (Fig. 7) (Swanson, 1976; Simerly and Swanson, 1988). The only distinct nucleus in the preoptic part of the lateral zone is the magnocellular preoptic nucleus (MCPO), which is differentiated from the dorsally adjacent LPA and substantia innominata by its large darkly staining neurons. This cell group has been called the “nucleus interstitialis septo-hypothalamicus” by Gurdjian (1927)

The LPA is characterized by lightly stained (thionin) medium-sized neurons scattered amongst the fibers of the dorsomedial division of the medial forebrain bundle. It is bordered dorsally by the BST and the substantia innominata, and caudally it merges imperceptibly with the lateral hypothalamic area. The medial border of the LPA (designated Lpo in Fig 7) corresponds rather precisely with the medial border of the medial

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FIGURE 7 Lateral zone projections. (A) Bright-field photomicrograph to illustrate the appearance and location of a PHA-L injection site centered in the lateral preoptic area (LPO). (B, C) Dark-field photomicrographs that illustrate projection axons and terminals, emanating from the injection site shown in panel A, observed in the lateral hypothalamic area (LH) and lateral habenula (LHb; B and C) and in the posterior hypothalamic area (C; original magnification, ×50). Note the relative absence of labeled fibers in the periventricular and medial zones of the hypothalamus. See Fig. 1 for other abbreviations.

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and the “nucleus of the horizontal limb of the diagonal band” by Price and Powell (1970). However, the MCPO is clearly separated from the ventrolateral tip of the nucleus of the diagonal band which forms the medial border of the MCPO. Because the MCPO consists mostly of large cholinergic neurons that project to the olfactory bulb and isocortex (Swanson, 1976; Woolf et al., 1984), it may be considered a differentiated component of the substantia innominata in the rat that corresponds in part to the basal nucleus of Meynert of primates (Saper, 1984, 2000; Saper and Chelimsky, 1984). The projections of the LPA travel almost exclusively in the medial forebrain bundle. In contrast to the projections of the medial preoptic nuclei, the LPO provides widespread inputs to the cerebral cortex and hippocampus, as well as inputs to the mediodorsal and reticular nuclei of the thalamus (Swanson, 1976; Simerly and Swanson, 1988). In addition, the dorsomedial hypothalamic nucleus, supramammillary nucleus, and ventral tegmental area receive significant inputs from the LPO. Descending projections extend into the medulla with particularly dense terminal fields in the parabrachial and pedunculopontine nuclei. Lateral Hypothalamic Area (LHA) The LHA may be further divided into three rostrocaudal subregions that correspond to the anterior (LHAa), tuberal (LHAt), and mammillary (posterior) (LHAp) levels of the hypothalamus. Despite containing several cell condensations the LHA has not been further subdivided, although each major subdivision appears to have a unique set of connections (Saper et al., 1979). The LHAa is continuous with the LPA and is bounded medially by the anterior hypothalamic area and the descending columns of the fornix. Laterally it merges with the substantia innominata and the amygdala. The LHAt replaces the LHAa at tuberal levels and is bordered laterally by the optic tract and the internal capsule and, more caudally, by the subthalamic nucleus. The TU is embedded in the LHAt and consists of an oval cluster of neurons lying lateral to the VMH. Many TU neurons express estrogen receptors (Simerly et al., 1990) and have connections that are similar to those of the VMH (Canteras and Simerly, 1994). At the level of the mammillary complex, the LHAt is replaced by the LHAp which merges caudally with the ventral tegmental area. Medially the LHAp is limited by the fornix, the mammillothalamic tract, and the posterior hypothalamic area and is separated dorsally from the thalamus by the zona incerta and the fields of Forel. The connections of the LHA are extremely complex and, although it is clear that fibers in the medial forebrain bundle innervate neurons in the lateral zone

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(Millhouse, 1979; Veening et al., 1982), for the most part the region-specific pattern of lateral zone afferents has not been established. The results of retrograde tracttracing studies have proven difficult to interpret because of the possibility of retrograde labeling produced by uptake of tracers by fibers of passage. This problem is severe in the lateral zone because all regions that contribute fibers to the medial forebrain bundle also project to one or more nuclei in the medial zone (see Swanson, 1987, for references). Information derived from anterograde tract-tracing studies suggests that restricted parts of the amygdala, septum, and hippocampal formation supply strong inputs to the LHA, as do the substantia innominata and lateral preoptic areas. In general, cell groups in the periventricular zone do not send many projections to the LHA, but the arcuate nucleus is an exception and sends direct inputs to peptidergic neurons in the LHA that are thought to play an important role in feeding (Elias et al., 1999). Other parts of the periventricular zone, such as the anteroventral periventricular nucleus, have relatively weak descending projections to the lateral zone that appear to end primarily in the perifornical region and in the ventrolateral part of the LHAt (Gu and Simerly, 1997). Although brain stem regions also appear to provide inputs to the LHA, the detailed organization of these pathways is yet to be defined. Evidence from systematic anterograde tract-tracing studies indicates that the outputs of the lateral zone of the hypothalamus closely resemble those of the medial zone as a whole. Thus, widespread projections have been demonstrated that include inputs to the entire cerebral cortical mantle (including the hippocampal formation), parts of the amygdala and septum, the substantia innominata, parts of the thalamus, nearly every part of the periventricular and medial zones of the hypothalamus, numerous brain stem cell groups, and the spinal cord. As a whole the hypothalamus provides the largest nonthalamic input to the cerebral cortex. This input is derived primarily from neurons in the lateral zone of the hypothalamus (Saper, 1985, 2000), in addition to the cortical inputs supplied by the tuberomammillary and supramammillary nuclei alluded to above. The lateral zone also projects to many parts of the septum and amygdala, but this projection is much more diffuse than the massive input it receives from these telencephalic regions, suggesting that it represents a possible feedback pathway. With respect to projections from the lateral zone to the thalamic nuclei, both the LPA and the LHA appear to provide inputs to the lateral habenula, the paraventricular nucleus of the thalamus, the nucleus reunions, and the zona incerta, as well as to the intralaminar nuclei (Swanson, 1976; Saper et al., 1979; Berk and Finkelstein,

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1982; Simerly and Swanson, 1988). These thalamic cell groups, in turn, may relay afferent information from the lateral zone to the hippocampal formation, amygdala, septum, prefrontal cortex, and nucleus accumbens. Neurons in the lateral zone send projections to widespread regions in the brain stem and most of these connections are bidirectional. These regions include nuclei that relay visceral sensory information to the forebrain, such as the nucleus of the solitary tract and the parabrachial nucleus (Saper and Loewy, 1980; Ter Horst et al., 1989; Bester et al., 1997), and nuclei involved in mediating somatomotor control mechanisms, such as the ventral tegmental area and substantia nigra, the posterior hypothalamus, and periaqueductal gray, as well as the locus coeruleus and various nuclei of the raphe (Swanson, 1976; Saper et al., 1979). Finally, there are direct projections from the LHA to the spinal cord that are distributed such that it may directly influence sensory, somatomotor, and autonomic spinal mechanisms (Saper et al., 1976a; Hosoya and Matsushita, 1983).

HYPOTHALAMIC INTEGRATION At the outset of this review the hypothalamus was described as providing a suitable neurological substrate for coordinating the needs of the individual animal with dynamic changes in its environment. The appropriate display of adaptive responses to specific cues from the environment involve complex endocrine, autonomic, and somatomotor mechanisms that must be integrated for these responses to be of optimal benefit to the animal. Moreover, because the simultaneous expression of multiple behaviors is not adaptive, these responses must be coordinated and be subject to a hierarchy of homeostatic priorities, a process termed “motivational time-sharing” by McFarland (1974). For example, animals do not generally feed while copulating or simultaneously display defensive and courtship behaviors. These functional requirements, therefore, imply not only integration of endocrine, autonomic, and somatomotor aspects of each adaptive response but also coordination and communication between neural systems mediating different functional responses. Behaviors are generally considered to consist of three phases: initiation, procurement, and consummation (see Beach, 1967; Swanson and Mogenson, 1981, for reviews). The initiation phase involves the perception of a stimulus, while the procurement phase consists of behaviors that are directed toward a specific goal. The consummatory phase is characterized by preprogrammed motor responses thought to be established by central pattern generators. By analogy, endocrine responses can also be divided into three distinct phases.

Hormone secretion from the anterior lobe of the pituitary gland (functional output or consummation) is controlled by neuroendocrine neurons in the hypothalamus whose intrinsic activity is controlled by neural circuits that regulate the pattern of hormone secretion into the portal circulation (pattern generation). These neural circuits are in turn activated, or modulated, by both neural and humoral afferent information (initiation). Thus, a prerequisite to understanding the neurobiology of hypothalamic integration is the characterization of not only the neural pathways that convey specific signals to the hypothalamus but also of circuits that mediate the procurement and consummatory aspects of goal-directed behaviors or, alternatively, the secretion patterns of hypothalamic hormones. Superimposed on the organization of these neural pathways are patterns of neural activity that are regulated by changes in physiological state (e.g., estrous cycle, availability of metabolic fuels, circulating hormones, etc.) that alter the transmission of information through hypothalamic circuits. Although hypothalamic circuits are considerably more complex than the three neuron reflex arcs studied by Sherrington (1906), hypothalamic circuits can be viewed as essentially consisting of sensory and motor components, with intrahypothalamic integrative circuits interposed between the sensory and motor parts of the circuit. As outlined above, the hypothalamus receives sensory activation from visceral sensory regions such as the nucleus of the solitary tract and subfornical organ, as well as olfactory and other sensory information from limbic regions such as the amygdala and hippocampal formation. Superimposed on hypothalamic regions receiving these sensory pathways are projections from the brain stem reticular formation that may effect a more generalized arousal. With respect to motor pathways, the hypothalamus sends direct and massive projections to preganglionic autonomic neurons and all parts of the pituitary gland, as well as projections to regions that mediate somatomotor responses. Thus, the hypothalamus represents an interface between sensory and motor pathways that has intimate connections with the pituitary gland, thereby providing an effective means of coordinating stimulus-specific behaviors with appropriate autonomic and endocrine responses. The integrative mechanisms underlying this process occur at the level of intact neural systems, individual hypothalamic nuclei, and single cells and at the molecular level. Integration at the Neural Systems Level A great deal remains to be learned regarding the organization of the neural systems that mediate homeostatic responses, but certain patterns are emerging that

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suggest ways in which essential behaviors and endocrine responses are coordinated (Risold et al., 1997; Swanson, 2000). One such mechanism is for separate sensory pathways to converge onto functionally distinct parts of the hypothalamus. For example, pheromonal information is relayed to the hypothalamus from the vomeronasal organ by the medial amygdala and principal nucleus of the BST (Fig. 8). Each of these limbic regions provide strong inputs to both the anteroventral periventricular nucleus and the parvicellular division of the paraventricular nucleus. The anteroventral periventricular and paraventricular nuclei also receive strong inputs from the ventral part of the lateral septal nucleus, which is innervated topographically by the ventral subiculum and hippocampal field CA1. Thus, these two periventricular zone nuclei receive convergent sensory information: olfactory cues conveyed to the hypothalamus by the medial amygdala and BST, and information related to exploratory behavior conveyed to the hypothalamus by the ventral lateral septum. In a similar way, parallel limbic inputs to sexually dimorphic nuclei in the hypothalamus from the hippocampal formation and accessory olfactory pathway converge onto subcircuitries that regulate reproductive behavior and neuroendocrine responses (Simerly, 2002). In this way sexually relevant sensory cues can coordinate display of reproductive behavior with accompanying changes in reproductive physiology. Analysis of hypothalamic connections suggests that there are functionally distinct sets of pathways that mediate different homeostatic functions. Reproductive behavior is mostly dependent on the activities of the medial preoptic nucleus and ventrolateral part of the ventromedial hypothalamic nucleus, which play decisive roles in male and female copulatory behavior, respectively (Meisel and Sachs, 1994; Pfaff et al., 1994). Agonistic behavior is primarily controlled by a circuit between the anterior hypothalamic nucleus, the dorsomedial part of the ventromedial nucleus, and the dorsal premammillary nucleus (Canteras and Swanson, 1992a; Canteras et al., 1997). In contrast to the predominate role of these medial zone nuclei in social behaviors, nuclei of the periventricular zone are chiefly involved in neuroendocrine regulation and visceromotor responses (Markakis and Swanson, 1997). The major components of this system are the anteroventral periventricular nucleus, the periventricular and paraventricular hypothalamic nuclei, and the arcuate nucleus. Strong intrahypothalamic connections link these cell groups and they receive similar sensory inputs, either directly by regions such as the ventral lateral septal nucleus and the principal BST or via relays from medial zone nuclei such as the medial part of the medial preoptic nucleus and dorsomedial hypothalamic nucleus. These same

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FIGURE 8 Integration at the neural systems level. Pheromonal information from the accessory olfactory bulb (AOB) is relayed to the hypothalamus by cell groups in the amygdala and the BST. This information differentially impacts subsystems in the hypothalamus involved in defensive and reproductive behaviors. The reproductive subsystem has strong connections with periventricular nuclei that control endocrine and autonomic responses, whereas the defensive subsystem projects to several nuclei in the rostral thalamus thought to participate in attention and learning (modified from Fig. 2 of Risold et al., 1997, with permission ).

major sensory routes provide input to the medial preoptic nucleus and the ventrolateral part of the ventromedial nucleus. Therefore, a potential mechanism for functional integration is for a single sensory modality to affect multiple functional neural systems. For example, olfactory influences emanating from the vomeronasal organ play an important role in modulating gonadotropin secretion and the display of copulatory behavior (Wysocki, 1979; Johns, 1986). This olfactory information is relayed to the hypothalamus primarily by the medial nucleus of the amygdala and the principal nucleus of the BST (see Simerly, 1990), both of which provide strong inputs to the medial preoptic nucleus and ventromedial nucleus (Fig. 6) (Swanson and Cowan, 1979). The medial nucleus of the amygdala and the principal nucleus of the BST also provide strong inputs to the anteroventral periventricular and ventral premammillary nuclei, which in turn project to regions

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such as the paraventricular and arcuate hypothalamic nuclei, which contain high densities of neurons that secrete hypothalamic-releasing factors. Thus, the distribution of a distinct sensory input to neural systems modulating different functions provides a direct means of coordinating behavioral and endocrine responses to a single stimulus. Similarly, regions involved in mediating motor aspects of behavior can receive inputs from hypothalamic cell groups involved in initiating different behaviors, as illustrated by the strong projections from the medial preoptic nucleus (male copulatory behavior) and the ventromedial nucleus (female copulatory behavior) to the periaqueductal gray. Although the same major regions that supply telencephalic sensory inputs to the subsystem of hypothalamic nuclei involved in reproduction also send inputs to the subsystem that mediates agonistic behavior, these inputs are topographically organized such that each subsystem is differentially regulated by the telencephalon (Fig. 8) (see Risold et al., 1997). Connections between various parts of functionally distinct neural systems, such as those that exist between the medial preoptic and ventromedial nuclei, or between dorsomedial hypothalamic and anteroventral periventricular nuclei, may serve to coordinate the activity of these systems and provide for the display of appropriate responses. An important illustration of this principle relates to how animals integrate intrinsic physiologic stressors with those that require interpretation by the animal. The paraventricular nucleus acts as the final common pathway for the regulation of glucocorticoid secretion and receives convergent inputs that transmit visceral sensory information, via brain stem afferents, and telencephalic afferents from the limbic region of the forebrain that convey information from the external environment requiring interpretation before elaborating the appropriate adaptive response (Herman and Cullinan, 1997). Integration at the Level of Hypothalamic Nuclei Insight into the function of a hypothalamic nucleus can be gained by considering the functions of the structures it innervates, the types of information it receives from regions that supply afferents to it, and the amount of information processing that occurs within the nucleus. Most hypothalamic nuclei can be divided into components that share a distinct pattern of connections or cytoarchitectural and neurochemical characteristics. The spatial segregation of functionally distinct cell types in the periventricular zone is best illustrated by the paraventricular nucleus of the hypothalamus (PaV) (Swanson and Kuypers, 1980b; see Fig. 9). The projections of essentially separate populations of neurons in the PaV to the posterior pituitary, to the anterior pituitary,

and to autonomic preganglionic neurons divide the PaV into three distinct components. However, on homological, cytoarchitectonic, and neurochemical grounds these components can be subdivided further to form eight distinct subdivisions which represent an unusual degree of anatomical complexity and suggest that each subdivision may play a unique role in mediating visceral responses (Swanson and Sawchenko, 1983). This compartmentalization also suggests several possible mechanisms whereby disparate visceral responses are integrated. For example, different cell types in the PaV may receive a common input, such as the brain stem afferents from the nucleus of the solitary tract, which convey visceral sensory information to both the oxytocin neurons in the magnocellular division of the PaV and the corticotropin-releasing hormone neurons in the parvicellular division of the nucleus (Sawchenko et al., 1996). Additional integrative mechanisms operating at the level of the PaV are for different cells to have outputs that converge onto the same motor effector circuit or to be coupled by recurrent collaterals or connections with interneurons (Rho and Swanson, 1987). The organizational principals identified for the PaV may also apply to medial zone nuclei. As reviewed above, a variety of anatomical criteria indicate that the MPO is composed of three distinct sexually dimorphic subdivisions: a cell-sparse lateral subdivision and celldense medial and central subdivisions that are larger in males (Simerly et al., 1984b). As is the case for the PaV, sensory inputs are not uniformly distributed throughout the subdivisions of the MPO. Olfactory information relayed to the MPO by the medial amygdala and principal nucleus of the BST is primarily distributed to the medial and central parts of the MPO. Other classes of cortical information reach the MPO via the infralimbic cortex, subiculum, and rostral septal nucleus and end primarily in the lateral part of the MPO (Simerly and Swanson, 1986; Swanson, 2000), although projections from the ventral lateral septal nucleus are mostly distributed in the medial part of the MPO (Risold and Swanson, 1997a). Ascending noradrenergic inputs from the brain stem, which probably carry visceral sensory information, are distributed primarily to the central and medial parts of the MPO, whereas serotonergic afferents are largely localized to the lateral part of the nucleus. Thus, each compartment of the MPO receives a unique complement of inputs and may therefore respond differently to specific sensory modalities. However, because dendritic fields of one compartment may extend into an adjacent compartment, there may be some degree of overlap in sensory responsiveness. Nevertheless, afferents that terminate on distal dendrites have a functional impact distinctly different than that of inputs that terminate

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FIGURE 9 Integration at the level of hypothalamic nuclei. Schematic drawing of a midsagittal section through the rat brain to illustrate possible convergence of major sensory pathways onto CRH neurons in the paraventricular nucleus. Abbreviations used: C1, adrenergic cell groups; CG, central gray; HYP, hypothalamus; IGL, intergeniculate leaflet; LIMBIC, limbic region; LDT, laterodorsal tegmental nucleus; MePO, median preoptic nucleus; NTS, nucleus of the solitary tract; PB, parabrachial nucleus; PIN, posterior intralaminar nucleus; PP, peripeduncular nucleus; PPN, pedunculopontine nucleus; PVT, paraventricular nucleus of the thalamus; SFO, subfornical organ. (Reproduced from Sawchenko, P. E., Imaki, T., Potter, E., Kovacs, K., Imaki, J., and Vale, W. (1993) The functional neuroanatomy of corticotropin-releasing factor. Ciba Foundation Sympasium 172, 5–29, 1998, with permission).

directly onto the cell soma. Because each component of the MPO also displays a unique pattern of projections (Simerly and Swanson, 1988), the functional impact of information conveyed by these outputs depends at least in part on the distribution and intensity of afferents to each part of the MPO. Convergent inputs to a region that arise from multiple parts of a nucleus, such as those from the central and medial parts of the MPO to the arcuate nucleus, may provide for the integration of processed sensory cues. Each compartment of a nucleus may also display a unique neurochemical organization with the majority of neurons that display a particular transmitter phenotype localized to one compartment. This is especially true for the magnocellular neurons of the PaV, although under certain hormonal conditions vasopressin is induced in parvicellular neurons (Sawchenko et al., 1984). In a similar way, each neurotransmitter-specific cell type found in the MPO is largely localized to one of its three subdivisions (Simerly et al., 1986). For instance, cholecystokinin- and thyrotropin-releasing hormoneimmunoreactive neurons are nearly entirely localized to the central part of the MPO, whereas neurotensin and corticotropin-releasing factor-immunoreactive cells are localized to the lateral part of the nucleus. Similarly, neurotransmitter-specific terminals appear to be distributed in accordance with the three subdivisions of the MPO as illustrated by substance P- and enkephalin-immunoreactive fibers, which are localized primarily to the medial and lateral parts of the MPO, respectively. In addition, each compartment of a nucleus may contain unique patterns of neurotransmitter

receptors adding additional complexity to the complete response profile of hypothalamic nuclei. The hypothalamus is particularly responsive to the regulatory effects of circulating factors. Steroid and thyroid hormones have free access to hypothalamic neurons, and many aspects of hormone responsiveness depend on the expression of specific nuclear receptors by discrete subpopulations of cells in the hypothalamus. Autoradiographic, immunocytochemical, and in situ hybridization methods have been used to demonstrate the distribution of neurons that express steroid and thyroid hormone receptors including androgen, estrogen, progesterone, corticosteroids, and thyroid hormone receptors (see Simerly, 1993, for review). That hormone-sensitive neurons are often distributed in accordance with cytoarchitectonic subdivisions of hypothalamic nuclei (Simerly et al., 1990) suggests that functions mediated by such nuclear compartments may be differentially modulated by changes in levels of circulating hormones. Conversely, expression of hormone receptors by otherwise separate populations of neurons may serve to coordinate the activity of these hormone-sensitive neurons. Strong projections from telencephalic regions that contain high densities of neurons that express steroid hormone receptors to hypothalamic nuclei that also receive hormonesensitive intrahypothalamic inputs indicate that the regulatory effects of hormones can be amplified by convergent inputs from hormone-sensitive regions. For example, the medial part of the medial preoptic nucleus receives strong inputs from the posterodorsal part of the medial amygdaloid nucleus and principal

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nucleus of the BST, as well as from the anterior periventricular, arcuate, ventral premammillary, and ventromedial nuclei of the hypothalamus, all of which have numerous cells that express high levels of receptors for sex steroid hormones. Taken together with the fact that most neurons in the medial part of the MPO express high levels of sex steroid receptors it is clear that the influence of sex steroids acting locally can be complemented by that of the same hormones acting on afferent populations of neurons, thereby bringing together disparate hormonally regulated sensory influences at the level of the MPO. In addition to steroid and thyroid hormones other factors secreted in the periphery travel in the blood to regulate the activity of hypothalamic circuits. The adipocyte-derived leptin hormone acts to inhibit food intake and stimulates catabolic autonomic and neuroendocrine responses that tend to direct nutrient stores away from the fat compartment (Woods et al., 1998).

The effectiveness of leptin in regulating energy stores is due to its direct access to hypothalamic neurons that control feeding behavior and other aspects of energy metabolism (Fig. 10) (see Sawchenko, 1998; Elmquist et al., 1999; Watts, 2000; Woods et al., 2000). Distinct subsets of hypothalamic neurons may respond to humoral factors such as leptin by virtue of their location in a region where the blood–brain barrier is compromised or through connections with circumventricular organs. The arcuate nucleus of the hypothalamus meets both of these features since it resides above the median eminence and shares connections with regions such as the subfornical organ and ovlt. The arcuate nucleus has long been associated with obesity (Olney, 1969), expresses high levels of leptin receptor (Fei et al., 1997; Elmquist et al., 1998a; Hakansson et al., 1998), and has high densities of neurons that express fos protein following intravenous injection of leptin (Woods and Stock, 1996; Elmquist et al., 1998b). Moreover, the

FIGURE 10 Neurohumoral integration in feeding pathways. Schematic representation of the organization and chemical specificity of projections from leptin–sensitive neurons in the arcuate nucleus to the dorsomedial nucleus of the hypothalamus (DM), the paraventricular nucleus (PVH), and the lateral hypothalamic area (LHA). Neurons in the arcuate nucleus that express neuropeptide Y (NPY) and agouti-related protein (AgRP; black circles), or that express proopiomelanocortin (POMC) products (gray circles), project directly to discrete populations of LHA neurons that express melanin-concentrating hormone (MCH; black circles) or hypocretin/orexin (H/O; gray circles), which have been implicated in the control of feeding behavior. MCH and H/O neurons are thought to send widespread projections to regions involved in generalized arousal and sensorimotor integration. Abbreviations used: CRF, corticotropin-releasing factor; IML, intermediolateral cell column; PAG, periaqueductal gray; TRH, thyrotropin-releasing hormone; VMH, ventromedial nucleus; ZI, zona incerta. (Reproduced from Sawchenko, 1998, with permission)

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projections of the arcuate nucleus to other sites implicated in the control of feeding such as the paraventricular and dorsomedial nuclei that also show leptin-induced increases in fos expression suggest that the arcuate nucleus is a principal monitor of leptin signaling in the brain. Integration of leptin signals with other systems mediating a variety of homeostatic mechanisms, such as reproduction (Mantzoros, 2000; Schneider et al., 2000), may take place at the level of the arcuate nucleus by virtue of common populations of peptidergic neurons with divergent connections. For example, injections of leptin activate both NPY and POMC neurons in the arcuate nucleus, which project to regions such as the paraventricular and dorsomedial nuclei and appear to convey leptin-based signals (Elias et al., 2000), but arcuate neurons also project to gonadotropin-releasing hormone cells in the preoptic region (Chen et al., 1989; Li et al., 1999). Thus, different physiological functions may be coordinated by interactions between neurons located in the arcuate nucleus through common activation by leptin, or possibly through intercellular communication. Similarly, projections to the anterior and ventral parts of the dorsomedial hypothalamic nucleus from the anteroventral periventricular nucleus, presumably carrying information sensitive to sex steroid regulation, overlap with those from the arcuate nucleus that may be influenced by leptin. Thus, convergent humor signals may regulate the activity of neurons in the dorsomedial nucleus via these convergent neural pathways (Gu and Simerly, 1997; Thompson and Swanson, 1998; Qiu et al., 2001). However, since the leptin-induced foscontaining neurons are in the posterior part of the dorsomedial nucleus, coupling of these different hormonally regulated sets of signals must take place at the level of the dorsomedial nucleus, perhaps through local connections between nuclear subdivisions. Therefore, it is important to bear in mind that some of the distinct classes of neurons defined by the criteria outlined above may be coupled within a nucleus by gap junctions, recurrent collaterals, or connections with interneurons, thereby providing for an additional level of information processing and integration. Integration at the Cellular Level Despite the organization of hypothalamic nuclei into compartments with distinct connections, rarely are such compartments homogeneous with respect to the neurochemical phenotype of individual cells. For instance, neurons in the medial parvicellular part of the PaV that contain corticotropin-releasing hormone (CRH) can coexpress at least seven other neuropeptides (enkephalin, vasopressin, neurotensin, angiotensin II, vasoactive intestinal polypeptide, PHI, and cholecystokinin). Technical limitations have prevented deter-

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mining if single CRH neurons are capable of expressing all of these molecules, but triple-labeling studies have shown that at least some CRH neurons coexpress two other peptides (Swanson et al., 1987; Sawchenko, 1991). Little is currently known, however, regarding the cellular mechanisms responsible for patterns of coexpression within subpopulations of CRH neurons. Moreover, these patterns of coexpression are often dynamic with fluctuations in the expression of some, but not other, coexpressed peptides within individual neurons (Sawchenko et al., 1992; Kovács and Sawchenko, 1996). Such differential regulation of coexpressed neuropeptides may represent a cellular mechanism for “chemical switching” of information through hypothalamic circuits (Swanson, 1983b, 1991). For example, neurons in the medial nucleus of the amygdala that project to the hypothalamus coexpress cholecystokinin (CCK) and substance P, yet changes in levels of circulating sex steroid hormones affect only levels of cholecystokinin (Simerly et al., 1989). Thus, when circulating levels of estrogen are high, as they are during proestrus, cellular levels of cholecystokinin are induced and decrease in response to lowered levels of circulating hormone. In contrast, cellular expression of substance P in these cells remains unchanged during the estrous cycle, or in response to acute changes in circulating estrogen. Taken together with similar finding in other pathways (Swanson, 1991), these observations suggest a possible cellular mechanism for gating sensory information through hormone-sensitive circuits. If we assume that release of coexpressed peptides at individual synapses is proportional to cellular content, then the functional impact of this release might preferentially promote activation of postsynaptic neurons that express cholecystokinin receptors, relative to those that express only receptors for substance P. If similar events occur in the vomeronasal pathway, activation of neurons in the medial amygdala by olfactory stimuli during periods when cholecystokinin levels are increased will lead to a preferential activation of postsynaptic cells that express high levels of CCK receptors, thereby effecting a hormone- dependent “switching” of sensory information to discrete subpopulations of hypothalamic neurons (Simerly, 1990). Similarly, endocrine changes bring about alterations in the expression of neuropeptides contained in neurons that express CRH (Swanson et al., 1987; Sawchenko, 1991) and gonadotropinreleasing hormone (Marks et al., 1992), which may alter the impact of these releasing factors on hormone secretion from the anterior pituitary. The distribution of neurotransmitter receptors in the hypothalamus has been studied extensively with autoradiographic methods. This approach has contributed greatly to clarifying which hypothalamic cell

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groups are responsive to various neurochemical signals. However, these binding techniques do not afford cellular resolution and it is likely that considerable heterogeneity exists with respect to the receptor profiles of subpopulations of hypothalamic neurons. For example, the AVPV receives a dense innervation by fibers that contain substance P immunoreactivity that is distributed throughout the nucleus, but it is not known if each of the many neurotransmitter-specific cell types found in the AVPV express substance P receptors. The cloning of cDNAs that encode neurotransmitter receptors and the use of in situ hybridization histochemistry, together with generation of receptor subunit-specific antibodies, has made possible the localization of receptor expression to individual neurons. It is a common observation in histochemical studies that some, but not all, neurons in a hypothalamic nucleus express particular receptors, but more colocalization studies are needed in order to determine if the pattern of expression represents distinct populations of neurons. In addition, cell-type-specific patterns of receptor regulation may subdivide further the functional classes of hypothalamic neurons and provide for additional mechanisms whereby the flow of information through functionally distinct circuits is regulated. Because it is likely that individual neurons receive multiple neurochemically distinct inputs it will be important to determine if synapse-specific regulation of neuronal responsiveness such as that demonstrated for neurons in the hippocampus (Malinow et al., 2000) exists in hypothalamic circuits. It has been shown that hormonal modification of synaptic organization (Matsumoto, 1991; Segarra and McEwen, 1991; Garcia-Segura et al., 1994; Mong et al., 2001) and intracellular receptor distribution (Coirini et al., 1989; Eckersell et al., 1998) occur, demonstrating that such cellular plasticity is likely to be a common aspect of cellular integration of multiple signals in the hypothalamus. Neural pathways from neurons responding to leptin may converge onto neurons in the paraventricular nucleus and contribute to cellular integration of chemically specific signals related to food intake. Neuropeptide Y (NPY) and agouti-related peptide (AGRP) are coexpressed within arcuate neurons and their projections. Together with melanocortin peptides, such as α-MSH, NPY, and AGRP represent key regulatory peptides since they are orexigenic and melanocortins are anorexigenic. AGRP is a potent antagonist of melanocortin signaling at MC4 melanocortin receptors, and in the paraventricular nucleus this interaction is likely presynaptic since MC4 receptors do not appear to be expressed on paraventricular neurons in abundance (Cowley et al., 1999). Moreover, melanocortins and NPY appear to be functional antagonists in the

paraventricular nucleus and have opposing actions on GABA-evoked currents within individual paraventricular nucleus neurons (Cowley et al., 1999). Thus, it has been proposed that POMC and NPY/AGRP neurons in the arcuate nucleus project to GABAergic neurons in the paraventricular nucleus to regulate GABA release. Since both NPY/AGRP and POMC neurons respond to leptin, their coordinate action on GABA release in the paraventricular nucleus provides an efficient means of integrating orexigenic and anorexigenic signals at the level of individual paraventricular nucleus neurons. Despite the complexity of cellular interactions in the hypothalamus some of the mechanisms underlying neurohumoral integration are beginning to emerge. It is already clear that subpopulations of neurotransmitterspecific neurons have unique signal transduction properties. For instance, progesterone treatment appears to cause an induction of the immediate early gene c-fos in a subpopulation of gonadotropin-releasing hormone neurons (Lee et al., 1990), and estrogen and progesterone cause changes in noradrenergic neurotransmission at both the receptor and the second messenger level (Petitti and Etgen, 1990). Estrogen can also alter neuronal responsiveness to GABA and glutamate (Wong and Moss, 1992; Moss et al., 1997; Gu et al., 1999a, 1999b), indicating that information transmitted by these neurotransmitters can have quite different effects on populations of hypothalamic neurons depending on whether they possess the ability to respond to changes in circulating levels of estrogen. Similarly, the leptin-sensitive NPY and POMC containing neurons in the arcuate nucleus that innervate melanocortin and hypocretin/orexin neurons in the lateral hypothalamus show different cellular responses to leptin (Elias et al., 1999). Thus, hypothalamic nuclei may be composed of distinct compartments that are made up of subpopulations of neurons with different regulatory profiles and response characteristics. It is also important to keep in mind that different parts of the hypothalamus may have unique compliments of glial cells. The increasing appreciation of the diverse roles played by glial cells in regulating microenvironments and cell–cell communication (Chowen et al., 2000; Araque et al., 2001; Bezzi and Volterra, 2001) provides a broad array of cellular mechanisms for specifying regionally distinct aspects of cellular integration. Integration at the Molecular Level Given the diverse array of environmental stimuli that affect neuropeptide gene expression in the hypothalamus, the molecular mechanisms underlying transcriptional activation in hypothalamic neurons will prove to be exceedingly complex. This is not surprising since

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hypothalamic neural circuits are made up of neurons that have much in common with neurons in other parts of the brain, and progress toward understanding how neurons integrate diverse signals at the molecular level is proceeding rapidly. The ability of a particular stimulus to alter gene expression depends on the presence of the appropriate signal transduction pathway, including the expression of molecules to initiate signaling (e.g., neurotransmitter or hormone receptors), an appropriate complement of transcription factors that couple the stimulus with the genome, and the presence of corresponding enhancer elements in the promoter region of the target gene (Goodman, 1990; Habener, 1990; Herdegen and Leah, 1998). For example, responsiveness to direct activation of gene expression by steroid and thyroid hormones can be dependent on the presence of the hormone receptor and a hormone response element in the hormone-sensitive gene (Yamamoto, 1985; Evans, 1988; Tsai and O’Malley, 1994; Beato and Sanchez-Pacheco, 1996; Jenster et al., 1997; Zhang and Lazar, 2000; Aranda and Pascual, 2001). Similarly, genes that display transynaptic transcriptional activation generally contain binding sites for sequence-specific trans-acting factors that respond to changes in levels of intracellular second messengers such as cAMP and calcium (Montminy, 1997; De Cesare and Sassone-Corsi, 2000). This traditional view of hormonal and transynaptic regulation of gene expression remains essentially valid, but the isolation of proteins that function as coregulators of transcription through multiple protein–protein interactions has revealed regulatory mechanisms that profoundly complicate our understanding of transcriptional control. Although little is known regarding the role of transcription factors and coregulators in hypothalamic pathways, considerable progress is being made toward understanding stimulus transcription coupling in clonal cell lines, and emerging evidence suggests new molecular mechanisms for the integration of diverse stimuli at the level of individual genes. Just as diverse neural pathways can converge onto single neurons, distinct signal transduction cascades transmitting a variety of stimuli can converge at the molecular level. One of the most dramatic examples of this type of convergence leads to the activation of the cAMP response element binding protein (CREB), which is a stimulus-induced transcription factor that plays an important role in transynaptic regulation of gene expression (Montminy, 1997). CREB is a member of the bZip superfamily of proteins and is a target of multiple signaling pathways activated by a diverse array of stimuli through phosphorylation at a specific serine residue (see Shaywitz and Greenberg, 1999, for review). In addition, CREB can integrate signaling events by

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forming heterodimers with other bZip factors, such as ATF-1 and CREM, or through protein–protein interactions with CREB binding protein (see below). Cross talk between signal transduction pathways that link extracellular signals with the nucleus represents another important mechanism for integration of homeostatic processes. For example, CREB phosphorylation is induced by protein kinase A in response to elevated levels of cAMP caused by G protein-coupled receptor activation. CREB phosphorylation is also induced by calcium acting through multiple calcium/calmodulindependent kinases, thereby providing a means of linking membrane depolarization with CREB activation. CREB also plays an important role in mediating growth factor induction of transcription by serving as a target for members of the Ras–MAP kinase cascade or through cooperation with other growth factorinduced transcription factors such as those that act at serum response elements. An additional level of communication between distinct signaling pathways is suggested by the observation that estrogen can induce phosphorylation of MAP kinases and extracellularsignal-regulated kinases in vitro and in vivo (Singer et al., 1999; Singh et al., 1999; Toran-Allerand et al., 1999). Thus, it is likely that in hypothalamic cells the nucleus is linked to afferent signals conveyed by hormones, growth factors, and neurotransmitters through multiple interacting signal transduction pathways such as those that lead to CREB phosphorylation. Of particular importance for the control of gene expression in the hypothalamus is the role of steroid and thyroid hormones. Because the hypothalamus regulates secretion of the tropic hormones that control the activity of the peripheral endocrine organs, steroid and thyroid hormones secreted by the adrenal gland, gonad, or thyroid gland exert some of their strongest central effects on hypothalamic circuits. Steroid and thyroid hormone receptors bind to hormone response elements, which function like transcriptional enhancers. But, unlike other enhancers, their activity depends on the presence or absence of ligand. For instance, when estrogen and thyroid hormone receptors bind their cognate hormones the hormone–receptor complex acts as a trans-acting transcription factor that binds to cisacting enhancer-like hormone response elements located within or near responsive genes to influence promoter activity. Because the capacity of these ligand-activated hormone receptors to recognize and regulate target genes is often determined by their ability to bind hormone response elements of hormone-sensitive genes, similarities in the structure of the DNA binding domains of related receptors implies that such receptors may activate overlapping sets of genes by acting through common response elements. For instance, the thyroid

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hormone receptor can bind to the estrogen response element with high affinity in vitro, but fails to activate transcription from these same elements in vivo (Umesono et al., 1988). Moreover, thyroid hormone receptors are capable of inhibiting the ability of estrogen to activate transcription (Glass et al., 1988). In vivo data supporting similar receptor interactions have been reported in the hypothalamus with clear implications for the molecular control of behavior (Pfaff et al., 2000). For example, pretreatment of female rats with thyroid hormone inhibits the actions of estrogen on proenkephalin gene expression (Zhu et al., 2001), and receptor-specific activities of thyroid hormone receptor subtypes influence copulatory behavior (Dellovade et al., 2000). On the other hand, two distinct adrenal steroid hormone receptors, the glucocorticoid receptor and the mineralocorticoid receptor, bind corticosterone with different affinities and can activate gene expression from common promoters (Arriza et al., 1988; Evans and Arriza, 1989; Reichardt and Schutz, 1998). Estrogen can also affect gene expression by binding to different forms of the estrogen receptor, which are coexpressed in several hypothalamic sites, but differ somewhat in their ability to regulate hormoneresponsive genes (Paech et al., 1997; Shughrue et al., 1997; Pettersson and Gustafsson, 2001). Thus, the codistribution of neurons that express related receptors with such interactive properties suggests that coexpression of these trans-acting factors by individual neurons provides considerable combinatorial potential for regulation of gene expression by different endocrine systems. Despite documented effects of steroid hormones on gene expression, in many cases the hormone-responsive genes lack consensus hormone response elements, suggesting that observed regulatory patterns are not due to direct action of the hormone-bound receptors on transcription of such genes. The lack of distinct hormone response elements in hormone-sensitive neurotransmitter genes may relate to the diversity and cellspecific regulation of interactions between steroid hormone receptors and other nuclear trans-acting factors, neurotransmitter receptors, and second messengers that respond to a variety of hormonal and neural cues. For example, estrogen and progesterone have been shown to induce expression of Fos immunoreactivity in the hypothalamus (Hoffman et al., 1990; Insel, 1990; Le et al., 1999), and together with the coordinately expressed protooncogene c-jun, are thought to be representative of general transcription factors (designated Fos or Jun) that are induced by environmental signals and which bind to DNA at regulatory sites termed activator-protein-1 (AP-1) regulatory elements (Morgan and Curran, 1991). CREB may also mediate hormonal regulation of gene expression since phosphorylation of

CREB can be regulated by estrogen (Gu et al., 1996; Zhou et al., 1996), progesterone (Gu et al., 1996), and glucocorticoids (Kovács and Sawchenko, 1996). Thus, the induction of trans-acting factors such as Fos or CREB may mediate the induction of hormone-sensitive genes that lack conventional hormone response elements. The action of hormones like estrogen may converge with transduction pathways that act at AP-1 sites through molecular interactions between estrogen receptors and Fos or Jun (Kushner et al., 2000; Pettersson and Gustafsson, 2001). More central to the action of steroid hormone receptors are coregulators that function as coactivators to increase transactivation by steroid hormone receptors or as corepressors to lower transcriptional activity of target genes (Shibata et al., 1997; McKenna et al., 1999). Cell-type-specific patterns of expression of these coregulators allow different populations of hypothalamic cells to integrate the activity of hormone-activated gene networks with a diverse array of environmental signals. Interaction between families of transcription factors that coordinately regulate gene expression is perhaps best illustrated by the CREB binding protein (CBP) and its close relative p300 (Goodman and Smolik, 2000). These important proteins have binding sites for a wide variety of transcription factors, including not only CREB, but also Fos and Jun (Vo and Goodman, 2001). The observation that CBP can also bind estrogen and thyroid hormone receptors indicates that these protein–protein interactions represent a powerful means of integrating diverse molecular signals at the transcriptional level. Although these mechanisms of molecular integration are only now beginning to be clearly defined, the evidence to date indicates that the activity of a given gene is controlled by changes in the relative ratios of interacting transcription factors expressed within a cell, each of which may respond to distinct cellular signaling pathways in a cell-type-specific manner according to the activity of coexpressed factors. Thus, neurons within the hypothalamus of the rat (and likely most other mammals) possess the potential to receive convergent sensory information from the environment and to integrate this information with neural and humoral signals related to the internal state of the animal. This process is mediated through a variety of complex neurological, cellular, and molecular mechanisms that collectively provide the required breadth and accuracy of adaptive responses necessary for the survival and propagation of the species.

Acknowledgments I thank Drs. Larry Swanson and Paul Sawchenko for providing illustrations and Mara Nelson for preparing the manuscript. Work in

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the author’s laboratory is supported by NIH Grants NS37952, DK55819, and RR00163.

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15 Hypothalamic Supraoptic and Paraventricular Nuclei WILLIAM E. ARMSTRONG Department of Anatomy and Neurobiology,University of Tennessee College of Medicine,Memphis, Tennessee, USA

Magnocellular Neurosecretory System

nucleus (responsible for most of MEE dopamine), a principal source of these neurons lies in the medial aspects of the Pa. Additional parvocellular neurosecretory neurons are found in the hypothalamic and preoptic periventricular stratum (containing dopamine and somatostatin), medial septal nucleus, and nucleus of the diagonal band of Broca (containing luteinizing hormone-releasing hormone). Parvocellular neurosecretory neurons influence the secretion of anterior lobe hormones (e.g., growth hormone, adrenocorticopin hormone, or ACTH, luteinizing hormone, thyroidstimulating hormone) via the release of stimulatory and inhibitory factors from axonal terminals in the MEE. Among these are parvocellular AVP and OX neurons found in the medial portions of the Pa (Vandesande et al., 1977).

Neurons in the supraoptic (SO) and paraventricular nuclei (Pa) projecting to the neural lobe of the hypophysis are large relative to nearby hypothalamic neurons and are traditionally referred to as magnocellular neurosecretory cells. These magnocellular neurons synthesize the neurohormones arginine vasopressin (AVP) and oxytocin (OX) and transport these peptides (l1000 MW) to axonal swellings near fenestrated capillaries in the neural lobe, thus forming the hypothalamoneurohypophysial system (Bargmann and Scharrer, 1951) (Figs. 1, 3, and 4). This system constitutes the final common path for the action potential-dependent release of these hormones to the bloodstream in response to various stimuli ranging from lactation, labor, and nausea/gastric distension (selective for OX) to perturbations in water balance and blood pressure (both AVP and OX) (Renaud and Bourque, 1991; Armstrong, 1995, Poulain and Wakerley, 1982).

PITUITARY GLAND The pituitary gland lies beneath the brain at the mesodiencephalic junction (Atlas Figs. 79–83) and is formed by two distinct parts: a neural part, the neurohypophysis, and a glandular component deriving from oral epithelium called the adenohypophysis (Fig. 1). The neurohypophysis derives from diencephalic tissue and consists of the neural, or posterior lobe (pars nervosa), the pituitary stalk, and the median eminence. The neural stalk (or infundibular stem) is the distal continuation of axons in the internal layer of the median eminence (MEI) (Atlas Fig. 79), which originate primarily from AVP- and OX-secreting magnocellular neurons of

Parvocellular Neurosecretory System Neurons from several hypothalamic nuclei form the parvocellular neurosecretory system (Fig. 1), often called the tuberohypophysial system because of the prominent involvement of the arcuate nucleus and its location in the floor of the hypothalamus (tuber cinereum). In contrast to the magnocellular system, axons of parvocellular neurosecretory neurons project to the external layer of the median eminence (MEE) (Lechan et al., 1982; Wiegand and Price, 1980). In addition to the arcuate

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FIGURE 1 A schematic, midsaggital view of the pituitary gland and its major relationships with neuroendocrine hypothalamus. Magnocellular neurons of the paraventricular (PaMC), supraoptic (SO), and accessory neurosecretory (Acc) nuclei send axons through the internal layer of the median eminence (MEI) and infundibular (neural) stalk (InfS) to terminate in the posterior (neural) lobe of the pituitary (PPit) and release oxytocin (OX) and vasopressin (AVP) into the general circulation. Parvocellular neurons of the paraventricular (PaPC), periventricular (Pe), arcuate (Arc), medial septum (MS), and diagonal band of Broca (DB) project axons to the portal capillary plexus of the external layer of the median eminence (MEE). From here, hypothalamic releasing and inhibiting factors are carried into the anterior lobe (APit) via portal veins to modulate release of the APit hormones from secretory cells. Additional innervations of the MEE, PPit, and intermediate lobe (IPit) are not shown. In addition, a few neurons may innervate both the MEE and the PPit.

the SO and Pa. Neural lobe morphology is characterized by the terminal arborizations of these axons, a rich, fenestrated vasculature, and glial cells (pituicytes), the latter of which form intimate and dynamic morphological relationships with the terminals (Hatton, 1990). Additional inputs to the neural lobe arise from brain stem catecholaminergic neurons in the nucleus of the solitary tract (Sol) (Garten et al., 1989) and rostral periventricular dopamine neurons (Kawano and Daikoku, 1987). Finally, small numbers of neural lobe axons have been found to contain a great variety of neuroactive peptides, GABA, and serotonin, among other substances, which may modulate hormone release at the terminal level (Falke, 1991). The adenohypophysis consists of the anterior lobe (pars distalis), the intermediate lobe (pars intermedia), and the pars tuberalis. The anterior lobe is the largest part and in the rat is not strictly located anterior to the neural lobe, but rather surrounds all but its dorsal aspect. The anterior lobe also contains fenestrated vessels, is composed principally of various hormonesecreting cells, and connects to the MEE by small portal vessels running along the neural stalk. These vessels deliver the releasing and inhibiting factors to the anterior lobe upon their secretion from hypothalamic axons into the fenestrated capillary plexus of the MEE, which feeds into portal veins. The intermediate lobe is unique in containing a large population of gland cells derived from neural crest which synthesize members of the proopiomelanocortin family of peptides, including β-endorphin and α-melanocyte-stimulating hormone (Mains and Eipper, 1979). These cells receive a substantial neural innervation from dopamine (Björklund et al., 1973), serotonin (Mezey et al., 1984), and GABAcontaining axons (Tappaz et al., 1983), including direct

synapses onto the gland cells (see Saland, 2001, for review). The pars tuberalis is relatively small in rat, forming a rostral extension of anterior lobe cells along the neural stalk.

SUPRAOPTIC NUCLEUS Cytoarchitecture and Efferent Projections Distribution of Principal (Projection) Neurons The largest component of the SO is the anterior, or principal, division. This division begins rostrally as scattered, large and darkly staining neurons lateral to the optic chiasm (Atlas Fig. 21) and condenses caudally to abut the lateral border of the optic tract (Figs. 2 and 3; Atlas Figs. 22–27). Posteriorly, a retrochiasmatic continuation (SOR), separated from the SO by the optic tract, is a thin sheet of neurons lying along the pial surface between the optic tract and the third ventricle (Fig. 3; Atlas Figs. 28–32). Both the SO and the SOR project heavily and primarily to the neural lobe (Sherlock et al., 1975). Most SO neurons produce either OX or AVP and project to the neural lobe. Although AVP neurons slightly outnumber OX neurons, the large size of the SO relative to the Pa makes it quantitatively the most important site for production of both hormones. While, in general, neurons producing OX lie more anterodorsally and AVP neurons posteroventrally in the nucleus, there is considerable overlap of the two types within the SO (Swaab et al., 1975; Vandesande and Dierickx, 1975; Hou You et al., 1986). In the rat, the ratio of AVP to OX neurons in the SOR is greater than that in the principal SO (Rhodes et al., 1981).

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FIGURE 2 The anterior, or principal, supraoptic nucleus (SO) in coronal section. (A) Nissl-stained, 5-μm paraffin section illustrates the magnocellular neurons of the SO situated lateral to the junction of the optic chiasm (ox) with the optic tract (opt). The perinuclear zone (PZ) is a cell-poor region surrounding the SO whose neurons may relay inputs from limbic and other structures to magnocellular neurons. The approximate region of the PZ is indicated with small arrowheads, but the boundaries of this region are currently undefined. Note another cellpoor zone at the base of the SO, abutting the pial surface (large arrowheads). This region contains glial cells and, as shown in panel (B), the dendrites of SO neurons. (B) Immunocytochemical localization of AVP in a coronal vibratome (i.e., unembedded) section (30 μm) through the SO. The section has been counterstained for Nissl substance. The AVP-reactive neurons are concentrated ventrally. Note the extent of the ventral dendritic lamina (large arrowheads). The axons of SO neurons are thin and beaded and emit dorsally. One such process is seen arising from a dendrite to join the dorsally emerging supraopticoneurohypophysial tract (small arrowheads). AVP antibody courtesy of Dr. G. Nilaver.

Neurons in the SO producing OX and AVP may colocalize a variety of other peptides and potential neuroactive substances release (Meister, 1993), some of which act within the neural lobe (Falke, 1991) or locally within the SO (Brown et al., 1998) to modulate neurohypophysial hormone release. These include galanin and dynorphin, which primarily coexist with AVP, cholecystokinin (CCK), and corticotropin-releasing hormone (CRH), which primarily coexist with OX neurons and nitric oxide synthase (Nylen et al., 2001; Yamada et al., 1996) and cocaine–amphetamine-regulated transcript (CART) (Vrang et al., 1999), which are found in both OX and VP cells. A caveat is that OX and AVP coexist in 15–20% of magnocellular neurons in the lactating female rat (Mezey and Kiss, 1991), suggesting that the distribution of OX and AVP may be state-dependent. Indeed, single cell analysis of amplified mRNA suggests a potential for OX and AVP coexpression in most SO neurons (Xi et al., 1999). The same may be true of other coexpressed peptides. For example, neuropeptide Y (NPY) expression is revealed in the magnocellular neurons only after osmotic stimulation (Larsen et al., 1992). Morphology and Efferent Path of Magnocellular Supraoptic Neurons Supraoptic neurons exhibit a relatively simple cytoarchitecture (Armstrong, 1995). Neurons in both the SO and the SOR have round- to egg-shaped somata

(20–35 μm) and possess two to three primary dendrites (range, 1–5) that typically project ventrally and branch sparsely (Figs. 2B and 4A). The dendrites collect in bundles in a lamina between the cell bodies and the pial surface to form a receptive plexus for incoming axons (Sofroniew and Glasmann, 1981; Armstrong et al., 1982b) (Fig. 2B). The somata have a rough appearance caused by spines and irregularly shaped appendages (Randle et al., 1986; Smith and Armstrong, 1990; Stern and Armstrong, 1998). The dendrites are not heavily spinous, but do exhibit irregularly shaped appendages and are often varicose. Dendritic varicosities sometimes contain the dense core granules housing OX or AVP. The evidence for dendritic release of OX, AVP, and dynorphin, and the autoregulatory effects of these peptides, is substantial (Ludwig, 1998). The axons of SO neurons arise from a primary dendrite or directly from the soma and course dorsally before turning medially in a wide arc (tract of Greving) to project posteriorly to reach the MEI (Randle et al., 1986; Smith and Armstrong, 1990). Posteriorly, the axons become more varicose and within the neural lobe are punctuated by numerous small and occasionally large (10–15 μm) swellings traditionally known from neurosecretory stains as Herring bodies (Figs. 4B–4D). The apposition of these swellings to the basal lamina near fenestrated vessels forms the neurohemal contact zone (Fig. 4D). Within the neural lobe, OX fibers are preferentially distributed peripherally while AVP fibers are

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FIGURE 3 The distribution of magnocellular neurosecretory neurons is shown with the major divisions of the supraoptic (SO), paraventricular (Pa), and accessory nuclei (Acc) and their efferent projections. On the left are tracings from coronal sections of the distribution of magnocellular neurons containing either vasopressin (AVP) or oxytocin (OX) revealed with immunocytochemistry using an antibody crossreacting with both AVP- and OX-associated neurophysins (anti-rat neurophysin, courtesy of Dr. A. G. Robinson). The reactive cells are plotted in the four representative sections and shown in relation to the subdivisions of the Pa, SO, and Acc. Solid areas are virtually filled with reactive neurons. On the right, groups of neurons are marked to show the relative density of OX and AVP types according to the literature. Horizontal lines indicate a majority of OX neurons, vertical lines a majority of AVP neurons, and hatched areas a more equal mixing of the two. The great majority of Acc neurons in panels (A) and (B) are OX-containing, while further caudally (C and D) the Acc groups are more mixed. Also on the right, some major efferent connections of the nuclei and their subdivisions are shown, with distal targets enclosed by dashed lines. In the Pa, it should be pointed out that most of the neurons in the dorsal medial cap (PaDC), medial parvocellular (PaMP), ventral (PaV), and posterior (PaPo) groups project to the targets indicated, but not all of the OX and AVP neurons shown in each division contribute to the projection. In addition, many neurons in these subdivisions do not contain OX or AVP. AC, anterior commissural nucleus; Bs, brain stem; BSTV, bed nucleus of the stria terminalis; Cir, nucleus circularis; MEE, external layer of the median eminence; f, fornix; ic, internal capsule; LH, lateral hypothalamus; LPO, lateral preoptic area; MFB, nuclei of the medial forebrain bundle; opt, optic tract; ox, optic chiasm; PaLM, paraventricular nucleus, lateral magnocellular division; PaMM, paraventricular nucleus, medial magnocellular division; PeM, magnocellular periventricular nucleus; PPit, posterior pituitary; PoF, posterior fornical nucleus; sm, stria medullaris; SpC, spinal cord; 3V, third ventricle.

localized centrally (van Leeuwen et al., 1979; Tian et al., 1991). When viewing axons traced to the MEI from intracellularly labeled supraoptic neurons, short (~10 μm), filiform processes are occasionally seen, but not collateral arbors (Randle et al., 1986; Smith and Armstrong, 1990). However, branches within the lateral hypothalamus from SO axons have been observed with other techniques and may account for some of the collaterals seen in AVP-positive axons in this region (Mason et al., 1984; Ray and Choudhury, 1990). Immunochemically, synapses positive for AVP (Choudhury and Ray, 1990)

and OX (Theodosis, 1985) have been observed in the SO, but it remains uncertain whether these emanate from magnocellular axons. Supraoptic Interneurons A very small number of multipolar neurons with small somata are found within the SO and could function as interneurons (Dyball and Kemplay, 1982; Bruni and Perumal, 1984). An additional locus of interneurons is the perinuclear zone (PZ) immediately adjacent to the SO (Figs. 2A and 5). Many neurons in the PZ

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FIGURE 5 Photomontage of a PZ neuron filled with Neurobiotin and reacted with avidin–biotin peroxidase complex. A short axon from this neuron displays terminal-like varicosities in close proximity to SO neurons stained with thionine. Modified from Armstrong and Stern (1997).

FIGURE 4 Neurosecretory neuron and axons. (A) Photomontage of an SO neuron filled with Neurobiotin and reacted with avidin– biotin peroxidase complex (see Stern and Armstrong, 1998, for method). The axon originates from the soma and is indicated by an arrowhead. Arrows point to dendritic branches. The montage was created from digital micrographs through several focal planes in a 100-μm vibratome section. (B, C). Neural lobe axons labeled anterogradely from a pressure injection of biotinylated dextran in the SON, reacted with avidin–biotin peroxidase complex. (B) Axon with several small branches in a small area (arrowheads). (C) Axons with swellings present the classical image of strings of pearls. Note the thin intervaricose segments (arrows) and the large range of swelling sizes. (D) Electron micrograph of the neurohemal contact zone in the neural lobe. Note the large neurosecretory terminal (T) filled with dense core granules and surrounded by pituicyte (Pit) processes. The terminal abuts the basal lamina of the perivascular space (arrow) of a fenestrated capillary (Cap). Modified from Tian et al. (1991).

project to the SO (Iijima and Ogawa, 1981; Tribollet et al., 1985; Jhamandas et al., 1989; Roland and Sawchenko, 1993; Levine et al., 1994), and this zone is a target of many inputs known to influence the firing rate of OX and VP neurons. Thellier et al. (1994) place the densest population of locally projecting PZ neurons to a medial

region dorsal to the optic chiasm rather than immediately dorsal to the SO. PZ neurons are morphologically diverse (Armstrong and Stern, 1997) and can express many neuroactive substances including GAD or GABA (Tappaz et al., 1983; Theodosis et al., 1986; Okamura et al., 1990), choline acetyltransferase (Mason et al., 1983), substance P (Larsen, 1992), somatostatin (Mezey et al., 1991), estrogen receptor (Herbison et al., 1994), pituitary adenylate cyclase-activating polypeptide (PACAP) (Hannibal et al., 1995) and probably glutamate (Boudaba et al., 1997). All of these substances have been identified in fibers and some even in synaptic terminals within the SO. Much of the somatostatinergic input to the SO derives from the PZ (Mezey et al., 1991). PZ neurons are electrophysiologically distinct from SO neurons (Armstrong and Stern, 1997) and contribute functional inhibitory (Wuarin, 1997) and excitatory synapses (Boudaba et al., 1997) to the nucleus. The GABAergic neurons are thought to be important in mediating the response of AVP neurons to hypertension (Nissen et al., 1993; Grindstaff and Cunningham, 2001).

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Afferent Inputs The first chemically identified input to the SO was that from brain stem catecholamine neurons (Carlsson et al., 1962). This projection is primarily noradrenergic and originates from the A1 (the ventrolateral medulla) and A2 (caudal part of the Sol) cell groups (Sawchenko and Swanson, 1982a; Tribollet et al., 1985; Cunningham and Sawchenko, 1988; Day and Sibbald, 1988; Raby and Renaud, 1989). Cardiovascular and other visceral afferent (e.g., gut distension) signals controlling AVP and OX release could be routed through these brain stem nuclei (Renaud and Bourque, 1991). The projection from A1 is thought to favor AVP neurons and contacts both the SO (primarily the ventral region) and the SOR. While involved in the response of AVP to cardiovascular stimuli (both hypo- and hypertension), the precise action of noradrenaline in these responses is debatable. Noradrenergic neurons in A1 may colocalize NPY (Sawchenko et al., 1985), prolactinreleasing peptide or glucagon-like peptide-1 (Morales et al., 2000), PACAP (Shioda et al., 2000), and substance P (Bittencourt et al., 1991). Importantly, the corelease of ATP with noradrenaline could mediate some excitatory AVP cell responses to cardiovascular challenges (Buller et al., 1996). Some of the colocalized peptides can modulate noradrenaline’s actions on AVP release and are potent agents by themselves (Renaud and Bourque, 1991; Kapoor and Sladek, 2001; Shibuya et al., 2000). The A2 projection to the SO prefers more dorsally located OX neurons and is involved in OX release to gastric stimulation (Onaka et al., 1995). Some noncatecholaminergic neurons in the Sol contain the peptides activin/inhibin-β and somatostatin-28 and directly contact OX neurons (Sawchenko et al., 1990; Sawchenko and Pfeiffer, 1995). Smaller brain stem inputs to the SO have been summarized elsewhere (Hatton, 1990; Morris et al., 1987) and include the serotonergic innervation from raphe nuclei that projects to the dorsal (primarily OX-containing) SO. Brain stem afferents from the lateral parabrachial nucleus innervate the PZ and affect OX and AVP neurons through this relay (Jhamandas et al., 1991). The subfornical organ (SFO), the median preoptic nucleus (MnPO), and the organum vasculosum of the lamina terminalis (OVLT) (Camacho and Phillips, 1981; Sawchenko and Swanson, 1983; Tribollet et al., 1985) provide dense monosynaptic inputs important for the regulation of water balance and AVP secretion, although OX neurons may be contacted as well. These regions contain neurons sensing plasma osmolality (MnPO and OVLT) or peripheral AII (SFO) and provide powerful inputs to the SO (e.g., Richard and Bourque, 1995). Some of the SFO projection may utilize AII as an excitatory

neurotransmitter (Jhamandas et al., 1989). Afferents from the MnPO/OVLT are distributed uniformly within the SO and make largely symmetrical synapses with both dendrites and somata (Armstrong et al., 1996). However, both inhibitory and excitatory projections have been identified (Yang et al., 1994), and the osmosensing component of the OVLT projection to SO is mediated by glutamate (Richard and Bourque, 1995). Direct inputs to the ventral dendritic lamina of the SO from the olfactory bulb may be involved in mediating olfactory cues for the synaptic and dendritic reorganization of the SO occurring during pregnancy and lactation (Hatton, 1990; Meddle et al., 2000). A direct retinal input to the SO has been reported (Levine et al., 1994). Additional retinal information may pass to the SO through the PZ (Levine et al., 1994; Cui et al., 1997), which receives input from the suprachiasmatic nucleus (Stephan et al., 1981) and thus information regarding circadian rhythms and their entrainment to the diurnal cycle. Additional forebrain and hypothalamic inputs come from a wide variety of structures, including the bed nucleus of the stria terminalis, dorsomedial nucleus, anteroventral periventricular nucleus, medial preoptic area, rostral periventricular area, and tuberomammillary region (Hatton, 1990; Morris et al., 1987; Sawchenko and Swanson, 1983; Simerly and Swanson, 1988; also see Chapter 14 by Simerly, this volume). The diagonal band of Broca (Jhamandas et al., 1989) and lateral septum (Poulain et al., 1980) have a powerful, inhibitory influence on SO neurons, yet these regions send only sparse monosynaptic inputs. Instead, they more heavily contact the PZ (Jhamandas et al., 1989; Oldfield et al., 1985). Although a variety of neurotransmitters and neuromodulators have been located within the SO and SOR and are undoubtedly represented by the inputs discussed above, quantitatively the most important transmitter in the SO is GABA, which accounts for ~50% of all synapses in the SO (Theodosis et al., 1986; Decavel and van den Pol, 1990). Traditionally, GABA is considered to be of local origin, but recent studies indicate only sparse GABAergic inputs may arise from the area immediately adjacent to the SO (Roland and Sawchenko, 1993). The forebrain and hypothalamic distribution of neurons containing mRNA for GAD (Okamura et al., 1990; Cullinan et al., 1993; Roland and Sawchenko, 1993) suggests that other regions known to project to the SO also contain GABAergic neurons, including the periventricular preoptic area, bed nucleus of the stria terminalis, the tuberomammillary complex, the OVLT, the SFO, and the MnPO. The majority of the remaining synapses probably utilize the excitatory amino acid transmitter glutamate (Meeker et al., 1989; van den Pol, 1991; DeCavel and van den Pol, 1992). Glutamatergic

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inputs to the SON have been shown to derive from a number of nuclei, including, among others, the PZ, the dorsomedial nucleus, and the bed nucleus of the stria terminalis (Csaki et al., 2002). Following GABA and glutamate, noradrenaline accounts for ~10% of all SO synapses (Michaloudi et al., 1997). Histamine-containing neurons comprise much of the tuberomammillary input to the SO and SOR (Hatton, 1990). Dopamine has been localized in axons within the SO in the region of OX neurons, but its precise source is not clearly identified (Buijs et al., 1984; DeCavel et al., 1987).

PARAVENTRICULAR NUCLEUS The Pa is cytoarchitectonically more complex than the SO (Armstrong et al., 1980; Swanson and Kuypers, 1980; Swanson et al., 1986; Kiss et al., 1991). Unlike the SO, the Pa contains medium- and small-sized neurons that project to other CNS areas or to the MEE in addition to magnocellular OX and AVP neurons projecting to the neural lobe. The basis for Pa subdivisions is primarily the segregation of these projection neurons into groups with common outputs, but differences often are correlated with differences in biochemical types

TABLE 1

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and afferent inputs (Swanson et al., 1986), cell size and packing density (Kiss et al., 1991), and dendritic morphology (Armstrong et al., 1980; van den Pol, 1982; Ju et al., 1986, 1992; Rho and Swanson, 1989). Table 1 lists the cytoarchitectonic divisions used herein and their relationship to those of other schemes.

Magnocellular Neurosecretory Component Distribution of Principal Neurons The majority of Pa neurons sending axons to the neural lobe of the pituitary lie in two contiguous groups (Hatton et al., 1976): the medial magnocellular portion (PaMM) contains mostly OX neurons and lies anteromedially in the nucleus; the lateral magnocellular division (PaLM) contains primarily AVP neurons and forms a distinct, ball-shaped mass posterior and dorsolateral to the PaMM (Figs. 3 and 6; Atlas Figs. 26 and 27). A ring of OX neurons also lies around the densely packed PaLM. Like SO neurons, these cells are relatively large (20–35 μm). Less densely packed neurons containing OX and AVP, some of which project to the neural lobe, are found posterior in the Pa, medial and anterior to these divisions. Most of the substances found to colocalize with SO neurons can be found in Pa neurons projecting

Nomenclature for Paraventricular and Associated Nuclei

Terminology used herein

Other names

Magnocellular groups Medial (PaMM) (Armstrong et al., 1980; Hatton et al., 1976; Swanson et al., 1986)

Posterior magnocellular (Swanson and Kuypers, 1980) and magnocellular (Kiss et al., 1991)

Lateral (PaLM) (Armstrong et al., 1980; Hatton et al., 1976) Periventricular (PeM) (Armstrong et al., 1982a) Anterior commissural (AC) (Armstrong et al., 1980, 1982a; Kiss et al., 1991)a

Anterior magnocellular (Swanson and Kuypers, 1980; Swanson et al., 1986)

Median eminence projecting groups Anterior parvovellular (PaAP) (Swanson and Kuypers, 1980; Swanson et al., 1986) Medial parvocellular (PaMP) (Armstrong et al., 1980; Kiss et al., 1991; Swanson and Kuypers, 1980; Swanson et al., 1986)b Periventricular parvocellular (PeP) (Kiss et al., 1991; Swanson and Kuypers, 1980; Swanson et al., 1986) Extrahypothalamically projecting groups Ventral (PaV) (Swanson et al., 1986)c

Posterior (Armstrong et al., 1980, 1982a; Kiss et al., 1991)

Posterior (PaPo)

Lateral parvocellular (Swanson and Kuypers, 1980; Swanson et al., 1986)

Dorsomedial cap (PaDC) (Armstrong et al., 1980)

Dorsal parvocellular (Swanson and Kuypers, 1980; Swanson et al., 1986), dorsal (Kiss et al., 1991)

a

Referred to as Peterson’s AC. Kiss et al. (1991) divide the PaMP into anterior, medial, lateral, and caudal parts which encompass the PaAP, PaMP, and PeP as used herein. c Referred to as Koh and Ricardo’s ventral Pa. b

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FIGURE 6 Some divisions of the paraventricular nucleus (Pa) are shown in horizontal sections (30 μm) after reacting with an antibody to neurophysin that cross-reacts with both OX- and AVP-associated neurophysins (courtesy of Dr. A. G. Robinson). The most dorsal section is in panel (A); the most ventral in panel (D). These sections also illustrate the general orientation of dendrites (small arrowheads) from these neurons. (A) The anterior commissural (AC) and periventricular magnocellular (PeM) divisions, which contain almost exclusive populations of OX neurons in the dorsal, anterior hypothalamus are shown. Note the strong medial and posteromedial orientation of dendrites. (B) More ventrally, the top of the medial aspect of the posterior division of the Pa (PaPo) is visible. Some dendrites of both PaPo and AC neurons are indicated. Note some of the PaPo dendrites emit laterally, and some appear to cross the midline. (C) Further ventral, the lateral magnocellular Pa (PaLM) is visible as a dense ball of cells. The PaLM is mostly AVP-containing, and its dendrites are directed medially into the medial parvocellular zone (PaMP) and periventricular parvocellular (PeP) zones. This section is fully through the PaPo, which has many neurophysinpositive neurons whose morphology is quite different from those of the PaLM. Again, note the diverse orientation of PaPo dendrites. (D) In the most ventral section, the medial magnocellular Pa (PaMM) is visible. This group is mostly composed of OX neurons. Immediately caudal it is replaced by the ventral Pa (PaV), which is relatively poor in neurophysin-positive neurons.

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to the neural lobe (see above). Two OX-containing cell groups often are included in the Pa (Swanson and Kuypers, 1980): the periventricular magnocellular group (PeM) and the anterior commissural nucleus (AC). Both the PeM and the AC are considered herein as accessory neurosecretory nuclei and are dealt with in the final section of this chapter (also see Figs. 3A and 6; Table 1). A more detailed map of OX and AVP neurons in various regions of the Pa is available in HouYou et al. (1986). Morphology and Efferent Path of Magnocellular Paraventricular Neurons Magnocellular Pa neurons in the PaMM and PaML of the rat have an appearance similar to that of their SO counterparts, with the typical cell having two or three dendrites that branch relatively simply (Armstrong et al., 1980; van den Pol, 1982; Hatton et al., 1985; Rho and Swanson, 1989). These initially thick dendrites taper and invade the medial and ventromedial parts of the nucleus and form a dense plexus, which occasionally encroaches upon the third ventricular wall (Fig. 6) (Sofroniew and Glasmann, 1981; Armstrong et al., 1982a; Ju et al., 1986, 1992). While in lower vertebrates subependymal Pa dendrites can form ciliated, sensory type endings and contact the cerebrospinal fluid (Vigh and Vigh-Teichmann, 1998), such contact has not been verified for Pa dendrites in mammals. The medial parvocellular region is rich with axonal inputs (van den Pol, 1982), including a prominent catecholaminergic component (McNeill and Sladek, 1980). Thus, synaptic contacts upon OX and AVP neurons innervating the neural lobe can occur outside the confines of the subnucleus (see below). Like their SO counterparts, the dendrites of PaMM and PaML neurons are variably spinous, but can be quite varicose (Sofroniew and Glasmann, 1981; van den Pol, 1982). Many of the appendages are longer filiform processes or are irregularly shaped (van den Pol, 1982; Rho and Swanson, 1989). Like those in the SO, the dendrites of Pa neurons are capable of local secretion of peptide (Ludwig, 1998). Axons of the PaMM and PaLM egress laterally from either the soma or a primary dendrite (Armstrong et al., 1980; van den Pol, 1982; Hatton et al., 1985; Rho and Swanson, 1989) and course in a wide arc, passing over or beneath the fornix before turning medially above the SO to join the tract of Greving before reaching the MEI and the neural lobe. In the neural lobe, Pa axons are thought to distribute peripherally to those arising from the SO (Alonso and Assenmacher, 1981). Pa axons from magnocellular neurons may occasionally branch (van den Pol, 1982; Hatton et al., 1985; Ray and Choudhury, 1990), but the final destination and synaptic relationship of collaterals remain to be determined.

377

Parvocellular Neurosecretory Component Distribution of Principal Neurons Small- to medium-sized neurons in the anterior (PaAP) and medial (PaMP) parvocellular portions of the nucleus (Fig. 3), including the parvocellular periventricular stratum (Swanson and Kuypers, 1980), give rise to a substantial input to the MEE (Armstrong and Hatton, 1980; Wiegand and Price, 1980; Lechan et al., 1982; Kawano and Daikoku, 1988; Kawano et al., 1991). The PaAP and PaMP are chemically diverse and contain neurons immunoreactive (with varying degrees of colocalization) for angiotensin II, atrial natriuretic peptide, bombesin, AVP, CART, CCK, CRH, PACAP, GRF, tyrosine hydroxylase (presumptive dopamine neurons), GABA, enkephalins, galanin, growth hormone-releasing hormone, neurotensin, peptide histidine leucine, somatostatin, thyrotropin-releasing hormone (TRH), and vasoactive intestinal peptide (Swanson et al., 1986; Swanson, 1987; Hökfelt et al., 1990; Hannibal et al., 1995; Broberger, 1999). Many of these cell types project to the MEE or extrahypothalamically (see below). Most of the CRH (Antoni et al., 1983) and TRH (Brownstein et al., 1982; Kawano et al., 1991) present in the MEE derives from the Pa. Approximately 70% of all somatostatin neurons in the rostral periventricular region and PaMP project to the MEE (Kawano and Dikoku, 1988; Merchenthaler et al., 1989). Some somatostatin neurons innervate OT neurons, perhaps by collaterals (Hisano et al., 1993a). Peptidergic neurons are roughly segregated within the PaMP on cytoarchitectonic grounds (Kiss et al., 1991), such that CRH neurons are found lateral to TRH neurons, which are in turn found lateral to periventricular dopamine and somatostatin neurons (Swanson et al., 1986). A projection of AVP axons from the Pa to the MEE has been known since at least 1977 (Vandesande et al., 1977) and is related to ACTH and glucocorticoid secretion (Silverman and Zimmerman, 1982). The AVP derives from parvocellular neurons in the PaMP and PaAP and its expression is normally weak. In colchicine-treated rats, AVP is expressed in over half of the CRH neurons (Whitnall and Gainer, 1988), and after adrenalectomy the majority of parvocellular CRH neurons colocalize AVP (Kiss et al., 1984; Sawchenko et al., 1984). In addition to receiving stress signals through brain stem and limbic pathways (Herman and Cullinan, 1997; see below), these neurons also possess glucocorticoid receptors for direct feedback (Uht et al., 1988). Morphology and Efferent Pathway In Golgi studies, the dendritic architecture of Pa neurons in parvocellular neurosecretory subnuclei appears similar to that of the general population of

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parvocellular Pa neurons. Like magnocellular neurons, parvocellular neurons in the PaMP have a relatively restricted dendritic arbor showing few branches, with a moderate endowment of appendages (van den Pol, 1982). In addition to a strong medial orientation of the dendritic tree, some PaMP neurons have ventrally and dorsally directed dendrites that run along the ventricular wall. In more recent studies, parvocellular neurosecretory neurons have been identified by combining retrograde tracing, intracellular injection, and immunocytochemical identification for CRH (Rho and Swanson, 1987, 1989). These studies have confirmed the relatively simple dendritic tree for CRH neurons and the presence of filiform spines. Like magnocellular neurons, the dendrites of the PaMP seldom leave the Pa, but do cross into adjacent subnuclei. The axons of the parvocellular neurosecretory cells in the Pa follow closely those of the magnocellular group, emitting laterally toward the fornix before turning ventrally and posteriorly en route to the MEE (Swanson et al., 1986; Rho and Swanson, 1989; Larsen et al., 1991). Clear examples of axonal branching have been observed for CRH neurons, with local arborizations within and just outside the Pa (Liposits et al., 1985; Rho and Swanson, 1987). Synaptic contacts positive for CRH (Liposits et al., 1985; Shioda et al., 1985; Silverman et al., 1989) could arise from these collaterals, although other sources of CRH cannot be ruled out. A similar argument and reservation could be made for TRH-positive (Kiss and Hálász, 1990; Hisano et al., 1993b) and somatostatin-positive (Hisano et al., 1993a) synapses in the Pa. Golgi stains of PaMP neurons have revealed locally arborizing axons (van den Pol, 1982), but it is uncertain whether or not these represent neuroendocrine neurons.

Nonendocrine Projection Neurons Distribution of Principal Neurons A third major group of variably sized Pa neurons projects caudally to the brain stem and spinal cord and is distinctly separate from the other two groups (e.g., Hoysoya and Matsushita, 1979; Armstrong et al., 1980; Swanson and Kuypers, 1980). These neurons primarily contact autonomic preganglionic and related nuclei and are treated in detail in Chapter 24 of this volume. The dorsomedial cap (PaDC), the ventral Pa (PaV), and the posterior subnucleus (PaPo) are the primary sources of these projections (Figs. 3 and 6). The PaDC is more involved with the projection to the intermediolateral cell column, and the PaV and PaPo contribute fibers to both the spinal cord and several brain stem regions, most notably the dorsal motor nucleus of the vagus,

other preganglionic autonomic nuclei, the Sol, the periaqueductal gray, the dorsal raphe nuclei, the locus coeruleus, the area postrema, the parabrachial nucleus, the pedunculopontine nucleus, and the ventrolateral reticular nucleus (region of A1 noradrenergic neurons) (Saper et al., 1976; Luiten et al., 1985; Shapiro and Miselis, 1985; Hoysoya et al., 1991; Shafton et al., 1998; Pyner and Coote, 2000). A very small proportion of this descending projection derives from paraventricular AVP neurons and a more substantial, but still small, element from OX neurons (Sofroniew and Schrell, 1982; Sawchenko and Swanson, 1982b; Jansen et al., 1995). Other peptides make contributions to the descending pathway, including bombesin (Costello et al., 1991), CRH (Sawchenko, 1987; Milner et al., 1993; Jansen et al., 1995), and angiotensin II (Jansen et al., 1995). Functionally, descending Pa projections may operate either via these peptides or with classical amino acid transmitters such as glutamate (e.g., Coote et al., 1998). With few exceptions, descending Pa axons do not branch to the neural lobe (Swanson and Kuypers, 1980), but there is overlap between brain stem- and spinal cord-projecting components (Swanson and Kuypers, 1980; Shafton et al., 1998; Pyner and Coote, 2000). Less well examined is the degree to which median eminence-projecting and extrahypothalamic-projecting Pa neurons overlap, but this is predicted to be small based on the topographic segregation of projection neurons in the Pa. In addition to the projection to the median eminence, neurons from the PaMP project to two other circumventricular organs, the SFO and the OVLT, as well as to the pineal gland (Larsen et al., 1991). Additional pathways to di- and telencephalic regions are a dorsal pathway to the region of the thalamic paraventricular nucleus; a ventrolateral projection to the medial and central nucleus of the amygdala; a rostroventral projection to the anteroventral periventricular nucleus (Larsen et al., 1991); and possible intrahypothalamic connections to the anterior hypothalamus, the dorsoand ventromedial nuclei, the arcuate nucleus, and the perifornical regions (Luiten et al., 1985; Ter Horst and Luiten, 1987; Larsen et al., 1991). Morphology and Efferent Pathway The size of PaV, PaDC, and PaPo neurons varies considerably, but more medium- and even large-sized neurons are found than are present in the PaMP and PaAP (Kiss et al., 1991). These neurons have longer dendrites that branch more often but are less spinous than either group of neuroendocrine cells (Rho and Swanson, 1989). Medially oriented dendrites of the PaPo may project anteromedially, posteromedially, and even across the midline (Figs. 6B and 6C), but many have a strong perpendicular orientation with respect to the 3V

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(Fig. 7) (Stern, 2001). In addition, fusiform neurons in the PaPo also have one or more lateral dendrites projecting toward and sometimes over the fornix. These lateral dendrites branch and can give rise to an axon, or one may gradually taper to appear axonal (Armstrong et al., 1980; van den Pol, 1982; Hatton et al., 1985; Shapiro and Miselis, 1985). Neurons in both the PaV and the PaPo with descending projections are heterogeneous in their dendritic morphology and their physiology, suggesting functional subsets within these divisions (Stern, 2001). The axonal projection paths of this group of Pa neurons are diverse (Saper et al., 1976; Luiten et al., 1985; Larsen et al., 1991; Ranson et al.,1998). Many axons in the PaV, PaDC, or PaPo emit from a primary dendrite or soma and project laterally or ventrolaterally (Armstrong et al., 1980; Van den Pol, 1982; Rho and Swanson, 1989; Stern, 2001) (Fig. 7). In some cases, these axons branch and the collateral returns to the Pa (Rho and Swanson,

379

1989). Fibers innervating the dorsal thalamus emit directly dorsally, or course posteriorly along the midline before turning dorsally. Fibers innervating the amygdala take a ventrolateral course beneath the internal capsule or cerebral peduncle. Descending projections have been divided into a midline pathway innervating the periaqueductal gray, raphe, parabrachial nuclei, and locus coeruleus and another pathway which originates from the caudal portion of the Pa and stays medial until the midbrain, where it moves laterally and hugs the medial aspect of the cerebral peduncle (Luiten et al., 1985; Ranson et al., 1998). From the midbrain, this second pathway moves ventrally, and the medial pathway shifts laterally and ventrally. The two paths converge in the ventral medulla and from there send fibers to the area postrema, the Sol, the dorsal vagal complex, and the ventrolateral medulla (A1 region). The main projection to the spinal cord is through the dorsolateral funiculus, with branches innervating the intermediolateral cell column. Rostral projections (Larsen et al., 1991) travel along the ventricle or course anterolaterally to invade the bed nucleus of the stria terminalis and the medial preoptic area. Rostral periventricular fibers project to the OVLT and some turn dorsal to innervate the subfornical organ.

Paraventricular Interneurons

FIGURE 7 Drawing of a PaPo neuron. This neuron was identified as preautonomic following retrograde tracing from the dorsomedial medulla. The retrograde label was observed in a coronal slice in vitro, and the neuron was visually patched for whole cell electrophysiological recording and filled with biocytin. The dendrites had a strong mediolateral orientation, with medial dendritic branches (arrowheads) projecting near the third ventricle (3V). The axon projected ventrolaterally. Inset shows the location of the neuron in the Pa in coronal section. Modified from Stern (2001).

It is not certain whether some Pa cells are true interneurons (i.e., their axon solely innervates within the nucleus), but at the least there are axon collaterals from Pa neurons that make intrinsic connections. Some Golgi-impregnated parvocellular neurons in the PaMP have extensive axon collateral arbors that terminate in both the PaMP and the magnocellular PaLM groups (van den Pol, 1982). Some identified projection neurons, including CRH neurons, have recurrent axon collaterals (Rho and Swanson, 1989) that could be in part responsible for CRH-positive synapses within the periventricular zone (Silverman et al., 1989) and the magnocellular regions (Liposits et al., 1985), onto TRH neurons (Hisano et al., 1993b), and even recurrently onto CRH neurons (Liposits et al., 1985) Extensive intrinsic connections may account for the many axon terminals that are immunoreactive for peptides present in other Pa-projecting cell groups (e.g., TRH) and that are presynaptic to homo- and heterotypic elements (e.g., Kiss and Halasz, 1990; Hisano et al., 1993b), but an extrinsic origin cannot be discounted. In addition to peptides, classical neurotransmitters are likely involved with intrinsic Pa circuits. Like the SO, the Pa receives a dense GABAergic and glutamatergic input, part of which comes from neurons in or near the Pa as indicated by both anatomical (Roland and Sawchenko, 1993)

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and physiological studies (Tasker and Dudek, 1993; Boudaba et al., 1996, 1997).

Afferent Inputs Brain Stem Inputs Consistent with its diverse population of principal neurons, the Pa has a diverse source of inputs (Hatton, 1990; Armstrong, 1995; also see Chapters 14 and 24, by Simerly and by Saper, this volume). Like the SO, the most well described are dense projections from the brain stem nuclei rich in catecholamine neurons (Tribollet and Dreifuss, 1981; Sawchenko and Swanson, 1982a; McKellar and Loewy, 1981). As some of these connections relate to the role of the Pa in autonomic function, they are treated in detail in the chapter by Saper (Chapter 24). Different catecholamine groups have relatively different but overlapping targets within the Pa. The medullary A1 projection favors the AVP cells of the PaLM (but also innervates the PaMP), the A2 group favors the PaMP (but does not exclude the PaLM and PaMM), and the locus coeruleus (A6) projects to the more medial periventricular zone (Swanson et al., 1986; Cunningham and Sawchenko, 1988). While the majority of these fibers contain noradrenaline, with some colocalization in A1 with neuropeptide Y and galanin (Levin et al., 1987), there are smaller inputs from nearby adrenaline-containing nuclei (C1–C3) to the parvocellular parts of Pa, which colocalize neuropeptide Y (Sawchenko et al., 1985; Alonso, 1993). Within the PaMP, A1/C1 neurons undoubtedly target both TRH (Shioda and Nakai, 1993; Diano et al., 1998) and CRH (Kitazawa et al., 1987) neurons. In addition to the functions mentioned above for catecholaminergic pathways in the SO relative to OX and VP release, brain stem adrenergic and noradrenergic pathways also play important roles in physiological functions related to the parvocellular neurosecretory system. For example, these pathways are activated by both intero- and exteroceptive stress stimuli which activate CRH neurons and result in corticosterone release. In some cases, however, the activation of catecholaminergic cell groups may be secondary to the response of neurons (such as those in Pa) with which they make reciprocal connections (Li et al., 1996). A small number of Sol neurons projecting to the Pa (and probably to SO) are separate from catecholamine neurons. One group is immunoreactive for the gonadal peptide inhibin-β (activin) and favors the region of OX neurons (Sawchenko et al., 1988b), and a second group contains somatostatin and innervates parvocellular zones and also OX regions (Sawchenko et al., 1988a). The peptide PACAP is also present in several Sol and

ventrolateral medullary neurons (Legradi et al., 1994; Shioda et al., 2000). Some PACAP-positive terminals in the PaMP that innervate TRH- and CRH-positive neurons (Legradi et al., 1997, 1998) may derive from these inputs. Enkephalinergic inputs to the Pa also derive from lateral reticular and, to a small extent, Sol neurons (Beaulieu et al., 1996). While some afferents are relatively selective for Pa subgroups, the input topography also can reflect differences in dendritic vs somatic inputs. For example, the dendrites of OX and AVP neurons in the PaMM and PaML can cross into other Pa regions (Fig. 6), particularly into the PaDC, PaV, and PaMP, and it is known that noradrenergic synapses occur largely on Pa dendrites (Olschowka et al., 1981), some of which have been identified as AVP-containing and in the medial zone (Silverman et al., 1983; Ochiai and Nakai,1990). Second, local axon collaterals terminate in both parvo- and magnocellular regions. Third, some axons innervating the Pa bifurcate to supply both parvo- and magnocellular regions (van den Pol, 1982). Thus, while a particular input may prefer one cell type (e.g., Sol to OX and PaMP neurons, A1 to CRH and AVP neurons), a significant overlap may exist (Renaud and Bourque, 1991). Additional brain stem regions that provide a significant innervation of the Pa are the lateral parabrachial nucleus (Saper and Loewy, 1980), the median and dorsal raphe (containing serotonergic neurons) (Sawchenko et al., 1983; Petrov et al., 1992), and the ventral periaqueductal gray (Floyd et al., 1996). Forebrain and Hypothalamic Inputs SFO and MnPO inputs to the Pa are extensive (Miselis, 1981; Tribollet and Dreifuss, 1981; Silverman et al., 1981; also see Oldfield and Mckinley, Chapter 16, this volume) and may contact many subregions (Sawchenko and Swanson, 1983). Additional forebrain and hypothalamic inputs seem primarily directed to the nonmagnocellular subnuclei. These include afferents from most preoptic and hypothalamic nuclei, including the anteroventral periventricular nucleus, the arcuate nucleus, the dorsomedial nucleus, the lateral hypothalamus, the medial preoptic area, and the rostral periventricular area (Berk and Finkelstein, 1982; Sawchenko and Swanson, 1983; Simerly and Swanson, 1988; Standaert and Saper, 1988; Thompson et al., 1996; Csaki et al., 2000; Mihaly et al., 2001). A large portion of the galanin innervation of Pa comes from the dorsomedial nucleus (Levin et al., 1987). The arcuate nucleus provides ACTH/MSH (Sawchenko et al., 1982; Kiss et al., 1984) and NPY (Kalra and Crowley, 1992; Legradi and Lechan, 1998). The input from the anteroventral periventricular nucleus is immunoreactive for atrial natriuretic peptide (Standaert and Saper, 1988). Another prominent

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afferent source to the Pa is the bed nucleus of the stria terminalis (Silverman et al., 1981; Tribollet and Dreifuss, 1981; Sawchenko and Swanson, 1983), the projections of which can be divided according to the originating subdivision. The lateral division of the dorsal bed nucleus projects primarily to the PaAP, whereas denser and more widespread projections to many Pa subdivisions emanate from the fusiform subnucleus (Dong et al., 2001). Some forebrain inputs to the Pa, like those from the lateral septum and the ventral subiculum, may largely outline the nucleus rather than project within it (Sawchenko and Swanson, 1983). Nevertheless, direct projections to both OX and AVP dendrites from these regions have been confirmed with the electron microscope in this perinuclear position (Oldfield et al., 1985). The central and medial amygdaloid nuclei project largely to parvocellular regions of the Pa, including both MEE- and brain stem/spinal cord-projecting groups (Silverman et al., 1981; Tribolett and Dreifuss, 1981; Gray et al., 1989), but again, synapses with AVP and OX dendrites have been noted (Oldfield et al., 1985). This reinforces the need to consider dendritic cytoarchitecture and ultrastructure to fully explore synaptic connectivity. According to Prewitt and Herman (1998), direct amygdaloid projections to the Pa may be weak relative to central and medial nucleus projections to neurons in the bed nucleus of the stria terminalis, which in turn project to the Pa. Many of the hypothalamic and forebrain inputs to the Pa are thought to be glutamatergic (Csaki et al., 2000) or GABAergic (Roland and Sawchenko, 1993), including local inputs (Tasker et al., 1998). The perifornical and lateral hypothalamic regions also contain substantial numbers of neurons colocalizing CART and melanin concentrating hormone which project to the parvocellular parts of the Pa (Broberger, 1999; Peyron et al., 1998). The same regions house, in separate neuronal populations, hypocretin/orexin-containing neurons (Peyron et al., 1998; Broberger, 1999) which project to the Pa and widely in the CNS. Both peptides, and the Pa, have been implicated in energy balance (Meister, 2000), but hypocretin/orexin is also strongly associated with arousal state, and its absence correlates with narcolepsy (Sutcliffe and de Lecea, 2000). Originally described by Sofroniew and Weindl (1978), the suprachiasmatic nucleus (SCh) provides projections primarily to the parvocellular Pa (Berk and Finkelstein, 1981; Watts et al., 1987; Watts and Swanson, 1987). These projections give the Pa, and its hormonal and preautonomic outflow, direct access to circadian clock outputs. The SCh consists of two distinct regions with different connections and cell types. A dorsomedial shell, which contains VP neurons, is largely responsible for the Pa

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inputs (Sofroniew and Weindl, 1978; Vrang et al., 1995; Larsen, 1999; Teclemariam-Mesbah et al.,1999; Leak and Moore, 2001). The ventrolateral core, which receives direct retinal input and contains vasoactive intestinal polypeptide (VIP) and gastrin-releasing peptide neurons, projects much more strongly to the adjacent subparaventricular zone. However, a few direct Pa projections from the ventrolateral SCh, including VIP fibers, are suggested by close proximity of terminal-like profiles and the somata and dendrites of spinal cordprojecting neurons (Teclemariam-Mesbah et al., 1997; Vrang et al., 1997) and by transneuronal tracing studies (Teclemariam-Mesbah et al.,1999). Thus, outputs from both divisions of the SCh are important for circadian influences on sympathetic outflow (e.g., to the pineal for control of melatonin) by way of the intermediolateral cell column. Light microscopic evidence also supports direct SCh projections to CRH neurons in the PaMP (Vrang et al., 1995). Electrophysiologically, Pa neurons (including those in magnocellular subdivisions) can get conventional inhibitory or excitatory inputs from the SCh depending on their location in the Pa and their output target (Hermes and Renaud, 1993). Inputs to spinal cord-projecting neurons appear to be direct and mediated by GABA or glutamate (Cui et al., 2001). As with the SO, a variety of neurotransmitters and neuromodulators, and their receptors, have been located within the Pa (Hatton, 1990; Renaud and Bourque, 1991). Of these, GABA and glutamate are particularly important (Decavel and van den Pol, 1990; Decavel and van den Pol, 1992; Cullinan, 2000; Csaki et al., 2000). Considered with some of the above evidence, GABA synapses could arise from (1) interneurons; (2) collaterals of MEE-projecting neurons, including those colocalizing CRH; (3) local projection neurons in the subparaventricular zone, the perinuclear zone of the SO, and the anterior perifornical region; or (4) as discussed above for the SO, the several areas, such as the bed nucleus of the stria terminalis, which have many GABAergic neurons and which project to the Pa (van den Pol, 1982; Meister et al., 1988; Okamura et al., 1990; Rho and Swanson, 1989; Decavel and van den Pol, 1992; Cullinan et al., 1993; Roland and Sawchenko, 1993). Glutamatergic projections are thought to arise from a wide number of Pa inputs, but forebrain and hypothalamic inputs are particularly important (Csaki et al., 2000). In addition to adrenaline and noradrenaline, dopamine also provides a more substantial and homogenous synaptic input to the Pa than to the SO (Buijs et al., 1984; DeCavel et al., 1987). The source of dopamine remains to be clarified, but probably stems from some intrinsic parvocellular neurons and from other diencephalic sources (e.g., zona incerta, periventricular hypothalamus).

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ACCESSORY MAGNOCELLULAR NEUROSECRETORY NEURONS As illustrated in Figs. 3 and 6, about one-third of the magnocellular OX and AVP neurons lie outside the confines of the SO and the Pa (Rhodes et al., 1981). Two of these groups are almost exclusively composed of OX neurons and deserve special mention because they are often treated as part of the Pa (Swanson and Kuypers, 1980; Swanson et al., 1986). These are the AC and the PeM, both of which project to the neurohypophysis, as do additional accessory groups (e.g., Sherlock et al., 1975; Armstrong et al., 1980; Ju et al., 1986). The AC lies some 300–400 μm rostral to, and is discontinuous with, the other clusters of magnocellular Pa neurons (Armstrong et al., 1980; Kiss et al., 1991). Its dendritic orientations are strongly medial, toward the third ventricular wall, and posteromedial (Fig. 6) (Armstrong et al., 1980, 1982a; Ju et al., 1986, 1992). The PeM is closely associated with the AC, beginning at the same level but extending caudally (Armstrong et al., 1980; Ju et al., 1986). Neurons of the PeM have dendritic ramifications that form a dense subependymal plexus (Sofroniew and Glasmann, 1981; Ju et al., 1986, 1992). There are additional magnocellular accessory cell groups, the more prominent being the posterior and anterior fornical nuclei, the nucleus circularis, neurons within the medial forebrain bundle in the lateral hypothalamus, scattered neurons in the lateral preoptic area and lateral hypothalamus, and neurons within the boundary of the bed nucleus of the stria terminalis. The morphology of individual accessory neurons has been well studied by Sofroniew and Glasmann (1981), who noted that some of these cells had thick processes with terminal swellings and, occasionally, two axon-like processes. Accessory nuclei neurons are often associated with vasculature (Ambach and Palkovits, 1979) and their processes often project along blood vessels (Ju et al., 1986; Riva et al., 1999). Each cluster has a unique appearance. For example, the nucleus circularis contains cells that are tightly packed around blood vessels, exhibit extensive somatic–somatic appositions, and have only a sparse dendritic tree (Tweedle and Hatton, 1976; Ju et al., 1986). While accessory nuclei share some inputs with the SO and the PA, like that from the SFO (Miselis, 1981), their differing topography and dendritic structure may be associated with differences in connectivity and neurochemical expression. Examining a large variety of neuropeptides/neurotransmitters in two structures, Duan et al. (1998) found that the terminal-like patterns observed in the SO and the Pa were found in an accessory nucleus of the lateral hypothalamus, whereas nucleus circularis somata were seldom contacted. Some accessory nuclei show Fos responses to peripherally

administered CRF and urocortin that are different from those of the SO and Pa (Wang et al., 2000) and have different levels of coexpression of NADPH-diaphorase and acetylcholinesterase (Crespo et al., 1998). But in general, accessory nuclei respond to physiological stimuli such as dehydration (Ding et al., 1994) and hypovolemic challenge (Shen et al., 1992) in a fashion similar to that of the SO and Pa. In other species, some magnocellular accessory groups do not project to the neural lobe and may have central functions (Ferris, 1992).

CONCLUSION Understanding the functional circuitry of the Pa, SO, and related hypothalamic systems remains challenging on several fronts. The relationship between the release of hormones such as VP and OX and peripheral stimuli is well understood, even if many details of connectivity (e.g., how is the suckling stimulus transferred to OX neurons?) are not. For questions such as these, transneuronal tracing studies in combination with functional neuroanatomical probes (e.g., Fos localization) hold great promise. But perhaps the greatest general challenge is for detailed studies of connectivity to keep apace with new neurochemical discoveries. It is critical that colocalization of neuroactive substances in these nuclei, and their activity-dependent expression, not be studied at the expense of determining precise neuroanatomical relationships, especially synaptic connectivity. The necessity to combine these efforts is nowhere more important than in the Pa, where the wide variety of neurons, functions, and connectivity is truly daunting. The functional plasticity of this system and the meaning of functionally labile chemical maps remains a challenge for interpreting anatomical data. For example, more is known of the relationships of GABAergic and glutamatergic synaptic terminals in the SO than is known about their precise source. The ability to visually identify anatomically (through prior tracer placement) or neurochemically (with markers such as green fluorescent protein) defined neurons, or terminals, in complex electrophysiological recording experiments is another promising multidisciplinary effort. In this regard, the precise topographic distinctions among major Pa cell types and their separable efferent targets discovered over 20 years ago remain a solid foundation for expanding functionally related neuroanatomical studies in this nucleus.

Acknowledgments This chapter was completed while the author was supported by NIH Grants NS 23941 and HD 41002.

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in glucocorticoid-sensitive vasopressin and corticotropin-releasing factor neurons in the hypothalamic paraventricular nucleus. J. Neurosci. Res. 19, 405–411. Van den Pol, A. N. (1982). The magnocellular and parvocellular paraventricular nucleus of rat: Intrinsic organization. J. Comp. Neurol. 206, 317–345. Van den Pol, A. N. (1991). Glutamate and aspartate immunoreactivity in hypothalamic presynaptic axons. J. Neurosci. 11, 2087–2101. Vandesande, F., and Dierickx, K. (1975). Identification of the vasopressin producing and of the oxytocin producing neurons in the hypothalamic magnocellular neurosecretory system of the rat. Cell Tissue Res. 164, 153–162. Vandesande, F., Dierickx, K., and DeMey, J. (1977). The origin of the vasopressinergic and oxytocinergic fibers of the external region of the median eminence of the rat hypophysis. Cell Tissue Res. 180, 443–452. Van Leeuwen, F. W., De Raay, C., Swaab, D. F., and Fisser, B. (1979). The localization of oxytocin, vasopressin, somatostatin and luteinizing hormone releasing hormone in the rat neurohypophysis. Cell Tissue Res. 202, 189–201. Vigh, B., and Vigh-Teichmann, I. (1998). Actual problems of the cerebrospinal fluid-contacting neurons. Microsc. Res. Tech. 41, 57–83. Vrang, N., Larsen, P. J., Clausen, J. T., and Kristensen, P. (1999). Neurochemical characterization of hypothalamic cocaineamphetamine-regulated transcript neurons. J. Neurosci. 19, RC5, 1–8. Vrang, N., Larsen, P. J., Müller, M., and Mikkelsen, J. D. (1995). Topographical organization of the rat suprachiasmatic-paraventricular projection. J. Comp. Neurol. 353, 585–603. Vrang, N., Mikkelsen, J. D., and Larsen, P. J. (1997). Direct link from the suprachiasmatic nucleus to hypothalamic neurons projecting to the spinal cord: A combined tracing study using cholera toxin subunit B and Phaseolus vulgaris-leucoagglutinin. Brain Res. Bull. 44, 671–680. Wang, L., Martinez, V., Vale, W., and Tache, Y. (2000). Fos induction in selective hypothalamic neuroendocrine and medullary nuclei by intravenous injection of urocortin and corticotropin-releasing factor in rats. Brain Res. 855, 47–57. Watts, A. G., and Swanson, L. W. (1987). Efferent projections of the suprachiasmatic nucleus. II. Studies using retrograde transport of fluorescent dyes and simultaneous peptide immunohistochemistry in the rat. J. Comp. Neurol. 258, 230–252. Watts, A. G., Swanson, L. W., and Sanchez-Watts, G. (1987). Efferent projections of the suprachiasmatic nucleus. I. Studies using anterograde transport of Phaseolus vulgaris leucoagglutinin in the rat. J. Comp. Neurol. 258, 204–229. Wiegand, S. J., and Price, J. L. (1980). Cells of origin of the afferent fibers to the median eminence in the rat. J. Comp. Neurol. 192, 1–19. Whitnall, M. H., and Gainer, H. (1988). Major pro-vasopressinexpressing and pro-vasopressin-deficient subpopulations of corticotropin-releasing hormone neurons in normal rats. Neuroendocrinology 47, 176–180. Wuarin, J. P. (1997). Glutamate microstimulation of local inhibitory circuits in the supraoptic nucleus from rat hypothalamus slices. J. Neurophysiol. 78, 3180–3186. Xi, D., Kusano, K., and Gainer, H. (1999). Quantitative analysis of oxytocin and vasopressin messenger ribonucleic acids in single magnocellular neurons isolated from supraoptic nucleus of rat hypothalamus. Endocrinology 140, 4677–4682. Yamada, K., Emson, P., and Hokfelt, T. (1996). Immunohistochemical mapping of nitric oxide synthase in the rat hypothalamus and colocalization with neuropeptides. J. Chem. Neuroanat. 10, 295–316. Yang, C. R., Senatorov, V. V., and Renaud, L. P. (1994). Organum vasculosum lamina terminalis-evoked postsynaptic responses in rat supraoptic neurones in vitro. J. Physiol. 477, 59–74.

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Seven circumventricular organs (CVOs) are recognized in the brain of the rat and earn this classification because they are all located in the walls of the lateral, third, or fourth cerebral ventricles (Hofer, 1958). The subfornical organ, vascular organ of the lamina terminalis (OVLT), pineal gland, subcommissural organ, and median eminence/neurohypophysial complex are located at various positions in the wall of the third ventricle (Fig. 1A). The area postrema is found in the wall of the fourth ventricle (Figs. 1A and 2B). The choroid plexus can be found within the lumen of the lateral, third, or fourth ventricle (Figs. 1A, 1C, and 2B).

The ependymal cells forming the ventricular surfaces of the CVOs also are different in appearance from the regularly apposed cuboidal ependyma lining the rest of the ventricular surface. Ependymal cells of the CVOs can be elongated or columnar and of irregular appearance. They either are devoid of or have very few cilia on their luminal surface (Weindl and Joynt, 1971; Phillips et al., 1974; Dellman and Simpson, 1979; McKinley et al., 2003). This contrasts with the densely ciliated surfaces of normal ventricular ependyma. Another way in which the ependymal cells of the CVOs are different from normal ependyma is the presence of tight junctions between adjacent cells. This restricts the passage of marker molecules, such as horseradish peroxidase (HRP), between cerebrospinal fluid and the CVO interstitium and vice versa (Weindl, 1973; Krisch et al., 1978; Broadwell and Brightman, 1976; McKinley et al., 1990). Thus, it is thought that the blood–brain barrier is shifted from the level of the capillary endothelium to the ependyma in the CVOs. The choroid plexus and subcommissural organ are composed of one or more layers of modified ependymal cells and do not contain nerve cell bodies. These were termed ependymal CVOs by Kuhlenbeck (1970), whereas the remaining CVOs were termed paraependymal CVOs because they have subependymal elements that differ greatly from those of the ependymal cells. Only the subfornical organ, OVLT, and area postrema contain neuronal perikarya. They have many common features and have been classified as the sensory CVOs (Johnson and Gross 1993). These three CVOs have

GENERAL FEATURES Although there is not a uniformity of structure, several features distinguish CVOs from other regions of the brain. All are highly vascularized structures and possess unusual vascular arrangements, with many capillary loops reaching near to the ventricular surface (Duvernoy and Koritke, 1964). From a functional point of view, probably the most interesting aspect of the vascular morphology is the presence of capillaries having fenestrated endothelial cells in all CVOs except the subcommissural organ (Weindl, 1973; Weindl and Joynt, 1973). This results in the blood–brain barrier being disrupted in all but one of the CVOs, and, unlike the rest of the brain, there can be bidirectional movement of polar molecules between the hemal and the neural environments of the CVOs.

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FIGURE 1 (A) Location of the circumventricular organs shown on a sagittal section through the third ventricle of the rat brain. The choroid plexus of the lateral ventricle is not shown. The lateral part of the area postrema appears in the section because at this level the section is just lateral to the midline. The bar represents 1 mm. Stain; cresyl violet/luxol fast blue. (B) Enlarged view of the rat OVLT. Bar represents 250 μm. Stain: cresyl violet. (C) Enlarged view of the subfornical organ. Bar represents 100 μm. Stain, cresyl violet/luxol fast blue. Abbreviations used: ac anterior commissure; AP, area postrema; ChP, choroid plexus; ME, median eminence; MnPO, median preoptic nucleus; ox, optic chiasm; OVLT, vascular organ of the lamina terminalis; pc, posterior commissure; Pi, pineal gland; PiRe, pineal recess of the third ventricle; NH, neurohypophysis; SCO, subcommissural organ; SFO, subfornical organ; vhc, ventral hippocampal commissure; 3V, third ventricle.

extensive afferent and efferent neural connections and a distinctive cytoarchitecture and cytochemistry. The remainder of this chapter considers the structure, connectivity, and neurochemical characteristics of the three sensory CVOs. Brief individual descriptions of the other CVOs then follow.

SUBFORNICAL ORGAN The subfornical organ (also known as the intercolumnar tubercle in the older literature) was recognized initially in the brain of an Australian bat, Nictophylus timoriensis, by Elliot Smith (1898) and first

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FIGURE 2 (A) Enlarged view of the rat subcommissural organ in midsagittal section shown in Fig. 1A. Arrows indicate the extent of the subcommissural organ at the entrance to the aqueduct of Sylvius. The bar represents 100 μm. (C) Enlarged view of the area postrema at the entrance to the central canal in the midsagittal section. The bar represents 100 μm. Abbreviations used: AP, area postrema; Aq, aqueduct of Sylvius, cc, central canal; ChP, choroid plexus; hbc, habenular commusure; pc, posterior commissure; PiRe, pineal recess of the third ventricle; SCO, subcommissural organ; SFO, subfornical organ; 4V, fourth ventricle.

described in detail (in the human brain) by Putnam (1922). It is a prominent feature in Nissl-stained sections of the rat brain. Situated in the anterior dorsal wall of the third ventricle, at the confluence of the two interventricular foramina and the third ventricle, the subfornical organ of the rat lies ventral to the hippocampal commissure and caudoventral to the junction of the two fornical columns (Figs. 1A, 1C, and 3E). Its dorsal extremity connects to the tela choroidea, whereas the ventral stalk of the subfornical organ merges into the median preoptic nucleus. Rostrally the septal triangularis nucleus bounds the subfornical organ (Akert et al., 1961). The subfornical organ has been subdivided into several subregions by Sposito and Gross (1987). Two major subregions can clearly be defined on the basis of neural connectivity, receptor densities, vasculature, and function as shown by expression of the immediate early gene c-fos (McKinley et al., 2003). These are an inner “ventromedial core” and a peripheral “outer shell” (Figs. 3E and 3F; also see Figs 146 and 147 of Paxinos et al., 1999) As with other CVOs, the subfornical organ is highly vascularized (Fig. 3A). It contains an extensive network of capillaries taking the form of sinusoids and capillary loops (Spoerri 1963; Duvernoy and Koritke, 1964). The perivascular space is extensive and labyrinthine, and both fenestrated and nonfenestrated capillary endothelia are observed. Intravascularly administered horseradish peroxidase enters the total intercellular space of the subfornical organ, indicative that the entire organ is exposed to the hemal milieu (Broadwell and Brightman, 1976; McKinley et al., 1990). The ventrome-

dial core of the subfornical organ is reported to have the greatest density of capillaries and neurons (Sposito and Gross 1987). Neurons are distributed throughout the subfornical organ and have been classified into two types in the rat depending on whether or not they have vacuoles (Dellmann and Simpson, 1979; Dellman 1998). At the ventricular surface, the ependymal cells are flattened. Cilia are absent from many of the ependymal cells in the central part of the subfornical organ, whereas isolated ependymal cells around the periphery exhibit tufts of cilia (Phillips et al., 1974; McKinley et al., 2003).

Afferent Neural Connections Early studies in the rat, utilizing lesions and Golgi techniques, showed afferent neural input to the subfornical organ from the adjacent median preoptic nucleus and the triangular septal nucleus (Hernesniemi et al., 1972). A later study using anterogradely transported amino acids as tracers did not confirm the projection from the septal triangularis nucleus (Miselis et al., 1979). Other investigations, which utilize conjugates of HRP or fast blue injected into the subfornical organ, have confirmed that the median preoptic nucleus provides the richest afferent input to the subfornical organ (Lind et al., 1982, 1984; Saper and Levisohn, 1983). Approximately 20% of neurons in the median preoptic nucleus with projections to the subfornical organ also exhibit collateral branches projecting to the supraoptic nucleus (Oldfield et al., 1992). Another CVO in the lamina terminalis, the OVLT, also has neurons projecting to the subfornical organ (Lind et al., 1982; Gu and Simerly,

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FIGURE 3 (A, B) The circumventricular organs are the most vascularized structures in the brain. The extensive network of blood vessels in the rat subfornical organ is shown in A and the area postrema in B. Many looping and intertwining capillaries can be seen. The blood vessels have been filled with a mixture of Indian ink in gelatine that was injected through the heart postmortem. Bars = 200 μm. (C–F) Immunohistochemical detection of Fos expression in response to intravenous (iv) infusion of either angiotensin II or relaxin reveals functional subdivisions in the rat OVLT and subfornical organ. Fos-labeled cell nuclei are observed mainly in the lateral zone of the OVLT in response to infusion of angiotensin II (1 μg/h). The dorsal cap region of the OVLT shows considerably less Fos in response to iv infusion of angiotensin II (C), whereas the dorsal cap is the main site of Fos expression in response to iv infusion of relaxin at 25 μg/h (D). (E). Infusion of angiotensin II (0.3 μg/h) stimulates Fos expression in the ventromedial core of the subfornical organ, while infusion of relaxin (25 μg/h) stimulates Fos expression in the outer shell of the subfornical organ (F). Bars = 100 μm. Abbreviations used: AP, area postrema; cc, central canal; ChP, choroid plexus; cp, capillary plexus; dc, dorsal cap of the OVLT; Vhc, hippocampal commissure; lz, lateral zone of the OVLT; Sol, nucleus of the solitary tract; ox, optic chiasm or optic recess of the third ventricle; os, outer shell of the subfornical organ; vc, ventromedial core of the subfornical organ; 3V, third ventricle; 4V, fourth ventricle.

1997; McKinley et al., 2003), with up to 30% showing collaterals connecting to the supraoptic nucleus (Oldfield et al., 1992). Additional sites of afferent input to the subfornical organ are the paraventricular hypothalamic nucleus (Larsen et al., 1991); the medial preoptic,

dorsal preoptic, and anterior hypothalamic regions; and the medial septum, but these are present in densities lower than those in the lamina terminalis (Lind et al., 1982). Neurons in the zona incerta and reuniens nucleus of the thalamus also have projections to the subfornical

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organ, and along with the perifornical region of the anterior hypothalamus these connections appear to terminate in the ventromedial core of the subfornical organ and contain angiotensin II (Lind et al., 1984). Afferent neural connections to the subfornical organ from more caudal brain regions have also been identified. These projections are from the locus coeruleus (Miselis et al., 1987), the dorsal and median raphe (Lind, 1987), the lateral parabrachial nucleus (Gu and Ju, 1995), and the nucleus of the solitary tract (ZardettoSmith and Watson, 1987; Ciriello et al., 1996) to the subfornical organ.

Efferent Neural Connections As a result of studies using anterogradely transported amino acids (radiolabeled), three major efferent projections from subfornical organ neurons were initially identified. These were the median preoptic nucleus, the OVLT, and the supraoptic nucleus (Miselis et al., 1979). Subsequently, these findings have been confirmed, and connections to the paraventricular hypothalamic nucleus (both parvocellular and magnocellular regions), the anterior periventricular preoptic region, the dorsal perifornical area, the ventromedial region of the lateral preoptic region, and the lateral hypothalamus have been reported by two independent studies (Miselis, 1981; Lind et al., 1982). Some subfornical neurons have been found to have axons with branches projecting to both the supraoptic and the hypothalamic paraventricular nuclei (Weiss and Hatton, 1990), although these are not common and most neurons in the subfornical organ project only to the side of the supraoptic nucleus ipsilateral to the side of the subfornical organ in which they are located (Renaud et al., 1993). Other sets of terminal fields were identified when the anterogradely transported lectin Phaseolus vulgaris leucoagglutinin was injected into the subfornical organ. Neurons in the substantia innominata, the rostroventral parts of the bed nucleus of the stria terminalis, the rostral zona inserta, and the infralimbic area of the prefrontal cortex were the additional targets of subfornical efferents identified with this tracer (Swanson and Lind, 1986). Studies with retrogradely transported tracers show that, within the subfornical organ, neurons have a topographic distribution, depending on the destination of their projections. Neurons of the subfornical organ projecting to the median preoptic nucleus, supraoptic and paraventricular hypothalamic nuclei, zona incerta, prefrontal cortex, and substantia innominata are distributed mostly in an annular arrangement in the periphery of the subfornical organ, with very few retrogradely filled neurons in its ventromedial core (Miselis et al.,

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1979; Lind et al., 1982, 1984; Swanson and Lind, 1986). Two regions have been identified as the main targets of the subfornical organ neurons that occupy the ventromedial core of this CVO and these were the bed nucleus of the stria terminalis and the parvocellular subdivisions of the hypothalamic paraventricular nucleus (Swanson and Lind, 1986; McKinley et al., 2003). Recent neural tract-tracing studies in the rat which used the neurotropic virus pseudorabies injected into peripheral sites such as the kidney, gut, salivary glands, heart, forelimb muscle, and sympathetic ganglia, show that there are polysynaptic efferent neural pathways from the subfornical organ to these peripheral targets (Hubschle et al., 1998; Sly et al., 1999; Westerhaus and Loewy, 1999; Yang et al., 1999; McKinley et al., 2003). These results indicate that the subfornical organ has the potential to influence sympathetic nerves supplying many peripheral organs and tissues, as proposed by Miselis et al. (1987). Fibers emanating from the subfornical organ to the aforementioned terminal fields have two main trajectories. The first group of fibers exits from the anterior aspect of the subfornical organ and travels in a ventral direction rostral to the anterior commissure and along the anterior edge of the median preoptic nucleus to enter this nucleus as well as the OVLT and the preoptic periventricular and supraoptic nuclei. Other efferent fibers take a postcommissural route via the fornix and diverge with the stria terminalis to the medial and lateral hypothalamus, with terminations in the supraoptic and paraventricular nucleus. Some postcommissural fibers turn rostrally and join precommissural fibers bound for the median preoptic and supraoptic nuclei (Miselis, 1981; Lind et al., 1982).

Neuroendocrine Aspects Consistent with proposals that the subfornical organ is a site at which blood-borne humoral agents exert central actions are observations that this CVO contains receptor binding sites for a number of circulating hormones. These include amylin (Sexton et al., 1994), angiotensin II (Mendelsohn et al., 1984; Gehlert et al., 1986; Lenkei et al., 1997; Allen et al., 2000) (mostly the AT-1 receptor subtype), atrial natriuretic peptide (Quirion et al., 1984; Mendelsohn et al., 1987), calcitonin (Rouleau et al., 1984), glucagon-like peptide-1 (GLP-1) (Göke et al., 1995), relaxin (Osheroff and Phillips, 1991), somatostatin (Patel et al., 1986), and vasopressin (V1 receptor; Phillips et al., 1988). As well, the calcium receptor that responds to small changes in extracellular Ca2+ concentration (Rogers et al., 1997) and the lipopolysacharide CD14 receptor, which is implicated in neuroimmune responses such as fever (Lacroix et al.,

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1998), have been identified in the subfornical organ. Nitric oxide synthase (NOS) is abundant in neurons throughout all subdivisions of the subfornical organ (Jurzak et al., 1994; Alm et al., 1997). The actions of angiotensin II on the subfornical organ have been studied extensively (see Ferguson and Bains, 1997), and it is clear that this octapeptide acts on receptors in the subfornical organ to stimulate neural pathways subserving water drinking and pressor responses in the rat (Simpson et al., 1978; Simpson, 1981). Studies of c-fos expression show that circulating angiotensin II stimulates Fos production in neurons throughout the subfornical organ (McKinley et al., 1992; Rowland et al., 1996); however, neurons in the ventromedial core are more sensitive to angiotensin II and there appears to be a higher density of angiotensin II receptors within the ventromedial core of the subfornical organ than its periphery (Lenkei et al., 1997; McKinley et al., 1998; Allen et al., 2000). This distribution contrasts with the peripheral distribution within the subfornical organ of osmoresponsive neurons that project to the supraoptic nucleus (Oldfield et al., 1994). Of relevance to the actions of angiotensin II on the subfornical organ are observations of high concentrations of angiotensin-converting enzyme in this CVO (Brownfield et al., 1982; Chai et al., 1987), which may allow circulating angiotensin I to be converted to angiotensin II in the vicinity of subfornical neurons. Atrial natriuretic peptide antagonizes the dipsogenic action of angiotensin II (Masotto and Negro-Vilar, 1985), and these two peptides have an overlapping distribution of their receptors in the subfornical organ, as well as in the OVLT and the area postrema (Mendelsohn et al., 1987), suggesting that the inhibitory influence of atrial natriuretic peptide could be at the primary angiotensin-sensitive neuron in CVOs. The ovarian hormone relaxin may influence oxytocin secretion in pregnant rats via receptors in the subfornical organ (Summerlee et al., 1987). Circulating relaxin has been shown to increase the expression of c-fos in many neurons in the periphery of the subfornical organ that have efferent neural connections with the supraoptic and paraventricular nuclei (Sunn et al., 2001), and electrophysiological evidence confirms that relaxin stimulates neuronal activity in this CVO in the rat (Sunn et al., 2002). Systemically infused relaxin stimulates water drinking as well as vasopressin secretion in rats (Parry et al., 1994; Sinnayah et al., 1999) and the dipsogenic effect is abolished by ablation of the subfornical organ (Sunn et al., 2002). It is likely that the relaxin binding sites observed in the subfornical organ (Osheroff and Phillips, 1991) represent specific relaxin receptors that mediate water drinking induced by blood-borne relaxin.

In addition to receptors for angiotensin II, neurons containing this peptide have been immunohistochemically identified in the rat subfornical organ (Lind et al., 1985). These neurons are distributed mainly in the periphery of the subfornical organ and some have been shown to project to the median preoptic nucleus (Lind et al., 1984). Fibers containing angiotensin II (Lind et al., 1984, 1985; Oldfield et al., 1989), somatostatin (Krisch et al., 1978), and luteinizing hormone-releasing hormone (LHRH) (Krisch and Leonhardt, 1980; McKinley et al., 1990) terminate in the subfornical organ. Some terminals are found in the vicinity of perivascular spaces and do not make synaptic contact with other neurons, suggesting neurosecretion of peptides into the circulation of the subfornical organ (Krisch and Leonhardt, 1980; Oldfield et al., 1989).

VASCULAR ORGAN OF THE LAMINA TERMINALIS The vascular organ of the lamina terminalis forms the ventral part of the midline anterior wall of the third cerebral ventricle (Figs. 1A, 1B, and 3C). First described by Behnsen in 1927, it is commonly abbreviated as OVLT, from the Latin organum vasculosum laminae terminalis, and is also known as the supraoptic crest, medial prechiasmatic gland, or optic recess organ in the older literature. In the rat, it extends dorsally, approximately 1 mm from the optic chiasm and roof of the optic recess, the small anteroventral extension of the third ventricle. It is bounded both rostrally and caudally by liquor spaces, namely, the prechiasmatic cistern and the optic recess of the third ventricle, respectively. It is bounded by the diagonal band of Broca at its lateral edges, and the median preoptic nucleus is immediately dorsal. The OVLT and subfornical organ form the respective ventral and dorsal poles of a continuum of tissue that includes the intervening median preoptic nucleus and is collectively termed the lamina terminalis. Embryologically the subfornical organ and OVLT are derived from the same anterior pole of the neural tube and are progressively divided during development by the ingrowth of fibers that ultimately forms the anterior commissure. Therefore, it is not surprising that, in addition to the absence of a blood-brain barrier, those two regions share a commonality of structure, neural connections, receptor types, neurochemical content, and function (Landas and Phillips, 1987). The ependymal cells overlying the OVLT have tight junctions (both zonnula occludens and zonula adherens) near their apical surfaces. As previously mentioned, this is distinct from the cuboidal ependymal cells lining the remainder of the ventricular system, which lack

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junctions and therefore offer no impediment to the flow of substances between cerebrospinal fluid CSF and the parenchyma of the brain (Weindl and Joynt, 1971; McKinley et al., 1987; Landas and Phillips, 1987). The most prominent morphological feature of the OVLT, as with most CVOs, is that it contains a rich vascular plexus with specialized arrangements of the blood vessels (Duvernoy and Koritke, 1964; Mergner, 1961; Yamaguchi et al., 1993). It is this complex arrangement of blood vessels that has given rise to the compartmentalization of the OVLT into an external zone and an internal or parenchymal zone. Small branches of the preoptic artery (which are distributed from the anterior communicating artery) in the prechiasmatic cistern break up into a dense network of small vessels in the pia mater at the cisternal edge of the OVLT. Invaginations of pia mater that envelope capillary loops extend farther into the organ in a Y-shaped configuration when observed in horizontal section (Landas and Phillips, 1987), and these two vascularized regions compose the external zone. The internal zone contains a complex neuropil with neurons, fibers, glia, capillary loops, and perivascular spaces extending to near the ependyma. Many of the capillaries exhibit fenestrated endothelial cells (Rohlich and Wenger, 1969; Landas and Phillips, 1987; McKinley et al., 1987). The tight junctions normally present between the endothelial cells (the basis of the blood–brain barrier) are effectively shifted in part to the ventricular surface and partly to capillaries in the tissue at the boundary between the OVLT and adjacent brain regions (Krisch et al., 1978, 1987). The net effectiveness of junctions between specialized ependymal cells (tanycytes) at the ventricular surface and between convoluted basal processes extending to the boundaries of the OVLT is shown by the spread of blood-borne substances, such as horseradish peroxidase, through particular parts of the OVLT but not into the surrounding neuropil or into the ventricular CSF (Krisch et al., 1987). Based on neural connectivity, neuronal phenotype, and c-fos expression, three functional subdivisions of the rat OVLT can be made: a central capillary plexus, a dorsal cap region, and a lateral zone which extends caudally around the central capillary plexus into the periventricular tissue (Figs. 3C and 3D). The arterial blood supply to the fenestrated plexus of capillaries in the OVLT most commonly arises from one or two horizontally directed arterioles, which branch off the anterior communicating artery, and occasionally an arteriole that branches from an anterior cerebral artery, at the rostral margin of the organ (Ambach et al., 1970; Grafe and Weindl, 1987; see also Scremin et al., Chapter 34, for a depiction of the major arteries). The venous drainage of the OVLT is

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achieved by four to eight veins that run in a caudal direction, with the dorsal veins flowing into vessels directed toward the anterior commissure; the others extend caudally and ventrally into a venous sinus (Grafe and Weindl, 1987). This venous drainage is continuous with a surface network that eventually drains the preoptic and retrochiasmatic regions (Szabo, 1983). The arrangement of a fenestrated capillary plexus in the OVLT and the presence of neurohormones and neurotransmitters in a secretory configuration around perivascular spaces (see below) have led several workers to suggest a neuroendocrine role for the OVLT, analogous to the portal system of the median eminence (Weindl, 1973; Palkovits et al., 1978). The fact that the major venous drainage of the OVLT is to the medial preoptic region, where blood vessels are not fenestrated in the rat (Palkovits et al., 1978), is not consistent with the concept of a portal system. In this regard there is a poorly understood anastomosing vascular network that interconnects the OVLT with the subfornical organ, median preoptic nucleus, and choroid plexus (Szabo, 1983). The function for this network as a conduit for neurohormones is not clear, because the direction of blood flow is unknown. Between the centrally positioned vascular plexus and the outer margins of the OVLT is a complex internal zone consisting of a labyrinth of tanycyte processes, glial cells, and neural elements. The last are characterized by small, “primitive” neuronal cell bodies and nerve terminals. Often these terminals contain neurosecretory granules and are aligned without synaptic specializations along the basement membrane, which forms the outer margin of perivascular spaces surrounding fenestrated blood vessels. The growing list of neuropeptide releasing factors and putative transmitters contained within these terminals includes LHRH and somatostatin (Baker et al., 1975; Krisch and Leonhardt, 1980), angiotensin II (Oldfield et al., 1989), vasopressin and oxytocin (Weindl and Sofroniew, 1985), and atrial natriuretic peptide (Kawata et al., 1985). These terminals are derived from cell bodies, located largely outside the OVLT.

Afferent Neural Connections LHRH terminals arise from cell bodies in the adjacent preoptic area (Silverman et al., 1979; Palkovits et al., 1978). Similarly oxytocin fibers arise from cell bodies in the areas surrounding the OVLT (Weindl and Sofroniew, 1985). Vasopressin fibers project to the OVLT from parent cell bodies located in the suprachiasmatic nucleus (Buijs, 1978). Other areas known to project to the OVLT with nonspecified phenotype include the other components of the lamina terminalis, the subfor-

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nical organ, and the median preoptic nucleus (Miselis, 1981; Saper and Lewisohn, 1983), as well as the medial and lateral preoptic areas, the anterior lateral and dorsomedial hypothalamus, the locus coeruleus, the midbrain central gray (Camacho and Phillips, 1981; Palkovits et al., 1978), and the hypothalamic paraventricular nucleus (Larsen et al., 1991).

Efferent Neural Connections Efferent pathways from the OVLT have been studied by mapping the distribution of anterogradely transported tracer following its microinjection into the OVLT of the rat (Camacho and Phillips, 1981; Phillips and Camacho, 1987; Gu and Simerley, 1997; Uschakov et al., 2001). The major efferent pathway from the OVLT descends in the medial forebrain bundle, although a lesser number of fibers pass along the periventricular region of the hypothalamus (Gu and Simerley, 1997). There are reciprocal connections with the median preoptic nucleus and subfornical organ and hypothalamic paraventricular nucleus. A few efferent fibers ascend dorsally to reach the parastrial nucleus and the intermediate part of the lateral septal nucleus (Gu and Simerley, 1997). There also are direct neural inputs to the supraoptic nucleus which arise mainly from a compact group of neurons in the dorsal cap of the OVLT (as do the neurons in the OVLT that project to the magnocellular subdivisions of the paraventricular nucleus) and this has been confirmed by the retrograde transport of tracer molecules injected into the supraoptic nucleus. These neurons are activated by systemic hypertonicity or dehydration and may represent the major group of physiological osmoreceptors regulating vasopressin secretion (Oldfield et al., 1994; McKinley et al., 1994; Larsen and Mikkelsen, 1995; McKinley et al., 2003). Efferent connections to parvocellular regions of the paraventricular nucleus are also present (Gu and Simerley, 1997; Uschakov et al., 2001), and it is likely that the majority of these fibers come from the lateral zone of the OVLT (Sunn et al., 2001; McKinley et al., 2003). A strong projection from the OVLT to the lateral hypothalamic region has been confirmed by retrograde tracing experiments (Uschakov et al., 2001). Other regions of the hypothalamus that receive minor inputs from the OVLT are the arcuate and periventricular nuclei (Camacho and Phillips, 1981; Gu and Simerley, 1997). There are also connections to specific parts of the limbic system. These include efferent projections to the bed nucleus of the stria terminalis, which have been confirmed by retrogradely transported tracer, with this projection arising mostly from the more lateral parts of the OVLT rather than from the dorsal cap (Uschakov et al., 2001; McKinley et al., 2003).

Tracer retrogradely transports from the cingulate cortex to the dorsal cap of the OVLT (Uschakov et al., 2001) confirming that a direct projection from the OVLT to this region exists (Camacho and Phillips, 1981). More caudal brain regions that receive efferents from the OVLT are the periaqueductal gray and the lateral parabrachial nucleus (Gu and Simerley, 1997). Like the subfornical organ, tract-tracing investigations with the neurotropic virus pseudorabies show that there may be polysynaptic pathways from the OVLT to many peripheral organs and tissues (Hubschle et al., 1998; Sly et al., 1999; Westerhaus and Loewy 1999; McKinley et al., 2003). These data suggest that the OVLT connects to sympathetic nerves supplying the periphery, and this may be another means by which circulating factors may influence sympathetic vasomotor pathways. Such a scheme may provide the anatomical substrate for the effects of anteroventral third ventricle wall (AV3V) lesions attenuating experimentally induced hypertension (Brody et al., 1978).

Neuroendocrine Aspects As mentioned above, the OVLT is a site of dense LHRH containing fibers and terminals (Silverman et al., 1979) and may play a role in regulating estrous cycles (Wenger and Leonardelli, 1980). NO synthase is also present in neurons and fibers in the OVLT (Jurzak et al., 1994), although its function in the OVLT has yet to be delineated. Similar to the subfornical organ, the OVLT is rich in binding sites for several circulating peptides, such as amylin (Sexton et al., 1994), angiotensin II (Mendelsohn et al., 1984; Giles et al., 1999; Allen et al., 2000), calcitonin gene-related peptide (Sexton et al., 1986); atrial natriuretic peptide (Saavedra et al., 1992), and relaxin (Osheroff and Phillips, 1991), suggesting that neurons in the OVLT may be influenced by these circulating peptides. With regard to vasopressin secretion, as well as mediating osmotically stimulated vasopressin secretion, neurons in the dorsal cap of the OVLT (Fig. 3D) that project to the supraoptic and paraventricular nuclei may play a role in mediating influences of the ovarian pregnancy hormone relaxin on vasopressin secretion (Sunn et al., 2002). By contrast, neurons in the OVLT that exhibit angiotensin II AT1 receptors and respond with increased Fos production to intravenous angiotensin II (Fig. 3C) are situated mainly near the lateral boundaries of the organ and in its most caudal periventricular region (McKinley et al., 1992; Giles et al., 1999), and it is possible that they have a role in the centrally mediated pressor effect and sodium appetite induced by circulating angiotensin II (Simpson, 1981; Fitts and Masson, 1990).

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AREA POSTREMA The area postrema, first described in detail (in the human brain) by Wilson (1906), is situated at the apex of the calamus scriptorius in the dorsomedial medulla oblongata (Figs. 1A and 2B). In most species it appears as bilateral rounded eminences on either side of the fourth ventricle at its entrance to the central canal. In the rat the major part of these bilateral eminences have merged so that, in coronal sections, the area postrema has the appearance of a hump or quadrant of tissue dorsal to the nucleus of the solitary tract (Fig. 3B). The ependymal cells of the area postrema are flattened, and small neurons surrounded by astrocytic cells and processes are found throughout this CVO. Axodendritic synapses are common and axosomatic synapses are also present (Spacek and Parizek, 1969; Dempsey, 1973; Armstrong et al., 1982). Because of its rich vascularity, the area postrema appears spongy. On describing this vasculature, Duvernoy and Koritke (1964) commented on groups of capillaries that were spiral in form and that gave rise to many subependymal capillary loops (Fig. 3B). Two types of capillaries have been observed in the area postrema: large sinusoidal vessels and smaller capillaries. Fenestrated endothelial cells line many capillaries in the area postrema, causing the blood–brain barrier to be circumvented. Perivascular spaces that contain large nerve cells and processes containing both clear and dense-core vesicles are also observed (Roth and Yamamoto, 1968; Spacek and Parizek, 1969; Dempsey, 1973; Armstrong et al., 1982).

Afferent Neural Connections Afferent neural connections to the area postrema come from only a few regions of the rat brain. The most abundant afferent input comes from the hypothalamic region in the vicinity of the paraventricular and dorsomedial nuclei (Hosoya and Matsushita, 1981; van der Kooy and Koda, 1983; Shapiro and Miselis, 1985; Larsen et al., 1991). This connection appears to be a continuous group of cells extending from the lateral parvocellular subnucleus of the paraventricular hypothalamic nucleus to the perifornical region and dorsomedial nucleus of the hypothalamus. In horizontal sections, this group of cells and dendrites appears as oval rings extending from a periventricular to a perifornical location at each extremity (Shapiro and Miselis, 1985). A small number of cells in the lateral parabrachial nucleus have also been shown to project to the area postrema. There is also the possibility that the dorsal raphe and a few neurons in the A1 region of the ventrolateral medulla and A2 neurons within commissural and medial subnuclei of the nucleus of the solitary tract innervate the

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area postrema (Torack et al., 1973; Vigier and Portalier, 1979; Shapiro and Miselis, 1985). However, Shapiro and Miselis (1985) considered it unlikely that neurons in these regions were retrogradely labeled from injections of tracer into the area postrema because the tracer had probably diffused past the edge of the area postrema. In addition to this central input, the area postrema receives some fibers from the periphery. Vagal and glossopharyngeal fibers have been shown to terminate in the area postrema of the rat (Kalia and Sullivan, 1982; Altschuler et al., 1989).

Efferent Neural Connections Area postrema neurons connect mainly to adjacent regions of the nucleus of the solitary tract, particularly the dorsal part of its medial subnucleus (Torack et al., 1973; van der Kooy and Koda, 1983). The most dense terminal field is found in the C2 adrenergic group of neurons situated midway between the solitary tract and the area postrema. A dense terminal field of fibers from the area postrema also surrounds the A2 noradrenergic group (Cunningham et al., 1994). There are few if any connections within the dorsal motor nucleus of the vagus, but it has been suggested that some area postrema connections within the nucleus of the solitary tract are terminating on dendrites of vagal motor neurons with cell bodies in the vagal motor nucleus (Shapiro and Miselis, 1985). Other medullary targets of area postrema efferents may be the ventrolateral medulla, the dorsal aspect of the spinal trigeminal tract and nucleus, and the paratrigeminal nucleus (Shapiro and Miselis, 1985), although Cunningham et al. (1994) did not confirm these last two projections in a later study. They also doubted that fibers from the area postrema terminated in the ventrolateral medulla. However retrogradely transported tracers that were microinjected into the rostral ventrolateral medulla (RVLM) are transported back to noradrenergic cell bodies in the area postrema (Blessing et al., 1987; Badoer et al., 1994; Polson et al., 1996), evidence for a projection from the area postrema to the RVLM. The most prominent efferent projection of the area postrema outside the medulla is to the dorsal pontine region, the target of these fibers being the middle third of the lateral parabrachial nucleus (van der Kooy and Koda, 1983; Shapiro and Miselis, 1985; Cunningham et al., 1994). Two groups of neurons, one in the core and the other in the lateral zone of the area postrema, project to the specific subnuclei of the lateral parabrachial nucleus. In the case of the core, its projections are to the inner part of the external subnucleus, and the central and dorsal subnuclei, while the lateral zone projects to the outer part of the external subnucleus (Herbert et al.,

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1991). Cholecystokinin- and galanin-containing neurons are among those in the area postrema that provide this neural input to the lateral parabrachial nucleus (Herbert and Saper, 1990). The pericentral dorsal tegmental and dorsolateral tegmental nuclei may also receive inputs from the area postrema (van der Kooy and Koda, 1983).

Neuroendocrine Aspects Fuxe and Owman (1965) identified catecholamineand serotonin-containing neurons in the rat area postrema. Subsequently immunohistochemical investigations of tyrosine hydroxylase and dopamine βhydroxylase confirm the presence of a group of noradrenaline containing neurons in this CVO (Torack et al., 1973; Armstrong et al., 1982). Neurons containing GABA, substance P, enkephalin, neurotensin, and cholecystokinin, as well as serotonin-containing neurons, have also been identified by immunohistochemistry (Newton and Maley, 1984; Armstrong et al., 1981; Newton and Maley, 1985a, 1985b; Newton et al., 1985). Some of the serotonin-containing neurons have been shown to connect to the lateral parabrachial nucleus (Lanca and van der Kooy, 1985). Unlike the other sensory CVOs, little if any nitric oxide synthase is observed in the area postrema (Jurzak et al., 1994). Because the neuropil of the area postrema is accessible to circulating humoral agents, results from in vitro autoradiographic binding studies provide insights into possible hormonal influences that may be involved in the functions of the area postrema. Binding sites for amylin (Sexton et al., 1994), cholecystokinin (Moran et al., 1986), angiotensin (Lenkei et al., 1997; Allen et al., 2000), atrial natriuretic peptide (Mendelsohn et al., 1987; Saavedra et al., 1992), GLP-1 (Göke et al., 1995), somatostatin (Patel et al., 1986), vasoactive intestinal polypeptide (Shaffer and Moody, 1986), neuropeptide Y (Martel et al., 1986), vasopressin (Phillips et al., 1988), substance P (Helke et al., 1984), and insulin (Werther et al., 1987) have been observed in the area postrema. With regard to cholecystokinin, it has been shown that systemic infusion of this hormone induces Fos production in neurons of the area postrema, indicative that it is a receptor zone for circulating cholecystokinin (Luckman, 1992), although its major site of action appears to be on distal vagal afferents (Day et al., 1994). Like the other sensory CVOs, the area postrema exhibits the CD14 lipopolysacharide receptor (Lacroix et al., 1998). Neurons in the area postrema may participate in baroreflex and cardiovascular control (Undesser et al., 1985; Skoog and Mangiapane, 1988; Ferguson, 1991), with both angiotensin II and vasopressin acting on this CVO to influence these functions (Hasser et al., 2000).

The area postrema probably has a role in the control of food intake, body weight, and fluid homeostasis (Miselis et al., 1987). Earlier, it was proposed that the area postrema was a trigger zone for the vomiting reflex in cats (Borison and Brizzee, 1951). This idea has persisted since that time, with recent interest in the role of the area postrema in the actions of antinausea drugs such as the 5-HT3 antagonists (see Miller and Leslie, 1994, for review). Although rats do not vomit, there is evidence that the area postrema in this species has a role in the development of conditioned taste aversions associated with nausea (Berger et al., 1973; Coil and Norgren, 1981).

MEDIAN EMINENCE AND NEUROHYPOPHYSIS The median eminence is a circumventricular organ that would be inappropriate to consider in isolation from the associated pituitary gland. The internal lamina of the median eminence is continuous with the posterior pituitary or neurohypophysis, and both share embryological and functional properties. The median eminence, or more specifically the ependyma overlying it, forms the floor of the third ventricle immediately caudal to the optic chiasm. Similar to other CVOs, the ependymal cells of the median eminence are irregular in shape (generally flattened), lack cilia, and have intercellular junctions that restrict the passage of substances to and from the CSF (Kobayashi and Matsui, 1969; Page, 1986). In broad terms, the median eminence is divided into inner and outer laminae (see Armstrong, Chapter 15, for a diagram). The internal zone is immediately subjacent to the ependyma and consists of subependymal glial cells, pituicytes, the radially directed processes of tanycytes, and most importantly fibers of the supraopticoneurohypophysial tract, which traverse the inner zone of the median eminence en route to the posterior pituitary (Daniel and Pritchard, 1975; Haymaker et al., 1969; Page, 1986). The unmyelinated fibers of this tract contain vasopressin and oxytocin and are derived from magnocellular neurons in the paraventricular and supraoptic nuclei of the hypothalamus (see Armstrong, Chapter 15, for a detailed description). In addition to fibers en passage, nerve terminals are present in the internal zone, although the extent to which they form neurohemal or other contacts remains uncertain (Page and Dovey-Hartman, 1984). These terminals are derived from catecholaminergic neurons in the brain stem or arcuate nucleus and form the reticuloinfundibular and tuberohypophysial pathways (Björkland et al., 1973; Everett and Hokfelt, 1986). Peptidergic axons containing corticotropin-releasing

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factor (CRF) or products of the proopiomelanacortin pathway (Merchenthaler et al., 1982; Mezey et al., 1985) also terminate in the internal zone. The external zone contains a complement of glial cells, ependymal cells, and nerve terminals similar to that described for the internal zone, although terminals are distinctly neurohemal. Importantly, there is a fenestrated vascular network called the primary portal plexus, and, in close association with these vessels, an abundance of terminals containing neurohormones and releasing factors (Page, 1986). The pars tuberalis forms the ventral surface of the external zone of the median eminence and vessels of the portal system are found within this region. The arterial blood supply to the neurohypophysis arises entirely from the internal carotid artery (Wislocki and King, 1936). The adenohypophysis, on the other hand, does not receive a direct arterial blood supply (Green, 1951; Page and Bergland, 1977) but is perfused exclusively by long portal vessels emanating from the primary portal plexus and to a lesser extent from short vessels derived from the neurohypophysis (Wislocki and King, 1936; Xuereb et al., 1954; Page and Bergland, 1977). This is a critical feature of the structure and function of the anterior pituitary, because it is via these portal vessels that hypophysiotrophic hormones reach the anterior pituitary gland. This vascular anatomy, together with the lack of local nerve terminals in the adenohypohysis, led early workers to suggest that the glandular pituitary is under the control of humoral factors released into the vicinity of the portal vasculature (Green and Harris, 1947). Since these early observations, considerable emphasis has been placed on defining the nature and origins of the various hypophysiotrophic factors released into the portal vessels and ultimately responsible for the release of anterior pituitary hormones, including thyroid-stimulating hormone, luteinizing hormone, prolactin, and adrenocorticotropic hormone (ACTH). During the last 10–15 years, attempts to elucidate the different inputs to the median eminence and particularly the external lamina have relied on the use of retrogradely transported tracers (Weigand and Price, 1980; Lechan and Jackson, 1982; Silverman et al., 1987), immunocytochemistry (Hokfelt et al., 1975), and occasionally combinations of both (Silverman et al., 1987). Fibers in the supraopticoneurohypophysial tract of the internal zone are clearly derived from the supraoptic nucleus and to a lesser extent from the paraventricular nucleus. Other afferent inputs, more likely to contain hypophysiotrophic factors, arise from the parvocellular paraventricular nucleus, periventricular hypothalamus, arcuate nucleus, OVLT, preoptic region, diagonal band of Broca, medial septum, and brain stem (Weigand and Price, 1980). Combinations of immunocytochemistry

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and tracing studies, for example, have localized LHRH neurons projecting to the median eminence in the medial septum, the diagonal band of Broca, the preoptic region surrounding the OVLT, and the lateral hypothalamus (Silverman et al., 1987). Although combined experimental data of this type may not always be available, the anatomical overlap of regions supplying afferents to the median eminence with the distribution of hypophysiotrophic factors suggests the possible origins of many of these factors. Thyrotropinreleasing factor, for example, is derived from cells in the diagonal band of Broca, the medial septum, the arcuate nucleus, and the paraventricular nucleus (Hokfelt et al., 1975; Lechan and Jackson, 1982; Ishikawa et al., 1984). Dopamine, which has an inhibitory influence on prolactin release (Neill, 1985), is derived from cells in the arcuate nucleus, as are the cells of origin of fibers in the external lamina containing growth hormone-releasing hormone. The distribution of cells containing somatostatin is widespread but cells projecting specifically to the median eminence are situated in the periventricular hypothalamus (Eppelbaum et al., 1977). Corticotropinreleasing factor, the principal agent responsible for the release of ACTH, is contained in parvocellular neurons of the paraventricular nucleus (Vale et al., 1981; Sawchenko et al., 1984; Sawchenko, 1987). Vasopressin, which is also contained within parvocellular neurons, has long been known to be present in nerve terminals in the external lamina (Parry and Livett, 1973) and to have effects on ACTH release analogous to those of CRF in a variety of species, including the rat (Rivier and Vale, 1983). Those agents listed above are some of the major neurohormones contained in projections to the external lamina of the median eminence; however, there are others, such as cholecystokinin, neurotensin, oxytocin, neuropeptide Y, and dynorphin, that may have regulatory effects on the anterior pituitary and axonal trajectories to the median eminence, but are less well defined (Roth et al., 1983; Sawchenko et al., 1984; Sutton et al., 1988; Mezey et al., 1986).

SUBCOMMISSURAL ORGAN The subcommissural organ, located in the posterior wall of the third ventricle, forms the dorsal roof of the entrance to the aqueduct of Sylvius (Fig. 2A) and is immediately ventral to the posterior commissure of the rat (Rodriguez et al., 1987; Severs et al., 1987). This CVO consists of layers of elongated columnar ependymal cells, which appear to have a secretory function. Secretion of a glycoprotein into the CSF by subcommissural organ cells causes the formation of Reissner’s fiber. This noncellular condensation of glycoprotein con-

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tinues from the aqueduct of Sylvius through the whole length of the central canal of the spinal cord (Rodriguez et al., 1987; Sterba et al., 1987; Gobron et al., 1999). There are no neurons in the subcommissural organ, but there is neural innervation of the specialized ependyma. Serotonergic fibers from the raphe provide some of this neural input (Mollgard et al., 1978; Bouchard, 1979; Rodriguez et al., 1987). Like other CVOs, the subcommissural organ exhibits a dense capillary network running beneath the ependymocytes, with some capillaries extending into the ependyma (Duvernoy and Koritke, 1969). Unlike the other CVOs, capillaries in the subcommissural organ do not exhibit fenestrated endothelial cells; thus the blood–brain barrier is maintained in this region (Weindl and Joynt, 1973; Broadwell and Brightman, 1976; Severs et al., 1987; McKinley et al., 1990). The function of the subcommissural organ (and Reissner’s fiber) remains obscure, although a number of earlier speculations regarding a role in fluid and electrolyte homeostasis or gonadotropin secretion can be found in the literature (see Severs et al., 1987, for a review). It is also proposed that the molecules secreted in Reissner’s fiber may have a role in guiding the development of the brain and spinal cord (Gobron et al., 1999). Although the subcommissural organ is a distinctive feature in the brain of the rat and many other vertebrates, it seems to be absent from the human brain from soon after birth, although a remnant of it may remain in vestiges of the mesocelic recess (Rakic, 1965; McKinley and Oldfield, 1990).

PINEAL GLAND This endocrine gland, found throughout the vertebrate phylum, has been a subject of interest for many centuries. In mammals, the pineal gland is a midline, cone-like structure located in the dorsocaudal roof of the third ventricle. It develops as a thickening of the ependyma around the pineal recess of the third ventricle and is attached by peduncles to the habenular and posterior commissures (Kappers, 1981; Korf et al., 1998). In the rat, most pineal cells have migrated in a dorsocaudal direction so that the rat pineal is mostly superficial, with a relatively long stalk that connects to the habenular and posterior commissures (Fig. 1A). This stalk contains nerve fibers, blood vessels, connective tissue, and parenchymal cells (Pevet, 1983). The pineal gland of the rat does not contain any neurons, but has many secretory cells known as pinealocytes. The other major cell type is glial (Pevet, 1981). Phylogenetically, pinealocytes developed from photoreceptor neurons; however, in the rat these cells have no direct

sensitivity to light. Instead, a neural pathway from the eye via the sympathetic nervous system influences the secretion of melatonin and other indolamines from the pinealocytes (Kappers, 1981). This pathway originates in the retina, from where neural signals are relayed in the optic nerve via the retinohypothalamic pathway, which branches directly from the optic chiasm to reach the suprachiasmatic nucleus. Here, synaptic connections are formed with neurons projecting to the hypothalamic paraventricular nucleus. Another junction occurs here, and these neurons connect with sympathetic preganglionic neurons in the intermediolateral cell column of the spinal cord via a pathway that travels in the medial forebrain bundle and through the reticular formation. These sympathetic preganglionic neurons innervate the superior cervical ganglion, which in turn provides the nerve supply to the pineal and its melatonin-secreting cells through the nervi conari (Moore and Klein, 1974; Pickard and Turek, 1983; Tamarkin et al., 1985). Stimulation of melatonin by the pinealocytes is dependent on intact sympathetic innervation from the superior cervical ganglion, with darkness-stimulating and light-inhibiting melatonin secretion (Korf et al., 1998). Studies in which the neurotropic virus pseudorabies was injected into the rat pineal to trace the polysynaptic pathways to it from the hypothalamus confirm the above pathway and show that the neurons in the paraventricular nucleus that are involved in this pathway are situated in its dorsal, medial and lateral parvocellular subdivisions (Larsen et al., 1999; TeclamariamMesbah et al., 1999). Viral tracing from the pineal in rats in which the superior cervical ganglion had been bilaterally destroyed showed that neural inputs from the nucleus of the solitary tract and salivatory nucleus via a synaptic connection in the sphenopalatine ganglion may provide a parasympathetic input to the pineal (Larsen et al., 1999). Nerve fibers that probably come from the habenular and posterior commissures have been observed to travel in the pineal stalk and enter the gland, and there is evidence suggesting a direct central innervation from several brain regions (Guerillot et al., 1982; Daffly, 1983). Such central innervation of the pineal could not be confirmed when care was taken to prevent any spillage of horseradish peroxidase injected into the pineal for retrograde tracing studies (Patrickson and Smith, 1987).

CHOROID PLEXUS This ion transporting tissue has been grouped with the CVOs and is responsible for producing much of

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the cerebrospinal fluid formed in the central nervous system (Davson, 1960). The choroid plexus is found in the lateral, third (dorsal recess), and fourth ventricles and is attached to the ventricular walls (Fig. 1). It is a highly vascularized connective tissue that is enveloped by an extensively folded epithelial cover, giving it a “cauliflower-like” appearance when viewed with the scanning electron microscope. The epithelium consists of cuboidal cells that sit on a basement membrane. These cells exhibit microvilli on the luminal surface and, together with the folding, provide a large surface area for water and electrolyte transfer between the hemal milieu of the choroidal interstitium and the ventricular lumen. The core of the choroid plexus contains numerous capillaries with fenestrated endothelial cells. Vessels are surrounded by perivascular spaces, which are continuous with interdigitating intercellular spaces of the choroidal epithelia (McKinley et al., 1990). Although there is no barrier for the movement of tracer molecules such as horseradish peroxidase from blood to the perivascular spaces of the choroid plexus, the zonula occludens between adjacent epithelial cells provides the basis for a blood–cerebrospinal fluid barrier (Brightman et al., 1975). Sympathetic innervation from the superior cervical ganglion and a vagal input provide neural regulation of the vasculature of the choroid plexus (Edvinsson et al., 1974; Lindvall et al., 1977). The observation that the choroid plexus of the rat is rich in binding sites for many peptides (Chodobski and SzmydyngerChodobska, 2001) including insulin (Werther et al., 1987), vasopressin (Phillips et al., 1988), and atrial natriuretic peptide (Quirion et al., 1984) suggests that these peptides influence the function of this CVO, possibly as local paracrine factors. It is also possible that they are being taken up from the circulation by choroid plexus epthelium and secreted into the CSF. There is evidence that this occurs for thyroid hormones (Schreiber et al., 1990), leptin (Banks et al., 1996), and insulin-like growth factor I (Walter et al., 1999). Similar to some other CVOs, the choroid plexus throughout the rat brain contains high concentrations of angiotensin-converting enzyme (Yang and Neff, 1972). This enzyme is located on the microvilli at the ventricular surface of epithelial cells and also on the membranes of vascular endothelium (Brownfield et al., 1982; McKinley et al., 1990). The functional consequences of these histochemical findings remain to be determined.

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16. CIRCUMVENTRICULAR ORGANS

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17 Thalamus HENK J. GROENEWEGEN and MENNO P. WITTER Research Institute Neurosciences Vrije Universiteit (RIN-VU) Department of Anatomy, VU University Medical Center (VUmc) Amsterdam, The Netherlands

The thalamus of rats, like in most other vertebrate species, forms the largest part of the diencephalon. The diencephalon, situated between the two cerebral hemispheres rostrally and the brain stem caudally, consists of several main nuclear cell groups: the epithalamus, the dorsal thalamus, the ventral thalamus, and the hypothalamus. The dorsal thalamus has close connectional and functional relationships with the cerebral cortex and, to a lesser extent, the basal ganglia and amygdala in the telencephalon and is the main subject of this chapter. The ventral thalamus, ontogenetically originating from an anlage ventral to that of the dorsal thalamus, is a heterogeneous constellation of nuclear groups, i.e., the reticular thalamic nucleus, the ventral lateral geniculate nucleus, the zona incerta, and the subthalamic nucleus. With respect to the ventral thalamus, in view of their close association with the circuitry of the dorsal thalamus, only the reticular thalamic and ventral lateral geniculate nuclei are dealt with in this chapter. For a discussion of the hypothalamus and the subthalamic region see Simerly (Chapter 14 of this volume) and Gerfen (Chapter 18 of this volume), respectively. A discussion of the epithalamus, i.e., the epiphysis, the habenular complex, and the stria medullaris, falls outside the scope of this chapter. An important issue that should be mentioned at the very beginning of this chapter concerns the traditional distinction between the dorsal and ventral thalamus on the basis of developmental arguments. As indicated above, the nuclei that make up the ventral thalamus are generally described as developing from part of the early diencephalon situated ventral to the dorsal thalamic

The Rat Nervous System, Third Edition

anlage. However, as argued by Puelles et al. (for more details see Chapter 1 and references therein), what has traditionally been called “dorsal thalamus” is actually “caudal thalamus” and what has been called “ventral thalamus” is actually “rostral thalamus.” This welldocumented reinterpretation of the development and “layout” of the nuclei that derive from the diencephalic brain vesicle on the basis of gene maps is not trivial and we accept Puelles” argument (as explained in Chapter 1 of this volume) as valid. However, we remain using the terms dorsal and ventral thalamus to keep in continuity with the literature until the new terminology gains ascendancy. The dorsal thalamus forms a complex of cytoarchitectonically, chemoarchitectonically, and hodologically different nuclei, their common characteristic being a reciprocal connectional relationship with distinct parts of the cerebral cortex. Traditionally, the dorsal thalamus is viewed as the final synaptic relay station before extrinsic and intrinsic information can reach the cerebral cortex. On the basis of the content of the information that reaches the various thalamic nuclei through the inputs to these thalamic nuclei, as well as the organizational aspects of the thalamic projections to the cerebral cortex, the dorsal thalamic nuclei have been characterized as specific versus nonspecific; the specific nuclei encompass sensory, motor, and associational relays and the nonspecific nuclei subserve arousal and attentional mechanisms. A global view of the function of the thalamus is that various streams of information that are directed toward the cerebral cortex are gated and modulated at the level of the thalamus. However,

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the precise nature and the mechanisms of these gating and modulatory functions still remain largely elusive. The main characteristics of the ventral thalamus, i.e., the reticular thalamic (Rt) and ventral lateral geniculate (VLG) nuclei, are its projections to the dorsal thalamus and, unlike other thalamic nuclei, the lack of projections to the cerebral cortex. Furthermore, all afferent and efferent projections of the dorsal thalamus, among them prominently the reciprocal fibers with the cerebral cortex, have to pass through the ventral thalamus. Most of these fiber systems, but certainly not all (see below), give off collaterals to the reticular nucleus. In this way the Rt forms feedback and feedforeward loops with the dorsal thalamic nuclei. Thus, the reticular and ventral lateral geniculate nuclei are strategically placed to modulate all incoming and outgoing information of the dorsal thalamus (e.g., Jones, 1985; Sherman and Guillery, 2001). While the VLG and the reticular nuclei have several characteristics in common, the VLG has a number of additional features that distinguishes it from the reticular nucleus and, therefore, the VLG is considered later in this chapter in conjunction with the lateral geniculate complex as a whole (see page 411). Moreover, the reticular nucleus has a specific visual sector like all other sensory modalities (Coleman and Mitrofanis, 1996). Given the restrictions in approach and length, the present chapter can only provide a global overview of the functional anatomy of the thalamus, with emphasis on data obtained from studies in rats. In the past two decades comprehensive monographs have been published on the thalamus (e.g., Jones, 1985; Steriade et al., 1997; Sherman and Guillery, 2001); for more detailed discussions the reader is referred to these and the original publications. Simple as the thalamic relay function to the cerebral cortex may seem, the precise functional role of the thalamus still remains largely unknown. The chapter by Price in the 2nd edition of The Rat Nervous System (Price, 1995) gave an excellent and comprehensive overview of the architecture and connections of the rat thalamus. The present chapter aims to update the knowledge of the anatomy of the rat thalamus on the basis of studies in the past decade and will attempt to place this knowledge in a functional context. In the first section, a number of recent ideas and insights of general interest to thalamic structure and function are reviewed, in subsequent sections specific aspects of functionally different groups of thalamic nuclei are addressed. Emphasis is placed on those nuclei or nuclear groups for which recent research has unveiled new insights. This is in particular the case for the reticular thalamic nucleus and the midline and intralaminar complex of the thalamus. Important details of the organization of various thalamic nuclei have come about through the application

of modern neuroanatomical techniques such as the juxta- or intracellular filling of neurons which show the fine details of single neurons and their (collateral) projection patterns. Likewise, combinations of anatomical tracing, immunocytochemistry, and/or the revelation of activity markers in specific behavioral paradigms have revealed important functional aspects of specific thalamic nuclei. These exciting recent results are discussed in the context of the description of the individual thalamic nuclei below.

SOME GENERAL ASPECTS OF THALAMIC ORGANIZATION The delineation of nuclei in the dorsal thalamus is primarily based on cytoarchitectonic, chemoarchitectonic, and connectional features. The entire complex of nuclei is rostrally, laterally, and ventrally “encapsulated” by fiber bundles, i.e., the external medullary lamina, in which the reticular thalamic nucleus is embedded. The caudal borders of the dorsal thalamus are less clearly defined, the nuclei of the posterior thalamic complex merge with cell groups of the central tegmental field of the mesencephalon. Medially and dorsally the thalamus borders the third and lateral ventricles, respectively, although in rats the third ventricle is largely obliterated as a consequence of the merging of the thalamic halves on both sides of the midline. An internal medullary lamina divides the thalamus into a dorsal and medial group, and a ventral, lateral and posterior group of nuclei. Embedded in the internal medullary lamina are the intralaminar nuclei which, with some of the nuclei in the midline regions of the thalamus, form a separate group on the basis of their dual projections to the cerebral cortex and the basal ganglia, i.e., the midline/intralaminar complex. The reticular thalamic nucleus, as part of the ventral thalamus, forms a thin sheet of neurons at the rostral and lateral borders of the dorsal thalamus, at the outside mostly bordered by fiber bundles of the internal capsule.

Classical Functional Subdivision of the Thalamus The classical categorization of thalamic nuclei is primarily based on the kind of information that is transferred through a particular nucleus or group of nuclei to the cerebral cortex (for a discussion of corticothalamic relationships in the context of specific cortical areas, see Chapter 23, this volume, by Palomero-Gallagher and Zilles). The main category is formed by the principal “relay” nuclei which receive specific sensory, motor, or associational information through ascending or

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descending fiber pathways and transmit this information to specific areas and layers of the cortex. The group of relay nuclei includes among others the anterior and mediodorsal complexes and the ventral and lateral group of nuclei, encompassing among others the ventrolateral, ventromedial, ventral anterior, lateral dorsal, lateral posterior, and posterior nuclei. In addition, the medial and lateral geniculate nuclei belong to the main category of relay nuclei. Another category is formed by the midline and intralaminar nuclei, also indicated as the nonspecific thalamic nuclei, which are considered to receive more general information, in particular from arousal and visceral regions in the brain stem and forebrain. These nuclei project to both the cerebral cortex, although in a somewhat less strictly topographical manner and to cortical layers different from the principal thalamic nuclei, and to subcortical areas like the basal ganglia and the amygdala. As already indicated above, the reticular thalamic nucleus must be considered as a different category in view of its specific position between the dorsal thalamic nuclei and the cerebral cortex and the absence of cortical projections.

Categorization of Thalamic Afferents in “Drivers” and “Modulators” Before dealing in more detail with the structure, connections, and functional aspects of the various thalamic nuclei within this classical categorization in subsequent sections, it is important to briefly discuss recent ideas about the classification of afferents of the thalamus into two main categories, i.e., drivers and modulators (Sherman and Guillery, 1996, 1998, 2001). This classification provides a basis for distinguishing between afferents that carry specific information to the thalamus, which is subsequently transmitted to the cerebral cortex, and other systems that modulate the transfer of (specific) information at the thalamic level. Sherman and Guillery (2001; for the in-depth discussion of this topic and the supporting literature, the reader is referred to this lucidly written monograph) make a strong case for such an organization in the firstorder, sensory thalamic nuclei, in particular the lateral geniculate nucleus, but they propose to explore and test the generality of these organizational principles for other, in particular, higher order thalamic nuclei. The distinction between thalamic afferent fibers as drivers and modulators is based on several criteria, i.e., light microscopical aspects, ultrastructural characteristics, and physiological properties of these fibers and their terminals in the thalamus (Sherman and Guillery, 2001). Sensory afferents, whether from somatosensory, visual, or auditory origin, are all very similar with respect to their light microscopical appearance: relatively thick,

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richly branching terminal fibers with, in most cases, large boutons, although variable in shape and number. These fibers have been indicated as the type II afferent fibers of the thalamus and are considered to be the “drivers” for the primary sensory relay nuclei. At the ultrastructural level, these driving afferents are associated with large terminals containing round vesicles (RL = round vesicles and large terminals). The RL-type terminals are often found in triadic arrangements; i.e., they are presynaptic to the dendrite of a thalamic relay neuron and an axoniform dendritic profile of an interneuron, the latter being also presynaptic to the dendrite of the relay neuron. These triads regularly form part of glomerular structures in which various presynaptic and postsynaptic elements are arranged, typically, but not in all cases, ensheathed by astrocytic processes. The receptive fields of the thalamic afferents are rather faithfully mapped through the thalamus onto the cortical neurons and cortical layers in receipt of these thalamic fibers. In general, driver inputs are excitatory and exert their effect mainly via ionotropic glutamate receptors (Sherman and Guillery, 2001). Cortical afferents in the primary sensory relay nuclei have a different light microscopical appearance: these fibers are thinner and carry small, drumstick-like appendices with boutons along the course through the thalamic nucleus in which they terminate. They are categorized as type I thalamic afferents. These fibers are much more sparsely branched and exhibit fewer terminallike structures, and individual fibers distribute their terminals over a wider area than the type II afferents. The type I fibers are considered to be modulatory. Ultrastructurally, the terminals of these fibers are associated with the category of smaller terminals with round vesicles (RS = round vesicles and small terminals). The receptive field properties of these afferents are generally much wider and less defined compared to those of the type II afferents and, in the case that these afferents are glutamatergic, they probably exert their effects primarily via metabotropic glutamatergic receptors. Apart from the primary sensory relay nuclei, the ventral anterior and the anterior thalamic nuclei receive type II afferents from subcortical sources, i.e., the deep cerebellar nuclei and the mammillary nuclei, respectively. The cerebellar and mammillary afferents are, therefore, considered to be the driving afferents of these nuclei. Interestingly, cortical afferents of, at least a number of, higher order thalamic relay nuclei are of the type II category of afferent thalamic fibers, possibly indicating that these cortical afferents serve as drivers for these association thalamic nuclei. Whereas the type I corticothalamic axons derive mainly from layer 6 of the cerebral cortex, the type II corticothalamic afferents have shown, at least in a number of cases, to arise from

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cells in cortical layer V (Fig. 1). However, the generality of this organization remains to be largely established. Type I afferents, the “modulatory” category of thalamic afferents, not only originate in the deeper layers of the cerebral cortex but also derive from a wide array of subcortical structures in the basal forebrain, diencephalon, and brain stem, carrying various neurotransmitter signals (Sherman and Guillery, 2001). The concept of drivers and modulators provides a challenging framework for the discussion and interpretation of thalamic structure and function. It provides the basis for a further understanding of what specific messages are transferred through the thalamus to the cerebral cortex and which systems modulate this information transfer. It further emphasizes that not only the sheer number or volume of afferents to the thalamus determines their significance but also the character of the message. In that respect, numerically smaller inputs might be more determinant for the functional aspect of a particular afferent system and, by inference, of a particular thalamic nucleus, than more numerous inputs (Sherman and Guillery, 2001). The driver/ modulator concept has been briefly outlined above to serve as a reference for the discussion of the individual thalamic nuclei in subsequent parts of this chapter. In doing so, a final caveat must be placed here. The present chapter focuses on the situation in the rat; several of the data that are used by Sherman and Guillery (2001) to support their concept of drivers and modulators derive from species other than the rat (e.g., cat and primates). Therefore, it remains to be largely established whether the thalamus of the different species are organized according to these same principles, even though this seems very likely. Nevertheless, important species differences have been noted, for example, with

respect to the presence of interneurons (Price, 1995) or of various neurons expressing calcium-binding proteins (Jones, 1998) in the thalamus of rats versus other species.

Reciprocity of the Thalamocorticothalamic Relationships It is almost a dogma that the thalamocorticothalamic relationships are organized such that each cortical area receiving an input from a specific thalamic nucleus faithfully reciprocates this input through a topographically organized cortical projection to the same thalamic nucleus. Apart from the fact that the relationships between the thalamus and the cortex are certainly not exclusively reciprocal (see below), it is also far from clear what this reciprocity functionally means and why the corticothalamic fibers outnumber their thalamocortical counterparts (Jones, 1985). In reference to the discussion above about the distinction between driver inputs and modulator inputs, it is important to realize that the cortical inputs form a mixed population. Whereas most cortical inputs to the primary relay nuclei have the characteristic of type I, modulatory fibers, cortical inputs to higher order thalamic nuclei have the type II characteristics and may form the main, driving input to these nuclei (Sherman and Guillery, 2001). Driver inputs, by terminating on more proximal parts of the dendrites of thalamic relay neurons, might have a stronger influence on these neurons compared to the (cortical) modulator inputs, which in general contact more distal parts of the dendrites of thalamic neurons. Therefore, like Sherman and Guillery argue, it may not be the numbers that count, but more the (ultrastructural) type of input and the position of the terminations on the dendritic tree.

FIGURE 1 Camera lucida drawings of terminal branches of corticothalamic neurons that originate in layer VI (A) or in layer V (B) of the barrel cortex (primary somatosensory cortex) and terminate in the VPM of the thalamus. The axons were filled following small injections of biocytin in specific layers of the barrel cortex, filling only a small number of neurons of which the projection fibers could be individually traced. The corticothalamic fibers originating in layer VI are thin with many short side-branches and small-sized boutons. In contrast, the fibers from layer V neurons, arising as collaterals from axons that are directed toward the brain stem and/or spinal cord, are thicker and have relatively few, but large boutons. From Fig. 2 of Bourassa et al. (1995).

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Another approach of the issue of reciprocity has been entertained by Deschenes et al. (1998) in comparing the pattern and strength of the cortical inputs to a particular thalamic nucleus with those of prethalamic projections to that same thalamic nucleus. As indicated above, thalamocorticothalamic relationships appear not to be strictly reciprocal (e.g., Hoogland et al., 1987; Murphy and Sillito, 1996; Deschenes et al., 1998; McFarland and Haber, 2002). Deschenes et al. (1998) have formulated the “rule of parity” which states that “. . . the distribution of corticothalamic projections across and within thalamic nuclei is determined by the branching patterns of the different classes of prethalamic afferents.” In general, the rule of parity leads to reciprocity in the thalamocorticothalamic relationships, but this is not true in all cases. Deschenes et al. (1998) base their hypothesis on various aspects of the central organization of the vibrissal system in rats, transmitted through the ventrobasal thalamic complex. The functional consequence of the rule of parity, i.e., a matching of prethalamic and corticothalamic fibers at the level of a particular thalamic nucleus, might be that there is not an exclusive feedback role for the corticothalamic system. Thus, depending on the behavioral context, either the corticothalamic or the thalamocortical system might play a feedback role, modulating the input from the alternate system. How and to what extent these varying roles of the corticothalamic system are related to their origin from distinct cortical layers (see above) remain to be established. Whether a rule of parity applies to all thalamic systems is unclear, but this hypothesis provides an important new perspective on the bottomup and top-down functional arrangements of the corticothalamic system. A consequence of the nonreciprocity of the thalamocorticothalamic system might also be that thalamic nuclei play a role in transmitting information from one cortical area to the other, either from sensory areas to association cortical areas (Guillery, 1995) or from association cortical areas to motor areas (McFarland and Haber, 2002).

PRINCIPAL THALAMIC “RELAY” NUCLEI Sensory Nuclei Lateral Geniculate Nucleus The lateral geniculate nucleus in rats is a relatively flattened, more or less oval-shaped nucleus on the dorsolateral surface of the caudal thalamus. It can be subdivided into a dorsal lateral geniculate nucleus (DLG) and a ventral lateral geniculate nucleus (VLG). Between the DLG and the VLG a third component is recognized,

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the intergeniculate leaflet (IGL). The DLG constitutes the main thalamic relay of visual information to the primary visual cortex, the VLG has a number of characteristics in common with the thalamic reticular nucleus, and only its lateral part receives direct retinal input. The IGL constitutes a relay between the retina and the hypothalamus and is involved in circadian functions. For a comprehensive treatment of the anatomical and functional aspects of the lateral geniculate complex, including the DLG, VLG, and IGL, see Sefton et al. (Chapter 32, this volume). Dorsal lateral geniculate nucleus (DLG) The dorsal lateral geniculate nucleus can be readily identified in Nissl-stained sections (Plate 37 in Paxinos and Watson, 1998) as well in acetylcholinesterase-stained sections in which the DLG shows moderate activity (Plate 39 in Paxinos and Watson, 1998). The cytoarchitecture of the DLG of rats is rather homogeneous. The majority of dorsal lateral geniculate neurons are thalamocortical projection cells. Unlike most other principal thalamic nuclei in the rat, the DLG contains several types of interneurons, namely, GABAergic, most of which coexpress NADPH diaphorase, and solely GABAergic or NADPH diaphorase-containing neurons (Ohara et al., 1983; Jones, 1985; Gabbott and Bacon, 1994). In contrast to many other mammalian species, the rat DLG is not clearly laminated, although fiber bundles running in a ventrolateral to dorsomedial orientation, parallel to the optic tract, impose a certain orientation on the neurons in the dorsal lateral geniculate nucleus. However, the largely segregated terminations of the optic fibers from the contralateral and ipsilateral eyes reveal a form of “hidden lamination” in the rat DLG (Reese, 1988; also Jones, 1985; Price, 1995). The lateral lamina directly adjacent to the optic tract, also called the “outer shell,” receives input from the contralateral eye. The medial part of the DLG, called the “inner core,” consists of two regions, the most medial region receives input from the contralateral eye and the lateral region is innervated by the ipsilateral eye (Reese, 1988; Jones, 1985). This organization refers to the caudolateral part of the DLG in which there is a binocular representation of the visual field; in the rostroventral part of the DLG retinal fibers from only the contralateral eye terminate, representing a large part of the temporal visual field (Reese, 1988). Calcium-binding proteins are differentially distributed in fibers and neurons in the dorsal lateral geniculate nucleus. Whereas calretinin and parvalbumin are present only in fibers, calbindin D28K is also expressed in (inter)neurons, in particular in the outer shell (Luth et al., 1993; Figure 230 in Paxinos et al., 1999). The plexus of calretinin fibers is most dense also in the outer shell (Figure 230 in Paxinos et al., 1999). Parvalbumin fibers

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are most probably derived from the retina and the reticular thalamic nucleus, and calretinin fibers from the retina (Luth et al., 1993; Arai et al., 1992). Calbindin D28K-containing fibers may be derived from the superior colliculus (Lane et al., 1997). Affernt and efferent Projections The DLG forms the main relay between the retina and the primary visual cortex (area 17 or V1), the optic terminations in the dorsal lateral geniculate nucleus being retinotopically organized (see above; Jones, 1985; Reese, 1988). The inner core receives, in its two ocular laminae, inputs from the contralateral nasal and the ipsilateral temporal retina, mapping the contralateral visual hemifield. The outer shell receives a projection from the complete contralateral visual hemifield only. These retinotopic maps in the contralateral and ipsilateral laminae of the DLG are in complete register. Lines of projection are oriented rostroventromedially from the optic tract at the thalamic surface through the different laminae of the DLG (Reese, 1988). Retinal inputs terminate as RL-type boutons, indicating that the retinal fibers are the “drivers” of the DLG (Jones, 1985; Sherman and Guillery, 2001; also II). The retinal afferents contact the dendrites of both the thalamocortical neurons and the interneurons, and they take part in glomerulus-like formations separated from the rest of the neuropil by glial elements. Cortical inputs to the dorsal lateral geniculate nucleus are primarily derived from the primary visual cortex (area 17). The different cortical layers of the primary visual cortex have differential projection patterns to the visual thalamus (Bourassa and Deschennes, 1995). Fibers originating in the upper part of layer VI project to the DLG and they terminate in rostrocaudally oriented bands or “rods” that run parallel to the lines of projection of retinal afferents. Neurons in the deeper part of layer VI project to the lateral part of the lateral posterior thalamic nucleus and these fibers give off collaterals to the DLG where they take part in the formation of the rods. Neurons in layer V of the visual cortex do not target the DLG, but their main axon descends to the brain stem while giving off collaterals to the ventral lateral geniculate nucleus and to the lateral posterior and lateral dorsal thalamic nuclei (Bourassa and Deschennes, 1995). Layer VI corticothalamic fibers issue collaterals to the reticular thalamic nucleus while layer V axons do not. The terminating fibers in the DLG, originating from layer VI have small “en passant” varicosities which show at the ultrastrutural level the characteristics of the RStype boutons and may be considered as “modulators” (Jones, 1985; Price, 1995; Bourassa and Deschennes, 1995; Sherman and Guillery, 2001) Subcortical inputs to the DLG arise in the ventral lateral geniculate nucleus, reticular thalamic nucleus,

superior colliculus, and several brain stem nuclei (Reese, 1988; Coleman and Mitrofanis, 1996; Moore et al., 2000). The inputs from the superior colliculus terminate in the peripheral zone of the outer shell and are probably associated with the calbindin DD28K-positive fiber plexus (Reese, 1988; Lane et al., 1997). Brain stem inputs include those from retinal input receiving nuclei like the nucleus of the optic tract, the olivary pretectal nucleus, and the parabigeminal nuclei (Schmidt et al., 1995). Further inputs are derived from the locus coeruleus (noradrenaline), the dorsal raphe nucleus (serotonin), and the laterodorsal tegmental nucleus (acetylcholine) (e.g., Papadopoulos and Parnavelas, 1990). The output of the DLG is primarily directed at the primary visual cortex (area 17), terminating in layer IV, while there are lesser inputs to layers I and VI (Ribak and Peters, 1975; Jones, 1985). The peristriate area 18 also receives a weak input from the DLG (Sanderson et al., 1991). The geniculocortical pathway uses glutamate as neurotransmitter (Kharazia and Weinberg, 1994; Saez et al., 1998). Ventral lateral geniculate nucleus The VLG, like the reticular thalamic nucleus, is embryologically derived from the ventral thalamus. The ventral lateral geniculate nucleus could be viewed as a caudodorsal extension of the reticular thalamic nucleus. Cytoarchitectonically, the VLG can be subdivided into a lateral, magnocellular part (VLGMC) and a somewhat smaller, medial parvicellular part (VLGPC). The two parts are separated by a fiber-rich, cell-free zone (Jones, 1985). Neurons in the magnocellular part contain nitric oxide synthase and enkephalin; those in the parvicellular part substance P and calretinin (Meng et al., 1998; for review see Harrington, 1997). Enkephalinergic neurons have also been identified in the VLG (Hermanson et al., 1995). Afferent and efferent projections Afferents to the ventral lateral geniculate nucleus make a clear distinction between the medial VLGPC and the lateral VLGMC. The VLGPC receives extensive inputs from the brain stem, in particular from the reticular formation, the deep layers of the superior colliculus, the periaqueductal gray matter, peribrachial regions, the laterodorsal tegmental nucleus, the locus coeruleus, the substantia nigra pars reticulata, and deep cerebellar nuclei (Kolmac and Mitrofanis, 2000; Vaudano and Legg, 1992). The VLGMC receives strong projections mainly from the retina and layer V of the visual cortex (Hickey and Spear, 1976; Takahashi, 1985; Bourassa and Deschennes, 1995) and very few fibers from the brain stem (Kolmac and Mitrofanis, 2000). Unlike the dorsal lateral geniculate nucleus, there are no projections from the VLG to

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the cerebral cortex. Instead, the VLG has rather extensive projections to the dorsal thalamus, comparable with the intergeniculate leaflet (see below). Thus, the medial, parvicellular part of the VLG projects to the parafascicular and lateral dorsal thalamic nuclei as well as to the reuniens and rhomboid nuclei in the midline. The lateral, magnocellular VLG sends fibers to the dorsal lateral geniculate and lateral posterior thalamic nuclei (Moore et al., 2000; Kolmac et al., 2000). Hypothalamic projections from the VLG reach the lateral and posterior hypothalamus and the perifornical area. The ventral lateral geniculate nucleus further projects to the zona incerta, the pretectal nuclei, the deep and intermediate layers of the superior colliculus, the dorsal and medial terminal nuclei of the accessory optic system, the periaqueductal gray, the peripeduncular region, and the accessory inferior olive (Moore et al., 2000). Intergeniculate leaflet The IGL is a distinct, dorsoventrally narrow region between the dorsal and ventral lateral geniculate nuclei, which extends virtually over the entire rostrocaudal length of the lateral geniculate complex (Hickey and Spear, 1976; Moore and Card, 1994). Like the ventral lateral geniculate nucleus, it is a derivative of the ventral thalamus. Precise borders of the IGL are difficult to establish in Nissl-stained sections but can more readily be identified with the staining for glial fibrillary acidic protein (GFAP) and several peptides like neuropeptide Y, substance P, and enkephalin, as well as the neurokinin1 receptor. The IGL contains several types of small- to medium-sized neurons most of which have their dendritic arborizations within the nucleus (Moore and Card, 1994; Piggins et al., 2001). Afferent and efferent connections The main sources of input to the intergeniculate leaflet are fibers from the retina and from the contralateral IGL. Retinal fibers terminate as RL-type terminals and may be considered the driving afferents of the IGL (Mikkelsen, 1992; Moore and Card, 1994). The neurons that give rise to the commissural connections contain enkephalin (Card and Moore, 1989). Further inputs to the IGL originate in the suprachiasmatic nucleus, posterior hypothalamic area, superior colliculus, and several brain stem nuclei, among them the locus coeruleus, the raphe, and the laterodorsal tegmental area (Moore et al., 2000). There is a dense substance P-immuoreactive plexus in the intergeniculate leaflet, the origin of which is unknown, however (Piggins et al., 2001). The efferents of the IGL primarily reach the hypothalamus, in particular the suprachiasmatic nucleus and anterior hypothalamic regions, forming the so-called geniculohypothalamic

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tract. Neurons in the IGL projecting to the suprachiasmatic nucleus contain neuropeptide Y (Card and Moore, 1989; Harrington, 1997). Mikkelsen (1994) showed direct projections from the intergeniculate leaflet to the subcommissural organ and the pineal gland. Further outputs of the IGL reach the midline thalamic nuclei, in particular, the paraventricular, but also the reuniens and rhomboid nuclei, as well as the dorsal and lateral hypothalamus and the zona incerta. Brain stem targets include the superficial layers of the superior colliculus, the periaqueductal gray, and several accessory optic nuclei (Moore et al., 2000). Functional aspects of the lateral geniculate complex The DLG must be considered as the main thalamic gateway for visual information from the retina to reach the cerebral cortex. This nucleus rather faithfully maps the external visual field onto the primary visual cortex (area 17, V1). The functional aspects of the DLG have been the subject of a vast body of literature and have recently been elegantly reviewed by others (e.g., Sherman and Guillery, 2001). For a further elaboration on this subject see Sefton et al. (Chapter 32, this volume). The functions of the VLG have been less wellestablished. In view of its origin from the ventral thalamus, as well as some of its connectional characteristics, i.e., the rather extensive projections to the dorsal thalamus (see above), the functions of the VLG might, at least in part, be compared with the those of the reticular thalamic nucleus (see also section “Reticular Nucleus”; Sefton et al., Chapter 32, this volume). However, the functional aspects of the VLG are probably much more differentiated. As a result of their differential sets of afferent and efferent fibers, the medial and lateral parts of the VLG (VLGPC and VLGMC, respectively) are bound to have different, although possibly related, functions. Whereas the lateral VLGMC is more intimately related to the dorsal lateral geniculate nucleus and visual cortices, the medial VLGPC is more closely associated with the hypothalamus, in particular, the suprachiasmatic nucleus. Yet, these two parts of the VLG also share a number of afferents and efferents, as well as with the IGL (see below). Therefore, in a recent review, Harrington (1997) has argued that this entire ventral lateral geniculate complex may be regarded to fulfill distinct, yet interrelated, functions in controlling visuomotor responses and circadian rhythms. Details of these functions, however, remain to be established. Functional aspects of the IGL seem to have been studied in slightly more detail (see also Sefton et al., Chapter 32, this volume). The retinal input to the IGL originates from a specific set of retinal ganglion cells that convey luminance information (Moore et al., 1995). Furthermore,

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the strong reciprocal connections of IGL with the suprachiasmatic nucleus as well as its connections with midline thalamic nuclei and the pineal gland fit in with an important role for the IGL in circadian functions (Moore and Card, 1994; Harrington, 1997). While the IGL projections to the suprachiasmatic nucleus provide an indirect way to influence autonomic and neuroendocrine circadian rhythms, there are also direct connections of IGL fibers with neuroendocrine neurons in other parts of the hypothalamus (Horvath, 1998). Ventral Posterior Complex The ventral posterior complex (VP) in rats occupies an extensive ventrolateral thalamic area, appearing immediately rostromedial to the medial geniculate complex in the caudal third of the thalamus and extending into the rostral third of the thalamus. The VP is bordered ventrally and laterally by the medial lemniscus and the reticular thalamic nucleus and dorsomedially by the posterior complex (see page 417). At rostral levels the posterior complex is gradually replaced by the ventrolateral nucleus, which is situated medial to the VP (see Figs. 28–38 in Paxinos and Watson, 1998). The ventral posterior complex is the main relay for sensory inputs to reach the cerebral cortex and it can be divided into at least three main parts: the ventral posterolateral nucleus (VPL) receiving spinal somatosensory inputs, the ventral posteromedial nucleus (VPM), receiving trigeminal somatosensory inputs, and the parvicellular medial parts of both the VPL and VPM, i.e., the VPPC (see page 419), which forms the main thalamic relay for gustatory and visceral ascending pathways. The VPL and VPM are distinguishable not only based upon their connectivity (see below) but also on the basis of cyto- and chemoarchitectonics. The VPM stands out as a more densely packed nucleus in Nissl-stained sections as compared to the VPL and the posterior complex (e.g., Plates 33 and 37 in Paxinos and Watson, 1998). The VPL and the posterior complex demonstrate a moderate level of activity for acetylcholinesterase (AChE), while the activity is very low in the VPM (e.g., Plates 31 and 35 in Paxinos and Watson, 1998). Most neurons in the VPM and VPL are medium-sized thalamocortical neurons. In contrast to other species, only very few GABAergic interneurons are present in the ventral posterior complex of rats (Harris and Hendrickson, 1987; Price, 1995). The neurons in the VPL are arranged in rostrocaudal and dorsoventral rows that are roughly parallel with the external medullary lamina and these rows curve partially around the rostral pole of the VPM (McAllister and Wells, 1981). In both the VPL and the VPM a dense plexus of parvalbumin fibers is present, while only few calretinin-

or calbindin D28K-positive fibers occur. In subregions of both the VPL and VPM, calbindin D28K-positive neurons are present (Arai et al., 1994; e.g., Figs. 188, 194, 202, and 208 in Paxinos et al., 1999). The VPM is for the most part organized in so-called barreloids, first described in mice and later also identified in rats (Van der Loos, 1976) (Fig. 2). Barreloids are the representations of individual whiskers at the level of the thalamus (see below) formed by cellular aggregates; barreloids can best be visualized using mitochondrial markers such as cytochrome oxidase (Land and Simons, 1985; Haidarliu and Ahissar, 2001). These microstructures in the VPM are most apparent in young rats, but can also be demonstrated in adults. Whereas thalamic barreloids are considered to convey the information from single whiskers, the dendrites of neurons in these barreloids may cross the boundaries into neighboring barreloids providing a neural substrate for crosstalk between barreloids (Desilets-Roy et al., 2002). Afferent and efferent connections The ventral posterolateral and ventral posteromedial nuclei receive their main ascending inputs from somatosensory afferents originating in the spinal cord, dorsal column nuclei, and trigeminal complex (see also Tracey, Chapters 7 and 25, and Waite, Chapter 26, this volume). These somatosensory projections are somatotopically organized such that afferents from the trunk and limbs terminate in the VPL and those from the head terminate in the VPM. Spinal and trigeminal fibers not only reach the VPL and VPM but also distribute over much wider areas of the thalamus, including the posterior nucleus (see page 417) and the intralaminar nuclei (see section “Midline and Intralaminar Thalamic Nuclei”). Spinothalamic fibers, transferring among others nociceptive signals, originate from different laminae of the dorsal horn as well as the central gray of the spinal cord and terminate as large boutons in the ventral posterolateral nucleus (e.g., McAllister and Wells, 1981; Burstein et al., 1990; Dado et al., 1994; Katter et al., 1996; Kobayashi, 1998). Nociceptive information may reach the VPL also indirectly via the caudal medullary reticular formation (medullary dorsal reticular nucleus) in addition to the direct spinothalamic pathway (Villanueva et al., 1998). Dorsal column nuclear afferents likewise terminate with large boutons in the VPL (McAllister and Wells, 1981; Villanueva et al., 1998). Spinal and lemniscal fibers, in part, converge on individual thalamocortical neurons, the lemniscal fibers on the soma, and more proximal parts of dendrites of the neurons than the spinothalamic fibers (Peschanski and Ralston, 1985). Lemniscal, but also spinothalamic, fibers probably use glutamate as neurotransmitter (De Biasi et al., 1994). Spinothalamic fibers have also been

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FIGURE 2 Schematic, three-dimensional reconstruction of the barreloids in the VPM of the rat taken from Haidarliu and Ahissar (2001). (a, b) Low-power and high-power photomicrographs of two perpendicularly cut, cytochrome-oxidase stained sections of the VPM of young rats (6 days postnatal), showing the darkly staining barreloids and the lighter interbarreloid areas. The precise planes of cutting are important for optimal visualization of the barreloids and are described in detail in Haidarliu and Ahissar (2001). (c) In this reconstruction, the orientation of the long axis of the VPM, viewed from caudodorsal to ventrorostral, is shown in a schematic drawing of a coronal section through the right hemisphere at the caudal pole of the VPM. (d) Three-dimensional reconstruction of the barreloids in the VPM. A, B, C, D, and E represent the five barreloid rows; α, β, γ, and δ represent straddlers. Abbreviations: D, dorsal; DLC, dorsolaterocaudal; L, lateral; LC, laterocaudal, R, rostral; RL, rostrolateral; SP, saittal plane; V, ventral. Slightly modified from Fig. 12 of Haidarliu and Ahissar (2001).

shown to contain substance P (Battaglia et al., 1992; Nishiyama et al., 1995). The spinal and principal trigeminal nuclei both project to the ventral posteromedial nucleus as well as, less densely, to the posterior complex (cf. also Waite, Chapter 26, this volume). Projections to the VPM are more focussed, those to the medial part of posterior nucleus more diffuse (Chiaia et al., 1991a). The whiskersensitive parts of the spinal and principal trigeminal nuclei appear to give rise to two trigeminothalamic pathways that reach different parts of the barreloids. These pathways possibly play different functional roles in the relay of information from the whiskers to the somatosensory cortex (Williams et al., 1994; Veinante and Deschenes, 1999; Veinante et al., 2000a; Pierret et al., 2000). From the principal trigeminal nucleus two types of thalamic projection fibers originate. Fibers of the first type, arising from small neurons confined to the socalled barrelettes in the principal trigeminal nucleus

which each have a receptive fields for a single whisker, project densely to the “core compartments” of single barreloids in the dorsomedial two-thirds of the VPM. The second type of fiber originates from larger neurons with receptive fields for multiple whiskers. These fibers terminate in much wider thalamic regions, including the ventral part of the VPM containing the so-called “tail compartments” of the barreloids (Pierret et al., 2000; Veinante and Deschenes, 1999). Specific parts of the spinal trigeminal complex, in particular its interpolar division but also the rostral part of the caudal division, reach the ventral one-third of the VPM, as well as in the interbarreloid areas in the dorsal VPM (Williams et al., 1994; Veinante and Deschenes, 1999). The fibers from the interpolar trigeminal division are thin and form small-sized bushy arbors within the ventral posteromedial nucleus. The VPL and VPM receive afferents from various other subcortical areas, among them a serotonergic

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input from the dorsal raphe nucleus and a GABAergic input from the thalamic reticular nucleus (see section “Reticular Nucleus”) (Cox et al., 1996; Kirifides et al., 2001). Reticular thalamic afferents originate from different types of neurons which have different termination patterns in the ventral posterior complex: some reticular fibers terminate as clusters; others have a wide and more diffuse termination pattern. This indicates that the inhibitory influences of the reticular thalamic nuceus serve different roles in the ventrobasal complex (Cox et al., 1996). The barreloids in the VPM are the thalamic modules in the pathways that lead from the whiskers via the barrelettes in the trigeminal complex to the barrels in the somatosensory cortex (S1). There is an almost exclusive one-to-one relationship from the individual peripheral whiskers to the corresponding cortical barrels. Thus, the thalamocortical projections from the barreloids in the VPM to the barrel cortex, like their corticothalamic counterparts, are very strictly topographically organized (Lu and Lin, 1993; Land et al., 1995; see below). In contrast, thalamocortical fibers from the posterior complex terminate only in the interbarrel areas (Lu and Lin, 1993). Thalamocortical axons from the VPL and VPM terminate predominantly in layer IV of the primary sensory cortex and they most likely use glutamate as neurotransmitter (Kharazia and Weinberg, 1994). Apart from the main terminations in layer IV, thalamocortical axons from the VPM issue also terminal branches in layers I and V/VI (Lu and Lin, 1993; Zhang and Deschennes, 1998). The corticothalamic projections related to the VPL and VPM are organized in a complex way and reflect the hypothesized way in which the afferents of the thalamus are organized as drivers and modulators (see section “Some General Aspects of Thalamic Organization). Corticothalamic projections from the barrel cortex originating from pyramidal neurons in the upper part of layer VI terminate exclusively in the VPM where they arborize in long rostrocaudally oriented bands or “rods.” A rod originating from a single barrel in the cortex makes contact with a series of barreloids that together represent an arc of whiskers (Hoogland et al., 1987; Bourassa et al., 1995). Neurons in the deeper parts of layer VI of the barrel cortex primarily project to the medial part of posterior thalamic nucleus but, in addition, they give off collaterals to the VPM where they participate in the formation of rods (Fig. 3). Neurons in layer VI located in the interbarrel areas project exclusively to the posterior nucleus (Wright et al., 2000). All corticothalamic fibers originating in layer VI give off collaterals to the reticular nucleus and they have long branches that have numerous en passant boutons (Bourassa et al.,

1995; Levesque et al., 1996; Wright et al., 2000). Corticothalamic fibers originating from pyramidal neurons in layer V neurons in the barrel field have main axons descending to the brain stem. They give off collaterals to the posterior and intralaminar thalamic nuclei, but there are no collaterals to the reticular nucleus and VPM. These layer V fibers terminate in a few clusters with large boutons in the posterior thalamic nucleus (Fig. 4) (Hoogland et al., 1991; Bourassa et al., 1995; Veinante et al., 2000b). Functional aspects The VPL and VPM are the primary thalamic relays for somatic sensory, i.e., nociceptive and tactile/kinestetic, information from the body and the head, respectively. For a comprehensive treatment of the pain system and the role of the thalamus in transmitting and modulating pain signals, see Chapter 27 by Willis et al. in this volume. In the rat VPM, the barreloid area representing the whisker field occupies a large part of the nucleus, signifying the importance of the rodent whisker system for navigation and exploration of the environment. This is in part an active process during which the whiskers are being moved in exploratory activities. It has been hypothesized that one of the two functionally different “channels” that reach the VPM, i.e., the pathway that originates in the spinal trigeminal complex, may play a role in this active “whiskering,” while the second channel originating in the principal trigeminal nucleus conveys information about passive deflections (Pierret et al., 2000). As indicated above, there exists an almost exclusive one-to-one relationship of individual whiskers with single barreloids in the ventral posteromedial nucleus. However, for the whisker system to function as an integrated system, which is of course necessary in order to interprete the detailed environmental information, integration of information from the different whiskers is taking place at several levels along the ascending pathway from the periphery to the barrel cortex. The ventral posteromedial nucleus is thought to play an important role in this process. This is signified by the complex interrelationships that exist between the VPM and the barrel cortex, as well as by the differentiated position of the reticular thalamic nucleus. For example, as indicated above, whereas individual barreloids project to single cortical barrels, return projections from the cortex take the form of rods in the VPM, “interconnecting” barreloids representing several or all whiskers in an arc (Hoogland et al., 1987; Bourassa et al., 1995). In the reticular and posterior thalamic nuclei efferents from a row of cortical barrels converge to a common termination site (Welker et al., 1988). The reticular nucleus, sending at least two types of fibers with distinct distribution patterns to the VPM, plays a signifi-

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FIGURE 3 Camera lucida reconstruction of the intrathalamic axonal arborizations of a corticothalamic fiber arising from a pyramidal neuron in the lower part of layer VI of the barrel cortex (primary somatosensory cortex). Individual axons were filled following small injections of biocytin in specific layers of the cortex, filling only a small number of neurons of which the projection fibers could be selectively traced. Note the collaterals in the reticular thalamic nucleus (TR) and the extensive arborizations in the posterior thalamic nucleus (Pom). Terminal fibers in the ventral posterior medial nucleus (VPm) take the form of a “rod.” This fiber and its terminals are reconstructed from several horizontal sections through the thalamus. The drawing in the right-hand corner shows the location of the terminals in a horizontal section of the thalamus. Slightly modified from Fig. 5 of Bourassa et al. (1995).

cant and differentiated role in the modulation of information at the level of the VPM (Cox et al., 1996). There is an extensive body of literature about the electrophysiological and functional aspects of the thalamic relay of somatic sensory information in rats, in particular in relation to the whisker system, but further elaboration on this subject is beyond the scope of this chapter. For a more comprehensive view on this subject, the reader is referred to recent reviews (e.g., Moore et al., 1999; Ahissar and Zackenhouse, 2001). Posterior Nucleus The posterior thalamic nucleus (Po; also indicated as posteror complex) is situated in the caudal part of the

thalamus, bordered caudally by the pretectal nuclei. In its caudal aspect, the Po is situated medial to the posterior intralaminar (PIL) and the medial geniculate (MGM) nuclei, more rostrally this position is taken by the ventral posterior complex, in particular the ventral posteromedial nucleus (Figures 30–42 in Paxinos and Watson, 1998). Dorsally, the Po is bordered by the lateral dorsal and lateral posterior nuclei; medial to the Po the intralaminar nuclei are situated. The posterior nucleus in rats is a heterogeneous area, in particular in its caudal aspects. Within the region generally considered to belong to the Po, several subnuclei have been identified on the basis of differential staining patterns, e.g. the ethmoid, scaphoid, and retroethmoid nuclei

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FIGURE 4 Camera lucida reconstruction of the intrathalamic axonal arborizations of a corticothalamic fiber arising from a pyramidal neuron in layer V of the barrel cortex (primary somatosensory cortex). The axons were filled following small injections of biocytin in the barrel cortex, filling only a limited number of neurons of which the projection fibers could be individually traced. The terminal fibers in the thalamus are issued as collaterals from axons that are directed toward the brain stem. There is strong clustering of axon terminals in the posterior thalamic nucleus (Po) and a few collaterals reach the central lateral nucleus (CL) of the intralaminar complex. Note the absence of collateral fibers in the reticular thalamic nucleus (TR). The drawing in the right-hand corner shows the location of the terminals in a horizontal section of the thalamus. Slightly modified from Fig. 8A of Bourassa et al. (1995).

(Paxinos et al., 1999). The Po appears as a relatively cell-sparse area in Nissl-stained sections, standing out against the cell-dense ventral posteromedial nucleus medially. In acetylcholineseterase-stained sections, the Po shows moderate activity for this enzyme and also on the basis of this staining the posterior nucleus can be reasonably well-delineated from its neighboring nuclei (Plates 31, 33, 35, and 37 in Paxinos and Watson, 1998). With respect to calcium-binding proteins, the Po contains a population of calbindin D28K-positive neurons, while calretinin and parvalbumin are present only in a light plexus of fibers (Arai et al., 1994; e.g., Figs. 195, 201, 202, 208, and 209 in Paxinos et al., 1999). Afferent and efferent connections Like the ventral posterior complex, the posterior complex receives a significant input from the spinal cord and brain stem trigeminal complex (Cliffer et al., 1991; Chiaia et al., 1991a, 1991b; see also Tracey, Chapter 25, and Waite,

Chapter 26, this volume). However, most of these ascending projections are less dense and they terminate in a more diffuse way in the Po than in the adjacent ventral posterior nuclei (e.g., Chiaia et al., 1991a; Villanueva et al., 1998). Moreover, projections to the posterior nucleus from the spinothalamic tract and the spinal trigeminal nucleus appear to be relatively more dense than those from the dorsal column nuclei and the principal trigeminal nucleus (Chiaia et al., 1991a; McAllister and Wells, 1981). Further inputs to the Po come from the inferior colliculus (LeDoux et al., 1987) as well as from the vestibular nuclei (Shiroyama et al., 1999). Reciprocal connections have been described between the posteromedial part of the Po and the region of the nucleus ruber (Roger and Cadusseau, 1987). Recent studies have demonstrated that the trigeminal complex gives rise to several types of fibers ascending to the thalamus which differentially distribute over the ventral posteromedial nucleus and the Po, as well as

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over other extrathalamic nuclei. While most fibers originating from the principal trigeminal nucleus terminate in a very specific manner in the barreloids in the ventral posteromedial nucleus, a smaller contingent of principal trigeminal fibers ascends to the Po. In addition, these fibers give off collaterals to the tectum, the zona incerta, the ventral part of ventral posteromedial nucleus, and the medial part of medial geniculate nucleus. The neurons in the principal trigeminal nucleus giving rise to the projections to the Po are sensitive to stimulation of multiple whiskers, in contrast to those that project to the barreloids in the ventral posteromedial nucleus which transmit signals from single whiskers only (Veinante and Deschenes, 1999). Thalamic projection fibers originating in the spinal trigeminal complex can also be categorized in several types: one type of fiber projects primarily to the ventral part of ventral posteromedial nucleus; the other type of fiber projects predominantly to the caudal part of the Po while giving off collaterals to, among others, the tectum, the zona incerta, and the ventral lateral thalamic nucleus (Veinante et al., 2000a). The Po, like the ventral posterior complex, projects to the primary and secondary sensory cortices, largely following a somatotopical organization (Fabri and Burton, 1991). Thalamocortical fibers from the posterior nucleus terminate in the upper part of layer VI as well as in layer I of the cortex (Zhang and Deschenes, 1998). In the barrel cortex, the projection fibers from the Po do not terminate within the barrels, but they preferentially target the interbarrel areas (Lin and Lu, 1993). Cortical projections to the Po primarily arise from the somatosensory areas S1 and S2, but additional projections originate from motor, premotor (frontal eye field), and insular cortices (Veinante et al., 2000b; Guandalini, 2001). While S1 projects predominantly to dorsal parts of the posterior complex, S2 targets more ventral and medial parts, although there exist substantial areas of overlap (Shi and Cassell, 1998b; Veinante et al., 2000b). The corticothalamic projections to Po arise predominantly from layer V pyramidal neurons as collaterals from axons that project to the striatum and descend into the brain stem (Fig. 4) (Levesque et al., 1996; Veinante et al., 2000b). The fibers originating in layer V of the barrel cortex, possibly predominantly originating from neurons in interbarrel cortical areas, exhibit giant terminals of the RL-type in the Po and they may be considered drivers in this nucleus (see section “Some General Aspects of Thalamic Organization”; Hoogland et al., 1987, 1991; Bourassa et al., 1995; Wright et al., 2000; Veinante et al., 2000b). The other cortical areas mentioned above, as well as the deep layer VI neurons in the cortical barrel field, project fibers with much smaller boutons in the

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Po and these may be considered modulators (Welker et al., 1988; Hoogland et al., 1991; Bourassa et al., 1995). Functional aspects It may be clear that the Po receives ascending inputs from various different modalities (see above), i.e., somatosensory, auditory, visual, and vestibular. Although there is probably convergence within the posterior nucleus of fibers carrying information about these different modalities, the somatosensory modality strongly dominates. Whereas the Po might be considered a primary relay thalamic nucleus, the driving input from layer V of the sensory cortex places this nucleus also in the category of higher order thalamic nuclei (see section “Some General Aspects of Thalamic Organization”). As may be clear from the discussions of the afferents and efferents of both the ventral posteromedial nucleus and the Po, the whisker system in rats occupies a large part of the somatosensory system, and the functional anatomy of this system has been analyzed in great detail (e.g., Moore et al., 1999; Ahissar and Zackenhouse, 2001). Therefore, the most concrete suggestions for a functional role of the Po have been made with respect to the behavioral functions of the whisker system. The whisker system not only provides information about passive movements of individual or groups of whiskers but the whiskers are also actively moved in certain frequencies to “explore” the surroundings. The fact that the Po receives its main cortical input from layer V neurons probably indicates that this concerns an “efference copy” of decending (motor) cortical signals to the brain stem. This has led Veinante et al. (2000b) to the suggestion that the Po plays a role at the interface between the sensory and motor aspects of the whisker system. More specifically, the posterior complex receives the motor signals for active whisker movements as well as the sensory feedback. The Po, together with parts of the trigeminal complex, might thus be involved in monitoring the dynamics of the self-initiated whisker movements, a mechanism that may be necessary for a sensory organ such as the whiskers that lack proprioceptors (Veinante et al., 2000b). Whether the Po has other functions at the interface between the sensory and motor systems remains to be established. The reciprocal connections between the posterior nucleus and the red nucleus provide an indication in that direction (Roger and Cadusseau, 1987; Cadusseau and Roger, 1990; Arnault et al., 1994). Gustatory and Visceral Nuclei The ventromedial part of the ventral posterior complex represents the relay for gustatory and visceral information from the periphery to the insular cortex (Norgren and Leonard, 1973; Cechetto and Saper, 1987).

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The medial part of the ventral posterior complex contains neurons that are smaller than those in the other parts of the VP, and it is therefore referred to as the parvicellular ventral posterior nucleus (VPPC). The VPPC is ventrolaterally bordered by the medial lemniscus and extends medially just ventral to the parafascicular thalamic nucleus and, more rostrally, ventral to the paracentral and central medial nuclei. The VPPC is rostrally “replaced” by the ventromedial nucleus (see Plates 34–37 in Paxinos and Watson, 1998). The VPPC is a rather thin sheet of neurons and it is generally divided into medial and lateral parts, i.e., the parvicellular part of the ventral posteromedial nucleus and the parvicellular part of the ventral posterolateral nucleus (e.g., Cechetto and Saper, 1987; Shi and Cassell, 1998a). Since the medial part of VPPC relays gustatory information (Cechetto and Saper, 1987; Lundy and Norgren, Chapter 28, this volume), this part of the nucleus is indicated by some authors as the gustatory nucleus (e.g., Shi and Cassell, 1998a). Apart from a moderate number of calbindin D28Kpositive neurons, the VPPC only contains moderate to low amounts of calcium-binding proteins in fibers (Arai et al., 1994). The medial part of the VPPC contains a dense plexus of calcitonin gene-related peptide-containing fibers (Yasui et al., 1989). Afferent and efferent connections Major ascending inputs to the VPPC are derived from the parabrachial nucleus (Norgren and Leonard, 1973; Cechetto and Saper, 1987; Bester et al., 1999; Krout and Loewy, 2000a). Other brain stem inputs to the VPPC arise from specific parts of the spinal and principal trigeminal nuclei, as well as from the laterodorsal tegmental nucleus, locus coeruleus, nucleus of the tractus solitarius, the A5 region, and the cuneate nucleus (Krout et al., 2002). The VPPC, in turn, projects to the lateral and central nuclei of the amygdala, the amygdalostriatal transition area directly dorsal to the central amygdaloid nucleus, and the more rostrally located ventral parts of the caudate– putamen (fundus striati). Cortical projections are primarily directed at the granular and dysgranular insular areas of both the posterior insular and parietal insular cortices, while the anterior insular cortices receive only a minor projection from the VPPC (Kosar et al., 1986a, 1986b; Cechetto and Saper, 1987; Turner and Herkenham, 1991; Nakashima et al., 2000). In most of these efferent VPPC projections to cortical and subcortical targets a mediolateral topographical arrangement exists. Cortical afferents to the VPPC are primarily derived from the insular cortical areas to which it projects. Thus, the granular and dysgranular posterior insular cortices project predominantly to the medial VPPC, while the granular and dysgranular parietal insular cortices send fibers primarily to the lateral part

of the VPPC (Shi and Cassell, 1998a, 1998b). The rostrally located dysgranular and agranular insular cortices have only very few projections to the VPPC (Shi and Cassell, 1998a). Functional aspects The main function of the VPPC is the transfer of gustatory and visceral information to the granular and dysgranular insular cortices. These insular areas are situated between olfactory cortical areas more ventrally, i.e., around the rhinal sulcus, and the sensorimotor cortices involved with mouth/tongue, head, and forelimbs in the frontoparietal areas just dorsal to the insular cortices. Combined anatomical and physiological studies have demonstrated that the gustatory area is situated more medially in the VPPC, and the visceral area is located more laterally (Kosar et al., 1986a, 1986b; Cechetto and Saper, 1987). In a recent, detailed tracing study, it has been shown that the parabrachial projections to the medial part of the VPPC originate from the two main gustatory relay nuclei in the parabrachial complex. Based on light microscopic observations, the parabrachial fibers in the VPPC have large terminals, suggesting primary sensory relay of information (Bester et al., 1999). The subdivision of the VPPC into a medial gustatory and a lateral visceral part is in line with the functional topography that has been established in the granular and dysgranular posterior insular cortices (Kosar et al., 1986a, 1986b; Cechetto and Saper, 1987; Shi and Cassell, 1998a, 1998b). Behavioral studies, using specific lesions of the gustatory part of the VPPC, suggest that this thalamic region is not essential for taste discrimination per se, but rather that it is important for more complicated behavioral functions in which the performance is dependent upon “manipulation” of gustatory information (Reilly and Pritchard, 1997). This could be in line with anatomical observations that parabrachial projections bypass the thalamus to reach directly the amygdala and insular cortices (Shipley and Sanders, 1982; Yasui et al., 1989). For a more comprehensive review of the rat gustatory system and the role of the thalamus in this system, see Chapter 28 by Lundy and Norgren in this volume. Medial Geniculate Nucleus The medial geniculate nucleus (MG) forms the most caudal extension of the thalamus and its caudal half is situated laterally alongside the mesencephalon. For a detailed discussion of the auditory thalamus see Malmierca and Merchán (Chapter 31, this volume). The rostral part of the MG is located ventromedially to the lateral geniculate complex. The MG is the principal auditory relay nucleus of the thalamus and consists of several subnuclei that all have different functions within

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the auditory system. The medial geniculate nucleus can be subdivided into medial (MGM), ventral (MGV), and dorsal (MGD) subnuclei (Jones, 1985; Clerici and Coleman, 1990). The “auditory” thalamus further consists of a number of smaller nuclei that are positioned medially and ventromedially to the main MG complex, i.e., the suprageniculate nucleus (SG), the posterior limitans thalamic nucleus (PLi), the posterior intralaminar thalamic nucleus (PIL), and the lateral part of the parvicellular subparafascicular nucleus (SPF; see section “Midline and Intralaminar Thalamic Nuclei”). A marginal zone (MZMG) “covers” the dorsal, lateral, and ventral aspects of the MGD and MGV. The MGD and MGV are separated by the so-called midgeniculate bundle which is mainly derived from the inferior colliculus. The MGV can be further subdivided into a ventral and an ovoid part based on fiber architectonics and cytoarchitectonics (Clerici and Coleman, 1990). The main cell type in MGV is small- to medium-sized and has bushy tufted dendrites forming fibrodendritic laminae that are regularly oriented in association with the fiber bundles from the inferior colliculus in a dorsolateral to ventromedial direction (Winer et al., 1999a). In the oval part of the MGV, the orientation of cells and colliculular afferent fiber bundles is more spiral-like (Clerici and Coleman, 1990). The MGD is rather heterogeneous in its neuronal composition and can be subdivided into several subnuclei. Neurons in the MGD have radiating, tufty dendrites (Winer et al., 1999a). Also in the MGM, the neuronal population is rather heterogeneous, ranging from small to magnocellular. Few interneurons (about 1%) have been identified in the MG complex, most of them being GABAergic (Winer and Larue, 1988; Winer et al., 1999a). Afferent and efferent projections Ascending afferents to the MG primarily originate in the inferior colliculus. While the MGV receives its afferents mainly from the central nucleus of the inferior colliculus, the MG complex as a whole collects fibers from the cortices of the inferior colliculus. Like the MGV, the MGM receives, in addition, collicular fibers from the central nucleus (Jones, 1985; LeDoux et al., 1987). The central nucleus of the inferior colliculus is strictly tonotopically organized, and the MGV transfers this highly organized information to the primary auditory cortex Te1 in the temporal lobe (Winer et al., 1999b). As such, the MGV, in particular the ovoid subnucleus, forms the main thalamic input for area Te1. The projections from the MGV terminate primarily in layers III and IV of Te1, but there are also projections to the junction of layers V and VI and to superficial layer I (Romanski and LeDoux, 1993; Cetas et al., 1999; Winer et al., 1999b). The projections from the MGV in Te1 are strongly

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convergent, highly topographic, and spatially focal, and they originate from only one type of neuron in the MGV. Its caudal part excepted, the MGV does not project outside area Te1. The cortical projections from the MGM and the MGD are more divergent, not only directed toward Te1 but, in a much denser pattern, also to surrounding cortical areas Te2 and Te3. These thalamocortical projections arise from several types of geniculate neurons and mostly terminate in the middle cortical layers (Roger and Arnault, 1989; Arnault and Roger, 1990; Winer et al., 1999b). Corticothalamic fibers from the temporal cortical areas Te1 (primary auditory cortex) and Te2 and Te 3 (together indicated as the “auditory belt” cortex) terminate in different parts of the medial geniculate complex (Shi and Cassell, 1997). Cortical area Te1 projects primarily to the MGV and more moderately to the ventral part of the MGD. Area Te3 sends its most dense projections to the dorsal part of the MGD, directly adjacent to the lateral posterior nucleus which also receives a dense input from Te2 (Shi and Cassell, 1997). Strikingly, the ovoid subnucleus of MGV is largely avoided by these corticothalamic projections, indicating a lack of reciprocity in the cortical connections of this part of the MGV. At the ultrastructural level, corticothalamic projections arising from the primary auditory cortex terminate as small and large (“giant”) terminals in the medial geniculate complex. The smaller, most numerous corticothalamic terminals are present throughout the MG complex, the larger terminals have been observed in the ventral part of the MGD (Rouiller and Welker, 1991) as well as in the dorsal part of the marginal division (MZMG; Bartlett et al., 2000). Excitatory collicular terminals are rather variable in size and include small and large profiles, the larger ones being predominant in the MGV (Bartlett et al., 2000). This heterogeneity in the morphology of the ascending, colliculogeniculate projections indicates a higher complexity in the auditory pathways than in the ascending projections to other sensory thalamic nuclei. Furthermore, part of the collicular afferents of the MGV and MGD appear to be inhibitory (Bartlett and Smith, 1999). Yet, the general organization of ascending driving inputs and descending modulatory cortical inputs seems to hold also for the MGV. Driving cortical inputs from the primary auditory cortex may reach parts of the medial geniculate complex (MGD and MZMG) other than that from which it receives its primary thalamic input, i.e., the MGV (see section “Some General Aspects of Thalamic Organization”). All medial geniculate nuclei, except the MGV, project to other temporal association cortices, like the perirhinal cortex, as well as to the amygdala. A common feature of the medially located MGM, SG, and PIL nuclei is that

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they target the upper part of layer I of the temporal association cortices, in addition to more specific projections to deeper layers of these cortices (Linke and Schwegler, 2000). The SG, in addition, projects to the medial agranular frontal cortex (Kurokawa and Saito, 1995). Projections to the amygdala, primarily terminating in the lateral nucleus and, to a lesser degree, the basal nuclei, arise from the medially located subnuclei MGM, SG, and PIL and the lateral part of the parvicellular subparafascicular nucleus (LeDoux et al., 1985, 1990; Turner and Herkenham, 1991; Namura et al., 1997; Doron and LeDoux, 1999, 2000). Apart from projections to the amygdala, the medial subnuclei of the medial geniculate complex also have reciprocal connections with basal ganglia structures. In particular the caudal parts of the striatum, including the amygdalostriatal transition zone, and the caudal globus pallidus participate in these connections with the MGM, SG, PIL, and lateral subparafascicular nucleus (Moriizumi and Hattori, 1992; Shammah-Lagnado et al., 1996). The MGM, as well as the SG and PIL, not only receives ascending auditory information from the inferior colliculus but is also reached by fibers from the superior colliculus (Linke, 1999). In addition, the MGM receives direct projections from the dorsal cochlear nucleus, bypassing the inferior colliculus. In view of the fact that both the superior colliculus and the dorsal cochlear nucleus transfer multimodal information, it seems likely that the medially located subnuclei of the MG complex do not only process auditory information (Malmierca et al., 2002). Functional aspects of the medial geniculate complex It is evident from anatomical and physiological data that the MGV is the main thalamic relay to the primary auditory cortex, subserving the specific tonal analysis of sounds (e.g., LeDoux et al., 1987; Romanski and LeDoux, 1993; Bordi and LeDoux, 1994a; Winer et al., 1999b). The functions of the MGD are less clear, but are most probably concerned with nontonal aspects of sounds and the integration with other sensory modalities. The projections from the inferior colliculus stem from a part of the colliculus that is not tonotopically organized and in the MGD the projections to the temporal cortices are also less strictly organized, terminating primarily in the auditory association cortices (LeDoux et al., 1987; Clerici and Coleman, 1990). The superficial, dorsal region of the MGD may even be a visual-recipient rather than an auditory-recipient area of the medial geniculate complex (Sun et al., 1996; Shi and Cassell, 1997). For a more detailed discussion of the functional aspects of the auditory thalamus, see Chapter 31 by Malmierca and Merchan in this volume. The functions of the MGM, as well as those of the

PIL, SG, and SPF nuclei, must be interpreted in the context of their connections with the temporal association cortices, the amygdala, and the basal ganglia (see above). In particular the relationships of these MG nuclei with the amygdala have been the focus of electrophysiological and behavioral studies. It is clear from such studies that these pathways, which connect the auditory system primarily with the limbic association areas of the brain, are associated with the emotional and mnemonic aspects of sounds. The extensive and elegant functional anatomical studies of LeDoux and colleagues have demonstrated that the MGM, the PIL, and the lateral part of the parvicellular subparafascicular nucleus play an important role in the association of emotionally negative, e.g., noxious, stimuli and the context in which these stimuli occur (fear conditioning; LeDoux, 1993, 2000). The convergence of auditory and somatosensory inputs, but possibly also of visual inputs via the superior colliculus (Linke et al., 1999), in these nuclei of the MG complex appear to form the neuronal basis for such associations that are behaviorally expressed via the amygdala (LeDoux et al., 1987; Bordi and LeDoux, 1994a, 1994b; Linke et al., 1999; LeDoux, 2000). In the context of fear conditioning, plastic changes have been demonstrated in both the MG complex and the amygdala (LeDoux, 2000; Maren et al., 2001; cf. also Komura et al., 2001). The MGM and the PIL have also been shown to form a crucial link in pathways that lead to the neuroendocrine expression of audiogenic stress via the amygdala and, ultimately, the hypothalamic paraventricular–hypophysis system (Campeau et al., 1997; Campeau and Watson, 2000). Thus, the medially located nuclei of the medial geniculate complex are important for the association of auditory and other sensory inputs and the “translation” of this information, via the amygdala and temporal association cortices, in behavior and emotional reactions. Finally, it is important to note that several nuclei of the medial geniculate complex not only project to the cerebral cortex or subcortical forebrain structures, but that in particular the medially located MGM, PIL, SPF, and SG also have descending projections to the inferior colliculus (Senatorov and Hu, 2002; Winer et al., 2002). Thus, parallel to descending cortical projections to brain stem auditory centers, these medially located MG nuclei are also in a position to provide feedback to the early relays in the ascending auditory pathways.

Motor Nuclei Vental Lateral and Ventral Anterior Nuclei The ventral lateral (VL)–ventral anterior (VA) complex occupies an extensive nuclear area intercalated between the ventral posterior nuclei ventrolaterally,

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the posterior and lateral dorsal nuclei dorsally, and, adjacent to its rostral extension, the intralaminar and anterior nuclei dorsomedially. Ventromedially, the VA/VL complex abuts the ventromedial nucleus (VM) (Figs. 25–32 in Paxinos and Watson, 1998). Cytoarchitectonically, the VA/VL complex is cell sparse and contains relatively large neurons. In rats, a distinction between the ventral anterior and the ventral lateral nuclei on the basis of cytoarchitectonics is difficult to make (Jones, 1985). In acetylcholinesterase-stained sections the VA exhibits very low activity for this enzyme and is therefore easily delineated from its surrounding nuclei, e.g., the reticular nucleus and the anterior complex (Plate 26 in Paxinos and Watson, 1998). In the VL, acetylcholinesterase activity is slightly higher than that in the VA. Calcium-binding proteins are sparse in both the VA and the VL (Arai et al., 1994). Neurons containing calbindin D28K are present in some parts of both nuclei, parvalbumin is only present in a moderately dense fiber plexus in VA and VL (e.g., Figs. 173, 174, and 180 in Paxinos et al., 1999). Afferent and efferent connections The VL/VA complex receives afferents mainly from the cerebellum and the basal ganglia, and it is in reciprocal contact with somatomotor and premotor cortical areas. Additional subcortical inputs derive from the vestibular nuclei (Shiroyama et al., 1999). Cerebellar inputs originate from the deep cerebellar nuclei (Faull and Carmen, 1978; Angaut et al., 1985; Sawyer et al., 1994a, 1994b; Aumann et al., 1994, 1996). The cerebellar projections stem primarily from the lateral and interpositus nuclei and they spread virtually over the entire VL. There exists a topographical arrangement in the cerebellothalamic pathways such that the dorsoventral and mediolateral axes of the lateral cerebellar nucleus correspond to the dorsoventral and mediolateral axes in the VL, respectively (Angaut et al., 1985). In a single axon tracing study, Aumann et al. (1996) observed different types of cerebellothalamic fibers. Although all axons project to both the ventral lateral and the intralaminar and adjacent thalamic nuclei, within the VL one type of fiber has very restricted, clustered terminal areas, whereas a second fiber type has a more widespread and diffuse termination pattern. In some cases the latter fibers even reach the VL in both hemispheres (Aumann et al., 1996). Cerebellothalamic inputs are most probably the driving inputs for the ventral lateral nucleus since their terminals exhibit large round boutons with closely packed small round vesicles, i.e., resembling the RL-type terminal (Sawyer et al., 1994a; see section “Some General Aspects of Thalamic Organization”). This morphology fits with the excitatory nature of the cerebellothalamic projections

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that have been established in electrophysiological experiments (e.g., Sawyer et al., 1994b). The large terminals originating from the cerebellum make up only a minority of the presynaptic terminals in VL, the majority being of the RS-type originating from the cerebral cortex. A third category is formed by mediumsized boutons with pleomorphic vesicles forming symmetrical synapses. The latter represent most probably the GABAergic terminals from the reticular thalamic nucleus and the substantia nigra pars reticulata and/or the entopeduncular nucleus (Sawyer et al., 1994a). However, in an anterograde tracer study combined with electron microscopy Finkelstein et al. (1996) could not confirm the symmetrical morphology of entopeduncular terminals in VA/VL but, instead, demonstrated asymmetrical synapses with round vesicles to be associated with pallidal projections. In line with the latter finding, it has recently been shown that cholinergic and glutamatergic fibers possibly make up a large part of the nigral and pallidal projections to the VL (as well as to the ventral medial nucleus), although a GABAergic projection could also be established (Kha et al., 2000, 2001). The basal ganglia input to the VA/VL complex, represented by the projections from the entopeduncular nucleus and the pars reticulata of the substantia nigra, is concentrated in the rostral and medial parts of the complex, i.e., the area corresponding mostly to VA (Sakai et al., 1998). In agreement with findings in other species, there appear to be only limited areas of overlap of cerebellar afferents with entopeduncular and nigral terminations in the VA/VL complex of rats (Carter and Fibiger, 1978; Deniau et al., 1992). The VA/VL complex receives a third strong input from the somatomotor cortex and, in addition, from a wide array of frontal and parietal cortical areas. In particular the rostromedially located ventral anterior nucleus is the target of (pre)frontal cortical afferents (e.g., Sesack et al., 1989; Conde et al., 1990; Reep and Corwin, 1999; Vertes, 2002). As mentioned above, these cortical inputs are most likely the source of the RS-type of inputs, i.e., modulatory inputs. In return, the ventral lateral nucleus is the main thalamic source of inputs to the somatomotor cortex, but the VL also projects to premotor cortical areas (Aldes, 1988). These fibers target layers III–V, but they terminate most densely in superficial layer I (Yamamoto et al., 1990; Aumann et al., 1998; Mitchell and Cauller, 2001). Functional aspects The VA/VL complex must be considered the main motor thalamic relay to the cerebral cortex. The largest volume of VA/VL is devoted to the cerebellar influences on the motor thalamocortical system. This cerebellothalamocortical pathway is most

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probably devoted to the coordination of multijoint movements (Sawyer et al., 1994b). The area of the VA/VL receiving basal ganglia inputs is smaller, including predominantly rostral and medial parts of the complex, i.e., the area that is best characterized as the ventral anterior nucleus (Sakai et al., 1998). The basal ganglia and thalamocortical system are involved in a number of parallel, functionally segregated circuits (e.g., Deniau et al., 1994; Groenewegen et al., 1990, 1999). In the circuits that are closest related to motor functions in rats, the VM is generally considered to be the thalamic relay. However, as discussed below (see following section), the VM might have a different position in these circuits than generally assumed: as a result of the rather widespread cortical projections the VM presumably plays an integrating role for the various parallel, functionally segregated basal ganglia–thalamocortical circuits (Groenewegen et al., 1999). In that respect, it would be interesting to gain more insight into the “position” of the ventral anterior nucleus in these circuits. The VA nucleus has a projection to the (pre)motor cortical areas more concentrated than that of the VM and may therefore be considered as the thalamic relay in the motor basal ganglia–thalamocortical circuit (Sakai et al., 1998). Ventral Medial Nucleus The ventromedial thalamic nucleus (VM) is a slender, rostrocaudally elongated nucleus in the rostromedial part of the ventral thalamic complex. Medially, the VM can be clearly distinguished from the cell-sparse nucleus submedius. The mammillothalamic tract ascends just medially to and through the most medial parts of the VM. Dorsally and laterally the VM is bordered by the VA and VL nuclei, as well as, more caudally, by the ventral posterior lateral and ventral posterior medial (VPL/VPM) complex (Figs. 26–34 in Paxinos and Watson, 1998). The VM can be distinguished from these nuclei in Nissl-stained sections since it is made up of slightly smaller neurons that are more densely packed (see also Herkenham, 1979). The presently defined VM largely corresponds with the basal VM as defined by Jones (1985). In acetylcholinesterase-stained sections the ventral medial nucleus only exhibits low activity. With respect to calcium-binding proteins, the VM contains calbindin D28K-positive neurons throughout the nucleus while calretinin-positive neurons are restricted to its medial part (Arai et al., 1994; e.g., figures 174 and 175 in Paxinos et al., 1999). A low density of calbindin D28K- and parvalbumin-positive fibers is present in the ventral medial nucleus. Afferent and efferent connections Subcortical inputs to the VM originate to a large degree in the entope-

duncular nucleus (the internal segment of the globus pallidus) and the substantia nigra pars reticulata. These afferents preferentially target medial parts of the VM, while the lateral part is less densely innervated (Carter and Fibiger, 1978; Herkenham, 1979; Deniau et al., 1994; Groenewegen et al., 1999). Recently, a strong projection from the medullary subnucleus reticularis dorsalis to the VM, with a dominance for its lateral part, has been described (Villanueva et al., 1998). Other substantial subcortical inputs are derived from the deep layers of the superior colliculus, the deep mesencephalic nucleus, the periaqueductal gray matter, the raphe nuclei, the peripeduncular region, the laterodorsal tegmental nucleus, the locus coeruleus, the parabrachial region, and the trigeminal complex (Herkenham, 1979; Krout and Loewy, 2000a, 2000b; Krout et al., 2001, 2002). Afferents from the cerebellum have also been identified (Angaut et al., 1985). Cortical afferents of the VM originate in widespread cortical areas of the frontal lobe (Herkenham, 1979; Seseack et al., 1989; Hurley et al., 1991; Vertes, 2002). The projections from the ventral medial nucleus reach widespread cortical areas, in particular in the rostral, frontal part of the hemisphere, but to a lesser degree also in more caudal areas (Herkenham, 1979; Conde et al., 1990; Reep et al., 1996). The laminar distribution of the thalamocortical fibers arising in the VM is remarkable in that they reach primarily the most superficial part of layer I of almost the entire cortical mantle (Herkenham, 1979; Desbois and Villanueva, 2001; Mitchell and Cauller, 2001). Some VM fibers reach also the deeper layers III and V of the dorsolateral part of the frontal cortex, including the motor cortex (Arbuthnott et al., 1990). Functional aspects Together with the VA/VL complex, the VM is in general considered to be part of the motor thalamus. In view of its position between, on the one hand, the basal ganglia output structures, i.e., the entopeduncular nucleus and the substantia nigra reticulata and, on the other hand, the frontal cortex including motor and premotor cortices, this appears as a logical assumption. However, as has already been concluded by Herkenham in 1979, the VM has a rather widespread projection to layer I of a large part of the cortical mantle, in particular in the frontal cortical areas. It might therefore be conceived that the VM plays a particular integrative rol. The ventral medial nucleus projects to the superficial layers of all (pre)frontal cortical areas that form the starting point for the above-mentioned (see page 422) basal ganglia–thalamocortical systems, indicating that the VM potentially has an influence on all these functionally distinct circuits (Groenewegen et al., 1999).

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This influence might be of attentional significance, preparing the various circuits for a specific motor or behavioral output. In this respect, it is of interest to note that the projection from the substantia nigra to the VM primarily originates from the dorsomedial aspect of the pars reticulata, an area which receives strong inputs from the core of the nucleus accumbens (Deniau and Chevalier, 1992; Deniau et al., 1994, 1996; Montaron et al., 1996). The nucleus accumbens has also been implicated in attentional mechanisms. This proposed attentional functional role of the VM may even be further specified in that the VM receives a strong input from caudal reticular brain stem nuclei conveying nociceptive signals (Villanueva et al., 1998). It has recently been confirmed that the VM transmits this nociceptive information from the whole body surface to the frontal cortex (Monconduit et al., 1999). This might imply that the VM is important for allowing pain signals to gain direct access to widespread cortical areas. A possible interpretation is that a fast access route for vital signals serves to prime the cerebral cortex for attentional reactions that should lead to coordinated motor and behavioral responses in reaction to painful stimuli (Monconduit et al., 1999; cf. also Herkenham, 1980). A similar role could be played by the caudal ventral medial nucleus with respect to deep pain (Floyd et al., 1996). It is of interest to note that in his caudal region of the VM calbindin D28K-positive neurons might be involved (Floyd et al., 1996; cf. also Jones, 1998).

ASSOCIATION THALAMIC NUCLEI Mediodorsal Nucleus The mediodorsal thalamic nucleus (MD) is situated medially and dorsally to the internal medullary lamina of the thalamus in which the intralaminar nuclei are embedded and ventrally to the stria medullaris/ habenular complex. In between the left and the right MD nuclei, the midline paraventricular (PV) and intermediodorsal (IMD) nuclei are situated. The mediodorsal nucleus is bordered rostrally by the parataenial nucleus (PT) and caudally by the parafascicular nucleus (PF), two nuclei also belonging to the midline/intralaminar thalamic nuclei. Therefore, the MD is virtually “encircled” by the midline and intralaminar thalamic complex with which it shares inputs from many cortical and subcortical sources (cf. Section VI). The MD in rats is generally subdivided into three parts, the medial (MDM), central (MDC), and lateral (MDL) segments (Leonard, 1969, 1972; Krettek and Price, 1977). The distinction between these segments is based on

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cytoarchitectonics and fiber architectonics, as well as on distinct input–output relationships (see below). The central segment is richest in myelinated fibers and, in that respect, stands out against the two other segments. The lateral segment has the highest activity for acetylcholinesterase (Plate 31 in Paxinos and Watson, 1998). The differential distribution of calcium-binding proteins also confirms a distinction between the three MD segments. This is most apparent with respect to calbindin D28K, which is present in both cell bodies and fibers in the medial and lateral segments, but much less so in the central segment (Arai et al., 1994; see also Figs. 181, 188, and 195, Paxinos et al., 1999). Stellate and fusiform cells make up the majority of the neuronal population of the MD, both considered to be thalamocortical neurons (Kuroda et al., 1992, 1998). Stellate cells are most abundant in the MDC, whereas in the MDM and the MDL fusiform neurons are relatively more numerous. Interneurons are very rare if existent at all in the rat mediodorsal nucleus. Dendrites of the stellate and fusiform neurons largely remain within the segment in which their parent cell body resides, stressing the morphological distinction between the three segments (Kuroda et al., 1992). Further heterogeneities are present within the three segments. On the basis of differences in fiber densities and connections, Ray and Price (1992) suggested that the MDL may be further subdivided into a dorsolateral and a ventrolateral part. Groenewegen (1988), primarily based on fiber connections, recognized a paralamellar part of the MD, comparable with a similar segment in primates. In general, the MDM and MDC together are thought to be homologuous with the medial magnocellular segment of the primate MD, and the rat MDL may be homologuous with the primate parvocellular, lateral segment of the mediodorsal nucleus. Afferent and Efferent Projections The different segments of the MD are characterized by different input–output relationships. The MD as a whole is reciprocally connected with the frontal cortex rostral to the motor and premotor cortices. The MD– cortical projections are considered to define the “prefrontal cortex” (cf. Uylings and Van Eden, 1990). In this respect, the prelimbic and dorsal anterior cingulate (Cg1) areas, located in the medial wall of the hemisphere, as well as the dorsal and ventral agranular insular areas, situated rostrally in the dorsal bank of the rhinal sulcus, receive the strongest input from the MD (e.g., Leonard, 1969, 1972; Groenewegen, 1988; Conde et al., 1990; Ray and Price, 1992). However, also the infralimbic area, the various different orbital areas, the ventral anterior cingulate (Cg2) area, and the medial agranular cortex (“shoulder” cortex) receive a consid-

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erable input from the MD (Groenewegen, 1988; Ray and Price, 1992; Reep et al., 1996). The MD–thalamocortical projections are to a large degree reciprocated by corticothalamic projections with the notable difference that the latter distribute bilaterally while the thalamocortical projections are strictly unilateral. There exists a striking specificity in the reciprocal MD–prefrontal connections, which is as follows. The rostral part of the MDM is primarily connected with the ventral prelimbic and infralimbic areas, while the caudal part of the MDM communicates with the dorsal agranular insular area. The MDC is reciprocally connected with the ventral agranular insular area. Finally, the MDL has reciprocal connections with the dorsal prelimbic, dorsal anterior cingulate and medial agranular areas (Groenewegen, 1988; Conde et al., 1990; Ray and Price, 1992; Guandalini, 2001; Vertes, 2002; Heidbreder and Groenewegen, 2003). However, relationships with these (prefrontal) cortical areas are by no means “exclusive” for the mediodorsal nucleus. The dorsal anterior cingulate, prelimbic, and infralimbic areas project to and receive input from a rather extensive medial zone of the thalamus. This medial thalamic zone encompasses, in addition to the MD, parts of the midline and intralaminar complex, prominently including the nucleus reuniens, and further the anteromedial (AM), submedius (Sub), and ventromedial (VM) nuclei, as well as the medial parts of the lateral dorsal (LD) and lateral posterior (LP) nuclei (see also corresponding sections in this chapter; Conde et al., 1990; Reep et al., 1996; Vertes, 2002). Likewise, the orbital areas have reciprocal relationships with different segments of the MD, but these areas are also extensively connected with the–nucleus submedius–(see below) and, to a lesser degree, the VM (Ray and Price, 1992; Reep et al., 1996). Anatomical and electrophysiological experiments have shown that the reciprocal corticothalamic MD projections use excitatory amino acids as neurotansmitters (Ray et al., 1992; Pirot et al., 1994; Kuroda et al., 1998). The projections from the MD to the prefrontal cortex terminate predominantly in layer III and to a lesser degree in layer I of the cortex, while the return projections to the ipsilateral mediodorsal nucleus originate in layer VI (Krettek and Price, 1977; Groenewegen, 1988). Nevertheless, the MD thalamocortical neurons are in direct reciprocal contact with the layer VI pyramidal neurons that give rise to the corticothalamic projections. The thalamocortical neurons contact also pyramidal neurons in layer III that give origin to projections to the contralateral MD (Kuroda et al., 1993, 1998). The reciprocal circuitry between the mediodorsal nucleus and the prefrontal cortex may be modulated by dopaminergic inputs from the ventral tegmental area, as well as through local GABAergic cortical interneu-

rons that, in turn, receive direct excitatory inputs from the mediodorsal nucleus (Kuroda et al., 1998; Fig. 5). The MD receives inputs from a wide variety of subcortical sources (Groenewegen, 1988; Ray et al., 1992), many of which have been characterized at the ultrastructural level (for review, Kuroda et al., 1998). The MDM receives fibers from the amygdala, different nuclei in the basal forebrain, like the nucleus of the diagonal band of Broca and the ventromedial part of the ventral pallidum, the lateral preopic area, the ventral tegmental area, and several other brain stem nuclei. Amygdaloid inputs to the MDM derive from the central, cortical, and basal nuclei of the amygdaloid complex (Reardon and Mitrofanis, 2000), and these fibers form RL-type terminals establishing asymmetric synapses on complex dendritic appendages and large dendritic shafts from which spines arise (glomeruli; Kuroda and Price, 1991a). No data are available on the possible transmitter content of the amygdaloid inputs. In the MDC, terminals similar to those described for the amygdaloid inputs to the MDM and containing neurotensin have been identified originating from the prepiriform cortex (Kuroda and Price, 1991a; Ray and Price, 1990). Both the MDC and the MDM further receive inputs from pallidal neurons located in the deep layers of the olfactory tubercle (Groenewegen, 1988). Ventral pallidal terminals (in the MDM and the MDC) are large, contain pleomorphic vesicles, and establish symmetric synapses (Kuroda and Price, 1991a). These terminals are for the most part GABAergic although also other transmitters may be used in the projections from this heterogeneous basal forebrain region (Ray et al., 1992; Churchill et al., 1996). The MDL primarily receives inputs from brain stem structures like the reticular part of the substantia nigra, the deep layers of the superior colliculus, and the laterodorsal tegmental nucleus. Weaker inputs derive from the medial part of the globus pallidus (Groenewegen, 1988). Inputs from the reticular substantia nigra are GABAergic and have ultrastructural characteristics very similar to those of the pallidothalamic fibers (Kuroda and Price, 1991a). By contrast, collicular inputs establish small, asymmetric contacts of the RS-type. The afferents from the laterodorsal tegmental nucleus to the MDL, on the other hand, are large with round vesicles and they establish asymmetric RL-type contacts with proximal dendrites. These tegmental afferents most probably use acetylcholine as neurotransmitter. At the ultrastructure level, the inputs from layer VI of the prefrontal cortex to the mediodorsal nucleus are of the RS-type, the most numerous type of terminal located primarily on the more distal parts of the dendrites (Kuroda and Price, 1991b; Kuroda et al., 1998). All MD segments receive input from the reticular thalamic nucleus (Rt), terminals

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FIGURE 5 A diagrammatic representation of the synaptic connections between prefrontal cortical (PFC) neurons and MD cells. The projections from the MD end as asymmetrical (excitatory) terminals (open triangles) predominantly on the spines of pyramidal layer III and layer V/VI neurons in the prefrontal cortex. In addition, aspiny nonpyramidal neurons, presumably GABAergic interneurons, are contacted by MD terminals. Some of the pyramidal neurons in layer III give rise to callosal fibers. Pyramidal neurons in the deeper layers provide return projections to the MD. This scheme is reproduced from Kuroda et al. (1998). These authors hypothesize that, since interneurons in the rat MD are scarce or absent, excitatory influences of MD neurons on cortical pyramidal neurons that project back to the MD may lead to reverberatory activity in this MD–PFC thalamocortical circuitry. Via the callosal projections, such activity may spread to the contralateral MD–PFC circuit. Inhibitory inputs (symmetrical synapses, closed triangles) originating from the ventral tegmental area terminate on the dendritic shafts of pyramidal neurons in layers III, V, and VI. This may provide a morphological basis for inhibitory influences on the reverberatory excitation between the MD and the PFC and on the contralateral excitation. The inhibitory cortical interneurons are possibly also involved in the modulation of these excitatory circuits. For further discussion, see Kuroda et al. (1998).

which are small to medium in size, contain pleomorphic vesicles, and terminate as asymmetric synapses. On the basis of this brief survey of the afferents of the MD and their ultrastructural characteristics, it may tentatively be concluded that the different mediodorsal segments receive driving inputs (type II fibers; cf. see section “Some General Aspects of Thalamic Organiza-

tion) from different sources. Drivers for the MDM originate in the amygdala, for the MDC in the prepiriform cortex, and for the MDL in the laterodorsal tegmental nucleus. This may indicate that the MDM is primarily involved in “limbic-emotional” functions, the MDC in olfactory functions, and the MDL in attentional functions, an interpretation that is supported by

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the different combinations of other inputs in the three MD segments (Groenewegen, 1988; Ray and Price, 1992; Kuroda et al., 1998) and their relationships with different parts of the (pre)frontal cortex (Heidbreder and Groenewegen, 2003). While neurons in layer VI and, to a lesser degree, in layer V of the frontal cortex project to the mediodorsal nucleus (Groenewegen, 1988; Cornwall and Phillipson, 1988b), it has not been studied in detail whether there exists a morphological distinction (i.e., type I [modulators] versus type II [drivers]) between the fibers and terminals from these different layers in the MD (Kuroda et al., 1998; Vertes, 2002). However, Negyessy et al. (1998) found, among the majority of ipsi- and contralateral small terminals of the RS-type (type I), large terminals with round vesicles in the MD originating from the ipsilateral but not the contralateral prefrontal areas. Therefore, the mediodorsal nucleus might derive cortical type II driver inputs from (parts of) the ipsilateral prefrontal cortex in addition to those from the prepiriform cortex to the MDC. The inputs to the various MD segments from different parts of the ventral pallidum and the reticular part of the substantia nigra must be viewed from the perspective of the involvement of the MD in a series of prefrontal cortex basal ganglia–thalamocortical circuits (Groenewegen et al., 1990, 1999; Ray and Price, 1992; Churchill et al., 1996; O’Donnell et al., 1997; Zahm, 2000). As indicated above, the ventral pallidal and nigral fibers make GABAergic contacts with MD thalamocortcial neurons (Kuroda and Price, 1991a, 1991b; Churchill et al., 1996). In view of the high spontaneous activity of their neurons, the ventral pallidum and substantia nigra exert a tonic inhibitory influence on the MD. However, as a result of corticostriatal activity which, via a higher activity of the GABAergic striatopallidal and striatonigral projections, leads to an inhibition of the pallidal and/or nigral neurons, the MD may be disinhibited (Deniau et al., 1994). Since the ventral pallidal and nigral terminals tend to end on the more proximal parts of dendrites of MD thalamocortical neurons (Kuroda et al., 1998), these inputs may have a major impact on the activity of the prefrontal cortical–mediodorsal thalamus reciprocal circuitry. Functional Aspects It seems likely that the different segments of the mediodorsal nucleus subserve different functions, but the “resolution” of behavioral studies using lesions or inactivations of MD is too coarse to reliably distinguish between different parts of MD. Furthermore, the MD is spatially very closely associated with the midline and intralaminar nuclei, which also hampers the interpretation of lesion studies.

Functions of the MD have been closely associated with those ascribed to the prefrontal cortex, i.e., in broad terms learning and “working” memory, behavioral control, and visceral functions. As such, the functions of the mediodorsal nucleus should preferably be considered in the context of “limbic” basal ganglia– thalamocortical circuits that are thought to subserve different functional roles (Groenewegen et al., 1990; Deniau et al., 1994; Zahm, 2000). Thus, the rostral MDM is involved in a circuit that encompasses the ventral prelimbic and infralimbic areas, the ventromedial striatum (including the shell of the nucleus accumbens), and the ventromedial part of the ventral pallidum. This circuit might be involved in behavioral control on the basis of visceral–limbic information. The caudal MDM is a way station in a circuit that includes the dorsal agranular insular area, the core of the nucleus accumbens, and the dorsomedial part of the substantia nigra pars reticulata. The MDC forms part of a circuit that includes the infralimbic, orbital, and ventral agranular insular areas of the prefrontal cortex and the pallidal neurons in the deep layers of the olfactory tubercle. This circuit is most probably involved in aspects of the olfactory guidance of behavior (see also Shipley et al., Chapter 29, this volume). Finally, the MDL is involved in a circuit that includes the dorsal prelimbic and dorsal anterior cingulate areas, as well as the medial agranular cortex (Fr2), the medial caudate putamen, and the medial globus pallidus. This circuit might be involved in attentional control of behavior. The medial thalamus, prominently including the mediodorsal nucleus, has since long been implemented in learning and memory (review, Markowitsch, 1982). Yet, it has been difficult to precisely establish the role of the MD in these functions, certainly also in distinction with the anterior thalamic nuclei and the midline/ intralaminar complex, including the nucleus reuniens (reviews, Aggleton and Brown, 1999; Van der Werf et al., 2002). Several authors suggest a role for the MD in spatial working memory, as determined in spatial delayed alternation tasks (e.g., Hunt and Aggleton, 1998; Kalivas et al., 2001). Other studies, employing lesions of the mediodorsal nucleus in tests of discrimination and reversal learning, showed an impairment of reversal learning, in particular when stimulus–reward contingencies were reversed (Chudasama et al., 2001). In line with these observations, electrophysiological studies show a subpopulation of neurons in the MDM that code for stimulus–reward associations which change their activity during extinction and reversal in the tasks used (Oyoshi et al., 1996). Such findings comply with the involvement of the MDM in a triangular relationship with the prefrontal cortex and the amygdala (cf. Price, 1995). Several studies indicate that the mediodorsal nucleus

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is part of a circuit through which epileptic activity is transferred. In models for epilepsy, i.e., through hippocampal kindling or chronic spontaneous limbic epilepsy, it was found that the medial thalamus, including the MD and reuniens nuclei, showed clear seizure activity (Bertram et al., 2001). In both these models as well as in a model of status epilepticus in immature rats (Kubova et al., 2001), loss of thalamic neurons was observed, in particular in the MD but also in other medial thalamic nuclei.

Nucleus Submedius The thalamic nucleus submedius (Sub) is a relatively small nucleus in the midrostrocaudal part of the thalamus. The Sub is situated in between the reuniens (Re) and rhomboid (Rh) nuclei medially and the VM laterally. In the literature, this nucleus has also been indicated as the nucleus gelatinosus (Krieg, 1944; Jones, 1985). The Sub is relatively cell-sparse and stands out against its neighboring nuclei also in neurofilamentstained sections (Fig. 184 in Paxinos et al., 1999). Although there exists a light plexus of parvalbuminpositive fibers in the Sub, calcium-binding proteins are virtually absent from this nucleus (Arai et al., 1994). In particular, calbindin D28K is conspicuously absent from the nucleus submedius making the nucleus readily identifiable in sections stained for this protein (Fig. 188 in Paxinos et al., 1999). The Sub is generally subdivided into rostromedial and caudodorsal parts on the basis of different fiber connections: the former part predominantly receives olfactory inputs and the latter somatic sensory inputs (see below). The caudodorsal part of the Sub exhibits a high acetylcholinesterase activity (see Plate 30 in Paxinos and Watson, 1998), most probably related to a substantial cholinergic input from the laterodorsal tegmental nucleus. Afferent and Efferent Connections Since the original description by Craig and Burton (1981) of afferents from the superficial layers of the spinal dorsal horn and spinal trigeminal nucleus, the Sub has been primarily associated with nociception and pain modulation. However, apart from these ascending sensory afferents, the Sub also receives input from olfactory structures (Price and Slotnick, 1983; Yoshida et al., 1992). The olfactory input terminates mostly in rostroventral parts of the nucleus and is derived from the deep layers of the prepiriform cortex and the endopiriform nucleus. These inputs are very comparable to those of the central segment of the MD (MDC), although the nucleus submedius receives input from a more restricted area in the olfactory cortex than MDC. However, the main cortical associations of

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the Sub are with the ipsi- and contralateral ventrolateral orbital area (VLO). Other cortical areas that receive projections from the Sub include the medial part of the lateral orbital cortex (Yoshida et al., 1992; Coffield et al., 1992; Reep et al., 1996). The dorsal peduncular cortex, located just ventral to the infralimbic cortex, and the dorsal subiculum send cortical fibers to the nucleus submedius (Witter et al., 1990; Yoshida et al., 1992). Forebrain inputs to the nucleus submedius are derived from the horizontal limb of the diagonal band, the lateral hypothalamus, and the reticular thalamic nucleus (Yoshida et al., 1992). Brain stem inputs originate in the deep layers of the superior colliculus, the brain stem reticular formation, the parabrachial nuclei, the laterodorsal tegmental nucleus, and the spinal trigeminal complex (Yoshida et al., 1991, 1992; Krout et al., 2001, 2002; Krout and Loewy, 2000a). In view of the presumed role of the Sub in nociception, the organization of afferents from the trigeminal complex and the dorsal horn of the spinal cord has received special attention (e.g., Craig and Burton, 1981; Dado and Giessler, 1990; Yoshida et al., 1991; Blomqvist et al., 1992; Iwata et al., 1992). In cats, it has been shown that the trigemino- and spinothalamic afferents of the Sub contain glutamate as transmitter (Ericson et al., 1995), while in rats trigeminothalamic neurons appear to express substance P (Li, 1999). The trigeminothalamic fibers, originating from the caudal spinal trigeminal complex, terminate as RL-type terminals on dendritic protrusions forming glomeruli, surrounded by glial elements (Ma and Ohara, 1987; Ma et al., 1988; Ericson et al., 1996). There are also more “simple” boutons-enpassage from both trigeminal and spinal origin in the nucleus submedius, at least in cats (Ericson et al., 1996). Cortical afferents most likely terminate as RS-type terminals while large boutons with flattened vesicles are also present (Ma et al., 1988). As in the sensory relay nuclei, the ascending spinal or trigeminal inputs are most probably the driving inputs for the Sub. In this respect, the nucleus submedius could also be considered to belong to the sensory thalamic nuclei. Functional Aspects Although it may have still not been definitely settled in rats to what extent superficial layer I neurons in the dorsal horn of the spinal cord or the spinal trigeminal complex contribute to the ascending projections to the Sub, functional studies clearly demonstrate a role for the nucleus submedius in nociceptive functions. Electrophysiological studies have shown nociceptive neurons in the Sub following peripheral noxic stimulation (Miletic and Coffield, 1989; Fu et al., 2002). Lesions of the Sub indicate that this nucleus may play a role in supraspinally mediated inhibition of nociceptive input

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(Roberts and Dong, 1994). It has been suggested that the nucleus submedius is a way station in a feedback loop that modulates nociceptive inputs. This loop would run via the Sub to the ventrolateral orbital cortex and, subsequently, descend to the periaqueductal gray matter through which inhibition of nociceptive input at the spinal or trigeminal level can be established (Zhang et al., 1995, 1996, 1999). Behavioral studies may be in line with this interpretation: for example, morphine injections in the Sub depress nociceptive behavior (Yang et al., 2002). However, other studies show that the nucleus submedius may exert its nociceptive modulatory effects at supraspinal levels (Okada et al., 1999). Finally, it has been suggested that the Sub is a crucial nodal point in the brain for the inhibitory effect of acupuncture manipulations in the suppression of certain visceromotor reflexes (Sumiya and Kawakita, 1997).

Anterior Nuclei Three distinct nuclei in the rostral one-third of the thalamus belong to the anterior thalamic compex, i.e., the anterodorsal (AD), anteroventral (AV), and anteromedial (AM) nuclei (Figs. 22–28 in Paxinos and Watson, 1998). The AM nuclei of both thalamic halves fuse in the midsagittal plane and form the interanteromedial nucleus (IAM) which extends slightly further caudally then the other anterior nuclei. Because of great similarities in afferent and efferent connectivity, the LD is often also included in the group of anterior nuclei (Price, 1995). The LD rostrally begins as a small nucleus (as a cap) dorsal to the AV and expands more caudally to occupy an extensive dorsal region in the thalamus, lying lateral to the habenular complex. Caudally, laterodorsal nucleus ends at the same level as the lateral habenula and is gradually replaced medially by the LP which extends and expands into the most caudal parts of the thalamus (Figs. 34–42 in Paxinos and Watson, 1998). In Nissl-stained sections, the anterodorsal nucleus is most conspicious because of its large, darkly staining cells. The anteromedial and anteroventral nuclei have smaller and lighter staining neurons and, although the cell density in the AV is lower than that in the AM, in Nissl-stained sections the border between these nuclei is difficult to draw. However, in sections stained for AChE activity, a clear distinction can be made between the anterior nuclei, the AM exhibiting the lowest and the AV the highest activity of this enzyme (see Plate 26 in Paxinos and Watson, 1998). Neurons positive for calcium-binding proteins, in particular for calbindin D28K, are present in the AM, but not in the other anterior nuclei. The three anterior nuclei contain to a varying degree positive neuropil for the calcium-binding proteins calbindin D28K, calretinin, and parvalbumin,

but they are not conspicuous thalamic structures in that respect (Arai et al., 1994). Within the anteroventral nucleus a distinction can be made between a ventrolateral (AVVL) and a dorsomedial (AVDM) part on the basis of calbindin immunoreactivity; AVVL (like AM) contains a moderately dense calbindin-positive neuropil, AVDM (like AD) is almost blank (Plate 174 in Paxinos et al., 1999). Afferent and Efferent Connections Inputs to the anterior thalamic nuclei are mainly derived from the limbic cortices in the medial wall of the cerebral hemisphere (see below), hippocampal region, and mammillary nuclei (Swanson and Cowan, 1977; Witter et al., 1990; Seki and Zyo, 1984; Shibata, 1992). In addition, serotonergic inputs originate from dorsal and median raphe nuclei (Gonzalo-Ruiz et al., 1995) and cholinergic fibers from the dorsolateral tegmental and pedunculopontine tegmental nuclei in the brain stem (Gonzalo-Ruiz and Lieberman, 1995). Reticular thalamic fibers originate from specific parts of the rostral, limbic portion of the reticular nucleus (Lozsadi, 1995; Shibata, 1992; Seki and Zyo, 1984). Inputs from the (para)hippocampal region arise mainly from the subiculum proper and the presubiculum and parasubiculum, while much smaller contributions derive from other parahippocampal areas (Swanson and Cowan, 1977; Witter et al., 1990; Van Groen and Wyss, 1990a, 1990b). The entire anterior thalamic complex, including the LD, receives subicular fibers. The presubiculum projects massively and bilaterally to the anterior nuclear complex of the thalamus, in particular to the anteroventral, anterodorsal, and laterodorsal nuclei. A marked topographical difference is present between the dorsal and ventral presubiculum in that the dorsal part, also called postsubiculum, preferentially projects to the LD and AD, whereas the ventral portion projects to the LD and AV (Seki and Zyo, 1984; Van Groen and Wyss, 1990a, 1990b). The parasubiculum sends a modest projection to the anterodorsal nucleus (Van Groen and Wyss, 1990a). The mammillothalamic projections are highly topographically organized, such that medial parts of the medial mammillary nuclei project to the IAM and medial AM, while progressively more lateral parts of the mammillary complex project to successively more lateral and dorsal parts of the anterior thalamic complex (Shibata, 1992). Mammillothalamic neurons contain glutamate and/or aspartate as neurotransmitter, while a subpopulation of these neurons expresses enkephalin (Gonzalo-Ruiz et al., 1998). The anterior nuclei as a complex project to the entire rostrocaudal extent of the medial prefrontal and anterior and posterior cingulate cortices (for a more detailed

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overview of the relationships of the anterior nuclei with the cingulate cortical areas, see Vogt et al., Chapter 22, this volume). Whereas the AD and the AV have their main projections to the caudal parts of the cingulate cortex (Sripanidkulchai and Wyss, 1986; Van Groen et al., 1995), the AM projects mainly to the rostral areas, including the dorsal and ventral anterior cingulate, prelimbic, and infralimbic areas and, like the AD and AV but less densely, also to the retrosplenial areas (Van Groen et al., 1999). Further projections from the anteromedial nucleus have been described to visual area 18b, the entorhinal and perirhinal areas, the subiculum and presubiculum, and the lateral and basolateral amygdaloid nuclei (Van Groen et al., 1999). The projections from AD, AV, and AM to different parts of the retrosplenial cortex as well as to the various subicular and parahippocampal areas are organized topographically, reciprocating the afferent patterns of origin, and to a large degree terminate in complementary patterns (Van Groen et al., 1999). The different nuclei distribute their fibers over different cortical layers: in the retrosplenial cortex the anterodorsal nucleus projects equally to layers I, III, and V, while the anteroventral nucleus projects most densely to layer Ia and less densely to layer V; the anteromedial nuclear projections reach layers I and V. In the dorsal part of the presubiculum, the AD and the AV both project to layers I and V, while the AD, in addition, targets layer II/III. In the anterior cingulate and medial prefrontal areas, the AM reaches layers I, V, and VI; in the entorhinal and perirhinal cortices layer V is targeted (Van Groen et al., 1995, 1999). Cortical projections to the anterior thalamic nuclei arise in virtually all limbic cortical areas that receive a projection from these nuclei and these corticothalamic projections to a large degree reciprocate the thalamocortical projections. However, this reciprocity is not strict. For example, the AM projects to a part of the anterior and posterior cingulate areas much wider than that from which it receives projections. The anteromedial nucleus projects to area 18b and the subiculum, but does not receive projections from these cortices (Van Groen et al., 1999). Further, the frontal eye field has a substantial projection to virtually the entire anterior complex (Guandalini, 2001). At the ultrastructural level, mammillothalamic fibers terminate through large boutons with round vesicles that make asymmetric synapses on the proximal dendrites, reminiscent of type II afferents (Dekker and Kuijpers, 1976). Subicular and cingulate terminals are small to medium size, also have round vesicles, and terminate via asymmetrical synapses on more distal dendrites (Oda, 1997). These afferents may therefore be categorized as type I afferents. According to Sherman and Guillery (2001), the mammillary afferents might

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be the drivers of the anterior thalamic nuclei, whereas the afferents from the other cortical regions have more the characteristics of modulators. This is an interesting conclusion in view of recent functional insights in the anterior nuclei (see below). However, at present electrophysiological evidence is lacking to support this “morphological” conclusion. Functional Aspects As can be concluded from the previous paragraph, the anterior thalamic nuclei are included in a neural circuit that involves the anterior and posterior cingulate cortices, the hippocampal region, in particular the subiculum and presubiculum, and the mammillary bodies. In the terminology of Aggleton and Brown (1999), the anterior thalamic nuclei are part of an “extended hippocampal system.” The connections of the anterior nuclei with the cingulate and hippocampal cortical areas are reciprocal, while the input from the mammillary complex is unidirectional. Functional studies have shown that this circuit, most probably in conjunction with afferent brain stem connections of the mammillary nuclei originating in the dorsal tegmental nuclei, is crucial for spatial orientation and spatial memory (for a review, see Aggleton and Brown, 1999). Behavioral studies using lesions of different way stations in the anterior thalamic–hippocampal circuitry, in which also the mammillary nuclei and the cingulate cortices are incorporated, indicate that this circuit is important for attentional processes and various kinds of learning and memory processes, in particular spatial, allocentric memory (Aggleton et al., 1996; Warburton et al., 2001; Van Groen et al., 2002). Lesions of the anterior thalamic nuclei, but also of the fornix and the mammillary nuclei disrupt specific aspects of spatial learning, but not of object learning (Aggleton et al., 1995; Sziklas and Petrides, 1999). The exact contribution of the anterior thalamic nuclei to spatial memory and attentional functions remains to be established, but is seems clear that these nuclei play an important role in the network between the hippocampus, the mammillary bodies, and the cingulate cortex. None of the distinct anterior nuclei appears to have a preeminent role, since lesions of all nuclei have an effect on spatial memory tasks; however, complete anterior thalamic lesions have the greatest effect (Aggleton et al., 1996). These recent, behavioral findings have renewed the interest in the functional anatomy of the classic “Papez circuit” and its role in episodic memory (Aggleton and Brown, 1999; Van Groen et al., 2002). A physiological correlate of the role of the anterior thalamic nuclei in spatial tasks might be the coding by anterior thalamic neurons for the spatial orientation of the head of the animal (e.g., Aggleton et al., 1996; Blair

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et al., 1997, 1999; Vann et al., 2000; Zugaro et al., 2000; Wilton et al., 2001). Thus, electrophysiological studies have shown that neurons in the anterior thalamic complex, as well as in the laterodorsal nucleus, signal the rat’s directional heading in the horizontal plane; these neurons are indicated as head direction (HD) or “directional” cells (Taube, 1995; Taube et al., 1990). Such HD cells are most prominently present in the anterodorsal and laterodorsal thalamic nuclei, but they are also found in anteroventral and anteromedial nuclei (Blair et al., 1997; Mizumori and Williams, 1993), as well as in the dorsal part of the presubiculum and lateral mammillary nucleus (Taube et al., 1990; Stackman and Taube, 1998). Interestingly, the activity of HD cells in the anterior thalamus is not dependent upon an intact hippocampus (Goodridge and Taube, 1997; Golob and Taube, 1997). By contrast, (bilateral) lesions of the lateral mammillary nucleus do abolish the HD activity in the anterodorsal nucleus (Blair et al., 1999). This indicates that the lateral mammillary nuclei play a crucial role in transmitting the directional signal to the anterior thalamus, a signal that may be related to the actual head direction, but also to the anticipated directioning of the head (Blair et al., 1997). This directional information is dependent upon visual inputs (Goodridge et al., 1998), but may also be derived, via the lateral mammillary nuclei and the dorsal tegmental nuclei in the brain stem, from the vestibular system (Stackman and Taube, 1997; Blair et al., 1999). However, it is clear that also other cues are important for the activity of HD cells, such as tactile and olfactory (Goodridge et al., 1998). In this context, it is of interest to note that Risold and Swanson (1995) have demonstrated a pathway from the lateral hypothalamus via the anteromedial thalamic nucleus to the retrosplenial cortex that might mediate pheromonal influences on attentional mechanisms and eye and head movements. Whereas the anterior thalamic nuclei and postsubiculum contain HD cells that monitor the direction of the rat”s head, the hippocampus itself contains socalled “place cells” which code for the position of the animal in a particular space. It has been argued by Muller et al. (1996) that the hippocampal “positional system” might act, under particular circumstances, as a cognitive map and that the role of the presubicular– anterior thalamic “directional system” is to put the map into register with the actual environment while acting. Thus, through interactions between both systems, the animal would be able to properly orient and execute behaviors in a learned environment (Muller et al., 1996).

Lateral Nuclei A large dorsolateral part of the rat thalamus can be recognized as the lateral thalamic complex, consisting of

the caudally located lateral posterior nucleus (LP) and the more rostrally and medially situated lateral dorsal nucleus (LD). The two nuclei constitute the dorsal surface of the thalamus, lateral to the habenular complex and, caudally, medial to the dorsal lateral geniculate nucleus (DLG). The LD stretches from the caudal part of the anterior thalamic complex to the rostral pole of the DLG. The lateral posterior nucleus starts at midthalamic levels in between the habenular complex medially and the laterodorsal nucleus laterally, and in the posterior direction the LP gradually replaces the LD. Posteriorly, the lateral posterior nucleus becomes situated medial to dorsal lateral geniculate and, in its most posterior extent, dorsal to the medial geniculate nuclei (see Figs. 26–42 in Paxinos and Watson, 1998). In Nissl stained sections the lateral nuclei appear rather homogeneous, but both the LD and the LP can be subdivided into several subnuclei (see Paxinos and Watson, 1998). These subdivisions are based on cytoarchitectonics and distinct patterns of connectivity with the cerebral cortex, and they can, to a certain degree, be recognized in acetylcholinesterase-stained sections showing slight differences in staining intensity in the various subnuclei (Takahashi, 1985; Plates 26, 31, 35 and 39 in Paxinox and Watson, 1998). Acetylcholineserase activity in the lateral posterior nucleus is higher than that in the laterodorsal nucleus in which there is only moderate activity of this enzyme. Although not throughout the entire LD and LP, the calcium-binding proteins calretinin and calbindin D28K are present in neurons in certain parts of these nuclei, while parvalbumin as well as the other two proteins are present in fibers in both nuclei (Arai et al., 1994; Paxinos et al., 1999). Afferent and Efferent Connections Like the anterior thalamic nuclei, the LD is strongly related to the retrosplenial cortex (Van Groen and Wyss, 1990a, 1992a; Shibata, 2000; see also Chapter 22, this volume). The projections from different parts of the retrosplenial cortex, originating predominantly from layer VI, but also from neurons in layer V, distribute in a topographical way over the various subnuclei of the LD (Shibata, 2000). Thus, a rostral-to-caudal axis in the retrosplenial cortex (area 29c to area 29a) corresponds to the ventromedial-to-dorsolateral axis of the laterodorsal nucleus (Fig. 6). Further cortical inputs to the LD originate in visual areas 17 and 18, the presubiculum, and entorhinal cortex (Thompson and Robertson, 1987a, 1987b; Shibata, 1996). Several other cortical areas project to different parts of the LD, for example, medial prefrontal areas provide an input to its most medial part of laterodorsal nucleus (e.g., Sukekawa, 1988; Vertes, 2002), but such inputs have not been studied systematically. Subcortical inputs to the LD arise in the reticular

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FIGURE 6 Schematic representation of the topographic arrangements between the retrosplenial areas 29a–c and 29d with the laterodorsal thalamic nucleus (LD). Area 29a projects to the dorsolateral part of the LD; while moving from area 29a via area 29b and the caudal part of 29c to the rostral part of 29c, the termination pattern in the LD shifts from dorsolateral to ventromedial in the LD (A, B). Area 29d projects to the ventrolateral part of the LD; a rostrocaudal gradient in area 29d corresponds with a mediolateral gradient in the LD (C, D). The terminal fields of area 29c and area 29d overlap in the central part of the LD. Reproduced from Shibata (2000).

thalamic nucleus, the pretectal nuclei, the intermediate layers of the superior colliculus, and the ventral lateral geniculate nucleus (Thompson and Robertson, 1987b; Coleman and Mitrofanis, 1996; Kolmac et al., 2000). The efferent projections from the LD reach various limbic and visual cortical areas, i.e., the cingulate and retrosplenial cortices, the subicular complex, and the entorhinal cortex. These projections are topographically organized and originate from different subnuclei of the laterodorsal nucleus (Van Groen and Wyss, 1992b). Compared to the LD, the occipital, parietal, and temporal sensory association cortices have inputs to the lateral posterior nucleus much stronger than those of the limbic cingulate, retrosplenial, and subicular cortices (e.g., Vaudano et al., 1991; Coleman and Mitrofanis, 1996; Shi and Cassell, 1997; Kolmac et al., 2000; Shibata, 2000). Inputs from the primary visual cortex to the LP arise mostly from collaterals of neurons in layer V that primarily project to the brain stem (Bourassa and Deschenes, 1995). The LP receives a dense projection from temporal area Te2, an auditory and visual association area, but not from areas Te1 and Te3 (Shi and Cassell, 1997). Functional Aspects Both the laterodorsal and the lateral posterior nuclei are considered to be association thalamic nuclei in that they do not receive direct motor or sensory inputs. However, as indicated above these nuclei receive strong inputs from sensory and motor association cortical areas as well as from limbic association areas. The LP is most

strongly related to visual association areas, the LD is more related to limbic cortices. In view of similarities in their connectivity patterns, the laterodorsal nucleus is strongly associated with the anterior thalamic complex. As a result of the strong relationships with cortical and subcortical visual structures, the LP is often considered the visual association thalamus and compared with the pulvinar in other species like cats and primates. A pulvinar, however, is lacking in rats (Jones, 1985). Like in the anterior thalamic complex, head direction cells have been identified in the LD (see page 430). However, the role of the LD in head orientation and spatial navigation appears to be different from that of the anterior thalamic nuclei and the pre- and parasubiculum (Golob et al., 1998). Yet, lesions of the LD clearly demonstrate that the laterodorsal nucleus serves a task in spatial learning and memory (Van Groen et al., 2002).

MIDLINE AND INTRALAMINAR THALAMIC NUCLEI The intralaminar and midline nuclei form a conspicuous collection of nuclei in the medial and dorsal part of the thalamic complex (Figs. 22–38 in Paxinos and Watson, 1998). The midline nuclei are located medially in the thalamus as a narrow band of small nuclei that are distributed over the entire dorsal-to-ventral extension of the thalamus. The intralaminar nuclei are located lateral to the mediodorsal nucleus and “embedded”

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within the internal medullary lamina. Several of the nuclei of the midline and intralaminar complex have a high activity for acetylcholinesterase (see Plates 24, 25, 30, 31, and 34 in Paxinos and Watson, 1998).

Midline Nuclei The midline nuclei include the paraventricular, parataenial, intermediodorsal, reuniens, and rhomboid nuclei. These nuclei occupy the midline of the rat thalamus from its very rostral tip to approximately twothirds of the rostrocaudal length of the thalamus. The paraventricular nucleus (PV) is located medially, spanning the entire rostrocaudal length of the midline/intralaminar complex. The PV lies directly ventral to the third ventricle, dorsal and medial to the mediodorsal nucleus. Rostrally, it follows the surface of the massa intermedia and curves ventrally to form a wedge between the anterior poles of the nucleus reuniens (Krieg, 1944). More caudally, the PV curves slightly laterally, ventral to the habenula, and ends just rostral to the posterior commissure. The paraventricular thalamic originates phylogenetically from a pronuclear mass that gives rise also to the pineal and the habenular nuclei which has led Jones (1985) to consider the PV as part of the epithalamus. The parataenial nucleus (PT) forms a slender, elongated nucleus located anteriorly in the thalamus in close proximity and lateral to the paraventricular nucleus. At its posterior end, the PT fuses with the mediodorsal nucleus. The PT is thought to originate together with the mediodorsal and reuniens nuclei from a common nuclear mass. In contrast to Price (1995), we include the parataenial nucleus in the group of midline/ intralaminar nuclei, primarily on the basis of its projections’ patterns (see below). The intermediodorsal nucleus (IMD) is located more posteriorly, in between the left and right mediodorsal nuclei. An IMD is not recognized in all species, and it is described in most detail in rats (Jones, 1985; Berendse and Groenewegen, 1990). In many studies, the IMD is considered as the medial part of the mediodorsal nucleus. However, on the basis of the slightly darker staining of the neurons as well as its afferent and efferent relationships (see below), the IMD can be clearly distinguished from this nucleus. The nucleus reuniens thalami (Re) is located in the anterior one-third of the thalamus. Anteriorly, it is divided into a left and a right component by the third ventricle; toward its tail the two structures fuse and become a mass of cells in the midline of the thalamus, lying immediately dorsal to the third ventricle. The Re consists of a conglomerate of loosely packed cells (Jones, 1985). Caudally, the main mass of the Re is bordered

by the so-called perireuniens or ventral reuniens nuclei on both sides. The rhomboid nucleus (Rh) is located ventral to the internal medullary lamina (Berendse and Groenewegen, 1990). At its rostral end, it is confluent with the anteromedial nucleus and has two wing-like lateral extensions. Caudally, these two structures merge in the midline. The nucleus is easily distinguished by its conspicuous shape and its large and darkly staining cells (Plate 29 in Paxinos and Watson, 1998). The midline nuclei most probably use excitatory amino acids as neurotransmitter. In addition, calciumbinding proteins are present in both cell bodies and neuropil. Calretinin-positive neurons are present virtually throughout the midline nuclei, with a preference for the PV and the Re (Arai et al., 1994; see e.g. Plates 175 and 188, Paxinos et al., 1999). Calbindin-positive neurons are predominantly located located in the reuniens and intermediodorsal nuclei. Parvalbumin is conspicuously absent from the midline nuclei (Arai et al., 1994). The neuropil of the paraventricular nucleus is very rich in neurotransmitters and neuropeptides, most of which are contained in afferent fibers (see below). In addition, a subpopulation of paraventricular neurons expresses preproenkephalin mRNA (Hermanson et al., 1995). Afferent Connections The paraventricular nucleus (PV) receives afferent fibers from a wide variety of cortical and subcortical structures (e.g., Cornwall and Phillipson, 1988a; Chen and Su, 1990; Hurley et al., 1991; Otake et al., 1995; Ruggiero et al., 1998; Krout and Loewy, 2000a, 2000b; Krout et al., 2002). Thus, the PV receives strong aminergic inputs consisting of a histaminergic input from the tuberomammillary nucleus (Panula et al., 1989), a dense dopaminergic input from the ventral tegmental area and the retrorubral region (Takada et al., 1990), a noradrenergic input from the locus coeruleus and the nucleus of the solitary tract, and a serotonergic input from the dorsal raph (Otake and Nakamura, 1995; Otake and Ruggiero, 1995; Krout et al., 2002). In addition, the paraventricular nucleus receives input from the parabrachial nucleus, extensive parts of the brain stem reticular formation, deep layers of the medial part of the superior colliculus, the bed nucleus of the stria terminalis, the dorsomedial and posterior hypothalamus, and the supramammillary nuclei (Vertes, 1992; Vertes et al., 1995; Bester et al., 1999; Krout et al., 2001, 2002). Whereas most of these inputs are shared with several of the other midline nuclei, inputs from the suprachiasmatic nucleus and the intergeniculate leaflet are virtually unique for the PV (Moore et al., 2000; Kawano et al., 2001). Inputs from the amygdalar complex originate in the central nucleus. Cortical input is derived from

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the infralimbic cortex and the subiculum (Groenewegen, 1988; Sesack et al., 1989; Hurley et al., 1991). Much less is known about the afferents of the parataenial nucleus, (PT), presumably as a result of its small size and the difficulty in selectively targeting the PT in tracing studies. In anterograde tracing studies, the PT is frequently overlooked or not mentioned. Cortical input to the parataenial nucleus arises from the infralimbic cortex (Hurley et al., 1991; Sesack et al., 1989) reaching mainly the dorsomedial part of the PT. Chen and Su (1990) described input from the ventral subiculum and the rostral half of the claustrum. Further, relatively weak inputs derive from the lateral septal nuclei, the bed nucleus of the stria terminalis, the medial amygdala, the amygdalohippocampal area, a band between the ventral pallidum and the nucleus accumbens, the reticular thalamic nucleus, the zona incerta, and the suprachiasmatic nucleus as well as scattered regions throughout the hypothalamus (Chen and Su, 1990). The PT appears to receive moderate to weak brain stem input from the dorsal and median raphe nuclei, the periaqueductal gray, the locus coeruleus, the parabrachial nucleus, the laterodorsal tegmental nucleus, and the nucleus of the solitary tract (Bobillier et al., 1979; Cornwall and Phillipson, 1988a; Chen and Su, 1990; Newman and Ginsberg, 1994; Krout and Loewy, 2000a; Krout et al., 2002). Inputs to the intermediodorsal thalamic nucleus (IMD) have been the focus of only a few studies; in most tracing studies the IMD is either ignored or taken as part of the larger group of midline nuclei. Recently, Loewy and colleagues (Krout and Loewy, 2000a, 2000b; Krout et al., 2002) have shown that the pattern of brain stem inputs to the IMD is largely comparable to that of the paraventricular nucleus except that the IMD hardly receives any input from medullary structures or from motor-related structures in the mesencephalon such as the substantia nigra or the pedunculopontine region. Cortical inputs, in particular from the infralimbic and ventral agranular insular cortices, are sparse (Hurley et al., 1991). Inputs to the nucleus reuniens (Re), originally described by Herkenham (1978), are derived from several cortical and subcortical sources. Cortical inputs originate from deep layers of the infralimbic, prelimbic, and perirhinal cortices (see also Sesack et al., 1989; Witter et al., 1990; Hurley et al., 1991; Vertes, 2002). The subicular input is topographically organized (Witter et al., 1990; Wouterlood et al., 1990). Amygdalar inputs are sparse and arise from the medial and anterior nuclei (Herkenham, 1978). Basal forebrain inputs come from the nucleus of the diagonal band and the bed nucleus of the stria terminalis. Diencephalic inputs include projections from the reticular nucleus of the thalamus,

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the lateral geniculate nucleus, the zona incerta, the medial and lateral preoptic area, the medial and lateral hypothalamus, and the premammillary and supramammillary nuclei (Risold et al., 1997; Vertes and Martin, 1988). Brain stem areas projecting to the Re are the ventral tegmental area; the mesencephalic reticular formation; the laterodorsal tegmental nucleus; the superior colliculus; the periaqueductal gray matter; the dorsal, median, and central raphe; the locus coeruleus; and the parabrachial nucleus (Krout and Loewy, 2000a, 2000b; Krout et al., 2001, 2002). Weak projections to the nucleus reuniens originate in the medullary reticular nuclei and the cuneate nucleus (Peschanski and Besson, 1984; Villanueva et al., 1998; Krout et al., 2002). Data on inputs to the rhomboid nucleus (Rh) are sparse. Subcortical inputs to the Rh arise mainly in the brain stem (Krout et al., 2002), including very prominently the raphe nuclei (also: Bobillier et al., 1979); the mesencephalic, pontine, and medullary reticular formation (also: Peschanski and Besson, 1984); the lateral dorsal tegmental nucleus; and several other mesencephalic nuclei, including the substantia nigra. In addition, the supramammillary nucleus sends fibers to the rhomboid nucleus (Vertes, 1992).

Intralaminar Nuclei The intralaminar thalamic nuclei are made up of a rostral group, consisting of the central medial, paracentral, and central lateral nuclei. The caudal group is composed of the parafascicular–center median complex, in rats appearing as a single nuclear mass, briefly referred to as the parafascicular nucleus. In addition, in the caudal one-third of the thalamus and the subparafascicular and posterior intralaminar nuclei are considered to belong to the posterior intralaminar complex. In Nissl-stained sections, most of the intralaminar nuclei can be rather easily recognized on the basis of packing of the neurons that is denser than that in the adjacent thalamic nuclei (Plates 29 and 33 in Paxinos and Watson, 1998). The central medial nucleus (CM) is easily identifiable as a centrally located group of large, deeply staining, and flattened cells clearly distinct from the midline nuclei lying dorsal and ventral to it. Laterally, however, the CM is continuous with the paracentral nucleus on both sides. In rats, the left and right parts of the CM nuclei fuse with each other to form a single, centrally positioned nucleus (Plate 29 in Paxinos and Watson, 1998). The paracentral nucleus (PC) is a thin strip of cells that is continuous with the CM medially and the central lateral nucleus (CL) laterally. Its cells are difficult to distinguish from those of the CL, but appear more flattened. The PC lies in the anterior and middle por-

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tion of the internal medullary lamina, intercalated between the MD and the ventral nuclei. In the caudal PC, a distinct oval subnucleus can be recognized, the oval paracentral nucleus. The central lateral nucleus (CL) is the most dorsal component of the intralaminar nuclei, located posteriorly and dorsally to the PC with which it is confluent. The CL is larger than the PC, but these nuclei share many connectional characteristics. Therefore, the central lateral and paracentral nuclei are often regarded as a single structure. The parafascicular nucleus (PF) represents a caudal component of the intralaminar complex. The PF stands out as a darkly staining nucleus, surrounding the fasciculus retroflexus. The lateral, larger part of the rat parafascicular nucleus, containing darker staining neurons than the medial part, is considered to be equivalent to the center median nucleus, whereas the medial part of the rat PF is homologous to the primate parafascicular nucleus (Jones, 1985). Ventrally, the PF rests almost directly upon the mesencephalon; dorsally it is bordered by the central lateral nucleus and the habenular complex. The parafascicular nucleus lies just lateral to the most rostral extension of the peraventricular gray matter. The subparafascicular nucleus (SPF) is a flat, horizontally oriented nucleus which stretches from a position just ventral to the parafascicular nucleus rostromedially toward the posterior intralaminar and peripeduncular nucleus caudolaterally. Neurons in the SPF have a horizontal orientation. Medially, the subparafascicular nucleus lies just dorsal to the medial lemniscus. The nucleus consists of a medial magnocellular part and a lateral parvocellular part (Faull and Mehler, 1985; LeDoux et al., 1987; Coolen et al., 2003a). The principal neurotransmitters used by the intralaminar nuclei are excitatory amino acids. Neurons containing calcium-binding proteins are relatively sparsely present in the intralaminar nuclei (Arai et al., 1994; Coolen et al., 2003a). Enkephalinergic neurons, as indicated by the presence of preproenkephalin mRNA, are present in the central medial and central lateral nuclei and to a much lesser degree in the paracentral and parafascicular nuclei (Hermanson et al., 1995). Afferent Connections The main sources of input to the rostral intralaminar nuclei, i.e., the central medial (CM), paracentral (PC), and central lateral (CL), are subcortical structures, in particular a wide variety of brain stem nuclei as well as the spinal cord. Thus, different parts of the mesencephalic, pontine, and medullary reticular formation, several nuclei within the raphe complex which include serotonergic cell groups, the cholinergic pedunculopontine and laterodorsal tegmental nuclei, the nucleus

prepositus hypoglossi, the spinal trigeminal nucleus, the medial and lateral vestibular nuclei, several nuclei of the parabrachial complex, the locus coeruleus, the nucleus incertus (nucleus O), distinct regions of the periaqueductal gray matter, deep layers of the superior colliculus, the nucleus of Darkschewitsch, and the pars reticulata and pars compacta of the substantia nigra send projections to the entire or to specific parts of the rostral intralaminar complex (Peschanski and Besson, 1984; Yamasaki et al., 1986; Hallanger et al., 1987; Vertes and Martin, 1988; Villanueva et al., 1998; Bester et al., 1999; Groenewegen et al., 1999; Shiroyama et al., 1999; Krout and Loewy, 2000a, 2000b; Krout et al., 2001, 2002; Goto et al., 2001; for review see Van der Werf et al., 2002). Indeed, it has recently been elegantly demonstrated by Loewy and colleagues (Krout and Loewy, 2000a, 2000b; Krout et al., 2001, 2002), using small injections of retrograde tracers in the CM, PC, and CL, that these nuclei receive a different combination of inputs from the various brain stem nuclei, as well as from different subregions within these nuclei or regions, suggesting a certain degree of specificity of these three nuclei. Compared to the central medial and paracentral nuclei, the central lateral nucleus seems to receive the strongest inputs from the brain stem (Van der Werf et al., 2002). In addition, the rostral intralaminar nuclei receive inputs from the deep cerebellar nuclei, the supramammillary nuclei, the zona incerta, and the reticular thalamic nucleus (Haroian et al., 1981; Vertes, 1992; Kolmac and Mitrofanis, 1997; Power et al., 1999; Power and Mitrofanis, 2001; Van der Werf et al., 2002). Cortical inputs to the rostral intralaminar nuclei have not been specifically studied in rats, but it may be concluded from the literature that in particular (pre)frontal cortices, including the anterior cingulate cortex, send projections to the rostral intralaminar nuclei (e.g., Sesack et al., 1989; B. F. Jones and M. P. Witter, unpublished data). Compared to the adjacent higher order thalamic nuclei, the cortical inputs to the intralaminar complex are relatively weak. The subcortical inputs from brain stem nuclei and spinal cord to the parafascicular thalamic nucleus (PF) are to a large degree comparable to those to the rostral intralaminar nuclei (see above; Haroian et al., 1981; Cornwall and Phillipson, 1988a; Lai et al., 2000; for overview see Van der Werf et al., 2002; Krout et al., 2002). As described, a distinction should be made between the lateral and the medial part of the PF. The lateral parafascicular nucleus receives relatively strong inputs, both in comparison with the medial PF as well as with the rostral intralaminar nuclei, from sensorimotor-related brain stem nuclei, such as the vestibular nuclei, the spinal trigeminal complex, the superior colliculus, and the substantia nigra (Shiroyama et al.,

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1999; Krout et al., 2002). The medial PF receives more autonomic and visceral-related inputs, e.g., from the nucleus solitarius, the periaqueductal gray matter, and the parabrachial complex (Bester et al., 1999; Krout et al., 2000a, 2000b, 2002). Compared to the rostral intralaminar nuclei, the parafascicular nucleus receives relatively strong inputs from forebrain structures, in particular structures involved in the basal ganglia circuitry such as the entopeduncular nucleus (e.g., Gonzalo et al., 2002). Cortical inputs to the PF are derived from layer V and, to a lesser extent, layer VI of the frontal and parietal cortices (Cornwall and Phillipson, 1988a). The subparafascicular nucleus (SPF), as indicated above, can be subdivided into a medial and a lateral part (Coolen et al., 2003a), each receiving a different set of inputs (Coolen et al., 2003b). Thus, the medial SPF, characterized by galanin-immunoreactive fibers, receives inputs from lumbar spinothalamic neurons and visceral-related brain stem and forebrain regions. In contrast, the lateral SPF, containing calcitonin generelated peptide (CGRP)-immunoreactive neurons and fibers, receives inputs from auditory- as well as visualrelated brain stem and forebrain regions (LeDoux et al., 1987; Coolen et al., 2003b). The lateral subparafascicular nucleus is strongly related to the auditory thalamus, including also the posterior intralaminar and the medial geniculate nuclei.

Efferent Projections of the Midline and Intralaminar Complex The efferent projections of the midline and intralaminar thalamic nuclei are collectively discussed in this paragraph since the outputs of both groups of nuclei are in a number of respects very similarly organized. Whereas other thalamic nuclei have their main projections to the cerebral cortex, most of the midline and intralaminar nuclei project extensively to both cortical and subcortical structures. Subcortical structures that are reached by midline and intralaminar thalamic nuclei include several components of the basal ganglia (Berendse and Groenewegen, 1990; Feger et al., 1994), the amygdala (LeDoux et al., 1990; Turner and Herkenham, 1991), and preoptic and hypothalamic regions (Coolen et al., 1998; review Van der Werf et al., 2002). Another common finding is that most of these nuclei project to one cortical or subcortical target only; i.e., except for collaterals within the thalamus axons distribute preferentially to one target (Su and Bentivoglio, 1990). Virtually all nuclei that belong to the midline and intralaminar complex send projections to both the striatum and the cerebral cortex (Berendse and Groenewegen, 1990, 1991; Groenewegen and Berendse,

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1994), although there are differences among the individual nuclei in the strength of the projections to either the cortical or the subcortical targets. Midline and intralaminar nuclei like the PV, IMD, CM, PC, CL, and PF nuclei have relatively strong projections to the striatum, while other nuclei, such as the Re and PT nuclei, have rather weak projections to the striatum. The projections from the first-mentioned group of nuclei to both the striatum and the cerebral cortex are highly topographically organized such that the projections from individual thalamic nuclei reach restricted parts of the striatum and specific regions of the cerebral cortex, in particular in the frontal lobe (Fig. 7). This topography can be broadly described as follows. The PV reaches the most ventromedial parts of the striatum, in particular the shell of the nucleus accumbens, as well as the ventral part of the medial prefrontal cortex, including the infralimbic and ventral prelimbic areas. The IMD projects to more lateral parts of the ventral striatum, including the core of the nucleus accumbens and ventral parts of the caudate–putamen, as well as the lateral prefrontal cortex, including the dorsal agranular insular cortex. Moving within the intralaminar complex from the CM, via the PC and the CL to the PF, the projection field in the striatum shifts from a ventromedial to a dorsolateral position in the caudate– putamen and, in the cerebral cortex, from the prelimbic, via the anterior cingulate (Cg1) cortices in the medial wall to the somatomotor cortex (M2 and M1) in the dorsal aspect of the hemisphere. Combined with the topographical arrangement of the (pre)frontal corticostriatal projections, this means that individual midline or intralaminar thalamic nuclei target cortical and striatal areas that are associated with each other via corticostriatal projections. Moreover, on the basis of single-cell tracing, Deschennes et al. (1996) concluded that the striatal and cortical projections from the caudal intralaminar nuclei arise from collaterals of the same neuron (see also Bubser and Deutch, 1998; Otake and Namura, 1998). Thus, the PV, IMD, CM, PC, CL, and PF may have a strong influence on the level of activity of a series of parallel, functionally segregated (pre)frontal cortical basal ganglia–thalamocortical circuits that also involve the mediodorsal, ventromedial, and ventral anterior thalamic nuclei (Fig. 8) (Groenewegen and Berendse, 1994). The “central position” of the mentioned midline and intralaminar nuclei in cortical–subcortical circuits is further emphasized by the fact that these thalamic nuclei project also to other forebrain structures that are involved in these circuits. For example, the PV and IMD project to specific parts of the amygdaloid complex and the medial temporal lobe that, in turn, project to the prefrontal and ventral striatal targets of the PV and IMD (e.g., Wright and Groenewegen,

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FIGURE 7 Schematic illustration showing the distinctive and common inputs of the midline paraventricular–intermediodorsal thalamic nuclei (PV–IMD) and the intralaminar parafascicular nucleus, in particular its lateral part (PFl). In addition, the relationships of these midline and intralaminar nuclei with the parallel, functionally segregated basal ganglia–thalamocortical circuits are shown. (A) The involvement of the PV–IMD in the prefrontal cortical and ventral striatal way stations of “limbic circuits” is illustrated. The projections of these thalamic nuclei to the basal amygdaloid complex which, in turn, projects directly to the same cortical and striatal targets as the thalamic PV and IMD, are also indicated (cf. Wright and Groenewegen, 1995, 1996). (B) The influence of the PFl on the cortical and striatal way stations of the “motor circuits” is shown. Note that the PFl also projects to the lateral part of the subthalamic nucleus that is intimately involved in the motor circuits. Abbreviations: ABL, basal amygdaloid complex; ac, anterior commissure; DStr, dorsal striatum; fr, fasciculus retroflexus; GP, globus pallidus; GPe, external segment of GP; GPi, internal segment of GP; HF, hippocampal formation; IMD, intermediodorsal thalamic nucleus; MD, mediodorsal thalamic nucleus; PF, parafascicular nucleus; PFC, prefrontal cortex; PFl, lateral part of PF; PV, paraventricular thalamic nucleus; SMC, somatomotor cortex; STh, subthalamic nucleus; VA, ventral anterior thalamic nucleus; VL, ventral lateral thalamic nucleus; VP, ventral pallidum; VStr, ventral striatum. Slightly modified from Groenewegen and Berendse (1994).

1995, 1996). Likewise, the projections from the different parts of the parafascicular nucleus to the subthalamic nucleus (Berendse and Groenewegen, 1991; Feger et al., 1994) “fit into” the topographical organization of the (pre)frontal cortical basal ganglia–thalamocortical circuits (Groenewegen and Berendse, 1994; Van der Werf et al., 2002). The efferent projections of the Re, Rh, and PT nuclei, in contrast with those of the nuclei discussed in the previous paragraph, are primarily directed toward the cerebral cortex and to a much lesser degree to the striatum (Van der Werf et al., 2002). All three nuclei send fibers to the prelimbic, infralimbic, and anterior cingulate areas, as well as to the (ventral) striatal regions associated with these cortices. The Rh has, in addition, rather widespread projections to motor, sensory, visceral, and temporal association cortical areas, in particular directed to layer I of those cortices. This pattern of projection is shared with the caudal part of the CM which, as argued by Van der Werf et al. (2002), might be more related to the Rh than to the rostral CM. While

the Rh has very sparse projections to the entorhinal cortex and the hippocampus, these areas are a main target for the projections of the Re (Herkenham, 1978; Wouterlood et al., 1990). Thus, the Re has heavy projections to the CA1 area and the subiculum of the hippocampal formation. The pre- and parasubiculum, as well as the entorhinal cortex, receive fibers predominantly in their superficial layers. Different parts of the Re, including the ventral reuniens nuclei, have slightly different projection patterns (Van der Werf et al., 2002). Apart from basal ganglia structures, several of the midline and intralaminar nuclei reach other subcortical structures. Most midline and intralaminar nuclei send collaterals to the rostral part of the reticular thalamic nucleus; the reuniens nucleus and the caudal part of the central medial nucleus, in addition, have some bilateral intrathalamic projections (Van der Werf et al., 2002). Different subnuclei of the amygdaloid complex are reached by individual (sub)nuclei of the midline and intralaminar complex, including its posterior representatives (Turner and Herkenham, 1991). The

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FIGURE 8 Schematic drawing of the topographical organization of the projections from the midline– intralaminar thalamic complex to the striatum (cf. Berendse and Groenewegen, 1990) and the frontal cortex (cf. Berendse and Groenewegen, 1991) and of the projections from the frontal cortex to the striatum (cf. Berendse et al., 1992). Interconnected parts of the midline and intralaminar complex, the cerebral cortex, and the striatum are indicated by corresponding colors. Solid lines indicate the convergence of connected parts of the thalamus, cerebral cortex, and striatum; broken lines mark the path of parallel circuits leading from the cerebral cortex via the basal ganglia and the thalamus back to the cortex. Abbreviations: ac, anterior commissure; CeM, central medial thalamic nucleus; CL, central lateral thalamic nucleus; DStr, dorsal striatum; fr, fasciculus retroflexus; GP, globus pallidus; GPi, internal segment of GP; IMD, intermediodorsal thalamic nucleus; LHb, lateral habenula; MD, mediodorsal thalamic nucleus; PC, paracentral thalamic nucleus; PF, parafascicular nucleus; PFC, prefrontal cortex; PFl, lateral part of PF; PFm, medial part of PF; PT, parataenial thalamic nucleus; PV, paraventricular thalamic nucleus; sm, stria medullaris; SMC, somatomotor cortex; SNR, pars reticulata of the substantia nigra; VA, ventral anterior thalamic nucleus; VL, ventral lateral thalamic nucleus; VP, ventral pallidum; VStr, ventral striatum. Slightly modified from Groenewegen and Berendse (1994).

paraventricular nucleus has projections to a wide variety of preoptic and hypothalamic nuclei. The medial part of the parvicellular subparafascicular nucleus projects to the medial preoptic area (Coolen et al., 1998). Descending projections, in particular from the parafascicular and subparafascicular nuclei, to viscero- and somatomotor-related structures in the brain stem and spinal cord (Yasui et al., 1992; Gaytan and Pasaro, 1998; Marini et al., 1999) The inferior colliculus (Winer et al., 2002), as well as to have been described.

Functional Aspects of the Midline and Intralaminar Complex Because of the small size of the individual midline/ intralaminar nuclei, the complex configuration of this collection of nuclei as a whole, and the intricate relationships of these nuclei with the neighboring thalamic nuclei, it has been very difficult up till now to experimentally and specifically probe their functions. Yet, on the basis of the specific input–output relationships, supplemented with the relatively scarce animal experimental and clinical data, inferences have been made about the functional role of the midline and

intralaminar thalamic nuclei. Thus, on the basis of the patterns of afferent connections from the brain stem (Krout et al., 2002) and a synthesis based on afferent and efferent connections (Van der Werf et al., 2002), it has been concluded that the midline/intralaminar complex is involved in visceral functions and has a role in arousal and awareness (Fig. 9). As is discussed below, the midline/intralaminar complex most probably does not act as a unitary structure in these functions. In a recent review, Van der Werf et al. (2002) proposed four functionally distinct clusters of individual midline and intralaminar nuclei on the basis of similarities in connectivity. Although all midline and intralaminar nuclei share a number of connectional characteristics (see above), each group of nuclei that is recognized has a set of afferent and efferent connections that clearly distinguishes it from the other groups. Thus, a dorsal group, having connections predominantly with the medial nucleus accumbens, the medial prefrontal cortex, and the amygdala, includes the PV, the IMD, and the PT. This group of midline nuclei, labeled as “viscerolimbic”, is suggested to play a major role in the awareness of viscerosensory stimuli. The PV is strongly influenced by monoaminergic inputs and

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FIGURE 9 Schematic representation of the broad influence of arousal-related brain stem nuclei on the midline and intralminar thalamic nuclei. Midline nuclei, in addition, receive visceral-related inputs. Through the midline and intralaminar nuclei, arousal- and visceral-related information may reach extensive areas of the cerebral cortex. This summary figure is based on a large set of tracing experiments with small injections of retrograde tracers in individual midline and intralaminar thalamic nuclei reported by Krout et al. (2002). Reproduced from Fig. 21 in Krout et al. (2002).

corticotropin-releasing hormone (Otake and Namura, 1995; Otake and Ruggiero, 1995) and receives input from the suprachiasmatic nucleus and the intergeniculate leaflet (Moore et al., 2000; Kawano et al., 2001), and is therefore thought to be involved in state-setting properties. A role of the PV in stress and fear has been documented (Beck and Fibiger, 1995; Chastrette et al., 1991), while also visceral processing and visceral feedback regulation have been attributed to the PV (Moga et al., 1995; Ruggiero et al., 1998). A lateral group, consisting of the PC and the CL and the anterior part of the CM, is concidered to have its predominant connections with the dorsal striatum and the anterior cingulate cortex. This group of nuclei is thought to be involved in executive functions, in particular in cognitive awareness (Van der Werf et al., 2002). This assumption is primarily based on human

studies. However, in line with this, lesions of the PC and CL in rats result in deficits of working memory rather than reference (long-term) memory (Young et al., 1996; Savage et al., 1997; Burk and Mair, 2001). A ventral group of nuclei, which is composed of the Re and the Rh and the posterior part of the CM, only sparsely projects to the striatum. Instead, the nuclei in this ventral group have rather widespread and strong projections to superficial and deep layers of gustatory, visceral, insular, auditory, and motor cortical areas (see above). In addition, the Re and, more sparsely, the Rh project to the hippocampus and entorhinal cortices. This group of nuclei is thought to play a role in polymodal sensory awareness (Van der Werf et al., 2002). In electrophysiological studies it has been shown that stimulation of the reuniens nucleus can modulate the information transfer in the hippocampus (Dolleman-

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Van der Weel et al., 1997; Dolleman-Van der Weel and Witter, 2000). Preliminary results of lesions of the Re support a role of this nucleus in awareness rather than in memory per se (Dolleman-Van der Weel et al., 1994). A posterior group consists of the centromedian– parafascicular complex but, as discussed above, in rats includes only the PF. The PF primarily projects to basal ganglia structures and, in addition, reaches the sensory and motor cortices. It is being postulated that the PF has an important role in the generation of motor responses following awareness of salient stimuli and, in this way, serves a role in limbic-motor functions (Van der Werf et al., 2002). Indications for an involvement of the parafascicular nucleus in the generation of motor responses follow from studies that show a role in the modulation of intracranial self-stimulation behavior and of active avoidance in which a motor response is needed to avoid a foot shock (GuillazoBlanch et al., 1999; Massanes-Rotger et al., 1998). Electrophysiological studies show that lesions of the PF may interrupt neocortical oscillatory activity in the theta range which is important for attentional, orienting, and ongoing motor responses (Marini et al., 1998). Finally, the functions of the posterior intralaminar nuclei, in particular the SPF, deserve attention. It has been known for some time that the lateral part of the parvicellular SPF is a way station in ascending auditory signals to the cerebral cortex and the amygdala (e.g., Yasui et al., 1990; Coolen et al., 2003a). The medial part of the parvicellular SPF, by contrast, appears to be involved in sexual behavior. As shown by Veening and Coolen (1998), employing the expression of c-fos as a marker for neuronal activity, the medial parvicellular SPF is specifically involved in a circuit underlying ejaculation behavior in male rats. This circuit involves an ascending spinothalamic pathway containing galanin as neurotransmitter, terminating in the medial parvicellular SPF (Coolen et al., 2003b) and a number of forebrain regions like the medial amygdala and the medial preoptic area (Veening and Coolen, 1998). In female rats, activity in the medial parvicellular SPF is related to vaginocervical stimulation (Coolen et al., 1996).

RETICULAR NUCLEUS The reticular thalamic nucleus (Rt) forms a thin neuronal sheet at the rostral, dorsolateral, lateral, and ventrolateral margins of the dorsal thalamus. The Rt is strategically “placed” between the dorsal thalamus and the cerebral hemisphere, such that all incoming and outgoing fibers of the thalamus have to pass through it, most of them giving off collaterals to a restricted part of the reticular nucleus (but see below). Reticular thal-

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amic neurons have a relatively extensive dendritic tree extending in a disk-like form in the same plane as the thin sheet formed by the Rt itself (Jones, 1985; Spreafico et al., 1991; Ohara and Havton, 1996). Frequent dendrodendritic junctions, i.e., synapses and puncta adhaerentia, have been noted in parallel bundles of dendrites of Rt neurons, indicating a special form of interneuronal communication within the reticular nucleus (Pinault et al., 1997; however, see also Liu and Jones, 1999). Reticular thalamic neurons are all GABAergic and express parvalbumin (Mitrofanis, 1992), while a subset of Rt neurons contains calretinin (Lizier et al., 1997). The full extent of the reticular nucleus can be very well appreciated in parvalbumin-stained sections (see relevant Plates in Paxinos et al., 1999). The Rt in rats is a relatively well-developed nucleus that provides an important GABAergic control of the dorsal thalamus; in rats, GABAergic interneurons are relatively sparse (Price, 1995). Arcelli et al. (1997) have noted that in different species there is an inverse relationship between the number of GABAergic interneurons in the dorsal thalamic nuclei and the cellular density of the Rt. Their interpretation is that an increase in complexity of local information processing in different thalamic nuclei is accompanied by a more extensive interneuronal circuitry and a relatively small reticular nucleus. The ventral lateral geniculate and subgeniculate nuclei, which seem to form a dorsolateral and caudal extension of the caudal reticular thalamic nucleus, may be considered the “reticular” part of the visual thalamus. These nuclei have a ventral thalamic embryonic origin and their structural and connectional characteristics are very similar to those of the Rt (Jones, 1985). A specific visual sector of the Rt, however, also exists (Coleman and Mitrofanis, 1996). During prenatal development, a considerable number of neurons exist within the internal capsule, directly laterally adjacent to the reticular thalamic nucleus, forming the so-called perireticular nucleus (Mitrofanis and Guillery, 1993). These neurons have been shown to project to the dorsal thalamus (Mitrofanis et al., 1995), but most probably not to the cerebral cortex (Coleman and Mitrofanis, 1999). The perireticular neurons are thought to play a role in guiding axons during development; the function of the relatively few surviving perireticular remaining neurons in adulthood remains to be established (Amadeo et al., 1998; Coleman and Mitrofanis, 1999).

Efferent and Afferent Projections Unlike the dorsal thalamic nuclei, the Rt does not project to the cerebral cortex but sends its fibers almost exclusively to the dorsal thalamus (see also Vaccaro

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and Mitrofanis, 1997). There might be a small projection from the Rt to the lateral hypothalamus (Barone et al., 1994). The projections from the Rt to the dorsal thalamic nuclei are highly specific and topographically organized. Individual dorsal thalamic nuclei receive projections from a specific subset of spatially closely related reticular thalamic neurons (e.g., Jones, 1985; Kolmac and Mitrofanis, 1997; Pinault and Deschennes, 1998). Based on a large number of juxtacellularly filled reticular thalamic neurons, Pinault and Deschennes (1998) have shown that the terminal fields of these neurons in dorsal thalamic nuclei are well-focused with a patchy distribution of boutons and that the Rt– dorsal thalamic projections are organized largely in parallel with only a slight degree of divergence. For the relationship of the ventrobasal complex with the Rt this divergence appears to lead to an intrathalamic pathway which links this first-order thalamic nucleus via the Rt with the higher order posterior thalamic nucleus (Crabtree et al., 1998). Whereas such interconnections between different dorsal thalamic nuclei via the the reticular nucleus might exist, the high degree of topography between the Rt and individual thalamic nuclei is remarkable. The result of this strict topography is that each individual, functionally distinct thalamic nucleus is represented in a restricted sector of the Rt (Jones, 1985; e.g., anterior nuclei: Gonzalo-Ruiz et al.,) 1995; intralaminar and midline nuclei: Kolmac and Mitrofanis, 1997; visceral thalamus: Hayama et al., 1994; Stehberg et al., 2001). The relationships with the midline thalamic nuclei appear to be the least strictly organized (Kolmac and Mitrofanis, 1997). Afferents to the Rt originate from the dorsal thalamic nuclei and cerebral cortex, but also from the basal forebrain and brain stem. The thalamic inputs to the Rt, collaterals of the thalamocortical axons, are strictly topographically organized and rather faithfully reciprocate the Rt–dorsal thalamic projections (Jones, 1985; Mitrofanis and Guillery, 1993). Likewise, corticothalamic axons issue a collateral to the Rt sector associated with the dorsal thalamic nucleus to which the corticothalamic fibers are directed. In this way, the different sectors of the Rt receive cortical and thalamic glutamatergic inputs (Kharazia and Weinberg, 1994; Eaton and Salt, 1996). The terminal fields of both cortical and thalamic afferents mostly form narrow disklike patterns that are oriented perpendicular to the parent axons and conform to the shape and orientation of the dendrites of reticular neurons (Mitrofanis and Guillery, 1993). However, even though the topography of projections between the cerebral cortex and the reticular nucleus as well as between the dorsal thalamic nuclei and the reticular nucleus shows such a strict point-to-point relationship, it is unclear how

precise the relationships are at the microcircuit level (Pinault and Deschennes, 1998; Guillery et al., 1998; Sherman and Guillery, 2001). Thus, it is, for example, unclear whether the point-to-point relationships are such that strict reciprocal relationships exist between individual dorsal thalamic and reticular neurons, whether cortical afferents project to reticular neurons that target precisely the same dorsal thalamic neurons as these corticothalamic fibers, or whether “adjacent” neurons in the dorsal thalamus are innervated by these reticular thalamic neurons. The actual organization of specific cortical–reticular–dorsal thalamic microcircuits, which may even differ for different dorsal thalamic nuclei, may make the difference between a precise inhibitory feedback, an inhibitory feedforward, or different forms of lateral inhibition in dorsal thalamic nuclei (cf. also Pinault and Deschennes, 1998b; Sherman and Guillery, 2001). As discussed by Sherman and Guillery (2001), an important aspect of the innervation of the Rt may be that the modulatory, but not the driving, afferents of the dorsal thalamus have a collateral projection to the reticular nucleus. For cortical inputs this means that corticothalamic afferents from layer VI, which terminate as type I fibers in the dorsal thalamus, do have a collateral to the Rt, whereas corticothalamic fibers from layer V, which are mostly collaterals of corticofugal fibers proceeding to the brain stem or spinal cord, do not project to the Rt (Fig. 10) (Deschennes et al., 1994; Bourassa et al., 1995). Likewise, ascending driving afferents, for example from somatosensory systems in general, do not have a collateral to the Rt. Whether this distinction between type I and type II afferents in relation to the reticular nucleus can be regarded as a general rule remains to be established.

Functional Aspects The prevailing interpretation of the functional role of the Rt is that it serves attentional brain mechanisms (e.g., “searchlight hypothesis”) (Crick, 1984; Guillery et al., 1998, McAlonan et al., 2000). The abovedescribed topographical and specific arrangement of the afferents and efferents of different sectors of the Rt indicates a specific role of the reticular nucleus for functionally distinct thalamic nuclei, contrasting with the general, more global reticular influence on thalamocortical mechanisms that has been proposed originally (Scheibel and Scheibel, 1966; Steriade et al., 1993). The Rt is important for the control of the firing mode of thalamocortical projection neurons and, in this way, for the selection of the information that is transferred from the thalamus to the cerebral cortex. The reticular nucleus plays an important role as pacemaker during

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FIGURE 10 Schematic representation of the intricate relationships of the reticular thalamic nucleus (Rt) with the dorsal thalamus and specific layers of the cerebral cortex (from Guillery et al., 1998). The reticular thalamic nucleus (Rt) receives collaterals from thalamocortical fibers originating in virtually all nuclei of the dorsal thalamus, while most of the corticothalamic fibers likewise issue a collateral to the Rt. However, as such collaterals are common for the corticothalamic fibers originating in layer VI, the “modulatory” corticothalamic projections (see page 408 of this chapter), collaterals are absent from corticothalamic fibers arising from more superficial layers. This is correlated with a distinction in firstorder and higher order thalamic nuclei and their cortical associations. So-called first-order (FO) thalamic nuclei receive their main “driving” afferents from ascending specific afferents, such as somatosensory or visual modalities, while so-called higher order (HO) thalamic nuclei receive their main driving afferents from layer V of the cerebral cortex (see page 408 of this chapter). Thalamic afferents originating from layer V of the cortex and primary ascending fibers are morphologically very comparable, terminating as RL-type boutons, and thought to be the drivers of the various thalamic nuclei (Sherman and Guillery, 2001; see page 408 of this chapter). However, neither descending layer V axons nor ascending sensory afferents issue a collateral to the reticular thalamic nucleus. Apart from the dorsal thalamic and deep cerebral cortical layers, the reticular thalamic nucleus is targeted by several brain stem and basal forebrain inputs. Slightly modified from Fig. 2 in Guillery et al. (1998).

synchronized firing of thalamocortical cells. Basically, two different firing patterns exist in the thalamocortical system, i.e., a “tonic” and a “burst mode” (Jahnsen and Llinás, 1984; McCormick and Fraser, 1990). In the tonic mode of thalamocortical activity, information from, for example, ascending sensory pathways, is almost linearly transferred through the thalamus to the cerebral cortex. Burst firing occurs during sleep, as well as during epileptic seizures. In the bursting mode, relay of information to the cerebral cortex is either prevented or nonlinearly transformed. Thus, in the latter case, thalamocortical neurons may still respond during burst firing but the message is not transferred to the cortex in the same form as during tonic firing (Guido et al., 1995; Guillery et al., 1998). During burst firing there is a higher signal-to-noise ratio, providing a mechanism to select specific, novel information to reach the cortex (Guido and Weynand, 1995; Guillery et al., 1998).

The precise physiological mechanisms by which the Rt exerts its gating role remain to be established. There might be a prominent role for the descending corticothalamic fibers, at least in the visual system (Montero, 2000), and this may provide a very topical mechanism within a specific sensory system to attend to a specific stimulus (see also Hartings et al. (2000) for the somatosensory, vibrissal system). Numerically, cortical inputs to the Rt dominate other inputs (Liu and Jones, 1999). Subcortical afferents, originating from the brain stem reticular formation, the basal forebrain, or the dorsal thalamus (i.e., the parafascicular nuleus) also exert their influence on particular sectors of the Rt, although apparently less focused (Kolmac and Mitrofanis, 2001). Notwithstanding the recently elucidated specificity in the connections and functions of the Rt, the nucleus might also act as a unit and generate general synchronized thalamocortical activity, in this way “closing” the thalamocortical gate like during

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sleep. The mechanisms by which the synchronicity of reticular neurons is achieved are largely unknown, but electrical synapses may play an important role (Landisman et al., 2002).

Acknowledgments The authors thank Ingrid Riphagen for performing an extensive literature search and Sanderien van Oudenaren for her help in preparing the reference list. Dr. Piet Hoogland gave critical and supportive comments on parts of the manuscript. The authors are also grateful for the assistance of Han Verbeek and Dirk de Jong who helped with the illustrations.

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Yoshida, A., Dostrovsky, J. O., and Chiang, C. Y. (1992). The afferent and efferent connections of the nucleus submedius in the rat. J. Comp. Neurol. 324, 115–133. Yoshida, A., Dostrovsky, J. O., Sessle, B. J., and Chiang, C. Y. (1991). Trigeminal projections to the nucleus submedius of the thalamus in the rat. J. Comp. Neurol. 307, 609–625. Young, H. L., Stevens, A. A., Converse, E., and Mair, R. G. (1996). A comparison of temporal decay in place memory tasks in rats (Rattus norvegicus) with lesions affecting thalamus, frontal cortex, or the hippocampal system. Behav. Neurosci. 110, 1244–1260. Zahm, D. S. (2000). An integrative neuroanatomical perspective on some subcortical substrates of adaptive responding with emphasis on the nucleus accumbens. Neurosci. Biobehav. Rev. 24, 85–105. Zhang, Z. W., and Deschenes, M. (1998). Projections to layer VI of the posteromedial barrel field in the rat: A reappraisal of the role of corticothalamic pathways. Cereb. Cortex 8, 428–436.

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C H A P T E R

18 Basal Ganglia CHARLES R. GERFEN Laboratory of Systems Neuroscience, NIMH

The basal ganglia connect the cerebral cortex with neural systems that effect behavior. Most cortical areas provide inputs to the basal ganglia, which in turn provide outputs to brain systems that are involved in the generation of behavior. Among the behavior effector systems targeted are thalamic nuclei that project to those frontal cortical areas involved in the planning and execution of movement; midbrain regions including the superior colliculus, which is involved in the generation of eye movements; the pedunculopontine nucleus, which is involved in orienting movements; and hypothalamic systems involved in autonomic functions. Two points concerning the function of the basal ganglia are emphasized. First, while the basal ganglia connect the cerebral cortex with a wide range of behavior effector systems, the basal ganglia operates in parallel with other output systems of the cerebral cortex. These other corticofugal systems may have a more primary role in the actual generation of behavior. For example, the frontal cortical areas involved in the planning and execution of movement behavior provide direct projections, via direct corticospinal projections, that are responsible for the generation of movement. Thus, the role of the basal ganglia in affecting cortically generated behavior remains unclear. Second, while the basal ganglia are connected with a wide range of behavior effector systems, not all regions of the basal ganglia are connected with all of the output systems. In other words, there is a conservation of regional functional organization of the cerebral cortex in the connections of the basal ganglia. In considering the neuroanatomical organization of the basal ganglia there are differing views. On the

The Rat Nervous System, Third Edition

one hand, the basal ganglia have been proposed to provide for interactions between disparate functional circuits, for example, between so-called “limbic” and “nonlimbic” functions. Another view holds that there are parallel functional circuits, in which distinct functions are for the most part maintained, or segregated, one from the other. This review is biased toward the view that there is maintenance of functional parallel circuits in the organization of the basal ganglia, with considerable interaction between adjacent circuits. The organization of the basal ganglia is intimately linked to that of the cerebral cortex, with distinct differences between those regions of the basal ganglia that receive inputs from neocortical, six-layered cortex, compared with those receiving inputs from allocortical areas. This review focuses primarily on the neocortical part of the basal ganglia. A general canonical organizational plan of the neocortical-related basal ganglia is described. Understanding the organization of the neostriatal part of the basal ganglia provides a framework for determining the general organizational principles of the parts of the basal ganglia connected with allocortical areas and the amygdala. The components of the canonical basal ganglia system, which is the subject of this review, include the neocortex, the striatum, which includes the caudate– putamen and the core of the nucleus accumbens, the globus pallidus (lateral segment), the subthalamic nucleus, the medial globus pallidus (also termed the medial globus pallidus), and the substantia nigra (Fig. 1). The major input to this system comes from layer 5 glutamatergic neurons from nearly all areas of the

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FIGURE 1 Main components of the basal ganglia shown on coronal diagrams of The Rat Brain in Stereotaxic Coordinates (Paxinos and Watson, 1998). The striatum, which includes the caudate–putamen (Cpu) and the core of the nucleus accumbens (acc), is the main input nucleus of the basal ganglia. The output nuclei of the basal ganglia are the medial globus pallidus (MGP) and substantia nigra pars reticulata (SNR). The striatum is connected through direct and indirect pathways with the output nuclei. The indirect pathway includes the globus pallidus (GP) and subthalamic nucleus (STh). Targets of the output of the basal ganglia include the mediodorsal (md), ventromedial (vm), ventrolateral (vl), and parafascicular (pf) thalamic nuclei; the intermediate gray layer (InG) of the superior colliculus (sc); and the pedunculopontine tegmental nucleus (PPT). Dopamine neurons are located in the ventral tegmental area (VTA), substantia nigra pars compacta (dorsal tier, SNCD; ventral tier, SNCV). Levels are taken from Paxinos and Watson (1998).

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FIGURE 1, cont’d Main components of the basal ganglia shown on coronal diagrams of The Rat Brain in Stereotaxic Coordinates (Paxinos and Watson, 1998). The striatum, which includes the caudate–putamen (Cpu) and the core of the nucleus accumbens (acc), is the main input nucleus of the basal ganglia. The output nuclei of the basal ganglia are the medial globus pallidus (MGP) and substantia nigra pars reticulata (SNR). The striatum is connected through direct and indirect pathways with the output nuclei. The indirect pathway includes the globus pallidus (GP) and subthalamic nucleus (STh). Targets of the output of the basal ganglia include the mediodorsal (md), ventromedial (vm), ventrolateral (vl), and parafascicular (pf) thalamic nuclei; the intermediate gray layer (InG) of the superior colliculus (sc); and the pedunculopontine tegmental nucleus (PPT). Dopamine neurons are located in the ventral tegmental area (VTA), substantia nigra pars compacta (dorsal tier, SNCD; ventral tier, SNCV). Levels are taken from Paxinos and Watson (1998).

neocortex. The output of this system is the projection of GABAergic neurons in the medial globus pallidus and the substantia nigra pars reticulata. This basal ganglia output targets thalamic nuclei, which project to frontal cortical areas involved in the planning and execution of movement behavior; the intralaminar thalamic nuclei, which provide inputs to the neocortex and the striatum; the intermediate layers of the superior colliculus, which are involved in the generation of eye and head movements; and the pedunculopontine nucleus, which is involved in orienting movements of the body. In between the neocortical inputs and the GABAergic output systems are the neuroanatomical circuits that comprise the prototypical basal ganglia. The main input structure of the basal ganglia is the striatum. Those regions of the striatum that receive inputs from neocortical areas are the caudate–putamen and core of the nucleus accumbens. The targets of the neocortical input are medium-sized spiny GABAergic

projection neurons, which account for over 90% of neurons in the striatum. These neurons are divided into two types, which give rise to the two main components of the prototypical basal ganglia circuit, the “direct” and “indirect” striatal projection systems. The “direct” striatal projection system is so-named as these neurons provide direct inputs to the output neurons of the basal ganglia in the medial globus pallidus and the substantia nigra pars reticulata. Indirect striatal projection neurons provide inputs to the globus pallidus, which together with the subthalamic nucleus compose the major components of the “indirect” basal ganglia circuit. GABAergic neurons in the lateral globus pallidus project back to the striatum, to the output neurons of the basal ganglia in the medial globus pallidus and substantia nigra, and to the subthalamic nucleus. The subthalamic nucleus, which itself receives inputs from the neocortex, provides excitatory projections to the output neurons of the basal ganglia.

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In the canonical basal ganglia circuit (Fig. 2), the neocortex provides excitatory inputs to the striatum, whose output, through the direct and indirect projection systems, provide both inhibitory and excitatory regulation of the output of the basal ganglia. The output neurons of the basal ganglia, GABAergic neurons in the medial globus pallidus and substantia nigra pars reticulata, display a relatively high level of tonic activity. In a long held model of basal ganglia function, the excitatory input from the cortex has been demonstrated to function through a disinhibitory mechanism. Thus, activation of the direct output neurons of the striatum by the excitatory input from the neocortex results in inhibition of the tonic inhibitory output of the basal ganglia. The role of the indirect circuit is more complex. On the one hand, the target of the indirect striatal output neurons are GABAergic neurons in the lateral globus pallidus, which project to the output neurons of the basal ganglia and to the subthalamic nucleus. Thus, cortical excitation of the indirect pathway inhibits the GABAergic pallidal output, resulting in disinhibition of the output neurons of the basal ganglia and the subthalamic nucleus. The subthalamic nucleus, which receives direct excitatory inputs from the cerebral cortex, provides excitatory inputs to the output neurons of the basal ganglia. Additionally, it has been demonstrated that the interconnections between the lateral globus pallidus and subthalamic nucleus generate an oscillatory pattern of activity that is conveyed to the output neurons of the basal ganglia. Given the complexities of the organization of these circuits at this time the specific mechanisms responsible for regulating the output of the basal ganglia remain to be established. However, in general terms the activity in the direct and indirect striatal output pathways may be viewed as providing counterbalanced or antagonistic regulation of the output of the basal ganglia. Overlain on the canonical basal ganglia circuit are a number of additional neuroanatomical features that add to the complexity of the organization of this system. Notable among these is the dopaminergic nigrostriatal system, which provides a massive dopaminergic input to the striatum from the midbrain dopamine neurons in the ventral tegmental area and substantia nigra pars compacta. In addition, this review describes the following features of basal ganglia organization: (1) the organization of the corticostriatal system, which incorporates both a general topographic organization with considerable overlap of corticostriatal inputs from cortical areas that are interconnected; (2) the patch and matrix compartmental organization of the striatum, which is related to the laminar organization of the cerebral cortex and provides differential inputs to the output systems of the basal ganglia and the nigros-

triatal dopaminergic system; and (3) the dual representation of striatal outputs in the lateral globus pallidus and output nuclei of the basal ganglia.1

CORTICAL INPUT TO THE STRIATUM The cerebral cortex provides a major input to the striatum. Corticostriatal afferents arise from cortical pyramidal neurons, located primarily in layer 5, with some being distributed in layer 3 and rarely in layer 6. These neurons utilize glutamate as neurotransmitter and provide the major excitatory input to the striatum. Subtypes of corticostriatal neurons may be characterized on the basis of their axonal collateral connections with cortical and subcortical areas and features of their axonal arborization within the striatum.

Subtypes of Corticostriatal Neurons Two main subtypes of corticostriatal neurons have been identified based on the distribution of their subcortical axon collaterals. One type may be considered a corticocortical/corticostriatal neuron, in that it provides axon collaterals that distribute only within the striatum and cerebral cortex (Wise and Jones, 1977; Wilson, 1987; Cowan and Wilson, 1994; Zheng and Wilson, 2002). This type of cell is very numerous in agranular cortical regions giving rise to bilateral corticocortical and corticostriatal projections. These neurons are located in a band at the superficial half of layer 5 and in the deep half of layer 3. 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 second type of corticostriatal neuron is classified as a pyramidal tract or brain stem projecting neuron. These neurons are 1

In this review the entopeduncular nucleus is referred to as the medial globus pallidus. In the primate, the globus pallidus is divided into two parts, the lateral or external globus pallidus and the medial or internal segment of the globus pallidus. In the rat, the term globus pallidus generally refers to the lateral or external globus pallidus, whereas the entopeduncular nucleus is the equivalent of the internal or medial globus pallidus. As The Rat Brain in Stereotaxic Coordinates of Paxinos and Watson (1998) uses the term medial globus pallidus to refer to the entopeduncular nucleus, this terminology is used in this review.

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FIGURE 2 Connections of the basal ganglia associated with the neocortex shown on a sagittal diagram of the brain. (A) The major input and output connections of the basal ganglia. Layer 5 pyramidal neurons of most areas of the cerebral cortex provide a major input to the striatum, which comprises the caudate–putamen (CP) and the core of the nucleus accumbens (Acc). The output of the basal ganglia arises from GABAergic neurons in the medial lateral globus pallidus (MGP) and substantia nigra pars reticulata (SNR). These neurons provide inhibitory inputs to thalamic nuclei, including the ventrolateral (vl), mediodorsal (md), and ventromedial (vm) nuclei, as well as to the superior colliculus (SC) and pedunculopontine nucleus (PPN). (B) The direct and indirect striatal projection pathways from two subsets of striatal medium spiny neurons. Direct projecting neurons provide an axon with collaterals to the lateral globus pallidus (LGP) and to the medial globus pallidus (MGP) and substantia nigra pars reticulata (SNR). Indirect striatal projection neurons project to the lateral globus pallidus (LGP). These neurons are indirectly connected to the medial globus pallidus and substantia nigra through connections that involve the lateral globus pallidus and subthalamic nucleus (STN). (C) Feedback pathways of the basal ganglia include the dopaminergic nigrostriatal dopamine (DA) pathway from the substantia nigra pars compacta, a pallidostriatal pathway from the lateral globus pallidus to the striatum, and a thalamostriatal pathway from the intralaminar and parafascicular (pf) thalamus to the striatum and from the vl, md, and vm thalamic nuclei, which project to the frontal cortex.

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located primarily in frontal cortical areas and give rise to pyramidal tract projections to the brain stem or the spinal cord, which have been shown to contribute axon collaterals to the striatum (Cowan and Wilson, 1994; Donoghue and Kitai, 1981; Landry et al., 1984). Within the cortex, these neurons are distributed, for the most part, deeper than the first type of corticostriatal neuron, in the deeper parts of layer 5, and occasionally in layer 6. 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. This projection has attracted some interest because of its potential for providing the neostriatum with a copy of the cortical motor signal. A subset of these neurons also appears to provide projections to the thalamus (Royce, 1983; Levesque et al., 1996, 1998). Corticostriatal neuron subtypes are also distinguished by having either a focal or extended distribution of axonal arborizations within the striatum (Cowan and Wilson, 1994; Kincaid et al., 1998; Wright et al., 1999). One subtype is characterized as having a focal axonal arborization pattern within the striatum (Fig. 3A). Such axons often arise as a collateral of the major axon of a pyramidal tract neuron as it courses through the striatum in white matter fascicles. Individual collaterals may extend up to 1 mm from the main axon, with a few side branches that may end with a small cluster of bouton-bearing terminal branches. A single axon may give rise to several of these striatal collateral branches, which may be separated by more than a millimeter in some cases. Synaptic varicosities are distributed at regular intervals on these types of collaterals; however, the number of varicosities is extremely sparse. Axon collaterals of neurons providing focal striatal afferents distribute focal axon collaterals within the cortex as well. Axon collaterals often spread within layer 5, with individual axon collaterals spreading into nearby cortical areas ascending into supragranular layers with focal arborizations in layer 1. The other subtype of corticostriatal afferent forms an extended axonal arborization pattern in the striatum, occupying an area with dimensions of 1 mm or greater (Fig. 3B). Within that volume the axon occupies space in a very sparse fashion, with individual branches running approximately parallel and separated by large uninnervated areas. These afferent fibers have very irregular distribution of synaptic varicosities, which, although sparse, appear clustered in particular regions. The intracortical distribution of axon collaterals of cortical neurons providing extended striatal afferents was rather widespread, arborizing locally within the area of the parent neuron, while other collaterals distributed to adjacent cortical areas, with extensive arborizations in layers 5 and 3. Thus, the patterns of intracortical and striatal

distributions of the focal and extended types of corticostriatal neurons appear to be somewhat correlated. There does not appear to be a simple classification scheme relating the subtypes of corticostriatal neurons based on their axonal arborizations within the striatum and whether they arise from collaterals of pyramidal tract axons or as afferents of neurons with pure cortical and striatal distributions. In an early paper it was reported that focal and extended corticostriatal afferents arose, respectively, from the pyramidal tract subtype and pure corticostriatal subtypes of corticostriatal neurons (Cowan and Wilson, 1994). However, a more recent study, analyzing a larger number of neurons, demonstrated that both extended and focal corticostriatal afferents arose from pyramidal tract neurons (Zheng and Wilson, 2002). As the determination of these different subtypes requires data gathered from individual neurons, at this point there is no way to estimate the relative number of the different subtypes. Nonetheless, there appear to be subtypes of corticostriatal neurons, whose distinct patterns of connections determine the organization of the patterns of information that the cerebral cortex provides to the striatum.

Patterns of Organization of Corticostriatal Afferents Cortical input to the striatum originates from most cortical areas, including primary and higher order sensory areas; motor, premotor, and prefrontal regions; and 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; McGeorge and Faull, 1989; Ebrahimi et al., 1992). 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 has 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. Topographic Corticostriatal Organization The topographic organization of corticostriatal inputs is well established with the orderly mapping of inputs

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FIGURE 3 Tracings of dendrites (black) and cortical (A and B) and striatal axons (A′ and B′) of two subtypes of corticostriatal neurons, which had been intracellularly labeled. (A) The corticostriatal neuron depicted provides an axonal projection to the pyramidal tract. Axon collaterals within the cortex are distributed in relatively close proximity to the parent neuron. (A′) The pyramidal tract axon of this neuron extends collaterals in the striatum, which display a focal terminal arborization. (B) The corticostriatal neuron depicted, located in the medial agranular cortex (AGm), is a bilaterally projecting corticocortical neuron, with axon collaterals projecting bilaterally to the striatum. Axon collaterals of this neuron within the ipsilateral striatum are distributed both locally around the parent neuron in the AGm and extend to the adjacent lateral agranular motor cortical area (AGl). (B′) This neuron provides an axon that provides an extensive arborization pattern within the striatum and does not provide collaterals that extend beyond the striatum. These neuronal tracings are from Cowan and Wilson (1994).

of regions of the cortex into topographically related regions of the striatum. Thus, for example, frontal areas provide inputs to the rostral regions of the striatum, sensorimotor cortex provides inputs to the dorsal lateral region, and parietal cortex provides inputs to more caudal regions. The conservation of this general topographic organization of corticostriatal inputs through the rest of the basal ganglia circuits is the basis of the popular concept of the existence of functional parallel loops (Alexander et al., 1986). An example of the regional topographic organization of the corticostriatal projections is provided by the mapping of the

various motor cortical area projections (Ebrahimi et al., 1992). Such mappings demonstrate that the spatial relationship of motor cortical areas are preserved in the organization of inputs of these areas into the striatum, for example, frontal eye fields are medial to limb and body areas. The projections from functionally defined cortical areas may be rather precise in terms of representing somatotopic organization. Thus, areas such as the somatosensory cortex, which have a somatotopic organization, provide a similar somatotopic organization of corticostriatal afferents preserving the spatial

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organization of hindlimb and forelimb areas, for example (Brown et al., 1998). Such somatotopically organized corticostriatal representations may be quite precise. The striatal projections from adjacent “barrels” in the somatosensory barrel cortex have been shown to map to adjacent sites within the striatum (Wright et al., 1999). This study, using analysis of individual corticostriatal axons showed two types of corticostriatal afferent systems. One is discrete, which maintained, rather precisely, the relationship between barrels. The other is more diffuse, which provided for considerable overlap of the projections of different barrels within the striatum. Thus, the corticostriatal inputs map not only in a general manner but also in a quite precise topographic manner to the striatum; there also exists a corticostriatal system, which provides for overlapping and converging projections from different cortical areas. Convergent Corticostriatal Organization In addition to the topographic organization of the corticostriatal system, another organizational feature of the corticostriatal system is the fact that a given cortical area projects to an area 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 rostrocaudal dimension. From their work in primates, 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 rostrocaudal area. As these areas are connected by corticocortical connections they suggested that corticostriatal organization is related to the cortical connections of the area from which the corticostriatal inputs arise. They formulated a rule, which suggested that areas of cortex that are interconnected by corticocortical connections provide overlapping inputs to the striatum, whereas areas of cortex that are not interconnected do not. Theirs was the first formulation of a concept to explain the widespread nature of corticostriatal projections from a given region. The problem of relating the organization of corticocortical 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 rostrocaudal 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 the studies of Kunzle in the 1970s (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 organized 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). In primates, Graybiel and her colleagues have examined the connectional basis of this organization in a set of studies in which they examined the relationship of corticostriatal inputs from cortical areas that had been mapped in terms of somatosensory and motor function (Flaherty and Graybiel, 1991, 1993; Parthasarathy et al., 1992). In one study the projections from different somatosensory areas were examined (Flaherty and Graybiel, 1991, 1993b). Electrophysiologically mapped regions of somatosensory areas 3a, 3b, and 1 were injected with anterograde tracers (Flaherty and Graybiel, 1991). They found that injections into matched body part representation sites in different cortical 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 nonoverlapping striatal regions. A similar organization was also found in motor cortical areas. Parthasarathy et al. (1992) examined the corticostriatal 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 nonoculomotor 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 (S1) and primary motor (M1) 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 were directed to the putamen and distributed in multiple, discontinuous zones. What they also found was that when somatotopically related areas of M1

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and S1 were injected, such as the hand representation, that the discontinuous zones of each cortical projection zone overlapped in the ipsilateral putamen.

General Organization of Corticostriatal Inputs A working model of the organization of corticostriatal inputs is that each cortical area provides a principal input to a topographically related area of the striatum and secondary inputs to the principal topographic striatal areas of those cortical areas with which it is connected. This organization is apparent in the results described above for corticostriatal afferents from somatotopically organized cortical areas, which provide multiple somatotopically organized projections to the striatum. Moreover, projections of different somototopically organized cortical areas provide overlapping inputs of related body part representations. These relationships appear to apply not only to the somatotopic representations of multiple sensory or motor cortical areas but also to overlap of somatotopically related areas of motor and sensory corticostriatal afferents. The organization of corticostriatal inputs from somatotopically organized cortical areas displays some relatively straightforward principles of convergent organization. Individual cortical areas provide inputs to multiple striatal zones, which are somatotopically organized. Convergence occurs in the somatotopically organized overlap from other cortical areas that are somatotopically organized. However, even in these corticostriatal inputs there are other patterns of overlap. An example is provided in the organization of the striatal inputs from the barrel cortex (Wright et al., 1999). It has been shown that projections of individual cortical barrels provide inputs that target discrete zones of the striatum, with adjacent barrels projecting to adjacent zones of the striatum. Additionally, there are other projections from interbarrel areas of the barrel cortex that provide a diffuse projection that overlaps several of the more discrete barrel projections. Thus, even in this very precisely somatotopically organized cortical area, there are both precise somatotopically organized corticostriatal inputs and diffuse corticostriatal inputs. In these cases the different patterns of corticostriatal inputs mirror the organization of the intracortical connections of the parent neurons. In the case of those corticostriatal inputs from individual barrels, the intracortical connections are restricted within the barrel, whereas those interbarrel cortical neurons that provide diffuse corticostriatal inputs have intracortical connections that span several barrels. Data from the corticostriatal input patterns from individual corticostriatal neurons suggest a relationship between the pattern of intracortical connections

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and the pattern of distribution of corticostriatal inputs (Cowan and Wilson, 1994). Thus a corollary of the working model of the organization of corticostriatal afferents stated above is that patterns of corticostriatal axon distributions are related to the pattern of the intracortical connections of the parent corticostriatal neuron. While the organizational patterns underlying convergence of corticostriatal inputs from somatotopically organized cortical areas provide some comprehensible principles, the divergent/convergent principles underlying the organization of corticostriatal inputs from areas that do not have such straightforward functional organization are more difficult. Nonetheless, there are clearly patterns of overlap of inputs from so-called associational cortical areas, such as the parietal and prefrontal cortical areas. It is most likely that similar organization principles apply, namely, that cortical areas that are interconnected provide convergent inputs to the striatum as originally suggested by Yeterian and Hoesen (1978). The rules governing patterns of convergence corticostriatal inputs of such associational cortical areas likely reflect that rules governing the corticocortical connections between such areas. In summary, the organization of corticostriatal inputs appears to be governed by two general principles. The first is that there is a topographic organization of inputs. The second is that there are secondary patterns of inputs that appear to mirror the patterns of corticocortical connectivity. These secondary patterns of corticostriatal inputs provide for patterns of convergence among distributed sets of corticostriatal neurons. Such patterns of convergence of corticostriatal inputs are as varied as the patterns of connectivity between cortical areas.

Quantitative Data Regarding Corticostriatal Inputs 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. Thus, the inputs of an individual corticostriatal neuron to the striatum may be considered extremely sparse in terms of the number of contacts it makes with a single striatal neuron. Wilson and his colleagues

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(Kincaid et al., 1998; Zheng and Wilson, 2002) have provided some informative quantitative information that provides some informative boundaries for the type of information processing that may be taking place within the basal ganglia. First, there appears to be roughly a 6:1 ratio in terms of the numbers of corticostriatal neurons (17,000,000) and striatal projection neurons (2,800,000). Second, the area over which the dendrites of a single medium spiny projection neuron spreads (400 μm in diameter) contains approximately 2850 other neurons. Third, approximately 380,000 corticostriatal neurons innervate the area of the dendritic field of a single medium spiny projection neuron, which contains 2850 neurons. Fourth, the axon of a single corticostriatal axon traversing this area has on average 40 synaptic boutons. If, as is estimated, each axon makes only a single, or a few contacts, with an individual medium spiny neuron, then each corticostriatal input makes contact with about 1% of the striatal neurons in the area across which it extends. Taken together these quantitative estimates indicate that the cortical input to a single striatal medium spiny projection neuron is rather unique; that is, no two striatal neurons share common inputs from the cortex. Based on their quantitative analysis of corticostriatal inputs Wilson and his colleagues (Kincaid et al., 1998; Zheng and Wilson, 2002) make several observations on the constraints of the information processing within the striatum. First, they make the point that individual striatal projection neurons receive inputs from a large number of cortical neurons, such that the cooperative effect of activity of those cortical neurons is required to provide sufficient excitatory input for the projection neuron to generate an action potential. Thus, each striatal neuron’s activity represents the coordinated activity of a distributed ensemble of a large set of cortical neurons, whose input converges on that neuron. Second, such an organization has been suggested to represent a competitive network, in which different patterns of cortical activity compete for representation in the striatum. Typically, in such competitive models, synaptic plasticity is proposed to be involved in the resolving competing representations. However, such models require considerable overlap of corticostriatal inputs to individual neurons rather than the observed unique inputs that striatal neurons receive. Thus, the organization of corticostriatal inputs appears to represent the end stage of a competitive network rather than a dynamic one in which different patterns of corticostriatal inputs are competing for synaptic sites. Third, the possibility that the striatum generates a compact representation of patterns of cortical activity is not supported by the organization of corticostriatal inputs. Wilson argues that since striatal neurons each

receive a unique set of corticostriatal inputs the 6 to 1 ratio of cortical input neurons to striatal target neurons does not allow for all patterns of cortical activity to be represented in the patterns of activity in the striatum. Thus, rather than removing redundancy in patterns of corticostriatal input activity, a large proportion of cortical patterns of activity must either be lost or treated as though they are similar when they are not.

STRIATUM The striatum comprises the caudate, putamen, and nucleus accumbens. The striatum is composed of one principal neuron cell type, the medium spiny projection neuron (Bishop et al., 1982; DiFiglia et al., 1976; Wilson and Groves, 1980). This medium 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 striatum 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 (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 (Fig. 4). The remaining striatal neurons are interneurons (Bishop et al., 1982; DiFiglia et al., 1976), in that they do not provide projection axons out of the striatum, 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 and Kitai, 1988). 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; Kawaguchi et al., 1995).

Medium Spiny Projection Neuron Medium 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), with a cell body approximately 20–25

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μm 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 μm 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). Medium spiny projection neurons also provide an axon collateral that projects out of the striatum to the lateral globus pallidus and/or medial globus pallidus/ 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 lateral globus pallidus. The other subset provides an axon collateral to the lateral globus pallidus and additional collaterals to the medial globus pallidus and/or the substantia nigra. These latter neurons constitute the “direct striatal projection pathway” as they, are directly connected to the output of the basal ganglia, which are GABAergic neurons in the medial globus pallidus and substantia nigra pars reticulata. The former neurons constitute the “indirect striatal projection pathway” as they are connected indirectly, through connections of the lateral globus pallidus and subthalamic nucleus, with the output neurons of the basal ganglia. Medium 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 lateral 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 are discussed in further detail below. Medium spiny projection neurons receive inputs from the cortex, thalamus, and amygdala, which make

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asymmetric synapses on dendritic spines and, to a lesser degree, dendritic shafts (Fig. 5). 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. Inputs to Medium Spiny Neurons Cortical inputs 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, 1971; Somogyi et al., 1981). According to a 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, as discussed above, input from any individual corticostriatal axon to an individual striatal spiny neuron is very sparse (Cowan and Wilson, 1994; Kincaid et al., 1998; Zheng and Wilson, 2002). Consistent with the asymmetric character of corticostriatal synapses onto spiny neurons electrophysiologic studies have demonstrated that corticostriatal input evokes a monosynaptic excitatory postsynaptic 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 brain stem and evokes an EPSP with a latency of 3 ms. 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 ms. Thalamic inputs Thalamic inputs from the intralaminar nuclei, including the parafascicular/centromedian complex, provide inputs to the striatum that are similar to cortical afferents in that they form asymmetric synaptic contacts and have strong excitatory effects on

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D FIGURE 4 The medium spiny striatal projection neuron. (A) Photomicrograph of a single medium spiny projection neuron filled with biocytin. (A′) High magnification of intracellularly filled medium spiny neuron. (B) Tracings of an indirect and direct striatal projection neuron drawn in place on a sagittal brain diagram. The indirect striatal pathway neuron has a projection axon that extends into the lateral globus pallidus, where it arborizes, and does not extend beyond this nucleus. The direct striatal pathway neuron has a projection axon that extends some collaterals into the lateral globus pallidus (LGP) and extends to the medial globus pallidus (MGP) and substantia nigra. Higher magnification of the indirect and direct striatal pathway neurons show their dendrites (red/green) and local axon collaterals within the striatum (orange/blue). (C) Diagrammatic representation of the direct and indirect striatal pathway neurons. Both neurons are GABAergic and receive a glutamatergic corticostriatal input. Direct pathway neurons express the D1 dopamine receptor subtype, the Gs and Golf stimulatory G proteins, as well as the peptides substance P (SP) and dynorphin (DYN). These neurons project to the lateral globus pallidus (LGP), medial globus pallidus (MGP), and substantia nigra pars reticulata (SNR). Indirect striatal pathway neurons express the D2 dopamine receptor, which is coupled to the inhibitory Gi G protein, as well as the α2-adenosine receptor, which is coupled to the stimulatory Golf G protein. These neurons also express the peptide enkephalin (ENK). (D) Functional dissociation of direct and indirect striatal projection neurons. In dopamine-depleted striatum, D1 receptor agonist treatment results in phosphorylation of ERK1/2/MAPkinase (immunoreactive green neurons) selectively in direct striatal projection neurons, but not in indirect pathway neurons, which are labeled by ISHH localization of mRNA encoding ENK (red neurons). One neuron in this field is double labeled (yellow). From Gerfen et al. (2002).

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input to cell body/proximal dendrites from interneurons

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B connections of striatal medium spiny neurons FIGURE 5 Diagram of the connections of medium spiny striatal projection neurons. (A) Synaptic input to medium spiny striatal projection neurons showing the location spine the cortical glutamatergic synapses on the head of medium spiny neurons, inputs to the neck and interspine dendritic shafts from nigrostriatal dopamine inputs, or GABAergic input from other medium spiny neurons. Inputs to the cell body and proximal dendrites are from striatal GABAergic and cholinergic interneurons. (B) Diagram of the inputs to the proximal and distal parts of striatal medium spiny neurons. Inputs to distal parts of the dendrites are from the cerebral cortex, nigrostriatal dopamine afferents, and thalamus, whereas inputs to the proximal part of the neurons are from GABAergic parvalbumin (Pv) and cholinergic (ChAT) neurons.

the spiny cells. Early studies of Powell and Cowan (1956) suggested that the thalamostriatal pathway consisted of a single topographically organized projection. More recent studies (Dube et al., 1988; Xu et al., 1989) have shown that this pathway, like the corticostriatal

projection, is heterogeneous in nature. There are actually two independent thalamostriatal projections of the intralaminar nuclear complex, one originating from the parafascicular/centromedian nuclei and a separate one from rostral parts of the complex including the

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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 with the spines. In one study (Xu et al., 1989), 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 not receive a cortical input or at least do not receive as dense a 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. (1989), 93% 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. Dopamine inputs Inputs from midbrain dopamine neurons that project to the striatum make synaptic contact with medium spiny neurons. These afferents have been identified at the ultrastructural level with immunohistochemical localization of either dopamine (Voorn et al., 1986) or the dopamine-synthesizing 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 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 dopamine-containing synapses are sometimes seen to be in close apposition with the presynaptic part of asymetric forming boutons, presumably of cortical or thalamic origin. However, these close appositions lack synaptic specializations, and they

have not been shown to be more common than appositions between any other parts of the neuron. Medium spiny cell local collateral input Medium 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 identified 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 necks 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 lateral globus pallidus. Substance P is perhaps a more selective marker for labeling of afferents originating from other striatal spiny neurons, although substance P is also localized in some striatal interneurons. Nonetheless, immunohistochemical localizations 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 those of the dopaminergic input. Izzo and Bolam (1988) 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 described, medium spiny neurons are subdivided into different subpopulations, which give rise to the direct and indirect striatal output pathways. 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 (also substance P-positive), which establishes that, at

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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. Cholinergic interneuron 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 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, and resemble afferents from the cerebral cortex and thalamus. GABA interneuron input In addition to the GABAergic striatal spiny projection neuron, a GABAergic interneuron has been identified within the striatum that makes up 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 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 innervate the somata of spiny neurons (Kita et al., 1990). 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

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physiological criteria (Kawaguchi, 1993). Somatostatinpositive 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. 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 lateral 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 lateral 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 brain stem at least two other forebrain projection systems 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 are not dealt with in this review, which does not reflect their probable important contribution to basal ganglia function. Interactions between Medium Spiny Neurons As described above, regulation of the relative activity in direct and indirect striatal pathway neurons may be affected 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 has 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. The distribution of such collaterals does not appear to cover the same area as the dendrites of

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its parent, and in some cases the distribution is complementary. This would suggest that one neuron’s 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 is 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 colocalized 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 synaptic contact dependent on the receptors on the postsynaptic neuron. In this case it is suggested that medium spiny neurons might affect 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 μm in diameter. 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. 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. Models of striatal function have often invoked inhibition between striatal medium spiny neurons as pro-

viding lateral inhibition. Although physiologic evidence for such interactions has been lacking, recent studies have demonstrated direct inhibition between medium spiny neurons (Tunstall et al., 2002). 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 is 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 affects striatopallidal neurons. The select effect that stimulation of D1 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 D1 and D2 receptors are coactivated result from receptors coexpressed 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 of 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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Striatal Interneurons 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; Kawaguchi et al., 1995). This class of neuron presents a variety of morphologically and neurochemically distinct subtypes (Fig. 6). Two major subtypes are identified on morphologic and neurochemical grounds. One is the large aspiny striatal neuron, which utilizes acetylcholine as a neurotransmitter (Bolam et al., 1984; Kawaguchi, 1992, 1993; Wilson et al., 1990). The other is the medium aspiny GABAergic striatal interneuron, of which there are several varieties (Kita, 1993; Kawaguchi et al., 1995). 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 axons 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 Kölliker and his observations on the human striatum, have described an aspiny neuron with a large cell body and radiating, sparsely branched dendrites. Kölliker reported that this cell had a short axon, but it was described by Ramon y Cajal (Ramón and Cajal, 1911) as the principal projection neuron of the striatum, and this view was held by most authors until the late 1970s and 1980s 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

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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 Golgistained neuron and is justifiably met with a degree of skepticism, but the axons of these two classes of large cells are also proposed to have dramatically different axonal arborizations. While one is the giant interneuron of Kölliker, 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 (Grofová, 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 Ramon and 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).

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A

large aspiny neuron

B ChAT neurons

100 μm

1 mm

C. ChAT-IR neurons

D

E parvalbumin and somatostatin-IR

medium aspiny neuron 100 μm

F parvalbumin-IR neurons

parvalbumin-IR neurons

G somatostatin-IR neurons

1 mm

somatostatin-IR neurons

FIGURE 6 Striatal interneurons. (A) Tracing of dendrites (black) and axon collaterals (gray) of a large aspiny neuron in the striatum (from Wilson et al., 1990). (B) Distribution of cell bodies (black) of striatal neurons immunoreactive for choline acetyltransferase (ChAT). (C) Photomicrograph of ChAT-immunoreactive neurons in the striatum. (D) Tracing of dendrites of a medium aspiny neuron in the striatum. (E) Distribution of cell bodies of parvalbumin (black dots) and somatostatin (white dots) within the striatum. There is an inverse gradient of these two types of striatal interneurons: parvalbumin neurons are more numerous dorsolaterally and somatostatin neurons are more numerous ventrally. (F) Photomicrograph of parvalbumin-immunoreactive neurons in the striatum. (G) Photomicrograph of somatostatin-immunoreactive neurons in the striatum.

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Large Aspiny Cholinergic Neurons Striatal neurons, which utilize acetylcholine as a neurotransmitter, constitute an important type of interneuron population (Bolam et al., 1984; Kawaguchi, 1993; Wilson et al., 1990). 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 acetylcholinesterase (Fibiger, 1982), by immunohistochemical studies employing antibodies directed to the synthetic enzyme choline acetyltransferase (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 μm 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 1 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 that 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 inputs (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, identification 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 contains 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 direct pathway neurons. Dopamine-containing afferents to cholinergic neurons have not been conclusively identified. 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 mediated through activation of substance P receptors. Such a mechanism is supported by anatomical evidence, not only with the demonstration of synaptic contact between substance

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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). Medium Aspiny GABAergic Striatal Interneurons The second major subtype of striatal interneurons are characterized morphologically as medium aspiny neurons and utilize GABA as their main neurotransmitter (Pasik et al., 1988; Smith et al., 1987; Oertel and Mugnaini, 1984; Ribak et al., 1979). Combined Golgi stain and 3H-labeled GABA uptake studies (Bolam et al., 1983) demonstrated that GABAergic interneurons were morphologically characterized as medium-sized aspiny interneurons. Interestingly, isoforms of the GABA synthesizing enzyme glutamic acid decarboxylase (GAD), are differentially expressed by striatal medium spiny projection neurons and striatal medium aspiny interneurons, with GAD67 being expressed at higher levels in striatal interneurons, compared with GAD65, which is expressed in striatal projection neurons. Further classes of GABAergic striatal interneurons are characterized on the patterns of coexpression of neuropeptides, such as somatostatin and neuropeptide Y, and calcium-binding proteins such as parvalbumin and calretinin. Parvalbumin striatal interneurons The most abundant type of GABAergic striatal interneuron expresses the calcium-binding protein parvalbumin (Cowan et al., 1990; Gerfen et al., 1985; Kita et al., 1990; Kubota and Kawaguchi, 1993). Parvalbumin striatal interneurons have very distinct neurophysiologic characteristics, marked by a hyperpolarized resting potential, lower input resistance, shorter duration action potential spikes, and abrupt repetitive firing (Kubota and Kawaguchi, 1993). Due to these physiologic features, this type of neuron is often referred to as a fastspiking interneuron, which is similar to fast-spiking interneurons in the cerebral cortex. These neurons receive inputs from the cerebral cortex and provide inputs to medium spiny projection neurons. As a result of gap junctions between them, and their high level of activity, these neurons may provide synchronized inhibitory feedforward inhibition to restricted regions of the striatum. Although distributed throughout the striatum, parvalbumin neurons are more frequent in the dorsolateral region and display a dorsolateral to ventral gradient. Somatostatin/NPY striatal interneurons A second clear class of GABAergic interneuron coexpresses somatostatin, neuropeptide Y, or nitric oxide synthetase (Dawson et al., 1991; Pasik et al., 1988; Smith and

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Parent, 1986; Vincent et al., 1983a, 1983b; Chesselet and Robbins, 1989). These neurons may be visualized by NADPH diaphorase histochemical staining. This staining appears to be a consequence of nitric-oxide synthase (NOS), independent of NOS activity, to transfer electrons from NADPH to nitroblue tetrazolium. A subset of these neurons (about 20%) express the calciumbinding protein calbindin (28 kDa). The numbers of somatostatin neurons appear to follow a gradient, with more being distributed in ventral areas than in dorsal areas. Moreover, while somatostatin neurons are located in both patch and matrix compartments, their axons within the striatum are preferentially distributed in the matrix compartment. Calretinin/GABA interneurons Another class of striatal GABAergic interneurons coexpress the calciumbinding protein calretinin (Bennett and Bolam, 1993). Calretinin is a calcium binding protein with a strong homology to calbindin. Neurons of this type are distributed mainly in the dorsal medial striatum.

Direct/Indirect Striatal Output Systems Medium spiny striatal projection neurons make up some 90% of the neuron population of the striatum. These neurons have a common morphology in terms of their size, dendritic organization, and local axon collaterals, which extend within the striatum around the parent neuron. Each of these neurons provides an axon that projects out of the striatum. Medium spiny neurons are divided into two subsets, of approximately equal numbers, which contribute to a projection pathway that provides either direct or indirect input to the output neurons of the basal ganglia in the medial globus pallidus and substantia nigra. Connectional Basis Studies in which individual medium spiny striatal neurons were intracellularly filled provide the clearest evidence of subsets of striatal medium spiny projection neurons on the basis of the projection axons (Kawaguchi et al., 1990). One type extends an axon collateral into the lateral globus pallidus, which does not arborize extensively, and extends other collaterals into either the medial globus pallidus and/or the substantia nigra. This type of neuron is referred to as a direct striatal pathway neuron in that it provides direct inputs to the output nuclei of the basal ganglia. A second type provides an axon that extends into the lateral globus pallidus and arborizes extensively, usually in two separate domains within this nucleus. These neurons do not project beyond the lateral globus pallidus and are thus termed “indirect” striatal

projection neurons, in that they connect indirectly to the output of the basal ganglia, through transynaptic connections of the lateral globus pallidus and subthalamic nucleus. It is noteworthy that “direct” projection neurons also contribute inputs to the lateral globus pallidus and thus contribute inputs to the “indirect” pathway system (Kawaguchi et al., 1990). The extent of arborization of this axon collateral is less than that of the indirect projection neuron; however, it exists and appears to 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 or indirect projection neuron input is not yet known. However, in terms of the numbers of synapses, direct projection neuron axon collaterals in the lateral globus pallidus make as many as half the number of synaptic contacts as the axons of indirect striatopallidal neurons. Some direct projection neurons provide inputs to the medial globus pallidus only, some to the substantia nigra only, and some to both. Nonetheless, each of these types are considered direct projection striatal neurons. The medial globus pallidus and the substantia nigra are 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 medial globus pallidus 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 the topographic organization of striatal outputs. Historically, direct and indirect striatal projection neurons were identified with retrograde axonal tracing experiments, in which tracers injected into the target nuclei demonstrated that striatal neurons projected to either the lateral globus pallidus or the substantia nigra (Beckstead and Cruz, 1986; Gerfen and Young, 1988; Loopuijt and Kooy, 1985). Such studies established several features of striatal organization. First, there appeared 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 lateral globus pallidus, have shown most neurons to be labeled from only one injection site (Beckstead and Cruz, 1986; Loopuijt and Kooy, 1985). Such a labeling pattern points to an inherent limitation

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of the technique, as, based on the existence of a collateral of striatonigral neurons in the lateral 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 lateral 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 two to five neurons projecting to one site. In other cases, neurons projecting to separate sites may be nearest neighbors, often appearing to be in close apposition. Neurochemical Basis That subtypes of striatal output neurons provide direct and indirect projection pathways was first suggested by the immunohistochemical labeling of opiate and tachykinin peptides in striatal terminals. Striatal 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; Hokfelt et al., 1977; Pickel et al., 1980) and dynorphin (Vincent et al., 1982a, 1982b) or the tachykinin substance P (Brownstein et al., 1977; Hong et 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 lateral 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 medial globus pallidus) and only a sparse distribution in the lateral globus pallidus (Brownstein et al., 1977; Hong et al., 1977; Kanazawa et al., 1977; Vincent et al., 1982b). Whereas such studies had established 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

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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 reflect 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. Dopamine Receptor Subtypes in Direct and Indirect Striatal Projection Neurons Direct and indirect striatal projection neurons selectively express the D1 and D2 dopamine receptor subtypes, respectively (Gerfen et al., 1990). The mRNA encoding the D1 dopamine receptor subtype was shown to be selectively localized in neurons that project to the substantia nigra and colocalized with substance P and dynorphin, which are selectively expressed by direct striatal projection neurons. Conversely, the mRNA encoding the D2 dopamine receptor is selectively localized in neurons that project to the lateral globus pallidus and is colocalized with enkephalin, which is selectively expressed in indirect striatal projection neurons. This finding has been confirmed by other in situ hybridization studies (Le Moine et al., 1990, 1991). These studies are consistent with prior receptor binding studies, which demonstrated a differential localization of D2 and D1 receptor binding, of striatal origin, in terminals in the lateral globus pallidus and substantia nigra, respectively (Beckstead, 1988; Richfield et al., 1989). Immunohistochemical studies of D1 and D2 receptor protein expression have confirmed the segregation of these receptor subtypes in different populations of

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striatal neurons (Hersch et al., 1995; Levey et al., 1993; Yung et al., 1995, 2000). The selective distribution of receptor expression in the terminal areas confirms the selective localization of D1 receptors in direct pathway neurons and D2 receptors in indirect pathway neurons. Moreover, this selective segregation of D1 and D2 dopamine receptors was demonstrated in rodents as well as primates. The segregation of D1 and D2 dopamine receptor studies was challenged on the basis of single cell mRNA expression studies, which reportedly demonstrated near complete colocalization of both dopamine receptor subtypes in all striatal neurons (Surmeier et al., 1992). While they argued against segregation of D1 and D2 dopamine receptor subtypes (Suremeier et al., 1993), they have now reversed their position (Suremeier et al., 1996), and there is now a consensus concerning this issue. Dopamine Receptor-Mediated Gene Regulation The functional significance of the segregation of D1 and D2 dopamine receptors in direct and indirect striatal projection neurons has been demonstrated by gene regulation studies. Prior to the determination of the localization of D1 and D2 dopamine receptors 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, 1978b; Young et al., 1986). Dopamine depletion of the striatum or neuroleptic blockade of D2 dopamine receptors results in an elevation of enkephalin peptide and mRNA levels in indirect striatal projection neurons (Hong et al., 1978b; Hong et al., 1985; Mocchetti et al., 1985; Tang et al., 1983) and a decrease in substance P levels in direct striatal projection neurons (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 direct striatal projection neurons (Gerfen et al., 1990, 1991; Hanson et al., 1987; Li et al., 1986, 1988). These opposite effects that dopamine has on the peptides in striatal output neurons was demonstrated to be related to the differential expression of the D1 and D2 dopamine receptor subtypes by the neurons that express these peptides (Gerfen 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 the dopamine-depleted striatum, levels of genes contained in indirect striatal projection neurons are increased, including mRNAs encoding enkephalin and the D2 receptor. Conversely, in direct striatal projection 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 receptor 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 (SKF-38393) 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 receptor-selective 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 × 1 mg/day) quinpirole treatment. Conversely, reversal of the lesion-induced decrease in substance P and D1 receptor mRNA was effected with intermittent (1 × 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 activation or blockade of receptor subtypes do 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 effects of D1 and D2 dopamine receptor subtype effects on direct or indirect striatal projection 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 direct striatal projection neurons and not in indirect striatal projection neurons (Robertson et al., 1989, 1990). Thus, the immediate effect of activation of D1 receptor activation appears to have a selective effect on direct striatal projection neurons. While the dopamine-depleted striatum provides a good model for study of the selective effects of D1 and

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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 D1 and D2 receptor subtypes to direct and indirect striatal projection neurons occurs in both the lesioned and the intact 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 receptormediated 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

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systems, most notably the corticostriatal and thalamostriatal systems. Other (Nondopaminergic) Regulatory Receptor Systems in Striatum The organization of D1 and D2 dopaminergic receptors among the direct and indirect striatal projection 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 among 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 pattern similar to that of the dopamine receptor distribution is the α2-adenosine receptor. The mRNA encoding this receptor 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 changes in peptide similar to those that occur 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). There are similar opposite effects of α2-adenosine receptor-mediated immediate early gene regulation (Le Moine et al., 1997; Svenningsson et al., 1999). These α2-adenosine effects appear to involve G protein coupling through the Golf subtype of stimulatory G protein (Herve et al., 2001). Thus, the localization of both the α2-adenosine and D2 dopamine receptor subtypes is expressed in a similar restricted set of striatal output neurons, and activation of these receptors produces selective changes in gene regulation in these neurons. Although the changes produced by α2-adenosine and D2 dopamine receptor stimulation appear to be similar as regards changes in gene

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regulation of peptides in these neurons, the effects of costimulation 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 direct and indirect pathway 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 cannibioid receptors in striatal output neurons in terms of the regulation of receptor gene products (Mailleux and Vanderhaeghen, 1992, 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 hybridization 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 m1 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 (striatoplallidal) and 80% of the D1 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 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 descriptions 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).

INDIRECT PATHWAY Indirect striatal projection neurons project an axon to the lateral globus pallidus, but do not extend axon collaterals to either the medial globus pallidus or the substantia nigra (Fig. 7). Consequently, indirect striatal projection neurons are connected with the output of the basal ganglia indirectly, through transsynaptic connections of the lateral globus pallidus and subthalamic nucleus. GABAergic neurons of the lateral globus pallidus, which are the target of the indirect striatal projection neurons, provide inputs to both the substantia nigra pars reticulata and the medial globus pallidus, as well as to the subthalamic nucleus. The subthalamic nucleus provides a glutamatergic, excitatory input to the medial globus pallidus and substantia nigra. The main components of the indirect pathway, lateral globus pallidus, and subthalamic nucleus are interconnected with each other in such a manner that they do not simply relay information from the striatum (Smith et al., 1998). In combinatorial organotypic culture preparations, in which the connections between the lateral globus pallidus and the subthalamic nucleus are established, an oscillatory pacemaker pattern of neurophysiologic activity is generated in neurons of these structures (Plenz and Kitai, 1999). Such activity occurs

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LGP

LGP

B globus pallidus

A striatopallidal inputs

0.5 mm

neurons

CPu

100μm

LGP STh

1.0 mm

C globus pallidus

MGP

neuron

SNR

CPu 100μm

LGP 1.0 mm

D subthalamic nucleus

MGP

neurons

STh

SNR

FIGURE 7 Components of the indirect striatal output pathway. (A) Tracing of the striatopallidal axon of a single indirect striatal neuron in the sagittal plane. Of note are the double arborization zones within the lateral globus pallidus (LGP) from a single striatal neuron. (B) Tracings of the dendrites of two globus pallidus neurons in the lateral globus pallidus in the sagittal plane. Of note is the distribution of the dendrites, which conform to the same pattern as the striatal afferent axons. Thus, the two regions of the LGP, which are defined by the dual terminal patterns of striatal afferents, appear to have distinct populations of pallidal neurons. (C) Tracing of a single LGP neuron (dendrites in white in sagittal section) and its axon (black) which provides collaterals to the striatum (CPu), subthalamic nucleus (STh), and substantia nigra (SNR). A larger tracing of the dendrites of this neuron is shown on the right. (D) Tracing of a single subthalamic nucleus (STh) neuron (dendrites in white in sagittal section) and its axon (black) which provides collateral inputs to the striatum (CPu), lateral globus pallidus (LGP), and substantia nigra (SNR).

in the absence of striatal or cortical input. It has been suggested that the underlying oscillatory activity in this circuit is important to the normal function of the basal ganglia, its unmasking may be responsible for the tremors associated with Parkinon’s disease and other basal ganglia disorders (Bevan et al., 2002; Plenz and Kitai, 1999).

Globus Pallidus (Lateral Segment) The morphology of the lateral globus pallidus has been well studied at both the light and the 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.,

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1984; Yelnik et al., 1984), immunohistochemistry (DiFiglia et al., 1982), and intracellular labeling (Falls et al., 1982; Kita and Kitai, 1994). These studies are only briefly described in the present paper in the context of issues concerning basal ganglia organization. More thorough treatment of the organization of the lateral 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 lateral globus pallidus (Kita and Kitai, 1994). One type has a moderate to large cell soma from which radiate three to five 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 μm in length, giving a total maximal dendritic coverage of over 1 mm in some cases. Some aspiny neurons display a discoid dendritic field in that the dendrites spread mainly in a two-dimensional field parallel to the border between the lateral globus pallidus and the striatum. Other aspiny neurons have dendrites that cover a volume with a more three-dimensional distribution. While neurons at the border region between the lateral globus pallidus and the striatum often display a discoidal pattern, neurons with discoid dendritic fields are distributed in the central medial regions of the lateral globus pallidus as well. A second type of lateral 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 the spiny neurons did not display 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 discoid type dendrite bearing neurons contain the calcium-binding 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; Hoover et al., 2002). Parvalbumin-positive/discoid dendrite-bearing neurons provide axon collateral projections to the subthalamic nucleus, medial globus pallidus, 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, medial globus pallidus, substantia nigra, and 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 lateral globus pallidus, which are the 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 projection profile somewhat different than that of more dorsal pallidal neurons (Haber et al., 1985, 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 lateral globus pallidus, there are a scattering of cholinergic neurons within the body of the lateral globus pallidus as well as a large number of cholinergic neurons ventral to the lateral 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 the 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, 1986; Saper, 1984). Synaptic Input Neurons in the lateral 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 1 μm in diameter. These afferents have been demonstrated to contain both GAD and enkephalin immunoreactivity, consistent with their origin from the striatum. Some

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estimates place the percentage of such synapses at over 80% of those within the lateral globus pallidus. The manner in which these afferents entwine dendrites forming a mosaic pattern of large synapses gives the lateral globus pallidus the appearance of being composed 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 lateral 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 lateral 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 organization 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). Output Descending output of the lateral globus pallidus to other components of the basal ganglia is directed principally to the subthalamic nucleus and to the medial globus pallidus and the substantia nigra (Haber et al., 1985, 1993; Kita and Kitai, 1994). Ascending outputs of the lateral 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, 1994). 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 are directed to the cell soma and proximal dendrites, whereas the striatal afferent input is directed to the same

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neurons’ more distal dendrites (Smith and Bolam, 1989, 1990, 1991).

Subthalamic Nucleus Based on cellular and dendritic morphology, neurons in the subthalamic nucleus appear to be of one main type, but 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 soma is ovoid or polygonal with a medium size ranging 11–18 μm in diameter. Most subthalamic neurons extend three or four 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 the sagital 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 and in the rostrocaudal 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 that the synapses of subthalamic afferents to neurons in the lateral globus pallidus, the medial globus pallidus, and the 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 Féger, 1990) Synaptic Input Neurons in the subthalamic nucleus receive inputs from the lateral 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 principally to the dendrites of the neurons. Inputs from the lateral globus pallidus make large symmetric contacts which are directed relatively equally to the cell soma (30%), proximal dendrites (39%), or distal dendrites (31%) (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).

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Output

outside the basal ganglia, in particular the thalamus and midbrain structures including the superior colliculus and the pedunculopontine nucleus. The neurons that provide these output projections utilize GABA as a transmitter and form a nuclear complex that is continuous from the medial globus pallidus 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.

Neurons in the subthalamic nucleus project axons that target neurons in the lateral globus pallidus and in the medial globus pallidus and substantia nigra, as well as having 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 Féger, 1990).

Substantia Nigra/Medial Globus Pallidus

BASAL GANGLIA OUTPUTS

The substantia nigra is composed of two main neuron cell types (Fig. 8), those that utilize dopamine (Bjorklund and Lindvall, 1984) and those that utilize GABA as a neurotransmitter (Oertel and Mugnaini,

Together, the substantia nigra and the medial globus pallidus may be considered output nuclei of the basal ganglia in that they provide the interface with brain areas

Summary of inputs to Substantiia nigra pars retiiculata neurons

SNCD SNR SNCV A

SNR sagittal section

input from subthalamic input from nucleus striatum

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input from subthalamic nucleus input from striatum input from LGP

B FIGURE 8 Summary of inputs to substantia nigra pars reticulata neurons. (A) Diagram of a sagittal section of the substantia nigra depicting the double organization of dopamine neurons of the pas compacta (SNC) over GABAergic neurons of the pars reticulata (SNR). Tracings of the dendrites of pars reticulata neurons display how they are oriented along the rostrocaudal dimension and conform with the double regional organization. (B) Schematic of the synaptic locations of inputs to GABAergic neurons of the substantia nigra pars reticulata. Inhibitory inputs from the striatum, and excitatory synaptic inputs from the subthalamic nucleus are both targeted to the distal dendrites. Significantly, individual axons make multiple synaptic contacts with individual dendrites. Inhibitory inputs from the lateral globus pallidus make multiple large synaptic inputs to the neuron’s perikarya and proximal dendritic shafts.

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FIGURE 9 Basal ganglia output pathways arise from GABAergic neurons of the medial globus pallidus (MGP/medial globus pallidus) and substantia nigra pars reticulata (SNR). The MGP output is directed to the ventrolateral (vl) thalamus and to the lateral habenula (lh). The SNR output is directed to the paralamellar mediodorsal (md), intralaminar and parafascicular (pf), and ventromedial (vm) thalamic nuclei; to the intermediate layers of the superior colliculus; and to the pedunculopontine tegmentum nucleus (PPT). Numbers at the bottom refer to the coronal levels shown in Fig. 1 from Paxinos and Watson, 1998.

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 sizes and shapes. Neurons in the substantia nigra have a medium- to large-sized irregularly shaped cell soma with axis dimensions ranging from 16 to 50 μm (long axis) and 8 to 32 μm (short axis). Several (two to four) main dendrites radiate from the cell soma and extend over a generally large domain. Dendrites may divide into secondary or tertiary branches similar in 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 of afferent fibers (Francois et al., 1987). In the rat, dendritic fields may extend as much as 1–1.5 mm in the rostrocaudal axis, 700 μm mediolaterally, and 400 μm in the dorsoventral 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 axes, 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 has been described in the primate as well. As is discussed in

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some detail below, the organization of the dendritic distributions of pars reticulata neurons is related to the organization of afferent inputs from the striatum, lateral globus pallidus, and subthalamic nucleus (Gerfen, 1985; Kita and Kitai, 1987). Dopamine-containing neurons in the substantia nigra in many respects have morphology similar to that 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 appears 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 possesses 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 possess dendrites that extend into the pars reticulata. These two populations are also distinct in their efferent projections and neurochemical content, which is discussed in terms of striatal patch–matrix compartmental organization below.

Synaptic Input to Pars Reticulata Neurons The major sources of input to the substantia nigra neurons are GABA inhibitory inputs from the striatum and lateral globus pallidus and excitatory inputs from the subthalamic nucleus (Fig. 8B). 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 lateral globus pallidus has more recently been found 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 inputs to identified pars reticulata neurons projecting to the superior colliculus was examined at light and electron microscopic levels. 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 lateral globus pallidus for symmetric, relatively large, synapses were directed principally to the perikarya (54% of such input) or to proximal dendrites (32%). The differential distribution of inputs from the striatum and lateral 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.

Projections of Pars Reticulata Neurons Output targets of the substantia nigra include the following: the thalamus, the superior colliculus, and the pedunculopontine nucleus (Fig. 9) (Beckstead, 1979; Deniau and Chevalier, 1992; Gerfen et al., 1982; Kita and Kitai, 1987; Nakanishi et al., 1987b). Nigral inputs to the thalamus are directed to two main parts of the thalamus. The first is the set of nuclei, including the intralaminar nuclei, which project back to the striatum. The second thalamic target is 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 the 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 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 medial globus pallidus are the ventral lateral pars oralis and the 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.

DUAL OUTPUT SYSTEMS OF STRIATAL OUTPUT PATHWAYS A distinctive feature of striatal output organization is the dual projections from the striatum to subdivisions of the lateral globus pallidus and substantia nigra (Fig. 10) (Chang et al., 1981; Gerfen, 1985; Wilson and Phelan, 1982). This organization has also been observed in the primate (Parent and Hazrati, 1994). Striatal projections to the lateral globus pallidus have extensive axon arborizations in a region immediately adjacent to the striatum and a second arborization

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zone in the central part of the lateral 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 associated neurons. These dual projection systems are not to be confused with the patch–matrix projections.

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FIGURE 10 Diagram illustrating the longitudinal, topographic, and dual features of striatal connections. (A) Sagittal diagram of the brain showing the corticostriatal projections and striatal projections to the lateral globus pallidus (LGP) and substantia nigra (SNR). The cerebral cortex projects to the striatum in a general topographic manner when viewed in the coronal plane, but projects to an extensive rostrocaudal domain of the striatum. Projections of individual neurons in the striatum provide axons with extensive distribution of terminals in the rostrocaudal dimension of the lateral globus pallidus (LGP, indirect pathway neurons) or SNR (direct pathway neurons). (B) Diagram of neurons injected in the striatum with two different anterograde tracers (1 and 2) to show projections to the LGP and SNR. In the coronal plane, there is a topographic organization of afferent terminal fields in the lateral globus pallidus LGP that extends from rostro (LGPa) to caudal (LGPc) and in from rostral (SNRa) to caudal (SNRc) in the substantia nigra. In both nuclei there is also a dual representation of these topographically organized projections. In the LGP there is one topographically organized zone just medial to the striatum and a second zone in the more medial part of the nucleus. In the SNR there are also dual zones, one dorsal and one ventral, with the topography of the ventral zone mirroring that of the dorsal zone. Diagram adapted from Gerfen, 1985.

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1.0 mm FIGURE 11 Illustration of the dual projections from (A) the striatum (CPu) to the lateral globus pallidus (LGP) and substantia nigra pars reticulata (SNR) and from (B) the subthalamic nucleus to the lateral globus pallidus (LGP) 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 a 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 Figs. 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 the 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.

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The dual nature of inputs to the lateral globus pallidus and substantia nigra is not only observed in the striatal projections to these nuclei (Fig. 11) . 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 among the nuclei that are the targets of this striatal projection. In both the lateral globus pallidus and the substantia nigra the dendritic morphology of neurons in these nuclei conforms to the dual innervation patterns from the striatum (Gerfen, 1985). Thus in the lateral 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 lateral globus pallidus, whereas neurons in the central region of the lateral 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 dopamine neurons in the pars reticulata. Again, as in the lateral globus pallidus there is one region that forms a shelllike 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. Neurons in these two regions have dendrites that are distributed so as to conform with the shape of the regions (Grofova et al., 1982). This organization was first described by Grofova et al. (1982) based on the morphology of the dendrites of pars reticulata neurons. The organization of the substantia nigra pars reticulata into subregions appears to be related not only to the organization of inputs from the striatum and subthalamic nucleus but also to the organization of its outputs. The organization of the projections of the substantia nigra pars reticulata to the thalamus and to the superior colliculus appears to maintain a rough topography. This topographic organization has been described by Gerfen et al. (1982) and in considerable detail by Deniau and Chevalier (1992). 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 topography involves both the central and peripeduncular shell region of the pars reticulata neurons. Neurons 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 output neurons was described by

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Deniau and Chevalier (1992) 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 (1992), and Redgrave et al. (1992). The functional significance of the dual projection systems of striatal outputs may be related to the organization of the target structure of the nigral outputs (Fig. 11). 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 the peripeduncular “shell” project to more caudal regions. In addition, each of these nigrotectal projections maintains a mediolateral topography. Redgrave et al. (1992) suggested that the organization of the nigrocollicular organization reflects differences in both afferent and efferent organization of the intermediate layer of the superior colliculus that is the target of these inputs. An alternative organization within the superior colliculus that might be the basis for the organization of the nigrocollicular 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 that 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 lateral 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 lateral 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 lateral 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 remain to be worked out. Several possibilities might be investigated. One is that

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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 nonhomogeneous innervation pattern (Holsapple et al., 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, 1991b). It is possible that one part of the medial globus pallidus innervates one of the compartments, whereas the other zone innervates the other compartment. If VLo is organized in a similar manner similar to that of the 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 pallidothalamic output might target different thalamic nuclei, VLo and the centromedian 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 that distinguish this organization from other aspects of striatal output organization. First, individual striatal neurons innervate both zones of the external segment of the lateral globus pallidus. It is also likely that individual striatal neurons also innervate the two zones of the medial 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, which might also be viewed as providing dual projection systems to separate regions of the target nuclei. As is discussed below, the dual projections from the patch–matrix compartments arise from separate neuron populations. Second, the separate zones of the substantia nigra provide topographically organized inputs to the superior colliculus, and a similar organization of nigrothalamic and pallidothalamic 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 nigrotectal 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.

NIGROSTRIATAL DOPAMINE SYSTEM Dopamine neurons in the ventral midbrain, which may be labeled with tyrosine hydroxylase immunoreactivity, are the origin of the nigrostriatal dopamine system (Fig. 12). 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 densely packed; the pars reticulata, which is relatively cell sparse compared to the pars compacta; and the retrorubral area, which lies caudal and dorsal to the substantia nigra (Bjorklund and Lindvall, 1984; Gerfen et al., 1987). The designation of the subgroupings of dopamine neurons 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 (Bjorklund and Lindvall, 1984). 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. Dopamine innervation of the striatum (Bjorklund 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 patchand matrix-directed systems (Gerfen et al., 1987a, 1987b; 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 that arise from distinct sets of dopamine neurons in the substantia nigra (Gerfen et al., 1987a). 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 distribution 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 the matrix

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FIGURE 12 Dopaminergic neurons in the midbrain. (A–C) Tyrosine hydroxylase (TH) immunoreactivity shows distribution of dopamine neurons at rostral (A), middle (B), and caudal (C) levels of the ventral midbrain in coronal section (medial is to the left). Adjacent sections showing calbindin immunoreactivity (A′,B′,C′) delineate subgroups of dopaminergic neurons. The ventral tegmental area (VTA) and dorsal tier of the substantia nigra pars compacta (SNCD) contain dopamine neurons, which coexpress calbindin. Ventral tier substantia nigra pars compacta neurons (SNCV), contain dopamine neurons, which are calbindin-negative. At rostral levels (A,A′) the SNCD and SNCV are both located dorsal to the substantia nigra pars reticulata (SNR), whereas at caudal levels there are two parts of the SNCV, one just subjacent to the SNCD and the other within the SNR. At caudal levels part of the dopamine cell group extends dorsally into the retrorubral nucleus (RRF).

striatal compartments (Gerfen et al., 1987a, 1987b). 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 a ventral tier.

Dorsal Tier Dopamine Neurons 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 tegmental 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

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Synaptic Input to Pars Compacta Neurons

FIGURE 13 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 dark-field 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 calbindin-poor 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 dorsolateral striatum, which nonetheless contains opiate receptor patches.

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.

Ventral Tier Dopamine Neurons 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 compacta 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 reticulata projecting to the dorsal striatal patch compartment. It is worthwhile to note that the number 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.

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 (Smith and Bolam, 1990). However, in the case of input from the lateral 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 inhibitory input (Christoph et al., 1986).

STRIATAL PATCH/MATRIX COMPARTMENTS Within the complexity of the striatum it is important to identify those aspects of organization that provide underlying mechanisms that 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 lateral globus pallidus and to the medial globus pallidus/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 is 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 direct and indirect output neurons and function to regulate these output neurons. One such system is the so-called “patch– matrix” striatal compartments, which are involved in

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FIGURE 14 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) are depicted to show the innervation of the patch and matrix compartments from different subsets of midbrain dopamine neurons from three levels (B,C,D). Neurons providing inputs to the striatal matrix compartment (white in B,C,D) are located in the ventral tegmental area (VTA, A10 DA cell group), in the dorsal tier of the substantia nigra pars compacta (in B: SNc, A9) and in the retrorubral area (in D: 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 B,C,D: SNc, A9 DA cell group) and from A9 DA cells located in the substantia nigra pars reticulata (in C and D). 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.

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1 mm FIGURE 15 Organization of the striatal patch–matrix compartments provides parallel pathways from the cerebral cortex through the striatum that provide differential input to the dopamine and GABA neurons in the substantia nigra. Deep layer 5 corticostriatal neurons provide selective inputs to the striatal patch compartment, whose neurons provide inputs targeting dopamine neurons in the substantia nigra pars compacta. Superficial layer 5 corticostriatal neurons provide inputs to the striatal matrix compartment, whose neurons project to the substantia nigra pars reticulata, which contains the GABAergic output neurons of the basal ganglia. This organization arises from most neocortical areas, although there is a gradient such that those areas closer to the allocortex provide a greater input to the patch compartment, whereas primary sensorimotor areas provide a greater input to the matrix compartment.

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 com-

partments may be defined precisely on the basis of connections of the neurons in these compartments (Gerfen, 1984, 1985, 1989; Gerfen et al., 1987a). 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 striatum. Thus, although we 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 is 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 regulationdependent 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). 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 it 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 in most regions of the striatum and have been useful in establishing the connectional basis of patch–matrix organization.

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 medial globus pallidus (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 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

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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, 1987). 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 both in the area in which dopamine neurons are located in 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 neuron 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 medial globus pallidus (Rajakumar et al., 1993). The medial globus pallidus 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, medial globus pallidus neurons provide a projection to the thalamus (van der Kooy and Carter, 1981). The thalamic targets of the medial globus pallidus 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 medial globus pallidus and the substantia nigr pars reticulata appear to form a continuous somatotopically organized output of the basal ganglia. The medial globus pallidus is also distinct from the substantia nigra in lacking an associated dopamine cell group. However, like the substantia nigra the medial globus pallidus may be divided into two parts on the basis of output neurons. In addition to those medial globus pallidus neurons that project to the ventral lateral thalamus, there is a medially situated part of the nucleus that 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.

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Recent studies have shown that the striatal matrix compartment provides inputs directed to the thalamic projecting part of the medial globus pallidus, 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 medial globus pallidus suggests an organization similar to, though different from, the organization 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 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 lateral globus pallidus. Whereas the projection of patch neurons to the substantia nigra and medial globus pallidus has been described, these neurons account for only half of the projection neurons in the patch compartment. The other half provides inputs to the lateral globus pallidus (Gerfen and Young, 1988). One possibility might have been that patch and matrix neurons provide differential inputs to the striatopallidal border region versus the central region of the lateral 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 lateral globus pallidus. These neurons, which are scattered in the dorsal lateral globus pallidus and are 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

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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 the matrix compartments. This organization of the dendrites of medium spiny neurons suggests that afferents from outside the striatum target a particular compartment.

Cortical Input Cortical inputs to the patch and matrix compartments originate in different sublayers of layer 5, from most cortical areas (Gerfen, 1989). Cortical inputs were among 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 compartment 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 into a specified cortical area, such as the prelimbic area (Gerfen, 1989). 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 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 the pattern of thalamic labeling of 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 cingluate and medial agranular cortices, the areas adjacent to the prelimibic cortex. The third criterion was that the pattern of the crossed corticocortical 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 prelimibic 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 that 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 layers 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

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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 corticocortical and corticostriatal labeling is of interest in the context of the relationship between corticocortical and corticostriatal organization. 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 an input to the matrix relatively greater than that 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 are 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 periallocortical 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.

Thalamic Afferents Thalamic afferents in the striatum from the parafascicular/intralaminar nuclei are organized relative to

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the patch/matrix compartments (Beckstead, 1985; Berendse et al., 1988; Gerfen et al., 1984; Herkenham and Pert, 1981). 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.

General Patch–Matrix Organization The general organization of the patch–matrix compartments provides separate pathways from the cortex through the striatum to differentially affect dopamine and other basal ganglia feedback circuits or to affect basal ganglia GABAergic output neurons in the medial globus pallidus 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 the 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 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.

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

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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 among 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 three types of corticostriatal neurons; pyramidal tract neurons with collaterals into the striatum, corticothalamic neurons with collaterals into the striatum, and bilaterally projecting corticostriatal 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 corticocortical projections of the injected neuron population. Injections, which selectively labeled inputs to the patch compartment, labeled both local and contralateral corticocortical connections that were preferentially distributed in deeper cortical layers. Conversely, injections, which selectively labeled inputs to the matrix compartment, labeled both local and contralateral corticocortical connections that were preferentially distributed to superficial cortical layers. With this method it is not possible to attribute the corticocortical pattern of labeling to neurons that project also to 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 corticocortical connectivity. For example, at least two distinct types of layer 5 neurons have been distinguished on the basis of their local axon collaterals: one type has axon collaterals that distribute longitudinally in layers 5 and 6, and another neuron type has axon collaterals that distribute to superficial layers 2 and 3 (ChagnacAmitai et al., 1990). These latter axon collaterals display a much more restricted distribution in the longitudinal axes. If such differences in local cortical axon 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 (1966, 1978) that the basal ganglia was a site of integration of limbic and nonlimbic 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 olfactoryrelated cortical areas, provided the main input to the dopamine neurons in the substantia nigra pars compacta, which projected back to both ventral, limbic striatal and also dorsal, nonlimbic 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 the 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 to only limbic-connected cortical areas. Thus, given the fact that the patches in the dorsal striatum receive inputs from neocortical, nonlimbic areas of cortex and that these patches nonetheless provide

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inputs to the dopamine nigrostriatal feedback system, further modification of the concept of the integration of so-called limbic and nonlimbic integration within the basal ganglia is required. Rather than consider that the striatal patch–matrix systems are related to limbic and nonlimbic 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 alloand periallocortical and neocortical areas because of the obvious conceptual difficulties of defining limbic and nonlimbic cortices. Allo-, periallo-, and neocortical 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 corticocortical 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 corticocortical 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 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 allocortical 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 striatal targets of allocortical areas, which may be the shell region of the nucleus accumbens, and the striatal target of the 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 vary 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 is carried through the striatum to provide segregated inputs to dopamine neurons and GABA output neurons in the substantia nigra.

SUMMARY A canonical organizational scheme for the part of the basal ganglia that receives input from the neocortex is

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described. Neocortical areas provide excitatory, glutamatergic inputs from layer 5 neurons to the striatum. The main target of this cortical input are medium spiny projection neurons in the striatum, which includes the caudate–putamen and the core of the nucleus accumbens. Striatal medium spiny projection neurons give rise to two separate output pathways, one provides a direct pathway and the other an indirect pathway to the output nuclei of the basal ganglia. The indirect pathway involves connections with the lateral globus pallidus and subthalamic nucleus. The output of the basal ganglia are GABAergic neurons in the medial globus pallidus and substantia nigra pars reticulata, which project to the thalamus, superior colliculus, and pedunculopontine nucleus. Dopamine neurons in the substantia nigra pars compacta provide input to the striatum, which provides differential regulation of the striatal direct and indirect pathways, due to the segregation of the D1 and D2 dopamine receptor subtypes on the neurons giving rise to these projections.

such that an individual axon makes contact with only 1% of the striatal medium spiny neurons in the area across which it is distributed. In addition, there is a 6 to 1 ratio of corticostriatal neurons to striatal projection neuron targets. Together, these quantitative estimates indicate that the cortical input to a single striatal medium spiny projection neuron is rather unique; that is, no two striatal neurons share common inputs from the cortex. This organization of corticostriatal inputs appears to represent the end stage of a competitive network rather than a dynamic one in which different patterns of corticostriatal inputs are competing for synaptic sites. Additionally, Wilson (Zheng and Wilson, 2002) has argued that rather than providing a compact representation of patterns of cortical activity, which would remove redundancy in patterns of corticostriatal input activity, a large proportion of cortical patterns of activity must either be lost or treated as though they are similar when they are not.

Direct and Indirect Striatal Projection Systems Corticostriatal Projection System Corticostriatal inputs arise mostly from layer 5 neurons in the neocortex. Subtypes of inputs have been identified on the basis of whether the axon is a collateral of a cortical axon projecting to the brain stem or a collateral of axons that are distributed only to the cortex and the striatum. Other subtypes are identified by the distribution of axons within the striatum. One subtype provides a focal axon within the striatum while another provides a distributed axon within the striatum. The distribution of corticostriatal inputs provides a primary topographically organized input to the striatum. This topographically organized system provides for parallel functional circuits through the basal ganglia. In addition, corticostriatal inputs provide a divergent input such that there is a convergence within the striatum of inputs from dispersed cortical areas. Somatotopically organized cortical areas provide inputs to multiple regions within the striatum, with convergence of the inputs of similar body parts in these multiple striatal regions. The patterns of corticostriatal inputs are related to the organization of the corticocortical connections of the parent corticostriatal neurons. Such divergent/convergent patterns of corticostriatal inputs provide for overlap within functional circuits and between functional circuits. Quantitative analysis of the cortical inputs to the striatum provides informative constraints in considerations of the potential functional information processing that occurs in this pathway. Most distinctively corticostriatal axons provide an extremely sparse input to individual medium spiny projection neurons

Over 90% of neurons of the striatum are medium spiny projection neurons, which are the major target of cortical and thalamic inputs. Medium spiny neurons are divided into two roughly equal subsets which give rise to the direct and indirect striatal projection pathways. In addition to differences in their connections, these neurons are neurochemically distinct, expressing different neuropeptides and G protein receptors. Clinical hypokinetic and hyperkinetic movement disorders have been attributed to imbalances between the relative activity in these two output pathways (Albin et al., 1989; DeLong, 1990). The components of the indirect pathway have received recent interest in their involvement in such movement disorders. The finding that connections between the component parts of the indirect pathway, the lateral globus pallidus and the subthalamic nucleus, support an oscillatory pattern of neural activity may be related to the tremors often unmasked in movement disorders. Both types of neurons receive input from the cerebral cortex and respond to activity in the corticostriatal input with excitatory postsynaptic potentials (Kawaguchi et al., 1990). The different expressions of G protein-coupled receptors and, in particular, the D1 and D2 dopamine receptor subtypes in the direct and indirect pathway neurons, respectively, are responsible for differences in the signal transduction pathways regulating gene expression. A recent study has shown that protein kinase signal transduction systems are differentially regulated in direct and indirect striatal pathway neurons (Gerfen et al., 2002). ERK1/2/MAPkinase appears to be selectively activated in indirect striatal

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FIGURE 16 Diagram illustrating the maintenance of a general topographic organization 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 (corticostriatal) maintain the spatial relationships within the cortex in the targeted regions in the striatum. Similarly the projections of the striatum to the lateral globus pallidus and substantia nigra/MGP 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 lateral globus pallidus and the substantia nigra, 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 MGP 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).

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projection neurons, whereas protein kinase A is utilized in the direct striatal projection neurons. This differential regulation of these two protein kinase signal transduction systems appears to be a consequence of the expression of the D1 and D2 dopamine receptors in the direct and indirect striatal projection neurons, respectively. Changes in gene regulation resulting from activation of protein kinase signal transduction systems are responsible for synaptic plasticity that alters synaptic efficacy. The differential regulation of protein kinase-mediated changes in gene regulation that occur in the direct and indirect striatal neurons suggests that different forms of synaptic plasticity may occur in these striatal neurons. Dopamine appears to be involved in such differences. Dopamine depletion of the striatum, in animal models of Parkinson’s disease, results in a dramatic switch in the regulation of ERK1/2/ MAPkinase in direct striatal projection neurons (Gerfen et al., 2002). This switch in regulation of ERK1/2/ MAPkinase is responsible for the supersensitive response of direct striatal projection neurons to D1 dopamine agonist treatments.

Patch/Matrix Striatal Compartments The striatum contains two distinct compartments, termed the patch and matrix compartments, which may be identified with a number of neurochemical markers. Although the neurochemical markers are subject to regulation that alters their compartmental-specific expression, the patch matrix compartments are morphologically based and distinct on the basis of their ipnut and output connections. Patch and matrix neurons appear to be physically separated, such that the dendrites of patch and matrix neurons remain within their respective compartments. On the input side, corticostriatal inputs are differentially distributed to the two compartments, with neurons in deeper parts of layer 5 providing inputs directed to the patch compartment, while neurons in superficial layer 5 provide inputs to the matrix. Most areas of neocortex provide inputs to both compartments, but the relative contribution to each displays a gradient such that cortical areas adjacent to allocortex have a greater input to the patch compartment. Different subsets of dopamine neurons in the midbrain innervate the two compartments. Dorsal tier, calbindin-containing dopamine neurons provide inputs to the matrix compartment, while ventral tier, calbindin-negative neurons provide input to the patches. On the output side, patch neurons provide inputs to the dopamine neurons of the substantia nigra pars compacta, whereas matrix neurons provide inputs to the GABAergic neurons of the substantia nigra pars reticulata. Thus, the patch

and matrix compartments provide segregated pathways from different populations of corticostriatal neurons through the striatum, which differentially target the output neurons of the basal ganglia and the nigrostriatal dopamine feedback system.

Dual Representation of Striatal Outputs in the Output of the Basal Ganglia A distinct organizational feature of the basal ganglia is the dual projections of individual striatal neurons to the lateral globus pallidus and the substantia nigra. This organization provides a double representation of the striatum in each of these structures. In the lateral globus pallidus one zone is immediately adjacent to the striatum and the second zone is located medially. In the substantia nigra, one zone is located dorsally and the other ventrally. Each striatal target zone is topographically organized in a manner similar to that of the corticostriatal inputs. In addition, the subthalamic nucleus connections with these nuclei also provide a double representation. While striatal target nuclei provide double mappings of the striatal output, the basal ganglia output nuclei appear to provide a single output mapping. This is seen in the projection of the substantia nigra to the superior colliculus. The two striatal mapping zones in the substantia nigra project to different parts of the superior colliculus, with the dorsal zone projecting to the rostral, short saccade zone of the superior colliculus while the ventral nigral zone projects to the caudal, long saccade zone of the superior colliculus. This raises the possibility that the dual representation of striatal outputs is related to gross and fine movements involved in orienting behavior. Determining the relationship of the dual representation of the striatum in the substantia nigra and medial globus pallidus to the organization of the connections with the thalamus will aid in determining the function of this organizational feature of the basal ganglia.

Conclusion The basal ganglia provide one of the output systems of the neocortex, which connects to neural systems involved in generating behavior. The main input structure of the basal ganglia is the striatum and the main outputs are GABAergic neurons in the medial globus pallidus and substantia nigra pars reticulata. These outputs are directed to thalamic nuclei providing feedback to the frontal cortical areas involved in the planning and execution of movements, to the superior colliculus and pedunculopontine nucleus. There are two main pathways, one direct and the other indirect, which connect the striatum with the output nuclei of

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the basal ganglia. While the neurons giving rise to these pathways are morpohologically similar and receive similar inputs from the cerebral cortex, they are connectionally and neurochemically distinct. In particular D1 and D2 dopamine receptors are segregated respectively to the direct and indirect striatal pathway neurons. The basal ganglia are organized in a general topographic manner such that the relationship of cortical areas is maintained in the connections through the striatum and output nuclei, providing parallel, functionally defined loops. However, there is considerable divergence and convergence of connections at each level of the system, which appear to reflect organizational features similar to those connecting cortical areas through intracortical connections. In addition, the striatum provides a double representation to each of its target zones. While the function of the basal ganglia is debated, its dysfunction results in profound movement disorders that are characterized by either too little movement (Parkinson’s disease) or uncontrolled (dyskinesia) movements.

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19 Amygdala and Extended Amygdala of the Rat: A Cytoarchitectonical, Fibroarchitectonical, and Chemoarchitectonical Survey JOSE S. DE OLMOS1, CARLOS A. BELTRAMINO1,2, and GEORGE ALHEID3 1

Instituto De Investigacion Medica "Mercedes Y Martin Ferreyra" Cordoba, Argentina 2

Dept. De Neurofisiologia Y Psicofisiologia, Facultad De Psicologia Universidad Nacional De Cordoba, Argentina 3

Department of Physiology and Institute for Neuroscience Northwestern University Feinberg School Of Medicine, Chicago, Illinois, USA

Fuchs and Siegel, 1984; Fuchs et al., 1985; Gloor et al., 1982, 1983; Mishkin and Aggleton, 1981; Sarter and Markowitsch, 1985; Schoel et al., 1981; Squire and ZolaMorgan, 1985; LeDoux, 1990; Gloor, 1997; Maren, 1999; Hurd, 2000). Such a modulation is exercised, at least in part, through a vast network of connections with other brain regions, such as the hypothalamus, the brain stem, and spinal cord autonomic cell aggregates. These other regions have been thought to have the most direct involvement in the regulation of these functions. Recent experimental anatomical, physiological, and behavioral data suggest, however, that the amygdala is more directly involved in such actions than was previously suspected. These actions are subserved by some of the nuclear groups into which the amygdala has been divided on the basis of cytoarchitectonic, histochemical, immunocytochemical, and hodological studies (de Olmos et al., 1985; Alheid et al., 1995; Heimer et al., 1999). Finally it seems important to note that in humans the amygdaloid complex has been shown to be significantly affected in a number of human neurodegenerative diseases, including Alzheimer’s disease, the chorea of Huntington, and neuronal ceroid lipofuscinosis (Brockhaus, 1938; Cipollini

The rat amygdaloid body is a relatively large conglomerate of gray substance lying in the depth of the anteromedial temporal lobe ventral to the lentiform nucleus from which it is separated by the magnocellular basal forebrain neuronal complex (MBFNC) constituting the basal nucleus of Meynert and related cell groups, or the so-called substantia innominata sensu ampliori. Massive fiber systems separate the amygdala from the basal forebrain and form part of the telencephalic and diencephalotegmental radiations that connect bidirectionally the amygdala and neighboring gray formations in the temporal lobe with those in the basal and medial telencephalon, the diencephalon, the brain stem tegmentum, and even the spinal cord. Although the amygdala has been generally thought of as a single structure, it is actually a very heterogeneous gray complex which has been shown to be involved in the modulation of neuroendocrine functions, visceral effector mechanisms, and complex patterns of integrated behavior, such as defense, ingestion, aggression, reproduction, memory, and learning (Beltramino and Taleisnik, 1978, 1980; Benjamin and Jackson, 1974; Carrer, 1978; Carrer et al., 1973, 1980; De Vito and Smith, 1982; Dunn, 1987; Dunn and Whitener, 1986; Fahrbach et al., 1983;

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et al., 1988; Herzog and Kemper, 1980; Braak and Braak, 1983, 1987; Brashear et al., 1988; Brady and Mufson, 1989; Powers et al., 1987; Benzing et al., 1990, 1992; Emre et al., 1993; Hudson et al., 1993; Pitkänen and Amaral, 1998; Hurd, 2002; Perez et al., 2000).

GENERAL TOPOGRAPHY AND TERMINOLOGY This review of the main anatomical and cytochemical features of the rat amygdala departs from previous cytoarchitectonical studies by Johnston (1923), Gurdjian (1928), Brodal (1947), Yu (1969), Price (1973), and Krettek and Price (1978b) as well as from reviews by de Olmos et al. (1985) Price et al. (1987), and Alheid et al. (1995). In analyzing the nuclear arrangement of the rat amygdala and in critically reviewing the literature on the hodological relationships of this griseum with the cerebral cortex, the basal ganglia, the diencephalic nuclei, and the tegmental grisea in the brain stem and the spinal cord, the nomenclature employed here is based substantially on the seminal comparative neuroanatomical work by Johnston (1923), who examined the basal forebrain, but more particularly the amygdala of the opossum, bat, rat, rabbit, macaque monkey, and human embryos. With few modifications, the terminology proposed in this newer version of the chapter about the rat amygdala coincides almost entirely with that used in previous versions of this chapter (de Olmos et al., 1985; Alheid et al., 1995) and is consistent with the terminology used in the last edition of Paxinos and Watson’s (1997) stereotaxic atlas of the brain of this macrosmatic mammal. In our view, this is a well-developed nomenclature supported not only by well-established cytoand fibroarchitectonical criteria but also by ontogenetic, histo- and immunocytochemical, and hodological data (Bayer, 1980; de Olmos et al., 1985, 1999; Grove, 1988a, 1988b; Barbas and de Olmos, 1991; Alheid et al., 1990; Heimer et al., 1999). By following the simple topographical criteria once proposed by Brockhaus (1938), the gray nuclei composing the amygdaloid complex can be grouped into two major nuclear groups, superficial and deep (Fig. 1). Within the superficial group and since some of its members show a stratified arrangement (in both molecular (subpial) and cell layers) very much resembling that seen in neighboring truly cortical fields, they have been named so to address such a condition either by considering them to represent a single structure, the cortical amygdaloid nucleus, or, by adopting topographical and cytoarchitectonical criteria and considering them to represent several different nuclei or areas, i.e.,

the anterior, (ACO), posterolateral (PLCO), and posteromedial (PMCO) cortical amygdaloid nuclei. In addition, the existence of the rather shallow amygdaloid fissure has also contributed to the distinction of two corticallike transition areas, the amygdalohippocampal (AHI) and amygdalopiriform transition areas. Rostrally, a wellstratified but quite confined cortical-like cell aggregate, the nucleus of the lateral olfactory tract (LOT), together with a transversely oriented column of more or less tightly packed cells, the bed nucleus of the accessory olfactory tract (BAOT), help to delimit the rostral pole of the superficial amygdala, a delimitation that is complemented by the presence at that level of a more or less undifferentiated griseum, the anterior amygdaloid area (AA). This area constitutes the most rostrally located representative of the amygdaloid body superficial grisea (Figs. 1–3). Other authors (Krettek and Price, 1978b; Haberly and Price, 1978; Price, 1981; Price et al., 1987; Kemppainen and Pitkänen, 2000, etc.), somewhat in line with early neuroanatomists (Rose, 1927; Popoff and Popoff, 1929), utilized the term “periamygdaloid cortex” to name the superficial cortical-like amygdaloid structures including under that term both the PLCO and the APIR. However, against the full cortical nature implicitly conveyed by this terminology is the fact (shown in the thymidine autoradiographic studies of Bayer (1980) in the rat) that during its ontogenetic development the cortical nucleus or nuclei show a pattern of ontogenetic features totally coincident with those of a subcortical or nuclear gray formation. Furthermore, as will be seen later in this chapter, there are cytoarchitectonical, histo- and immunocytochemical, and hodological data that are compelling enough to consider APIR as an anatomical entity by itself, completely separated from the PLCO nucleus, which also reaffirms the nuclear (rather than cortical) nature of the so-called “periamygdaloid cortex.” In the deep aspects of the amygdaloid body several cell aggregates can be identified among which a laterobasal nuclear complex, consisting of three major divisions, the lateral (LA), basolateral (BL), and basomedial (BM) nuclei, can be clearly identified. Another less clearly differentiated griseum, the ventral basolateral (BLV) nucleus, named as the BL nucleus by Brodal (1947), is here assigned to the laterobasal nuclear complex on account of its clear cytoarchitectonical, histo- and immunocytochemical, and hodological distinction from the more scattered-celled ventral endopiriform nucleus (VEn). Apart from these major cell aggregates, the deep sector of the amygdaloid body contains other less conspicuous groups of cells that on account of their peculiar morphological characteristics cannot be assigned to any of the main amygdaloid cell groups and it will be necessary to obtain more information than is presently

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FIGURE 1 Schematic diagram of the amygdala and the extended amygdala. (A) The block diagram approximates a horizontal view of the nuclei composing the amygdala and extended amygdala. The most superficial amygdaloid nuclei have been rotated toward the midline as shown in B in order to artificially place all the amygdaloid nuclei in a single plane. It should be noted that the extension of the dorsal pallidum (globus pallidus) into the ventral pallidum (DP–VP) divides the cell columns of the extended amygdala. Not shown are the dorsal cell pockets of the extended amygdala that accompany the axons of the stria terminalis over the internal capsule (BSTS, supracapsular bed nucleus of the stria terminalis) and the interconnecting column of the large intercalated cell mass that runs beneath the central amygdaloid nucleus. For abbreviations, see the list at the end of this chapter.

FIGURE 2 Amygdala subdivisions. (A) Extended amygdala. The central and medial divisions of the extended amygdala are shown with bold outlines. The central division is identified by the central amygdaloid nucleus at the caudolateral end of the outline and the lateral bed nucleus of the stria terminalis at the rostrolateral end. Similarly, the medial subdivision is identified by the medial amygdaloid nucleus at the caudomedial end and the medial bed nucleus of the stria terminalis at the rostromedial end. (B) The cortical-like nuclei. The cortical-like nuclei of the amygdala are shown with bold outlines. It should be noted that the intramedullary gray, and intercalated cell masses have not been included within this designation (see text). For abbreviations, see the list at the end of this chapter.

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FIGURE 3 Olfactory and vomeronasal amygdala. (A) The shaded areas represent the amygdaloid nuclei with reciprocal relations with the main olfactory bulb. (B) The shaded areas represent the amygdaloid nuclei with reciprocal relations with the accessory olfactory bulb. It should be noted that the narrow column interconnecting the medial amygdala with the medial bed nucleus represents scattered cells within the medial sublenticular extended amygdala that project to the accessory olfactory bulb, as well as scattered cells in the medial part of the supracapsular bed nucleus of the stria terminalis that are potential recipients of accessory bulb efferents. For abbreviations, see the list at the end of this chapter.

available in order to establish its affiliation in a correct manner. These cells aggregates are the intercalated masses (IM), the amygdalostriatal zone (Astr), the granular and parvicellular interfascicular islands (INFI), and the intramedullary griseum (IMG) (Fig. 1). Finally, two major amygdaloid grisea, one superficially located, the medial amygdaloid nucleus (Me), and the other, situated in the deep dorsal sector of the amygdala, the central amygdaloid nucleus (Ce), present a number of morphological characteristics that set them apart from any of the amygdaloid cell groups discussed above on account of their cytological, cytochemical, and functional continuity into basal telencephalic structures that today are known to represent their extraamygdaloid counterparts, i.e., the central (SLEAc) and medial (SLEAm) sublenticular extended amygdala and the paraseptal lateral (BSTL) and medial (BSTM) bed nuclei of the stria terminalis, respectively (Fig. 2A). With these remarks in mind, we recognize in the rat amygdala four major supranuclear divisions: (1) a superficial cortical-like nuclear group (CONG), (2) an extended amygdala (EXA), (3) a laterobasal nuclear complex (LBNC), and (4) an unclassified cell group (UNCG) (Figs. 1A, 2A, and 2B). By taking into consideration the connectional relationships between the olfactory bulb and the amygdala of the rat (Scalia and Winans, 1975; de Olmos et al., 1978, 1985; Switzer et al., 1985; Shipley et al., 1995, 1996) and by applying the morphological criteria mentioned

above with respect to the parcellation of the superficial nuclei of the amygdala, the olfactory amygdala in the rats is considered here to comprise the following structures: (1) the anterior amygdaloid (AA), (2) the nucleus of the lateral olfactory tract (LOT), (3) the anterior (ACO) (Fig. 3A) and (4) posterolateral (PLCO) cortical nuclei, and (5) the amygdalopiriform transition (APIR) area; while the vomeronasal cortical-like amygdala includes the nucleus of the accessory olfactory tract (BAOT), the posteromedial (PMCO) cortical nucleus, and the amygdalohippocampal area (AHI) (Fig. 3B). The fact that only a very small subfield of the medial amygdaloid nucleus (Me) in the rat receives a direct afferent supply from the main olfactory bulb (de Olmos et al., 1985; Switzer et al., 1985), while the bulk of this nucleus is innervated by the terminal arborizations of the accessory olfactory tract (de Olmos et al., 1978, 1985; Scalia and Winans, 1975; Shipley et al., 1995, 1996), places this nucleus in a sort of ambiguous anatomical and functional position concerning its relationships with the MOB system. For most purposes, however, like it was done in the previous versions of this chapter (de Olmos et al., 1985; Alheid et al., 1995), the Me nucleus is included as part of the so-called “vomeronasal amygdala” following the suggestion of Kevetter and Winans (1981). Furthermore, as has been mentioned previously, recent morphological, cytochemical, and hodological evidence indicates that the Me nucleus is only one of the components of the major supranuclear structure termed

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the medial extended amygdala (MEXA) (Figs. 3B and 4B; see also Alheid et al., 1995; Heimer et al., 1999). As already mentioned and as is described in a more detailed fashion subsequently, this supranuclear entity comprises also the medial sublenticular extended amygdala (SLEAm) and the medial (BSTM) division of the bed nucleus of the stria terminalis and its supracapsular (BSTSm) and intraamygdaloid (BSTIA) divisions. Within the MEXA, however, only the rostroventral and midcaudal sectors of the Me nucleus (caudal MAD, MPV, MPD) and the medial small-celled subnucleus (BSTMPm) of the posterior part of the medial BST nucleus receive direct inputs from the AOB, a condition which would leave those sectors of the MEXA not directly innervated by the accessory olfactory tract outside of what could be a strictly defined vomeronasal amygdala. However, due to the massive intranuclear bidirectional connections that are shared by every other member of the MEXA, such a restrictive interpretation may not be applicable. The central extended amygdala group (CEXA), on the other hand, consists of the central amygdaloid nucleus (Ce), the central sublenticular extended amygdala (SLEAc), the lateral division of the bed nucleus of the stria terminalis (BSTL), its lateral supracapsular division (BSTSl), and the interstitial nucleus of the posterior limb of the anterior commissure (IPAC) (Alheid et al., 1995; Heimer et al., 1999). On account of the profuse afferent supply that the CEXA receives from secondary olfactory areas such as the PIR cortex, the APIR transition area, and the lateral Ent area (see below), and by

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applying hodological criteria similar to those in the case of the Me nucleus, this major division of the EXA could be in turn affiliated to the main olfactory system (Figs. 1A, 2A, and 4A). As mentioned before, the laterobasal nuclear gray complex (LBNC) is constituted by the lateral (La), basolateral (BL), basomedial (BM), and ventral basolateral (BLV) amygdaloid nuclei. Each of the three first named members of the LBNC can in turn be divided into subnuclei possessing cytoarchitectonical, histo- and immunocytochemical, and hodological characteristics that differentiate them, but it can be said, although with some restrictions, that they are closely related to the striatopallidal complex and those cortical fields subserving different sensory modalities and motor functions (Figs. 1A and 2B). Finally, those amygdaloid nuclei that have been enlisted with the fourth group, including the amygdalostriatal transition zone (Astr), the granular and parvicellular interface islands (INFI), the intercalated masses (ICM), and the intramedullary griseum (IMG), cannot be readily enlisted with any of the three previous groups of nuclei since they do not seem to share many of the morphological, cytochemical, or hodological features that characterize each of these major cell groups. Instead the Astr and the INFI cell groups may represent transition areas to pallidostriatal formations (Heimer et al., 1999), while the latter two mentioned cell groups, may do it with respect to the extended amygdala (de Olmos, 1990; Martin et al., 1991) (Fig. 1A).

FIGURE 4 (ex Fig. 7) Sagittal semi-schematic views of (A) central division and (B) medial division of the extended amygdala. ac, anterior commissure; Acb, accumbens. For other abbreviations, see the list at the end of this chapter.

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DESCRIPTION: OBSERVATION PROCEDURES This section is concerned with some of the main architectonic and histochemical features that characterize each one of the main nuclei and their respective subnuclei, as well as those main areas and corresponding subareas that compose the amygdaloid body. This description is based on the study of frontal, sagittal, and horizontal sections of normal rat brains that were alternatively stained for Nissl substance (thionine, cresyl violet), myelinated fibers (Weigert), neurofibrillary frameworks (Bielschowsky-Gross modifications) cuproargentic technique (CuAg, 1969, 1972) acetylcholinesterase (AChE) activity (Geneser-Jensen and Blackstad’s (1971) modification of the Koelle method), and heavy metals by the Danscher’s modification of the Timm method (Danscher, 1981) or immunostained for neuropeptides such as cholecystokinin (CCK), leuenkephalin (ENK), somatostatin (SOM), substance P (SP), calbindin (CB), calretinin (CR), parvalbumin (PAV), dopamine-β-Hydroxylase (DBH), Tyrosine hydroxylase (TH), etc. To describe cytoarchitectonically all the anatomical components of the present chapter, it is mandatory to include a series of Nissl figures. Because of a strict space limitation, we will refer in this aspect to Fig. 15 of the previous edition of this chapter (Alheid et al., 1995). Therefore, in every section entitled as topographical landmarks and cytoarchitectonics, we refer the reader to the mentioned figure as Alheid et al., 1995, Fig. 15, adding the corresponding figure letter of the series.

extended amygdala. Since the data provided by the Golgi technique has also been described in the previous chapters it is not included here.

Olfactory Amygdala: Superficial Cortical-like Nuclei The Anterior Amygdaloid Area (AA) This area, which constitutes the rostral undifferentiated pole of the amygdala, is classified here as a member of the “olfactory amygdala” (Kevetter and Winans, 1981) not only because it receives and reciprocates afferent projections from the MOB but also because of its profuse interconnections with other members of the “olfactory amygdala” as well as with cortical and subcortical secondary olfactory centers such the PIR cortex and the anterior olfactory nuclei (de Olmos et al., 1985; Switzer et al., 1985; Shipley et al., 1995, 1996). But it should not be ignored that this part of the amygdala, more particularly its ventral medial subdivision (AAVm), contains a neuronal pool that not only projects to the AOB but shares with those AOB projecting cells in the SLEAm and BSTMPl similar morphological characteristics, a feature that segregates them from other neurons constituting the AA area (Fig. 5). Furthermore, evidence has been presented that these cells and others of different morphological characteristics located in the AA area appear to be involved, like those in the MEXA, in the regulation of salt appetite (Johnson et al., 1999). It is for these reasons that this part of the amygdala

ORGANIZATION OF THE RAT AMYGDALOID COMPLEX AND EXTENDED AMYGDALA A significant portion of the morphological, histochemical, immunocytochemical, and hodological data analyzed in the following sections of the present chapter has been obtained from an exhaustive survey of the almost overwhelming amount of information produced by different laboratories of the world. For that reason, and due to the limitations of space made available to the present writers, the reader is advised to consult the previous versions of this chapter (de Olmos et al., 1985; Alheid et al., 1995) in order to obtain a more complete listing of bibliographic references as well as a more detailed comparison of the present nomenclature with that proposed by other authors. It is also advisable to consult those previous versions with regard to the numerous data supporting the existence of a symmetry among the components of the two divisions of the

FIGURE 5 (ex Fig. 6) Axis of symmetry. In a broad sense, the extended amygdala may be thought of as symmetrically disposed about the axis indicated in the schedule (see text). For abbreviations, see the list at the end of this chapter.

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could well be affiliated either anatomically with the “vomeronasal amygdala” or functionally, at least, with the MEXA. Topographic landmarks The anterior amygdaloid area represents a considerable territory in the rat amygdala (Figs. 15C–15F in Alheid et al., 1995). Few studies have been directed to understand its complex anatomical associations. The AA area appears as a transition region inserted between the amygdala and the rostral basal forebrain structures. Moreover, in contrast with other amygdaloid recipients of direct olfactory inputs from the mitral cells of the MOB, it lacks a defined stratified arrangement, although it bears a molecular layer of varying thickness that separates it from the pial surface of the brain. Ventrally, this irregularly shaped amygdaloid griseum starts abruptly just caudally to the olfactory tubercle lying intercalated between this latter and the LOT nucleus, a cell aggregate which AA surrounds laterally, medially, and dorsally (Figs. 12, 16, 17, 19, and 21). Through a somewhat more extended medial wing, the ventral or superficial sector of the AA area lies also interposed between the LOT nucleus, on one side, and the large-celled preoptic nucleus and the interstitial nucleus of the posterior limb of the anterior commissure IPAC (Figs. 19 and 21), on the other, a topographical relationship that is maintained until the BAOT and Me nuclei appear beside the caudal end of the LOT nucleus. Through a shorter and narrower lateral wing, the ventral sector of the AA area also intervenes between the LOT nucleus and the deep layers of the Pir cortex, but this time it does it until the ACO nucleus makes its appearance just a little rostrally to the caudal end of the LOT nucleus. Medially and mediodorsally, both ventral and dorsal sectors of the AA area become continuous with the medial component of the SLEA, while rostrodorsally only the dorsal sectors of the AA area merge with the ventral pallidum (Figs. 29A and 29B), medially, and the medial division of the IPAC nucleus, laterally (Figs. 21A–21C). Finally, dorsal sectors of the AA area cupping the LOT nucleus become replaced by the BMA nucleus and the most rostral and ventral aspects of the medial division of the Ce nucleus caudally (Fig. 17). Cytoarchitectonics On the basis of cytology and histochemical data, the AA area can be divided into a ventral or superficial medium-celled part (AAV) and a mixed-celled dorsal or deep part (AAD). The first mentioned division, in turn, can be subdivided, on the basis of both its topographical relationships to the LOT nucleus and cytoarchitectonic criteria, into two major subareas, medial (AAVm) and lateral (AAVl), and a very narrow shell-like one (AASH; Figs. 12A, 16A, and 17A).

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In the AAVm subdivision, which composes the previously mentioned ventral medial wing of the AA area, the cell population is more homogeneous than that in the AAVl subdivision constituting the ventral lateral wing, and it shows a predominant presence of slender and vertically oriented perikarya as compared with the more heterogeneous and haphazardly oriented neuronal perikarya present in the second named subdivision. In the “shell” subdivision, on the other hand, the long axis of the neuronal perikarya making up this perinuclear formation is predominantly oriented parallel to the surface of the LOT nucleus, with the cells in the innermost row being tightly apposed to it. An additional characteristic of the ventral divisions of the AA area is the presence of cells of different sizes and shapes within the molecular layer (L1) as far superficially as the boundary line between sublamina 1α and sublamina 1A. Such an arrangement is shared only with the ACO and BAOT nuclei from which they can be delimited by their different cell compositions, i.e., more homogenous in ACO with respect to the heterogeneously populated AAVl subdivision, and the far more compact cell arrangement of the BAOT nucleus as compared with that of the AAVm subdivision (Figs. 12, 16, and 17). The dorsal division of the AA area, on the other hand, can be characterized by the presence of large isolated neurons showing morphological features typical of cells commonly found in the ventral pallidum or in the interstitial nucleus of the horizontal limb of the diagonal band (HDB) or those displayed by the basal forebrain cholinergic magnocellular neurons. Most of them are ectopic in nature and, therefore, should not be confused with true AA area neurons, since even the largest of them are smaller in size and lighter in staining (Nissl) compared with those belonging to any of the previously mentioned gray formations. Finally, with respect to the delimitation of the boundary between the AAD division and the IPAC nucleus this is facilitated by the fact the AA area cells are larger than and show a packing density lower than that of those in the IPAC nucleus. Fibroarchitectonics In neurofibrillar preparations, the AA area is characterized by the presence of a very dense, homogeneously arranged fiber network along it whole extent. The fibers invade the molecular layer (L1) of the AA area and show no layered arrangement except for sublamina 1α which corresponds to fibers of the lateral olfactory tract (lot), and for a capsule-like arrangement which forms around the LOT nucleus coincidentally with the topographic arrangement of the AA cell shell (Fig. 12B). Heavy metals In Timm/Danscher’s preparations, the molecular layer (L1) of the AA area shows a common

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FIGURE 6A (ex Fig. 4) Dark-field image of normal cupric silver stain of the central division of the extended amygdala in a horizontal section. When this method, which also stains degenerating fibers, is applied to unlesioned brains, the argyrophilic normal fibers are the last to be suppressed by the preincubation step or removed by subsequent bleaching. Note the lack of staining in the anterodorsal part of the medial central amygdaloid nucleus. Acb, accumbens; aca, anterior limb of anterior commissure; GP, globus pallidus; f, fornix; opt, optic tract. For other abbreviations, see the list at the end of this chapter.

FIGURE 6B (ex Fig. 5) Horizontal section with degenerating fibers in the central division of the extended amygdala after a large electrolytic lesion of the basolateral and lateral nucleus. Modified from de Olmos (1972) to include current terminology. Acb, accumbens; f, fornix; GP, globus pallidus; opt, optic tract. For other abbreviations, see the list at the end of this chapter.

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clear-cut negative reaction at the level of the two ventral subdivisions (Figs. 16B and 17C), with the only difference made by its greater thickness at the level of the AAVl subdivision. On the rostrodorsomedial pole of the AAD a more pronounced staining reaction as well as a tendency for the granular silver sulfide precipitate to form thin aggregates around unstained slender stripes can be observed. Such arrangements provide the sector with a blurred striated appearance (Figs. 21B and 29B), which contrasts with the light to moderately diffuse granular staining pattern in the other sectors. This disposition resembles that seen in the VP and it is feasible that these zinc-containing aggregates correspond to the ectopic cell profiles described above regarding the AAD division cell composition. Apart from this, a semiquantitative densitometric analysis of the distribution of the Timm/Danscher’s reaction (TD-R) in this part of the amygdala reveals the presence of similar densities of silver precipitate in both the major AAD division and in the AAVl subdivision, which, by comparison, are denser than those in the AAVm subdivision. This lesser density of the TD-R in the AAVm subdivision contrasts also with the more compact silver precipitate present in the BAOT and MeAD nuclei. On the contrary, a lighter TDR in both the ACO and BMA nuclei as compared with that in the AAVl support the parcellation suggested by our Nissl and neurofibrillar preparations. Acetylcholinesterase (AChE) In AChE preparations (Figs. 29A and 30A), the AA area shows a weak to moderate patchy staining reaction. Such a staining pattern contrasts sharply with the very strong reactivity of the horizontal limb of the diagonal band dorsomedially, the olfactory tubercle rostroventrally, the IPAC nucleus rostrodorsally, and the ventromedial end of the PIR laterally. Caudally, the slightly higher and patchy staining reactivity of the AA area allows it to be distinguished from the ACO nucleus and the Me nucleus; at the same time the AA area is nearly completely delineated by the very strong staining reaction in the adjacent LOT nucleus. In contrast, caudodorsally and dorsomedially, its delineation from the CeM division of the Ce nucleus and SLEA becomes hardly traceable. No lamination can be observed in the molecular layer, which presents the same staining pattern as the rest of the AA area. At variance with the complete absence of AChE-positive somata in the superficial sectors of the AA area, some large AChE-positive cells can be detected in its deeper portions, probably representing ectopic diagonal band or related cholinergic projection neurons. It is feasible that the AA area neurons projecting to places like the AHI or other amygdaloid nuclei (Canteras et al., 1992) could be ectopic cholinergic elements

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detached from the MBFGC as suggested above and, therefore, not true AA neurons participating in intraamygdaloid interconnections. The fact that the neurons in the ventral sector of the AA (equivalent to the most rostral sectors of their BMAa and COAa nuclei (their Figs. 4C and 4D) do not seem to be a source of projections to the AHI seems to provide support for this speculation. Choline acetyltransferase (ChAT) The AA area displays a medium density of ChAT-immunoreactive (ChAT-IR) fibers interrupted by small sectors of lesser density (Heckers and Mesulam, 1994). The Nucleus of the Lateral Olfactory Tract (LOT) Topographic landmarks The nucleus of the lateral olfactory tract is a conspicuous ovoid collection of neurons creating an eminence on the surface of the rostromedial pole of the amygdala, just at the caudomedial end of the main body of the lateral olfactory tract (Figs. 8, 12, 16, 17, 19, 21, 25, 29, 30, and 34A). It is surrounded by a superficial extension of the AA area, which separates the LOT nucleus from the olfactory tubercle (Tu) and the BAOT and Me nucleus caudomedially. Cytoarchitectonics In Nissl sections (Alheid et al., 1995, Figs. 15C–15F), three layers are evident in the LOT: layer 1, a molecular layer (L1); layer 2, a superficial dense cell layer (L2); and layer 3, a deep multiform cell layer (L3). L1 contains a few scattered small- to medium-sized cells. L2 consists of a circumscribed oval cell aggregate composed of tightly packed, deeply staining, mediumsized pyramidal neurons. L3 is formed by slightly larger, more loosely arranged, multiangular cells, which cover the top of L2 like a cap. Based on the differential lamination of superficial olfactory bulb projection fibers and terminals within the L1, this layer can be further subdivided into (i) a sublamina 1α, containing lateral olfactory tract fibers; (ii) a sublamina 1A, containing their terminals, and (iii) a sublamina 1B, which is slightly wider than 1A (Figs. 12A, 16A, and 19). Fibroarchitectonics In neurofibrillar preparations layer 1 of the LOT is very distinctly stained and it can be subdivided into: (i) a sublamina 1α containing lateral olfactory tract fibers very sharply delimited in the caudolateral two-thirds, but rapidly decreasing in density rostromedially; (ii) a sublamina 1A, not clearly separated from sublamina 1B because of its relative fiber-rich content (unlike the other olfactory bulb (OB) recipient superficial amygdaloid nuclei); and (iii) a sublamina 1B, wider than 1A, found at the edges of the LOT, which becomes continuous with the fiber capsule surrounding it (Fig. 1A). Sublamina 1B is distinguished from 1A by

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FIGURE 7 (A, B, C, D) Normal cupric silver-stained sections cut transversally to display other views of the central division of the extended amygdala when depicted by the mentioned procedure. Other abbreviations not related to the amygdala or the extended amygdala are the same as those used for the Paxinos and Watson 1997 version of The Rat Brain in Stereotaxic Coordinates. For abbreviations, see list at the end of this chapter.

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FIGURE 8 Sagittal view of a section stained immunohistochemically for angiotensin II (direct print). The angiotensin II is relatively rich in the amygdala and extended amygdala, while avoiding pallidal areas. In this figure, it can be seen that the extended amygdala territory surrounds the immediate subcommissural ventral pallidum. ac, anterior commissure; HDB, horizontal limb of diagonal band; VP, ventral pallidum. For other abbreviations, see list at the end of this chapter.

its lighter background staining in neurofibrillar preparations and by its richer fiber plexus. Layer 2 is characteristically fiber poor; the few fibers present are mainly oriented perpendicularly to the pial surface. Layer 3 by contrast shows a very dense fiber network which stains even more heavily than the fiber network in the surrounding AA (Figs. 12B and 25). Heavy metals In Timm/Danscher’s preparations (Figs. 16B, 21B, 29B, and 30B) the LOT nucleus has a more complex and unique staining pattern. Sublamina 1α shows at its peripheral, subpial edge a thin rim of silver precipitate, which decreases sharply in intensity in deeper parts of the sublamina, until it becomes quite light, but never as light as that in the similar sublaminae of the other superficial amygdaloid cell groups. Sublamina 1A is hardly differentiated from 1α except for interrupted small and very narrow nonreactive segments. Sublamina 1B shows, in contrast to 1A, a much darker staining reaction. This is not as heavy as that in the capsule-like rim formed by the heavy precipitate that surrounds the nonreactive cell bodies of the entire

periphery of L2. The core of L2, on the other hand, is even less reactive than sublamina 1B. The capping layer 3 has a similar distribution of Timm/Danscher’s silver precipitate but is lighter and has fuzzy contours compared with the distribution of layer 2. Acetylcholinesterase In acetylcholinesterase preparations (Fig. 29A) the LOT nucleus presents a differential staining in its three layers and appears to be sharply delimited from surrounding structures. Sublaminae 1α and 1A show a homogeneous and weak, but consistent, staining reaction, which diminishes quite sharply at the borders of the nucleus. Sublamina 1B shows a stronger staining reaction, which abuts the very reactive neuropil that surrounds the otherwise AChE-negative cell bodies in L2. Choline acetyltransferase According to Hellendall et al. (1986), together with the BLA nucleus, L2 of the LOT nucleus contains the highest ChAT activity in the amygdaloid complex. The deeper, less densely packed cell layer (L3) has a substantially lower activity, but still

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FIGURE 9 Horizontal view of a section stained immunohistochemically for angiotensin II. As seen in Fig. 8, the angiotensin II is relatively rich in the amygdala and extended amygdala, while avoiding pallidal areas. Note the lateral wing of the central division of the extended amygdala formed by the interstitial nucleus of the anterior commissure (IPAC), which courses rostral to the globus pallidus (EGP). aca, anterior limb of anterior commissure; AcbSh, shell of nucleus accumbens; acp, posterior limb of anterior commissure; cst, commissural component of stria terminalis. For other abbreviations, see list at the end of this chapter.

is much higher than in most other regions of the brain. The ChAT-IR fibers in L2 of the LOT nucleus stand out as a relatively dark structure but do not not seem to reach the levels observable in the BLA nucleus. The deep L3, on the other hand shows ChAT-IR fibers comparable to those present in the surrounding structures.

The Anterior Cortical Amygdaloid Nucleus (ACO) Topographic landmarks The anterior cortical amygdaloid nucleus occupies the rostral third of the cortical amygdaloid area (Figs. 15D–DI in Alheid et al., 1995) and begins anteriorly as a caudal replacement of the ventral

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FIGURE 10 Horizontal view of choline acetyltransferase-positive cells just dorsal to the centromedial amygdala (direct print). Note the intermingling of these cells within the globus pallidus and the sublenticular extended amygdala. The nucleus of the ansa lenticularis (NAL) is a cluster of cholinergic cells wedged between the cerebral peduncle and the optic tract. aca, anterior limb of the anterior commissure; acp, posterior limb of the anterior commissure; CPu, caudate–putamen. Photograph courtesy of Drs. William E. Cullinan and Laszlo Zaborszky. For abbreviations, see list at the end of this chapter.

AA area, wedged between the medial border of the PIR cortex laterally and the LOT nucleus medially. Just caudal to the LOT nucleus, the ACO nucleus expands sharply to become interposed between the piriform cortex and the PLCO nucleus laterally and the Me nucleus medially. Farther caudally, the ACO nucleus borders the PLCO nucleus laterally and the PMCO nucleus medially (Figs. 8, 14B, 15B, 16, 17, 21C, 22, 29, 30, and 33A). Cytoarchitectonics A reduction of the cell density in the depth of the nucleus marks its dorsal boundary, which separates it from the more densely packed BMA

nucleus (Figs. 15D–15I in Alheid et al., 1995). The small, slender, and loosely packed cells of the ACO nucleus show no obvious lamination except for a slightly higher superficial crowding, which constitutes a relatively wide superficial cell layer. Dorsal to this poorly defined cell stratum appears a deep cell layer made up of more loosely arranged cells. Ventral to these two cell strata is a rather wide molecular layer in which sublaminae 1A and 1B can be easily identified. A very distinctive morphologic characteristic of the molecular layer of the ACO nucleus is the occurrence of a local widening of the subpial glial layer. Neuronal perikarya of differ-

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FIGURE 11 (ex Fig. 13) Retrograde labeling in basal forebrain after Fast Blue injection in central amygdaloid nucleus (CeA); composite tracing from two adjacent sections. Note the large number of cells in the extended amygdala areas, including the interstitial nucleus of the posterior limb of the anterior commissure (IPAC), the central division of the sublenticular extended amygdala (SLEAc), the bed nucleus of the stria terminalis (BST), and the caudomedial accumbens (Acb). aca, anterior limb of the anterior commissure; acp, posterior limb of the anterior commissure. Modification of Fig. 5A in Heimer et al., 1993, with permission. For abbreviations, see list at the end of this chapter.

ent sizes and shapes invade sublamina 1B and even the deepest aspects of sublamina 1A (Fig. 15H of Alheid et al., 1995). This is a feature almost entirely absent in the molecular layer of the PMCO nucleus. Within the molecular layer, a very narrow sublamina 1α occupied by the LOT is also evident, as is a wider sublamina 1A and a narrow sublamina 1B (Figs. 16A and 22A). Fibroarchitectonics In neurofibrillar preparations (Fig. 13E) the molecular layer (L1) of the ACO offers again some distinctive features: a very narrow subla-

mina 1α occupied by the LOT; a much wider sublamina 1A traversed its whole width by darkly stained fibers which run parallel to the pial surface; and a relatively narrow sublamina 1B containing fewer, finer, and more lightly stained fibers, with the majority oriented perpendicular to the pial surface. A rich fiber plexus encloses the neuronal somata of the superficial cell layer (L2) while a richer plexus of more densely stained fibers surrounds the cells located deeper in the ACO. The latter plexus is directly continuous with the deep layer of the PIR cortex and helps to delimit the ACO from

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FIGURE 12 Nissl (A) and Neurofibrillar (B) stained coronal sections through the anterior amygdaloid area (AA) and nucleus of the lateral olfactory tract (LOT). For abbreviations, see list at the end of this chapter.

the more dorsal anterior part of the BM nucleus. However this layered disposition of the fiber plexus in the cell portions of the ACO is poorly differentiated when compared with that seen in other subdivisions of the superficial cortical-like amygdaloid region. Heavy metals In Timm/Danscher’s preparations (Figs. 16B, 21C, 29B, and 30B), the ACo nucleus shows predominantly a homogeneous, weak to moderate neuropil staining. This contrasts markedly with the dense staining reaction in most of the surrounding structures, except for the more palely stained BMA nucleus dorsally. The outer zones (sublaminae 1α and 1A) of the ACO nucleus molecular layer are completely unreactive, whereas the innermost zone (sublamina 1B) exhibits a homogeneous staining, slightly denser than that of the deeper portions of the nucleus, but distinctly weaker than its homologs in the Me nucleus and the PMCO nucleus. Acetylcholinesterase In acetylcholinesterase preparations (Figs. 29A and 30A) the ACO nucleus displays a weak staining reaction that contrasts with the dense staining of the LOT nucleus and the anteromedial corner of the temporal primary olfactory cortex (the cortical amygdala transition area indicated by Paxinos and Watson (1986)). The ACO nucleus shows more AChE activitiy than the medial amygdala, but its molecular layer lacks the characteristic staining pattern found in layer 1 of the ME nucleus and of the PMCO nucleus. The staining reaction in the ACO nucleus does not differ substantially from that of the PIR cortex and PLCO

nucleus laterally and farther caudally or the BMA nucleus dorsally. Choline acetyltransferase The ACO, like the other cortex-like amygdaloid nuclei, i.e., PLCO and PMCO, are not remarkable for their ChAT activity, which is, however, higher than that in the neocortex, but less than that at the adjacent medial edge of the PIR cortex. Nevertheless, its ChAT-immunoreactivity appears to be practically nonexistent as compared with that in surrounding structures (Hellendall et al., 1986). However, to judge from the illustrations by Hecker and Mesulam (1994), it seems that the amount of ChAT-IR fibers in the ACO nucleus, though clearly less than in the LOT nucleus or at the medial edge of the PIR cortex, is still somewhat higher than that in the Ce, BSTIA, or Me (dorsal sectors) nuclei. The Posterolateral Cortical Amygdaloid Nucleus (PLCO) Topographic landmarks This superficially located amygdaloid nucleus, termed by some authors as the periamygdaloid cortex (PAC) (=PLCo, in Figs. 15H–15M in Alheid et al., 1995), occupies an intermediate external position with regard to the other components of the so called “olfactory amygdala”; i.e., it is found intercalated among the ACO nucleus, rostrally and rostromedially, the PMCO nucleus, medially and posteromedially, the APIR transition area, caudolaterally, and the PIR cortex, laterally. A shallow sulcus, termed the amygdaloid fissure, separates the PLCO nucleus from the last two named structures. Dorsally, it blends in a rostrocaudal

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succession with the BLV nucleus, the BM nucleus, the BLP nucleus, and the AHI AL subarea (Figs. 13E, 16, 17, 22, 23, 26, 27C, 29, 30, and 33A). Cytoarchitectonics In Nissl preparations, the PLCO nucleus appears as a complex collection of cells clustered into five cell subgroups: oral medial (OM), oral lateral (OL), caudal medial (CM), small-celled (PC), and caudal lateral (CL) (Figs. 15H–15M in Alheid et al., 1995). The most conspicuous differentiating features are those shown by the oral OM, CM, and PC subgroups. The PLCO OL subnucleus sits at the rostral border between the PLCO nucleus and the PIR cortex and is a relatively small but distinct cell aggregate. Its superficial cell layer is made up of chromophilic, tightly packed pyramidal neurons arranged in a band-like fashion. Underlying this is a rarefied deep cell layer, a feature that allows the delineation of the PLCO OL nucleus from the adjacent PIR cortex and from the other cortical subgroups. The caudal medial PLCO subnucleus (PLCO CM) appears as a distinct wedge between the ACO nucleus and the PMCO nucleus and is the most medially located of all the PLCO nucleus subgroups It shows a well-defined cell stratification and, unlike the PLCO OL subnucleus, has a broad and compact deep cell layer that contains cells more darkly staining and larger than those in the neighboring cell groups, except for the large neurons of the AL AHI area. Within the PLCO CM subnucleus and almost adjacent to the PLCO CL subnucleus appears a very clearly circumscribed ovoid neuronal aggregate, the PLCO PC subnucleus. This is formed by small, tightly packed neurons that invade the depth of the PLCO nucleus in an oblique rostromedial direction. The layering in this tiny gray formation, as in the adjacent caudal lateral PLCO subnucleus (PLCO CL) and oral medial PLCO subnucleus (PLCO OM), is less defined than that in the other two PLCO nucleus cell subgroups. Further, both the caudal lateral and the oral medial subdivisions of the PLCO nucleus show very few features distinguishing them from the surrounding nuclear and cortical formations, except for a less ordered arrangement of their cell components (Figs. 16A, 17A, and 22A). Fibroarchitectonics In neurofibrillar preparations (Fig. 13E), the PLCO nucleus offers a picture that differs from other subdivisions of the cortical amygdaloid nucleus, and in the PIR cortex the superficial sublamina 1α is reduced to a narrow layer of olfactory fibers more particularly in the caudal and medial sectors of the nucleus. L1A, being more poorly fibered, contrast markedly with its richly fibered equivalent in the PMCO and Me nuclei. Although L1B is relatively rich in

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tangentially running fibers it shows fewer fibers than its equivalent in the PIR cortex. The L2, which corresponds to the superficial cell layer, contains a quite prominent fiber plexus, denser in the superficial aspect of this layer. This superficial plexus deepens around the periphery of the PLCO CM and small-celled subnuclei of the PLCO nucleus forming a basquet-like fiber formation. The plexus is loosely arranged in the CM, but very tightly packed in the small-celled subgroups. In both cases, these fiber formations are continuous with the fiber systems which join the stria terminalis. In the PLCO OL subnucleus, the tangential fiber plexus in L1B is clearly continuous with its equivalent in the PIR cortex and fuses with the peripheral aspect of the superficial cell layer. In contrast with the predominantly transverse orientation of the very rich fiber plexus in the deep cell layer of the OM and CL subdivisions of the PLCO nucleus, the deep fiber plexus in the OL PLCO subnucleus is dominated by vertically oriented fibers forming small bundles of fibers in many places. The fiber bundles eventually become continuous with the deep fiber plexus of the temporal PIR cortex. Thus, fibroarchitectonically, the PLCO OL subnucleus constitutes a transitional gray formation between the PLCO proper and the medial aspect of the temporal PIR cortex. Heavy metals In Timm/Danscher’s stained sections (Figs. 16B, 17C, 22B, 23B, 26A, 29B, and 30B), the staining pattern of the PLCO nucleus is more complex. Common to all the PLCO nucleus subdivisions, a clear trilamination of the molecular layer appears as a continuation of the layers observed in the primary olfactory cortex. It consists of an outer nonreactive zone (1α and 1A), a middle weakly staining zone (the superficial part of 1B), and a deep strongly reactive zone (in the deep part of 1B), which outline the practically nonreactive perikarya in the superficial cell zone. This is particularly noticeable in the oral lateral subgroup. In the deep cell layer, the neuropil shows a bilaminar staining pattern. A very deeply staining “hilar” zone outlines the basal surface of the unstained perikarya in the superficial cell layer, and a deeper zone shows a homogeneous slightly weaker staining reaction, which is continuous with the adjacent olfactory cortex. This reaction stops quite sharply at the border with the ACO nucleus and the BMA nucleus. At the level of the oral lateral PLCO nucleus, a lighter and narrower zone separates the PLCO nucleus from the darkly staining neuropil in the BLV nucleus. More caudally, the staining reaction in the deeper zone becomes homogeneously continuous with that in the PLBL nucleus, BMP nucleus, AL AHI area, and PMCO nucleus. In contrast with the rest of the PLCO subgroups, the small-celled subdivision shows a homogeneous and less strong staining reaction.

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FIGURE 13 (A–E) Neurofibrillary stained frontal sections through the paraseptal, sublenticular, and amygdaloid components of the extended amygdala in the rat. Approximate Atlas distances from interaural line: (A) IA = 8.74 mm; (B) IA = 8.60 mm; (C) IA = 8.20 mm; (D) IA = 8.20 mm; and (E) IA = 6.70 mm. For abbreviations, see list at the end of this chapter.

Acetylcholinesterase In AChE preparations (Figs. 17B, 26B, 26C, 22C, 29A, and 30A), the PLCO nucleus shows quite a uniform staining reaction comparable to that in the ACO nucleus and the adjacent primary olfactory cortex. Caudally and starting approximately at the level of the cephalic pole of the AHI transition area, the PLCO nucleus becomes slightly darker in the caudal lateral and the caudal medial subgroups, but is almost completely negative for AChE in the small-celled subgroup, outlining sharply its boundary with the surrounding structures. Immediately lateral to the smallcelled subgroup, the most lateral part of the caudal medial PLCO nucleus shows a slightly stronger staining reaction, forming a bottle-like shape, with the neck connecting to the darker staining in the AHI transition area. The adjacent caudal lateral PLCO nucleus shows a staining reaction sufficient to be differentiated dorsally from the BMP nucleus, which is poor in AChE activity. However, common to all the PLCO nucleus subdivisions, the molecular layer shows a poor, practically unstratified, AChE reactivity (Figs. 17B, 26B, 26C, 22C, 29A, and 30A).

Choline acetyltransferase According to Hellendahl et al. (1986). The PLCO nucleus, like other members of the superficial cortical-like amygdala, is not remarkable for its ChAT activity, which is higher than in the neocortex but lesser than in the PIR cortex. The ChATimmunoreactivity in this nucleus according to Heckers and Mesulam (1994) is however of medium density. Amygdalopiriform Transition Area (APIR) Topographic landmarks The APIR transition area is the gray formation situated in the fundus and lateral lip of the shallow amygdaloid or semiannular fissure (Figs. 15L–15N´ in Alheid et al., 1995) which is densely connected with the olfactory system (Scalia and Winans, 1975; Krettek and Price, 1977; Kosel et al., 1981; de Olmos et al., 1978; Switzer et al., 1985). On the basis of cyto- and chemoarchitectonic data as well as its connectional relationships, but also by applying topographical criteria, this transitional periallocortical field can be divided into anterolateral (AL APIR) and posteromedial (PM APIR) divisions, a parcellation somewhat eclectic suggested by the work of previous authors

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FIGURE 14 (ex Fig. 16) (A, B) Coronal sections through the posterior medial bed nucleus of the stria terminalis. (A) Timm’s–stained section, sodium selenite perfusion. Note the continuity of the medial (BSTMPm), intermediate (BSTMPi), and lateral (BSTMPl) columns of the posterior medial bed nucleus. The small-celled medial column of the posterior medial bed nucleus is distinguished by a somewhat lighter reaction. It is encapsulated by the denser staining of the intermediate column. Also evident is the continuity of the BSTMPl with the sublenticular extended amygdala (SLEA). (B) Angiotensin II-immunostained section. In this section the continuous columns of the posterior medial bed nucleus are immunoreactive for angiotensin II; BSTMPl is less densely stained than the adjacent intermediate column. Also apparent is the immunoreactivity in the interstitial nucleus of the posterior limb of the anterior commissure (IPAC). AH, anterior hypothalamus. For other abbreviations, see list at the end of this chapter.

FIGURE 15 (ex Fig. 17) Coronal sections comparing angiotensin II immunoreactivity (A, C, and E) with immunodetection of retrograde trans-synaptic pseudorabies transport from the stomach (B, D, and F). Note the labeling of the interstitial nucleus of the posterior limb of the anterior commissure (IPAC) by both the angiotensin II antibodies and by antibodies to the pseudorabies virus. Also evident is the rather selective viral labeling of the central division of the extended amygdala, rather than the medial division. Modified from Fig. 2 in Alheid et al. (1994), with permission. For abbreviations, see list at the end of this chapter.

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FIGURE 16 Horizontal sections through the amygdala stained for either (A) Nissl or (B) Timm/Danscher’s histochemistry (sodium thiosulfide perfusion). For abbreviations, see list at the end of this chapter.

(de Olmos et al., 1985; Alheid et al., 1995; Jolkonnen et al., 2001). The anterolateral APIR subarea is narrow and is interposed between the temporal (posterior) piriform cortex, laterally, and the most lateral representative of the superficial amygdala (that is, the PLCO nucleus), medially (Figs. 17 and 23). The expanded posteromedial APIR subarea (Figs. 16, 17, and 23), on the other hand, lies intercalated between the ventromedial (Krettek and Price, 1977c (VMEA); Haug, 1976 (28M´)) and ventrolateral (Krettek and Price, 1977c (VLEA); Haug, 1976 (28L´)) subdivisions of the entorhinal periallocortex, caudally, and the caudolateral pole of the superficial amygdala (here, the PMCO nucleus) and small-celled BLP, rostrally. Cytoarchitectonics The APIR transition area (Figs. 15L–15N´ in Alheid et al., 1995) differentiates

itself from the other structures that surround it on the basis of the greater amplitude of its molecular layer (L1) and the lack of a defined segregation of its superficial and deep cells. Rostrally, in the anterolateral APIR subarea, both the superficial (L2) and the deep (L3) layers are typically displaced toward the depth of this cortical sector. Cells in the deeper stratum are, however, larger than those in the superficial one. Laterally and caudally, the superficial stratum appears to be broken up in roughly spherical clusters of cells. Caudally, the neuronal population in the posteromedial APIR subarea appears more homogeneously and densely arranged than in the AL division, being separated from the pial surface by a less thick molecular layer (L1). At its level the separation of layers 2 and 3 becomes practically indistinguishable, a feature that facilitates its distinction from the more clearly layered rostral pole of the lateral entorhinal periallocortex (L Ent, Figs. 16A, 17A, and 23A).

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C FIGURE 17 Horizontal sections through the amygdala stained for (A) Nissl, (B) acetylcholinesterase, or (C) Timm/Danscher’s histochemistry (sodium thiosulfide perfusion). For abbreviations, see list at the end of this chapter. Note the continuous large intercalated masses (IM).

A

B FIGURE 18 (ex Figs. 18 and 19) Horizontal sections through the amygdala stained for either (A) acetylcholinesterase or (B) Timm’s histochemistry (sodium thiosulfide perfusion). For description of various other nuclei see text. For abbreviations, see list at the end of this chapter.

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FIGURE 19 Frontal Nissl-stained section through the level of the posterior medial division of the bed nucleus of the stria terminalis at a plane where it is joined by the sublenticular extended amygdala (SLEA) (IA = 8.20 mm), showing also the most rostral pole of the temporal amygdala represented by the anterior amygdaloid area (AA) and the nucleus of the lateral olfactory tract (LOT) and fragmented parts (arrows) of the bed nucleus of the accessory olfactory tract (AOT). For abbreviations, see list at the end of this chapter.

Heavy metals In Timm/Danscher’s stained sections (Figs. 16B, 17C, and 23B), the APIR area shows a differential stratified staining of its neuropil, which, at the level of the deep cell stratum, displays a staining reaction more intense than that at the superficial one. Furthermore, the staining reaction in the anterolateral APIR subarea is stronger than that in the posteromedial APIR subarea.

Acetylcholinesterase In AChE preparations, the whole APIR transition area shows an intense AChE reactivity that is very characteristic of this transitional periallocortical field. However, in APIR AL the AChEpositive fibers form islands (de Olmos et al., 1985; Jolkonnen et al., 2001) which appear to overlap with that conformed by parvalbumin-IR cells, a pattern that

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FIGURE 20 Frontal Nissl (A) and Timm/Danscher’s silver sulfide (B) stained sections through the supracapsular segment of the stria terminalis (st), showing the topographical location of two clusters of cells representing the lateral (BSTSl) and medial (BSTSm) divisions of the supracapsular extended amygdala. For abbreviations, see list at the end of this chapter.

is not present in APIR PM (Jolkonnen et al., 2001). The source of this AchE-positive neuropil presumably is extrinsic coming most probably from cholinergic neurons in the basal forebrain (Fig. 17B). Choline acetyltransferase According to Hellendall et al. (1986), the region of their periamygdaloid cortex corresponding to the APIR transition area would share little ChAT activity with other members of the superficial cortical-like amygdala there, but again this activity would be higher than that in the neocortex but lesser than that in the PIR cortex. Under similar terminological grounds, the ChAT-IR in the APIR transition area would be, however, of medium density (Heckers and Mesulam, 1994).

Vomeronasal Amygdala The Bed Nucleus of the Accessory Olfactory Tract (AOT) Topographic landmarks The AOT nucleus is a small column of small, loosely packed cells closely associated with the accessory olfactory tract (Figs. 15F and 15G in Alheid et al., 1995). The nucleus accompany the accessory olfactory tract from just prior to the entrance of the tract to the molecular layer of the anterior part of the amygdala until the tract assumes a position immediately caudal to the lateral olfactory tract (LOT) nucleus. At this caudal level, the AOT nucleus lies ventral to another small group of larger, more darkly staining neurons, the anterior ventral dense-celled subdivision of the Me nucleus (Fig. 8, 21, 22, and 29).

Cytoarchitectonics The cells in the AOT nucleus are slightly larger and more darkly stained than those in the remaining subdivisions of the medial amygdaloid nucleus (Me) or those in the ACO nucleus. Further, the lack of stratification in the AOT nucleus provides an easy means of distinguishing it from the LOT nucleus (Figs. 15F and 15G in Alheid et al., 1995). Fibroarchitectonics In neurofibrillar preparations the AOT nucleus and its neighbors in the superficial amygdala present a wide molecular layer (L1) which is completely occupied by tangentially running fibers arranged in two sublayers: (i) a superficial or subpial sublayer made of lateral olfactory tract fibers that are extensions of those in the ACO nucleus and are destined to innervate the more caudal and medial edge of the AA area and (ii) a deep sublayer made up of fibers and terminal branches of the AOT. The cell layer that lies immediately deep to the AOT contains a moderately rich fiber plexus arranged in a nonlayered pattern. On this basis, the most distinguishing features of AOT in this type of preparation are the arrangement in a cup-like fashion of the AOT fibers embracing the ventromedial surface of the nucleus and the presence of the LOT immediately superficial to it. Heavy metals In Timm/Danscher’s preparations (Figs. 21C and 29B), although still less reactive than the LOT nucleus, the AOT nucleus shows a strong staining reaction, which allows it to be distinguished from the dorsally adjacent, anterior ventral Me nucleus, as well as from its lateral neighbor, the ACO nucleus. In contrast with the layered pattern in Timm’s stained sections

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in the LOT nucleus, the AOT nucleus cell layer shows a homogeneous staining reaction. Its molecular layer displays, however, a trilaminar pattern with a negative superficial or subpial layer, a moderately staining narrow intermediate strip, and a negative deeper sublamina. This deeper layer coincides entirely with the area of distribution of moderate AChE activity seen in the molecular layer of this nucleus, which corresponds with the same type of histochemical reactions occurring in other members of the vomeronasal amygdala. Acetylcholinesterase (AChE) In AChE preparations, the AOT nucleus is easily distinguished from the LOT nucleus because there is almost no AChE activity, a property that it shares with the medial amygdala and the ACO nucleus (Fig. 21A). The Posteromedial Cortical Amygdaloid Nucleus (PMCO) Topographic landmarks The PMCO nucleus constitutes the caudal third of the superficial amygdala, lying posterior to the Me nucleus and caudomedial to the PLCO nucleus. The posterior limb of the amygdaloid fissure marks its separation from the caudomedial tail of the AHI area and from the ventromedial transitional L Ent cortex. Dorsally, it is capped almost completely by the remainder of the AHI area and the hippocampal formation. Cytoarchitectonics In Nissl preparations, the posteromedial cortical nucleus appears as a homogeneous, relatively well-circumscribed ovoid mass of small- and medium-sized, palely staining cells that is separated from the pial surface by a molecular layer. A closer examination reveals, however, the presence of a narrow superficial cell layer composed of many neuronal perikarya with their long axes oriented parallel to the pial surface. This arrangement fades out at the anterior third of the nucleus. Heavy metals In Timm/Danscher’s silver-stained sections the PMCO nucleus shows quite a uniform and strong staining reaction throughout most of its extent, except rostrally, where it is more dense. Its molecular layer shows a differential staining pattern, with a completely nonreactive subpial sublamina 1A neatly outlined against a very heavily and homogeneously impregnated sublamina 1B. The wider part of the entirely white sublamina 1A and the homogeneity of the staining reaction in sublamina 1B are useful landmarks for differentiating the PMCO nucleus from the PLCO and ACO nuclei. Acetylcholinesterase In AChE preparations (Figs. 29A and 30A), the PMCO nucleus presents a relatively

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weak staining reaction. Although this reaction is not very different from that in the greater part of the PLCO nucleus, it contrasts with the very weak reaction in the small-celled part of the PLCO nucleus and the Me nucleus on one side and with the higher reactivity of the AHI area on the other. The molecular layer in the PMCO nucleus presents a moderate AChE reaction, which contrasts with the almost complete lack of reactivity in the deep cell layer. This staining in the molecular layer stops sharply at its boundary with the PLCO nucleus, but is continuous with the AChE reaction in the molecular layer of the ME nucleus and, as in the latter nucleus, it involves the entire surface of the PMCO nucleus. The Amygdalohippocampal Transition Area (AHI) The AHI transitional area is a prominent member of what has been classically defined as the corticomedial nuclear complex of the amygdaloid body, and it is located in the caudal ventromedial angle of the amygdaloid body, immediately caudal to the posterior part of the medial amygdaloid nucleus (Figs. 15L–15N´ in Alheid et al., 1995). In this account, however, the AHI transitional area is listed together with the group of superficial cortical-like amygdaloid nuclei regardless its predominantly deeper location and the fact that it does not receive direct afferent projections from either the MOB or the AOB. However, the AHI transition area, apart from being the target of intraamygdaloid afferents coming from the cortical-like and noncortical amygdaloid nuclei directly linked either to the MOB and to the AOB (see below), surfaces to the pial surface at its caudomedial ventral end by way of a deep though narrow molecular layer (L1) devoid of neuronal bodies. This anatomical feature, shared with other members of the group, as well as comparative anatomical data (see below), strongly supports its inclusion among the superficially located cortical-like amygdaloid gray formations. Topographic landmarks The AHI transition area is a prominent member of the corticomedial nuclear complex located in the caudal ventromedial angle of the amygdaloid body immediately caudal to the posterior part of the medial amygdala (Figs. 15J–15M in Alheid et al., 1995). It is in direct continuity with the ventral subiculum caudomedially and with the medial extension of the APIR transition area caudally, which separates it from the ventrolateral subdivision of the L Ent periallocortex. The AHI transition area forms a shallow inverted basket-like formation around the dorsal and caudomedial aspect of the PMCO nucleus, from which, in some places, it is indistinctly separated. On its lateral aspect, the AHI transition area merges with the BMP nucleus rostrally, and successively with the BLP nucleus, and with the main body of the APIR transition area

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C FIGURE 21 (A–C) Frontal sections through the level of the sublenticular extended amygdala (SLEA) to show in a Fluoro Gold-immunostained section (A) the presence of accessory olfactory bulb-projecting neurons along the ventral sublenticular bridge of the medial division of the extended amygdala formed by the rostral anterodorsal medial amygdaloid nucleus (not shown), the medial division of the ventral anterior amygdaloid area (AA VM), the medial sublenticular extended amygdala (SLEAm), and the ventral posterior part of the large-celled subdivision of the posteromedial division of the BST nucleus (BSTMPl). In B and C, the Timm/Danscher’s silver sulfide precipitate displays both medial and central sublenticular bridges of the extended amygdala. For abbreviations, see list at the end of this chapter.

caudally. Dorsally, it is in successive topographic relationship with the Me PD nucleus, the floor of the temporal horn of the lateral ventricle, and the region (field CA1) superior of Ammon’s horn. It is separated from the last two by fiber bands joining the stria terminalis (Figs. 17, 23, 29, and 30).

more densely packed neurons, bearing a close resemblance to those in the subiculum. Caudally and medially, the PM AHI forms the bottom and medial lip on the posteromedial limb of the amygdaloid fissure, a level at which there is a relatively wide overlying molecular layer with a low cell density (Figs. 17 and 23A).

Cytoarchitectonics The AHI area (Figs. 15J–15M in Alheid et al., 1995) is readily distinguished from the neighboring amygdaloid nuclei by its larger, more deeply staining, and tightly packed cells. A closer examination reveals internal cytoarchitectonical differences that provide for its subdivision into anterolateral (deep lying) and posteromedial (superficial) parts. The anterolateral AHI area contains cells that slightly resemble those in the PLBL nucleus, whereas the posteromedial AHI area is made up of smaller, more lightly stained, and

Fibroarchitectonics In neurofibrillar preparations, the AHI transition area is characteristically pierced by a multitude of small fiber bundles mainly longitudinally oriented. Many of these bundles, in conjunction with those coursing along its boundary with the temporal hippocampus and with the BMP nucleus, converge to form the dorsal component of the stria terminalis. Many others contribute to the formation of a fence-like plexus at its rostral pole where it borders the main body of the Me nucleus. This disposition provides for the striated

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FIGURE 22 (ex Fig. 26) Frontal sections through the amygdala at the bed nucleus of the accessory tract (AOT). (A) Nissl stain. (B) Timm’s stain (sodium thiosulfide perfusion). See text for description of various nuclei. For abbreviations, see list at the end of this chapter.

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A

B FIGURE 23 (A, B) Frontal Nissl and Timm/Danscher’s (B) stained sections through the caudal sectors of the amygdala to show the cytoarchitectonical and histochemical (zinc) arrangement of the amygdalohippocampal transition area (AHi Al and PM), the posteromedial (PMCo) and posterolateral cortical amygdaloid (PLCo) nuclei, and the amygdalopiriform transition rea (APir AL). Note that the caudal tip of the posterobasal amygdaloid nucleus (BLP) lies intercalated between the APir AL and the AHi AL. For more details see the text. IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

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FIGURE 24 (ex Fig. 28) Comparison of the bed nucleus of the stria terminalis and the centromedial amygdala in horizontal sections. (A,B) Nissl stain. (C,D) Acetylcholinesterase histochemistry. (E,F) Timm’s stain, sodium thiosulfide perfusion. For discussion see text. For abbreviations, see list at the end of this chapter.

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appearance of the AHI transition area in this type of preparation. The molecular layer of the AHI PM division, which extends dorsally and wedges between its cell layer and the dorsal border of the PMCO nucleus, is a rather poor fiber area with few axons oriented transversely and caudally to continue within the molecular layer of the adjacent subiculum.

FIGURE 25 (ex Fig. 29) Frontal section immunostained for tyrosine hydroxylase. In this section the medial division of the sublenticular extended amygdala (SLEAm) is evident as an area poor in tyrosine hydroxylase immunoreactivity. itp/mfb, fibers collected in the inferior thalamic peduncle (itp) and medial forebrain bundle (mfb). These also include descending axons from the stria medullaris.

A

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Heavy metals The Timm/Danscher’s staining pattern (Figs. 16B, 17C, 23B, 29B, and 30B) in the AHI transition area follows quite closely the distribution pattern of the previously described AChE reaction. The AHI AL division displays strong silver deposits indistinguishable from those observed in the BMP nucleus, the PLBL nucleus, and the anterolateral APIR area. In contrast, the AHI PM division shows a much weaker reaction. The molecular layer (L1) in the PM AHI division shows a uniformly strong staining reaction (although slightly weaker than that in the PMCO nucleus), but this does not provide any indication for a subdivision of this layer. Such disposition clearly contrasts with the presence in both the PMCO (the posterior cortical nucleus of Canteras et al., 1992) and PLCO amygdaloid nuclei (de Olmos et al., 1985; Alheid et al., 1995; Swanson, 1992) of a very sharply and negatively stained subpial laminae 1α intervening between brain surface and an otherwise densely stained sublamina 1B in the molecular layer of all these structures including the PIR cortex. This staining pattern in

C

FIGURE 26 (ex Fig. 30) Frontal sections through the caudal part of the posterolateral cortical amygdaloid nucleus (PLCo) stained for (A) Nissl or (B) acetylcholinesterase. Note the dense cholinesterase reaction (arrow) in the medial portion of the caudolateral part of the posterolateral cortical nucleus (PLCoCL). For abbreviations, see list at the end of this chapter.

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FIGURE 27 (ex Fig. 27) Substance P-stained sections through posterior medial bed nucleus of the stria terminalis (A,B) and the posterodorsal medial amygdala (C,D). Note the similarity between the medial and intermediate subdivisions of the medial posterior bed nucleus and the respective medial and intermediate divisions of the posterodorsal medial amygdala. The lateral subdivision of the posterior medial bed nucleus appears to be paired with the anterodorsal medial amygdala, which is not shown in these sections (see text). For abbreviations, see list at the end of this chapter.

FIGURE 28 (ex Fig. 11) Frontal sections through the amygdala from similar sections stained for (A) glutamic acid decarboxylase or (B) angiotensin II (direct prints). In both cases the dense staining in the centromedial amygdala (arrows) is in contrast with that in the basolateral complex which stains similarly to that in the adjacent cortex (see text). IGP, internal globus pallidus; opt, optic tract. For other abbreviations, see list at the end of this chapter.

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the molecular layer of the AHI area provides also a means for identifying the border of the AHI PM division from the medial part of the ventrolateral Ent cortex because the latter shows a clear lamination pattern similar to those detectable in other cortical olfactory areas. Acetylcholinesterase In AChE preparations (Figs. 17B, 29A, and 30A), the AHI area shows moderate activity, similar to the reaction in the neighboring Ammon’s horn but substantially less than that in the PLBL nucleus. The distribution pattern of the reaction is not uniform and is more pronounced at the level of the anterolateral AHI area. The strength of the AChE staining reaction in the posteromedial AHI area greatly resembles that seen in the medial extension of the APIR area. Choline acetyltransferase According to the microassay quantitative studies of Hellendall et al. (1986) the AHI transition area has the third highest average ChAT activity in the amygdaloid complex after the BLA and LOT nuclei. With regard to the distribution of ChAT IR in the rat AHI transition area, this is described as containing a ChAT IR as dense as that in the BLP nucleus (Heckers and Mesulam, 1994). Its distribution, however, is not homogeneous, but, as shown by their mapping (see their Fig. 4), it tends to be according to the parcellation suggested here, i.e., much more dense in the AHI AL subfield than in the AHI PM one.

Neurotransmitters and Neuromodulators within the Cortical Amygdala: Olfactory and Cortical-like Vomeronasal Amygdalae Cells The olfactory amygdala like the other parts of the amygdala has a rich variety of potential transmitters. Neurotransmitters In the olfactory amygdala glutamate/aspartate (GLU/ASP) may be the transmitter used for projections to the main olfactory bulb and for targets in the olfactory peduncle (Watanabe and Kawana, 1984). Neuropeptides Cholecystokinin (CCK) and vasoactive intestinal peptide (VIP cell bodies are found throughout the nuclei of the olfactory amygdala (Roberts et al., 1982; Shiosaka et al., 1983; Zaborszky et al., 1985a). Somatostatin (SOM) cell bodies are found in all members of the olfactory amygdala except for the LOT nucleus (Bennett-Clarke et al., 1980; Finley et al., 1981b), while neurotensin (NT) perikarya are found in the AHI transition and AA areas and in the ACO and PLCO

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nuclei (Roberts et al., 1982; Shiosaka et al., 1983). NPYIR perikarya are present in all cell layers of the ACO and PLCO nuclei and in the AHI transition area after colchicine administration (De Quidt and Emson, 1986). Enkephalin (ENK) cell bodies are found in the ACO and PLCO nuclei and in the AHI transition area, but not in the remaining nuclei of the olfactory amygdala (Finley et al., 1981c). Corticotropin-releasing factor (CRF) is occasionally found in the cells of the AA area (Swanson et al., 1983), but has not been found in the rest of the olfactory amygdala. Fibers Neurotransmitters In addition to fibers related to the preceding list of potential transmitters, fibers reactive for several other transmitters are found in the olfactory amygdala but no cell containing these transmitters has been identified yet within a particular subnucleus. For example, serotonin terminals reach all the nuclei of the olfactory amygdala (Fallon, 1981), and this is probably also the case for norepinephrine (NE) (Fallon, 1981) and acetylcholine (ACh). Dopamine (DA) fibers reach the AA and APIR, but provide little or no contribution to the remaining nuclei of the olfactory amygdala (Fallon, 1981). Neuropeptides Neurotensin fibers reach the APIR (Jennes et al., 1982) as do fibers that contain pancreatic polypeptide (Olschowska et al., 1981) (also neuropeptide Y; NPY). Ranatensin (RT) (Knight et al., 1983) and luteinizing hormone releasing factor (LHRH) (Barry, 1979) fibers can be seen in the PLCO nucleus, but have not been reported for the other nuclei of the olfactory amygdala. Similarly, arginine–vasopressin (AVP) and oxytocin (OX) fibers have been observed in the ACO nucleus alone (Sofroniew et al., 1981) (Table 1).

Intraamygdaloid, Interamygdaloid, and Extraamygdaloid Connections of the Olfactory and Vomeronasal Cortical-like Amygdaloid Nuclei (OLF/VM-CORLN) The hodological relationships of this nuclear group are presented in a tabular form (Tables 2–4) that summarizes its main afferent and efferent connections. However, it should be pointed out that on account of the approach followed for the compilation of Tables 2–5, as well as for Tables 6–15, and in order to present the data in the most efficient way, it has not been possible to specify the authorship of the data. For that reason, and also on account of obvious space restrictions, the following reduced list of authors, prepared in alphabetic order, should provide, however, sources for obtaining punctual bibliographic information concerning

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FIGURES 29 AND 30 (ex Figs. 21 and 20, respectively) Sagittal sections through the amygdala stained either for (A) acetylcholinesterase or (B) Timm’s histochemistry (Fig. 20, sodium thiosulfide perfusion; Fig. 21, sodium selenite perfusion). For description of various nuclei see text. For abbreviations, see list at the end of this chapter.

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FIGURE 30

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FIGURE 31 (ex Figs. 31 and 32) Frontal (A,B) and horizontal (C,D) sections through the bed nucleus of the stria terminalis and the central amygdaloid nucleus, immunostained for tyrosine hydroxylase (TH). Note that the capsular subdivision of the dorsal part of the lateral bed nucleus (BSTLDc) is poor in tyrosine hydroxylase staining, in contrast to that in the central subdivision of the dorsal part (BSTLDcn). In the horizontal tyrosine hydroxylase-immunostained sections through the intermediate subdivision of the lateral bed nucleus of the stria terminalis (BSTLI) and the anterodorsal subdivision of the medial central amygdaloid nucleus (CeM ADd) from a preadolescent male rat it is possible to detect a preferential location (CeM and BSTLi) of the TH immunostained neurons in both of these nuclei. For abbreviations, see list at the end of this chapter.

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FIGURE 32 (ex Fig. 33) Frontal Timm’s stained section through the bed nucleus of the stria terminalis, 3 days after cutting the stria terminalis. Note the disappearance of metal sulfide staining, except for the nucleus of the commissural component of the stria terminalis. MnPO, median preoptic nucleus. For other abbreviations, see list at the end of this chapter.

such authorship, i.e., Bernard et al., 1990; Canteras et al., 1992, 1995; Cassell et al., 1986, 1999; de Olmos et al., 1985; Dong et al., 2001a, 2001b; Freedman and Cassell, 1993; Gray and Magnuson, 1987, 1992; Gray et al., 1989; Eray et al., 1984 Grove, 1988a, 1988b; Hasue and Shammah-Legnado, 2002; Hopkins, 1975; Harley et al., 1991; Jia et al., 1992a; Krukoff et al., 1993; LeDoux et al., 1985: Li et al., 1990; Majak et al., 2002; McDonald, 1987, 1991, 1992, 1998; McDonald et al., 1999; Moga and Gray, 1985a, 1985b; Moga et al., 1989, 1990; Nitecka and Frotscher, 1989; Ottersen, 1981; Perez-Clausell et al., 1989; Petrovich et al., 2001; Prewitt and Herman, 1998; Ricardo and Koh, 1978; Riche et al., 1990; Risold et al., 1994; Rizvi et al., 1991; Roder and Ciriello, 1993; Roeling et al., 1993; Roland and Sawchenko, 1993; Romanski and Le Doux, 1993; Rosen et al., 1991; Sakanaka et al., 1981, 1986; Saper and Loewy, 1980; Savander et al., 1995; Schwaber et al., 1980, 1982, 1988; Seroogy and Fallon, 1989; Seroogy et al., 1989; Sesack et al., 1989; Shammah-Lagnado et al., 1999, 2000, 2001; Simmerly and Swanson, 1986, 1988; Stefanacci et al., 1992; Sun and Cassell, 1993; Sun et al., 1991, 1994; Switzer et al., 1985; Takada, 1990; Takayama and Miura, 1991; Turner and Herkenham, 1991; Van der Kooy et al., 1984; Veening et al., 1984; Vertes, 1991; Wallace et al., 1989; Woulfe et al., 1988; Yamano et al., 1988a, 1988b; Yasui et al., 1991a, 1991b; Zahm et al., 1999).

THE EXTENDED AMYGDALA (EXA) A large amount of morphological, histochemical, immunocytochemical, and hodological data has been gathered, particularly in the last 18 years to support the theory of a neuronal continuum, known as the extended amygdala, within the basal forebrain (de Olmos et al., 1985) The concept started with the pioneering comparative and developmental work reported by J. B. Johnston (1923), who suggested the existence of close anatomical relationship between the central and medial amygdaloid nuclei and the bed nucleus of the stria terminalis. He pointed out that in the human embryos and in lower vertebrates these two territories form a continuum, and he proposed that this continuity is still present in adult mammals through a more or less continuous cell column embedded in the stria terminalis. In 1969 and 1972, de Olmos proposed a revival of this concept by reporting, on the basis of a specific and sensitive cupric-silver histochemical reaction, the existence even in adult mammals of a continuity between the central amygdaloid nucleus and the lateral bed nucleus of the stria terminalis that includes columns of gray matter located in the sublenticular (subpallidal) substantia innominata (see below). Thus, this amygdalosublenticulostrial continuum or extended amygdala comes to form a large ring formation around

IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

548 JOSE S. DE OLMOS ET AL.

IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

FIGURE 33 Timm/Danscher-stained frontal sections of experimental material intended for the tracing of zinc-rich projections to the extended amygdala after microinjections of sodium selenite in the central amygdaloid nucleus (A) or the bed nucleus of the stria terminalis (B). In both cases, the accumulation of zinc in the cell bodies of neurons by retrograde transport from either injection places revealed several sources of zinc-containing afferents to the extended amygdala. For abbreviations, see list at the end of this chapter.

B FIGURE 34 Frontal sections of brains microinjected with biotin dextran amine (BDA) in the caudal sector of the anterior division of the medial amygdaloid nucleus (MeAD) showing the distribution of BDA-immunostained axons running in the stria terminalis and ventral amygdalofugal pathway to reach their final destination in the medial preptic region (A) and tuberal hypothalamus (B). Note the very strong stream of BDA-immunostained axons at the level of the medial sublenticular extended amygdala (SLEAm). For abbreviations, see list at the end of this chapter.

ANG II

ANP

CART

Distribution in the Extended Amygdala Continuum of Perikarya of Neuropeptide-Synthetizing Neurons on the Basis of the Data Available in the Pertinent Literature

CCK

BSTMA

ⵧ

BSTMV

ⵧ

CRF

ⵧⵧ

ENK

ⵧⵧ

GAL

NKB-2P

ⵧⵧ

ⵧⵧⵧ

ⵧⵧ

BSTMPm

ⵧⵧⵧ

ⵧⵧⵧ

ⵧ

ⵧ

ⵧⵧ

ⵧⵧⵧⵧ

ⵧ

ⵧⵧⵧ ⵧⵧ

BSTIA

ⵧⵧⵧⵧ

ME AV

ⵧⵧⵧ

ME AD

ⵧⵧⵧ

ME PV ME PD BSTSm

ⵧⵧⵧ

ORX

ⵧⵧ

VIP

ⵧⵧⵧ

BSTMP l

ⵧⵧ

ⵧⵧⵧ

BSTMP i

ⵧⵧⵧ

ⵧⵧⵧ

ⵧⵧⵧ

BSTMPm

ⵧⵧ

ⵧⵧⵧ

ⵧⵧⵧ

BSTIA

ⵧ

ⵧ

ⵧⵧⵧ

ⵧⵧⵧⵧ

BSTIm

ⵧ

ⵧ

ⵧ

SLEAm

ⵧⵧⵧ

ⵧⵧ

ⵧⵧ ⵧ

ⵧ ⵧ

ⵧⵧⵧⵧ ⵧⵧⵧⵧ

ⵧ

ⵧⵧ

ⵧ

BSTLD

ⵧⵧⵧⵧ ⵧⵧⵧⵧ ⵧⵧ

ⵧⵧⵧ

BSTLV

ⵧ

BSTLP R

ⵧ

BSTLP C

ⵧ

BSTl l

ⵧⵧⵧ

BSTS l

ⵧⵧ

ⵧⵧⵧ ⵧⵧⵧ

ⵧ

ⵧ

ⵧⵧ

ⵧⵧ

ⵧⵧ

ⵧ

ⵧⵧⵧ

ME AV

ⵧⵧ

ⵧⵧⵧ

ⵧⵧⵧ

ⵧⵧⵧⵧ

ME AD

ⵧ

ME PV

ⵧⵧ

ME PD BSTSm

ⵧⵧⵧⵧ ⵧ

ⵧⵧⵧ

ⵧⵧ

ⵧ

ⵧ ⵧⵧⵧ

ⵧⵧⵧ

ⵧⵧⵧ

ⵧ

BSTLD

ⵧⵧⵧ

ⵧⵧ

ⵧⵧ

BSTLV

ⵧ

ⵧⵧ

ⵧⵧ

BSTLP R

ⵧⵧ

ⵧⵧ

ⵧⵧ

BSTLP C BSTl l

ⵧⵧⵧ ⵧ

CeM AD

ⵧ

ⵧ

ⵧⵧ

CeM AV

ⵧ

ⵧ

ⵧⵧ

CeM PV

ⵧ

ⵧ

ⵧⵧ

ⵧ ⵧ

ⵧⵧ

ⵧⵧⵧ ⵧⵧ

ⵧⵧ

BSTS l

ⵧⵧ

SLEA c

ⵧⵧⵧ

ⵧ

ⵧⵧⵧ

ⵧⵧ

CeM AD

ⵧⵧⵧ

ⵧ

ⵧ

ⵧⵧ

ⵧⵧⵧ

CeM AV

ⵧⵧⵧ

ⵧ

ⵧ

ⵧ

ⵧⵧⵧ

CeM PV

ⵧⵧ

IPAC

IPAC

CE I

ⵧ

ⵧⵧⵧ

ⵧ

ⵧⵧ

CEL

ⵧ

ⵧⵧⵧⵧ ⵧⵧⵧⵧ ⵧⵧⵧ

ⵧ

ⵧⵧⵧⵧ

CEC

ⵧ

ⵧ

ⵧ

ⵧ weak; ⵧ ⵧ moderate; ⵧ ⵧ ⵧ dense; ⵧ ⵧ ⵧ ⵧ very dense.

VP

BSTMV

ⵧⵧ

ⵧ

SLEA c

TRF

ⵧⵧ ⵧⵧ

ⵧⵧ

SP

ⵧ

ⵧ

ⵧⵧ

ⵧ

SOM

BSTMA

ⵧ

ⵧ

SN

ⵧ

ⵧⵧⵧ

ⵧⵧ

OT

ⵧⵧⵧ

ⵧ

ⵧⵧⵧ

ⵧⵧⵧ

ⵧⵧⵧ

ⵧ

ⵧⵧ

ⵧ

ⵧ

CE I

ⵧ

CEL CEC

JOSE S. DE OLMOS ET AL.

IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

ⵧ

SLEAm

PACAP

ⵧ

ⵧ

ⵧ

BSTIm

NT

ⵧⵧ

BSTMP l BSTMP i

NPY

550

TABLE 1

551

19. AMYGDALA AND EXTENDED AMYGDALA OF THE RAT

TABLE 2

Intraamygdaloid Connections of the Olfactory and Vomeronasal Cortical-like Amygdaloid Nuclei (OLF/VM-CORLN) OLF/VM-CORLN Intrinsic connections

Efferents to MEXA

Efferents to CEXA

Efferents to LBNC

Olfactory cortical-like

Olfactory cortical-like

Olfactory cortical-like

Olfactory cortical-like

Olfactory cortical-like

nuclei:

nuclei:

nuclei:

nuclei:

nuclei:

Lateral olfactory tract

LOT → only to

LOT → Me (bilaterally?!)

LOT → bilaterally

AA → BMA (vd), BLP, BLV

(LOT)

contralateral LOT

to CeM AA→BSTMA, BSTMV,

Anterior amygdaloid

AA → ipsilateral ACO

area (AA) divided into

and PLCO

AA→ CeL and CeM AD

neurons scattered in AAD

(large-celled). These

deep dorsal (AAD) and superficial ventral (AAV);

AA (contralaterally)

the AAV is subdivided

→AA and PMCO

into lateral AAVl) and (medial (AAVm) parts.

BSTMPl

These projections may originate in cholinergic

ACO → Me AD (vd),

projections may originate

LOT→ BM ipsilaterally

Me PV, not MePD

in cholinergic neurons

and bilaterally to BL

scattered in AAD. ACO → lateral nucleus,

ACO→BSTMA, rostral AA → PMCO and AHI.

BSTMPm, BSTMPi,

AA→ SLEA

These projections may

lBSTMPl, and SLEAm

BSLTP, BSTLV

BLV PLCO and APIR → all the

Anterior cortical

arise in the cholinergic

amygdaloid nucleus

elements scattered

PLCO →almost all the

ACO → ventral and

(ACO)

in AAD.

members of the MEXA

most rostral CeC

amygdaloid complex,

and SLEAc

BM included

Posterolateral cortical

ACO → bilaterally to

amygdaloid nucleus

AA (m)

nuclei of the basolateral

PLCO→ vomeronasal and nonvomeronasal

PLCO→ all of the

sectors of the MeAD

subnuclei of the central

Vomeronasal superficial

ACO →ipsilaterally to

(vd) including a PLCO

amygdaloid group

cortical-like nuclei

LOT, PLCO, and APIR

→ SLEAm

except BSTV and BSTS

divided into anterolateral

PLCO → all the nuclei

PLCO→ BSTMPl (large-

PLCO →CeM and

(APIR AL) and

of the olfactory

celled) and BSTMPi

(m) but more particularly CeC (Vd)

(PLCO) Amygdalopiriform

PMCO → BMP, BMA,

transition area (APIR)

posteromedial

cortical-like amygdala

(medium-celled)

(APIR PM) parts

including APIR PM

subdivisions of medial

Anterior tip of the

ACO and PLCO →

principal division of

vomeronasal PMCO

the medial amygdala

nucleus and AHI area

division of the

PLCO → SLEAc and

BST nucleus

BSTLP, this latter bilaterally

PLCO → medial (BSTMAm) and lateral

APIR → CeL, CeC, CeM,

(BSTMAl) and ventral

BSTL, and SLEAc

PLCO and PMCO nuclei

(BSTMV) subdivisions

APIR → all members

the BSTM nucleus

of the anterior part of

Vomeronasal superficial cortical-like nuclei:

AHI does not seem to supply the LBNC

PLCO → contralateral

(MeR)

BLV (massively)

of the olfactory and

Vomeronasal superficial cortical-like nuclei:

vomeronasal cortical-like

APIR → MeAD,

Bed Nucleus of the

nuclei including AHI

MeAV, and BSTIA

Accessory Tract (BAOT)

area Vomeronasal

PMCo terminates Ce

superficial cortical-like

PMCO → CeMav

nuclei:

(massively)

(l) light; (m) moderate; (d) dense; (vd) very dense; (vl) very light; (pm) poor to moderate.

Continued

IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

552

JOSE S. DE OLMOS ET AL.

TABLE 2

Intraamygdaloid Connections of the Olfactory and Vomeronasal Cortical-like Amygdaloid Nuclei (OLF/VM-CORLN)—cont’d OLF/VM-CORLN Intrinsic connections

Efferents to MEXA

Efferents to CEXA

Posteromedial Cortical

BAOT → PMCO and

Vomeronasal superficial

AHI → CeM (vl) and

amygdaloid nucleus

AHI (vd)

cortical-like nuclei:

SLEA (vl), BSTLP (vl)

Efferents to LBNC

(PMCO) Amygdalohippocampal

PMCO → AAD, AAVl,

PMCO → ventral part

AAVM (massively)

of Me, Me AD, SLEAm,

Transition Area (AHI)

BSTMA, BSTV, BSTMPi, PMCO → BAOT

and BSTIA

(vd), AHI AHI → Me (all) and PMCO

BSTMAm, BSTMPm,

(contralaterally) →

BSTMV, and BSTI pm

PMCO PMCO → AA, APIR, and caudal PLCO PMCO → AHI area only to the contralateral side but does not seem to send any projections to the ipsilateral AHI PMCO → small-celled PLCO subnucleus AHI → PMCO (l) AHI → AOB and AA (l) light; (m) moderate; (d) dense; (vd) very dense; (vl) very light; (pm) poor to moderate.

the internal capsule. Later de Olmos et al. (1985), on the basis of experimental anatomical data obtained by applying retrograde tract-tracing procedures, described that neurons projecting to the medial preoptic–hypothalamic continuum conformed to a continuous field extending from the intermediate and medial divisions of the BST nucleus through the sublenticular sector of the so-called substantia innominata (now the SLEAm) into the medial amygdaloid nucleus and involving also scattered neurons in the supracapsular part of the stria terminalis. On account of these initial findings and on a wealth of diverse morphological, cytochemical, and functional data it is possible to recognize in the EXA at least two major parallel divisions or ring formations: (A) a medial division (MEXA), which is named after the Me nucleus and its rostral partner, the BSTM nucleus, which also includes cell columns within the stria terminalis and in

the subpallidal area, and (B) a central division (CEXA) that includes the Ce and lateral BST nuclei, in addition to cell columns that bridge the gap between these main structures both within the stria terminalis and in the subpallidal region. To this ring, more recent data incorporates to the CEXA the interstitical nucleus of the posterior limb of the anterior commissure (IPAC) which maintains close structural, cytochemical, and hodological relationships with both the Ce and the BSTL (Figs. 1–4, 6A, 6B, 7A, 9A, and 9B).

The Medial Division of the Extended Amygdala (MEXA) The medial division of the extended amygdala is a large complex of nuclei more or less recognizable by their extensive connectional relations to the medial

IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

553

19. AMYGDALA AND EXTENDED AMYGDALA OF THE RAT

TABLE 3

Olfactory and Vomeronasal Cortical-like Amygdaloid Nuclei Afferents Subcortical Afferents

Cortical afferents

Telencephalon

Diencephalon

Brain stem afferents

Olfactory cortical-like

Input from mitral cells

AONv/p and AONd →

Hypothalamic inputs

To olfactory cortical-like

nuclei:

of the MOB→ lamina

Lamina 1B of LOT and

to the AA:

nuclei:

1A of the superficial

superficial part of AA

Lateral preoptic lateral hypothalamic

Dopaminergic (DA9, DA8)

Diagonal band nuclei

continuum (LPO-LH),

afferents from the VTA and

Lateral olfactory

molecular layer of each

tract (LOT)

of these nuclei of the olfactory amygdala

Anterior amygdaloid area (AA) divided in

MOB mitral cells→ AA,

deep dorsal (AAD)

COA, PLCO, APIR, and

and superficial ventral

MeR

(AAV); the AAV is

(HDB/VDB) → all the

tuberal division of the

RRF → Olfactory amygdala

nuclei of the olfactory

lateral hypothalamic

except LOT

amygdala

area (LHTC), and possibly the

Locus coeruleus (NE 5) →

Only LOT has shown to

ventromedial hypothal.

all the nuclei of the

be a ChAT-IR projection.

nuc. (VMH) as well as

olfactory amygdala except APIR

subdivided into lateral

MOB linked DTT

This projection comes

the medial preoptic area

(AAVl) and medial

(taenia tecta 2) → LOT,

from the HDB and

(MPA) and anterior

(AAVm) parts

AA, ACO

terminates in lamina 1B.

hypothalamic (AH)

Serotoninergic

areas; paraventricular

(5-HT) (B7 and B5) afferents

Anterior cortical

AON→LOT and PLCO

Dorsal (En D) and

hypothalamic (PAVH)

from DRN and MRN raphe

amygdaloid nucleus

PIR cortex→ all

ventral (En V)

and arcuate (Arc) nuclei

nuclei → olfactory amygdala

(ACO)

olfactory cortical-like

endopiriform nuclei →

→ AA, ACO, and PLCO

nuclei and areas

all the nuclei of the

(ipsilateral) →PLCO

olfactory amygdala but

and APIR

Practically all the nuclei

more particularly

of the olfactory

from EnD

Posterolateral cortical amygdaloid nucleus (PLCO)

corticoal-like Amygdalopiriform

amygdala receive

To vomeronasal

transition area

imputs from Ammon’s

superficial cortical-like

(APIR) divided into

Horn related areas:

amygdaloid nuclei:

anterolateral (APIR AL)

LPB and MPB → AA but do not terminate in any other

VMH and

nucleus of the olfactory

premammilary

amygdala. NEergic cells

nuclei → PLCO (bilat)

in the ventral lateral medulla oblongata (MVL) → AA PLCO

To Vomeronasal

and posteromedial

Ventral CA1 field of

(APIR PM) parts

Ammon’s horn → AA, APIR

AON d →PMCO

superficial cortical-like nuclei:

Dorsal (En D) and ventral

Anterior tip of the

(En V) endopiriform nuclei → PMCO

To vomeronasal superficial cortical-like nuclei:

Medial Preoptic

principal division of the

Archicortex of the

Anterior

VTA, PAG, and

medial amygdala (MeR)

ventral S →LOT, AA,

Hypothalamic

DRN → AHI

PLCO, APIR, LOT

Continuum →AHi

Vomeronasal superficial

LEnt. periallocortex

VMH → AHi (bilat.)

cortical-like nuclei:

→LOT, AA, PLCO, ACO and APIR

Bed nucleus of the accessory tract (BAOT)

Dorsal (DPM) and (VPM) ventral

Caudal portion AIV

premammillary

periallocortex → ACO

nuclei → PMCO,

and PLCO

AHI (bilat)

Posteromedial cortical

Rostral portion of the

Arcuate nucleus →

Amygdaloid Nucleus

AIp cortex →ACO,

AHI (bilat)

(PMCO)

APIR, and PLCO

(l) light; (m) moderate; (d) dense; (vd) very dense; (vl) very light; (pm) poor to moderate.

Continued IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

554

JOSE S. DE OLMOS ET AL.

TABLE 3

Olfactory and Vomeronasal Cortical-like Amygdaloid Nuclei Afferents—cont’d Subcortical Afferents

Cortical afferents

Telencephalon

Diencephalon

Amygdalohippocampal

PRh periallocortex →

Perivetricular

transition area (AHI)

APIR

hypothalamic nucleus (Pe) → AHI

Caudal IL and PrL periallocortices →

The lateral

AA and ACO

preoptic–lateral hypothalamic

PDC periallocortex →

continuum → PMCO

AAD and AAV, PLCO

and AHI

To vomeronasal

Thalamic inputs:

superficial cortical-like nuclei:

Mediodorsal (MD), interanteromedial

Input from AOB tufted

(IAM), paratenial,

cells →BAOT and PMCO

(IAM), paratenial, RE, and RH thalamic

Olfactopeduncular TrD

nuclei → ACO

and TrL → PMCO

PAV, PT, and RE thalamic nuclei → APIR

Anterior and posterior PIR cortex → layer 1B

Paraventricular

of PMCO

thalamic nucleus → (PaV) AHI

Almost all nuclei of vomeronasal cortical-like amygdala receive inputs from Ammon’s Horn related areas: Ventral CA1 field → PMCO Archicortex of the ventral S → PMCO and AHI LEnt. periallocortex → PMCO (vd) and AHI Ventral MEnt. periallocortex → AHI PRh periallocortex → PMCO (only) (l) light; (m) moderate; (d) dense; (vd) very dense; (vl) very light; (pm) poor to moderate.

IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

Brain stem afferents

555

19. AMYGDALA AND EXTENDED AMYGDALA OF THE RAT

TABLE 4

Olfactory and Vomeronasal Amygdaloid Nuclei Efferents To subcortical targets

Olfactory cortical-like nuclei:

To cortical targets

Telencephalon

Diencephalon

To brain stem targets

Olfactory cortex-like

PLCO → AON (medial,

Olfactory connections

AHI → rostral

like amygdala

dorsal, and

with the

PGA

reciprocates inputs

posteroventral parts)

preoptic/hypothalamic

Nucleus of the lateral

from the main

olfactory tract (LOT)

olfactory bulb (MOB).

Anterior amygdaloid

Mainly, the LOT and

area (AA)

PLCO nuclei

All olfactory amygdala

(bilaterally) →MOB

ventral striatum:

Anterior cortical

AA → (ipsilaterally)

ipsilareral

amygdaloid nucleus

→ MOB

continuum: ACO and APIR→ AON (posteroventral part)

(ACO) Posterolateral cortical

LOT →medial edge

preoptic area AA → LH area LOT→ retrochiasmatic

of primary olfactory

LOT (bilat) →islands of

cortex

Calleja

amygdaloid nucleus (PLCO)

AA → periventricular medial

area, VMH, and zone between VMH and arcuate nucleus

AA, ACO → anterior

AA → lateral Tu PLCO → few axons

and posterior PIR cortex Amygdalopiriform transition area (APIR)

PLCO → deep

Anterior tip of the

posterior PIR cortex

ACO, PLCO → islands

seem to terminate in the

of Calleja and L3

preoptic region where

APIR → posteromedial

the undifferentiated part

Tu

of the medial preoptic

layers adjacent principal division of

they distribute mainly to

the medial amygdala

APIR → lateral caudal

(MeR)

sector of anterior PIR

ACO, PLCO, APIR

part of the medial

cortex

(ipsi)) → IPAC 1 (fundus

preoptic nucleus (MPO)

striati)

Vomeronasal cortical-like nuclei:

area (MPA) and lateral

PLCO → massive

Hippocampal archicortex receives

LOT (bilat) → IPAC 1

noticeable terminal fields in

Bed nucleus of the

inputs from all

(fundus striati)

the ventrolateral

accessory tract (BAOT)

except LOT

Posteromedial cortical

ACO, PLCO,

amygdaloid nucleus

APIR → ventral S

(PMCO)

(molecular layer)

ventromedial nucleus VMH PLCO (bilat) →

(VL) (both the cellular core

caudomedial Acb

and the shell), lateral tuberal nucleus (LT), and ventral

APIR (ipsi) →

premammillary nucleus

posteromedial Acb

(VPM) → while other

Amygdalohippocampal

APIR PM → CA1,

transiton area (AHi)

CA3, and DG

ACO, PLCO, APIR → (ventral En) and

capsule around the

ACO, PLCO, and

dorsal (En D)

mammilary nucleus

APIR → tenia tecta (2/3)

endopiriform nucleus

Vomeronasal MEXA:

fiber contingents seem to

Medial amygdaloid nucleus (Me)

terminate in the cell-sparse

With the thalamus: AA →only to En D

ACO → MD thalamic nucleus

(l) light; (m) moderate; (d) dense; (vd) very dense; (vl) very light; (pm) poor to moderate.

Continued

IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

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JOSE S. DE OLMOS ET AL.

TABLE 4

Olfactory and Vomeronasal Amygdaloid Nuclei Efferents—cont’d To subcortical targets

To cortical targets

Telencephalon

Diencephalon

PosteroMedial

Periallocortex: Insular

ACO and PLCO → HDB,

PLCO → light

subnucleus or column

and frontal ACO, PLCO,

VDB, and ventral part

innervationto the

of the posterior medial

APIR → agranular

of SL nucleus

paraventricular (PAV)

division of the bed

insular (AIV and AIP

nucleus of the stria

cortices, and infralimbic

AHi → posteromedial

thalamic nuclei as well

terminalis (BSTMPM)

(IL) cortex

Acb and Tu

as medial sectors of MD

ACO → prelimbic

AHi → En D

With the epithalamus:

To brain stem targets

and parathaenial (PT)

AA and ACO → LHb

cortex AHI and PMCO → Periarchicortex:

ventral SL and SFi

Entorhinal (Ent) and

nuclei, etc.

Amygdaloid/vomeronasal

perirhinal (PRh)

projections to the

cortices: ACO, PLCO,

preoptic–hypothalamic

APIR → all layers

continuum:

of the L Ent. and PRh cortices

Few neurons from PMCO → periventricular hypothalamic (Pe)

Vomeronasal cortex-like

nucleus

amygdaloid nuclei: PMCO → to the area BAOT, PMCO → AOB

around but not in the

AHI → ventral S and

hypothalamic nucleus

paraventricular adjacents parts of the Ammon’s horn field CA1

PMCO, AHi → cell-poor shell surrounding VMH

AHI and PMCO → DTT (precommissural

PMCo and AHi (bilat) →

hippocampus)

premammillary nuclei

Entorhinal cortex: PMCO

AHI→ Anteroventral

axons terminate →

periventricular nucleus

lamina 1B and 2 of dorsal

of the preptic region

L Ent and ventra M

(AVP), the MPO nucleus,

Ent cortex

the VMH VL, and the

AHI and PMCO →

part of the LHA and the

IL(d) prefrontal

VPM nucleus are all

periarchicortex

densely innervated and

immediately adjacent

in a capsule around the medial mammillary body (MM). (l) light; (m) moderate; (d) dense; (vd) very dense; (vl) very light; (pm) poor to moderate.

Continued

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TABLE 4

Olfactory and Vomeronasal Amygdaloid Nuclei Efferents—cont’d To subcortical targets

To cortical targets

Telencephalon

Diencephalon

To brain stem targets

To the thalamus: PMCO → MD thalamus AHI→ PAV, PT, and medial MD thalamic nuclei

hypothalamus and their somewhat extensive network of intrinsic associative connections. Several of the corticallike nuclei of the amygdala, notably the PLCO nucleus, the AHI transition area, and the BM nucleus, send relatively massive projections to the MEXA (Figs. 1–4), with only a few terminals reaching the adjacent areas in the CEXA. In describing the various structures belonging to the MEXA, we begin with the bed nucleus of the stria terminalis (BST) for the same reasons that were thoroughly discussed in the previous version of this chapter (Alheid et al., 1995). The Medial Bed Nucleus of the Stria Terminalis (BSTM) The BST has been traditionally divided into medial and lateral parts. Of these, the lateral part is described within the context of the CEXA. The medial part of the BST may be further divided into anterior, ventral, and posterior parts. Topographic landmarks The anterior part of the medial bed nucleus of the stria terminalis (BSTMA) The anterior part is located dorsal to the crossing fibers of the anterior commissure, lying within the course of the dorsal component of the stria terminalis as it traverses the griseum surrounding the dorsal and rostral aspects of the commissure (Figs. 15A and 15B in Alheid et al., 1995). It borders the ventral lateral septum rostromedially and reaches the rostrodorsal border of the ventral part of the medial bed nucleus of the stria terminalis ventrocaudally. Dorsally, the stria terminalis separates the anterior part of the medial bed nucleus from its posterior part, whereas ventrally the anterior and posterior parts of the medial bed nucleus are separated by the commissural component of the stria terminalis. Medially, the cells of the BSTMA are medium sized and irregular in shape and are dispersed among the fibers of the stria terminalis. More laterally, the cells are smaller and more tightly packed (Figs. 7, 13, 14, 15, 20, 24A, 24C, 24E, 31, and 33B). The ventral part of the medial bed nucleus of the stria terminalis (BSTMV) The ventral part of the medial bed nucleus is located ventral to the anterior commis-

sure and is composed of medium-sized cells interposed between the parastrial nucleus (i.e., the “round nucleus” of Raisman and Fields (1973)) and the ventral part of the lateral bed nucleus of the stria terminalis (Fig. 15B in Alheid et al., 1995). Rostrodorsally, it reaches the anterior part of the medial bed nucleus, whereas it borders the intermediate part of the lateral bed nucleus caudally. The cells are loosely arranged, contrasting with those of the adjacent parastrial nucleus, and are larger and more chromophilic than the more closely spaced neurons of the ventral lateral bed nucleus of the stria terminalis. Ventrally, a cell-poor zone separates it from the preoptic area (Figs. 7, 11, 13, 15, 31, 32, and 33B). The posterior part of the medial bed nucleus of the stria terminalis (BSTMP) The BSTMP is a heterogeneous group of cells arranged in columnar fashion, following the postcommissural fibers of the st (Figs. 15C–D´ in Alheid et al., 1995). It is the largest and longest of the major divisions of the BST, extending continuously from the uppermost caudal part of the paraseptal BST, behind the ac, to the hypothalamus, where it merges with the anterior hypothalamic area. For most of its dorsoventral extent, it borders the sm caudally, and the fx medially. Rostrally, at the paraseptal level, it is apposed to the intermediate part of the BSTL (Figs. 11, 13, 14, 19, 21A, 21B, 24A, 24C, 24E, 27A, 27B, 31C, 34A, and 34B). Cytoarchitectonics Based on its cytoarchitecture, the posterior part of the BSTM can be subdivided into three cell columns: (i) a medial small- or dense-celled column (BSTMPm), (ii) an intermediate medium-celled column (BSTMPi), and (iii) a lateral large-celled column (BSTMPl) (Figs. 15A and 15B Alheid et al., 1995). The medial dense-celled column of the posterior part of the medial bed nucleus of the stria terminalis (BSTMPm) This column is composed of homogeneous, densely packed, darkly stained, round to oval neurons, located in the extreme medial aspect of the posterior part of the medial bed nucleus (Fig. 24A). The small-celled column extends ventromedially and stops at a level adjacent to the posterodorsal preoptic nucleus. In the

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ventral part of the small-celled column, the densely packed cells form an oval cylinder in horizontal sections, whereas dorsally this column is larger and more rounded. The cells in the dorsal subventricular portion of this column expand to form a wider head, which appears to be partially encapsulated by a cell-poor zone of fibers. These cells are more loosely packed and even more homogeneous in appearance than those in the deeper portions of this column, where larger cells are sometimes intermixed with the predominantly small-cell population. The cells in the dorsal part of the BSTMPm lie adjacent to the AOT as it separates from the stria terminalis. The fact that the AOT reaches only the dorsal part of the BSTMPm suggests that a dorsal to ventral differentiation may be warranted within this nucleus, although this is not readily apparent on other cytoarchitectural grounds. The BSTMPm is almost completely surrounded by the BSTMPi, which extends more dorsally and laterally, as well as more ventrally and medially, than the former cell group. The intermediate medium-celled column of the posterior part of the medial bed nucleus (BSTMPi) The BSTMPi (Fig. 24A) is composed of loosely arranged, more lightly stained, medium-sized, spindle- and angular-shaped cells. Throughout most of its course, this column lies just rostrally apposed to the stria medullaris thalami (sm in Figs. 13, 21A,B, 24A,C,E, 31C). At its caudal end, it separates from the dispersing fibers of the sm, but at the same time comes to lie rostral and ventral to the descending columns of the fornix (f) (Fig. 21B). Caudally, the cells of the BSTMPi merge imperceptibly with the medium-celled portion of the anterior hypothalamus (Fig. 16A), whereas, dorsally and laterally, the cells of the medium-celled column form the floor of the lateral ventricle. The lateral column of the posterior part of the medial bed nucleus of the stria terminalis (BSTMPl; Fig. 24A) The lateral column contains a more heterogeneous population of neurons than the other two divisions of the BSTMP. Most of the cells in theBSTMPl are loosely arranged in rows oriented parallel to the incoming fibers of the stria terminalis. These cells range from medium size to large, with some of the largest cells of the BSTMP occurring in this column. Fiberarchitectonics In fiber preparations the anterior part of the BSTM division is characterized by a rich plexus of obliquely running fibers representing the supracommissural component of the stria terminalis. In the posterior BSTM division, the small-celled medial column stands out as relatively fiber poor in comparison with the medium-celled intermediate and large-celled

lateral columns. The thick fiber bundles seen in the medium-celled intermediate and large-celled lateral subdivisions mainly represent stria terminalis fibers which parallel the course of these two cell columns. The large-celled lateral subdivision of the BSTM division is also invaded by transverse running fibers ascending from the lab fiber system (Fig. 13C). Heavy metals In Timm/Danscher’s silver sulfide preparations (Figs. 14A, 24E, and 32), the BSTMA subdivision is very densely stained but is still more lightly stained than the adjacent BSTL division. There is a gradual increase in silver precipitate from the mediocaudal to the rostrolateral parts of the BSTMA subdivision. The BSTMV subdivision stains lightly in Timm/ Danscher’s preparations, which can be distinguished from the dense reaction in the BSTLV subnucleus. The BSTMPm column appears to be densely stained by the Timm/Danscher’s procedure, but is surrounded by a capsule of still darker, very dense staining in the BSTMPi subdivision (Fig. 21). The BSTMPl column also stains densely, but to a lesser extent than the BSTMPi subnucleus. At its lateral edge it merges with the moderately stained medial division (posteroventral part) of the SLEA. The presumably zinc-containing terminals identified by Timm/Danscher’s stain in the BSTM appear to depend almost entirely on the morphologic integrity of the stria terminalis and on that of the AHI (de Olmos et al., 1985; Perez-Clausell et al., 1989). Acetylcholinesterase In AChE sections (Fig. 24C), the BSTMA subdivision contains poorly stained neuropil compared with the stronger reaction in the adjacent BSTL division, whereas the BSTMV subnucleus is lightly stained and indistinguishable from the BSTLV subnucleus. In the posterior part of the BSTM subdivision, the BSTMPm and the surrounding BSTMPi subnuclei are only poorly stained by the AChE reaction. In the BSTMPi subnucleus this staining gradually becomes more dense laterally as the BTMPi and BSTMPl subnuclei blend. It is difficult to discriminate a clear-cut border between the BSTMPi and the BSTMPl columns because of this gradual increase in AChE reactivity. The BSTMPl column provides a light reaction, the staining of which merges with that of the moderate reaction in the medial division (posteroventral) of the SLEA, with which it is continuous. The Medial Amygdaloid Nucleus (Me) Topographic landmarks The Me nucleus is a prominent member of the superficial amygdala and occupies its rostromedial aspect (Figs. 6, 8, 13, 16–18, and 21C). It begins just caudal and medial to the LOT, overlying for a short distance the BAOT nucleus, and extends

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caudally until the temporal horn of the lateral ventricle appears. At this level, it lies dorsomedial to the cephalic pole of the AHI and forms the medial aspect of the anterior wall of this portion of the ventricle. Along its entire extent, the medial amygdala lies in direct apposition to the ventrolateral side of the optic tract. It is not clearly delimited as it merges rostrally with the AA area and laterally with the deep layer of the ACO nucleus. More caudally, the nucleus becomes more sharply defined. At midlevels the Me nucleus is separated from the medial division of the Ce nucleus, located dorsolaterally to the medial amygdala, and by a cellpoor zone, which is replaced more ventrally and caudally by the BSTIA. At caudal levels, the sagittal limb of the main group of the IC nuclei separates the Me nucleus from the LBNC laterally. Caudally, bands of fibers ascending into the stria terminalis are interposed between the medial amygdala and the other amygdaloid nuclei (Figs. 22, 23C, 23D, 24B, 24D, 24F, 26–30, 31D, and 33A). Cytoarchitectonics The Me nucleus consists almost entirely of small- and medium-sized lightly stained cells of various shapes (Figs. 15G–15J in Alheid et al., 1995). It also contains some large cells, which are more abundant toward the rostrodorsal aspect of the nucleus. A cytoarchitectonical analysis reveals that the Me nucleus is subdivided into a principal part or body, a small anteroventral part with densely packed medium-sized cells (MeAV) (Fig. 16A), and a posterodorsal part (MePD) (Figs. 17A and 24B) at the caudal end of the nucleus. The principal part of the nucleus can be subdivided further into a mixed-cell anterodorsal part (MeAD) (Figs. 17A, 22A, 23C, and 24B) and a posteroventral part (MePV) (Fig. 16A) with isomorphic cells. In the posterodorsal part of the Me nucleus, particularly toward the ventral aspect, cells aggregate into three columns oriented roughly parallel to the lateral surface of the nucleus. The first is a superficial or medial column (MePDm) made up of small- to medium-sized, tightly packed cells, with a second deeper and more lateral column of medium-sized cells (MePDl, Fig. 24B) that are densely packed but less than those in the medial superficial column. A third, more loosely arranged, intermediate column of medium-sized cells (MePDi) forms a matrix encompassing and separating the other two columns. The cellular part of the Me nucleus is surrounded on its ventral and medial sides by a molecular layer that becomes narrow dorsorostrally until it disappears completely at the level of the anterodorsal subdivision of the Me nucleus. Fibroarchitectonics In silver-stained preparations (Fig. 13E), the Me nucleus shows fibroarchitectonical

559

variations harmonizing closely with the cytoarchitectural features described above. Thus the MePD subdivision stands out by the thinness of its fiber plexus which contrasts with the richness and heavier staining of the fiber plexus in the MeAD subdivision and the more complex fiber arrangements in the MeAV and MePV subdivisions. The molecular layer (L1) in the Me nucleus is occupied by transversely and sagittally running fibers which fill it almost entirely except for a narrow, fiber-free peripheral rim. Rostrally and dorsally, the dense staining sagittally running fibers in L1 form a bundle that spreads out like a fan within the LPO area and the horizontal limb of the diagonal band of Broca. At the level of the MePD a stream of loosely arranged fibers intervenes between columns of cell aggregates in its course toward the stria terminalis. Heavy metals Alternate sections stained with the Timm/Danscher’s procedure (Figs. 16B, 17C, 18B, 21C, and 28B) show a staining pattern in the Me nucleus more complex than the subdivisions mentioned earlier suggest. The Timm/Danscher’s pattern at the level of the molecular layer of the Me nucleus helps to differentiate it from the ACO and PLCO nuclei. Thus, the Me nucleus is covered by a wide poorly stained subpial lamina (1A) that starts sharply at its border with the ACO nucleus and PLCO nucleus. The remaining width of the Me nucleus molecular layer (lamina 1B) is homogeneously and densely impregnated, and this also contrasts with the bilaminated and generally weaker staining pattern of lamina 1B of the ACO nucleus and PLCO nucleus. In the PDMe subnucleus, the two columnar cell aggregations (MePDm and MePDl) show a dense Timm/Danscher’s reaction, but this reaction is significantly weaker than the very dense reaction in the surrounding neuropil (Figs. 26A, 29B, and 30B) of the intermediate part (MePDi). Another differential Timm/ Danscher’s stain reaction is appreciable at the level of the MeAV subnucleus, where superficial cell L2 is poorly to lightly covered with silver precipitate, whereas deep cell L3 displays a very dense reaction. The MeAD also stains very densely with the Timm/Danscher’s reaction, but this staining is still less dense than the even darker staining in the MePV subnucleus and intermediate column of the MePD subnucleus. Considered as a single entity, the Me nucleus can be differentiated from the ACO nucleus and BSTIA division by its stronger Timm/Danscher’s reaction and from the PLCO nucleus, PMCO nucleus, AHI area, and main IC nucleus by its weaker staining reaction. Acetylcholinesterase In AChE preparations (Figs. 17B, 18B, 22C, 26B, 26C, 29A, and 30A), the Me nucleus displays a light to poor reaction, especially in its posterior

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subnuclei. An exception to this is the consistent, although moderate, staining reaction throughout its molecular layer (Figs. 26B and 26C). This laminar reactivity starts ventrally just at the border of the Me nucleus with the ACO nucleus and the PLCO nucleus and constitutes an easily distinguished landmark for differentiating the Me nucleus from both of them. At the level of the anterodorsal subdivision of the Me nucleus, where the molecular layer disappears, the stained band becomes reduced to a thin but darkly staining rim. This rim outlines the lateral margin of the optic tract before blending with the very highly AChE-positive neuropil in the sublenticular field of the magnocellular cholinergic projection neurons. Choline acetyltransferase According to Hellendall et al. (1986), the Me nucleus is remarkable because it has the lowest ChAT activity in the amygdala, less than half that in the neocortex and less than 5% of that in the anterior division of the BL nucleus. According to Heckers and Mesulam (1994), the ventral parts of the Me nucleus display a low to medium density of ChATIR fibers while the dorsal parts of this nucleus are almost totally devoid of these types of fibers. The Intraamygdaloid Bed Nucleus of the Sria Terminalis (BSTIA) Topographic landamarks The BSTIA division of the BST nucleus (Figs. 13E, 22, 24B, 24D, 24F, 27C, 27D, 29, and 30) is a relatively cell-poor area located lateral to the dorsal part of the Me nucleus and is traversed by fibers destined to the stria terminalis. Cytoarchitectonics The BSTIA nucleus cells are medium sized, but are larger and more lightly stained than those of the Me nucleus (Figs. 15I–15J´ in Alheid et al., 1995; also Figs. 22A and 24B of present chapter). Fibroarchitectonics In neurofibrillar preparations, no particular arrangement of fibers can be appreciated other than passing axons (Fig. 13E). Heavy metals In Timm/Danscher-stained sections the BSTIA division of the BST nucleus (Figs. 22B, 24F, and 30B) appears as an area with weak to moderate silver precipitate compared with the darker Me nucleus and the very dense precipitate in the main IC mass, whose sagittal limb invades the BSTIA division. Acetylcholinesterase In AChE-stained sections (Figs. 22C, 24D, and 30A), the BSTIA division of the BST nucleus has a light to moderate reaction compared with that of the surrounding neuropil, except for that of the BL nucleus which is the strongest.

Choline acetyltransferase According to Heckers and Mesulam (1994), this area, lying medially to the Ce nucleus displays a thick bundle of ChAT-IR fibers which according to their illustrations (their Fig. 2) curves ventrolaterally to become embedded in the rich ChATIR fiber plexus present in the BL nucleus. The Medial Sublenticular Extended Amygdala (SLEAm) Topographic landmarks The medial portions of the SLEA (Figs. 6, 7C, 8–11, 13D, 14, 15, 19, 21, 25, 29, and 34) is a subdivision of the territory that has been previously referred to as the posterior, or sublenticular, substantia innominata (Figs. 15D and 15E in Alheid et al., 1995). The anterior, or subcommissural, part of the substantia innominata has been more or less completely identified with the ventral extension of the globus pallidus mostly below the anterior commissure, i.e., the ventral pallidum VP in Figs. 6A, 15A. The area known as the sublenticular substantia innominata includes several corridors of cells that are associated with the continuum stretching from the centromedial amygdala to the BST nucleus, as well as the more difficult to categorize basal forebrain complex of cholinergic cells. Owing to the fact that, together, the ventral pallidum, the central and medial divisions of the extended amygdala, and the cholinergic projection neurons occupy practically the entire field designated by the term substantia innominata, it can be argued that this term may be superfluous (Alheid and Heimer, 1988) and that its persistence is mainly supported by the inconvenience and sometimes real difficulties associated with applying a more anatomically accurate identification of these territories. With respect to the medial division of the extended amygdala (MEXA), it is clear that some of the neurons traversing the sublenticular zone are associated with this complex, rather than with the central division of the extended amygdala (CEXA) or for that matter with the cholinergic complex. However, positive cytoarchitectonical identification of the cells belonging to the medial sublenticular division of the extended amygdala is difficult, given most normal histochemical procedures available. However, this identification may be partly accomplished by exclusion. For example, immunohistochemical methods for angiotensin II stain the entire sublenticular extended amygdala, whereas the cupric silver normal stain (i.e., adjusted to stain normal elements of the extended amygdala) is more or less confined to the central division of the extended amygdala (Figs. 6 and 7). The medial part of the sublenticular extended amygdala, therefore, would be the cell territory that is relatively poor in silver deposited by the normal cupric silver method, but still relatively

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rich in angiotensin II immunoreactivity. Another alternative is to make use of the fact that the medial division of the extended amygdala is relatively poorly stained for tyrosine hydroxylase immunoreactivity compared with the adjacent central division or the medially adjacent lateral hypothalamus, so that the posteroventral sublenticular band that is essentially free of tyrosine hydroxylase terminal staining corresponds more or less to the medial division of the SLEA (Fig. 25). The column of neurons associated with the medial SLEA can be also identified hodologically, when cells from targets also innervated by the Me nucleus or BSTM division are retrogradely labeled or when sublenticular terminal fields from brain areas that specifically target the medial portions of the EXA are labeled (Grove, 1988a, 1988b). In the following paragraphs, we describe the general appearance of the SLEA, with an emphasis on its medial division, and return to this topic later as we discuss the sublenticular parts of the central division of the extended amygdala. Cytoarchitectonics The medial SLEA in the rat is a relatively homogeneous collection or palely staining, medium-sized, spindle-shaped neurons that are dispersed throughout a diagonally oriented arch occupying the gray area just dorsal to the inferior thalamic peduncle (Figs. 15D and 15E in Alheid et al., 1995). It is bounded by the globus pallidus dorsally and to some degree by the interspersed aggregates of large cholinergic neurons, which form part of the cholinergic projection system. At its rostromedial pole, it expands to merge with the BSTMP subdivisions, whereas at its caudolateral pole it arches downward behind the magnocellular neurons of the nucleus of the horizontal limb of the diagonal band in order to merge with the cells in the anterodorsal portions of the Me nucleus ventrally. The majority of the neurons making up the SLEA have their longer axes arranged parallel to its general diagonal orientation (Figs. 13D, 14A,B, 15C, 21A,B, 25). However, particularly along its border with the globus pallidus and related structures, larger, dark-staining multipolar cells, singly or grouped in loosely packed clusters, that do not obey this orientation pattern can be observed. The presence of these cells also suggests the subdivision of the sublenticular extended amydala into anterodorsal and posteroventral portions (corresponding to the central and medial divisions of sublenticular extended amygdala, respectively) (Fig. 19). Fibroarchitectonics In neurofibrillar preparations, the general orientation of the cellular components corresponds with that of the fiber system in which they are embedded, with the most conspicuous contribution

561

coming from the longitudinal association bundle (lab) and from the ansa peduncularis. The ventral and medial sectors of the basal forebrain region, encompassed by the SLEAm, are broken up by fiber packages of the medial forebrain bundle interfering although partially with the course of the above-mentioned fiber systems. By doing so, the two above-mentioned fiber systems come to fill the celled, mostly diagonally oriented, arched corridors extending between the more dorsally located medial forebrain bundle before reaching the ventrolateral aspects of the BST nucleus (Figs. 13C and 13D). Heavy metals In Timm’s-stained preparations, the SLEAm (Figs. 14A, 21B, 21C, and 29B) can be differentiated from some of its prominent neighboring structures by a consistent, light to moderate, granular staining reaction along its full extent. This staining fuses gradually with that resulting from a considerably stronger reaction produced in the BST nucleus rostrally and in the Ce nucleus, Me nucleus, BSTIA, and AA area caudally. This positive staining reaction contrasts markedly with the almost total absence of silver deposits in the horizontal limb of the diagonal band and lateral preoptic area, on one side, and with the patchy staining reaction characteristic of the globus pallidus and ventral pallidus, on the other (Haug, 1973; Ottersen, 1980). In the selenite version of Timm’s stain (Danscher, 1981), the contrast is even greater, because the patchy staining produced by the Timm’s sulfide method within the globus pallidus and ventral pallidus does not occur (Fig. 14A, 24B,C). Comparison of the medial division of the SLEA with the central division indicates that the former stains somewhat darker in the Timm’s preparations. Acetylcholinesterase In AChE-stained sections, the SLEA (Fig. 29A) appears as a passageway exhibiting a light to moderate staining reaction, which contrasts with the heavier AChE activity in the horizontal limb of the diagonal band, lateral preoptic–lateral hypothalamic continuum, and accompanying forebrain cholinergic neurons. The moderate AChE staining in the SLEA is confined to the neuropil that surrounds negatively reacting perikarya; except that in the anterodorsal or central division of SLEA it is possible to detect the presence of scattered, strongly AChE-positive (Johnston et al., 1979; Lehman et al., 1980; Parent, 1979) or ChATpositive (Mesulam et al., 1983; Rye et al., 1984), larger multipolar neurons. The Medial Division of the Supracapsular Bed Nucleus of the Stria Terminalis (BSTSm) The supracapsular part of the BST nucleus is a term that—for the rat—encompasses interrupted cell columns accompanying the stria terminalis as it traverses the

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dorsal part of the internal capsule (ic). As with the SLEA, the BSTS division of the BST nucleus includes elements of both the medial and the central divisions of the EXA. Incidental observations, made in the course of other retrograde labeling experiments, indicate that there is also a medial to lateral segregation of the cells associated with the medial and central divisions of the extended amygdala, respectively. As with the SLEA, a general description of this territory is given here, with a return to specific comments relative to the central division of the extended amygdala further below. Topographic landmarks and cytoarchitectonics The neurons of the BSTS division of the BST in some mammals may form a continuous column following the dorsal course of the stria terminalis (Johnston, 1923; Strenge et al., 1977; Alheid et al., 1994). In the rat, however, intermittent cell pockets may be observed along the course of this tract. These neurons are fusiform and oriented parallel to the axons of the stria terminalis. Although two larger pockets of cells (one medial and one lateral) are found in continuity with the caudodorsal tip of the BST (Geeraedts et al., 1990), such aggregations become scarce in the retrocapsular portion of the stria terminalis, but increase in number as the stria enters the amygdala. With respect to the two divisions of the EXA, it appears that the neurons related to the central division may be more numerous and are located in fiber-free pockets at the lateral edge of the stria terminalis (e.g., Fig. 20). Cells related to the medial division of the EXA are found embedded within the medial part of the stria terminalis and are most readily identified by retrograde labeling from medial hypothalamic targets.

The Central Division of the Extended Amygdala (CEXA) The nuclei that compose the central division of the extended amygdala owe this designation to their overall similarity to the central nucleus of the amygdala. In this instance, the various subnuclei are characterized by their close connections with the lateral, rather than the medial, hypothalamus and with the caudal brainstem nuclei in both the dorsal and the ventral tegmenta. This is particularly evident in the reciprocal connections of the central division with the parabrachial nucleus and with the neurons in the solitary nuclear complex. However, reciprocal projections to the retrorubral area as well as descending projections to the pontine reticular formation may also be typical of the central division of the extended amygdala, but at this point are less clearly defined for the entire continuum than for the central amygdala itself. As with the medial divi-

sion of the extended amygdala, the central division is characterized by an extensive network of intrinsic connections, and this as much as any other feature serves to distinguish it from the nearby and often overlying striatal territories, because the latter appear to adhere much more strictly to a vertical or columnar scheme of connections, in which a few long, horizontal intrinsic connections are found. Of the deep cortical-like nuclei, some of them seem to favor the central division of the extended amygdala as a corridor for their descending projections. These include the BL nucleus and particularly its posterior part, in addition to the BLV nucleus. The central division of the extended amygdala defines a particularly complex macrostructure when its many subdivisions are considered. These include components stretching just beneath the globus pallidus through the central division of the SLEA (Fig. 6), but also additional columns of neurons that more or less accompany the posterior limb of the anterior commissure as it curves rostrally from the temporal lobe to the point at which it is straddled by the BST. As with our description of the medial division of extended amygdala, we begin with the central division’s more rostral counterpart in the BST. The Lateral Bed Nucleus of the Stria Terminalis (BSTL) The BSTL division of the BST nucleus in general forms an irregular truncated pyramid, with its base apposed to the medial side of the internal capsule, which forms its lateral boundary (Figs. 15B and 15C in Alheid et al., 1995). Rostrally, it adjoins the posterior aspect of the nucleus accumbens ventrally and the caudate–putamen dorsally. The medial boundary of the BSTL nucleus is marked by the compact fibers of the stria terminals. It rests atop and around the anterior commissure and extends slightly rostrally along its anterior limb. Caudally, it dips ventrally to surround the posterior limb of the anterior commissure (Figs. 6, 7, 11, 13–15, 19, and 24). Moving from rostrodorsal to caudoventral, the BSTL nucleus tapers somewhat until it fuses with the rostromedial end of the SLEA. In general, cells in the BSTL nucleus are somewhat larger and more loosely packed than those in the BSTM nucleus (Figs. 31A, 31C, 32, 33B, and 34A). Cytoarchitectonics Based on recognizable cytoarchitectural and histochemical features, the lateral bed nucleus may be readily divided into five subdivisions: dorsal, posterior, ventral, intermediate, and juxtacapsular (Figs. 15B–15C´ in Alheid et. al., 1995; Figs. 31A, and 31C, this chapter). The dorsal part of the lateral bed nucleus (BSTLD) The dorsal part of the lateral bed nucleus is composed of pale-staining, rounded, medium-sized cells and is

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relatively homogeneous in appearance. It has a teardrop shape, whose long axis is oriented along the course of the stria terminalis dorsocaudally to rostroventrally, and because of this it resembles a double convex lens in frontal and horizontal sections (Figs. 15B and 31B). On histochemical grounds this may be further subdivided into (i) capsular (BSTLDC) (Fig. 24A) and (ii) central (BSTLDcn) parts, with the capsular portion demonstrating a richer substance P terminal field, but much poorer tyrosine hydroxylase terminals (Fig. 31A). The posterior part of the lateral bed nucleus (BSTLP) The posterior part of the lateral bed nucleus (BSTLP) accounts for the majority of the cells in this complex. It surrounds the ventral and caudal aspects of the dorsal part, with its main mass found posterior to the dorsal part of the lateral bed nucleus, where it replaces the dorsal part, but also extends ventrally and laterally, where it merges with the anterodorsal or central division of the SLEA (Figs. 19 and 24A). In Nissl-stained sections (Figs. 15B and 15B´ in Alheid et al., 1995), the cell composition of the posterior lateral bed nucleus resembles that of the dorsal lateral bed nucleus, except that it is somewhat more heterogeneous, containing smaller elongated cells that are more closely packed. The ventral part of the lateral bed nucleus (BSTLV) The ventral part of the lateral bed nucleus, as its name implies, occupies the ventrolateral quadrant of the lateral bed nucleus, starting just below the anterior commissure (Figs. 15B–15C in Alheid et al., 1995) and extending caudally until it merges with the posterior part of the lateral bed nucleus. It is continuous on its lateral edge with extended amygdaloid neurons, which envelop the posterior limb of the anterior commissure in the territory we have termed the interstitial nucleus of the posterior limb of the anterior commissure (IPAC). The ventral part of the lateral bed nucleus is composed of a mixed population of neurons, including many that are larger, more angular, and more darkly stained than those in the dorsal part. The intermediate part of the lateral bed nucleus (BSTLI) The intermediate part of the lateral bed nucleus of the stria terminalis is composed of predominantly angular, dark-staining, large (15–20 m) neurons (the largest of all neurons in the BST nucleus), which are interposed between the remaining subnuclei of the BSTL division and the BSTMP division of the bed nucleus. The cells form a narrow wedge in sagittal sections, but appear similar to a flattened crescent in horizontal sections (Figs. 15C and 15C´ in Alheid et al., 1995). The long axes of these cells are oriented in an oblique angle, which is

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parallel to fibers traversing the BSTI nucleus in a lateroventral to mediodorsal direction. The BSTI nucleus extends dorsally until it reaches the dark-staining cell groups that accompany the cst (that is, the bed nucleus of the commissural component of the stria terminalis). Ventrally, the BSTI merges with the lateral preoptic area, whereas laterally it merges with the SLEA (Fig. 24A). The juxtacapsular part of the lateral bed nucleus (BSTLJ) The juxtacapsular part of the BSTL division is a small lens-shaped cell aggregate made up of small ovoid cells that fill a shallow concavity made by the medial surface of the internal capsule at the same plane as the crossing fibers of the anterior commissure. Its medial edge borders the BSTLDl and BSTLP subnuclei, but is distinguishable from these areas by its smaller and more densely packed cells (Fig. 24A). Cupric silver stain The normal cupric silver method is especially useful in differentiating several subdivisions of the BSTL (Figs. 6 and 7). The neuropils of both BSTLA and BSTLV subnuclei display an equally dense granular argyrophilia with the cupric silver technique. On the other hand, the BSTLJ is completely negative with the cupric silver technique, whereas the BSTLP subnucleus shows a considerably lighter granular argyrophilia than either the BSTLD or the BSTLV subnuclei. The BSTLI stains poorly, if at all, with the cupric silver method. A striking feature of the BSTSL subnucleus is the presence of neurons with granular argyrophilia (in somata and dendrites), which is not the case for the remaining divisions of the BSTL subnuclei. These neurons, however, appear to be identical to those in the central component of the lateral part of the Ce nucleus of the amygdala. The granular argyrophilia in the neuropil of the BSTLD subnucleus is continuous with that present in the anterolateral portion of the SLEA. Heavy metals In Timm/Danscher’s preparations, the BSTLD, the BSTLJ, and the BSTLP subnuclei are all very densely stained (Figs. 14A and 24E). Of these, the BSTLP subnucleus tends to stain more lightly than the BSTLD subnucleus, and the BSTLJ subnucleus appears to be somewhat darker. The BSTLV subnucleus stains densely; this staining is generally lighter than that in the preceding divisions. The Timm/Danscher’s reaction in the BSTLI is only moderate and distinguishes this part from the BSTLP subnucleus and from the BSTM division nuclei. The Timm/Dancher’s stain in the BSTL division appears to be dependent on the integrity of the stria terminalis (de Olmos et al., 1985; PerezClausell et al., 1990) and on that of the BLP subnucleus (observations by J. S. de Olmos and C. A. Beltramino cited in de Olmos et al., 1985).

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Acetylcholinesterase In AChE-stained sections, the BSTLD subnucleus stands out because of its light to poor staining in its capsular subdivision, in contrast to the slightly darker but still light staining in the BSTLP subnucleus and the very dense staining in the juxtacapsular part (Fig. 24C), although this staining is not as dark as that in the caudate–putamen. The BSTLV subnucleus is lightly stained for AChE, but is characteristically surrounded ventrally and laterally by an AChE-rich capsule. The BSTI division of the BST nucleus is also lightly stained to about the same degree as that in the BSTLP subnucleus and in the central component of the BSTLD subnucleus. The Central Amygdaloid Nucleus (CeA) The central amygdaloid nucleus (Ce) is an ovoid mass of cells in the dorsal central part of the amygdaloid complex, bordered dorsolaterally by the striatum and dorsomedially by the caudal extension of the network of cholinergic projection neurons (this part is also known as the interstitial nucleus of the ansa peduncularis), which separates it from the globus pallidus. At rostral levels, the Ce nucleus is flanked by the LA and BL nuclei laterally and ventrally and by the transverse limb of the main intercalated cell mass ventrally, which intervenes between the central nucleus and the BMA subnucleus (Figs. 15G–15J in Alheid et al., 1995). Farther caudally, the BSTIA becomes interposed between the Ce nucleus and the caudal limb of the main intercalated cell mass. At its rostral end, the Ce nucleus lacks clear boundaries as it blends with its sublenticular extension and with the AA area. Caudally, a cell-poor lamina and the small islands of intercalated cells practically separate the Ce nucleus from the anterior wall of the temporal horn of the lateral ventricle. The traditional two major divisions of the central nucleus, the medial and lateral, are heterogeneous enough in cell composition and in connectivity to be susceptible to further parcellations. Such subdivisions have been proposed on the basis of ontogeny (Bayer, 1980), structure (McDonald, 1982a), histochemistry (de Olmos et al., 1985), and immunohistochemistry (Wray and Hoffman, 1983; Cassell et al., 1986). The intercalated cell groups, the AStr, and the IMG are gray formations that appear to have some association with the Ce nucleus (Figs. 6, 7D, 9, 10, 11, 13E, 15E, 18, 21C, 22, 24B, 24D, 24F, 27, 28, 30, 31B, 31D, and 33A). The medial part of the central amygdaloid nucleus (CeM) Topographic landmarks The medial part of the central nucleus characteristically envelops the upper segments of the intraamygdaloid course of the nucleus of the commissural component of the stria terminalis (Figs. 6, 9, 7D, 13E, and 15E), and, in Nissl prepara-

tions (Figs. 15F–15I´ in Alheid et al., 1995), it can be differentiated from its lateral counterpart by its larger, more heterogeneous and darkly staining neurons. The borders of the CeM division with other neighboring grisea are, on the contrary, not well defined. This nucleus merges gradually with the AA area and sublenticular territories rostrally, with the nucleus of the ansa lenticularis dorsomedially and with the BSTIA division of the BST caudally (Figs. 21C, 22, 24B, 24D, 24E, 27C, 27D, 30, 31B, 31D, and 33A). Cytoarchitectonics On purely cytoarchitectural grounds, three further subdivisions (Figs. 15F–15I´ in Alheid et al., 1995) can be recognized in the medial central nucleus: (i) a comparatively narrow and largecelled anterodorsal part (CeMad) that occupies the rostral pole of the medial central nucleus; (ii) a prominent dense-celled anteroventral part (CeMav) located immediately caudal to the large-celled part of the CeM division; and (iii) a heterogeneous, loose-celled posteroventral part (CeMpv), which replaces the more anterior portions of the CeM division caudally and ventrally. Fibroarchitectonics In neuroibrillary or myelinstained preparations, a discontinuous fence made up of small fiber bundles running dorsally toward the boundary line between the CeM and the CeL divisions can be detected (Fig. 13E). The existence of a rich fiber plexus in the CeM divisions and the almost complete absence of it in the CeL constitutes a major differentiating landmark between these two divisions. However, this characteristic is not as useful in separating the CeM division from some of its neighbors, as the CL division of Ce, and the BSTIA division of the BST. As can be seen below, this distinction is better achieved in cupric silver (Cu–Ag)- or Timm’s/Danscher-stained sections. Cupric silver stain The differentiation between the CeM and CeL divisions is better achieved in cupric silver (de Olmos, 1969, 1972; de Olmos et al., 1994) and Timm’s impregnated sections. In cupric silver preparations of normal (i.e., unlesioned) brains, the CeM division only shows scattered granular argyrophilia, a type of impregnation not seen in the AA area, in adjacent cholinergic cell groups, or in the bulk of the BSTIA division of the BST nucleus, but which appears to be similar to that in the sublenticular areas rostrally (in the anterodorsal SLEA; see Figs. 6 and 7). As an exception, the large-celled part of the CeM division appears to be practically free of any silver deposits (Figs. 6 and 7D). Heavy metals With the Timm/Danscher’s reaction the CeM division is moderately stained in the anterodorsal part, densely stained in the anteroventral

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part, and very densely stained in the posteroventral part. The medial division of the Ce nucleus is generally darker and more homogenous than the AA area, the adjacent cholinergic neurons, the BSTIA division of the BST nucleus, and the SLEA. It is this division of the Ce nucleus that forms on topographical basis the caudal end of the central division of the SLEA (Figs. 18B, 22B, 21C, 24F, and 30B). The lateral part of the central amygdaloid nucleus (CeL) Topographic landmarks The lateral part of the central amygdaloid nucleus extends along the entire anteroposterior length of the central nucleus, forming a relatively homogeneous ovoid mass of medium-sized cells with appearance and chromophilia similar to those of the adjacent striatum (Figs. 15F–15J in Alheid et al., 1995). Within the CeL division two main subdivisions can be recognized: (i) central (CeL) and (ii) capsular (CeC). The central part of the CeL division appears in the caudal two-thirds of the Ce nucleus (Figs. 6, 7D, 10, 11, 13E, 15F, and 18) as an ellipsoid cell condensation surrounded in a cup-like fashion by the lateral capsular part, which intervenes between the central part and the IMG rostrally, laterally, and caudolaterally, between the central part of the CeL division and the main ICM; and between the CeL and the BMA and BMP subnuclei of the basomedial amygdaloid gray aggregate ventrally. Medially, the CeM division of the Ce nucleus closes the cup. At its rostral end, this central core becomes further condensed into a spherical cell mass of reduced dimensions (around 400 m) (Figs. 21C, 22, 24B, 24D, 24F, 27, 28, 31B, 31D, and 33A). Cytoarchitectonics Neurons in the rostral condensation of the CeL subnucleus are not only more tightly packed but also significantly more basophilic than those in the main body of the central part of the CeL division (Figs. 15F–15J in Alheid et al., 1995). This cellular condensation, mentioned in our previous versions of this chapter, does not receive any designation in the Paxinos’s 1997 atlas. However, because of its configuration and chemoarchitectonical characteristics, this cell condensation has been considered a separate entity by McDonald (1982a), who named it as the central intermediate nucleus. This author has offered some evidence for the existence of a distinctive (although not exclusive) projection from his intermediate zone to the dorsal hypothalamus and the BST nucleus. The lateral capsular part of the CeL division (CeC), which is the largest of the two subdivisions of the CeL division of the Ce nucleus, is characterized by the loose arrangement of its cells. In a plane passing through the rostral border of the transverse limb of the main ICM,

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the CeC subnucleus splits into the anterior and posterior parts by the most rostral fascicles of the medially shifting fibers of the lab. This partition is clearly seen in sagittal sections stained by neurofibrillary or myelin procedures. The anterior portion of the CeC subnucleus stands out from the rest of the CeL division on the basis of its higher content of fibers. These intrinsic fibers are organized in close correspondence with the cell grouping occurring in the anterior CeC subnucleus (that is, a rostral anterior subgroup, made up of densely packed, deeply staining cells, and a caudal anterior subgroup morphologically similar to the rest of the CeC subnucleus). On the basis of Nissl and neurofibrillary preparations, the posterior part of the CeC subnucleus can be subdivided into dorsal and ventral parts, separated from one another by numerous bundles of fibers that mark the boundary between the CeL and the CeM divisions of the Ce nucleus farther dorsally. The posteroventral CeC subnucleus differs from its dorsal counterpart by the more diffuse arrangement of its cells and its more fibrous appearance (Figs. 22A and 24B). Fibroarchitectonics In fiber preparations, the bulk of the CeL division appears to be characteristically fiber poor and practically encapsulated by small bundles of fibers that separate it from its medial sibling and from the IMG and LBNC laterally and ventrally (Fig. 13E). Dorsally, these fibers mark a boundary between the CeL division and the overlying transition area between it and the striatum. Cupric silver stain In normal cupric silver preparations, the CeL division is characterized by a homogenous and dense granular argyrophilia (Figs. 6 and 7D). A distinction between the CeL and the CeM parts of the Ce nucleus is the occurrence of granular silver deposits within neurons for the CeL division. This is comparable to the situation in the lateral bed nucleus of the stria terminalis, where such cell staining occurs in the BSTLD subnucleus rather than in the ventral or posterior subdivisions of the BSTL division of the BST nucleus. Heavy metals In Timm/Danscher’s preparations (Figs. 21C, 22B, and 24F), both the CeC and the CeL subnuclei stain very densely. However, it is apparent that the CeL subnucleus stains somewhat more lightly than the adjacent capsular part. Acetylcholinesterase In AChE-stained material (Figs. 22C and 24B), while the CeM division exhibits light to dense staining depending upon the various subdivisions under consideration. The anterodorsal subdivision stains densely, as does the anteroventral part, whereas a weak reaction occurs in its posteroventral

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part. On the other hand, throughout the CeL division there is a relatively homogeneous, but poor to weak AChE reaction. The Central Sublenticular Extended Amygdala (SLEAc) As with the medial division of the extended amygdala, the central counterpart extends into the sublenticular zone (Fig. 6). The designation “central sublenticular extended amygdala” is used to distinguish this territory from the medial SLEA, from the network of cholinergic projection cells, and from the ventral pallidum, all of which have been encompassed by the somewhat magnetic term substantia innominata. The general Nissl, cholinesterase, Timm’s, and Golgi appearances of the central SLEA correspond to those described in this section we supplement that description with those elements that are distinctive for the central division of this structure. Topographic landmarks The central part of the SLEA (Figs. 15D and 15E in Alheid et al., 1995) emanates from the BSTLP part of the BST nucleus and runs in a dorsomedial to ventrolateral direction until it merges with the rostromedial quadrant of the Ce nucleus (Figs. 6, 7C, 9, 10, 11, 13C, 13D, and 14). Again, although both the medial and the central parts of the SLEA are immunoreactive for angiotensin and stained by the Timm’s method, only the Ce nucleus shows a significant and dense immunoreactivity for TH (Figs. 31 B,C). A part of this TH staining presumably includes adrenergic and noradrenergic terminals, which at the rostral and caudal ends of the SLEA merge with similar staining in the BSTLP subdivision of the BST nucleus and in the CeM subdivision of the Ce nucleus, respectively (Fig. 31) (Fallon and Ciofi, 1992). This suggests that a major portion of the central division of the SLEA is related to the CeM part of the Ce nucleus (Figs. 15D, 19, 21, and 25). Cytoarchitectonics The neurons that form the central sublenticular extended amygdala (SLEAc) are loosely packed, medium-sized, fusiform- to triangularshaped, and more elongated than those in the SLEAm. Like in the primates (Freedman and Shi, 2001), the rat SLEAc appears to be made up of a cell population that shows many morphological similarities with those of the cells present in the medial division of Ce nucleus. However, in the rat, perhaps more than in primates, the SLEAc seems to be more heteromorphic. Such an apparent heterogeneity is probably due to the interdigitation of the true SLEAc cell constituents with other basal forebrain neuronal subpopulations (Fig. 6).

Cupric silver stain In cupric silver preparations of normal brains, the anterodorsal (central division) of the SLEA exhibits a typical granular argyrophilia in its neuropil similar to and continuous with that seen in the Ce nucleus, caudally, and in the BSTL division of the BST nucleus, rostrally (Figs. 6 and 7C). The posteroventral (medial division) of the SLEA, in contrast, does not demonstrate this reaction. In the anterodorsal SLEA the granular argyrophilic reaction occurs in a perisomatic and peridendritic arrangement, which outlines the contours of the spindle-shaped neurons, but it does not appear to contain granular cells showing intracytoplasmatic granulations as do the CeL division of the Ce nucleus and the dorsal part of the BSTL (the BSTLD). Heavy metals On the basis of a careful analysis of all the data so far gathered by us concerning a histochemical and immunocytochemical characterization of the central division of the SLEA, it is possible to state that the dorsal sectors of the wedge-like Timm/ Danscher-positive granular material zone interposed between the inferior border of the internal capsule, on one side, and the inferior thalamic peduncle and dorsal aspects of the medial forebrain bundle, on the other, most probably are overlapped by the central division of the SLEA (Figs. 14A, 21B, 21C, and 29B). The Central Division of the Supracapsular Bed Nucleus of the Stria Terminalis (BSTSc) Topographic landmarks The portions of the BSTS division that are related to the CEXA are found exclusively in the fiber-poor pockets present in the lateral edge of the stria terminalis, and, concurrently with it, its retrograde labeling occurs only after injections that also label retrogradely the lateral BST nucleus, the central SLEA, and the Ce nucleus. This type of labeling also includes intrinsic connections among the various components of the CEXA. Furthermore, the supracapsular cells appear to be embedded within an anngiotensinII-rich neuropil with the same characteristics as those seen in the rest of the CEXA (Figs. 15D, 15E, and 20). Cytoarchitectonics In the rat, the neurons of the BSTSl are found in intermittent cell pockets distributed along the course of the stria terminalis. The neuronal population present in these pockets is made up of spindle-shaped neurons oriented parallel to the axons of the stria terminalis. They become scarcer in the retrocapsular portion of this bundle but increase in number again at the entrance of the stria terminalis to the amygdala (Fig. 20A). Cupric silver stain While these neurons are difficult to locate in sections stained by conventional histo-

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chemical procedures, many of the neurons in the BSTSl division exhibit a granular argyrophilia similar to that of the neurons in the BSTLD subdivision of the lateral BST nucleus and in the CeL division of the Ce nucleus. Like in these latter two nuclei the BSTSl cells are surrounded by an argyrophilia neuropil that varies between having either a rather homogenous granular appearance or showing a “rail-rod”: arrangement, a disposition presents particularly at the periphery of the nucleus. Heavy metals and acetylcholinesterase (AChE) In Timm/Dansher-stained sections, the BSTSl pockets appear filled with a dense nonhomogenous silver precipitate that is significatively more dense than that present in the immediately adjacent striatum. By contrast, although in AchE-stained sections these pockets also show a granular AChE reactivity, a reactivity that never comes close to that seen in the neighboring striatum. The Interstitial Nucleus of the Posterior Limb of the Anterior Commissure (IPAC) Topographic landmarks and cytoarchitectonics The interstitial nucleus of the posterior limb of the anterior commissure is a term that was earlier used to describe a narrow band of cells accompanying the posterior limb as it arches laterally, caudally, and ventrally toward the amygdala in the rat (Figs. 15A–15G in Alheid et al., 1995) (de Olmos, 1972; de Olmos and Ingram, 1972) and that seems to be the recipient of particular afferents from the amygdala, which were unusual in the context of the nearby basal forebrain. In the earlier version of this chapter (de Olmos et al., 1985), we included this territory within our discussion of the central division of the extended amygdala under both the “fundus striati” and the anterior amygdalostriatal zone and as part of the rostral column of IC clusters. At that time, however, it was difficult to ascertain the degree to which these structures could be related to the remainder of the central division of the amygdala. We have acquired a better appreciation of some of the complex relations of the area (Alheid and Heimer, 1988; Heimer et al., 1990, 1993; Heimer and Alheid, 1991; Alheid et al., 1994) and have gradually come to use the term “interstitial nucleus of the posterior limb of the anterior commissure” (IPAC) in an expanded sense. This applies to a broad core of neurons found above and below the posterior limb of the anterior commissure that seem to share histochemical features and connections that most resemble those of the central division of the extended amygdala. This, in fact, seems to constitute a lateral wing of this structure that is separated from the central division of the sublenticular extended amygdala, by virtue of the fact that it courses across

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the rostral and lateral face of the dorsal and ventral pallidum in the area just above and below the level of the posterior limb of the anterior commissure (Figs. 6A, 9, 14B, and 15). In one respect, the term amygdalostriatal transition zone (Astr) might still be appropriate for this territory, insofar as the cellular composition of this area and the dense staining for AChE make it difficult to define the boundary of this area with the adjacent ventral parts of the caudate–putamen and nucleus accumbens (Figs. 18A and 30A), and there is the likelihood that some basal ganglia elements are located within its borders. Few reports, however, have applied the term anterior amygdalostriatal zone to their descriptions of the rostral forebrain territory we are currently describing. The term, moreover, is firmly associated with the caudal area interposed between the Ce nucleus and the LBNC, portions of which appear to be of a distinctive character, more closely related to the striatum than to the area currently under discussion. Nor does the term “fundus striati” appear to be an appropriate designation for the rostral transition area, because it has been used to designate just that pocket of cells below the posterior limb of the anterior commissure, and it ignores the dominant relation of the IPAC nucleus with the amygdala, rather than with the striatum. Moreover, the term fundus striati has classically been used to refer to the rostral ventral portions of the caudate–putamen and nucleus accumbens in the human and monkey, a combination of territories that is not homologous with the topography of the area as it is currently designated in the rat. In contrast, the term IPAC nucleus, although lengthy, is appropriate on several grounds. First of all, it is historically accurate. As applied here, it is reasonably to designate the same area as that in its original usage. Second, it is topographically descriptive, because the area under consideration is more or less associated with the posterior limb throughout most of its course. Third, it is probable that the homologous territory will occupy approximately the same position in other species, such as the primate (Alheid et al., 1994). Fourth, the term is essentially neutral with respect to adjacent neural territories, precluding any unintended connotations about its functional–anatomical role in the forebrain (Figs. 7D, 8, 9, 11, 13E, 15, 18, 21B, 21C, 22, 25, 29, 30, and 34). The IPAC nucleus appears to encompass at least two, more or less continuous, columns of cells, one both medial and ventral and the other both lateral and dorsal to the first. The medial column includes medium-sized to small neurons encircling the posterior limb of the anterior commissure as it diverges from the anterior limb of this structure and runs posterolaterally in the forebrain (Figs. 15A–15G in Alheid et al., 1995). These cells take on a somewhat fusiform shape in the areas

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just above and below the anterior commissure as it crosses the rostral face of the pallidum (Fig. 9), whereas, laterally, these cells appear to be more rounded in frontal sections, but can be seen as fusiform in horizontal sections. On the basis of retrograde labeling of deep brainstem targets such as the solitary complex, the cells in this column drop below the anterior commissure laterally and form the medial surface of the pocket of gray matter that is ventral to the posterior limb of the anterior commissure. The medial column and the ventral part of the dorsolateral column of the IPAC nucleus together seem to compose this pocket of gray matter, which appears also to include pockets of the ICM.

nucleus and the solitary complex, whereas this does not seem to be the case for the medial, lateral, and caudal clusters of smaller neurons belonging to the IC groups. Thus, in our estimate, most reports of long projections from the IC groups (Moga and Gray, 1985a; Moga et al., 1990) appear to involve the medium and large neurons within the IPAC nucleus rather than any of the remaining small-celled groups identified with the IC masses.

The lateral part of the interstitial nucleus of the posterior limb of the anterior commissure (IPACl) The lateral (and dorsal) column of cells belonging to the IPAC nucleus is continuous across the posterior limb, and its dorsomedial part occupies a pocket adjacent to the dorsal pallidum just above the anterior commissure (Figs. 15D–15F in Alheid et al., 1995; and Fig. 14B, this chapter). On the basis of the evidence from tract-tracing and histochemistry studies, it appears that the medial column (see below) is continuous caudally with the CeL division of the Ce nucleus, whereas the dorsolateral column is continuous with the territory just dorsal to the CeL division, which has been previously identified with the anterior part of the Astr area (de Olmos et al., 1985) or identified as the dorsal component of the CeC subnucleus of the Ce nucleus of the amygdala (McDonald, 1992). Finally, it is possible that the dorsolateral column might be further divided into two columns on the basis of intrinsic connections with the EXA. The ventrolateral part of this column seems to have a greater number of cells projecting to the Ce or to the BST nucleus, whereas the dorsomedial sector of this column appears to have a greater concentration of cells projecting to the mesopontine tegmentum (Berendse et al., 1991; Alheid et al., 1994) (Fig. 3). However, at this time it is not possible to draw a boundary between these two sectors, and it appears that these two populations of cells overlap to a considerable degree.

Heavy metals In Timm/Danscher preparations, the IPAC nucleus is very densely stained and somewhat more densely than the adjacent striatum (Figs. 4, 29A, and 30B).

The medial part of the interstitial nucleus of the posterior limb of the anterior commissure (IPACm) The cells of the medial column have sometimes been included within the rostral chain of the ICM. However, many of the cells within the medial column of the IPAC nucleus are larger and less densely packed than the typical cells of the IC masses, although clusters of small neurons do occur within this column. Moreover, based on our incidental observations in tract-tracing experiments, it appears that the larger cells of the medial column project to deep brain-stem targets such as the parabrachial

Cupric silver stain With the normal cupric silver stain, the dorsolateral column of the IPAC nucleus is relatively free from stain, whereas the medial column is positive (Fig. 6).

Acetylcholinesterase (AChe) When moderate development of the AChE reaction is used, the IPAC nucleus is observable as a lighter area compared with the ventral caudate–putamen and the remainder of the nucleus accumbens, although it is still densely stained (Figs. 18, 29A, and 30A). It should be noted, in this context, that neither the connections of this territory nor its neurohistochemistry supports the likelihood that it is merely one of the AChE-poor striosomal compartments that are especially populous within the medial dorsal striatum.

Neurotransmitters and Neuromodulators within the Medial and Central Extended Amygdala Cells Already from a cytoarchitectonical point of view the EXA continuum appears as a very heterogeneous gray mass. However, the existence of certain common structural, connectional, morphofunctional, and histochemical features, which distinguish the neuronal pools that form part of it, have allowed us to differentiate, as already described, two major divisions, the MEXA and CEXA continua. Both of them, in turn, and according to the same morphological premises have been subdivided into subgroups whose temporal (amygdala), paraseptal (BST nucleus), and substriatal (IPAC nucleus) ends display features sufficiently similar to suggest the existence of a pairwise symmetry, a disposition which finds many times reinforcement in the cell composition of the cell bridges, sublenticular and supracapsular, that interconnect them. The complexity of the cell arrangement of the EXA continuum becomes even more difficult to interpret

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when the examiner is confronted with the vast heterogeneity of neuropeptides synthesized by the neurons that participate in the formation of the EXA continuum, or even that of each one of its successive subdivisions. To make such a task even more difficult, it seems now that at least some of the diverse neuronal phenotypes so far identified in the amygdalosublenticulostrial continuum undergo modifications that can be transient or become permanent depending on the strain; age; sex; and physiological, behavioral, or pathophysiological conditions of the animal under study. To illustrate this, and just with the perhaps incomplete data available to the present writers, the following neuropeptides have been identified in the rat EXA continuum: angiotensin II (ANG II), atrial natriuretic peptide (ANP), calcitonin gene-related peptide (CGRP), cholecystokinin (CCK), corticotropin-releasing factor (CRF), enkephalins (ENK), galanin (GAL), gastrin-releasing peptide (GRP), neurokinin B (NKB-P2), neuromedin B (NMB), neuropeptide Y (NPY), neurotensin (NT), pituitary adenylate cyclase activating polipeptide (PACAP), oxytocin (OT), secretoneurin (SN), somatostatin (SOM), substance P (SP), tyrotropin-releasing Factor (TRF), vasointestinal peptide (VIP), and vasopressin (VP) (Arluison et al., 1990; Asan, 1997; Bayer et al., 1991; Buijs et al., 1978; Cassell and Gray, 1989; Casell et al., 1999; Ciriello et al., 2003; Day et al., 1999; De Quidt and Emson, 1986; Eiden et al., 1985; Emson et al., 1978; Fallon and Leslie, 1986; Gray, 1989, 1990; Gray and Magnuson, 1987, 1990; Gray et al., 1989; Gustafson et al., 1986; Hallbeck et al., 1999; Harrigan et al., 1994; Higgins and Schwaber, 1983; Ju and Swanson, 1989b; Kozicz et al., 1998; Larsson et al., 1976; Lind et al., 1985; Ljungdahl et al., 1978; Lorén et al., 1979; Lucas et al., 1992; Marksteiner et al., 1992; McDonald, 1987; Micevych, 1988; Moga and Gray, 1985b; Moga et al., 1989; Piggins et al.,1996; Rao et al., 1987; Riche et al., 1990; Roberts, 1992; Roberts et al., 1982; Sakanaka et al., 1981, 1986; Shimada et al., 1989, 1992; Sofroniew, 1985; Veening et al., 1984; Veinante et al., 1997; Woodhams et al., 1983; Wray and Hoffman, 1983). To this long list of neuropeptides (20) which, in many cases, are colocalized in some of the neuronal pools making up the CEXA subcontinuum, the presence in some of them of at least two classical neurotransmitters, one inhibitory, γ-aminobutyric acid (GABA), and the other excitatory, glutamic/aspartic amino acid (GLU/ASP), should be added. Furthermore, and as an example of the transient expression of a monoaminergic neurotransmitter, neurons synthesizing both tyrosine hydroxylase (TH) (Fig. 31) and dopamine (DA) have been identified at the level of the Ce nucleus and the BSTI subdivision of the BST nucleus, while no data are available concerning the cytochemical identity of the “granular argyrophilic

569

neurons” present in the CeL division of the Ce nucleus, and in the BSTLD and BSTSl divisions of the BST nucleus, a cupric-silver histochemical reaction which reinstated the soundness of the morphological theory about the existence of an anatomical continuity between all these amygdaloid, strial, and paraseptal forebrain structures proposed by Johnston (1923) 80 years ago. According to the findings reported by several research groups (Mugnaini and Oertel, 1985; Christie et al., 1987; Nitecka and Ben-Ari, 1987; Sun and Cassell, 1993) GABAergic cells are present all along both major divisions of the EXA continuum, but with a majority of them being located apparently in the CeL and BSTLD subnuclei of the CEXA subcontinuum and a much lesser proportion located in the CeM and BSTLP. Moreover, a large amount of the EXA GABAergic neurons would be involved in establishing intrinsic connections within the EXA continuum which can be of short (strictly local), intermediate (just linking neighboring amygdaloid, or strial subnuclei), or long (interamygdalostrial or interamygdalo–IPAC, and vice versa) nature (Sun and Casell, 1993; Casell et al., 1999). The latter two appear to link subdivisions with similar cytochemical and connectional characteristics imitating their arrangements. Other EXA GABAergic neurons are extrinsic in nature; i.e., they belong to the category of long projection neurons targeting neuronal formations situated not only outside the EXA continuum but also outside the amygdaloid complex. Such would be the case of the long GABAergic projections to the nucleus accumbens (Christie et al., 1987), to the region of the paraventricular hypothalamic nucleus (Roland and Sawchenko, 1993), to the subretrofacial nucleus (Takayama and Miura, 1991), to the nucleus of the solitary tract (NTS) (Pickel et al., 1995; Saha et al., 2000), to the ventrolateral medulla (Jia et al., 1997), or to the dorsal vagal motor neurons (Takeuchi et al., 1983). Other targets of the projections that arise in either major division of the EXA continuum appear to be non-GABAergic but rather purely peptidergic such as would be the case for the CEXA projections to the parabrachial nuclear complex (Sun and Cassell, 1993; Cassell et al., 1999). However, in situ hybridization studies in the monkey (Pitkannen and Amaral, 1994) indicate that the number of GABAergic neurons may be greater than suggested by immunohistochemical procedures, raising the possibility that many of the long projecting GABAergic neurons in the EXA continuum may escape detection on account of a very fast turnover of the neurotransmitter. Finally, it should be pointed out that a considerable number of the GABAergic cells has been found to colocalize with CCK, NPY, SOM, and SP (McDonald, et al., 1995; Shimada et al., 1989), while at the level of the CeL– BSTSl–BSTLD subcontinuum, every CRF-, ENK-,

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and NT-containing cell would be GABAergic in nature (Day et al., 1999). As shown in Table 1, the distribution of neuropeptides in the EXA continuum follows different patterns, with few exceptions, it may involve both of its major divisions, or just one of them, or even one semi-ring or subcontinuum. Fiber Networks Concerning the type of neurotransmitter or neuromodulators contained by the fiber networks that are present in the EXA continuum, and regardless of whether their origin is intrinsic, i.e., issued by the neuronal pools present there, or extrinsic, that is, issued by neuronal aggregates situated outside this gray continuum, it is now possible to ascertain that such networks may contain and release through their axon terminals not only classical neurotransmitters, such as GLU/ASP, GABA (Fig. 28A), acetylcholine (ACh), monoamines, or heavy metals, that interact with their corresponding receptors but also different types of neuropeptides. These neuropeptides may be, or not, colocalized with some of the above-mentioned neurotransmitters, or with other neuropeptides, presumably acting as neuromodulators of the release of the above-listed neurotransmitters or interacting with each other. On the basis of their origin, the fiber plexuses found in the EXA continuum can be classified as properly intrinsic, intraamygdaloid and extraamygdaloid, or extrinsic. As their name implies the intrinsic plexuses in the EXA continuum are made up by the axons issued by intrinsic neurons of the extended amygdala, i.e., short and semi-short axon neurons whose projections remain confined to the continuum or even more strictly within the confinements of a given component of it. The fiber networks of intraamygdaloid nature can be considered to represent fiber plexi formed exclusively by axons of cells located in amygdaloid nuclei that do not belong to the EXA continuum. On the other hand, the plexi contributed by fiber systems of cortical and subcortical origin which reach and innervate the continuum can be considered extraamygdaloid or properly extrinsic. According to the data so far available in the literature, the intrinsic plexuses of the EXA continuum would be formed mainly by the axons of GABAergic (Fig. 28A), GABA/peptideergic, and purely peptidergic neurons whose cell bodies are located within the boundaries of this gray continuum. By contrast there is no clear-cut evidence that supports the existence of intrinsic GLU/ ASPergic neurons and, therefore, of properly intrinsically generated GLU/ASPergic plexuses. However, it is not possible to discard that terminal

ramifications of axon collaterals issued by long projecting putatively GLU/ASPergic neurons seemingly present in the EXA continuum may contribute to some extent to the fiber plexuses present in this continuum, since cells of this connectional type lacking such collateral branching have not been reported to be present in any of the many components of the EXA gray continuum (McDonald, 1983; de Olmos et al., 1985). On the other hand, although until today there was no data concerning the cytochemical basis for the so called “granular argyrophilic neuronal and neuropil” staining revealable with the cupric silver (Figs. 6 and 7) technique (de Olmos, 1969; de Olmos et al., 1994), unpublished lesion experiments and combined CuAg staining and retrograde tract-tracing results in our laboratory (Figs. 11, 15B, 15D, and 15F) suggest that these argyrophilic profiles that are detectable along the CEXA continuum correspond mostly to an intrinsic system of perikarya and axonal processes, even though a contribution from equally argyrophilic cells located in the dorsolateral hypothalamus seems to be present. As already mentioned previously, by using immunohistochemical procedures, it has been possible to identify a long list of putative neuropeptides in the neuropil of the EXA, such as Ang II (Figs. 8, 14B, 15A, 15C, 15E, and 28B), CGRP, CCK, CRF, ENK, GAL, NPY, NT, SOM, SP (Fig. 27), VIP, etc. If it is true that such a wealth of neuropeptides is by itself remarkable, it is even more outstanding the fact that many of the fiber systems containing them, whose sources reside either within and/or outside the extended amygdala, appear to adopt in their distribution within this continuum a pairwise symmetry pattern. Furthermore, it should be stressed that, as happens with the distribution within the extended amygdala of the perikarya of neurons synthesizing some of the above-mentioned neuropeptides, the peptidergic axons and terminals originating intrinsically and/or extrinsically in the EXA continuum are not evenly distributed throughout the continuum. Even so, they seem to parallel the pairwise symmetry displayed by the cytoarchitectonical organization of the EXA continuum. Thus, it is possible to verify that such patterns are also consistent with the completion of whole rings or of semi-rings of gray matter around the internal capsule, which taken conjunctly, constitute a subcortical gray cingulum, i.e., the EXA continuum. It is also important to point out, however, that the phenotypic expression of these neuropeptides seems to be very plastic and, for that reason, they may show variations in their density and/or distribution not only with age and sex but also while subjected to some persistent physiological and/or behavioral conditions. These variations, mostly transients, may well become consolidated, i.e., permanent, when

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they transpose certain limits and, as a result of this, a pathophysiological condition finally develops. Among the fiber plexuses of intraamygdaloid origin (see Tables 2–15), it seems that most of them would be of GLU/ASPergic nature, born mainly in neurons located in the posterior subdivisions of the LBNC or in the amygdalocortical transition areas (APir, Ahi, and CxP areas). A significant proportion of them would be apportioned by zinc-containing neurons which give rise to the Timm/Danscher’s silver reaction filling different components of the continuum with variable densities (Howell et al., 1991; Christensen and Frederickson, 1998; de Olmos and Beltramnino, unreported results) (see Figs. 33A and 33B). Furhermore, the apparent lack of participation in this type of projection from part of the remaining amygdaloid grisea and of practically all the gray components of both the MEXA and the CEXA would presumably be due to their complete lack of zinc-containing neurons or, if present, that they do not project to the EXA continuum but rather possess a projection field not related to the amygdaloid complex itself. Furthermore, concerning the distribution pattern of the zinc-rich neuropil present with variable magnitude throughout the EXA, there seems to exist a close correspondence between its density (see previous version of this chapter by Alheid et al., 1995) observable at both ends, amygdaloid and paraseptal, of the EXA continuum. Thus, as it can be appreciated in Figs. 6B, 7, the silver precipitate appears to be lighter in the MEXA subcontinuum than in the CEXA one. Concomitantly, within this latter subcontinuum, the MePD–BSTMPm pair contains less silver precipitate than the MeAD/ MePV–BSTMPi/l pair, whereas in the second subcontinuum, the CeL/CeC–BSTLD pair contains densities higher than those of the CeM– BSTLV/BSTLP pair. Finally, it seems quite plausible that the gradual thinning out and/or discontinuity displayed by the silver precipitates in both dorsal and ventral bridges (i.e., BSTS and SLEA, respectively) of the EXA continuum correspond to parallel thinning out and/or to discontinuations undergone by the neuronal pools representing either central and medial subcontinua caused by the passage of fiber through them. According to the bibliographic information available to the present writers, the greater part of the fiber plexuses of extraamygdaloid origin present in the EXA continuum is constituted by axons of neurons synthesizing classical neurotransmitters, such as GLU/ASP, ACh, DA (Figs. 25 and 31), NE, epinephrine (E), serotonin (5HT), and histamine (HA), while a much smaller part of them is apportioned by peptidergic neurons. The main sources of GLU/ASPergic axons would be of cortical origin, while the remaining neurotransmitters would be conveyed by fibers systems of subcortical origin.

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With respect to the afferent supply that the EXA continuum receives from the cerebral cortex, it should be pointed out that at least part of it originates from zinc-containing neurons, the majority of which are located in allo- and periallocortical (mesocortex) fields such as the temporal Pir paleocortex; the ventral S, the ventral ProS, and the caudal IL archicortices; and the L Ent, Cg, and PRh periallocortices (Howell et al., 1991; Christensen and Frederickson, 1998; de Olmos and Beltramino, unpublished observations) (Fig. 33A). As revealed by histochemical and immunohistochemical procedures, the cholinergic input reaching either major division of the rat EXA appears to be, in general, remarkably weak. However, as has been described in preceding sections of this chapter, there are some exceptions to this broad pattern. Such is the case of the IPACm, which displays a moderate to dense cholinergic innervation, or, though not so pronouncedly, of the CeM–BSTLP subcontinuum with which the IPACm nucleus shares many connectional features (Shammah-Lagnado et al., 2001). Moreover, it is also worth stressing the almost complete absence of a cholinergic input to the CeL–BSTLD/BSTLV subcontinuum, or to the MeAD/MePD–BSTMPi/m one. In all of these cases such particular disposition appears to go along with the pairwise symmetry that provides the basis for the concept of the extended amygdala. Concerning the source of this cholinergic input, the experimental data so far available indicate that its major source of such type of innervation is the magnocellular cholinergic neuronal complex of the basal forebrain, more particularly its anterolateral Ch4 group. Some of the cholinergic supply may also come from the pontine lateral tegmental nucleus. In this context it is interesting that part of this cholinergic projection arises from neurons containing also the p75 nerve growth factor (NGF) receptor providing a heterogeneous cholinergic supply to the EXA continuum which sharply contrasts with those directed to the BLA nucleus which do not contain such receptor (Heckers and Mesulam, 1994). The monoaminergic supply to the EXA continuum comes from the DA-, NE, E-, 5HT-, and HAergic cell groups distributed in the hypothalamus (DA 11 (Figs. 25 and 31); tuberomammillar HA) and in the mesencephalic (DA/A8, DA/A9, and DA/A10; 5-HT/B7), pontine (NE/A 5, 5-HT/B8, and 5-HT/B5), and medullary (NE/A2; E/C2) tegmentum (see Table 1). Finally, the extraamygdaloid peptidergic supply to the EXA continuum seems to be provided by diverse sources in the basal telencephalon, diencephalon, and brain-stem tegmentum. Notable among them are those coming from the OXC- and VP-synthesizing cells in the so-called hypothalamic neurosecretory nuclei, as well

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as those originating in the LHRF- and β-endorphinsynthesizing cells of the periventricular and infundibular arcuate hypothalamic nuclei, respectively; in CGRPcontaining neurons in the subparafascicular (SPFF) and parabrachial (PBR) nuclei, in VIP- and CCKproducing cells of the midbrain PAG matter; in NT/DAproducing cells in the mesencephalic VTA–SN region; in DYN-, ENK-, and SOM-containing neurons in the PBR nuclear complex; and, finally, in the DYN-, ENK, NPY- NT-, and SOM-containing neurons in the dorsal vagal complex.

Intraamygdaloid, Interamygdaloid and Extraamygdaloid Connections of the Medial and Central Extended Amygdala TABLE 5

THE LATEROBASAL NUCLEAR COMPLEX (LBNC) On the basis of cytoarchitectonic and neurochemical criteria, four major nuclei can be recognized as constituents of the LBNC: lateral (LA), basolateral (BL), ventral basolateral (BLV), and basomedial (BM). This heterogeneous assembly of neurons occupies approximately 48.8% of the total volume of the monkey temporal amygdala (Herzog, 1982). It is set off from other amygdaloid nuclei and from the interstitial nucleus of the posterior limb of the anterior commissure (IPAC) by the longitudinal association bundle that runs along the medial border of the LBNC complex. This bundle continues ventrally and tends to separate the medially

Intraamygdaloid Connections of the MEXA MEXA efferents to the

Intrinsic connections

OLF/VM-CORLN

CEXA

LBNC

The medial extended

MeAD → BSTMPl,

MEXA innervates

Several efferent

Sparce connections from

amygdala (Me) can be

BSTMPi, BSTMV

several nuclei of the

pathways from the

the MEXA to the LBNC

MEXA to the CEXA

except for the caudal and

divided into:

possibly BSTIA, and

olfactory amygdala:

anteroventral MeAV),

caudal pole of

AA, ACO, PLCO,

anterodorsal (MeAD),

BSTMPm, but not

AHi and APIR

MeAD → BSTLP and

posteroventral (MePV),

to BSTMA

BSTLV and CEM AD

MeAV → posterior

MeAD → AAD, AAVm

and PV

part of the LA

and posterodorsal (MePD) parts.

MePV→ME AD,

(vd), AA Vl (m), ACO,

MePD is subdivided

MeAV and BAOT

APIR AL (vd), APIR PM

into medial column

MePV →

MePDm), lateral

MePV → caudal end of BSTMA, BSTMPl,

and rostral almost none

BSTIA → CeM (vd),

intermediate column

caudal end of BSTMPm

MePV → AAD, AAV (l),

and BSTMV but

ACO (l/m), Me PD (l/m)

not BSTMPi

APIR AL (m)

The medial bed nucleus

MePD → Me

BSTMA, BSTMP CST,

of the stria terminalis

(whole), BSTMPm

BSTIA → AA

(BSTM) is divided into:

(whole, vd), and

anterior (BSTMA),

caudal BSTMA

BSTMP → BM SLEAm→ BSTL (l)

SLEAm → AAV and APIR

BSTMP → Me

The latter is subdivided

and LA VL/VM

CeL(m) and BSTL

(MePDi)

ventral (BSTMV), and

MeAD, MePV and MePD → BMA and BMP

(l/m), and rostral (l/m),

column (MePDl), and

posterior (BSTMP) parts.

ventral sector of it

MeAD and MePV

into three cell columns:

BSTIA→ Me and

→ BAOT,

medial small celled

BSTM (m)

PMCO, AHi

medium celled

It is important to stress

BSTIA → AHi

(BSTMPi), lateral large

here that the CeM and

celled (BSTMPl)

CeC divisions of the Ce

(BSTMPm), intermediate

nucleus do contribute to

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TABLE 5

Intraamygdaloid Connections of the MEXA—cont’d MEXA efferents to the

Intrinsic connections Intraamygdaloid bed

the intrinsic circuitry of

nucleus of the stria

the MEXA. However, this

terminalis (BSTIA)

does not detract from the

OF/VM-CORLN

CEXA

LBNG

fact that for many other Medial sublenticular

reasons these two parts

extended amygdala

of the EXA should be

(SLEAm)

considered separately.

Medial division of the

CeM → rostral BSTMA,

supracapsular bed

BSTMV, BSTIal, BSTMPl,

nucleus of the stria

and BSTMPi

terminalis (BSTSm) CeC → BSTMA, BSTMV, and BSTI (l) light; (m) moderate; (d) dense; (vd) very dense; (vl) very light; (pm) poor to moderate.

located temporal components of the MEXA from the superficial cortical-like amygdaloid nuclei (Stephan and Andy, 1977). Clusters of small neurons, the perinuclear intercalated masses (IM and IP), occur along the medial and ventral margins of LBNC toward its rostral pole, and similar neuronal nests within the longitudinal association bundle are inserted between the LBNC and the representatives of the EXA and between the LBNC and the external capsule (Figs. 1 and 2).

capsule (Fig. 19). Caudally, the dorsolateral and ventromedial subnuclei of the LA nucleus become separated from the BL nucleus by a horizontal band of cells and fibers (that is, the ventrolateral LA nucleus) that intervenes between the BL nucleus and the rest of the LA nucleus. Despite their contiguity, the smaller and fainter stained cells as well as a much poorer fiber content in the LA nucleus allow it to be easily distinguished from the BL nucleus (Figs. 9, 13E, 16, 17, 18, 21C, 22, 23, 24B, 24D, 24F, 30, 31B, 31D, and 33A). Cytoarchitectonics

The Lateral Amygdaloid Nucleus (La) Topographic Landmarks The lateral amygdaloid nucleus is a relatively homogeneous gray mass that extends from a level just caudal to the cephalic pole of the BLA subnucleus until nearly the caudal end of the amygdaloid complex, occupying the dorsal aspect of the LBNC and enlarging at more caudal levels (Figs. 15G–15M in Alheid et al., 1995). Throughout its entire extent, it rests atop the BL nucleus, flanked laterally by the external capsule, which separates it from the endopiriform nucleus, and medially by neurons of the amygdaloid IMG and by a subventricular fiber stratum that intervenes between the LA nucleus, on the lateral side, and the Astr zone, the capsular part of the CeL division of the Ce nucleus, and the temporal horn of the lateral ventricle on the medial side. At rostral levels the LA nucleus sends out a ventral, laminar cell extension that separates the BLA subnucleus of the BL nucleus from the external

The LA nucleus is made up of palely staining smalland medium-sized neurons (Figs. 15G–15M in Alheid et al., 1995). On the basis of cell morphology and topography, the LA nucleus can be divided into dorsolateral (LaDL), ventromedial (LaVM), and ventrolateral (LaVL) subnuclei. Small, densely packed cells occupy the entire extent of the dorsolateral LA subnucleus, whereas the ventromedial LA subnucleus, the main division, is made up of slightly larger, more scattered, and palely staining cells. As can be seen below, this cytoarchitectonic distinction is further accentuated by the poorer fiber content in the ventromedial lateral subnucleus. The ventrolateral LA subnucleus, a horizontal band of cells separating the dorsolateral and ventromedial LA subnuclei from the BL nucleus, is first observed concurrently with the ventromedial lateral subnucleus and extends almost up to the caudal pole of the ventromedial LA subnucleus. In frontal sections, this subdivision of the LA nucleus appears for most of its length as a triangular wedge, with its wide end resting against the

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TABLE 6

Medial Extended Amygdala Afferents Subcortical afferents

Medial extended

Cortical afferents

Telencephalon

Diencephalon

Brain stem afferents

AOB→ Me

Anterior olfactory

Hypothalamus has

MEXA is a focus for brain stem afferents

amygdala (Me) can be

nucleus, horizontal limb

several amygdalopetal

divided into:

AOB terminals →

of the diagonal band,

fiber systems that

anteroventral (MeAV),

cell-poor zone that

endopiriform

innervate practically

Pars compacta substantia

anterodorsal (MeAD),

encapsulates the dorsal

nuclei → MEXA

the entire MEXA.

nigra, retrorubal fields,

posteroventral (MePV),

small-celled column

and posterodorsal

of BSTM.

Axons from dorsal

VMH→ Me (Bilat),

and caudal linear

anterior olfactory

AHi (Bilat), BSTMPi,

nucleus → Me

nucleus → layer 1B of

SLEAm, and maybe

Me and PMCO

BSTIA (Bilat)

Horizontal limb of the

Paraventricular

Anterior and posterior

diagonal band, →

hypothalamus → ME,

Dorsal tegmental nucleus,

olfactory cortex →

Me, SLEAm

AHi, BSTM, SLEAm, and

laterodorsal tegmental

(MePD) parts. MePD is subdivided

MOB → anterior tip Me

into: medial column

dorsal and medial raphe

(MePDl), intermediate column (MePDi)

Parabrachial area axons → Me

(MePDm), lateral column

Medial bed nucleus of the

→MEXA terminating in

stria terminalis (BSTM) is

1b Me

maybe BSTI Dorsal and ventral

divided into: anterior

nucleus, and dorsal nucleus of the lateral

endopiriform nuclei

Medial preoptic

(BSTMA), ventral

Neurons from the

→ Me and PMCo

terminals → Me, AHi,

(BSTMV), and posterior

posterior part of the

(layers 1 and 2)

BSTMPi, SLEAm

(BSTMP) parts. The

agranular insular cortex

latter is subdivided in

→ Me, BSTIA and

three cell columns:

SLEAm

Lemniscus → Me Locus coeruleus axons (noradrenergic) → Me

Anterior hypothalamus → Me, AHi, BSTM,

Ventral tegmental area →

medial small celled

SLEAm

BSTMPi, and PMCo

(BSTMPm), intermediate

Dorsal and Ventral

medium celled (BSTMPi),

premammillary axons

A8 (Retrorubal) → SLEAm

and lateral large

→ Me, PMCo, AHi

rostral and Caudal

celled (BSTMPI).

(Bilat), BSTMPi,

Linear nuclei → BSTMPi

possibly BSTIA

and Me

Entorhinal cortex → Intraamygdaloid bed

MEXA → sending

nucleus of the stria

terminals to PMCo,

Supramammillary →

Dorsal and median raphe

terminalis (BSTIA)

and BSTIA

only to posterior BSTM

→ BSTMP, PMCo, SLEAm

Medial sublenticular

Hippocampal areas →

and perhaps BSTIA Parabrachial nuclei →

extended amygdala

Me, BSTM, and BSTIA

Arcuate nucleus

(SLEAm)

Subiculum → Me, BSTM

terminate inMe, AHI

and BSTIA

(bilat), → BSTMP, SLEAm

Medial division of the supracapsular bed

Me, BSTMPl, SLEAm Locus coeruleus → PMCo, SLEAm

CA3→BSTM and BSTIA

Lateral preoptic area →

nucleus of the stria

Me, PMCo, AHi, BSTMPi

In the caudal medulla,

terminalis (BSTSm)

Lateral preoptic-lateral

nucleus of the solitary

hypothalamic continuum

tract → SLEAm

innervates → Me, PMCo Lateral hypothalamus → Me, PMCo, SLEAm, BSTIA

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TABLE 6

Medial Extended Amygdala Afferents—cont’d Subcortical afferents

Cortical afferents

Telencephalon

Diencephalon

Brain stem afferents

Tuberal and posterior LH→Me Periventricular hypothalamus is also a potential source of afferents to BSTIA and AHi. Dorsal and posterior Hypothalamus → SLEAm Dorsomedial hypothalamus → BSTM and BSTIA Medial amygdaloid nucleus is the main gateway for thalamic afferents: Paraventricular nucleus (bilat), paratenial nucleus, ventral posteromedial and posterolateral nuclei →Me Medial geniculate, suprageniculate nuclei and peripeduncular → Me Paraventricular nucleus of the thalamus terminals → posterior BSTM Rhomboid nucleus, subparafascicular nuclei of the thalamus and medial habenula → BSTM (l) light; (m) moderate; (d) dense; (vd) very dense; (vl) very light; (pm) poor to moderate.

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TABLE 7

Medial Extended Amygdala (MEXA) Efferents Subcortical efferents

Medial amygdaloid

Cortical efferents

Telencephalon

Diencephalon

Brain stem efferents

Me→ through band of

The MEXA sends axons

MEXA has reciprocal

Me is the main source of projections to brain stem

nucleus (Me) divided

axons of SLEAm→to

to horizontal limb of the

connections with medial

into: anteroventral

BSTMPm→ MEnt, LEnt

diagonal band, septum,

hypothalamus, mainly

and ventral striatum

VMH and PM nuclei:

(MeAV), anterodorsal (MeAD), posteroventral

MeAD→ AOB

(MePV), posterodorsal (MePD)

(RF) formation, PAGl BSTIA→HDB MS

BSTIA→ AOB BSTMPl→ AOB

into medial, lateral, and

Me→MPO, AHA, PM, RCh, SON, area around

BSTM→LSV, HDB MePD is subdivided

Me→ VTA, PAG, reticular

BSTI→RF (bilateral)

PVH, DMH, LHTC, Arc.(l)

BCST→RF

MeAV → VMH (core,

SLEAm→RF

BSTI→LS

intermediate columns

SLEAm→AOB, MEnt,

(MePDm, MePDl,

LEnt

SLEAm→ IPAC, Acb

bilateral), PMD/PMV

MePDi)

(bilateral), PP

Intraamygdaloid bed

MePD→MPO, PVH, SuM

nucleus of the stria terminalis (BSTIA)

BSTMA→MPO

Medial bed nucleus of the

BSTMA→VMH shell,

stria terminalis (BSTM)

PMV, PMD, AHA, PVH,

divided into: anterior,

LHTH, SON

ventral, and posterior (BSTMA, BSTMV, BSTMP) BSTMP: subdivided in three cell columns:

BSTSm→ MPO, AHA

medial small size (BSTMPm), intermediate

SLEAm→ AHA

medium size (BSTMPi), lateral large size (BSTMPl)

To thalamus:

Medial division of the

Me→ Re, MD, PP (bilateral)

supracapsular bed nucleus of the stria

BSTIA→ PP

terminalis (BSTSm) BSTMP→ PV, PT, PP Bed nucleus of

(bilateral)

commissural component of stria terminalis (BCST)

BSTS→ PP

Medial sublenticular

BCST→ PP

extended amygdala (SLEAm) (l) light; (m) moderate; (d) dense; (vd) very dense; (vl) very light; (pm) poor to moderate.

IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

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19. AMYGDALA AND EXTENDED AMYGDALA OF THE RAT

TABLE 8

Intraamygdaloid Connections of the Central Extended Amygdala (CEXA) CEXA efferents to

Forms a gray

CEXA intrinsic connections

Olfactory cortical-like amygdala

MEXA

LBNC

Unclassified amygdaloid nuclei

The nuclei of the

The CEXA sends

Few pathways

Few nuclei of the

BSTLD → Fu (vd) BSTLP→ Fu

continuum made of

CEXA possess a

axons to several

from CEXA to the

CEXA gray

the lateral division

rich complement

targets in the

MEXA group have

continuum project

of the BST nucleus

of associational

olfactory amygdala.

been identified.

(BSTL), subdivided

connections.

to the laterobasal

into dorsal (BSTLD),

CeC, CeL, CeM →

Ce → SLEAm,

posterior (BSTLP),

AA area

however, this lateral

CeCv → CeM (vd)

(LBNC).

projection may be

CeC, CeL, CeM →

CeM → AHi

directed, at least in

caudolateral BLP (l)

BSTL → AA area

neurons lying within

CeC, CeL, CeM

with preferential

and adjacent to the

→ BMA

termination in AAV

SLEAc.

divided into a medial Rostral and caudal

SLEAc → AA area

BSTLD → BSTMA

part (CeM) which

and also to ACO

rostral and caudal (d)

ventral (BSTLV), intermediate (BSTLI), and juxtacapsular

CeL → CeM (vd)

(BSTLJ) CeL → CeCv Central anygdaloid nucleus (CeA)

part to cholinergic

CeL and CeC → BSTLP

is subdivided into: anterodorsal

SLEAc → BL, BLV,

CeM → CeC

BSTLD → BSTMAl

(retrograde

Small celled part

(d) but not to

labeling from BL)

BSTMAm

anteroventral

CeC, CeL, CeM

CeM → PLCO

(CeMav),

→ SLEAc

nucleus and APIR

posteroventral

transition area

BSTLP → central BSTLD → far lat.

(CeMpv); and in a

CeM → BSTLD,

lateral (CeL) also

rostral and caudal

BSTLD → APIR AL

subdivided into:

(vd) BSTLP, BSTLV

transition area

BSTLD→ BSTI pm

central (CeL)

(d), BSTMA (m), IPACm → BSTMA

IPACm → BMA,

BSTMV (m), BSTI

CeL and BSTLD →

(CeC) subnuclei

(m) and BSTMPl (m)

ChAT-IR small cells

but rest of MEXA

in SLEA and in AAD

avoided

Central sublenticular

CeL → BSTLD (vd),

extended amygdala

and caudal BSTLP

Ce → PMCO

CeCv → caudal

ventral CeL)

BLP (vd)

(from the

(SLEAc) BSTLP (vd), BSTLV

the supracapsular bed (vd), rostral BSTMA, BSTMV, BSTI

IPACm → LOT, AAD/AAVl, APIR

terminalis (BSTSc)

AL transition area (vd)

Rostral BSTLP Interstitial nucleus of

BMA and BLP

BSTMPL (m)

and capsular

nucleus of the stria

and BM BSTVL → BL

nucleus (diffusely)

(CeMad),

Central division of

CeC, CeL, CeM → Fu

nuclear complex

→ caudal BSTLP

posterior limb of the anterior commissure

BSTSc → BSTL Continued

IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

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JOSE S. DE OLMOS ET AL.

TABLE 8

Intraamygdaloid Connections of the Central Extended Amygdala (CEXA)—cont’d CEXA efferents to Olfactory cortical-like amygdala

EXA intrinsic connections of which the medial

SLEAc (SID) → all

part (IPACm)

other nuclei of the

belongs to the

CEXA

MEXA

LBNC

CEXA BSTLD → BSTLV (m) and BSTI al BSTLD → BSTLP (vd) but not to BSTLjxc BSTLD → SLEAc (vd) BSTLD → st → BSTSc and CeA [CeM (vd), CeL (m), CeC (m)] BSTLP → BSTLD BSTLP→ BSTLV BSTLP → BSTMV, BAC (bed nucleus of the anterior commissure) BSTLP → SLEAc IPACm → SLEAc, BSTLP, BSTLV, but not BSTLD nor BSTLJ IPACm → CeM (vd), CeC (l/m), but not CeI or CeL (l) light; (m) moderate; (d) dense; (vd) very dense; (vl) very light; (pm) poor to moderate.

IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

Unclassified amygdaloid nuclei

579

19. AMYGDALA AND EXTENDED AMYGDALA OF THE RAT

TABLE 9

CEXA Afferent Connections Subcortical afferents

Cortical afferents

Telencephalon

Diencephalon

Brain stem afferents

Forms a continuum

The Ce nucleus receives

Few noncortical

The medial preoptic

The central amygdaloid

composed of lateral

terminals from the

telencephalic areas

(MPA) and anterior

group receives a striking

division of the bed

piriform cortex (PIR),

project to central

hypothalamic (AH)

set of afferents from the

nucleus of the stria

dorsal peduncular

amygdaloid group.

areas and hypothalamic

brain stem.

terminalis (BSTL),

(DPC), Ammon’s horn

paraventricular (PAVH)

subdivided into dorsal

(CA), and subicular

Interstitial nucleus of

nucleus all → both BSTL

(BSTLD), posterior

areas (S), as well as

the horizontal limb of

and SLEAc and to

complex and caudal

(BSTLP), ventral

from entorhinal (Ent),

the diagonal band (HDB)

CeMad

reticular medulla

(BSTLV), intermediate

perirhinal (PRh),

send possibly → to CeM,

(BSTLI), and

posterior agranular

BSTV, and SLEAc but

The lateral preoptic

juxtacapsular (BSTLJ)

insular (AIp), anterior

not to BSTL.

hypothalamic

Parabrachial nuclear

oblongata → BSTL, BSTV, SLEAc, Ce, and IPACm

continuum → the whole

Main projection areas of

Central nucleus of the

granular insular (GI),

Nucleus of the vertical

Ce, as well as the BSTV,

the pontine parabrachial

amygdala (Ce)

dorsal agranular

limb of the diagonal

BSTL, and SLEAc

nuclear complex (PB) →

subdivided into Medial

insular(AID), ventral

band may also project

part (CeM), subdivided

agranular insular

to SLEAc.

into anterodorsal

(AIV), infralimbic

(CeM ad), anteroventral

(IL), and prelimbic

Nuc. accumbens (Acb) →

hypothalamic (LH) area

The medial subnucleus

(CeM av), posteroventral

(PrL) cortices.

BSTV and BSTL as

also innervates → Ce,

(MPB) of the pontine

well as SLEAc

BSTL, and SLEAc

disgranular (DI),

(CeM pv); and lateral

Ce–SLEAc–BSTL The rostral and tuberal

continuum

part of the lateral

parabrachial nuclear

(CeL) which is also

Neurons from posterior

subdivided into two

sectors of PIR cortex

The rest of the dorsal and

LH and PeF

main parts: central (CeL)

are retrogradely

ventral striatum does not

hypothalamic neurons,

ventral, lateral subnucleus

and capsular (CeC)

labeled from FG

participate in the

→ orexin-B-like IR

(VL PB), and especially the

injections involving

projection to the CEXA

neurons within the Ce

external, lateral subnucleus

CeI and CeL.

gray continuum

and rostral sectors of

(EL PB) → BSTLD and CeL

the BSTLP

(d)

Medial hypothalamus

Finally, the central lateral

Central sublenticular extended amygdala (SLEAc or SID)

complex, the parabrachial waist area (PBwa), the

DPC → CeC → CEXA gray continuum: subnucleus (CL PB) →

Central division of the

Ventral S, ventral CA1

supracapsular bed

field, and DTT division

nucleus of the stria

of the precommissural

VMH and arcuate nuclei

terminalis (BSTSc)

hippocampus → CeI

innervate Ce, BSTL,

There are no significant

and CeL (FG inj.)

and SLEAc.

projections from the

Ventral S → CeC

Afferents from VMH

parabrachial area (mpPB)

terminate in the

to the BSTL division.

BSTLD and CeL (d).

Interstitial nucleus of the posterior limb of the

mesencephalic

anterior commissure (IPAC) which is divided

Dorsolateral lateral Ent

capsule-like area

Medial (MPB) and lateral

into lateral (IPACl) and

cortex → CeC (d),

surrounding Ce.

parabrachial (LPB) nuclei,

medial (IPACm), this

ventral rostral IPACm

latter being member

as well as from the nucleus Both dorsal (PMD)

of the solitary tract (NTS)

of the CEXA while the

Ventrolateral lateral Ent

and ventral (PMV)

→ bilaterally into the Ce as

former is a transitional

cortex → CeC (d),

premammillary nuclei

well as ipsilaterally to the

gray (Alheid et al., 1995;

ventral rostral IPACm

→ medial part of Ce,

SLEAc and BSTL Continued

IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

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JOSE S. DE OLMOS ET AL.

TABLE 9

CEXA Afferent Connections—cont’d Subcortical afferents

Cortical afferents

Telencephalon

Diencephalon

Brain stem afferents

but not any other nuclear

Dopaminergic (DA)

de Olmos and Heimer,

Rostral AIp cortex

1999; Heimer et al., 1999)

→ throughout the Ce

member of the CEXA

neurons in ventral

nucleus (diffusely)—

gray continuum

tegmental area (VTA/A10),

except CeC—BSTI (l) BSTLP (l), and IPAC (vd)

compact subst. nigra Dorsal tuberal area → BSTL

(SNc/A9), and retrorubal area (RR/A8) → Ce, SLEAc, BSTL, and IPACm

Caudal AIp (or PaRH) cortex → bilaterally →

Periventricular A11 group

throughout CeC and

of DAergic neurons →

DA neurons (A10cd) in

sparse to IPAC and Ce M

Ce–BSTL continuum

the mesencephalic

DI cortex→ CEXA:

Extensive array of

raphe linear (Li), and dorsal

BSTLD (vd), BSTSc (d),

thalamic amygdalopetal

raphe nucleus (DRN) →

periaqueductal gray (PAG),

BSTLJ (d), BSTLP

axons → central nucleus

whole CEXA gray

(l.), SLEAc (SID),

of the amygdala:

continuum (vd)

CeM, Ce I, CeL (d),

To a greater or lesser

IPACm (m)

extent, from the anterior

Noradrenergic (NE)

(PVA) and posterior

neurons in the locus

Rostral parts of DI cortex

(PVP)periventricular thal. coeruleus (LC) → Ce, BSTV,

avoid the core of Ce but

Nuclei, as well as from

rather terminate in the

the paratenial (PT),

surrounding neuropil.

reuniens (Re),

NE neurons in lateral dorsal

anteromedial (AM),

tegmentum → Ce, SLEAc,

Cells in the AIv cortex

interanteroventral (AIV),

BSTL, and BSTV

may also be labeled from

parafascicular (PF),

BSTL, SLEAc, and IPACm

retrograde injections

subparafascicular (SPF),

in CeI/CeL But AI

and suprageniculate (SG) neurons in dorsal raphe

v cortex → CeC (m)

thalamic nuclei, and

nucleus (NRD) → Ce,

the medial geniculate

BSTL, SLEAc and IPACm

Rostral IL cortex →

body (MGB)

whole CeC nucleus (l) and rostral IPACm

Serotoninergic (5-HT)

5-HT neurons in the raphe Afferent projection

nucleus and in the median

originated in the

raphe nucleus (MRN) →

Caudal IL cortex → CeM,

mediodorsal thalamic

Ce and SLEAc

CeI (l/m), CeL BSTL

nucleus (MD) and the

(l/m), BSTLD (m), and

lateral habenula (LHb)

Periaqueductual gray → Ce

BSTLP (m)

may also exist, but

and BSTLD

PrL cortex → CeC (l),

peripeduncular nucleus

5-HT neurons in PAG,

IPACm (l), BSLP (m),

(PP) → Ce nucleus

DRN, linear nuclei→

more securely from the

and BSTLV (m) but??? not BSTLD or BSTLJ

BSTLD CRF neurons Periventricular thalamic nuclei (PVA/PVP) → SLEAc

IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

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19. AMYGDALA AND EXTENDED AMYGDALA OF THE RAT

TABLE 9

CEXA Afferent Connections—cont’d Subcortical afferents

Cortical afferents

Telencephalon

Diencephalon

Brain stem afferents

PT and IAV thalamic nuclei bilaterally → SLEAc PP nucleus → ipsilaterally to SLEAc

TABLE 10

To cortical targets

CEXA Efferent Connections

SLEAc and corticopetal cholinergic cell groups

Subcortical efferents ———————————————————— Telencephalon Diencephalon

To brain stem targets

Form a gray

Except for SLEAc

Large cholinergic

Few subcortical

To the

The most

continuum made of:

which contains

neurons follow

telencephalic

hypothalamus:

characteristic

Lateral division

scattered probably

SLEAc along part

projections have

of the BST nucleus

ectopic cholinergic

of its course, but it

been shown to

CEXA innervates

CEXA →

(BSTL), subdivided

neurons, cortical

is not clear if they

the CEXA.

nuclei in both

mesencephalic PAG,

medial and lateral

pontine PPTg, and

hypothalamus.

parabrachial nuclear

projections of the

into: dorsal (BSTLD), efferents from

are an integral part

posterior (BSTLP),

CEXA appear to be

of this bed nucleus

ventral (BSTLV),

minimal.

of the longitudinal

the medial dorsal

association bundle.

striatum (heavier on

Ce, BSTV, BSTL,

medullary dorsal

contralateral side)

and SLEAc all

vagal nuclear complex

intermediate (BSTLI) and juxtacapsular

Ce → perirhinal

(BSTLJ)

cortex

Central anygdaloid

IPACm → PIR cortex, an important source

nucleus (CeA)

prefrontal (vl)

Ce → bilaterally to

send → lateral

The CEXA, including SLEAc is potentially

complex (PBNC) and

Ce, SLEAc → IPAC 1

hypothalamiarea

CeM, BSTLV, SLEAc,

Only Ce → lateral

ventrolateral

preoptic area

PAG and PPtg

Ce, BSTV, SLEAc

Ce, BSTL, BSTSI,

and IPACm →

(fundus striati)

of afferents to

divided into a

corticopetal

BSTLP →

medial part (CeM)

cholinergic cell

ventromedial

which is subdivided

groups located

and caudal Acb

into:anterodorsal

within the SLEAC

and BSTS → medial

SLEAc, IPACm →

(CeMad),

itself. These cells

preoptic area

PBNC

anteroventral

are, however,

(CeMav),

smaller than

IPACm → ventral

Ce and BSTV →

Ce, BSTLD, BSTLV,

posteroventral

those of the MCBFC

AcbSh,caudal Tu

paraventricular

BSTSI, SLEAc and

hypothalamus,

IPACm → dorsal motor nuc. of the

(CeMpv); and a lateral part (CeL) also

IPACm → VP,

VMH and DMH

subdivided into:

SIB (CeXA?)

hypothalamic nuclei

central (CeL) and

BSTLP → Fu

capsular (CeC)

IPACm → not to Fu

vagus, nuc. of the solitary tract (NTS),

SLEAc→ VMH

and nuc. ambiguous

Ce→ dorsal

Ce→ intercalated nuc

subnuclei Central sublenticular

hypothalamic

of the medulla and

extended amygdala

area (DA)

subpostrema area

(SLEAc) (l) light; (m) moderate; (d) dense; (vd) very dense; (vl) very light; (pm) poor to moderate.

Continued

IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

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TABLE 10

To cortical targets

CEXA Efferent Connections—cont’d

SLEAc and corticopetal cholinergic cell groups

Subcortical efferents ———————————————————— Telencephalon Diencephalon

To brain stem targets

Central division of the

SLEAc, Ce, and

SLEAc → central

supracapsular bed

IPACm→ PH

linear nucleus

nucleus of the stria terminalis (BSTSc)

(LiC) of the raphe SLEAc and IPACm → SuM and Zi

Interstitial nucleus

(subthalamus)

of posterior limb of

Ce, BSTLP, BSTLD, BSTLV, and IPACm → DRN

anterior commissure

IPACm and BSTLD

of which the medial

→ anterior PaAP

Ce and BSTLD →

part (IPACm)

and PeF nuc.

locus coerulus

BSTLD→

CE, BSTL, SLEAc, and

parasubthalamic.

IPACm→ SNc and

Hypothalamus

VTA

To the thalamus

Ce, BSTLD,

(which is only

BSTLV, SLEAc,

sparcely supplied

and IPACm →

by the CEXA):

lateral RRF

Ce→ VM

Ce, BSTL, SLEAc, and

thalamic nuc.

IPACm → medullary

belongs to the CEXA

dorsal reticular nuc. SLEAc→ medial part of the MD thalamic

BSTLD → lateral LTg

nuc.

nucleus

However, the thalamopetal neurons in SLEAc may only represent caudal-most elements of the VP. BSTLD → midline thalamus IPACm→ medial MD, IMD, RE, PAV, RE, and VM thal. nuc. To epithalamus (lateral habenula) which is only sparsely supplied by the CEXA

IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

19. AMYGDALA AND EXTENDED AMYGDALA OF THE RAT

external capsule (Figs. 13E, 18, 22, 24B, 24C, 24F, 31B, 31D, and 33A). Fibroarchitectonics In neurofibrillar preparations (Fig. 13E), the LA nucleus appears encapsulated by bundles of fibers laterally (external capsule), medially (the longitudinal association bundle among others), dorsomedially, rostrally, and even caudomedially where a very thin plate of fibers separates the nucleus from the ventricular wall. These clear-cut separators are so neatly defined ventrally where it borders the BLA, but from which it can be distinguished by the lesser density of its fiber network, more particularly caudally and medially, where the La nucleus appears clearly fiber poor. Caudally, at the level of the rostral tip of the temporal lateral ventricle, a very dense transversely coursing group of fibers intervenes between the LaVL division of the LA nucleus and the BLP division of the BL nucleus. Furthermore, the fiber networks within the nucleus are not homogeneously distributed, i.e., they are more dense in the LaDL and LaVL subdivisions than in the LaVM subdivision. Heavy Metals and Acetylcholinesterase The distinction between the dorsolateral and the ventromedial LA subnuclei is also verifiable in AChEstained and Timm/Danscher-impregnated sections. The dorsolateral LA1 subnucleus gives a moderate AChE reaction (Figs. 18A, 22C, and 24D), but a weak Timm/ Danscher’s staining (Figs. 18B, 22B, and 24E), which contrasts with the very weak AChE staining and the very intense Timm/Danscher’s impregnation in the neuropil of the ventromedial LA subnucleus. The AChE and Timm’s staining patterns described above are not completely uniform throughout either subdivision of the LA nucleus, but show some topographic variations. Thus, the Timm/Danscher’s reaction is even weaker in the ventral part of the dorsolateral LA subnucleus, contrasting sharply with that in both the ventromedial LA subnucleus and the BLP nucleus. The ventromedial LA subnucleus in turn shows patches of increased AChE staining toward its medioventral aspect and gradually a more intense Timm/Danscher’s staining caudally (Ben-Ari et al., 1977; Haug, 1976; Jacobowitz and Palkovits, 1974). Choline Acetyltransferase According to Hellendall et al. (1986), the LA displays less ChAT-IR than the BL nucleus, being that this IR is still higher than that in other areas around it. This contrast is even more obvious in the microdissected and assayed material, the average ChAT activity in the LA nucleus being roughly half of that in the BL nucleus. Both methods indicate a small gradient of ChAT

583

activity from anterodorsal (high) to posteroventromedial (low). Heckers and Mesulam (1994) reported a medium ChAT-IR density for the LA nucleus equivalent to that present in the BM and PLCO nuclei and the AA area.

The Basolateral Amygdaloid Nucleus (BL) Topographic Landmarks The BL nucleus is a collection of large, strongly staining cells in Nissl sections (Figs. 15G–15M in Alheid et al., 1995). It spans the entire rostrocaudal extent of the LBNC, appearing rostrally before its dorsolateral, ventromedial, and ventral companions, the LA nucleus, BM nucleus, and BLV nucleus, respectively, and extending farther caudally than any of them (Figs. 9, 13E, 16, 17, 18, 21C, 22–24B, 24D, 24F, and 33A). Cytoarchitectonics Because of the larger size of its cells, the largest found in the amygdala (Figs. 15G–15M in Alheid et al., 1995), the BL nucleus can be easily delineated from neighboring structures. Rostrolaterally, a ventral prolongation of the LA nucleus appears to intervene between the BL nucleus and the external capsule (ec) (Figs. 22A, 23A, and 24B). This extension of the LA nucleus alternatively has been suggested as the homolog of the primate paralaminar subdivision of the LBNC (Price et al., 1987). In our view it is not likely that these particular neurons are the counterpart of those in the primate. An alternative may be some of the small cells that occur more caudally along the external capsule. We have included all of these caudal cell clusters within our definition of the intercalated cell masses (see below). The external capsule separates, more ventrally and caudally, the BL nucleus from the ventral olfactory cortex and, through a medially directed branch, the BLV nucleus. At a level slightly caudal to the exit of the stria terminalis from the amygdala, the BL nucleus is separated from the rest of the LA nucleus by the medially orientated band of cells and fibers that form the ventrolateral LA nucleus. Before reaching its caudal end, the BL nucleus adopts a typical crescent shape, blending mostly with the APir area. Rostrally, the BL nucleus is composed of cells slightly larger and darker staining than those observed caudally, and for this reason the nucleus may be subdivided into anterior (BLA) and posterior (BLP) parts (Krettek and Price, 1978b). The rostral pole of the anterior BL nucleus appears just dorsocaudal to the anterolateral groups of the intercalated cell masses (IM), it expands into an avoid mass whose longer axis is oriented parallel to the medial margin of the main nucleus, occupying in this manner the rostromedial third of the BL nucleus. Due to the small size and light staining of

IV. DIENCEPHALON, BASAL GANGLIA, AMYGDALA AND SEPTUM

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JOSE S. DE OLMOS ET AL.

its cells, the BLP nucleus cannot be easily distinguished from the ventromedial LA nucleus at levels at which the ventrolateral LA nucleus does not intervene between these two nuclei. As pointed out by Yu (1969), small groups of intercalated cells interposed intermittently between the LA nucleus and the posterior BLP nucleus help to trace their common boundary. Fibroarchitectonics In neurofibrillar preparations (Fig. 15E), the BL nucleus appears to be clearly delimited from surrounding cell groups by a capsule-like fiber formation. The fibers envelop the BL nucleus on most sides and especially in its anterior subdivision, the core of which also contains a characteristically dense fiber network. Caudal to the BLA nucleus, the posterior BL nucleus is striated because of a multitude of small bundles of fibers that course through this subdivision to join the longitudinal association bundle. The presence of these bundles also helps to differentiate at this level the posterior division of the BL nucleus from the LA nucleus, which appears to be almost free of them. More caudally, the posterior BL nucleus loses its striated appearance to become separated from the LA nucleus by the previously mentioned horizontal fiber lamina, whereas ventromedially another laminar fiber formation intervenes between the posterior BL nucleus and the BM nucleus. On the whole, the fiber architecture in the BL nucleus not only confirms its cytoarchitectonical subdivisions but also suggests a further subdivision of its posterior portion. Heavy Metals In Timm/Danscher’s preparations (Figs. 18B, 21C, 22B, 23B, and 24F), the BLA subnucleus stands out because of its lighter staining compared with that of the LA nucleus laterodorsally, the amygdaloid IMG and the capsular subdivision of the CeL nucleus medially, and the ventral LA nucleus ventrally. A rim of darker staining separates the BLA subnucleus ventromedially from the very lightly stained BMA subnucleus. Caudally, the boundary with the BLP subnucleus is not sharp but changes rather gradually to a quite darker staining. The Timm/Danscher’s staining in the BLP subnucleus does not reach the density observed in the posterior LA nucleus and the posterior BM nucleus. The transition zone between the anterior and the posterior encapsulated portions of the BL nucleus shows an intermediate staining reaction in the neuropil, which is independent of the bundles of fibers traversing it. Acethylcholinesterase In acetylcholinesterase preparations (Figs. 17B, 18A, 22C, and 24D), the BL nucleus is homogeneously and

densely stained throughout its whole extent, a feature that also contributes to its sharp delineation from the surrounding cell groups (Ben-Ari et al., 1977; Parent, 1971). Choline Acetyltransferase According to Hellendall et al. (1984), the BL nucleus, together with the L2 of the LOT nucleus, displays the highest ChAT-IR and the highest ChAT activity in microdissected and assayed material. However, there exists a clear anterior to posterior gradient, which is substantially lower in the posterior part of the nucleus.

The Ventral Basolateral Amygdaloid Nucleus (BLV) Topographic Landmarks This irregular mass of large- and medium-sized, originally deeply staining cells (Figs. 15H–15J in Alheid et al., 1995) makes its appearance at a transverse plane just caudal to the LOT nucleus and extends caudally until the BLV merges with the rostral unfolding of the APir area (Figs. 13E, 16, 17, 22, and 33A). Throughout most of its rostrocaudal extent, the BLV nucleus lies deep to the ventromedial edge of the primary olfactory cortex, interposed between it and the BL nucleus. It is continuous laterally with the ventral EN nucleus (that is, in the restricted sense described below) and medially with the BM nucleus, from which it can be differentiated on the basis of its larger and more deeply staining cells. Toward the caudal end of the amygdala, its topographic relations change, and it comes to occupy a position ventromedial to the BL nucleus and dorsal to the PLCO nucleus. Cytoarchitectonics The population of large and deeply staining neurons in the BLV nucleus remains quite homogeneous throughout its extent (Figs. 15H–15J in Alheid et al., 1995), which it is not the case with the ventral division of the endopiriform nucleus (VEn), which has a more heterogeneous cellular composition with abundant neurons that are smaller and more irregularly shaped than those in the BLV nucleus (Figs. 16A, 17A, and 22A). Fibroarchitectonics In neurofibrillar prreparations (Fig. 13E), the BLV cells appear embedded in a dense plexus of transversely coursing fibers which become denser toward the dorsal and ventral boundaries of the nucleus, but lighter medially. The general orientation of this plexus as well as a less dense arrangement of its fibers allows us to differentiate the BLV nucleus from the more dense obliquely arranged fiber plexus in the VEn.

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Heavy Metals The BLV nucleus shows a rather strong Timm/ Danscher’s reaction (Figs. 16B, 17C, and 22B) that is of a magnitude greater than that in the territory of the VEn nucleus. Acetylcholinesterase Thoughout its extent the BLV nucleus displays a AChE activity that is rather low but still greater than that in the VEn nucleus (Figs. 17B and 22C). Choline Acetyltransferase The BLV nucleus, although not recognized as such by Hellendall et al. (1984) who define it as the medial part of the VEn nucleus, would contain 50% higher activity than the DEn and the neocortex; however, in contradiction with this the ChAT-IR in the BLV would be less than that in the latter-mentioned nucleus. A close examination of the illustrations presented by these authors (Fig. 1D) indicates that the median and caudal sectors of the BLV would display a ChAT-IR at least equivalent to that in the LA and BM nuclei. Like the just-mentioned authors, Heckers and Mesulam (1994) do not mention the BLV nucleus, which they seem to include within their accessory basal nucleus which these authors consider to display a medium density of ChAT-IR.

The Basomedial Amygdaloid Nucleus (BM) Topographic Landmarks The BM nucleus is formed by two ovoid cell masses succeeding one another in the rostrocaudal direction (Figs. 15G–15M in Alheid et al., 1995). The rostral collection of cells, or the anterior subdivision (BMA), is bordered throughout its anteroposterior extent by the BLV nucleus laterally, the Ce nucleus and main intercalated cell mass dorsally, and the ACO nucleus and PLCO nucleus ventrally. The posterior subdivision (BMP) is flatter and narrower than the BMA subnucleus; it is surrounded dorsally, laterally, and caudally by the BLA and BLP subnuclei, whereas medially and ventrally it is bordered by the AHI area and PLCO nucleus, respectively (Figs. 13E, 16, 17, 21C, 22, 30, and 33A). Cytoarchitectonics The BMA subnucleus (Figs. 15G–15M in Alheid et al., 1995) is made up predominantly of small- and mediumsized, lightly staining neurons that become densely packed dorsally and caudally; rostrally and ventrally they mingle diffusely with those belonging to the ACO nucleus and the AA area. In comparison with the BMA subnucleus, the BMP subnucleus is made up of

585

slightly larger and more darkly staining neurons. These neurons show a more scattered arrangement, and as a result it is sometimes difficult to delineate the posterior BM nucleus from its dorsolateral escort, the BL nucleus, or ventrally from the more superficial PLCO nucleus, and the more laterally and deeply lying cells of the AHI area (Figs. 16A, 17A, and 22A). Fibroarchitectonics In neurofibrillary preparations (Fig. 13E), the two subdivisions of the BM nucleus show relatively poorer fiber networks than the majority of the neighboring nuclei. This characteristic facilitates the distinction of the BMA subnucleus from the deeper ventral layer of the ACO nucleus, which contains a much denser fiber plexus which appears to be a continuation of that in the deep layers of the primary olfactory cortex. Medially, a loose longitudinal system of fibers runs immediately above and below the intercalated cell masses and is interposed between the BMA subnucleus and the deep cell layer of the anteroventral and anterodorsal Me nucleus. This medial, sagitally oriented fiber system thins out caudally and drifts dorsally, and as a result it becomes difficult to trace a clear boundary between the BMA subnucleus and its medial escort, the Me nucleus. The BMP subnucleus not only possesses a fiber network richer than that of the BMA subnucleus but also lies semiencapsulated by subventricular fiber bundles dorsally and by a loose collection of fibers running tangentially to its medial and lateral borders. These separate it from the AHI area and the BLP nucleus, respectively. Ventrally, however, no special fiber arrangement assists the observer in tracing its boundary with the PLCO nucleus. Heavy Metals In Timm/Danscher’s preparations (Figs. 16B, 17B, 21C, 22B, and 30B), the BMA subnucleus is, with the AA area, the least reactive of all the amygdaloid nuclei and is even less reactive than the BLA, AHIPM area, and anteroventral Me nucleus. The BMP subnucleus shows exactly the opposite staining pattern: it accumulates a very dense silver sulfide precipitate in its neuropil, comparable to the one seen in the deep layer of the primary olfactory cortex, posterior LAVL nucleus, PMCO nucleus, PLCO nucleus, and lateral capsular subdivision of the Ce nucleus. Acetylcholinesterase In AChE-stained preparations (Figs. 17B, 22C, and 30A), theBM nucleus shows, in general, a very weak staining reaction, which contrasts markedly with the very strong one in the BL nucleus (Fig. 18A). Examination, under dark-field illumination, of cholinesterase

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preparations that have not been reacted with silver indicates a differential staining reaction pattern in the two subdivisions of theBM nucleus. The BMA subnucleus shows a reaction somewhat stronger than that of the BMP subnucleus, although weaker than that of the ACO nucleus and the BLV nucleus. The BMP subnucleus, because of its very low AChE activity, can be easily distinguished from the PLCO nucleus and, certainly, from the BLP subnucleus and the AHI area. Choline Acetyltransferase According to Hellendall et al. (1984), the ChAT activity in the BM nucleus would be equivalent to that present in the LA nucleus despite that its ChAT-IR appears to be rather light. By contrast Heckers and Mesulam (1994), in agreement with the microassay results of the previous authors, evaluated the ChAT-IR in the BM nucleus to be of medium intensity.

General Hodological Relationships of the LBNC A wide range of reciprocal connections with the cortex as well as the extensive efferents to the striatum seems to distinguish the LBNC from the rest of the amygdala. All of the nuclei of the basolateral amygdaloid group receive afferents from the PIR cortex, the AI cortex, and the PrL cortex. While only the BLV nucleus seems to project back to the PIR cortex, all the nuclei of the basolateral amygdaloid group project back to the AI cortex and the PrL cortex. In addition, all of the nuclei of the LBNC project to the Ent cortex. Subcortically, all of the nuclei of the LBNC including the BM nucleus, project to the dorsal and ventral striatum. Although this projection appears to be poorly reciprocated, neurons in the dorsal and ventral pallidum may send striatal information back to the BL nucleus (Carlsen, 1988; Groenewegen et al., 1984a). Together with pallidal areas, neurons in the NHDB and in the SLEA project into the LBNC, carrying presumably cholinergic and noncholinergic terminals (Carlsen, 1988). As mentioned previously, the BM nucleus appears exceptional among the LBNC in that it sends a rather distinctive group of projections to media1 hypothalamic targets. Efferents to the medial hypothalamus have also been ascribed to the BL nucleus, but in our view such hypothalamopetal neurons do not belong to the BL nucleus. Rather, they fall either in the most caudal aspect of the BM nucleus, which intervenes between the AHI transition area and the most caudal pole of the BL nucleus, or in an area that we have termed the ventrolateral part of the LA (LaVL). This latter subnucleus has connections that are not typical of either the LA or the BL nucleus and has been

loosely grouped with the LA nucleus only because it appears to follow the rostral–caudal extent of the ventromedial part of the LA (LaVM). Interestingly, it is the LA and, possibly, the BL nuclei that receive afferents from the medial hypothalamic nuclei, while only a few terminals from the medial hypothalamus terminate in the BM nucleus. Considering the thalamus, all of the nuclei of the LBNC receive inputs from the nucleus reuniens (Re) and paratenial nucleus (PT). However, thalamic afferents to the LBNC are not limited to these two nuclei. A diverse array of thalamic areas apparently projects into the LBNC with a slight preference for targets in the LA and BL nuclei as opposed to the BM nucleus. Only a sparse projection from the BL and its BLV nuclei appears to innervate the thalamus, and this efferent terminates in the mediodorsal thalamic nucleus (MD). Within the amygdala the LBNC receives the projections from all the nuclei of the olfactory amygdala, while afferents from the other amygdaloid groups are somewhat sporadic. The LBNC reciprocates the projection to the olfactory amygdala but does not seem to have systematic efferents to the MEXA. The LBNC has a rather dense projection to the continuum formed by the central amygdala–SLSI–BSTL or CEXA. In some respects, the LA and BL nuclei resemble the cortex (Hall, 1972a, 1972b; Millhouse and de Olmos, 1983). A majority of the neurons in these nuclei resemble pyramidal neurons with their apical dendrite directed rostrally. Similar to the cortex, they possess a diverse input from the thalamus which, however, is only poorly reciprocated, and they send a widespread projection to the striatum (dorsal and ventral). Interestingly, glutamate may be the transmitter used in the amygdalostriatal projection (Fuller et al., 1987), as it is also in corticostriatal pathways. The striatal terminals from the LBNC tend to be most dense in the ventral striatum, to be less dense in the dorsomedial parts of the striatum, and to avoid the dorsolateral sectors of the striatum altogether. This topography is consistent with the idea that the LBNC projects most densely to the same zones of the striatum that receive cortical inputs from areas of the cortex that are reciprocally connected with the LBNC. This feature of related cortical areas projecting to similar zones in the striatum (Yeterian and Van Hoesen, 1978) has been seen to be as characteristic for the rat (Donoghue and Herkenham, 1985; Gerfen, 1984; Veening et al., 1984) as it is for other species. As with cortical areas, the LBNC receives afferents from the cholinergic cells of the ventral forebrain and although the projections to the BM nucleus may be slight, the cholinergic projection to the BL nucleus is one of the densest in the brain. Finally, as with association areas of cortex, the LBNC sends efferents to the primary motor cortex (Sripanidkulchai

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et al., 1984), so that the LBNC may have both direct and indirect influences on the organization of movement.

Neurotransmitters and Neuromodulators of the LBNC Cells The laterobasal amygdaloid nuclear complex does not appear to have the rich diversity of transmitters that is so striking for nuclei in the medial and central extended amygdala. Nonetheless, several neurotransmitter systems have been identified in the laterobasal complex and this includes both cells and fibers of neuropeptides. Cholecystokinin, VIP, NT, and SOM cells are found throughout the laterobasal nuclear complex (including the BM) (CCK, Cho et al., 1983, Greenwood et al., 1981; Innis et al., 1979; Roberts et al., 1982; Vanderhaeghen et al., 1980, 1981; Zaborsky et al., 1985a; VIP, Lorén et al., 1979; Roberts et al., 1982; Sims et al., 1980; NT, Jennes et al., 1982; Roberts et al., 1982; SOM, Bennet-Clarke et al., 1980; Finley et al., 1981b; Gray, 1983.) However, Shiosaka et al. (1983) do not describe VIP neurons in the BLV. Enkephalin neurons are found in the LA, BL, and BM but have not been described in the BLV (Finley et al., 1981c; Khachaturian et al., 1983; Roberts et al., 1982; Sar et al., 1978; Shiosaka et al., 1983), however, do not

TABLE 11

report enkephalin neurons in the BM. Substance P, βEND, NPY, and Ang II apparently do not occur in the neurons of the laterobasal nuclear complex while CRF neurons have only been reported in LA (Pilcher and Joseph, 1984; Swanson et al., 1983). Finally, neurons in the BL selectively transport glutamate retrogradely from injections sites in the striatum (Fuller et al., 1984) suggesting that this transmitter is used in the extensive amygdalostriatal projection. Fibers Fibers of various neuropeptides are found thoughout the laterobasal nuclear complex and, of course, in all the nuclei that contain the relevant neuropeptide-positive cell bodies. In addition to this, SP fibers can be seen in the LA and BL despite the absence of SP-positive cells. Similarly, CRF fibers are observed in the BM as is Ang II (in BMP), while βEND and pancreatic polypeptide fibers are found in the LA (Finley et al., 1981a; Olschowska et al., 1982) As mentioned in the description of the laterobasal amygdaloid nuclear complex, the LA, BL, and BLV stain well with AChE histochemistry, suggesting the presence of cholinergic terminals in all these nuclei.

Intraamygdaloid, Interamygdaloid, and Extraamygdaloid Connections of the LBNC

Laterobasal Amygdaloid Nuclear Complex (LBNC) Intraamygdaloid Connections Efferents to

Lateral amygdaloid

Intrinsic connections of LBNC

Olfactory amygdala

Medial amygdala (MEXA)

LA → BL, BM

The basolateral (BL)

Several Zn-rich pathways The CEXA seems to be the

nucleus (LA) subdivided

Central amygdala (CEXA)

Amygdaloid nucleus:

come from the LBNC to

principal recipient of

terminate in the MEXA

Zn-rich afferents from

into dorsolateral,

Apparently BLV does

main source of efferents

ventromedial,and

not → BM

from the LBNC to the

ventrolateral (LADL,

LA→(Zn) to LAP,

olfactory amygdala (Zn:

LAVM, LAVL)

BLP, BM

zinc-rich efferents)

the LBNC. All nuclei of LBNC → Ce

BMA→LA, BL Basolateral nucleus

BMP→ LAvl(d), BLA,

(BL) subdivided into

BLP, BLV →

anterior and posterior (BLA, BLP)

LAa→AHI(Zn)

BLA/BLP/BLV →BSTL BLV→ BSTLPr

LAp→PLCO, → BMP, LA

APIR(PM)(Zn)

BLV→ layers 1 and 2

(contralat.)

PMCO(Zn)

of the Me, SLEAm.

BL → all subnuclei of

BSTMV, BSTIpm.

BLP/BLV→ SLEAm

→BSTMP, BSTMA,

Ventral basolateral nucleus (BLV)

BLA →Me (bilateral)

BLP→BSTPc, BSTLV

olfactory amygdala (l) light; (m) moderate; (d) dense; (vd) very dense; (vl) very light; (pm) poor to moderate.

Continued

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TABLE 11

Laterobasal Amygdaloid Nuclear Complex (LBNC) Intraamygdaloid Connections—cont’d Efferents to Intrinsic connections of LBNC

Basomedial nucleus (BM)

Olfactory amygdala

Medial amygdala (MEXA)

Central amygdala (CEXA)

BLV →APIR, PMCO

BLP→(Zn) to APIR,

BM→SLEAm

PMCO. →BSTMA/MV

subdivided into anterior and posterior (BMA, BMP)

BMA→ACO, PLCO(Zn),

BSTMP, BSTL

BMA

LBNC has strong

→ BAOT, LOT

BMA

→CeLCn, CeI(l).

→Temporal: MeAD,

→Paraseptal: BSTLP,

cortical areas and striking

BMP→APIR (m), AA,

MeAV, MePV (d),

BSTLV, BSTLD.

output to dorsal and

LOT, PLCO(Zn), PMCO,

MePD (l).

ventral striatum.

ACO. (l) AHi

→ Paraseptal:

BMP

BSTMPl, BSTMPi, (d)

→ Temporal: CeLCn, CeM.,

BSTMV, BSTI, and

IPAC.

SLEAm, SLEAc, IPAC.

→Paraseptal: BSTLP,

→Temporal: CeLC, CeM(d).

PMCO, APIR(d) interconnections with

(See Table 12)

BSTLV, BSTLD. BMP→MeAD, MePV(l), and BSTIA. →BSTMA/MV /MP, BSTImp, →BMP(contr.) Legends: (l) light; (m) moderate; (d) dense; (vd) very dense; (vl) very light; (pm) poor to moderate.

UNCLASSIFIED CELL GROUPS IN THE AMYGDALA AND BED NUCLEUS OF THE STRIA TERMINALIS The difficulty with including the large intercalated cell mass (IM) within the symmetrical structures (Fig. 5) of the central or medial division of the EXA highlights a problem that generally confronts any classification schedule. Specifically, some of the areas or items under consideration are often different enough from the proposed classification schedule that to arbitrarily include them within an ill-fitting category may obscure their unique nature. To avoid this we have left several unclassified nuclei for consideration in this section.

The Intercalated Cell Masses (IM and I) and the Extended Amygdala Topographic Landmarks In frontal Nissl sections (Figs. 15F–15I´ in Alheid et al., 1995) the intercalated nuclei appear to be constituted by oval and elongated plate-like clusters of smallto medium-sized, tightly packed neurons located within the spaces existing: (i) among various amygdaloid nuclei; (ii) between some amygdaloid nuclei and the striatum;

and (iii) along the longitudinal association bundle, in the intraamygdaloid portion of the stria terminalis, and between the LA or BL nuclei and the external capsule, that is, the paracapsular (I) groups (Figs. 24B, 24D, 24F, and 31B). Serial horizontal sections reveal, in addition, that the most conspicuous representatives of the intercalated cells are constituted in reality by a continuous irregular Z-shaped cell column, here termed the main intercalated mass, or nucleus (IM) (Figs. 1, 13E, 17, 22, and 30). The lateral arm of the IM gray mass starts rostral and ventral to the cephalic pole of the anterior BL nucleus and medial to the ventral EN nucleus and maintains this topographical relationship until it turns medially (its transverse arm) and then again caudally (its medial sagittal arm). Along this route, it lies interposed successively (from rostral to caudal): (i) between the Ce nucleus and the anterior BM nucleus; (ii) between the anterior BL/anterior BM and the Me nucleus; (iii) and between the anterior BM/posterior BM nuclei and the anterolateral AHI transition area (Figs. 15G–15J in Alheid et al., 1995). By contrast with this long, continuous column of intercalated cells, those appearing in a dorsal paracapsular position or intervening between the ventromedial LA and the BL nuclei do not maintain any continuity among themselves, with the IM, or with other less prominent small cell clusters which

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TABLE 12

Laterobasal Amygdaloid Nuclear Complex (LBNC) Afferents Subcortical afferents

Lateral amygdaloid

Cortical afferents

Telencephalon

Diencephalon

Brain stem afferents

→ LA:

Ventral forebrain

From hypothalamus:

Dopamine afferents:

nucleus (LA) subdivided

PRh, PIR, Ect, DEnt,

→ all LBNC

into dorsolateral,

LEnt, AI, IL, PRL,

(cholinergic and

→ LA:

→ all LBNC:

ventromedial, and

AU, SM, PrL, CgI, VI,

non-cholinergic)

VMH, DMH, PM, SuM,

VTA, SN

ventrolateral LA (LADL,

DIv, DIg, PAR → LA:

LAVM, LAVL) → LAVM: Basolateral (BL)

DB, VP, Acb(1)

LEnt, MEnt

hypothal.

→ BL:

→ LADL:

DB, VP,

nuclei

RRF (A8)

GI, SM, PAR, DIv

AcB(1), VDB

→ LAVLm:

→ BLV:

VI, SM, S

DB, VP, PIR

→ BM:

subdivided into anterior

VMH, PMD, PMV

5HT afferents: DG, RMg, CLi

From thalamus:

and posterior (BMA,

→ BL:

→ BM:

BMP)

PIR, LEnt, AI, MO, MEnt,

HDB, VDB, VP

Laterobasal nuclear

RRF (A8)

Some of above

Ventral basolateral (BLV) Basomedial nuclei (BM)

→ LA: → BL:

→ BL:

subdivided into anterior and posterior (BLA, BLP)

PeF, LPO, PH.

→ → LA:

LADL, LAVM,

PRL, Cg1, VI, PAR, DIg,

Re, PT, PV,

LAVL, BL (vd)

DIv, SM, M1, M2, CA1

IAM(bilat),

complex (LBNC) has strong interconnections

→ BLA:

with cortical areas and

PIRM, DEnt, Ect, LEnt(vd)

CM, CL, VPM, MG,

→ BL:

PP

PAG, dtg, PPt, MPB, LPB

→ BL:

→ BLV:

and ventral striatum

→ BLP:

Re, IAM, MD, SPFC,

PB

(See Table 11).

AID, DIg, S. PIR,

MG, PP

striking output to dorsal

→ BM:

DEnt(vd), LEnt(vd), → BLV:

DRD, MnR, VTA, MPB,

IAM, Rh, MD(1)

LPB

PIR(bilat), DEnt(vd) LEnt,

→ BM:

Noradrenaline afferents:

DPC, AI, SM, PAR, IL, Sv

Re, PT, PV, PP

IL, DIg, Sv → BLV:

From LC to all LBNC → BM: → LA: LDTg

PIR, DEnt(m), AI, LEnt(m), PRI, SM,

→ BL: SubC

AU(1), DIg, Div, VI, Sv (l) light; (m) moderate; (d) dense; (vd) very dense; (vl) very light; (pm) poor to moderate.

accompany the IMG (Figs. 1, 13E, 17, 21, 22, 24B, 24D, 24F, 30, 31B, 31D, and 32). Cytoarchitectonics The intercalated cell masses, as generally defined, represent a heterogenous population of scattered, tightly packed, darkly staining cell clusters and one large

nucleus which are found interspersed among the various nuclei of the amygdala (Figs. 15F–15J in Alheid et al., 1995; and Figs. 17A and 22A, this chapter). Fibroarchitectonics In neurofibrillar preparations, the intercalated cell masses appear typically encapsulated by dense

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TABLE 13

Laterobasal Amygdaloid Nuclear Complex (LBNC) Efferents Subcortical targets

Lateral amygdaloid

Cortical targets

Telencephalon

Diencephalon

Brain stem targets

LA→ID, AIP, IL, Prh, M2,

LA→VStr, DStr

→Hypothalamus:

LA→VTA

LA→VMH, LH

BLV→VTA

nucleus (LA) subdivided

Cg, Tev, VIll, AUd,

into dorsolateral,

Ptp, RSd, Prh, Ect, Ent

BL→DStr, VStr:

ventromedial, and

cortices

TuO, Acb, CP,

ventrolateral (LADL,

LAVL→LEnt, IL, MEnt,

IPACl

LAVM, LAVL)

PRh, Ect, AID, AIP

→VP, HDB

LAVL→LPO, LH BMA→ VTA, CCLi, BL→LPO, LH, MH(sc)

LAVM→LEnt, MEnt, PRh, BLV→DStr, VStr,

BLP→LPO, LH

Basolateral nucleus (BL)

Ect IL, AID, AIV cortices.

TuO, Acb, CP,

subdivided into anterior

→Ammon’s Horn:

IPACl

BMA→AH, PV, LH, RCh,

and posterior (BLA, BLP)

CA1, CA2, CA3

→VP, HDB

SuM, Pe, PMV, PMD

→Sv, PaS

BLP→IPACl

BMP→By precommis. ST to → AH, PV, LH,

Ventral basolateral

BL→AID, AIP, Prl, M1,

nucleus (BLV)

M2, SM, PRh, and

BM→DStr, VStr: IPACl,

IL cortices

Acb, TuO, CP

Basomedial nuclei

PAG, and PBr

→Sv.

RCh, SuM, Pe, PMV, PMD, VMHC(d), VMHA (l), VHMVL,

(BM) subdivided into

→Ammon’s Horn:

LPA, MPA, MnPO,

anterior (BMA) and

CA1, CA3

Pc, SPa

posterior (BMP)

BLA→LEnt, IL AID, AIP BLP→LEnt(l), MEnt, IL.

By Postcommis. st:

Laterobasal group has

AID, AIP

RCh, LH, Mtu, PH, MM.

strong interconnections

→Sv, PaSpv

→Thalamus:

with cortical areas and

BLV→PIR, LEnt, MEnt,

BL→MD

striking output to ventral

AID, AIP, PRh cortices

BLV→MD

and dorsal striatum (VStr,

→ Sv. S, PaS

DStr), caudate–putamen,

→Ammon’s Horn: CA1,

BMA→same of BMP,

(CP), and olfactory

CA3

plus PT, CM, SPFPC

tubercle (TuO)

BMA→ Same of BLP

BMP→RE, PVT, MDm

(but rostral), BMP→ Paleocortex: PIR, SUB, CA1 Periallocortex: SUBv, LEnt, IL, PrL, AId, AIp Proisocortex: VISC., PRh, Ect, SM, TE, AU, PtAP (l) light; (m) moderate; (d) dense; (vd) very dense; (vl) very light; (pm) poor to moderate.

networks of fibers, allowing their easy identification (Fig. 13E). Heavy Metals The main IM cell mass and the cell masses separating the Ce nucleus from the AStr zone, as well as the cell masses separating the LBNC from the stria termi-

nalis and lateral ventricle, are very dense in Timm/ Danscher’s stain (Figs. 17C, 21C, 22B, and 30B). However, in contrast with it, the cell clusters between the LA nucleus and the external capsule, or between the striatum and the BLI nucleus, and those integrating the I group show a dense to very dense Timm/ Danscher’s reaction.

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Acetylcholinesterase

Acetylcholinesterase

In this type of preparation, the main IM and the cell masses separating the Ce nucleus from the AStr zone, as well as those separating the LBNC from the stria terminalis and from the lateral ventricle, are very poor in AChE reactivity. By comparison, a strong AChE reactivity is displayed by the intercalated cell clusters between the LA nucleus and the external capsule, or between the striatum and the BL nucleus, and the IP group (Figs. 17B, 22C, and 30A).

AChE material reveals also a strong activity in the AStr zone, although lower than in the rest of the striatum. The rostrally adjacent IPAC nucleus is similar in AChE staining, but perhaps slightly denser, in between that of the AStr area and that of the striatum proper. Comparing the AStr area with the CeL division, the almost complete lack of AChE activity, the consistently stronger Timm’s precipitate, and the exclusive cupric silver granular argyrophilia present in the CeL division preclude confusion about these different territories (Figs. 18A, 22C, and 24D).

The Amygdalostriatal Transition Zone (Astr) Topographic Landmarks

The Amygdaloid Intramedullary Gray (IMG)

The AStr transition zone encompasses the region of the striatum that is wedged between the Ce nucleus and the LBNC (Figs. 15I–15K in Alheid et al., 1995). In our earlier description of this area (de Olmos et al., 1985), we distinguished both an anterior and a posterior part of the AStr. For the reasons discussed in the most recent version of this chapter (Alheid et al., 1995), we have included the anterior (large-celled) part within the IPAC nucleus. The remaining posterior part has only an uncertain alliance with the central division of the extended amygdala, mainly on the basis of proximity, with little in the way of demonstrable interconnections (Gray et al., 1989; Jolkonnen et al., 1998) (Figs. 7D, 13, 18, 21C, 22, 24B, 24D, 24F, 31B, 31D, and 33A).

Topographic Landmarks What appears to be a ventral prolongation of the AStr zone, together with the numerous small clusters of intercalated cells, constitutes a significant portion of the IMG. The IMG is a thin cell and fiber plate that fills the corridor extending between the CeL division of the Ce nucleus and the LA nucleus and between the CeL division and the basolateral nucleus. An extension of this network appears to also run caudally and ventrally, where it is interposed between the BM and the Me nuclei, as well as between the BM nucleus and the AHI area. Fibers belonging to the lab run through the IMG and also highlight its medial boundary (Fig. 1, 7D, 22, 24, 31B, 31D, and 33A).

Cytoarchitectionics

Cytoarchitectonics

Its cell population consists mostly of cells that are smaller, more loosely packed, and more lightly stained than those in the remaining striatum (Figs. 15I–15K in Alheid et al., 1995), from which the AStr zone is discontinuously separated by small, transversely oriented bundles of fibers running along the border between the two structures. Ventromedially, it is also difficult to trace a boundary with the CeL division of the Ce nucleus except for the presence of small bundles of fibers interrupting the continuity between the two grisea. Ventrolaterally, the border of the AStr zone with the LBNC is easier to trace because of the interposition of the fiber-encapsulated IMG (Figs. 22A and 24B).

In addition to some of the intercalated cell masses, the IMG contains medium-sized, palely staining fusiformor oval-shaped neurons very loosely arranged in the spaces left free by the clusters of intercalated cells (Figs. 22A and 24B). As with the intercalated cell masses, the association of the intramedullary gray with a particular division of the amygdala is complicated. At the present time, insufficient data exist on the basis of histochemistry or tract-tracing experiments to clarify this point. The significance of understanding this difficult structure is underscored by the fact that it is apparently even more evident in the primate brain (de Olmos, 1990; Heimer et al., 1999).

Heavy Metals

Heavy Metals

Timm staining was useful for differentiation of the intensely labeled amygdalostriatal transition area from the dorsally located, lightly stained caudoputamen. The large-celled rostrolateral part of the amygdalostriatal transition area was also lightly stained (Figs. 21C, 22B, and 24F).

In Timm/Danscher’s preparations (Figs. 22B and 24F), this narrow plate of gray matter, whose thickness suffers a thinning in dorsoventral and rostrocaudal directions, shows variable degrees of stainability but still can be differentiated from the heavier pattern of silver deposits present in the capsular part of the Ce

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nucleus. However, within its dorsal and rostral expansions, where it merges dorsomedially with IPAC, or dorsorostrally with the AStr zone, it presents patches showing relatively light staining, lying beside others containing a denser silver precipitate. Laterally the much lighter Timm/Danscher staining of the BLA neuropil helps to trace its boundary with this nucleus.

Heavy Metals and Acetylcholinesterase

Acetylcholinesterase

Topographic Landmarks and Cytoarchitectonics

In AChE-stained preparations (Figs. 22C and 24D), rostrally, at the level of the anterior division of the BL nucleus, the IMG can be differentiated with difficulty from the very vigorous reactivity present in this division of the BL nucleus which overshadows the much lighter AChE activity detectable in the IMG. By contrast, such differentiation can be easily done with respect to the lateral division of the Ce nucleus which displays a completely negative staining reaction. More caudally, when the ventromedial subdivision of the LA nucleus makes its appearance, the IMG plate appears clearly delineated from both this subdivision and from the lateral division of the Ce nucleus, on account of its AchE reactivity which, although light to moderate, contrasts clearly with the almost totally negative picture shown by both neighboring structures.

The SV nucleus is a small ovoid group of cells located just beneath the floor of the lateral ventricle (i.e., the dorsal nucleus of Ju and Swanson (1989) or the subventricular nucleus of Moga et al. (1989)). In Nissl-stained sections, the cells are small, dark staining, and more or less closely packed together.

The Parastrial Nucleus (PS) Topographic Landmarks and Cytoarchitectonics The PS nucleus is a small lens-shaped nucleus located beneath the anterior commissure at the medial edge of the BSTMV subnucleus of the medial division of the BST nucleus.This nucleus consists of small- to mediumsized neurons oriented in a dorsomedial to ventrolateral direction (Fig. 15B in Alheid et al., 1995; and Figs. 13A, 13B, 31A, 32, and 33, this chapter).

The Bed Nucleus of the Anterior Commissure (BAC) Topographic Landmarks and Cytoarchitectonics The BAC nucleus is a dark-staining, compact group of cells located just lateral to the fornix as it crosses behind the anterior commissure (Fig. 15C in Alheid et al., 1995). In Nissl stains this group of small neurons appears to be similar in appearance to the intercalated cell clusters or to the juxtacapsular subnucleus of the BSTL division of the BST nucleus, although the cells of the former appear to be somewhat more darkly stained. However, mixed with the small cells are larger multipolar neurons, a feature that differentiates this conspicuous cell aggregate from the previously mentioned nuclei (Figs. 13B, 24, and 32).

These cells appear to be densely stained in both AChE and Timm/Danscher’s histochemically processed sections (Figs. 24C and 24E and 32, respectively).

The Subventricular Nucleus (SV)

Heavy Metals and Acetylcholinesterase In acetylcholinesterase sections, little or no reactivity is seen in this nucleus, whereas in Timm’s-stained sections a strong reaction comparable to that in the surrounding subdivisions of the BST nucleus is observed.

The Nucleus of the Commissural Component of the Stria Terminalis (CST) The CST nucleus is the same as our earlier bed nucleus of the commissural component of the stria terminalis (BCST) (de Olmos et al., 1985). We have made only a minor adjustment in the term used in order to apply the term used by Paxinos and Watson (1986). Topographic Landmarks and Cytoarchitectonics This nucleus, as its name indicates, appears as cell groups interspersed among the fibers of the commissural component of the stria terminalis (Fig. 32). These cell masses are made up of small, medium, and large cells that stain more lightly than those in the surrounding areas. The CST nucleus extends from the dorsolateral surface of the anterior commissure, becoming interposed between the posterior subdivision of the BSTM and the intermediate part of the BSTL. These cells maintain a close relationship with the commissural fibers of the stria terminalis, but do not possess any distinctive boundary. Rather, they are scattered among and around the fibers from which their name is derived. Heavy Metals With the Timm/Danscher’s stain, the cells of this nucleus appear to be embedded within a very dense band of silver precipitate. The zinc-rich terminals of the CST nucleus do not reach this cell corridor via the stria terminalis because the Timm’s-rich neuropil survives the transection of the ipsilateral stria terminalis (Fig. 32).

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Acetylcholinesterase The AChE histochemistry only lightly stained the CST nucleus.

The Fusifonn Nucleus (Fu) Although the fusiform nucleus has been considered to represent a component of the BST nucleus (Ju and Swanson, 1992; Moga et al., 1989), and some of its connectional relationships with some of its subnuclei may speak in favor of such an affiliation (Dong et al., 2001a), the fact that this cell mass does not receive a direct supply from the stria terminalis cannot be overlooked. Topographic Landmarks and Cytoarchitectonics The Fu nucleus (Figs. 13A, 13B, 31A, 32, and 33B) is a small cluster of dark-staining, medium to small neurons located ventral to the medial portion of the ventral part of the BSTM and below the BSTLV. The

TABLE 14

cells appear round to ovoid when viewed in frontal sections, but are elongated in horizontal sections, with their major axis oriented from rostromedial to caudolateral. This orientation follows the course of the nucleus, which stretches for a short distance across the ventral surface of the BSTL division (Ju and Swanson, 1989). Heavy Metals and Acetylcholinesterase In AChE sections, this small area is difficult to identify, but appears to have a moderately dense reaction. In Timm/Danscher-stained sections, the FS nucleus appear slightly stained.

Some Connectional Relationships of the Unclassified Nuclei Located within the Amygdaloid Complex or Maintaining Close Topographical Relationships with It or the Extended Amygdala

Unclassified Amygdaloid Nuclei Afferents Subcortical afferents

Cortical afferents

Telencephalon

Diencephalon

Brain stem afferents

Amygdalostriatal zone

Auditory belt (TeA

Posterior intralaminar

Dopaminergic cells in the

(AStr)

and AU) and visual

nuc. and MGM → AStr

VTA, SNc, and RRF →

Oc2L/Te2 → AStr

Astr, IMG, IM

griseum (IMG)

PRh, → Astr, IMG

Noradrenergic neurons

Intercalated cell masses

Dorsolateral and,

ventrolateral medulla

(IMm and IMp) and

ventrolateral (but not

oblongata → Astr, IMG, IM

interface islands

ventromedial) L. Ent

Amygdaloid intramedullary in the pons and

cortex → Astr, IMG Parastrial nucleus (PS)

and IM

Bed nucleus of the anterior

AIV/LO, PrL → Astr

commissure (BAC)

and IMG IL→ Astr, IMG and IM

Subventricular nucleus (SV) Nucleus of the commissural component of the stria terminalis (CST) Fusiform nucleus (Fu)

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TABLE 15

Unclassified Amygdaloid Nuclei Efferents To subcortical targets

Amygdalo striatal zone

Cortical targets

Telencephalon

Diencephalon

To brain stem targets

Fu → IL periallocortex

Astr → LA DL

Fu → rostral LH, LH

Fu → ventrolat. PGA,

BM, and PMCO

TC, PaAP, PaV, PaPo,

DRN, RRF, LDTg, etc.

No structure other

nuclei

etc., PeF, DMH, SuM

(AStr) Amygdaloid intramedullary

than the Fu nucleus

griseum (IMG)

would seem to send

Intercalated cell masses and

Fu → SL v/r, caudal

projections to any

AcbSH (d),

type of cortical field.

caudomedial Tu (vl)

(Im and Ip) and interface islands

Fu → APIR PM

Parastrial nucleus (PS)

Fu → CeM (Vd), CeL

Bed nucleus of the anterior

SLEAc (vl)

(vl), CeC (vl), and commissure (BAC) Fu → BSTMAm (vd), Subventricular nucleus (SV)

BSTMPl (vl), caudal BSTLP (m), BSTLV (l),

Nucleus of the commissural

PS,

component of the stria terminalis (CST)

Fu → PBL nuc.

Fusiform nucleus (Fu)

PS does not have significative projections. → amygdala

(l) light; (m) moderate; (d) dense; (vd) very dense; (vl) very light; (pm) poor to moderate.

Abbreviations Used in All Figures for Amygdala and Extended Amygdala AA, anterior amygdaloid area AA VL, ventrolateral part AA VM, ventromedial part AA D, dorsal part AA Sh, shell area Acb, nucleus accumbens AcbSh, shell area of accumbens ACo, anterior cortical nucleus AHi, amygdalohippocampal area AHiAL, anterolateral subdivision of amygdalohippocampal area AHiPM, posteromedial subdivision of amygdalohippocampal area AOT, nucleus of the accessory olfactory tract APir, amygdalopiriform transition area APir AL, anterolateral part APir PM, posteromedial part BAC, bed nucleus of the anterior commissure BLA, anterior basolateral nucleus BLP, posterior basolateral nucleus BLV, ventral basolateral nucleus BM, basomedial nucleus BMA, anterior basomedial nucleus

BMP, posterior basomedial nucleus BSTIA, intraamygdaloid bed nucleus of the stria terminalis BSTL, lateral bed nucleus of the stria terminalis BSTLI, lateral bed nucleus of the stria terminalis, intermediate part BSTLJ, lateral bed nucleus of the stria terminalis, juxtacapsular part BSTLP, lateral bed nucleus of the stria terminalis, posterior part BSTLV, lateral bed nucleus of the stria terminalis, ventral part BSTMA, anterior medial bed nucleus of the stria terminalis BSTMP, posterior medial bed nucleus of the stria terminalis BSTMPi, posterior medial bed nucleus of the stria terminalis, intermediate part BSTMPl, posterior medial bed nucleus of the stria terminalis, lateral part BSTMPm, posterior medial bed nucleus of the stria terminalis, medial part BSTMV, ventral medial bed nucleus of the stria terminalis BSTS, supracapsular bed nucleus of the stria terminalis BSTSl, lateral part BSTSm, medial part CeA, central amygdaloid nucleus CeL, lateral subdivision of central amygdaloid nucleus CeL, central part CeC, capsular part

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CeM, medial subdivision of central amygdaloid nucleus CeMad, anterodorsal part CeMav, anteroventral part CeMpv, posteroventral part Fu, fusiform nucleus IM, large intercalated cell mass IMG, Intramedullary gray I, Intercalated masses Ip, paracapsular, intercalated cell clusters IPAC, interstitial nucleus of the posterior limb of the anterior commissure IPACl, lateral part IPACm, medial part La, lateral nucleus LaDL, dorsolateral part of lateral nucleus LaVL, ventrolateral part of lateral nucleus LaVM, ventromedial part of lateral nucleus LOT, nucleus of the lateral olfactory tract MBFGC, Magnocellular Basal Forebrain Gray Complex MeA, medial amygdaloid nucleus MeAD, anterodorsal part MeAV, anteroventral part MePD, posterodorsal part MePDi, intermediate subnucleus of posterodorsal part MePDl, lateral subnucleus of posterodorsal part MePDm, medial subnucleus of posterodorsal part PMCo, posteromedial cortical nucleus PLCo, posterolateral cortical nucleus PLCoOL, posterolateral cortical nucleus, oral lateral part PLCoOM, posterolateral cortical nucleus, oral medial part PLCoCL, posterolateral cortical nucleus, caudolateral part PLCoCM, posterolateral cortical nucleus, caudomedial part PLCoPC, posterolateral cortical nucleus, parvocellular part SLEA, sublenticular extended amygdala SLEAc, central division of sublenticular extended amygdala SLEAm, medial division of sublenticular extended amygdala SV, subventricular nucleus

Aknowlegdments The authors thank Dr. George Paxinos, School of Psychology, The University of South Wales, Sydney, Australia, and Dr. John Jane and the Department of Neurological Surgery, University of Virginia Health System, Charlottesville, VA, USA, for their constant support and encouragement. We are especially grateful to Soledad De Olmos for her excellent technical support. This work was supported by the Instituto de Investigacion Médica Mercedes y Martín Ferreyra (INIMEC), Consejo Nacional de Investigaciones Científicas y Técnicas of Argentina (CONICET) PIP 2908/1073/01, the Fundación Interior Argentina (FUNINAR), and Secretaria de Ciencia y Técnica, Universidad Nacional de Córdoba, Argentina (Grant P 004/2001–2002).

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Woodhams, P. L., Roberts, G. W., Polak, J. M., and Crow, T. J. (1983). Distribution of neuropeptides in the limbic system of the rat: The bed nucleus of the stria terminalis, septum and preoptic area. Neuroscience 8, 677–703. Woulfe, J. M., Hrycyshyn, A. W., and Flumerfelt, B. A. (1988). Collateral axonal projections from A1 noradrenergic cell group to the paraventricular nucleus and bed nucleus of the stria terminalis in the rat. Exp. Neurol. 102, 121–124. Wray, S., and Hoffman, G. E. (1983). Organization and interrelationship of neuropeptides in the central amygdaloid nucleus of rat. Peptides 4, 525–541. Yamano, M., Hillyard, C. J., Girgis, S., MacIntryre, L., Emson, P. C., and Tohyama, M. (1988a). Presence of substance P-like immunoreactive neurone system from the parabrachial area to the central amygdaloid nucleus of the rat with references to coexistance with calcitonin gene-related peptide. Brain Res. 451, 179–188. Yamano, M., Hillyard, C. J., Girgis, S., MacIntryre, L., Emson, P. C., and Tohyama, M. (1988b). Projection of neurotensin-like immunoreactive neurons from the lateral parabrachial area to the central amygdaloid nucleus of the rat with reference to the coexistence with calcitonin gene-related peptide. Brain Res. 451, 179–188. Yasui, Y., Breder, C. D., Saper, C. B., and Cechetto, D. F. (1991a). Autonomic responses and efferent pathways from the insular cortex in the rat. J. Comp. Neurol. 303, 355–374. Yasui, Y., Saper, C. B., and Cechetto, D. F. (1991b). Calcitonin generelated peptide (CGRP) immunoreactive projections from the thalamus to the striatum and amygdala in the rat. J. Comp. Neurol. 308, 293–310.. Yeterian, E. H. and Van Hoesen, G. W. (1978). Cortico-striate proyections in the rhesus monkey: The organization of certain cortico-caudate connections. Brain Res. 138, 43–63. Yu, H. (1969). “The Amygdaloid Complex in the Rat” Master thesis, University of Ottawa, Ottawa, Canada. Zaborszky, L., Alheid, G. F., Beinfeld, M. L., Eiden, L. D., Heimer, L. and Palkovits, M. (1985) Cholecystokinin innervation of the ventral striatum. A morphological and radioimmunological study. Neuroscience 14, 427–453. Zahm, D. S., Jensen, S. L., Williams, E. S., and Martin, J. R. (1999). Direct comparison of projections from the central amygdaloid region and nucleus accumbens shell. Eur. J. Neurosci. 11, 1119–1126. Zardetto-Smith, A. M., Beltz, T. G., and Johnson, A. K. (1994). Role of the central nucleus of the amygdala and bed nucleus of the stria terminalis in experimentally induced salt appetite. Brain Res. 123–134.

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C H A P T E R

20 The Septal Region P. Y. RISOLD Laboratoire d’Histologie, Embryologie, Cytogénétique Faculté de Médecine et de Pharmacie Université de Franche-Comté Besançon Cedex, France

Many parts of the brain have been named on the basis of their shapes (hippocampus, amygdala), texture (striatum), location (paraventricular nuclei), and even color (substantia nigra), often at times when investigators had little idea of what are the basic constituents of this organ, namely, the neural cells (see Swanson, 2000a, 2000b). Thus, conforming to this general trend, the medial interventricular wall of the telencephalon, displaying no other particular shape or color, received the Latin name for “wall,” that is “septum” (septum pellucidum in human because of its translucent appearance; see Swanson et al., 1987; Jakab and Leranth, 1995; Swanson and Risold, 2000; Swanson, 2000a, 2000b, for additional historical overviews). At the turn of the 20th century, Cajal (1909–1911, 1995) identified several nuclei within this structure, but, most importantly, this brain area was recognized as bidirectionally connected with the hippocampus, although the ascending projections from the septum to the Ammon’s horn were only clearly demonstrated decades later (Daitz and Powel, 1954). The borders of the septal region are still discussed even if many controversial points have been settled during the last century. For example, it is now very widely accepted that the accumbens nucleus is not part of the septal region, but is a component of the striatum. The general scheme adopted here for the parceling of the septal region is based on Swanson and Cowan (Swanson and Cowan, 1979; Paxinos and Watson, 1986) with some slight modifications (Risold and Swanson, 1997a; Swanson

The Rat Nervous System, Third Edition

and Risold, 2000). Four groups of nuclei are described again based on their anatomical location within the septal region. The lateral group contains the lateral septal, septofimbrial, and septohippocampal nuclei. The medial septal complex (medial septal nucleus and nucleus of the diagonal band) forms the medial group. The posterior group is made of the triangular nucleus and the bed nuclei of the anterior commissure and of the stria medullaris. Finally, the bed nuclei of the stria terminalis (BST) form the ventral group. In a broader morphofunctional context, the septal region has been attached to the “limbic system” (Fig. 1), while more recently, parts of this region (BST) have been included into the “extended amygdala” that can be regarded as a particular division of the limbic system. Furthermore, cholinergic cells of the medial septal complex (medial septal and diagonal band nuclei) with cholinergic neurons in the substantia innominata and in the globus pallidus/basal nucleus of Meynerts form the basal forebrain cholinergic corticopetal system which projects topographically throughout the cortical mantle. Thus, from the preceding, two main considerations can be assessed: • We are still very confused about the organization of the septum. • Most of the septal region develops topographical morphofunctional links with, at least, two other parts of the telencephalon: the hippocampus and the amygdala.

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Copyright 2004, Elsevier (USA). All rights reserved.

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LIMBIC SYSTEM SEPTOHIPPOCAMPAL SYSTEM

PFC

Isocortex

HiF cortico

deep

Strv

Strd

ACB shell

Amy

EXTENDED AMYGDALA

LSC BST

GP

B

VP

SI

MSC SEPTAL REGION

CHOLINERGIC SYSTEM OF THE BASAL FOREBRAIN

FIGURE 1 Schematic representation of some of the current “systems” described in the telencephalon. See text for details. Amy, amygdala; B, basal nucleus of Meynert; BST, bed nuclei of the stria terminalis; GP, globus pallidus; HiF, hippocampal formation; LSC, lateral septal complex; MSC, medial septal complex; PFC, prefrontal cortex (symbolizing all cortical areas projecting in the ventral striatum); Strd, striatum (dorsal part); Strv, striatum (ventral part); VP, ventral pallidum

DEVELOPMENT OF THE SEPTAL REGION The majority of the data found in the literature concerns the lateral and medial septal nuclei. Both cell groups evolve from the septal ridge which forms the rostroventromedial wall of the telencephalic vesicle (Bayer, 1979a, 1979b, 1987; Alvarez-Bolado et al., 1995; Swanson and Risold, 2000). At early stages (e12) this septal anlage lies rostral to the striatal and pallidal ridges and ventral to the rostromedial cortical germinal neuroepithelium. The BST originates from a distinct anlage localized in the caudal portion of the anterior limb of the ventral angle (made by the ventral tip of the rostral horn of the lateral ventricle, see in Alvarez-Bolado et al., 1995). From Bayer (1987), the posterior part of the BST, corresponding roughly to the cell group posterior to the decussation of the anterior commissure, is generated from the very rostral portion of the posterior limb (just behind the interventricular foramen). Fusion of the right and left septal regions (e15) follows the dramatic development of the rest of the basal nuclei and precedes the development of the anterior commissure (e17, Bayer, 1979a, 1979b), as well as that of the other commissures that pass through or tether the septal region. The potential influence of septal neurons on the establishment of these commissures is largely unclear. Bayer and Altman, Chapter 2 (this volume), present in graphic form data for the birth dates of all forebrain regions. Most of the septal neurons are generated from e12 to e18 (Fig. 2) (Swanson and Cowan, 1976; Bayer,

1979a; Semba and Fibiger, 1988; Semba, 1992; AlvarezBolado et al., 1995). However, neurons of the medial, posterior, and ventral nuclear cell groups are born before the neurons of the lateral septal complex. Broadly, cells of the former are born between e12 and e16, while those in the later are generated between e14 and e18. The spatiotemporal patterns of genesis of these cell groups are distinct as well (Bayer, 1979a, 1979b, 1987). The medial septal nucleus and the BST expand following rostral and caudal gradients, while the lateral septal nucleus develops following a strictly outside-in gradient (thus, from medial to lateral), maybe benefiting of the enlarged septal germinal zone resulting from the genesis of the medial septal nucleus. The adult medial septal complex contains GnRH-producing neurons (Barry et al., 1973, 1985). These cells are born in the olfactory placode and migrate in association with the nervus terminalis to reach their final destinations in the septal region and the preoptic hypothalamus. In the rat medial septal complex, the first GnRH neurons are seen on e17 and reach the adult number on the day of birth (Jennes and Schwanzel-Fukuda, 1992). The development of the fiber tracts that characterize the septal region has been the object of relatively few studies. The stria terminalis and the stria medullaris appear early (e13) on the templates of atlases of the developing rat brain (Alvarez-Bolado and Swanson, 1996; Foster, 1998). The first component of the fornix could mostly be made of descending projections (around e16/e17; Linke and Frotscher, 1993). The postcommissural fornix has been shown to reach the

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e11 e12 e13 e14 e15 e16 e17 e18 e19 e20

p0

p5

p20

MS /DB TS / BAC BST LS fornix baskets FIGURE 2 Diagram schematizing the time laps of neuronal birth leading to the formation of the nuclei of the septal region, and the maturation time laps of the fornix system and of most ascending afferents to the lateral septal nucleus (baskets). BAC, bed nucleus of the anterior commissure; DB, nucleus of the diagonal band; LS, lateral septal nucleus; MS, medial septal nucleus; TS, triangular nucleus.

posterior hypothalamus a day or two before birth (Stanfield et al., 1987; Horton and Levitt, 1988; Stanfield and O’Leary, 1988). The precommissural component of the fornix appears to develop from e16 and to become abundant in the lateral septal nucleus only after birth (Linke et al., 1995). The first ascending monoaminergic (serotoninergic) fibers from the brain stem enter the septum from the medial forebrain bundle, and have been noticed on e15/16 in the ventral medial septal complex (Wallace and Lauder, 1983, 1992). However, most of these projections develop later (from e18) and sequentially following a medial to lateral gradient, particularly in the lateral septal nucleus (Verney et al., 1987; Kalsbeek et al., 1992; Dinopoulos et al., 1993). A putative relationship between the maturation of those projections arising mainly from the hypothalamus and the midbrain, with the development of the descending hippocampal inputs is unclear. It has been suggested that lateral septal cells begin differentiation as hippocampal afferents enter this nucleus (Bayer, 1979b). But extension of certain catecholaminergic and peptidergic projections occurs a few days later (from P0 to P6 – Verney et al., 1987; Dinopoulos et al., 1993) and appears concomitant with lateral septal cell maturation (decrease in neuronal density, increase in cell body size, differentiation of some intracytoplasmic organites) (Verney et al., 1987). The adult aspect of basketlike terminals is reached by the end of the third postnatal week. However, this last phenomenon can be modified by lesion of hippocampal afferents, suggesting that direct or indirect interactions may exist between both types of afferents for their harmonious development (Raisman, 1969; Moore et al., 1971; Ueda et al., 1991).

Correlation between the establishment of these connections and expressions of particular behaviors in the young rat is not clear. Topographically organized connections between the hippocampus and the amygdala with respectively the lateral septal nucleus and the BST, and between these structures and the brain stem (hypothalamus and mesencephalon), suggest that specific circuits are involved in the expression of specific behaviors (Swanson and Risold, 2000; see further). It could be interesting to verify how early social interactions with the mother and littermates help in shaping these circuits.

MORPHOLOGICAL OVERVIEWS AND CYTOARCHITECTURE OF THE SEPTAL NUCLEI There is no doubt about the telencephalic, but subcortical nature of the septal region, therefore implying that this structure is part of the cerebral nuclei. Several myelinated fiber tracts (Fig. 3) reach, pass through, or bound the septal region (Paxinos and Watson, 1986; Swanson et al., 1987; Swanson, 1998). These fiber tracts often carry afferents to, or efferents from, the septal nuclei. They, in part, shape this region. As previously mentioned, some of these fiber tracts are commissures. Usually, they originate and end in other telencephalic (cortical) structures. These commissures are (i) the corpus callosum that bounds rostrally and dorsally the septal region, apparently without any functional interaction; (ii) the ventral hippocampal commissure that passes (parts of it) through the triangular nucleus, but functional relationships between these

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IG

A

IG

B

cc cc SHi LS SHi

df

LS pf

f

pf LV

SFi

SFi

LV

vhc

fi

TS TT

C

ic

fi

vhc

st Acb

D

SI Acb

SI

ICjM db LS

db

CPu

LV

LPO MS

MS

f D3V

ac BST st sm

GP

st

D3V

ic THd

f

BST st BAC ic sm

Rt

Rt

FIGURE 3 A series of line drawings of horizontal sections through the septal region illustrating its main nuclear organization. From dorsal (A) to ventral (D). ac, anterior commissure; Acb, nucleus accumbens; BAC, bed nucleus of the anterior commissure; BST, bed nuclei of the stria terminalis; cc, corpus callosum; CPu, caudoputamen; db, diagonal band; df, dorsal fornix; f, fornix; fi, fimbria; GP, globus pallidus; ic, internal capsule; ICjM, major island of calleja; IG, induseum griseum; LPO, lateral preoptic area; LS, lateral septal nucleus; LV, lateral ventricle; MS, medial septal nucleus; pf, precommissural fornix; Rt, reticular nucleus thalamus; SFi, septofimbrial nucleus; SHi, septohippocampal nucleus; SI, substantia innominata; sm, stria medullaris; st, stria terminalis; THd, thalamus dorsal; TS, triangular nucleus; TT, tenia tecta; vhc, ventral hippocampal commissure; D3V, third ventricle.

elements are unclear, although Cajal (1909–1911, 1995) described some collaterals ending in this nucleus from that commissure (see as well Swanson and Cowan, 1979); and (iii) finally, the anterior commissure that passes through the BST. The anterior commissure is made of one component originating and ending in the anterior olfactory nuclei and with which the BST appears to have little interaction, one originating from more temporal telencephalic structures including the piriform cortex, and a third from several amygdalar nuclei (nucleus of the lateral olfactory tract and basolateral, basomedial, and cortical nuclei) (Horel and Stelzner, 1981; Jouandet and Hartenstein, 1983; Yajeya et al., 1987). These last projections usually course through the stria terminalis and then join the decussation of the anterior commissure to innervate the contralateral BST (Krettek and Price, 1978; Canteras et al., 1992a, 1995; Petrovich et al., 1996; Dong et al., 2001).

Afferents and efferents of the septal region take four main routes: the fornix, the stria terminalis, the medial forebrain bundle, and the stria medullaris (Fig. 4). The lateral and medial groups are connected with the hippocampus through the fimbria/fornix, the ventral group is connected with the amygdala through the stria terminalis, and the caudal group projects through the stria medullaris. All septal nuclei receive abundant afferents from the medial forebrain bundle, and most of them send descending projections through it as well. Finally, the BST is bound laterally by the internal capsule and caudally by the postcommissural fornix and the dorsally directed fibers of the stria medullaris originating in anterior parts of the lateral hypothalamic area and external parts of the globus pallidus. Nine nuclei (or collection of nuclei in the case of the BST) are registered in the septal region that can be gathered into four groups. A complete description of

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precommissural fornix

fornix fimbria

stria terminalis

? Lateral Group

Medial Group

Ventral Group

Posterior Group

medial forebrain bundle

stria medullaris

FIGURE 4 General organization of the connections of the septal region. See text for detail.

the cytoarchitecture of the septal region can easily be found in the literature, and only the prominent features are discussed here.

The Lateral Group The lateral septal and septohippocampal nuclei have long been thought to be the only members of this group. However, the septofimbrial nucleus, on the basis of its connections, can also be categorized as a nucleus of the lateral group (Risold and Swanson, 1997b). The lateral septal nucleus is the largest nucleus of the septum. On Nissl-stained sections, authors have recognized that this nucleus is not homogeneous (Swanson and Cowan, 1979). Analysis of cell size and density shows differences following the dorsoventral or the lateromedial axis, and usually authors distinguish in this nucleus a dorsal, an intermediate, and a ventral part (Swanson and Cowan, 1979; Paxinos and Watson, 1986). However, some authors have acknowledged that “the boundaries between the various parts are not always obvious in routine histological preparation” (Swanson and Cowan, 1979). Furthermore, chemoarchitecture and patterns of connections do not respect most of these boundaries (Staiger and Nürnberger, 1991a, 1991b; Risold and Swanson, 1997a, 1997b). Other types of organization have been recently proposed which I will come back to later. Several cell types have been described in the lateral septal nucleus after Golgi impregnation (Alonso and Frotscher, 1989a, 1989b; Jakab and Leranth, 1995). All of them have their dendrites covered with spines. It is unfortunately unknown whether the septofimbrial and septohippocampal neurons share the same characteristic. A particular class of lateral septal neurons exhibits somatic spines, and they have been called “somatospiny neurons” (Jakab and Leranth, 1990a, 1990b). No

strictly local interneurons have been described in the lateral septal nucleus, but axons of some neurons in dorsal parts of this nucleus exhibit local collaterals, suggesting local inhibition circuits (Phelan et al., 1989). Very little is known about the cytoarchitecture of the septofimbrial and septohippocampal nuclei. Septofimbrial neurons are on average larger than those of the lateral septal nucleus and are scattered within myelinated bundles of the precommissural fornix (Swanson and Cowan, 1979). The septohippocampal nucleus is a very elongated structure localized adjacent to the midline, dorsal in the lateral septal nucleus throughout the length of this nucleus (Swanson and Cowan, 1979; Paxinos and Watson, 1986; Swanson, 1998).

The Medial Group At least five cell types have been described in the medial septal complex (Brauer et al., 1988; Dinopoulos et al., 1988; Jakab and Leranth, 1995). Most of them are dendritic spine free; however, some of these cells are characterized by their large size and dark thionine stain. They correspond to the cholinergic neurons contained by those nuclei. The medial septal complex contains as well a population of small spindle-shaped neurons that may correspond to the GnRH neurons found there. Cytoarchitectonic boundaries between the medial septal nucleus and the nucleus of the diagonal band are not clear, and both nuclei are traversed by myelinated axons of the diagonal band (Swanson et al., 1987).

The Posterior Group All three nuclei of this group (triangular nucleus and bed nuclei of the anterior commissure and of the stria medullaris) have been the object of very few studies

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and are still enigmatic (Sturrock, 1977; Swanson and Cowan, 1979). The bed nucleus of the anterior commissure is sometimes related to the ventral group because this nucleus is adjacent to the BST. However, connectional data clearly indicate that this nucleus is more likely part of the posterior group as it projects, along the triangular nucleus, through the stria medullaris (Swanson and Cowan, 1979; Staines et al., 1988). The triangular nucleus and the bed nucleus of the anterior commissure are made of densely packed cells between small bundles of the ventral hippocampal commissure for the former, and at the angle formed by the fornix and the anterior commissure for the latter. Cells of the bed nucleus of the stria medullaris, localized at the angle formed by the fornix and the stria medullaris, are small and their cytoplasm is lightly stained by thionine (Cajal, 1909–1911, 1995; Risold and Swanson, 1995).

The Ventral Group The BST is the only component of this group. Many studies have focused on the cytoarchitecture of this nucleus or group of nuclei, and it is obvious that there is a great diversity of opinion concerning its (their) organization. One of the main reasons for this is that nobody has yet made a thorough analysis of the connections of the BST with regard to its fine cytoarchitecture as well as with regard to its chemoarchitecture. In particular, the exact topography of connections between many nuclei or subnuclei of the BST, the amygdala, the hypothalamus, and the brain stem is still unclear. There is no doubt that such a study will come out shortly, which will shed a new light on this structure (see Dong et al., 2001). But for now, the author does not wish to add to this confusion, this position being reinforced by the fact that elements for a juxtaposition of the existing literature are provided in another chapter in this book (De Olmos et al., Chapter 19).

THE CHEMOARCHITECTURE OF THE SEPTAL REGION A huge amount of information has been published concerning the distribution of neuropeptides and neurotransmitters in the septal region. Most of these molecules are expressed by neurons or by afferent projections following patterns that do not respect cytoarchitectonic boundaries, even more so than in many other brain regions. As a first general statement, it is important to note that the whole septal region, like the rest of the cerebral nuclei, is rich in GABAergic neurons that express glutamic acid decarboxylase (Köhler and Chan-Palay,

1983; Panula et al., 1984; Onteniente et al., 1986; Feldblum et al., 1993; Risold et al., 1997a). However, obviously with regard to the cytoarchitecture, GABAergic neurons do not form a homogeneous population, and non GABAergic neurons containing other neurotransmitters are found in the septal region.

The Lateral Group Since 30 years ago, the band-shaped distribution patterns of neurotransmitters, neuropeptides, and several proteins in the lateral septal nucleus has attracted the attention of many investigators. Most of these patterns do not respect the classical cytoarchitectonic divisions of the nucleus and are a clear indication of the extreme heterogeneity of this structure. In recent analyses of the chemoarchitecture of the rat lateral septal nucleus (Risold and Swanson, 1997a; Paxinos et al., 1999), up to 20 divisions were identified while cytoarchitecture allows the description of only three. Paxinos et al. (1999) divided the caudal intermediate part of the lateral septal nucleus into four ventrodorsally oriented layers, corresponding to the alternate distribution of calbindin and tyrosine hydroxylase. Risold and Swanson (1997a, 1997b) confronted distribution patterns of around 15 neuropeptides or proteins in the lateral septal nucleus and patterns of connections of this nucleus with the hypothalamus. There is neither need nor space here to enter deep into these analyses and the reader is referred to the original paper or book (Risold and Swanson, 1997a; Paxinos et al., 1999). Broadly, both parcelations are not only based on cytoarchitectonic ground (after a Nissl stain) on which, then, authors fit in observed patterns, but also they derive from these patterns. As more work is done in this field, modifications of these nomenclatures may be needed. Nonetheless, to associate experimental events with a particular chemically defined part of the nucleus may yield a more reproducible estimation of the location of that event (for example, distribution of c-fos-labeled nuclei) in this otherwise large and poorly differentiated, but extremely heterogeneous, nucleus. Following the chemoarchitectural analysis of Risold and Swanson (1997a), three large parts (rostral, caudal, and ventral) were described that only partially overlap with the three classic dorsal, intermediate, and ventral parts (Table 1; Figs. 5 and 6). The distribution of several molecules helps in characterizing each one of them. In brief, using in situ hybridization, the rostral part is best characterized as containing abundant enkephalinergic and/or neurotensinergic neurons, the caudal as containing abundant somatostatinergic neurons, and the ventral as containing a significant labeling for the estrogen receptor α (Table 1). Compared with

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TABLE 1

Major Subdivisions of the Lateral Septal Complex

Subdivisions of the LS

Ascending inputs (immuno.)

Cells (+GABA) (in situ hybridization)

Part

Zone

Region

LSc

d

r

—

SS, MR

d

—

SS, DYN, MR

l

DA

SS, MR

v

—

SS, SP, MR

d

DA, SP

SS, MR

v

SP, DA

SS, MR, AR

—

SS, MR

d

SP, VAS, GAL

SS, AR, MR

v

m

Domain

i l

v LSv LSr

m

vl

d

DA

ENK, NT, SP ENK, NT

c

NE/E

ENK, NT

m

—

ENK, NT

—

ENK, NT, GRH

r

d

l m l

SS, AR, MR ER, AR

NE/E

v

v dl

DA, VAS, GAL, SP SP

SP

ENK, NT, GRH

d

CGRP, SP

ENK, NT, SS

v

CGRP, SP

ENK, NT, SS

d

ENK, CGRP

ENK, NT, SS

v

ENK

ENK, NT, SS

Note. LS, lateral septal nucleus; LSc, caudal part of LS; LSc.d, dorsal zone of LSc; LSc.d.d, dorsal region of LSc.d; LSc.d.l, lateral region of LSc.d; LSc.d.r, rostral region of LSc.d; LSc.d.v, ventral region of LSc.d; LSc.v, ventral zone of LSc; LSc.v.i, intermediate region of LSc.v; LSc.v.l, lateral region of LSc.v; LSc.v.l.d, dorsal domain of LSc.v.l; LSc.v.l.v, ventral domain of LSc.v.l; LSc.v.m, medial region of LSc.v; LSc.v.m.d, dorsal domain of LSc.v.m; LSr, rostral part of LS; LSr.dl, dorsolateral zone of LSr; LSr.dl.l, lateral region of LSr.dl; LSr.dl.l.d, dorsal domain of LSr.dl.l; LSr.dl.l.v, ventral domain of LSr.dl.l; LSr.dl.m, medial region of LSr.dl; LSr.dl.m.d, dorsal domain of LSr.dl.m; LSr.dl.m.v, ventral domain of LSr.dl.m; LSr.m, medial zone of LSr; LSr.m.d, dorsal region of LSr.m; LSr.m.v, ventral region of LSr.m; LSr.m.v.c, caudal domain of LSr.m.v; LSr.m.v.r, rostral domain of LSr.m.v; LSr.vl, vemtrolateral zone of LSr; LSr.vl.d, dorsal region of LSr.vl; LSr.vl.d.l, lateral domain of LSr.vl.d; LSr.vl.d.m, medial domain of LSr.vl.d; LSr.vl.v, ventral region of LSr.vl; LSv, ventral part of LS. (Adapted from Risold, P. Y., and Swanson, L. W., 1997, Chemoarchitecture of the rat lateral septal nucleus, Brain Res. Rev. 24, 91–113, with permission from Elsevier Science.)

the cytoarchitectonic divisions of the lateral septal nucleus, the rostral part encompasses the cytoarchitectonic rostral–ventral and rostral–intermediate parts, the caudal encompasses the cytoarchitectonic dorsal and caudal–intermediate parts, and the ventral is limited to the cytoarchitectonically very well differentiated caudal–ventral part. The caudal and rostral parts are further divided (Table 1, Fig. 5). It would be very fastidious here to describe all these patterns, and so the reader is referred to the original paper (Risold and Swanson, 1997a). However, it is worth mentioning that all these divisions are based on the laminated distributions of chemically defined afferents (e.g., CGRP, enkephalin, DBH, TH, substance P; Fig. 7) that superimpose on patterns of expression of receptors (e.g., receptors to sexual and mineralocorticoid hormones— in Risold and Swanson, 1997a). Furthermore, a good correlation appears to exist between the layers reported by Paxinos et al. (1999) and several of the divisions

adopted by Risold and Swanson (1997a). In particular, the LSc.v.l, LSc.v.i, and LSc.v.m of Risold and Swanson (1997a—see Fig. 5 and the legend of Table 1 for the explanation of these abbreviations) mostly correspond to the layers 1, 2, and 3 of Paxinos et al. (1999). It is a matter of fact that, although the two studies are mostly based on different chemical markers, the patterns of these markers are often very similar (for example, compare the distribution of the peptide substance P in the study of Risold and Swanson (Fig. 7a) and the distribution of the protein calbindin in the work of Paxinos et al., 1999). Like the caudal part of the lateral septal nucleus, the septohippocampal nucleus contains many somatostatinexpressing neurons, but both nuclei can easily be differentiated because the latter contains many neurons expressing neuropeptide Y (Risold and Swanson, 1997a). Finally, little is known about the chemoarchitecture of the septofimbrial nucleus. Like the rostral part of the

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FIGURE 5 Series of drawings arranged from rostral (A) to caudal (I). The right side of each drawing illustrates the classical (from the cytoarchitecture) dorsal, intermediate, and ventral parts of the lateral septal nucleus; the left side illustrates the new divisions (from the chemoarchitecture). See Table 1 for the nomenclature. (Adapted from Risold, P. Y., and Swanson, L. W., 1997, Chemoarchitecture of the rat lateral septal nucleus, Brain Res. Rev. 24, 91–113, with permission from Elsevier Science.)

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FIGURE 6 Distribution of in situ hybridization labeling with probes for enkephalin (A–E), neurotensin (B–F), and somatostatin (C–G) on adjacent sections through a rostral (A–D) and a caudal (E–H) levels of the lateral septal nucleus. (D–H) Adjacent Nissl-stained sections. Bar = 500 μm. (Adapted from Risold, P. Y., and Swanson, L. W., 1997, Chemoarchitecture of the rat lateral septal nucleus, Brain Res. Rev. 24, 91–113, with permission from Elsevier Science.)

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FIGURE 7a Series of line drawings illustrating the distribution of immunohistochemical labeling for substance P (7a, left), enkephalin (7b, left), CGRP (7b, right), DBH (7c, left), and TH (7c, right). In 7a, the right side of the drawings illustrates the classical (from the cytoarchitecture) dorsal, intermediate, and ventral parts of the lateral septal nucleus. ac, anterior commissure; Acb, accumbens nucleus; BST, bed nucleus of the stria terminalis; cc, corpus callosum; CPu, caudoputamen nucleus; f, fornix; IG, induseum griseum; LV, lateral ventricle; MS, medial septal nucleus; SFi, septofimbrial nucleus; TS, triangular nucleus of the septum; TTd, tenia tecta (dorsal). (Adapted from Risold, P. Y., and Swanson, L. W., 1997, Chemoarchitecture of the rat lateral septal nucleus, Brain Res. Rev. 24, 91–113, with permission from Elsevier Science.)

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left : ENK ; right : CGRP Fibers

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FIGURE 7b, cont’d Series of line drawings illustrating the distribution of immunohistochemical labeling for substance P (7a, left), enkephalin (7b, left), CGRP (7b, right), DBH (7c, left), and TH (7c, right). In 7a, the right side of the drawings illustrates the classical (from the cytoarchitecture) dorsal, intermediate, and ventral parts of the lateral septal nucleus. ac, anterior commissure; Acb, accumbens nucleus; BST, bed nucleus of the stria terminalis; cc, corpus callosum; CPu, caudoputamen nucleus; f, fornix; IG, induseum griseum; LV, lateral ventricle; MS, medial septal nucleus; SFi, septofimbrial nucleus; TS, triangular nucleus of the septum; TTd, tenia tecta (dorsal). (Adapted from Risold, P. Y., and Swanson, L. W., 1997, Chemoarchitecture of the rat lateral septal nucleus, Brain Res. Rev. 24, 91–113, with permission from Elsevier Science. Continued

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left : DBH ; right : TH Fibers

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FIGURE 7c, cont’d Series of line drawings illustrating the distribution of immunohistochemical labeling for substance P (7a, left), enkephalin (7b, left), CGRP (7b, right), DBH (7c, left), and TH (7c, right). In 7a, the right side of the drawings illustrates the classical (from the cytoarchitecture) dorsal, intermediate, and ventral parts of the lateral septal nucleus. ac, anterior commissure; Acb, accumbens nucleus; BST, bed nucleus of the stria terminalis; cc, corpus callosum; CPu, caudoputamen nucleus; f, fornix; IG, induseum griseum; LV, lateral ventricle; MS, medial septal nucleus; SFi, septofimbrial nucleus; TS, triangular nucleus of the septum; TTd, tenia tecta (dorsal). (Adapted from Risold, P. Y., and Swanson, L. W., 1997, Chemoarchitecture of the rat lateral septal nucleus, Brain Res. Rev. 24, 91–113, with permission from Elsevier Science.

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lateral septal nucleus, it contains many enkephalinergic neurons (Fig. 6), but this nucleus is quite large and may not be homogeneous.

The Medial Group The medial septal nucleus and the nucleus of the diagonal band contain an abundant population of cholinergic neurons (Fig. 8). However, if the existence of this population appears to be the marking trait, the chemoarchitecture of the nuclei of the medial group is far more complex. Broadly, at least three heterogeneous neuronal types can be described. The cholinergic neurons, which contain the enzymes choline acetyltransferase and acetylcholinesterase, form one of these populations. Their existence was first suggested by Shute and Lewis (1967), but their cholinergic nature was ascertained later (for review Kàsa, 1986; Woolf, 1991; Butcher, 1995; Butcher and Woolf, Chapter 35, this volume). Various proportions of the corresponding neurons coexpress other neurotrans-

SHi

LS

MS ac

FIGURE 8 Line drawing illustrating the distribution of cholinergic neurons (black dots), parvalbumin-containing cells (open dots), and GnRH-producing neurons (black squares) at one mid-rostrocaudal level of the medial septal complex. ac, anterior commissure; LS: lateral septal nucleus; MS, medial septal nucleus; SHi, septohippocampal nucleus.

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mitters/neuropeptides and/or receptors (Semba, 2000). For example, cholinergic neurons contain glutamate (Forloni et al., 1987), nitric oxide (Schober et al., 1989; Geula et al., 1993; Kitchener and Diamond, 1993), neuropeptides (in particular galanin; Melander et al., 1985; for a recent review, see Semba, 2000), and receptors (for example, to NGF, Gibbs and Pfaff, 1994; Sobreviela et al., 1994). Several studies have attempted to analyze the coexpression of these molecules in cholinergic neurons (Melander et al., 1985; Pasqualotto and Vincenty, 1991; Sobreviela et al., 1994, 1998) or tried to determine their possible interactions with acetylcholine (Lamour and Epelbaum, 1988; De Wied, 1997; Planas et al., 1997). However, the general picture concerning these coexpressions and interactions is still rather unclear (Semba, 2000). GABAergic neurons form a second chemically defined large group of neurons. Most of them express peptides and/or other proteins. Of those, calciumbinding proteins and trophic factors have been the most studied (Jakab and Leranth, 1995; Semba, 2000). Parvalbumin and calbindin D-28k as well as NGF are specific of subsets of GABAergic neurons (Freund, 1989; Celio, 1990; Kiss et al., 1990; Brauer et al., 1991; Jakab and Leranth, 1995; Lauderborn et al., 1995; Paxinos et al., 1999). Some GABAergic neurons are local interneurons while others project in the cortex (Brauer et al., 1991; Jakab and Leranth, 1995), further implying that GABAergic neurons do not form a homogeneous cell “class” or “population.” Finally, GnRH neurons can be gathered in a third class. They are scattered within the medial septal complex and, more caudally, are found in the hypothalamic preoptic region. Many of them are neuroendocrine (Barry et al., 1973, 1985). The medial septal complex, through the medial forebrain bundle, receives from the brain stem abundant afferents containing GABA, glutamate, acetylcholine, serotonine, dopamine, and norepinephrine (Vertes, 1988; Panula et al., 1989; Carnes et al., 1990; Cullinam and Zaborzky, 1991; Zagon et al., 1994; Gaykema and Zaborszky, 1996; Kia et al., 1996; Leranth and Vertes, 1999). Many of these projections contain peptides as well (Woodhams et al., 1983). The topographical arrangement of these afferents is not yet clear. Many of them pass through these nuclei to reach the lateral septal nucleus or continue through the fornix/fimbria to enter the hippocampus, but often they innervate en passant neurons in the medial septal complex.

The Posterior Group These nuclei appear to be far less rich in terms of chemical content than the rest of the septal region. The

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triangular nucleus contains a large population of neurons expressing calretinin (Fig. 9), calbindin, and the enzyme adenosine deaminase (Staines et al., 1988; Wilson and Kawaja, 1996; Paxinos et al., 1999). Enkephalin has been described in neurons of all three nuclei, and, indeed, this peptide is at the origin of the description of the bed nucleus of the stria medullaris (Risold and Swanson, 1995). The posterior nuclei receive several solid afferents from the brain stem. In particular, the bed nucleus of the anterior commissure contains very abundant noradrenergic projections (labeled by an antibody against DBH; Fig. 7; Swanson and Hartman, 1975). Furthermore, this nucleus and the triangular nucleus contain very dense cholinergic projections labeled for the enzymes choline acetyltransferase and acetylcholinesterase (Paxinos and Watson, 1986; personal observation).

The Ventral Group The chemoarchitecture of the BST, as is its cytoarchitecture, is very complex (Woodhams et al., 1983; Moga et al., 1989; Ju et al., 1989; Arluison et al., 1994) and would need a comprehensive new review. At least as many peptides are expressed in the BST as in the lateral septal nucleus. However, they often respect bound-

A

aries of specific cell groups of the BST. For example, substance P is intensely expressed in the BSTPM (nomenclature from Paxinos and Watson, 1986, or principal nucleus from Swanson, 1998). Interestingly, this nucleus is tightly connected with the posterodorsal part of the medial nucleus of the amygdala, and with the hypothalamic anteroventral periventricular nucleus (ventromedial preoptic), the medial preoptic nucleus, and the ventral premammillary nucleus (Simerly and Swanson, 1988; Canteras et al., 1992b; Hutton et al., 1998). These cell groups express high levels of substance P, but each one of them also expresses high levels of estrogen and androgen receptors and has been experimentally involved in reproductive functions (Simerly et al., 1990; Larsen, 1992; Risold et al., 1997; Simerly, 1998). More lateral parts of the BST are connected with other amygdalar and hypothalamic/brain stem nuclei. Unfortunately, the organization of their connections and their functions are still often unclear.

CONNECTIONS OF THE SEPTAL REGION General connectional principles resulting from our actual understanding of the septal organization were

B

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ac

FIGURE 9 (A) Photomicrograph of the distribution of calretinin-containing neurons in the triangular nucleus and in the bed nucleus of the anterior commissure. (B) Photomicrograph of the adjacent Nissl-stained section. Scale bar = 200 μm. ac, anterior commissure; BAC, bed nucleus of the anterior commissure; BST, bed nuclei of the stria terminalis; f, fornix; TS, triangular nucleus.

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specified previously (Fig. 4). However, before considering functions of the septal region, a more detailed account of its afferents and efferents is obviously necessary.

The Lateral Group Nuclei of the lateral group are characterized by massive glutamatergic afferents from the pyramidal cell layers of the Ammon’s horn and subiculum and by massive bidirectional connections with the rostral brain stem, particularly with the hypothalamus and the ventral midbrain (Raisman, 1966a, 1966b; Meibach and Siegel, 1977; Swanson and Cowan, 1979; Swanson et al., 1987; Staiger and Nürnberger, 1989, 1991a; Risold and Swanson, 1996, 1997b). However, the different nuclei or parts of the lateral group are differentially connected with these structures. This has been well demonstrated for the lateral septal nucleus already during the 70s (Meibach and Siegel, 1977; Swanson and Cowan, 1976, 1979) and more recently refined using modern tract-tracing procedures in the rat and guinea pig (Staiger and Nürnberger, 1989, 1991a; Risold and Swanson, 1996, 1997b). Descending Projections from the Hippocampal Formation Descending hippocampal afferents to the lateral septal nucleus essentially originate in the Ammon’s horn and subiculum and run through the fimbria and then the precommissural limbs of the fornix (see as well Chapter 21, this volume, Witter and Amaral). It is now well established that these projections are excitatory and use glutamate as neurotransmitter (Walaas and Fonnum, 1980; Joëls and Urban, 1984a, 1984b), as well as maybe CCK (Greenwood et al., 1981). These projections are topographically organized following both a dorsoventral (Siegel et al., 1974; Swanson and Cowan, 1976; Swanson, 1977; Swanson and Cowan, 1979) and a rostrocaudal (Staiger and Nürnberger, 1989; Risold and Swanson, 1996, 1997b) axis. In particular, the septal half of the hippocampus innervates the dorsomedial pole of the lateral septal nucleus, maybe including the septohippocampal nucleus. The ventral two-thirds of the lateral septal nucleus receive afferents from the temporal half of the hippocampus. These projections appear to respect the organization of the lateral septal nucleus in at least two main parts, with a rostral part receiving mainly ipsilateral afferents from the hippocampal field CA1 and the subiculum and a caudal part targeted by bilateral afferents from the field CA3 (Risold and Swanson, 1996, 1997b). However, projections from the fields CA1 and CA3 overlap in the most lateral parts of the rostral lateral septal nucleus (Risold and Swanson, 1997b). The

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chemoarchitectonic ventral part of the lateral septal nucleus is targeted by axons originating in the temporalmost (or ventral-most) tip of the fields CA1/ subiculum (see as well Canteras and Swanson, 1992). Finally, both the septohippocampal and septofimbrial nuclei are in a good position to receive inputs from the precommissural fornix, unfortunately, they have not yet been clearly shown. Bidirectional Connections with the Hypothalamus and Midbrain The ventral part of the lateral septal nucleus appears to be directly connected with the periventricular zone of the hypothalamus (Fig. 10) (Oldfield and Silverman, 1985; Staiger and Wouterlood, 1990; Staiger and Nürnberger, 1991a; Varoqueaux and Poulain, 1994; Risold and Swanson, 1997b), and, through these projections, to be directly involved in the control of neuroendocrine and autonomic responses. More extensive projections have been described in the medial and lateral zones of the hypothalamus (Fig. 10). These projections are topographically organized from the rostral and caudal parts of the lateral septal nucleus. Recent evidences suggest that the nuclei of the hypothalamic medial zone and adjacent parts of the perifornical region of the lateral hypothalamic area are massively interconnected and form intrahypothalamic circuits involved in ingestive, reproductive, and defensive behaviors (Risold et al., 1997; Swanson, 2000a). Recent anatomical and fos studies have shown that distinct zones or regions of the rostral part of the lateral septal nucleus are selectively connected with specific components of these intrahypothalamic circuits (Fig. 11) (Risold and Swanson, 1997b; Varoqueaux and Poulain, 1998). The caudal part of the lateral septal nucleus, along the septohippocampal and septofimbrial nuclei, projects massively through lateral components of the lateral hypothalamic area and terminates in the lateral supramammillary nucleus, as well as, to a lesser extent, in the ventral tegmental area (Fig. 10). The lateral hypothalamic area and the supramammillary nucleus reciprocate these projections, at least to the caudal lateral septal nucleus because the specific afferents to the septohippocampal and septofimbrial nuclei are still unclear (Risold and Swanson, 1997b). The caudal part of the lateral septal nucleus is characterized by massive afferents from several nuclei of the brain stem including the ventral tegmental area and the laterodorsal tegmental nucleus. These afferents are well differentiated because they correspond to the dense dopaminergic and cholinergic band-shaped terminal fields in the caudal lateral septal nucleus (Staiger and Nürnberger, 1989; Risold and Swanson, 1997a, 1997b).

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arousal, behavioral activation

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FIGURE 10 Model of circuitry involving the hippocampal formation, LS, and hypothalamus. See text for details. CA1–3, Ammon’s horn, field 1 or 3; CING, cingular cortex; DG, gyrus dentatus; ENT, entorhinal cortex; LC, locus ceruleus; LDT, laterodorsal tegmental nucleus; LH, lateral hypothalamic area; LPO, lateral preoptic area; LSc-r-v, caudal–rostral–ventral parts of the lateral septal nucleus; MEZ, medial zone of the hypothalamus; MS, medial septal nucleus; NDB, nucleus of the diagonal band; Palv, ventral pallidum; PVz, periventricular zone of the hypothalamus; SN, substantia nigra; Strv, ventral striatum; SUB, subiculum; SUMl, lateral part of the supramammillary nucleus; TH, thalamus; VM, ventral medulla; VTA, ventral tegmental area. (Reprinted from Risold, P. Y., and Swanson, L. W., 1997, Connections of the rat lateral septal nucleus, Brain Res. Rev. 24, 115–195, with permission from Elsevier Science.)

Other Connections of the Lateral Group Connections with other telencephalic structures and with the thalamus have been described for the nuclei of the lateral group: • Weak projections from the whole lateral septal nucleus have been noticed in the hippocampus (Staiger and Nürnberger, 1991b; Jakab and Leranth, 1995; Risold and Swanson, 1997b), maybe reciprocating the massive hippocampal inputs. Little is known about the physiological importance of these projections. • A ventrolateral region of the caudal lateral septal nucleus (LSNc.v.l) receives abundant projections from the medial amygdala and the BST (Caffé et al., 1987). These projections contain vasopressin and galanin and are more abundant in the male rat than in the female (sexual dimorphism), and their distribution in the

caudal lateral septal nucleus follows the pattern of expression of the androgen receptor mRNA (DeVries and Buijs, 1983; DeVries et al., 1985, 1994; Skofitsch and Jacobowitz, 1985; Caffé et al., 1987; Wang et al., 1993; Risold and Swanson, 1997a). These projections have been involved in several social behaviors and are worth mentioning. Axons from the lateral septal nucleus innervate several nuclei of the thalamus (Staiger and Nürnberger, 1991a; Risold and Swanson, 1997b). Most of these projections are found in midline nuclei, in particular the nucleus reuniens, the thalamic paraventricular nucleus, and the paratenial nucleus (from the rostral and ventral parts), and in an enigmatic cell group called the median interanteromedial nucleus (from the caudal part; Risold and Swanson, 1997). In turn, the reuniens

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FIGURE 11 Illustration of the complex patterns of connections between the LSr and AH/LHApfx. (A) Bright-field photomicrograph of a PHA-L injection site in the central AH. (B) Dark-field photomicrograph of a fluorogold injection centered in the same level of the AH. (C, D) Dark-field photomicrographs illustrating the distribution of PHA-L-labeled axons (C) and fluorogold-labeled neurons (D) after injections of these tracers in the central AH (injection sites in A and B). Both tracers are distributed in a very similar way (in the LSr.vl.d.l (a) and LSr.vl.d.m (b), See table 1 for nomenclature) indicating that the central AH is bidirectionally connected with these two domains of the LSr. (E, F and G, H) After PHA-L and fluorogold injections centered in posterior and dorsal parts of the AH (injection sites not shown here, see Risold and Swanson, 1997b), both PHA-L and fluorogold labeling are observed in the LSr.vl.d.m (E, F) and LSr.dl. (G, H). Scale bar = 200 μm. AH, anterior hypothalamic nucleus; f, fornix; LHpfx, perifornical region of the lateral hypothalamic area; Pa, paraventricular nucleus of the hypothalamus. (Reprinted from Risold, P. Y., and Swanson, L. W., 1997, Connections of the rat lateral septal nucleus, Brain Res. Rev. 24, 115–195, with permission from Elsevier Science.)

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and paraventricular nuclei send a few fibers in the rostral part of the lateral septal nucleus. Most interestingly, these nuclei project heavily in the hippocampal formation (entorhinal cortex, temporal field CA1, and ventral subiculum; see in Risold et al., 1997). Finally a small input from the whole lateral septal complex has been described in the lateral habenular nucleus (Staiger and Nürnberger, 1991a; Risold and Swanson, 1997b).

The Medial Group Connections of the medial septal nucleus and of the nucleus of the diagonal band are characterized by massive telencephalic outputs as well as by bidirectional connections with the brain stem, in particular with several nuclei of the posterior hypothalamus and of the reticular formation. The cholinergic nature of projections from the medial septal complex to many cortical areas was presumed more than 30 years ago (Lewis and Shute, 1967; Shute and Lewis, 1967; Butcher and Woolf, Chapter 35, this volume). Septocortical projections innervate all fields of the hippocampus, entorhinal, cingulate, medial prefrontal, olfactory (olfactory bulb, piriform cortex, corticoamygdaloid nuclei), and insular areas, as well as, to a lesser extent, occipital, somatosensory, and orbital areas of the cerebral cortex. These projections are topographically organized (Divac, 1975; Domesick, 1976; Swanson and Cowan, 1979; Price and Stern, 1983; Alonso and Köhler, 1984; Woolf et al., 1984, 1986; Amaral and Kurz, 1985; Kasa, 1986; Luiten et al., 1985, 1987; Zaborszky et al., 1986; Swanson et al., 1987; Gaykema et al., 1990; Woolf, 1991). The medial septal nucleus projects essentially in the hippocampal formation. Gaykema et al. (1990) have shown that lateral cells in this nucleus mostly send their axons in ventral (temporal) hippocampal fields, while medial neurons project in the dorsal (septal) hippocampus. The nucleus of the diagonal band innervates the hippocampus, but also sends heavy projections in other cortical fields, including olfactory, cingulate, and entorhinal areas. Acetylcholine is a major neurotransmitter in these pathways, but many medial septal complex cortically projecting cells are not cholinergic but GABAergic (Köhler et al., 1984; Rye et al., 1984; Semba, 2000). Several peptides are also involved such as substance P, enkephalin, galanin, and neurotensin (Lamour et al., 1988; Lamour and Epelbaum, 1988; Gonzalo-Ruiz and Morte, 2000; Semba, 2000). Depending on the cortical area that they innervate, these projections use distinct medial or lateral routes (Peterson, 1994; Jakab and Leranth, 1995). Descending projections from the medial septal complex travel through ventral aspects of the medial

forebrain bundle to reach the brain stem (Meibach and Siegel, 1977; Swanson and Cowan, 1979; Veening et al., 1982; Tomimoto et al., 1987). However, some of them arch dorsally to enter the stria medullaris and terminate in the lateral habenula (moderately) or innervate several nuclei of the thalamus including the reticular and lateral mediodorsal nuclei. Reportedly, some projections in the stria medullaris continue in the fasciculus retroflexus and innervate the interpeduncular nucleus. Parts of them may be cholinergic (Albanese et al., 1985; Motohashi et al., 1987). Other medial septal efferents innervate en passant the lateral hypothalamic area. Most of those terminate in the lateral part of the supramammillary nucleus. A few of them, however, innervate the medial and lateral mammillary and tuberomammillary nuclei, the ventral tegmental area, and the raphe and laterodorsal tegmental nuclei (Meibach and Siegel, 1977; Swanson and Cowan, 1979; Veening et al., 1982; Tomimoto et al., 1987; Wouterlood et al., 1988; Kalén and Wiklund, 1989; Cornwall et al., 1990; Jakab and Leranth, 1995). Many of these descending projections to the brain stem are noncholinergic (Kalén and Wiklund, 1989), although cholinergic projections to the interpeduncular nucleus may use this path as well (Groenewegen and Wouterlood, 1988; Vertes and Fass, 1988). The medial septal complex receives descending and ascending afferents. Emphasis has been put on the direct projections from the hippocampus (Gaykema et al., 1991a). Projections originating in the “septal” and temporal end of the hippocampus are segregated, with septal afferents innervating medial parts of the medial septal nucleus and the nucleus of the diagonal band, while temporal projections are found in lateral parts of the medial septal nucleus and the rostromedial nucleus of the diagonal band. Projections from other cortical fields have also been described, particularly from entorhinal and prefrontal areas (Sesack et al., 1989; Gaykena et al., 1991b; Hurley et al., 1991; Takagishi and Chiba, 1991; Totterdel and Meredith, 1997; Leranth et al., 1999). During the 70s, using the anterograde transport of 3 H-labeled amino acids, Swanson and Cowan (1979) described dense projections from the lateral septal nucleus to the medial septal complex. Later, Leranth et al. (1992) challenged this view. Using the Phaseolus vulgaris-leucoagglutinin (PHA-L) method, the last authors described weak afferents to the medial septal nucleus from small injection sites centered dorsally in the lateral septal nucleus. An actual view of the projections from the lateral septal complex to the medial septal complex could be in between these two positions. After multiple injections of PHA-L in the lateral septal nucleus and adjacent nuclei, such as the

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Groenewegen et al., 1986; Montone et al., 1988; Vertes, 1988; Panula et al., 1989; Cornwall et al., 1990; Cullinam and Zaborsky, 1991; Zagon et al., 1994; Gaykema and Zaborzky, 1996; Kia et al., 1996; Leranth and Vertes, 1999). A clear topography has not yet been reported for these last projections.

septofimbrial and septohippocampal nuclei, Risold and Swanson (1997b) described topographically organized outputs from these cell groups in the medial septal complex (Fig. 12). From this, it is tempting to postulate that septal or temporal parts of the hippocampus project to or receive afferents from interconnected divisions of the lateral and medial septal complexes. However, this hypothesis needs to be investigated. Finally, the medial septal complex receives ascending inputs from the brain stem (Vertes, 1988). More is said about these afferents in the functional section of this chapter. The hypothalamus, in particular the supramammillary nucleus, the posterior hypothalamic nucleus, and the lateral hypothalamic area, appears to constitute one of the main sources of ascending projections (Cullinam and Zaborzky, 1991; Vertes, 1992; Oddie et al., 1994). Evidence has been published that they may respect a certain topography (Cullinam and Zaborzky, 1991). However, it is still very difficult to relate these patterns to those involving the medial septal complex projections toward the cerebral cortex. Finally, many other brain stem nuclei (interpeduncular, ventral tegmental area, raphe, and laterodorsal tegmental nuclei and locus ceruleus) project in the medial septal complex (Segal and Landis, 1974;

The Posterior Group Very few studies have investigated the connections of the nuclei of the posterior group (triangular nucleus, bed nuclei of the anterior commissure and of the stria medullaris). Swanson and Cowan (1979) described these nuclei as the main source of septal afferents through the stria medullaris to the medial habenular nucleus (mostly bilaterally; Swanson and Cowan, 1979; Contestabile and Villani, 1984; Kawaja et al., 1990). Some of these projections may continue in the fasciculus retroflexus to reach the interpeduncular nucleus (Swanson and Cowan, 1979; Villani, 1996). Subsequent works showed that most of the septal region sends at least a few fibers into the habenular nuclei (Staiger and Nümberger, 1991a; Risold and Swanson, 1997b). Other connections of the nuclei of the posterior group appear less clear. In particular, very little is known

A

d

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FIGURE 12 Schematic representation of topographically organized projections from the lateral septal complex (on A, gray areas labeled a to e) onto the medial septal complex. ac, anterior commissure; f, fornix; LS, lateral septal nucleus; MS, medial septal nucleus; SFi, septofimbrial nucleus; SHi, septohippocampal nucleus.

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about their afferents. Dense fibers labeled for DBH or acetylcholinesterase have been described, respectively, in the bed nucleus of the anterior commissure and in the triangular nucleus (Swanson and Hartman, 1975; Paxinos and Watson, 1986). Furthermore, Cajal (1911, 1995) indicated that a few collaterals from the ventral hippocampal commissure enter the triangular nucleus, and Swanson and Cowan (1979) suspected weak hippocampal afferents to the posterior nuclei. However, these observations need to be reinvestigated using modern neuroanatomical methods.

The Ventral Group Nuclei of the lateral and medial groups are related to the fornix system. On the other hand, the BST is related to the stria terminalis (Alheid et al., 1995; Swanson and Petrovich, 1998; for a recent review, see Dong et al., 2001), even though some projections between the amygdala and the BST travel through the ansa peduncularis (Dong et al., 2001). Descending inputs from the amygdala to the BST may be GABAergic as well as glutamatergic (see Swanson and Petrovich, 1998). Furthermore, it has been suggested that medial parts of the BST receive afferents from amygdalar nuclei associated with the accessory olfactory system, while anterior and lateral components are targeted by the central nucleus and some nuclei associated with the main olfactory pathways (Krettek and Price, 1978; Weller and Smith, 1982; Canteras et al., 1992a; Aldheid et al., 1995; Petrovich et al., 1996; Dong et al., 2001). Only the lateral and anterior basolateral nuclei do not provide significant projections to the BST (Dong et al., 2001). In addition, the BST is bidirectionally connected with the brain stem. Past and recent anatomical data suggest that the medial nuclei of the BST are connected mainly with the hypothalamic nuclei of the medial zone, while lateral cell groups are essentially linked to autonomic centers (Sofroniew, 1983; Aldheid et al., 1995; Hutton et al., 1998; Saper, Chapter 24, this volume). But descending projections from some cell groups of the BST may not fit in this simple medial– lateral partition (Dong et al., 2000).

FUNCTIONAL ORGANIZATION OF THE SEPTAL REGION Septal nuclei have been implicated in a large range of motor (autonomic, neuroendocrine), behavioral, and cognitive responses (Swanson et al., 1987; Sheehan and Numan, 2000; Swanson and Risold, 2000). The septohippocampal system, which includes the largest component of the septal region with the nuclei of the

lateral and medial groups, has attracted most of the attention of investigators. Lesions of these nuclei have long been associated with changes in emotional behaviors (Brady and Nauta, 1953), resulting in the description of the septal rage. Subsequent refined experimental works have shown that the lateral septal nucleus is the main structure responsible for the septal rage. On the other hand, lesions of the medial septal complex produce several cognitive deficits (for example, disruption of forms of memory or of attention) and desynchronize the electrical activity (theta activity) of the hippocampal formation (Wenk, 1997; Apartis et al., 1998; Leutgeb and Mizumori, 1999; Durkin, 2000; Numan, 2000). Furthermore, the progressive loss of cholinergic inputs to the hippocampal formation in patients suffering from Alzheimer’s disease is suspected to contribute to the impairment of memory characteristic of this disease (Perry et al., 1978a, 1978b). The body of work linking the medial septal complex to these functions is enormous, and the reader is referred to several reviews (Swanson et al., 1987; Jakab and Leranth, 1995; Hörtnagl and Helleweg, 1997; Wenk, 1997; Bland, 2000; Hasselmo, 2000; Walsh, 2000; Whishaw, 2000; Wu et al., 2000). In brief, GABAergic and cholinergic septal inputs contribute to synchronize hippocampal neurons. GABAergic septal afferents target GABAergic interneurons throughout the hippocampal formation (including the entorhinal cortex) and synchronize these neurons in faster frequency domains (40–200 Hz). Cholinergic inputs target pyramidal cells and interneurons and drive them to discharge upon an oscillatory theta frequency (Chrobak, 2000). Thus, the medial septal complex is often perceived as the pacemaker of the hippocampus, but the firing pattern of medial septal neurons is under the control of ascending projections from the brain stem (Stewart and Steven, 1990; Kirk, 1998; Bland, 2000; Denham and Borisyuk, 2000; Leranth and Vertes, 2000). Several circuits have been described, but there is a general agreement on the fact that the ascending brain stem hippocampal synchronizing pathway originates mainly in rostral parts of the pontine reticular nucleus (or part oralis), the general area encompassing the pedunculopontine tegmental nucleus. Caudal diencephalic neurons in the supramammillary nucleus as well as in the posterior hypothalamic nucleus appear to play a key role in relaying the signal from the pontine region (Kirk, 1998; Leranth and Vertes, 1999, 2000). Finally, serotoninergic afferents from the raphe (median or central superior nucleus) may have a desynchronizing effect on the hippocampal EEG through projections in the medial septal complex (Leranth and Vertes, 2000).

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The roles of the theta rhythm are still under investigation (Hasselmo, 2000). Theta activity has been linked to aspects of memory formation in the hippocampus, but its amplitude varies with the behavioral state of the individual (Feder and Ranck, 1973; King et al., 1998; Oddie and Bland, 1998; Apartis et al., 2000; Hirase et al., 2001; Shin and Talnov, 2001; Wiebe and Staubli, 2001). For example, theta oscillation amplitude increases during voluntary locomotion, but decreases during automatic behaviors such as grooming (Kemp and Kaada, 1975; Maloney et al., 1997; Oddie et al., 1997). Paxinos and Bindra (1970) argued that the theta rhythm was simply the accompaniment of gross movement. It is worth mentioning here that the pedunculopontine tegmental nucleus is part of the mesencephalic locomotor region (Skinner and Garcia-Rill, 1984). Experimental data tend to implicate the lateral septal nucleus in many responses and behaviors, including feeding, water intake, autonomic responses, sexual and emotional behaviors, or aspects of adaptative navigation (Jakab and Leranth, 1995; Risold and Swanson, 1997b; Sheehan and Numan, 2000; Mizumori et al., 2000). As already mentioned, lesions of this nucleus result in the septal rage syndrome (Brady and Nauta, 1953). At first, this behavioral change was believed to correspond to an enhanced aggressiveness. However, subsequent studies favored the hypothesis of an exaggerated defensive behavior (Blanchard et al., 1979; Albert and Chew, 1980; Sparks and Ledoux, 1995, 2000; Sheehan and Numan, 2000). A number of studies have indicated a role for the lateral septal nucleus in other social behaviors related to dominant–subordinate relationships, or parental care. The huge descending inputs from the lateral septal nucleus to the hypothalamus support the involvement of this nucleus in such responses (Swanson and Cowan, 1979; Staiger and Nümberger, 1991a; Staiger and Wouterlood, 1990; Risold and Swanson, 1997). Several circuits have been described within the hypothalamus during the last 15 years using recent tract-tracing procedures (Risold et al., 1997; Swanson, 2000a). These circuits are critical for the expression of such behaviors. In particular, the hypothalamic sexual dimorphic circuit, connecting anteromedial preoptic areas, the ventrolateral part of the ventromedial nucleus, and the ventral premammillary nucleus, is involved in reproductive (including parental) behaviors. A defensive circuit connecting the hypothalamic anterior, dorsomedial part of the ventromedial, and dorsal premammillary nuclei has been described (see as well Canteras et al., 1997; Comoli et al., 2000). A complex network is wiring the ventral part or several divisions of the rostral part of the lateral septal nucleus, respectively, with intercon-

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nected divisions of these hypothalamic circuits (Risold and Swanson, 1997b). Thus, both parts appear involved in the control of hypothalamic circuits that initiate social behaviors. The caudal lateral septal nucleus projects heavily in the lateral hypothalamic area and the lateral supramammillary nucleus. It receives abundant afferents from the laterodorsal tegmental nucleus, the locus ceruleus, and the raphe. These last cell groups are implicated in arousal and the sleep/wake cycle (Jones, 1991; Shouse et al., 2000). The supramammillary nucleus is a key structure for the control of the hippocampal theta rhythm (Leranth and Vertes, 2000; Vertes and McKenna, 2000; Nakanishi et al., 2001). Thus, the caudal lateral septal nucleus may very well be involved in such responses. The intrinsic circuitry of the hippocampal formation (trisynaptic circuit) is organized transverse to the septal temporal axis of the hippocampus (Swanson et al., 1987; Amaral and Witter, 1989, 1995; Tamamaki and Nojyo, 1995; Moser and Moser, 1998). Projections from the fields CA1 and CA3 onto the lateral septal complex are topographically organized so that pyramidal cells in progressively more temporal parts of the Ammon’s horn and subiculum project in more ventral regions of the lateral septal complex, themselves interconnected with distinct hypothalamic nuclei or areas (Risold and Swanson, 1996). Thus, this anatomical evidence completed by the hypotheses of topographical inputs from the lateral septal complex to the medial septal complex and from the hypothalamus to both suggests the existence of complex but very organized networks linking the hippocampus, septum, and hypothalamus (Fig. 10). For example, temporal parts of the hippocampal fields CA3 and CA1/subiculum project, respectively, in caudal or ventral/rostral parts of the lateral septal nucleus. In turn, these last cell groups influence or control distinct hypothalamic functions, including arousal, and social behaviors such as reproductive and defensive behaviors. Finally, the same cell groups and the hypothalamus may send information back to the hippocampus through parts of the medial septal complex. However, it has been demonstrated that hypothalamic medial zone nuclei can influence the ventral hippocampal regions (including temporal Ammon’s horn/subiculum and medial entorhinal cortex) through dense projections onto midline nuclei of the thalamus, in particular the nucleus reuniens (Risold et al., 1994, 1997). These pathways resemble somewhat the Papez (1937) circuit that involves the mammillary nuclei, the anterior thalamic nuclei, and the cingulate/retrosplenial cortical areas as well as dorsal hippocampal regions. The dorsal subiculum projects in the mammillary body through the postcommissural fornix (Swanson and Cowan, 1975). The septal pole of

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the hippocampus projects in the dorsomedial tip of the lateral septal nucleus. Few direct links have been found between this part of the lateral septal nucleus and the mammillary body, except for dense projections in the supramammillary nucleus (Risold and Swanson, 1997b). However, this septal cell group innervates the nucleus of the diagonal band (Fig. 10, and see Leranth et al., 1992; Risold and Swanson, 1997b) which in turn innervates the cingulate/retrosplenial cortical regions. The BST and the amygdala, like the lateral septal nucleus, have been involved in a large range of functions that, in part, are the results of descending projections from the BST to the hypothalamus and caudal brain stem. However, the BST is a very complex structure with an organization that is still the object of many controversies. In the hypothalamus, projections from the BST and the lateral septal nucleus often overlap, although a lot still needs to be learned about the complete extent and significance of such convergent informations. In conclusion, structural evidence shows that the septal region does not form a functional unit by itself, neither does the septohippocampal system nor the cholinergic basal forebrain system. The hippocampal formation, septal region, and hypothalamus compose complex and parallel circuits that may form loops (for example, the Papez circuit), which are largely open as they are often interconnected (Fig. 10). Similar remarks have been made less than 15 years ago, when describing

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the functional organization of the “basal ganglia circuitry” (involving the adjacent striatum) (Alexander et al., 1990). Indeed, in a recent review, Swanson (2000a) has proposed a new wiring diagram for the motor neural network that controls motivated behaviors. In this work (see as well Risold et al., 1997; Swanson and Petrovich, 1998; Swanson and Risold, 2000), it is proposed that the lateral septal complex be viewed as a medial component of the striatum, as it receives abundant glutamatergic afferents from the cortex that innervate GABAergic spiny neurons in this structure (Fig. 13). The medial septal complex shares many particularities with pallidal cell groups, as it contains cholinergic neurons projecting in the cortex, receives topographical inputs from the medial striatum, and develops projections with the brain stem, in particular with the reticular formation. A similar line of thinking allowed the same authors (Swanson and Petrovich, 1998; Swanson, 2000a) to assimilate the cortical nuclei of the amygdala as cortex, several deep nuclei (central and medial amygdalar nuclei) as the caudal striatum, and the BST as the caudal pallidum.

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PFC

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BRAINSTEM FIGURE 13 Schematic representation of the organization of the telencephalon. See text for details. Amyc, cortical nuclei of the amygdala; BST, bed nuclei of the stria terminalis; HiF, hippocampal formation; LSC, lateral septal complex; MSC, medial septal complex; PFC, prefrontal cortex (symbolizing all cortical areas projecting in the ventral striatum); Pald-v-m-c, dorsal–ventral–medial–caudal pallidum; Strd-v-m-c, dorsal–ventral–medial–caudal striatum.

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21 Hippocampal Formation MENNO P. WITTER Graduate School Neurosciences Amsterdam, Research Institute Neurosciences Vrije Universiteit Medical Center, Department of AnatomyAmsterdam, The Netherlands

DAVID G. AMARAL Department of Psychiatry, California National Primate Research Center and The M.I.N.D. Institute, University of CaliforniaDavis, California, USA

The hippocampal region is a prominent component of the rat nervous system that has attracted the attention of neuroanatomists since the beginning of formal study of the nervous system. Much of the available information concerning the cellular organization and intrinsic connectivity of the hippocampal formation dates from the classical Golgi studies of Ramon y Cajal and Lorente de Nó (Lorente de Nó, 1933, 1934; Ramón y Cajal, 1911). Another golden age of hippocampal investigation was ushered in with the use of the Nauta degeneration method to map the intrinsic and extrinsic pathways of the rat hippocampal formation. The careful and comprehensive studies of Blackstad (1956, 1965; Blackstad et al., 1970), his students (HjorthSimonsen, 1972a, 1973; Hjorth-Simonsen and Jeune, 1972; Storm-Mathisen and Blackstad, 1964; Zimmer, 1971; Zimmer and Haug, 1978; Zimmer and HjorthSimonsen, 1975), and others, refined concepts concerning the regional organization of the hippocampal formation and revealed new information on the organization of connections between the different fields of the hippocampal formation and between the hippocampal formation on each side of the brain. But progress on unraveling the neurocircuitry of the hippocampal formation has not stopped and every advance in neuroanatomical methodology has fostered new insights into the structure and functional organization of the rat hippocampal formation and the adjacent parahippocampal region. With the renaissance in neuroanatomical methodology in the last part of the previous century, there has been a massive expansion of

The Rat Nervous System, Third Edition

data generated from studies employing techniques such as immunohistochemistry, receptor autoradiography, in situ hybridization, sensitive new anterograde and retrograde tracing methods, intracellular labeling, and computer-aided reconstruction combined with newly developed protocols for light, confocal, and electron microscopical assessment.

Synopsis of the Chapter An overview chapter such as this cannot hope to comprehensively review all of the historical and recent findings on the neuroanatomy of the hippocampal region. The interested reader should consult a series of older yet still relevant comprehensive accounts and books (Amaral, 1991, 1993; Amaral and Witter, 1989; Chan-Palay and Köhler, 1989; Lopes da Silva et al., 1990; Ribak et al., 1992; Seifert, 1983; Storm-Mathisen et al., 1990; Swanson et al., 1987; Witter et al., 1989). There are also a number of more recent publications and informative publications that focus on topics such as the organization of local neuronal networks (Alonso, 2002; Freund and Buzsaki, 1996; Parra et al., 1998; Wouterlood, 2002) or overall connectivity (Burwell, 2000; O’Mara et al., 2001; Pitkänen et al., 2000; Witter et al., 2000a, 2000b; see also Scharfman et al., 2000; Witter and Wouterlood, 2002). An important area of advance since the last version of this chapter is related to knowledge on interneurons (see Freund and Buzsaki, 1996; McBain and Fisahn, 2001; Parra et al., 1998; Sloviter et al., 2001, and references therein for further details).

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In this chapter, we do not go into great detail trying to cover all aspects of interneurons; we limit ourselves to a description of the distinguishing features of the main populations present. The intent of this chapter is to provide an overview of the constituent regions, cytoarchitectonic organization, connectivity, and chemoanatomy of the rat hippocampal region. Among the topics that are covered, we describe the principles of the organization of intrinsic circuitry of the hippocampal region which provide constraints on how information is processed and thus have important functional implications. We do not deal in great detail with aspects of either hippocampal function or computational models of hippocampal function.

Definition of the Hippocampal Region Despite extensive study, there is still substantial confusion concerning the terminology of the various components of the hippocampal region and even which components should be included under this term. The confusion is due, in part, to the fact that criteria for grouping structures related to the hippocampus are often not functionally grounded but are based, for example, on arbitrary structural traits such as the number of cortical layers. What is clear is that when a histological section is prepared through the hippocampal region and stained for the demonstration of neuronal cell bodies, a number of cytoarchitectonically distinct regions are apparent. Various terminologies have been applied to these different regions and several synonymous terms are still commonly employed for some of them. The terms used in this chapter are based on converging evidence from cytoarchitectonic, histochemical, connectional, and functional data that justify their usage. The locations of the various fields of the hippocampal region are indicated in Figs. 1 and 2. The hippocampal region is taken to include two sets of cortical structures, the hippocampal formation on the one hand and the parahippocampal region on the other hand. The major defining differences between the two are the number of cortical layers present and the overall principles of connectivity. The hippocampal formation comprises three cytoarchitectonically distinct regions: the dentate gyrus; the hippocampus (or hippocampus proper), which is subdivided into three fields (CA3, CA2, and CA1); and the subiculum. All three hippocampal regions share the characteristic three-layered appearance that in older accounts has been considered the defining feature of the so-called allocortex. Moreover their connectivity is largely unidirectional. The dentate granule cells project via their distinctive mossy fibers to the CA3 field of the hippocampus. While some CA3 cells contribute axon

collaterals to the deep or polymorphic layer of the dentate gyrus, these axons do not innervate the granule cells. A similar preferential unidirectional pattern holds for the other major intrinsic connections (CA3to-CA1; CA1-to-subiculum) of the hippocampal formation. One justification for using this cascade of unidirectional connections as a principal criterion for inclusion of structures in the hippocampal formation is that these one-way projections are extremely atypical for corticocortical connections; in most other cortical regions, reciprocal connections are the norm. The entorhinal cortex provides the dentate gyrus with its major input via the so-called perforant pathway. The entorhinal to dentate gyrus projection is not reciprocated either, since none of the cells in the dentate gyrus project back to the entorhinal cortex. Therefore, one might argue that the entorhinal cortex should be considered part of the hippocampal formation (cf. Amaral and Witter, 1995; Witter, 2002). Aside from the fact that within the entorhinal cortex one can distinguish more than three layers, its position is uniquely different from the ones that constitute the hippocampal formation in that the entorhinal cortex sends direct projections into all subfields of the hippocampal formation. Moreover, those reaching field CA1 and the subiculum are reciprocated. The feature of reciprocal connections with CA1 and the subiculum has also recently been described to hold true for cortical regions adjacent to the entorhinal cortex, including the perirhinal and postrhinal cortices. Therefore, these latter two cortical areas, together with the entorhinal cortex are included in the parahippocampal region (para = alongside or near). The parahippocampal region, or retrohippocampal region (retro = behind) as it has been referred to, also includes the presubiculum and parasubiculum since these latter areas share the laminar features of having more than three layers and they are reciprocally connected with the subiculum (cf. Scharfman et al., 2000; Witter, 2002). The reader should be aware that our definition of the terms “hippocampal region” and “hippocampal formation” is not universally accepted in the hippocampal literature. Some authors would, as indicated above, emphasize the unidirectionality of connectivity as the key-defining characteristic and include entorhinal, presubiculum, and parasubiculum into the hippocampal formation (cf. Amaral and Witter, 1995). The subiculum, presubiculum, and parasubiculum are sometimes grouped together as the subicular complex. Aside from the fact that it is controversial whether the presubiculum and parasubiculum should be considered as having only three layers, a characteristic which obviously does not hold true for the entorhinal cortex, we argue in the overview section of the Subiculum, Presubiculum and

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Parasubiculum that most connectional features of the pre- and parasubiculum do not seem to support linking them with the subiculum into a separate conglomerate called the “subicular complex.”

The Position of the Hippocampal Formation and Parahippocampal Region in the Rat Brain The three-dimensional position of the rat hippocampal formation in the brain is rather complex (Fig. 1). The hippocampal formation appears grossly as an elongated structure with its long axis extending in a Cshaped fashion from the septal nuclei of the basal forebrain rostrodorsally, over and behind the diencephalon, into the incipient temporal lobe caudoventrally. The long axis of the hippocampal formation is referred to as the septotemporal axis (with the septal pole located dorsally and rostrally) and the orthogonal axis is referred to as the transverse axis. What is not obvious from the surface view of the hippocampal formation is that different fields make up the structure at different septotemporal levels (Figs. 1A and 1D). At extreme septal levels, for example, only the dentate gyrus and the CA1–3 subdivisions of the hippocampus are present. About 15% of the way back toward the temporal pole, the subiculum first appears. The parahippocampal region should be envisaged as the most caudal and ventral portion of the cortical mantle, wrapping around the caudal and ventral portions of the hippocampal formation (Figs. 1B and 1C). Thus, the fields that make up the parahippocampal region can be found caudal and ventral to the other fields of the hippocampal formation. The presubiculum and parasubiculum form the most medial rim of this cortical covering, the central part of which is formed by the entorhinal cortex. The perirhinal and postrhinal cortex form the outer lateral and caudodorsal rim. The border of the entorhinal cortex with the perirhinal and postrhinal cortices is approximately located at the rhinal sulcus (Figs. 1B and 1C), which forms one of the major sulci of the rat cerebral cortex. Two additional sets of terms must be mentioned in order for the following descriptions to be understood. In describing locations within the transverse axis of a hippocampal field, we use the dentate gyrus as the proximal extreme of the hippocampal formation (Fig. 2C) and transverse locations are stated with reference to this. As an example, the portion of CA1 located closer to CA2 is called the proximal portion and the portion closer to the subiculum is called the distal portion. Regarding the description of the parahippocampal region, we use the standard dorsal-to-ventral, rostral-to-caudal, and lateralto-medial nomenclature, in which lateral means close to the rhinal fissure and medial is taken to imply close to

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the border between parahippocampus and hippocampus (Figs. 1C and 1D). In discussing the radial organization of a particular field, layers located closer to the pia or hippocampal fissure are called superficial whereas those located closer to the ventricle are called deep. In the hippocampus, for example, the apical dendrites of pyramidal cells (see below) are located superficially and the basal dendrites are deep.

Fiber Bundles of the Rat Hippocampal Region The deep or ventricular surface of the subiculum and hippocampus are covered by a thin sheet of mainly myelinated afferent and efferent fibers called the alveus. In part, these are fibers originating from the pyramidal cells of the hippocampus and subiculum that are in route to subcortical termination sites or to the contralateral hippocampal formation. The fibers extend obliquely over the surface of the hippocampus and collect in an increasingly thicker fiber bundle (as one progress from temporal to septal) located at the lateral extreme of the hippocampus called the fimbria. As the fibers leave the hippocampus and descend into the forebrain they are usually referred to as the columns of the fornix. The fornix splits around the anterior commissure to form a rostrally directed precommissural component, which innervates the septal nuclei and other basal forebrain structures, and a caudally directed postcommissural component that is directed toward the diencephalon. As the postcommissural fornix begins its course into the diencephalon (ultimately to reach the mammillary region of the posterior hypothalamus), two smaller bundles split off. One, the so-called medial corticohypothalamic tract, travels medially to innervate a number of anterior hypothalamic areas. The other, which has no formal name but has been called the subiculothalamic track (Swanson et al., 1987) carries fibers to the anterior thalamic nuclei. Returning to the fimbria, a large number of fibers cross the midline before entering the columns of the fornix. These fibers cross in a position just caudal to the septal area in the ventral hippocampal commissure. Many of these fibers are true commissural fibers and are directed to fields in the contralateral hippocampal formation. A smaller number of the fibers are directed into the contralateral descending column of the fornix and ultimately innervate the same structures that receive the ipsilateral pre- and postcommissural fornix. We should emphasize that the fornix and fimbria carry both efferent fibers from the hippocampal formation and subcortical afferent fibers to the hippocampal formation. Thus, at all points along these fiber bundles, including the ventral hippocampal commissure, there will be axons flowing in both directions. Within the fimbria–fornix and com-

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FIGURE 1 Schematic representations showing the three-dimensional organization of the hippocampal region in the rat brain. (A) The C-shaped hippocampus is shown in a transparent shell of the rat brain. Note the indication of the septotemporal or longitudinal axis. (B) Lateral surface reconstruction illustrating the positions of the perirhinal (PER), postrhinal (POR), and entorhinal (EC) cortices. (C) Slightly rotated reconstruction of caudal half of the left hemisphere, detailing the subdivision of the entorhinal cortex into the caudal (CE), medial (ME), dorsolateral (DLE), dorsal-intermediate (DIE), ventral-intermediate (VLE), and amygdaloentorhinal (AE) areas. (D) The surface of the hippocampal formation has been pictured outside of the brain. Three horizontal sections (numbered 1–3) at different dorsoventral levels of the hippocampal formation are shown. Note that the entorhinal cortex (EC) is not seen at the most dorsal level but is apparent at the mid and ventral levels of the hippocampal formation. Also three coronal sections (numbered 1´–3´) are shown at different rostrocaudal levels through the hippocampal formation. Additional abbreviations: CA1, CA3, fields of the hippocampus; CPu, caudate–putamen; DG, dentate gyrus; f, fornix; S, subiculum.

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missural system a topographical organization of fiber tracts appears to exist such that the diffusely terminating septohippocampal projections are loosely distributed throughout the fimbria–fornix, while those projections that are known to terminate in specific laminae of the hippocampal formation (commissural projection, crossed entorhinohippocampal projection) form fiber bundles within the fimbria and the ventral hippocampal commissure (Adelmann et al., 1996). There is a second commissural bundle associated with the hippocampal formation. This dorsal hippocampal commissure crosses the midline just rostral and ventral to the splenium of the corpus callosum. It carries fibers mainly originating from, or projecting to, the presubiculum, parasubiculum, and entorhinal cortex. The fibers of the dorsal hippocampal commissure are continuous laterally with a bundle of fibers that is interposed between the entorhinal cortex and the preand parasubiculum. This collection of fibers is referred to as the angular bundle and, in addition to carrying commissural fibers of entorhinal and pre- and parasubicular origin, is the main route by which fibers from the ventrally situated entorhinal cortex travel to all septotemporal levels of the other hippocampal fields, particularly the dentate gyrus, hippocampus, and subiculum. The angular bundle further contains fibers to and from a variety of cortical and subcortical structures that are connected with the entorhinal cortex. Examples are fibers connecting the entorhinal cortex with the thalamus and with cingular areas.

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devised by Lorente de Nó (1934), has remained inconsistent and is an area of substantial confusion in the hippocampal literature. This is due, in part, to the difficulty in determining the boundary, in standard Nisslstained sections, of the polymorphic cell layer and the hippocampal pyramidal cell layer; the large, darkly stained cells of the polymorphic cell layer often closely approach the pyramidal cell layer and this close juxtaposition of two groups of large cells tends to blur the border between the two fields. Connectional data (see below), however, provide a reliable distinction between cells in the hippocampus and those in the polymorphic layer. Moreover, a variety of histochemical and immunohistochemical staining techniques have repeatedly indicated that the region of the polymorphic layer is clearly distinct from the pyramidal cell layer of the hippocampus (Amaral, 1978; Blackstad, 1956). Thus, to avoid the confusion that is engendered in the term CA4, it is best if this term is not used. It is often useful to be able to distinguish one portion of the granule cell layer from another. Again, a variety of terms have been used for this purpose. We call the portion of the granule cell layer that is adjacent to CA1 the enclosed blade (which is generally taken to be synonymous with suprapyramidal blade or limb, or dorsal limb) and the remaining portion of the granule cell layer is called the free blade (which is synonymous with infrapyramidal blade or limb, or ventral limb). The region of the “V” or “U” that unites the two blades is called the crest.

Neurons of the Dentate Gyrus

Cytoarchitectonics

The Granule Cell Layer

The dentate gyrus comprises three layers (Fig. 2). Closest to the hippocampal fissure is a relatively cellfree layer called the molecular layer. The principal cell layer or granule cell layer lies deep to the molecular layer and is made up primarily of densely packed columnar stacks of granule cells. The granule cell and molecular layers form a “V” or “U” shaped structure (depending on the septotemporal position) that encloses a cellular region, the polymorphic cell layer (or polymorphic layer for short), which constitutes the third layer of the dentate gyrus. The term hilus is often used synonymously with polymorphic layer. The polymorphic layer is sometimes given the incorrect name CA4 as if it were a portion of the pyramidal cell layer of the hippocampus. The term CA4 has also been used to designate the terminal portion of the hippocampal pyramidal cell layer that inserts within the limbs formed by the granule cell layer of the dentate gyrus. The use of the CA4 term, which was originally

The principal cell type of the dentate gyrus is the granule cell. Estimates of the total number of granule cells in one rat dentate gyrus range from approximately 0.6 × 106 to 2.2 × 106 (Amaral et al., 1990; Boss et al., 1985; Gaarskjaer, 1978a; Seress and Pokorny, 1981; West et al., 1988) and the number appears to depend on the age (Bayer, 1982; Bayer et al., 1982) and strain of the animal and on the method by which the counts are carried out. In part, the variability in cell counts is due to the fact that proliferation of granule cells continues at a slow rate well into adult life (Bayer, 1982; Bayer et al., 1982; Kempermann and Gage, 2000). Interestingly, such newly born cells appear to become functionally integrated in the hippocampal network (Hastings and Gould, 1999; Markakis and Gage, 1999; Van Praag et al., 2002). The packing density of granule cells is higher septally than temporally (Gaarskjaer, 1978a). Because the packing density of CA3 pyramidal cells follows an inverse gradient, the net result is that

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FIGURE 2 This figure shows various histological views of the rat hippocampal formation. (A, B) These photomicrographs show Nissl (A) and Timm’s sulfide silver (B) stained horizontal sections through the rat hippocampal formation. The accompanying line drawings (C, D) indicate various regions, layers, and fiber pathways of the rat hippocampal formation. Note in D that the three bands of the molecular layer of the dentate gyrus are labeled. The outer band demonstrates the lateral perforant path (lpp), the middle unstained region corresponds to the terminal zone of the medial perforant pathway (mpp), and the inner dark band corresponds to the zone of termination of the associational and commissural pathways of the dentate gyrus. Calibration bar in A equals 500 μm and also applies to B. Additional abbreviations: ab, angular bundle; PaS, parasubiculum; PrS, presubiculum; ML, GL, and PoDG, molecular, granule cell, and polymorphic layers, respectively, of the dentate gyrus; so, stratum oriens; pcl, pyramidal cell layer; sl, stratum lucidum; sr, stratum radiatum; s l-m. stratum lacunosum-moleculare; ab, angular bundle.

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at septal levels of the hippocampal formation the ratio of granule cells to CA3 pyramidal cells is something on the order of 12:1 whereas at the temporal pole this drops to 2:3. Since a major terminal field of the granule cell axons is the CA3 portion of the hippocampus (see below), the implication is that contact probability is much lower septally than temporally. The dentate granule cell has an elliptical cell body with a width of approximately 10 μm and a height of 16 μm. Each granule cell is closely apposed to other granule cells and in most cases there is no glial sheath intervening between the cells. The granule cell has a very characteristic cone-shaped tree of spiny dendrites with all of the branches directed toward the superficial portion of the molecular layer; most of the distal tips of the dendritic tree end just at the hippocampal fissure or at the ventricular surface (Fig. 6; Claiborne et al., 1990; Desmond and Levy, 1982). The dendritic trees of granule cells located in the enclosed blade tend, on average, to be larger than those of cells located in the free blade. Thus, the summed length of all of the dendrites of an average granule cell in the enclosed blade is approximately 3500 μm whereas a cell in the free blade has a dendritic tree of approximately 2800 μm. Desmond and colleagues (Desmond and Levy, 1982, 1985) have provided estimates for the number of dendritic spines on the granule cell dendrites. They found that cells in the enclosed blade had 1.6 spines/μm whereas cells in the free blade had 1.3 spines/μm. With these numbers and the mean dendritic lengths given above, an estimate for the number of spines on the average enclosed granule cell would be 5564 and for a free cell 3630. Along the deep surface of the granule cell layer, socalled basket cells with pyramidal-shaped cell bodies are found wedged between the granule cell and polymorphic layers (Fig. 3). The basket portion of the name refers to the fact that the axon of these cells forms pericellular plexuses that surround the somata of granule cells. Basket cells, with bodies 25–35 μm in diameter, were described by Ramon y Cajal (Ramón y Cajal, 1911) as having a single, principal aspiny apical dendrite directed into the molecular layer that divides into several aspiny branches and two or three primary basal dendrites that ramify and extend into the polymorphic layer. A second type, having a dendritic tree restricted to the hilus has been reported as well. The majority of these cells are GABAergic and many are immunoreactive for other substances such as the calcium-binding protein parvalbumin. Seress and Pokorny (1981) found that the number of basket cells was not constant throughout the transverse and septotemporal extents of the dentate gyrus. At septal levels, the ratio of basket cells to granule cells was 1:100 in the

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enclosed blade and 1:180 in the free blade. At temporal levels, the number was 1:150 for the enclosed blade and 1:300 for the free blade. These data emphasize the point that, despite the apparent homogeneity of the hippocampal fields along their long axis, there are several regional differences in connectivity, especially as regards neurochemical innervation. Within the same subgranular region occupied by the cell bodies of the basket cells are several other cell types with distinctly different somal shapes and dendritic configurations. Some of these are multipolar with several aspiny dendrites entering the molecular and polymorphic layers while others tend to be more fusiform shaped with a similar dendritic distribution. As Ribak and colleagues have pointed out (Ribak and Seress, 1983; Seress and Ribak, 1983, 1984, 1985, 1990), all of these cells share fine structural characteristics such as infolded nuclei, extensive perikaryal cytoplasm with large Nissl bodies, and intranuclear rods. These cells either contribute to the innervation within the granular cell layer or send their axons out into the molecular layer (see page 639 for further details; Freund and Buzsaki, 1996). It has become clear that many of these neurons are immunoreactive for GABA and most likely modulate granular cell activity. Cells of the Molecular Layer The molecular layer is mainly occupied by the dendrites of the granule, basket, and various polymorphic cells as well as terminal axonal arbors from several sources. However, a few neuronal cell types are also present in the molecular layer. The first is located deep in the molecular layer, has a multipolar or triangular cell body, and gives rise to an axon that appears to contribute to the basket plexus within the granule cell layer. This neuron, which stains positively for VIP, has aspiny dendrites that remain mainly within the molecular layer. It would appear that this is another form of basket cell located above, rather than below, the granule cell layer (Hazlett and Farkas, 1978; Kosaka et al., 1984, 1987; Ribak and Seress, 1983). A second type of neuron has been observed in the molecular layer of the dentate gyrus that resembles the so-called “chandelier” or axoaxonic cell (Fig. 3) originally found in the neocortex (Kosaka, 1983; Somogyi et al., 1985; Soriano and Frotscher, 1989). The axoaxonic cell is named for the fact that its axon descends from the molecular layer into the granule cell layer, collateralizes profusely, and then terminates exclusively on the axon initial segments of granule cells. These cells have a dendritic tree, which generally spans the width of the molecular layer, and basal dendrites are not welldeveloped or even absent. This dendritic organization suggests that the main inputs of these cells are

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FIGURE 3 Summary diagram of interneurons of the dentate gyrus. Filled circles indicate the location of the cell bodies and thick lines indicate the predominant orientation and laminar distribution of the dendritic tree. Indicated in the background are the principal neurons, providing an indication which domain is innervated by the different interneuron groups. Also indicated is the laminar distribution of different inputs, often showing a striking match with interneuron type or axon distribution. (A) Morphological classification scheme. Interneuron types are identified according to dendritic and axonal arborization. The hatched boxes indicate the preferential axonal distribution for each cell type. (B) Laminar distribution of dendritic and axonal arborizations of chemically defined types of interneurons. The hatched boxes indicate the preferential axonal distribution for each cell type. The vertically striped boxes indicate that other interneurons, rather than principal cells, are the primary targets. Note that morphologically and chemically defined classes of interneurons largely correspond. Adapted with permission from Freund and Buzsaki (1996).

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excitatory perforant path and commisural/associational inputs. Since these cells are immunoreactive for markers of GABA, such as GAD and parvalbumin, and reportedly form symmetrical synapses with their targets, it is likely that they provide a second means of inhibitory control of granule cell output. A third cell type, that has its cell body in the deep molecular layer, has been named the molecular layer perforant path-associated cell (MOPP) on the basis of its preferential axonal distribution within the outer two-thirds of the molecular layer (Han et al., 1993).

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fibers (Gulyas et al., 1992) and distributing a main axon into the underlying white matter, suggesting that these cells project outside the hippocampus (see Amaral, 1978; Freund and Buzsaki, 1996 for a more detailed description of hilar cells). Finally, chandelier cells have also been described that have a dendritic tree confined to the polymorphic layer and may receive mossy fiber inputs. The axons most likely terminate on the axon initial segments of mossy cells (Martinez et al., 1996).

Connections within the Dentate Gyrus

Cells of the Polymorphic Cell Layer The polymorphic cell layer harbors a variety of neuronal cell types (Amaral, 1978) but little is know about many of them. The most common cell type, and certainly the most impressive, is the mossy cell. The cell bodies of the mossy cells are large (25–35 μm) and are often triangular or multipolar in shape. Three or more thick dendrites originate from the cell body and extend for long distances within the polymorphic layer; occasionally a dendrite will also extend through the granule cell layer and into the molecular layer. The mossy cell dendrites virtually never leave the confines of the polymorphic layer to enter the adjacent CA3 field. Each principal dendrite bifurcates once or twice and generally gives rise to a few side branches. The most distinctive feature of the mossy cell is that all of the proximal dendrites are covered by the very large and complex spines called “thorny excrescences” that are the sites of termination of the mossy fiber axons (Frotscher et al., 1991; Ribak et al., 1985). Although thorny excrescences are also observed on the proximal dendrites of pyramidal cells in CA3, they are far denser on the mossy cells. The distal dendrites of the mossy cell have typical pedunculate spines that are less dense than on the pyramidal cells in the hippocampus. There are also a number of fusiform type cells in the polymorphic layer (Fig. 3). The main difference between the types is the number and density of spines (Ribak and Seress, 1988). The dendrites of these cell types run mainly parallel to the granule cell layer and can extend for nearly the entire length of one blade of the granule cell layer. There is also a group of small, round or multipolar cells in the polymorphic layer that gives rise to a stellate plexus of thin, aspiny dendrites and a local, ramifying axonal plexus. In some instances, such cells have been reported to distribute axons out into the hippocampus proper, even reaching as far as the subiculum (Sik et al., 1997). Also, so-called long-spined multipolar cells have been described with a dendritic tree confined to the hilus, similar to the cells described above, that project to the outer two-thirds of the molecular layer (Han et al., 1993), apparently innervated by mossy

Projections from the Granule Cells The granule cells give rise to distinctive unmyelinated axons which Ramony Cajal called mossy fibers. Each principal mossy fiber (which is on the order of 0.2–0.5 μm in diameter) gives rise to about seven thinner collaterals within the polymorphic layer before entering the CA3 field of the hippocampus (Claiborne et al., 1986). These dentate-to-CA3 projections are described in detail on page 644). Within the polymorphic layer, the mossy fiber collaterals bear two types of synaptic varicosities. There are about 160 small (approximately 2 μm) varicosities distributed throughout the axonal collateral plexus of a single granule cell and these form either thin filopodial-like extensions or en passant contacts on spines and dendritic shafts of presumed interneurons located in the polymorphic layer (Acsady et al., 1998; Claiborne et al., 1986). At the ends of each of the collateral branches there are usually also single, larger (3–5 μm), irregularly shaped varicosities that resemble the mossy fiber terminals found in the CA3 field and contact the hilar mossy cells. It is now clear that the mossy fiber terminals of a single granule cell in the polymorphic layer establish contacts with the proximal dendrites of about 7–12 mossy cells and with the basal dendrites of a much greater number of interneurons such as the pyramidal basket cells and other cells, including cells identified with immunostaining for substance P receptor and mGluR1a receptor (Acsady et al., 1998; Ribak and Seress, 1983; Ribak et al., 1985; Scharfman et al., 1990). Projections from Basket Cells and Other “GABAergic” Interneurons As noted previously, there are a variety of basket cells located close to the granule cell layer. These all appear to contribute to the very dense terminal plexus that is confined to the granule cell layer (Fig. 3). The terminals in this basket plexus are GABAergic and form symmetric, presumably inhibitory, contacts primarily on the cell bodies and shafts of apical dendrites of the granule cells (Kosaka et al., 1984). There is growing

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evidence that GABAergic neurons in the polymorphic layer are themselves innervated by GABAergic terminals (Misgeld and Frotscher, 1986). Given the small number of basket cells relative to granule cells, the question arises as to how widespread is the influence of a single basket cell. Analysis of Golgi-stained axonal plexuses as well as of intracellularly filled basket cells indicates that the axon of a single basket cell may well distribute between 400 μm and 1 mm in the transverse plane and between 1 and 1.5 mm along the septotemporal axis (Sik et al., 1997; Struble et al., 1978). It is conceivable, therefore, that a single basket cell has influence over a very large number of granule cells. A second inhibitory input to granule cells originates from the axoaxonic or “chandelier-type” cells located in the molecular layer (Han et al., 1993; Kosaka, 1983; Soriano and Frotscher, 1989). The axonal plexus, which extends more than 400 μm in all directions, is mainly present in the granular cell layer and adjacent deep portions of the hilus, where symmetric contacts have been reported exclusively onto the axon initial segment (see Freund and Buzsaki, 1996, for further details). Another intrinsic projection within the dentate gyrus was uncovered when antibodies directed against the peptide somatostatin demonstrated a population of immunopositive neurons scattered throughout the polymorphic layer (Bakst et al., 1986; Morrison et al., 1982). These cells give rise to a dense plexus of somatostatin-immunoreactive fibers and terminals in the outer two-thirds of the molecular layer thus coinciding with the terminal distribution of the perforant path (see page 674). Intracellular fillings of such somatostatin-positive neurons that colocalize GABA have shown that their axon may distribute over 3 mm along the septotemporal axis of the dentate gyrus. They form symmetrical contacts exclusively onto the distal dendrites of presumed granule cells (Bakst et al., 1986; Halasy and Somogyi, 1993). This system of fibers, which forms contacts on the apical dendrites of the granule cells, provides a third means for inhibitory control of granule cell activity. More recent findings underscored this idea and showed that the well-known lateral inhibition in the dentate gyrus most likely is brought about by somatostatin-positive interneurons (Boyett and Buckmaster, 2001). Another class of cells with their parent cell bodies close to the granular cell layer or in the polymorphic layer has axons that are distributed specifically to the inner one-third of the molecular layer. The axon from any particular neuron may span as much as 20% of the septotemporal length of the dentate gyrus and forms symmetrical synapses with proximal portions of spiny dendrites of presumed granule cells (Buckmaster and Schwarzkroin, 1995; Sik et al., 1997). These inhibitory axons thus coincide with the terminal

field of the commissural and associational fibers (see section below). Finally, the MOPP cells, described earlier, have axons that distribute within the outer two-thirds of the molecular layer (Han et al., 1993). These cells have a dendritic tree spanning over 800 μm and a slightly more extensively distributed axonal plexus that forms symmetrical synapses onto distal dendrites of presumed granule cells (Freund and Buzsaki, 1996; Halasy and Somogyi, 1993). Several of these interneurons may also contribute collaterals to the commissural connections of the dentate gyrus as well, having a distribution similar to that of their ipsilateral counterpart. (Deller et al., 1995; Zapone and Sloviter, 2001; see section below). The Ipsilateral Associational/Commissural Projection The inner third of the molecular layer receives a projection that originates exclusively from cells in the polymorphic layer (Blackstad, 1956; Laurberg, 1979; Laurberg and Sorensen, 1981; Swanson et al., 1978, 1981; Zimmer, 1971). Since this projection originates both on the ipsilateral and contralateral sides, it has been called the ipsilateral associational/commissural projection. Earlier studies raised the possibility that this projection originated, in part, from cells of the CA3 portion of the hippocampus. The notion that CA3 projects to the dentate molecular layer was based on neuroanatomical studies which employed relatively large injections of anterograde tracers that apparently involved both the CA3 field and cells of the polymorphic layer (see Laurberg (1979) and Laurberg and Sorensen (1981) for a more complete discussion of this issue). It is now clear that CA3 does not send projections to the molecular layer of the dentate gyrus on either side of the brain. The associational/commissural projection appears to be somewhat more complex than initially appreciated. At least four different commissural fiber types have been differentiated on the basis of their laminar termination pattern (Deller et al., 1995, 1996a, 1996c). In addition to the traditionally known fibers to the inner molecular layer, Deller noted fibers to the outer molecular layer, fibers terminating throughout the molecular layer, and fibers terminating in both the granule cell layer and the molecular layer (see also Deller, 1998). Therefore, the overall commissural projection largely mimics the ipsilateral associational projection taking into account the previously described projections of interneurons to the molecular layer (see page 639). With respect to the densest projection to the inner onethird of the molecular layer, it has been shown that many of the axons originate from the large mossy cells of the polymorphic layer and individual mossy cells contribute a projection to both the ipsilateral associational and commissural projections (Frotscher et al.,

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1991; Laurberg and Sorensen, 1981; Ribak et al., 1985; Swanson et al., 1981). Since the mossy cells are immunoreactive for glutamate (Soriano and Frotscher, 1994), and the majority of terminals of this pathway form asymmetric synaptic terminals on spines of the granule cell dendrites (Kishi et al., 1980; Laatsch and Cowan, 1966, 1967), it is likely that they release this excitatory transmitter substance at their terminals within the ipsilateral associational/commissural zone of the molecular layer. There is an interesting facet to the topography of this “feedback” projection from the polymorphic layer to the molecular. Not only does the projection from one septotemporal level distribute widely, both septally and temporally from the point of origin, but also the projection at the level of origin appears to be substantially lighter than to more distant levels of the dentate gyrus (Amaral and Witter, 1989). If we remember that the mossy cells receive a dense innervation from the granule cells at the same level (via the mossy fiber collaterals into the polymorphic layer) it would appear that the mossy cells pass on the collective output of granule cells from one septotemporal level to granule cells located at distant levels of the dentate gyrus. The functional significance of the longitudinal distribution of the associational projection cannot be fully appreciated without one further piece of information. In addition to contacting the spines of dentate granule cells, the associational as well as the commissural fibers also contact the dendritic shafts of GABAergic/parvalbumin-positive basket cells (Deller et al., 1994; Frotscher and Zimmer, 1983; Seress and Ribak, 1984). Thus, the associational and commissural projections may function both as a feed-forward excitatory pathway and as a disynaptic feed-forward inhibitory pathway. It is also interesting to point out that the somatostatin/GABA pathway described above appears to have a more local and limited terminal distribution. Thus, mossy fiber collaterals terminating on GABA/somatostatin cells in the polymorphic layer may lead predominantly to direct inhibition of granule cells at the same septotemporal level and to inhibition of excitation (via the ipsilateral associational connection) at more distant levels of the dentate gyrus. Interestingly, the overall organization of the commissural projections appears strikingly similar. As mentioned before, some of the interneurons of the dentate gyrus reportedly contribute to the commissural projection. Although initially thought to be rather weak (Bakst et al., 1986; Ribak et al., 1986; Seress and Ribak, 1983), more recent findings seem to indicate that this commissural inhibitory projection is more prominent. In contrast to the commissural projections that originate in the hilar mossy cells and show quite extensive septotemporal distributions, these interneuron-

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originating projections exhibit a more restricted commissural distribution along the septotemporal axis (Deller et al., 1995; Zappone and Sloviter, 2001).

Extrinsic Inputs and Outputs The dentate gyrus gives rise to no extrinsic projections other than the mossy fiber projection to the CA3 field of the hippocampus. The major input to the dentate gyrus is from the entorhinal cortex (this is discussed in page 674) and the dentate gyrus receives no direct inputs from other cortical structures. Therefore, the remainder of this section deals with the subcortical inputs to the dentate gyrus, which originate mainly from the septal nuclei, from the supramammillary region of the posterior hypothalamus, and from several monoaminergic nuclei in the brain stem, especially the locus coeruleus and the raphe nuclei (Fig. 5). Projections from the Septal Region The septal projection to the hippocampal formation was first experimentally demonstrated in the early 1950s (Daitz and Powell, 1954; Morin, 1950) and numerous studies have subsequently been conducted to further define the projection (Amaral and Kurz, 1985; Baisden et al., 1979, 1984; Fibiger, 1982; Gage et al., 1984; Lubke et al., 1997; Meibach and Siegel, 1975, 1977a; Milner and Amaral, 1984; Mosko et al., 1973; Swanson, 1978; Swanson and Cowan, 1977; for septohippocampal relations see also Risold, Chapter 20, this volume). The septal projection arises from cells of the medial septal nucleus and the nucleus of the diagonal band of Broca and travels to the hippocampal formation via four routes: the fimbria, dorsal fornix, supracallosal stria, and via a ventral route through and around the amygdaloid complex. Septal fibers terminate in essentially all fields of the hippocampal formation and are particularly prominent in the dentate gyrus (Fig. 4A). In the dentate gyrus, fibers heavily innervate the polymorphic layer, particularly in a narrow infragranular band and terminate more lightly in the molecular layer. In the hilus, these septal fibers impinge on proximal and distal dendrites of hilar mossy cells but do not seem to innervate the cell body. In contrast, aspiny hilar neurons, presumably GABAergic interneurons, receive a septal innervation on their somata and proximal primary dendrites (Lubke et al., 1997). The septal projection to the dentate gyrus and remainder of the hippocampal formation is topographically organized. Cells located medially in the medial septal nucleus tend to project preferentially to septal levels of the hippocampal formation whereas cells located laterally in the septal area tend to project temporally in the hippocampal formation (Gaykema et al., 1990;

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FIGURE 4 (A) This illustration shows line drawings of two coronal sections through the hippocampal region at a rostral (1) and a more caudal (2) level. The distribution of PHA-L-labeled fibers in the hippocampal formation resulting from an injection focused in the medial portion of the medial septal nucleus is diagrammed in these drawings. Reproduced with permission from Gaykema et al. (1990). (B) Line drawings of coronal sections through the hippocampal region at a rostral (1) and a caudal (2) level, illustrating the distribution of PHA-L-labeled fibers resulting from an injection of the supramammillary nucleus. Note the dense plexus of fibers located just superficial to the granule cell layer and in the CA2 region of the hippocampus (between the small arrows). Adapted with permission from Haglund et al. (1984).

Nyakas et al., 1987; Swanson, 1978). Retrograde tracer studies support this notion of a topographical organization, indicating that septal levels of the dentate gyrus and other hippocampal fields obtain most of their input from the nucleus of the diagonal band, in particular from the rostral half of the vertical limb of the diagonal band (VDB) and the core part of the horizontal limb of the diagonal band (HDB), whereas temporal levels of these areas receive their cholinergic innervation primarily from the medial septal nucleus, in particular from the caudal portions of the medial septum and the ventral diagonal band (Amaral and Kurz, 1985; Yoshida

and Oka, 1995). Moreover, Yoshida and Oka (1995) reported that the projection to the hilus preferentially originated from the medial septum, whereas septal projections to the hippocampus proper arose from both the medial septum and the diagonal band. The ventral septal pathway to the hippocampal formation appears to arise mainly from the nucleus of the diagonal band of Broca (Gage et al., 1984; Milner and Amaral, 1984). Lewis and Shute (1967) were the first to propose that the septohippocampal projection was cholinergic after showing that transsection of the fimbria led to a substantial loss of histochemical staining of the enzyme

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acetylcholinesterase. Using immunohistochemical procedures to demonstrate the specific cholinergic marker, choline acetyltransferase, in conjunction with retrograde tracer techniques several investigators (Amaral and Kurz, 1985; Wainer et al., 1984, 1985) have found that 30–50% of the cells in the medial septal nucleus and 50–75% of the cells in the nucleus of the diagonal band that project to the hippocampal formation are cholinergic. These retrograde tracer studies emphasized the fact that only a portion of the septal cells that project to the dentate gyrus and other (para)hippocampal structures are cholinergic. Many of the septal cells that project to the dentate gyrus and other hippocampal fields contain glutamic acid decarboxylase and are presumably GABAergic (Köhler et al., 1984a). These GABAergic projections terminate in the dentate gyrus and also in the fields of the hippocampus and of the parahippocampal region with an apparent preference for the GABAergic nonpyramidal cells (Freund and Antal, 1988). Many of the GABAergic cells that receive GABAergic septal projections are also immunoreactive for cholecystokinin, somatostatin, or VIP (Gulyas et al., 1990) or contain one of the calcium-binding proteins (calretinin, calbindin, or parvalbumin; Acsády et al., 1993; Freund and Antal, 1988). Taken together, these findings indicate that the cholinergic and GABAergic septal projections not only innervate different cell types but also innervate them differently. A final observation that is of interest is that cells in the medial septum as well as their potential target cells in the dentate gyrus receive collaterals from the same cells in the medial raphe nucleus (McKenna and Vertes, 2001; see also below). Inputs from the Supramammillary Region The major hypothalamic projection to the dentate gyrus arises from a population of large cells that cap and partially surround the mammillary nuclei in a zone that has been named the supramammillary area (Dent et al., 1983; Haglund et al., 1984; Riley and Moore, 1981; Segal and Landis, 1974; Vertes, 1993; Wyss et al., 1979a, 1979b). The supramammillary projection terminates heavily in a narrow zone of the molecular layer located just superficial to the granule cell layer (Fig. 4B); there is only light innervation of the polymorphic layer or remaining parts of the molecular layer. The neurons in the supramammillary nucleus that originate these projections are to a large extent calretin-positive and their asymmetrical, presumably excitatory, terminals (Borhegyi and Leranth, 1997; Kiss et al., 2000; Magloczky et al., 1994; Nitsch and Leranth, 1996) reportedly target not only dentate granule cells but also inhibitory interneurons such as basket cells and calbindin-positive interneurons (Nitsch and Leranth, 1996; see, however, Magloczky et al., 1994). Moreover,

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the supramammillary projections to the dentate gyrus are bilateral and partially originate as collaterals from cells that also project to the medial septum (Borhegyi et al., 1998; Vertes and McKenna, 2000). In addition to the supramammillary cells, there are cells scattered in several hypothalamic nuclei, many of which are in a perifornical position or within the lateral hypothalamic area, that project to the hippocampal formation. Taken together, these cells constitute a sizable input to the hippocampal formation. However, their diffuseness and lack of any distinguishing biochemical marker has made it difficult to study their projections. A subset of these cells demonstrate α-melanocytestimulating hormone-like immunoreactivity and give rise to a diffusely distributed pathway to several fields of the hippocampal formation (Köhler, 1984). Similarly distributed diffuse projections from this region are immunoreactive for substance P (Davies and Köhler, 1985; Gall and Selawski, 1984) or for angiotensin II (Wayner et al., 1997). Monoaminergic Inputs from the Brain Stem The dentate gyrus receives a particularly prominent noradrenergic input primarily from the pontine nucleus locus coeruleus (Haring and Davis, 1983, 1985a, 1985b; Koda et al., 1978a, 1978b; Moore et al., 1978; Pickel et al., 1974; Swanson and Hartman, 1975). As illustrated in Fig. 5, the noradrenergic fibers terminate mainly in the polymorphic layer of the dentate gyrus. An additional input originates from the subcoeruleus nucleus (Datta et al., 1998) which plays a prominent role in generating pontogeniculooccipital (PGO) waves during sleep. The serotonergic projection that originates from several subdivisions of the raphe nuclei also terminates most heavily in the polymorphic layer but the projection tends to be limited to an immediately subgranular portion of the layer (Conrad et al., 1974). It is important to point out that the raphe serotonergic fibers preferentially terminate on a class of interneurons in the dentate gyrus which primarily influence the distal dendrites of the granule cells (Halasy et al., 1992). As with the cholinergic projection, many of the cells in the raphe nuclei that project to the hippocampal formation appear to be nonserotonergic (Köhler and Steinbusch, 1982; Montone et al., 1988). Raphe projections to the dentate gyrus, preferentially originate from the medial raphe nucleus (Vertes et al., 1999). Interestingly, median raphe cells also project to the medial septum where they may be able to modulate hippocampal EEG activity (Acsady et al., 1996; McKenna and Vertes, 2001). The dentate gyrus receives a minor and diffusely distributed projection from cells located in the ventral tegmental area, with minor components originating in the retrorubral field A8 and in the substantia nigra (Gasbarri et al.,

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FIGURE 5 Line drawing of horizontal sections through the hippocampal formation showing the distribution of noradrenergic (A), serotonergic (B), or dopaminergic (C) fibers. Adapted with permission from Swanson (1987).

1994a, 1994b, 1996, 1997; Swanson, 1982; Fig. 5C). Some of the cells in these regions are likely to be dopaminergic (Gasbarri et al., 1994b).

Intrahippocampal Connections (Mossy Fibers) As noted above, the only extra dentate projection of the dentate gyrus is the mossy fiber projection to the CA3 field of the hippocampus. Once the mossy fiber enters the CA3 region, it has few collaterals. Rather, as Claiborne et al. (1986) noted, the large (3–6 μm in diameter) presynaptic varicosities characteristic of mossy fiber/CA3 pyramidal cell contacts (mossy fiber expansions) are distributed at approximately 140-μm intervals along the course of the mossy fiber axons. It should be noted that regardless of the location of the granule cell of origin, all mossy fibers extend throughout the full transverse extent of the CA3 field. Proximally in CA3, some mossy fibers travel deep to the pyramidal cell layer in what has been called the infrapyramidal bundle. Other fibers travel within the pyramidal cell layer in the intrapyramidal bundle and yet others take a course superficial to the pyramidal cell layer in the

suprapyramidal bundle that forms the major constituent of stratum lucidum. Fibers from all bundles, however, ultimately attain stratum lucidum and stop at the CA3/CA2 border. The mossy fiber presynaptic expansion forms a unique synaptic complex with an equally intricate postsynaptic process called the “thorny excrescence.” These spine-like processes are large, multilobulated entities (with as many as 16 branches) that are surrounded by a single mossy fiber expansion. A single mossy fiber expansion can make as many as 37 synaptic contacts with a single CA3 pyramidal cell dendrite (Chicurel and Harris, 1992). Because of the large size and proximal dendritic location of the mossy fiber synapse, the granule cells are in a unique position to influence the activity of hippocampal pyramidal cells. As summarized more completely elsewhere (Amaral et al., 1990) mossy fibers contact relatively few pyramidal cells. Claiborne et al. (1986) have estimated that each mossy fiber demonstrates approximately 14 synaptic expansions along its course in stratum lucidum and since each of these expansions contacts a single pyramidal cell (or at most two dendrites from adjacent pyramidal cells)

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each granule cells is likely to influence only 14–28 pyramidal cells. Acsady et al. (1998) have confirmed and extended these findings and noted that, while a single granule cell has 11–15 mossy fiber expansions within the CA3 region, there are 7–12 contacts with mossy cells in the polymorphic layer. Conversely, each pyramidal cell has been estimated to receive contacts from something on the order of 50 granule cells. In addition to contacting principal CA3 neurons, the mossy fibers also contact dendrites of a number of GABAergic interneurons via thin filopodial extensions of the mossy fiber terminals as well as en passant synaptic varicosities (Acsady et al., 1998) The mossy fibers have a relatively “lamellar” trajectory and in this regard are atypical for hippocampal intrinsic connections. Throughout much of their course through the stratum lucidum, they remain at approximately the same septotemporal level as the cells of origin (Claiborne et al., 1986; Gaarskjaer, 1978a, 1978b; Swanson et al., 1978). Near the CA3/CA2 border, however, the mossy fibers make an abrupt turn temporally and extend for 1 mm or more toward the temporal pole. The significance of this component of the mossy fibers has never been adequately addressed. On the face of it, granule cells from a much wider septotemporal extent of the dentate gyrus may influence the most distal portion of CA3 than is the case for more proximal portions of CA3. However, it has as yet to be established that these temporally directed mossy fibers do make contact with pyramidal cells at more temporal levels. As in the other principal intrinsic connections of the rat hippocampal formation, the mossy fibers are thought to use glutamate (Storm-Mathisen and Fonnum, 1972) as a primary transmitter substance. However, it is also clear that some mossy fibers harbor opiate peptides such as dynorphin (Gall, 1984; Gall and Moore, 1984; McGinty et al., 1983, 1984; Roberts et al., 1983; van Daal et al., 1989).

HIPPOCAMPUS Definitions and Distinctions of Fields CA3, CA2, and CA1 The hippocampus proper can clearly be divided into two major regions, a large-celled proximal region and a smaller-celled distal region. Ramón y Cajal called these two regions regio inferior and regio superior, respectively. The terminology of Lorente de Nó, however, has achieved more common usage. He divided the hippocampus into three fields (CA3, CA2, and CA1). His CA3 and CA2 fields are equivalent to the large-celled regio inferior of Ramon y Cajal and his CA1 is

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equivalent to regio superior. In addition to differences in the size of the pyramidal cells in CA3 and CA1 there is a clear-cut connectional difference. The CA3 pyramidal cells receive a mossy fiber input from the dentate gyrus and the CA1 pyramidal cells do not. The CA2 field has been a matter of some controversy. As originally defined by Lorente de Nó, it was a narrow zone of cells interposed between CA3 and CA1 that had large cell bodies like CA3 but did not receive mossy fiber innervation like CA1 cells. Although the validity of this subdivision has often been questioned, the bulk of available evidence indicates that there is a narrow CA2 that has both connectional and perhaps even functional differences with the other hippocampal fields. In many respects, CA2 resembles a terminal portion of the CA3 field. In other ways, however, CA2 is quite distinct from either CA3 or CA1 (see below).

Laminar Organization The laminar organization is generally similar for all of fields of the hippocampus. The principal cellular layer is called the pyramidal cell layer. The narrow, relatively cell-free layer located deep to the pyramidal cell layer is called stratum oriens and deep to this is the fiber-containing alveus. In the CA3 field, but not in CA2 or CA1, a narrow acellular zone located just above the pyramidal cell layer is occupied by the mossy fiber axons originating from the dentate gyrus. This narrow layer is called stratum lucidum. At the distal end of the stratum lucidum there is a slight thickening of the layer where the mossy fibers bend temporally. This “end bulb” (which is more prominent in species such as the guinea pig) marks the CA3/CA2 border. Superficial to the stratum lucidum in CA3, and immediately above the pyramidal cell layer in CA2 and CA1, is the stratum radiatum. The stratum radiatum can be defined as the suprapyramidal region in which CA3 to CA3 associational connections and CA3 to CA1 Schaffer collateral connections are located. The most superficial portion of the hippocampus is called the stratum lacunosummoleculare. It is in this layer that perforant pathway fibers from the entorhinal cortex travel and terminate although afferents from other regions, such as those from the nucleus reuniens of the midline thalamus, also terminate in this layer.

Cell Types and Local Interconnections The principal neuronal cell type of the hippocampus is the pyramidal cell, which makes up the vast majority of neurons in the pyramidal cell layer. Pyramidal cells have a basal dendritic tree that extends into stratum oriens and an apical dendritic tree that extends to the hippocampal

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fissure (Fig. 6). Lorente de Nó (1934) noted a variety of subtle differences in the dendritic organization of pyramidal cells in different parts of CA3 and CA1 and used these distinctions, in part, to further subdivide these fields into three subareas each (CA3a,b,c; CA1a,b,c). The length and organization of CA3 pyramidal cells vary consistently related to the position along the transverse axis of CA3. Cells in the proximal part, close to the dentate gyrus, have the smallest overall dendritic trees with about 10 mm in total length. Cells in the distal part of CA3 (near CA2) have the largest dendritic tree showing total dendritic lengths on the order of 16 mm. CA2 contains a mixed population of both large cells, similar in dimensions to those of the distal CA3 as well as cells having smaller dendritic trees, resembling CA1 pyramidal neurons. The dendritic trees of CA1 pyramidal cells have an average length of 13.5 mm (Ishizuka et al., 1995). In Fig. 6, computer reconstructions of a CA3 pyramidal cell and a CA1 pyramidal cell are drawn on a schematic diagram of the hippocampus. The lengths of different components of the dendritic tree are indicated on this illustration and it serves to point out that a substantial portion of the dendritic tree is located in

the stratum oriens. In the case of CA3 pyramidal cells, 42–51% of the dendritic tree is in the stratum oriens; in the case of CA1 pyramidal neurons this is around 34%. Moreover, nearly 18% of the dendritic tree of the typical CA1 pyramidal cell is located in the stratum lacunosummoleculare, i.e., in the terminal region of the perforant pathway. The amount of dendrites in the stratum lacunosum-moleculare ranged from none for those pyramidal cells located within the confines of the hilus to about the same percentage as in the stratum radiatum for the distal CA3 and CA2 pyramidal cells (Ishizuka et al., 1995). In addition to the pyramidal cells, there is a heterogenous population of basket cells of various sizes and shapes which have their cell bodies located in the pyramidal cell layer (Freund and Buzsaki, 1996; Seress and Ribak, 1985). These also have apical and basal dendritic trees (Fig. 7), like the pyramidal cells, but their dendrites have few if any dendritic spines and have an overall beaded appearance. The pyramidal basket cell axon extends transversely from the cell body of origin and forms a basket plexus that innervates the cell bodies of the hippocampal pyramidal cells.

FIGURE 6 Line drawing illustrating the size and shape of granule cells in the dentate gyrus and pyramidal cells in the CA3 and CA1 fields of the hippocampus. Numbers located near two of the granule cells in the dentate gyrus indicate total dendritic length. Numbers in boxes beside the pyramidal cells indicate total dendritic length while smaller numbers indicate summed dendritic length in the stratum oriens, the stratum radiatum, or the stratum lacunosum-moleculare. Data are derived from intracellular HRP injections in the in vitro slice preparation by Brenda Claiborne (granule cells) and Norio Ishizuka (pyramidal cells).

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FIGURE 7 Summary diagram of interneurons of CA3 and CA1. Filled circles indicate the location of the cell bodies and thick lines indicate the predominant orientation and laminar distribution of the dendritic tree. Indicated in the background are the principal neurons, providing an indication of which domain is innervated by the different interneuron groups. Also indicated is the laminar distribution of different inputs, often showing a striking match with interneuron type or axon distribution. (A) Morphological classification scheme. Interneuron types are identified according to dendritic and axonal arborization. The hatched boxes indicate the preferential axonal distribution for each cell type. (B) Laminar distribution of dendritic and axonal arborizations of chemically defined types of interneurons. The hatched boxes indicate the preferential axonal distribution for each cell type. The vertically striped boxes indicate that other interneurons, rather than principal cells, are the primary targets. Note that morphologically and chemically defined classes of interneurons largely correspond. Adapted with permission from Freund and Buzsaki (1996).

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The early Golgi studies of Ramon y Cajal and Lorente de Nó made it abundantly clear that there are a variety of nonpyramidal cell types in the stratum oriens, stratum radiatum and stratum lacunosum-moleculare of the hippocampus (Fig. 7). The vast majority of these neurons are immunoreactive for markers of GABA (Ribak et al., 1978) and most are considered to be local circuit neurons. Many of these neurons can also be visualized with antibodies to other neuroactive substances such as peptides (Morrison et al., 1983) or calciumbinding proteins or can be characterized by their specific dendritic and axonal distribution (cf. Freund and Buzsaki, 1996, for a comprehensive review). The overall composition of the classes of interneurons in the hippocampus is quite similar to that described for the dentate gyrus (compare Figs. 3 and 7). One class of these cells, the so-called O-LM cells, has dendrites in the stratum oriens and an axon that innervates the stratum lacunosum-moleculare (Blasco-Ibanez and Freund, 1995), Like their dentate gyrus counterparts, these cells generally appear to contain somatostatin and in some instances NPY. In contrast to the cells in the dentate gyrus, the axonal distribution of these cells in the hippocampus is quite limited along the septotemporal (less than 700 μm) and transverse axes (less than 400 μm; Han et al., 1993; Sik et al., 1995). These cells form symmetrical synapses with the spines of dendrites of presumed pyramidal cells. Most of these spines have additional asymmetrical synapses onto them, indicative of excitatory input to converge onto a single spine with inhibitory input from local interneurons (Gulyas et al., 1993; Sik et al., 1995). A second type of interneuron commonly found in the hippocampus, the so-called bistratified neuron, distributes its major axonal plexus to the stratum radiatum and stratum oriens, innervating dendrites of pyramidal cells, coinciding with the major intrinsic hippocampal projection systems (see Section III.D.1; Buhl et al., 1994; Gulyas and Freund, 1996; Miles et al., 1996; Sik et al., 1995). The transverse and septotemporal axonal spread ranges between 1 and 2 mm (Sik et al., 1995). Also, interneurons with a comparable axonal distribution and an additional terminal plexus in the stratum pyramidale (trilaminar neurons) have been described. These so-called trilaminar cells, of which the intrinsic axonal distribution may extend for 2.5 mm in the septotemporal and transverse axes, have been reported to give rise to axonal collaterals targeting extrahippocampal targets as well (Toth and Freund, 1992). Both the bistratified and trilaminar cells form symmetrical synapses onto dendrites of presumed pyramidal cells (Halasy et al., 1996; Sik et al., 1995). Other interneurons have their cell bodies in either the stratum pyramidale or the stratum radiatum and give rise to an axonal projection that terminates mainly in

the stratum radiatum; these have a rather limited local axonal distribution with symmetrical synapses onto spiny dendrites of pyramidal cells (Gulyas et al., 1993). All hippocampal fields also contain chandelier cells with dendrites running parrallel to the dendrites of pyramidal cells and thus likely to receive inputs similar to those terminating on pyramidal cells. The axons of these cells innervate the initial axonal segment of pyramidal cells (Martinez et al., 1996). Finally, interneurons having their cell bodies as well as most of their dendritic arbor within the stratum lacunosum-moleculare have been reported and therefore these interneurons most likely receive excitatory inputs from the entorhinal cortex (Desmond et al., 1994; Witter et al., 1992) as well as from the nucleus reuniens thalami (Dolleman-van der Weel et al., 1997; Dolleman-van der Weel and Witter, 2000). It is of interest to note that some of the interneurons in CA1 tend to distribute a rather extensive axonal arbor along the transverse axis of the hippocampus, reaching not only into CA1 but also into CA3 and the hilus of the dentate gyrus. Such cells typically occur at the stratum oriens/alveus border, having a horizontal dendritic tree mainly confined to the stratum oriens. The axons of these cells reportedly form symmetrical synapses onto dendrites of principal cells and provide inhibitory feedback which is contrary to the overall unidirectionality of the main hippocampal connectivity (Freund and Buzsaki, 1996; Sik et al., 1994). Although a lot of detailed information on the organization of hippocampal interneurons has become available over the last decade, their overall function in the neuronal network of the hippocampus is still not fully understood. Interneurons may be excited or inhibited by an as yet incompletely understood but obviously highly variable number of excitatory and inhibitory inputs that may be different for different subclasses of interneurons as shown for parvalbumin, calbindin, or calretinin containing neurons (Gulyas et al., 1999). Moreover, the overall effect of interneuron transmission may be strikingly different depending on the temporal dynamics of interneurons and the postsynaptic target with which they make synaptic contact (Gupta et al., 2000). Also it appears that in CA1, the proximal portion of the apical dendrite of pyramidal cells, up to 200 μm away from the soma, receives preferential input from GABAergic interneurons whereas more distal portions receive increasing densities of excitatory inputs (Papp et al., 2001). To further complicate the issue, recent findings seem to indicate that even in the adult brain GABAergic transmission does not necessarily result in inhibition of the postsynaptic target. In some instances, GABAergic transmission may actually lead to local depolarization and thus may enhance the probability for action potential generation (Gulledge and Stuart, 2003).

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Connections of the Hippocampus Intrahippocampal Connections of CA3 The CA3 pyramidal cells give rise to highly collateralized axons that distribute fibers both within the hippocampus (to CA3, CA2, and CA1) and to the same fields in the contralateral hippocampus (the commissural projections). CA3 cells, especially those located proximally in the field, and CA2 cells contribute a small number of collaterals that innervate the polymorphic layer of the dentate gyrus. While claims of other hippocampal connections are to be found in the literature (Swanson et al., 1978), it is now quite clear that CA3 does not project to the subiculum, the presubiculum, the parasubiculum, or the entorhinal cortex. All of the CA3 and CA2 pyramidal cells give rise to highly divergent projections to all portions of the hippocampus (Ishizuka et al., 1990). The projections to CA3 and CA2 are typically called the associational connections and the CA3 projections to the CA1 field are typically called the Schaffer collaterals. Using the discrete anterograde tracer PHA-L a highly ordered pattern of projections from CA3 to CA3 and to CA1 has been discovered (Ishizuka et al., 1990). A full description of the various gradients of axonal distribution of the CA3-

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to-CA1 and CA3-to-CA3 connections is beyond the scope of this chapter (a more detailed description is available in Ishizuka et al., 1990). The essential elements of the organization of these connections would include the following (a schematic representation of the connections is illustrated in Fig. 8). All portions of CA3 and CA2 project to CA1 but the distribution of terminations in CA1 depends on the transverse location of the CA3/CA2 cells of origin (Fig. 8). The older notion that a typical CA3 pyramidal cell sends a single axon to CA1 that travels linearly through the field with equal contact probability at all regions within CA1 is clearly incorrect. The topographic organization of projections from CA3 to CA1 determines a network in which certain CA3 cells are more likely to contact certain CA1 cells. CA3 cells located close to the dentate gyrus (proximal CA3), while projecting both septally and temporally for substantial distances, tend to project to levels of CA1 located septal to their location. CA3 cells located closer to CA1, in contrast, project more heavily to levels of CA1 located temporally. At, or close to, the septotemporal level of the cells of origin, those cells located proximally in CA3 give rise to collaterals that tend to terminate superficially in the stratum radiatum with a clear preference for the

FIGURE 8 This diagram illustrates the organization of projections from the CA3 to the CA1 fields of the hippocampus. The location of cells of origin is indicated in the middle coronal section and the distribution of fibers and terminals resulting from cells in the positions marked by triangles is indicated by shading patterns similar to those in the triangles.

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more distal portions of CA1 (near the subicular border). Conversely, cells of origin located more distally in CA3 give rise to projections that terminate deeper in the stratum radiatum and in the stratum oriens, with a preference for portions of CA1 located closer to the CA2 border (Fig. 8). Regardless of the septotemporal or transverse origin of a projection, the highest density of terminal and fiber labeling in CA1 shifts to deeper parts of the stratum radiatum and stratum oriens at levels septal to the cells of origin and shifts out of the stratum oriens and into superficial parts of the stratum radiatum at levels temporal to the origin (Fig. 8). Moreover, the highest density of fiber and terminal labeling in CA1 shifts proximally (toward CA3) at levels septal to the origin and distally (toward the subiculum) at levels temporal to the origin. Beyond the orderliness apparent in these hippocampal intrinsic connections, these gradients of connectivity have a number of functional implications. It is clear, for example, that each individual CA3 neuron gives rise to a complex, multibranched axonal plexus that terminates over a broad region of the ipsilateral and contralateral CA1 (Li et al., 1994). This axonal plexus may make several synapses on individual CA1 pyramidal cell dendritic trees (Sorra and Harris, 1993). It is also apparent that individual CA3 axon collaterals take both linear and oblique courses through CA1 and that the same CA3 axon may terminate on the apical dendrite of one neuron and the basal dendrite of another. Although Schaffer collaterals are typically illustrated as extending only through the stratum radiatum, it should be emphasized that both the stratum radiatum and the stratum oriens of CA1 are heavily innervated by CA3 axons. Thus, the Schaffer collaterals are as highly associated with the apical dendrites of CA1 cells in the stratum radiatum as they are with the basal dendrites in the stratum oriens. Associational Connections of CA3 The associational projections from CA3 to CA3 are also organized in a highly systematic fashion. One somewhat idiosyncratic facet of this projection is that cells located proximally in CA3 only communicate with other cells in the proximal portion of CA3 of the same and adjacent septotemporal levels. Associational projections arising from mid and distal portions of CA3, however, project throughout much of the transverse extent of CA3 and also project much more extensively along the septotemporal axis (Ishizuka et al., 1990). The density of CA3 associational projections also shifts along the septotemporal axis. The radial gradient of termination (superficial to deep in the stratum radiatum and stratum oriens) is similar to that described for the CA3-to-CA1 projection. The transverse gradient, however, is the

reverse; CA3 projections shift proximally in CA3 at levels located temporal to the cells of origin and shift distally in CA3 at more septal levels. An important feature of the CA3-to-CA3 associational and CA3-to-CA1 Schaffer collateral projections is that they are both divergently distributed along the septotemporal axis. In a technically demanding series of studies, Tamamaki et al. (1984, 1988) have demonstrated that single CA3 and CA2 pyramidal cells give rise to highly arborized axonal plexuses that distribute to as much as 75% of the septotemporal extent of the ipsilateral and contralateral CA1 fields. These data argue strongly against the notion that information processing within the hippocampus progresses in a chiplike or lamellar fashion. Rather, the distribution of information flow appears to be as extensive in the septotemporal axis as it is in the transverse axis. It should be noted however that the overall functional connectivity of the network still may favor a more restricted, i.e., lamellar, transfer of activity in the CA3 and CA1 pathways (Bernard and Wheal, 1994; Hampson et al., 1999; see page 685). As mentioned before, there is evidence that CA3 is innervated not only by the massive mossy fiber input from the dentate gyrus but also receives additional sparse inputs from interneurons in CA1 (Sik et al., 1994). Commissural Projections of CA3 In the rat, but not in the monkey (Amaral et al., 1984; Demeter et al., 1985), the CA3 pyramidal cells give rise to commissural projections to the CA3, CA2, and CA1 regions of the contralateral hippocampal formation (Swanson et al., 1978). The same CA3 cells give rise to both ipsilateral and commissural projections (Swanson et al., 1980). While the commissural projections roughly follow the same topographic organization and generally terminate in homologous regions on both sides, there are minor differences in the distribution of terminals. If a projection is heavier to the stratum oriens on the ipsilateral side, for example, it may be heavier in the stratum radiatum on the contralateral side (Swanson et al., 1978). As with the commissural projections from the dentate gyrus, CA3 fibers to the contralateral hippocampus form asymmetric synapses on the spines of pyramidal cells in CA3 and CA1 (Gottlieb and Cowan, 1972) but also terminate on the smooth dendrites of interneurons (Frotscher et al., 1984). Extrinsic Inputs and Outputs of CA3 Until the mid 1970s, it was commonly assumed that the hippocampal fields (CA1–CA3) gave rise to both the precommissural and postcommissural hippocampal projections that terminate in the basal forebrain and diencephalon. But Swanson and Cowan (1975)

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demonstrated that most of these projections actually originate from the subiculum. It is now quite clear that the only sizable subcortical projection from CA3 is to the lateral septal nucleus (Swanson and Cowan, 1977). The CA3 projection to the septal complex is distinct from other hippocampal projections to the septal region in that it is bilateral. Some CA3 fibers cross in the ventral hippocampal commissure to innervate the homologous region of the contralateral lateral septal nucleus. This pathway is topographically organized; septal portions of CA3 project dorsally in the lateral septal nucleus and progressively more temporal portions of CA3 project more ventrally; proximal CA3 cells tend to project medially in the lateral septal nucleus and distally situated CA3 cells terminate more laterally (Gaykema et al., 1991; Risold and Swanson, 1997; Siegel et al., 1974, 1975; Swanson and Cowan, 1977). Interestingly, essentially all of the CA3 cells give rise to projections both to-CA1 and to the lateral septal nucleus (Swanson et al., 1980). Moreover, the distribution of the projections from CA3 to the lateral septum differs from that of the CA1 projections, in that those of CA3 preferentially project to more caudal levels (Risold and Swanson, 1997). Finally, it should be noted that at least some of the hippocampal neurons that project to the septal region are GABAergic (Toth and Freund, 1992). The septal nucleus also provides the major subcortical input to CA3. As with the dentate gyrus, the septal projection originates mainly in the medial septal nucleus and nucleus of the diagonal band of Broca, both the horizontal and vertical limbs (Yoshida and Oka, 1995). The projection appears to terminate most heavily in the stratum oriens and to a lesser extent in the stratum radiatum (Fig. 4A) (Gaykema et al., 1990; Nyakas et al., 1987). As in the dentate gyrus, the GABAergic component of the septal projection to the CA3 field terminates mainly on GABAergic interneurons (Freund and Antal, 1988; Gulyás et al., 1990). Sensitive tracing methods have recently shown that CA3, in particular the temporal parts, also receives inputs from the amygdaloid complex, traditionally thought to distribute projections mainly to CA1 and subiculum. These inputs originate mainly from the caudomedial portion of the parvicellular division of the basal nucleus and terminate heavily in the stratum oriens and the stratum radiatum (Pikkarainen et al., 1999). Additional but very weak inputs to the temporal part of CA3 have been described to originate from the piriform nucleus (Behan and Haberly, 1999). The CA3 field also receives inputs from the noradrenergic nucleus locus coeruleus (Swanson et al., 1987) as well as from the nucleus subcoeruleus (Datta et al., 1998). Noradrenergic fibers and terminals are most densely distributed in the stratum lucidum and in the

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most superficial portion of the stratum lacunosummoleculare (Fig. 5). A much thinner plexus of axons is distributed throughout the other layers of CA3. Serotonergic fibers are diffusely and sparsely distributed in CA3 (Acsady et al., 1996; Vertes et al., 1999) and at least to some extent arise as collaterals from cells targeting the medial septum as well, similar to what has been reported for the raphe projections to the dentate gyrus (McKenna and Vertes, 2001). In CA3, there are few, if any, dopaminergic fibers (cf. Swanson et al., 1987). As in the dentate gyrus, the serotonergic fibers, while diffusely distributed, nonetheless appear to terminate preferentially on interneurons (Freund et al., 1990) with axons that innervate the distal dendrites of pyramidal cells. Connections of CA2 As noted above, there has been substantial controversy concerning whether a CA2 region is identifiable in the rat hippocampus. Several lines of evidence indicate that the CA2 region can be distinguished on a number of grounds. The field CA2 is a relatively narrow field that is located distal to the end bulb of the mossy fiber projection and is typically no longer than approximately 250 μm. It is made up of large, darkly staining pyramidal cells, like those of CA3. However, the pyramidal cells of CA2 lack the thorny excrescences that are characteristic for CA3 pyramidal cells (Lorente de Nó, 1934; Tamamaki et al., 1988). A number of immunohistochemical studies have also demonstrated differential labeling of CA2. This region demonstrates denser acetylcholinesterase staining (Paxinos and Watson, 1982) and much denser labeling for the calcium-binding protein parvalbumin than adjacent regions of CA3 or CA1 (Baimbridge and Miller, 1982; Leranth and Ribak, 1991; Sloviter et al., 1991). This is of interest since the calcium-binding proteins are considered to be protective of ischemic or excitotoxic cell death and the CA2 region is purported to be the “resistant sector” described in the human epilepsy literature (Corsellis and Bruton, 1983). The intrahippocampal connections of CA2 resemble, in part, those of the distal portions of CA3 but there are also some distinguishing characteristics. Like CA3, the CA2 cells give rise to a projection to CA1 (Ishizuka et al., 1990). The projection is rather sparse and diffuse, however, and does not clearly follow the gradient rules established by the CA3-to-CA1 projection. Interestingly, more collaterals from CA2 are distributed to the polymorphic layer of the dentate gyrus than from any portion of CA3. There has been little work dealing specifically with the extrinsic inputs and outputs of CA2. In general CA2 appears to share the connections of CA3. However, as illustrated in Fig. 4B, the CA2 field appears to receive

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a particularly prominent innervation from parts of the posterior hypothalamus, in particular from the supramammillary area (Haglund et al., 1984), and from the tuberomammillary nucleus (Köhler et al., 1985). Interestingly, a number of the supramammillary cells projecting to CA2/CA3 distribute collaterals to either the medial septum or the contralateral CA2/CA3 region (Borhegyi et al., 1998; Vertes and McKenna, 2000). This projection, which is most likely excitatory and glutamatergic, (Kiss et al., 2000), appears to terminate exclusively on principal neurons in the hippocampus, similar to what has been reported for the dentate gyrus (Magloczky et al., 1994). These neurons also stain positively for either substance P or calretinin, and it has been shown that the neurons that are immunopositive for substance P selectively project to CA2, whereas the calretinincontaining neurons distribute axons more widely throughout the hippocampus (Borhegyi et al., 1997). Intrahippocampal Connections of CA1 Certain interneurons in CA1 project extensively to CA3 and the hilus of the dentate gyrus. These cells typically are located at the stratum oriens/alveus border and have a horizontal dendritic tree that is mainly confined to the stratum oriens. The axons of these cells form symmetrical synapses onto dendrites of principal cells and thus provide inhibitory feedback (Sik et al., 1994; Freund and Buzsaki, 1996). The major projection arising from the CA1 field, however, is a topographically organized projection to the adjacent subiculum. Axons of CA1 pyramidal cells descend into the stratum oriens or the alveus and bend sharply toward the subiculum (Amaral et al., 1991; Finch and Babb, 1981; Finch et al., 1983; Tamamaki et al., 1988). The fibers reenter the subiculum and ramify profusely in the pyramidal cell layer and in the deep portion of the molecular layer. Unlike the CA3 to CA1 projection that distributes throughout CA1 in a gradient fashion, the CA1 projection ends in a highly topographic columnar fashion in the subiculum (Fig. 9). CA1 cells located proximally in the CA1 field project to the distal third of the subiculum whereas CA1 cells located distally in the field project just across the border into the proximal portion of the subiculum; the mid portion of CA1 projects to the mid portion of the subiculum (Amaral et al., 1991). Intracellular labeling of single CA1 pyramidal cells with horseradish peroxidase demonstrates that individual axonal plexuses distribute to about onethird of the width of the subicular pyramidal cell layer (Tamamaki et al., 1988). It would appear, therefore, that the CA1-to-subiculum projection segments these structures roughly into thirds. Interestingly, this organization of CA1-to-subicular connectivity appears in register with the origin and terminal distribution of the

reciprocal connections with the entorhinal cortex (see pages 675 and 688). Although the CA1-to-subiculum projection has been described as lamellar (Swanson et al., 1987), more recent analyses have indicated that any particular level of CA1 projects to about one-third of the septotemporal extent of the subiculum (Fig. 9). Thus, the CA1 to subiculum projection, like the CA3to-CA1 projection, appears to be organized in a divergent fashion. Finally it should be noted that CA1 receives a small input originating in the subiculum (Commins et al., 2002; Harris and Stewart, 2001; Harris et al., 2001; Seress et al., 2002; see page 659). Associational/Commissural Connections of CA1 Unlike the CA3 field, pyramidal cells in CA1 do not appear to give rise to collaterals that distribute within CA1; i.e., there is only a weak associational connection (Amaral et al., 1991; Tamamaki et al., 1987). As the CA1 axons travel in the alveus or in the stratum oriens toward the subiculum, occasional collaterals arise and appear to enter the stratum oriens and the pyramidal cell layer. It is conceivable, therefore, that these collaterals might be the substrate for projections from CA1 pyramidal cells to the basal dendrites of other CA1 cells. Among the receiving cells, most likely are interneurons such as basket cells in the stratum oriens (Fig. 7) that have been shown to receive direct input from CA1 pyramidal cells, in turn inhibiting CA1 pyramidal cells (Lacaille et al., 1989; Schwartzkroin and Kunkel, 1982, 1985; Schwarzkroin and Mathers, 1978; Schwartzkroin et al., 1990; cf. Freund and Buzsaki, 1996; see page 645). What is clear, however, is that the massive associational network, which is so apparent in CA3, is missing in CA1. Similarly, although a weak commissural projection to the contralateral CA1 appears to be present (Van Groen and Wyss, 1990b), no extensive commissural projection like that existing in the case of CA3 exists. This striking difference in intrinsic organization between CA3 and CA1 has been taken to indicate that they may serve different functional processes within the hippocampal formation related to learning and memory (Treves and Rolls, 1992). Extrinsic Inputs and Outputs of CA1 The CA1 field receives both cortical and subcortical inputs from a variety of structures. The main cortical inputs arise from the parahippocampal region and are described in the sections on “Entorhinal Cortex” and “Perirhinal and Postrhinal Cortices,” as are the projections from CA1 to the parahippocampal region. With respect to subcortical inputs, CA1 receives a substantially lighter septal projection than CA3 but the fibers are also most densely distributed in the stratum oriens (Nyakas et al., 1987).

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FIGURE 9 Schematic representation of the organization of CA1 projections to the subiculum. In the coronal section shown on the left side of the figure, cells of origin and respective terminal fields are indicated by similar shading patterns. The right side of the illustration shows a twodimensional unfolded map of CA1 and the subiculum. The septal pole of these fields is at the top of the figure and the temporal pole is at the bottom. The left-hand limit of the figure is the border of CA1 with CA2. It should be noted that discrete injections of PHA-L in CA1 lead to labeling of a terminal field that extends for approximately one-third of the septotemporal distance of the subiculum. Both parts of the figure indicate that an injection located proximally in CA1 (near the CA2 border) labels a column of the subiculum located distally, whereas cells located distally in CA1 project just across the border into the proximal subiculum.

The temporal two-thirds of CA1, more in particular the distal half, adjacent to the subiculum, receive a fairly substantial input originating in the amygdaloid complex (Krettek and Price, 1977). Using modern sensitive anterograde tracing, Pitkänen and her colleagues (Pitkänen et al., 2000) unraveled amygdaloid projections in great detail (Fig. 10). They showed that the caudomedial portion of the parvicellular division of the basal nucleus projects heavily to the stratum oriens and the stratum radiatum of CA1. In addition, the accessory basal nucleus as well as the posterior cortical nucleus of the amygdala projects to the stratum lacunosum-moleculare of CA1 (Kemppainen et al., 2002; Pikkarainen et al., 1999). An additional though weak input originates in the endopiriform nucleus (Behan and Haberly, 1999). The thalamic inputs to the hippocampal formation have been fairly well established using a variety of different anatomical approaches, and their presence has been corroborated by electrophysiological findings. Fairly prominent projections from midline (or “nonspecific”) regions of the thalamus to several fields of the hippocampal formation have been described (cf. Van der Werf et al., 2002, for a recent review; see also Groenewegen and Witter, Chapter 17, this volume). In particular, the small midline nucleus reuniens gives rise to a prominent projection to the stratum lacunosummoleculare of CA1 (Herkenham, 1978; Wouterlood

et al., 1990; Zheng, 1994). Interestingly, the nucleus reuniens projections to CA1, as well as to the subiculum (see sections on “Hippocampus” and “Subiculum”), travel via the internal capsule and cingulum bundle rather than through the fimbria/fornix. The nucleus reuniens projection terminates massively in the stratum lacunosum-moleculare and innervates all septotemporal levels, with a preference for the intermediate septotemporal levels (Fig. 11). The projection is topographically organized such that the dorsal portion of nucleus reuniens tends to project to more septal portions of CA1 whereas the ventral portion of the nucleus reuniens projects more temporally in CA1. Electron microscopic analysis indicates that the nucleus reuniens fibers terminate with asymmetric synapses on spines of principal neurons and thin dendritic shafts, most likely representing dendrites of inhibitory interneurons (Dolleman-van der Weel and Witter, 2000) in the stratum lacunosum-moleculare. This is in line with recent findings using retrograde tracing with 3 D-[ H]aspartate that showed that most of the reuniens neurons projecting to CA1, and also to the subiculum, use excitatory transmitters and colocalize up to 50% calretinin or calbindin (Bokor et al., 2002). It should also be noted that projections to the hippocampus arise from cells different from those projecting to other targets such as the medial septum or entorhinal cortex (Bokor et al., 2002; Dolleman-Van der Weel and Witter, 1996).

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FIGURE 10 Schematic summary of the reciprocal connections between the amygdala and the hippocampal region. Reproduced with permission from Pitkänen et al. (2000).

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FIGURE 11 This line drawing illustrates the distribution of fibers and terminals originating from cells in the nucleus reuniens of the midline thalamus. The locations of the fields shown in the higher magnification drawings (B–D) are indicated in Panel A. See text for details. Adapted with permission from Wouterlood et al. (1990).

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Like CA3, the CA1 field receives light noradrenergic, serotonergic, and dopaminergic projections (Fig. 5; cf. Swanson et al., 1987). Projections from the different dopaminergic cell groups distribute their axons along the septotemporal axis, but only about 15% of these neurons appear to be dopaminergic as shown with retrograde tracing combined with tyrosine hydroxylase (TH) immunocytochemistry (Gasbarri et al., 1994a, 1996). The projections from the raphe nucleus to CA1 arise, at least to some extent, from neurons that also innervate the medial septum (Acsady et al., 1996; McKenna and Vertes, 2001; Vertes et al., 1999). These inputs terminate, in part, on VIP-positive interneurons, known to innervate both principal neurons as well as other interneurons (Papp et al., 1999). Also inputs have been described from the nucleus subcoeruleus or pontogeniculooccipital wave-generating region in the brain stem (Datta et al., 1998). The CA1 field has substantially more extrinsic projections than the CA3 or CA2 fields. Field CA1 projects not only to the lateral septal area, which also receives inputs from field CA3, but also to a number of other subcortical and cortical areas. As such field CA1 is much more similar to the subiculum, with which it shares most of its projections (Naber and Witter, 1998; Jay and Witter, 1991; Van Groen and Wyss, 1990b). The organization of projections to the lateral septal area is similar to that of those originating in CA3 in that a septal-totemporal axis in CA1 is mapped onto a dorsal-to-ventral axis in the lateral septum. However, the projection from CA1 overall terminates at levels more rostral than that from CA3 (Naber and Witter, 1998; Risold and Swanson, 1997). This septotemporal organization of CA1 projections appears to be a general phenomenon (Naber and Witter, 1998; Van Groen and Wyss, 1990b). Septal levels of CA1 give rise to extrinsic connections to retrosplenial cortex, dorsal portions of the lateral septum and the nucleus of the diagonal band of Broca, as well as to rostrolateral parts of the nucleus accumbens. Midseptotemporal levels of CA1 do not project to the retrosplenial cortex but do distribute fibers to the ventral taenia tecta (or dorsal peduncular cortex) and the pre- and infralimbic cortex. Temporal levels project strongly to infralimbic cortex, ventral taenia tecta, anterior olfactory nucleus, anterior and medial parts of the hypothalamus and to more medial portions of the nucleus accumbens. The temporal two-thirds also originate most of the projections to the amygdaloid region, in particular to the basal nucleus. Weak projections to the olfactory bulb have been reported (Van Groen and Wyss, 1990b). CA1 generates prominent projections to the medial prefrontal cortex. The projections are quite similar to, but generally less dense than, those originating from the

subiculum (see page 661). In all innervated frontal regions, the CA1 projections innervate all layers diffusely (Conde et al., 1995; Jay et al., 1989; Jay and Witter, 1991; Ruit and Neafsey, 1990; Verwer et al., 1997; White et al., 1990). Interestingly, the direct inputs from the hippocampal CA1 field to prelimbic and infralimbic cortices in the rat, innervate not only “spiny” (presumed pyramidal) neurons but also NADPH diaphorase reactive cells and parvalbumin-containing local circuit neurons. The CA1 projections do not appear to innervate either calbindin or calretinin cells (Gabbot et al., 2002).

OVERVIEW OF THE SUBICULUM, PRESUBICULUM, AND PARASUBICULUM Should the subiculum, presubiculum, and parasubiculum be grouped together as the “subicular complex?” The term “subicular complex” is used by a number of authors to indicate a conglomerate of cytoarchitectonically different, relatively small cortical fields, that are located between CA1 and the entorhinal cortex, ventrally, and CA1 and the retrosplenial cortex, dorsally. In the literature, the following five cortical areas have been included as components of the subicular complex: the prosubiculum, the subiculum, the presubiculum, the postsubiculum, and the parasubiculum. Following earlier descriptions by the Vogts (Vogt and Vogt, 1919) and by Rose (Rose, 1926), Lorente de Nó (Lorente de Nó, 1934) described a transitional region between field CA1 and the subiculum proper and called it the prosubiculum. The prosubiculum was defined generally as an area where the stratum radiatum was no longer visible and where the pyramidal cell layer of CA1 merged and overlapped with subicular pyramidal cells. Although the term prosubiculum is still occasionally seen in the literature, many contemporary researchers agree that this region is better characterized as an oblique transitional region where field CA1 gradually replaces the subiculum. In this chapter, therefore, the term prosubiculum is not used (Fig. 2). The subiculum was originally described by Ramón y Cajal (Ramón y Cajal, 1911), but its characteristics were not clearly differentiated from the transitional area with CA1 nor was it clearly distinguished from the presubiculum. The subiculum suffered relative anonymity until the discovery that it is the major origin of the fornix (Swanson and Cowan, 1975). This has sparked renewed interest in this area which has been more extensively studied in the last several years (see below). The presubiculum is located distal to the subiculum (Fig. 2) and has been divided into ventral and dorsal

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portions. Some confusion surrounds the exact location and extent of the presubiculum since the dorsal portion is sometimes given the separate name “postsubiculum.” Brodmann (1909) originally described the postsubiculum in several species and called it area 48 (regio postor retrosubicularis) which was distinct from his area 27 or the presubiculum proper (ventral presubiculum). Based on cytoarchitectonic and connectional criteria, several authors have concluded that, in the rat, the postsubiculum is indeed an entity distinct from the presubiculum (Rose and Woolsey, 1948; Swanson and Cowan, 1977; Van Groen and Wyss, 1990c; Vogt and Miller, 1983). Other authors, however, view the great similarities in connectivity and histochemical appearance as being unifying characteristics and thus consider the postsubiculum to simply be a dorsal extension of the presubiculum (Blackstad, 1956; Haug, 1976; Witter, 2002; Witter et al., 1989). We have adopted the latter view and thus the term postsubiculum is not used in this chapter. The final component of the subicular complex is the parasubiculum. Although the cytoarchitectonic distinctness of the parasubiculum was appreciated by early investigators, until very recently there has been no connectional data that supported the separation from either the presubiculum or the entorhinal cortex. The parasubiculum has been subdivided into areas 49a and 49b (Blackstad, 1956; Haug, 1976) based largely on histochemical data; for the purposes of the present description, we do not subdivide the parasubiculum. One last area that should be mentioned is area 29e. This small wedge-shaped region was initially described in the rat by Blackstad (1956) and later characterized in more detail by Haug (1976). Although the neuroanatomy of area 29e remains sketchy, Stephan (Stephan, 1975) and others who have studied it consider it to be a part of the parasubiculum and we shall treat it as such in this chapter. From the cytoarchitectonic perspective, the subiculum, the presubiculum, and the parasubiculum fall into one of two categories. The first, to which the subiculum belongs, shares with the other hippocampal subfields the typical cytoarchitectonic characteristics of allocortex, i.e., three layers. The second category, to which the presubiculum and the parasubiculum belong, is more similar to the entorhinal cortex or adjacent parts of the retrosplenial cortex in that, by some accounts, these cortical regions are multilaminate. Typically the layers of these “periallocortical” regions are subdivided into external and internal principal lamina, separated by a cell free lamina dissecans. The external principal lamina comprises the molecular layer and cell layers II and III. The internal lamina, which appears to be the continuation of the cell layer of the subiculum, on the

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one hand, and the continuation of layers V and VI of the entorhinal cortex, on the other, are designated layers V and VI. As mentioned earlier, it is not clear whether the cell layers located deep to the external layer of the presubiculum and parasubiculum are more closely related to these structures or to the subiculum or entorhinal cortex. More evidence is clearly needed to resolve this issue and in the following discussions of the pre- and parasubiculum we generally summarize information that relates to the superficial layers. It is now generally accepted in the rat that the subiculum is the major subcortical output structure of the hippocampal formation. The entorhinal cortex, in contrast, appears to serve a dual role in that the superficial layers provide a major cortical input to all hippocampal subfields, whereas its deep layers receive a substantial output projection from CA1 and the subiculum. As described more fully below, the deep layers of the entorhinal cortex also generate a prominent projection back to the neocortex. The question arises whether the pre- and parasubiculum should be considered as portions of the output side of the hippocampal formation, i.e., more like the subiculum, or as part of the input side, similar to the entorhinal cortex. An interesting issue in this respect is whether the pre- and parasubiculum should be considered as links along the unidirectional pathway through the hippocampal formation. Although the pre- and parasubiculum receive inputs from the subiculum (see below), these are neither as robust as, for example, the CA1 projections to the subiculum or as unique since the subiculum also projects to the entorhinal, perirhinal, retrosplenial, and medial prefrontal cortices. Thus one might argue that they are not important continuations of the “trisynaptic” circuit. Like the entorhinal cortex, the presubiculum receives a fairly heavy direct neocortical input, particularly from the adjacent retrosplenial cortex (cf. Witter et al., 1989a; Wyss and Van Groen, 1992). Perhaps the most unique characteristic of the pre- and parasubiculum is their strong interconnectivity with the anterior thalamic nuclear complex (see below). The significance of the pre- and parasubiculum to hippocampal functioning, therefore, may lie in the fact that together they form the major route through which the anterior thalamus may influence the hippocampal formation. In recent years, it has been proposed that this pathway provides information specifically dealing with the orientation of the animal in space producing the so-called head-direction cells, present preferentially in the presubiculum and anterior thalamus (Taube, 1998). As we describe more fully below, the pre- and parasubiculum also send a major projection to the superficial layers of the entorhinal cortex, i.e., the layers which receive most of the other

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sensory inputs and which give rise to the major portion of the perforant pathway. For all these reasons, we consider the pre- and parasubiculum to be “input” structures rather than “output” structures and in this way they differ from the subiculum. Based both on the cytoarchitectonic and on the connectional characteristics reviewed above, we believe that it is unreasonable to lump the subiculum, presubiculum, and parasubiculum under the rubric “subicular complex.” We have therefore chosen not to use this term in this chapter and treat the subiculum separately as a hippocampal area, whereas the presubiculum and parasubiculum are included in the parahippocampal region (see also Lopes da Silva et al. (1990) and O’Mara et al. (2001) for a detailed discussion on this topic).

SUBICULUM Cytoarchitectonic Organization Although the cytoarchitectonic characteristics of the rat subiculum have not been studied in great detail, there are clear features that help to demarcate its borders with CA1 on the one hand and with the presubiculum on the other. The CA1/subicular border is marked mainly by a rather abrupt widening of the pyramidal cell layer (Fig. 2A) as well as by an abrupt loss of staining for calbindin, which is present in the superficial pyramidal cells of CA1, and an increased staining intensity with the Timm stain (Fig. 2B). The stratum radiatum is no longer present at this border and is continuous with and replaced by the wide molecular layer of the subiculum. The molecular layer of the subiculum can be subdivided into a deeper portion that is continuous with the stratum radiatum of CA1 and a superficial portion that is continuous with the molecular layer of the presubiculum and CA1 (Blackstad, 1956; Haug, 1976; Swanson et al., 1987). The stratum oriens of CA1 is not present in the subiculum. The border of the subiculum with the presubiculum is not easily discriminated but is generally positioned in the internal lamina where a quite striking decrease in overall size of the pyramidal cells can be seen, just deep to the proximal tip of the superficial lamina (external/ layers II and III) of the presubiculum (Figs. 2A and 2B).

Cell Types Much of the current information on subicular cell types comes from the studies of Ramón y Cajal and Lorente de Nó in young mice. Although the neuroanatomical, neurophysiological, and functional properties of the subiculum have been summarized recently

(O’Mara et al., 2001; see also Swanson et al., 1987), the main conclusion is that the subiculum is still an underinvestigated part of the hippocampal system. The principal cell layer of the subiculum is populated by large pyramidal neurons that end just beneath the distal end of CA1 and continue underneath the proximal portion of layers II/III of the presubiculum. These cells are relatively uniform in shape and size and extend their apical dendrites into the molecular layer; the basal dendrites extend into deeper portions of the pyramidal cell layer. Intermingled among the pyramidal cells are many smaller neurons, presumably representing the interneurons of the subiculum. Subpopulations of these neurons appear to have characteristics similar to those described for field CA1 (Witter, unpublished observations). Among those are GABAergic cells that stain for the calcium-binding protein parvalbumin. As in the dentate gyrus and hippocampus, these cells appear to be contacted by perforant path fibers (Bakste Bulte et al., 2003; Vinkenoog et al., 2003). With respect to the principal spiny pyramidal cells of the subiculum, a subdivision into at least two groups has been proposed on the basis of firing characteristics (O’Mara et al., 2001). Although different nomenclatures are used, a commonly accepted dichotomy is between regular spiking cells (nonbursters) and intrinsically bursting cells (bursters). Although these two cell types cannot be correlated with any distinguishing morphological characteristic, the two cell types show a differential distribution within the pyramidal cell. Bursting cells are more numerous deep in the layer whereas regular spiking cells are more common superficially (Greene and Totterdell, 1997). However, it has also been reported that bursting cells are preferentially found further away from CA1, i.e., in the distal subiculum, and that both cell types may actually reflect only a single class of neurons sharing a burst mechanism that is stronger in some cells (Staff et al., 2000). It thus appears that both distribution and subtyping are not all-or-none phenomena and that much overlap has been observed (Harris et al., 2001). The two populations can be related to preferential staining for either somatostatin (the bursters) or NADPH diaphorase/nitric oxide synthetase (NOS) (nonbursting regular spiking neurons) (Greene and Mason, 1996; Valtschanoff et al., 1993; Greene et al., 1997; Lin and Totterdell, 1998). Both of these cell types are projection neurons but they appear to differ with respect to their local connectivity (see following section). Neurons of the subiculum give rise to projections to the presubiculum, entorhinal cortex (Harris et al., 2001), and CA1 (Harris et al., 2001; Seress et al., 2002; Stewart, 1997). Interestingly, there is evidence indicating that only bursting cells project to the entorhinal cortex (Gigg et al., 2000).

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Connections of the Subiculum Associational Connections The subiculum gives rise to a longitudinal associational projection that extends from the level of the cells of origin to much of the subiculum lying ventrally to the level of origin. Interestingly, this projection seems to be largely unidirectional since few if any associational projections have been reported to course dorsally from their point of origin. The association fibers terminate diffusely in all layers of the subiculum (Köhler, 1985a). Despite the early demonstration by Blackstad (1956) that large hippocampal formation lesions result in anterograde degeneration in the contralateral subiculum, more recent studies have consistently found that the rat subiculum does not give rise to commissural connections (Köhler, 1985a; Swanson and Cowan, 1977; Witter et al., 1989). On the basis of intracellular labeling, it is apparent that subicular pyramidal cells provide a strong local input within the pyramidal cell layer and just superficial to it, targeting proximal portions of the apical dendrites. Interestingly, the density of this local intrinsic connection, as estimated by the number of varicosities on locally distributed axon collaterals is much higher than in CA1 (Harris et al., 2001). In addition, the two types of principal neurons differ with respect to their local connectivity. Intracellular labeling of electrophysiologically identified bursting cells generally show an axonal distribution that remains within the region circumscribed by their apical dendrites, whereas the regular spiking cells generally give rise to an axon that shows more widespread distribution along the transverse axis. Since these data have been generated in in vitro slices, it is not known whether similar differences exist with respect to a possible dorsoventral spread. Although much work needs to be done, available data indicate that the intrinsic organization of the subiculum might be different from that of the CA1 and CA3 fields. Local connections indicate the presence of both a crude columnar and a laminar organization (Fig. 12) such that bursting cells form a set of columns and the regular spiking neurons serve to integrate columnar activity. This characteristic organization of the subiculum, which appears strikingly different from the neighboring CA1, is of interest in view of the specific input–output organization of the subiculum (see following section; pages 673, 688). Hippocampal and Parahippocampal Connections As indicated earlier, the subiculum receives its major intrahippocampal input from the CA1 field of the hippocampus. It is also heavily innervated by fibers from the entorhinal cortex, with a less dense innervation

FIGURE 12 Schematic representation of the proposed columnar and laminar intrinsic organization of principal neurons of the subiculum. Deep cells have ascending axon collaterals that remain in close proximity to their apical dendrites, thus providing for a columnar organization. Superficial cells have axon collaterals that run long distances in the transverse, laminar direction. Reproduced with permission from Harris et al. (2001).

originating in the perirhinal and postrhinal cortices (see sections “Entorhinal Cortex” and “Perirhinal and Postrhinal Cortices”). Relatively weak inputs originate from the pre- and parasubiculum. There are no interconnections between the subiculum and the dentate gyrus or fields CA2/CA3 of the hippocampus. The subiculum gives rise to a minor intrahippocampal projection to field CA1. This projection mainly originates from superficially located pyramidal cells, including a subset of NOS-positive presumed regular spiking neurons (Commins et al., 2002; Harris and Stewart, 2001; Harris et al., 2001; Seress et al., 2002). The axon collaterals of these neurons specifically target dendrites of presumed pyramidal cells as well as interneurons present in the stratum oriens and stratum radiatum where they from symmetrical synapses onto dendritic shafts and asymmetrical synapses onto dendritic spines (Seress et al., 2002). Projections to the parahippocampal region largely reciprocate the inputs. The subiculum gives rise to a projection to the presubiculum, where fibers preferentially terminate in layer I. The projection to the dorsal part of the presubiculum terminates somewhat differently because the densest projection is to layer V, with weaker projections to layers I and II. The projections from the subiculum to the parasubiculum mainly terminate in layer I and the superficial part of layer II (Köhler, 1985a; Swanson et al., 1978; Van Groen and

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Wyss, 1990a, 1990c). These projections are topographically organized such that septal parts of the subiculum project to dorsal and caudal parts of both the pre- and parasubiculum, and temporal parts of the subiculum project to ventral and rostral parts (Swanson and Cowan, 1977; Van Groen and Wyss, 1990a, 1990c). Projections to the remaining portions of the parahippocampal region, i.e., to entorhinal, perirhinal, and postrhinal cortices, are described in sections “Entorhinal Cortex” and “Perirhinal and Postrhinal Cortices”. It is of interest to note that for all these projections, as well as the extrinsic ones (see page 661), a transverse topography has become apparent with cells in the proximal subiculum projecting differently from those in the distal subiculum (Naber and Witter, 1998; Naber et al., 2000b), which is discussed in more detail in page 668. Afferent Connections The subiculum has afferent and efferent connections with various nonhippocampal cortical and subcortical structures. While several of these connections are reciprocal, at least a few are not. In the first portion of this section, we deal with the afferent connections of the subiculum and in the next section we provide an overview of its efferent projections. In the rat, there is a paucity of detailed information regarding direct cortical inputs to the subiculum except those originating in parahippocampal regions. Many of the cortical regions that project fairly heavily to the entorhinal cortex (see section “Entorhinal Cortex”) have not been found to project to the subiculum. No inputs, for example, have been reported from the pre- and infralimbic cortices (Beckstead, 1979; Sesack et al., 1989; Jones and Witter, unpublished results; Witter, 2003). With respect to inputs from the retrosplenial cortex, some conflicting data have been reported. According to Wyss and Van Groen (1992) no portions of the retrosplenial cortex project to the subiculum. In contrast, according to a more recent study, the retrosplenial granular area provides a weak input to the molecular and pyramidal cell layers of the ipsilateral subiculum (Shibata, 1994; Jones and Witter, unpublished observations). With respect to the anterior cingulate cortex (area 24), reports of subicular projections have also been somewhat contradictory. A combined electrophysiological and neuroanatomical study reported a projection from the cingulate cortex to the subiculum (White et al., 1990). This projection has not been observed, however, in other neuroanatomical studies (Sesack et al., 1989; Vogt and Miller, 1983). In recent comprehensive studies also no evidence has been found for projections from area 24 to the subiculum (Jones and Witter, unpublished observations). In contrast to the rather meager complement of cortical inputs to the subiculum, those from subcortical

structures are quite numerous and robust. The origin and distribution of these inputs as well as their physiological role have been reviewed extensively and the reader is referred to these papers for a more detailed description (Lopes da Silva et al., 1990; Swanson et al., 1987). Because, in general, the subcortical projections to the subiculum arise from the same sources that innervate the other hippocampal fields, we confine ourselves to brief descriptions of these inputs. From the basal forebrain, a projection originates from the septal complex and fibers from the medial septal nucleus and the nucleus of the diagonal band terminate in the pyramidal cell and molecular layers (Chandler and Crutcher, 1983; Nyakas et al., 1987; Swanson and Cowan, 1977; see also in this volume Risold, Chapter 20, and Butcher and Woolf, Chapter 35). Inputs from the amygdaloid complex have been described in detail (Fig. 10) (see Pitkänen et al., 2000, for a detailed review). Substantial input arises from the caudomedial parvicellular division of the basal nucleus (Krettek and Price, 1977; Pikkarainen et al., 1999; Price et al., 1987) and from the posterior cortical nucleus and adjacent amygdalohippocampal area (Canteras et al., 1992a; Kemppainen et al., 2002). These amygdaloid inputs terminate mainly at the CA1/subiculum border region where they preferentially innervate the molecular layer of the subiculum and stratum lacunosummoleculare of CA1. Finally, the temporal part of the subiculum receives a weak input from the endopiriform nucleus (Behan and Haberly, 1999). The thalamic inputs to the subiculum are also comparable to those directed to CA1. Thalamic inputs originate mainly in the nucleus reuniens, the paraventricular nucleus and the parataenial nucleus (see Van der Werf et al., 2002, for a recent review). The septal and temporal extremes of the subiculum appear to be devoid of an input from the nucleus reuniens. The reuniens projections terminate mainly in the molecular layer of the subiculum (Herkenham, 1978; Wouterlood et al., 1990) where they are coextensive with the projections from the entorhinal cortex. It is also of interest that the projections to the subiculum and to CA1 originate from different but intermingled populations of neurons in the nucleus reuniens (Dolleman-Van der Weel and Witter, 1996). However, like the projections to CA1, those to the subiculum are most likely mainly excitatory (Bokor et al., 2002). Projections from the paraventricular and parataenial nuclei mainly target the molecular layer of the temporal subiculum. Also, there appears to be a minor input from the rhomboid nucleus to the more septal portions of the subiculum and adjacent part of field CA1 (Van der Werf et al., 2002). With respect to inputs from the anterior nuclear complex, it is clear that these projections are distributed

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mainly to the pre- and parasubiculum. However, the anteromedial and anteroventral nuclei weakly innervate the subiculum (Shibata, 1993; Thompson and Robertson, 1987; Van Groen and Wyss, 1990a, 1990c; Van Groen et al., 1999). Monoaminergic ascending pathways from the noradrenergic locus coeruleus, the dopaminergic ventral tegmental area, nigra A10 and retrorubral A8 group, and the serotonergic median and dorsal raphe nuclei reach the subiculum but do not show a preferential innervation of this region (Fig. 5). These inputs mainly target the more ventral portions of the subiculum. As is the case in general for most aminergic projections, it appears likely that only part of the projections from these different nuclei do contain monoamines as their transmitter (cf. Gasbarri et al., 1994a, 1994b, 1996). The supramammillary region (Haglund et al., 1984) projects heavily to the subiculum (Fig. 4B) particularly the temporal subiculum and this portion of the subiculum also receives an input, probably histaminergic, from the premammillary nucleus (Canteras et al., 1992b). Finally, inputs from the nucleus incertus reach the subiculum, preferentially terminating in the deep part of the molecular layer in the temporal two-thirds (Goto et al., 2001). Efferent Connections The subiculum is one of the major output regions of the hippocampal formation and projections are generated to a number of cortical and subcortical regions (Fig. 13; Naber and Witter, 1998; Ishizuka, 2001; O’Mara et al., 2001). The subiculum gives rise to a prominent projection to portions of the medial prefrontal cortex, particularly the medial and ventral orbitofrontal cortex, and to the prelimbic and infralimbic cortices. In the orbitofrontal and prelimbic cortices, the subicular fibers mainly innervate the deep layers, whereas in the infralimbic cortex the superficial layers also receive a projection. Some of the subicular fibers that reach the infralimbic cortex make contact with cells that project to the nucleus of the solitary tract (Ruit and Neafsey, 1990). Subicular projections also reach medial portions of the anterior olfactory nucleus and the agranular insular cortex (Conde et al., 1995; Jay and Witter, 1991; Jay et al., 1989; Ruit and Neafsey, 1990; Verwer et al., 1997; White et al., 1990; Witter et al., 1989a, 1990). The subiculum provides a meager projection to the anterior cingulate cortex (area 24) (White et al., 1990) whereas the projection to the retrosplenial cortex is very substantial (Witter et al., 1990; Wyss and Van Groen, 1992). The subiculoretrosplenial projection terminates predominantly in layers II and III with a modest innervation of layer IV. The projections to the retrosplenial cortex originate predominantly from the septal twothirds of the subiculum (Witter et al., 1989, 1990),

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although Wyss and Van Groen (1992) have suggested that the portion of the subicular projection to the most ventral subdivision “a” of the granular retrosplenial cortex originates in the ventral one-third of the subiculum. The perirhinal and postrhinal cortices receive a moderately strong input from the subiculum which terminates in both superficial and deep layers that are described in more detail in section “Perirhinal and Postrhinal Cortices” (Deacon et al., 1983; Swanson and Cowan, 1977; Swanson et al., 1978; Witter et al., 1989). Although it has been claimed that the subicular projection to the perirhinal cortex extends dorsally into the auditory temporal cortex (Swanson and Köhler, 1986), more recently findings do not support this (Kloosterman et al., 2003a; Vaudano et al., 1991). Also projections to the presubiculum have been described (see section “Presubiculum”). The most prominent subcortical subicular projections are those to the septal complex and adjacent nucleus accumbens as well as to the mammillary nuclei and adjacent hypothalamic regions. The projection to the septal area terminates predominantly in the lateral septal nuclei (to all subdivisions), although a few fibers also terminate within the medial septal nucleus (Gaykema et al., 1991; Ishizuka, 2001). Closely associated with the septal projection is the equally robust projection to the nucleus accumbens and adjacent parts of the olfactory tubercle. Subicular fibers distribute throughout the nucleus accumbens, with the projection to its caudomedial part being most dense (Groenewegen et al., 1987; Ishizuka, 2001; Kelley and Domesick, 1982; Witter et al., 1990). Whereas the subicular projections to the lateral septal nucleus are almost entirely confined to the ipsilateral side (which is in marked contrast to the septal projections from CA3), those to the nucleus accumbens show a weak contralateral component. Subicular projections to the mammillary nuclei are quite robust and are distributed bilaterally in nearly equal density. Additional projections target the ventromedial hypothalamic nucleus as well as the medial preoptic region, the anterior and tuberal hypothalamic regions, and the subparaventricular zone and its caudal continuum around the dorsal and medial aspects of the ventromedial nucleus, as well as the suprachiasmatic nucleus (Canteras and Swanson, 1992; Donovan and Wyss, 1983; Ishizuka, 2001; Kishi et al., 2000; Krout et al., 2002; Shibata, 1989; Swanson and Cowan, 1977; Witter et al., 1989a, 1990; Witter and Groenewegen, 1990). The projections to the mammillary nuclei are distributed throughout the medial nuclei; the lateral mammillary nucleus is only sparsely innervated by the subiculum. Subicular fibers also distribute to the lateral hypothalamic region, just adjacent to or in the lateral mammillary region, where fibers may interact with the cells of

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FIGURE 13 Line drawing illustrating the septotemporal (dorsoventral) and transverse origin of different subicular projections, supporting the proposed partition of the subiculum in septal(dorsal)–distal, septal(dorsal)–proximal, temporal(ventral)–distal, and temporal(ventral)–proximal areas. Reproduced with permission from Naber and Witter (1998).

origin of the histaminergic projection to the hippocampal formation (Wouterlood and Tuinhof, 1992). The thalamus also receives a subicular input (Canteras and Swanson, 1992; Meibach and Siegel, 1977b; Witter et al., 1990). Subicular fibers terminate bilaterally in the nucleus reuniens, the nucleus interanteromedialis, the paraventricular nucleus, and the nucleus, gelatinosus (submedius). With respect to subicular projections to parts of the anterior thalamic complex the story remains rather confusing. Initially described by Swanson and Cowan (1977) to arise from the subiculum, later studies clearly showed that these projections arise largely from the adjacent part of the presubiculum (Van Groen and

Wyss, 1990a, 1990c; Witter and Groenewegen, 1990; Witter et al., 1990). However, Ishizuka (2001) reported that a large number of neurons in the subiculum do indeed project to the anterior complex, with a possible preference for the anteromedial nucleus. Finally, the temporal one-third of the subiculum gives rise to projections that reach the amygdaloid complex (Pitkänen et al., 2000), primarily the lateral, basal, and accessory basal nuclei with more moderate or light projections reaching several other nuclei (Fig. 10; cf. Price et al., 1987). Interestingly, although quite a few of the amygdaloid nuclei also receive projections from CA1, all CA1 projections, except those to the basal nucleus, are

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much less prominent than those arising from the subiculum. The subiculum also projects heavily to the bed nucleus of the stria terminalis and moderately to the endopiriform nucleus (Canteras and Swanson, 1992; Witter et al., 1990). The subiculum and CA1 project to many of the same brain regions. However, most subicular neurons tend to distribute their axon to only a single target structure or to a very limited number of structures, whereas the axon of CA1 neurons collateralizes to a much larger number of brain structures (Naber and Witter, 1998; Ishizuka, 2001). The projections to the septum may be an exception to this overall rule, since quite a few studies indicate that up to 50% of subicular neurons that project to the lateral septum also project to at least one other brain area such as the the entorhinal cortex (Calderazzo et al., 1996; Donovan and Wyss, 1983; Swanson et al., 1981).

Topographical Organization of the Subicular Connections As with the projections from CA3 to CA1 and from CA1 to the subiculum, the subicular efferent projections are also topographically organized. In large part, the subicular projections honor the transverse topography established by the CA1 to subiculum projection; different projections originate from the proximal, middle, and distal thirds of the subiculum. The subiculum also demonstrates a marked septotemporal topography such that projections that arise from the septal or dorsal regions of the subiculum are different from those that arise from the temporal or ventral portions. Turning first to the septotemporal topography, it appears that the projections to the entorhinal cortex, the lateral septal complex, the nucleus accumbens, and the medial mammillary nucleus originate from the entire septotemporal extent of the subiculum (Ishizuka, 2001). Different septotemporal levels of the subiculum, however, project to different portions of these fields. In the entorhinal cortex, for example, the septal-to-temporal origin in the subiculum is related to a lateral-to-medial termination within the entorhinal cortex. Septal levels of the subiculum project preferentially to lateral and caudal parts of the entorhinal cortex, i.e., the parts that lie adjacent to the rhinal sulcus. Progressively more temporal levels of the subiculum project to more medially located parts of the entorhinal cortex. Although addressed in more detail below, it is important to point out that this topography is completely in register with the reciprocal projections from the entorhinal cortex to the subiculum (Kloosterman et al., 2003a). In the nucleus accumbens, the septotemporal axis of origin in the subiculum determines a caudomedial to

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rostrolateral axis of termination (Groenewegen et al., 1987). Dorsomedial portions of the lateral septal complex receive inputs from septal levels of the subiculum while ventral portions of the lateral septal complex are innervated by fibers originating in more temporal parts of the subiculum (Swanson and Cowan, 1977). A similar septotemporal topography has also been described for the subicular projections to the presubiculum and to the medial mammillary nuclei (Swanson and Cowan, 1977; Witter et al., 1990). The septal-to-temporal axis of origin corresponded to a dorsomedial-to-ventrolateral axis of termination in the medial mammillary nuclei (Kishi et al., 2000; Naber and Witter, 1998; Witter et al., 1990). Similarly, projections to the medial preoptic region, the ventromedial nucleus, and adjacent anterior and tuberal portions of the hypothalamus, which mainly originate from the temporal two-thirds of the subiculum, are topographically organized such that the septotemporal origin determines a dorsolateral-to-ventromedial terminal distribution in this region (Kishi et al., 2000; Witter and Groenewegen, 1990). This apparent dichotomy between the septal and temporal portions of the subiculum is reflected in the organization of other projections. Projections to the amygdala and the bed nucleus of the stria terminalis, for example, originate preferentially from the temporal one-third of the subiculum whereas the projections to the retrosplenial and perirhinal cortices originate predominantly from the septal two-thirds. The subicular projections to the midline thalamus also demonstrate a septotemporal topography. The most septal part of the subiculum projects preferentially to the interanteromedial nucleus, mid septotemporal levels of the subiculum preferentially project to the nucleus reuniens, and the temporal third of the subiculum projects most heavily to the paraventricular nucleus (Dolleman-van der Weel et al., 1993; Naber and Witter, 1998; Otake et al., 2002). Whereas the septotemporal topography appears to be organized in a gradient or gradual fashion, the transverse organization of subicular efferents is remarkably discrete. Along the transverse axis of the subiculum, three essentially nonoverlapping populations of cells have been differentiated that give rise to projections to a specific set of brain structures. This transverse organization of the outputs of the subiculum is consistently observed along its entire septotemporal axis although it is clearer dorsally than ventrally (Witter and Groenewegen, 1990; Witter et al., 1990; cf. Naber and Witter, 1998; see, however, Ishizuka, 2001). Neurons in the proximal third of the subiculum (closest to CA1) project to the infralimbic and prelimbic cortices, the perirhinal cortex, the nucleus accumbens, the lateral septum, the amygdaloid complex, and the core of the

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ventromedial nucleus of the hypothalamus. These projections originate not only from the subiculum but also from the adjacent distal one-third of the CA1 field. Cells in the central one-third of the subiculum project rather selectively to the midline thalamic nuclei. Cells in the most distal portion of the subiculum project mainly to the retrosplenial cortex and the presubiculum (these projections are summarized in Fig. 13). It should be mentioned here, however, that in a recent study using retrograde tracing of the origin of projections to the nucleus accumbens, the mammillary complex, and the anteroventral nucleus of the thalamus more emphasis has been put on laminar differences in the origin of these different projections (Ishizuka, 2001). Although this matter is still unclear, it appears that both organizational principles may hold true to some extent, which might be in line with the proposed columnar and laminar intrinsic organization of the subiculum (Harris et al., 2001) as described above. The subicular projections to the entorhinal cortex and to the medial mammillary nucleus do not follow a strict transverse organization since cells in all proximodistal portions of the subiculum project to these areas. However, the topography of these projections indicates a more subtle transverse organization and conflicting reports have been published. Proximal portions of the subiculum tend to project to more caudal portions of the medial mammillary nuclei and distal portions of the subiculum project more rostrally (Shibata, 1989). Other recent reports indicate a reversed gradient (cf. Naber and Witter, 1998). A more gradient-like topographical organization appears to exist for the subicular projections to the entorhinal, perirhinal, and postrhinal cortices (Kloosterman, 2003; see sections “Entorhinal Cortex” and “Perirhinal and Postrhinal Cortices”).

PRESUBICULUM Cytoarchitectonics and Cell Types The presubiculum, Brodmann’s area 27, is relatively easily differentiated from the subiculum (Fig. 2). The presubiculum has a distinct, densely packed external cell layer that is located just superficial to a relatively cell-free zone, which is continuous with lamina dissecans in the entorhinal cortex. This densely packed layer consists mainly of darkly stained small pyramidal cells. The most superficial cells are the most densely packed (layer II), while the deeper cells have a somewhat looser arrangement (layer III). The differentiation between layers II and III is clearer at dorsal levels of the presubiculum. Cells located deep to layers II and III of the pre-

subiculum are considered, by some, to be deep layers of the presubiculum. Cells in these deep layers, however, appear to be continuous with the deep layers of the entorhinal cortex and with the principal cell layer of the subiculum. Deep to the lamina dissecans, one finds one or two layers of large, darkly stained pyramidal cells. Deep to these neurons is a rather heterogeneous collection made up of pyramidal cells and polymorphic cells (Lorente de Nó, 1934). In both superficial and deep layers stellate as well as pyramidal cells have been recently reported using intracellular filling of electrophysiologically defined neurons. The stellate cells in the superficial layers tend to have spiny dendrites radiating into layer I. Neurons in the deep layers showed either sparsely spiny or nonspiny dendrites that in all cases did not reach superficial layers. Regarding the pyramidal cells, the dendritic characteristics are remarkably similar to those of the stellate cells, with the exception that the dendrites of layer V pyramidal cells do reach into layers II and III (Funahashi and Stewart, 1997a, 1997b; see also Wouterlood, 2002). For further details on the cytoarchitectonic organization of the presubiculum as well as the description of several histochemical characteristics of the presubiculum that help to differentiate it from the parasubiculum the reader is referred to the study by Haug (1976) (see also Insausti et al., 1997; Van Groen and Wyss, 1990a).

Connections of the Presubiculum Associational Connections Associational connections are rather well developed in the presubiculum and interconnect all dorsoventral levels. Ventral portions of the presubiculum project to more dorsal levels. These dorsally directed projections originate mainly from layer II cells and, to a lesser extent, from deeper layers as well (Van Groen and Wyss, 1990a). Dorsal portions of the presubiculum also project to ventral levels but these projections arise preferentially from cells in the deep layers (Van Groen and Wyss, 1990c). Regarding the terminal distribution of these dorsal-to-ventral projections there are some discrepancies in the literature. According to Van Groen and Wyss, the projections terminate in layers II, III, and V, similar to the associational connections that run in the opposite direction. Köhler (1985a) indicated, however, that they distribute mainly to the deep portion of layer I and to layer II. Additional data on intrinsic connectivity generated in in vitro slices using electrophysiology and intracellular filling indicate that whereas neurons in superficial layers exert strong monosynaptic effects onto cells in the deep layers, the return projection from deep to superficial appears virtually absent (Funahashi and Stewart, 1997a).

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According to these authors, a similar situation is found in the parasubiculum. Commissural Connections Strong commissural projections originate in the presubiculum and distribute to layers I and III of the homotopic part of the contralateral presubiculum (Köhler, 1985a; Van Groen and Wyss, 1990a). Commissural projections from the dorsal portion of the presubiculum are relatively sparse (Van Groen and Wyss, 1990c). Hippocampal and Parahippocampal Connections Cells throughout the presubiculum project to layers I (deep) and II of the parasubiculum (Köhler, 1985a; Van Groen and Wyss, 1990a) bilaterally. The presubiculum is reciprocally connected with the subiculum (Funahashi and Stewart, 1997b; Harris et al., 2001; Van Groen and Wyss, 1990a, 1990c). The presubiculum-to-subiculum projection that is moderate in density appears to be bilateral (Köhler, 1985a), mainly originating from the dorsal two-thirds of the presubiculum, whereas only a weak ipsilateral projection reaches the temporal portions of the subiculum (Van Groen and Wyss, 1990a, 1990c). There is also a weak projection to all fields of the hippocampus and to the molecular layer of the dentate gyrus (Köhler, 1985a; Witter et al., 1988). The presubicular fibers to the dentate molecular layer tend to be radially arranged which is quite distinct from the predominantly transverse orientation of the entorhinal perforant pathway fibers. The most prominent projection from the presubiculum is the bilateral projection to the entorhinal cortex. This projection has a number of interesting features. First, the presubiculoentorhinal projection is directed only to the medial division entorhinal cortex. Second, the projection terminates almost exclusively in layer III and to a much lesser extent in the deep part of layer I. Third, the crossed projection from the presubiculum to the contralateral entorhinal cortex is every bit as dense as the ipsilateral projection. The projection to the entorhinal cortex is topographically organized and although some discrepancies are present in the literature (Caballero-Bleda and Witter, 1993; Köhler, 1985a; Swanson and Cowan, 1977; Van Groen and Wyss, 1990a) it is clear that a dorsal-toventral and a transverse topography are present. The proximodistal position of the cells of origin along the transverse axis determines the mediolateral location of the terminal plexus in the entorhinal cortex and the dorsoventral position of the cells of origin determines the dorsoventral level of entorhinal cortex that is innervated (Honda and Ishizuka, 2001). In the medial entorhinal cortex, presubicular axons contact both spiny dendrites of neurons, most likely projecting to CA1 and the subiculum (Caballero-Bleda and Witter,

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1994; Van Haeften et al., 1997), as well as dendrites of inhibitory interneurons and of layer V pyramidal cells (Van Haeften et al., 1997, 2000). Among the interneurons receiving presubicular inputs are most likely parvalbumin-positive GABAergic neurons that presumably form inhibitory axonal baskets around the somata of principal neurons, as well as calretinin-positive interneurons that are potentially involved in feedforward excitatory microcircuits (Wouterlood, 2002). Interestingly, it has been shown that 20–30% of the ipsilaterally projecting presubicular neurons are GABAergic. The projections to the contralateral MEA, however, are not GABAergic. Moreover, GABAergic projection neurons were observed only in the dorsal part of the presubiculum, which, when taking into account the topography of presubicular projections to the MEA, indicates that only the dorsal part of the MEA receives a GABAergic input. The GABAergic projection neurons constitute approximately 30–40% of all GABAergic neurons present in the superficial layers of the dorsal presubiculum. The ipsilateral and contralateral presubiculoentorhinal projections originate from different populations of neurons. The presubiculum receives inputs from the entorhinal cortex (Beckstead, 1978; Köhler, 1986, 1988). The subiculum also gives rise to a projection to the presubiculum with fibers terminating mainly in layer I. The projection to the dorsal part of the presubiculum terminates somewhat differently since the densest terminal distribution is to layer V, with weaker termination in layers I and II (Köhler, 1985a; Swanson et al., 1978; Van Groen and Wyss, 1990a, 1990c). These projections are topographically organized such that septal parts of the subiculum project to dorsal and caudal parts of both the pre- and parasubiculum and temporal parts of the subiculum project to ventral and rostral parts (Swanson and Cowan, 1977; Van Groen and Wyss, 1990a, 1990c). Extrinsic Inputs and Outputs The presubiculum receives relatively few extrahippocampal cortical inputs. The most prominent one originates in the retrosplenial cortex. Cells located in layer V of the retrosplenial cortex give rise to projections that terminate in layers I and III–V of the presubiculum (Swanson and Cowan, 1977; Van Groen and Wyss, 1990a, 1990c). Subtle differences have been reported between subdivisions of the retrosplenial cortex with respect to their preferential laminar termination. Whereas projections from the retrosplenial granular area terminate predominantly in layers I, III, V, and VI of the presubiculum, projections from the retrosplenial agranular area terminate predominantly in layers I and III of the presubiculum (Shibata, 1994). A

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second cortical input originates from layer V of the visual area 18b (Vogt and Miller, 1983). This projection mainly distributes to the dorsal half of the presubiculum and terminates in layers I and III. Minor cortical inputs appear to originate in the prelimbic cortex (Beckstead, 1979) and in a dorsal part of the medial prefrontal cortex (Beckstead, 1979; Deacon et al., 1983; Van Groen and Wyss, 1990c). Whether this latter cortical region should be considered to be area 8 (Swanson et al., 1987) or part of area 24 of the cingulate cortex (Van Groen and Wyss, 1990c) is not clear. The presubiculum receives a number of subcortical inputs. The subcortical relationship that most uniquely defines the pre- and parasubiculum is their interconnections with dorsal parts of the thalamus, primarily the anteroventral and anterodorsal nuclei, as well as the laterodorsal nucleus (Shibata, 1993; Van Groen and Wyss, 1995; see also Groenewegen and Witter, Chapter 17, this volume). Although earlier studies have provided conflicting information, recent tracing studies indicate that in particular layer V of the presubiculum also receives inputs from the anteromedial nucleus, although the strongest projections are distributed elsewhere (Thompson and Robertson, 1987; Van Groen et al., 1999). It seems most likely that these differences in observations are related to subtle differences in connectional topography within the nucleus (Van Groen et al., 1999). Such topographical features are common in the thalamic connections of the ventral and dorsal parts of the presubiculum. The ventral part receives most of its input from the laterodorsal and anteroventral nuclei whereas the dorsal part receives its inputs from the laterodorsal and anterodorsal nuclei (Van Groen and Wyss, 1990a, 1990c). The thalamic projections mainly terminate in layers I, III, and IV but subtle differences in laminar termination have been described, apparently related to the origin in different quadrants of the anteroventral nucleus (Shibata, 1993; Van Groen and Wyss, 1995). Minor thalamic inputs to the presubiculum, predominantly to layer I, originate from the nucleus reuniens (Herkenham, 1978; Wouterlood et al., 1990). The presubiculum receives a particularly prominent cholinergic input. The medial septal nucleus and the vertical limb of the diagonal band of Broca give rise to projections that terminate predominantly in layer II of the presubiculum. The presubiculum also receives projections from the endopiriform nucleus which terminate preferentially in the external principal lamina (Behan and Haberly, 1999; Eid et al., 1996; Van Groen and Wyss, 1990a, 1990c). Additional projections originate from the hypothalamus, in particular from the area surrounding the mammillary nuclei. Fibers from the supramammillary nucleus terminate preferentially

in the deeper cell layers of the presubiculum (Haglund et al., 1984; Saper, 1985), although those that are characterized as being α-MSH positive terminate in the molecular layer of the presubiculum (Köhler et al., 1984b). Although older tracing studies have reported a minor input from the basal nucleus of the amygdala, recent, more detailed tracing experiments using sensitive anterograde tracing techniques have not provided evidence for any marked input from the amygdaloid complex to the presubiculum (Fig. 10; Kemppainen et al., 2002; Pitkänen et al., 2000). The presubiculum also receives inputs from various nuclei in the brain stem. A particularly dense innervation arises from the dorsal and median raphe nuclei; at least a component of this projection is serotonergic and innervates layer I (Köhler et al., 1981; Köhler and Steinbusch, 1982; Van Groen and Wyss, 1990a, 1990c). There is a fairly substantial plexus of noradrenergic fibers in the plexiform layer that arise primarily from the locus coeruleus (Swanson et al., 1987). Projections from layer V of the presubiculum reach the granular retrosplenial cortex, where they terminate preferentially in layers I and II (Van Groen and Wyss, 1990a). These projections are topographically organized such that the ventral presubiculum projects mainly to the ventral part of the granular retrosplenial cortex (subdivision a) and the dorsal part of the presubiculum projects preferentially to the more dorsally located subdivision b. These projections also exhibit a rostrocaudal organization such that rostral parts of the presubiculum project to rostral parts of the retrosplenial cortex and caudal portions of the presubiculum project to caudal parts of the retrosplenial cortex. It has been suggested that the dorsal presubiculum projects to the deep layers of the most caudal portion of the perirhinal cortex (Deacon et al., 1983; Van Groen and Wyss, 1990c). In recent studies, this area, which is now generally referred to as postrhinal cortex, has indeed been shown to receive inputs from dorsal portions of the presubiculum (Burwell and Amaral, 1998b). The presubiculum gives rise to a massive projection to the anterior nuclear complex of the thalamus, in particular to the anteroventral, anterodorsal, and laterodorsal nuclei. The projections are bilateral and the ipsilateral component closely reciprocates the anterior thalamic inputs to the presubiculum. These presubicular projections mainly arise from cells located deep to lamina dissecans (layer VI) (Seki and Zyo, 1984; Swanson et al., 1987; Van Groen and Wyss, 1990a, 1990c). The deep layers of the presubiculum also project bilaterally to the medial and lateral nuclei of the mammillary complex (Donovan and Wyss, 1983; Meibach and Siegel, 1975; Shibata, 1989; Swanson and Cowan, 1977). The projections to the medial mammillary

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nuclei are topographically organized similar to those that originate in the subiculum. Projections from the presubiculum to the amygdaloid complex appear to be nonexistent in the rat (Pitkänen et al., 2000).

PARASUBICULUM Cytoarchitectonic Organization and Cell Types Layers II and III of the parasubiculum (Brodmann’s area 49) consist of large, rather densely packed, lightly stained pyramidal cells. This, and other characteristics such as the distinctive staining for heavy metals observable with the Timm’s sulfide silver method (Haug, 1976) are the major features that differentiate the parasubiculum from its neighboring cortical areas (Fig. 2). There is no clear differentiation between layers II and III of the external lamina, and, as with the presubiculum, the deep layers are continuous with those of the entorhinal cortex.

Connections of the Parasubiculum Associational and Commissural Connections The Parasubiculuma gives rise to associational connections that distribute both dorsally and ventrally from the cells of origin. The dorsally directed projections extend only for short distances and are generally quite weak. The ventrally directed projections are substantially denser and extend for long distances along the long axis of the parasubiculum (Caballero-Bleda and Witter, 1993; Köhler, 1985a). The parasubiculum gives rise to a particularly dense projection to the dorsally located area 29e (Köhler, 1985a). The parasubiculum also gives rise to a commissural projection that terminates most densely in layer I (Köhler, 1985a), layer III (Van Groen and Wyss, 1990a), or both. Additional data on intrinsic connectivity generated in in vitro slices have been taken to indicate a preferential superficial-to-deep local connectivity whereas projections from deep to superficial are virtually absent (Funahashi and Stewart, 1997a). Hippocampal and Parahippocampal Connections The parasubiculum receives hippocampal input from the subiculum and an additional, though rather weak, parahippocampal input from the entorhinal cortex. The subicular projection mainly terminates in layer I and the superficial part of layer II (Köhler, 1985a; Swanson et al., 1978; Van Groen and Wyss, 1990a, 1990c). These projections are topographically organized such that septal or dorsal parts of the subiculum project to dorsal and caudal parts of the parasubiculum and temporal or ventral parts of the subiculum project to ventral and

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rostral parts (Swanson and Cowan, 1977; Van Groen and Wyss, 1990a, 1990c). With respect to the entorhinal fibers innervating the parasubiculum it has been reported that these arise predominantly from cells in layer Va and terminate predominantly in layer I (Köhler, 1986, 1988; Van Groen and Wyss, 1990a, 1990c). With respect to parasubicular outputs, it is not generally appreciated that the parasubiculum gives rise to a fairly substantial projection to the molecular layer of the dentate gyrus. Like the lighter projection from the presubiculum, this projection occupies the superficial two-thirds of the molecular layer (with a preference for the mid portion of the molecular layer) and fibers have a predominantly radial orientation (Köhler, 1985a; Witter et al., 1988). Since the parasubiculum receives a prominent projection from the anterior thalamic nuclei, its projection to the molecular layer provides a route by which thalamic input might influence very early stages of hippocampal information processing. The parasubiculum also gives rise to a weak to moderate projection to stratum lacunosum-moleculare of the hippocampus and to the molecular layer of the subiculum (Köhler, 1985a; Van Groen and Wyss, 1990a). The parasubiculum innervates all septotemporal levels of the hippocampus with a clear-cut septotemporal topographic organization. The parasubiculum also projects to layers I and III of the presubiculum (Köhler, 1985a); both ipsilateral and contralateral projections have been reported by Van Groen and Wyss (1990a). The most impressive projection of the parasubiculum selectively innervates layer II of the entorhinal cortex (Caballero-Bleda and Witter, 1993; Köhler, 1985a; Van Groen and Wyss, 1990a), most likely targeting the dendrites of principal neurons (Caballero-Bleda and Witter, 1994). In contrast to the presubiculum, the parasubiculum projects both to the medial and lateral entorhinal areas, though the projection to the lateral entorhinal area is slightly less robust (Caballero-Bleda and Witter, 1993; Van Groen and Wyss, 1990a; see, however, Köhler, 1985a). Projections from the parasubiculum to the contralateral entorhinal area have been reported but are weak (Caballero-Bleda and Witter, 1993; Köhler, 1985a; Van Groen and Wyss, 1990a). The topographical organization of the parasubicular projection to the entorhinal cortex is comparable to that of the presubiculoentorhinal projection (Caballero-Bleda and Witter, 1993). Extrinsic Inputs and Outputs With the exception of the relatively light projections from the retrosplenial cortex and the occipital visual cortex (Van Groen and Wyss, 1990a; Vogt and Miller, 1983), there are no other known extrahippocampal cortical inputs to the parasubiculum. Cortical inputs from the retrosplenial granular area terminate predominantly

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ipsilaterally in layers I and IV–VI, whereas those originating in the agranular area terminate predominantly in layers IV–VI of the ipsilateral parasubiculum (Shibata, 1994). With respect to the subcortical inputs of the parasubiculum, one might simply state that they are quite similar to those of the presubiculum. One significant difference is that the parasubiculum receives a much more prominent input from the amygdala, particularly from the posterior cortical nucleus and the caudal portion of the medial division of the lateral nucleus, all divisions of the basal nucleus, but most dense from its caudomedial parvicellular division, and the accessory basal nucleus, terminating in layers I and superficial II and III (Fig. 10). With respect to the latter projection it has been reported that the parvicellular division of the accessory basal nucleus mainly projects to ventral portions of the parasubiculum, the dorsal portions receive stronger inputs from the magnocellular subdivision (Kemppainen et al., 2002; Krettek and Price, 1977; Petrovich et al., 1996; Pikkarainen et al., 1999; Van Groen and Wyss, 1990a; see also Pitkänen et al., 2000). With the exception of a modest projection to the anterodorsal nucleus of the thalamus, no other extrahippocampal projections of the parasubiculum have been reported in the rat (Van Groen and Wyss, 1990a).

ENTORHINAL CORTEX The rat entorhinal cortex makes up the ventroposterior convexity of the rat cerebral hemisphere. It extends ventromedially to border either the parasubiculum medially or the piriform cortex and the amygdaloid complex rostrally; it also extends dorsolaterally to approach the rhinal fissure (Fig. 1). At rostral levels, the entorhinal cortex ends just ventral to the rhinal fissure and, at these levels, the perirhinal cortex forms the fundus and dorsal bank of the rhinal fissure. At caudal levels, the entorhinal cortex extends within and slightly above the rhinal fissure.

Lamination It is important to point out that there are two schemes of cortical lamination currently applied to the entorhinal cortex. As one might expect, this causes substantial confusion especially to the neophyte hippocampologist. One nomenclature, which divided the entorhinal cortex into seven layers, was first suggested by Ramón y Cajal (Ramón y Cajal, 1911) and later modified to more closely resemble the standard six-layer scheme applied to the isocortex. According to this scheme, there are four cellular layers (layers II, III, V, and VI) and two acellular or plexiform layers (layers I and IV).

The acellular layer IV is also called lamina dissecans. The Ramón y Cajal scheme, with slight modification, has been employed in several primate studies (e.g., Amaral et al., 1987). The other commonly used scheme was proposed by Lorente de Nó (Lorente de Nó, 1933) who also differentiated six layers. Five of Lorente de Nó’s layers were cellular (layers II, III, IV, V, and VI) with a cell-free lamina dissecans or IIIb in between layers III and IV; as in Ramón y Cajal’s nomenclature, layer I was the most superficial, acellular layer. The latter is the scheme that has been used in most of the older studies of the entorhinal cortex but has been replaced by the nomenclature as proposed by Ramón y Cajal in most of the contemporary studies. In this description, we have labeled lamina dissecans as the cell-poor layer IV to emphasize the lack of an internal granular cell layer in the entorhinal cortex. The resulting nomenclature that we have employed thus resembles the one initially proposed by Ramón y Cajal with some modifications (Fig. 14) (see also Insausti et al., 1997, for a detailed cytoarchitectonic description): • Layer I is the most superficial plexiform or molecular layer and is relatively devoid of neurons and rich in transversely oriented fibers (Blackstad, 1956; Haug, 1976). • Layer II mainly contains medium- to large-sized stellate cells. These cells tend to be grouped in clusters (cell islands), although caudally the cell islands coalesce and the layer is thus more continuous. • Layer III contains cells of various sizes and shapes, thus rendering a heterogeneous aspect to the layer. Pyramidal cells are predominant in layer III. The appearance of the layer varies from lateral to medial and from rostral to caudal. • Layer IV (or lamina dissecans) is located between layers III and V. It is most obvious at caudal levels of the entorhinal cortex. In the remainder of the entorhinal cortex, patches of cells invade the layer so that it has an incomplete or dashed appearance. • Layer Va forms a band of large, darkly stained pyramidal neurons. This layer is most conspicuous in the central parts of the entorhinal cortex. At other levels, the packing density of cells is not high and the smaller cells of the deeper part of this layer (Vb) intermingle with it. Rostrally, layer Vb exhibits a rather loose arrangement of smaller pyramidal cells and polymorphic cell types; caudally, this sublayer takes on a more columnar appearance. • Layer VI contains a very heterogeneous population of cell sizes and shapes. Cell density decreases toward the limit with the white matter. The cells of layer VI appear to blend, in a gradual way, both into the subjacent subcortical white matter and into the

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FIGURE 14 This line drawing illustrates several of the major cell types of the entorhinal cortex and some of their major inputs and outputs. The superficial layers of the cortex (I–III) are at the top of the illustration and the deep layers (V–VI) are at the bottom. An exclamation mark (!) indicates that synaptic contacts between the involved elements have indeed been reported. See text for details about the various cells pictured. Adapted with permission from Wouterlood (2002).

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overlying layer V. As in layer V, layer VI has a more columnar organization at caudal levels.

Cell Types Our current knowledge of the cytology of the entorhinal cortex is based largely on the classical Golgi studies of Ramón y Cajal and Lorente de Nó. More recent studies have focused on determining which particular neuronal cell types project to the dentate gyrus, hippocampus, and subiculum (Caballero-Bleda and Witter, 1994; Germroth et al., 1989a, 1989b; Schwartz and Coleman, 1981), what types of interneurons can be differentiated, or whether principal neurons have local intrinsic collaterals. Our current understanding of the individual neuronal elements of the entorhinal neuronal network, their interconnections, and major functional characteristics has recently been summarized and the reader is referred to those summaries for a more detailed account (Wouterlood, 2002; Alonso, 2002). The following description provides a short overview of the various cell types of each of the entorhinal layers and a brief comment on the major characteristics of their dendritic and axonal organization (Fig. 14). Layer 1 is populated by a low number of widely dispersed neurons that have been further differentiated using various criteria. Using morphological criteria, this layer contains stellate and horizontal cells. In general, the stellate cells possess smooth dendrites that radiate into layer 1 and are more numerous in the superficial portion of layer I. Although initially thought to be all GABAergic, more recent findings have indicated that quite a few of these neurons do stain for calretinin but not for GABA (Miettinen et al., 1997; Wouterlood et al., 2000). This implies that layer I neurons comprise both inhibitory and excitatory interneurons. The horizontal cells, which are less abundant than the stellate cells, are found preferentially in more deeper portions of the molecular layer at the transition with layer II. It is preferentially this class of GABAergic layer I neurons that has been shown to project to the dentate gyrus and hippocampus. Layer 1 neurons also target the dendrites of layer II cells that project to the dentate gyrus (Caballero-Bleda and Witter, 1994; Germroth et al., 1989a, 1989b; Witter et al., 1989a). Layer II is populated mainly by stellate neurons (or modified pyramidal cells). The stellate cells are considered to be the principal source of fibers for the perforant pathway projection to the dentate gyrus and CA3. However, layer II also contains pyramidal, multipolar, and horizontal cells, all of which apparently contribute to the perforant pathway projection. In general, axons of layer II cells have relatively few collaterals distrib-

uting to deeper layers of the entorhinal cortex. Some collaterals innervate layer III, but layers V and VI receive little or no innervation (Köhler, 1985a, 1986; Lingenhohl and Finch, 1991). Within layer II, two disparate patterns of collateralization have recently been described. Some stellate cells contribute their main axon to the perforant pathway with little collateralization in the entorhinal cortex, while others (which also direct their main axon to the perforant path) give rise to an extensive collateral plexus within layers I and II (Klink and Alonso, 1997; Lingenhohl and Finch, 1991; Tamamaki and Nojyo, 1993). Whether this differential collateral pattern is related to the currently used distinction between spiny and sparsely spiny stellate cells is not clear (Wouterlood, 2002). The collaterals of the layer II stellate cells are mainly oriented parallel to the pial surface and on the basis of electrophysiological observations it is assumed that these local collaterals mainly contact inhibitory interneurons, thus providing a strong feedback inhibition (Alonso, 2002). Although there are some exceptions, the majority of layer II cells have a dendritic tree that is confined to layers I and II, radiating essentially in all directions; however there is evidence indicating that superficially radiating dendrites are longer and more divergent (Klink and Alonso, 1997; Van der Linden and Lopes da Silva, 1998). It has recently been shown that a class of axoaxonic cells, similar to the cortical chandelier cell, is located in layer II. While some of these cells are also present in layer III, they have not been observed in the deeper layers (Soriano et al., 1993). Such cells, which apparently contain GABA and stain for parvalbumin, have their candle-like terminal arrangements predominantly in the upper portions of layers II and III, where they contact neurons projecting to the hippocampal formation (Martinez et al., 1996). Two types of chandelier cells have been described (Soriano et al., 1993), a vertically oriented cell only present in the medial entorhinal cortex and a horizontally oriented one present throughout all subfields of the entorhinal cortex. A second type of interneuron, also quite characteristic for layers II and III is the GABAergic, parvalbumin-positive neuron that distributes a dense axonal plexus within its parent cell layer. These axons ramify profusely forming basketlike terminal plexi onto the somata of principal neurons (Wouterlood et al., 1995; Wouterlood, 2002). In layer III, the most numerous cell type is the pyramidal cell which gives rise to the projections to field CA1 and to the subiculum. Their axons give rise to collaterals that distribute mainly to layers III and I. Only a few collaterals are present in layer II and layer Vb. Apparently no collaterals are given off to layer Va (Köhler, 1985a, 1986; Lingenhohl and Finch, 1991). The apical dendrites of the layer III pyramidal cells ascend

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radially, give off branches to layer III, and ultimately form a terminal tuft in layer I (Van der Linden and Lopes da Silva, 1998). In addition, multipolar, stellate, fusiform, horizontal and bipolar cells in layer III, all of which appear to contribute to the perforant pathway, have been described. This large, heterogeneous group of cell types undoubtedly coincides with what Lorente de Nó described as “atypical” layer III neurons. Layer IV, the lamina dissecans, although generally referred to as a cell-free layer, does contain scattered cell bodies described as fusiform or pyramidal cells (Lingenhohl and Finch, 1991) having apical dendrites reaching up to layer I and an axon reaching the deep white matter, a characteristic of projection neurons. Some of the cells in the lamina dissecans stain positively with antibodies against GABA, neuropeptide Y, calretinin, calbindin, or somatostatin (Köhler et al., 1986; Wouterlood and Pothuizen, 2000; Wouterlood et al., 2000). According to the Golgi-based description of Lorente de Nó, layer V contains three classes of neurons, pyramidal cells, small spherical cells, and fusiform neurons. This description to a large extent has been corroborated by a more recent in vitro intracellular filling study in which it was reported that layer V contains pyramidal , horizontal, and polymorph neurons (Gloveli et al., 2001; Hamam et al., 2000). In the latter study, most of the filled neurons apparently have their cell bodies in the deeper portions of layer V, i.e., layer Vb. The more superficial portion, layer Va, has not been studied in much detail, but Nissl-stained as well as Golgi-stained sections indicate that the predominant cell type is a large, darkly staining pyramidal cell, which has large apical spiny dendrites that, without giving off branches, ascend vertically toward the most superficial parts of layer II and into layer I where they have their terminal tuft (Van Haeften et al., 2003). These cells resemble the pyramidal cells described in layer Vb (Hamam et al., 2000). The axons of all the layer V pyramidal cells run predominantly toward the deep white matter, but collaterals are generated both within the entorhinal cortex, as well as in the angular bundle. According to Lorente de Nó, the collaterals of such cells may form a column-like plexus close to the location of the parent cell body, which spans the entire thickness of the entorhinal cortex. Although in one intracellular labeling study (Lingenhohl and Finch, 1991) it was reported that these collaterals mainly distribute to the deep cell layers (V/VI) with only an occasional collateral reaching layer II, more recent data support the original notion of Lorente de Nó indicating that at least some cells in layer V provide strong inputs to layers II and III (Gloveli et al., 2001; Van Haeften et al., 2003). With respect to the other small

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neurons in layer V, conflicting data have been reported. Golgi studies (Lorente de Nó, 1933) indicate that the axonal plexus of the latter group either remain in layer V or extend to the superficial layers I–III. According to the study of Hamam et al. (2000), however, most if not all of these cells do send an axon to the underlying white matter and may thus be considered projection neurons. Gloveli et al. (2001) found that layer V cells also provide axon collaterals to the superficial layers. The overall picture emerging from these recent studies is thus that all three types of layer V neurons should be considered projection neurons in that they send an axon to the white matter. They also may function as local circuit neurons, connecting deep layers to superficial layers. Finally these results indicate that the morphologies of these cells do not relate in a simple way to a subdivision into projection neurons versus interneurons. There is a wide variety of neuronal cell types in layer VI. Based on the predominant distribution of their axonal plexus, V and VI cells in both layers can be grouped into three categories: (1) cells that mainly influence other cells in layer Vb or VI, (2) cells that by means of their highly collateralized axons can influence a vertical column of cells in layers I–III, and (3) cells whose axons are directed toward the deep white matter and are therefore likely to be projection neurons. Some of these cells also contribute to the projections to the dentate gyrus and the hippocampus (Caballero-Bleda and Witter, 1994; Köhler, 1985a; Lingenhohl and Finch, 1991; Lorente de Nó, 1933). On the basis of their restricted axonal distribution, several of the smaller cell types in the entorhinal cortex have been classified as interneurons. The majority of these interneurons appear to be GABAergic (Swanson et al., 1987), although there is also a class of calretininpositive, GABA-negative neurons that may form asymmetrical synapses onto principal neurons (Wouterlood, 2002). GABAergic neurons are found in all layers of the entorhinal cortex, although they are most abundant in the superficial layers. GABAergic interneurons of the entorhinal cortex can be subcategorized on the basis of their colocalization with a variety of substances such as peptides or one of the various calcium-binding proteins (for a detailed review, see Wouterlood, 2002). The superficial layers contain a high number of cells that are immunoreactive for parvalbumin and all of these neurons colocalize GABA (Miettinen et al., 1996; Wouterlood et al., 1995). The number of parvalbuminpositive neurons in layers V and VI is much lower (Wouterlood et al., 1995). In contrast to earlier studies (Celio, 1990) indicating that the number of calbindinD28-positive neurons in layers II and III is much lower, the actual density of such neurons appears to be related to the different subdivisions of the entorhinal cortex.

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Layers II and III of the lateral entorhinal cortex contain high numbers of calbindin-positive neurons, whereas these cells are only sparsely present in layers II and III of the medial subdivision (Wouterlood, 2000). In the deep layers, calbindin cells are less numerous and preferentially found in layer V. Calretinin-, substance P-, and somatostatin-positive cells are quite abundant in the deep layers (Davies and Köhler, 1985; Köhler and Chan-Palay, 1983; Miettinen et al., 1997; Swanson et al., 1987; Wouterlood et al., 2000), whereas neurons positive for neuropeptide Y are most numerous in layer VI (Köhler et al., 1986).

Subdivisions There have been several published attempts at subdividing the rat entorhinal cortex, and unfortunately there are almost an equal number of different opinions concerning the number and terminology of the subfields (see Stephan, 1975, for a detailed review of the older literature and Witter, 2002, for a short historical summary). In his extensive comparative studies, Brodmann (1909) parceled the entorhinal cortex into two fields, a lateral area 28a and a medial area 28b. Subsequent usage of this terminology was confounded by the reports of Krieg (1946a, 1946b) in the rat in which he applied the term 28a to the medial subdivision and 28b to the lateral part of the entorhinal cortex. It is now generally accepted that the entorhinal cortex can be subdivided into two general areas, the lateral entorhinal area (LEA) and the medial entorhinal area (MEA; Fig. 1C). Layer II in the LEA is more clearly demarcated than in the MEA and the cells are very densely packed and tend to be clustered in islands. The cells in layer II of the MEA are somewhat larger and do not show a distinct clustering into islands, and the border between layers II and III is not as sharp as in the LEA, although in both entorhinal areas the all differences in cell sizes between layers II and III facilitate the delineation of the two layers. It should be pointed out, however, that there is some vagueness in the delineation of layer II in the most lateral parts of the LEA where layer II appears to be separated from the deeper cell layers by a cell-free zone. According to several authors (Blackstad, 1956; Caballero-Bleda and Witter, 1993; Swanson et al., 1987), layer II in the LEA actually consists of a layer IIa that is separated from layer IIb by a cell-free zone. Layer IIb is thus not clearly separated from layer III but is formed by cells in the most superficial part of the deeper cell plate. The other cell layers, in particular layers IV–VI, can be better differentiated from each other in the MEA than in the LEA, and cells in the MEA generally show a more radial or columnar arrangement. The lamina

dissecans of the MEA is sharply delineated but is less clear in the LEA. Although the notion of two major subdivisions of the entorhinal cortex is well accepted, it is also true that several authors have felt the need to recognize more than two subdivisions of the entorhinal cortex (Blackstad, 1956; Haug, 1976; Insausti et al., 1997; Krettek and Price, 1977; Ruth et al., 1982, 1988; Swanson et al., 1987; Wyss, 1981). It should also be stressed that the terms lateral and medial entorhinal areas do not relate in a simple manner to the cardinal transverse plane of the rat brain. Both the LEA and the MEA have a more or less triangular shape. The LEA occupies the rostrolateral part of the entorhinal cortex; its base is oriented rostrally and its tip caudolaterally, next to the rhinal fissure. The MEA occupies the remaining triangular area which has its base caudally and its tip rostromedially such that the tip lies medial to the LEA (Figs. 1C and 15). To accommodate the oblique orientation of the entorhinal cortex and to address the obvious need for subdemarcations of the LEA and MEA, a different nomenclature for the rat entorhinal cortex has been proposed (Insausti et al., 1997). The areas that fit the general cytoarchitectonic features of the LEA and MEA as described above have been designated “caudal entorhinal” (CE—formerly a major portion of the MEA) and “dorsal intermediate entorhinal” fields (DIE—formerly a major portion of the LEA; see Fig. 11). According to several authors (Steward, 1976; Steward and Scoville, 1976; Wyss, 1981), the transitional area between the LEA and the MEA should be considered a third subdivision which has been called the intermediate entorhinal area (IEA); it has been claimed to share cytoarchitectonic features that are common to both the MEA and the LEA. In the nomenclature proposed by Insausti et al. (1997), this transitional area has been further subdivided into a part that cytoarchitectonically looks more like the CE and is called the “medial entorhinal” field (ME) and a part that looks more like the LEA, which is called the “ventral intermediate entorhinal field” (VIE). The most lateral part of the entorhinal cortex, i.e., the part lying closest to the rhinal sulcus, is cytoarchitectonically distinct from the rest of the LEA. This region is characterized by a loose arrangement of cells in layer II and this gives the impression that they extend into the molecular layer. Other differences are that no clear lamina dissecans can be distinguished and the deeper cell layers are relatively well differentiated when compared to the rest of the LEA. This area lies parallel to the rhinal sulcus and extends caudally along the rostral portion of the CE; it has been designated the “dorsolateral entorhinal cortex” (DLE). A comparably placed dorsolateral zone of the entorhinal cortex has been differentiated by several previous authors (Krettek

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FIGURE 15 Line drawing illustrating the laminar and topographic organization of the perforant projection to the dentate gyrus, the hippocampus, and the subiculum. The image on the left side of the figure represents the surface of the rat entorhinal cortex. The rhinal sulcus would be located to the left. The surface of the entorhinal cortex has been divided into medial (MEA) and lateral (LEA) areas. The illustration indicates that the perforant path projection to the dentate gyrus terminates in a laminar fashion with the LEA projecting superficially in the molecular layer and the stratum lacunosum-moleculare and the MEA projecting deeper. In CA1 and the subiculum, in contrast, the LEA terminates at the CA1/subiculum border whereas the MEA terminates more proximally in CA1 and more distally in the subiculum. Laterally situated portions of the entorhinal cortex project to septal levels of the hippocampal formation whereas progressively more medial portions of the entorhinal cortex project to more temporal levels of the hippocampal formation. Projections from the presubiculum terminate exclusively in the medial entorhinal area and projections from the retrosplenial cortex (RSP Ctx.) terminate in the caudal portions of the medial entorhinal area. The entorhinal cortex receives a prominent input from the perirhinal cortex and this terminates in the lateral aspects of both the LEA and the MEA.

and Price, 1977; Ruth et al., 1982, 1988). Since the terms LEA and MEA are well established in the current literature and most of the studies describe entorhinal connectivity with respect to these two subdivisions, we use the term LEA to include the DLE, DIE, and VIE, whereas the MEA includes the CE and ME (see below for further discussion). The last field of the entorhinal cortex, which has been labeled “amygdaloentorhinal cortex” (AE), is found at the transition between the ventrorostral part of the entorhinal cortex and the amygdala (Haug, 1976; Insausti et al., 1997). This area has been included by these authors on the basis of the observation that cells in the superficial layers project to the dentate gyrus. More recently the efferent projections of this area have been studied in detail using anterograde tracing techniques (Jolkkonen et al., 2001) and although projections to CA1 and the subiculum were reported, similar to other cortical areas belonging to the amygdaloid complex, no projections to the dentate gyrus were observed. This latter finding may thus be taken to imply that area AE is not part of the entorhinal cortex.

Connections of the Entorhinal Cortex Connections of the Entorhinal Cortex with the Hippocampal Formation The major input to the dentate gyrus and a prominent input to the hippocampus and the subiculum arises from the entorhinal cortex. Fibers of the so-called perforant pathway take their origin mainly from neurons located in layers II and III, although a smaller component of the projection originates in the deeper layers. Although it has been assumed for some time that the stellate cells in layer II and the pyramidal cells in layer III were the exclusive cells of origin for the perforant pathway, recent studies have indicated that many other cell types, including at least a few GABAergic neurons and calbindin (Wouterlood, 2002), also project to various hippocampal subfields (Caballero-Bleda and Witter, 1994; Germroth et al., 1989a, 1989b). The perforant path terminates in all subdivisions of the hippocampal formation (Köhler, 1985a, 1985b, 1986, 1988; Ruth et al., 1982, 1988; Steward, 1976; Steward and Scoville, 1976; Witter, 1989; Witter et al., 1989). After

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leaving the entorhinal cortex, the fibers enter the underlying white matter and the angular bundle (Fig. 2C). They then traverse the pyramidal cell layer of the subiculum and cross the hippocampal fissure to enter the dentate gyrus or distribute to the molecular layer of the subiculum and the hippocampus. The entorhinal cortex fibers also take alternative routes to the other hippocampal fields such as projecting through the alveus before entering the hippocampus or by traversing the molecular layers of the entorhinal cortex and the pre- and parasubiculum. A recent analysis of the alvear pathway has shown that entorhinal fibers heading toward increasingly more septal levels of CA1 and the subiculum preferentially travel by way of this pathway. Moreover, the crossed projection to CA1 travels exclusively by way of the alvear path (Deller et al., 1996a). In the present description, we only employ the term perforant pathway to encompass all of the projection systems from the entorhinal cortex to the hippocampal formation (for a detailed description of these connections, see Witter et al., 2000b; see also Witter, 1989). Dentate gyrus The dentate gyrus is the “traditional” target of the entorhinal–hippocampal fibers, but there is ample evidence that the entorhinal cortex also projects to the hippocampus and the subiculum as well (Naber et al., 2001a; Steward, 1976; Steward and Scoville, 1976; Witter, 1989). The projection to the dentate gyrus arises mainly from layer II of the entorhinal cortex (Ruth et al., 1982, 1988; Schwartz and Coleman, 1981; Segal and Landis, 1974; Steward and Scoville, 1976). In the molecular layer of the dentate gyrus, the terminals of the perforant path fibers are strictly confined to the outer or superficial two-thirds where they form asymmetric synapses (Nafstad, 1967). These occur most frequently on the dendritic spines of dentate granule cells, although a small proportion of perforant path fibers terminate on parvalbumin/GABA-positive neurons (Zipp et al., 1989). Within the outer two-thirds of the molecular layer, perforant path synapses make up at least 85% of the total synaptic population (Nafstad, 1967). There is evidence that a minor component of the projection also comes from the deep layers (IV–VI) of the entorhinal cortex (Köhler, 1985b). In contrast to the superficial pathway, this deep-originating projection preferentially distributes to the inner portion of the molecular layer, the granule cell layer, and the subgranular zone, where they form asymmetrical synapses onto granule cell dendrites as well as on their somata and onto spine-free dendrites in the subgranular zone. The latter most likely represent dendrites of interneurons (Deller et al., 1996b). Two fundamental organizational features of the entorhinal projection to the dentate gyrus have been

described in the literature, one related to the distribution along the radial axis of the molecular layer and one related to its transverse axis (Fig. 15). With respect to the radial organization, it has been well established that fibers originating in the LEA terminate in the outer one-third of the molecular layer and fibers from the MEA terminate in the middle onethird of the molecular layer (Fig. 15; Hjorth-Simonsen, 1972b; Steward, 1976; Witter, 1989; Wyss, 1981). On the basis of experiments with anterograde transport of tritiated amino acids, Steward (1976) suggested that the perforant path could actually be subdivided into lateral, intermediate, and medial components (Wyss, 1981). The intermediate pathway was thought to originate in the intermediate entorhinal area (see above) and terminate in the molecular layer in between the projections from the LEA and the MEA. However, a study conducted with the anterograde tracer PHA-L (Witter, 1990) indicates that there is probably not a discrete intermediate perforant path. Rather, the perforant pathway is organized such that, as the position of the cells of origin move from lateral to medial in the LEA or in the MEA, the densest portion of the terminal field in the molecular layer correspondingly shifts from superficial, i.e., closer to the hippocampal fissure, to deep, i.e., closer to the granule cell layer. There are conflicting papers on the transverse distribution of the perforant path projection. Wyss (1981) reported that the lateral perforant pathway preferentially projects to the enclosed blade of the dentate gyrus whereas the medial component either does not show a preference or predominantly targets the free blade. In a PHA-L study (Witter, 1990), projections from both subdivisions of the entorhinal cortex were clearly seen to reach the entire transverse extent of the molecular layer though the projection does have a topography that again relates to the lateral-to-medial axis of origin within the entorhinal cortex. Lateral parts of the LEA project almost exclusively to the enclosed blade and more medial parts of the LEA preferentially project to the free blade. Lateral and caudal parts of the MEA project more heavily to the enclosed blade whereas medial parts of the MEA provide a denser projection to the free blade. Finally, in a more recent report that only studied inputs to the septal portions of the dentate gyrus, Tamamaki (1997) used anterograde tracing with the fluorescent lipophylic tracer DiI and reported that the LEA projects with thick fibers to the enclosed blade and with thin fibers to the free blade. The fiber profile of fibers originating in the MEA was thick in the free blade and thin in the enclosed blade. The perforant path is most likely glutamatergic (Fonnum, 1970), which is in line with the overall observations that the majority of entorhinal fibers in the

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dentate gyrus, as well as in the hippocampal fields and the subiculum, form asymmetrical synapses (Nafstad, 1967; Desmond et al., 1994; Witter et al., 1992b; Bakst-te Bulte et al., 2003). With respect to the projection to the dentate gyrus, the terminals of the lateral perforant pathway are also enkephalin immunoreactive, whereas those of the medial pathway are immunoreactive for CCK (Fredens et al., 1984). Finally it should be noted that no cells in the dentate gyrus project back to the entorhinal cortex, which is in line with the overall unidirectional flow of information thought to be characteristic for the hippocampal system. Hippocampus—Fields CA3/CA2 Although the entorhinal projection to CA3 is mentioned in most studies of the perforant path, the organization of this component of the projection has not been documented in much detail. The origin and terminal distribution appear to be very similar to those described for the dentate gyrus. Like the projection to the dentate molecular layer, projections from the LEA terminate superficially in the stratum lacunosum-moleculare and those from the MEA terminate more deeply in the same layer (Fig. 15). The types of synaptic contacts and numbers of synapses appear to be similar to those found in the dentate gyrus as well (Nafstad, 1967; Witter et al., 1992). In line with evidence from retrograde tracing studies indicating that the projections to CA3/CA2, like those to the dentate gyrus, originated from cells in layer II (Steward and Scoville, 1976), an intracellular labeling study by Tamamaki and Nojyo (1993) has convincingly demonstrated that collaterals of the same layer II cells reach both the dentate gyrus and fields CA3/CA2. This entorhinal projection appears to provide for a strong and highly modifiable excitatory input to CA3 (Urban et al., 1998). As in the dentate gyrus, cells of CA3 do not project to the entorhinal cortex. Hippocampus—Field CA1 The entorhinal projections to CA1 originate in layer III rather than layer II (Steward and Scoville, 1976). Not only are the cells of origin different for this component of the perforant path, but also the pattern of terminal distribution for this component of the perforant path projection is different from the layer II projection. In CA1, the medial and lateral components of the perforant pathway each terminate throughout the full radial extent of stratum lacunosum-moleculare (Fig. 15). In contrast to the projections to the dentate gyrus and CA3, and similar to what is described for the subiculum (see below), the lateral and medial components of the perforant path terminate at different transverse positions within CA1. The fibers from the MEA terminate in the proximal part of CA1, i.e., in the part that borders CA2. Fibers from

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the LEA, in contrast, terminate more distally in CA1, i.e., close to the subiculum (Fig. 15). Although Nafstad (1967) did not find much evidence for synaptic termination of perforant path fibers in CA1, which was most likely the result of the strikingly restricted distribution along the transverse axis, the size of fibers and terminals and the morphology of the terminals are similar to those in CA3 and the dentate gyrus (Desmond et al., 1994; Witter et al., 1992). The majority of entorhinal terminals, irrespective of their origin in the LEA or MEA, make asymmetrical synaptic contacts (>96%) with both dendritic spines (93%) and dendritic shafts (7%). These findings are interpreted as an indication that the perforant path projection to CA1 provides a strong excitatory input to CA1 pyramidal cells as well as a minor feed-forward inhibitory input onto these neurons by way of local interneurons, including parvalbumin-containing ones representing both basket cells and chandelier cells (Kiss et al., 1996). Although this excitatory input to CA1 has been strongly debated (see Naber et al., 1999, for further discussion), it has been convincingly shown, using in vivo electrophysiological approaches, that this input is indeed excitatory and that it is focally distributed (Canning and Leung, 1997; Naber et al., 1999). This latter feature, taken together with the quite strong innervation of basket and chandelier cells (Kiss et al., 1996), may easily explain why so many electrophysiological studies have failed to detect this input (Naber et al., 1999; Witter et al., 2000b). Interestingly, the direct entorhinal–CA1 pathway has been put into a functional context, suggesting that place field activity specific for CA1 neurons can be maintained without the CA3-to-CA1 input in the presence of intact entorhinal-to-CA1 input (Brun et al., 2002). CA1 is the first hippocampal field that originates a return projection to the entorhinal cortex and is thus different from the dentate gyrus and fields CA3/CA2 in this respect. Projections from CA1 to the entorhinal cortex originate from the full septotemporal and transverse extent of CA1 and appear to terminate more densely in the MEA than in the LEA. The CA1 projections to the entorhinal cortex terminate predominantly just below lamina dissecans in layer V (Finch and Babb, 1980, 1981; Finch et al., 1983; Naber et al., 2001a; Swanson and Cowan, 1977; Van Groen and Wyss, 1990b). Subiculum Because the perforant path fibers traverse the subiculum on their way to the dentate gyrus and hippocampus, the question of whether some of these fibers terminate in the subiculum has long been a matter of controversy. The consensus has been that fibers probably do not terminate in the subiculum but simply passed through it. Anterograde tracer studies

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indicated that perforant path fibers are directed toward the molecular layer of the subiculum but proof that these fibers form a terminal plexus among the subicular pyramidal cells was lacking (Steward, 1976; Wyss, 1981). Whereas Köhler (1986, 1988) failed to observe any marked projections to the subiculum from either the LEA or the MEA with the anterograde tracer PHA-L, more recent tracing studies, both at the light and electron microscopic levels, reported that the subiculum receives a strong projection from the entorhinal cortex (Baks-te Bulte et al., 2003; Naber et al., 2001a; Witter et al., 2000b) and these findings have been corroborated electrophysiologically (Behr et al., 1998). Fibers of both the lateral and the medial component preferentially target dendritic spines of presumed principal neurons with asymmetrical synapses (80%), whereas between 5 and 10% of the asymmetrical synapses terminate on dendritic shafts, most likely belonging to interneurons in the subiculum. Interneurons may also receive a minor inhibitory perforant path input in view of the symmetrical synapses onto dendritic shafts (10%) (Baks-te Bulte et al., 2003). As with the CA1 projection, depending on the location of the cells of origin in the entorhinal cortex, the fibers are directed toward restricted transverse portions of the subiculum and terminate in the outer two-thirds of the molecular layer. The lateral component of the perforant pathway preferentially projects to the part of the subiculum that is adjacent to CA1, i.e., the proximal part of the subiculum, and the medial component distributes to more distal portions of the subiculum, i.e., closer to the presubiculum (Naber et al., 2001a; Witter et al., 2000b; Fig. 15). The projection originates mainly from layer III, although the axons of layer II cells, which cross the subiculum on their way to the dentate gyrus and CA3, appear also to give off some collaterals that terminate in the subiculum (Lingenhohl and Finch, 1991; Tamamaki and Nojyo, 1993). The subiculum reciprocates the entorhinal input. Projections from the subiculum reach all parts of the ipsilateral entorhinal cortex where they terminate within and deep to the lamina dissecans; termination is particularly dense in the layer of the large pyramidal neurons just deep to the lamina dissecans (Bartesaghi et al., 1989; Beckstead, 1978; Finch et al., 1983, 1986; Kloosterman et al., 2003a, 2003b; Köhler, 1985a; Naber and Witter, 1998; Naber et al., 2001a; Swanson and Cowan, 1977). A minor component of the subicular projection also extends superficial to the lamina dissecans, predominantly innervating layer III. Therefore, the overall organization of the subiculoentorhinal projection mimics that of the CA1–entorhinal projection. Subicular fibers generally form asymmetrical synapses with spines (67.5%) and dendritic shafts (23.5%) of cells that

most likely have their cell bodies in the deep layers of the EC. A minority of these fibers form symmetrical synapses, taken to indicate a small inhibitory input from the subiculum to layer V (Van Haeften et al., 1995). The presence of asymmetrical synapses at the termination of this pathway is consistent with its reported excitatory influences on the entorhinal cortex (Jones, 1993; Kloosterman et al., 2003b). Topography of entorhinal–hippocampal reciprocal pathways In addition to the distribution of the lateral and medial components of the perforant path along the superficial-to-deep gradient in the molecular layer of the dentate gyrus and stratum lacunosummoleculare of CA3 and along the transverse axis of CA1 and the subiculum, both components of the perforant pathway projection also demonstrate a similar septotemporal organization. Cells located laterally in the entorhinal cortex project to septal levels of the hippocampal fields while cells located progressively more medially project to more temporal levels of the hippocampal subfields (Fig. 15). This organization leads to a pattern of connections such that the septal parts of the dentate gyrus, the hippocampus and the subiculum, receive inputs from lateral parts of the LEA and lateral and caudal parts of the MEA (Fig. 15), whereas the temporal portions of the dentate gyrus, the CA fields and the subiculum, receive input from more medial portions of both the LEA and the MEA (Ruth et al., 1982, 1988; Witter, 1989; Witter et al., 2000b). This topographical organization has been convincingly demonstrated in a series of retrograde tracing experiments, in which discrete injections in septal, midseptotemporal, and temporal levels of the dentate gyrus labeled populations of entorhinal neurons, in both the LEA and the MEA, with a different lateral to medial position (Dolorfo and Amaral 1998a; Fig. 15). Although the domains of the entorhinal cells projecting to septal, midseptotemporal, and temporal levels do not show much overlap, one should take into account that a single entorhinal layer II neuron, as shown with intracellular filling, may distribute an axon along as much as 2 mm (20–25%) of the septotemporal extent (Tamamaki and Nojyo, 1993), such that overlapping terminal zones in the dentate gyrus of these populations of layer II neurons may exist. The projections from CA1 and the subiculum back to the entorhinal cortex are also topographically organized such that septal portions of CA1 and the subiculum project chiefly to lateral parts of the entorhinal cortex and more temporal parts of CA1/subiculum project to more medial parts of the entorhinal cortex (Naber and Witter, 1998; Kloosterman et al., 2003a). The transverse location of the cells of origin in CA1

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and the subiculum determines whether they terminate in the MEA or the LEA. The projections from the proximal part of CA1 and the distal part of the subiculum distribute exclusively to the MEA whereas cells located in the distal part of CA1 and the proximal part of the subiculum project mainly to the LEA (Naber et al., 2001a; Tamamaki and Nojyo, 1995; Witter et al., 2000b). This organization along the septotemporal and transverse axes indicates that the projections from the entorhinal cortex to CA1 and the subiculum are in register, i.e., are point-to-point reciprocal, with the projections from CA1 and the subiculum back to the entorhinal cortex. Interestingly, the overall organization of these reciprocal entorhinal-to-CA1/subiculum connections is also in register with the projections interconnecting CA1 to the subiculum as described in page 652 (Naber et al., 2001a). Crossed Hippocampal Connections The entorhinal cortex also gives rise to a crossed projection to components of the contralateral hippocampal formation. The largest component of this projection is directed toward the dentate gyrus, but fields CA3 and CA1 of the hippocampus and the subiculum also receive a contralateral input. The crossed entorhinal projection is most prominent to the more septal portions of the hippocampal subfields and rapidly diminishes in strength at more temporal levels (Goldowitz et al., 1975; Köhler, 1988; Steward, 1976; Van Groen and Wyss, 1990a, 1990c). With respect to the laminar origin of the crossed projection, Steward and Scoville (1976) reported that they arose exclusively from cells of layer III of the entorhinal cortex. No crossed projections to the pre- and parasubiculum have been described. As mentioned above, the crossed projection to fields CA1 and the subiculum preferentially travel by way of the alvear pathway (Deller et al., 1996a); the crossed dentate projection mainly takes the more common perforant path trajectory, crossing the midline through the ventral hippocampal commissure. Associational and Commissural Connections The entorhinal cortex contains a substantial system of associational connections. Available data indicate that intraentorhinal fibers are directed mainly in a longitudinal direction and there are relatively modest connections that link different transverse or mediolateral regions of the entorhinal cortex (Dolorfo and Amaral., 1998b; Köhler, 1986, 1988). Regarding the longitudinal projections, one organizational principle is that cells in a particular layer innervate more superficial layers. A second principle is that associational projections tend to go from caudal to rostral in the entorhinal cortex. Finally, whereas projections that originate in the LEA

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decrease in density with increasing distance from their origin, those from the MEA show an increase in density with increasing distance from their origin. It thus appears that the associational connections that originate in the MEA are more pronounced than those from the LEA. It is worth mentioning that the overall predominance of deep-to-superficial connectivity is in line with all reports based on Golgi studies and intracellular fillings as described previously on page 670, an observation that seems very different from reports in the preand parasubiculum where the preferential direction is from superficial to deep (see sections “Presubiculum” and “Parasubiculum”). With respect to the transverse connections, Köhler (1986, 1988) reported that superficial layers of the entorhinal cortex preferentially project medially while projections from deeper layers preferentially travel in a lateral direction. The finding that needs to be stressed is that the transverse connections are very limited as mentioned above. Dolorfo and Amaral (1998b) further showed that the transverse spread of the longitudinally oriented connections are more or less confined to the three lateral-to-medial zones that project to different septotemporal levels of the dentate gyrus. The overall organization of the intrinsic network of the entorhinal cortex thus suggests that integration may take place within a band, projecting to a particular septotemporal segment of the dentate gyrus, but that relative independence of bands is the key feature of entorhinal information processing. Relatively strong commissural connections, arising from all portions of the entorhinal cortex, terminate predominantly in layers I and II of the homotopic area of entorhinal cortex (Köhler, 1986, 1988). Connections of the Entorhinal Cortex with the Parahippocampal Region Perirhinal and postrhinal cortices The entorhinal cortex sends projections to both the perirhinal and postrhinal cortex (Fig. 16B; Insausti et al., 1997; Burwell and Amaral, 1998a, 1998b). These projections preferentially originate in the deep layers of the entorhinal cortex, although the projections to the perirhinal area 35 (see section “Perirhinal and Postrhinal Cortices”) originate quite massively from superficial layers of the LEA. Moreover, both anterograde as well as retrograde data apparently show that these projections are strongest from the lateral and caudal rim of entorhinal cortex, i.e., those parts of both the LEA and MEA that are adjacent to the perirhinal and postrhinal cortex and that are reciprocally connected to the septal portion of the hippocampus. Although this does not imply that more central and medial portions of the entorhinal cortex do not project to the surrounding rim of the parahip-

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FIGURE 16 Unfolded maps of the parahippocampal region of the rat showing the patterns of intrinsic connections (left) and interconnections (right). The postrhinal cortex (POR) is shown in dark grey, and the perirhinal cortex (35 and 36) in middle grey. The entorhinal cortex (LEA and MEA) is indicated in three shades of light grey, indicating the lateral-to-medially located dentate projecting bands as reported by Dolorfo and Amaral (1998a). Reproduced with permission from Burwell (2000).

pocampal cortex, these projections are strikingly less strong. The connections from perirhinal and postrhinal cortex into the entorhinal cortex show a more or less reciprocal distribution, although the overall terminal distribution is more widespread. Data presented in different studies, however, are somewhat conflictive. According to Burwell and Amaral (1998a, 1998b) the perirhinal cortex preferentially projects to the most lateral and caudal strip of the entorhinal cortex, comprising portions of both the LEA and MEA, and the postrhinal cortex preferentially projects to more central portions of both the LEA and MEA. In contrast, according to Naber et al. (1997) the densest projections to the caudal part of the entorhinal cortex, i.e., the caudal rim of the MEA, originate in the postrhinal cortex. However, there is general agreement that the strongest input to the LEA originates in the perirhinal area 35, whereas that to the MEA comes from the postrhinal cortex (Burwell and Amaral, 1998b). Projections from the perirhinal and postrhinal cortices mainly terminate in layer III of the entorhinal cortex with a weaker innervation of deep layer I at the border with layer II. No detailed studies have been carried out with respect to the potential postsynaptic targets of these inputs. Preliminary data indicate that perirhinal inputs target both principal neurons as well as parvalbuminpositive interneurons in the dorsolateral portions of the entorhinal cortex (Wouterlood et al., 1998). Pre- and parasubiculum As described above, a minor component of the perforant pathway courses through the molecular layer of the para- and presubicu-

lum. In addition, fibers from cells in layer Va and, to a lesser extent, in layers III, Vb, and VI of the entorhinal cortex terminate weakly in layer I of the presubiculum and parasubiculum (Köhler, 1986, 1988; Van Groen and Wyss, 1990a, 1990c). This modest projection from the entorhinal cortex to the pre- and parasubiculum stands in marked contrast to the previously described dense projections to the entorhinal cortex from these two areas (see sections “Presubiculum” and “Parasubiculum”), indicating that the pre- and parasubiculum should be considered functionally as input structures to the entorhinal cortex. Extrinsic Inputs and Outputs Cortical afferents The entorhinal cortex of the rat receives inputs from a variety of cortical regions (Burwell and Amaral, 1998b). These cortical inputs form two groups: those that terminate in the superficial layers (I–III), and those that preferentially distribute to the deep layers (IV–VI). The first category delivers information to the entorhinal neurons that are the main source of projections to the dentate gyrus, hippocampus, and the subiculum. The second group of inputs terminates on the deeper cells of the entorhinal cortex which receive processed information from the other hippocampal fields and also give rise to projections back to certain cortical regions; the second class of cortical inputs might then be considered to have influence on the output side of the hippocampal formation. In general, the cortical afferents that reach the deep layers terminate rather diffusely, whereas the affer-

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ents that terminate superficially have a more restricted mediolateral and/or rostrocaudal distribution. A substantial input to the superficial layers of the entorhinal cortex originates from olfactory structures of the telencephalon, in particular from the olfactory bulb, the anterior olfactory nucleus, and the piriform cortex (Haberly and Price, 1978; Kosel et al., 1981). Using a semiquantitative retrograde tracing approach, Burwell and Amaral (1998b) indicate that roughly one-third of the cortical input originates in layer II of the piriform cortex. Olfactory projections terminate throughout most of the rostrocaudal extent of the LEA and the MEA and selectively in layer I and the superficial part of layer II. Only the most caudal portion of the rat MEA does not receive olfactory inputs. Whereas the rostrolateral part of the LEA receives 45% of its inputs from the piriform cortex, these percentages drop dramatically in more caudomedial portions receiving only 16% piriform input. A similar trend has been described comparing rostrolateral versus caudomedial MEA (Burwell and Amaral, 1998b). Olfactory fibers terminate on cells in layers II and III of the entorhinal cortex. Olfactory fibers also terminate on GABAergic neurons in layer I that presumably interact with principle cells in layers II and III (Carlsen et al., 1982; Wilson and Steward, 1978; Wouterlood and Nederlof, 1983; Wouterlood et al., 1985). Interestingly, the projection neurons in the olfactory bulb are to a large extent calretinin positive and these fibers form asymmetrical contacts in the molecular layer, including contacts onto calretinin-positive, GABA-negative neurons (Wouterlood, 2002). Cortical afferents to the deep layers of the entorhinal cortex arise from a variety of cortical areas that together can be considered as limbic or paralimbic cortices (Lopes da Silva et al., 1990). Although detailed anterograde tracer studies of cortical inputs to the entorhinal cortex are not available in the rat, the following inputs have been described in the literature. Projections from the agranular insular cortex distribute preferentially to the ventral bank of the rhinal sulcus (perirhinal cortex) and to directly adjacent parts of the entorhinal cortex (Deacon et al., 1983; Markowitsch and Guldin, 1983). Additional inputs arise from the medial prefrontal region, in particular from the infralimbic, prelimbic, and anterior cingulate cortices (Beckstead, 1978, 1979; Sesack et al., 1989; Takagishi and Chiba, 1991; White et al., 1990; Witter 2003). Finally, the retrosplenial cortex (area 29) also projects to the entorhinal cortex. These projections originate from both the granular and the dysgranular subdivisions of the retrosplenial cortex and terminate almost exclusively in the most caudal portions of the MEA (Wyss and Van Groen, 1992; Jones and Witter, unpublished observations).

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The most detailed analysis of these cortical inputs is presently the study by Burwell and Amaral (1998b; Figs. 17 and 18). Their results largely corroborate these earlier scant reports. Inputs from the prefrontal cortex to both the LEA and the MEA make up about 10% of the total cortical input, but there are some differences with respect to the overall composition of those inputs. The origins of inputs to the LEA include both medial prefrontal and orbital regions, whereas the largest input to the MEA originates in the medial orbital region. Most of these inputs originate in layer II, although cells in layer V of the prelimbic cortex and superficial V and VI of the medial orbital region contribute as well. Regarding inputs from the insular cortex, these make up a larger proportion of the LEA inputs (20%) compared to the MEA (6%) inputs. The densest projection to both entorhinal subdivisions arises from layers II and III of the agranular insular cortex. Both the LEA and the MEA appear to receive weak inputs (3%) from ventral parts of the temporal cortex, adjacent to the perirhinal and postrhinal cortex. The MEA receives three to four times more input from retrosplenial, parietal,

FIGURE 17 Schematic representation of main cortical inputs (expressed as percentages of total) to the lateral (A) and medial (B) entorhinal cortex. For both lateral and medial entorhinal cortex the differential inputs to the lateral-to-medially located dentate projecting bands as reported by Dolorfo and Amaral (1998a) are indicated as well (compare with Fig. 20). Reproduced with permission from Burwell (2000).

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FIGURE 18 Diagram representing the pattern and strength of cortical connectivity of the hippocampal formation and the parahippocampal region as well as the intrinsic connections. The thickness of the solid lines represents the relative strength of these connections based on densities of retrograde labeled neurons. Open lines reflect connections that have been reported but for which no comparable quantitative data are available. ACAd and v, dorsal and ventral anterior cingulate cortices; AId,v, and p, dorsal, ventral, and posterior agranular insular cortices; AUD, primary auditory cortex; AUDv, auditory association cortex; DG, dentate gyrus; GU, gustatory cortex; HPC, hippocampus proper; LEA, lateral entorhinal cortex; MEA, medial entorhinal cortex; MOp and MOs, primary and secondary motor areas; POR, postrhinal cortex; RSPd and v, dorsal and ventral retrosplenial cortices; SSp and SSs, primary and supplementary somatosensory cortices; Sub, subiculum; VISC, visceral granular insular cortex; VISl and m, visual association cortex; VISp primary visual cortex. Reproduced with permission from Burwell (2000).

and occipital regions compared to the LEA (on average 10% versus 3%, respectively), and the weak input from the anterior cingulate cortex is confined to the MEA. As can be seen in Fig. 17, the three rostrocaudal bands in the entorhinal cortex, defined on the basis of the topographical distribution of the perforant pathway along the hippocampal septotemporal axis do receive different sets of cortical inputs (see page 686). Subcortical afferents The entorhinal cortex receives subcortical inputs from several of the structures that innervate the other hippocampal and parahippocampal

fields as well. Although we provide a short description of the most relevant data, more detailed accounts are available in the literature (Lopes da Silva et al., 1990; Pitkänen et al., 2000; Swanson et al., 1987; Witter et al., 1989a). A rather prominent projection to the entorhinal cortex originates from the medial septal complex (Alonso and Köhler, 1984; Beckstead, 1978; Milner and Amaral, 1984; Saper, 1985; Swanson, 1978). This projection is topographically organized such that cells in the horizontal limb of the nucleus of the diagonal band preferentially distribute fibers to the most lateral part of the entorhinal cortex, whereas the medial septal

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nucleus and the vertical limb of the nucleus of the diagonal band project to more medial parts of the entorhinal cortex (Alonso and Köhler, 1984; Beckstead, 1978; Gage et al., 1984; Milner and Amaral, 1984; Saper, 1985; Swanson, 1978). Septal afferents terminate densely in the cell-sparse lamina dissecans and less densely in layer II. The entorhinal cortex also receives a substantial topographically organized input from the amygdaloid complex, in particular from the lateral, basal, accessory basal, medial, and posterior cortical nuclei (Fig. 10). This input mainly targets the rostrolateral and central portions of the entorhinal cortex, including most of the LEA and the rostromedial part of the MEA (area ME according to Insausti et al., 1997). The remaining caudodorsal part of the MEA appears almost devoid of amygdaloid inputs. Details of the topographic and laminar distribution of inputs from the various amygdaloid region can be found in the excellent review by Pitkänen et al. (2000; Price et al., 1987; see also De Olmos et al., Chapter 19, this volume). Amygdaloid inputs terminate preferentially in layers III and V. Efferents from the medial part of the lateral nucleus distribute most intensely to the deep part of layers III and V of dorsal and ventral parts of the LEA, but end also between the cell islands of layer II and in layer I. The fibers from the basal and accessory basal nuclei terminate diffusely in layers III to V, whereas those from the cortical nuclei and the periamygdaloid cortex preferentially project to layers I–III (Canteras et al., 1992a, 1995; Petrovich et al., 1996). Additional, moderately dense inputs originate from the ventral part of the claustrum or the endopiriform nucleus (Behan and Haberly, 1999; Eid et al., 1996; Krettek and Price, 1977; Wilhite et al., 1986), terminating preferentially in layer V of both the LEA and the MEA (Eid et al., 1996). The major thalamic input to the entorhinal cortex originates in the nucleus reuniens and in the nucleus centralis medialis. Minor inputs arise from the rhomboid, the paraventricular, and the parataenial nuclei (Van der Werf et al., 2002). Although initially no inputs from the anterior thalamic complex or the mediodorsal nucleus have been reported (Beckstead, 1978; Segal, 1977; Wyss et al., 1979b), according to more recent studies, layer V of the entorhinal cortex receives weak inputs from the anteromedial, anterodorsal, and anteroventral nuclei (Van Groen and Wyss, 1995; Van Groen et al., 1999). Fibers from the nucleus reuniens terminate densely in the deep part of layer I (layer Ib) and in layer III, with minor innervation of neurons in layer II (Herkenham, 1978; Wouterlood, 1991; Wouterlood et al., 1990). Interestingly, a separate population of nucleus reuniens cells projects to the entorhinal cortex and to CA1 and the subiculum (Dolleman-Van der Weel and Witter, 1996).

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The entorhinal cortex receives additional, rather diffuse inputs from various structures in the hypothalamus and the brain stem (Beckstead, 1978; Köhler and Steinbusch, 1982; Segal, 1977; Swanson, 1982). These include afferents from: (i) the supramammillary nucleus that terminate rather diffusely with some preference for layers III–VI (Haglund et al., 1984); (ii) the tuberomammillary nucleus (Köhler, 1985a; Saper, 1985; Wyss et al., 1979b), distributing diffusely in the the LEA and the MEA; (iii) the lateral hypothalamic area (Köhler et al., 1984b) reaching preferentially the deep layers of the entorhinal cortex; (iv) the ventral tegmental area (Fallon et al., 1978; Swanson, 1982), terminating preferentially in a restricted rostrolateral part of the LEA, where the presumably dopaminergic fibers are arranged in dense, columnar patches in layers I–III; (v) the central and dorsal raphe nuclei, terminating diffusely in all layers, with a preference for the superficial layers (Azmitia and Segal, 1978) [the latter nuclei most probably supply the entorhinal cortex with its serotonergic innervation (Köhler and Steinbusch, 1982)]; and (vi) the locus coeruleus, which is a major source of input to the entorhinal cortex from the pontine region. It supplies the entorhinal cortex with a light, diffusely organized noradrenergic input that exhibits a slightly more dense termination in layer I (Moore et al., 1978). An additional moderate input originates from the nucleus incertus (Goto et al., 2001), distributing diffusely throughout all layers of both the LEA and the MEA. Minor pontine projections arise from the parabrachial nucleus, the dorsal tegmental nucleus (Groenewegen and Van Dyk, 1984), and the nucleus subcoeruleus (Datta et al., 1998). Cortical efferents Efferents of the entorhinal cortex reach widespread parts of the limbic, paralimbic, and olfactory regions of the cortex (Insausti et al., 1997). The projections to olfactory areas predominantly originate from layers II, III, and Va of the central portions of both the LEA and the MEA (DeOlmos et al., 1978; Insausti et al., 1997). The same layers and regions emit rather strong projections to the infralimbic cortex; the ventral taenia tecta; and the prelimbic, orbitofrontal, and agranular insular cortices, although at more caudal levels layer V cells contribute as well. Only moderate projections reach the retrosplenial cortex and those arise preferentially from layer V neurons in the most caudal part of the MEA (Conde et al., 1995; Insausti et al., 1997; Wyss and Van Groen, 1992). An important issue is whether or not the entorhinal cortex of the rat, like that of the monkey, (Van Hoesen, 1982), gives rise to prominent and widespread projections to the multimodal association cortex. Initially conflicting reports had been published, suggestive of

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either very limited projections to cortex directly adjacent to the parahippocampal region (Kosel et al., 1982) or alternatively much more widespread projections (Swanson and Köhler, 1986). The detailed study of Insausti et al. (1997), however, confirmed the report by Sarter and Markowitsch (1985) that only cells in the deep layers of the dorsolateral area (DLE) give rise to the extensive projections reported by Swanson and Köhler (1986). Based on the distribution of immunoreactivity for parvalbumin (Wouterlood et al., 1995; Burwell et al., 1995; Witter et al., 2000) and various subunits of glutamate receptors (Martin et al., 1993), the border between the entorhinal and perirhinal cortices appears to be oblique such that those cells most likely belong to the entorhinal cortex. It should be stressed though that entorhinal projections to these widespread neocortical regions are rather weak and that these projections increase in strength quite dramatically when the origin shifts from the dorsolateral entorhinal rim into the adjacent portions of the perirhinal and postrhinal cortices (see following section). In general, entorhinal– cortical projections terminate preferentially in layers I, II, and III, but particularly in the more densely innervated areas projections also terminate in layer V. Moreover, most of the entorhinal cortical efferents are bilaterally distributed though the contralateral component is much weaker. Subcortical efferents Like the hippocampal formation, but unlike the pre- and parasubiculum, the entorhinal cortex projects to the septal region (Alonso and Köhler, 1984; Swanson and Cowan, 1977). Fibers mainly arise from cells in layer Va of the LEA and MEA, although in the most medial part of the LEA and MEA many layer II cells contribute to these projections (Alonso and Köhler, 1984). The fibers from the entorhinal cortex preferentially terminate in the lateral septal complex, although a minor component distributes to the medial septal complex as well. The entorhinal cortex also projects widely to the amygdala especially to the basal nucleus, although medial portions of the lateral nucleus, the accessory basal nucleus, and the posterior cortical nucleus are among the targets of entorhinal fibers (Fig. 10). The fibers predominantly originate from cells in layers V of the LEA, although a few cells in more superficial layers may also contribute to the projection (McDonald and Mascagni,1997; Ottersen, 1982; Shi and Cassell, 1999; Veening, 1978). No projections appear to originate from the MEA (Pitkänen et al., 2000). The entorhinal cortex projects bilaterally to the striatum, in particular to the ventral portion, i.e., the nucleus accumbens and adjacent parts of the olfactory tubercle (Phillipson and Griffiths, 1985; Wyss, 1981). These

projections originate mainly from layer V and are topographically organized (Phillipson and Griffiths, 1985) such that medial parts of both the LEA and MEA project to the caudomedial portion of the nucleus accumbens and more lateral portions of the entorhinal cortex project to more lateral parts of the nucleus accumbens. Finally, there have been no reports of entorhinal projections to the thalamus or brain stem (Herkenham, 1978; Wyss, 1981; see also Van der Werf et al., 2002).

PERIRHINAL AND POSTRHINAL CORTICES General Description and Topology The perirhinal and postrhinal cortices of the rat have been differentiated from each other only recently (Burwell et al., 1995). Initially, both areas were taken together as perirhinal areas 35 and 36 (or 35 and ectorhinal cortex).The overall appearance of these regions is such that they can be classified as agranular and/or dysgranular cortex, such that there is a general homogeneous transition from layers III to V with a variably developed layer IV in between. As implied by their respective names, these regions are strongly related to the rhinal fissure. The perirhinal cortex is situated more rostrally along the posterior half of the rhinal fissure, whereas the postrhinal cortex is related to the posterior and extremely shallow portion of the fissure (Figs. 1B and 1C). The borders of these two regions have been quite controversial and it was only recently that the cortical area associated with the posterior portion of the rhinal fissure was subdivided consistently into perirhinal and postrhinal cortices (Burwell, 2000, 2001; Burwell and Amaral, 1998a, 1998b; Burwell et al., 1995).

Subdivisions, Lamination, and Cell Types The perirhinal cortex of the rat is composed of the agranular area 35 and the dysgranular area 36. Although each area can be further subdivided (Burwell, 2001; Burwell and Witter, 2002), for the purpose of this chapter the present subdivisions suffice. Both regions have large, heart-shaped neurons in layer V. In area 36, the overall organization of layer V is more radial than that of area 35. Area 35 shows a laminar differentiation much poorer than that of area 36. In particular, the border between layers II and III is very hard to discern. In area 36, layer II tends to be irregular and patchy (Burwell et al., 1995; Naber et al., 1997). Not much is known about the cell types present in the different subdivisions of the perirhinal cortex but a number of recent studies have reported a number of interesting

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features with respect to electrophysiological and morphological characteristics. Layers II and III contain a high number of so-called late-spiking pyramidal cells (Beggs et al., 2000). Such cells can delay the onset of their spike trains by several seconds. Similar cell types have been reported in layer V (Moyer et al., 2002) and in layer VI as well (McGahn et al., 2001). In layer V, these cells are pyramidal cells, as in the superficial layers, and in layer V they form about 14% of the total population. The rest of the recorded pyramidal cells are either regular spiking or burst spiking neurons. Burst spiking neurons have a nontapering thick apical dendrite spanning the entire width of the cortex and have a tuft in layer I. Both the late spiking and regular spiking pyramidal cells have thin apical dendrites and a somewhat variable morphology, not always showing an apical dendrite reaching into the superficial layers. The axons of all layer V cells reach layer VI and enter the underlying white matter. Some of the late spiking neurons distribute axon collaterals to layers II and III (Moyer et al., 2002). In layer VI, the vast majority of neurons are of the late spiking type (86%) and the population comprises both pyramidal and nonpyramidal cells. The second most common cell type in layer VI is single spiking neurons (7%), which as yet have never been reported in cortex. Finally, regular spiking neurons are virtually absent in layer VI (McGahn et al., 2001). It has been suggested that the presence of late spiking neurons might permit a recurrent network to hold information or encode temporal intervals. This finding is of interest in light of recent suggestions that these specific neuronal characteristics of the perirhinal cortex may be specifically involved in the integration of cortical information with inputs from the amygdaloid complex (Kajiwara et al., 2003). The postrhinal cortex is a rather primitive looking cortical region, which can also be subdivided into dorsal and ventral subdivisions (Burwell and Witter, 2002). The ventral region is dorsal to the most caudolateral part of the entorhinal cortex and has an overall agranular to dysgranular appearance. In Timm sulfide silver-stained sections, it looks quite similar to perirhinal area 35. The rostral border of this region is with the perirhinal cortex. This border can be easily demarcated by the presence of ectopic layer II cells, located near the border with the entorhinal cortex. The dorsal postrhinal cortex has a much more developed layer IV and, overall, the cortical lamination appears to be well developed. Its dorsal border with the adjacent dorsal association cortex is characterized by a sudden change from a rather poorly differentiated layer IV into a well-developed granule cell layer. No detailed description of cell types, principal neurons, or interneurons is currently available.

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Hippocampal and Parahippocampal Connections Connections of the Peri- and Postrhinal Cortices with the Hippocampal Formation and Parahippocampal Region Neither the perirhinal nor the postrhinal cortices project to the dentate gyrus and CA3 (Canning and Leung, 1997; Naber et al., 1999, 2001a; see, however, Liu and Bilkey, 1997). Both tracing studies as well as electrophysiological data have convincingly indicated that both these cortical regions provide a strong and mainly excitatory input to CA1 and the subiculum. These projections are strongest to the subiculum, whereas the projection to CA1 is much weaker (Naber et al., 2001a, 2001b). Strong reciprocal connections originate in both the subiculum and CA1 (Burwell and Witter, 2002; Kloosterman et al., 2003a). Projections from the peri- and postrhinal cortices, like those from the entorhinal cortex, terminate in stratum the lacunosum-moleculare of CA1 and the stratum moleculare of the subiculum, similar to the projections from the entorhinal cortex. Interestingly, these inputs exhibit a striking topographical organization strongly reminiscent of the entorhinal–hippocampal organization. The main targets of these projections are the septal two-thirds of CA1 and the subiculum; along the transverse axis, the perirhinal inputs mainly terminate in the most proximal extreme of the subiculum (and adjacent distal CA1; Canning and Leung, 1997; Kosel et al., 1983; Naber et al., 1999), whereas those from the postrhinal cortex preferentially target the most distal extreme of the subiculum and to a much lesser extent the most proximal CA1 region, close to the border with CA2 (Naber et al., 2001a). This transverse organization is of interest in view of the preferential connectivity of the perirhinal cortex with the lateral entorhinal cortex and of the postrhinal cortex with the medial entorhinal cortex (see page 677). Although return projections originating from CA1 and the subiculum have been reported, details are still lacking and some conflicting data have emerged. Projections from CA1 are moderate to the perirhinal cortex, but quite strong to the postrhinal cortex. Retrograde tracing has indicated that whereas those to the perirhinal cortex originate preferentially from the septal portion of CA1, those to the postrhinal cortex preferentially originate from intermediate septotemporal and temporal levels of CA1 (Burwell and Witter, 2002). The projections from the subiculum to both cortical areas have recently been described using anterograde tracing techniques and it was observed that projections originate preferentially from the septal part of the subiculum, although, comparable to the CA1 inputs, those to the postrhinal cortex tend to find their origin slightly

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more temporally. Subicular projections to both the perirhinal and postrhinal cortices terminate exclusively in layers V and VI (Kloosterman et al., 2003a). There are no significant connections from the peri- and postrhinal cortices and the pre- and parasubiculum. Associational and Commissural Connections Area 36 shows quite extensive associational projections such that any part appears connected to the entire region (Fig. 16A). In contrast to what has been observed in the entorhinal cortex, these associational connections do not show a clear rostrocaudal preference. There is, however, a marked dorsal-to-ventral gradient (Burwell and Amaral, 1998a), such that area 36 projects quite densely to area 35, but the reciprocal connection is less dense. The ventrally directed connection preferentially targets the same rostrocaudal level at which it originates. Cells of origin are in layers II, III, and VI, and the terminal distribution is in all layers. In contrast, the ventral-todorsal projection has a preferential origin in layers II and VI and terminates in layers I, II, and VI. Connections within the postrhinal cortex are extensive as well, but unlike the perirhinal cortex no clear directionality is present (Fig. 16A). The origin and terminal distribution is similar to that in area 36 in that cells in layers II, V, and VI are the major source, distributing projections in all directions, terminating in layers I and V/VI. The perirhinal areas 35 and 36 both project to the postrhinal cortex, with the strongest projection coming from area 36. Interestingly, an overall “reversed” topography is present in this connection. Rostral parts of perirhinal cortex preferentially project to caudal parts of the postrhinal cortex, whereas caudal perirhinal cortex projects more strongly to anterior postrhinal cortex. These projections originate from neurons in deep layer V and VI and terminate in layers I/II and V/VI. The postrhinal cortex projects to the perirhinal cortex, most strongly to area 36, and this projection appears stronger than the perirhinalto-postrhinal projection. The projection, which originates in layers II and V, terminates in a columnar fashion throughout all layers of the perirhinal cortex. The overall organization of these interconnections indicates that there is no strict reciprocity. Moreover, on the basis of laminar patterns it can be suggested that the projections from the perirhinal cortex to the postrhinal cortex, like those from area 35 to 36, have the features of feedback projections.

Extrinsic Connections Cortical Connections Cortical inputs to the perirhinal and postrhinal cortex have only recently been studied in some detail. Using retrograde tracing techniques, Burwell and Amaral

(1998b) provided a quantitative assessment of the major cortical inputs (Fig. 18) to the perirhinal and postrhinal cortices. These results are largely in line with the fragmentary anterograde and retrograde data found scattered throughout the literature (see also Burwell and Witter, 2002). For the perirhinal cortex, there is a striking difference between areas 35 and 36. Area 36 receives much more higher-order cortical input than 35. Area 36 receives about 35% of its input from adjacent ventral temporal cortex; additional strong inputs originate in the lateral entorhinal area and the postrhinal cortex. Minor inputs arise from auditory, somatosensory, and gustatory regions (Burwell and Amaral, 1998b; Naber et al., 2000a). In contrast, area 35 receives its predominant input from piriform, lateral entorhinal, and insular cortices. For both regions though, the inputs terminate preferentially in the superficial layers. These findings are largely in agreement with earlier reports on connections of the rat perirhinal cortex (Deacon et al., 1983). Most of these connections appear reciprocal, although details are lacking. The postrhinal cortex receives its main cortical inputs (in descending order of magnitude) from visual association cortex, parietal cortex, retrosplenial cortex, and ventral temporal domains. Similar to the connections of the perirhinal cortex, the postrhinal cortex cortical connections appear to be reciprocal. These findings are in line with previous reports indicating retrosplenial inputs to posterior portions of the perirhinal cortex (Wyss and Van Groen, 1992) and reports of connections of posterior perirhinal cortex with visual association cortex (Vaudano et al., 1991; Miller and Vogt, 1984; cf. Burwell and Amaral, 1998b). Subcortical Connections No systematical studies of the subcortical connections of the perirhinal and postrhinal cortices of the rat have been carried out. However, data from different studies taken together lead to the following conclusions. Reciprocal connections with the amygdala are quite extensive and dense (Pitkänen et al., 2000; Fig. 10). Area 35 receives its major input in layers I through V from the lateral and accessory basal nuclei. Return projections terminate densely in the magnocellular basal nucleus. This projection arises from layers II and V. Weaker projections reach the lateral and accessory basal nuclei (McDonald and Jackson, 1987; McIntyre et al., 1996; Shi and Cassell, 1998a, 1998b; Shi and Cassell, 1999). Although area 36 receives inputs from the amygdala, in general these are light compared to the innervation of area 35. However, layer I of area 36 receives a dense innervation that originates in the dorsolateral part of the lateral nucleus and the magnocellular basal

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nucleus (Pitkänen et al., 2000). In contrast, all layers of area 36 contribute to a strong projection to the lateral nucleus, whereas moderate projections reach the accessory basal nucleus and layer III of the periamygdaloid cortex. Regarding projections of area 36 to the central nucleus, some conflicting data have been reported (for further details see Pitkänen et al., 2000). With respect to connections between the postrhinal cortex and the amygdala almost no data are currently available. According to Pitkänen et al. (2000), the postrhinal cortex projects to the lateral nucleus and receives inputs from the lateral and accessory basal nuclei. The perirhinal cortex also projects to the nucleus accumbens (McIntyre et al., 1996) and light return projections originate from the endopiriform nucleus and the claustrum (Behan and Haberly, 1999; McIntyre et al., 1996). Thalamic inputs to both the postrhinal and perirhinal cortices, as well as corticothalamic projections, have been described although systematic studies are lacking. The postrhinal cortex appears to be reciprocally connected with the lateral posterior nucleus (Deacon et al., 1983). The perirhinal cortex receives input from the anteromedial nucleus (Van Groen et al., 1999), the reuniens/perireuniens nuclei (Dolleman-van der Weel and Witter, 1996), the paraventricular nucleus (Moga et al., 1995), the posterior intralaminar nucleus, the suprageniculate nucleus, the medial division of the medial geniculate nucleus, the peripeduncular nucleus, and the posterior nucleus (Linke and Schwegler, 2000; Linke, 1999; McIntyre et al., 1996; Romanski and LeDoux, 1993). Finally, an input to the perirhinal cortex from the posterior nucleus of the hypothalamus has been reported (Vertes et al., 1995), whereas the suprachiasmatic nucleus apparently receives input from the perirhinal cortex (Krout et al., 2002). A connection has been reported from the perirhinal cortex to the raphe complex and this is reciprocated (Hermann et al., 1997; Swanson, 1987; Fig. 5B).

CONCLUSIONS: THE ORGANIZATION OF HIPPOCAMPAL CIRCUITRY AND THE FLOW OF INFORMATION PROCESSING In the following sections we raise a number of issues concerning the topography of information processing within the hippocampal region. In many respects this discussion oversimplifies the various connections described above in order to conclude with a number of principles that may provide insight into the flow of information through, and thus the function of, the various regions of the hippocampal region. Some

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assumptions, warranted or not, are made concerning certain of the hippocampal connections. For example, although the hippocampal formation receives both cortical and subcortical inputs, information received from cortical sources is regarded to be the major substrate by which the hippocampal formation carries out its cognitive/mnemonic functions. Subcortical inputs are regarded generally to be modulatory and may reflect, in a very broad sense, the behavioral state of the organism. These modulatory inputs arise predominantly from the medial septal complex, the hypothalamus, and the brain stem. While the return hippocampal projections to these subcortical brain regions through the fornix were long thought to be the sole output of the hippocampus, little is known concerning their function. Lopes da Silva et al. (1990) have proposed that these projections provide feedback about ongoing hippocampal activity to the modulatory systems, and a discussion concerning the functional relevance of these modulatory or control systems can be found in their review.

The Lamellar Concept of Hippocampal Information Flow Is Not Compatible with Neuroanatomical Data The lamellar concept, which was developed in the early 1970s on the basis of existing neuroanatomical and electrophysiological data (Andersen et al., 1971), suggested that the major excitatory pathways of the hippocampal formation were all oriented perpendicular to the long axis of the structure and had a restricted septotemporal spread. Thus, like slices from a banana, the hippocampal formation comprised a number of similarly organized but connectionally isolated slices stacked along the long axis (see Amaral and Witter, 1989, for a more detailed discussion). If this view were correct, the hippocampal formation could be conceived of as containing a series of independent processing chips. But as has been emphasized in nearly all of the sections on intrinsic hippocampal connections, the idea that the flow of neural activity preferentially takes place within a lamella is no longer tenable. Many of the intrinsic hippocampal connections are as extensively distributed in the septotemporal axis as in the transverse axis. A view more consistent with the known neuroanatomy is that the hippocampal formation contains a series of threedimensional networks of connections. Moreover, the rules of connectivity appears to be different for each of the networks; the dentate-to-CA3 projection is organized in a lamellar fashion, the CA3-to-CA1 projection appears to be organized in a gradient fashion, whereas the CA1to-subiculum projection is organized in a columnar fashion. Interstingly, computer simulations as well as

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recent sophisticated electrophysiological data indicated that the hippocampal network, at least in part, functions in a lamellar fashion (Bernard and Wheal, 1994; Andersen et al., 2000). In particular, the recent finding that alternating lamellea appear to code for spatial and nonspatial features of a particular event is of particular interest in this respect (Hampson et al., 1999). It is clear that more research is needed to be able to settle the issue whether or not the complete hippocampal formation in vivo functions in a lamellar fashion. An interesting set of observations that may indicate the way to pursue this is that rats that store a particular event using their intact hippocampal system need that complete septotemporal structure in order to retrieve that event. However, animals with only restricted parts of the functional network can still store and retrieve the event (Moser and Moser, 1998). It is clear that a minimal volume of hippocampal tissue is needed and that this minimal volume is still much larger than a slice, but these findings do indicate that a minihippocampus constitutes a functional structure.

The Concept of the Trisynaptic Circuit and Serial/Parallel Information Processing A unique feature of the hippocampal intrinsic circuitry is the largely unidirectional organization of the projections that interconnect the various hippocampal regions. A popular notion is that these unidirectional projections also imply an exclusively serial or sequential flow of information from the entorhinal cortex to the dentate gyrus then to the CA3 field of the hippocampus, etc. But the data summarized in the body of this chapter indicate that the intrinsic hippocampal circuitry has both serial and parallel projections. The entorhinal cortex, in particular, contributes parallel projections to all fields of the hippocampal formation (Fig. 15). The same layer II entorhinal cells give rise to projections that terminate both in the dentate gyrus and in the CA3 field of the hippocampus. Thus, whatever information is conveyed by the entorhinal cortex would arrive both monosynaptically and disynaptically (through mossy fiber intermediaries) at the CA3 field. It is still not known, however, whether information from a single entorhinal cell reaches a particular CA3 cell both monosynaptically and disynaptically. This will be an important question to resolve in future studies. Likewise, parallel pathways are present in the projections from layer III cells to CA1 and the subiculum. In addition, the recently described direct inputs from the perirhinal and postrhinal cortices, particularly targeting the subiculum, add yet another level of complexity. Prominent associational connections in the dentate gyrus, in the hippocampus, in particular in CA3 and

the subiculum, and in some of the parahippocampal components (Fig. 16A) also provide the substrate for polysynaptic activation within hippocampal circuits. The functional implication of this more complex circuitry is that each hippocampal region is not entirely dependent on the preceding region for input and thus raises the prospect that each region may be acting independently as well as in concert with other hippocampal fields. Hippocampal neuroanatomy is thus entirely consistent with the electrophysiological finding, for example, that CA1 place fields are apparently normal even after the pharmacological inactivation of the dentate gyrus (Mizumori et al., 1989) or selective destruction of the CA3-to-CA1 projection (Brun et al., 2002). An additional issues that remains is why the overall output connections originating from both CA1 and the subiculum are also organized in parallel. An interesting approach might be to compare more specifically the shared and different projections of these two hippocampal domains as well as the observation that the two output pathways differ strikingly with respect to their amount of collateralization. The serial organization of the circuitry has also been taken to indicate that both CA1 and the subiculum may subserve actions in different time domains, such that ongoing processing is mediated by CA1–hippocampal– entorhinal networks, whereas longer-lasting processes that need a temporal linkage between events may depend on the subiculum (Hampson et al., 2000). Finally it is worth mentioning that strong associational networks are present in the parahippocampal regions and these indicate that complex associations might already occur at the level of these cortical regions. This added complexity is as yet not fully functionally understood, but has been interpreted such that increasing levels of complex associative processing may be dealt with in a hierarchical order from parahippocampal up to hippocampal processing.

Functional Implications of Septotemporal Topography of Perforant Path Projections Because the entorhinal cortex is the major relay for incoming sensory information, the septotemporal topography of its projections to other hippocampal fields largely determines the kinds of processing that will take place. On the one hand, the divergent nature of the perforant path projections makes it likely that information originating focally in the entorhinal cortex will be distributed fairly widely along the septotemporal axis. One focal group of entorhinal neurons might innervate as much as one-third of the septotemporal extent of the dentate gyrus and other innervated hippocampal fields. On the other hand, because of the topo-

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graphic organization of the perforant path projections, whatever information arrives in the lateral portions of the entorhinal cortex will tend to have greater influence over the septal end of the dentate gyrus, hippocampus, and subiculum, whereas inputs to the medial portions of the entorhinal cortex will tend to be relayed mainly to the temporal end of these fields. Because the lateral portion of the entorhinal cortex receives the major input from other neocortical areas, it is reasonable to assume that septal levels of the hippocampal formation will be more highly involved with the processing of exteroceptive sensory information. Because the medial portions of the entorhinal cortex are preferentially innervated by structures such as the amygdaloid complex, the temporal portion of the hippocampal formation may preferentially deal with interoceptive or emotional information. In recent years, more detailed behavioral comparative data have become available. In a recent review (Moser and Moser, 1998), it was concluded that functional differences along the lines developed above are truly present. Most convincing are the findings that the septal hippocampus is a necessary structure for spatial learning and memory (Moser et al., 1993), whereas the temporal hippocampus appears to be essential for normal fear-related behavior in rats (Kjelstrup et al., 2002). It is still unclear whether this is related to the differences in entorhinal inputs to septal and temporal hippocampus or to the fact that connections between the hippocampal formation and the amygdala are most strong in the temporal hippocampus or whether a combination of both is the critical factor. Also, differences between local hippocampal circuitry cannot be excluded. In this respect, the recent findings that the perirhinal cortex may be a critical structure for amygdala–cortical interactions, facilitating transfer through the entorhinal cortex into the hippocampal system, further complicates the picture (Kajiwara et al., 2003). Finally, most of the electrophysiological analyses of the rat hippocampal formation have been and still are conducted on the septal portion. Fortunately, in recent years more interest has been paid to the more temporal portions indicating that some of the characteristics of the network differ between the septal and temporal portions. Still it needs to be established whether these different response patterns result from anatomically different networks, functionally different inputs, or the interactions between those two variables.

The Layer II and Layer III Perforant Path Projections Are Organized Differently As noted in the previous section, the septal portion of the hippocampal formation receives input from a

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laterally situated band of entorhinal cortex. This band encompasses a rostrally situated portion of the lateral entorhinal area and a caudally situated portion of the medial entorhinal area (Fig. 15). The layer II cells in this band project throughout the full transverse extent of the dentate gyrus and CA3/CA2 fields of the hippocampus. Thus, all of the granule and pyramidal cells at the innervated levels are potentially privy to all of the information entering the laterally situated band of the entorhinal cortex. Since the perirhinal cortex projects preferentially to the LEA whereas the postrhinal cortex, the presubiculum, and retrosplenial cortex project mainly to the MEA, cells in the dentate gyrus and CA3/CA2 (which are innervated by both portions of the entorhinal cortex) might be viewed as a further stage of convergence. A comparable situation holds for the intermediate and medial bands of the entorhinal cortex projecting, respectively, to midseptotemporal and temporal levels of the dentate gyrus and CA3/CA2. The situation is quite different for the layer III projection to CA1 and the subiculum. The same septal portion of these fields receives input from the laterally situated band of entorhinal cortex. But the rostrally situated LEA projects to the border region of CA1 and the subiculum whereas the caudally situated MEA projects more proximally in CA1 and more distally in the subiculum. This characteristic feature of the layer III projection also stays constant along the septotemporal extent of CA1 and the subiculum. The implication of this organization is that CA1 and subicular cells potentially receive a more limited complement of entorhinalderived information via their direct inputs than via the disynaptic and trisynaptic inputs from CA3 and the dentate gyrus. Similar to the situation described for the layer II projection, it is likely that information carried by layer III neurons in the LEA is different from that conveyed by layer III neurons in the MEA. Most of the major inputs to the entorhinal cortex exhibit a clear laminar distribution, such that layer II receives inputs different from those innervating layer III. This may imply that the quality of information carried by the layer II projection may be substantially different from that carried by the layer III projection. However, as indicated above (see section “Entorhinal Cortex”), principal neurons of layers II and III, and of layer V as well, have apical dendrites reaching superficially into the molecular layer; these dendrites thus cross other layers and may receive inputs at that level as well. For example, olfactory inputs to layer I most likely target dendrites of both layer II and layer III cells. Similarly, presubicular fibers in layer III not only target dendrites of layer III cells but also synapse onto apical dendrites of layer V neurons. No clear conclusions regarding the information conveyed by

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layers II and III can be formulated yet. It is clear though that whereas the full transverse extents of the dentate gyrus and CA3/CA2 are involved in integrating or selecting sensory information derived from the layer II entorhinal projections, the different portions of CA1 and the subiculum use the output of CA3 to perform more selective computations involving the conjunction of a raw associative sensory map from the perirhinal and postrhinal cortices and a modified version of that same map, but now processed by the entorhinal cortex Naber et al., 2000b). These latter inputs have also been interpreted as providing the hippocampal system with a constant update of information, such that differences between predicted or reconstructed input and real input can be fed back through the layer II projections (Lörincz and Buzsaki, 2000). These notions, while speculative, do highlight the emerging principle that at least CA1 and the subiculum do have a transverse organization, distinctly different from that in the dentate gyrus and CA3/CA2, such that neurons in different proximodistal portions of these fields may be carrying out distinctly different tasks.

The Transverse Topography and Its Functional Implications Part of the intrinsic circuitry of the hippocampal formation appears to be organized such that cells located at a particular transverse position within a field are much more likely to be connected with cells located at a particular transverse position of the innervated field (Fig. 19). This allows for the possibility, therefore, that there is “channeling” of information processing through the various hippocampal fields (Amaral, 1991, 1993; Witter et al., 2000a, 2000b). This tendency for a transverse organization appears to become more apparent when moving from CA3 to CA1 to the subiculum. As described fully above, cells located proximally in CA3, near the dentate gyrus, tend to project to the most distal CA1 cells where the terminal plexus tends to be heavier in the stratum radiatum than in the stratum oriens, whereas projections from the distal portion of CA3 terminate mainly in the proximal portion of CA1 and most heavily in the stratum oriens and the deep portion of the stratum radiatum. The mid portion of CA3 fills in the spaces between these two projections. The CA1 projection to the subiculum demonstrates an even more striking transverse topography. The CA1 projection divides the transverse extent of the subiculum into roughly three equal parts. The proximal portion of CA1 projects to the distal third of the subiculum, the distal portion of CA1 projects just across the border into the proximal third of the subiculum, and the middle portion of CA1 projects to the midregion of

the subiculum (Fig. 19). This columnar organization of the CA1 to subiculum projection is all the more impressive in that the axons of single intracellularly labeled CA1 pyramidal cells demonstrate the same type of topographic organization and the same transverse spread of their terminal axonal arbors (Tamamaki and Nojyo, 1990). The notion of transverse topography of hippocampal connections is made all the more compelling when it is appreciated that both the subicular intrinsic network as well as its output are also organized in a columnar fashion. Both the distribution of dendrites of subicular pyramidal cells and the columnar organization of local axon collaterals appear to indicate the presence of a substrate for columnar modules along the transverse axis of the subiculum, although there is an integrating intrinsic network as well (Harris et al., 2001). The outputs are organized such that projections to different brain regions, or different parts of the same brain region, originate from the proximal, middle, and distal thirds of the subiculum (Swanson and Cowan, 1977; Witter and Groenewegen, 1990; Witter et al., 1990). Neurons in the proximal third of the subiculum project to the infralimbic and prelimbic cortices, the nucleus accumbens, and the lateral septal region. Projections from this portion of the ventral part of the subiculum also project to the ventromedial nucleus of the hypothalamus and to the amygdala. The mid transverse portion of the subiculum projects mainly to the midline thalamic nuclei and neurons in the distal portion of the subiculum project to the retrosplenial portion of the cingulate cortex and to the presubiculum. Although all portions of the subiculum project to the entorhinal cortex, the pattern of projections reciprocates the topography of the perforant path projections to the subiculum. Thus, the proximal portion of the subiculum projects to the lateral entorhinal area and more distal portions of the subiculum project to the medial entorhinal area. In addition, we described that even the relationship between the reciprocal connections between the entorhinal cortex and CA1/subiculum are in register with the CA1-to-subiculum connections, indicating the formation of very precisely organized parallelprocessing circuits (Naber et al., 2001a). A final piece of information that should be considered is the recently reported columnar organization of the entorhinal deepto-superficial connectivity (Stewart, 1999; Van Haeften et al., 2003), providing further support for this suggestion of parallel circuits.

Acknowledgments We thank our current and previous collaborators and colleagues for sharing data and figures to be used in this chapter. We are indebted to Mrs. I. I. Riphagen for conducting the literature searches

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FIGURE 19 Summary diagram illustrate the transverse organization of connections through the hippocampal formation. This figure highlights the possibility that information is segregated or “channeled” through the hippocampal formation and ultimately reaches different recipients of hippocampal output. See text for details.

that form the foundation of this chapter. We further thank Ms. S. van Oudenaren for secretarial assistance and Mr. D. de Jong for assisting us with the preparation of the figures. Original research reported in this chapter was funded by several grants from the Netherlands Organization for Scientific Research (NWO), an EC Grant QLG3-CT1999-00192, and Grant NS 16980.

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22 Cingulate Cortex and Disease Models BRENT A. VOGT and LESLIE VOGT Cingulum NeuroSciences Institute and Cingulate NeuroTherapeutics and Department of Neuroscience and Physiology SUNY Upstate Medical University, Syracuse, New York

NURI B. FARBER Department of Psychiatry, Washington University St Louis, Missouri

of the anterior and posterior cingulate regions that must be considered when using rats and the midcingulate region differs in both species. For example, there is no cingulate sulcus in rodent and the cingulate motor areas that form sulcal cortex in primates are not present; yet corticospinal connections characteristic of these latter areas arise from the most anterior cingulate areas in rat. In addition, there are no apparent rodent equivalents to areas 23 and 31 on the posterior cingulate gyral surface of primate brain. Thus, one of the main goals of the present chapter is to identify relationships of cingulate areas between the rat and a primate species and evaluate the extent to which similar areas share afferent and efferent connections. Cytoarchitectural, electrical stimulation, and functional imaging studies of human cingulate cortex confirm early suggestions that anterior cingulate cortex (ACC) is not uniform but composed of at least two divisions: perigenual ACC (pACC) and a caudal midcingulate cortex (MCC) (Vogt, 1993; Vogt et al., 2003). This review considers the regional definition of pACC and MCC and the cytology of each in the rat brain with immunohistochemical methods and relates them to previous atlases. Progress over the past decade in parcellating ACC has been made mainly in primate and this approach to cingulate cortex must now undergo a major revision in rodent brain. This theme is used to consider opioid architecture and cortical and thalamic connections. A detailed consideration of the

Human imaging and neuropathology research has implicated cingulate cortex in many neurological and psychiatric disorders. In some instances, as in Alzheimer’s disease, the earliest changes in brain metabolism occur in the posterior cingulate gyrus (Minoshima et al., 1997). In most studies of acute and chronic pain, a significant alteration in cingulate function has been identified (Casey, 1999; Derbyshire, 2000; Peyron et al., 2000). In the context of a growing recognition of the relevance of cingulate cortex to human disease, rat cingulate cortex is playing an important role for models of human diseases. This includes potential circuitry disturbances and the mechanisms of neurodegeneration in schizophrenia and Alzheimer’s disease (Farber et al., 1995b, 2002c; Olney et al., 1997), alterations in different models of chronic pain (Donahue et al., 2001), and changes associated with prenatal exposure to ethanol (Miller and Robertson, 1993). To assure accurate determination of which rodent models contribute to understanding the mechanisms of human disease, it is necessary to define relationships between each cytoarchitectural area in rodent with those in the primate medial cortex. If, for example, most changes associated with chronic pain occur in human midcingulate cortex and the rodent has no such region, the value of this species would be reduced. Fortunately, there is a midcingulate region in rodent and it is described in detail herein. There are, however, substantial differences in the structure and organization

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pathomorphological response to N-methyl-D-aspartate (NMDA) receptor antagonists including MK-801 is made in the context of the cytology and connections of retrosplenial cortex. As a prelude to considering general issues of modeling human disease, the pathomorphological response is related to changes in the schizophrenic and Alzheimer’s diseased brains and regional relations are considered between rat and monkey cingulate cortices. The final section raises explicit issues about the use of rat cingulate cortex to model human diseases.

REGIONAL ORGANIZATION The terms anterior and posterior cingulate cortex are used to designate general regions of cingulate cortex rather than particular cytoarchitectural areas, although there is an underlying morphological basis for this distinction. This difference is based on a granular layer IV in posterior cortex and the lack of a layer IV forming an agranular architecture in anterior cortex. This simple dichotomy, however, leads to confusion on three fronts. First, functional imaging studies in human medial cortex suggest there are many functional specializations in the cingulate gyrus that cannot be accommodated by a simple dichotomy in cingulate structure. Second, ACC is not a single entity but two based on structural and functional assessments as noted above (pACC and MCC) and they are present in rodent. Third, posterior cingulate cortex (PCC) in rat is not equivalent to the same general region in primates. In rodents it is composed entirely of retrosplenial areas 29 and 30 and there is no cingulate gyrus because a delimiting cingulate sulcus is not present. In primates, retrosplenial areas 29 and 30 are in the callosal sulcus on the ventral bank of the cingulate gyrus, while the exposed gyral surface is composed of

areas 23 and 31. Thus, the concept of a PCC is not interchangeable in rodent and primate species. The duality of rat ACC was first proposed to account for structural, connection, and limited functional observations (Vogt, 1993). The pACC receives the most prominent amygdala input (Sripanidkulchai et al., 1984) and the MCC projects strongly to the pontine nuclei, while the pACC does not (Wiesendanger and Wiesendanger, 1982). The MCC transiently expresses oxytocin receptors and neurotrophin-3 (Triboll et al., 1989; Friedman et al., 1991) and adults have an opioid architecture that differs from pACC (discussed below). Finally, extensive human imaging studies have defined a border between pACC and MCC (Bush et al., 2000). These fundamental differences in connections, transmitter systems, and functions require redefinition of rat cingulate cytology in a manner compatible with our evolving understanding of primate medial cortex including that in human brain. The three regions of rat cingulate cortex are shown in Fig. 1 in a modification of our original rat map (Vogt and Peters, 1981). This revision includes localization of each region (pACC, MCC, and retrosplenial cortex, RSC), shows the distribution of each area and subarea, redefines dysgranular area 29d as area 30 to draw a comparison with an area of the same cortical moiety in monkey, and provides an approximation of the anterior/posterior coordinates for each major region and area in relation to the Paxinos and Watson atlas (1986). The pACC comprises areas 25, 32, and 24a/b. Realizing that the MCC is a posterior division of area 24, it is designated area 24′ and includes areas 24a′ and 24b′. A similar strategy has been applied to the divisions of “Cg” by Zilles and Wree (1995). Although PCC is equivalent to RSC in rat, confusion is generated by the use of PCC for rat cortex because primates have a massive expanse of posterior cingulate areas 23 and 31

FIGURE 1 Overview of the rat medial cortex including the cingulate areas within the heavy line and adjacent areas 10, AGm, and 18b. The three regional designations identify approximate anterior–posterior levels (A/P) of the pACC, MCC, and RSC regions. The pACC includes areas 25, 32, and 24; the MCC is comprised of area 24’; please note figure and the RSC is comprised of areas 29 and 30. The A/P coordinates are approximate levels from Paxinos and Watson (1986). ac, anterior commissure; Post, postsubiculum (area 48).

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that are not present in rodents. Since RSC is equivalent to PCC in rat but not primate PCC, the regional designation of RSC is used here for the posterior region in rat brain. Finally, Chapter 23 provides a comparison of the nomenclatures used for each cingulate and retrosplenial area in rat.

IS “INFRA” LIMBIC AREA IL VENTRAL TO LIMBIC CORTEX? Although the nomenclature of Rose (1927) has been used frequently for studies of rodent cingulate cortex, it raises important questions about the organization of this region. It suggests that an “infra” limbic area, which is similar to Brodmann’s area 25, lies below limbic cortex, rather than being limbic cortex itself. Although there have also been proposals that all cingulate cortex is “para” limbic rather than limbic (Mesulam, 1995), to characterize part of the ACC as “infra” or “para” requires a definition of what constitutes limbic cortex. For the designation of paralimbic, a cortex needs to abut the indusium griseum; a structure that is less than 0.01% the size of the human cingulate gyrus. Indeed, referring to cingulate cortex as paralimbic says nothing about what the cingulate cortex is or what it does. We prefer a functional definition for a limbic area that includes any cortex with a specific role in regulating autonomic responses, dense projections to the hypothalamus, and subserving emotion (positive or negative internal states and associated memories). To the extent the posterior hippocampus and indusium griseum are involved in general short-term memory formation including emotional memories, they do not have a specific role in emotion and are not limbic by this definition. This functional definition identifies area 25 as a major limbic area. It has been termed visceromotor cortex and has projections to the nucleus of the solitary tract and dorsal motor nucleus of the vagus (Neafsey et al., 1993) and may mediate autonomic activity through the amygdala (Fisk and Wyss, 2000). Direct modulation of autonomic activity assures that area 25 is limbic cortex rather than infralimbic. If one accepts the functional definition for a limbic structure, it becomes clear the posterior hippocampal, posterior cingulate, and retrosplenial cortices are not limbic because they are not known to have a specific role in regulating emotion and associated autonomic responses. The confusion over these areas as being limbic derives from a century-old notion that cortices with a “simple” laminar architecture are limbic without consideration of their roles in brain function. An irony of this view is that even on this early anatomical criterion, area 25 is more characteristic of a limbic cortex

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than area 24 because area 25 has less differentiated layers than does area 24. To demonstrate this point, we begin the cytological analysis of cingulate cortex below with a side-by-side comparison of these two areas. Thus, area 25 is not “infra” (below) or “para” (adjacent) to limbic cortex but, rather, it is limbic in more direct ways than area 24. Furthermore, the evidence from human studies is very compelling that area 25 has a direct role in regulating autonomic functions and generating emotional states (Vogt et al., 2003). Thus, area 25 is limbic cortex and use of the IL concept leads to misuse of the term limbic.

CYTOLOGY OF LIMBIC AREA 25 Area 25 is one of the least differentiated cingulate areas in rat; the other being area 29a in the RSC. An early stage of differentiation is usually associated with large pyramidal somata, high neuron densities in layers V and II, and poor laminar differentiation; i.e., laminar subdivisions are difficult to detect. Figure 2 shows examples of all cingulate areas with an antibody to neuron-specific nuclear binding protein (NeuN) and at four levels there are silver-stained sections from another animal to show the distribution of axons in similar areas. Since the NeuN antibody does not label glial or vascular elements, it provides a clear picture of laminar architecture. Levels A and B in this figure can be used to evaluate areas 25 and 24a, respectively, and there are higher magnification photographs for comparison of the layers. Area 25 has much larger neuronal somata throughout all layers than does area 24a. Area 25 has a very broad layer II and a poorly differentiated layer III, layer V is uniform and relatively thick, and layer VI is thin and hard to detect. In contrast, area 24a is generally thicker and has smaller neuronal somata than area 25, it has a thinner layer II, broader layers III, V, and VI, and many more parvalbumin-expressing neurons than does area 25 (i.e., areas Cg and IL; Paxinos et al., 1999). Also, the superficial part of layer V has larger neurons and this further enhances laminar differentiation. Thus, on anatomical grounds, area 25 is the least differentiated area of the ACC.

MODIFIED BRODMANN NOMENCLATURE Most functional studies of human cortex employ the Brodmann (1909) nomenclature to designate sites of activity and none use the Rose nomenclature that is so frequently used in rodent studies. Therefore, nomenclatures often used in rodents have not been directly

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FIGURE 2 Structural organization of cingulate cortex at different rostrocaudal levels. A–E each have a NeuN-stained, coronal section to show the laminar distribution of neurons unconfounded by costaining of glial and vascular elements and B–E have an additional silver pyridinestained section to show the distribution of large caliber axons. Of particular note are the following: area 25 has a very poorly differentiated laminar architecture when compared to area 24a (higher magnification photographs at bottom), area 24 is not uniform and the area 24/24′ dichotomy is supported, and the internal plexiform layer of area 29c is pronounced (D, silver stain).

applied to human medial cortex. The resulting paradox is that findings in rat cortex cannot be directly related to the structure and functions of primate cortex and a disconnect has developed between research in rat brain and this makes modeling human diseases difficult in terms of an extensive and growing body of human research in normal and pathological states. Taking from our own experience, how does one relate the distribution of neurodegeneration in rat cortical areas following MK-801 exposure to the pattern of cell death in human Alzheimer’s disease cases, if the former is to be considered a model of the latter? If there is no similarity among areas in rat and human brains, it will be difficult to model human diseases in the rat. Indeed, although no area in rat can be truly equivalent to that in human, one can characterize areas that share structural features. The hippocampus, for example, is often analyzed in transgenic mouse models of Alzheimer’s disease and it is suggested that changes in both species are related. If this strategy can

be used in the hippocampus, why should it not be applied to medial cortex? Consider area 29, for example. Area 29 in both species lies between allocortical and isocortical regions and contains poorly differentiated granular layers. This does not mean, however, that they are the same. Area 29 in rat receives direct inputs from primary and secondary visual cortices (i.e., areas 17 and 18, respectively) and this is not the case in monkey (Vogt and Miller, 1983; Vogt and Pandya, 1987). The rat granular layer has both fusiform and extraverted pyramids, while the monkey has neither of these types of neurons (Vogt and Peters, 1981; Vogt, 1976). Thus, rat area 29 is similar in ways to monkey area 29 and there may be instances when neurodegeneration in rat area 29 may be a model of cell death in some human diseases. This does not mean, however, they are equivalent. When Brodmann’s scheme was originally modified for rat cingulate cortex (Vogt and Peters, 1981), it was done in relation to work in monkey (Vogt et al., 1987) and it was eventually related directly to human cortex

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(Vogt et al., 1995). Each cingulate area in rodent can be evaluated for counterparts in primate cortex determined from relative cortical differentiation, phenotypic expression of particular peptides, and common connections and functions. Although this undertaking is an active area of research, an example of this type of undertaking is provided below under Section, “Comparison of Medial Cortex in Rat and Monkey.”

CYTOLOGY OF THE PERIGENUAL ANTERIOR AND MIDCINGULATE REGIONS One of the most important current considerations is the differentiation of ACC into pACC and MCC regions at the cytological, connection, and transmitter system levels of organization. Area 24′ designates the posterior division of area 24 or MCC and this emphasizes that area 24′ is an agranular cortex and not simply a narrow “transitional” region with a mixture of both anterior and posterior cortical features. Differentiation of areas 24 and 24′ is shown in Fig. 2 with NeuN immunoreactivity for neurons and differentiated, silver pyridine sections for large axons. The dorsal part of pACC is

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shown in Fig. 2B, and MCC is shown in Fig. 2C, for both stains, respectively. These macrophotographs show that layer II in area 24 is thick relative to that in area 24′ and that the thickness from pia to white matter is greater in area 24. The detailed differences in laminar architecture between areas 24 and 24′ are shown in Fig. 3 for the “b” subdivisions. Although there are occasional large neurons in layer II of area 24′, they are generally smaller and more numerous than in area 24. Comparison of both divisions to area 29c in the same figure suggests there is a transition throughout cingulate cortex that culminates in small and densely packed neurons in the external layers. In addition to this trend that is most pronounced in layer II, layer III is thicker in area 24b′ than in area 24 and layer Va in area 24 is more neuron dense than in area 24′. The silver-stained tissue is informative at low magnification. Although an axonal plexus is in layer I of both divisions of area 24, it is thicker and more neuron dense in area 24′ than in area 24. Also, the relative differentiation of the a/b subdivisions is more pronounced in area 24′. The layer I plexus in area 24a′ extends into layers II and III, while in area 24b′ the plexus clearing is more pronounced in layer II and the top of layer III. Finally, there is a substantially greater

FIGURE 3 Differential architectures of areas 24b and 24b′ and their comparison to area 29c. In addition to the generally higher density of neurons in area 24 than in area 24′, the neurons in layer II of area 24b are larger than those in area 24b′. Although some size differences may also exist in layers III and V, they will require a quantitative analysis.

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density of large axons in layers V and VI of area 24′ than is the case for area 24. Axonal staining, therefore, confirms cytoarchitectural observations of a division between pACC and MCC. The full range of GABAergic interneurons recognized in other cortical areas has been observed in cingulate cortex (Vogt and Peters, 1981), there are distinct subsets of calcium-binding, immunoreactive interneurons (Hof et al., 1993; Paxinos et al., 1999), and Chapter 23 includes a detailed analysis of GABAergic and other transmitters in cingulate cortex. Since the contribution of each class of interneuron to cingulate functions is not known, they are not reviewed here. Instead, calretininimmunoreactive neurons are considered in each cingulate region to evaluate regional differentiation. There are generally more calretinin+ neurons in areas 24 and 24′ than in RSC and these neurons are in all layers, though not at similar densities. Calretinin is also expressed by thalamic afferents to cingulate cortex as shown in monkey (Vogt et al., 1997). These latter axons appear to be mainly in layer Ia (i.e., the outermost onethird of this layer) of all cingulate areas in rat (Paxinos et al., 1999). Two main classes occur: long bipolar cells with slender dendritic trees extending across layers and multipolar cells with spherical dendritic trees that may be either intra- or interlaminar in their distribution. Although there tend to be many bipolar neurons in layer II of areas 24 and 24′, they flank layer II in area 29. Multipolar neurons in area 29c often have dendrites that branch throughout layer I, somata at the surface of layer II, and descending axons with extensive terminal arbors in layer Va. In contrast, the multipolar

neurons in area 24 are located in deep layer I and have long and descending dendrites that reach throughout layers II and III.

CYTOLOGY OF RETROSPLENIAL CORTEX Retrosplenial cortex forms approximately the posterior one-third of the medial surface. Beginning with the most ventral area 29a, each subdivision contributes to a progressive elaboration that focuses mainly on the granular layers, but also involves differentiation of layers V and VI. Area 29a abuts the postsubiculum (area 48) as shown in Fig. 2. The transition to area 48 is characterized by smaller somata in area 29a. The latter area has an external pyramidal layer composed of a very thin layer II and a layer III/IV (Fig. 4), whereas layer IV is hardly perceptible. Nissl stains with their costaining of glial elements originally suggested this is a homogeneous granular layer (Vogt and Peters, 1981). The deep pyramidal layer is quite homogeneous, although a thin layer VI of small neurons can be detected. Area 29b, in contrast, has a very dense and thick layer II of usually three to five neuronal somata in depth, a dispersed and granular layer III, and a thin layer IV that is neuron sparse. It might be argued a layer IV is not present; however, areas 29b and 29c receive a thin layer of thalamic terminals in this layer and a silver-stained axonal plexus can also be detected therein (Fig. 2). Thus, this distinction is based on more than cell structure considerations. Finally, layer V has

FIGURE 4 Laminar architectures in the four divisions of area 29 shown with NeuN immunohistochemistry. The progressive elaboration of laminar differentiation and cortical thickness is apparent beginning with the least differentiated area 29a. Although there is some differentiation of the external layers into layers II and III/IV, the neurons in layer II are larger and less dense in area 29a than they are in area 29c. In addition to the highest level of external pyramidal layer differentiation in area 29c, layer Vb has the largest neurons in layer V and there appear to be two divisions of layer VI. Finally, dysgranular area 30 has larger layer II and III pyramidal neurons and layer IV has clumps of small neurons. Notice in Fig. 10 that the LD projects heavily to layer IV in area 30, while the AV projects heavily to layer IV in area 29c.

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large typical pyramids and there is a slight size differential with deeper neurons being larger than superficial ones. Layer VI, though thin, is quite clear. Area 29c cytology is shown in detail in Figs. 3 and 4. It has the most differentiated granular layers and, overall, the neurons tend to be smaller in diameter than those in areas 29a and 29b. The predominant neuron types are the fusiform and star pyramids (Vogt and Peters, 1981). Examples of these small neurons are also shown later in Fig. 8 from a preparation in which they were backfilled from contralateral cortex. The apical dendrites of these neurons form bundles as do their apical tufts in layer Ia and they are sensitive to aging processes and form fewer branches than in young adult animals. One consequence of this aging process could be impaired visuospatial learning that is mediated in part by area 29 (van Groen et al., 1993). Other examples of the apical tuft distributions in layer Ia for fusiform and star pyramids are shown later in Fig. 10 in the context of the pathomorphological response to NMDA receptor antagonists. In the deeper layers of area 29c, differentiation of layer V is very pronounced (Fig. 3) where layer Va is formed by medium-sized pyramidal neurons more diffusely packed than those in layer Vb. This layer V organization is characteristic of “motor” cortices where the large layer V corticospinal projection neurons are mainly in deep layer V and may suggest the important role of this region in behavioral performance in addition to acquisition. Finally, Layer VI is quite thick and supports a heavy interaction of area 29c with the anterior thalamic nuclei. Dorsal to area 29c lies Brodmann’s area 29d, which Rose (1927) referred to as agranular and is often termed “RSA” (see also Chapter 23). Although an equivalent area in primate is termed area 30, it is not agranular but rather dysgranular (Vogt et al., 2001). Since the rat and monkey share this dysgranular cortex, the rat area 29d is now termed area 30 for consistency between species and to recognize its fundamentally different architecture from that of the granular parts of area 29. In the rat, the posterior cingulate region undergoes almost continual transition as shown in Figs. 2D/E and 4. Layers II–III are composed of larger neurons that are progressively more dispersed in the dorsal cortex. The silver-stained axons show a disappearance of the layer IV plexus and a progressive widening of these layers and reduction in the overall density of large diameter axons. At higher magnification (Fig. 4), all of these features are apparent and it can be seen there is a small and dysgranular layer IV. Thus, area 30 is not truly agranular as is area 24. Indeed, in rat, even motor cortex can have a layer IV and is not truly agranular (Donoghue and Wise, 1982). It is for these reasons that

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early anatomists may have overlooked the dysgranular nature of area 30. Finally, the presence of layer IV is confirmed by a very dense projection of the laterodorsal nucleus to layer IV of this area as discussed below.

OPIOID ARCHITECTURE: REGIONAL DIFFERENCES AND NEURONAL EXPRESSION PATTERNS It has long been known that rat anteromedial cortex has the highest density of opioid receptors and enkephalin expressing interneurons (Sar et al., 1978). The highest binding in human brain is in pACC with lesser amounts in MCC, and the least in PCC including RSC (Vogt et al., 1995). Experimental studies in rat have provided much information about the organization of cortical opioid systems including its regional differentiation in cingulate cortex. In the context of the motor functions of ACC discussed below, including those associated with pain processing, it is important to consider the distribution of opioid receptors and how their activation might modulate motor functions. Expression of the μ-opioid receptor agonist Tyr–DAla–Gly–MePhe–Gly–ol (DAMGO) has been used for receptor binding and DAMGO-stimulated GTPγS stimulation with autoradiography to assess the overall composition of opioid circuits and provides insight into the differentiation of pACC and MCC (L. Vogt et al., 2001). This study showed the highest binding in area 32, intermediate amounts in areas 24 and 24′, and the least in area 29 (Fig. 5), thus mimicking the regional differentiation observed in human brain. While area 24′ shared a similar laminar pattern of μ-opioid binding with area 24 (highest in layer V and moderate in layers I and VI), area 24′ shared a similar pattern of GTPγS stimulation with area 29 (moderate to low levels in layers I and layers V and VI). A correlation analysis of DAMGO binding and DAMGO-stimulated GTPγS activity confirmed that area 24′ has an opioid architecture different from that of either areas 24 or 29. This confirms the regional differentiation of rodent cingulate cortex into pACC, MCC, and PCC. Early studies reported a layer I concentration of μ-opioid receptors in ACC; however, experimental studies were needed to identify those components of the cortical neuropil that expressed the receptors in their dendrites and axonal terminals. Undercut lesions that remove all afferent axons, and therefore presynaptic receptors, show that about 30% of layer II–VI binding is lost in area 24 and 50% is lost in layers I and V in area 29 (LJ Vogt et al., 2001; Vogt et al., 1995). The remaining binding following undercut lesions is expressed by the soma/dendritic membranes and

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the midline and intralaminar thalamic nuclei and some nuclei that project to the RSC express opioid receptors (LJ Vogt et al., 1992) and thalamic lesions reduce binding throughout the cingulate gyrus including a 24% reduction in layer Ia of area 29c (Vogt et al., 1995). Finally, nociceptive neurons in rat are in highest density in deep layers and most are pyramidal neurons with apical dendrites that distribute apical tufts in layer I (Yamamura et al., 1996). A nociceptive region is located in areas 32 and 24b in rat (Hsu et al., 2000) and rabbit (Sikes and Vogt, 1992) and this region is driven by electrical stimulation of the medial thalamus (Hsu and Shyu, 1997). Thus, there are circuits in cingulate cortex for regulating motor behaviors as noted below and many of the cortical motor projection systems arise in layer V where there are the highest levels of opioid receptor binding and the most nociceptive neurons.

AREA 24b: MOVEMENT, VISION, AND PAIN BEHAVIORS

FIGURE 5 Distribution of G-opioid receptor binding with DAMGO autoradiography and activation of G-proteins with DAMGO and their assay with autoradiography for GTPγS. The highest binding and stimulation is in area 32 with progressively less in the rostrocaudal extent of the cingulate cortex. This is similar to the topographic distribution of opioid receptor binding in primates. It is important that, although binding in area 24b higher than that in area 24b′, the relative level of G-protein stimulation in area 24b′ is higher than that in area 24b, suggesting that the opioid architecture and function in these two cingulate divisions is quite different. Although area 29c does not appear to have any direct role in pain processing, it does have μ-opioid receptors and G-protein stimulation and these regulate, among other systems, thalamocortical inputs because thalamic lesions greatly reduce ligand binding to these receptors (Vogt et al., 1995b).

possibly some glia. The question remains, however, which afferent axonal systems express μ-opioid receptors. There are at least two such sources. First, the locus coeruleus synthesizes μ-receptors and they are transported to presynaptic terminals in cingulate cortex. This was shown using the neurotoxin saporin conjugated to dopamine β-hydroxylase to kill neurons in the locus coeruleus followed by autoradiographic assay of binding and G-protein stimulation. This produced a 31% decrease in DAMGO binding in layer I of area 24 but not in areas 24′ and 29 (LJ Vogt et al., 2001). Second,

Area 24b lays ventral to the medial agranular field (AGm) of Donoghue and Wise (1982) or Fr2/M2 in the rat (Zilles, 1985; Paxinos and Watson, 1986). Miller (1987) found corticospinal neurons mainly in areas 24b and 32 and these may be the rodent precursors of the cingulate motor areas in primates described by Morecraft and Van Hoesen (1992) and Dum and Strick (1993). The AGm has strong connections with other motor areas, visual cortex, and retrosplenial area 29d (Reep et al., 1990) and there have been reports of low-threshold, contralateral head turning produced by electrical stimulation of area 24/24′ (Sinnamon and Galer, 1984). Unilateral lesions of the AGm and area 24b impair approach to contralateral visual cues and transient sensory neglect (Vargo et al., 1988). In light of the major and reciprocal visual inputs to area 24b′ (Vogt and Miller, 1983; Miller and Vogt, 1984; Paperna and Malach, 1991) as well as posterior AGm (Reep et al., 1990), it is quite likely that area 24b is part of a visuomotor integration system. Since lesions of AGm and area 24b′ alter hot plate reflexes in a condition termed “nocifensive apraxia” by Pastoriza et al. (1996), this rostral shoulder cortex may also be employed in generating avoidance behaviors in the context of nociceptive processing. Electrical stimulation of area 32 inhibits cardiovascular reactions in rats (Maskati and Zbrozyna, 1989) and Vaccarino and Melzack (1989) were able to produce analgesia (i.e., reduced responses to tonic and phasic noxious stimulation) by injecting lidocaine into the cingulum bundle. Finally, Donahue et al. (2001) selectively blocked responses to inflammatory pain versus neuropathic pain with lesions in this region. Since area 24b has

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nociceptive neurons (Sikes and Vogt, 1992; Yamamura et al., 1996), it appears this area is involved in visual nocifensive processing as first suggested by Pastoriza et al. (1996). This also places the role of rat ACC (specifically area 24b) in pain processing in the domain of modulating motor functions. Since it has been suggested in monkey (Shima and Tanji, 1997) and human (Bush et al., 2002) that ACC is critical to changing the reward properties of behavior, including those associated with pain processing, area 24b appears to be pivotal to establishing the reward properties of a range of visually guided behaviors and may be critical to the prediction and avoidance of painful outcomes.

CORTICAL CONNECTIONS OF RETROSPLENIAL CORTEX AND ROLE IN VISUOSPATIAL FUNCTION The distribution of retrogradely labeled neurons following a horseradish peroxidase (HRP) injection into areas 29c/30 is presented in Fig. 6. This case provided

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the first compelling demonstration of direct visual and retrosplenial cortex interactions and those with parahippocampal cortices (Miller and Vogt, 1983) and it now serves as the first to show differential inputs to these dorsal retrosplenial areas from the pACC and the MCC. There are three classes of cortical input to this region. First, direct visual sensory input arrives from the deep layers of areas 18a and 18b, but also from area 17 as well as from the auditory cortex (Paperna and Malach, 1991). Second, the subiculum, deep layers of area 48, also termed the postsubiculum (Ps), and entorhinal cortex project to this region. Third, area 25 of pACC and MCC areas 24a′ and 24b′ project massively to areas 29c/30. It is surprising to see how little input to this region arises from areas 24a/b and how much arises from areas 24a′/b′. The HRP findings have been validated with injections of biotinylated dextran amine (BDA) into the same region and their cellular origin is shown in Fig. 7. In particular there is the heavy and reciprocal connection between areas 24′ and RSC and much weaker input from area 24 (Figs. 7B and 7C). There also is massive

FIGURE 6 Distribution of neurons retrogradely labeled with HRP following an injection into areas 29c/30 (each dot represents about three labeled neurons). In addition to the three classes of cortical inputs (cingulate, parahippocampal, and sensory), notice that area 24′ has a substantially greater projection than does area 24. There is a high level of input from the basal forebrain (DBB, diagonal band of Broca), anterior thalamic nuclei (AV, anteroventral; AD, anterodorsal; AM, anteromedial), and laterodorsal (LD) and superior centrolateral (Csl) nuclei. The lateral hypothalamus (LH), ventral tegmental area (VTA), and raphe nuclei (DR, rorsal raphe; MR, median reaphe) also project prominently to this region. AC, anterior commisure; AGm, medial agranular motor cortex; VB, ventrobasal nucleus; MD, mediodorsal nucleus; MGB, medial geniculate nucleus; PAG, periaqueductal gray; Ps, postsubiculum; Sub, subiculum.

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intraretrosplenial inputs from areas 29b and 29a (Fig 7D), and a major input from parahippocmpal cortex that includes the subiculum and the entorhinal cortex (Figs. 7D and 7E). Many observations support a significant role of area 29 in visuospatial functions. There is massive visual input to areas 29 and 30 and major projections from the postsubiculum which is involved in coding for head position in space (Taube et al., 1990). Indeed, RSC and visual area 18b both receive major inputs from the anteromedial nucleus of the thalamus (Fig. 8) (Rieck and Carey, 1985). This shared input likely assures that both areas have coordinated visuospatial processing. In addition, there are massive projections from the anterodorsal thalamic nucleus to area 29, the former of which is involved in coding spatial orientation according to background cues and radial-maze learning (Zugaro et al., 2001; Byatt and Dalrymple-Alford, 1996). Finally, many studies of visually guided behaviors support the visuospatial hypothesis of RSC function including those with the Morris water maze (Sutherland et al., 1988; Sutherland and Hoesing, 1993; Harker and Whishaw, 2002) and those showing that the late stages of acquisition of a visuospatial conditional discrimination are dependent on the RSC (Bussey et al., 1997). Thus, parahippocampal, anterior thalamic, and visual cortical inputs to the RSC provide the visual cues and orientation inputs necessary for neurons in this region to code for position of the body in space.

THALAMIC AFFERENTS Regional Differentiation The dense innervation of the RSC by the anterior thalamic nuclei is well documented and the heavy labeling shown in Fig. 6 confirms the heavy inputs from the anteromedial (AM), anteroventral (AV), and anterodorsal nuclei (AD) as well as the laterodorsal nucleus (LD) to this region. In the context of the regional divisions of cingulate cortex, it is crucial that these inputs differentiate each region. Horikawa et al. (1988) injected retrograde tracers into the “a” and “b” divisions of anterior and posterior area 24 (i.e., areas 24 and 24′) and their summary diagram is particularly instructive as to the differential projections of the anterior and laterodorsal thalamic nuclei. Area 24 receives primarily AM input, while area 24′ receives mainly AM and AD afferents. Area 29 receives both of these plus a large input from AV and LD. Further support for the pACC/MCC distinction comes from Shibata (1993) who shows that area 24 receives more input from the interanteromedial nucleus, while area 24′ has a higher density of input from AM proper, although AM input is also shown to be extensive throughout the cingulate cortex. Finally, the midline and intralaminar thalamic nuclei differentiate between the pACC and the MCC areas. The reuniens nucleus projects most intensely to areas 25 and 24 and less so to area 24′

FIGURE 7 An injection of biotinylated dextran amine into areas 29c/30 (white arrows in A indicate cannula track) and retrogradely labeled neurons throughout cingulate and parahippocampal cortices. Of particular note are the level of input from area 24′ that is higher than that from area 24, the dense innervation from area 29a and parahippocampal areas 48 and subiculum (D), and the less pronounced input from the entorhinal cortex (E).

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FIGURE 8 Laminar distribution of terminals from AD, AV, and LD in areas 29c and 30 following injections of Phaseolus vulgaris leucoagglutinin into each nucleus. The differential projections to layers in each area confirm key cytoarchitectural observations such as the presence of a layer IV in both areas based on AV projections to area 29c and LD projections to area 30. The clumps of terminals in layer Ia of area 29c provide a critical rationale for hypotheses that relate nonglutamate receptors with thalamic afferents. The arrows show a magnification of part of area 29c after labeling a single AV axon with fluororuby. Its arborization may be juxtaposed onto the horizontally dispersed arbors of dendrites in the same layer that are here retrogradely labeled with fluorogold in layer II of area 29c following a contralateral injection. The bundles of apical dendrites and their apical arborization throughout layer Ia may be associated with similarly shaped clumps of transmitter receptors in layer Ia (Fig. 9) and suggest that thalamic afferents are under the control of a number of heteroreceptor systems. Modified from van Groen et al. (1993).

(Herkenham, 1976), while the parafascicular nucleus projects to the deep layers of pACC (Marini et al., 1996). Thus, the three-region model demonstrated above with immunohistochemical, neurotransmitter receptor binding, and connection methods is supported by thalamic afferents.

Projections to Area 29 Reports by van Groen et al. (1993) and van Groen and Wyss (1995) provide important new details of the thalamoretrosplenial projection system and serve as the basis for interpreting the distribution of many neurotransmitter receptors in this region. Figure 8 docu-

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ments the distributions of axons and dendrites from the former publication. The figure shows that each nucleus has a different area and laminar projection pattern with the AD and AV nuclei projecting mainly to granular area 29c and LD projecting to both area 29c and dysgranular area 30. The AD projects diffusely throughout layer I and the projection is more dense, although still diffuse, throughout layers II and III. In contrast, the AV projects mainly to layer Ia in coneshaped clusters, an example of which is shown with individual axonal labeling with fluororuby. Other classes of axons terminate diffusely throughout the remainder of layer I and in a tight band in layer IV. Finally, LD projects mainly to layer I in areas 29c and 30, lightly to layer IV in area 29c, and densely to layer IV in area 30. The fusiform pyramids of layer II have primary apical dendrites that form bundles in layers Ib and Ic and they splay out in layer Ia to form tangentially dispersed aggregates (Fig. 8, dendritic bundles). It appears that these bundles of apical tuft dendrites are targeted by axons from the AV as also shown in this figure. Indeed, the second major source of acetylcholinesterase activity in RSC, after that originating from the diagonal band of Broca (Fig. 6), is associated with anterior thalamic afferents. Figure 9 (AChE) shows high expression of AChE in layer Ia where AV axons terminate and greater than 50% loss of this activity following thalamic lesions (right side of Fig. 9) (Vogt, 1984).

Axon Terminal Morphology and Multiple Heteroreceptor Regulation Projections of the anterior thalamic nuclei to RSC are glutamatergic (Gonzalo-Ruiz et al., 1997) and they form large axon terminals and asymmetric synapses with dendrites in layer Ia of area 29c that are consistent with a major excitatory pathway (Vogt et al., 1981). The high level of AChE activity in RSC is due to the fact that these terminals are postsynaptic to cholinerigic inputs rather than to acetylcholine being a transmitter in this system. Transmitter receptors localized to these thalamocortical axon terminals that are not selective for glutamate and, therefore, are termed heteroreceptors. The laminar distribution of ligand binding autoradiography, layer Ia clumping, and effects of unilateral lesions in rat have suggested there are many heteroreceptors expressed by these glutamatergic terminals. Although the functions of these multiple heteroreceptor systems are not known, it is clear they can modify thalamocortical processing even before postsynaptic activation at the dendritic level (Vogt et al., 1995b). Furthermore, this complex presynaptic regulation does not appear to occur in area 29 of monkey (Vogt et al.,

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lesions, suggesting the muscarinic M1 receptors are expressed by intracortical neurons rather than extrinsic afferent axons (Vogt, 1984, for more details). The second binding pattern has binding peaks in layers Ia and III–IV similar to the distribution of AV axons (Fig. 8) and this bilaminar binding can be abolished with thalamic lesions as for oxotremorine-M and cyanopindolol binding (Fig. 9). The association of these latter populations of receptors with thalamic inputs is confirmed in the same cases with the loss of acetylcholinesterase activity and similar losses occur in DAMGO binding. Thus, M2, β-adrenoceptor, and μ-opioid receptors are expressed by these thalamocortical terminals, although they may not all be expressed on the same axon terminal. Nonetheless, these multiple heteroreceptors provide for significant presynaptic regulation and might be relevant to interventions that engage gluatamatergic systems.

NMDA RECEPTOR ANTAGONIST-INDUCED NEUROTOXICITY IN RETROSPLENIAL CORTEX FIGURE 9 Superficial layer distribution of neurons (top), acetylcholinesterase (AChE), pirenzepine binding (PZ) autoradiography for M1 binding, oxotremorine-M binding in the presence of unlabeled PZ (OXO-M) autoradiography for M2 binding, and cyanopindolol binding in the presence of unlabeled isoproterenol (CYP/IPT) autoradiography for β-adrenoceptor binding in area 29c. Each layer is labeled in the top section for both hemispheres and there is a thalamic ablation on the right side. Note high activity of AChE in layers Ia and IV and high binding of OXO-M and CYP/IPT in the same layers. The latter ligands form clumps in layer Ia just like those of AV axons in Fig. 8 and ablation of this input to area 29c abolished much AChE activity and binding of OXO-M andCYP/IPT but not PZ which is likely expressed by intrinsic cortical neurons.

1997) and could be crucial to attempts to model primate diseases. The unique clustering of AV thalamic input to area 29c, bilaminar projections to layers Ia and IV, and matched dendritic/axonal aggregates (Fig. 8) provide valuable markers for assessing receptor localization in experimental autoradiographic ligand binding studies. The bundles of axons from the thalamus have been shown to express M2 binding with oxotremorine-M in the presence of unlabeled pirenzepine, β-adrenoceptor binding with cyanopindolol in the presence of isoproterenol, and μ-opioid binding with DAMGO as discussed above (Vogt et al., 1995b). Two binding patterns are shown for area 29c in Fig. 9. In one pattern for pirenzepine, there is a modest peak in binding in layers II and III that is not abolished with thalamic

Extensive research focusing on the amino acid glutamate (Glu) has documented the central role played by this compound in both the normal and the abnormal functioning of the CNS. Glu is the main excitatory neurotransmitter in the CNS and is released at up to half of the synapses in the brain. The Glu receptor family is composed of two major subfamilies (ionotropic and metabotropic), and within these subfamilies there are many additional subdivisions. The ionotropic receptors are further subdivided into three major categories, each being named for an agonist molecule to which it is preferentially sensitive (NMDA; AMPA, amino-3hydroxy-5-methyl-isoxazole-4-proprionic acid; kainic acid, KA). Chapter 23 shows the distribution of binding to the glutamate receptors in cingulate cortex. Within each of these categories, multiple subunits and splice variants have been identified, and it is believed that these form heteromeric assemblies, the exact number and types of which remain to be determined. The most widely and densely distributed of the Glu receptor subtypes is the NMDA receptor. Several decades of work have shown that excessive activation of NMDA receptors (NMDA receptor hyperfunction, NRHyper) plays an important role in the pathophysiology of acute CNS injury syndromes such as hypoxia–ischemia, trauma, and status epilepticus (Olney, 1990). More recently it has become apparent that excitation of NMDA receptors (NMDA receptor hypofunction, NRHypo) also can injure CNS neurons.

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Pathomorphological Response At low doses, NMDA antagonists (e.g., MK-801, phencyclidine, ketamine, nitrous oxide, CPP, CPP-ene, CGS-19755) induce reversible pathomorphological changes (Olney et al., 1989) in layer IV–V pyramidal neurons of RSC. Figure 10 shows the laminar pattern of neuron death in deOlmos silver-stained sections. The greatest number of degenerating somata is in layers IV and Va and the associated degenerating dendrites form bundles in layers II and III and massive terminations throughout layer I. The loss of neurons in layers IV and

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Va is prominent in thionin-stained sections, particularly when compared to a control case stained for NeuN. Although the thionin section shows a small amount of damage to the top of layer Vb via pale staining and some shrinkage of neurons, most of the degeneration in layer Vb (Fig. 10) is associated with descending axons as noted in a subsequent figure. One type of neuron destroyed by this reaction is the medium-sized pyramidal neuron of layer Va. Less obvious are the types of neurons that degenerate in layer IV. The Golgi illustrations in Fig. 10D show that layer II is composed of fusiform pyramids (a, b), layer III of star pyramids

FIGURE 10 Systemic injections of MK-801 produce neuronal palor and gliosis in layers IV and Va of area 29c (A, thionin) when compared to the distribution of normal neurons (C, NeuN control). Silver-stained dendrites are prominent in layers I and IV–Va as are ascending dendritic bundles in layers II/III and somata in layers IV and Va. Although some neurodegeneration is in layer Vb, most of the argyrophilia is associated with descending axons as shown in detail in Fig. 11. In addition to the medium-sized neurons that express the pathomorphological response in layer Va, layer IV neurodegeneration is mainly by the star pyramids in this layer (D, neurons g and h). Neurons in superficial layer II include the fusiform pyramids and in layer III the star pyramids with mainly descending basal dendrites; however, neither of these latter two groups of neurons have the pathomorphological response.

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with basal dendrites that project into layer IV (c–f), and layer IV star pyramids with horizontally dispersed, basal dendritic trees (g, h). The apical dendrites of these latter neurons ascend throughout layers III, II, Ib, and Ic and arborize primarily in layer Ia. It is these latter neurons along with the medium-sized pyramids in layer Va that express much of the pathomorphological response following NMDA antagonist exposure. The pathomorphological injury consists of swollen mitochrondria and endoplasmic reticulum. If NMDA receptor blockade is maintained for a prolonged interval, as occurs following a single high dose or repeated treatment with lower doses of an NMDA antagonist, neurons in the RSC and several other cerebrocortical and limbic regions of the adult rat brain undergo irreversible degeneration as reviewed recently (Farber et al., 2002a).

Area 30 Deafferentation Following the Pathomorphological Response The precise localization of the pathomorphological response provides a unique opportunity to view an important intracingulate connection from granular to dysgranular retrosplenial areas. Figure 11 shows a low magnification of the RSC in a deOlmos silver-stained section as well as a higher magnification of the distribution of degenerating axons. In the rectangle of tissue in area 29c, there are bundles of degenerating axons that descend beneath the lesion in layer V. Projecting from these bundles at oblique angles and oriented toward area 30 are many individual axons that likely do not penetrate into the white matter but make a brief excursion directly to the adjacent area. Since there are

FIGURE 11 Selectivity of the pathomorphological response to granular area 29a–c provides an opportunity to evaluate intracingulate projections from these areas to area 30. This is from the same case shown in Fig. 10 that received an injection of MK-801. The obliquely oriented rectangle shows a high magnification of the descending bundles of axons emitted from the lesion (black arrows) and the obliquely oriented axons that are oriented toward area 30 (white arrows). Termination in area 30 (larger rectangle) is mainly in layer I but also in layers III and IV. The cytoarchitecture of area 30 is shown in the thionin section and this emphasizes that area 30 does not express the pathomorphological response.

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no degenerating neurons in area 30 and amino acid injections into area 29c have a halo of transported proteins around the injection site and extending into area 30 (Vogt and Miller, 1983), there is evidence in this material for a direct projection from area 29a–c to area 30. Support for this observation comes from analyzing the perilesion cortex in the MK-801-injected animals. Bundles of descending axons are emitted from beneath the area 29c pathomorphological response and branches penetrate superficial layers where they terminate in layers I, III, and IV of area 30 as shown in the higher magnification rectangle in Fig. 11 where a thioninstained section is provided for demonstration of the laminar boundaries in the silver-stained section. The direct connection demonstrated in the MK-801ablated tissue raises two issues. First, although these areas are structurally very different and both project independently to visual cortex, they are reciprocally connected (see amino acid injections in Vogt and Miller, 1983 for reciprocal projection to area 29c) and their functions are not independent. Indeed, they may contribute differently but in parallel to visuospatial processing. Second, functional deficits following MK-801 toxicity are not solely the result of damage to granular areas 29a–c but also of deafferentation of other cortices including area 30. Other likely candidates for deafferentation include the postsubiculum and area 24b′.

POLYSYNAPTIC CIRCUIT DISINHIBITION UNDERLIES NRHYPO NEUROTOXICITY Cholinergic System After the initial report of reversible neurotoxicity, it was found that GABAA receptor agonists and muscarinic receptor antagonists blocked the reversible neurotoxic reaction (Olney et al., 1991) and it was proposed that certain GABAergic inhibitory neurons receive tonic glutamatergic input via NMDA receptors and that these neurons form inhibitory synapses onto cholinergic neurons. NMDA antagonists, by blocking stimulation of GABAergic inhibitory neurons, would cause a loss of GABAergic inhibitory control over excitatory cholinergic neurons that innervate the RSC. The resultant excessive cholinergic stimulation of RSC neurons would be the proximal event to produce neuronal injury (Olney et al., 1991, Fig. 14). Based on the relative potencies of a large number of antimuscarinic compounds to prevent NRHypo neurotoxicity, it was determined that an m3 receptor was the most likely subtype of muscarinic receptor that was overstimulated on the injured neuron (Farber et al., 2002a); however, an m1 subtype could

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not be excluded. Consistent with the latter proposal, NMDA antagonists produce excessive release of acetylcholine (Kim et al., 1999) in the cerebral cortex and GABAergic agents can reverse this increase (Kim et al., 1999). In addition scopolamine, when injected directly into the RSC, prevents the damage caused by systemic injection of MK-801 (Farber et al., 2002a), confirming that excessive activation of muscarinic receptors in the RSC is necessary to produce the damage. Because there are cholinergic neurons in the RSC (Johnston et al., 1981; Olney et al., 1993), this circuit was conceived initially as being intrinsic to the RSC. However, MK-801 directly applied to the RSC did not cause an increase in acetylchoine release (Kim et al., 1999) nor did it produce the neurotoxicity (Farber et al., 2002a), indicating that the cholinergic neurons and the NMDA-receptor-bearing GABAergic neurons that control them are not in the RSC. Injection of muscimol, a GABA agonist, directly into the diagonal band of Broca, where cholinergic neurons that project to the RSC are located (Fig. 6), prevents NRHypo neurotoxicity (Jiang et al., 2001). Thus, the disinhibition of diagonal band cholinergic neurons by NMDA antagonists may result in the excessive stimulation of muscarinic receptors in the RSC.

Adrenergic System A large number of α2-adrenoreceptor agonists administered systemically prevent the neurotoxic reaction, and this protection can be reversed by α2-adrenergic antagonists (Farber et al., 1995a). The ability of α2adrenoreceptor agonists to prevent the increase in acetylcholine release induced by NMDA antagonists (Kim et al., 1999) indicates that α2-adrenoreceptor agonists also control cholinergic neurons in the diagonal band as shown in Fig. 12. Consistent with this conclusion, the injection of clonidine, an α2adrenoreceptor agonist, directly into the diagonal band can prevent the neurotoxicity whereas injection of clonidine into the RSC does not (Farber et al., 2002a).

Non-NMDA Glutamatergic System While these data confirm that disinhibition of the cholinergic system is a necessary component underlying NRHypo neurotoxicity, it is not sufficient because the injection of carbachol, a muscarinic agonist, directly into the RSC does not reproduce the damage (Farber et al., 2002a). NMDA antagonists also produce excessive release of Glu in the cerebral cortex suggesting that excessive stimulation of glutamatergic receptors might also be involved in the neurotoxic process. NBQX, an antagonist of AMPA and KA receptors, protects

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FIGURE 12 Circuitry mediating NRHypo neurotoxicity. Glu appears to act through NMDA receptors on GABAergic and noradrenergic (NE) neurons and maintains tonic inhibitory control over two major excitatory pathways that convergently innervate neurons in granular RSC, areas 29a–c. Systemic administrations of NMDA antagonists would block NMDA receptors, thereby abolishing inhibitory control over both excitatory inputs to the RSC. The disinhibited excitatory pathways would then simultaneously hyperactivate RSC neurons, possibly disrupting multiple intracellular signaling systems and thereby causing immediate derangement of the cognitive functions of the RSC and reversible or irreversible neuronal injury, depending upon the length of the exposure. It is postulated that the glutamatergic cell bodies that project to the AMPA/KA receptors in the RSC are located in the anterior thalamus. Although this diagram emphasizes the RSC, a similar disinhibitory mechanism and similar but not necessarily the same circuits and receptor mechanisms may mediate damage in other corticolimbic regions by sustained NRHypo. Excitatory (+) and inhibitory (−) inputs are shown. ACh, acetylcholine; GA, GABAA receptor; m3, muscarinic receptor subtype; σ, sigma site; 5-HT, serotonin.

against NRHypo neurotoxicity, when applied systemically or when injected directly into the RSC (Farber et al., 2002a), indicating that the excessively stimulated glutamatergic receptors are likely of the AMPA/KA subtype. Although injection of KA or AMPA directly into the RSC does not reproduce the damage (Farber et al., 2002a), coinjection of KA and carbachol does reproduce the neurotoxicity (Farber et al., 2002a). The need for both agents to produce the damage indicates that the combined excessive activation of both muscarinic and non-NMDA, glutamatergic receptors is necessary and sufficient to produce the neurotoxicity. Injection of muscimol into either the AD/AV or LD nucleus of the thalamus, where thalamic input into the RSC arises (Figs. 6 and 9), protects against NRHypo neurotoxicity (Jiang et al., 2001), indicating that thalamic glutamatergic neurons are the likely source of the excessive release of Glu in the RSC and that these neurons also are under tonic inhibition from NMDAreceptor-bearing GABAergic neurons (Fig 12).

Additional Evidence That NMDA Antagonists Produce Disinhibition Based on this disinhibition model, agents that reduce the ability of these excitatory projections to release excessive neurotransmitter and stimulate the vulner-

able RSC neuron should protect against the neurotoxic reaction. Activation of voltage-gated sodium channels is necessary for propagation of the action potential down the axon and inhibitors of these channels, e.g., tetrodotoxin, valproic acid, and carbamazepine, prevent NRHypo neurotoxicity (Farber et al., 2002b). NMDA antagonists also acutely increase metabolism in certain corticolimbic regions (Farber et al., 1999, 2002a). In general the corticolimbic regions experiencing hypermetabolism tend to be the same corticolimbic regions that also develop either the reversible or irreversible forms of NRHypo neurotoxicity. The increase in metabolism in these corresponding regions could be a reflection of a disinhibition syndrome in which acetylcholine and Glu are excessively released at certain corticolimbic neurons that are injured in the NRHypo neurotoxic syndrome. Consistent with this proposal, clozapine and halothane reverse the hypermetabolism induced by NMDA antagonists (Duncan et al., 1998b) just as they reverse NRHypo neurotoxicity (Ishimaru et al., 1995; Farber et al., 1996).

Other Markers of NRHypo-Induced Pathology and Disinhibition Circuit NRHypo produces several other effects. Dragunow and Faull (1990) reported that MK-801 induced the

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production of c-Fos protein in these same neurons and, not only c-Fos, but other immediate-early genes, including c-Jun, Jun-B, NGFI-A [a.k.a. zif268, krox-24], NGFI-B, NGFI-C, and Nurr1 (Farber and Newcomer, 2002), are activated by NRHypo. In addition, the heat shock protein HSP70 and its mRNA are induced by NRHypo (Sharp et al., 1991; Olney et al., 1991). Lastly, NRHypo induces the expression of brain derived growth factor mRNA (Hughes et al., 1993; Castren et al., 1993). The ability of some of the same pharmacological treatments, which have been shown to prevent NRHypo neurotoxicity, to prevent these other responses (Farber and Newcomer, 2002) suggests that these other responses may be secondary to activation of the same NRHypo disinhibition mechanism. Consistent with this proposal, PCP’s induction of c-Fos and HSP70 has a similar age-dependency profile (Sharp et al., 1992; Sato et al., 1997), as does MK-801 induction of the reversible form of NRHypo neurotoxicity (Farber et al., 1995b).

NRHYPO-INDUCED PSYCHOSIS A variety of NMDA antagonists (e.g., ketamine, PCP, CPP, CPP-ene, CGS19755, CNS 1102) cause a psychotic state in humans (Farber and Newcomer, 2002). These findings suggest that a NRHypo state might be involved in the pathophysiology of psychotic disorders. While schizophrenia has received the most attention as the disorder in which an NRHypo state might exist (e.g., Olney and Farber, 1995), the fact that NMDA antagonists can produce maniacal excitation, catatonic signs and euphoria suggests that such a NRHypo state also could be responsible for some of the signs and symptoms of bipolar and schizoaffective disorder (Farber and Newcomer, 2002b). Based upon several intriguing parallels between NRHypo neurotoxicity and NRHypo-induced psychosis, it has been proposed (Olney and Farber, 1995; Farber et al., 1999; Farber and Newcomer, 2002) that the complex polysynaptic disinhibition mechanism that underlies the neurotoxic action of NMDA antagonists also underlies their psychotomimetic effects. This model proposes that mild elevations in the release of acetylcholine and Glu induced by mild NRHypo result in functional overactivation of cerebrocortical neurons and their projection fields, producing cognitive and behavioral disturbances without neurotoxicity. More severe NRHypo causes greater increases in the amount of excessive transmitter release and in the degree of postsynaptic m3 and non-NMDA receptor overstimulation, resulting in neurotoxicity. While the exact role that a NRHypo-disinhibited state plays in idiopathic

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psychotic disorders like schizophrenia is mostly hypothetical, the data in rodents point to the importance of NMDA-receptor-bearing GABAergic interneurons in certain cortical and thalamic regions. Consistent with this conclusion are reports of deficiencies in GABAergic and NMDA/glutamatergic systems in cortical and thalamic regions of subjects with schizophrenia and other idiopathic psychotic disorders (Woo et al., 1998; Benes, 1999; Guidotti et al., 2000; Ibrahim et al., 2000).

NRHYPO AND NEURODEGENERATION IN ALZHEIMER’S DISEASE One of the first sites of impaired glucose metabolism in Alzheimer’s disease (AD) patients with early memory impairment is in posterior cingulate cortex (Minoshima et al., 1997) and this includes the RSC. An important basis for postulating that NRHypo may play a role in AD is that the disseminated pattern of irreversible neuronal degeneration induced in the adult rat brain by NMDA antagonists (Corso et al., 1997; Wozniak et al., 1998) resembles the pattern of neurofibrillary degeneration in AD. In addition, pyramidal neurons are most vulnerable to NRHypo degeneration and they are also the cell type most vulnerable in AD. Thus, the NRHypo disinhibition model of neurotoxicity could offer a partial explanation for the distribution pattern of neurodegeneration in AD. Hypofunction of the NMDA receptor system, which is the condition that triggers neurodegeneration in the NRHypo model, is a condition present in the normal aging brain and may be present, to a more exaggerated degree, in the brains of AD patients (Olney et al., 1997). Moreover, it is generally agreed that loss of synaptic complexes is the specific neuropathological change that correlates most closely with cognitive deterioration in AD. The neurotoxicity induced by NMDA antagonists involves the selective deletion of dendritic spines and large numbers of synaptic complexes (Corso et al., 1997; Wozniak et al., 1998) and these changes induced by NMDA antagonists are associated with memory loss in rodents (Wozniak et al., 1996; Brosnan-Watters et al., 1996, 1999). In addition, although hyperphosphorylation of tau protein has been proposed as a mechanism to link neurofibrillary tangle formation, only limited headway has been made in understanding the mechanisms that initiate and drive the hyperphosphorylation process. The NRHypo mechanism entails excessive activation of transmitter receptors on the surface of the types of neurons that degenerate in AD and these receptors are linked to second-messenger systems which, if hyperactivated, might provide the driving force for a

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hyperphosphorylation process. Based on these considerations it has been proposed that the NRHypo mechanism acts in concert with amyloid deposition in a multiphase process that results in AD (Olney et al., 1997; Farber et al., 2002b).

COMPARISON OF MEDIAL CORTEX IN RAT AND MONKEY One of the reasons for employing a modification of Brodmann’s original scheme for rodent and primate species is to assure that direct comparisons can be made among species and support a rational process for devising models of human disease. This type of analysis does not presume evolutionary or developmental relationships, although homologies may exist. Rather, it states that areas on the medial surface in all mammalian species undergo a series of architectural transitions and that each area evaluated in this context provides for a direct comparison and areas with the same relative position may be similar among species. Demonstration of similarities between areas in different species and use of a common nomenclature does not imply that two areas with the same designation are exactly equivalent, only that they share enough similarity to explore common mechanisms of disease. Here we consider relations between rat and rhesus monkey. Figure 13 shows the medial surface of both animals with the areas delimited. Areas in monkey cortex that do not appear to have a rodent counterpart are mainly in the cingulate sulcus and include areas 24c, 24c′, and 24d as well as the gyral areas 23 and 31. The cingulate sulcus in the monkey was opened so this point could be better appreciated. Although the cingulate motor areas on monkey in areas 24c′ and 24d are not present in rat, there is a part of the cingulate cortex in rat that projects to the spinal cord including areas 32 and 24b and it overlaps with AGm (Miller, 1987). Area 24b could be homologous to the rostral cingulate motor area in primates; however, this conclusion suggests that cingulate skeletomotor activity in rat is mediated by the pACC, while that in monkey is associated with the MCC indicating a role for these projections to spinal cord in rat very different from their role in monkey. The massive posterior cingulate gyral surface of primates has no equivalent in rodents, since monkey areas 23a, 23b, and 31 on the gyral surface and area 23c in the caudal cingulate sulcus cannot be identified in rat. The RSC in the rodent is composed of granular area 29 and dysgranular area 30 and this cortex forms the entire PCC in this species. While areas 29 and 30 in the rat are similar to those in primates, these latter areas are actually buried in the callosal sulcus in monkey

FIGURE 13 The ultimate success in modeling human disease with rodents depends on determining similarities among the medial surfaces of rat and different primate species. Here the cingulate areas in rat and monkey are outlined in photographs at the same magnification. In the monkey the cingulate sulcus was separated (double arrow) to expose the depths of the cingulate sulcus. The splenium of the corpus callosum was also warped ventral from the point marked with small dots so the depths of the callosal sulcus can be appreciated. Area 25 in both species is shadowed as are areas 29 and 30 in both species. Although similar cortical regions are smaller in rat, the pericallosal areas in monkey are shown: areas 25, 32, 24a/b, 24a′/b′, 29a–c, and 30. The two regions that do not appear to have counterparts in the rat include monkey areas in the cingulate sulcus (24c, 24c′, 24d) and on the posterior cingulate gyrus (23, 31). The greatest similarity between rat and monkey is in the structure of the pericallosal areas.

rather than forming the gyral surface as in rat. The corpus callosum in the monkey in Fig. 13 was warped ventrally to expose the depths of the callosal sulcus and demonstrate the retrosplenial areas therein. Areas 23a, 23b, and 31, which are on the surface of the posterior cingulate gyrus in monkey, and area 23c in the caudal cingulate gyrus together form the PCC in primates and do not appear to have counterparts in the rat. Rose and Woolsey (1948) emphasized this fact by lauding M. Rose’s observations with their following observation: “Area 23 as determined by Brodmann in the rabbit, by Krieg in the rat and by virtually all others except M. Rose, who denied its existence in the rodents, is not likely to exist in any of the loci which have been labeled 23 on rodent cortical maps. M. Rose was obviously right in maintaining that in the rodent’s cortex there is nothing resembling area 23 of carnivores and primates. What appears to be its equivalent area in the rodents has such an outspoken “retrosplenial” appearance that no student of architectonics ever has suggested that it may be equivalent to area 23 in higher forms.” Approximately equivalent areas between the rat

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and monkey include the following. (1) Area 25 (shaded in both in Fig. 13) has a subgenual position. (2) Area 32 is rostral to area 24 in both species. (3) Areas 24a/b and 24a′/b′ have equivalents, although in the rat these areas comprise the entire perigenual and midcingulate regions. (4) Although there is no direct equivalent for the primate sulcal cingulate motor areas 24c′ and 24d in the rat, there are a moderate number of corticospinal projection neurons in rat cingulate cortex as noted above and this supports the notion of a spinal connection, though not from an area that is similar in these species. The different origins of corticospinal projections underscores the less differentiated functions of each rodent cingulate area in contrast to monkey cortex where the corticospinal projections are differentiated into motor areas that are separated from gyral divisions of area 24′. (5) Areas 29a–c appear similar to areas 29l and 29m in monkey and rat area 30 is similar to monkey area 30. Thus, similarities between rat and monkey medial cortices are most prominent in the pericallosal areas. Even when an area appears to have a similar laminar organization and position in cortical differentiation trends (i.e., periallocortex, proisocortex, isocortex), differences can exist at the connection and cellular levels. At the level of extrinsic connections, it was noted above that corticospinal projections arise from areas in the pACC in rat rather than in the MCC as in monkey. Also, the primary and secondary visual cortices have major and reciprocal connections with area 29 in rat; however, these do not exist in monkey (Vogt and Pandya, 1987). At the cellular level, even though granular area 29 in rat has a similar counterpart in the monkey, they are not cytologically equivalent. Indeed, the fusiform and extraverted pyramids in rat layers II and III in area 29 have not been observed in monkey (Vogt, 1976; Vogt and Peters, 1981). Finally, at the receptor expression level, presynaptic heteroreceptor organization appears to be different. The presynaptic M2 binding in layers Ia and IV that is so clear in rat has not been observed in monkey (Vogt et al., 1997) where layer I has little dendritic arborization and only weak overall binding for many transmitter receptors due to the presence of a myelin-rich fiber tract passing through layer I (taenia tecta). Despite the differences between rat and monkey, the essential architecture of pericallosal areas is similar in both species and the rat cortex has significant value as a potential model for certain diseases including neurodegenerative and pharmacological models of psychiatric disease. Given the less differentiated connections, intrinsic organization, and functions of rodent areas, it is unlikely these differences can be overlooked when assessing the mechanisms of cell death and dysfunction. Indeed, rodent models must be considered in the context

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of these unique morphological and functional properties.

RODENT MODELS OF DISEASE Morphology at all levels of analysis provides a basis for assessing the extent to which the rat can be used to model primate diseases. If the pericallosal areas play an important role in the onset and/or progression of a disease, the rat is an appropriate choice for model development. However, if the cingulate sulcal or posterior cingulate gyral areas are the primary target, the rodent is not an appropriate animal. Even when pericallosal areas are the primary region of interest, intrinsic differences among rat and primate species could restrict the value of the rat as a model system. For example, although the rat is often used to study the mechanisms of diseases of the basal ganglia, the human has substantially more calretinin-expressing neurons than does the rat and, to the extent that mechanisms of neurodegeneration in movement disorders depend on the calcium-buffering properties of these neurons in human, rat may not be a useful model of these diseases (Wu and Parent, 2000). Indeed, the cytology, connections, and transmitter receptors expressed by afferent axons to area 29 are not the same in rodent and primate and there does not appear to be, for example, heteroreceptor regulation of thalamic afferents in monkey as there is in rat. Thus, defining animal models for cortical diseases involves determining which areas are similar and the extent to which similar areas have the same organization. Although the rat has areas 25 and 24a/b that have a relative degree of laminar differentiation similar to that of the monkey, the rodent has cingulospinal projections that originate from the pACC rather than the cingulate motor areas, which it does not have. Aspects of neurodegeneration in multiple systems atrophy, therefore, may not be ideal candidates for study in rodent cortex. Furthermore, although the RSC receives anterior thalamic afferents in rodents and primates, muscarinic, presynaptic heteroreceptors regulate these terminals in rat but not in primate (Vogt et al., 1997). The import of this difference is currently unclear; however, this difference provides a unique opportunity to determine the importance of these receptors in the rodent and what beneficial or detrimental consequences their absence has for primates. In this chapter we discussed the neurotoxic effects of NMDA antagonists in rodents and how these effects could shed light on certain diseases like schizophrenia and Alzheimer’s disease. An important step that remains is to determine whether a similar neurotoxicity can be induced in nonhuman primate brain. Obviously, finding

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similar damage in similar brain regions would be important for advancing our understanding of human physiology and pathophysiology and would testify to the importance of using the NRHypo rodent model to study primate CNS function and disease. However, not finding similar damage in similar brain regions would also provide beneficial information and potentially could be important over the long run for understanding of the primate brain. Finding NRHypo neurotoxicity in non-RSC areas would provide information about the importance of NRHypo neurotoxicity for human biology and may shift interpretations of the psychogenic properties of dissociative anesthetics. Neurotoxicity in other regions in primates might be responsible for the cognitive and behavioral changes (e.g., psychosis) seen with NMDA antagonists. Ultimately, specifying animal models of human diseases is a dynamic process of refining the cellular and molecular mechanisms of brain structure and function. For example, identifying the distribution of amyloid-β peptide in early cases of Alzheimer’s disease led to in vitro and in vivo studies of its neurotoxic properties. This led to analysis of its deposition in murine transgenic models that deleted either or both presenilin genes and mutating the amyloid precursor protein gene. Although no one would suggest that behavioral and structural changes in the mouse are equivalent to those in human, the actions of these genes and previous studies in rat and monkey are now serving as a basis for exploring new therapeutic interventions in human clinical trails. Continued progress in understanding the mechanisms of human disease will depend upon hypotheses and mechanistic findings generated via the dynamic interchange among research activities using many mammalian species and a systematized nomenclature serves as a platform for this process.

Acknowledgments This work, including developing rodent and primate models of Alzheimer’s disease and other neurological and psychiatric disorders, is supported by grants from the National Institutes of Health (NINDS NS38485 and NS44222; NIA PO1-AG11355).

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C H A P T E R

23 Isocortex NICOLA PALOMERO-GALLAGHER Institute of Medicine, Research Centre Jülich, Germany

KARL ZILLES Institute of Medicine, Research Centre Jülich, Germany C. and O. Vogt Institute of Brain Research Heinrich-Heine University Düsseldorf, Germany

The rat isocortex has been subject of numerous mapping studies, which revealed a considerable degree of regional differentiation into functionally and structurally specialized regions (Burwell, 2001; Droogleever Fortuyn, 1914; Krieg, 1946a, 1946b; Paxinos and Watson, 1986; Paxinos et al., 1999; Schober, 1986; Svetukhina, 1962; Swanson, 1992, 1998; Von Volkmann, 1926; Zilles, 1985, 1990; Zilles and Wree, 1985; Zilles et al., 1980). However, the resulting maps differ considerably regarding the number and size of identified cortical areas. Furthermore, considerable discrepancies exist concerning the nomenclature of isocortical regions. One system contains numerous terms formulated according to Brodmann’s (1909) classification of the human cortex (Krieg, 1947), although homologies between primate and rat brains were not demonstrated. The use of terms such as “striate” and “peristriate” areas (Montero, 1973, 1981; Montero et al., 1973a) suggest architectonical similarities between primate and rodent cortical areas. However, since the primary visual cortex of the rat does not have a Gennari stripe, which is the reason for the term “striate area” in the cerebral cortex of higher primates, these terms are incorrect. Aiming at a comprehensive, neutral, and self-consistent nomenclature for the rat isocortex, Zilles and Wree (1985) proposed a topographic division into frontal (Fr), parietal (Par), temporal (Te), and occipital (Oc) regions, which, in turn, can be subdivided into several areas, designated by numbers. Some of these areas can be further subdivided (e.g., the primary visual cortex Oc1 into the

The Rat Nervous System, Third Edition

monocular and binocular subfields Oc1M and Oc1B; Zilles et al., 1984). Mapping studies are based on a wide range of methods, which include axonal tracing, electrophysiology, and immunohistochemistry, as well as other diverse histological stainings. However, classical studies relied mostly on extensive observations of cellbody-stained sections and were based mainly on an observer dependent use of cytoarchitectonic criteria. In recent years, quantitative in vitro receptor autoradiography has proven to be a powerful mapping tool (for a review see Zilles et al., 2002a, 2002b). Receptors for GABA, glutamate, acetylcholine, noradrenaline, and serotonin are heterogeneously distributed throughout the cerebral cortex. They show regional differences in both their mean densities and their laminar distribution patterns. Furthermore, these variations are also present between different receptor types for a single neurotransmitter, and although each receptor does not indicate all possible areal borders, there is a perfect agreement in the location of those borders visualized by several receptors. Therefore, receptor autoradiography reveals the brain’s chemoarchitectonic organization, which is correlated with its cyto- and myeloarchitectonical as well as its functional organization (Zilles et al., 2002a). The aim of the present chapter is to present a parcellation scheme of the rat isocortex based on cyto- and myeloarchitectonical criteria as well as on recently registered receptor-architectonical data. We have used cell-body-stained sections as a “template” to present

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the results achieved by comparisons of the regional and laminar distribution patterns of various transmitter receptors with cyto- and myeloarchitectonical borders. Since a single neuron expresses a variety of receptor subtypes of different neurotransmitter systems, a single architectonical area will contain many different receptor subtypes, which may interact with each other. Therefore, it would be advantageous to examine as many different subtypes of receptors from as many different neurotransmitter systems as possible, in order to obtain a comprehensive view of such a complex system as that constituted by a cortical area. We have, therefore, concentrated on receptors for the classical neurotransmitters glutamate, GABA, acetylcholine, noradrenaline, and serotonin (for a review, see Schleicher and Zilles, 1988; Zilles and Schleicher, 1995; Zilles et al., 2002b). The site-specific balance between different receptors in a single architectonically defined brain region can be visualized as a “receptor fingerprint” (Zilles et al., 2002a, 2002b; Zilles and PalomeroGallagher, 2001), which is based on the quantification of the mean (averaged across all cortical layers) receptor density in femtomoles per milligram protein of each examined receptor (Fig. 1). The shapes and sizes of a fingerprint are specific for each area. Cortical areas with different functions and architecture differ in the shapes of their fingerprints (Fig. 1A), whereas cortical areas with a similar function build a neurochemical family, which results in similarly shaped receptor fingerprints (Fig. 1B). Thus, the receptor fingerprint represents multimodal organizational aspects of a cortical area. The information contained in a fingerprint can also be expressed as a feature vector and further processed using a hierarchical clustering procedure. This analysis was used to group cortical areas into clusters, each of which comprise regions with similar receptor distributions, i.e., fingerprints (Fig. 2). Based on differences in its laminar structure, the rat cortex, like that of all mammals, can be subdivided into isocortex and allocortex, which are separated by a transition zone. The allocortex is the phylogenetically oldest cortex; it encompasses those regions showing a highly variable laminar structure and comprises both the paleo- and the archicortex (Vogt and Vogt, 1919). The term isocortex, or neocortex (Vogt and Vogt, 1919), describes the phylogenetically youngest cortical regions, which show a homogeneous lamination into six layers. The transition zone encompasses the regions bordering the isocortex and displays gradual changes in its architectonic pattern, ranging from an isocortical–proisocortical zone to an allocortical–periallocortical structure (Stephan, 1975). The present chapter focuses on the parcellation of the rat isocortex and its neighboring regions (Fig. 3). For a list of abbreviations, see Table 1.

ISOCORTEX The rat isocortex is characterized by a typical laminar organization consisting of six layers that run parallel to the cortical surface. Regional differences in laminar architecture enable a parcellation of the isocortex into areas which can be further characterized by their predominant connectivity and function as either motor or unimodal or multimodal associative sensory regions. Areas with which motor functions have been associated are characterized by a poorly developed or even absent inner granular layer (Brodmann, 1909). Conversely, sensory areas have a conspicuous inner granular layer, which is the target of numerous afferents from modality-specific thalamic nuclei. Most isocortical regions exhibit packing densities of glial fibrillary acidic protein-immunopositive cells lower than those of the adjoining periallocortical areas (Zilles et al., 1991).

Frontal Cortex The frontal isocortex represents the motor cortex of the rat (Donoghue and Wise, 1982; Donoghue et al., 1979; Hall and Lindholm, 1974; Neafsey, 1990; Neafsey and Sievert, 1982; Wiesendanger and Wiesendanger, 1982a; Wise, 1975) and is an architectonically inhomogeneous region (Fig. 4). Based on their cyto- and myeloarchitecture, as well as on their chemoarchitecture, local cerebral glucose utilization (LCGU), and connectivity patterns, three areas can be defined: Fr1, Fr2, and Fr3. Fr1 represents the primary motor cortex of the rat brain, with Fr3 as a somatotopical subfield, and Fr2 is the putative anatomical equivalent of the primate premotor, supplementary motor, and frontal eye field areas (Donoghue and Parham, 1983; Hicks and Huerta, 1991; Van Eden et al., 1992). Cytoarchitectonically (Fig. 4), the laminar pattern of the frontal regions is characterized by the lack of a prominent layer IV and the presence of large and densely packed pyramidal cells in the outstanding inner pyramidal layer, thus enabling its delineation from the adjacent parietal cortex. Fr1 and Fr2 have conspicuous layers II and V. Layer III is clearly narrower in Fr2 than in Fr1. Fr3 has broader layers II–V than Fr1 as well as a slightly higher cell packing density in the lower range of layer III. Fr3 has a broader layer V than of Fr1 and Fr2, but has the lowest cell packing density and also the lowest gray level index (GLI) values (Zilles and Wree, 1995). The delineation of the frontal areas varies markedly in different studies (Krieg, 1946a, 1946b; Swanson, 1992, 1998; Zilles, 1985; Zilles and Wree, 1995; Zilles et al., 1980). Zilles et al. (1980) originally described four

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FIGURE 1 (A) Receptor fingerprints of the primary motor (Fr1), somatosensory (Par1), auditory (Te1), and visual (Oc1) areas of the rat cortex. (B) Receptor fingerprints of the three frontal (Fr1, Fr2, Fr3) areas of the rat cortex. The mean densities of AMPA (labeled with [3H]AMPA), kainate (labeled with [3H]kainate), NMDA (labeled with [3H]MK-801), GABAA (labeled with [3H]muscimol), GABAB (labeled with [3H]CGP 54626), BZ (labeled with [3H]flumazenil), M1 (labeled with [3H]pirenzepine), M2 (labeled with [3H]oxotremorine-M), M3 (labeled with [3H]DAMP), nicotinic (labeled with [3H]epibatidine), α1 (labeled with [3H]prazosin), α2h (labeled with [3H]UK-14, 304), 5-HT1A (labeled with [3H]8-OH-DPAT), and 5-HT2 (labeled with [3H]ketanserin) binding sites are displayed in polar coordinate plots (binding site densities in 0–5000 fmol/mg protein). The lines connecting the mean densities of the receptor types measured in each cytoarchitectonically determined cortical area define the contour of the fingerprint. The shapes and sizes of the fingerprints are specific for each area.

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FIGURE 2 Dendrogram resulting from the hierarchical clustering analysis of the mean regional densities of glutamatergic (AMPA, NMDA, and kainate), GABAergic (GABAA, GABAB, and benzodiazepine binding sites), cholinergic muscarinic (M1, M2, and M3), cholinergic nicotinic (N), adrenergic (α1 and α2h), and serotoninergic (5-HT1A and 5-HT2) receptors in the isocortical regions of the rat brain. The mean densities of all examined receptor types in a given area can be expressed as a feature vector. The feature vectors of all examined areas averaged across animals can subsequently be statistically evaluated by means of a hierarchical clustering analysis. In order to do so, Euclidean distances are calculated for all possible combinations of pairs of areas, and the final clustering of neurochemically related areas is determined by a Ward linkage algorithm. The greater the similarity in the shape and size of the fingerprints of two areas, the smaller the resulting Euclidean distance. The occipital regions Oc1M and Oc1B are an example of two highly related functional regions that are members of the same cluster. They represent the monocular and binocular parts of the primary visual cortex of the rat and can be clearly delineated from each other based on their cyto- and myeloarchitectonical characteristics, as well as on their differential connectivity patterns (for details see text). However, they do not differ significantly in their chemoarchitecture, a fact which is reflected in the smallest Euclidean distance measured between all isocortical areas. It is interesting to note that temporal areas Te2D and Te2C do not cluster with the remaining temporal areas, but with the occipital, visual areas. This is in accordance with the involvement of Te2 in visual attention tasks (for details see text). Furthermore, the hierarchical clustering clearly segregates Fr3 from the parietal regions, thus further supporting our classification of this area as a frontal cortical region and not a parietal one (see text for details).

regions, Prcm, Prc1, Prc2, and Prc3, within the frontal cortex, but in subsequent studies merged Prcm and Prc3 into the present Fr2 (Zilles, 1985, 1990; Zilles and Wree, 1995; Zilles et al., 1990). This definition of Fr2 is in agreement with findings of microstimulation and tracing studies (Donoghue and Wise, 1982; Leong, 1983; Wiesendanger and Wiesendanger, 1982a). Although Donoghue and Wise (1982) do not differentiate

between Fr1 and Fr2 (they only defined one region, their Ag 1), these two regions differ considerably in their cell packing densities, in their GLI values (Zilles and Wree, 1995), and in their mean receptor densities and distribution patterns (Figs. 1 and 5–7). Swanson (1992, 1998) included a region comparable to our Fr3 in his SS (our Par1). The presence throughout the frontal cortex of some very small cell bodies similar to the

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FIGURE 3 Schematic drawings of the lateral, dorsal, and medial surfaces of the rat brain showing the parcellation scheme of the isocortex and of the regions of transition between the isocortex and the allocortex. See Table 1 for abbreviations.

neurons in layer IV of Par1 and the slightly higher cell packing density in the lower range of layer III of Fr3 (Zilles, 1990), which may be comparable to layer IV in the parietal regions, could lead to the inclusion of Fr3 in the parietal cortex. However, the general agranular to dysgranular architecture of the frontal areas, including Fr3, supports the hypothesis that Fr3 must be considered as part of the frontal isocortex (Zilles

and Wree, 1995). Moreover, Fr3 and Par1 differ considerably in their GLI values (Zilles and Wree, 1995). Furthermore, neurochemically, Fr3 resembles Fr1 and Fr2 more closely than it does Par1, as statistically determined by the hierarchical cluster analysis (Fig. 2). The regional and laminar cerebral glucose utilization patterns within the frontal cortex were registered by means of quantitative 2-DG autoradiography (Zilles

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TABLE 1

List of Abbreviations

ac

Anterior commissure

AID

Agranular insular cortex, dorsal part

AIP

Agranular insular cortex, posterior part

AIV

Agranular insular cortex, ventral part

AO

Anterior olfactory nucleus

cc

Corpus callosum

Cg1

Cingulate cortex, area 1, rostral part

Cg1′

Cingulate cortex, area 1, caudal part

Cg2

Cingulate cortex, area 2, rostral part

Cg2′

Cingulate cortex, area 2, caudal part

Cg3

Cingulate cortex, area 3

Ent

Entorhinal cortex

FL

Parietal cortex, forelimb area

Fr1

Frontal cortex, area 1

Fr2

Frontal cortex, area 2

Fr3

Frontal cortex, area 3

HL

Parietal cortex, hindlimb area

IL

Infralimbic area

LO

Lateral orbital area

LS

Lateral septal nucleus

MO

Medial orbital area

Oc1B

Occipital cortex, area1, binocular part

Oc1M

Occipital cortex, area1, monocular part

Oc2L

Occipital cortex, area2, lateral part

Oc2ML

Occipital cortex, area2, mediolateral part

Oc2MM

Occipital cortex, area2, mediomedial part

ox

Optic chiasm

Par1

Parietal cortex, area 1

Par2

Parietal cortex, area 2

ParPC

Parietal cortex, posterior area, caudal part

ParPD

Parietal cortex, posterior area, dorsal part

ParPR

Parietal cortex, posterior area, rostral part

ParVC

Parietal cortex, ventral area, caudal part

ParVR

Parietal cortex, ventral area, rostral part

Pir

Prepiriform cortex

PRh

Perirhinal area

RSA

Agranular retrosplenial cortex

RSG

Granular retrosplenial cortex

Te1

Temporal cortex, area 1

Te2C

Temporal cortex, area 2, caudal part

Te2D

Temporal cortex, area 2, dorsal part

Te3R

Temporal cortex, area 3, rostral part

Te3V

Temporal cortex, area 3, ventral part

TeV

Temporal cortex, ventral area

TTd

Taenia tecta, dorsal part

TTv

Taenia tecta, ventral part

Tu

Olfactory tubercle

VLO

Ventrolateral orbital area

VO

Ventral orbital area

FIGURE 4 Cyto- and myeloarchitectonical structure of the frontal isocortical areas Fr1, Fr2, and Fr3 visualized in adjacent coronal sections. Roman numerals indicate cortical layers.

and Wree, 1995). The registration of LCGU can provide a map of the functional activity of cortical areas or layers, since functional activity and energy metabolism of the normal adult brain are entirely dependent on glucose metabolism (Elliott and Heller, 1957; Lassen, 1959; Sokoloff, 1982). LCGU in conscious animals is closely linked to synaptic (Nudo and Masterton, 1986) and ion pump (Mata et al., 1980) activity. Therefore, LCGU mapping can be looked upon as representing a “metabolic encephalography” (Sokoloff, 1982). Fr1 shows the highest LCGU values within layer V. However, it contains a LCGU lower than that of the medially adjoining Fr2, especially in layers II–V. Fr3 has a high LCGU predominantly in the lower part of the supragranular layers. Fr1, Fr2, and Fr3 show considerable differences concerning their mean densities of receptors for classical neurotransmitters (Fig. 1). Glutamatergic AMPA (Fig. 7) and NMDA receptors show the highest densities in layers I–III and the lowest values in layers V–VI (Zilles et al., 1990). Kainate receptors (Figs. 5–7) show the opposite laminar distribution pattern, with highest densities located in layers V–VI. The laminar distribution pattern of kainate receptors in the frontal cortex nicely reflects the distribution of zinc-containing vesicles revealed by the Timm stain (Zilles et al., 1990). Fr2 has the lowest AMPA and NMDA receptor densities (Fig. 1), especially in layers I–III, whereas layers V–VI of Fr3 contain the lowest AMPA and NMDA concentrations of the three frontal areas. Mean regional kainate densities are high in Fr2, intermediate in Fr1, and low in Fr3 (Fig. 1). GABAergic receptors and benzodiazepine binding sites show equal laminar distribution patterns

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FIGURE 5 Neighboring coronal cryostat sections (15 μm thick) through the rat brain at two different rostrocaudal levels (A, B, and C rostral to D, E, and F) processed for silver cell-body (A, D) and myelin (B, E) staining as well as for the visualization of glutamatergic kainate receptors (C, F) by means of [3H]kainate. The scale bar indicates the color coding of kainate binding site densities in femtomoles per milligram protein, and changes in kainate receptor density and laminar distribution pattern coincide with the cyto- and myeloarchitectonically defined borders. See Table 1 for abbreviations.

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FIGURE 6 Neighboring coronal cryostat sections (15 μm thick) through the rat brain at two different rostrocaudal levels (A, B, and C rostral to D, E, and F) processed for silver cell-body (A, D) and myelin (B, E) staining as well as for the visualization of glutamatergic kainate receptors (C, F) by means of [3H]kainate. For further details, see Fig. 5. See Table 1 for abbreviations.

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FIGURE 7 Neighboring coronal cryostat sections (15 μm thick) through the rat brain processed for silver cell-body staining (A) or the visualization of AMPA (B), kainate (C), GABAA (D), M2 (E), nicotinic (F), α2h (G), and 5-HT2 (H) receptors. Color coding indicates binding site densities in femtomoles per milligram protein. AMPA receptors: dark blue, less than 950 fmol/mg protein; red, more than 2650 fmol/mg protein. Kainate receptors: dark blue, less than 1340 fmol/mg protein; red, more than 3650 fmol/mg protein. GABAA receptors: dark blue, less than 1000 fmol/mg protein; red, more than 2490 fmol/mg protein. M2 receptors: dark blue, less than 410 fmol/mg protein; red, more than 1040 fmol/mg protein. Nicotinic receptors: dark blue, less than 150 fmol/mg protein; red, more than 590 fmol/mg protein. α2h receptors: dark blue, less than 280 fmol/mg protein; red, more than 1520 fmol/mg protein. 5-HT2 receptors: dark blue, less than 290 fmol/mg protein; red, more than 975 fmol/mg protein. The regional and laminar distribution patterns of a single transmitter receptor reveal interareal borders that coincide with cyto- and myeloarchitectonically defined borders. A single receptor does not necessarily reveal all borders; rather, it can define the neurochemical family of cortical areas with a similar function. However, there is a perfect agreement in the location of those borders that are displayed by several receptors. See Table 1 for abbreviations.

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throughout the three frontal regions, reaching the highest values in layers I–III. The laminar distribution pattern of GABAA receptors in Fr1 is in accordance with that of previous reports (Zilles et al., 1990). Fr2 contains BZ binding site densities clearly higher than those of Fr1, and Fr3 shows the lowest BZ concentrations. Fr2 contains GABAB receptor densities slightly higher than those of Fr1 or Fr3. These two regions show comparable mean GABAergic densities (Fig. 1). Muscarinic cholinergic M1 and M3 receptors show higher densities in layers I–III than in layers V–VI, whereas M2 receptors (Fig. 7) present a bilaminar distribution pattern, with lower values in layers I–III than in layers V–VI (Zilles et al., 1990). The laminar distribution patterns of the cholinergic receptors reflect the laminar distribution of cholinergic axons and terminals (Eckenstein et al., 1988). Fr2 contains M1, M2, and M3 receptor densities slightly higher than those of Fr1 (Fig. 1). Fr1 and Fr3 show comparable M1 and M2 mean receptor densities (Fig. 1). Fr1 contains lower M3 receptor densities in layers I–II, but higher concentrations in layers V–VI, than Fr3. Nicotinic cholinergic receptors, as described for the muscarinic M2 receptors, also show alternating layers of intermediate and low densities. The lowest nicotinic concentrations are located in layers I, III, and VI, whereas layers II and V show intermediate values. Mean regional nicotinic receptor densities are higher in Fr2 than in Fr1 or Fr3 (Fig. 1). Of the examined receptor types, only the noradrenergic α1 receptors present differential laminar distribution patterns between the frontal regions. Furthermore, Fr1–3 also differ in their mean regional α1 and α2h receptor densities (Fig. 1). The α1 receptor densities in Fr1 are higher in layers I–II and V than in layers III and VI (Jones et al., 1985; Palacios et al., 1987). Fr2 contains the most α1 receptors, and these are located mainly in layer III. The α1 receptor densities in Fr3 are highest in layers I–II and diminish gradually throughout the cortex, reaching the lowest values in layer VI. The α2h receptor densities are highest in layers I–II and V and show no significant regional differences in their mean densities (Fig. 7). The frontal areas of the rat contain the highest cortical α1 receptor densities, which is in accordance with the presence of the highest noradrenaline concentrations in the frontal cortex and their gradual decrease throughout the rostrocaudal axis of the hemisphere, reaching the lowest values in the occipital cortex (Palkovits et al., 1979). Throughout the frontal cortex, the highest serotoninergic receptor densities are found in layer V. Layers I–III contain lower 5-HT1A receptor densities than layer VI. Conversely, layers I–III and VI do not differ in their 5-HT2 densities. Fr2 contains higher mean 5-HT1A

densities than of Fr1 or Fr3 (Fig. 1). The frontal regions do not differ significantly in their mean 5-HT2 densities (Fig. 7). The frontal areas Fr2 and Fr3 contain the highest cortical 5-HT1A receptor densities, which is in accordance with the presence of the highest concentrations of serotonin in the frontal cortex and a gradual decrease throughout the rostrocaudal axis of the hemisphere, reaching the lowest values in the occipital cortex (Reader, 1981).

Parietal Cortex Anterior Parietal Cortex The anterior parietal region of the rat occupies the dorsolateral middle third of the cerebral cortex, is characterized by the presence of a prominent inner granular layer IV, and has the strongest degree of myelination of all isocortical areas (Zilles, 1985, 1990; Zilles et al., 1980). It constitutes the somatosensory cortex of the rat (Welker, 1971, 1976; Welker and Sinha, 1972; Woolsey and leMessurier, 1948) and is bordered anteromedially by the frontal region, rostroventrally by the ventral parietal region, caudoventrally by the temporal region, and caudally by the occipital region. There are considerable differences concerning the number of areas within the parietal cortex. Krieg (1946a) defined six regions, which he named 1, 2, 3, 7, 39, and 40, following Brodmann’s (1909) numerical nomenclature of the human cortex. However, the existence of a direct homology between the human posterior parietal association regions and areas 7, 40, and 39 of the rat brain remains highly speculative. Swanson (1992, 1998) defined two areas, denominated SSp and SSs based on their functional properties (primary and supplementary somatosensory areas, respectively), and further described the existence of somatotopic subdivisions within the SSp. Swanson’s (1992, 1998) SSp and SSs correspond to Zilles and Wree’s (1985) primary (SmI) and secondary (SmII) somatosensory areas, respectively. However, in subsequent quantitative architectonical studies, these authors separated the forelimb area (FL) and the hindlimb area (HL) from Par1 based on significant differences in GLI values (Zilles and Wree, 1995). Thus, Par2 corresponds to SmII, FL and HL cover the dorsomedial part of SmI, and Par1 covers its dorso- and ventrolateral part (Zilles and Wree, 1995). Our area Par1 corresponds in position and topology to Area j of Droogleever Fortuyn (1914), whereas FL and HL together roughly correspond to his Area n. Furthermore, the topology and extent of our FL correspond to those of Welker (1971, 1976) and Donoghue and Wise (1982). Par1, FL, and HL represent the primary somatosen-

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sory cortex (Welker, 1971, 1976; Welker and Sinha, 1972; Woolsey and leMessurier, 1948). FL and HL, however, also exhibit architectonic (Donoghue et al., 1979) and functional (Donoghue and Wise, 1982; Donoghue et al., 1979; Hall and Lindholm, 1974; Wise and Jones, 1977) characteristics of a motor cortex. Par2 is the supplementary somatosensory cortex (Welker, 1971, 1976; Welker and Sinha, 1972; Woolsey and leMessurier, 1948). Cytoarchitectonically (Fig. 8), the parietal region is characterized by a highly developed inner granular layer IV, which is reflected in the highest GLI value determined in the whole cortex of the rat brain (Zilles, 1990). Structural differences in layer IV enable the delineation of Par1, Par2, FL, and HL. Layer IV of Par1 shows higher GLI values than of FL, HL, and Par2 (Zilles, 1990; Zilles and Wree, 1995). Furthermore, HL shows GLI higher values than those of FL (Zilles, 1990; Zilles and Wree, 1995). Par1 and Par2 can be clearly

FIGURE 8 Cyto- and myeloarchitectonical structure of the anterior parietal isocortical areas Par1, Par2, HL, and FL visualized in adjacent coronal sections. Roman numerals indicate cortical layers.

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delineated based on the pale layer Va of Par1, containing medium-sized pyramidal cell bodies of low packing density (Welker, 1971; Welker and Sinha, 1972; Zilles et al., 1980), and the higher GLI values of the supragranular layers of Par1 (Zilles and Wree, 1995), but higher GLI values in the infragranular layers of Par2 (Zilles, 1990). Par1, FL, and HL can be further differentiated from each other based on the size and packing density of their layer Vb pyramids. The pyramids of FL are more densely packed than those of Par1 and are even larger and more densely packed in HL (Droogleever Fortuyn, 1914; Zilles and Wree, 1995). The somatotopic subdivisions of the parietal cortex, which have been revealed by electrophysiological studies (Chapin and Lin, 1990; Donoghue and Parham, 1983; Donoghue and Wise, 1982), are also reflected in differences in the densities of receptors for classical neurotransmitters (Figs. 5–7 and 9). This is particularly true for Par1, where architectonical differences in layer IV also enable the determination of subareas. Layer IV has a “cloudy” appearance (Droogleever Fortuyn, 1914) in the posterior part of Par1, where vertically oriented cell-dense patches are surrounded by narrow dysgranular strips. This subarea corresponds to the barrel field, which is the cortical representation of the mystacial vibrissae (Welker, 1971, 1976; Welker and Sinha, 1972; Woolsey and leMessurier, 1948). The remaining part of Par1 is arranged in granular regions surrounded by perigranular and dysgranular regions (Chapin and Lin, 1990; Donoghue and Wise, 1982), defined according to their higher or lower packing density of the small cell bodies in layer IV. These regions also differ in their functionality. The granular and perigranular regions show a unimodal, finely detailed, map of the cutaneous representation, whereas the dysgranular regions exhibit a multimodal convergence of information from proprioceptors located in the skin and joints (Chapin and Lin, 1984). Some studies describe the most rostral part of our Par1 as a representation of the mouth and nose (Swanson, 1992, 1998) and the most caudal part of our Par1 as a representation of the trunk and tail (Hall and Lindholm, 1974; Welker, 1971, 1976). Quantitative 2-DG autoradiography was applied in order to register the regional and laminar LCGU patterns within the parietal region, revealing slightly higher values in Par2 than in Par1 as well as high values in all layers of FL and HL with the exception of layer VI (Zilles and Wree, 1995). The parietal region is characterized by high AMPA and NMDA receptor densities in the supragranular layers (Fig. 7). Conversely, kainate concentrations are highest in the infragranular layers (Figs. 5–7 and 9). The laminar distribution pattern of kainate receptors in the parietal cortex reflects the distribution of zinc-containing

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FIGURE 9 Neighboring coronal cryostat sections (15 μm thick) through the rat brain at two different rostrocaudal levels (A, B, and C rostral to D, E, and F) processed for silver cell-body (A, D) and myelin (B, E) staining as well as for the visualization of glutamatergic kainate receptors (C, F) by means of [3H]kainate. Asterisks indicate sectioning artifacts. For further details, see Fig. 5. See Table 1 for abbreviations.

vesicles revealed by the Timm stain (Zilles et al., 1990). Par1 contains the lowest mean regional glutamatergic receptor densities, particularly in the infragranular layers. Par2 shows the highest AMPA and NMDA receptor densities, especially in the supragranular layers,

whereas the highest kainate concentrations were measured in HL (Figs. 5–7 and 9). The border between FL and HL is visible due to the slightly higher AMPA and NMDA densities in the infragranular layers of HL as well as its overall higher kainate densities (Figs. 5–7 and 9).

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GABAergic receptors and BZ binding sites are present in higher concentrations the supragranular layers than in the infragranular layers of the parietal region. Par1 contains the lowest infragranular GABAA and GABAB receptor concentrations, and HL the highest values (Fig. 7). Mean BZ binding site concentrations are highest in HL, and diminish throughout FL and Par1, reaching the lowest values in Par2. This preferential location of GABAA receptors in the supragranular layers (Fig. 7), in particular the high densities in layer IV, is in accordance with the presence of numerous GABA-containing presynaptic terminals in layers II–IV (Chmielowska et al., 1988). The laminar distribution patterns of the muscarinic and nicotinic cholinergic receptors differ from each other in the parietal cortex. M1 and M3 receptors show high densities in the supragranular layers and significantly lower values in the infragranular layers. Muscarinic M2 and the nicotinic receptors show alternating bands of high and low densities (Fig. 7). Layers I–II and VI contain intermediate M2 receptor densities, layers III–IV contain the highest densities, and layer V contains the lowest concentrations. Nicotinic receptors are present in lower concentrations in layers I–II and V than in layers III–IV. Layer VIa contains the lowest nicotinic receptor densities, whereas the concentrations in layer VIb are comparable to those of layers I–II. The laminar distribution patterns of the cholinergic receptors reflect the laminar distribution of cholinergic axons and terminals (Eckenstein et al., 1988). Par1, Par2, FL, and HL contain comparable M1 receptor densities. Conversely, they differ in their infragranular concentrations of M3 receptors, with the highest values located in Par2 and HL and the lowest density in Par1. Par2 contains lower nicotinic receptor densities in the supragranular layers but higher ones in the infragranular layers than Par1 (Fig. 7). HL and FL do not differ in their mean nicotinic receptor densities or laminar distribution patterns (Fig. 7). The regional distribution pattern of M2 receptor densities in the rat brain differs significantly from that of primate brains. M2 receptors are present in significantly higher densities the primary visual, auditory, and somatosensory cortices of humans and macaque monkeys than in the neighboring secondary sensory and multimodal association cortices (Zilles et al., 2002a). Although the rat brain does not display this specific regional distribution pattern of exceptionally high M2 receptor densities in primary sensory areas—rather, all isocortical regions contain comparable mean M2 receptor concentrations—it does present a singular laminar distribution pattern, since layer IV of Par1 contains M2 significantly higher receptor densities than the adjoining cortices (Fig. 7).

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The noradrenergic α1 and α2h receptors show alternating layers of high and low densities throughout the four areas of the parietal region (Fig. 7). Layers I–III contain considerably higher α1 receptor densities than layers IV–VI. Layer V has slightly higher concentrations than the adjacent layers. The α1 receptor densities of layer VI diminish gradually, reaching the lowest values at the border with the white matter. The α1 receptor densities in layers I–II and IV–V are higher than those in layers III and VI. The α2h receptor densities are high in layers I–II and IV, low in layer III, and reach the lowest values in layers V–VI (Fig. 7). HL contains the highest and the Par1 lowest α1 receptor densities, particularly in the infragranular layers. Par2 contains the highest and Par1 the lowest α2h receptor densities. HL and FL do not differ in their mean α2h receptor concentrations. Throughout the parietal cortex, the highest serotoninergic 5-HT1A receptor densities are found in the infragranular layers, whereas the highest 5-HT2 concentrations are restricted to layer IV (Fig. 7). Furthermore, layers I–III contain higher 5-HT2 receptor densities than layers V–VI in Par1, HL, and FL. Conversely, layers I–III of Par2 show lower 5-HT2 receptor densities than layers V–VI. The infragranular layers of Par1 and Par2 have lower 5-HT1A receptor densities than HL or FL. HL and FL do not differ significantly in their mean 5-HT1A receptor densities. 5-HT2 receptor densities are highest in Par2. Par1, HL, and FL cannot be clearly delineated from each other based on their 5-HT2 receptor distribution patterns or densities (Fig. 7). For further information concerning connectivity and functionality of the somatosensory areas, see Chapter 25. Ventral Parietal Cortex The ventral part of the parietal cortex, abutting the insular cortex, shows a dysgranular isocortical structure (Fig. 10) and was, therefore, separated from the ventrolaterally adjacent agranular insular cortex (Zilles, 1990; Zilles and Wree, 1995). This area was originally designated visceral cortex (Vi) by Zilles and Wree (1995), since they considered that it may correspond to a visceral cortical field implicated in functions as varied as taste (Ogawa et al., 1990, 1991; Yamamato et al., 1990), visceral motility (Allen et al., 1991; Lasiter and Glanzmann, 1983; Yasui et al., 1991), and cardiovascular functions (Oppenheimer et al., 1991). Adhering to the principle of a neutral nomenclature, we have now replaced the term visceral cortex with the designation parietal ventral area (ParV). Based on differences in mean regional GABAB, α1, and 5-HT1A receptor densities as well as in BZ binding site concentrations, we have subdivided ParV into a rostral (ParVR) and a

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caudal (ParVC) part. ParVR is bordered dorsally by Par1, whereas ParVC adjoins Par2. Ventrally, both areas are delimited by the agranular insular cortex. Our definition of ParVR and ParVC is compatible with Swanson’s (1992, 1998) gustatory and visceral areas, respectively. Furthermore, our definition of ParV is also compatible with the description of a “gustatory insular cortex” (Benjamin and Akert, 1959; Benjamin and Pfaffmann, 1955; Braun et al., 1982; Guldin and Markowitsch, 1983; Lasiter et al., 1982; Van der Kooy et al., 1982; Wolf, 1968), located between the insular cortex at the ventral borders of Par1 and Par2. The ventral parietal cortex can be clearly delineated from the agranular insular cortex and the anterior

FIGURE 10 Cyto- and myeloarchitectonical structure of the ventral parietal isocortical areas ParVR and ParVC visualized in adjacent coronal sections. Roman numerals indicate cortical layers.

parietal cortices based on the mean regional densities and laminar distribution patterns of receptors for classical neurotransmitters (Figs. 5–7). ParVR and ParVC do not differ in their mean regional glutamatergic receptor densities. The AMPA and NMDA receptors are present in higher densities in the supragranular layers of ParVR and ParVC than in the infragranular ones (Fig. 7), whereas kainate receptors show the opposite laminar distribution pattern (Figs. 5–7). The GABAergic receptors and the BZ binding sites are present in higher concentrations in the supragranular layers of ParVR and ParVC than in the infragranular ones. Although these two regions do not differ in their mean GABAA receptor densities, ParVR contains significantly higher GABAB receptor and BZ binding site densities than of ParVC. The muscarinic cholinergic M1 and M3 receptors are present in higher densities in the supragranular layers of ParV than in the infragranular ones. M2 receptors show the highest concentrations in layers IV and VI and the lowest values in layers I–III and V (Fig. 7). The highest nicotinic cholinergic densities are present in layer IV, and the lowest concentrations are in layers I–III (Fig. 7). As in the case of the glutamatergic receptors, the mean regional densities and laminar distribution patterns of muscarinic and nicotinic cholinergic receptors do not highlight the border between ParVR and ParVC. The noradrenergic α1 receptors of ParVR and ParVC reach their highest densities in layers I–II and IV–V, whereas α2h receptors reach maximal values in the supragranular layers (Fig. 7). ParVR contains higher α1 receptor densities than ParVC. These two regions do not differ in their mean α2h receptor densities.

FIGURE 11 Cyto- and myeloarchitectonical structure of the posterior parietal isocortical areas ParPD, ParPR, and ParPC visualized in adjacent coronal sections. Roman numerals indicate cortical layers.

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The serotoninergic 5-HT1A receptors are found in higher densities in layers IV–VI of ParVR and ParVC than in layers I–III. The 5-HT2 receptors are present at high concentrations in layer IV and at intermediate values in the remaining layers (Fig. 7). ParVR contains higer 5-HT1A receptor densities than ParVC. These two regions cannot be delineated on the basis of their mean 5-HT2 receptor concentrations or laminar distribution patterns. For additional connectivity and functional considerations of ParV, see Chapter 29. Posterior Parietal Cortex Krieg (1946a) described the existence of three posterior parietal regions, Areas 7, 39, and 40, which he delineated from the adjacent somatosensory and visual cortices due to their thinner layers I–III than the somatosensory cortex and more myelinated fibers than the visual cortex. Based on differences in myelination and acetylcholinesterase staining intensity, Kolb (1990) delineated a posterior parietal region, PPC, comparable to Krieg’s (1946a) Area 7. Pérez-Clausell (1996) applied a staining method visualizing zinc in the terminal fields of the rat neocortex and defined two posterior parietal areas, PPC and Par2P. Paxinos and coworkers (1999) delineated two posterior parietal areas, PtA and MPtA, which topographically coincide with the most posterior portions of our Fr1, Fr2, and HL and with the most anterior parts of our Oc2MM and Oc2ML. Furthermore, Swanson (1992, 1998) defined the existence of posterior parietal areas located between the unimodal somatosensory and visual areas and receiving inputs from the lateral posterior thalamic nucleus, which he grouped under the term PTLp. However, other authors have described putative visual areas in this position (Schober, 1986; Zilles, 1985, 1990; Zilles and Wree, 1995; Zilles et al., 1980), although Zilles (1990) did not discard that part of the visual areas in general, and Oc2L in particular may be implicated in multimodal processing. Furthermore, recent functional findings support the existence of a distinct visual processing route in the rat posterior parietal cortex involved in visuospatial guidance (DiMatia and Kesner, 1988; Kametani and Kesner, 1989; Kolb et al., 1983; Save and Poucet, 2000) and cross-modal matching (PintoHamuy et al., 1987). Based on the regional and laminar distribution patterns of neurotransmitter receptors (Figs. 9 and 12), we have subdivided Oc2L, which was described as being cytoarchitectonically heterogeneous (Zilles and Wree, 1995), along its rostrocaudal axis into a dorsal part, for which we maintain the designation of Oc2L (see below), and a ventral part, which we classify as posterior parietal cortex (ParP). These two areas are difficult to

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delineate based on cyto- and myeloarchitectonical criteria (Fig. 11). Layer IV of ParP is not as conspicuous as that of Oc2L, and cell packing density in layer V of ParP is slightly higher than that in Oc2L. Furthermore, based also on neurochemical differences (Figs. 9 and 12), we have further subdivided ParP into three subareas, which we have designated ParPD (posterior parietal cortex, dorsal part), ParPR (posterior parietal cortex, rostral part), and ParPC (posterior parietal cortex, caudal part). Topographically, ParPD and ParPR could correspond to the lateral part of Krieg’s (1946a) Area 7, to area Par2P of Pérez-Clausell (1996), and to the rostrolateral lateral part of Swanson’s (1992, 1998) PTLp areas, whereas ParPC covers the caudal part of Swanson’s (1992, 1998) PTLp. The posterior parietal cortex receives extensive afferents from the central lateral, ventrolateral (Giannetti and Molinari, 2002), lateral dorsal, and lateral posterior thalamic nuclei (Giannetti and Molinari, 2002), but none from the ventrobasal complex or the dorsal geniculate nucleus (Chandler et al., 1992; McDaniel et al., 1978). Corticocortical connections of the posterior parietal cortex were determined by means of retrograde degeneration and axonal tracing techniques. They include efferents to as well as afferents from the retrosplenial cortex, Fr2, and Oc2M (Corwin and Reep, 1998; Kolb and Walkey, 1987; Reep et al., 1994). Furthermore, the posterior parietal cortex receives afferents from Te1, Par1, Par2, VLO, and MO (Kolb and Walkey, 1987; Reep et al., 1994). ParPD, ParPR, and ParPC have comparable laminar distribution patterns of the examined receptors for classical neurotransmitters. They differ, however, in their mean receptor densities, particularly those of the GABAB receptors. ParPD contains higher AMPA concentrations than ParPR, but lower values than ParPC. ParPD contains slightly higher kainate receptor densities than ParPR, and clearly lower concentrations than ParPC (Figs. 9 and 12). The lowest NMDA receptor densities were measured in ParPD, whereas the highest values were located in ParPC. Although the posterior parietal areas do not differ significantly in their mean GABAA receptor densities, they show their most conspicuous differences regarding the GABAB receptors, which are present at the lowest concentrations in ParPC. ParPR contains the lowest and ParPC the highest BZ binding site densities. Posterior parietal regions contain comparable mean M1, M2, and nicotinic cholinergic receptor densities. ParPD and ParPR do not differ in their mean M3 receptor densities, but contain lower concentrations than ParPC. ParPR shows higher α1 receptor densities than either ParPD or ParPC. These three areas do not differ in their noradrenergic α2h or serotoninergic 5-HT1A and 5-HT2 receptors.

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FIGURE 12 Neighboring coronal cryostat sections (15 μm thick) through the rat brain at two different rostrocaudal levels (A, B, and C rostral to D, E, and F) processed for silver cell-body (A, D) and myelin (B, E) staining as well as for the visualization of glutamatergic kainate receptors (C, F) by means of [3H]kainate. Asterisks indicate sectioning artifacts. For further details, see Fig. 5. See Table 1 for abbreviations.

Temporal Cortex The cortex of the temporal region is thinner than that of the parietal region, in particular concerning layers V and VI. It has been the subject of numerous

connectivity and architectonical studies (Arnault and Roger, 1990; Droogleever Fortuyn, 1914; Krieg, 1946a, 1946b; Miller and Vogt, 1984; Roger and Arnault, 1989; Schober, 1986; Swanson, 1992, 1998; Zilles and Wree, 1985, 1995; Zilles et al., 1980). The resulting maps show

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FIGURE 13 Cyto- and myeloarchitectonical structure of the temporal isocortical areas Te1, Te2, Te3, and TeV visualized in adjacent coronal sections. Roman numerals indicate cortical layers.

both similarities and discrepancies with our present parcellation scheme. Based on differential patterns of thalamic and callosal input, a division of the rat auditory cortex into a primary auditory field, or “core cortex”, and a nonprimary auditory belt, or “belt cortex”, was proposed. This parcellation scheme was in accordance with an earlier subdivision of the temporal cortex into two regions, Te1 and Te2, by Zilles et al. (1980). However, later studies with improved measuring techniques led to further subdivisions (Zilles and Wree, 1985), resulting in the parcellation of the temporal region into three areas (Te1–3), with areas Te2 and Te3 forming a ring-like belt around Te1 (Zilles and Wree, 1985). Te2 can be further subdivided into a dorsal (Te2D) and a ventral (Te2V) part (Zilles and Wree, 1995). Te3 can be subdivided into a rostral (Te3R)

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and a ventral (Te3V) part (Zilles and Wree, 1995). Since the identification of these borders was based on differences in the degree of myelination which are visible on tangential sections through the lateral and dorsal surface of the hemisphere in flattened tissue preparations, and not on cytoarchitectonical characteristics, they were not indicated by solid lines (Zilles and Wree, 1995). However, the subdivision of Te2 and Te3 is further supported by differences in the mean densities and laminar distribution patterns of receptors for classical neurotransmitters. Area p of Droogleever Fortuyn (1914) topologically resembles our areas Te1–3, but also seems to cover parts of Par1 and Par2. Krieg (1946a, 1946b) and Miller and Vogt (1984) delineated four areas within the temporal region—Areas 20, 36, 39, and 41—in positions comparable to those on our map. Their Area 41 is comparable to our Te1, but this is not true for the other areas. Swanson (1992, 1998) defined three auditory regions, AUDd, AUDp, AUDv, which he delineated as three rostrocaudally oriented bands running parallel to each other. Our parcellation scheme is in agreement with the subdivision of the temporal region into a “core cortex”, comparable to our Te1, and a “belt cortex”, consisting of two areas, comparable to our Te2–3, as proposed by Arnault and Roger (1990) and Roger and Arnault (1989). Our parcellation scheme is further supported by the observations of Schober (1986). Swanson (1992, 1998) delineated a field within the temporal cortex of the rat brain, which he designated ventral temporal association areas (TEv). He considered TEv may be homologous to the ventral temporal association areas located on the dorsal, middle, and inferior temporal gyri of the human brain (Swanson, 1992, 1998). Furthermore, Paxinos and co-workers (1999) defined their TeA at a comparable topographical location. Dorsally, TEv is delimited by the auditory and visual cortices and ventrally by the ectorhinal area (Swanson, 1992, 1998). Based on the distribution patterns of neurotransmitter receptors, we have delineated a cortical region (TeV) with a topographical location similar to that described by Swanson (1992, 1998) for his area TEv. Te1 has been identified as the primary auditory cortex (Cipolloni and Peters, 1979; Krieg, 1947; leMessurier, 1948), and Te2–3 are the secondary regions (Cipolloni and Peters, 1979; Guldin and Markowitsch, 1983). TeV has been only sparcely examined. It has been classified as a multimodal cortex and is involved in visual discrimination learning tasks (Kolb et al., 1994; Wortwein et al., 1994). The connectivity pattern of Swanson’s (1992, 1998) TEv resembles that of the monkey’s inferotemporal association cortex (Kolb, 1990) in that it receives

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FIGURE 14 Cyto- and myeloarchitectonical structure of the occipital isocortical areas Oc1M, Oc1B, Oc2MM, Oc2ML, and Oc2L visualized in adjacent coronal sections. Roman numerals indicate cortical layers.

projections from the posterolateral thalamic nucleus (Mason and Gross, 1981), from visual cortical areas (Miller and Vogt, 1984), and from the entorhinal cortex (Kosel et al., 1982) and projects to the perirhinal cortex (Deacon et al., 1983). Te1–3 differ in their architectonical patterns (Fig. 13). Te2–3 have a lower content of myelinated fibers than Te1. Furthermore, layer IV of Te1 is more prominent and has higher GLI values than layer IV of Te2–3 (Zilles and Wree, 1995). The presence of a welldeveloped and prominent layer IV is a characteristic shared by all primary sensory areas (Par1, Te1, and Oc1). Within Te1, the supragranular layers show a higher cell packing density than the infragranular ones. Layers V–VI of Te1 have a lower packing density than that of Te2–3. Layer VI of Te2 is narrower than that of Te3. Te1 is characterized by an exceedingly high local glucose metabolism (Sokoloff et al., 1977; Zilles and Wree, 1995). Te2–3 have lower LCGU values than Te1,

but higher ones than the adjoining cortical regions (Zilles and Wree, 1995). The temporal areas have different mean regional receptor densities and laminar distribution patterns. The glutamatergic AMPA and NMDA receptors are present in high densities in the supragranular layers and in low densities in the infragranular layers throughout the temporal region. Conversely, the kainate receptors are present in higher concentrations in the infragranular layers than in the supragranular ones (Figs. 9 and 12). The laminar distribution pattern of kainate receptors in the temporal cortex reflects the distribution of zinccontaining vesicles revealed by the Timm stain (Zilles et al., 1990). Te1, Te3R, Te3V, and TeV show comparable mean AMPA receptor densities, but values lower than those measured in Te2C or in Te2D, which is the area containing the highest AMPA receptor densities. Te1, Te3R, and Te2D do not differ in their mean kainate receptor densities (Figs. 9 and 12). Layer V of Te3V contains higher kainate concentrations than Te1, but

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lower than those of TeV (Figs. 9 and 12). Te2V shows higher kainate densities than T3V, but lower ones than TeV (Fig. 12). All temporal areas contain comparable mean NMDA receptor densities and laminar distribution patterns. The GABAergic receptors are present in higher densities in the supragranular layers of the temporal cortex than in the infragranular layers. The BZ binding sites show alternating bands of high and low densities throughout the temporal cortex, with significantly higher concentrations in layers IV and VI than in layers I–III and V. TeV contains slightly higher GABAA receptor densities than the remaining areas of the temporal cortex, which cannot be delineated from each other based on their mean GABAA receptor concentrations or laminar distribution patterns. Conversely, the mean GABAB receptor densities vary considerably between the temporal areas. TeV shows the highest measured GABAB densities. Te1 contains lower GABAB concentrations than Te3R and Te3V, but significantly higher ones than Te2D or Te2C. Neither Te3R and Te3V nor Te2D and Te2C differ in their mean GABAB concentrations. Te1 and Te3V show comparable BZ densities, which are significantly lower than those measured in Te2C, Te3R, or TeV, which do not differ in their mean BZ binding site densities either. Muscarinic cholinergic M1 and M3 receptors are present in higher concentrations in the supragranular layers than in the infragranular layers of the temporal region. In all temporal areas, with the exception of TeV, with the highest values in the infragranular layers, M2 receptors show higher densities in layers IV and VI in layers I–III and V. Nicotinic cholinergic receptors show the highest densities in layers III–IV of Te1, Te3R, and Te3V, but are homogeneously distributed throughout all layers of Te2D, Te2C, and TeV. The laminar distribution patterns of the cholinergic receptors reflect the laminar distribution of cholinergic axons and terminals (Eckenstein et al., 1988). TeV contains higher M1 receptor densities in the infragranular layers than Te2D or Te2C, but lower concentrations in the supragranular layers than Te1, Te3R, or Te3V. As described for the parietal cortex, the M2 receptors present a specific laminar distribution pattern in the temporal cortex, since layer IV of Te1 contains significantly higher M2 receptor densities than in the adjoining regions. Te3R, Te3V, Te2D, and Te2C contain comparable mean nicotinic receptor densities, which are lower than those measured in Te1. This is specially true for the layer IV densities. Laminar distribution patterns and mean regional densities of the noradrenergic α1 and α2h receptors vary throughout the temporal region. The α1 receptors are

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homogeneously distributed throughout all layers of Te2D and Te2C, but show the highest densities in the supragranular layers of Te1, Te3R, and Te3, as well as in layer IV of TeV. Despite these differences in their laminar distribution patterns, all temporal regions show comparable mean α1 receptor densities. The α2h receptors are present in highest densities in layer IV and in lowest concentrations in layers V–VI of the temporal cortex. Te1 and Te3R contain the lowest receptor densities and TeV contains the highest α2h receptor densities. Te3V and Te2D contain comparable α2h concentrations, which are lower than those found in Te2C. The highest serotoninergic 5-HT1A receptor densities of the temporal region are located in layer V, and layer IV contains the highest 5-HT2 receptor concentrations. The lowest 5-HT1A receptor densities are located in Te1 and Te3V, whereas the highest concentrations are in Te2D, Te2C, and TeV. Te2D, Te2C, Te3R, Te3V, and TeV contain comparable 5-HT2 receptor densities, which are slightly higher than those measured in Te1. For further detailed considerations of the rat auditory cortex, see Chapter 31.

Occipital Cortex The occipital region of the rat contains a conspicuous inner granular layer which is in accordance with its sensory function. The occipital region has been the subject of numerous studies based on anatomical, physiological, and behavioral techniques (Adams and Forrester, 1968; Droogleever Fortuyn, 1914; Krieg, 1946a, 1947; Montero, 1973, 1981; Montero et al., 1973a, 1973b; Ribak and Peters, 1975; Swanson, 1992, 1998; Thomas and Espinoza, 1987; Winkelmann et al., 1972; Zilles, 1985; Zilles et al., 1980, 1984). Despite disagreements concerning the number, location, and extension of separate visual areas, it is accepted that the visual cortex of the rat is not homogeneous, but seems to be organized in a modular fashion, containing various distinct representations of visual space (Dean, 1990). Electrophysiological and connectivity studies (Dean, 1990; Montero, 1973; Montero et al., 1973a, 1973b; Swanson, 1992, 1998; Thomas and Espinoza, 1987) reveal a complex pattern of distinct visuotopically organized areas. These areas, however, do not differ significantly in their architectonical structure. Therefore, studies based on cyto- or myeloarchitectonical criteria reveal a much simpler parcellation scheme (Schober, 1986; Zilles, 1985, 1990; Zilles and Wree, 1985, 1995). Zilles and coworkers (Zilles, 1985; Zilles et al., 1980, 1984) created a comprehensive cyto- and myeloarchitectonic map in which the occipital region of the rat brain was divided into four areas, denominated Oc1, Oc2MM, Oc2ML, and

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Oc2L. Oc1, the primary visual cortex, is surrounded at its rostromedial border by the secondary visual areas Oc2MM and Oc2ML and at its rostrolateral border by the secondary visual area Oc2L (Zilles, 1990). Intraocular injection of [3H]proline and its subsequent transneuronal transport enabled the subdivision of Oc1 into the monocular (Oc1M, covering the medial part of Oc1) and binocular (Oc1B, covering the lateral part of Oc1) subfields (Zilles et al., 1984). Oc1M and Oc1B can also be differentiated based on their LCGU levels as well as on their cyto- and myeloarchitecture (Zilles et al., 1984). Oc2L was described as being a cytoarchitectonically heterogeneous area composed of several subareas (Zilles and Wree, 1995). This cytoarchitectonical heterogeneity (Figs. 11 and 14) is mirrored by a chemoarchitectonical heterogeneity, based on quantitative receptor autoradiography (Figs. 9 and 12). Thus, Oc2L of Zilles and Wree (1995) can be subdivided into a dorsomedial part, for which we maintain the designation of Oc2L, and a ventrolateral part, which we have included here in the posterior parietal cortex (see above). Droogleever Fortuyn (1914) defined a single visual region, Area w, which corresponds approximately to the total of our visual areas. It is difficult to compare our map of the visual cortex with that of Krieg (1946a, 1946b), since in his map the primary visual cortex does not reach the occipital contour of the hemisphere, this being occupied by the caudal part of his Area 18a. However, our delineation of Oc1 is not only in accordance with other architectonical maps (Ribak and Peters, 1975; Winkelmann et al., 1972; Zilles, 1985; Zilles et al., 1980, 1984), but has also been supported by means of electrophysiological studies (Adams and Forrester, 1968; Montero, 1973, 1981; Thomas and Espinoza, 1987). Krieg’s (1946a, 1946b) Area 18 resembles our Oc2ML and Oc2MM. Cytoarchitectonically (Fig. 14), the visual cortex is characterized by a conspicuous layer IV, though it is not as prominent as that of the somatosensory cortex. Within the visual cortex, layer IV shows the highest cell packing density in Oc1. Furthermore, although layer V of Oc1 has lower GLI values than Oc2M or Oc2L, Oc1 shows an overall higher cell packing density than the adjoining visual areas (Zilles, 1990; Zilles and Wree, 1995). Oc1 is further characterized by a more prominent layer IV than Oc2MM, Oc2ML, or Oc2L (Zilles and Wree, 1995). Within Oc1, GLI values of layers IV and V in Oc1M are higher than those in Oc1B (Zilles et al., 1984). The different regions of the occipital cortex can be further differentiated based on their LCGU levels. The primary visual areas contain clearly higher LCGU levels, particularly in layer IV, than the adjoining secondary regions, and Oc2L shows higher LCGU

values than the medial (Oc2ML and Oc2MM) secondary visual areas (Zilles and Wree, 1995). Oc2MM shows the lowest LCGU levels throughout the occipital cortex (Zilles and Wree, 1995). The glutamatergic AMPA and NMDA receptors are present higher densities in in the supragranular layers than in the infragranular ones throughout the visual cortex. The kainate receptors, conversely, show maximal concentrations in the infragranular layers (Figs. 9 and 12). The laminar distribution pattern of kainate receptors in the occipital cortex reflects the distribution of zinc-containing vesicles revealed by the Timm stain (Zilles et al., 1990). Oc1M contains only slightly higher AMPA and kainate, but lower NMDA recepter densities than Oc1B (Figs. 9 and 12). Oc1M contains slightly lower AMPA, kainate (Figs. 9 and 12), and NMDA receptor densities, particularly in the infragranular layers, than Oc2ML. Oc1B contains lower AMPA and NMDA receptor densities, particularly in layer IV, than Oc2L. Furthermore, Oc1B shows lower kainate receptor densities in the supragranular layers than Oc2L (Fig. 12). Oc2MM and Oc2ML do not differ significantly in their mean AMPA, kainate, or NMDA receptor densities or laminar distribution patterns. GABAB receptors are present in higher densities in the supragranular layers than in the infragranular layers of the occipital cortex. GABAA receptors and BZ binding sites are also present in highest densities in the supragranular layers of Oc1M, Oc1B, and Oc2L, but are homogeneously distributed throughout all layers of Oc2MM and Oc2ML. GABAA receptor densities are higher in the supragranular layers but lower in the infragranular layers of Oc1M, Oc1B, and Oc2L than in Oc2MM or Oc2ML. Therefore, mean regional GABAA receptor densities are comparable in all occipital areas. Oc2MM and Oc2ML contain clearly higher GABAB receptor and BZ binding site densities, particularly in the infragranular layers, than Oc1M, which shows the same mean GABAB density as Oc1B, but higher BZ concentrations. Oc2L contains lower GABAB but higher BZ concentrations than Oc1B, particularly in the infragranular layers. The described laminar distribution pattern of GABAA receptors is in accordance with the distribution of the neurotransmitter GABA, which was determined enzymatically in the occipital cortex of the rat (Ishikawa et al., 1983). Furthermore, as described for the somatosensory cortex, the high GABAA receptor densities in layer IV are correlated with the laminar distribution of GABAergic terminals (Lin et al., 1986). Muscarinic cholinergic M1 and M3 receptors are present in higher densities in the supragranular layers than in the infragranular ones throughout the occipital

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cortex. M2 and nicotinic receptors are present at highest concentrations in layer IV, followed by intermediate values in layer VI and low values in layers I–III and V. Nicotinic receptors are also present at highest concentrations in layer IV of the occipital cortex, but show comparable values in the remaining layers. The laminar distribution patterns of the cholinergic receptors reflect the laminar distribution of cholinergic axons and terminals (Eckenstein et al., 1988). Oc1M and Oc1B do not differ in their mean M1 concentrations or laminar distribution patterns. However, they contain lower M1 receptor densities in the infragranular layers than Oc2L, Oc2MM, or Oc2ML. The latter regions also show similar mean M1 receptor concentrations and laminar distribution patterns. As described for the primary somatosensory and auditory cortices, the primary visual cortex (Oc1M and Oc1B) can be distinguished from the adjacent secondary areas due to the significantly higher M2 receptor densities in layer IV of Oc1M and Oc1B. However, overall M2 receptor densities do not vary throughout the occipital cortex, due to the higher M2 concentrations in the infragranular layers of Oc2MM, Oc2ML, and Oc2L than in the infragranular layers of Oc1M or Oc1B. The areas of the occipital cortex do not differ significantly in their mean regional M3 or nicotinic receptor densities, nor in their laminar distribution patterns. Noradrenergic α1 receptors are present in higher densities in layers I–III and V of the occipital cortex, whereas α2h receptors show higher values in layers I–II and IV. Oc2MM and Oc2ML contain comparable α1 and α2h receptor densities, which are slightly higher than those measured in Oc1M or Oc1B. Although Oc2MM and Oc2ML contain higher α1 receptor densities than Oc2L, these three regions do not differ in their mean α2h concentrations. The occipital areas of the rat contain the lowest cortical α1 receptor densities, which is in accordance with the presence of the highest noradrenaline concentrations in the frontal cortex, and their gradual decrease throughout the rostrocaudal axis of the hemisphere, reaching the lowest values in the occipital cortex (Palkovits et al., 1979). The serotoninergic 5-HT1A receptors are present in higher densities in the infragranular layers than in the supragranular ones throughout the occipital region, with the exception of Oc2MM, which has overall high 5-HT1A concentrations. The 5-HT2 receptors are present at high concentrations in layer IV and in intermediate values in the remaining layers. Oc2MM shows the highest 5-HT1A receptor concentrations, followed by Oc2ML, whereas the lowest values were measured in Oc1M, Oc1B, and Oc2L. The occipital areas do not differ in their mean 5-HT2 receptor densities. The occipital

areas of the rat contain the lowest cortical 5-HT1A receptor densities, which is in accordance with the presence of the highest concentrations of serotonin in the frontal cortex, and a gradual decrease throughout the rostrocaudal axis of the hemisphere, reaching the lowest values in the occipital cortex (Reader, 1981). See Chapter 32 for additional consideration of the visual cortex.

TRANSITION REGIONS BETWEEN ISOCORTEX AND ALLOCORTEX Orbitofrontal Cortex The orbitofrontal region has a periallocortical laminar structure and was defined as the cortical projection area of the mediodorsal thalamic nucleus by means of retrograde degeneration studies (Divac, 1972; Krettek and Price, 1977a; Reep et al., 1996). It covers the medial, basal, and lateral surfaces of the frontal contour of the hemisphere and can be clearly delineated from the laterally adjoining insular cortex and the medially adjoining cingulate region due to their significantly lower LCGU levels (Zilles and Wree, 1995). Four areas can be delineated within the orbitofrontal cortex on horizontal or sagittal sections: the medial orbital area (MO), the ventral orbital area (VO), the ventrolateral orbital area (VLO), and the lateral orbital area (LO). These regions show differential connectivity patterns. MO receives projections from the hippocampal formation (Thierry et al., 2000), from the anteromedial thalamic nucleus (Van Groen et al., 1999), and from the cingulate, frontal (Fr2), and posterior parietal cortices (Reep et al., 1996). VO receives corticocortical afferents from the cingulate cortex and from Fr2, Par2, ParP, Oc2M, and Oc2L (Reep et al., 1996). VLO receives cortical input from the postrhinal (Delatour and Witter, 2002) and insular cortices as well as from Fr2, Par1, Par2, ParP, and Oc2L (Reep et al., 1996). LO receives afferents from the insular cortex and from Par2 (Reep et al., 1996). The connectivity pattern of VLO supports the hypothesis that this region is implicated in directed attention and allocentric spatial localization tasks (Reep et al., 1996). Furthermore, the orbital cortex of the rat is thought to play a crucial role in olfactory sensory processing and in odor-guided motivational behaviors (Yonemori et al., 2000). The orbitofrontal areas differ in their mean densities of receptors for classical neurotransmitters. The glutamatergic AMPA and NMDA receptors are present in higher densities in layers I–III of the orbitofrontal cortex, whereas maximal kainate receptor concentrations are located in layers V–VI (Fig. 5). MO and VO

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show the highest AMPA, kainate, and NMDA densities within the orbitofrontal cortex, whereas LO and VLO contain the lowest values. The GABAergic receptors are present in higher densities in layers I–III and V of the orbitofrontal cortex. The highest BZ binding site densities are present in layer V. The orbitofrontal areas contain comparable mean GABAA receptor concentrations. MO contains the highest and VLO and LO the lowest GABAB receptor densities. MO contains higher BZ binding site densities in layers I–III, but lower concentrations in layers V–VI than VO, thus resulting in comparable mean receptor densities. VO and VLO do not differ in their mean BZ densities, which are lower than those measured in MO and VO. The muscarinic cholinergic M1 and M3 receptors are present in highest densities in layers I–III of the orbitofrontal cortex. The M2 receptors show high densities in layers V–VI, and the highest nicotinic cholinergic receptor concentrations are found in layers I–II and V. Mean muscarinic receptor densities are slightly higher in MO and VO than in the lateral orbital areas. VLO and LO contain higher nicotinic receptor concentrations in layer V, but lower values in layer VI, than MO or VO. The noradrenergic α1 receptors are homogeneously distributed throughout all layers of the orbitofrontal cortex. The densities of α2h receptors are maximal in layers I–II and diminish throughout the cortex, reaching the lowest values in layer VI. MO and VO contain higher α1 receptor densities than LO or VLO. MO and LO show the highest mean α2h receptor concentrations, particularly in layers I–II of the latter region. The serotoninergic receptors are homogeneously distributed throughout all layers of the orbitofrontal cortex. MO and VO show 5-HT1A receptor densities higher than those of VLO or VO. The four orbitofrontal regions do not differ significantly in their mean 5-HT2 densities or laminar distribution patterns.

Agranular Insular Cortex The agranular insular cortex, or claustrocortex (Stephan, 1975; Zilles et al., 1980), lies mainly within the rhinal sulcus. This region was divided (Zilles and Wree, 1985) into three areas: the dorsal part of the agranular insular cortex (AID), the ventral part of the agranular insular cortex (AIV), and the posterior part of the agranular insular cortex (AIP). Cytoarchitectonically, AIP is classified as a periallocortical area, whereas AID and AIV are more similar to the isocortex and were, therefore, classified as proisocortical areas (Reep and Winans, 1982).

The insular cortex can be myeloarchitectonically identified by its low content of myelin and its threelayered cytoarchitectonical appearance: layers II–IV and VI appear darker than the intervening layer V, which is broad, composed of sparsely packed medium to large, darkly stained pyramidal cells and shows particularly cell-sparse gaps on either side. Layer IV is present as a discrete layer which contains small granular cells. The agranular insular cortex shows significantly lower LCGU levels s than the rostrally adjoining orbital cortex (Zilles and Wree, 1995). It can be clearly delineated from the adjoining isocortex and the piriform cortex based on mean regional receptor densities and laminar distribution patterns. Furthermore, receptors for classical neurotransmitters also enable the visualization of the three areas into which the agranular insular cortex is divided. AMPA receptors are present in higher concentrations in the supragranular layers of AID and AIV than in the infragranular ones, whereas they are equally distributed throughout all cortical layers of AIP (Fig. 7). All three areas of the agranular insular cortex show their highest kainate receptor densities in the infragranular layers (Figs. 5–7) and their highest NMDA concentrations in the supragranular layers. The lowest mean AMPA densities were measured in AIP, and the highest values were found in AIV. The highest mean kainate receptor densities are located in AID (Figs. 5–7); AIP shows kainate concentrations slightly higher than those of AIV. NMDA densities are lowest in AIV, and AIP shows concentrations slightly higher than those of AID. GABAB receptors and BZ binding sites are present in higher densities in layers I–III of AIV and AIP. GABAA receptors are homogeneously distributed throughout all cortical layers of AID and AIP (Fig. 7), but are restricted to layers I–III of AIV. AIV shows overall lower GABAergic and BZ concentrations than AID or AIP, particularly due to differences in layers I–III. AIP contains higher mean GABAB receptor densities higher than AID, but these two regions do not differ in their mean GABAA receptor on BZ binding site concentrations. Muscarinic cholinergic M1 and M3 receptors are present in higher contrations in layers I–III of the agranular insular cortex than in layers V–VI. Conversely, the M2 and nicotinic cholinergic receptors show higher densities in layers V–VI (Fig. 7). AID, AIV, and AIP do not differ significantly in their mean regional muscarinic or nicotinic cholinergic receptor densities. Noradrenergic α1 receptors reach the highest densities in layer V of the agranular insular cortex, whereas the highest α2h receptor densities are located in layers V–VI. AID and AIP do not differ in their mean α1

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receptor densities, which are slightly higher than those present in AIV. AIP shows mean α2h receptor densities higher than those of either AID or AIV. Serotoninergic 5-HT1A and 5-HT2 receptors show maximal concentrations in layers V–VI of the agranular insular cortex (Fig. 7). AIP contains lower 5-HT1A receptor densities than AIV or AIP. AID and AIV show the same mean 5-HT2 densities, which are slightly higher than those measured in AIP.

Perirhinal Cortex The term perirhinal cortex (PRh) was coined (Zilles, 1990; Zilles and Wree, 1985, 1995; Zilles et al., 1980, 1990) to designate an architectonically heterogeneous cortical region located along the caudal half of the rhinal sulcus and extending further caudally, thus reaching the occipital contour of the hemisphere. PRh is delimited rostrally by the insular cortex, dorsally by the temporal cortex, and ventrally by the entorhinal cortex. It has been implicated in memory processes (Wiig et al., 1996; Zhu et al., 1995) and participates in the integration of polymodal sensory information, since it receives input from more than one sensory modality as well as from other polymodal regions (Burwell and Amaral, 1998a). PRh receives afferents from Cg1–3, RSA, the agranular insular cortex, the hippocampal formation, and secondary sensory and motor isocortices, as well as from the anterior thalamic nucleus and the amygdala (Beckstead, 1979; Burwell, 2000; Burwell and Amaral, 1998a, 1998b; Deacon et al., 1983; Inagaki et al., 1990; Kosel et al., 1982; Krettek and Price, 1977b; Markowitsch and Guldin, 1983; Wouterlood et al., 1990). PRh projects to the ipsilateral hippocampal formation (Burwell and Amaral, 1998b; Kosel et al., 1983). Our definition of PRh is in agreement with the descriptions of Burwell and Amaral (1998a, 1998b), Deacon et al. (1983), Kosel et al. (1983), and Krettek and Price (1977a) and corresponds to the posterior part of Krieg’s (1946a) Area 35 and to Swanson’s (1992, 1998) areas ECT and Peri. The rostral part of our PRh encompasses Areas 35 and 36 of Burwell (2001), whereas the caudal part of our PRh includes Burwell’s (2001) areas PORd and PORv. PRh can be clearly delineated from the insular, temporal, and entorhinal cortices due to its low degree of myelination. Furthermore, the rostral border of PRh can be clearly defined due to the disappearance of claustral cells below layer VI of the cortex. The trilaminar architectonical pattern described for the insular cortex is not apparent in PRh, since layer V pyramids are more densely packed and there are no cell-sparse gaps. Layer IV of PRh is rather inconspicuous and

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appears to merge with layer V. Conversely, layer VI is prominent and can be subdivided into two sublayers. PRh shows higher glutamatergic AMPA and NMDA receptor densities in layers I–III, whereas maximal kainate receptor densities are located in layers V–VI (Figs. 9 and 12). PRh contains lower mean AMPA, kainate (Figs. 9 and 12), and NMDA receptor densities than the adjoining temporal areas. The laminar distribution patterns of GABAergic receptors and BZ binding sites differ between PRh and the isocortex. GABAA receptors and BZ binding sites reach their highest concentrations in layer VI of PRh, and their densities diminish throughout the cortex, reaching their lowest values in layers I–II. GABAB receptors show the opposite laminar distribution, with their highest concentrations in layers I–II. PRh shows lower densities of GABAA and BZ binding sites but higher GABAB concentrations in layers I–III than in the adjoining areas. However, PRh contains lower overall GABAA and GABAB concentrations than the temporal cortex. Furthermore, PRh contains higher BZ binding site densities, particularly in layers V–VI, than the temporal cortex. Muscarinic cholinergic M1 and M3 receptors reach the highest densities in layers I–III. Conversely, maximal M2 and nicotinic receptor densities are found in layers V–VI. PRh cannot be clearly delineated from the adjacent temporal cortex on the basis of muscarinic cholinergic receptors, but contains lower nicotinic densities in layers I–III than the TeV. Noradrenergic α1 receptors show the highest concentrations in layers V–VI of PRh, whereas α2h receptors are homogeneously distributed throughout all layers. PRh contains lower α1 receptor densities than the temporal cortex. Conversely, PRh shows significantly higher mean α2h receptor densities than TeV. Serotoninergic receptors are present in highest densities in layers V–VI of PRh. PRh contains higher 5HT1A densities than the temporal cortex Te2, but slightly lower 5-HT2 concentrations.

Cingulate Cortex The cingulate cortex covers the frontomedial half of the rat cortex situated above the corpus callosum, it is one of the largest components of the limbic system and is characterized by diffuse projections from the anteromedial thalamic nucleus (Bentivoglio et al., 1990; Van Groen et al., 1990; Musil and Olson, 1990; Vogt, 1993). It is involved in motivational aspects of learning tasks (Gabriel et al., 1980; Porrino, 1990; Vogt et al., 1990) and contributes to motor functions via numerous efferents to subcortical motor systems (Dum and Strick, 1990; Hoesen van et al., 1990; Neafsey et al., 1990).

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The cingulate cortex can be subdivided into five areas: Cg1, Cg1´, Cg2, Cg2´, and Cg3. Some authors (Groenewegen, 1988; Uylings and Van Eden, 1990; Van Eden et al., 1992) include the infralimbic area (IL) in their definition of the cingulate cortex. However, the lamination pattern of IL is quite primitive compared with that of the other cingulate areas and does not resemble a proisocortical or isocortical type; rather, IL was identified as a periallocortical area, is classified as infralimbic or prelimbic cortex (Vogt, 1993), and therefore is not discussed further in the present chapter. For details see Chapter 22. Areas Cg1 and Cg1´ correspond to the rostral and caudal parts of Area Cg1, respectively (Zilles and Wree, 1985, 1995). Similarly, Areas Cg2 and Cg2´ correspond to the rostral and caudal parts of Area Cg2, respectively (Zilles and Wree, 1985, 1995). Areas C1–4 of Zilles et al. (1980) correspond to Areas Cg1, Cg2, IL, and Cg3, respectively (Zilles and Wree, 1995). The present delineation of Areas Cg1–3 is equivalent to Areas 23 and 24 of Krieg (1946a), to Area c of Droogleever Fortuyn’s (1914), and to Areas Cg1–3 of Schober (1986). Areas Cg1 and Cg2 were also delineated by Paxinos and co-workers (1999), and Areas Cg1´ and Cg2´ correspond to their Areas Cg and Rs. Areas Cg1 and Cg1´ were identified as Area ACd by Krettek and Price (1977a), as Area 24b by Vogt and Peters (1981), and as Area ACAd by Swanson (1992, 1998). Areas Cg2 and Cg2´ correspond to Area ACv of Krettek and Price (1977a), to Area 24a of Vogt and Peters (1981), and to Area ACAv of Swanson (1992, 1998). Vogt (1993) further subdivided Areas 24a and 24b into rostral (24a, 24b) and caudal (24a´, 24b´) parts, which correspond to our Areas Cg1, Cg2, Cg1´, and Cg2´, respectively. Our Area Cg3 is in accordance with Area 32 of Richter and Kranz (1979) and of Vogt (1993). The cingulate cortex is characterized by being agranular and having a particularly thick layer I and a prominent layer V. Cg1 shows an isocortical lamination pattern, with a homogeneous layer III, relatively large pyramids in layer V, and a double layer VI. Layer VIa neurons are smaller than those of layer VIb. Cg1 and Cg1´ have a poorly differentiated cytoarchitecture. However, they differ considerably in their connectivity patterns as well as in their mean densities of receptors for classical neurotransmitters (Figs. 5 and 6). Cg1, but not Cg1´, receives afferents from the mediodorsal thalamic nucleus and from the amygdala (Krettek and Price, 1977a; Sripanidkulchai et al., 1984). Furthermore, Cg1´, but not Cg1, projects to the pontine nuclei (Wiesendanger and Wiesendanger, 1982a, 1982b). Cg2 and Cg3 show rather homogeneous lamination patterns and have, therefore, been classified as proisocortical areas (Richter and Kranz, 1979). Layer V of Cg2 is pro-

minent, whereas Cg3 shows poorly differentiated inner and outer pyramidal layers. The cingulate cortex shows clearly lower LCGU levels than the adjoining orbitofrontal cortex (Zilles and Wree, 1995). Furthermore, Cg1 can be easily delineated from Areas Cg2 and Cg3 due to its higher mean LCGU values (Zilles and Wree, 1995). The glutamatergic AMPA and NMDA receptors are present at highest densities in layers I–III throughout the cingulate cortex, whereas kainate receptors show their maximal values in layers V–VI (Figs. 5 and 6). Cg2 contains clearly higher AMPA receptor densities in layers I–III than Cg1, Cg2´, and Cg3. Cg1 and Cg1´ do not differ significantly in their mean AMPA densities or laminar distribution patterns. Whereas Cg1, Cg2´, and Cg3 do not differ in their mean kainate receptor densities or laminar distribution patterns, layers I–III of Cg1 show higher kainate concentrations than Cg1´, but lower values than those present in Cg2 (Figs. 5 and 6). Cg1, Cg2´, and Cg3 show comparable mean NMDA receptor densities and laminar distribution patterns. Cg3 contains the highest and Cg1´ the lowest mean NMDA receptor densities. GABAergic receptors and BZ binding sites are present in higher concentrations in layers I–III than in layers V–VI of the cingulate cortex. Cingulate areas show comparable mean regional GABAA receptor densities and laminar distribution patterns, but they differ significantly in their mean regional GABAB receptor and BZ binding site densities. The highest GABAB receptor concentrations are located in Cg1´ and Cg2´, whereas Cg3 contains the lowest densities. The highest mean BZ binding site densities were measured in Cg1. The muscarinic cholinergic M1 receptors show maximal densities in layers I–III of the cingulate cortex, whereas M2 receptors reach their highest values in layers V–VI, and M3 receptors are homogeneously distributed throughout all layers. Nicotinic cholinergic receptors are present in higher densites in layers I–II and V than in layer VI; layer III shows extremely low values. Although Cg3 contains lower M1 receptor densities in layers I–III lower than Cg1, it shows higher concentrations in layers V–VI, resulting in comparable mean regional densities. The remaining cingulate areas do not differ significantly in their mean regional M1 receptor densities or laminar distribution patterns. Cg1, Cg2, and Cg1´ present comparable mean M2 receptor densities, whereas Cg3 contains lower M2 receptor densities in layers I–III than Cg1. Similarly, layers I–III of Cg2´ show lower M2 concentrations than Cg1´. Although Cg1 and Cg1´ contain higher M3 receptor densities in layers I–III than the adjacent cingulate areas, the mean regional M3 receptor

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concentrations do not differ throughout the cingulate cortex. The mean regional densities and laminar distribution patterns of the nicotinic cholinergic receptors do not enable a delineation of Cg1, Cg1´, Cg2, Cg2´, or Cg3. The noradrenergic α1 receptors show highest concentrations in layers I–II and V of the cingulate cortex, whereas α2h receptors reach maximal values in layers I–III. Cg1 shows slightly lower α1 receptor densities in layer VI and clearly higher α2h concentrations in layers I–III than does Cg2. Furthermore, Cg1 contains significantly higher α1 receptor densities than Cg1´ or Cg3, but lower α2h values in layers V–VI. Cg2 and Cg3 present the same mean α1 receptor densities and laminar distribution patterns, but differ in the higher α2h concentrations of Cg2. The serotoninergic 5-HT1A receptors are equally distributed throughout all cortical layers of the cingulate cortex, whereas maximal 5-HT2 densities are restricted to layer V in Cg1–3 and to layers I–III in Cg1´ and Cg2´. The lowest mean regional serotoninergic receptor densities are located in Cg1´ and Cg2´, whereas Cg2 and Cg3 contain the highest values. For a detailed account of the cingulate cortex, see Chapter 22.

Retrosplenial Cortex The retrosplenial cortex covers the mediocaudal surface of the hemisphere and is subdivided into a granular retrosplenial area (RSG) and an agranular retrosplenial area (RSA), although the RSA has also been described as dysgranular (Vogt, 1993). RSG is located on the medial surface of the hemisphere, whereas RSA covers the dorsomedial surface. Together with the anterior cingulate cortex, the retrosplenial cortex is involved in the motivational aspects of learning tasks (Porrino, 1990; Vogt et al., 1990) and contributes to motor functions via numerous efferents to subcortical motor systems (Dum and Strick, 1990; van Hoesen et al., 1990; Neafsey et al., 1990). Our delineation of the retrosplenial cortex is in accordance with previous observations (Krettek and Price, 1977a; Schober, 1986; Vogt, 1993; Vogt and Peters, 1981). RSA corresponds to Area 29d of Vogt (1993), Area RSA of Schober (1986), and part of Area RsAg of Krettek and Price (1977a). RSG is the equivalent of Vogt’s (1993) Areas 29a, 29b, and 29c, Schober’s (1986) Area RSG, and Krettek and Price’s (1977a) Area RsG. Cytoarchitectonically, RSG is characterized by a conspicuous layer II, but a poorly differentiated layer IV, and a subdivision of layer V into layers Va, with medium-sized pyramids, and Vb, with larger pyra-

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mids. Layers II–III of RSA are wider than those of the RSG, but poorly differentiated; layer V is prominent and contains large neurons. Furthermore, RSA is characterized by a degree of myelination slightly higher than that of RSG. The retrosplenial regions can be clearly delineated from the adjacent visual and cingulate regions based on their regional and laminar receptor distribution patterns. The retrosplenial cortex shows higher AMPA and NMDA receptor densities in layers I–III and VI than in layer V. Kainate receptors show a relatively homogeneous distribution throughout the retrosplenial cortex, although layer VI contains slightly higher densities than the remaining layers. RSA shows higher AMPA and NMDA but lower kainate concentrations than RSG (Fig. 7). GABAergic receptors and BZ binding sites are present in higher densities in layers I–III than in layers V–VI of the retrosplenial cortex (Fig. 7). Layers I–III of RSA contain clear higher GABAA and GABAB receptor densities as well as BZ binding site densities than RSG (Fig. 7). RSG shows slightly higher BZ binding site densities in layers V–VI than RSA. Muscarinic and nicotinic cholinergic receptors are present in higher densities in layers I–III than in layers V–VI of the retrosplenial cortex (Fig. 7). Although RSA and RSG do not differ significantly in their mean M1, M2, and M3 receptor densities, there are slight differences in the laminar distribution patterns of the M2 and M3 receptors. RSA shows higher M2 and M3 receptor concentrations than RSG in layers I–III, but lower values in layers V–VI (Fig. 7). RSG contains clearly higher nicotinic receptor densities in layers I–III (Fig. 7). The noradrenergic α1 and α2h receptors are present in higher densities in layers I–III than in layers V–VI of the retrosplenial cortex. RSA shows slightly higher α1 receptor densities in layers I–III than RSG. These two regions do not differ in their mean α2h receptor concentration or laminar distribution pattern (Fig. 7). The retrosplenial cortex shows the highest 5-HT1A receptor densities in layers V–VI, whereas 5-HT2 receptors are homogeneously distributed (Fig. 7). RSA and RSG do not differ in their mean 5-HT1A or 5-HT2 receptor concentrations or laminar distribution patterns (Fig. 7). For further details concerning the retrosplenial cortex, see Chapter 22.

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C H A P T E R

24 Central Autonomic System CLIFFORD B. SAPER Department of Neurology and Program in Neuroscience Beth Israel Hospital and Harvard Medical School Boston, Massachusetts, USA

Autonomic control must be closely related not only to the ongoing metabolic demands of the individual but also to behavioral and emotional responses. For this reason, the central autonomic control system is tightly linked to sensory systems monitoring the internal environment, including both visceral sensation and direct monitoring of various chemical and physical qualities of the bloodstream (such as temperature and osmolality). At the same time, the autonomic control system receives massive inputs from forebrain structures that are involved in every aspect of behavior (from salivation during eating to blood pressure elevation during exercise) and is especially important in emotional response. The output of the autonomic control system involves both the regulation of sympathetic and parasympathetic preganglionic neurons in the medulla and spinal cord, as well as the level of behavioral arousal. The latter feature is not often included within the spectrum of autonomic responses, but from the perspective of survival of the individual it is clearly the single most important aspect of what is often called the “fight or flight” response. Recent research indicates that the neuronal systems controlling autonomic response are inseparable from those regulating behavioral arousal. The earliest modern formulation of the mechanism for behavioral regulation of the autonomic nervous system evolved from the work of Cannon and Britton (1925) with the decorticate cat preparation. Innocuous stimuli, such as lightly stroking the flank of the animal, would produce a massive rage response, including both somatomotor components such as arching of the back, hissing and spitting, and autonomic responses

The Rat Nervous System, Third Edition

such as retraction of the nictitating membrane, elevation of blood pressure and heart rate, and piloerection. Cannon’s student, Philip Bard (1928), later performed serial transections through the remaining neuraxis of these animals, demonstrating that the “sham rage” response (sham, presumably, because the animal lacked the cognitive component of rage) was eliminated with cuts that separated the diencephalon from the mesencephalon. On the basis of these experiments, a hierarchical model of autonomic control evolved, in which the cortex was thought to inhibit the sympathoexcitatory activity of the hypothalamus, which in turn was thought to play upon the reflex autonomic control circuitry of the brainstem. Observations during the 1970s and 1980s indicated that the organization of the autonomic control system is much more complex than the hierarchical model would suggest. In fact, the interconnectivity of its components, located at virtually every level of the neuraxis, is more similar to a network than a strict hierarchy. The data supporting this network model now come from both physiological and neuroanatomical studies. However, the outline of this network emerged largely on the basis of the availability of the modern axonal tracer methods that became available in the 1970s. The first strategy that was used to identify the central autonomic system was the injection of the preganglionic cell groups of the medulla and the spinal cord with retrograde tracers (see Fig. 1). These studies identified a series of sympathetic and parasympathetic premotor cell groups in the hypothalamus, pons, and medulla. Subsequent anterograde transport studies demonstrated

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FIGURE 1 Visceral afferent (A) and efferent (B) systems. In A, the cell groups are identified that receive visceral afferent information either directly from the nucleus of the solitary tract (NTS) or via a relay in the parabrachial nucleus (PB). Note that there are two limbs to the ascending pathway. The projections to the ventroposterior parvocellular nucleus of the thalamus (VPpc) and the insular cortex (IC) originate exclusively from the parabrachial nucleus in rats, whereas the innervation of the hypothalamus [including the anteroventral third ventricular region (Av3v), paraventricular nucleus (Pa), and lateral hypothalamic area (LH)], infralimbic cortex (ILC), and basal forebrain [including the bed nucleus of the stria terminalis (BST) and central nucleus of the amygdala (CeA)] arises from both the parabrachial nucleus and nucleus of the solitary tract. In B, the main sources of inputs to preganglionic neurons in the medulla and spinal cord are identified. Pathways providing direct input to preganglionic cell groups are illustrated by a solid line. Of all the forebrain cell groups, only the infralimbic cortex (ILC) and tuberal hypothalamic cell groups (Pa, LH, and dorsomedial and arcuate/retrochiasmatic nuclei, not shown) contribute to this pathway. Other forebrain structures exert autonomic control (dotted pathway) by projecting to visceral premotor sites that, in turn, innervate the preganglionic systems. A5, A5 noradrenergic cell group; AMB, nucleus ambiguus; DMV, dorsal motor vagal nucleus; IML, intermediolateral cell column; VM, ventral medullary reticular formation.

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that these neurons directly innervate the preganglionic autonomic cell groups in the medulla and the spinal cord. The second strategy was to trace the distribution of visceral sensory information in the brain. By examining the output from the nucleus of the solitary tract and those cell groups that receive its projections, it was possible to identify an array of cell groups that receive visceral sensory information, with representations in the cortex, amygdala, thalamus, hypothalamus, midbrain, pons, and medulla. Tracing the connections of these groups has indicated that they form an incestuous web, in which most of the central autonomic cell groups are connected to most of the others in the network. Furthermore, tracer studies have identified the entry into this system of interoceptive information derived from brain monitoring of the bloodstream, as well as inputs from and outputs to structures involved in other aspects of cognitive, emotional, and behavioral response. This chapter, which is focused on the anatomy of the central autonomic control system, takes a largely segmental approach to discussing connectivity. A recent review on functional aspects of this system covered the organization of pathways that relay visceral perception to the cerebral cortex and the organization of autonomic pattern generators (Saper, 2002). The reader is referred to this source for a more functional approach to the system.

MEDULLOSPINAL LEVEL: REFLEX CONTROL Although integrative activity occurs at all levels of the nervous system, it is worthwhile in a heuristic sense to consider the main focus of activities at different levels of the neuraxis. This approach risks comparison with the outdated hierarchical view of autonomic control, but helps put into perspective the connectivity of the central autonomic system. The organization of the preganglionic neurons themselves is not discussed. In recent years these have been covered in a series of articles that have used retrograde tracing with conventional and transneuronal viral tracers to identify sources of sympathetic and parasympathetic input to specific peripheral organs (see., e.g., Bieger and Hopkins, 1987; Strack et al., 1989; Loewy and Haxhiu, 1993; Hopkins et al., 1996; ter Horst et al., 1996; Huang and Weiss, 1999; Cano et al., 2001). This chapter instead focuses on cell groups that provide inputs to the preganglionic neurons.

Nucleus of the Solitary Tract (NTS) This structure is also covered in Chapter 28 on gustatory systems (Lundy and Norgren, this volume) and

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so is considered here mainly from the perspective of its position as an entry point for visceral sensory information arising from the vagal, glossopharyngeal, and facial nerves. The different visceral nerves end in a complex topographic distribution within the NTS, with the taste afferents occupying the most rostral portion and gastrointestinal afferents synapsing in the intermediate portion of the nucleus, including the central and gelatinous subnuclei (Altschuler et al., 1989; Broussard and Altschuler, 2000; Hayakawa et al., 2001; see Chapter 28 and Fig. 2 for cytoarchitecture). Cardiovascular afferents terminate in the caudal half of the nucleus, in the dorsomedial, medial, parvocellular, and commissural subnuclei, as well as in the area postrema (Wallach and Loewy, 1980; Davies and Kalia, 1981; Ciriello, 1983; Seiders and Steusse, 1983; Erickson and Millhorn, 1991; Chan et al., 2000) and respiratory afferents end mainly in the ventrolateral, intermediate, interstitial, and commissural subnuclei (Kalia and Richter, 1985, 1988; Otake et al., 2001). The NTS also receives afferents from the superficial layers of the spinal and trigeminal dorsal horns (Menetrey and Basbaum, 1987). Many of the dorsal horn neurons contributing to this pathway are activated by visceral stimuli and contain glutamate (GamboaEsteves et al., 2001). The NTS receives additional afferents from virtually every other component of the central autonomic system (see below), but these other inputs do not appear to have the exquisite topographic specificity of the peripheral visceral nerves. The area postrema is a circumventricular organ (see Chapter 16, Oldfield and McKinley, this volume) that sits along the dorsal surface of the NTS at the level of the obex and has major projections into the NTS. Some NTS neurons send dendrites into this neurohemal contact zone (Herbert et al., 1990). The projections from the area postrema to the medullary reticular formation and parabrachial nucleus are virtually identical with those of the underlying NTS (Shapiro and Miselis, 1985; Herbert et al., 1990). For this reason, the area postrema can be considered a chemosensory portion of the dorsal vagal complex. The NTS has three major classes of projections (Loewy and Burton, 1978; Ricardo and Koh, 1978; Norgren, 1978; Ross et al., 1985; Cunningham and Sawchenko, 1989; Herbert et al., 1990). First, it sends a descending projection to autonomic preganglionic neurons. The projection to the brainstem preganglionic cell groups, including the dorsal motor vagal nucleus, the nucleus ambiguus, and the superior and inferior salivatory nuclei, is quite extensive, and each portion of the NTS contributes to this set of pathways. The descending projection to the spinal cord emerges exclusively from a small population of neurons in the ventrolateral NTS. The spinal targets of this pathway include respiratory

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FIGURE 2 Cytoarchitecture of the general visceral sensory components of the nucleus of the solitary tract. The panels on the left (from rostral, A, to caudal, D) illustrate the subnuclei in Nissl-stained material. NADPH-diaphorase staining, on the right, demonstrates some of the subnuclear borders (e.g., the central subnucleus, ce) more clearly. Other subnuclei of the nucleus of the solitary tract are as follows: com, commissural; dm, dorsomedial; g, gelatinous; I, intermediate; m, medial; pc, parvocellular; vl, ventrolateral. The neurons within the solitary tract (ts) constitute the interstitial nucleus. Adjacent structures include the following: AP, area postrema; 10, dorsal motor vagal nucleus. Scale = 0.5 mm. Reprinted from Herbert et al. (1990a) with permission.

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motor neurons as well as the sympathetic preganglionic column (Loewy and Burton, 1978; Dobbins and Feldman, 1994), suggesting that it may be most closely involved in respiratory-related responses. Second, the NTS projects into the medullary reticular formation, including a parvicellular zone that integrates gustatory sensation with oropharyngeal and gastrointestinal reflexes, and the rostral and caudal ventrolateral areas, which regulate cardiovascular and respiratory reflexes. The third output from the nucleus of the solitary tract is its ascending projection, which defines most of the remainder of the central autonomic control system. The largest single projection from the NTS is to the parabrachial nucleus. This projection is topographically organized (see Fig. 3) and has been extensively reviewed (Herbert et al., 1990; Karimnamazi et al., 2002). The taste, general visceral, and respiratory components of the NTS mark out distinct terminal fields in the parabrachial nucleus, indicating that some topographic organization is maintained in the ascending projection. This ordering is especially well preserved in the visceral sensory projection from the parabrachial nucleus to the thalamus and from there to the insular cortex (see below). The NTS also sends a major projection into the periaqueductal gray matter that arises in large part from noradrenergic neurons (Herbert and Saper, 1992). The NTS projects less heavily to a number of forebrain sites that receive more extensive inputs from the parabrachial nucleus, including the central nucleus of the amygdala; the bed nucleus of the stria terminalis; the median preoptic, paraventricular and dorsomedial hypothalamic nuclei; and the lateral hypothalamic area (Ricardo and Koh, 1978; ter Horst et al., 1989). It even sends a few axons as far as the subfornical organ (Zardetto-Smith and Gray, 1987; Ciriello et al., 1996; Tanaka et al., 2001) and the infralimbic cortex (Tucker and Saper, 1984). In primates, the far rostral NTS may send a direct projection to the visceral sensory relay nucleus in the thalamus for taste (Beckstead et al., 1980); a comparable projection in rats has not been identified.

Rostral Ventrolateral Reticular Nucleus (RVL) The RVL is a cytoarchitecturally distinct, pyramidshaped area (see, e.g., Paxinos and Watson, 1986, plate 67), with its apex at the compact part of the nucleus ambiguus and its base along the ventrolateral surface of the medulla. The RVL is now recognized as playing an extraordinarily important role in the integration of cardiovascular and respiratory reflexes. Within the RVL are contained two populations of respiratory neurons, the Bötzinger complex rostrally and dorsally, which contains a high proportion of expiratory-related

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neurons, and the ventral respiratory group caudally and ventrally, which contains many inspiratory-related cells (Otake et al., 1987; Saether et al., 1987; Ezure et al., 1988; Schwarzacher et al., 1991). At more slightly more caudal levels is a population of neurons termed by Smith and colleagues (1991) the “pre-Bötzinger complex,” whose firing appears to play an important role in driving the respiratory cycle (Gray et al., 2001). The RVL also contains the external formation of the nucleus ambiguus, which includes many cardiomotor vagal neurons, and it lies directly ventral to the compact formation of the nucleus ambiguus, which contains esophageal motor neurons (Bieger and Hopkins, 1987; Miselis et al., 1989; Corbett et al., 1999; Broussard and Altschuler, 2000). Intertwined with the external formation is a population of neurons roughly coinciding with the most rostral part of the C1 adrenergic cell group, whose firing rates increase during sympathoexcitatory stimuli and that project to the sympathetic preganglionic column (Ross et al., 1984; Barman and Gebber, 1985; Guyenet and Brown, 1986; Morrison et al., 1988). Some of these neurons probably belong to the C1 group, whereas others are probably nonadrenergic (Tucker et al., 1987; Sun et al., 1988; Guyenet et al., 2001). Lesions of this area cause blood pressure to fall to nearly the same levels as spinal transection (i.e., complete loss of sympathetic descending tone) and abolish a variety of cardiovascular reflexes and centrally evoked responses, including the cardiovascular and vasopressin responses to baroreceptor stimulation (Ross et al., 1984; Yamada et al., 1984; Shreihofer and Guyenet, 2002). Selective lesions of the C1 neurons impair baroreflexes, but do not eliminate descending tone for maintenance of blood pressure (Schreihofer et al., 2000). The RVL also receives ascending inputs from the caudal ventrolateral medulla (Schreihofer and Guyenet, 2002) as well as collaterals from neurons in the spinal and trigeminal dorsal horns (Mehler et al., 1960; Panneton and Burton, 1985). In addition the RVL receives descending afferents from the infralimbic cortex (Hurley et al., 1991); central nucleus of the amygdala (Takayama et al., 1990; Cassell and Gray, 1991); median preoptic, paraventricular, and dorsomedial hypothalamic nuclei and lateral hypothalamic area (Saper et al., 1976; Saper and Levisohn, 1983; ter Horst et al., 1984; Luiten et al., 1985, 1987; Hosoya, 1985; Ciriello et al., 1985; Pyner and Coote, 2000); periaqueductal gray matter (Carrive et al., 1988; Carrive and Bandler, 1991; van Bockstaele et al., 1989, 1991; Farkas et al., 1998; Keay et al., 2000); and parabrachial nucleus (Ellenberger and Feldman, 1989; Herbert et al., 1990; Chamberlin and Saper, 1994). It sends ascending projections to the locus coeruleus; the parabrachial nucleus; the periaqueductal gray matter;

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FIGURE 3 Topographic arrangement of projections from the nucleus of the solitary tract and area postrema to the parabrachial nucleus. In this schematic figure, the nucleus of the solitary tract is illustrated at the top left from a dorsal perspective and below left in a plane taken at the level illustrated above. Different subregions with characteristic parabrachial projections are illustrated by different shading; the terminal fields of these subregions are demonstrated in the series of transverse sections through the parabrachial nucleus on the right (from rostral, top, to caudal, bottom) by the same shadings. Note that the gustatory zone (dark shading with light diagonal lines), general visceral zone (small and large dots), and respiratory zone (horizontal dark lines) are largely mutually exclusive. Regions that do not receive major inputs from the nucleus of the solitary tract receive afferents from the spinal cord (dorsal and internal lateral subnuclei) and forebrain (medial, ventral, and central lateral nuclei). AP, area postrema; calamus script., calamus scriptorius; com, commissural solitary subnucleus; Cu, cuneate nucleus; dm, dorsomedial solitary subnucleus; Gr, gracile nucleus; mNTS, medial solitary subnucleus; NTS, nucleus of the solitary tract; pyram. decuss., pyramidal decussation; rNTS, rostral solitary subnucleus; scp, superior cerebellar peduncle; vl, ventrolateral solitary subnucleus; 10, dorsal motor vagal nucleus; 12, hypoglossal nucleus. Reprinted from Herbert et al. (1990a) with permission.

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and the median preoptic, supraoptic, and paraventricular nuclei of the hypothalamus; and the central nucleus of the amygdala (Guyenet and Young, 1987; Tucker et al., 1987; Cunningham and Sawchenko, 1988; Pieribone et al., 1988; Weiss and Hatton, 1990; Herbert et al., 1990; Herbert and Saper, 1992; Petrov et al., 1993; Ciriello et al., 1994). Some of the RVL neurons projecting to the locus coeruleus and periaqueductal gray matter may also project to the spinal cord (Guyenet and Young, 1987), but few spinally projecting neurons in the RVL send collaterals to the hypothalamus (Tucker et al., 1987).

Caudal Ventrolateral Medulla The caudal ventrolateral medulla is much less well anatomically defined: the term has mostly been used as a physiological construct to identify an area of the ventrolateral medullary reticular formation, just caudal to the rostral ventrolateral medulla and surrounding the lateral reticular nucleus caudal to the obex, from which depressor responses may be obtained (i.e., a fall in arterial blood pressure when stimulated). The caudal ventrolateral medulla receives extensive afferents from the cardiovascular portion of the NTS (Ross et al., 1985) as well as the parabrachial complex (Chamberlin et al., 1994) and the ventrolateral periaqueductal gray matter (Chen and Aston-Jones, 1996; Henderson et al., 1998) and also contains the caudal expiratory-driven part of the ventral respiratory group (Otake et al., 1987; Saether et al., 1987; Ezure et al., 1988). It projects rostrally to the RVL and A5 region and is believed to be a critical relay for baroreceptor reflex bradycardia (Agarwal et al., 1990; Agarwal and Calaresu, 1991; Masuda et al., 1991; Li et al., 1992). Injection of GABA antagonists into the RVL prevents the baroreceptor initiated cardiovascular and vasopressin responses (Willette et al., 1983; Yamada et al., 1984; Blessing and Willoughby, 1985), but it has not been possible to identify GABA-ergic neurons in the caudal region that project to the RVL. Current models suggest that baroreceptor information is relayed from the NTS to GABAergic neurons in the caudal ventrolateral medulla, which in turn inhibit sympathoexcitatory neurons in the RVL (Ciriello et al., 1986; Blessing, 1988; Guyenet, 1990; Chan and Sawchenko, 1998; Schreihofer and Guyenet, 2002).

Ventromedial Medulla and Medullary Raphe Nuclei The medullary raphe nuclei, including the raphe obscurus and pallidus and a lateral extention into the roots of the hypoglossal nerve, provide a serotoninergic

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projection to the sympathetic preganglionic cell column (Loewy et al., 1981). Some of these same cells contain various peptides, including substance P and thyrotropinreleasing hormone (Appel et al., 1987; Chiba and Masuko, 1989; Sasek et al., 1990; Poulat et al., 1992). The role played by these neurons in autonomic regulation is controversial. Early physiological studies suggested that this is probably an inhibitory input for many sympathetic preganglionic neurons (Cabot et al., 1979; Gilbey et al., 1981). Subsequent work demonstrated that injections of excitatory amino acids into the rostral ventromedial medulla produced mainly hypertensive and vasoconstrictor responses (Cox and Brody, 1989, 1991; Varner et al., 1992). Consistent with these studies, recent observations have defined a rostral ventral component of the medullary raphe that excites specific sympathetic responses. Neurons in the raphe pallidus at the level of the facial motor nucleus, particularly those most ventral between the pyramids (parapyramidal group) have increased activity during hypothermia (Morrison et al., 1999). They in turn activate specific populations of sympathetic preganglionic neurons that increase body temperature (Rathner and McAllen, 1999; Blessing and Nalivaiko, 2001; Morrison, 2001). Among these are preganglionic neurons that activate brown adipose tissue, which can generate heat by metabolizing lipids, and others that constrict the tail artery, which reduces passive heat loss through the naked skin of the tail (the largest radiative surface in a rat). The parapyramidal region of the raphe receives afferents from the median preoptic nucleus and the paraventricular nucleus in the hypothalamus which are believed to play an important role in both thermoregulation and producing fever responses (Nakamura et al., 2002; Tanaka et al., 2002; Saper, 2002). These observations have suggested that the organizational pattern of sympathetic preganglionic control in the ventrolateral medulla may be along the lines of central pattern generators (Morrison, 2001; Saper, 2002). For example, neurons in both the ventrolateral medulla and ventromedial medulla are capable of causing constriction of the tail artery, but the former are called into play as part of a generalized response to support blood pressure by hypotensive baroreceptor stimuli, whereas the latter are activated as part of a generalized thermogenic response by stimuli that promote increased body temperature. This distinction suggests that the neurons that participate in a particular physiological function may cosegregate, rather than the premotor autonomic neurons sorting topographically, e.g., by tissue target. This concept has been recently received extensive attention in reviews (Morrison, 2001; Saper, 2002) to which the reader may turn for further discussion.

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MESOPONTINE LEVEL: MODULATION AND INTEGRATION OF REFLEX CONTROL AND AROUSAL Parabrachial Nucleus The parabrachial nucleus occupies a key position in the central autonomic network, as an interface between medullary reflex control and forebrain behavioral and integrative regulation of the autonomic system. The parabrachial nucleus has been divided into thirteen distinct subnuclei and regions (see Figs. 4–9), each associated with a unique set of afferents, efferents, and neurotransmitters (for review see Fulwiler and Saper, 1984; Herbert et al., 1990; Herbert and Saper, 1992; Moga et al., 1989, 1990a; Chamberlin and Saper, 1992). Although the ascending inputs from the different subdivisions of the NTS (gustatory, general visceral, and respiratory) mark out distinct terminal fields in the parabrachial nucleus (Herbert et al., 1990; Karimnamazi et al., 2002; see Fig. 3), the plan of the parabrachial subnuclei reflects a different level of organization. In general, the subnuclei seem primarily concerned with the relay of specific types of information (reflected by chemical specificity of their inputs and outputs) to their individual terminal fields. For example, the paraventricular nucleus of the hypothalamus receives afferents from several parabrachial subnuclei, including the superior, dorsal, and central lateral groups. However, the projection from the superior lateral nucleus originates from cholecystokinin-immunoreactive neurons and is shared with the ventromedial nucleus and lateral hypothalamic area (Zaborszky et al., 1984; Shimada et al., 1984; Fulwiler and Saper, 1985). The projection from the dorsal and central lateral subnuclei, by contrast, is shared with the median preoptic nucleus (Saper and Levisohn, 1983; Fulwiler and Saper, 1984). The latter projection originates from neurons containing a number of different peptides, including enkephalin, corticotropin-releasing factor, galanin,

somatostatin, and brain natriuretic peptide (Lind and Swanson, 1984; Chamberlin and Saper, 1989). Whether the paraventricular projection from these subnuclei contains the same peptides remains to be determined. The resorting of visceral projections from the PB along chemically coded lines suggests that the organization of the PB reflects bodywide regulatory considerations, such as control of body fluids, energy metabolism, and blood oxygenation, rather than specific organs or autonomic reflexes (as represented in the medulla) or behavioral contexts (as organized by the forebrain). The internal lateral parabrachial subnucleus receives afferents mainly from the spinal and trigeminal dorsal horn (Cechetto et al., 1985; Menetrey and de Pommery, 1991 Slugg and Light, 1993; Caous et al., 2001; Gauriau and Bernard, 2002) and projects primarily to the thalamic intralaminar nuclei (Fullwiler and Saper, 1984; Bester et al., 1999; Krout and Loewy, 2000a). As the inputs to this cell group are so closely related to pain modulation, it is thought that the internal lateral nucleus projection to the thalamus may play a role in the arousal associated with nociception. The medial parabrachial subnucleus also innervates the thalamic intralaminar nuclei, as well as the caudal ventral part of the mediodorsal nucleus of the thalamus (which projects to the agranular insular cortex) (Fulwiler and Saper, 1984; Slugg and Light, 1993). The projection to the ventroposterior parvicellular nucleus of the thalamus, which is the main visceral sensory relay to the insular cortex, originates predominantly in the contralateral external medial parabrachial subnucleus (Cechetto and Saper, 1987). This latter projection, which is the only component of the parabrachial projection to the forebrain that is not predominantly ipsilateral, arises largely from cells that are immunoreactive with antisera against calcitonin gene-related peptide (Yasui et al., 1989). The external and extreme lateral PB nuclei project mainly to the amygdala and to the associated portions of the substantia innominata and the bed nucleus of

FIGURES 4–9 A series of figures illustrating the cytoarchitecture of the parabrachial nucleus and the distribution of points at which electrical microstimulation produce increases or decreases in blood pressure. In each figure, A is a photomicrograph of a Nissl-stained section through the parabrachial nucleus, and B–D are drawings of those sections with superimposed stimulation results. Open circles are points at which 5 μA of stimulation current produced a 5–24 mmHg change in blood pressure; small open triangles, 25–49 mmHg; small filled triangles, 50–74 mmHg; large filled triangles, 75–90 mmHg. Parabrachial subnuclei are as follows: cl, central lateral; dl, dorsal lateral; el, external lateral; exl, extreme lateral; exm, external medial; il, internal lateral; KF, Kölliker-Fuse nucleus; sl, superior lateral; vl, ventrolateral. Other labeled structures include the following: B, Barrington’s nucleus; CnF, cuneiform nucleus; DLL, dorsal nucleus of the lateral lemniscus; LC, locus coeruleus; LDT, laterodorsal tegmental nucleus; mcp, middle cerebellar peduncle; Me5, mesencephalic trigeminal nucleus; me5, mesencephalic trigeminal tract; Mo5, motor trigeminal nucleus; mo5, motor trigeminal root; Pr5, principal sensory trigeminal nucleus; scp, superior cerebellar peduncle; Su5, supratrigeminal nucleus; vsct, ventral spinocerebellar tract; 4n, trochlear nerve. Scale = 0.5 mm. Reprinted from Chamberlin and Saper (1992) with permission.

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the stria terminalis (Fulwiler and Saper, 1984; Moga et al., 1990; Bernard et al., 1992). This series of pathways is topographically organized, with the projection to the lateral part of the central nucleus of the amygdala originating in the outer part of the external lateral nucleus; the projection to the medial part of the central nucleus and the adjacent substantia innominata from the inner part of the external lateral subnucleus and adjacent ventral lateral and waist subnuclei; the afferents to the laterocapsular part of the central nucleus from the external medial subnucleus; and the pathway to the bed nucleus of the stria terminalis originating most heavily from the extreme lateral subnucleus. Many of the neurons in the outer portion of the external lateral nucleus that project to the lateral part of the central nucleus of the amygdala contain calcitonin gene-related peptide (Shimada et al., 1985; Schwaber et al., 1988; Yasui et al., 1991). Finally, the descending projections from the parabrachial complex originate from its most lateral components. A crescent shaped region in the far lateral

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part of the lateral parabrachial region projects to the RVL and to the ventrolateral and intermediate part of the NTS (Chamberlin and Saper, 1992, 1994). The Kölliker-Fuse subnucleus, which occupies the far ventrolateral corner of the parabrachial complex, innervates the sympathetic preganglionic column of the spinal cord, the nucleus ambiguous, the ventrolateral and intermediate NTS subnuclei, the RVL, and the more caudal parts of the ventrolateral medullary reticular formation (Saper and Loewy, 1980; Fulwiler and Saper, 1984; Ellenberger and Feldman, 1989; Herbert and Saper, 1992; Chamberlin and Saper, 1992, 1994; Hayakawa et al., 1999). Microstimulation studies have demonstrated a variety of cardiovascular responses from the parts of the parabrachial complex that project to the medulla and the spinal cord. However, large (>50 mmHg) responses to low-threshold currents (5 μA) or glutamate injections (10–500 pmole) occur mainly along the lateral margin of the external lateral nucleus (Chamberlin and Saper, 1992). Somewhat smaller

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depressor responses occur during electrical or chemical microstimulation in the dorsal lateral subnucleus. The pathways and neurotransmitters responsible for these cardiovascular effects have not yet been identified. Microstimulation with glutamate produces mainly increased ventilatory responses from the lateral crescent and external lateral subnuclei (Chamberlin and Saper, 1994). Stimulation of the Kölliker-Fuse nucleus produces increased and prolonged inspiration. Because the Kölliker-Fuse nucleus is the only part of the parabrachial complex to innervate the NTS, and its main targets are the respiratory parts of the NTS, it is thought that the Kölliker-Fuse nucleus input may inhibit responses to pulmonary inflation, thus allowing increased and prolonged lung inflation. Apneic responses are mainly obtained from the region just ventral to the Kölliker-Fuse nucleus (Chamberlin and Saper, 1998). This area is termed the intertrigeminal region because its neurons lie between the principal sensory and motor trigeminal nuclei and are intermingled with the trigeminal root bundles. The intertrigeminal neurons can best be demonstrated by retrograde labeling from the ventrolateral medulla, to which they project extensively. They also receive inputs from each of the sensory regions in the NTS and spinal trigeminal nucleus that are thought to initiate apneic reflex responses (Chamberlin and Saper, 1998). A few of the most dorsal neurons of the intertrigeminal group invade the ventral border of the Kölliker-Fuse nucleus, so that apneic responses can be obtained from the ventral edge of the Kölliker-Fuse nucleus (see also Dutschmann and Herbert, 1996). However, stimulation sites confined to the central part of the KöllikerFuse nucleus only augment inspiration.

A5 Noradrenergic Group The A5 group provides the main descending noradrenergic projection to all levels of the sympathetic preganglionic cell column in the spinal cord (Loewy et al., 1978; Strack et al., 1989a, 1989b; Clark and Proudfit, 1993). Some A5 neurons innervate the NTS and others the caudal ventrolateral medulla or the periaqueductal gray matter, often as collaterals of individual spinally projecting A5 neurons (Loewy et al., 1986; Kwiat and Basbaum, 1990; Tavares et al., 1997). No ascending projections to the forebrain have been identified from the A5 group. Afferents to the A5 area have been identified from the caudal ventrolateral medulla, the spinal trigeminal nulceus, the Kölliker-Fuse nucleus, and the paraventricular and dorsomedial nuclei and lateral hypothalamus, but there have been no electron microscopic studies to date to determine whether these fibers synapse upon A5 neurons (Saper et al., 1976;

Saper and Loewy, 1980; ter Horst et al., 1986; Hosoya et al., 1990; Li et al., 1992; Tavares et al., 1997). However, spinally projecting A5 neurons are clearly modulated by baroreceptor inputs (Andrade and Aghajanian, 1982; Guyenet, 1984). Chemical stimulation of neurons in the A5 region causes a fall in blood pressure, but electrical stimulation at this site produces the opposite response, most likely due to activation of fibers of passage (Neil and Loewy, 1982; Stanek et al., 1984).

A7 Noradrenergic Group The A7 group also provides a descending noradrenergic projection to the spinal cord (Westlund et al., 1983; Kwiat and Basbaum, 1990). One problem in determining the projections of the A7 group has been its proximity to the Kölliker-Fuse nucleus, which projects to the sympathetic preganglionic column (Tan and Holstege, 1986). However, a study combining anterograde transport of PHA-L with immunocytochemistry for dopamine βhydroxylase showed that the A7 axons end primarily in the dorsal horn, where they provide intense innervation of the marginal zone, as well as layers II–IV (Clark and Proudfit, 1991).

Periaqueductal Gray Matter The periaqueductal gray matter (PAG) contains a prominent fiber system running through the PAG (the so-called dorsal longitudinal fasciculus of Schütz, now usually termed the periventricular fiber system) connecting forebrain and brainstem autonomic control nuclei. The PAG has been divided into longitudinal zones or columns (Fig. 10), on the basis of both cyto- and chemoarchitecture and its connections (see Herbert and Saper, 1992, Keay and Bandler, 2001). The dorsomedial zone straddles the midline, above the cerebral aqueduct; the dorsolateral zone contains smaller, more densely packed neurons that stain for NADPH diaphorase, occupying the dorsolateral quadrant of the PAG; the lateral zone contains somewhat larger, less densely packed neurons; and the ventrolateral zone, which occupies the remainder of the PAG, excluding the dorsal raphe, dorsal tegmental nucleus, and laterodorsal tegmental nucleus, is even less densely packed. These PAG columns have very different roles in autonomic function (see Keay and Bandler, 2001, for review). Electrical or chemical stimulation of the dorsolateral and lateral columns mainly invoke active coping strategies, including confrontational defense (fighting) when stimulated rostrally and flight when stimulated caudally. Autonomic concomitants of both strategies include hypertension, tachycardia, hindlimb

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FIGURE 10 Cytoarchitecture of the periaqueductal gray matter (PAG) in rat. The lower series of panels illustrates the appearance of the periaqueductal gray matter in Nissl-stained sections, from rostral (A´) to caudal (D´). The upper panels (A–D) demonstrate the same levels in sections that have been stained for NADPH-diaphorase. Notice that this method clarifies the borders of the dorsolateral subdivision of the PAG (PAGDL) with the dorsormedial PAG (PAGDM) and lateral PAG (PAGl). The ventrolateral PAG (PAGVL) is slightly more densely stained with NADPHdiaphorase than PAGL subdivisions, and laterodorsal tegmental (LDTg) and dorsal raphe (DR) nuclei stand out as well. DTg, dorsal tegmental nucleus; EW, Edinger-Westphal nucleus; me5, mesencephalic trigeminal tract; mlf, medial longitudinal fasciculus; Su3, supraoculomotor nucleus; 4, trochlear nucleus. Scale = 1.0. Modified from Herbert and Saper (1992) with permission.

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vasodilation, and visceral vasoconstriction. Stimulation of the ventrolateral column, by contrast, evokes passive coping, manifest as behavioral quiescence with autonomic responses, including hypotension and bradycardia. The connections of the different columns vary both between them and in a rostrocaudal gradient. The lateral and ventrolateral columns receive extensive afferents from the NTS and the RVL, arising in large part from noradrenergic and adrenergic neurons, respectively (Kwiat and Basbaum, 1990; Herbert and Saper, 1992). These same columns also are a major termination site of the spinomesencephalic tract, arising predominantly from lamina I of the spinal cord but to a lesser extent from lamina V and the lateral spinal nucleus (Swett et al., 1985; Pechura and Liu, 1986; Menetrey et al., 1992; Keay et al., 1997). Most of these afferents arise from high cervical segments, but all levels of the spinal cord contribute. Afferents from the forelimbs and medullary dorsal horn terminate more rostrally in the PAG, whereas inputs from the hindlimbs terminate more caudally. Afferents from laminae I and V intermingle in the ventrolateral PAG, but the lateral PAG is filled predominantly with afferents from lamina I, whereas the lamina V afferents mainly terminate near the aqueduct (Keay et al., 1997). The PAG also receives topographically ordered afferents from the prefrontal cortex (Hurley et al., 1991; Shipley et al., 1991; Floyd et al., 2000; Fisk and Wyss, 2000). The caudodorsal prelimbic and anterior cingulate areas mainly innervate the dorsolateral column, whereas the more rostroventral prelimbic and infralimbic areas and the agranular insular cortex innervate the ventrolateral column. In monkeys the dorsomedial prefrontal cortex also projects to the lateral PAG, but a similar projection has not been identified in rats (An et al., 1998; Keay and Bandler, 2001). It is interesting that the medial prefrontal cortex also projects mainly to medial hypothalamic fields, such as the dorsomedial and ventromedial nuclei of the hypothalamus (Floyd et al., 2001), which in turn project to the lateral and dorsolateral columns of the PAG, respectively (Canteras et al., 1996; Thompson et al., 1996; Keay and Bandler, 2001). By contrast, the agranular insular cortex projects mainly to the lateral hypothalamic area (Saper, 1982b; Allen et al., 1991), which projects to the ventrolateral PAG (Keay and Bandler, 2000). The median preoptic, paraventricular, and periventricular hypothalamic nuclei also innervate the lateral and ventrolateral PAG; the medial preoptic area has a more complex pattern of topographic innervation of both the dorsomedial and lateral/ventrolateral PAG (Saper et al., 1976; Saper and Levisohn, 1983; Luiten et al., 1985b; Rivzi et al., 1992). The central nucleus of the amygdala and components of the extended amygdala, including the bed nucleus

of the stria terminalis (Krettek and Price, 1978; Rizvi et al., 1991) and the substantia innominata (Grove, 1988) also project to both the dorsomedial and the lateral/ ventrolateral PAG. The PAG provides reciprocal projections back to the central nucleus of the amygdala (Li et al., 1990a; Luiten et al., 1985a; Rizvi et al., 1991), hypothalamus (Carstens et al., 1990; Reichling and Basbaum, 1991; Rizvi et al., 1992), NTS (Ross et al., 1981; Bandler and Törk, 1987; Farkas et al., 1997), RVL (van Bockstaele et al., 1991; van Bockstaele and Aston-Jones, 1992), and spinal cord (Skirboll et al., 1983). The ventrolateral PAG, especially near the border of the aqueduct, contains a population of dopaminergic neurons that have been identified as the dorsocaudal part of the A10 group (Hasue and Shammah-Lagnado, 2002). These neurons project to the nucleus accumbens (Hasue and ShammahLagnado, 2002), and they may also innervate forebrain sites involved in arousal (Lu and Saper, unpublished observations). The PAG innervates areas involved in modulation of nociception including an input from the ventrolateral PAG to the A7 noradrenergic group (Bajic et al., 2001) and several thalamic regions, including the ventrobasal, intralaminar, and reticular nuclei (Rinvik et al., 1990; Carstens et al., 1990; Krout and Loewy, 2000). The PAG projection to the nucleus raphe magnus has also been associated with modulation of nociception (Beitz et al., 1983; Lakos and Basbaum, 1988; Guimaraes and Prado, 1999), however, many of the inputs from the PAG to the raphe magnus end on or near sympathetic premotor neurons (Farkas et al., 1998). Farkas and colleagues (1998) also examined the range of pathways by which PAG efferents may influence sympathetic responses by injecting different parts of the PAG with the anterograde tracer PHA-L in animals in which the inputs to the stellate ganglion were retrogradely identified by the transneuronal viral tracer, pseudorabies virus. They found that terminals from the lateral or ventrolateral PAG apposed retrogradely labeled (presympathetic) neurons in greatest numbers in the medullary raphe, but that smaller numbers of appositions were found in the ventrolateral medulla, the locus coeruleus, and the A5 cell group in the brainstem and in the paraventricular hypothalamic nucleus and the lateral hypothalamic area. There is evidence in cats that different sites in the PAG innervate pools of neurons in the RVL concerned with blood flow in different organs (Carrive et al., 1989; Carrive and Bandler, 1991a, 1991b). Evidence for similar topographic organization of the PAG projections to the RVL in the rat (van Bockstaele et al., 1990) hints that the differential connections of the PAG may support these differentiated responses in rats as well.

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Locus Coeruleus Although it might be tempting to consider the locus coeruleus, the brain’s largest source of noradrenergic input, to be a component of the central autonomic control system, it has been remarkably difficult to provide evidence for this hypothesis. Careful studies of the inputs to the core of the locus coeruleus reveal that it receives a very restricted range of inputs, mainly from the rostral ventrolateral and paramedian medulla (coextensive with the C1 and C3 adrenergic cell groups but mainly from noncatecholaminergic neurons) and from the lateral hypothalamic area and ventrolateral preoptic nucleus (see Aston-Jones et al., 1986; Ennis and AstonJones, 1986, 1987; Astier et al., 1990; Guyenet and Young, 1987; Pieribone et al., 1988; Luppi et al., 1995; Sherin et al., 1998; Lu et al., 2002). However, more recent studies have focused on inputs to the more distal dendrites of the locus coeruleus neurons, demonstrating inputs from other components of the central autonomic system, including the bed nucleus of the stria terminalis, central nucleus of the amygdala, and nucleus of the solitary tract (van Bockstaele et al., 1996, 1999a, 1999b; Bajic et al., 2000) The locus coeruleus projects widely in the brain, but does not contribute much to the noradrenergic innervation of the sympathetic preganglionic column, the RVL, the NTS, the PAG, or the paraventricular nucleus of the hypothalamus (Loewy et al., 1978; Ross et al., 1981; Sawchenko and Swanson, 1982; Kwiat and Basbaum, 1990; Fritschy and Grzanna, 1990). The locus coeruleus is retrogradely labeled transneuronally after injection of pseudorabies virus into the stellate ganglion (Farkas et al., 1998), and it provides no noradrenergic afferents to the lateral hypothalamus, the bed nucleus of the stria terminalis, the central nucleus of the amygdala, and the cerebral cortex (Jones and Moore, 1977; Loughlin et al., 1982, 1986; Peschanski and Besson, 1984). Recordings from neurons in the locus coeruleus in cats and monkeys indicate that they fire in relationship to arousing stimuli, including novel sensory events, pain and autonomic stimuli, slow down during the deeper stages of slow-wave sleep and almost cease firing during REM sleep (Aston-Jones and Bloom, 1981; Foote et al., 1983; Rassmussen et al., 1986; Grant et al., 1988), an activity profile that suggests a role in the maintenance of alertness or attention. Consistent with this view, the phasic firing of locus coeruleus neurons is associated with good performance on tasks that require focused attention, whereas tonic firing is associated with poor performance (Aston-Jones et al., 1999). Studies on the effects of norepinephrine on the cerebral cortex indicate that it causes an increase in the

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signal-to-noise ratio of neuronal firing, suppressing background activity but enhancing the responses to specific sensory stimuli (Foote et al., 1975). This effect would be consistent with the locus coeruleus playing a major role in preparing the cerebral cortex for efficient processing of sensory stimuli during arousal. However, lesions of the locus coeruleus produce only minor changes in the amount of wakefulness and circadian rhythms of wake–sleep cycles in cats (Jones et al., 1977) and rats (Gonzalez et al., 2002). Hence, although the locus coeruleus may play an important role in arousal responses to novel stimuli (Aston-Jones et al., 2001), its role in setting arousal tone may be limited to, or at least redundant with, parallel pathways from other arousal-related cell groups.

Dorsal and Median Raphe Nuclei The midbrain raphe nuclei provide the major ascending serotoninergic projection to the forebrain. Although they have not been implicated in playing a major critical role in autonomic regulation, there is evidence that they may modulate the activity of cell groups in the hypothalamus that are involved in autonomic control (Benarroch et al., 1983; Robinson et al., 1985; Petrov et al., 1992; Bell et al., 1999). The dorsal and median (also called the superior central) raphe nuclei receive substantial afferents from the parabrachial nucleus (Saper and Loewy, 1980; Lee et al., 2003), as well as from many of the same hypothalamic nuclei that innervate the PAG (see above). The role of the neurons in the midbrain raphe nuclei in regulating sleep and wakefulness is controversial. Early studies showed that lesions of the dorsal and median raphe nuclei, or depletion of serotonin with parachlorophenylalanine, caused insomnia, which was reversed when serotonin was restored with 5hydroxytryptophan (see Saper, 1987; Jouvet et al., 1989, for review). However, later studies showed that the activity patterns of single neurons in the midbrain raphe are similar to those of locus coeruleus neurons (slowing down during the deeper stages of slow-wave sleep and all but ceasing during rapid eye movement sleep; see Trulson et al., 1981; Lydic et al., 1987). Recent studies, however, suggest that the serotoninergic neurons in the midbrain raphe are intermixed with dopaminergic neurons. The dopaminergic neurons in the raphe region show Fos-expression suggesting activity primarily during wakefulness, whereas the serotoninergic neurons in this region lack the Fos-response during wakefulness (Lu et al., 2002b). These observations may explain the paradoxical earlier findings (i.e., the extracellular recordings in waking animals may have inadvertantly included dopaminergic wakeactive

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neurons, thus obscuring the responses of the serotoninergic population).

Pedunculopontine and Laterodorsal Tegmental Nuclei These mesopontine cholinergic cell groups are covered in detail in Chapter 36 (Butcher, this volume), and so their autonomic connections will be described only briefly here. They receive autonomic afferents from most of the same sources as the adjacent parabrachial nucleus and PAG (Moon-Edley and Graybiel, 1983; Herbert et al., 1990a; Steininger et al., 1992), although this innervation is typically less intense. They provide only a minor cholinergic projection to the cerebral cortex but may be responsible for substantial inputs to the hypothalamus and basal forebrain (Vincent et al., 1983; Hallenger and Wainer, 1988) and to the RVL (Yasui and Saper, 1991). The largest outputs from the pedunculopontine and laterodorsal tegmental nuclei, however, are to the thalamus and to the medial pontine and medullary reticular formation (Moon-Edley and Graybiel, 1983; Rye et al., 1988; Hallenger et al., 1987; Hallanger and Wainer, 1988; Krout and Loewy, 2002). These projections have been implicated in initiating the switching process from slow-wave to rapid eye movement sleep (see Rye et al., 1987; Datta and Siwek, 2002), which is accompanied by an increase in sympathetic tone (see Yasui et al., 1991; Trinder et al., 2001).

Cerebellum The finding of large increases in blood pressure during electrical stimulation of the fastigial nucleus in rats focused interest in the role of the cerebellum in autonomic control (see Andrezik et al., 1984; Iadecola et al., 1990). Although the fastigial nucleus does project to brainstem sites involved in cardiovascular control, including the RVL, it has not been possible to reproduce the blood pressure changes using cell-body-specific chemical stimulation. Subsequently, Paton and Spyer (1989, 1990) demonstrated both pressor and depressor pathways, originating from different parts of the vermis in rabbits. They reported projections through the region of the fastigial nucleus into the region of the parabrachial nucleus in this species. The author has been unable to confirm the presence of similar projections in rats (Saper et al., unpublished observations). However, a projection from the posterior vermis in rats does innervate the superior vestibular nucleus, which is caudally adjacent to the parabrachial nucleus. Data from lesions of the cerebellar vermis in cats suggest that it is concerned with maintaining blood pressure during an upright posture (Holmes et al., 2002).

In addition, the profound cardiovascular responses seen during vestibular vertigo suggest that there must be a direct and potent input from the vestibular system, possibly including the vermis, into the autonomic control system. Identifying this link will be an important problem for future investigation.

FOREBRAIN LEVEL: BEHAVIORAL AND METABOLIC INTEGRATION OF AUTONOMIC CONTROL AND AROUSAL Thalamus Ventroposterior Parvocellular Nucleus (VPpc) The ventroposterior medial parvocellular nucleus of the thalamus (VPMpc) was initially identified as a relay for gustatory information involved in taste discrimination (see Chapter 28 by Lundy and Norgren, this volume). More recent work, however, has demonstrated that the gustatory afferents occupy only the most medial part of this nucleus and that general visceral inputs terminate in the more lateral parts of the cell group, originally termed the ventroposterior lateral parvocellular nucleus (VPLpc; see Cechetto and Saper, 1987). As there is no sharp division between the medial (special visceral sensory) and lateral (general visceral sensory) parts of this cell group, the term VPpc has been introduced to describe the entire nucleus (see Yasui et al., 1989). The topographic ordering of visceral sensory inputs to the VPpc is similar to that of the NTS, with the gustatory afferents terminating at one extreme (medially), the cardiovascular and respiratory inputs at the opposite extreme (laterally), and the gastrointestinal afferents in between (Cechetto and Saper, 1987). However, in rats the NTS does not project directly to the VPpc; rather, the visceral afferent projection is relayed through the parabrachial nucleus (Norgren, 1976; Ricardo and Koh, 1978). These observations suggest that the viscerotopic organization of the NTS is preserved in the parabrachial relay as well. It has been remarkably difficult to test this hypothesis. First, the cells of origin of the parabrachial projection to the VPpc has been difficult to establish. The gustatory part of the NTS projects to a large terminal territory in the parabrachial nucleus in the rat, including much of the medial division and the waist area (see Norgren, 1976; Herbert et al., 1990), and tasteresponsive neurons can be recorded throughout this field (Ogawa et al., 1987; DiLorenzo, 1988; Nishijo and Norgren, 1990). Furthermore, many neurons within these regions project to the thalamus.

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More detailed tracing studies, however, have demonstrated that neurons in the medial parabrachial region and in the waist area have a rather larger target area in the thalamus, including the Vppc bilaterally, but with an ipsilateral predominance. Other axons from this site innervate the mediodorsal and the centromedial and other intralaminar nuclei (Yasui et al., 1989; Cechetto and Saper, 1987; Karimnamazi and Travers, 1998; Bester et al., 1999). By contrast, the external medial parabrachial subnucleus projects predominantly to the contralateral VPpc. This latter projection is best seen in material stained with antiserum against CGRP (see Fig. 11; Yasui et al., 1989) in which the VPpc is outlined by a large population of CGRP-immunoreactive terminals that originate from CGRP-containing neurons in the external medial PB. A second problem has been identifying the topographic organization of connections with a cell group that is as small as the external medial subnucleus.

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However, retrograde transport from small injections of retrograde tracers into either the medial or the lateral extreme of VPpc demonstrates that the input to the medial VPpc originates from the rostromedial part of the external medial subnucleus, and the projection to the lateral VPpc arises from the caudolateral part of the external medial subnucleus (Cechetto and Saper, 1987, and unpublished observations). Anterograde tracing studies from the NTS confirm that the gustatory rostral NTS projects most rostromedially and the caudal NTS most caudolaterally in the external medial subnucleus (Herbert et al., 1990). Hence, the topographic ordering of the main visceral sensory system is preserved at all levels, up through the thalamus and even the cortex (see below). Mediodorsal and Paraventricular Thalamic Nuclei Although the mediodorsal nucleus is most closely identified with the prefrontal cortex, the caudomedial part of the nucleus is the thalamic relay for the agranular insular cortex (Krettek and Price, 1977; Saper, 1982a). In addition, the most dorsomedial extreme of the mediodorsal nucleus, along with the adjacent part of the paraventricular nucleus, is related to the infralimbic cortex (Hurley et al., 1991; Fisk and Wyss, 2000). These two cortical areas are thought to be involved in the integration of limbic and autonomic response (see below). The parabrachial nucleus projects to the portion of the mediodorsal nucleus that innervates the insular area, as well as to the entire paraventricular thalamic nucleus (Saper, 1982; Bester et al., 1999). In addition, the paraventricular thalamic nucleus receives afferents from the median preoptic nucleus and the periventricular, paraventricular, ventromedial, and dorsomedial hypothalamic nuclei, as well as the lateral hypothalamic area and the suprachiasmatic nucleus (Saper et al., 1976, 1979a, 1979b; Saper and Levisohn, 1983; ter Horst et al., 1986; Simerly and Swanson, 1988; Moga and Moore, 1997). The role played by these thalamic nuclei in autonomic control remains largely unexplored. Intralaminar Nuclei

FIGURE 11 A summary drawing of the contribution to the visceral thalamocortical pathways made by CGRP-like immunoreactive neurons in the parabrachial nucleus and the thalamus. See text for details. exm, external medial parabrachial subnucleus; Ins, insular cortex; PoI, posterior intralaminar thalamic nuclei; Prh, perirhinal cortex; scp, superior cerebellar peduncle; v, ventral lateral parabrachial subnucleus; Vppc, ventroposterior parvicellular thalamic nucleus. Reprinted from Yasui et al. (1991) with permission.

The intralaminar nuclei of the thalamus constitute a diverse group of structures that are best characterized by their location within the internal medullary lamina that separates the lateral and medial tiers of thalamic relay nuclei and by their diffuse cortical projections. The intralaminar nuclei are covered in greater detail in Chapter 17 (Groenewegen and Witter, this volume) and so only their relationship to the autonomic system is considered here. The parafascicular, centromedial, centrolateral, and paracentral nuclei all receive substantial afferents from the parabrachial nucleus, originating in

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the internal lateral, central, and ventral lateral and medial subnuclei (Fulwiler and Saper, 1984; Krout and Loewy, 2000) as well as inputs from many other forebrain and brainstem sources (van der Werf et al., 2002; Krout and Loewy, 2002). As the internal lateral cell group receives its major afferents from the spinal and trigeminal dorsal horn (Cechetto et al., 1985; Slugg and Light, 1993; Feil and Herbert, 1995), it has been suggested that the parabrachial input to the intralaminar thalamic nuclei may contribute to the arousal that accompanies pain. The neurons in the ventral lateral and medial parabrachial nucleus that project to the thalamus mainly fall within the zone that receives gustatory and oral somatosensory afferents (see discussion in Herbert et al., 1990a). It is possible that the arousing affects of oral stimuli may be mediated by this pathway. A posterior group of intralaminar cell groups, including the subparafascicular, lateral subparafascicular, and posterior intralaminar nuclei and the peripeduncular nucleus, have been implicated in the transmission of conditioned auditory stimuli to areas in the amygdala that organize conditioned behavioral and cardiovascular autonomic responses (see Reis et al., 1984; Ledoux et al., 1990). The posterior intralaminar group provides a topographically organized projection to the amygdala, much of which originates from glutamate—or CGRP—immunoreactive neurons (Reis et al., 1985; Yasui et al., 1991; LeDoux and Farb, 1991)

Hypothalamus Anteroventral Third Ventricular Area The region surrounding the anteroventral tip of the third ventricle comprises a number of distinct nuclei, including the median preoptic nucleus, the anteroventral periventricular nucleus, and parts of the periventricular preoptic nucleus, as well as a circumventricular organ, the organum vasculosum of the lamina terminalis (OVLT). These cell groups share certain afferents, for example, from the subfornical organ and the parabrachial nucleus (Lind et al., 1982; Lind and Swanson, 1984; Saper and Loewy, 1980; Saper and Levisohn, 1983; Saper and Fulwiler, 1984). The projections from the anteroventral third ventricular nuclei also overlap considerably, providing the largest single input to the magnocellular neurons in the paraventricular and supraoptic nuclei and including parvocellular portions of the periventricular and paraventricular hypothalamic nuclei and the lateral hypothalamic area and sending small numbers of axons descending through the periaqueductal gray matter and the pontine tegmentum to the parabrachial nucleus and

the nucleus of the solitary tract (Simerly and Swanson, 1984; Saper and Levisohn, 1983; Gu and Simerly, 1997). Simerly and colleagues (1985, 1986, 1998) have examined in detail the distribution of a variety of neurotransmitter-specific fiber types in this area. Many neurons in the anteroventral periventricular nucleus are immunoreactive with antisera against atrial natriuretic peptide (Standaert et al., 1986). These latter neurons are the main source of atrial natriuretic peptideimmunoreactive innervation of the paraventricular and supraoptic nuclei (Standaert and Saper, 1988; Hurley et al., 1992). Lesions in the anteroventral third ventricular area cause profound disruption of fluid and electrolyte balance, ranging from decreased drinking to dysregulation of plasma osmolality and blood volume and pressure (Brody and Johnson, 1981; Johnson, 1985; Colombari and Cravo, 1999). In addition, such lesions are associated with loss of thermoregulation and absence of a febrile response to immune stimulation (Stitt, 1985; Szymusiak et al., 1985; Blatteis et al., 1987; Whyte and Johnson, 2002). Pharmacological studies show that the fever response is due to the action of prostaglandins on neurons near the anteroventral tip of the third ventricle (Scammell et al., 1997; Nakamura et al., 2002) The elevation of body temperature depends critically on the redirection of blood flow from cutaneous to deep vascular beds, emphasizing the close relationship between thermoregulation and cardiovascular control. In addition, many neurons in the median and periventricular preoptic nuclei contain luteinizing hormone-releasing hormone (Witkin et al., 1982; King et al., 1982; Simerly et al., 1985) and participate in reproductive hormone and behavior regulation. Many aspects of reproduction are related to cardiovascular regulation, ranging from the redistribution of blood flow during sexual arousal to the fluid shifts that accompany hormonal cycles and pregnancy. Paraventricular Nucleus The paraventricular nucleus of the hypothalamus represents a microcosm of homeostatic control mechanisms. It consists of a number of distinct cell subnuclei, subserving a variety of neuroendocrine and autonomic functions. In general, it may be divided into magnocellular and parvocelluar divisions. The magnocellular neurons contain oxytocin (anterior and medial magnocellular and rim of the posterior magnocellular groups) or vasopressin (the main body of the posterior magnocellular group), which are released from axon terminals in the posterior pituitary gland. Many neurons in the medial parvicellular subnucleus contain releasing hormones (especially corticotropin-releasing hormone), which they release from axon terminals onto the hypo-

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physial portal vessels in the median eminence. The neuroendocrine aspects of the paraventricular nucleus are covered in detail in Chapter 15 (Armstrong, this volume). The remaining dorsal, ventral, and lateral parvicellular subnuclei project mainly to structures involved in central autonomic control, including the periaqueductal gray matter, parabrachial nucleus, NTS, RVL, and to both parasympathetic and sympathetic preganglionic populations in the medulla and the spinal cord (Saper et al., 1976; Sawchenko and Swanson, 1983; Luiten et al., 1985, 1987). Some of these neurons may send collaterals to several sites along this path (Pyner and Coote, 2000). The projections from the paraventricular nucleus to autonomic structures was first shown by Swanson (1977) to include many oxytocin-containing axons. Retrograde transport studies demonstrated that few if any of these cells also projected to the pituitary gland, and only a small percentage of paraventricular neurons projecting to the spinal cord were doublestained with antisera against oxytocin or vasopressin (Swanson and Kuypers, 1980a; Sawchenko and Swanson, 1983). Later studies using colchicine to enhance peptide immunoreactivity or combining retrograde tracing with in situ hybridization demonstrated that the majority of paraventricular neurons projecting to the spinal cord are either oxytocin- or vasopressin-immunoreactive (Cechetto and Saper, 1988). More recent studies, combining in situ hybridization with retrograde tracing indicate that at least 40% of the paraventriculospinal neurons contain oxytocin and 40% vasopressin (Hallbeck et al., 1999; 2001). These parvocellular neurons are distinct from the magnocellular endocrine cells, and they stain considerably less intensely with antisera against oxytocin and vasopressin. About 40% of the paraventriculospinal neurons contain dynorphin mRNA and 20% enkephalin (Hallbeck et al., 1999, 2001). Some paraventricular neurons that project to the parabrachial nucleus also stain with antisera against oxytocin, vasopressin, dynorphin, or enkephalin, but most remain uncharacterized (Moga et al., 1990) Both anterograde transport studies (Saper et al., 1976; Luiten et al., 1985) and immunocytochemical staining (Swanson, 1977; Swanson and McKellar, 1979; Sofroniew, 1983) have demonstrated the pathway taken by paraventricular descending fibers. Axons run both through the periventricular hypothalamus into the periaqueductal gray matter and through the lateral hypothalamic area into the lateral brainstem tegmentum. As this fiber pathway runs along the ventrolateral surface of the medulla, it provides projections to the parasympathetic preganglionic neurons in the dorsal motor vagal nucleus and the nucleus ambiguus. Paraventricular axons then run through the lateral funiculus of the spinal cord to

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innervate the entire length of the thoracic (sympathetic) and sacral (parasympathetic) preganglionic cell columns. However, this innervation is not evenly distributed. McKellar and Swanson (1979) found that oxytocinimmunoreactive axons terminated in a patchy distribution on clusters of preganglionic neurons, with much more intense innervation of some spinal levels than others. These observations hint at a chemoanatomicalfunctional organization to the paraventriculospinal projection, which remains a fertile area for future investigation. There is some connectional evidence as well for differentiation of function among the parvocellular subnuclei of the paraventricular nucleus. For example, all three subdivisions project to the spinal cord, but only the dorsal and lateral subnuclei innervate the dorsal vagal complex (Swanson and Kuypers, 1980a). Studies employing retrograde transneuronal transport of viruses have found that only selective clusters of paraventricular parvocellular neurons are labeled when injections are made into the adrenal medulla or into different sympathetic ganglia (Strack et al., 1989a, 1989b; Sved et al., 2001), suggesting a topographic organization. In addition, certain physiological stimuli produce Fos expression in only subsets of the hypothalamospinal projection. For example, injection of intravenous lipopolysaccharide cause Fos expression selectively in spinally projectingneurons in the dorsal parvicellular paraventricular nucleus (Zhang et al., 2000), whereas intravenous leptin causes Fos expression in neurons in the arcuate nucleus that project to the sympathetic preganglionic column, but not in any of the paraventriculospinal neurons (Elias et al., 1998). Interestingly, after lipopolysaccharide Fos-positive neurons were retrogradely labeled only in the dorsal parvicellular paraventricular nucleus, regardless of the level of the spinal cord at which the retrograde tracer injections were placed (Zhang et al., 2000). These observations suggest that the hypothalamic-autonomic projection may be organized in a functional-anatomic pattern rather than a strictly topographic-anatomic pattern. In other words, clusters of hypothalamospinal neurons that are specific for a particular response may contact a range of autonomic preganglionic targets that are involved in eliciting that response (see Saper, 2002). Electrical or chemical stimulation studies of the paraventricular nucleus likewise provide support for functional differentiation of its subnuclei. Either increases or decreases of blood pressure have been reported, depending on the exact site of stimulation (Porter and Brody, 1986; Darlington et al., 1989; Gelsema et al., 1989). However, the relationship of stimulation sites with specific cell populations or projections has not been investigated. More specific information is

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available concerning an oxytocin-containing projection from the paraventricular nucleus to dorsal vagal motor neurons controlling gastric motility and acid secretion (Rogers and Hermann, 1987). Injection of oxytocin into the dorsal motor vagal nucleus reproduced the response, and injection of an oxytocin antagonist into the dorsal motor vagal nucleus blocked the gastric response to paraventricular stimulation. Sympathetic preganglionic neurons are excited by direct application of either oxytocin or vasopressin (Gilbey et al., 1982) or by paraventricular nucleus stimulation (Caverson and Ciriello, 1988; Martin and Haywood, 1992). Dorsal Hypothalamic Area The dorsal hypothalamic area, lying just dorsal and caudal to the paraventricular nucleus, also provides a spinal projection, originating mainly from dopaminergic neurons of the A11 catecholamine group (see Fig. 12; Björklund and Skagerberg, 1979; Hökfelt et al., 1979; Cechetto and Saper, 1988). These neurons probably supply the dopaminergic innervation of the sympathetic preganglionic column as well as of the superficial dorsal horn (Skagerberg et al., 1982). Strack and colleagues (1989a) reported tyrosine hydroxylaseimmunoreactive neurons in the paraventricular nucleus of the hypothalamus contained transneuronally transported virus particles, after an injection into the superior cervical ganglion. However, it likely that they had identified the dorsal hypothalamic dopaminergic group as part of the paraventricular nucleus. A later article from the same group (Jansen et al., 1995) using the same method after injecting the stellate ganglion did not report any retrograde labeling of tyrosine hydroxylase-immunoreactive neurons in the paraventricular nucleus but did identify retrogradely labeled neurons (of unidentified chemical type) in the dorsal hypothalamic area. Dorsomedial Nucleus of the Hypothalamus The dorsomedial nucleus, like the paraventricular nucleus, receives substantial afferents from the NTS and the parabrachial nucleus and contributes to the descending projection to the parabrachial nucleus, vagal complex, and the spinal cord (Saper et al., 1976; Ricardo and Koh, 1978; Saper and Loewy, 1980; Fulwiler and Saper, 1984; ter Horst et al., 1986; Luiten et al., 1987; Moga et al., 1990a, 1990b). Some neurons along the lateral edge of the dorsomedial nucleus that contribute to these projections contain orexin or melaninconcentrating hormone (see next section), but the proportions of spinally projecting dorsomedial nucleus neurons that contain these neurotransmitters remains unknown. Chemical stimulation of the dorsomedial nucleus produces increases in blood pressure and heart

rate, whereas injections of GABA receptor agonists into the same region block cardiovascular responses to stress (Soltis and Dimicco, 1990, 1992). These cardiovascular responses are associated with hyperthermia and appear to be mediated by the raphe pallidus nucleus (Zaretskaia et al., 2002; Samuel et al., 2002). The dorsomedial nucleus receives extensive afferents from the suprachiasmatic nucleus and the subparaventricular zone, two critical components of the circadian system and mediates guide range of circadian responses (see Chou et al., 2003). Thus, the dorsomedial nucleus may mediate at least in part the circadian modulation of sympathetic tone (Ueyama et al., 1999). Lateral Hypothalamic Area The lateral hypothalamic area and zona incerta at the level of the tuberal hypothalamus provide projections both to the parabrachial nucleus and the spinal cord (Cechetto and Saper, 1988; Moga and Saper, 1990). These neurons include a perifornical population as well as neurons that invade the overlying zona incerta and some that extend as far laterally as the cerebral peduncle (see Fig. 12; Cechetto and Saper, 1988). The autonomic projections originate from two interdigitated populations of lateral hypothalamic neurons, most of which express either orexin (also known as hypocretin) or melanin-concentrating hormone (Peyron et al., 1998; Bittencourt and Elias, 1998; Elias et al., 1998), and sympathetic preganglionic neurons express orexin-2 receptors (Marcus et al., 2001 and unpublished results). Previous studies also demonstrated that some of the hypothalamospinal neurons are immunoreactive for dynorphin (Cechetto and Saper, 1988), but it is now recognized that virtually all of these neurons also contain orexin (Chou et al., 2001). Early studies had also identified a population of spinally projecting lateral hypothalamic neurons that were immunoreactive for α-melanocyte stimulating hormone or acetylcholinesterase (Köhler and Swanson, 1984; Cechetto and Saper, 1988), but these cells do not contain pro-opiomelanocortin, the precursor for α-MSH (Saper et al., 1986), and it now appears that these stains identified mainly neurons that contained orexin or melanin-concentrating hormone (Rotman, Chou, and Saper, unpublished observations). Orexin- and melaninconcentrating hormone-immunoreactive axons in the spinal cord innervate the sympathetic preganglionic column as well as lamina I (Skofitsch et al., 1985; van den Pol, 1999), consistent with earlier anterograde tracer studies (Saper et al., 1976; Luiten et al., 1987). Electrical or chemical stimulation in the lateral hypothalamic area produces increases in blood pressure, heart rate, and renal sympathetic nerve discharge (Spencer et al., 1988; Cechetto and Chen, 1992; Sun and Guyenet,

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FIGURE 12 Chemical composition of the hypothalamic projection to the spinal cord. In this series of schematic drawings of transverse sections illustrating four levels through the hypothalamus from rostral (upper left) to caudal (lower right), the approximate locations of the five different populations of hypothalamospinal neurons are indicated by different shading, and the approximate percentages of neurons that stain immunocytochemically for various putative neurotransmitters or their synthetic enzymes are indicated. AHc, central part of the anterior hypothalamic area; AHP, posterior part of the anterior hypothalamic area; ANP, atrial natriuretic peptide; Arc, arcuate nucleus; AVP, arginine vasopressin; DA, dorsal hypothalamic area; DC, dorsal cap parvicellular division of the paraventricular nucleus; DM, dorsomedial hypothalamic nucleus; DMC, dorsomedial nucleus compact division; DYN, dynorphin; ENK, enkephalin; f, column of the fornix; ic, internal capsule; LM, lateral magnocellular division of the paraventicular nucleus; MCH, melanin-concentrating hormone; MP, medial and ventral parvicellular divisions of the paraventricular nucleus; mt, mammillothalamic tract; opt, optic tract; ORX, orexin; OXY, oxytocin; Pa, paraventricular nucleus; Po, posterior lateral parvocellular division of the paraventricular nucleus; POMC/CART, neurons containing both pro-opiomelanocortic and CART (cocaine and amphetamine responseive transcript), RCh, retrochiasmatic area; SO, supraoptic nucleus; SOC, ventral supraoptic commissure; STh, subthalamic nucleus; VMH, ventromedial nucleus; ZI, zona incerta; 3V, third ventricle. Reprinted from Cechetto and Saper (1988), with permission.

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1986; Allen and Cechetto, 1992). However, the effects of the different neurotransmitters on specific populations of preganglionic neurons remains a fertile area for future investigation. Many lateral hypothalamic neurons that contain either orexin- or melanin-concentrating hormone also project to the cerebral cortex, but very few of cells project both to the cortex and the spinal cord (Köhler et al., 1984; Saper et al., 1986; Bittencourt et al., 1992; Peyron et al., 1998; Cechetto and Saper, unpublished observations). Arcuate Nucleus and Retrochiasmatic Area A modest but important population of neurons in the retrochiasmatic area and adjacent arcuate nucleus project either to the spinal cord (see Fig. 12; Swanson and Kuypers, 1980a; Cechetto and Saper, 1988) or to the parabrachial nucleus (Moga et al., 1990a). It has not been determined whether the latter pathway may represent collaterals of the spinal projection. The arcuate nucleus and retrochiasmatic area include many neurons that contain releasing hormones (Hökfelt et al., 1989), and many of the neurons contributing to the descending pathways from this region contain αmelanocyte stimulating hormone, as well as other products of the proopiomelanocortin gene, such as βendorphin and ACTH (Cechetto and Saper, 1988; Moga et al., 1990b). These neurons are activated by leptin (Elias et al., 1998a; Cowley et al., 2001), and they also contain cocaine and amphetamine-related transcript (CART), another peptide modulator. The α-melanocyte stimulating hormone/CART neurons project to the spinal cord, and they are believed to modulate sympathetic responses associated with satiety. The remarkable specificity of the hypothalamo-spinal neurons that are activated by leptin, for the α-melanocyte stimulating hormone/CART population supports the notion that the hypothalamic–autonomic projections may be primarily organized along functional lines (Saper, 2002). Posterior Lateral Hypothalamic Area The posterior lateral hypothalamic area, adjacent to the subthalamic nucleus, contains a discrete population of neurons that contain neither orexin- nor melaninconcentrating hormone, but provide projections both to the cerebral cortex and to the parabrachial nucleus (Saper, 1985; Saper et al., 1986; Moga et al., 1990a,b) and not to the spinal cord (Cechetto and Saper, 1988). This same region receives intense afferents from the parabrachial nucleus and from the infralimbic and insular cortical areas (Saper and Loewy, 1980; Saper et al., 1982; Yasui et al., 1991; Hurley et al., 1991). Injection of cobalt ions into this region blocks cardiovascular responses elicited from electrical stimulation of the insular cortex (Cechetto and Chen, 1990),

suggesting that, like the tuberal lateral hypothalamus, it may play an important role in the integration of autonomic mechanisms with behavior and ascending arousal responses. Tuberomammillary Nucleus The tuberomammillary nucleus is the main histaminergic cell group in the rat brain, providing widespread innervation ranging from the brainstem to the hypothalamus and the cerebral cortex (Vincent et al., 1983; Panula et al., 1984; Köhler et al., 1985). Many of its cells also contain other putative neurotransmitters, such as GABA, adenosine, brain natriuretic peptide, and galanin (Semba et al., 1985; Köhler et al., 1985; Saper et al., 1989). Like the locus coeruleus, its axons tend to diverge widely, innervating sites as far apart as the preoptic area and midbrain or even sending collaterals into both hemispheres (Vincent et al., 1983; Köhler et al., 1985; Inagaki et al., 1990). Although early studies had difficulty in identifying afferents to tuberomammillary neurons (Ericson et al., 1991), more recent work has found that they receive intense inhibitory inputs from the ventrolateral preoptic area (which is active during sleep; see Sherin et al., 1996, 1998) and the orexin neurons (which are active during wakefulness; Peyron et al., 1998; Chemelli et al., 1999; Estabrooke et al., 2001). Physiological studies have implicated the tuberomammillary nucleus in maintaining cortical arousal and a waking state, particularly in response to orexin signaling (Lin et al., 1988, 1989; Huang et al., 2001).

Amygdala Central Nucleus The cytoarchitecture and subdivisions of the central nucleus of the amygdala are covered in detail in Chapter 19 (de Olmos et al., this volume); this chapter concentrates on its connections with the central autonomic control system. The central nucleus receives substantial inputs both from the NTS and the parabrachial nucleus (Ricardo and Koh, 1978; Norgren, 1976; Saper and Loewy, 1980; Fulwiler and Saper, 1984; Ottersen et al., 1981; Bernard et al., 1993). Many of the cells in the NTS that contribute to this projection contain somatostatin, dynorphin, enkephalin, neuropeptide Y, or norepinephrine (Riche et al., 1990), whereas many of the parabrachial cells (particularly in the external lateral nucleus) contain substance P, neurotensin, or calcitonin gene-related peptide (Schwaber et al., 1988; Yamano et al., 1988; Block et al., 1989). The parabrachial projection is topographically organized, with the outer part of the external lateral parabrachial nucleus projecting to the lateral part of the central nucleus and the inner

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part of the external lateral nucleus projecting to the laterocapsular part of the central nucleus (Bernard et al., 1993). The central nucleus is a major termination site for descending projections from the agranular insular cortex (Saper, 1982; Yasui et al., 1991a) and the posterior intralaminar thalamic complex (Ledoux et al., 1990; Yasui et al., 1991b). The latter projection arises in large part from cells that are immunoreactive with antisera against calcitonin gene-related peptide. The major projections from the central nucleus descend via the lateral hypothalamic area to reach the periaqueductal gray matter, parabrachial nucleus, ventrolateral medulla, and NTS (Krettek and Price, 1978; Ross et al., 1981; Veening et al., 1984; Cassell and Gray, 1989; Moga et al., 1990a). These descending projections originate mainly from GABAergic neurons within the central nucleus (Jongen-Relo and Amaral, 1998; Saha et al., 2000), and many cells GABAergic cells in the central nucleus also contain peptides such as enkephalin or corticotropin-releasing hormone (Vienante et al., 1997). Hence, many neurons in the lateral part of the central nucleus that project to the parabrachial nucleus are immunoreactive with antisera against corticotropin-releasing hormone, neurotensin, or somatostatin (Moga and Gray, 1985) and those that project to the nucleus of the solitary tract stain with antisera against somatostatin, neurotensin, and vasoactive intestinal peptide (Batten et al., 2002). There have been reports of a few cells in the central nucleus projecting as far as the spinal cord in monkeys and cats (Mizuno et al., 1985; Sandrew et al., 1986) but it has not been possible to identify such a projection in rats (Saper, unpublished observations). Neurons in the central nucleus respond to cardiovascular baroreceptor information relayed through the parabrachial nucleus in the cat (Cechetto and Calaresu, 1985). A variety of cardiovascular responses have been obtained from stimulation in the central nucleus, but they have not been successfully localized to specific subnuclei (Iwata et al., 1987; Cox et al., 1987; Gelsema et al., 1987; Bakhlavadzhyan et al., 2000). The central nucleus appears to be a critical component in producing a conditioned cardiovascular fear response (Ledoux et al., 1990; Roozendaal et al., 1990; Markgraf and Kapp, 1991; Nader et al., 2001; Cain et al., 2002). Basolateral Complex The basal and lateral nuclei of the amygdala are mainly related to the cerebral cortex. They are discussed in considerable detail in Chapter 19 (de Olmos et al., this volume), and this chapter confines itself to its relationship to autonomic responses. The basolateral complex receives inputs from the cortex, including the insular region (Saper, 1982; Shi and Cassell, 1998), and

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from the posterior intralaminar thalamic complex (Reis et al., 1985; Yasui et al., 1991). Like the projection from this latter group to the central nucleus, many of the thalamic axons that innervate the basolateral complex are immunoreactive with antisera against calcitonin generelated peptide. The behavioral aspects of the conditioned fear response to a tone stimulus appear to be dependent on the integrity of this thalamic projection into the lateral nucleus (Ledoux et al., 1990; Nader et al., 2001). It is interesting that the basolateral complex also provides some intraamygdaloid input to the central nucleus (Krettek and Price, 1977; Otterson et al., 1982; Pare et al., 1995; Savander et al., 1995; Pitkanen et al., 1995), which may allow the integration of cognitive (and perhaps arousing) aspects of an emotional response with the autonomic concomitants of the experience. Extended Amygdala and Bed Nucleus of the Stria Terminalis The concept of the extended amygdala and its various subdivisions are discussed in Chapter 19 (De Olmos et al., this volume). One of the earliest pieces of evidence for this concept was provided by retrograde transport studies, which demonstrated that the neurons that project to the NTS extend from the central nucleus, through the substantia innominata, and into the bed nucleus of the stria terminalis in an unbroken chain (Schwaber et al., 1982; Ross et al., 1981). Similar observations have been made concerning the projection to the parabrachial nucleus (Moga et al., 1989, 1990a). Both the parabrachial and NTS projections arise from neurons containing corticotropin-releasing factor, somatostatin, and neurotensin in both the lateral part of the central nucleus of the amygdala and the dorsolateral subnucleus of the bed nucleus of the stria terminalis (Moga et al., 1985, 1989; Gray and Magnuson, 1987). The bed nucleus of the stria terminalis, substantia innominata, and central nucleus of the amygdala also receive afferents from the same parts of the NTS, VLM, and parabrachial complex (Ricardo and Koh, 1978; Saper and Loewy, 1980; Fulwiler and Saper, 1984; Grove, 1988a,b; Moga et al., 1990a; Woulfe et al., 1990; Riche et al., 1990). The projections from the dorsolateral nucleus have been studied in some detail (Dong et al., 2001) and are quite similar to those from the lateral part of the central nucleus. They contact both the lateral hypothalamus and ventrolateral part of the periaqueductal gray matter but end more heavily in the lateral parabrachial nucleus, particularly in the inner part of the external lateral subnucleus. Although there is some evidence that neurons in the bed nucleus of the stria terminalis respond to cardiovascular stimuli (Hilton and Spyer, 1981), and stimulation of the bed nucleus can cause

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increased gastric motility (Hermann et al., 1990), there has been no systematic attempt to relate the subnuclei of the bed nucleus of the stria terminalis or their connections with specific autonomic responses. The function of at least the dorsolateral subnucleus of the bed nucleus may be correlated with that of the lateral part of the central nucleus of the amygdala. For example, intraperitoneal injection of interleukin-1β causes expression of Fos in GABAergic neurons, most of which also contain enkephalin mRNA in both sites (Day et al., 1999).

Cerebral Cortex Insular and Perirhinal Cortex The insular cortex of the rat occupies the dorsal bank of the rhinal sulcus and extends dorsally to the borders of the primary and secondary somatosensory areas (see Fig. 13). It has been divided into an anterior region, which is mainly agranular and a posterior region with somewhat better developed dysgranular and granular regions occupying the dorsal portion of the field. Caudally, the granular areas disappear at about the

level of the foramen of Monro, and the agranular insular cortex merges imperceptibly with the perirhinal cortex. The dysgranular insular cortex, which is most prominent at about the rostral level of the genu of the corpus callosum, receives the bulk of the gustatory afferents in rat (see also Chapter 28, Lundy and Norgren, this volume), whereas the granular insular area, which is more prominent caudally, receives predominantly general visceral afferents (Cechetto and Saper, 1987; Kosar and Norgren, 1986a; Ogawa et al., 1991; Hanamori et al., 1998). Furthermore, the general visceral area is topographically organized, with the neurons responding to gastrointestinal stimuli placed most rostrally and dorsally, adjacent to the taste cortex, and the cardiovascular- and respiratory-responsive neurons located most caudally in the granular field. On the other hand, there is some convergence of different visceral stimuli to individual neurons, particularly between functionally related classes of stimuli such as taste inputs and gastric stretch (Cechetto and Saper, 1987; Hanamori et al., 1998). The general topographic pattern of organization, which is identical to that in the NTS, suggests the maintenance of strict

FIGURE 13 Subdivisions of the insular cortex. These photomicrographs taken at middle (A) and caudal (B) levels of the insular cortex demonstrate the laminar patterns that distinguish the subdivisions. Notice the presence of a relatively dense population of granule cells in layer IV of the granular field (GI); less dense granule cells and a more prominent layer V in the dysgranular field (DI); and near absence of layer IV granule cells in the agranular area (AI). CLA, claustrum; EN, endopiriform nucleus; PIR, piriform cortex; S II, secondary somatic sensory cortex. Scale = 0.5 mm. Reprinted from Cechetto and Saper (1987), with permission.

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topographic ordering throughout the visceral sensory pathway to the cerebral cortex. The same pattern of topographic ordering is also apparent in the thalamic input to the insular cortex. The thalamic relay for the granular insular cortex is the lateral part of the VPpc; the medial VPpc supplies the dysgranular insular cortex (Kosar and Norgren, 1986b; Cechetto and Saper, 1987; Stehberg et al., 2001; Barnabi and Cechetto, 2001). The medial division of the parabrachial nucleus, along with the ventral lateral and waist subnuclei, projects directly to all subfields of the insular cortex and to the perirhinal cortex (Saper, 1982b; Allen et al., 1991); the projection arises in part from neurons that are immunoreactive with antisera against calcitonin gene-related peptide (see Fig. 11; Yasui et al., 1989). Although the type of information relayed by the direct parabrachial input is not known, the densest part of the projection is to the agranular insular field, in which few neurons respond to visceral stimuli, but from which most descending projections of the insular cortex arise (see below). The thalamic relay for the agranular insular cortex is the posteromedial edge of the mediodorsal nucleus (Saper, 1982a), which also receives a medial parabrachial input (Saper and Loewy, 1980). In addition, neurons in the posterior intralaminar thalamic complex along the borders of the VPpc (including the subparafascicular and lateral subparafascicular nuclei) innervate the perirhinal cortex. As is the case with the posterior intralaminar projection to the amygdala (see above), this input to the cortex arises in large part from neurons that are immunoreactive with antisera against calcitonin gene-related peptide (Yasui et al., 1989). Descending projections from the insular cortex back to the hypothalamus, parabrachial nucleus, and nucleus of the solitary tract mainly originate in the agranular field (Saper, 1982; Moga et al., 1990a; Yasui et al., 1991a; Hayaaw and Ogawa, 2001). Electrical or chemical stimulation of the posterior part of the agranular insular cortex can produce a variety of autonomic responses: depressor-bradycardic responses are seen at the most caudal sites; increases in blood pressure with tachycardia can be evoked at slightly more rostral sites; and increased gastric motility is seen with stimulation even further rostrally (Ruggiero et al., 1987; Yasui et al., 1991a). Both the pressor-tachycardic and depressorbradycardic responses could be blocked by injection of cobalt ions, interrupting synaptic transmission in the posterior lateral hypothalamus (Cechetto and Chen, 1990). The agranular insular cortex also innervates the infralimbic area. It is interesting that the preponderance of visceral sensory afferents to the insular cortex terminate in the granular and dysgranular fields, whereas the prominent efferent projections from the insular cortex to autonomic structures arise from the

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agranular field (Allen et al., 1991). These observations suggest substantial communication and differentiation of function between the components of the insular cortex. Infralimbic and Prelimbic Cortex The infralimbic cortex is a poorly laminated region along the most medial part of the medial prefrontal cortex (see Vogt et al., Chapter 22, this volume). Although it was originally omitted from the prefrontal cortex, based on the absence of a mediodorsal nucleus projection to this region (Krettek and Price, 1977), later studies found it to be reciprocally related with neurons along the most dorsomedial edge of the mediodorsal nucleus and extending into the paraventricular nucleus of the thalamus (see Groenewegen, 1988; Hurley et al., 1991). The adjacent prelimbic cortex is related to the adjacent dorsomedial part of the mediodorsal nucleus. The prelimbic cortex receives extensive projections from limbic regions, including the cingulate, entorhinal, and subicular cortices (Reep, 1984). In contrast, the infralimbic cortex receives only selected limbic afferents, such as projections from the CA1 field of the hippocampus and from the prelimbic cortex (Swanson et al., 1981; Hurley et al., 1991), but it receives no afferents from central autonomic structures, such as the parabrachial nucleus (Saper, 1982b). The prelimbic cortex projects back to limbic cortical regions but has modest descending projections, mainly to the lateral hypothalamus, PAG, parabrachial nucleus and NTS (Sesack et al., 1989; Hurley et al., 1991). The infralimbic area, again in contrast, has only limited projections to cortical limbic areas, but instead provides an extensive descending projection to the central nucleus of the amydgala, the bed nucleus of the stria terminalis, the lateral hypothalamic area, the dorsomedial hypothalamic nucleus, the PAG, the parabrachial nucleus, the RVL, the NTS and the spinal cord, including both lamina I and the sympathetic preganglionic column (Hurley et al., 1991; Takagishi and Chiba, 1991; Fisk and Wyss, 2000). Electrical stimulation of the infralimbic cortex has been shown to affect gastric motility and to cause hypotension (Hurley-Gius and Neafsey, 1986; Pantelew and Grundy, 2000; Fisk and Wyss, 2000). On the basis of these experiments, it has been suggested that the infralimbic area may serve as a visceral motor cortex, as opposed to the visceral sensory area in the insular cortex. Interestingly, the hypotensive response can be largely eliminated by injection of cobalt chloride into the lateral hypothalamus, suggesting an obligate relay for this influence (Fisk and Wyss, 2000). However, the demonstration that the agranular insular field is more closely related to visceral motor than sensory function (see above) suggests that the analogy with the somatic sensory and motor system may not be entirely applicable.

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Motor and Sensory Cortex Although not usually considered to constitute a part of the autonomic control system, the close coordination of autonomic and somatomotor responses begins at the cortical level. For example, motor “central command” produces an increase in blood pressure during attempted exercise, even if the muscle involved has been paralyzed and no actual contraction occurs electrical stimulation of sensory or motor cortex also can produce blood pressure responses, although the mechanism for these changes is not known (Wall and Davis, 1951). Studies by Ito (2002) have identified a site in the anterior tip of the somatosensory cortex in rats that responds to stimulation of the cervical vagus nerve, but this may represent somatosensory fields from intraoral sites.

SUMMARY AND CONCLUSION In retrospect, it should not be surprising that so pervasive an aspect of everyday life as autonomic response should have such a widely distributed, complex, and redundant representation in the brain. Certainly, the striking degree of interconnectivity of the structures composing the central autonomic system, involving every level of the neuraxis, belies a heirarchical organization of autonomic control. In fact, the nuances of control of a series of interrelated systems that are responsible both for maintaining homeostasis, and for dealing with the exigencies of behavioral perturbations, would dictate the need for continuous and extensive feedback and communication. Now that the outlines of the central autonomic control system have been defined, the challenge for the future will be to define the specific connections responsible for different aspects of autonomic control and ultimately to determine their relationships with one another.

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25 Somatosensory System DAVID TRACEY School of Medical Sciences, University of New South Wales New South Wales, Australia

The somatosensory system involves those parts of the nervous system that provide information from somatosensory receptors to the cerebral cortex. This includes the sensory receptors and primary afferent neurons, ascending spinal pathways, somatosensory nuclei in the brainstem and thalamus, and the somatosensory regions of the cortex. This chapter provides an overview of that part of the system concerned with the limbs and trunk, whereas the trigeminal system is dealt with in Chapter 26.

and Ide, 1988), which help to determine the rate of adaptation. Merkel Endings Merkel endings are located in the epidermis of glabrous and hairy skin and are expanded nerve terminals associated with specialized cells, the Merkel cells (Fig. 1A). Merkel cells contain neuropeptides and serotonin (Weihe et al., 1998) and modulate the response of the Merkel ending and also have trophic effects on neurons and keratinocytes (Tachibana, 1995). The Merkel endings are slowly adapting mechanoreceptors (SAI) and have small receptive fields. Mechanical stimulation of Merkel endings results in increased levels of intracellular Ca2+ (Senok and Baumann, 1997), suggesting that Merkel cells play a role in the transduction process (Ogawa, 1996). Recent work provides evidence that glutamate acts as a neurotransmitter between Merkel cells and nerve terminals (Fagan and Cahusac, 2001). In hairy skin, Merkel endings occur as compact clusters of 50–70, associated with the terminals of a single myelinated axon and located beneath an elevation of the skin known as a touch dome or Haarscheibe (Casserly et al., 1994). Merkel endings are particularly dense in the follicles of sinus hairs on the face (Rice et al., 1997). In glabrous skin they are located at the base of rete ridges or pegs, and they are also present in palatine mucosa (Tachibana et al., 1997).

SOMATOSENSORY RECEPTORS These are sensory receptors that detect mechanical, thermal, or noxious stimuli and are generally located in skin, muscle, or joints. They are the peripheral terminals of primary afferent neurons, with cell bodies in the dorsal root ganglia or in the trigeminal ganglion. The sensory terminals of nociceptors and thermoreceptors are free nerve endings without accessory structures, and their axons are unmyelinated or thinly myelinated (see Willis et al., Chapter 27). The sensory terminals of mechanoreceptors are associated with accessory structures or cells that affect the rate of adaptation of the receptor to a constant stimulus, and their afferent fibers are myelinated.

Cutaneous Receptors

Ruffini Endings

Cutaneous receptors may be classified according to the morphology of their accessory structures (Munger

These are found in the dermis of the skin, where their unmyelinated terminal branches are intertwined

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sp

A

k

k

k d

k

k

k

dcv

k

bl

Merkel Receptor Complex

B Ruffini Corpuscle

Golgi Tendon Organ

FIGURE 1 Mechanoreceptor endings. (A) Merkel ending partially enveloped by a Merkel cell. Abbreviations: bl, basal lamina; d, desmosome; dcv, dense-cored vesicles; k, keratinocyte; sp, spine-like process. (B) A Ruffini corpuscle and a Golgi tendon organ, drawn to illustrate their similarity.

with bundles of collagen fibers; the innervated bundle is generally surrounded by a capsule (Fig. 1B). In the rat, Ruffini endings are found in hairy skin (Fundin et al., 1997; Rice et al., 1997), the hard palate (Arvidsson et al., 1995), periodontal ligaments (Wakisaka et al., 2000), and even in the dura mater (Andres et al., 1987). Ruffini endings are slowly adapting (SAII) mechanoreceptors which respond to tissue stress (Grigg, 1996). Pacinian Corpuscles Pacinian corpuscles are the largest of the encapsulated receptors. The capsule has 20 to 70 lamellae,

arranged like the layers of an onion (Munger and Ide, 1988). They are found in the deeper layers of the skin and are also associated with interosseous membranes and mesentery. Pacinian corpuscles are very rapidly adapting, extremely sensitive to vibration, and have very large receptive fields (Leem et al., 1993). Small Lamellated Corpuscles These are frequent in the rat, sometimes called paciniform, Krause, or Golgi–Mazzoni endings. These are rapidly adapting mechanoreceptors found in glabrous skin, with encapsulated endings which may be cylin-

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drical or spherical. They have been reported in oral mucosa (Tachibana et al., 1987) and in the genitalia (Patrizi and Munger, 1965; Johnson and Halata, 1991). Meissner corpuscles are somewhat similar, but are rare in the rat. They are found mainly in the ridged glabrous skin of primates and marsupials. Lanceolate Endings The lanceolate endings of hair follicles are the fine, flattened endings of myelinated fibers that run longitudinally in the follicles of down, guard, and sinus hairs or vibrissae (Mosconi et al., 1993; Fundin et al., 1997; Takahashi-Iwanaga, 2000). They are apparently coupled mechanically to the follicle by fine collagen fibers. The more superficially located lanceolate endings may be modified Meissner corpuscles and are rapidly adapting, whereas those around the bulb of the sinus hair are slowly adapting (Baumann et al., 1996).

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et al., 1988), where they are restricted to regions of the muscle containing slow fibers. Golgi Tendon Organs The receptor is spindle shaped, and the sensory fibers branch among spiral bundles of collagen fibers in an arrangement very similar to that of the Ruffini ending (Jami, 1992). The whole structure, which is about 600 μm long, is surrounded by a capsule (Fig. 1B). The development of Golgi tendon organs has been described in the rat (Zelená and Soukup, 1977). Free Nerve Endings Free nerve endings are found in muscle and tendon, where they may serve as nociceptors and thermoreceptors, as in the skin (Mense, 1996). A small proportion respond to ischemic contraction of the muscle.

Free Nerve Endings

Joint Receptors

Free nerve endings are found in the dermis of glabrous and hairy skin, as well as in muscles, joints, and viscera (Belmonte and Cervero, 1996; Kumazawa et al., 1996). They are the terminals of unmyelinated C-fibers or thinly myelinated Aδ fibers, and function as nociceptors or thermoreceptors (Perl, 1996; McCleskey, 1997). Nociceptors respond to intense mechanical, thermal, or chemical stimulation (Belmonte and Cervero, 1996; Khalsa et al., 1997). These responses are mediated by a range of ion channels, including the capsaicin (vanilloid) receptor VR1 (Caterina and Julius, 1999).

The joint capsule contains Ruffini endings (slowly adapting), small lamellated corpuscles or paciniform corpuscles (rapidly adapting), and free nerve endings, associated with nociceptors and thermoreceptors as in the skin (Messlinger, 1996; Heppelmann, 1997). No sensory endings are found in the synovial membrane. The slowly adapting mechanoreceptors, once thought to play an important role in kinesthesia, are more likely to signal when the limits of joint movement are reached. Articular receptors have been described in the knee and elbow joints of the rat (Strasmann et al., 1990; Marinozzi et al., 1991).

Muscle Receptors CELL BODIES AND CENTRAL PROCESSES OF SOMATOSENSORY RECEPTORS

Sensory receptors in muscle include muscle spindles, Golgi tendon organs, and free nerve endings. Muscle Spindles The muscle spindle is the central, encapsulated region of a group of modified muscle fibers, the intrafusal fibers. Spindle afferents terminate in this region, where they are enclosed by a connective tissue capsule. There is usually one primary or annulospiral ending, which winds around all the intrafusal fibers and is derived from a group Ia afferent axon. There is also a secondary or flowerspray ending, which terminates on a subset of intrafusal fibers and is derived from a group II afferent axon. Group II endings signal static muscle length (i.e., they are slowly adapting), whereas group I endings are sensitive to both static length and the rate of stretch (Hunt, 1990). The distribution of muscle spindles has been reported in the cervical musculature (Brichta et al., 1987) and muscles of mastication of the rat (Rokx et al., 1984; Rowlerson

The cell bodies of somatosensory receptors in the trunk and limbs are located in the dorsal root ganglia, in which two main groups of cell bodies can be distinguished on the basis of size, axonal diameter, and neurochemistry (Lawson, 1992). These are the large neurons that may be identified with antibodies against neurofilament protein (Lawson et al., 1984) and the small neurons, for which calcitonin gene-related peptide and isolectin B4 are often used as markers. The large neurons are associated with low-threshold mechanoreceptors, whereas the small neurons are associated with the unmyelinated or thinly myelinated axons of nociceptors or thermoreceptors (Harper and Lawson, 1985). Primary afferent neurons contain excitatory amino acids, including glutamate and aspartate (Tracey et al., 1991; Valtschanoff et al., 1994; Larsson

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et al., 2001); glutamate is well established as a neurotransmitter released by these neurons. Many of the small neurons also contain neuropeptides such as calcitonin gene-related peptide, substance P, somatostatin, galanin, and vasoactive intestinal polypeptide, which serve as neuromodulators and help to mediate changes in the responses of spinal neurons, such as those responsible for pain sensation (Yaksh et al., 1999; Xu et al., 2000). The central processes of primary afferent neurons enter the spinal cord in the dorsal roots. In the dorsal root entry zone, the fibers tend to be segregated so that the thin, unmyelinated fibers of nociceptors are located in the lateral part of the dorsal root and terminate in the superficial laminae of the dorsal horn, whereas the thicker, myelinated axons are located in the medial part of the root and terminate in deeper layers of the dorsal horn (Light and Perl, 1979). The majority of central processes bifurcate into ascending and descending branches. Branches of the unmyelinated fibers enter Lissauer’s tract as well as the dorsal columns and the dorsolateral fasciculus (Chung et al., 1979, 1987). Some unmyelinated afferents are found in the ventral roots, although these aberrant fibers still have cell bodies in the dorsal root ganglion and do not appear to enter the spinal cord via the ventral roots (Hildebrand et al., 1997; Elder et al., 2000). The terminals of unmyelinated fibers are found mainly in laminae 1 and 2 of the dorsal horn (Sugiura et al., 1993), whereas the terminals of myelinated fibers from cutaneous mechanoreceptors extend through laminae 3 to 6 (Woolf, 1987). Just lateral to the superficial laminae of the dorsal horn, there is a lateral spinal nucleus, which receives visceral primary afferents (Neuhuber, 1982, 1986). The lateral spinal nucleus contains neurons at the origin of the spinomesencephalic tract that probably contribute to nociception (Willis et al., Chapter 27, see also Grant and Robertson, Chapter 4; Grant and Koerber, Chapter 5; and Ribeiroda-Silva, Chapter 6).

nuclei. This view needs to be modified in several respects. First, many fibers in the dorsal columns are descending, not ascending. This is particularly so in the rat, in which the corticospinal tract descends in the base of the dorsal columns. Fibers also descend from the dorsal column nuclei to the spinal cord (see Tracey, Chapter 7). Second, many of the ascending collaterals of primary afferents in the dorsal columns are unmyelinated (Chung et al., 1987). Third, by no means all of the ascending fibers in the dorsal columns reach the dorsal column nuclei; many leave the columns to terminate in the gray matter of the spinal cord (Chung et al., 1987). Fourth, proprioceptive information from the hindlimb is not carried by the dorsal columns. Sensory information from proprioceptors in muscles and joints of the forelimb is carried by axons in the cuneate fasciculus of the dorsal columns. However, the ascending collaterals of hindlimb proprioceptors leave the gracile fasciculus and terminate on cells in the dorsal nucleus of the spinal cord (Clarke’s column), which give rise to the dorsal spinocerebellar tract. A small proportion of these spinocerebellar axons send collaterals to nucleus Z at the rostral pole of the gracile nucleus (Low et al., 1986). Fifth, an important group of axons that ascend in the dorsal columns are not the collaterals of primary afferents, but are instead the axons of spinal neurons, referred to as postsynaptic dorsal column (PSDC) neurons. Their axons constitute about 30–40% of those terminating in the dorsal column nuclei (Giesler et al., 1984). PSDC neurons are activated by primary afferents, many of which are probably unmyelinated. The final point is that information carried by the dorsal columns is not restricted to innocuous sensation or to information from receptors in skin, joint, and muscle. Recent work has shown that PSDC neurons provide the most important pathway for nociceptive signals from the pelvic viscera (Al-Chaer et al., 1996; Willis et al., 1999).

Spinothalamic Tract ASCENDING SPINAL PATHWAYS Several ascending pathways carry somatosensory information from peripheral receptors. The most important are the dorsal column pathways and the spinothalamic tract (see also Tracey, Chapter 7).

Dorsal Column Pathways The classic view of the dorsal columns is that they are made up of the ascending collaterals of myelinated primary afferents, carrying information on discriminative touch and proprioception to the dorsal column

The spinothalamic tract signals information to higher centers about pain and temperature (Willis and Westlund, 1997) and about innocuous stimuli. Several morphological groups of spinothalamic neurons can be distinguished (Kobayashi, 1998) that differ to some extent in their thalamic projections and their responses to noxious and innocuous stimuli. Lateral thalamic nuclei (including the ventrobasal thalamus) receive a strong projection from the internal basilar nucleus, whereas medial thalamic nuclei receive most terminations from neurons in the intermediate gray and ventral horn (Giesler et al., 1979; Kobayashi, 1998). There are relatively few spinothalamic neurons in the lumbar

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cord of the rat, and most of these are “low-threshold” neurons, responding well to innocuous mechanical stimuli (Menétrey et al., 1984; Dado et al., 1994; Leem et al., 1994). The numbers of spinothalamic neurons are higher in the cervical cord, where about half of the spinothalamic neurons are “wide dynamic range” neurons, and respond in a graded fashion over a wide range of stimulus intensities from the innocuous through the noxious (Dado et al., 1994). In rats with nerve damage, there is an increased sensitivity to mechanical stimuli that is reflected in an increased responsiveness of spinothalamic neurons to mechanical stimulation (Palecek et al., 1992). Most axons of the spinothalamic tract ascend in the ventrolateral quadrant (Giesler et al., 1981) and terminate in the ventrobasal thalamus, the posterior thalamic group, and intralaminar nuclei. Neuronal responses in the ventrobasal thalamus and behavioral responses to noxious stimulation of the hindpaw are interrupted by lesion of the ventrolateral quadrant on the side contralateral to the paw (Peschanski et al., 1985; Vierck et al., 1995; see also Tracey, Chapter 7; Willis et al., Chapter 27).

MEDULLARY RELAY NUCLEI The medullary nuclei that receive somatosensory afferents are the dorsal column nuclei, including the gracile, cuneate, and external cuneate nuclei, and nucleus Z. Some of the neurons in these nuclei project to the ventrobasal thalamus; their axons arch ventrally as the internal arcuate fibers cross the midline and enter the medial lemniscus, in which they ascend to terminate in the ventral posterolateral nucleus of the thalamus. The lateral cervical nucleus is located in the upper three segments of the spinal cord and relays information from all levels of the spinal cord to the thalamus (Lund and Webster, 1967; Giesler et al., 1979).

Cytoarchitecture In most mammals, three distinct zones can be distinguished in the cuneate nucleus—a rostral, middle, and caudal region. The middle part receives the densest terminations from primary afferent fibers and contains distinct clusters of cells that project to the thalamus. In the rat it is difficult to distinguish these three regions on the basis of cytoarchitecture. However, retrograde labeling of neurons by injections of HRP into the thalamus reveals clusters of cells that are concentrated in the middle third of the gracile and cuneate nuclei (Kemplay and Webster, 1989). Work based on primary afferent terminations and staining for cytochrome

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oxidase, synaptophysin, and calcium binding proteins supports the idea that three rostrocaudal zones can be distinguished in the rat gracile and cuneate nuclei (Maslany et al., 1992; Crockett et al., 1993; Crockett et al., 1996). Many of the projection neurons in the cuneate nucleus which terminate in the ventrobasal thalamus are positive for somatostatin (Wang et al., 2000) and appear to use glutamate as a neurotransmitter (De Biasi and Rustioni, 1990). Approximately 20–30% of the neurons in the dorsal column nuclei are positive for GABA or its synthetic enzyme, glutamic acid decarboxylase. These inhibitory interneurons are scattered throughout the dorsal column nuclei (Barbaresi et al., 1986; Roettger et al., 1989). They receive synaptic inputs from primary afferents (Lue et al., 1997a) and make synaptic contacts with primary afferent terminals and cuneothalamic relay neurons, implicating them in presynaptic and postsynaptic inhibition (Lue et al., 1996).

Somatotopic Organization and Plasticity Primary afferent fibers from cutaneous receptors terminate somatotopically in the dorsal column nuclei, with the tail represented in the medial part of the gracile nucleus, and the shoulder, neck, and ear represented in the lateral part of the main cuneate nucleus (Maslany et al., 1991). This is consistent with the dermatomal arrangement of fibers in the dorsal columns, where the afferents from the most caudal structures ascend in the midline, and afferents from more rostral structures are added laterally. The representation of the paws is considerably larger than that of the rest of the body, consistent with their density of innervation. Cutaneous afferents from the digits have a complex pattern of termination, with the footpads represented ventrally in the cuneate and dorsally in the gracile (Fig. 2). This representation differs from that found by determining the receptive fields of second-order neurons using electrophysiological techniques, which led to the idea that the digits were represented dorsally in both the gracile and cuneate nuclei (Nord, 1967). Electrophysiological techniques have also been used to demonstrate a detailed musculotopic organization in the external cuneate nucleus, where neck muscles are represented in the rostrolateral pole, whereas muscles of the forelimb and paw are represented caudomedially (Campbell et al., 1974). Plasticity The normal somatotopic representation in the dorsal column nuclei is disrupted by partial or complete denervation, so that parts of the cuneate deprived of their normal inputs tend to become responsive to

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Gracile Nucleus

Cuneate Nucleus

Dorsal

Lateral footpads wr i

D I G I T S

st

dorsaall ffoorreepaw

dorsal surface of the foot

D footpads

I

el

e a r

T

he

I G

t a il

h i p

s n h e o c u k l d e r

kn e e

S

FIGURE 2 Somatotopic organization of dorsal column nuclei, based on transganglionic transport of WGA-HRP injected into different regions of skin (after Maslany et al., 1991).

inputs from afferents that remain intact. In the dorsal column nuclei of normal rats, axons in the sciatic nerve terminate in a region restricted to the gracile nucleus. In adult rats whose forelimbs were removed as neonates, sciatic nerve axons extend their terminations into the cuneate nucleus (Lane et al., 1995). A similar reorganization was demonstrated in adult rats in which the dorsal roots projecting to the cuneate nucleus are cut. Within 3 months of deafferentation, aberrant terminations of hindlimb afferents were found in the cuneate nucleus. However, different mechanisms probably underlie the reorganization due to peripheral nerve injury and axotomy (Sengelaub et al., 1997).

Afferents Most of the fibers terminating in the gracile, cuneate, and external cuneate nuclei are the collaterals of primary afferents, with cell bodies in the dorsal root ganglia (Giuffrida and Rustioni, 1992; Rivero-Melian and Arvidsson, 1992; Ueyama et al., 1994; Cha and Tan, 1996). These primary afferent collaterals belong to lowthreshold mechanoreceptors. Ultrastructural studies demonstrated two types of primary afferent terminal, both of which appear to use glutamate as a neurotransmitter (De Biasi et al., 1994). Although about 25% of primary afferents in the dorsal columns are unmyelinated (Chung et al., 1987), they do not appear to terminate in the dorsal column nuclei (Giuffrida and Rustioni, 1992). The dorsal column nuclei also receive an important set of terminations from the ascending axons of postsynaptic dorsal column (PSDC) neurons. In the cat and monkey, the terminals of PSDC axons avoid the cell nest region and tend not to overlap with the terminal zones of primary afferents. However, in

the rat, the terminations of PSDC axons and primary afferents overlap throughout most of the gracile and cuneate nuclei; PSDC axons often have terminations that are closely apposed to cells retrogradely labeled from the thalamus (Cliffer and Giesler, 1989). A subgroup of PSDC neurons appears to be located in the central gray of the spinal cord and to terminate in the medial cuneate and lateral gracile (Wang et al., 1999). These neurons provide an important pathway for nociceptive signals from the viscera (Al-Chaer et al., 1996; Willis et al., 1999). Ultrastructural studies suggest that the terminals of PSDC axons do not use glutamate as a neurotransmitter; they differ both in morphology and in transmitter content from the terminals of primary afferents (De Biasi et al., 1995). Some fibers ascending in the dorsolateral fasciculus also terminate in the dorsal column nuclei (Tomasulo and Emmers, 1972). The dorsal column nuclei receive afferent fibers from the trigeminal nerve (Marfurt and Rajchert, 1991) and from various brainstem nuclei, including the red nucleus and the trigeminal, vestibular, and cochlear nuclei (Weinberg and Rustioni, 1989). The dorsal column nuclei also receive axonal terminations from the cerebral cortex (Antal, 1984; Chimelli et al., 1994; Desbois et al., 1999; Martinez-Lorenzana et al., 2001). In the cuneate, corticofugal axons terminate on the dendrites of glycinergic interneurons in the ventral part of the nucleus (Lue et al., 1997b). They modulate transmission of sensory data from the dorsal column nuclei to the ventrobasal thalamus (Malmierca and Nunez, 1998). Cutaneous afferents terminate separately from muscle afferents. Cutaneous afferents from the neck and forelimb terminate in the cuneate nucleus (Ygge, 1989; Maslany et al., 1991; Bolton and Tracey, 1992), whereas

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Dorsal Column Nuclei (Horizontal Section)

ECu

Thalamus (Coronal)

Z Cu

VPL

Po

Gr

VPM Rostral

Lateral

Dorsal

Medial

Medial

Caudal

Lateral

Ventral

Somatosensory Cortex (Cortical Surface)

inputs from muscle and joint

inputs from skin

Lateral

Rostral

Caudal

Medial

FIGURE 3 Representation of deep versus superficial receptors in the dorsal column nuclei, thalamus, and somatosensory cortex (see text for explanation).

muscle afferents from the same region terminate in the external cuneate nucleus (Ammann et al., 1983; Pfister and Zenker, 1984; Bolton and Tracey, 1992) (Fig. 3). This clear segregation is not found in the cat or monkey, where muscle afferents also terminate in the main cuneate nucleus. Cutaneous afferents from the hindlimb terminate in the gracile nucleus (LaMotte et al., 1991; Ueyama et al., 1994), whereas muscle afferents project to nucleus Z via axon collaterals of the dorsal spinocerebellar tract (Low et al., 1986). A small contingent of primary afferents from the hindlimb also terminates directly in nucleus Z (Leong and Tan, 1987).

Efferents The dorsal column nuclei have neurons whose axons travel as internal arcuate fibers, cross the midline in the sensory decussation, and enter the contralateral medial lemniscus. Some of these axons terminate in the ventroposterolateral nucleus (VPL), part of the somatosensory thalamus. The cells of origin of these axons are found throughout the gracile and cuneate nuclei, with the highest concentration just caudal to the obex (MantleSt. John and Tracey, 1987; Kemplay and Webster, 1989). Neurons projecting to VPL are also found in the

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external cuneate nucleus (Mantle-St. John and Tracey, 1987). These lemniscal afferents to the thalamus use glutamate as a neurotransmitter (De Biasi and Rustioni, 1990). The dorsal column nuclei also project to the posterior thalamic group (Feldman and Kruger, 1980). In addition to this main somatosensory pathway, a separate contingent of axons splits off the medial lemniscus at the obex to terminate in the pontine nuclei (Kosinski et al., 1986), parabrachial nuclei, dorsal reticular nuclei, and the inferior colliculus (Coleman and Clerici, 1987), as well as the mesencephalic reticular nuclei (Massopust et al., 1985). There are also projections to the inferior olive (Molinari et al., 1996) and cerebellum (Ji and Hawkes, 1994; Alisky and Tolbert, 1997; Tolbert and Gutting, 1997) as well as the spinal cord (Burton and Loewy, 1977; Leong et al., 1984; Villanueva et al., 1995).

SOMATOSENSORY THALAMUS The best studied part of the somatosensory thalamus is the ventrobasal complex (VB). This has two distinct parts: the ventroposterolateral nucleus (VPL) and the ventroposteromedial nucleus (VPM). VPL receives tactile inputs from the trunk and limbs via the dorsal column nuclei and is discussed here, whereas VPM receives similar inputs from the face and head via the trigeminal nuclei (see Waite, Chapter 26). Other parts of the thalamus that receive somatosensory inputs (primarily nociceptive) include the posterior group (Po) (Cadusseau and Roger, 1992; Diamond et al., 1992b), the centrolateral and parafascicular nuclei, the nucleus submedius or gelatinosus (Miletic and Coffield, 1989; Yoshida et al., 1992) and the lateral ventromedial thalamus (Monconduit et al., 1999; Desbois and Villanueva, 2001; see also Groenewegen and Witter, Chapter 17).

Ohara and Lieberman, 1993), and all VPL neurons appear to send their axons to the somatosensory cortex (Saporta and Kruger, 1977), where they use glutamate as a neurotransmitter (Kharazia and Weinberg, 1994). In other mammals, GABAergic interneurons are found scattered throughout VPL. However, GABAergic neurons that mediate inhibition within the ventrobasal thalamus in the rat are located in the reticular nucleus of the thalamus (De Biasi et al., 1988; Pinault and Deschênes, 1998a). This is a thin lamina of neurons surrounding the dorsal thalamus, which appears to play a role in selective attention (McAlonan et al., 2000).

Somatotopic Organization and Plasticity There is a somatotopic representation of the body in VPL, with a larger representation of the forelimb than of the hindlimb (Emmers, 1988). Neurons responding to joint rotation were found in a thin ventral lamina (Angel and Clarke, 1975). Some neurons in VB respond to noxious inputs; these neurons are scattered throughout VB, with the caudal part of the body represented in rostral VB and the rostral part of the body represented in caudal VB (Guilbaud et al., 1980). The somatotopic organization of the posterior thalamic group in the rat appears to be a mirror image of that found in VB. Neurons in the cortex respond to whisker displacement before neurons in medial Po, suggesting that sensory input reaches Po via the cortex (Diamond et al., 1992b). Most neurons in the posterior intralaminar region, including the centrolateral nucleus, respond to noxious mechanical stimuli. These neurons tend to have very large receptive fields and are likely to signal the existence of a noxious stimulus rather than its location or intensity (Peschanski et al., 1981). The same is true for nociceptive neurons in the nucleus submedius (gelatinosus)(Miletic and Coffield, 1989) and the ventromedial thalamic nucleus (Monconduit et al., 1999).

Cytoarchitecture

Plasticity

The ventroposterolateral nucleus of the thalamus is composed of medium-sized, multipolar neurons that are arranged in a series of concentric layers running parallel to the external medullary lamina. These neurons have round cell bodies with dendritic fields in the shape of a biconcave disk, with the soma at the center and the disks parallel to the external medullary lamina (McAllister and Wells, 1981). Intracellular labeling of physiologically identified thalamocortical neurons revealed no obvious morphological differences between neurons excited by different stimulus modalities (Peschanski et al., 1984; Harris, 1986). The rat VPL does not contain significant numbers of local interneurons (Barbaresi et al., 1986; Harris and Hendrickson, 1987;

Normal response properties and somatotopic organization of neurons in the ventrobasal thalamus are altered by reduction of normal inputs. Removal of hindlimb input to VPL by lesions of nucleus gracilis results in expansion of the shoulder representation in the rostral part of VPL (Parker et al., 1998), and removal of whisker input to VPM by blocking trigeminal nerve fibers results in changes in the response properties of neurons in VPM (Faggin et al., 1997). Corticothalamic input is apparently required for the reorganization of the hindlimb representation in VPL (Parker and Dostrovsky, 1999), but not for changes in response properties of neurons activated by whiskers in VPM (Krupa et al., 1999).

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Afferents VPL receives somatosensory information from the limbs and trunk via the dorsal column nuclei and spinal cord, whereas VPM receives somatosensory data from the face and head via the sensory trigeminal nuclei. Afferents from the dorsal column nuclei terminate throughout VPL (Peschanski et al., 1983; Massopust et al., 1985) with lemniscal axons from the gracile nucleus terminating in the rostral and dorsal parts of VPL and axons from the cuneate nucleus terminating in the caudal and ventral parts. Spinal terminations are concentrated in the rostral part of VPL (McAllister and Wells, 1981; Peschanski et al., 1983), but the separate spinothalamic and lemniscal projections from a given body part have overlapping terminations in the same region of VPL (Ma et al., 1986). The ventrobasal complex also receives afferent terminations from the thalamic reticular nucleus (Pinault and Deschênes, 1998a, 1998b) and the somatosensory cortex (Deschênes et al., 1998; Veinante et al., 2000), which modulate the activity of thalamocortical neurons. The ventrobasal thalamus also receives afferents from other regions of the brainstem, including the raphe nuclei and locus coeruleus (Peschanski and Besson, 1984a, 1984b; Carstens et al., 1990) The posterior thalamic group receives terminations from the spinal cord (Cliffer et al., 1991), medullary dorsal horn (Iwata et al., 1992), and the dorsal column and trigeminal nuclei (Cadusseau and Roger, 1992; Villanueva et al., 1998) as well as the zona incerta (Power et al., 1999), thalamic reticular nucleus (Pinault et al., 1995), and cortex (Veinante et al., 2000). The centrolateral and submedius (gelatinosus) nuclei receive nociceptive information directly from the spinal cord (Ma et al., 1987; Dado and Giesler, 1990; Cliffer et al., 1991), whereas the ventromedial thalamus receives nociceptive information relayed by the reticularis dorsalis nucleus in the medulla (Villanueva et al., 1998).

Marini et al., 1996). The nucleus submedius (gelatinosus) projects to the ventrolateral orbital cortex (Yoshida et al., 1992); neurons in this region of cortex respond to noxious stimuli (Backonja and Miletic, 1991). The ventromedial thalamic nucleus is also implicated in nociception and projects to a strip of dorsolateral frontal cortex (Desbois and Villanueva, 2001).

SOMATOSENSORY CORTEX The mammalian neocortex contains at least two somatosensory regions, the first and second somatosensory areas (SI and SII). In the primate, SI can be subdivided into areas 3, 1, and 2 following Brodmann’s scheme. Each of these areas has a distinct cytoarchitecture and contains a separate representation of the body surface. In the rat SI, there is only one representation of the body surface, dominated by the area devoted to the face and whiskers. Further, SI in the rat has a partial overlap with motor cortex. This region of overlap is defined by the zone in which movements can be elicited by low-intensity electrical stimulation, and neuronal responses can be recorded in response to cutaneous stimulation. It occupies a strip about 1 mm wide, containing the representation of the hindpaw and forepaw (Sapienza et al., 1981; Sanderson et al., 1984). This overlap zone can be recognized anatomically as the region which receives thalamocortical afferents from both VPL and VL (Donoghue et al., 1979). The second somatosensory area (SII) is located lateral to SI and contains another complete representation of the face and body. The parietal ventral area (PV) appears to be an additional somatosensory area (Li et al., 1990; Fabri and Burton, 1991a), corresponding to area Vi of Zilles (Palomero-Gallagher and Zilles, Chapter 23).

Cytoarchitecture Efferents In the rat, all neurons in the ventrobasal thalamus appear to send their axons to the somatosensory cortex (Saporta and Kruger, 1977). Neurons in the posterior thalamic nuclear group also project to SI, where they terminate in the “dysgranular” and “perigranular” zones (Koralek et al., 1988; Lu and Lin, 1993). Thalamocortical neurons from both VB and Po terminate in the second somatosensory area (SII; Par2) as well as in SI, but relatively few of these neurons send axon collaterals to both cortical areas (Spreafico et al., 1987). Neurons in the intralaminar nuclei (e.g., centrolateral and parafascicular nuclei) project to wide areas of frontal cortex (Berendse and Groenewegen, 1991;

The somatosensory cortex of the rat has the six layers typical of neocortex, with layer 4 rich in granule cells and a poorly marked boundary between layers 2 and 3 (Wise and Jones, 1978). SI has been divided into Par1, FL, and HL, where Par1 contains the representation of the head, whereas FL and HL contain the forelimb and hindlimb representation (Palomero-Gallagher and Zilles, Chapter 23). Par1 contains “barrels” or aggregations of granule cells, which can be visualized in living brain slices. The walls of the barrels are made up of granule cells in layer 4, whereas the centers contain only a low density of granule cells (Welker and Woolsey, 1974). Narrow channels of perigranular cortex (“septa”) separate the barrels, whereas supra-

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granular and infragranular neurons in register with a barrel together form a functional column (Chmielowska et al., 1986). Each barrel corresponds to a single vibrissa and receives its principal input from a single thalamocortical axon, although each such axon diverges to at least three cortical barrels (Arnold et al., 2001). About one-third of excitatory neurons within a given barrel are synaptically coupled, but there are very few connections between neurons of adjacent barrels, so that each barrel can be regarded as an independent excitatory neuronal network (Kim and Ebner, 1999; Petersen and Sakmann, 2000). However, neurons in the septa are more widely interconnected within Par1, and it has been suggested that the septal pathway is responsible for generating cues about spatial relationships in the environment that guide movement based on somatosensory information from whiskers (Kim and Ebner, 1999). The rat SI also contains granular aggregates in regions outside the whisker field such as FL and HL (Dawson and Killackey, 1987). Barrels and granular aggregates may be lumped together as “granular zones,” which have a dense population of granule cells in layer 4, and a layer 5 which is subdivided into a layer 5a containing few cells and a layer 5b containing numerous pyramidal cells (Chapin and Lin, 1984). Perigranular cortex separates the granular aggregates, and there are two “dysgranular zones,” a central one lying within FL and HL between the representations of head and paws and one lying between SI and SII (Chapin and Lin, 1990). It has been suggested that the dysgranular regions invading SI are at locations corresponding to sulci in other mammals such as marsupials and lorises (Johnson, 1990) The cytoarchitecture of the SII cortex in the rat (Par2 of Zilles) is comparable with that of SI. However, in SII there are no aggregates of granule cells and layer 4 is thinner than in SI, whereas in PV, layer 4 is thinner still (Fabri and Burton, 1991a).

Somatotopic Organization and Plasticity There is a single representation of the body surface in SI (Fig. 4). The map is inverted, with the paws medial, the whiskers caudolateral, and the jaws rostrolateral (Welker, 1971; Dawson and Killackey, 1987). Each vibrissa is represented by its own barrel, with the barrels arranged in rows that echo the arrangement of the vibrissae on the mystacial pad. The digits of the forepaw are also represented in an orderly sequence in FL cortex (Waters et al., 1995), and the same pattern is found for digits of the hindpaw in HL. Between the representation of paws and vibrissae is the central dysgranular zone. In the anesthetized rat, neurons in

this region are unresponsive, but in the conscious animal, they may be activated by muscle stretch, joint manipulation, or cutaneous stimulation (Chapin and Lin, 1990). In this sense, dysgranular cortex may be comparable with area 3a of higher mammals. It has also been suggested that granular cortex is analogous to area 3b of primates (Chapin and Lin, 1990). SII contains a separate representation of the body surface; this representation is described as upright in the mouse (Carvell and Simons, 1986) with the trunk and whiskers medial, the paws caudolateral, and the jaws rostrolateral (Fig. 4). In SII, neurons with whisker inputs are activated by several vibrissae, and neurons in the paw area have receptive fields that always include at least two adjacent digits, unlike corresponding neurons in SI (Carvell and Simons, 1986; Alloway et al., 2000). In PV, the representation of the body surface is inverted (Fabri and Burton, 1991a). Plasticity As in the thalamus and dorsal column nuclei, parts of the somatosensory cortex that are deprived of their normal inputs tend to become responsive to inputs from afferents that remain intact. Suggested mechanisms include disinhibition of corticocortical connections, reorganization of projections from subcortical levels to cortex, and sprouting. If the forelimb of an adult rat is amputated, some of the neurons within the forepaw representation become responsive to shoulder stimulation (Pearson et al., 1999). These new responses were thought to be due to changes at the subcortical level, presumably dorsal column nuclei or thalamus, rather than to plasticity of intracortical connections (Pearson et al., 2001). Application of a similar paradigm to neonatal rats has led to a different view. If the forelimb of a neonatal rat is amputated, some of the neurons within the forepaw representation become responsive to hindlimb inputs. Subcortical reorganization was originally thought to be responsible (Lane et al., 1995), but recent work suggests that disinhibition of a polysynaptic corticocortical pathway is more likely to be involved (Stojic et al., 2001). It remains to be seen whether this apparent difference in the mechanisms underlying plasticity is due to differences between the neonate and adult. It is worth noting that there is a critical period early in the life of the rat during which thalamocortical synaptic transmission in the rat’s primary somatosensory cortex is modified by sensory experience. There is some evidence suggesting that long-term potentiation (LTP) and depression (LTD) at thalamocortical synapses are involved in this plasticity. Recent work on in vitro slices has shown that thalamocortical synaptic responses exhibit N-methylD-aspartate (NMDA) receptor-dependent LTP and

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PV

trunk fore- foot paw upper lip

SII

trunk vibrissae

A B

8 7 6

DZ

C

5

E 4

FL

3

δ 2

γ

1

β

SI

DZ

AGl

AGm

α

D

HL Lateral Rostral

Caudal Medial

FIGURE 4 Somatotopic organization of SI. Granular zones are occupied by barrels; barrels representing the mystacial vibrissae are arranged in rows (A–E) corresponding to rows of vibrissae. Barrels are surrounded by perigranular cortex. Between the representations of the head and paws is the central dysgranular zone (DZ). Abbreviations: AGl, lateral agranular cortex corresponding to primary motor area; AGm, medial agranular cortex corresponding to supplementary motor area; FL, forelimb area; HL; hindlimb area. After Carvell and Simons (1986), Dawson and Killackey (1987), Chapin and Lin (1990), and Fabri and Burton (1991a).

LTD during a developmental period similar to the critical period in vivo, but LTP and LTD could not be induced after the critical period. It was suggested that thalamocortical synapses may be formed as silent synapses that can then be modulated by LTP or LTD and contribute to cortical plasticity (Feldman et al., 1999). A great deal of work has been done on plasticity of the somatotopic organization of the whisker representation, but this is beyond the scope of this chapter (see Waite, Chapter 26).

Afferents Thalamic Afferents The primary somatosensory cortex (SI) receives its dominant thalamic input from VPL and VPM (Saporta and Kruger, 1977). Axons projecting to primary somatosensory cortex (SI) from VB terminate mainly in layer 4 (Herkenham, 1980; Kharazia and Weinberg, 1994). These terminations are located in the “granular

zones,” which include the barrels of Par1 as well as granular regions of FL and HL (Lu and Lin, 1993). SI also receives terminations from neurons in the posterior thalamic nucleus (Fabri and Burton, 1991b); these terminations are found in layers 1 and 5a (Herkenham, 1980) and are concentrated in the “dysgranular” and “perigranular” zones (Koralek et al., 1988; Lu and Lin, 1993). The dysgranular zones contain neurons activated by stimulation of receptors in deep tissues like muscles and joints; this information is probably relayed by the dorsolateral part of the rostral Po (Chapin and Lin, 1990). SI receives additional thalamic input from the ventrolateral thalamic nucleus, VL. Thalamocortical axons from VL terminate not only in motor cortex, but also in HL and FL, the region which can be regarded as a zone of overlap between sensory and motor cortex (Donoghue et al., 1979). SI also receives axons from the ventromedial thalamic nucleus, VM, and sparse projections from the intralaminar thalamic nuclei (Herkenham, 1980; Berendse and Groenewegen, 1991).

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The secondary somatosensory cortex (SII) receives axonal terminations from VB and from Po (Carvell and Simons, 1987; Spreafico et al., 1987; Pierret et al., 2000); terminations from Po are found in layers 1 and 4 (Herkenham, 1980).

Po (Veinante et al., 2000), whereas those in layer 6 of SI project to the ventrobasal complex, the posterior group, and the reticular thalamic nucleus (Land et al., 1995; Deschênes et al., 1998). The function of the corticothalamic projection is unclear, although it seems to play a role in the response of neurons in Po, whose activation depends on the functional integrity of the barrel field cortex (Diamond et al., 1992a).

Other Afferents The somatosensory cortex receives afferents from regions of the brain outside the thalamus. These include serotonergic inputs from the raphe nuclei (Kirifides et al., 2001) that play a role in the development of somatotopic organization (Boylan et al., 2000). The locus coeruleus sends noradrenergic fibers to somatosensory cortex, which modulate synaptic inputs (Devilbiss and Waterhouse, 2000). There are also cholinergic inputs from the basal nucleus of Meynert (Baskerville et al., 1993) that modulate plasticity of the somatosensory cortex (Zhu and Waite, 1998) and may serve to increase the influence of extracortical inputs relative to intracortical afferents (Kimura, 2000). Somatosensory cortex also receives afferents from the zona incerta (Lin et al., 1997) but the role of this pathway is unclear.

Other Efferents The somatosensory cortex sends corticobulbar axons to the dorsal column nuclei (see section on “Somatosensory Thalamus”) and trigeminal nuclei. Corticotrigeminal projection neurons are located in layer 5b of the dysgranular portion of somatosensory cortex (Killackey et al., 1989; Desbois et al., 1999). Their terminations are found throughout the trigeminal sensory complex and are densest in septal regions between single whisker representations (Jacquin et al., 1990). Projections from SI cortex to sensory nuclei in the brainstem modulate the responses of neurons terminating in the somatosensory thalamus. Both SI and SII project to the caudate-putamen (Alloway et al., 2000; Wright et al., 2001). The major somatosensory projections form a latticelike grid in the striatum that allows the integration of information for sensorimotor and cognitive processing (Brown et al., 1998). The somatosensory cortex also projects to the pontine nuclei (Leergaard et al., 2000a, 2000b), red nucleus (Ebrahimi-Gaillard and Roger, 1993), vestibular nuclei (Nishiike et al., 2000), and spinal cord (see Tracey, Chapter 7).

Efferents The somatosensory cortex sends axons to the thalamus, to the dorsal column and trigeminal nuclei in the medulla, and to other regions of cortex. There are also significant projections to the striatum, red nucleus, pontine nuclei, and spinal cord (Akintunde and Buxton, 1992). Thalamic Efferents

Interconnections between Somatosensory Cortex and Other Cortical Areas

There are generally reciprocal connections between the somatosensory cortex and the thalamic nuclei that provide its inputs (Deschênes et al., 1998). Corticothalamic neurons are located in layers 5 and 6. Those in layer 5 of SI terminate exclusively in the dorsal part of

MCx

FL, HL

There are reciprocal connections between SI and the primary motor area MI or lateral agranular cortex AGl (Chapin and Lin, 1990; Paperna and Malach, 1991;

SI

SII

Par1

GZ Fr1, Fr3 (AGl)

Par2 DZ, PGZ

2, 13

2

VM

1,3, 4,5

VL

VL

1,6 12

Po

VPL

2,3, 7,10, 11,12

VPM

1,8 9

VP

8, 9

Po

FIGURE 5 Thalamic inputs to somatosensory cortex. SI is subdivided into areas representing the face (Par1) and the limbs (FL and HL). Each area has granular zones (GZ) and perigranular zones (PGZ). 1, Fabri and Burton, 1991b; 2, Donoghue et al., 1979; 3, Saporta and Kruger, 1977; 4, Wise and Jones, 1978; 5, Chapin and Lin, 1984; 6, Koralek et al., 1988; 7, Chmielowska et al., 1986; 8. Carvell and Simons 1987; 9, Spreafico et al., 1987; 10, Land et al., 1995; 11, Lu and Lin, 1993; 12, Pierret et al., 2000; 13, Wang and Kurata, 1998.

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References

Cauller et al., 1998). There are also reciprocal connections between SI and medial agranular cortex (Agm) corresponding Fr2 or the supplementary motor area (Reep et al., 1990; Paperna and Malach, 1991), between SI and SII (Koralek et al., 1990; Cauller et al., 1998; Kim and Ebner, 1999), and between SI and PV (Fabri and Burton, 1991a). Callosal axons connect SI of the left and right sides of the cortex. Callosal connections generally link representations of axial or midline parts of the body, and in the rat granular zones in the jaw representations of SI are linked by callosal connections (Hayama and Ogawa, 1997). Otherwise, callosal connections between left and right SI are confined to dysgranular and perigranular zones. They originate from pyramidal neurons in layers 3 and 5 and terminate in corresponding layers in SI of the contralateral side (Akers and Killackey, 1978; Olavarria et al., 1984). SII also sends callosal projections to contralateral cortex. The cells of origin of these callosal projections tend to be segregated from neurons with projections to ipsilateral SI (Koralek et al., 1990). The main corticocortical connections are summarized in Fig. 6.

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Acknowledgments We are grateful to Dr. Zdenek Halata, Dr. Klaus Baumann, and Dr. Jonathan Dostrovsky for constructive comments on the chapter. We also thank Ms. Alicia Fritchle for assistance with illustrations and Dr. Peter Grafe for providing facilities for preparation of the chapter.

Left

Midline

AGm

Right

AGm

AGl

2,7

AGl 1,6

,7

2,

SI

SI

7

1,3

,5

1,4

DZ

DZ

SII

3,4

SII

FIGURE 6 Corticocortical connections of somatosensory cortex. Abbreviations: AGl, lateral agranular cortex corresponding to primary motor area (MI); AGm, medial agranular cortex corresponding to supplementary motor area; SI, primary somatosensory area, incorporating granular zones and dysgranular zones (DZ); SII, secondary somatosensory area. 1, Akers and Killackey, 1978; 2, Reep et al., 1990; 3, Koralek et al., 1990; 4, Wise and Jones, 1978; 5, Chapin and Woodward, 1982; 6, Chapin and Lin, 1990; 7, Paperna and Malach, 1991.

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Maslany, S., Crockett, D. P., and Egger, M. D. (1991). Somatotopic organization of the dorsal column nuclei in the rat: transganglionic labelling with B-HRP and WGA-HRP. Brain Res. 564, 56–65. Maslany, S., Crockett, D. P., and Egger, M. D. (1992). The cuneate nucleus in the rat does have an anatomically distinct middle region. Neurosci. Lett. 139, 130–134. Massopust, L. C., Hauge, D. H., Ferneding, J. C., Doubek, W. G., and Taylor, J. J. (1985). Projection systems and terminal localization of dorsal column afferents: An autoradiographic and horseradish peroxidase study in the rat. J. Comp. Neurol. 237, 533–544. McAllister, J. P., and Wells, J. (1981). The structural organization of the ventral posterolateral nucleus in the rat. J. Comp. Neurol. 197, 271–301. McAlonan, K., Brown, V. J., and Bowman, E. M. (2000). Thalamic reticular nucleus activation reflects attentional gating during classical conditioning. J. Neurosci. 20, 8897–8901. McCleskey, E. W. (1997). Thermoreceptors: Recent heat in thermosensation. Curr. Biol. 7, R679–681. Menétrey, D., de Pommery, J., and Roudier, F. (1984). Properties of deep spinothalamic tract cells in the rat, with special reference to ventromedial zone of lumbar dorsal horn. J. Neurophysiol. 52, 612–624. Mense, S. (1996). Group III and IV receptors in skeletal muscle: are they specific or polymodal? Prog. Brain Res. 113, 83–100. Messlinger, K. (1996). Functional morphology of nociceptive and other fine sensory endings (free nerve endings) in different tissues. Prog. Brain Res. 113, 273–298. Miletic, V., and Coffield, J. A. (1989). Responses of neurons in the rat nucleus submedius to noxious and innocuous mechanical cutaneous stimulation. Somatosens. Mot. Res. 6, 567–587. Molinari, H. H., Schultze, K. E., and Strominger, N. L. (1996). Gracile, cuneate, and spinal trigeminal projections to inferior olive in rat and monkey. J. Comp. Neurol. 375, 467–480. Monconduit, L., Bourgeais, L., Bernard, J. F., Le Bars, D., and Villanueva, L. (1999). Ventromedial thalamic neurons convey nociceptive signals from the whole body surface to the dorsolateral neocortex. J. Neurosci. 19, 9063–9072. Mosconi, T. M., Rice, F. L., and Song, M. J. (1993). Sensory innervation in the inner conical body of the vibrissal follicle- sinus complex of the rat. J. Comp. Neurol. 328, 232–251. Munger, B. L., and Ide, C. (1988). The structure and function of cutaneous sensory receptors. Arch. Histol. Cytol. 51, 1–34. Neuhuber, W. (1982). The central projections of visceral primary afferent neurons of the inferior mesenteric plexus and hypogastric nerve and the location of the related sensory and preganglionic sympathetic cell bodies in the rat. Anat. Embryol. (Berl.) 164, 413–425. Neuhuber, W. L., Sandoz, P. A., and Fryscak, T. (1986). The central projections of primary afferent neurons of greater splanchnic and intercostal nerves in the rat. A horseradish peroxidase study. Anat. Embryol. (Berl.) 174, 123–144. Nishiike, S., Guldin, W. O., and Baurle, J. (2000). Corticofugal connections between the cerebral cortex and the vestibular nuclei in the rat. J. Comp. Neurol. 420, 363–372. Nord, S. G. (1967). Somatotopic organization in the spinal trigeminal nucleus, the dorsal column nuclei and related structures in the rat. J. Comp. Neurol. 130, 343–356. Ogawa, H. (1996). The Merkel cell as a possible mechanoreceptor cell. Prog. Neurobiol. 49, 317–334. Ohara, P. T., and Lieberman, A. R. (1993). Some aspects of the synaptic circuitry underlying inhibition in the ventrobasal thalamus. J. Neurocytol. 22, 815–825. Olavarria, J., Van Sluyters, R. C., and Killackey, H. P. (1984). Evidence for the complementary organization of callosal and thalamic connections within rat somatosensory cortex. Brain Res. 291, 364–368.

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26 Trigeminal Sensory System P. M. E. WAITE School of Medical Sciences, University of New South Wales New South Wales, Australia

The trigeminal sensory system is concerned with sensory inputs from the head and face and their central pathways. Several features of the system merit its consideration as a separate chapter from other somatic inputs. First, although some of the peripheral inputs, such as those from the glabrous and hairy skin, are similar to those from other body regions, others arise from structures unique to the head such as the cornea, teeth, and tongue. Second, the central pathway through the brainstem is distinct from that for spinal inputs, though sharing some common features. Third, the organization of one component of the trigeminal system, the whisker-barrel pathway, has provided advantages for studies on neural development and plasticity. Finally, the trigeminal system of the rat has proved a valuable model in our understanding of human craniofacial disorders such as headache and toothache. The first section of this chapter outlines the adult trigeminal system from the periphery, through the brainstem and thalamus to the cortex. The focus is on the pathways contributing to sensation; the trigeminal motor system (Travers, Chapter 12, this volume) and sensory projections to other areas (e.g., cerebellum, tectum, and hypothalamus) are discussed in the relevant chapters. Structure–function relationships are emphasized, where appropriate. The section on peripheral receptors has been extended to accommodate recent studies on their innervation. Since the 1990s there has been a marked increase in research on nociceptive mechanisms within the brainstem, and these sections have been significantly expanded. The second section describes the development of the

The Rat Nervous System, Third Edition

pathway, and the effects of injury, and outlines how the unique features of the system have been utilised in studies of ontogeny and plasticity.

ADULT SENSORY TRIGEMINAL SYSTEM Peripheral Nerves and Receptors Depending on their location, cranial tissues are innervated by one of the three branches of the trigeminal nerve, the ophthalmic, maxillary, or mandibular divisions (Fig. 1). In the rat, as in other animals, the ophthalmic branch supplies the dorsum of the head, upper eyelid, and supraorbital vibrissae; the cornea and conjunctiva; and the glabrous and hairy skin over the dorsum and tip of the nose and the intranasal mucosa. Ophthalmic fibers also innervate the pineal gland (Reuss, 1999). The maxillary division supplies the postorbital skin, the upper lip, mystacial vibrissae, and lateral nose, as well as the intraoral upper jaw mucosa and upper teeth. The mandibular branch supplies the temporomandibular joint; the external auditory meatus (Folan-Curran and Cooke, 2001); proprioceptors in the jaw muscles; the skin over the mandible and lower lip; and the intraoral lower jaw mucosa, teeth, and anterior tongue. The ear pinna and caudal head is innervated by C2 and C3 from both the dorsal rami and cervical plexus; the presence of C1 is variable (Pfaller and Arvidsson, 1988). Trigeminal afferents from all three branches supply the dura and cranial blood vessels (Andres et al., 1987).

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FIGURE 1 Sensory trigeminal innervation of the rat head. The trigeminal ganglion (5Gn) is shown with its three divisions supplying the ophthalmic (V1), maxillary (V2), and mandibular (V3) regions. The main nerves innervating cutaneous or mucosal surfaces are indicated diagrammatically. Dorsal rami of C2, C3, and C4 innervate the posteromedial surface of the pinna and the skin over the dorsum of the head and neck. The cervical plexus (ventral rami of C3, C4, and C5) innervates the posterior pinna and skin over the lateral and ventral neck and shoulder. (Ophthalmic) a, lacrimal; b, frontal; c, ethmoidal. (Maxillary) d, infraorbital; e, zygomaticotemporal; f, zygomaticofacial; g, anterior and posterior superior alveolar. (Mandibular) h, buccal; i, lingual; j, inferior alveolar; k, mylohyoid; l, auriculo-temporal. (Cervical plexus) m, great auricular; n, cutaneous cervical; o, supraclavicular [from Greene, 1959; Dorfl, 1985; Waite and de Permentier, 1991 and by dissection (four animals, PW) for the dorsal rami, auriculotemporal and frontal nerves].

Sensory receptors are found in the skin of the face; the oral and nasal mucosa; and deeper structures such as subcutaneous tissues, facial muscles, joints, and tendons and are similar in structure and function to those throughout the rest of the body. Facial hairy skin contains lanceolate, Ruffini, and free nerve endings (Rice et al., 1997; Muller, 2000), whereas Meissner-like corpuscles, Merkel cells, and free nerve endings are found in the glabrous snout skin (McIntosh, 1975; Verzé et al., 1999). Muscle spindles and Golgi tendon organs are present in jaw muscles (reviewed Cooper, 1960), although no muscle spindles occur in extraocular muscles in the rat (Daunicht et al., 1985; reviewed in Donaldson, 2000). The nasal mucosa receives trigeminal innervation from Aδ and C fibers, with substance P and calcitonin gene-related peptide(CGRP) positive fibers terminating as free nerve endings within the epithelium (Zhao and Tao, 1994; Hunter and Dey, 1998). Activation of trigeminal nasal nociceptors by irritants elicits protective reflexes such as sneezing as well as neurogenic inflammation and can modify cardiorespiratory rhythms (Takeda et al., 1998; Dutschmann and Paton, 2002). For the mouth, the rodent buccal mucosa and papilla around the incisor contain both encapsulated and unencapsulated

endings (Ichikawa and Sugimoto, 1997). Ruffini endings, Merkel cells, and intraepithelial terminations have been described in the hard palate as well as trigeminal fibers associated with taste buds (Arvidsson et al., 1995; Tachibana et al., 1997). In addition to these tissues, the head contains several specialized structures that are considered in more detail below.

Meninges and Cranial Vessels The cranial meninges and vasculature is richly innervated by trigeminal afferents from all three branches as well as cervical afferents (Andres et al., 1987). Both myelinated and unmyelinated afferents are present, along with sympathetic and parasympathetic efferents. A detailed study of rat ventral leptomeninges (Fricke et al., 1997) described CGRP and substance P fibers within the trabecular component of the arachnoid, the pia mater, and the adventitia of blood vessels (Fig. 2A). Similarly, CGRP and substance P fibers are plentiful supratentorially in the middle meningeal artery and branches, the dural sinuses, and dura mater (Keller and Marfurt, 1991; Knyihar-Csillik et al., 2001). Two types of nerve termination have been described; one associated with collagen fibers is

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similar to Ruffini endings and has been suggested to be mechanosensitive (Andres et al., 1987; Fricke et al., 1997). The other ending lies near the subarachnoid space, either just below the pial mesothelium or in the blood vessel adventitia (Fricke et al., 1997). This group of endings has been suggested to be chemoceptive or nociceptive. Stimulation of trigeminal afferents or ganglion releases neuropeptides such as CGRP (Knyihar-Csillik et al., 1995; Ebersberger et al., 1999) with increased levels detectible in the superior sagittal sinus (reviewed Buzzi et al., 1995). Such release has been considered to be responsible for neurogenic inflammation, associated with vasodilation and plasma extravasation. The possible activation and degranulation of mast cells has also been reported, further enhancing inflammation. Such neurogenic inflammation has been widely implicated in vascular headaches

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and the model of trigeminal stimulation has proved useful for investigating the mechanisms involved in headache, as well as potential pharmacological treatments (Knyihar-Csillik et al., 2001; Limmroth et al., 2001; Williamson and Hargreaves, 2001). Similarly, sensitisation of meningeal afferents has been reported in response to inflammatory mediators or exposure to vascular components, such as would occur in subarachnoid haemorrhage (Levy and Strassman, 2002).

Cornea and Conjunctiva The cornea receives small myelinated and unmyelinated trigeminal afferents as well as a sympathetic and modest parasympathetic innervation (Marfurt et al., 1998). Nerves enter the corneoscleral limbus radially and branch to give dense limbal and subepithelial plexuses (Fig. 2B). Fibers then enter the basal

A

FIGURE 2 (A) Schematic representation of the innervation of the rat meninges meninges. The segment shows the dura mater (dm), the dural neurothelium (ne), the leptomeninx, and the subarachnoid space (sas). Outer arachnoid cell layer (oa), inner arachnoid layer (ia), trabecular leptomeninx (tl), adventitia leptomeninx (al), pia leptomeninx (pl), cerebral artery (a), venous vessels (v), and postcapillary venules (vv) traversing the leptomeninx. Cortex, co. Afferent nerve fibers are found in the trabecular, pial, and adventitial leptomeninges. Reprinted from Fricke et al. (1997) Cell Tissue Research 287 p. 11, figure 1a, with permission. Continued

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C

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FIGURE 2, cont’d (B) Schematic drawing of the vibrissal sinus follicle. The follicle consists of an inner (IRS) and outer (ORS) root sheath surrounded by the glassy membrane (GM). At the mouth of the follicle the epidermis (Ep) thickens to form the rete ridge collar (RRC). A collagenous capsule (C) encloses the sinus and expands near the neck to become the outer conical body (OCB) containing sebaceous glands (SebG). The glassy membrane is surrounded by a mesenchymal sheath (MS) and two sinuses: a deep cavernous sinus (CS) partially filled with trabeculae (Tr) and a ring sinus (RS) containing a ringwulst (RW). The deep vibrissal nerve (DVN) can be seen entering the capsule and innervating the lower MS, GM, and ORS; superficial vibrissal nerves (SVN) innervate the upper MS, OCB, and RRC. Deep and superficial arterioles (Art) supply a capillary network (CN) and the sinuses, with separate vessels to the dermal papilla (DP). Reprinted from Rice et al. (1997) J. Comp Neurol. 385 pp. 149–184, Figure 1, with permission. (C) The distribution of afferent fibers in the periodontal ligament of the rat incisor. The ligament next to the dentine on the lingual side has two regions, the tooth-related portion (TRP) that moves with the erupting tooth and an alveolar portion (AP) that remains stationary. Afferent terminations are found in the AP zone only. Modified from Byers and Dong, (1989) J. Comp Neurol. 279 pp. 117–127, Figure 18c, with permission. (D) CGRP-positive fibers in a quadrant of the rat cornea. At the bottom of the figure a dense network of nerve fibers supplies the corneoscleral limbus. Nerves enter the corneal stroma in radially directed bundles (long arrowheads) or in superficial branches of the limbal plexus (short arrowheads). Nerves branch repeatedly to give rise to a dense stromal plexus (large arrows) and radially oriented epithelial leashes (small arrows) directed toward the central cornea (asterisk). Reproduced from Jones and Marfurt (1991) J. Comp. Neurol. 313 pp. 132–150, Figure 10, with permission.

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epithelium and run in bundles called “leashes” in the epithelium (Jacot et al., 1997). Individual fibers terminate as free nerve endings in all epithelial layers. Direct visualization over time has shown that individual terminals continually undergo morphological rearrangements (Marfurt, 2000). Rat corneal sensory fibers contain CGRP and substance P, which are often colocalized. Pituitary adenylate cyclase activating peptide (PACAP) is also present in many fibers and may colocalize with CGRP and substance P (Marfurt, 2000). Other sensory fibers are positive for galanin (Jones and Marfurt, 1998) and FRAP (Szönyi, 1979). Mechanosensitive, thermosensitive, and polymodal nociceptive responses have been described in cats and rabbits (reviewed Marfurt, 2000). In addition to contributing to protective reflexes, a trophic effect from the release of neuropeptides has been suggested (Garcia-Hirschfeld et al., 1994). The corneal sensory innervation also plays a role in maintaining epithelial integrity and in wound healing. In addition to the cornea, trigeminal afferents provide a modest innervation to the conjunctiva and eyelid (Simons and Smith, 1994). For the conjunctiva, CGRP and substance P afferents ramify in the subepithelial layers, with some fibers entering the epithelium (Elsås et al., 1994).

Vibrissae For rodents, the arrangement of the facial vibrissae is highly consistent; selective breeding has shown that the pattern is genetically determined (Van der Loos et al., 1984). The structure and innervation of vibrissal follicles are similar in a number of species (Rice et al., 1986) although follicles in aquatic rodents (e.g., water rats) are particularly large and densely innervated (Dehnhardt et al., 1999). Individual follicles are supplied by both deep and superficial nerves (Fig. 2C); in common laboratory rats each follicle receives approximately 250 nerve fibers. About one-third of this sensory innervation is unmyelinated (Klein et al., 1988; Waite and Li, 1993), with different types described based on location, lectin, and peptide content (Rice et al., 1997). The presence of Merkel cells, lanceolate endings and free nerve endings in these follicles is well established (Renehan and Munger, 1986; Fundin et al., 1995). Reticular and Ruffini endings have also been described (Rice et al., 1997). Functionally, slowly adapting (sinus hair) type 1 and type 2 responses, as well as rapidly adapting responses, have been recorded from vibrissal follicles. There is increasing evidence for type 1 responses being associated with Merkel cells (Baumann et al., 1996; Senok and Baumann, 1997). Lanceolate endings have

traditionally been thought to be rapidly adapting (Gottschaldt et al., 1973; Lichtentstein et al., 1990) but detailed morphological analysis indicates a complex structure whose response properties are still to be confirmed (Takahashi-Iwanaga, 2000). The finding that these lanceolate endings express degenerin/ epithelial Na+-channel subunits and stomatin, implicated in mechanotransduction, supports their role as mechanoreceptors (Fricke et al., 2000). Innervation of intervibrissal fur has also been described in detail, with transverse and longitudinal lanceolate endings and several types of unmyelinated endings (Fundin et al., 1997).

Temporomandibular Joint Aδ- and C-fiber afferents containing CGRP and substance P supply the joint capsule, peripheral regions of the disk especially anteriorly, and the synovium (Kido et al., 1995; Uddman et al., 1998; Liu et al., 2000). This innervation originates from trigeminal and cervical (C2 to C5) ganglia but not the mesencephalic nucleus (Casatti et al., 1999). Only nonencapsulated nerve endings were reported in rats (Kido et al., 1995), unlike cat or human where encapsulated endings have been described (reviewed Waite and Ashwell, 2003). Injections of irritant into the joints have provided useful models of arthritis and deep craniofacial pain (Iwata et al., 1999; Imbe et al., 2001). In addition to irritants, 80% of the trigeminal afferents are excited by intraarticular glutamate injection (Cairns et al., 2001). This excitation was greater in female rats and may contribute to the known gender differences in jaw pain in humans.

Teeth and Periodontal Ligaments Rat molars are typical of mammalian teeth in receiving trigeminal sensory innervation as well as sympathetic fibers (reviewed Byers and Närhi, 1999). The nerve supply to the rat mandibular dentition has been described in detail by Naftel et al. (1999). Approximately 230 myelinated, mainly Aδ fibers, and 800 unmyelinated fibers enter each rat molar. There is a rich coronal plexus in the pulp; nerve endings are found in the odontoblast layer, predentine and extending up to 0.2 mm into the dentinal tubules (Byers, 1994; Ibuki et al., 1996). CGRP-positive afferents are particularly numerous, some colocalized with substance P (Byers, 1994) and activation by capsaicin has also been noted. Fibers containing galanin are described (Wakisaka et al., 1996; Suzuki et al., 2002) and PEP-19 endings are found in the coronal pulp (Ichikawa and Sugimoto, 1999). In

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contrast to molar teeth, the rat incisor is continuously erupting and the innervation of the tooth and periodontium is modified to reflect this (Byers and Närhi, 1999). Innervation is less extensive than in molars and mainly unmyelinated. CGRP-positive fibers are common in the pulp and odontoblastic layer on the labial but not lingual side (Zhang et al., 1998) and do not invade the dentinal tubules. Incisal pulp stimulation has been reported to lead to substance P release into the CSF (Zubrzycka and Janecka, 2002). The periodontium receives a rich sensory innervation from somata in both the trigeminal ganglia and mesencephalic nucleus (reviewed Byers and Maeda, 1997). Receptors are free nerve endings and unencapsulated, branched, Ruffini-like terminals, associated with collagen fibers (Byers and Maeda, 1997; Takahasahi-Iwanaga et al., 1997). For periodontium of the rat molar, both types of endings are present throughout the periodontal ligament and are especially numerous periapically (Byers and Maeda, 1997). For rat incisors, most Ruffini endings were found on the lingual side in the nonerupting alveolar zone (Byers and Dong, 1989) (Fig. 2D). In contrast few terminations occur in the tooth related zone, where the ligament erupts with the tooth. Periodontal Ruffini endings are associated with specialized Schwann cells (Maeda and Byers, 1996). Ruffini endings are low-threshold mechanoreceptors, whereas the free nerve endings are primarily nociceptive (Ishii, 1997). Occlusal stimuli are necessary for maintaining the integrity of the periodontal ligament and its mechanoreceptors (Muramoto et al., 2000). Like other cranial structures, teeth have proved useful for studies on injury (Wheeler et al., 1998; Bongenhielm et al., 2000), dentinal hypersensitivity (Byers et al., 2000), and inflammation (Fristad, 1997; Chidiac et al., 2002). Response properties are modified by inflammation (Byers and Närhi, 1999). Even benign stimuli such as orthodontic tooth movement increase CGRP innervation in the pulp, periodontal ligament, and gingiva (Norevall et al., 1995).

Tongue Trigeminal afferents in the lingual nerve supply the anterior surface of the tongue and provide for general somatic sensation. Trigeminal afferents innervate the surface epithelium and filiform and fungiform papillae, mainly ipsilaterally (Suemune et al., 1992). Fibers containing CGRP, substance P, and neurokinin A have been described (Astbäck et al., 1997). For the fungiform papillae, fibers enter the connective tissue core and terminate in the epithelium both around and within the taste buds (Astbäck et al., 1997). In addition

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to providing somatic responses, lingual activity has been noted to modulate taste responses, perhaps through peptide release (Wang et al., 1995). Bitter tastants such as nicotine and caffeine can cause trigeminal activation (Liu and, Simon, 1998).

Trigeminal Ganglion The cell bodies of most trigeminal afferents lie in the trigeminal (semilunar or Gasserian) ganglion in the middle cranial fossa in the base of the skull. An exception to the location of trigeminal cells occurs for the muscle spindles of masticatory muscles and some periodontal receptors, which have their cell bodies in the mesencephalic trigeminal nucleus (see later) located in the brainstem. Not all masticatory proprioceptors have central somata; labeling of the masseter nerve indicates that about 10% of the muscle afferents have somata in the trigeminal ganglion (Zhang et al., 1991). Moreover, Golgi tendon organs and temporomandibular joint afferents have cell bodies in the trigeminal ganglion (Casatti et al., 1999). Estimates of total ganglion cell number are quite variable, ranging from 25,800 to 43,000 (mean 35,300; Lagares and Avendano, 2000) and 39,900 to 62,600 neurons (mean 52,400; Forbes and Welt, 1981) in the same strain (Sprague–Dawley). The ophthalmic and maxillary divisions of the ganglion are not separable, though the mandibular is clearly distinguishable, lying laterally (Fig. 3). Ganglion cell somata are pseudounipolar and are enveloped in satellite cells. Like those in spinal ganglia, trigeminal somata can be classified on the basis of ultrastructural and neurochemical differences (see below) into large, type A cells and smaller, type B cells, with subclasses of each (Kai-Kai, 1989; Pena et al., 2001). Lagares and Avendano (2000) report 66% are type A with an interesting laterality: A cells were 23% larger on the right side. There is an approximate correlation between cell type and function; for example, large vibrissal afferents mainly connect to type A cells (Zhou and Rush, 1995), whereas somata innervating cornea are mainly type B (Sugimoto and Takemura, 1993). Interestingly, tooth pulp afferents are commonly associated with type A cells, although generally considered to be nociceptive (Sugimoto and Takemura, 1993). Ganglion cells are encircled by a range of fibers, including noradrenergic sympathetic axons, VIPpositive parasympathetic fibers, serotonergic fibers, and a variety of peptidergic axons such as CGRP, substance P, CCK, galanin, and NOS (Lazarov, 2002). Synapses on the somata are rare although Yamamoto and Kundo (1989) describe occasional synapses from CGRP-positive pericellular fibers.

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Somatotopy Neurons within the trigeminal ganglion supply relatively localized regions of skin and are somatotopically organized, with the cells innervating ophthalmic skin lying anteromedially and those supplying mandibular skin lying posterolaterally. There is also a dorsoventral organization with dorsal peripheral regions (e.g., cornea and supraorbital vibrissae) innervated by dorsally situated somata and a similar correspondence for ventral periphery and somata (Arvidson, 1977). All studies agree that the somatotopy is only approximate with considerable intermingling of somata from adjacent regions. Neurochemistry This has been the subject of a comprehensive recent review by Lazarov (2002). Larger ganglion cells are immunopositive for NPY and peptide 19; many peptide 19-positive cells project to the tooth pulp (Ichikawa and Sugimoto, 1999). The localization of glutamate in large or small cells is controversial; Wanaka et al. (1987) found glutamate mainly in larger somata, whereas Azérad et al. (1992) reported it in small cells. Lazarov (2002) describes small to medium cells containing glutamate, substance P, CGRP, neurokinin A, CCK, somatostatin, VIP, and galanin. The peptides CGRP and substance P are commonly colocalized (Lee et al., 1985); similarly CGRP or substance P-positive cells may colocalize with NPY, CCK, or enkephalins (Lazarov, 2002). Two classes of small nociceptive-responsive cells have been described based on isolectin IB4 staining: substance P/CGRP positive, IB4 negative and substance P/CGRP negative, and IB4 positive (Ambalavanar and Morris, 1992). Both large and small cells containing GABA (Szabat et al., 1992) and NOS (Riemann and Reuss, 1999) have also been described. Like spinal ganglia, trigeminal cells contain calcium binding proteins and a variety of receptors (reviewed Lazarov, 2002; and see Tracey, Chapter 25, and Willis et al., Chapter 27, volume). Response Properties Although most of the trigeminal afferents have similar response properties to those in other body regions, certain structures are worthy of comment. Thus the cornea, dura, and tooth pulp give rise primarily to nociceptive sensations in humans (Anderson, 1975; Beuerman and Tanelian, 1979). In animal studies, low-threshold mechanical and thermal responses as well as nociceptive responses have been described from the cornea (cat and rabbit, Marfurt, 2000). Ganglion cells can be activated by stimulation of

intracranial vessels (cat, Dostrovsky et al., 1991). Rat periodontal receptors provide low-threshold responses to tooth displacement (Ishii, 1997) as well as highthreshold activation that is modified by inflammation (reviewed Byers and Maeda, 1997). Responses from periodondium of the incisor tooth are restricted to one tooth and show directional sensitivity (Tabata and Hayashi, 1994). The responses of trigeminal ganglion cells have been most extensively studied for the large vibrissae (Zucker and Welker, 1969; Gibson and Welker, 1983a, 1983b; Lichtenstein et al., 1990; Shoykhet et al., 2000). In rats most cells lack spontaneous activity but respond to movement of one vibrissa, often with very low thresholds. The majority of vibrissal responses (60–75%) are slowly adapting and many are highly directionally sensitive, having “best” deflection positions. SA (sinus hair) type 1 responses probably arise from Merkel cells in the root sheath (Baumann et al., 1996; Senok and Baumann, 1997), although the position of the receptors in relation to the direction of maximal sensitivity is unknown. Most of the remaining vibrissal neurons are rapidly adapting and show limited directional sensitivity (i.e., respond similarly to different directions of movement). Such responses probably correspond to lanceolate receptors (Rice et al., 1986; Litchenstein et al., 1990), although these have also been implicated in SA (sinus hair) type 2 responses (Baumann et al., 1996). No differences in responses occur for cells innervating the deep or superficial parts of the follicle (Waite and Jacquin, 1992). Finally, most studies report a small proportion of vibrissal cells (<10%) requiring high velocities of movement or high threshold stimuli for activation. These may correspond to free nerve endings, and the low proportion probably reflects a sampling bias toward the larger, more sensitive neurons.

Trigeminal Brain Stem Sensory Nuclei The trigeminal sensory nuclei comprise several groups extending from the midbrain to the upper cervical spinal cord (Fig. 3). The mesencephalic nucleus (Me5) contains somata of primary afferents from muscle spindles and some periodontal receptors. The trigeminal nuclear complex consists of the main or principal sensory nucleus (Pr5) and the spinal trigeminal nuclei (Sp5), which are subdivided into oral (Sp50), interpolar (Sp5I), and caudal (Sp5C) parts. The paratrigeminal nucleus (Pa5) has traditionally been thought of as a separate sensory component but should probably be considered as another subdivision of the spinal nuclei (see Usunoff et al., 1997, for discussion). Brainstem trigeminal nuclei also include the trigeminal motor nucleus (Mo5) medial to Pr5; the

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FIGURE 3 Diagrammatic representation of the trigeminal brainstem nuclei as seen from above. Inputs from the ophthalmic (V1), maxillary (V2), and mandibular (V3) divisions are shown on the left entering the trigeminal ganglion (5Gn). Their central processes project via the trigeminal root (s5) into the trigeminal tract (sp5) and give collaterals to the principal nucleus (Pr5) and the spinal subnuclei oralis (Sp5O), interpolaris (Sp5I), and caudalis (Sp5C) and to the paratrigeminal nucleus (Pa5). A proprioceptive input from the masticatory muscles is shown with its centrally located soma in the mesencephalic trigeminal nucleus (Me5). Mo5, motor trigeminal nucleus; Su5, supratrigeminal nucleus. The main outputs of each subnucleus are indicated on the right. 7, 12, facial, hypoglossal nuclei; Amb, nucleus ambiguus; APT, anterior pretectal nucleus; Cb cerebellum; IO, inferior olive; parabrachial nuclei, PB; PCRt, parvocellular reticular nucleus; Po, posterior nucleus; RVL, rostral ventrolateral reticular nucleus; SC, superior colliculus; Sol, nucleus of the solitary tract; VPM, ventroposteromedial nucleus; ZI, zona incerta.

supratrigeminal nucleus (Su5) on the dorsomedial border of Pr5; and the intertrigeminal nucleus; which lies between Pr5 and Mo5. Mo5, Su5, and the intertrigeminal nuclei are associated with masticatory reflexes and control of jaw movements (see Travers, Chapter 12, this volume). Afferent Organization within the Nuclear Complex Central processes of trigeminal ganglion cells enter the brainstem via the trigeminal sensory root, adjacent to the trigeminal motor root. Afferents with central somata pass rostromedially to Me5 (see below). All other afferents enter the trigeminal tract in the lateral pons. There is some organization within the root and

tract, with large myelinated afferents lying centrally, whereas small myelinated and unmyelinated fibers are found peripherally in the root and under the pial surface of the tract (Crissman et al., 1996). After entering the tract, some afferents bifurcate to give a rostral branch to Pr5 and a descending branch to the spinal tract; others simply descend in the tract. Terminations of trigeminal afferents are predominantly ipsilateral though some contralateral inputs occur for Sp5C, particularly from mandibular and ophthalmic afferents with midline receptive fields (Pfaller and Arvidsson, 1988; Jacquin et al., 1990b; Marfurt and Rajchert, 1991; Ellrich and Messlinger, 1999). Although the trigeminal ganglion provides the

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FIGURE 4 Diagrams of coronal sections through the brainstem at the level of the principal trigeminal nucleus (Pr5) and the spinal subnucleus interpolaris (Sp5I) indicating the somatotopic organization of each nucleus. In cross-section Pr5 is somewhat peanut-shaped and Sp5I kidney-shaped; both have cells responding to the mandibular division (V3) situated dorsally, ophthalmic (V1) ventrally and maxillary (V2) between 4V, 4th ventricle. Amb, nucleus ambiguus; icp, inferior cerebellar peduncle; IO, inferior olive; LPGi, lateral paragigantocellular nucleus; LRt, lateral reticular nucleus; LSO, lateral superior olive; me5, mesencephalic trigeminal tract; Me5, mesencephalic trigeminal nucleus; ml, medial lemniscus; mlf, medial longitudinal fasciculus; Mo5, motor trigeminal nucleus; PrH, repositus hypoglossal nucleus; py, pyramidal tract; pyx, pyramidal decussation; RVL, rostroventrolateral reticular nucleus; sol, solitary tract; sp5, sensory root of trigeminal nerve; sp5, spinal trigeminal tract; scp, superior cerebellar peduncle; Su5, supratrigeminal nucleus; 12, hypoglossal nucleus (from Paxinos and Watson, 1986; Waite, 1984).

major afferent input to the nuclear complex, spinal nuclei also receive inputs from ipsilateral C2 and C3 (Xiong and Matsushita, 2000). Trigeminal afferent fibers also project to other areas such as the nucleus of the solitary tract, the parvicellular reticular formation, Probst’s nucleus, and the upper cervical cord (Marfurt and Rajchert, 1991; Sugimoto et al., 1997a). Cyto- and chemoarchitecture Consistent with the range of neurochemistry present in the ganglion, brainstem afferents contain a variety of transmitters. Glutamate is commonly found in myelinated afferents at all levels (Bae et al., 2000). Small fibers containing CGRP and substance P are widely distributed to the complex, not only being particularly dense to laminae 1 and 2 of Sp5C and to Pa5 but also innervating dorsomedial Sp5O and medial Sp5I (Sugimoto et al., 1997b). At the ultrastructural level, synaptic glomeruli are common, as in the dorsal horn (Ide and Killackey, 1985; Falls, 1986; discussed by Ribeiro-da-Silva in Chapter 6, this volume). GABAergic synapses to afferent fibers are present in both Pr5 and the spinal nuclei (Bae et al., 2000) and NOS synapses to afferents in laminae 1 and 2 of Sp5C (Yeo et al., 1997). GABA and glycine are also commonly colocalized in glomeruli in lamina 2 of Sp5C (Dumba et al., 1998).

Somatotopy All afferent fibers give off a number of medially directed collaterals that synapse in the adjacent trigeminal nuclei. These collaterals terminate somatotopically, with mandibular afferents ending dorsally and ophthalmic fibers ventrally; in Sp5I mandibular endings become more dorsomedial and ophthalmic more ventrolateral (Fig. 4). In addition there is a mediolateral organization in the nuclei: for Pr5, Sp5O, and Sp5I caudal skin projects laterally in the nucleus, whereas the nose and rostral face end medially (Nord, 1967; Waite and Cragg, 1982; Takemura et al., 1991). However in Sp5C, referred to as the laminated nucleus or medullary dorsal horn, this pattern is modified. The nose and rostral face end rostrally, whereas inputs from more posterior facial areas terminate at increasingly more caudal levels within the nucleus (Arvidsson, 1982; Jacquin et al., 1986a). This rostrocaudal layering has been referred to as an “onion skin” arrangement (Renehan and Jacquin, 1993). For Sp5C, as in the dorsal horn, different receptors and afferent fiber types terminate in different laminae (see below). The precise topography of the sensory complex means that afferents from particular structures (such as the vibrissae and teeth) terminate at somatopically appropriate locations (in both dorsoventral and mediolateral planes),

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forming rostrocaudal columns throughout the nucleus (Bates and Killackey, 1985; Ma, 1991; Sugimoto and Takemura, 1993). The organization of terminations in the main and spinal nuclei is most clearly evident for vibrissal afferents (Nord, 1967; Arvidsson, 1982) where a pattern analogous to the peripheral arrangement of vibrissae can be discerned in coronal sections, especially in young animals. Each vibrissa is associated with a patch or “barrelette”. Three representations of the vibrissae are seen in Pr5, Sp5I, and Sp5C (Belford and Killackey, 1979); although Sp5O receives vibrissal terminations no patches are evident. In horizontal sections, terminations from each vibrissa are seen as long rostrocaudal columns throughout the nuclei (Bates and Killackey, 1985). Individual afferents from vibrissae, skin, or oral structures often provide collaterals to all the subnuclei (Hayashi, 1980, 1985). Most studies report that hair afferents (vibrissal and guard hairs) have more compact, circumscribed arbors than those from skin, mucosa, teeth, or nociceptors (Jacquin et al., 1986a, 1988; Toda and Hayashi, 1992). For vibrissal afferents the major predictor of terminal arbor morphology is the subnucleus of termination, although subtle receptive field differences exist, such as mandibular vibrissae tending to have more elongated arbors (Shortland et al., 1996). Like vibrissae, tooth pulp afferents also project to all four subnuclei, forming continuous columns (Sugimoto and Takemura, 1993). These not only are densest in Pr5 and rostral Sp5O but also innervate Sp5I and Sp5C, especially laminae 1, 2a, and deep 4 (Marfurt and Turner, 1984). In contrast to other inputs, the cornea projects predominantly to more caudal nuclei (Marfurt and Del-Toro, 1987). Mesencephalic Trigeminal Nucleus Me5 forms a narrow band of cells extending from the lateral margin of the periaqueductal gray at the level of the superior colliculus rostrally to the trigeminal motor nucleus caudally. The neurons are the cell bodies of primary afferent fibers from muscle spindles of jaw muscles and periodontal receptors of both maxillary and mandibular teeth (cat; Jerge, 1963; Alvarado-Mallart et al., 1975). Muscle spindle somata are found throughout Me5, whereas periodontal neurons are reported to be more common caudally (cat; Nomura and Mizuno, 1985; Gottlieb et al., 1984). Cyto- and chemoarchitecture In the rat the majority of the cells are pseudounipolar but small multipolar cells are also present (Luo et al., 1991; Mineff et al., 1998); these have been suggested to be GABAergic and glycinergic interneurons (Mineff et al., 1998). Most Me5

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cells contain glutamate (86%; Jacobs and Miller, 1999) and aspartate and express Glu R2/3 subunits of the AMPA receptor (Mineff et al., 1998). Unlike other primary afferents they do not express neuropeptides but are encircled by fibers immunoreactive for CGRP, substance P, CCK, encephalin, VIP, galanin, and orexin as well as receiving GABAergic, serotonergic, histaminergic, and dopaminergic synapses (Liem et al., 1997; Chen et al., 2001; reviewed in Lazarov, 2002). Inputs from Pr5 and all Sp5 as well as the amygdala, area postrema, hypothalamus, and parvocellular reticular formation have been noted (Manni et al., 1982; Nagy et al., 1986; Minkels et al., 1991; Buisseret-Delmas et al., 1997; Zhang, 1998). A variety of receptors have been described on Me5 cells (reviewed Lazarov, 2002). Many Me5 cells also express a range of neurotrophins and growth factors (e.g., NGF, NT-3, and FGF) and their receptors (Zhou and Rush, 1994; Jacobs and Miller, 1999). Cells show membrane oscillations, which may relate to gap junctions and electrophysiological coupling between them (Pedroarena et al., 1999). Projections and function The bifurcating Me5 axons project peripherally into the trigeminal root and centrally to motor regions such as the trigeminal motor nuclei and hypoglossal nuclei, where they synapse directly with motoneurons (Luo and Dessem, 1999; Zhang et al., 2001). Me5 also projects to sensory regions such as dorsomedial Pr5, Sp5O, and Sp5I; laminae 4 and 5 of Sp5C; the nucleus of the solitary tract, as well as to the supratrigeminal region, parvicellular reticular nucleus (PCRt), and the medullary reticular formation (Luo et al., 1991; Herbert et al., 1997). The sensorimotor connections of Me5 underpin its role in proprioception during mastication and the integration of jaw movements. Me5 also has connections with the spinal cord, vestibular nucleus and cerebellum, superior colliculus (Luo et al., 1991, 2001; Raappana and Arvidsson, 1993, 1995; Ndiaye et al., 2000; Zhang et al., 2001), and hypothalamus, suggested to influence satiety through dopaminergic modification of chewing speed (Fujise et al., 1998). Principal Sensory Nucleus Cyto- and chemoarchitecture Pr5 lies in the lateral pons and is peanut-shaped in coronal section (Fig. 4). Rostrally it begins level with the rostral pole of Mo5 and it extends caudally to the level of the rostral pole of the facial nucleus where it is contiguous with Sp5O. Its boundary with Sp5O is oblique (Fukushima and Kerr, 1979; Bates and Killackey, 1985). Pr5 contains a high density of medium and small neurons, some of which contain glutamate (45%; Jacobs and Miller, 1999), GABA (Haring et al., 1990) or glycine (Rampon

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et al., 1996), and the calcium binding protein parvalbumin (Bennett-Clarke et al., 1992). In addition there are a few large neurons, especially dorsally, some expressing immunoreactivity for calbindin (BennettClarke et al., 1992). Many neurons contain NGF, although only a few express the low- or high-affinity neurotrophin receptors (Jacobs and Miller, 1999). Responses Pr5 cells have responses on the ipsilateral face, such as guard hairs, skin, and teeth, with a large representation from the whiskers (Jacquin et al., 1988). Two classes of whisker-responsive cells have been described (Veinante and DeschÊnes, 1999). Sixty-seven percent are small cells with single whiskerreceptive fields, asymmetric dendrites localized to one barrelette, and projections to the ventral posteromedial nucleus of the thalamus (VPM). Others are multipolar cells with multiwhisker fields, widespread dendrites, and projections to posterior thalamus, Po. Projections The main projection of Pr5 is to the contralateral thalamus (VPM and Po; Fukushima and Kerr, 1979; Kemplay and Webster, 1989; Chiaia et al., 1991a; Bennett-Clarke et al., 1992) and arises from predominantly glutamatergic neurons (Magnusson et al., 1987). Veinante and DeschÊnes (1999) describe two classes of whisker-related projections, those from cells activated by single whiskers, which project to single barreloids in VPM, and those with multiple whisker fields that project to Po and the tectum. In addition to these thalamic connections, Pr5 projects to the zona incerta (Shammah-Lagnado et al., 1985), the superior colliculus (Bruce et al., 1987; Huerta et al., 1983; Ndiaye et al., 2002), and the cerebellum (Watson and Switzer, 1978), with different terminations from dorsal and ventral parts of Pr5 (Yatim et al., 1996). There are also projections to Mo5 (Travers and Norgren, 1983), VII (from ventral Pr5, Pinginaud et al., 1999a), and the hypoglossal nucleus (from dorsal Pr5; Pinginaud et al., 1999a). Pr5 has bilateral (especially ipsilateral) reciprocal connections with the parabrachial nuclei (Yoshida et al., 1997) and the vestibular nuclei (BuisseretDelmas et al., 1999). It receives dopaminergic fibers (Kitahama et al., 2000) and a serotonergic input from the dorsal raphe (Kirifides et al., 2001). Oral Subnucleus Cyto- and chemoarchitecture This subnucleus is approximately coextensive rostrocaudally with the facial nucleus. It contains some very large cells (30–50 μm) with widespread dendritic arbors (Jacquin and Rhoades, 1990) as well as loosely packed medium and small cells, some of which contain glutamate, GABA or glycine, and parvalbumin or calbindin. Intranuclear

fiber bundles, providing connections between the nuclei, are a characteristic feature of all the spinal nuclei (Renehan and Jacquin, 1993). Responses Sp5O receives a predominant input from intraoral and perioral regions (Takemura et al., 1991; Dallel et al., 1999) associated with both lowthreshold mechanoreceptors and high-threshold afferents, including from the tooth pulp (Turner and Marfurt, 1988; Dallel et al., 1990; Sugimoto and Takemura, 1993). Cells with restricted receptive fields (e.g., one whisker or tooth), as well as widespread fields (e.g., several whiskers), are described (Jacquin and Rhoades, 1990; Tabata et al., 2001). Periodontal responses are common in rostral Sp5O; most cells give a sustained response that shows directional sensitivity, with some of these cells (20%) projecting to VPM (Tabata et al., 2001). For nociceptive responses, both wide dynamic range and nociceptive-specific responses are present; receptive fields include the facial skin, tongue, or teeth and often encompass maxillary and mandibular regions (Dallel et al., 1990). Projections Sp5O is thought to be involved in orofacial reflexes and behaviors. It provides direct projections to Me5, facial, and hypoglossal nuclei (Erzurumlu and Killackey, 1979; Zhang, 1998; Pinganaud et al., 1999), with some to the contralateral facial nucleus (Dauvergne et al., 2002). There are also indirect projections to motor nuclei via pontomedullary reticular areas such as the parvocellular reticular field (for orofacial integration) and the gigantocellular reticular nucleus (for oculomotor and neck reflexes) (ZerariMailly et al., 2001). There are projections to cervical spinal cord (Dessem and Luo, 1999); projections to all levels have been described (Ruggiero et al., 1981). In addition to these, Sp5O has connections with Po and the tectum (Veinante et al., 2000a), the zona incerta (Shammah-Lagnado et al., 1985), and the anterior pretectal nuclei (Yoshida et al., 1992) as well as to Pr5 (Zhang and Yang, 1999) and more caudal trigeminal nuclei (Jacquin et al., 1990a). Interpolar Subnucleus Cyto- and chemoarchitecture This nucleus extends from the level of the caudal facial nucleus to just below the obex. Its caudal boundary with Sp5C is oblique (Fukushima and Kerr, 1979; Phelan and Falls, 1989). Sp5I is the widest of the sensory trigeminal nuclei and contains medium and small neurons and a few larger cells (Bates and Killackey, 1985; Phelan and Falls, 1989). Cells contain glutamate, GABA, or glycine as well as parvalbumin or calbindin (Haring et al., 1990; Magnusson et al., 1986; Bennett-Clarke et al., 1992;

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Rampon et al., 1996). Sp5I reacts intensely for the somatostatin receptor. Responses There is a rich intraoral input with tooth pulp afferents forming asymmetric synaptic contacts on intermediate and distal dendrites (Lapa and Bauer, 1992). As in Sp5O, and unlike Pr5, peripheral receptive fields of projection cells in Sp5I are typically widespread (e.g., 4–19 whiskers; Jacquin et al., 1986b; Jacquin et al., 1989). Local circuit neurons, which respond to vibrissae, guard hairs, skin, or periodontium, typically had small fields (Jacquin et al., 1989). In addition to low-threshold mechanoresponses, Sp5I has nociceptive responses; the corneal projection ends in caudal Sp5I and rostral Sp5C, with the majority of cells (85%) being nociceptive-specific (Pozo and Cervero, 1993). Projections The projections of Sp5I are to the thalamus (VPM and Po; Erzurumlu and Killackey, 1979; Kemplay and Webster, 1989; Jacquin et al., 1989; Chiaia et al., 1991a; Ohya et al., 1993), the superior colliculus (Jacquin et al., 1986b), and the cerebellum, both directly as well as indirectly, via the olivary nuclei and spinal cord (Watson and Switzer, 1978; Huerta et al., 1983; Bruce et al., 1987; Jacquin et al., 1989; Yatim et al., 1996). In addition there are projections to the zona incerta (Shammah-Lagnado et al., 1985), the anterior pretectal nuclei (Yoshida et al., 1992), parabrachial nuclei (Yoshida et al., 1997), and brainstem motor nuclei (Pinganaud et al., 1999). Mapping of single whisker cells by Veinante et al. (2000a) shows thick fibers innervating Po and the midbrain, with thin fibers supplying VPM. Some Sp5I cells provide widespread collateral projections, for instance, to the thalamus and superior colliculus (Jacquin et al., 1986b, 1990a), to the olivary nuclei and superior colliculus (Huerta et al., 1983), inferior colliculus and cochlear nucleus (Li and Mizuno, 1997), or thalamus and cerebellum (Bruce et al., 1987; but see Phelan and Falls, 1991). Caudal Subnucleus (Medullary Dorsal Horn) Cyto- and chemoarchitecture Sp5C continues from the level of the obex to the upper cervical dorsal horn, with which it is continuous. Its rostral border is oblique, Sp5C being displaced medially in this region by the caudal extension of Sp5I (Fig. 3). Like the dorsal horn (Grant and Koerber, Chapter 5, this volume) it has a laminar arrangement so is often referred to as the medullary dorsal horn (Renehan and Jacquin, 1993), although differences in organization between spinal and trigeminal dorsal horns have been noted (reviewed Bereiter et al., 2000). The layers consist of a marginal layer (lamina 1) and substantia gelatinosa (lamina 2), which together make up the superficial laminae, and a

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deeper magnocellular layer (laminae 3 and 4; a separate lamina 3 is not obvious in the rat but is usually included in the magnocellular layer). Some authors also distinguish lamina 5 in Sp5C. Lamina 1 contains a variety of cell types, predominantly fusiform in shape and aligned parallel to the spinal tract, with a few large multipolar cells. In addition to extrinsic projections, some cells send axon collaterals to deeper laminae (Li et al., 2000). Lamina 2 comprises mainly small oval or fusiform cells, some of which are intrinsic cells, with axonal arbors restricted to lamina 2, whereas others are projection cells sending collaterals to laminae 1 and 3 and rostrally and caudally within the trigeminal complex (Li et al., 1999). Small-diameter primary afferent fibers containing substance P, CGRP, neurokinin A, and enkephalin project heavily to lamina 1 and outer 2 and less densely to inner 2 and 5 (Sugimoto et al., 1997b; Martin-Schild et al., 1999). In comparison, somatostatin and isolectin B4-positive fibers are particularly dense in inner 2 (Ambalavanar and Morris, 1992). These superficial layers also receive dense serotonergic and noradrenergic inputs (Lopez Costa et al., 1994). Superficial cells contain GABA and glycine (Wang et al., 2000b) and express a range of receptors, as in the dorsal horn (Riberio-da-Silva, Chapter 6, this volume). The magnocellular layer contains small, medium, and large multipolar cells, often with dendrites extending into superficial laminae (Viosin et al., 2002). It is the main termination site for low-threshold afferents from regions like the vibrissae and facial skin (Hayashi, 1985; Jacquin et al., 1986b). As in lamina 1, some cells express NK1 (Brown et al., 1995). Cells containing GABA, glutamate or aspartate, NOS, and the calcium binding proteins are seen in all layers (Magnusson et al., 1986; Haring et al., 1990; Polgar and Antal, 1995; Esteves et al., 2000). Responses Cells in the magnocellular layer respond mainly to low-threshold mechanical or thermal stimuli, such as deflection of one or more vibrissae (Renehan et al., 1986; Hu, 1990; Hutchinson et al., 1997). In addition to this low-threshold activation, Sp5C is a major center for nociceptive responses from head regions such as the temporomandibular joint, teeth, tongue, cornea, and meninges (Tsai et al., 1999; Bereiter et al., 2000; Cairns et al., 2001). Nociceptive responses occur mainly in laminae 1, 2, and 5 and include both wide dynamic range and nociceptive specific patterns of activation (Renehan et al., 1986; Hu, 1990; McHaffie et al., 1994; Iwata et al., 1999, reviewed Sessle, 2000). Meningeal (Strassman et al., 1994) and corneal (Meng and Bereiter, 1996) activation, temporomandibular joint inflammation (Zhou et al., 1999),

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tooth pulp stimulation (Coimbra and Coimbra, 1994), and experimental tooth movement (Aihara et al., 1999) all cause c-Fos activation in Sp5C, particularly in superficial layers. Several studies indicate that nociceptive responses show convergence, for example, from facial skin and meninges (Ellrich et al., 1999), cornea (Hirata et al., 1999; Pozo and Cervero, 1993), teeth (Matsumoto et al., 1999), and the temporomandibular joint (Takeshita et al., 2001). Sensitization of low-threshold responses can occur as a result of nonciceptive activation of another site for example sensitization of vibrissal responses following mengingeal stimulation (Cumberbatch et al., 1999). As in the dorsal horn, both glutamatergic and peptidergic transmission is involved in nociceptive activation (see Willis et al., Chapter 28, this volume, for discussion). NMDA activation is implicated in wide-dynamic-range responses (Luccarini et al., 2001) and plasticity after tooth pulp stimulation (Chiang et al., 1998). Nociceptive responses from some regions are represented twice within Sp5C and upper cervical cord. For example, ophthalmic inputs (e.g., cornea, see Fig. 5, nasal mucosa, transverse dural sinus) are represented at the Sp5I/Sp5C transition zone and in caudal Sp5C, near the border with C1 (Strassman and Vos, 1993; Takeda et al., 1998; Hirata et al., 1999; Meng et al., 2000). Similarly inflammation of the temporomandibular joint causes c-Fos activation at the Sp5I/ Sp5C border and in C2 (Zhou et al., 1999). The rostral representation receives both substance P and CGRP

inputs but not isolectin B4-positive fibers, whereas the more caudal sites receive substance P, CGRP, and IB4positive afferents (Sugimoto et al., 1997a). There is also evidence that the rostral and caudal sites differ in their response properties and projections, suggesting different functional roles for the two areas in nociception (reviewed Bereiter et al., 2000). Responses in both regions are modulated by descending controls from the parabrachial nuclei, periaqueductal gray, and nucleus raphe magnus (Cahusac et al., 1995; Chiang et al., 1995; Meng et al., 2000). Sp5C also receives inhibitory inputs from somatosensory cortex and insular (Jacquin et al., 1990b; Gojyo et al., 2002). Projections Lamina 1 neurons project to a number of regions within the thalamus, including VPM, submedius nucleus (Dado and Giesler, 1990; Iwata et al., 1992; Li, 1999), and intralaminar and Po (Fukushima and Kerr, 1979; Shigenaga et al., 1979; Krout et al., 2002). In addition to thalamic sites, lamina 1 and 2 cells project to the periaqueductal gray (Li et al., 1998a), the dorsal reticular nucleus (Almeida et al., 2002), and the zona incerta (Shammah-Lagnado et al., 1985). There are projections to the nucleus of the solitary tract (Menétrey et al., 1992; Guan et al., 1998), the supratrigeminal region (Li et al., 1998b), and all layers have projections to the facial nucleus (Erzurumlu and Killackey, 1979; Pinganaud et al., 1999). Sp5C also gives projections to catecholaminergic cell groups in the pontine and medullary reticular formation, such as

FIGURE 5

Location of responses from the cornea only, low-threshold mechanoreceptors (LTM) from wide dynamic range (WDR), and nociceptive-specific (NS) units recorded at the Sp5I/SP5C and Sp5C/C1 transition zones (distance from obex indicated), as determined from electrolytic lesions at each recording site. These sites correspond to the rostral and caudal locations of corneal activity. Note the ventrolateral location of responses rostrally compared with their superficial location in the medullary dorsal horn. cu, cuneate fasciculus; gr, gracile fasciculus; IO, inferior olive; LRt, lateral reticular nucleus; py, pyramidal tract; sp5, spinal trigeminal tract. Reproduced from Meng et al. (2000) Pain 87, pp. 241–251, figure 1, with permission.

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to the rostral ventrolateral reticular nucleus (Esser et al., 1998), parabrachial nuclei (Feil and Herbert, 1995; Allen et al., 1996; Li and Li, 2000), and hypothalamus (Malick et al., 2000; Cavdar et al., 2001). These projections are thought to be involved in cardiovascular and visceral modifications, such as loss of appetite, which are associated with noxious stimulation (Malick et al., 2001). Although Sp5C has an integral role in cranial nociception, it also provides intranuclear projections to all other trigeminal nuclei (Jacquin et al., 1990a; to Sp5O, Voisin et al., 2002), important in modulating both nociceptive and innocuous responses of the more rostral levels (Hallas and Jacquin, 1990; Renehan and Jacquin, 1993). Paratrigeminal Nucleus

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tory reflexes (Lindsey et al., 1997). There are also projections to the dorsal vagal complex that may modulate autonomic function (Armstrong and Hopkins, 1998) and to the parabrachial nuclei and hypothalamus for visceral and autonomic integration (Feil and Herbert, 1995; Malick and Burstein, 1998).

Thalamus The main trigeminal pathway to somatosensory cortex projects through VPM (the medial part of the ventroposterior or ventrobasal complex) and part of Po (Fig. 6 and see Groenewegen and Witter, Chapter 17; Tracey, Chapter 25). Trigeminal inputs to VPM and Po arise predominantly from Pr5, Sp5I, and Sp5C. Po

Cyto- and chemoarchitecture Pa5 consists of groups of cells embedded in the dorsal part of the spinal tract at caudal levels of Sp5I, i.e., just rostral to the obex. It was originally referred to by Ramon y Cajal (1909) as part of the interstitial system and was described in detail by Phelan and Falls (1989). The nucleus stains moderately with cytochrome oxidase and cells are mainly small and scattered among the axons of sp5. Pa5 receives inputs from perioral and intraoral regions and the upper gastrointestinal tract via trigeminal and glossopharyngeal afferents (Contreras et al., 1982; Pfaller and Arvidsson, 1988; Altschuler et al., 1989) and possibly vagal fibers (Contreras et al., 1982, but see Altschuler et al., 1989). The region shows high immunoreactivity for CGRP and substance P (Sugimoto et al., 1997b) and enkephalin, with substance P present in both fibers and cells (Chan-Palay, 1978). The region is also positive for isolectin B4 (Sugimoto et al., 1997a) and NPY Y1 receptor (Migita et al., 2001; Kopp et al., 2002). Neurons containing substance P, encephalin, NOS, and calbindin have been described (Armstrong and Hopkins, 1998). Responses Inflammation of the temporomandibular joint or chemical irritation of the tongue leads to an increase of c-Fos and preprodynorphin in Pa5 (Carstens et al., 1995; Imbe and Ren, 2000; Imbe et al., 2001). This nociceptive activation and the connectivity of the nucleus have led to the suggestion that Pa5 may be an important integrating area for nociceptive somatovisceral reflexes involving the upper gastrointestinal tract (Altschuler et al., 1989). Projections The region projects to the nucleus of the solitary tract and the rostral ventrolateral reticular nucleus as well as adjacent reticular areas and nucleus ambiguous (de Sousa Buck et al., 2001), suggesting the involvement of the area in cardiovascular and respira-

FIGURE 6 Diagram illustrating the interconnections between the SI barrel cortex and the thalamic nuclei VPM and PoM. Cortical inputs from VPM terminate in layers 3, 4, and 6, particularly in the barrel centers. Inputs from PoM terminate in layers 1, 4, and 5a; those to layer 4 end in the septa between the barrels. Projections from layer 6 cortical cells innervate both VPM and PoM. In VPM termination areas are arc – shaped, encompassing several barreloids (a, Herkenham, 1980; b, Koralek et al., 1988; c, Hoogland et al., 1987; d, Zhang and DeschÊnes, 1998; e, Jensen and Killackey, 1987a; f, Chmielowska et al., 1989; g, Land et al., 1995; h, Bourassa et al., 1995.

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also receives from Sp5O (Veinante et al., 2000a). In addition the trigeminal system provides inputs to the intralaminar nuclei and nucleus submedius, arising primarily from Sp5C and analogous to those from the dorsal horn (see Tracey, Chapters 7 and 25, and Willis et al., Chapter 27, this volume). Ventroposteromedial Nucleus Cyto- and chemoarchitecture VPM is ovoid to crescent-shaped in coronal sections and lies dorsomedial to VPL. It contains a homogenous population of medium-sized multipolar neurons, all of which are thalamocortical relay cells. Unlike carnivores and primates, no interneurons or GABAergic cells are present in rat VPM (Barbaresi et al., 1986; Williams and Faull, 1987). Inhibitory inputs come from the reticular thalamic nucleus (Pinault and Deschenes, 1998; Desîlets-Roy et al., 2002) and are modified by group 2 metabotropic glutamate receptor activation (Salt and Turner, 1998). Inputs to VPM are somatotopically organized with nose responses medial and caudal face lateral. Inputs from the vibrissae supply a disproportionately large topographic region (Waite, 1973a; Van der Loos, 1976; Sugitani et al., 1990). As in the brainstem, inputs from each whisker end on cells in discrete clusters, named “barreloids” (mouse, Van der Loos, 1976; rat, Land and Simons, 1985; Haidarliu and Ahissar, 2001) and estimated to contain about 250–300 cells (Land et al., 1995). Dendritic fields of VPM cells are asymmetrical and extend outside one barreloid (Ohara and Havton, 1994; Zantua et al., 1996). Dorsal barreloid cells have more elaborate arbors but a functional difference is not yet established (Varga et al., 2002). Barreloids receive dense inputs from contralateral Pr5 (Veinante and Deschenes, 1999) that end on proximal dendrites (Varga et al., 2002). Distal dendrites receive synapses from Sp5 (Williams et al., 1994) and somatosensory cortex (Varga et al., 2002). Pierret et al. (2000) describe each barreloid as having two parts: a dorsal “core” region, which stains darkly with cytochrome oxidase, receives from Pr5, and projects to barrels in SI, and a ventrolateral “tail,” lightly stained with cytochrome oxidase, which receives from Sp5I and projects to SII and dysgranular SI. Whether these correspond to functional different parallel pathways is not yet determined. Responses Most cells in VPM are activated by glutamate via NMDA and AMPA receptors (Do et al., 1994). They have small receptive fields responding, for instance, to movement of one or a few whiskers (Waite, 1973b; Ito, 1988; Chiaia et al., 1991b; Brecht and Sakmann, 2002). As in SI, receptive field size is affected

by type and depth of anesthesia (Freidberg et al., 1999) as well as by feedback from SI cortex (Ghazanfar et al., 2001). Both rapidly and slowly adapting responses occur and often show directional sensitivity. VPM has also been shown to have cells responding to stimulation of the cornea (Hayashi, 1995), teeth (cat, Angus-Leppan et al., 1995), and the dura or cranial vasculature, often with convergence of inputs (cat, Dostrovsky et al., 1991). Projections VPM projects topographically to SI, ending predominantly in layer 4 but also supplying supragranular and infragranular layers (Herkenham, 1980; Jensen and Killackey, 1987a; Chmielowska et al., 1989). For the vibrissal area, cells in one barreloid project predominantly to the center of one barrel (Land et al., 1995). Collaterals of thalamocortical cells active the reticular thalamic nucleus and provide feedback to VMP and PoM (Crabtree et al., 1998). Posterior Nucleus Cyto- and chemoarchitecture Po surrounds VPM on the dorsomedial and caudal aspects. The more rostral, dorsomedial sector (PoM) receives inputs from the whole-body surface and is topographically organized (Fabri and Burton, 1991; Diamond et al., 1992a), whereas the caudal sector receives convergent somatic, auditory, and visual inputs; lacks topography; and projects predominantly to SII. Cells in PoM are multipolar and have a variable morphology and more widespread dendritic fields than in VPM (Chiaia et al., 1991b). PoM lacks GARAergic interneurons and, like VMP, receives inhibition from the reticular thalamic nucleus (Pinault et al., 1995). PoM also receives a dense projection from somatosensory cortex (Hooglund et al., 1988), which modulates its activity (Diamond et al., 1992b). Responses Trigeminal afferents to PoM arise from Pr5 and Sp5I but end more diffusely than those ending in VPM. Less than half of the PoM cells are activated by whisker movement; many cells respond to stimuli to the skin, mucosa, or muscle afferents and a few are nociceptive (Chiaia et al., 1991b). In general, cells in PoM have larger receptive fields and respond at longer latencies than those in VPM (Chiaia et al., 1991b; Diamond et al., 1992a; Sosnik et al., 2001). Projections Both VPM and PoM project to SI but to complementary regions (Fig. 6; Lu and Lin, 1993) and have been suggested to form lemniscal and paralemniscal paths, respectively (Ahissar and Zacksenhouse, 2001). Thus PoM avoids the barrel centers and projects to the septa between barrels and to the surrounding dys-

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granular cortex (see below) as well as to SII (Carvell and Simons, 1987).

Somatosensory Cortex SI: Barrel Field The overall organization and cytoarchitecture of SI and SII have been fully described by PalomeroGallagher and Zilles in Chapter 23 (this volume). The body representation is somatotopic with the face area lying laterally (area Par1) and occupying a disproportionately large area of S1 (66%, Welker, 1971; see Tracey, Chapter 25, Fig. 4, this volume). The facial region is dominated by inputs from the vibrissae to a specialized region, the “barrel field”; this region has proved a fruitful research area and is the focus of the remainder of this section. Cyto- and Chemoarchitecture Specialized cell groupings, called barrels, in layer 4 of Par1 replicate the pattern of vibrissae on the face (mouse, Woolsey and Van der Loos, 1970; rat, Petrovicky and Druga, 1972; Welker and Woolsey, 1974; and see Tracey, Chapter 25, Fig. 4, this volume). Each barrel is about 200–400 μm in diameter and consists of a cell-sparse center, rich in thalamic afferents and synapses, and a cell-dense wall (Fig. 7). Septa between barrels are densely packed with vertically oriented dendrites and intracortical fibers (White, 1976; Waite, 1977; White and Peters, 1993) although occasional cells are present (Woolsey et al., 1975). Barrels can be seen with Nissl stains and are readily visualized with histochemical reactions for succinic dehydrogenase and cytochrome oxidase. Each barrel is predominantly associated with responses from one whisker (Welker, 1976). Barrels size is roughly proportional to peripheral innervation density (Lee and Woolsey, 1975), with larger barrels associated with larger, caudal whiskers. The barrel contains both smooth and spiny stellate cells; these are more densely packed peripherally, in the wall, than in the central hollow (Fig. 7), though this difference in density is less marked in the rat than the mouse (Welker and Woolsey, 1974). The stellate cells commonly have dendrites restricted to one barrel, often asymmetrically directed toward the barrel center (mouse, Woolsey et al., 1975; rat, Petersen and Sakmann, 2000), although a small percentage projects to septa and neighboring barrels. Contacts between excitatory cells of the same barrel are common (Feldmeyer and Egger, 1999). Approximately 15% of the barrel cells contain GABA and are presumably inhibitory (Chmielowska et al., 1986; Keller and White, 1987). VIP-immunoreactive cells and fibers are present, especially in barrel sides, and may colocalize with

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GABA (Zilles et al., 1993). Layer 5 pyramidal cells are preferentially aligned with the periphery of the barrels (mouse, Crandall et al., 1986); apical dendrites of layer 5b cells pass predominantly through the barrel walls (Ito, 1992). The barrel field also receives inputs from PoM and contralateral cortex (Olavarria et al., 1984), which terminate mainly in the septa in layer 4 as well as in layers 1 and 5 and surrounding dysgranular cortex (Herkenham, 1980; Koralek et al., 1988; Lu and Lin, 1993). This pattern of PoM terminations appears complementary to that of VPM (Fig. 7) and project to different areas (see below), leading to the suggestion that they constitute parallel paths processing different aspects of whisker stimuli (Ahissar et al., 2001). In addition to thalamic fibers, layer 6 receives from the claustrum (Zhang and Deschenes, 1998). The barrel field also receives noradrenergic inputs from the locus coeruleus (Simpson et al., 1997), serotonergic fibers from the dorsal and median raphe (Bennett-Clarke et al., 1991), and cholinergic inputs from the basal forebrain nuclei (McKinney et al., 1983). Both adrenergic and serotonergic inputs often arise from cells giving collaterals to subcortical trigeminal centers (Simpson et al., 1997; Kirifides et al., 2001). Studies on these extrathalamic inputs indicate their roles in shaping responses and in cortical plasticity (e.g., Devilbliss and Waterhouse, 2000; Kimura, 2000; Ego-Stengel et al., 2001). Columnar Organization As in other cortical areas, a columnar organization is seen, with VPM not only providing inputs to barrel centers in layer 4 and extending into layer 3 but also giving collaterals to layers 5b and 6 (Jensen and Killackey, 1987a; Chmielowska et al., 1989; Lu and Lin, 1993). Individual thalamocortical afferents not only project primarily to one barrel but also extend their terminal arbors into neighboring barrels (Arnold et al., 2001). In addition to this restricted termination pattern, Arnold et al. (2001) describe other thalamocortical fibers from VPM that bifurcate to innervate several columns. Afferents from VPM synapse on both smooth and spiny stellate cells in layer 4 as well as on pyramidal neurons within the column. Direct contacts onto vertically oriented VIP neurons in all layers have also been noted (Bayraktar et al., 2000) and may be involved in inhibitory mechanisms for shaping receptive fields (Staiger et al., 1996a). Layer 4 stellate cells have axonal arbors in supragranular layers and contact pyramidal cells, mainly within one column (Feldmeyer et al., 2002). Pyramidal cells in supragranular layers have extensive arbors in their own column but can also extend contacts to several

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FIGURE 7 Neuronal components of a cortical barrel. (A) the distribution of neuronal somata as seen with Nissl staining, showing the high density of stellate cells in layer 4, particularly in the barrel sides (after Welker and Woolsey, 1974). (B) An afferent from VPM ends predominantly in the barrel center and also sends collaterals to the border between layer 5b and 6 and to layer 3 (after Jensen and Killackey, 1987a). (C) Morphology of barrel cells showing approximately equal numbers of smooth and spiny stellate cells, with dendrites projecting asymmetrically within the barrel and occasionally extending to neighboring barrels; a few somata are located in the septal region (Woolsey et al., 1975; Vercelli et al., 1999). (D) Apical dendrites of infragranular (mainly layer 5) cells and axons of supragranular cells pass preferentially through the barrel wall and septum (White, 1976; Waite, 1977; Ito, 1992; White and Peters, 1993).

adjacent columns (Gottlieb and Keller, 1997). Layer 5 cells usually contact adjacent columns, although their connections can extend up to 2 mm within the cortex. The columnar organization indicated by dendritic and axonal arbors is reflected in functional columns as seen with the 2-deoxyglucose technique (Kossut and Hand, 1984; McCasland and Woolsey, 1988) or c-Fos (Melzer and Steiner, 1997; Staiger et al., 2002) or by electrophysiology (Simons, 1978; Armstrong-James and Fox, 1987; Armstrong-James et al., 1992). Responses Cells within barrels respond best to one whisker, the principal whisker, and less effectively and at longer latency by several surrounding whiskers (Zhu

and Conners, 1999; Moore et al., 1999; Brecht and Sakmann, 2002). Different response properties have been noted for barrel versus septal cells (Brumberg et al., 1999; Brecht and Sakmann, 2002). Cells within one vertical column share the same principal whisker, although responses vary by layer (Staiger et al., 2000). For instance, cells within barrels or layer 5b have been reported to detect amplitude versus velocity components of the whisker movement, respectively (Ito and Kato, 2002). As in the thalamus, many units show a direction preference (Simons, 1978; Brecht and Sakmann, 2002). There is evidence of inhibition from adjacent whiskers, which may be asymmetrical, with caudal and ventral whiskers evoking more inhibition (Brumberg et al., 1996).

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Layer 4 stellate cells contact pyramidal cells in layers 2 and 3 (Feldmeyer et al., 2002). Careful measurements of latencies of responses for cells within a column have shown a sequence of activation from the thalamus starting with layers 4 and then spreading within the column before activating adjacent columns (Armstrong-James et al., 1992). Interestingly, the spread of excitability is not uniform in all directions; for instance, connections are stronger and of shorter latency for barrels within a row compared to those between rows (mouse, Bernardo et al., 1990a, 1990b; rat, Armstrong-James et al., 1992). Activation of most cortical cells, other than the early monosynaptic responses of layer 4 cells, is dependent on NMDA receptors (Armstrong-James et al., 1993). Responses are modified by anesthesia, with deeper levels of anesthesia associated with increased rhythmicity and decreased spontaneous activity and spread of evoked responses. Recently, Shuler et al. (2001) reported ipsilateral, as well as contralateral, responses in SI and assigned it an integrative role. Imaging A variety of methods have been used to map the spread of activity across the barrel field, in response to whisker stimulation. These techniques include electrical recordings from arrays of electrodes (Petersen and Diamond, 2000), optical imaging of intrinsic signals (Masino and Frostig, 1996; Peterson et al., 1998; Yazawa et al., 2001), changes in blood flow (Moskalenko et al., 1998; Lindauer et al., 2001), and voltage-sensitive dyes (Kleinfeld and Delaney, 1996). The initial response to movement of a single whisker occurs within 5 ms and is focused on one column; activity rapidly spreads laterally to cover 2 to 11 barrels by 15 to 35 ms and then diminishes (Petersen and Diamond, 2000). Optical imaging techniques indicate more widespread activation than extracellular recordings, probably due to detection of subthreshold activity (Takashima et al., 2001). Comparison of electrophysiological recordings and voltage-sensitive imaging in vitro shows a marked dependence of spread of activity on the level of GABAergic inhibition (Petersen and Sakmann, 2001). Barrel field injury models The quantifiable modules of the barrelfield have provided fruitful ground for investigating a wide variety of insults. Cortical responses show rapid and marked plasticity in response to changes in peripheral input, such as whisker trimming, pairing, lesions, or nerve cuts in adults. This extensive literature on adult plasticity is outside the scope of the present chapter, which has focused on normal structure and function; the interested reader is referred to reviews by Buonomano and Merzenich

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(1998), Kossut (2001), and Waite (2001). In addition to peripheral insults, the barrelfield has been useful for studies in many other areas such as cortical ischemia and stroke (Wei et al., 2001; Watanabe et al., 2001), focal brain injury (Hoane and Barth, 2002), tumor growth (Sherburn et al., 1999), and chemical insults such as methamphetamine challenge (O’Dell and Marshall, 2002) and lead exposure (Wilson et al., 2000). Projections Different intracortical connections have been described from cells in register with barrels and from septal regions between barrels (Kim and Ebner, 1999). Intracortical projections from above and below barrels are generally limited to the same and neighboring columns, whereas projections from septal regions extend to other columns within a whisker row. There are reciprocal corticocortical connections with adjacent dysgranular cortex, motor cortex, and SII (Welker et al., 1988; White and DeAmicis, 1977; Israeli and Porter, 1995; and see Tracey, this volume, Chapter 25, Fig. 6). Barrel cortex provides feedback projections to VPM, PoM, and the thalamic reticular nuclei, mainly from cells in layer 6 (Fig. 6), with the indication that corticothalamic cells to VPM are in upper layer 6 and those supplying both VPM and PoM are in lower 6 (Bourassa et al., 1995; Zhang and Deschenes, 1997). VPM-projecting cells also give axon collaterals to layers 4 and 5 of the same column (Staiger et al., 1996b). Projections for both VPM and PoM are somatotopic (Fabri and Burton, 1991; Land et al., 1995). For VPM the projections from one cortical column end in a curved arc encompassing several barreloids (Hoogland et al., 1988; Bourassa et al., 1995 and see Fig. 6). Cells projecting to VPM and PoM commonly send collaterals to the reticular thalamic nucleus, although this also receives projections from layer 5 cells (Wright et al., 2001). Layer 5 cells project to the striatum, superior colliculus, anterior pretectal nuclei, pontine nuclei, and the trigeminal sensory complex in the brainstem; those to the brainstem may give collaterals to Po (Bourassa et al., 1995; Veinante et al., 2000b). For the striatal projections, layer 5 cells below the septa project bilaterally, whereas those in register with barrels project ipsilaterally and are topographic (Wright et al., 2001). Barrelfield corticopontine projections are ipsilateral and preserve topography (Alloway et al., 1999; Leergaard et al., 2000). Cortical terminals in the brainstem end contralaterally at all levels of the trigeminal nuclei, being particularly dense in magnocellular Sp5C. Cortical ablations increase the size of the brainstem receptive fields and the degree of convergence.

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Behavioral Importance Although our understanding of the mechanisms linking sensory inputs to somatic sensation is still rudimentary, the vibrissal pathway is clearly behaviorally important for the rat. This is suggested by the rich peripheral innervation and disproportionately large central representation. Moreover, several studies have shown that rats actively whisk their vibrissae during exploratory behavior and texture discrimination (Welker, 1964; Carvell and Simons, 1990). The amount and type of whisking varies with the task (Harvey et al., 2001) and has been shown to modify responses in thalamus and cortex (Fanselow and Nicolelis, 1999). Vibrissae have been shown to be important in a variety of activities, including maze learning and spatial orientation (Riesenfeld, 1979), swimming (Ahl, 1982), depth perception (Schiffman et al., 1970), and roughness discrimination (Davis et al., 1989; GuicRobles et al., 1989). Rats trained to undertake a task with one whisker still know the task if the whisker is cut and replaced with a prosthesis. However, if the prosthesis is attached to adjacent whiskers, retraining time increases with distance of the prosthesis from the trained whisker (Harris et al., 1999). Ablation of barrel cortex impairs roughness discrimination (Guic-Robles et al., 1989) and has led to the idea that vibrissae are like outstretched fingertips, providing spatial mapping and tactile discrimination. However, Brecht et al. (1997) have suggested two subsystems, macrovibrissae (long whiskers of rows A and B and caudal rows C, D, and E) and microvibrissae (short rostral whiskers of rows C, D, and E and the upper lip). On the basis of spatial and recognition tasks with normal, blind, and whiskertrimmed animals, they argue that the macrosystem acts as a distance detector and is critical for spatial tasks but not recognition, whereas the microsystem is used for object recognition. Harvey et al. (2001) have shown that the macrosystem can be used for object discrimination in certain situations. Rats have been reported to learn a novel foraging pattern better using the right whiskers (LaMendola and Bever, 1997). Although no lateral asymmetry is present for barrelfield size in the two hemispheres on a population basis, there is a large variation in areas of barrels and barrel fields in individual animals and between the two sides of the same animal (Riddle and Purves, 1995; Chen-Bee and Frostig, 1996). The behavioral significance of these individual size and laterality differences remains to be explored.

Somatosensory Cortex SII This lies lateral to SI and contains a second somatotopic map of the periphery (see Tracey, Chapter 25,

Fig. 4). Trigeminal inputs arise from VPM and Po (Carvell and Simons, 1987; Pierret et al., 2000) but do not form discrete clusters or barrels. Receptive fields are mainly contralateral but more widespread than in SI and the vibrissae occupy a relatively smaller area. SII is reciprocally connected to ipsilateral SI and motor cortex and to contralateral SII (Carvell and Simons, 1987; Koralek et al., 1990; and see, Tracey, this volume, Chapter 25, Fig. 6). SII also has projections to VMP, PoM, and the striatum (Alloway et al., 2000).

DEVELOPMENT OF THE TRIGEMINAL SYSTEM Several features of the trigeminal system have been used to advantage for developmental studies (reviewed Davies, 1997). These include the dual location of primary afferent neurons in the ganglion and Me5, the large size of the ganglion and accessibility of target tissue, and the visible and quantifiable topography of whisker-related patterns. The aim here is to highlight selected features, unique to the trigeminal pathway.

Trigeminal Ganglion Cells and Periphery Neurons of the trigeminal ganglion arise from both ectodermal placodes and neural crest (mouse, Chan and Tam, 1988; Davies and Lumsden, 1990). In the rat they are born between embryonic day (E) 9.5 and E14.5 (Rhoades et al., 1991), with no correlation between birthdate and innervation territory. (E0 corresponds to first 24 h after mating; where other timing systems have been used they are given in italics.) Process outgrowth occurs rapidly; initial axons have reached the brainstem by E12–E13 and, within a day, fine fibers can be seen in the periphery, in association with the developing whisker pad (Van Exan and Hardy, 1980) and tongue (Mbiene and Mistretta, 1997). Comparison of these dates with those for the dorsal root ganglia suggests a craniocaudal sequence of development, with trigeminal cells preceding spinal ganglion cells at cervical and thoracic levels by 1–2 days and lumbar cells by 2–3 days (Altman and Bayer, 1984; Killackey et al., 1990). Trigeminal innervation of the mystacial pad, tooth, and tongue has proved useful for studying the role of neurotrophins and their receptors in the growth and survival of sensory nerves, with many afferents having complex sequential dependencies on several factors (Luukko et al., 1997a; Rice et al., 1998; Rochlin et al., 2000; Lillesaar et al., 2001; Cronk et al., 2002). Culturing techniques and genetic manipulations indicate that peripheral target tissue produces chemoattractant and repellent factors for the axons,

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aiding their guidance (Lumsden and Davies, 1986; Rochlin et al., 2000). During the outgrowth stage, cells are independent of trophic factors; such dependence does not develop until after axons reach their target tissue (reviewed Davies and Lumsden, 1990). Concomitant with the arrival of innervation, peripheral tissues synthesize several trophic factors and ganglion cells become dependent on these for their survival. Vibrissal follicles are first apparent as surface domes, associated with mesenchymal condensations, at about E14 in the rat (Erzurumlu and Jhaveri, 1992; Osada and Kobayashi, 2000). Culture studies have shown that follicles and hairs can develop independent of innervation (mouse, Andres and Van der Loos, 1982; rat, Lillesaar and Hillebrand, 1999). Similarly, initiation of tooth development is independent of afferent nerves (reviewed Fristad, 1997, but see Luukko et al., 1997b). Periodontal innervation shows changes in terminal morphology correlated with tooth eruption and onset of occlusion (Nakakura-Ohshima et al., 1995). There is no evidence of abnormal or widespread peripheral connectivity, even in the fetus (Rhoades et al., 1990b). Somatotopic organization is present in the ganglion from E12–E13 (Erzurumlu and Jhaveri, 1992) and, by E16, receptive fields of ganglion cells are of relatively similar size to those in the adult (Chiaia et al., 1993). At early embryonic stages (E14), the trigeminal ganglion lies anterior and adjacent to the pontine flexure. The ganglion is also in close proximity to the peripheral regions it innervates, so that neither the peripheral or central processes have to traverse long distances to reach their targets. The processes extend radially and relatively directly between metencephalon and periphery (Altman and Bayer, 1982; Erzurumlu and Killackey, 1983; Erzurumlu and Jhaveri, 1992) with mandibular cells lying ventrally and ophthalmic cells dorsally. Over the next few days rotations of the ganglion (by nearly 90º so that ventral mandibular cells become lateral) and of the maturing brainstem (by a further 90º, so that lateral mandibular afferents and cells become dorsal) result in a change from the direct topography of the early embryo to a nearly 180º inverted somatotopy in the neonatal brainstem.

Central Vibrissal Pathway Birthdates for cells of the brainstem trigeminal nuclei (E12–E15 [E11–E14], Altman and Bayer, 1982; Al-Ghoul and Miller, 1993a) and VPM thalamus (E14–E15 [E13–E14], McAllister and Das, 1977) slightly precede those for the cortex (E16–E21 [E15–E20]; layer 4, E17–E19 [E16–E18], Hicks and D’Amato, 1968). The earliest afferents arrive prenatally in the brainstem (E12–E13, Erzurumlu and Jhaveri, 1992), thalamus

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(E17, Leamey and Ho, 1998), and cortex (E17, Catalano et al., 1991). Trigeminal inputs to the nucleus of solitary tract are also present from E14 (Zhang and Ashwell, 2001). Synapses are seen in Pr5 from E20 (E19) although they are transiently present in the spinal tract from E18 (E17) (Al-Ghoul et al., 1993b). Synapses are also present in Sp5C by birth (Golden et al., 1997). For the brainstem, coculturing with the ganglion has shown that axonal elongation versus branching is dependent on target age, neurotrophin levels, and slit2 expression (Erzurumlu and Jhaveri, 1995; Ulupinar et al., 2000; Ozdinler and Erzurumlu, 2002). Vibrissa – related patterns develop postnatally and appear sequentially in the brainstem (E20–P0), thalamus (P2–3), and cortex (P3–5) (Rice and Van der Loos, 1977; Belford and Killackey, 1979; Chiaia et al., 1992). The development of afferent terminal patches precedes patterning of neuronal somata. Neuronal aggregations develop around the afferent terminal patches, probably as a result of asymmetric growth of dendrites toward the afferents (Jacquin et al., 1996; Woolsey et al., 1975; Ma, 1993) and uneven growth of neuropil. Patterns at all levels become less clear with maturation, likely to be due to extensive dendritic development within the regions (Greenough and Chang, 1988; Zantua et al., 1996) as well as the arrival of other inputs and the development of interneuronal connectivity (Rhoades et al., 1997). Responses can be recorded in the brainstem from E14 to E15, with NMDA and GABAA mediated activity present from early ages (Waite et al., 2000; Lo and Erzurumlu, 2002). Response properties of cells in neonatal Sp5C are similar in somatotopy and receptive field size to those in the adult, but have longer latencies and recovery periods (Waite, 1984). Thalamic responses can be recorded from E18, with GABAergic inhibition already present (Leamey and Ho, 1998). In SI cortex responses to stimulation of the face are present as early as 12 h after birth, preceeding the earliest responses from forepaw (24 h postnatal) and hindpaw (30 h) (McCandlish et al., 1993). These early responses may reflect activity in thalamocortical axons rather than cortical cells; however, by P2.5 activity of cortical origin can be identified (Waite and Cragg, 1982). Studies with 2-deoxyglucose indicate responses in the brainstem and thalamus by birth and the cortex by P6 (Wu and Gonzales, 1997). As at other levels, GABAergic activity is present from early ages, although it matures slowly, over at least 2 months (Micheva and Beaulieu, 1997). In the fetal and perinatal brainstem, GABA causes excitation (Waite et al., 2000), compared with inhibition in thalamus and cortex (Leamey and Ho, 1998; Agmon et al., 1996). By 1 week of age, cortical cells have longer latencies, larger receptive fields, and poorer following frequencies than in adults, although

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direction sensitivity is already present (ArmstrongJames, 1975). Also the overall columnar organization is apparent in the cortex at 1 week, both from electrophysiological results and from 2-deoxyglucose (Kossut and Hand, 1984).

Plasticity in the Developing Vibrissal System The importance of the periphery in instructing central connectivity is clearly indicated by the changes that follow damage to the vibrissae or their afferent nerve supply (the infraorbital nerve). Lesions made prior to the formation of the vibrissal-related patterns prevent their development at all levels, whereas lesions after formation leave the pattern intact (Jeanmonod et al., 1981, reviewed Miller et al., 2001). There is thus a critical period during which the central relays are dependent on the periphery for normal pattern development. This critical period correlates with the developmental period, being progressively longer from the brainstem (P0-1) through the thalamus (P2-3) to the cortex (P3-4). For the brainstem, failure to form aggregations at least partly reflects a simple loss of central whisker terminals, because follicle lesioning or infraorbital nerve section causes the death of the majority of injured ganglion cells. However, loss of afferent endings is unlikely to be the explanation for lack of patches in thalamus or cortex because lesioninduced cell death, although present, is less pronounced at these levels. This suggests that lack of patterns is due to loss of an essential signal from the periphery and evidence for a chemical or electrical mediator has been much discussed (see below). Thalamocortical terminations in the cortex are evenly distributed following peripheral lesions rather than in discontinuous patches, and terminal arbors of surviving afferents in both the brainstem and cortex are more widespread (Jensen and Killackey, 1987b; Waite and de Permentier, 1991; Chiaia et al., 1992a). Functional changes after nerve injury also occur at both cortical and subcortical levels, with responses showing modified receptive fields and increased convergence, as well as take over of denervated central territory by surviving inputs (reviewed Rhoades et al., 1990a; Waite, 1984; Kossut and Siucinska, 1996; Stern et al., 2001). In these peripheral lesion models, disruption of the sensory nerve between whiskers and brainstem is essential to prevent barrel formation; clipping the whiskers or preventing their movement has no effect on barrel patterns, although it results in other structural changes (Vees et al., 1998; Lendvai et al., 2000) as well as marked modifications in responses (Li et al., 1995; Nicolelis et al., 1996; Micheva and Beaulieu,

1997). Genetic mutations resulting in lack of whiskers and impaired follicle innervation also cause modified brainstem patterns (Jhaveri et al., 1998). There is some evidence for the periphery providing a chemical signal, which allows afferents to recognize each other and hence aggregate. Thus blocking axoplasmic flow in the infraorbital nerve prevents patterns at all levels, although it also causes death of ganglion cells (Chiaia et al., 1996). Despite lack of patterns, axoplasmic blockade causes limited functional changes in the brainstem, with normal somatotopy, enlarged receptive fields, and more phasic responses (Chiaia et al., 2000). In contrast, blocking electrical activity in the peripheral nerve to the whiskers for the first postnatal week has no effect on pattern formation in the brainstem, thalamus, or cortex (Henderson et al., 1992). Genetic deletion studies are making inroads on the molecular mechanisms underlying barrel formation and plasticity (reviewed Erzurumlu and Kind, 2001). In addition to the requirement for guidance mechanisms for thalamocortical axons to reach the barrelfield, both serotonergic and glutamatergic activity is required for barrel formation. Barrel cortex shows transient aggregations of serotonergic afferents for 2–3 weeks after birth (D’Amato et al., 1987). Thalamocortical afferents express 5HT1B receptors and the serotonin transporter and take up serotonin. Serotonin has a presynaptic inhibitory effect on thalamocortical glutamatergic transmission. Preventing serotonin breakdown, by monoamine oxidase inhibitors, increases thalamocortical arbors and prevents barrel formation (Cases et al., 1996). For glutamatergic transmission, blocking NMDA receptor activation also prevents barrel formation. Thus mutants lacking NMDA R1 or R2B subunits lack whisker-related brainstem patterns (Li et al., 1994; Kutsuwada et al., 1996). Iwasato et al. (1997) showed that the presence of patterns throughout the pathway depended on the level of transgene expression, with barrels absent when expression was reduced to about 30% but restored when high levels were present. Use of a telencephalon-specific promoter to restrict R1 depletion to excitatory cortical cells showed loss of cortical barrels, with patterning intact in the brainstem and thalamus (Iwasato et al., 2000). Metabotropic glutamate transmission is also implicated, with knock-outs for mglu5 lacking barrels, but having normal thalamocortical terminals (Hannan et al., 2001). Differential effects on thalamocortical arbors and cortical cells also occur in a spontaneous mutation affecting this system, “barrelless”; such mutations have shed light on possible downstream effects of glutamatergic and serotonergic transmission (reviewed in Erzurumlu and Kind, 2001).

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Acknowledgments We acknowledge the assistance of Alicia Fritchle and Cathy Gorrie in the design and construction of the figures used in this chapter. We are also grateful to Brooke Quigley and Angela Laird for technical assistance.

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C H A P T E R

27 Pain System WILLIAM D. WILLIS, KARIN N. WESTLUND, and SUSAN M. CARLTON Department of Anatomy and Neurosciences, and Marine Biomedical Institute University of Texas Medical Branch, Galveston, Texas, USA

The pain system can be defined as the set of neural pathways that is responsible for mediating the sensation of pain (Willis, 1985). Sensory-discriminative aspects of pain include the quality, intensity, duration, and location of pain. These components are most clearly associated with pain that originates from the body surface. Pain that arises from visceral organs and other deep structures of the body has a different quality than that associated with pain from the body surface, and visceral and other deep pain is usually poorly localized. Neurons that signal the sensory-discriminative components of pain sensation encode these aspects of painful stimuli in a fashion that is generally similar to that used by other sensory systems, such as the visual and auditory systems (Perl, 1971; Price and Dubner, 1977). However, pain is more often accompanied by strong motivational-affective responses, including emotional reactions (suffering, anxiety, depression), somatic and autonomic reflexes, and endocrine changes, and it more powerfully engages the mechanisms of attention and arousal (Hardy et al., 1952; Melzack and Casey, 1968; Price, 1999) than is generally the case for vision and audition. This is especially true for visceral pain. The neural pathways that mediate the motivationalaffective components of pain are likely to overlap to some extent those involved in pain sensation, but also include additional neural structures (Casey and Bushnell, 2000; Price, 1999). A case has been made that cerebral cortical circuits that are responsible for the sensory-discriminative aspects of pain are to some extent in series with those that underlie pain affect (Price, 1999).

The Rat Nervous System, Third Edition

Based largely on studies in experimental animals, but also in part on work involving human subjects, an understanding of some aspects of the sensory-discriminative components of the pain system is beginning to emerge (see Besson and Chaouch, 1987; Casey and Bushnell, 2000; Ploner et al., 1999; Willis, 1985; Willis et al., 2001a, 2000b). Less is known about the neural circuits underlying the motivational-affective component of pain, but progress is being made on this as well (Casey and Bushnell, 2000; Price, 1999). In the past 2 decades, a number of experimental models of pathological pain have been developed that permit the study of peripheral and central mechanisms of inflammatory pain (Dickinson and Sullivan, 1987; Dubuisson and Dennis, 1977; Lariviere and Melzack, 1996; Ruda et al., 1988; Sluka and Westlund, 1992a, 1993b; Sluka and Willis, 1997), neuropathic pain (Beattie et al., 1997; Bennett and Xie, 1988; Christensen et al., 1996; Hao et al., 1991; Kim and Chung, 1992; Selzer et al., 1990; Vierck and Light, 1999; Yezierski et al., 1998), and visceral hyperalgesia (Al Chaer et al., 2000). Most of these studies have been performed on rats, although some have been done in monkeys or cats, as well (Baumann et al., 1991; Carlton et al., 1994; Chen et al., 1988; Coggeshall et al., 1983; Dougherty et al., 1992b; Palecek et al., 1992a; Schaible and Schmidt, 1985, 1988; Simone et al., 1991; Vierck et al., 1990). An understanding of the pathophysiology in these experimental models may lead to improvements in our understanding of comparable human pain states and hopefully improved therapy. Because most pain models employ rats, a more detailed knowledge of the pain system in this species is needed.

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Information about pain is transmitted by peripheral nociceptors to spinal projection neurons, such as spinothalamic tract neurons, and then to sensory relay nuclei in the thalamus and on to the cerebral cortex (Fig. 1; Willis and Westlund, 1997; see Groenewegen and Witter, Chapter 17; Tracey, Chapter 26; Waite, Chapter 27). Although pain arising from somatic structures is transmitted to higher brain centers through the spinothalamic tract and accompanying pathways in the lateral funiculus of the spinal cord, pain arising from visceral structures is relayed in large part by postsynaptic dorsal column neurons in the central core of the spinal cord and transmitted by way of the dorsal column nuclei through the medial lemniscus to the thalamus (Fig. 2; Willis and Westlund, 1997). Thus, visceral pain information is transmitted, along with nonpainful sensory-discriminative information, to the thalamus by way of the dorsal column-medial lemniscus pathway. Despite this association with the dorsal column-medial lemniscus pathway, visceral pain is diffuse and poorly localized. Other ascending pathways that contribute to various aspects of the pain experience include the spinoreticular, spinoparabrachial, spinomesencephalic, and spinohypothalamic tracts (Fig. 1; see Tracey, Chapter 26).

In addition to activation by the neural pathways that transmit information from the periphery to the brain, the pain system is regulated by pathways that descend from the brain (Fig. 1; Willis and Westlund, 1997; see Keay and Bandler, Chapter 10). Such centrifugal control is typical of sensory systems in general (Willis, 1982). However, the clinical significance of pain suppression has led to a special research emphasis on the “endogenous analgesia system,” which includes several descending pathways whose overall effect is inhibition of nociceptive signaling (see Basbaum and Fields, 1978; Besson and Chaouch, 1987; Willis, 1982; see also Fields and Besson, 1988; Millan, 2002). There are also pathways descending from the brain that facilitate nociceptive transmission (Haber et al., 1980; Giesler et al., 1981b; Urban et al., 1996; Wei et al., 1998; Yezierski et al., 1983). The discussion to follow includes an overview of the neural pathways likely to mediate the sensorydiscriminative and motivational-affective components of somatic and visceral pain. Also included is a brief description of some of the descending control systems that affect nociceptive signaling and an overview of several experimental models of pathological pain states. Findings in rats are emphasized.

FIGURE 1

Somatosensory pathways that ascend in the spinal cord to the brain and endogenous analgesia pathways that descend from the brain to the spinal cord. Information about noxious insults in the periphery are relayed from cutaneous nociceptors by Aδ and C fibers to the spinal cord. Spinothalamic tract cells send axonal projections across the midline to ascend to many brain stem sites and to the thalamus. Nonnoxious input from the periphery is transmitted by Aβ afferent fibers to the dorsal column nuclei. This information is then relayed to the thalamus. The activation of brain stem sites by noxious events in the periphery triggers modulatory activity that is mediated by the descending analgesia systems through projections that descend to the spinal cord.

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FIGURE 2 Recently discovered visceral pain pathways in the dorsal column. Nociceptive visceral afferents that innervate internal organs are shown to enter the spinal cord and to terminate in the central region of the spinal cord. The somatotopically arranged second order axons ascend in the dorsal column. Postsynaptic dorsal column neurons in the region around the central canal at L6-S1 receiving visceral nociceptive input from the pelvis and lower abdomen are shown to ascend in the dorsal column near the midline in the fasciculus gracilis to end in the medial gracile nucleus (Gr). Comparable projection neurons in the thoracic spinal cord that receive visceral nociceptive input from internal organs in the upper abdomen send their axons along the border between the fasciculi gracilis and cuneatus to terminate at the junction of the gracile and cuneate (Cu) nuclei. The Gr and Cu nuclei project to the contralateral ventral posterolateral (VPL) nucleus of the thalamus (Th).

NOCICEPTORS There has been a long-standing controversy about the nature of the sensory receptors that signal pain. It is now generally accepted that specific nociceptors innervate the skin and other organs, including muscles, joints, and viscera (see Willis, 1985; Willis and Coggeshall, 2004). (For a description of various classes of sensory receptors, as well as of the general organization of the somatosensory system, see Tracey, Chapters 7 and 26, and Waite, Chapter 27.) Sherrington (1906) defined nociceptors as sensory receptors that signal damage or the threat of damage. Recordings have been made from nociceptors supplying the skin of rats (Fleischer et al., 1983; Handwerker et al., 1987; Leem et al., 1993; Lynn and Carpenter, 1982). As in a variety of species, cutaneous and other nociceptors are supplied by finely myelinated (Aδ) fibers and unmyelinated (C) fibers. In general, the adequate stimuli for nociceptors are stronger than those needed to activate mechanoreceptors or specific thermoreceptors, although threshold stimuli need not be overtly damaging. However, nociceptors respond progressively more vigorously to graded strengths of stimuli well into the noxious range. The adequate stimuli vary with

the type of nociceptor, but in general they may include strong mechanical, thermal (heat or cold), and/or chemical stimuli. A type of cutaneous nociceptor has recently been described that is initially insensitive to mechanical or thermal stimuli but that may become sensitized; it has been suggested that such “silent” nociceptors may contribute to the pain of inflammation (Handwerker et al., 1991; Kress et al., 1992). The peripheral endings of visceral primary afferent fibers that innervate internal organs are localized in the organ walls, in the parenchyma, in the vessels supplying the viscera, and in the serosal membranes that cover the internal organs (Gebhart, 2000). The cell bodies of visceral afferent fibers are located in spinal dorsal root ganglia, cranial nerve ganglia, and enteric plexus. Neurochemical mediators of visceral pain include peptides, such as calcitonin gene-related peptide (CGRP) and substance P (SP); studies on the receptors for these neuropeptides have been reviewed (Bueno et al., 1998; Westlund, 2000). It has been known for many years that the perception of visceral pain is diffuse, poorly localized, and often “referred” to another body site. Visceral nociceptive afferent fibers are considered “silent” afferent fibers because they are minimally responsive under normal conditions (Gebhart, 2000). Very little

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nociceptive information is transmitted from the visceral afferent fibers other than about the state of distension or muscle contraction even in the face of frank damage. Visceral nociceptors that have been further tested usually also respond to chemical and thermal stimuli (Su and Gebhart, 1998). Visceral afferent fibers become highly sensitized, however, in states of inflammation, as discussed in the final sections of this chapter. Dorsal root ganglion (DRG) cells that give rise to nociceptors belong to the “small, dark” class (LaMotte et al., 1991; Robertson and Grant, 1985; see Willis and Coggeshall, 2004). These DRG cells generally contain one or more peptides. Some of the peptides that have been identified in DRG cells by immunohistochemical staining include the following: substance P, somatostatin, cholecystokinin, calcium gene-related peptide, bombesin, vasoactive intestinal polypeptide, galanin, vasopressin, oxytocin, dynorphin, enkephalin, α-neoendorphin, corticotropin-releasing factor, and neurokinin A (see Willis and Coggeshall, 2004, for references). To date it is unclear if the presence of a particular set of peptides can predict the functional type of sensory receptor (Cameron et al., 1988; however, see Lawson et al., 1997). It seems likely that the peptides are neuromodulators that act in concert with fast-acting neurotransmitters, either to enhance or to diminish their action. Numerous types of neurotransmitter receptors for both peptides and amino acids have been localized on subpopulations of DRG cells and their peripheral processes (Carlton, 2001; Coggeshall and Carlton, 1997, for review). In addition, neurotrophins and their receptors have been associated with DRG cells, contributing to the survival as well as to the activation of nociceptors (Mendell, 1996). An amazing amount of plasticity has been described in small-diameter DRG cells, presumed to be nociceptors, in response to inflammation or injury, including the up- or down-regulation of various neurotransmitters, receptors, ion channels, neurotrophins, and cell-to-cell adhesion/recognition markers. These changes are reviewed in depth in the final sections of this chapter. Nociceptor plasticity is likely to contribute to peripheral sensitization, as well as to repair and regeneration of injured axons. DRG cells and their synaptic endings in the dorsal horn have been shown to contain glutamate (GLU) and its conversion enzymes (for a further discussion, see Otterson, Chapter 37). Immunocytochemical localization of GLU has been reported in both small and large DRG cells (Battaglia et al., 1988; Wanaka et al., 1987). GLU has also been demonstrated in fine fibers in dorsal roots (Westlund et al., 1989) and in the projections of these to the superficial dorsal horn (De Biasi and Rustioni, 1988; Weinberg et al., 1987). GLU may be

colocalized with aspartate (ASP) in DRG cells (Tracey et al., 1991) Colocalization of GLU with SP and CGRP has been demonstrated in some DRG neurons and in primary afferent terminals in the dorsal horn (Battaglia and Rustioni, 1988; Jessell and Dodd, 1986; WiesenfeldHallin et al., 1984), suggesting that corelease of and interactions between these substances may occur. Excitatory amino acids have been shown to be released from primary afferent neurons in an in vitro preparation (Jeftinija et al., 1991) and from their central (Skilling et al., 1988) and peripheral processes (deGroot et al., 2000). Although cutaneous afferent fibers may be either peptidergic or nonpeptidergic, visceral afferent fibers are predominantly peptidergic (Perry and Lawson, 1998). Nociceptors supplied by C fibers in adult rats can often be activated by the excitotoxin, capsaicin; the C fibers are then inactivated and may even be killed (see Lynn, 1990). Capsaicin works by activation of VR1 vanilloid receptors, which causes the opening of nonselective cation channels (Urban and Dray, 1991). VR1 receptors are also activated by noxious heat and by protons (acid pH) (Caterina et al., 1997), and so these receptors play a fundamental role in transduction of noxious heat stimuli, as well as the recognition of an acidic microenvironment, such as occurs, for example, during inflammation. VR1 receptors have been localized on small- to medium-diameter DRG cells and their central and peripheral processes (Michael and Priestley, 1999; Szallasi, 1995). Administration of capsaicin to neonatal rats can selectively destroy the population of C nociceptors (Lynn, 1984; Nagy et al., 1983). The central projections of the fine primary afferent fibers in rats, including nociceptors, have been traced into the dorsal horn using the transganglionic transport of wheat germ agglutinin conjugated to horseradish peroxidase (WGA-HRP; LaMotte et al., 1991; Robertson and Grant, 1985; Swett and Wolff, 1985) or isolectin B4 from Griffonia simplicifolia I conjugated to HRP (Wang, 1994). The axons pass through the dorsal roots into the dorsolateral fasciculus of Lissauer. The axons end chiefly in the superficial layers of the dorsal horn (Fig. 3A–C), although some fine axons terminate in deeper layers of the dorsal horn and near the central canal [for a description of the pattern of projection of various sized primary afferents to the spinal cord (see Grant and Robertson, Chapter 4) and for the laminar organization of the gray matter of the rat spinal cord (see Grant and Koerber, Chapter 5)]. The terminal zone in the substantia gelatinosa has a somatotopic organization, and different peripheral nerves project to characteristic regions within the substantia gelatinosa (Fig. 3D). Injections of single, identified Aδ nociceptors in cats show that the axons of these enter the marginal plexus, a system of nerve fibers that passes over the dorsal

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FIGURE 3 Somatotopic organization of projections of fine afferent fibers from the hindlimb of the rat onto the superficial dorsal horn. (A–C) Transverse sections of the spinal cord showing the dorsal horn. For A, the saphenous nerve was labeled anterogradely with wheat germ agglutinin conjugated with horseradish peroxidase (WGA-HRP). The section is from mid-L3. For B, the tibial nerve and the sural and lateral sural nerves were labeled, and the section was also at mid-L3. The label in the medial dorsal horn was from the tibial nerve and that seen more laterally from the sural and lateral sural nerves. For C, the tibial, sural and superficial peroneal nerves were labeled and the section is from caudal L4. (D) A horizontal reconstruction that shows the projection territories of the tibial (T), superficial peroneal (SP), sural (S), lateral sural (LS), saphenous (SA), and posterior cutaneous (PC) nerves. From Swett and Woolf, 1985.

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aspect of the dorsal horn and then terminate in the marginal zone and in the neck of the dorsal horn; there are also terminations in the gray matter around the central canal (Light and Perl, 1979). Individual cutaneous C-nociceptors have also been labeled in guinea pigs by injections of Phaseolus vulgaris leukoagglutinin into the cell bodies of DRG cells; the axons of these cells enter the dorsolateral fasciculus of Lissauer and distribute mainly to the substantia gelatinosa (Fig. 4A and B; Sugiura et al., 1986). Visceral C fibers have a much wider distribution (Sugiura et al., 1989), both with respect to terminations in different laminae (Fig. 4C) and also to divergence rostrocaudally within the spinal cord. The terminals of presumed nociceptors in the superficial dorsal horn are frequently of the glomerular type, consisting of a central terminal that is in synaptic contact with several dendrites, as well as with other axon terminals (Csillik and Knyihár-Csillik, 1986). There are several types of glomeruli, including dense sinusoidal, light, and large dense core vesicle types (see Willis and Coggeshall, 2004, for a detailed description and for references; for a description of these glomeruli, see also Ribeiro-da-Silva, Chapter 6). Many fine afferent fibers make simple terminals rather than glomerular ones.

DORSAL HORN INTERNEURONS Neurotransmitter Content and Presumed Functional Role The first level of processing of nociceptive information is in the dorsal horn (Fig. 5). The chemical neuroanatomy of the dorsal horn is a subject that is constantly being revised and updated as improved techniques reveal new insights into classical pathways or as substances are localized in the spinal cord for the first time. Several comprehensive reviews are available (see Ruda et al., 1986; Tohyama and Shiotani, 1986; Willis and Coggeshall, 2004). This section emphasizes the chemical neuroanatomy of dorsal horn interneurons in the rat.

Excitatory Transmitters and Circuits Excitatory Transmitters Excitatory amino acids GLU-labeled cells and terminations, some of which are believed to belong to interneurons, have been localized in the superficial laminae of the spinal cord (DeBiasi and Rustioni, 1988; Greenamyre et al., 1984; Miller et al., 1988; Weinberg et al., 1987). In addition to their presence in the glomerular terminals shown to be primary afferent endings

(Csillik and Knyihar-Csillik, 1986; Coimbra et al., 1984), GLU and ASP have been localized by electron microscopic studies in large dome-shaped terminals believed to originate from interneurons and/or descending pathways (Tracey et al., 1991). Light-level photomicrographs, in fact, demonstrate that both large and small diameter neurons in most laminae of the spinal cord contain excitatory amino acid transmitters. Half of the population of terminal profiles that contact the somas of spinothalamic tract neurons in deeper laminae in the monkey are dome-shaped profiles that contain glutamate immunoreactivity (Fig. 6C; Westlund et al., 1992a). GLU and ASP are prime candidates to be the fast neurotransmitters used in nociceptive transmission (Curtis et al., 1959a). It seems quite likely that these excitatory amino acids are released from the terminals of primary afferents, interneurons, and/or descending pathways (Fig. 5) and have actions in the dorsal horn on N-methyl-D-aspartic acid (NMDA) receptors in the case of ASP and both NMDA and non-NMDA receptors in the case of GLU. GLU also activates metabotropic glutamate receptors. The net effect is the excitation of spinothalamic and other ascending tract neurons that relay nociceptive information to higher centers. Peptides Inhibitory and excitatory modulators can alter the effects of the excitatory amino acid neurotransmitters. Two peptides that increase the effectiveness of excitatory amino acid transmission are SP and CGRP. For example, when GLU or SP is injected intrathecally in rats or mice, the animals exhibit nocifensive behaviors, including caudally directed biting and scratching (Aanonsen and Wilcox, 1987; Gamse and Saria, 1986; Raigorodsky and Urca, 1987; Wiesenfeld-Hallin et al., 1984). SP has less effect, and the time course of the SP action is delayed relative to the rapid action of GLU. Application of SP to slice preparations of the spinal cord causes a slow depolarization of interneurons in the dorsal horn of the spinal cord (Randic et al., 1987), whereas the action of excitatory amino acids is a rapid depolarization (Zieglgänsberger and Puil, 1973). Intrathecal injection of CGRP by itself is often without an obvious effect. However, CGRP knockout mice are insensitive to inflammatory pain (Zhang et al., 2000), whereas SP knockout mice have normal nociceptive responses. On the other hand, transgenic mice that lack the receptor for substance P do not develop hyperalgesia after some types of visceral inflammation and noxious chemical stimuli, whereas they do develop hyperalgesia after inflammation of somatic tissues (Cervero and Laird, 1999; Laird et al., 2000). If CGRP is injected intrathecally with SP, there is a potentiation of the behavioral effects of SP (Gamse and Saria, 1986; Wiesenfeld-Hallin, 1984). This potentiation

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FIGURE 4 Distribution of the terminals of individual cutaneous C fibers in the superficial dorsal horn of the guinea pig. The C fiber in A was labeled by injecting a dorsal root ganglion cell belonging to a C polymodal nociceptor intracellularly with Phaseolus vulgaris leukoagglutinin. The axon and its terminals were labeled anterogradely. (B) Details of one of the terminal arborizations, which were largely in the substantia gelatinosa. From Sugiura et al., 1986. (C) The abundant terminals of a visceral C afferent fiber in several laminae of the dorsal horn, as well as in the region of the central canal, bilaterally. From Sugiura et al., 1989. VI. SYSTEMS

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FIGURE 5 Schematic representation of the dorsal horn circuitry that processes sensory information, including pain, temperature and crude touch, from the periphery by primary afferents. Initial processing of nociceptive information occurs in the spinal cord dorsal horn by excitatory and inhibitory interneurons. The initial nociceptive information may be further modulated by signals that descend from higher centers. Spinothalamic tract cells that receive the nociceptive information are localized primarily in laminae I and V and send their axons across the midline to ascend in the contralateral anterolateral column. The spinothalamic tract conveys nociceptive information to higher centers and signals the localization and character of the nociceptive input. Inputs related to the subjective aspects of pain and to autonomic and autonomic control are also relayed to higher centers through projection systems that accompany the spinothalamic tract.

of the effects of SP by CGRP can be explained by the action of CGRP in blocking the degradation of SP (LeGreves et al., 1989; Mao et al., 1992a), thus prolonging the action of SP and allowing SP to spread into the deeper laminae of the dorsal horn (Schaible et al., 1992). Infusion of SP or CGRP increases the release of GLU from dorsal horn slices (Kangrga et al., 1990; Kangrga and Randic, 1990) and the deep dorsal horn in awake rats (Smullin et al., 1990b). SP potentiates the action of NMDA on NMDA receptors in dorsal horn interneurons (Murase et al., 1989), including spinothalamic neurons (Dougherty and Willis, 1991). Excitatory Circuits Nociceptive information is relayed through the dorsal horn either directly or indirectly by excitatory and inhibitory interneurons (Fig. 5). Increases in primary afferent activity that signal a nociceptive event result

in the release of GLU, SP, and CGRP in the dorsal horn, causing multiple events to occur, including (1) further release of excitatory amino acids, including GLU and ASP from interneurons (Paleckova et al., 1992; Sluka and Westlund, 1992a, 1993a; Sorkin et al., 1992); (2) an increase in the sensitivity of NMDA receptors (Dougherty and Willis, 1991; Murase et al., 1989); (3) a slow, prolonged depolarization of dorsal horn interneurons, probably due to a decreased potassium conductance (Randic et al., 1987; Urban and Randic, 1984); (4) activation of second messenger systems by increasing intracellular calcium levels (Womak et al., 1988); and (5) an increase in the sensitivity of NMDA receptors resulting in a long-term increase in the responsiveness of interneurons and also spinothalamic cells (Dougherty and Willis, 1991; Murase et al., 1989), presumably because of their phosphorylation by protein kinases (Zou et al., 2000, 2002).

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FIGURE 6 Electron micrographs that illustrate postembedding immunogold staining (arrows) for GABA (A and B) and for glutamate (C) in terminal profiles apposing lamina I spinothalamic tract cells retrogradely labeled with wheat germ agglutinin conjugated horseradish peroxidase (WGA-HRP) in the rat spinal cord. Open arrows indicate the WGA-HRP reaction product, and arrowheads indicate synaptic specializations. The scale bar represents 0.5 μm. Courtesy of H. A. Lekan.

Inhibitory Transmitters and Circuits Inhibitory Transmitters Inhibitory amino acids The classical fast inhibitory neurotransmitters that are found in dorsal horn interneurons include the amino acids γ-amino butyric acid (GABA; Fig. 6A and B) and glycine (GLY). Several studies describe the distribution of GABA-containing interneurons in the rat using antibodies generated against GABA itself (Magoul et al., 1987; Todd and McKenzie, 1989) or the enzyme that synthesizes GABA, glutamic acid decarboxylase (GAD; Barber et al., 1982; Fuji et al., 1985; Hunt et al., 1981; Perez de la Mora, 1981). It is generally accepted that GABAergic interneurons have a wide distribution in the spinal cord and are present in all spinal laminae except lamina IX (Barber et al., 1982; Carlton and Hayes, 1990). In the rat dorsal horn, GABA-containing interneurons are particularly concentrated in laminae I–III (Fuji et al., 1985;

Hunt et al., 1981; Magoul et al., 1987; Todd and McKenzie, 1989). Barber et al. (1982) reported that both stalked and islet cells contain GAD immunoreactivity. Todd and McKenzie (1989) confirmed that some lamina II islet cells are immunoreactive for GABA; however, they found no GABA-containing stalked cells. GABA has been shown to coexist with several other substances in dorsal horn neurons, including GLY (Todd and Sullivan, 1990), parvalbumin (Antal et al., 1991), met-ENK (Todd et al., 1992), and nitric oxide synthase (Valtschanoff et al., 1992). Iontophoresis of GABA results in the inhibition of activity not only of dorsal horn interneurons (Curtis, 1959b; Zieglgänsberger and Sutor, 1983) but also of spinothalamic tract cells (Willcockson et al., 1984; Lin et al., 1996). Furthermore, GABA is a prime candidate for the transmitter involved in primary afferent depolarization (PAD; Eccles et al., 1962a, 1962b, 1963a, 1963b, 1963c; see reviews by Rudomin and Schmidt, 1999; Willis, 1999).

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Using autoradiography, [3H]GLY was localized in neurons in lamina III (Ribeiro-Da-Silva and Coimbra, 1980). With the advent of antibodies raised against GLY, mapping studies confirmed a concentration of GLYcontaining cells in lamina III, with smaller populations noted in laminae I and II (Campistron et al., 1986; Todd, 1990; van den Pol and Gorcs, 1988). In a double labeling study, Todd and Sullivan (1990) reported that nearly all GLY-immunoreactive cells in laminae I, II, and III also contain GABA. Iontophoresis of GLY results in inhibition of activity of dorsal horn interneurons (Bruggencate and Engberg, 1968; Curtis et al., 1967a, 1967b, 1968; Werman et al., 1968; Zieglgänsberger and Sutor, 1983) and identified spinothalamic tract cells (Willcockson et al, 1984). Several lines of evidence indicate that the amino acid taurine (TAU) is a candidate inhibitory transmitter/ modulator (McBride and Frederickson, 1980; Oja and Lahdesmaki, 1974; Serrano et al., 1990), with a possible role in antinociception in the spinal cord (Beyer et al., 1988; Kuriyama and Yoneda, 1978; Smullin et al., 1990a, 1990b). TAU is present in cell bodies in laminae I and II of the rat dorsal horn and is found in terminals that contain pleomorphic vesicles and that make symmetrical synapses (Lee et al., 1992), the hallmark of inhibitory synapses. Peptides Several lines of evidence show that endogenous opioid peptides have inhibitory functions in the CNS, particularly in relation to pain pathways (Duggan and North, 1984; Frederickson and Geary, 1982; Martin, 1984; Olson et al., 1988; Yaksh and Noueihad, 1985). Opioid peptides that are present in dorsal horn interneurons include ENK and DYN. Recently, two newly recognized endogenous opioids, endomorphin1 and endomorphin-2, have been demonstrated in the central nervous system (Zadina et al., 1997). These peptides have a high affinity for the μ-opioid receptor and may be the natural ligands for this receptor (Zadina et al., 1997). Endomorphin-1 is more prevalent in the brain (including the periaqueductal gray, locus coeruleus, and amygdala) and endomorphin-2 in the superficial dorsal horn of the spinal cord and medulla (Martin-Schild et al., 1999). Endomorphin-2 is localized in primary afferent neurons (Martin-Schild et al., 1997, 1998; Pierce et al., 1998), and it is released from the isolated spinal cord following electrical stimulation (Williams et al., 1999). Intrathecal (Stone et al., 1997; Przewlocka et al., 1999) or intracerebroventricular (Ohsawa et al., 2000) administration of endomorphins results in antinociception; however, the mechanism involves interactions with a number of neurotransmitters and their receptors (Hao et al., 2000; Ohsawa et al., 2000).

Mapping studies of ENK-containing neurons have been performed in many species, including rats; such cells have been observed in laminae I–V, with a concentration in laminae I and II, particularly IIouter (Elde et al., 1976; Finley et al., 1981a; Gibson et al., 1981; Hökfelt et al., 1977a, 1977b; Hunt et al., 1980; Merchenthaler et al., 1986; Miller and Seybold, 1987; Sar et al., 1978; Senba et al., 1988; Simantov et al., 1977; Standaert et al., 1986). Iontophoretic studies indicate that ENK (or the analog, met-enkephalinamide) can produce a selective inhibition of nociceptive dorsal horn interneurons (Duggan et al., 1976, 1977, 1981; Randic and Miletic, 1978; Sastry and Goh, 1983; Zieglgänsberger and Tulloch, 1979b) and identified spinothalamic tract cells (Willcockson et al., 1986). This action can be blocked by naloxone. Both δ- and μ-opiate receptors are believed to mediate the inhibitory actions of ENK on dorsal horn interneurons (Dickenson et al., 1986; Jeftinija, 1988). A quantitative study documenting the distribution of ENK-containing cells in lamina I at different spinal cord segmental levels demonstrated significantly more immunoreactive cells in L6 and T9 (receiving visceral as well as somatic input) compared to the L4 segment (receiving principally somatic input) (Miller and Seybold, 1989). This finding suggests that opioids may play an important role in modulating the visceral input received by these segments. Double-labeling studies indicate that ENK is frequently colocalized with SP in dorsal horn neurons (Katoh et al., 1988; Ribeiro-da-Silva et al., 1991; Senba et al., 1988). The finding that an excitatory transmitter, SP, is colocalized and presumably coreleased with the inhibitory transmitter, ENK, is paradoxical. It has been hypothesized that SP cells can regulate their own excitation by means of ENK release via autoreceptors or postsynaptic mechanisms (Ribeiro-da-Silva et al., 1991). Interneurons containing DYN have a more restricted distribution in the dorsal horn, being located almost exclusively in laminae I, II, and V (Cho and Basbaum, 1988, 1989; Khachaturian et al., 1982; Ruda et al., 1988; Vincent et al., 1982; Weihe et al., 1988). More DYN Acontaining cells are present in cord segments L6 and T9 than in L4 (Miller and Seybold, 1989). As for ENK, it has been hypothesised that the differential distribution of DYN neurons in particular spinal segments may be related to visceral sensory processing. Application of DYN to the spinal cord surface can produce both excitation and inhibition of responses of nociceptive dorsal horn neurons to C-fiber volleys (Knox and Dickenson, 1987). Similar mixed results were obtained when DYN was iontophoresed onto identified STT cells (Willcockson et al., 1986). The number of cells in the dorsal horn that are DYN-immunoreactive increases in several pathological conditions. These

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include peripheral nerve and/or root transection (Cho and Basbaum, 1988, 1989) and painful conditions, such as peripheral inflammation (Iadarola et al., 1988; Weihe et al., 1988) and neuropathy (Kajander et al., 1990). The increase occurs in both local interneurons and in projection neurons (Nahin et al., 1989). The importance of this DYN up-regulation is unclear at this time; however, the up-regulation suggests that this peptide is intimately related to the processing of primary afferent input in the dorsal horn. Another peptide that has been shown to have inhibitory effects in both the periphery and the dorsal horn is SOM. Labeled cells are concentrated in lamina II, with fewer observed in lamina I (Dalsgaard et al., 1981; Finley et al., 1981b; Ho, 1988; Hunt et al., 1981; Johansson et al., 1984; Mizukawa et al., 1988; Rosenthal and Ho, 1989; Schroder, 1984; Seybold and Elde, 1980; Vincent et al., 1985). Those in lamina II have a morphology resembling islet cells (Schoenen et al., 1985). Noxious thermal stimulation of the skin results in SOM release in the superficial dorsal horn (Kuraishi et al., 1985; Morton et al., 1988, 1989). Iontophoretic application of SOM onto nociceptive dorsal horn interneurons results in their inhibition (Randic and Miletic, 1978). Thus, nociceptive input to the spinal cord dorsal horn can result in the inhibition of nociceptive transmission by release of SOM, as well as of other inhibitory substances (see below). Inhibitory Circuits The initial integrative processing of nociceptive information in the dorsal horn involves inhibitory circuits (Fig. 5). Knowledge of the involvement of opiates and endogenous opioids in pain and pain pathways has a long history (see Basbaum and Fields, 1978; Besson and Chaouch, 1987; Fields and Besson, 1988; Willis, 1982). Defining the neurocircuitry underlying inhibition of nociceptive input by opioids is complicated by the fact that there are several sources of opioid peptides in the spinal cord dorsal horn. ENK is present in interneurons, especially in lamina I and II (Bennett et al., 1982; Glazer and Basbaum, 1983); cell bodies projecting to supraspinal sites (Nahin and Micevych, 1986); fibers descending from the medulla (Finley et al., 1981a; Hökfelt et al., 1979; Hunt and Lovick, 1982; Millhorn et al., 1987b); and DRG cells (Senba et al., 1982). DYN has also been located in interneurons in laminae I, II, and V (Cho and Basbaum, 1988; Ruda et al., 1988); cell bodies projecting to supraspinal sites (Leah et al., 1988; Nahin et al., 1989; Standaert et al., 1986); and DRG cells (in guinea pig; Gibbins et al., 1987; Kummer and Heym, 1986). As mentioned earlier, endomorphin-2 is found in the superficial dorsal horn (Martin-Schild et al., 1999) and is localized in primary afferent neurons (MartinSchild et al., 1997, 1998; Pierce et al., 1998).

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Dorsal rhizotomy results in a decrease in opiate receptor binding in the dorsal horn (Besse et al., 1990, 1992; LaMotte et al., 1976; Ninkovic et al., 1981), suggesting presynaptic modulation of primary afferents by endogenous opiates. However, few axoaxonic synapses involving primary afferent terminals have been reported where ENK or DYN is present in the presynaptic element (Aronin et al., 1981; Carlton and Hayes, 1989; Cho and Basbaum, 1989; Glazer and Basbaum, 1983; Hunt et al., 1981; LaMotte and DeLanerolle, 1983). The presence of ENK- and DYN-containing terminals postsynaptic to identified primary afferents (Carlton and Hayes, 1989; Cho and Basbaum, 1989; LaMotte and DeLanerolle, 1983) and the identification of ENK- and DYN-containing synapses (Carlton, unpublished observations; Ruda, 1982) on thalamic projection neurons is further evidence of the important role that endogenous opioid systems play in the modulation of noxious input. Other important transmitters in inhibitory circuits in the dorsal horn are GABA and GLY. There is considerable evidence for a role of GABA and GLY in the generation of IPSPs at postsynaptic sites in the dorsal horn (Bruggencate and Engberg, 1968; Curtis et al., 1959b, 1967a, 1967b, 1968; Werman et al., 1968; Zieglgänsberger and Sutor, 1983). GABA-containing terminals have been observed in synaptic contact with spinothalamic tract cells (Carlton et al., 1992). GABA has also been implicated in presynaptic inhibition, because it causes primary afferent depolarization (Eccles et al., 1962a, 1962b, 1963a, 1963b, 1963c; see reviews by Rudomin and Schmidt, 1999; Willis, 1999). Desarmenien et al. (1984) have described the coexistence of GABAA and GABAB receptors on the membranes of identified Aδ and C primary afferents in vitro. The simplest spinal pathway for the production of primary afferent depolarization, based on a central latency of at least 2.0 ms, has been postulated to include at least two synapses (Eccles et al., 1962a). The existence of axoaxonic synapses in which GABA-labeled terminals are presynaptic to primary afferent terminals has been difficult to document; however, several examples of this type of anatomical arrangement have been published (Alvarez et al., 1992; Barber et al., 1978; Basbaum et al., 1986; Carlton and Hayes, 1990; Carlton et al., 1992; Maxwell and Noble, 1987; Maxwell et al., 1990; Todd and Lochhead, 1990). What appear with greatest frequency in the superficial dorsal horn are GABAergic dendrites postsynaptic to primary afferents (Hayes and Carlton, 1990, 1992). This arrangement suggests that primary afferents may serve as a driving force for an inhibitory system; they are in a position to activate GABAergic inhibitory circuits that contribute to modulation of nociceptive transmission. Hence, disruption of primary afferent input, which occurs following peripheral nerve

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injury, rhizotomy, evulsions, and so on, could lead to a loss or attenuation of this afferent-mediated inhibition.

ASCENDING NOCICEPTIVE PATHWAYS Spinothalamic Tract For a survey of the various ascending spinal cord sensory pathways, see Tracey (Chapters 7 and 26). For a description of comparable sensory pathways for the trigeminal system, see Waite (Chapter 27). The spinothalamic tract is regarded as one of the most important pathways responsible for mediating pain sensation in humans (Gybels and Sweet, 1989). There is evidence from behavioral experiments that a pathway ascending in the anterolateral quadrant of the spinal cord, presumably the spinothalamic tract, is also important for nociception in rats (Peschanski et al., 1986; Palecek et al., 2001). The cells of origin of the spinothalamic tract in rats have been labeled by several groups using retrograde transport of horseradish peroxidase (HRP; Giesler et al., 1979a, 1981a; Granum, 1986; Kemplay and Webster, 1986; Kevetter and Willis, 1982, 1983), fluorescent dyes (Burstein et al., 1990b; Harmann et al., 1988; Kevetter and Willis, 1982, 1983), or cholera toxin subunit B (Lima and Coimbra, 1989). The most effective labeling in rats has been with fluorogold (Burstein et al., 1990b). Figure 7 shows the distribution of fluorogold-labeled spinothalamic cells in the rat spinal cord after large injections into the thalamus. In the cervical and lumbosacral enlargements, spinothalamic cells are concentrated in the marginal zone, nucleus proprius, and medial intermediate gray on the side contralateral to the thalamic injection site. The population of neurons labeled from the medial thalamus is distributed preferentially in the medial base of the dorsal horn and intermediate region (Giesler et al., 1979a). The largest group of spinothalamic neurons is actually in the upper cervical spinal cord (Burstein et al., 1990b; Granum, 1986; Kemplay and Webster, 1986; Kevetter and Willis, 1983). This cell group projects bilaterally to the thalamus. Some of the neurons give off collaterals to other structures as well (Verburgh et al., 1990). In labeling studies using HRP, it was initially estimated that there is a total of about 1000 cellular profiles of spinothalamic cells in the rat spinal cord, most of which are at cervical levels (Granum, 1986; Kemplay and Webster, 1986). However, when the spinothalamic cells were labeled with cholera toxin subunit B, the estimate increased to 6000 (Lima and Coimbra, 1989), and with fluorogold the number exceeded 9500 (Burstein et al., 1990b). This is nearly

half the number of spinothalamic cells estimated to exist in macaque monkeys (Apkarian and Hodge, 1989), using WGA-HRP as a retrograde marker. Some spinothalamic cells in rats have been found to contain neuropeptides, such as enkephalin, dynorphin, cholecystokinin, and galanin (Battaglia and Rustioni, 1992; Coffield and Miletic, 1987; Ju et al., 1987; Leah et al., 1988; Nahin, 1988). It seems likely that these peptides are colocalized with a short-lasting neurotransmitter, such as an excitatory amino acid (Westlund et al., 1992a). The axons of spinothalamic cells in rats that ascend to the lateral thalamus are in the lateral funciculus, whereas those that project to the medial thalamus do so through the ventral funiculus (Giesler et al., 1981a). The rat spinothalamic tract gives off collaterals to a variety of neural structures that are thought to play a role in the motivational-affective component of pain or to contribute to the endogenous analgesia system (Fig. 1; see Willis and Coggeshall, 2004). For example, doublelabeling studies show that there are collaterals of the spinothalamic tract to the reticular formation (Kevetter and Willis, 1982, 1983) and to the periaqueductal gray (Harmann et al., 1988; Liu, 1986; see Keay and Bandler, Chapter 10). The targets of the axons of spinothalamic cells in the lateral thalamus include the ventral posterolateral (VPL) nucleus and the posterior thalamic nuclear group (Po), and those in medial thalamus include the central lateral nucleus and the nucleus submedius (Cliffer et al., 1991; Craig and Burton, 1981; Lund and Webster, 1967; Mehler, 1969; Peschanski et al., 1983; Zemlan et al., 1978; see Groenewegen and Witter, Chapter 17). The region of termination in the VPL nucleus overlaps that of the medial lemniscus (Lund and Webster, 1967; Ma et al., 1986). The thalamic terminals of spinothalamic neurons are large endings with round vesicles and the terminals make asymmetric contacts with the dendrites of thalamic neurons (Peschanski et al., 1985b). Other synaptic terminals are either small ones containing round vesicles or profiles containing pleomorphic or flattened vesicles. These are presumed to originate from the cerebral cortex and thalamic reticular nucleus, respectively. Because the VPL nucleus in rats has few interneurons (see Groenewegen and Witter, Chapter 17), the thalamic neurons receiving synaptic contacts from the spinothalamic tract are thalamocortical projection neurons (Ralston, 1983). Several laboratories have recorded the activity of spinothalamic cells in the rat dorsal horn (Giesler et al., 1976; Palecek et al., 1992b). Many spinothalamic cells in the superficial dorsal horn and in the lateral part of the neck of the dorsal horn respond most vigorously to noxious cutaneous stimuli, although some in the nucleus proprius are mechanoreceptive. Spinothalamic cells in

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FIGURE 7 Distribution of the cell bodies of spinothalamic tract cells in the rat following injection of the retrograde tracer, fluorogold, into the thalamus. (A) The site of multiple injections, which included much of the lateral and medial thalamus on one side. (B) The locations of 4887labeled neurons at various segmental levels. Most of the spinothalamic tract cells in the cervical and lumbosacral enlargements were contralateral to the injection. From Burstein et al., 1990b.

the intermediate region and ventral horn respond best to activation of receptors in tissue deep to the skin.

Postsynaptic Dorsal Column Pathway Visceral nociceptive information is relayed to the brain by cells located around the spinal cord central canal in lamina X and also in laminae III and IV (Palecek et al., 2003). These cells are referred to as postsynaptic dorsal column cells, because the cell bodies of the neurons are in the spinal cord gray matter, rather than in dorsal root ganglia, and the axons travel uncrossed in the dorsal column. The course of the axons of postsynaptic dorsal column neurons that transmit visceral nocicep-

tive information has been described (Hirshberg et al., 1996; Wang et al., 1999). Postsynaptic dorsal column neurons relaying information about pelvic visceral pain send their axons rostrally in the dorsal column near the midline, whereas abdominal and thoracic visceral pain is relayed by axons in a position between the gracile and cuneate fasciculi (Fig. 2). The primary termination of the visceral input of the postsynaptic dorsal column cells is in the dorsal column nuclei. The third-order neurons of the dorsal column nuclei relay the information to the contralateral thalamus by way of the medial lemniscus. Other major brain sites receiving direct spinal input from neurons revealed by these and other recent tract tracing studies include the

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medullary reticular formation, the raphe, the periqueductal gray, the parabrachial nucleus, the hypothalamus, the central nucleus of the amygdala, and medial regions of the thalamus (Wang et al., 1999). These brain regions are known to be concerned with control of visceral function and affective perception of pain. They have been shown to express Fos protein after activation of visceral afferent nerve fibers in a variety of animal models (Rodella et al., 1998). Fos protein expression has also been shown to appear in the ventral reticular formation and the central nucleus of the amygdala after cyclophosphamide induced inflammation of the bladder (Bon et al., 1998) and colorectal distension (Traub et al., 1996). Recordings have been made of the responses of neurons to noxious colorectal distention or colon inflammation at several levels of this pathway in rats: sacral spinal cord, nucleus gracilis, and ventral posterolateral (VPL) nucleus of the thalamus (Al-Chaer et al., 1996a, 1996b, 1997). The same neurons also receive an input from cutaneous receptive fields that are distributed in a region comparable to that to which colon pain is referred in disease. Lesions that interrupt the dorsal column near the midline or that destroy part of the gracile nucleus largely eliminate the nociceptive visceral responses of VPL neurons. Recently, behavioral experiments demonstrated that the responses to colon inflammation plus moderate colorectal distention are eliminated by a prior lesion of the dorsal columns (Palecek et al., 2002). Similar experiments have shown that the responses of VPL neurons to distention of the duodenum, as well as writhing responses to the same stimuli, are dramatically reduced by lesions of the appropriate part of the dorsal columns (Feng et al., 1998). Dorsal column lesions also attenuated behavioral changes induced by bradykinin activation of pancreatic afferent fibers (Houghton et al., 1997). In addition, dorsal column lesions reduced the excitatory effects of pancreatic afferent fibers activated by bradykinin, as shown by recordings from neurons in the dorsal column and VPL nuclei (Houghton et al., 2001; Wang and Westlund, 2001). However, in these studies of activation of visceral afferents from abdominal organs, lesions adjacent to the midline were ineffective, whereas more laterally placed lesions near the junction of the gracile and cuneate fasciculi were effective. This result is as predicted from the anterograde tracing study that demonstrated a somatotopic arrangement of visceral input in the dorsal column (Fig. 2; Wang et al., 1999).

Spinomesecephalic Tract The spinomesencephalic tract is also likely to be involved in nociception. However, it is not clear that this

tract contributes to the sensory-discriminative aspects of pain. Instead, it seems more suited to contribute to the motivational-affective aspects of pain, as well as to trigger activity in descending control systems. The cells of origin of the spinomesencephalic tract in rats have been mapped by retrograde labeling with HRP or with fluorescent dyes (Cechetto et al., 1985; Harmann et al., 1988; Lima and Coimbra, 1989; Liu, 1983; Menétrey et al., 1982; Pechura and Liu, 1986; Swett et al., 1985; Yezierski and Mendez, 1991). The cells are concentrated in the marginal zone, the lateral part of the neck of the dorsal horn, and in the area near the central canal (laminae I, V, and X), as well as in the lateral spinal nucleus. Some spinomesencephalic tract cells contain dynorphin or enkephalin (Standaert et al., 1986). The axons of many spinomesencephalic tract neurons project rostrally through the ventrolateral white matter of the spinal cord, along with the spinothalamic tract and the spinoreticular tract. However, in rats the axons of spinomesencephalic tract cells located in lamina I and the lateral spinal nucleus ascend in the dorsal part of the lateral funiculus (Baker and Giesler, 1984; McMahon and Wall, 1983, 1985; Swett et al., 1985; Zemlan et al., 1978). The spinomesencephalic tract projects to a number of midbrain nuclei, including the cuneiform nucleus, the parabrachial nuclei, the periaqueductal gray, the intercollicular nucleus, the deep layers of the superior colliculus, the nucleus of Darschewitsch, the anterior and posterior pretectal nuclei, the red nucleus, the Edinger–Westphal nucleus and the interstitial nucleus of Cajal (Antonetty and Webster, 1975; Lund and Webster, 1967; Mehler, 1969; Zemlan et al., 1978; see Willis and Coggeshall, 2004, for references to studies on mammals other than rats). Electrophysiological recordings indicate that spinomesencephalic tract cells in rats are often nociceptive (Menétrey et al., 1980). Cells of the spinomesencephalic tract that are located in the lateral spinal nucleus have poorly defined receptive fields that appear to lie in deep tissues. McMahon and Wall (1988) have proposed that spinomesencephalic tract cells in lamina I that project to the parabrachial nuclei have a special function. As already mentioned, the axons of these neurons ascend in the dorsal part of the lateral funiculus. Stimulation of this part of the spinal cord causes a long-lasting excitation of neurons in lamina I (following a brief inhibition). However, neurons deeper in the dorsal horn are inhibited by such stimulation. The suggestion was made that lamina I spinomesencephalic tract neurons activate descending pathways that (1) reexcite the lamina I neurons and (2) inhibit deeper lying neurons of the dorsal horn. The anterior pretectal nucleus may

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be involved in the circuitry of this system (Rees and Roberts, 1993), because lamina I spinomesencephalic tract cells also project to the anterior pretectal nucleus (Cliffer et al., 1991), and the descending inhibitory effects of stimulation in the anterior pretectal nucleus depend on pathways involving the parabrachial nuclei, the ventrolateral medulla, and the dorsal lateral funiculus (Rees and Roberts, 1987; Terenzi et al., 1991, 1992). Another function of neurons in the parabrachial nuclei is to relay nociceptive information to the amygdala. Lamina I neurons project to the parabrachial nuclei, and neurons in this area project to the central nucleus of the amygdala (Bernard et al., 1989). Recordings from parabrachial neurons in this pathway show that they are nociceptive (Bernard and Besson, 1990). It has been proposed that this pathway contributes to the motivational-affective components of pain.

Spinoreticular Tract The spinoreticular tract in rats originates from cells that are concentrated in laminae V, VII, VIII, and X (Chaouch et al., 1983; Kevetter and Willis, 1982, 1983; Nahin and Micevych, 1986; Nahin et al., 1986; Peschanski and Besson, 1984). There is a less prominent projection from lamina I than the one from this lamina to the thalamus. However, there is a separate projection from laminae I and X, as well as from deep layers of the dorsal horn and intermediate region, to the dorsal reticular nucleus of the medulla (Lima, 1990; Lima and Coimbra, 1990; McMahon and Wall, 1985; see Bing et al., 1990). Some spinoreticular tract cells have been found to contain enkephalin immunoreactivity (Nahin and Micevych, 1986). The axons of spinoreticular tract cells ascend to the brainstem in the ventrolateral white matter of the spinal cord (Kevetter and Willis, 1983). The spinoreticular tract projects to a number of nuclei in the reticular formation (see Willis, 1985; Willis and Coggeshall, 1994), including the dorsal reticular nucleus of the medulla, as mentioned above.

Other Nociceptive Pathways Several other pathways are likely to include a nociceptive component. These pathways may contribute to pain under some circumstances. However, because nociceptive responses that depend on supraspinal projections, such as vocalization, are dramatically reduced following anterolateral cordotomy in rats (Peschanski et al., 1986), nociceptive signals transmitted in pathways located in the dorsal part of the spinal cord white matter are more likely to help trigger descending control systems.

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In addition to the viscerosensitive component of the postsynaptic dorsal column pathway (see above), there is a somatic component. The somatic component of the postsynaptic dorsal column pathway in rats originates from neurons located in the nucleus proprius just beneath the substantia gelatinosa (de Pommery et al., 1984; Giesler et al., 1984). It has been estimated that there are around 1500 of these neurons in rats. Most of the axons project through the dorsal funiculus to the dorsal column nuclei, although some from the cervical enlargement are in the dorsal lateral funiculus (Cliffer and Giesler, 1989; Giesler et al., 1984). The projection is a somatotopic one to the dorsal column nuclei. Recordings from these neurons have shown that many respond to innocuous stimuli, and those that prefer strong mechanical stimuli do not develop responses to noxious heat even with repeated stimuli that would presumably sensitize most nociceptors (Giesler and Cliffer, 1985). It was suggested that this pathway is unimportant for somatic nociception in rats. However, there is evidence that this pathway does contain somatic nociceptive neurons in other species (Cliffer et al., 1992; Ferrington et al., 1988; Kamogawa and Bennett, G.J., 1986; Uddenberg, 1968). There is also a spinocervical pathway in rats. The cells of origin are in the nucleus proprius, and the projection is to the lateral cervical nucleus. The number of spinocervical tract cells in rats is small (Baker and Giesler, 1984). It is unclear what the response properties of these cells are like. It is estimated that there are some 300–500 neurons in the lateral cervical nucleus in rats (Baker and Giesler, 1984; Giesler and Elde, 1985). Hair movement in large receptive fields can activate the cells in this nucleus. However, some of the cells respond to noxious mechanical or thermal stimuli (Giesler et al., 1979b), and so it is possible that this pathway can play some role in nociception in rats. A spinohypothalamic tract (Burstein et al., 1987) has been described in rats. This pathway is accompanied by axons that project directly from the spinal cord to several regions of the telencephalon, including the amygdala, septal nuclei, nucleus accumbens, and other limbic system structures (Burstein and Giesler, 1989; Burstein et al., 1987; Cliffer et al., 1991). Cells projecting to one side of the hypothalamus are distributed bilaterally in the marginal zone, neck of the dorsal horn, and the area around the central canal, as well as in the lateral spinal nucleus (Burstein et al., 1987, 1990a). Spinohypothalamic tract cells are as abundant as spinothalamic tract cells. It is not clear what the functions of the spinohypothalamic and spinotelencephalic tracts are, but it is evident that they can provide information about spinal cord activity rather directly to the limbic system and so can potentially contribute importantly to the motivational-affective

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component of pain, as well as to mediate endocrine and autonomic responses to painful stimuli.

located in layers V and VI, whereas mechanoreceptive neurons are mainly in layers II–V.

THALAMUS AND CORTEX DESCENDING CONTROL SYSTEMS Thalamus Recordings from the VPL nucleus of the rat thalamus reveal that, in addition to mechanoreceptive neurons, there are many nociceptive neurons in this nucleus (Al-Chaer et al., 1996a, 1997; Guilbaud et al., 1980; Houghton et al., 2001; Peschanski et al., 1980b). In addition, there are thermoreceptive neurons in the rat VPL nucleus (Hellon and Misra, 1973). The somatic nociceptive responses of VPL neurons are not affected by lesions of the dorsal or dorsal lateral white matter of the spinal cord, but they are eliminated by transection of the anterolateral quadrant (Peschanski et al., 1985a; Palecek et al., 2002), indicating that the somatic nociceptive responses are dependent on transmission in the spinothalamic tract and/or other anterolateral pathways. Intracellular injections of HRP into VPL neurons failed to demonstrate obvious morphological differences between mechanoreceptive and nociceptive neurons (Peschanski et al., 1984). The responses of nociceptive neurons in the VPL nucleus are enhanced by inflammation produced by injection of carrageenan into the foot and in animals subjected to experimental peripheral neuropathy (Guilbaud et al., 1986, 1987a, 1987b, 1990). Aspirin reduces the hyperresponsiveness of VPL neurons in arthritic animals (Guilbaud et al., 1982) (see section “Plastic Changes in Pathological Conditions”). The enhanced responses of gracile neurons to colorectal distention or inflammation and of VPL neurons to bradykinin activation of pancreatic afferents are reduced by spinal morphine (Al-Chaer et al., 1996b; Houghton et al., 2001). Nociceptive responses have also been recorded in the posterior thalamic nuclear group (Guilbaud et al., 1980) and intralaminar thalamic nuclei in rats (Peschanski et al., 1981). Neurons of the reticular thalamic nucleus are inhibited by noxious stimuli (Peschanski et al., 1980a).

Cortex Nociceptive neurons have been recorded in the primary somatosensory cortex (SmI) of rats (Lamour et al., 1983a, 1983b). They are intermingled with cortical neurons that respond only to innocuous mechanical stimuli, and like the mechanoreceptive neurons, they are somatotopically organized. The receptive fields of nociceptive cortical neurons are larger than those of mechanoreceptive neurons, and there are often inhibitory receptive fields. Nociceptive neurons are generally

Brain Stem Analgesia System McMahon and Wall (1988) have speculated that lamina I nociceptive neurons have ascending input to the brainstem, exciting cells, mainly in the rostral parabrachial region, which in turn excite descending control systems that have been proposed to play a role in modulation of nociception (Basbaum and Fields, 1978; Millan, 2002). An overwhelming number of studies have appeared in the literature describing regions within the brainstem that exert analgesic influences on somatosensory processing (Fig. 1; see Tracey, Chapter 7). Immunocytochemical localization of the immediate, early gene, c-Fos, following electrical stimulation of the Zusanli acupuncture site on both hindlimbs identified many of the brainstem sites (Lee and Beitz, 1993) proposed previously by others. These include the lateral parabrachial nucleus, the substantia nigra, the raphe pallidus nucleus, the dorsal raphe nucleus, the locus coeruleus, the posterior pretectal nucleus, and the ventrolateral periaqueductal gray. Periaqueductal Gray The antinociceptive effects of stimulation in the PAG of the midbrain has been studied most extensively (see Keay and Bandler, Chapter 10). Although very few spinally projecting neurons have been described in the vicinity of the PAG (Blessing et al., 1981; Castiglioni et al., 1978; Hancock and Fougerousse, 1976; Westlund and Coulter, 1980; Westlund et al., 1983), more complex brainstem connectivity has been described that could provide the necessary circuitry (Bajic and Proudfit, 1999). The PAG projects to the locus coeruleus, the medullary reticular formation, and the raphe magnus nucleus (Cameron et al., 1995; Shah and Dostrovsky, 1980; Yezierski et al., 1982a, Van Bockstaele et al., 1991; see also Keay and Bandler, Chapter 10). Some of the neurons projecting to the raphe magnus nucleus from this region contain serotonin (Yezierski et al., 1982a). An important role for the nearby anterior pretectal nucleus in nociception has been proposed in light of the fact that lowest intensities of electrical stimulation are required for antinociception in this region (Rees and Roberts, 1993). Electrical stimulation of the PAG has been found to inhibit spinothalamic tract neurons in the monkey (Gerhart et al., 1984; Hayes et al., 1979; Yezierski et al., 1982b). Ascending projections from the PAG into limbic and intralaminar thalamic nuclei may

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also provide an anatomical substrate for a role in nociception and affective responses (Eberhart et al., 1985). Recent evidence indicates that the inhibition of dorsal horn nociceptive neurons in the spinal cord following stimulation in the PAG can involve the release of a number of neurotransmitters, including not only serotonin, but also norepinephrine, GABA, and glycine (Lin et al., 1996; Peng et al., 1996a, 1996b, 1996c; Cui et al., 1999), as well as endogenous opioids (Budai and Fields, 1998; Budai et al., 1998). Raphe Nuclei The monoamine systems have been shown to be an important part of the endogenous analgesia system. At first glance, the anatomical organization of the descending serotonergic and noradrenergic systems seem to display many parallels. Both are part of the brainstem reticular system (Scheibel and Scheibel, 1958), characterized by long axons that extend to distant brain regions and highly collateralized terminal fields. The dorsal horn of the spinal cord, the site of initial somatosensory integration of incoming primary afferent information, is heavily innervated by a latticework of serotonin- (Bowker et al., 1982; Steinbusch, 1981) and norepinephrine-containing (Dahlström and Fuxe, 1965; Westlund et al., 1981; see Aston-Jones, Chapter 11) terminal varicosities. Many serotonergic neurons in the medullary raphe nuclei and the adjacent ventromedial reticular formation have spinal projections (Bowker et al., 1981). These include cells of the raphe magnus, raphe obscurus, and raphe pallidus nuclei. It appears that almost all of the spinally projecting serotonergic neurons also contain GLU and perhaps ASP (Nicholas et al., 1992). Serotonin has also been shown to colocalize with GABA in neurons with spinal projections in this region (Millhorn et al., 1987a). Many of the spinally projecting serotonergic neurons also contain peptides, such as substance P, cholecytokinin, enkephalin, somatostatin, and thyrotropinreleasing hormone (Bowker et al., 1982; Hökfelt et al., 1979). A population of spinally projecting somatostatinand enkephalin-containing neurons, in particular, are located just dorsal to the raphe magnus nucleus, which is the site proposed by Lundberg and colleagues to produce a profound tonic descending inhibition (Lundberg, 1982; see Willis, 1982). Both α1 and α2 adrenergic receptors are involved in the modulation of nociception originating in the region of the raphe magnus nucleus (Fleetwood-Walker et al., 1985; Hammond et al., 1985; Sagen and Proudfit, 1985). In fact, inhibition of nociception by serotonin in the spinal cord is also dependent on adrenergic mechanisms (Post et al., 1985).

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The serotonergic terminations in the superficial laminae of the spinal cord arise from projection cells in the raphe magnus nucleus (Basbaum et al., 1978; Bowker et al., 1982). Dorsal raphe nucleus projections to the spinal cord are limited to cervical spinal levels, suggesting that they are not involved in the control of somatosensory systems but in head and neck movements. Rather their influence on nociception is believed to be mediated by other brainstem regions to which the dorsal raphe nucleus projects, including the raphe magnus nucleus and the locus coeruleus (Yaksh, 1979). Locus Coeruleus There are numerous studies that report that electrical stimulation of locus coeruleus neurons produces antinociception in anesthetized rats (Janss et al., 1987; Jones and Gebhart, 1986; Margalit and Segal, 1979; Mokha et al., 1985, 1986; Segal and Sandberg, 1977; Sandberg and Segal, 1978). The antinociception can be blocked by intrathecal administration of noradrenergic antagonists (Jones and Gebhart, 1987). Both ipsilateral and contralateral electrical stimulation are effective in reducing heat-evoked activity in spinal cord neurons (Jones and Gebhart, 1986). Antinociception has also been reported following electrical stimulation of the adjacent ventrolateral pons, including the area containing the noradrenergic cells of the subcoeruleus and Kölliker–Fuse nuclei (Yeomans and Proudfit, 1992). Many dorsal horn neurons are inhibited by direct iontophoretic application of norepinephrine, whereas others are excited (Howe and Zieglgänsberger, 1987; Mokha et al., 1985, 1986). It has been shown that most catecholaminergic fibers in the spinal cord originate in the brainstem (Carlsson et al., 1964) and that the noradrenergic projections to the spinal cord arise almost exclusively from cell groups in the pons (Westlund et al., 1981, 1982, 1983). Descending catecholaminergic projections from the medulla arise from adrenaline-containing neurons of the rostral medulla and project primarily to autonomic regions of the thoracic spinal cord (Ross et al., 1981). Other noradrenergic cells of the medulla have ascending projections to hypothalamic and limbic structures (Ungerstedt, 1971). Of the noradrenergic cells with projections to the spinal cord, at least 80% are located in the locus coeruleus and subcoeruleus nucleus (Westlund et al., 1981, 1983, 1984). Bilateral electrothermic lesions of the locus coeruleus and immediately adjacent region reduce NE levels 40–70% in different portions of the spinal cord (Commissiong et al., 1978; Nygren and Olson, 1977). It has been reported that 50% of the coeruleospinal projections cross the midline at the segmental spinal level innervated (Björklund and Skägerberg, 1982). Anterograde studies of the

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descending projections of these two regions indicate that projections from both nuclei descend in the ipsilateral ventrolateral funiculus and project extensively to the spinal gray at all levels of the spinal cord in rats and monkeys (Clark and Proudfit, 1991; Clark et al., 1991; Jones and Yang, 1985; Westlund and Coulter, 1980). Interestingly, it was shown that more lamina I terminations arise from the locus coeruleus in rats purchased from Harlan Sprague–Dawley (Fritschy and Grzanna, 1990; Sluka and Westlund, 1992b), whereas more lamina I terminations arise from neurons of the subcoeruleus nucleus in rats purchased from Sasco (Clark and Proudfit, 1991; Clark et al., 1991; Proudfit and Clark, 1991; Sluka and Westlund, 1992b, 1992c). Variations may also occur in rats from the same vendor but purchased from various farms. Although the incidence of synaptic contacts in the dorsal horn is considerably less than in the ventral horn of rat (Rajaofetra et al., 1992), examination of the population of somatic contacts on STT neurons revealed that 8% of the contacts were noradrenergic in the monkey (Westlund et al., 1990). To date, only a few studies have addressed the role of catecholamines in somatosensory processing in rats. Norepinephrine has been shown to be equally or more potent than serotonin in selectively reducing the responses of multireceptive neurons in spinal cord to noxious but not innocuous stimuli. It has been predicted by several groups (Belcher et al., 1978; Engberg and Marshall, 1971), based on physiological and pharmacological studies, that norepinephrine causes inhibition of nociceptive neurons by a postsynaptic action. The iontophoretic application of either norepinephrine or dopamine has been shown to reduce the responses of primate spinothalamic tract cells to pulsed applications of GLU, consistent with a postsynaptic inhibitory action of catecholamines on spinothalamic tract cells (Willcockson et al., 1984). In an early study, it was noted that an increased concentration of the noradrenergic breakdown product, normethanephrine, accumulated in the cord following treatments with analgesics (Shiomi and Takagi, 1974). Electrical stimulation of the locus coeruleus, which provides a major portion of the noradrenergic projections to the spinal cord (Nygren and Olson, 1977; Westlund et al., 1983), produces analgesic effects similar to those produced by electrical stimulation of the raphe magnus nucleus (Jones and Gebhart, 1989). Iontophoretically applied norepinephrine has recently been shown to reduce selectively the responses of multireceptive neurons in spinal cord laminae IV and V to noxious (but not those to innocuous stimuli) by an α2-mediated mechanism (Fleetwood-Walker et al., 1985). Yaksh (1979) has observed that intrathecal administration of either the serotonin antagonist methysergide

or the norepinephrine blocker phentolamine is capable of only partial blockade of the analgesia produced by intracerebral opiate microinjection. Complete blockade of opiate analgesia can be achieved, however, if both antagonists are applied concurrently. Thus, it is clear that the descending noradrenergic neurons are an integral part of a complex feedback circuit. This circuit includes neurons that contain serotonin and opioid peptides and that are involved in monitoring and altering spinal somatosensory input, including noxious stimuli. An inhibitory supraspinal loop has been found to be responsible for the system of “diffuse noxious inhibitory controls” (DNIC) first described in rats by LeBars and colleagues (LeBars et al., 1979a, 1979b). Noxious stimuli cause a long-lasting inhibition of “wide dynamic range” neurons of the dorsal horn, but not of low-threshold, high-threshold, or proprioceptive neurons. The amount of inhibition is graded with stimulus intensity (LeBars et al., 1981). The inhibition depends on the activation of a pathway that ascends in the anterolateral quadrant of the spinal cord to regions that include the dorsal medullary reticular formation and another pathway that descends in the dorsal lateral funiculus from the dorsal medulla (Villanueva et al., 1986a, 1986b, 1989; see Tracey, Chapter 7). It is likely that the DNIC system is responsible for the analgesic effects of many forms of peripheral stimulation, including acupuncture and acupuncture-like transcutaneous electrical nerve stimulation.

PLASTIC CHANGES IN PATHOLOGICAL CONDITIONS Although rapid changes in neuronal excitability may occur with increases in nitric oxide and neurotransmitters, changes in receptor numbers and other cellular events could in turn produce long-term modifications of synaptic responses as is seen in persistent pain states. The subcellular mechanisms proposed for long-term facilitation of nociceptive events (Meller and Gebhart, 1993; Willis, 2002) are similar to those shown for other systems displaying long-term potentiation and synaptic plasticity. The activation of voltage-gated NMDA receptors allows Ca2+ to enter the postsynaptic neuron triggering the following cascade of events: (1) stimulation of phospholipases to produce diacylglycerol and eicosanoids; (2) formation of inositol triphosphate (IP3); (3) activation of calcium/calmodulin kinase II, protein kinase C, and protein kinase A (Fang et al., 2002; Lin et al., 2002; Peng et al., 1997); and (4) activation of the constitutive form of nitric oxide synthase and subsequent production of the labile product nitric oxide (Dawson et al., 1992; Wu et al., 1998; Lin et al., 1999a,

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1999b, 1999c). Nitric oxide would readily diffuse back to the presynaptic neuron and adjacent glia, where it activates guanylate cyclase (Garthwaite and Boulton, 1995). Evidence is increasing to substantiate that sensitization events may occur at both central and peripheral (Aley et al., 1998, 2000) neuronal endings and contribute to persistent pain. A mechanism for producing long-term changes is through the expression of immediate, early genes, such as c-fos, c-jun, and others (Herdegan et al., 1991; Hunt et al., 1987; Menétrey et al., 1989). The interactive effects of released GLU, SP, and CGRP, thus, initiate long-lasting spinal cord events leading to sensitization of dorsal horn neurons, that is, increased responsiveness of dorsal horn neurons to peripheral sensory stimuli and other excitatory inputs.

Inflammatory Pain Several arthritis models imitating human joint inflammatory conditions have been developed in animals to permit experimental studies of nociceptive processes in deep tissue. In the adjuvant arthritis model in rats, a chronic model of inflammation, both SP (Minami et al., 1989; Oku et al., 1987; Weihe et al., 1988) and CGRP (Kuraishi et al., 1989; Takahashi et al., 1988, 1990; Weihe et al., 1988) are altered in the peripheral and central nervous systems. An increase in release of SP (Oku et al., 1987) into the dorsal horn is seen in adjuvant arthritic animals. In addition, increased mRNA levels for preprotachykinin A, the precursor to SP, and CGRP are observed in both DRG cells and in the spinal cord dorsal horn (Donaldson et al., 1992; Dubner and Ruda, 1992; Minami et al., 1989; Noguchi and Ruda, 1992). Immunoreactivity for CGRP is increased both in DRG cells (Kuraishi et al., 1989) and in fibers located in the spinal cord dorsal horn (Takahashi et al., 1988, 1990). Antibodies to CGRP and substance P antagonists have been shown to reduce the hyperalgesia associated with inflammation (Kuraishi et al., 1988; Sluka et al., 1997). Therefore, one could postulate that the initial barrage of afferent activity in the early phase of the arthritis results in a depletion of SP and an upregulation of the preprotachykinin mRNA levels, resulting in an increase in the synthesis, transport, and content of the peptide in the dorsal horn of the spinal cord. Presumably, these biological mechanisms maintain a ready supply of releasable SP, which is known to contribute to the protein/plasma extravasation observed in neurogenic inflammation (Brain, 1997). Induction of an experimental model of acute arthritis of the knee joint with kaolin and carrageenan, as another example, results in limping, guarding, and increased responses to heat stimuli (hyperalgesia). Altered staining

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patterns for GLU, SP, and CGRP appear in the spinal cord dorsal horn (Fig. 8; Sluka and Westlund, 1993c; Sluka et al., 1992), dorsal root ganglia (Hanesch et al., 1993), and medial articular nerve (Westlund et al., 1992b). With a microdialysis fiber threaded through the spinal cord, an initial transient release of amino acids at the time of knee joint injection is measureable followed by a persistent release of GLU and ASP (Sluka and Westlund, 1992a; Sorkin et al., 1992). Specific nonNMDA (CNQX) or NMDA (AP7) antagonists introduced into the dorsal horn by microdialysis block the amino acid release and the accompanying acute behavioral hyperalgesia (Sluka and Westlund, 1993a). Perhaps the most surprising finding in these studies was that in addition to elimination of the sensitivity of the paw to radiant heat, direct dorsal horn spinal cord application of the non-NMDA receptor antagonist CNQX also significantly reduced the extent of the inflammation (Sluka and Westlund, 1993b). Pretreatment with the NMDA receptor antagonist did not reduce the degree of joint inflammation. The mechanism for this reduction in inflammation was shown to be the blockade of dorsal root reflexes that were triggered by the inflammation (Sluka et al., 1993, 1994; Rees et al., 1994, 1995; see Willis, 1999). These studies suggest a major role for central excitatory amino acid mechanisms in the development of the hyperalgesic state and suggest a differential central processing of sensory and tissue extravasation mechanisms involved in the development of inflammation. Plasticity of peripheral glutamate receptors is also reported for inflammatory pain models. Knee joint injections of excitatory amino acids result in hyperalgesic behaviors and glutamate antagonists administered into the knee joint reduce hyperalgesia induced in the kaolin/carrageenan knee joint inflammation model (Lawand et al., 1997). Peripheral release of excitatory amino acids and nitric oxide precursors are measurable in the joints of these animals (Lawand et al., 2000). Another model of inflammatory pain is intradermal injection of capsaicin. Although considerable experimental work using this model has been done in human subjects and in monkeys, some studies have also been done in rats. One result of an intradermal injection of capsaicin is the induction of central sensitization of neurons in the dorsal horn of the spinal cord. This leads to behavioral changes that mimic mechanical allodynia and hyperalgesia (Sluka and Willis, 1997). Similar to the central sensitization of primate spinothalamic tract neurons following intradermal capsaicin injection, the enhanced behavioral responses can be blocked by inhibitors of several signal transduction pathways that utilize protein kinases C, G, and A, as

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FIGURE 8 Photomicrograph of sections of the spinal cord from an arthritic rat. The sections were stained immunohistochemically for glutamate (GLU). An increase in the staining density is seen in the superficial dorsal horn on the side ipsilateral to the inflamed knee. Note the cells lightly stained for GLU (arrows) located bilaterally.

well as NO release (cf. Lin et al., 1996, 1997, 1999a, 1999b, 1999c; Wu et al., 1998). The enhanced responses of spinothalamic cells to peripheral inputs following mechanical stimulation of the skin and also to iontophoretic release of excitatory amino acids (Dougherty and Willis, 1992) can be accounted for by the phosphorylation of glutamate receptors that occurs after intradermal capsaicin (Zou et al., 2000). Intradermal injection of capsaicin also induces peripheral neurogenic inflammation in the skin surrounding the injection site. Flare and neurogenic edema are the result of dorsal root reflexes triggered by the afferent C fiber volley evoked by the capsaicin (Lin et al., 1999d). The dorsal root reflexes are in both Aδ and C afferents (Lin et al., 2000), presumably including peptidergic nociceptive afferents. Capsaicin was found to cause Fos expression in GABAergic dorsal horn neurons following capsaicin injection, and this was blocked by pretreatment with glutamate receptor antagonists (Zou et al., 2001).

Neuropathic Pain

allowing behavioral assessments of sensory function in the affected limb (Bennett and Xie, 1988; Kim and Chung, 1992; Seltzer et al., 1990; Attal and Bouhassira, 1999; Decosterd and Woolf, 2000; Ralston, 1998). The injured limb develops mechanical allodynia, heat hyperalgesia, cold allodynia, and spontaneous pain (Attal et al., 1990; Bennett and Xie, 1988; Kim and Chung, 1991, 1992; Seltzer et al., 1990). There are many factors that influence the development of neuropathic pain behaviors in rats, including genetic factors (Devor and Raber, 1990; Mogil et al., 1999a, 1999b; Xu et al., 2001), diet (Shir et al., 1998; Shir et al., 2001), age (Lee and Chung, 1996; Chung et al., 1995; Kim et al., 1995), degree of neuroinflammation (Maves et al., 1993; Bennett, 1999), and cytokine involvement (Wagner et al., 1998; Cui et al., 2000). The expression of these pain behaviors involves both spinal and supraspinal mechanisms (Ossipov et al., 2000; Pitcher and Henry, 2000; Porreca et al., 2002). In addition to the development of neuropathic pain behaviors, many changes occur in the rat nervous system following nerve injury and these are discussed in the following sections. Neuroanatomical Plasticity

Mononeuropathy and Plasticity in the Dorsal Horn During the past 30 years, several animal models of experimental peripheral neuropathy have been developed in the rat that mimic the pathophysiological and behavioral manifestations seen in the human condition. All of these models involve varying degrees of injury to fibers that are components of the sciatic nerve. Wall et al. (1979) describe a model in which a complete sciatic transection is followed by the formation of a neuroma. In contrast to this neuroma model, in which all of the axons in the sciatic nerve were transected, subsequent models have involved axotomy of some of the fibers contributing to the sciatic nerve and spared fibers maintain their connection with the periphery,

Transsynaptic degeneration of what are believed to be spinal interneurons might be one CNS abnormality that contributes to aberrations in sensation that occur in these nerve-injured animals. Dark “pyknotic” neurons are observed in the dorsal horn following chronic constriction injury (Bennett et al., 1989; Sugimoto et al., 1989) and TUNEL-labeled neurons are seen following sciatic nerve transection (Azuke et al., 1998). It has been suggested that in both instances these cells are damaged and perhaps dying due to excess release of glutamate in the cord and excitotoxicity (Coggeshall et al., 2001). Furthermore, it is hypothesized that these dying cells are inhibitory interneurons. If so, interruption or loss of these neurons would lead to

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disinhibition in the dorsal horn and possibly to sensory abnormalities. Other contributors to sensory aberrations following nerve injury include transganglionic degeneration of central terminals (Knyihar-Csillik and Csillik, 1981) and sprouting of myelinated primary afferents into inappropriate dorsal horn laminae. Several lines of evidence indicate that myelinated Aβ fibers sprout out of their normal territory and into lamina II (Shortland and Woolf, 1993; Woolf et al., 1992), making synaptic contact with the dendrites of neurons of lamina II (Koeber et al., 1994; Lekan et al., 1996; Woolf et al., 1995). The presence of myelinated primary afferents in an area that normally receives only unmyelinated fiber input suggests that this circuitry might account for the touch-evoked allodynia that often arises following nerve injury. However, see Tong et al., 1999; and Bird et al., 2002. There is little evidence for sprouting of unmyelinated C fibers after peripheral nerve injury (Nakamura and Myers, 1999). Neurophysiological Plasticity Results from electrophysiological recordings in various nerve injury models have demonstrated that ectopic discharges occur in both injured and uninjured peripheral fibers (Ali et al., 1999; Scadding, 1981; Wu et al., 2001, 2002) and injured DRG (Liu et al., 1999; Kajander et al., 1992; Wall and Gutnick, 1974). The discharges arise mainly in Aβ and Aδ fibers (Han et al., 2000; Kajander and Bennett, 1992; Liu et al., 1999). This activity develops soon after the injury (11–13 h) and presumably lasts for weeks (Liu et al., 1999). The ectopic activity contributes to neuropathic pain behaviors because removal of the affected DRG attenuates mechanical and heat hyperalgesia (Sheen and Chung, 1993; Yoon et al., 1999). One mechanism underlying the generation of ectopic discharges is the up-regulation and/or mobilization of ion channels, including sodium channels (Devor et al., 1992; Liu et al., 2001; Matznar and Devor, 1994; Omana-Zapata et al., 1997; Porreca et al., 1999; Waxman et al., 1994; Gold et al., 2003), potassium channels (Kajander et al., 1992; Matzner and Devor, 1994; Rasband et al., 2001), and calcium channels (Xiao and Bennett, 1995). Laird and Bennett (1992) demonstrated a decrease in dorsal root potentials at 5 and 10 days postsurgery. They hypothesize that constriction injury compromises the central mechanism responsible for primary afferent depolarization, thus impairing presynaptic inhibition. Recordings from unidentified dorsal horn neurons demonstrated increased spontaneous activity and prolonged after-discharges following stimulation of receptive fields. Similarily, recordings from identified spinothalamic tract cells at 7 and 14 days postsurgery, ipsilateral to the injury, demonstrated increased

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background activity, increased activity to nonpainful stimuli, and an increase in occurrence and magnitude of after-discharges in response to mechanical and thermal stimuli (Palecek et al., 1992a, 1992b). These electrophysiological findings reflect the behavioral changes reported in neuropathic animals and are consistent with the occurrence of a central sensitization of dorsal horn neurons. These findings are also consistent with the report by Mao et al. (1992b) of an increase in the metabolic activity in neuropathic rats of neurons in the lumbar spinal cord, as measured with 2-deoxyglucose. Neurochemical Plasticity Immunohistochemical studies have demonstrated decreases in substances known to be present in primary afferent terminals in the superficial dorsal horn, including SP (Al-Ghoul et al., 1993; Bennett et al., 1989; Cameron et al., 1991; Garrison et al., 1993) and fluorideresistant acid phosphatase (FRAP; Bennett et al., 1989). Decreases in CGRP in neuropathic animals have been reported following immunocytochemical staining (AlGhoul et al., 1993; Cameron et al., 1991) and radioimmunoassay (RIA; Bennett et al., 1989). However, using computer-assisted image analysis, Garrison et al. (1993) reported no significant change in CGRP staining but a significant reduction in SP staining in the dorsal horn ipsilateral to the nerve injury. Reductions in somatostatin (Hökfelt et al., 1994), cholecystokinin (CCK, Zhang et al., 1999), and GABA (Ibuki et al., 1997; Ralston et al., 1997) have also been reported. Increases in immunostaining densities ipsilateral to the nerve injury have been reported for glutamate (AlGhoul et al., 1993), dynorphin (Bennett et al., 1989; Kajander et al., 1990), enkephalin (Sommer and Myers, 1995), galanin (Zhang et al., 1995), vasoactive intestinal peptide (Zhang et al., 1994), and neuropeptide Y (Ma and Bisby, 1998; Wakisaki et al., 1991). Astrocytic labeling with glial fibrillary acidic protein increases (GFAP; Garrison et al., 1991). Substances associated with cellto-cell recognition and/or adhesion, such as soybean agglutinin (SBA) and RL-29, and a marker of axonal sprouting, GAP-43, also increase in the superficial dorsal horn (Cameron et al., 1991). Similar changes occur in the DRG of these animals, suggesting that the changes documented in the dorsal horn are probably of primary afferent origin (Cameron et al., 1997; Marchand et al., 1994; Nahin et al., 1994). Receptor changes in the dorsal horn have also been described and include a decrease (deGroot et al., 1997), an increase (Goff et al., 1998), and no change (Porreca et al., 1998) in opiate receptor staining. The SP receptor (NK1; Goff et al., 1998) and the CCKB receptor (Bras et al., 1999) increase following nerve injury. Both an increase (Croul et al., 1998) and a decrease (Hama et al.,

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1995) in N-methyl-D-aspartate (NMDA) receptors, as well as an increase in AMPA receptors (Harris et al., 1996; Popratiloff et al., 1998) have been observed. A decrease in GABAB binding but an increase in GABAA binding has been reported in the dorsal horn on the nerve injured side. P2X3 purinoceptors (Novakovic et al., 1999) and α-adrenergic 2C receptors increase in the dorsal horn, but α2A receptors decrease (Stone et al., 1999). It has been hypothesized that following a peripheral nerve injury, some substances involved in neural transmission are down-regulated, whereas those necessary for growth and repair are up-regulated (Cameron et al., 1991; Nielsch and Keen, 1989). Although this general phenomenon appears to be true, several timecourse studies using nerve injury models have reported that peptide mRNA levels in the DRG or peptide staining densities (SP, CGRP, GAL, VIP, NPY) in the dorsal horn have little or no correlation with pain behaviors (Cameron et al., 1997; Munglani et al., 1996; Nahin et al., 1994). Two exceptions appear to be met-enkephalin and μ-opiate receptors, which both show increases in staining densities that coincide with resolution of the neuropathic pain behaviors (Goff et al., 1998). Nerve injury models have been very useful in elucidating the role of the sympathetic system in the development and maintenance of sensory abnormalities following nerve injury. Sympathectomy alleviates neuropathic symptoms not only in humans (Loh and Nathan, 1978; Richards, 1967) but also in these animal models (Kim et al., 1993; Shir and Seltzer, 1991; Seltzer and Shir, 1991). Extensive sprouting of sympathetic fibers into DRG of injured spinal nerves and the formation of “baskets” of noradrenergic fibers around large-diameter DRG cells has been observed (Fig. 9;

Chung and Chung, 2001; Chung et al., 1999, 1996). Furthermore, ectopic firing of DRG cells decreases after sympathectomy (Han et al., 2000), suggesting the DRG develop an adrenergic sensitivity following nerve injury. It is anticipated that the insights gained into mechanisms underlying the sensory abnormalities in these models will identify novel therapies and/or explain the mechanisms underlying existing successful therapies. For example, it has been demonstrated that the NMDA antagonist MK-801, delivered intraperitoneally (Davar et al., 1991; Garrison et al., 1993) or intrathecally (Mao et al., 1992b; Yamamoto and Yaksh, 1992), attenuates the hyperalgesia observed following this nerve injury. Furthermore, local anesthetics applied to the injury site (Dougherty et al., 1992a; Mao et al., 1992a, 1992b; Seltzer et al., 1991) or injected systemically (Devor et al., 1992) attenuate not only the painrelated behaviors but also the ectopic injury discharges. Dextrorphan, a noncompetitive NMDA receptor antagonist, attenuated the heat hyperalgesia in the Bennett and Xie model in a dose-dependent manner (Tal and Bennett, 1993). Acute and chronic treatment with tricyclic antidepressants, a class of drugs frequently used to treat human neuropathic pain, resulted in antinociception that was naloxone-reversible in the animal models (Ardid and Guilbaud, 1992). Gangliosides, substances that have been shown to be effective in treating symptoms of diabetic and toxic peripheral neuropathies (Gregorio et al., 1990; Triban et al., 1989), also reduced the nociceptive responses associated with the Bennett and Xie model of experimental peripheral neuropathy (Hayes et al., 1992). Gabapentin (Neurontin), an anticonvulsant, decreases mechanical allodynia in

FIGURE 9 (A and B) Sections taken through dorsal root ganglia from rats made neuropathic by a tight ligature of the L5 and L6 spinal nerves. The sections were stained immunocytochemically for tyrosine hydroxylase and demonstrate sprouts of sympathetic postganglionic axons that form whorls around the dorsal root ganglion cells. From Chung et al., 1996.

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nerve injured rats (Abdi et al., 1998; Field et al., 1999; J.X. Hao et al., 2000) and suppresses ectopic discharges (Pan et al., 1999). Gabapentin is currently being widely used in the treatment of human neuropathic pain.

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spinothalamic neurons following intradermal injection of capsaicin in rats. J. Neurosci. 20, 6989–6997. Zou, X., Lin, Q., and Willis, W. D. (2001). NMDA or non-NMDA receptor antagonists attenuate increased FOS expression in spinal dorsal horn GABAergic neurons after intradermal injection of capsaicin in rats. Neuroscience 106, 171–182. Zou, X. J., Lin, Q., and Willis, W. D. (2002). Role of protein kinase A in phosphorylation of NMDA receptor 1 subunits in dorsal horn and spinothalamic tract neurons after intradermal injection of capsaicin in rats. Neuroscience 115, 775–786.

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28 Gustatory System ROBERT F. LUNDY, JR., and RALPH NORGREN Department of Behavioral Science, College of Medicine The Pennsylvania State University Hershey, Pennsylvania, USA

The gustatory apparatus bridges the sensory systems that deal with the external world and those that report on the internal one. The receptors are located in the oral cavity, which brings food and fluids from outside of the body into the gastrointestinal tract. Taste buds work in concert with oral thermal and tactile receptors to evaluate these substances, but in the brain, gustatory afferent axons segregate more with the visceral than with the somatic sensory system. The central gustatory system parallels the other visceral afferent systems that reach the brain over the cranial nerves but also maintains a close relationship to the central trigeminal nuclei. Due to its transitional location at the beginning of the alimentary canal, taste has several advantages over other visceral afferent modalities. The receptors are accessible and the adequate stimuli, easily specified. As with many external stimuli, humans can describe both the quality and intensity of sapid chemicals with considerable accuracy. As with many internal stimuli, humans also describe tastes in affective terms; they are either pleasant or aversive. Thus, on a number of dimensions, the gustatory system spans the exteroceptive and interoceptive sensory domains (Sherrington, 1906). Because we know so much about the former and so little of the latter, within the brain at least, taste has served as a foil for the somatosensory systems, but as a guide to their visceral counterparts. C. J. Herrick (1931) classified taste as a special visceral afferent system and this chapter respects that conception by emphasizing what is distinctive about taste within the brain. Most of the features that are shared with the visceral and somatic afferent systems

The Rat Nervous System, Third Edition

are cross-referenced to the chapters in this book that deal with those subjects (see Saper, Chapter 24, and Waite, Chapter 27). After briefly considering the peripheral gustatory apparatus, the bulk of the chapter will summarize what is known, and unknown, about the central taste system, that is, the nuclear areas and their connections. Subsequently, the cytoarchitecture of the central relays is reviewed, followed by short sections on neurochemistry and function. Purely sensory issues, such as coding, are not considered. As will become obvious, studies of the central gustatory system are unevenly distributed. More deal with the brainstem than with the forebrain, and the data often are indirect, that is, gustatory function is inferred from anatomical localization rather than from electrophysiological responses. In sensory systems that command large volumes of the central nervous system, inferring function from location poses little risk. In the gustatory system, where central relays are only a few hundred microns in their largest extent, strictly anatomical inferences can be misleading. The possibilities for confusion based on size are exacerbated by the close anatomical and functional association of taste with other visceral sensory systems. Nevertheless, more studies of taste use rodents than animals of any other order, so that these experimental animals provide the most complete picture of the central gustatory system. The focus of this chapter is on the organization of the gustatory system within the rodent brain. More detailed accounts of other aspects of taste, including neural coding and its central organization in other species, are available in both books and review articles.

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The most recent book length treatments of taste are edited by Cagan (1989); Finger, Silver, and Restrepo (2000); Getchell et al. (1991); and Simon and Roper (1993). In addition to the relevant chapters in these books, a variety of different perspectives can be obtained from review articles (Frank, 2000; Gilbertson et al., 2000; Hettinger and Frank, 1992; Kruger and Mantyh, 1989; Norgren, 1984, 1985, 1990; Norgren et al., 1989; Scott, 1992; Smith and St. John, 1999; Smith and Margolskee, 2001; Spector, 2000; Travers et al., 1987).

PERIPHERAL ANATOMY As in other mammals, taste buds in rats are segregated into distinct subpopulations within the oral and pharyngeal cavities. These receptor subpopulations differ in size, density, innervation, chemical sensitivity, and presumably function. As far as is known, however, the axons that carry gustatory afferent activity all terminate in the nucleus of the solitary tract in the medulla. The peripheral anatomy of the gustatory system is complex, but reasonably well documented. Most taste

buds (75%) are in three subpopulations on the dorsal surface of the tongue, the fungiform, foliate, and circumvallate papilla (Fig. 1). The fungiform papilla are scattered on the anterior two-thirds of the tongue and their gustatory afferent activity is carried by the axons of the chorda tympani nerve. This branch of the facial nerve comes close to being purely gustatory. Only about half of its axons are sensory but, of those, the vast majority respond to sapid stimuli. The chorda tympani also innervates small subpopulations of taste buds on the buccal wall and the sublingual organ, as well as a few receptors in the foliate papilla. Most gustatory receptors in the foliate, and all those in the circumvallate papilla, are supported by axons from the glossopharyngeal nerve. This cranial nerve actually innervates more than 60% of the gustatory apparatus but, because it also carries the somatosensory fibers for the posterior tongue and part of the oropharynx, taste axons are in the distinct minority (Boudreau et al., 1987; Frank, 1991). The other subpopulations of taste buds are on the palate, pharynx, and larynx. A small number of buds (5%) occur in or near the oral opening of the

FIGURE 1 Schematic view of the rat oral cavity showing the individual subpopulations of taste buds (arrows). Their cranial nerve innervation is given in parentheses. Abbreviations: BW, buccal wall; CN VII/CT, chorda tympani branch of the facial nerve; CN VII/GSP, greater superficial petrosal branch of the facial nerve; CN IX, linguotonsilar branch of the glossopharyngeal nerve; CV, circumvallate papillae; FOL, foliate papillae; FUN, fungiform papillae; GS, geschmacksstreifen; NID, nasoincisor ducts; PPF, posterior palatal field. Adapted with permission from Norgren et al. (1989).

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nasoincisor ducts just posterior to the incisors on the hard palate. A larger subpopulation (11%) appears on the soft palate. About half are packed densely on a slightly elevated surface at the junction of the hard and soft palate, termed the Geschmacksstreipen by Kaplick (1953), and the remainder are scattered on the middle third of the soft palate back to the opening into the nasopharnyx (Fig. 1; Miller, 1977). After bilateral section of the greater superficial petrosal nerve, all of the taste buds in the nasoincisor ducts and up to 85% of those on the soft palate disappear, indicating that most of these receptors are innervated by this branch of the facial nerve (Miller and Spangler, 1982). Although technically continuous with the distribution on the soft palate, the taste buds found in the pharynx and larynx (10% of the total) are considered a separate subpopulation (Belecky and Smith, 1990; Travers and Nicklas, 1990). Most or all of these receptors are innervated by axons of the superior laryngeal branch of the vagus nerve. Both their anatomical distribution and their spectrum of chemical sensitivity make it unlikely that they function in food selection as do taste buds in the oral cavity (Smith and Hanamori, 1991). Some difficulty arises in differentiating chemical and somatosensory functions in these areas, but the sensory receptors of the pharynx and larynx do trigger swallowing and other reflexes that protect the airway. In addition, some evidence implicates them in the process of monitoring fluid ingestion, specifically water intake (see Norgren, 1991, and Bradley, 2000, for reviews). Few would dispute that the oral trigeminal system interacts with the gustatory apparatus in the appreciation of flavor. The nature and location of that interaction, however, remains unknown. Early on, taste was reputed to be a function of the trigeminal system. Even in the late 19th century, this position was bucking considerable contrary evidence and, by World War II, it had been all but abandoned (Norgren, 1990). Nevertheless, based on anatomical evidence, Liem et al. (1990) concluded “that pathways of taste in the palate are of trigeminal origin” (p. 72). As mentioned above, other anatomical studies indicate that the greater superficial petrosal nerve carries the bulk of the gustatory axons for the palate (Hamilton and Norgren, 1984; Miller et al., 1978; Miller and Spangler, 1982). Direct electrophysiological recordings of gustatory responses from the greater superficial petrosal nerve effectively refute the strongest version of the assertion by Liem et al., but cannot discount the possibility that palatal fibers traveling within the trigeminal nerve also respond to sapid stimuli (Harada and Smith, 1992; Nejad, 1986). In fact, trigeminal fibers innervate most taste buds, as do parasympathetic efferents. When trigeminal afferent axons (i.e., capsaicin-sensitive) in the anterior

tongue are electrically stimulated, the responsiveness of single chorda tympani fibers to NaCl is altered (Wang et al., 1995). The function of the parasympathetic efferents is unknown. In addition, intraoral trigeminal axons terminate in the nucleus of the solitary tract [Sol] (Liem et al., 1990; Hamilton and Norgren, 1984) and some Sol neurons respond to both sapid and somatosensory stimuli (Travers and Norgren, 1995). Finally, some taste fibers in the chorda tympani and glossopharyngeal nerves and many of those in the superior laryngeal nerve respond to tactile or thermal, as well as to chemical, stimuli. Thus, the substrate for the interaction of somatosensory and gustatory afferent activity exists at the level both of the receptor cells and at the first central synapse.

CENTRAL ORGANIZATION Nucleus of the Solitary Tract The cranial nerves that carry gustatory and somatosensory axons from the oral cavity distribute in an overlapping rostrocaudal order within the nucleus of the solitary tract (Sol) of rodents (Åstrom, 1953; Torvik, 1956). The afferent components of the facial nerve, called the intermediate nerve of Wrisberg, innervate the gustatory apparatus on the anterior tongue, as well as the hard and soft palate, and terminate densely in the rostral tip of the nucleus. A few fibers extend in a tail caudally for some distance. The glossopharyngeal nerve supplies the taste buds on the caudal tongue and its axons end caudal to the main facial nerve area in a distribution that does not begin to thin out until the left and right Sol join on the midline. Some of these axons also continue even further caudally and a few pass to the contralateral side. Caudal to the main glossopharyngeal area, the axons of the vagus nerve take over the remaining Sol, which coalesces on the midline and trails into the spinal cord. Axons from the trigeminal nerves also reach the lateral edge of the Sol, not at the tip, but back along side the glossopharyngeal distribution. More recent studies, prompted by ever improving anatomical techniques, have refined and extended these observations without controverting the overall pattern (Contreras et al., 1982). Transganglionic anterograde labeling, primarily with horseradish peroxidase, permitted tracing the axons of individual nerves, nerve branches, and receptor surfaces. Among other things, this revealed the central representation of apposing receptor surfaces, such as the tongue and the palate (Hamilton and Norgren, 1984). As mentioned above, the two branches of the facial nerve, the chorda tympani and the greater superficial petrosal, innervate the

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anterior tongue and the palate, respectively, and their sensory axons are primarily gustatory. Rather than segregate, the terminations of these two nerves are almost coextensive within the Sol (Fig. 2A). Similarly, the superior laryngeal branch of X, which serves the pharynx and larynx, is coextensive within the Sol with the caudal half of the distribution from the linguo-

tonsilar branch of IX. Neither of these nerves is purely gustatory in function, but they do innervate partially apposing surfaces. Similar patterns appear in hamster and rabbit (Hanamori and Smith, 1986, 1989; Whitehead and Frank, 1983). Injecting anterograde tracers directly into the receptor surfaces provides a complimentary view of central

FIGURE 2 Schematic horizontal representations of the nucleus of the solitary tract in the rat. Anterior is toward the top of the page. (A) The terminal distribution of the peripheral gustatory nerves. (Filled circles) Chorda tympani branch of VII, (open circles) greater superficial petrosal branch of VII, (open squares) linguotonsilar branch of IX, (closed triangles) superior laryngeal branch of X. The dashed line separates the medial from the lateral division of the Sol. Reproduced with permission from Hamilton and Norgren (1984). (B) The distribution of neurons that responded to oral stimulation. (Filled circles) Taste responses from the anterior oral cavity (ao), (filled triangles) taste responses from the posterior oral cavity (po), (filled diamonds) whole mouth taste responses, (closed triangles) anterior oral cavity mechanical stimulation, (open triangles) posterior oral cavity mechanical stimulation. Shaded areas further caudal indicate the termination of afferent axons from the pharynx (ph), larynx (la), esophagus (es), stomach (st), and cecum (ce). Reproduced with permission from Travers (1993). (C) Overlapping terminations of afferent axons from anterior alimentary tract. (Filled circles) Soft palate, (open squares) pharynx, (filled triangles) esophagus, (open circles) stomach. Reproduced with permission from Altschuler et al. (1989). Abbreviations: DC, caudal extent of the dorsal cochlear nucleus; IV, the level at which the medial border of the nucleus of the solitary tract meets the fourth ventricle. The scale bar (0. 5 mm) is for A only. The course of the solitary tract is outlined in B (dashed lines) and C (solid lines).

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sensory organization. The most extensive study of this sort by Altschuler et al. (1989), revealed a similar rostrocaudal, overlapping organization of the soft palate, pharynx, and esophagus (Fig. 2C). In contrast, the vagal sensory axons from the stomach exhibited little or no overlap with the sensory distributions from the other structures. On the tongue, this approach has only been reported for the circumvallate papilla (Bradley et al., 1985). Although their injections labeled only a small percentage of the surface innervated by the linguotonsilar branch of IX, the central distribution that they report appears to be essentially identical to that for the entire nerve. In both of these investigations, however, the anatomical tracer cannot distinguish between gustatory and somatosensory fibers. On the anterior tongue, this distinction is possible because the somatosensory axons travel separately in the lingual nerve, whereas the taste fibers follow the chorda tympani. In fact, much of the trigeminal distribution to the Sol originates from the lingual nerve and it does not overlap extensively with the termination of the chorda tympani nerve (Hamilton and Norgren, 1984; Whitehead and Frank, 1983). Except for the anterior tongue, separating gustatory from somatosensory function is not an anatomical but a physiological problem. When just the anterior tongue is stimulated, taste-responsive neurons occur primarily within the anatomically defined chorda tympani terminal area of the Sol (see McPheeters et al., 1990). When the anterior tongue is sequestered in a chamber and the remainder of the oral cavity is flushed with sapid chemicals, the anatomy and the physiology do not match as well. First, many neurons respond to stimuli applied to both fields. Second, although there is a rostrocaudal trend, interdigitation is the rule, that is, anterior tongue only, posterior tongue only, and dually responsive neurons are mixed together. Finally, posterior tongue taste neurons occur only in the anterior half of the anatomically defined glossopharyngeal field (Dickman and Smith, 1989; Halpern and Nelson, 1965; Sweazey and Smith, 1987). This form of stimulation separates only the anterior tongue from all of the other receptor field subpopulations. When the individual subpopulations are stimulated independently, the sequence of receptive fields in the Sol becomes more predictable, but it still fails to correspond precisely with the primary afferent terminal fields (Fig. 2B). Neurons that responded only to sapid chemicals applied to anterior tongue were concentrated in the anterior tip of the Sol, but a few turned up caudally well into the glossopharyngeal terminal area (Travers, 1993; Travers and Norgren, 1995). The posterior tongue-taste neurons were concentrated caudal to the anterior tongue cells and the circumvallate

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responses tended to be posterior to those from the foliates (Halsell et al., 1992). Nevertheless, the gustatory neurons did not occupy the entire glossopharyngeal zone in the Sol. In another sample, taste neurons that responded to stimuli in the anterior oral cavity, the anterior tongue and the nasoincisor ducts, on average, were further rostral in the Sol than those that responded to taste applied in the posterior oral cavity. This relationship only held, however, when cells with receptive fields on the soft palate were included (Travers et al., 1986). Because the soft palate is innervated by the greater superficial petrosal, and not the glossopharyngeal nerve, the anatomy and the physiology still remain somewhat out of register. Unlike the gustatory neurons, somatosensory responses from the anterior oral cavity did correspond quite closely to the distribution of lingual nerve axons in the Sol (Travers and Norgren, 1995). To summarize, the gustatory zone of the nucleus of the solitary tract can be defined either anatomically, based on the central distribution of the nerve branches known to carry taste axons, or physiologically, based on the location of neurons that respond to sapid stimuli. Overall, both measures produce similar outlines, although the internal details may differ. The taste area of the Sol extends from the rostral tip caudally to the point at which the medial border of the nucleus abuts the fourth ventricle, a distance of about 2.0 mm [Paxinos and Watson, 1998 (Plates 63–72)]. This area includes gustatory, thermal, and tactile sensory input from the oral cavity in a roughly somatotopic ordering. In the mediolateral plane, taste responses are confined to the middle third of the nucleus; somatosensory responses occur along the lateral edge and scattered among the taste neurons, particularly caudally. The distributions of the glossopharyngeal and superior laryngeal nerves continue further caudally in the Sol, ending in a tail confined to the interstitial subnucleus that extends to the level of the obex. Gustatory responses from the pharynx and larynx have not been documented in the Sol of rats, but in the sheep they overlap those from the posterior tongue, but extend further caudally as well (Sweazey and Bradley, 1988, 1989). The neurons in the rostral half of the nucleus of the solitary tract have one major synaptic target, the pontine parabrachial nuclei, and several minor, or at least less well documented, ones (Norgren and Leonard, 1973; Norgren, 1978). These latter projections are known primarily through anatomical investigations and thus cannot be termed gustatory with certainty. Nevertheless, the three studies that provide the bulk of this evidence are quite consistent with one another and with other relevant data (Beckman and Whitehead, 1991; Norgren, 1978; Travers, 1988). A few axons from the rostral Sol may reach the trigeminal and facial motor nuclei directly,

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but the vast majority spread into the parvicellular reticular formation ventral and rostral to the gustatory relay or course caudally within the nucleus itself to terminate in the hypoglossal nucleus and the adjacent reticular formation. When the tracer injection is confined to the gustatory areas of Sol, relatively few labeled fibers end in the caudal, visceral afferent aspect of the nucleus. Retrograde labeling studies confirm these projections. Injections in the hypoglossal nucleus resulted in labeled neurons in the anterior Sol, whereas injections into the facial or trigeminal motor nuclei produced few, if any, labeled cells (Travers, 1988; Travers and Norgren, 1983). Similar deposits into the parvicellular reticular formation labeled numerous cells in the Sol (400–500), 9% of these cells also expressed gustatory-evoked Fos-like immunoreactivity (Travers and Hu, 2000). The same parvicellular reticular areas that receive input from the anterior Sol contain preganglionic parasympathetic salivatory neurons and cells that synapse in all four of the oral motor nuclei (Contreras et al., 1980; Travers and Norgren, 1983). About half of the salivatory neurons in the reticular formation of cats were driven by electrical stimulation of the chorda tympani or glossopharyngeal nerves (Ishizuka and Murakami, 1992; Murakami et al., 1989). The primary, and probably the only, rostral target of axons from the anterior Sol of rodents is the parabrachial nuclei (PB) in the dorsal pons. In fact, neurons at all levels of the Sol synapse in these nuclei apparently in a fairly intricate pattern (Herbert et al., 1990; see also Fig. 3 of Saper, Chapter 24). Some neurons in the caudal Sol also distribute axonal processes widely in the forebrain, but neither anatomical nor electrophysiological evidence exists for any secondorder gustatory cells that project rostral to the pons (Norgren and Leonard, 1971, 1973; Ogawa et al., 1981; Ricardo and Koh, 1978). In primates, on the other hand, many neurons in the gustatory end of the Sol bypass the PB and ascend directly to the thalamic gustatory relay (Beckstead et al., 1980). Retrograde tracers confirm that many neurons in the anterior Sol synapse in the PB of rodents and that these connections are almost entirely ipsilateral. These data underscore several other features of the PB afferent supply. First, in the anterior Sol, the neurons that project to the PB are concentrated in the dorsal half of the nucleus, which also is where the bulk of the primary afferent taste axons terminate. Neurons with locally or caudally directed axons appear to be preferentially located in the ventral half of the Sol (Travers, 1988; Whitehead, 1990). Electrophysiological experiments find that from 35 to 80% of Sol taste neurons are antridromically invaded by stimulation in the PB (Cho et al., 2002b;

Monroe and DiLorenzo, 1995; Ogawa et al., 1984b; Ogawa and Hayama, 1984; Ogawa and Kaisaku, 1982). Second, the anatomical data emphasize that large numbers of neurons outside of the Sol, particularly in the subjacent reticular formation, but also in the trigeminal nuclei and the spinal cord, send axons to the PB (Cechetto et al., 1985; King, 1980).

Parabrachial Nuclei The parabrachial nuclei are obligate gustatory relays, but they also serve as relays for the other visceral afferent modalities found in the Sol (Cechetto, 1987). Although gustatory neurons have been documented repeatedly in the PB of rats, hamsters, and rabbits, few of these studies specifically mapped the distribution of taste within the nuclei (DiLorenzo and Schwartzbaum, 1982; see Norgren, 1984). This complicates differentiating taste and visceral afferent function within the nuclei, particularly when some overlap or convergence may exist between them (Hermann et al., 1983). In both the rat and hamster the most intense taste responses occur posteriorly in the medial parabrachial nucleus in a densely packed layer of cells just ventral to the brachium conjunctivum (Fig. 3A1–A4) (Halsell and Frank, 1991; Halsell and Travers, 1997; Norgren and Pfaffmann, 1975; Perotto and Scott, 1976; Van Buskirk and Smith, 1981). Responses also can be recorded from within, and just dorsal to, the brachium. The representation of the anterior and posterior oral cavity in the PB remains unsettled. Originally, Norgren and Pfaffmann (1975) felt that the posterior oral cavity taste receptors drove neurons that were more dorsal than those activated from the anterior tongue. The same arrangement was found for the palatal receptors versus the anterior tongue (Hayama et al., 1987). In a third study, the caudal tongue taste responses were, on average, 200 μ posterior in the PB to those elicited from the tongue tip (Miyaoka et al., 1997). Along the mediolateral axis, however, only gustatory cells responsive to posterior oral cavity stimulation were found in the external medial and lateral-inner subnuclei (Halsell and Travers, 1997). Not surprisingly, axonal projections from the anterior Sol terminate in the same areas of the PB that respond to sapid stimuli. Two other areas in the PB also appear to receive projections from anterior Sol, but have not been associated with taste function (Fig. 3B1 and B2) (Herbert et al., 1990; for more details on the Sol projections, see Fig. 3 of Saper, Chapter 24). These areas are rostral extensions from the electrophysiologcially identified gustatory zone in the posteromedial PB. These discrepancies may be explained by incomplete functional mapping studies or by Sol injection sites that involved both taste and nontaste neurons.

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FIGURE 3 Schematic coronal representations of the parabrachial nuclei. (A1–A4) The location of neurons responsive to gustatory stimuli applied to the anterior oral cavity (AO, anterior tongue and nasoincisor ducts) and posterior oral cavity (PO, posterior tongue and soft palate). A1 is approximately 700–850 μm posterior to A4. Gustatory neurons most responsive to AO stimulation are depicted as filled circles, whereas those to PO stimulation are depicted as open circles. Oral somatosensory neurons are depicted as filled (AO) and open (PO) triangles. Small dots represent cells unresponsive to gustatory and somatosensory stimulation. Scale bar is 0.5 mm. Adapted with permission from Halsell and Travers (1997). (B1 and B2) The distribution of afferent projections from different levels of the nucleus of the solitary tract. (Dark hatching) Anterior Sol, (large dots) medial and posterior Sol and central area postrema, (horizontal lines) lateral Sol, (fine dots) dorsomedial Sol and lateral area postrema. The left-hand panels is approximately 200 μm anterior to that on the right. Reproduced with permission from Herbert et al. (1990) (see also Fig. 3 of Saper, Chapter 24, this volume). Abbreviations: MPB, medial parabrachial nucleus; MPBE, medial parabrachial nucleus external; LPBC, lateral parabrachial nucleus central; LPBV, lateral parabrachial nucleus ventral; PBW, parabrachial nucleus waist part; scp, superior cerebellar peduncle (brachium conjunctivum).

The parabrachial nuclei are unique among brainstem sensory relays in that they project both to the dorsal thalamus and to the limbic system. In fact, the PB axonal ramifications in ventral forebrain are more extensive than those to the thalamus (Halsell, 1992; Norgren, 1976; Saper and Loewy, 1980). The diversity of the parabrachial efferent systems precludes considering these nuclei as traditional sensory relays. Parabrachial taste neurons have been linked to both of the ascending efferent systems, as well as the descending one. The descending PB axons related to taste project to the medullary parvicellular reticular formation and the rostral central and ventral subnuclei of the Sol (Karimnamazi and Travers, 1998). The pattern of label in the reticular

formation overlaps with that arising from gustatory responsive sites in the Sol. As mentioned earlier, this distribution includes preganglionic parasympathetic salivatory neurons and cells that synapse in all four of the oral motor nuclei. The anatomy of the ascending parabrachial connections is considered in more detail in Chapter 24 and is just summarized here, emphasizing the evidence for gustatory function. The ascending parabrachial axons remain largely ipsilateral and, with the exception of the thalamus, this holds true for their terminal distributions as well. In the midbrain, the PB fibers concentrate in two areas, ventrolateral to the central gray in the dorsal tegmental bundle and somewhat more ventral and lateral in

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the central tegmental tract (Paxinos and Watson, 1998, Fig. 43). The more ventral fascicles fan out into the substantia nigra pars compacta and continue rostrally through the posterior lateral hypothalamus while distinctly avoiding both the subthalamic nucleus and the cerebral peduncle. The more dorsal axons ascend into the thalamus to terminate in the medial tip of the ventral posteromedial nucleus and in the adjacent intralaminar nuclei (Bester et al., 1999; Krout and Loewy, 2000). Despite this extensive synaptic field, many PB axons extend ventrolaterally through the zona incerta and the internal capsule to distribute widely in the ventral forebrain. Based on the anterograde label, the central nucleus of the amygdala and the bed nucleus of the stria terminalis receive the most impressive synaptic input from the PB, followed by the lateral hypothalamus, lateral preoptic area, substantia innominata, and part of the diagonal band of Broca. With the largest PB injection sites, a few labeled axons reach the medial septum, olfactory tubercle, and even to the vicinity of the gustatory cortex (Norgren, 1976; Saper and Loewy, 1980). Retrograde labeling studies confirm the extensive forebrain connections of the parabrachial nuclei and provide information about their intranuclear sources (Fulwiler and Saper, 1984; Halsell, 1992). In the present context, the most salient feature of these data is that injections of retrograde tracers into the thalamus label many more cells in the posteromedial PB than injections in the ventral forebrain. This implies that the axons of PB gustatory neurons synapse preferentially in the thalamus rather than the hypothalamus or the amygdala. Nevertheless, anatomical evidence derived from small PB injections indicates that areas associated with gustatory function, that is, the medial nucleus and the “waist” areas, provide most or all of the axons that terminate in the medial aspect of the central nucleus of the amygdala (Bernard et al., 1993). In addition, antidromic invasion experiments found that almost as many PB taste neurons could be driven from the ventral forebrain as from the thalamus (Norgren, 1974, 1976). The difference was that only a single electrode was placed in the thalamus, whereas the ventral forebrain contained an array that spread over several millimeters. This emphasizes the anatomical and perhaps the functional difference between the two axonal systems. The PB taste neurons projecting to the thalamus are the afferent source for a specific sensory relay to the cortex. The PB taste neurons that project into the ventral forebrain reach many more, and much larger, neural structures, but the function of these nuclei apparently is neither gustatory nor even purely sensory. Gustatory responses have been recorded from neurons in the hypothalamus, amygdala, and the

intervening substantia innominata (Azuma et al., 1984; Nishijo et al., 1998, 2000; Yamamoto et al., 1989; see Norgren, 1984, and Norgren et al., 1989, for other references). These gustatory neurons are neither pure nor plentiful. Depending on the study, between 2 and 10% of the cells tested actually respond to sapid chemicals and the majority of these respond to other, unrelated sensory stimuli. Because these studies employed sapid stimuli applied to the oral cavity, the source of the gustatory afferent activity driving these neurons cannot be determined. Electrical stimulation in the PB does activate neurons in the ventral forebrain, sometimes with short latencies (Mogenson and Wu, 1982). Only one such study, however, has determined whether neurons that could be driven from the PB also responded to sapid stimuli (Block and Schwartzbaum, 1983). In their sample of 103 cells from the substantia innominata and amygdala of rabbits, 11 responded to taste stimuli and 6 of these also responded to electrical stimulation in the PB. Taken together these studies using antidromic, orthodromic, and sapid stimulation clearly establish that gustatory afferent axons from the parabrachial nuclei reach an extensive area in the ventral forebrain. The functions subserved by these gustatory projections to the limbic system remain to be deciphered. The parabrachial gustatory connections to the thalamus present a distinctly different arrangement (Fig. 4A and B). As mentioned earlier, these axons concentrate in a small area, the medial extension of the ventral posteromedial nucleus, but they do so bilaterally. Some fibers appear to cross in the pons, but many more do so within the thalamus itself (Norgren and Leonard, 1973). Retrograde labeling studies confirm the bilateral nature of the PB projections to thalamus, although there is some inconsistency in the number and location of the contralateral neurons. After horseradish peroxidase injections into the thalamic taste area, some studies depict relatively few labeled neurons in the contralateral PB, distributed more or less as a mirror image of those on the ipsilateral side [Ogawa et al., 1981; Fulwiler and Saper, 1984; Yasui et al., 1983 (cat)]. In hamsters, similar injections resulted in essentially equal bilateral label in the PB (Halsell, 1992). Yet other investigations also observed equal numbers of labeled neurons in the PB, but the distributions were not symmetrical. Ipsilaterally the cells were concentrated posteromedially, as well as in the external medial area at the lateral tip of the brachium conjunctivum, whereas contralaterally the cells were primarily located in the external medial area (Krout and Loewy, 2000; Ogawa and Akagi, 1978). Anatomical data underscore another point about the parabrachiothalamic gustatory system. In the hindbrain, taste is traditionally associated both

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FIGURE 4 Schematic coronal representation of medial parabrachial nucleus projections to the thalamus. (A) Location of parabrachial neurons labeled with the anterograde axonal tracer Phaseolus vulgaris leucoagglutinin (PHA-L). The distance of each section caudal to the coronal plane where the inferior colliculus merges with the pons is indicated in micrometers in each lower left corner. Scare bar is 1 mm. (B) The distribution of anterograde labeling (mainly terminal) in the gustatory and intralaminar regions of the thalamus. Scale bar is 1 mm. (C) The distribution of lingual sensory modalities in the thalamus of the rat. (Dark with open dots) Taste responses, (open with vertical line) tongue thermal responses, (open crosshatching) tongue tactile. Adapted with permission from Emmers (1977). Abbreviations: D3V, dorsal third ventricle; fr, fasiciculus retroflexus; ml, medial lemniscus; MPB, medial parabrachial nucleus; MPBE, medial parabrachial nucleus external; PBW, parabrachial nucleus waste part; PF, parafascicular nucleus; VPM, ventral posteromedial nucleus; VPPC, ventral posterior thalamic nucleus parvicellular (e.g., gustatory nucleus); scp, superior cerebellar peduncle. Source of A and B: Differential projections to the intralaminar and gustatory thalamus from the parabrachial area: A PHA-L study in the rat, Bester et al., The Journal of Comparative Neurology, Copyright © 1999 Wiley-Liss, Inc. Adapted by permission of Wiley-Liss, Inc., a subsidiary of John Wiley & Sons, Inc.

anatomically and functionally with the oral somatosensory system. The parabrachial projections to the thalamus, however, largely avoid the oral trigeminal areas (VPM). In addition to the gustatory area, which does include some tongue thermal and tactile responsive neurons, parabrachial cells terminate in the adjacent intralaminar nuclei, which are identified with somatic and perhaps viscerosomatic pain (Bester et al., 1999; Krout and Loewy, 2000). The electrophysiological data also support a bilateral projection from PB gustatory neurons to the thalamus. In the parabrachial nuclei, all gustatory and most mechanoreceptive neurons have ipsilateral receptive fields (Hayama et al., 1987). Somewhat more than half of the PB taste cells can be antidromically driven from

the thalamus (Ogawa et al., 1984a, 1987). Of these, approximately half were driven from both the ipsilateral and the contralateral side. Most of the rest were activated only from the ipsilateral thalamic electrodes, but a few were exclusively contralateral (Hayama and Ogawa, 1987). The implication is that most of the PB taste neurons that project to the thalamus synapse ipsilaterally, but as many as half of these have collaterals that terminate on the opposite side. A single, retrograde double-labeling study confirms this general arrangement (Voshart and van der Kooy, 1981). In addition to their projections to the thalamus and ventral forebrain, neurons in the dorsal pons also project monosynaptically to the insular cortex (i.e., to the vicinity of gustatory cortex) (Lasiter et al., 1982; Saper,

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1982b; Shipley, 1984; Whitehead et al., 2000). Three aspects of this projection are relevant to the central gustatory system. First, PB neurons apparently reach large expanses of cortex in addition to the area associated with taste (Saper, 1982a). Second, compared with what arrives from the thalamus, the parabrachial projection to insular cortex is sparse and quite posterior (Allen et al., 1991). Third, although taste has been associated with these projections, the evidence is entirely anatomical (Lasiter et al., 1982). Stimulating electrodes implanted in or near gustatory cortex reliably influenced 31% of a sample of PB taste neurons, but none of the cells were antidromically invaded (DiLorenzo and Monroe, 1992). Thus, although some PB axons reach insular cortex, the case for monosynaptic gustatory connections remains unproven.

Thalamic Taste Relay The location of the thalamic gustatory relay has been established by anatomical, electrophysiological, and lesion-behavioral techniques (Norgren, 1984). The name assigned to the taste relay varies from one publication to another, but its position relative to other thalamic nuclei does not. The nomenclature has been discussed elsewhere (Jones, 1985; Norgren, 1984, 1990). In Paxinos and Watson [1998 (Plates 34–37)], this issue was circumvented by calling the area the ventral posterior thalamic nucleus, parvicellular part (VPPC). The only other issues that require discussion are the extent of the area, the functions it subserves, and its projections (see also Simerly, Chapter 14). The first two issues are interrelated and probably contribute to the confusion about the third. The gustatory relay consists of a thin band of cells extending medially from the thalamic ventral posteromedial nucleus (VPM), the trigeminal somatosensory relay, between the parafascicular nucleus and the medial lemniscus (Fig. 4B). Its exact borders are difficult to define on histological grounds alone. Classically, the area was defined as a small celled or parvicellular subdivision of VPM, but that definition originally was based on cats and monkeys. At least one author asserts that, in rodents, the cells in this area are not smaller, but fewer or less dense than in the remainder of VPM (Halsell, 1992). Regardless of the reason, investigators outline the area differently—from less than 1.0 mm in mediolateral extent to more than 2.0 mm. These differences may account for some of the discrepancy in the functional localization of lingual sensory modalities. Electrical stimulation of the chorda tympani nerve activates the medial third of the area; similar stimulation of the lingual nerve, the lateral two-thirds (Emmers et al., 1962). A similar arrangement occurs using

physiological stimulation in that gustatory responses are most medial, lingual thermal activity somewhat more lateral, and tongue tactile neurons most lateral (Fig. 4C) (Blomquist et al., 1962; Kosar et al., 1986b). In other studies, however, single neurons that responded to sapid chemicals were concentrated in the middle third of the subdivision and were intermingled with tongue tactile cells (Ogawa and Nomura, 1988; Normura and Ogawa, 1985). In either case, the histologically distinct subdivision of VPM represents the thalamic relay for all lingual sensory modalities rather than just taste. In all likelihood, it contains neurons that respond to sensory stimulation of the entire oral cavity, including the tongue. The cortical projections of the thalamic gustatory relay follow a trajectory similar to that of other ventral thalamic radiations (Norgren and Wolf, 1975). Fibers pass rostrolaterally through the zona incerta, where some collateral synapses may occur, and then enter the internal capsule. They continue coursing through the ventral third of the caudate-putamen and reach the external capsule. At this point, they exhibit the only peculiarity in the projection. The axons turn dorsally in the external capsule, traveling up to 2.0 mm, before entering the deep layers of the cortex, often in a near hairpin turn. Although they terminate in cortex that overlies the claustrum, the gustatory fibers from the thalamus avoid passing through that structure (Figs. 5A1–A3). Thalamic gustatory projections become controversial only in relation to the amygdala. Earlier treatments of the system with anterograde tracing techniques do not mention the amygdala (Krettek and Price, 1977; Norgren and Wolf, 1975; Wolf, 1968). Nor was the issue raised when injection sites were placed under physiological guidance and kept small (Kosar et al., 1986b). Nevertheless, a number of both retrograde and anterograde labeling studies report connections between the thalamic gustatory relay (VPPC) and the amygdala. In several of the retrograde studies, the injections into the amygdala spread into the overlying caudate-putamen and thus could have labeled thalamocortical fibers en route. This possibility is acknowledged by Carter and Fibiger (1977) and is a plausible explanation for similar observations in a more recent study (Yasui et al., 1991b). Using a lateral approach circumvented this problem and, when the injections were in the central or medial amygdala, labeled neurons still appeared in the VPPC (Ottersen and Ben-Ari, 1979). In cats, a robust projection was observed from the vicinity of gustatory thalamus to the amygdala, but it terminated in the lateral nucleus (Yasui et al., 1984, 1987). Only functional data can resolve the issue, but a comprehensive examination of thalamoamygdaloid projections by Turner and Herkenham (1991) came up with a sort-of anatomical

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FIGURE 5 Gustatory cortex in the rat. (A2 & A3) The dorsal and ventral limits of terminal labeling on cortex after injection of an anterograde axonal tracer, biotinylated dextran amine, into the thalamic gustatory area (A1). The dashed lines in A2 and A3 approximate the boundaries of granular, dysgranular, and agranular insular cortex. The distance of each section caudal (A1) and rostral (A2 and A3) to bregma is indicated in micrometers in each lower right corner. Scare bars are 0.5 mm. Adapted with permission from Nakashima et al. (2000). (B) A higher power photomicrograph of a coronal section from one hemisphere of a rat brain. The marking lesion (arrow) marks the point at which the more dorsal tongue thermal responses ceased and gustatory responses began. It also marks the transition from cortex with a distinct layer of granule cells (IV) to cortex with few, if any granule cells. Reproduced with permission from Kosar et al. (1986a). (C) The distribution of tongue tactile (triangles), tongue thermal (squares), and gustatory (stars) responses projected onto the lateral surface of the rat’s cerebral cortex. Below the rhinal sulcus (dashed line) are neurons that responded to air blown into the nostrils. The dotted line represents the ventral limit of the granule cell layer. Adapted with permission from Kosar et al. (1986a). (D) Schematic lateral view of the rat cerebral cortex with major histological subdivisions. The shaded area is dysgranular (dorsal agranular) insular cortex that receives afferent input from the thalamic gustatory area (VPPC) and from the parabrachial nucleus (PB). Adapted with permission from Allen et al. (1991). Abbreviations: AI, agranular insular cortex; Cl, claustrum; DEn, dorsal endopiriform nucleus; DI, dysgranular insular cortex; ec, external capsule; fr, fasiciculus retroflexus; GI, granular insular cortex, ml, medial lemniscus; mt, mammillothalamic tract; OB, olfactory bulb; PF, parafascicular nucleus; Pir, piriform cortex; PRh, perirhinal cortex; rf, rhinal fissure; S1, primary somatosensory cortex; S2, secondary somatosensory cortex; SPF, subparafascicular thalamic nucleus; VPM, ventral posteromedial nucleus; VPPC, ventral posterior thalamic nucleus parvicellular (e.g., gustatory nucleus).

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compromise. First, they confirmed other reports documenting extensive projections to the amygdala from midline thalamic nuclei that are in close proximity to the gustatory relay. Confounding with these systems probably accounts for some data attributed to taste (Yasui et al., 1989, 1991b). Second, with one well-placed injection, they found evidence for a relatively weak projection from VPPC to the most dorsal aspect of the amygdala, including the central and lateral nuclei. Another study supports this general arrangement and provides evidence that separate populations of VPPC neurons project to the insular cortex and amygdala (Nakashima et al., 2000).

Cortical Gustatory Area In most respects consideration of the cortical gustatory area mirrors that of the thalamus. The general location has been known for more than 40 years and controversy on this topic is more apparent than real (Norgren, 1984). As in the thalamus, these apparent problems in localization involve differences in nomenclature and, in some cases, attributing functional significance to anatomical data. When thalamic gustatory sites are identified on the basis of electrophysiological responses to sapid stimuli and the lesions or injections are restricted to a physiologically meaningful size, the resulting cortical terminal field extends in a narrow band just dorsal to the rhinal fissure on either side of the middle cerebral artery (Fig. 5) (Kosar et al., 1986b; Norgren and Wolf, 1975). To be precise, the terminal fields are at most 0.5 mm wide dorsoventrally and up to 2.5 mm long anteroposteriorly. More of the field is rostral to the middle cerebral artery than caudal. At the rostral end, it is about 1.0 mm dorsal to the rhinal fissure, but caudally that decreases to less than 0.5 mm (Figs. 5A2 and A3). The terminal field maps onto a histologically distinct cortical subdivision that (1) overlies the claustrum and (2) lacks a distinct layer of granule cells (Fig. 5B) (see Fig. 13 of Saper, Chapter 24). Only minor differences occur in the delineation of this strip of cortex, but it has been given many different names. For present purposes, the only name differences that require comment are the (dorsal) agranular insular and the dysgranular insular cortex. The former term derives from Rose (1928) and was used by Kosar et al. (1986a) to designate the histologically distinct area from which they recorded gustatory responses (Fig. 5C). Cechetto and Saper (1987) located taste responses in the same zone but called it dysgranular, because “scattered granule cells can be observed” in layer IV (p. 30). This minor terminological difference is of little import unless it fosters confusion among those less schooled in the

mysteries of cytoarchitectonics. First, this histologically distinct layer extends both rostral and caudal to the area that contains taste neurons. Second, it is but one in a series of parallel zones that begins dorsally with the primary or secondary somatosensory representations, switches to granular insular cortex and then to the zone that contains taste neurons, and ends with (ventral) agranular insular cortex forming the banks of the rhinal sulcus (Fig. 5D). Using histological criteria alone, the borders of these zones are subtle or even arbitrary. Fortunately, their afferent and efferent projections also differentiate these histologically distinct strips (Allen et al., 1991; Guldin and Markowitsch, 1983; Krushel and van der Kooy, 1988; Shi and Cassell, 1998). More important to the present context, so do their sensory properties. On cortex, lingual sensory modalities are represented with touch most dorsal, then temperature, and taste most ventral (Kosar et al., 1986a; Yamamoto et al., 1981). As mentioned above, the gustatory neurons are located in the dysgranular zone. Tongue temperature responses occur immediately dorsal to the taste neurons in cortex with a distinct, but thin, granule cell layer (i.e., the granular insular zone). The tongue tactile neurons are still further dorsal where the granule cell layer reaches maximal thickness (Fig. 5C) (Kosar et al., 1986a). A similar ordering of sensory responses was observed while testing single units in awake, behaving rats (Yamamoto et al., 1988). Using a more detailed analysis, another study made the point that gustatory responses usually contain a somatosensory component and some lingual tactile responses contain a gustatory component (Katz et al., 2001). They located most of their gustatory responses in dysgranular and agranular insular cortex and some of the mixed taste– tactile responses in oral somatosensory cortex, but none in the interposed granular insular cortex. The histological definition of gustatory cortex has been confirmed by Cechetto and Saper (1987) using similar converging evidence. Rather than test other lingual stimuli, these investigators used procedures designed to increase visceral afferent activity. They found neurons that responded to these visceral stimuli were located in granular insular cortex. Although it may seem unlikely that tongue thermal neurons would be intermingled with visceral sensory neurons, as with the nomenclature, this discrepancy is more apparent than real. Both are in granular insular cortex, but most of the visceral responses are located caudal to the level of the tongue temperature responses (compare Fig. 3 in Kosar et al., 1986a, with Fig. 5 in Cechetto and Saper, 1987). This ordering also is consistent with the relative positions of primary and secondary somatosensory cortex, because the primary lingual area is at the rostroventral extreme of S1 and the visceral afferent

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area is subjacent to S2 (Fig. 5D) (Donaldson et al., 1975; Kaas, 1983; Welker, 1971; Welker and Sinha, 1972). Unfortunately, not all of the data fit easily into this story. The gustatory responses referred to above were elicited from the anterior tongue and they occurred in the region from which evoked potentials were recorded after chorda tympani nerve (CT) stimulation. The corresponding region for the glossopharyngeal nerve was immediately posterior to the CT zone, overlapping both the visceral afferent area and S2 (Yamamoto et al., 1980b; see Norgren, 1984, for earlier references). In fact, posterior tongue and pharyngeolaryngeal responses were located caudal to the anterior tongue area, and these neurons often responded to visceral afferent and painful stimuli (Hanamori et al., 1997, 1998a, 1998b). Using lightly anesthetized, paralyzed rats, other investigators stimulated the entire oral cavity with sapid stimuli and not only located taste neurons in both granular and agranular insular cortex but also found that they were intermixed with mechanoreceptive and thermoreceptive cells (Ogawa et al., 1990, 1992a, 1998b). Thus, one set of data places gustatory cortex in two orderly progressions. Oral sensory modalities are ordered dorsoventrally with tongue tactile neurons above in true granular cortex, tongue thermal cells next in cortex with a thinning granular layer that overlies the dorsal claustrum, and gustatory somata most ventral in cortex with at best scattered granule cells. In the other dimension, the anterior tongue taste representation is rostral to the posterior tongue, which in turn is rostral to visceral afferent neurons, all of them in a narrow strip of cortex sandwiched between the ventral borders of S1 and S2 and the dorsal lip of the rhinal fissure. Other data find these sensory responses in the same general area, but with modalities intermingled across several cytoarchitectonic zones and even converging on the same cells. It is possible to reconcile or at least rationalize some of these data based on differences in the protocols, for example, the anesthetic conditions, but the confusion underscores a broader issue, of which the cortex is but an example. Because the gustatory, somatosensory, and visceral sensory systems usually are treated in isolation and often not all that systematically, we know the locations of the central taste relays but not their boundaries or how gustatory neural activity relates to other oral or visceral sensory modalities.

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resentations, gustatory cortex projects back massively to its thalamic relay, but it also apparently sends axons directly to the parabrachial nucleus and the nucleus of the solitary tract (Norgren and Grill, 1976). In addition, neurons in or near taste cortex have reciprocal connections with ventral forebrain areas implicated in gustatory function (i.e., the amygdala and hypothalamus). The amygdala and hypothalamus, in turn, project back to the parabrachial nuclei and the nucleus of the solitary tract, but not to the gustatory thalamus (see Norgren, 1985, for a review; Krushel and van der Kooy, 1988; Shi and Cassell, 1998; van der Kooy et al., 1984; Veening et al., 1984; see also Saper, Chapter 24). The vast majority of these data are strictly anatomical and thus suffer from the same risks of functional inference encountered in the ascending limb of the taste system. Indeed, considerable functional evidence attests to the fact that these reciprocal projections influence the entire autonomic nervous system (see Yasui et al., 1991a, for example). Only a few studies document that the descending forebrain projections actually engage lower order gustatory neurons. Yamamoto and his colleagues (1980a) isolated 15 thalamic neurons that were activated by orthodromic stimulation from the chorda tympani nerve and antidromic stimulation from the vicinity of gustatory cortex. Of these cells, 9 also exhibited significant excitability changes that outlasted the cortical stimulation and were thus attributed to orthodromic effects. Similar effects were noted in an earlier study (Ganchrow and Erickson, 1972). A single study has examined the reciprocal relationship between taste cortex and the amygdala. More than half the neurons recorded in one area were influenced by stimulation in the other, but none were antidromically invaded (Yamamoto et al., 1984). Similarly, electrical stimulation of gustatory cortex, central nucleus of the amygdala, and lateral hypothalamus can influence both Sol and PB taste neurons (Cho et al., 2002a; DiLorenzo and Monroe, 1992; Li et al., 2002; Lundy and Norgren, 2001a; Matsuo et al., 1984; Murzi et al., 1986; Smith and Li, 2000). These forebrain-responsive neurons were either excited or inhibited. A few of the Sol neurons responsive to lateral hypothalamic stimulation showed signs of antidromic invasion (Cho et al., 2002a). Moreover, recent evidence indicates that descending projections from each of these forebrain sites converge to reach many of the same brainstem gustatory neurons (Cho et al., 2003; Lundy and Norgren, 2001b).

Reciprocal Projections The highest levels of the central gustatory system, cortical or subcortical, form reciprocal connections with each of the taste relays that are afferent to them, as well as with one another. As with other primary sensory rep-

Summary The cardinal points and caveats relating to the central gustatory system are as follows (Fig. 6). Even if the exact dimensions and internal organization remain

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unclear, the nuclear relays and their connecting pathways leading to the neocortical representation of taste are known with some certainty. The uncertainty derives from the small size of the gustatory relays and the relative inaccessibility of much of the peripheral gustatory apparatus. The first central relay for taste is in the anterior half of the nucleus of the solitary tract. It is arranged in a roughly somatotopic order with the taste buds on the anterior tongue represented most anteriorly in the nucleus. Lingual and intraoral somatosensory responsiveness also occur within the Sol, separate and distinct from those found in the central trigeminal system. Although the data are far from complete, these

two characteristics—the somatotopic arrangement of the taste receptors and the close association with lingual somatosensory modalities—appear to be characteristic at all levels of the central gustatory system. From the Sol, gustatory neurons project largely ipsilaterally to the pontine parabrachial nuclei. Based on electrophysiological evidence, the third-order taste neurons are concentrated posteromedially in the PB adjacent to and even within the brachium conjunctivum. These parabrachial gustatory cells send axons both to the ventral forebrain and to the thalamic taste relay on the medial tip of the lingual somatosensory area. Within the thalamus, gustatory neurons can have ipsilateral,

FIGURE 6 Schematic summary of the gustatory system in the rat brain. Outlines of coronal sections through the rostral medulla (lower right), pons, diencephalon, hypothalamus and amygdala, and cerebral cortex covering a rostrocaudal distance of about 12 mm. The solid lines connecting the panels represent axons known to convey gustatory information; dashed lines, axons associated with the taste system, but without documented sensory function. None of the lines follow actual pathways, nor do the bifurcations necessarily imply collateral projections. Abbreviations: AI, agranular insular cortex; Amyg, amygdala; Cl, claustrum; cp, cerebral peduncle; DC, dorsal cochlear nucleus; DI, dysgranular insular cortex; GI, granular insular cortex; LC, locus coeruleus; LH, lateral hypothalamic area; MD, mediodorsal thalamic nucleus; Me5, mesencephalic trigeminal nucleus; Mo5, motor trigeminal nucleus; Pr5, principal sensory trigeminal nucleus; Sp5, spinal trigeminal nucleus; Sol, nucleus of the solitary tract; Ve, vestibular nuclei; VPM, ventral posteromedial thalamic nucleus. Redrawn with permission from Paxinos and Watson (1998).

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contralateral, or even bilateral receptive fields. Thalamic taste neurons ascend in parallel with the trigeminal projections, but terminate in a histologically distinct strip of cortex squeezed between the primary somatosensory representation dorsally and the rhinal fissure ventrally. As mentioned earlier, the parabrachial nuclei project extensively into the ventral forebrain, including the hypothalamus, amygdala, preoptic area, and the bed nucleus of the stria terminalis. Electrophysiological confirmation does indicate that some of these axons are gustatory in nature, but their relative numbers, exact terminations, and functions are unknown. Finally, it should be noted that, at least in the forebrain, the gustatory system closely parallels the central ramifications of the other visceral afferent systems that first synapse in the nucleus of the solitary tract. Any differences appear to be more quantitative than qualitative. The gustatory system apparently has more substantial thalamocortical representation, whereas the (vagal) visceral afferent projections favor the ventral forebrain.

CYTOARCHITECTURE The cytoarchitectonic distinctions of the thalamus and cortex are described elsewhere (see Groenewegen and Witter, Chapter 17, and Zilles, Chapter 23, respectively). Those aspects that are directly relevant to the gustatory system are covered in the preceding section of this Chapter. Therefore, this section deals only with the cytoarchitectural characteristics of taste cells in the brainstem.

Nucleus of the Solitary Tract The same features of the central gustatory system that impede determining its boundaries and connections also interfere with descriptions of cellular morphology. The areas that contain taste neurons are small and, absent any functional identification, gustatory neurons are difficult to distinguish from other cells in the same nucleus or some adjacent structures. In fact, several widely cited cytoarchitectonic partitionings of the nucleus of the solitary tract do not extend to the rostral, largely gustatory part of the cell group (Kalia and Mesulam, 1980; Loewy and Burton, 1978). Earlier studies divided the nucleus into medial and lateral divisions with the medial part being small rostrally, but taking over the entire cross section posterior to the obex (Contreras et al., 1982). Subsequent analysis in rats and hamsters further subdivided the Sol on cytoarchitectonic grounds (Kalia and Sullivan, 1982; Whitehead, 1988). The hamster data are more relevant here because that treatment focuses on the rostral half of the

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nucleus. Five subdivisions are recognized—a rostrocentral area that is surrounded by the medial, ventral, rostrolateral, and dorsal areas (Fig. 7A). The cells in the rostral Sol are small, only a few are more than 25 μ in their long axis, and the distribution of shapes does not change dramatically. Thus the subdivisions are based as much on the density and clustering of the cells as on their size and shape. The cytoarchitecture of the caudal or visceral sensory half of the Sol is presented in Fig. 2 of Saper (Chapter 24). If the afferent and efferent projections of the Sol are factored in, however, these internal boundaries become less abstract. The rostrocentral subdivision and its caudal extension receive most of the primary afferent terminations from the chorda tympani and glossopharyngeal nerves and provide most of the efferent projections to the parabrachial nuclei (Fig. 7B and C) (Whitehead, 1990). The rostrolateral subdivision receives sensory terminations primarily from the lingual nerve; the other three areas get little peripheral input at all. The dorsal subdivision is histologically the most distinctive, because it contains relatively few, and uniformly small, cells. This thin band, barely 50 μ wide, forms an unmistakable boundary between the Sol and the overlying vestibular nuclei. The medial subdivision could be considered an extension of the dorsal except that it contains a few larger cells, some of which are preganglionic parasympathetic efferents of the chorda tympani and glossopharyngeal nerves (Contreras et al., 1980). Finally, the ventral and ventrolateral subdivisions contain more relatively large neurons than the other subdivisions. For this reason, the ventral and lateral borders of the Sol are difficult to distinguish without accompanying fiber stains. Anterior Sol neurons also can be distinguished on the basis of their nuclear profiles and their dendritic arborizations. Some 75% of Sol neurons have deeply invaginated nuclei. Two-thirds of these cells have larger, elliptical somata (average 18 μ long by 13 μ wide); the other third are round or cuboidal and smaller (average, 12 μ long by 10 μ wide) (Davis and Jang, 1986). The other 25% of Sol neurons have smooth nuclear envelopes and are about equally divided between the smaller and larger sizes. Based on their membrane characteristics, anterior Sol cells come in two varieties also in a roughly 70/30 ratio (Bradley and Sweazey, 1990). Current injection studies reveal four different characteristic patterns in anterior Sol neurons, but apparently they do not correlate with cell morphology (Bradley and Sweazey, 1992). Except for the fact that more of the larger cells are located ventrally, the differentiation based on nuclear characteristics does not obviously match the cytoarchitectonic subdivisions. Golgi studies of the same neuronal populations, how-

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FIGURE 7 Cross sections through the rostral nucleus of the solitary tract of the hamster. (A) Neuronal morphology in the subdivisions of the Sol. Cells: e, elongate; ls, large stellate; s, stellate; t, tufted; Subdivisions: D, dorsal; M, medial; RC, rostral central; RL, rostral lateral; T, solitary tract; V, ventral. (B) Termination of the chorda tympani (filled squares), glossopharyngeal (open circles), and lingual (open triangles) nerves with respect to the cytoarchitectonic subdivisions of the rostral Sol. (C) Distribution of neurons retrogradely labeled after a large injection of horseradish peroxidase into the ipsilateral parabrachial nucleus. Each panel is taken from approximately the same rostrocaudal level and reproduced at approximately the same scale. Adapted with permission from Whitehead (1988) (A and B) and Whitehead (1990) (C).

ever, are in substantial agreement and recognize two general categories of cells based on dendritic trees (Davis and Jang, 1988; Whitehead, 1988). Larger neurons with elliptical or fusiform somata have a few, long, relatively unbranched dendrites that can extend beyond the borders of the Sol. The other type of neuron is somewhat smaller, especially dorsally in the nucleus, and has more, shorter, and more branched dendrites spreading radially from the soma. A single,

intracellular recording and labeling study demonstrates that both morphological groups are in fact gustatory neurons (Renehan et al., 1994). This same group further divided these two types into six groups based on total internal cell volume, soma crosssectional area, mean dendritic branch segment length, swelling density, spine density, and number of primary dendrites. For all types, the dendrites usually are more widespread in the mediolateral plane, which

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also is the preferred orientation of the primary afferent axons as they leave the solitary tract (Davis, 1988; Renehan et al., 1994). This planar arrangement is consistent with observations that primary afferent axons of the facial nerve synapse mainly on small diameter (distal) dendrites within the Sol (Whitehead, 1986).

et al., 1990). The dendrites of the cells that failed to be invaded by thalamic stimulation more closely matched those of the Golgi material associated with the gustatory zone.

NEUROCHEMISTRY Parabrachial Nuclei As in the Sol, cells in the PB are small (8–20 μ in diameter) and not easily differentiated in Nissl stains. Traditionally, the nuclei were labeled medial, lateral, and Kolliker–Fuse, the former two being separated by the brachium conjunctivum. Histological analysis by Fulwiler and Saper (1984) recognized seven subdivisions in the lateral nucleus, two in the medial, but left the Kolliker–Fuse as is (see Figs. 4–9 of Saper, Chapter 24). Subsequent partitioning by others came up with somewhat fewer, but not substantially different, subdivisions (Halsell and Frank, 1991; Kolesarova and Petrovicky, 1987). Based on the electrophysiological evidence, gustatory neurons occur posteriorly in a part of both the medial and ventral lateral subnuclei, among the fascicles of the brachium conjunctivum itself, and in the external medial subdivision (Halsell and Frank, 1991; Halsell and Travers, 1997; Norgren and Pfaffmann, 1975). The cells in these areas share some characteristics. They are smaller (12 μ in diameter); lightly staining; and round, oval, or fusiform in shape. In the vicinity from which gustatory responses can be recorded, the two subnuclei are joined by thin bands of cells passing through the brachium conjunctivum, the so-called “waist area” (Fulwiler and Saper, 1984). The cells within the brachium are similar in size and Nissl staining to those in the adjacent subdivisions (Lasiter and Kachele, 1988a). The external medial subdivision of PB contains “large multipolar neurons whose long axis is oriented horizontally” (Fulwiler and Saper, 1984, p. 233). In Golgi material, only two types of neurons are obvious in the gustatory zone of PB, fusiform, and multipolar. In rats, the multipolar cells are larger than the fusiform variety, but in hamsters, the distribution overlaps (Davis, 1991; Lasiter and Kachele, 1988a). In both species, the dendritic trees of these neurons are neither extensive nor complicated, having few branches or spines. The dendrites do pass into and through the brachium, but seldom extend beyond the gustatory zone. In a study using intracellular labeling with horseradish peroxidase, however, lateral parabrachial neurons that were antidromically invaded from the posteromedial ventral thalamus did have extensive dendritic trees with many spines and terminal tufts. These dendrites often extended beyond the immediate area and even beyond the boundaries of the PB (Luo

For a chemical sense, taste has precious little (neuro)chemistry associated with it. The paucity of gustatory neurochemical data is all the more striking when compared with the other visceral afferent systems that pass through the Sol, because they apparently utilize a bewildering array of neurotransmitters and peptides. This close association between the two afferent systems means that an impressive neurochemical literature must be surveyed to uncover the small amount that is relevant to taste. A survey of such scope would be inappropriate here, because much of the information is covered in other chapters in this text. In addition, Kruger and Mantyh (1989) have sifted through the neurochemical literature and summarized the data related to the gustatory system. The synopsis that follows relies heavily on their review, simply updating it in places to cover newer observations.

Periphery Both substance P and calcitonin gene-related peptide (CGRP) are associated with axons that innervate taste buds. Although some controversy exists about substance P, the consensus seems to be that both peptides occur more in perigemmal rather than intragemmal axons and, thus, are not related to gustatory sensory function per se (Silverman and Kruger, 1990). The fact that neither peptide shows up prominently in either the geniculate ganglion or in the rostral Sol reinforces that notion. On the basis of their own data from Necturus taste buds and from the literature on other species, Roper and his colleagues concluded that it was unlikely that acetylcholine, histamine, or the catecholamines were acting as transmitters between the taste receptor cells and the primary afferent neurons (Jain and Roper, 1991; Roper, 1992; Welton et al., 1992), whereas serotonin, GABA, and glutamate are present either in the cells of the bud or in intragemmal axons and probably do play a role in transducing stimulus molecules into neural traffic. In addition, the following other peptides have been localized either in the cells of taste buds or in axons associated with them—bombesin, cholecystokinin (CCK), galanin, gastrin releasing peptide (GRP), histidine, isoleucine, neurokinin, neuropeptide Y (NPY), somatostatin, and vasoactive intestinal polypeptide (VIP; Welton et al., 1992). In a

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study that used antisera to many of these peptides (and others as well), only GRP immunoreactivity was observed in virtually all of the somata in the geniculate ganglion of the rat (Hardebo et al., 1992). Despite this impressive list of sightings, “there has yet to emerge a definitive identification for substances that mediate transmission between (gustatory) receptor cells and sensory afferents” (Welton et al., 1992, p. 518).

Nucleus of the Solitary Tract Of the dozen or more peptides identified in afferent axons entering the Sol, in Sol neurons, or in efferents leaving it, invariably the highest concentrations are associated with the caudomedial, visceral afferent half of the nucleus (Kruger and Mantyh, 1989). In fact, immunohistochemical surveys of the Sol often fail to illustrate its anterior half, presumably because the concentrations of reaction product are so low (Kalia et al., 1984; Yamazoe et al., 1984). Mantyh and Hunt (1984) not only noted this contrast but also observed that the number of peptides associated with the central visceral and gustatory system decreased further as they ascended the neuraxis. Some primary afferent axons that reach the anterior Sol in the facial and glossopharyngeal nerves contain both substance P and CGRP. Peripherally these peptides are localized more at the base of taste buds than within them, suggesting a nonchemosensory function. Substance P also is associated with both central and peripheral trigeminal neurons that project to the Sol (South and Ritter, 1986; Zhang et al., 1991). Nevertheless, substance P influences gustatory-elicited responses of neurons in the rostral Sol (Davis and Smith, 1997; Liu et al., 1991). Serotonin receptors are found in relatively high density in the anterior Sol and the adjacent oral trigeminal nucleus, implying that this transmitter plays a role in processing gustatory afferent activity (Thor et al., 1992). When applied iontophoretically to gustatory neurons in the Sol, 5-HT primarily produced sustained excitation (Thornton, Nicolaidis, and Norgren, unpublished observations, 1992). Although some serotonergic axons are present in the gustatory nerves, most of 5HT innervation is thought to arise centrally. Similarly, NPYimmunoreactive axons, but not somata, appear in some density in the Sol at least as far rostrally as the caudal gustatory zone. Unlike the serotonin receptors, however, NPY is more dense medially and absent from the adjacent trigeminal nucleus (Harfstrand et al., 1987). The origin of the NPY axons is not clear, but immunoreactive cells are present in the caudal Sol, the paratrigeminal nuclei, and the reticular formation. The excitatory amino acid glutamate does appear to mediate synaptic transmission between chorda tympani fibers

and taste neurons in the Sol. When an antagonist of the AMPA/kainite receptor, but not the NMDA receptor, is injected into the Sol, the responses to gustatory stimulation of the anterior tongue are blocked (Li and Smith, 1997). Nevertheless, no study has established that glutamate is present in chorda tympani axons or is released following nerve stimulation. Peptides are equally scarce within cells in the anterior Sol. Mantyh and Hunt (1984) screened the Sol for six different peptides and tyrosine hydroxylase (TOH). At caudal and intermediate levels, all six peptides and TOH were located within some neurons. Within the anterior Sol, only three peptides were observed—CCK, enkephalin, and somatostatin—and then only in a few neurons. When the rostral levels are displayed, similar patterns are evident in other studies (see ZardettoSmith and Gray, 1990), and Riche et al., 1990, for examples). Although neither acetylcholine nor the catecholamines figure prominently in the anterior Sol (Kruger and Mantyh, 1989), GABA is present in about one-fifth of the neurons. GABAergic-immunoreactive terminals are plentiful in the rostral Sol and, based on vesicular density, can be classified as GABA-LD (Low Density) and GABA-HD (High Density) (Wetherton et al., 1998). GABA-LD terminals apparently are preferentially found on the cell body and proximal dendrites of physiologically identified Sol taste neurons, whereas GABA-HD terminals are found on the distal dendrites (Leonard et al., 1999). One function of this synaptic arrangement is to inhibit the responsiveness of Sol neurons to sapid stimulation (Smith and Li, 1998). This population of GABAergic neurons apparently consists of interneurons, because they are not retrogradely labeled from either the caudal Sol or the parabrachial nuclei (Lasiter and Kachele, 1988b). Once again, although catecholamines and a variety of peptides have been detected in the axonal connections from the Sol to the parabrachial nuclei, they are seldom associated with gustatory projection neurons from that region (Herbert and Saper, 1990; Krukoff et al., 1992; Milner et al., 1984; Riche et al., 1990).

Parabrachial Nuclei Although the transmitters utilized by Sol taste neurons are unknown, monoamine and peptide labeled axons do terminate within the gustatory zone of the parabrachial nuclei. The fit is never perfect; the immunoreactive axons occur in other PB subdivisions as well and usually they do not even fill all of the area associated with taste. Thus, axons that stain for tyrosine hydroxylase are concentrated in the “waste area” of PB, which contains taste neurons. They also ramify throughout the lateral PB, most of which is not

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associated with taste, but not in the medial PB, which does contain gustatory cells (Block and Hofman, 1987). Galanin and corticotropin releasing factor (CRF) present similar patterns, although both of these peptides extend into the medial third of the medial nucleus (Herbert and Saper, 1990). Kruger and Mantyh (1989) list 12 other substances that have been localized in the PB, but assert that most of these are concentrated in the lateral nucleus. Some of these, particularly CGRP, substance P, and neuropeptide Y, also are associated with the external medial subnucleus, which as stated above, receives afferent axons from the rostral Sol and send axons to the vicinity of the thalamic taste relay (Fodor and Palkovits, 1991; Yamano et al., 1988). The same story holds for parabrachial neurons. Many cells contain peptides, sometimes as many as three localized in the same soma, but few are found in the areas associated with taste (Shinohara et al., 1988). Although galanin and CRF-positive fibers invade the PB gustatory area, few if any neurons of that type appear there. Cholecystokinin does occur in a few “waist” area cells, but the greatest concentration of CCKpositive neurons is in the superior lateral subnucleus (Herbert and Saper, 1990). Similarly, a few cells in the medial nucleus and the ventral lateral subnucleus, both of which are associated with gustatory function, contain CGRP, but far greater numbers occur in the external lateral and external medial subdivisions (Schwaber et al., 1988; Yasui et al., 1989). Otherwise, most other peptide containing neurons are concentrated in the lateral parabrachial nucleus, which receives afferent input from the caudal rather than the rostral Sol, and has been associated with visceral afferent functions and with pain (Bernard and Besson, 1990; Herbert et al., 1990). For additional information on the neurochemistry of the PB, see Saper (Chapter 24).

Thalamus The parvicellular extension of posteromedial ventral nucleus contains fewer peptides than either the Sol or the PB, but still a healthy variety. Manyth and Hunt (1984) observed at least modest concentrations of CCK, enkephalin, somatostatin, and substance P immunoreactivity in both cells and the neuropil. In addition, the same area contains some tyrosine hydroxylase and evidence of specific subunits of the neuronal nicotinic receptor (Kruger and Manyth, 1989). The problem here, as lower in the neuraxis, is differentiating taste from the visceral afferent relay. In the thalamus, however, the problem is exacerbated by the lack of precise definition of the visceral relay and the close association of taste with intraoral tactile and thermal sensibility. The issue is best illustrated by CGRP, the most

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well-documented peptide in this part of the thalamus. As mentioned earlier, cells containing CGRP are concentrated in the external medial and external lateral subnuclei of the PB and some of them project to VPM (Yasui et al., 1989). Axons containing CGRP preferentially terminate in the parvicellular VPM, including the most medial third that contains taste relay neurons. The afferent terminal character of the elements containing CGRP has been confirmed by electron microscopy (Williamson and Ralston, 1993). Cells containing CGRP, on the other hand, are not located in the gustatory relay, but medial to it in the subparafascicular nucleus. These CGRP neurons do not project to the gustatory cortex, but to the amygdala, striatum, and perhaps the perirhinal cortex (Yasui et al., 1989, 1991b). Based on one brief report, iontophoretically applied CGRP facilitated 5 of the 12 thalamic neurons that responded to sapid sodium (Liang and Cechetto, 1991). These authors also tested several visceral afferent modalities, as well as somatic pain stimuli, and found that CGRP increased the sensory responses in approximately the same proportion of thalamic cells regardless of the peripheral input. Thus, CGRP, which is associated with taste neurons from the periphery through to the cortex, may influence the system, but probably does not serve in gustatory coding per se.

Cortex The peptides found in or near the thalamic taste relay also occur in varying concentrations in the insular cortex. Although the “distributions have not been mapped in detail,” they are apparently more intense in the granular insular cortex and this associates them with visceral sensory functions rather than taste (Kruger and Mantyh, 1989). As alluded to above, CGRP immunoreactivity is actually more concentrated on both banks of the rhinal fissure caudal to the level of the gustatory representation. In addition, the CGRPpositive axons that do reach the vicinity of taste cortex appear to arise from cells in the PB rather than the thalamus (Yasui et al., 1989). Nevertheless, CGRP is more abundant in taste than in somatosensory cortex and the amount detected can be increased by subjecting the animal to aversive sapid stimuli (Yamamoto et al., 1990). When applied iontophoretically to taste neurons in the gustatory cortex, CGRP and substance P mainly produced inhibition of taste responses (Ogawa et al., 2000). Finally, another study examined the release of traditional neurotransmitters from gustatory cortex, but without defining the area precisely. When depolarized with KCl, these tissue samples released labeled acetylcholine, GABA, and glutamate, but not dopamine (Lopez-Garcia et al., 1990).

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FUNCTIONAL CONSIDERATIONS Taste-guided behaviors such as preference-aversion, conditioned taste aversion, and salt appetite exemplify the rudiments of dietary selection using both learned and unlearned analysis of gustatory afferent activity. This section summarizes evidence on the role of the nucleus of the solitary tract (Sol), the parabrachial nucleus (PB), the thalamic taste area (VPPC), and the gustatory cortex (DI) in the elaboration of these behaviors. When appropriate the function of peripheral gustatory nerves and other central nuclei, not traditionally considered as part of the gustatory system, are discussed. Purely sensory issues such as detection threshold and discrimination are beyond the scope of this Chapter. Virtually all the data derive from studies of behavioral deficits that follow bilateral lesions of taste nerves or central relays, or from the differential expression of c-Fos protein produced during behavioral tests. More detailed accounts of some of these issues are available (Bures et al., 1998; Norgren and Grigson, 1996; Reilly, 1998, 1999; Spector, 1995).

Preference–Aversion For present purposes, preference and aversion are considered to be the unlearned tendency for an animal to ingest or reject sapid stimuli usually with reference to water. To avoid the influences of nutritional feedback, most lesion-behavior studies used either briefaccess licking (10–30 s) or the taste reactivity tests as metrics. Taste reactivity refers to the stereotypic oromotor behaviors elicited by fluid stimuli infused directly into the oral cavity (Grill and Norgren, 1978). Surprisingly, bilateral section of the nerves serving individual taste receptor fields has little influence on concentration-dependent licking and oromotor behavior (Krimm et al., 1987; Markison et al., 1999; Spector et al., 1993b, 1996; St. John et al., 1994). Ingestive and aversive taste reactivity responses are reduced, but not eliminated, following bilateral transection of the chorda tympani (CT) and glossopharyngeal nerves (GL), respectively (Grill et al., 1992). Thus neither nerve exclusively controls either category of oromotor response. In fact, a combined transection of the CT and GL is required to abolish the concentration dependence of ingestive and aversive oromotor behaviors to sucrose, NaCl, and QHCI (Grill and Schwartz, 1992). When appetitive licking is the measure, transection of a combination of two peripheral nerves at best shifts normal preference-aversion functions to the right. This appears to be stimulus specific because bilateral removal of the CT and greater superficial petrosal nerves (GSP) disrupts normal licking to sucrose, but

not to QHCI, whereas removal of the CT plus the GL leads to the converse (Spector et al., 1996; St. John et al., 1994). Nevertheless, the rats still adjusted their licking as a function of concentration. Apparently the combined removal of the CT, GSP, and GL is required to flatten the concentration-response function for sucrose (Spector et al., 1996). Collectively, these investigations suggest that unconditioned responsiveness to a specific category of sapid stimuli is not the exclusive domain of any one gustatory nerve. Rather, normal responsiveness to changes in intensity of gustatory stimuli depends on centrally converging inputs. Electrophysiological recordings in the gustatory Sol support the existence of convergent information from different oral receptor populations (Travers and Norgren, 1995). As the nerve transection data suggest, the rostral Sol is crucial to an animal’s capacity to use afferent gustatory information to guide ingestion and rejection behavior. Extensive bilateral lesions of the gustatory zone of the Sol severely disrupt preference-aversion functions for a variety of gustatory stimuli (Blomquist and Antem, 1967; Flynn et al., 1991a; Shimura et al., 1997). This static sensitivity is not due simply to an inability to modify ingestive behavior because the same Sol-lesioned animals respond normally to increases in the concentration of the trigeminal stimulus capsaicin (Shimura et al., 1997). On the other hand, bilateral lesions centered on taste neurons in the PB can distort or blunt gustatory preference-aversion functions but fail to eliminate them (Flynn et al., 1991a; Spector et al., 1993a). Gustatory thalamic or cortical lesions have no obvious effects (Flynn et al., 1991a; Lasiter et al., 1985; Reilly and Pritchard, 1996a; Scalera et al., 1997). Thus, the thalamocortical connection is not critical for unconditioned responsiveness. After PB lesions, rats often ingest more rather than less of nutritive, normally preferred substances such as sucrose and polycose, particularly during consumption tests. When this effect occurs, it likely reflects a failure to appropriately integrate taste and visceral signals rather than an inability to respond to changes in taste intensity. When the lesions are aimed at the presumably viscerosensory, lateral PB or its major forebrain target, the central nucleus of the amygdala, long-term, but not short-term, intake tests exhibit impairments of concentration-dependent consumption of gustatory stimuli (Galaverna et al., 1993; Reilly and Trifunovic, 2000a; Touzani et al., 1997). The implication is that, during brief access tests, the behavior is driven primarily by taste, which is less affected by the lateral lesions. When significant consumption is involved, lateral PB lesions attenuate or block the attendant visceral afferent feedback and normal behavior is disrupted.

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Conditioned Taste Aversion The avoidance of foodstuff that previously produced illness is referred to as a conditioned taste aversion (CTA). This seemingly simple type of learning requires that the brain, at a minimum, detects the taste stimulus and the internal consequences of ingestion, integrates these signals to form an association, retains the association, retrieves the memory upon subsequent contact with the taste stimulus, and then expresses the learning in behavior. In contrast to the function of the rostral Sol in unconditioned responding, this medullary region is not critical for the behavioral expression of a CTA regardless of whether learning occurs before or after the lesions (Flynn et al., 1991b; Grigson et al., 1997a, 1997b). Moreover, acquisition of a CTA in Sol-lesioned animals is not dependent on olfactory cues (Grigson et al., 1997a). Although some of these lesions involve more than 90% of the rostral Sol, the residual gustatory neurons must be sufficient to make the gross discrimination between the absence and presence of a taste. Lesions of the gustatory PB, however, eliminate CTA (see Norgren and Grigson, 1996; Reilly, 1999; and Spector, 1995, for reviews). The deficit is profound in that learning is blocked both during multiple discrete trials or when LiCl is infused intragastrically whenever the rat ingests the CS over several days (Sclafani et al., 2001). Despite its severity, the deficit is quite specific on several dimensions. It affects acquisition of a CTA (after the lesions), but not the retention of an aversion learned prior to the damage (Grigson et al., 1997b; but see Sakai and Yamamoto, 1998). Although learned odor aversions appear to be blocked, rats with PB lesions can learn to avoid oral trigeminal stimuli, such as capsaicin (Grigson et al., 1998a, 1998b). Several additional tests indicate that the deficit reflects a specific inability to associate gustatory and visceral afferent cues (Reilly et al., 1993; Scalera et al., 1995; Spector et al., 1992). In addition to CTA, lesions centered on the gustatory PB block learned taste preferences in both the discrete or continuous trial paradigms (Sclafani et al., 2001). When the same paradigms use “flavors”, complex stimuli such as Kool Aid, the same lesions at best only attenuate learned preferences or aversions. In contrast to the specificity of the CS deficit after gustatory PB lesions, different USs are equally ineffective (Aguero et al., 1997). When similar lesions are aimed at the lateral PB, the presumed visceral afferent area, acquisition of a CTA, a “flavor” preference, and an oral trigeminal aversion are all blocked (Reilly and Trifunovic, 2000b). With lateral lesions, it is the aversive US that appears to be specific (Aguero et al., 1993; Cubero et al., 2001). That is, rats with lateral PB lesions fail to acquire a CTA

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induced with blood-borne toxins, but are unimpaired when induction occurs via aversive stimuli that seem to depend entirely on an intact vagus nerve. In the context of taste aversion learning, these findings have been interpreted as evidence for independent neural systems subserving visceral processing. In contrast to pontine lesions, similar lesions placed in the thalamic taste area (VPPC) produce no deficit in CTA acquisition (Flynn et al., 1991b; Scalera et al., 1997). Although VPPC lesions are without effect, lesions of gustatory cortex (DI) disrupt postoperative acquisition of a CTA and the retention or retrieval of a preoperatively learned CTA (Bermudez-Rattoni and McGaugh, 1991; Cubero et al., 1999; Dunn and Everitt, 1988). The encoding of the taste-visceral association in the DI relies on multiple neurotransmitters and neuromodulators, as well as protein kinase activity and protein synthesis (Berman et al., 2000; Gutierrez et al., 1999; Naor and Dudai, 1996; Yasoshima and Yamamoto, 1997). Some of these molecular mechanisms apparently do not play a role in the taste–malaise association per se, but rather contribute to the encoding of the novelty of the gustatory stimulus (Berman and Dudai, 2001). Other central nuclei relevant to CTA learning include the amygdala, lateral hypothalamus, and area postrema (Rabin et al., 1983; Ritter et al., 1980; Ruch et al., 1997, 1999; Schafe and Bernstein, 1996; Yamamoto, 1993). In the basolateral and lateral amygdaloid nuclei, protein kinase C activity and various glutamatergic receptor subtypes—AMPA, NMDA, and metabotropic—are involved in CTA acquisition (Yasoshima and Yamamoto, 1997; Yasoshima et al., 2000). Yasoshima and colleagues (2000) further showed that transient blockade of amygdaloid AMPA receptors disrupted the retrieval, but not the retention, of a learned taste aversion. This study is of import because it successfully used transient disruption of neural activity to distinguish between the different stages of the learning process. Similar to gustatory PB lesions, lateral hypothalamic and area postrema lesions made before, but not after, conditioning disrupts behavioral expression of a CTA (Rabin et al., 1984; Ruch et al., 1997, 1999). The involvement of the lateral hypothalamus seems to depend on a D1 dopaminergic mechanism (Caulliez et al., 1996). Overall, lesion-behavior studies demonstrate that thalamocortical projections are not required for the elaboration of a CTA and underscore an important interaction between the gustatory system, specifically the PB and DI, and the ventral forebrain. In fact, during acquisition the interaction appears to be entirely ipsilateral and nonlateralized (Bielavska and Roldan, 1996; Gallo and Bures, 1991). Many of the central structures mentioned above have been identified using the induction of c-Fos-like

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immunoreactivity (c-FLI) as a marker of neuronal activation following an intraperitoneal injection of LiCl, which is by far the most commonly used unconditioned stimulus in CTA. The densest expression of c-FLI following LiCl injection is observed in the area postrema, intermediate and commissural divisions of the Sol, lateral PB, and the central nucleus of the amygdala (Lamprecht and Dudai, 1995; Sakai and Yamamoto, 1997; Yamamoto et al., 1992). Of these areas, the intermediate Sol (SolIM) has received the most attention, because reexposure to a gustatory CS that had been previously paired with LiCl induced significant c-FLI that was not evident in unpaired controls or in animals responding to an innately aversive quinine solution. After CTA extinction, the sapid CS no longer elicited as much c-FLI (Houpt et al., 1994; Swank and Bernstein, 1994). Nevertheless, the functional significance of c-FLI in the Sol is unclear, because the behavioral expression of a CTA can occur without an increase in c-FLI and some c-FLI can occur in animals that fail to demonstrate CTA learning (Schafe and Bernstein, 1998; Spray et al., 2000). Some investigators have suggested that increases in c-FLI in SolIM during expression of a CTA might reflect some as-yet-unidentified conditioned response to the gustatory CS (Cubero et al., 1999; Spray et al., 2000). The neural substrates for CTA have been further complicated by the observation that the method of presenting the CS affects which systems are involved. Conditioned c-FLI is evident in the Sol only when the CS is presented by intraoral infusions (Spray et al., 2000). Similarly, excitotoxic lesions of the amygdala disrupt CTA expression with an intraoral CS, but not when the same CS is ingested from a tube hung on the cage (Schafe et al., 1998). Electrolytic lesions in the same area block CTA using both an intraoral or drinking tube presentation of the CS. Damage to the DI, on the other hand, disrupts CTA regardless of the lesion or conditioning procedure (Cubero et al., 1999). Thus, the picture that emerges from lesion-behavior and c-Fos studies is of a widespread neural network subserving CTA learning in which the critical neural structures differ for different components of the learning process, aversive visceral stimuli, and conditioning procedures.

Salt Appetite Salt appetite is triggered by a negative sodium balance and characterized by the vigorous consumption of previously avoided concentrations of sodium salt. Physiologically, sodium depletion activates the renninangiotensin system thereby stimulating the secretion of the mineralocorticoid aldosterone from the adrenal

glands and, consequently, peripheral Na+ reabsorbtion (Aguilera et al., 1978). In the brain, aldosterone interacts with angiotensin II to induce salt appetite (Fluharty and Epstein, 1983; Fregly and Rowland, 1985; Sakai et al., 1986). Thus, sodium regulation depends on at least the release of angiotensin II and aldosterone for sodium conservation and the apparent change in the behavioral significance of sodium salt. The expression of salt appetite is innate and does not require an animal to first learn an association between the sodium taste signal and the physiological consequence of ingestion. It is also critically dependent on the gustatory afferent information carried by a subset of axons in the chorda tympani (CT) and greater superficial petrosal nerves (GSP), but not the glossopharyngeal nerve (GL) (Bernstein and Hennessy, 1987; Breslin et al., 1993; Markison et al., 1995; Roitman and Bernstein, 1999). Given that the CT and GSP terminate densely in the anterior tip of the rostral Sol, the area that sustains the most complete damage produced by electrolytic lesions, it is not surprising that lesions centered on Sol taste neurons blunt or eliminate salt appetite (Flynn et al., 1991b; Grigson et al., 1997a). Similar lesions of the gustatory PB unequivocally eliminates the expression of salt appetite under a variety of circumstances— electrolytic or ibotenic acid lesions; dietary, diuretic, hormonal, and repeated induction—and appears to be permanent. Nevertheless, if animals experience induction prior to the PB lesions, their subsequent salt appetite is normal (Scalera and Norgren, 1993). A lateral PB serotonergic mechanism exerts an inhibitory influence on diuretic- and hormonal-induced salt appetite (Colombari et al., 1996; Menani et al., 2000). Lesions or blockade of 5HT receptors in lateral PB significantly increase both spontaneous and induced sodium intake. Naïve rats with lesions that include both medial and lateral PB, however, remain unable to express a sodium appetite (Norgren et al., 2001). Despite the profound effects of PB lesions, destruction of taste neurons in either VPPC or DI is without influence on salt appetite (Flynn et al., 1991b; Scalera et al., 1997; Wirsig and Grill, 1982; Wolf et al., 1970). Nevertheless, this taste-guided behavior requires some interaction between the PB and the ventral forebrain because chronically decerebrate rats that have intact Sol and PB, and near normal ingestion and rejection behaviors, fail to exhibit salt appetite (Grill et al., 1986). Lesions of forebrain circumventricular organs, specifically the subfornical organ (SFO) and the organum vasculosum of the lamina terminalis (OVLT), block sodium depletion-induced salt appetite (Fitts et al., 1990; Thunhorst et al., 1990). These two structures specifically mediate the salt appetite-inducing effect of

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circulating angiotensin II (Fitts et al., 2000; Menani et al., 2000). Lesions of the medial amygdala block steroid, but not sodium depletion-induced salt appetite (Schulkin et al., 1989; Zhang et al., 1993). When damage is in the central nucleus, where most of the PBN axons terminate, or the bed nucleus of the stria terminalis (BST), both steroids and sodium depletion fail to induce salt intake (Galaverna et al., 1992; Reilly et al., 1994; Zardetto-Smith et al., 1994). Sodium depletion also fails to evoke a salt appetite following lesions of the lateral hypothalamus (Ruch et al., 1997; Schulkin and Fluharty, 1985). In general, the results from lesion experiments are buttressed by investigations that evaluate the expression of Fos protein. Under conditions that deplete body sodium, increased c-FLI is observed in the SFO, OVLT, area postrema, the medial and central amygdaloid nuclei, medial and lateral nuclei of the BST, lateral PB, and rostral and caudal divisions of the Sol (Johnson et al., 1999; Thunhorst et al., 1998). When the central synthesis of angiotensin II is blocked with high does of captopril, sodium depletion-induced salt appetite and c-FLI in the circumventricular organs is abolished (Thunhorst and Johnson, 1994; Thunhorst et al., 1998). The effect of a high dose of captopril on sodium depletion induced c-FLI in the amygdala and BST remain to be investigated. In the medial amygdala and BST, tachykinin receptors, specifically NK-3 receptors, exert an inhibitory influence on sodium depletion, but not steroid induced salt intake (Massi et al., 1990; Pompei et al., 1991). Taken with the lesion data, it is surprising that the injection of NK-3 agonists into the medial amygdala block salt intake produced by sodium depletion. One interpretation is that activation of a tachykinergic mechanism in the medial amygdala inhibits other brain regions that are crucial for sodium depletion-induced salt appetite. Alternatively, the inhibition might be due to direct spread of the NK-3 receptor agonist into the nearby central nucleus. Nevertheless, these investigations clearly identify specific molecular machinery and structures of the lamina terminalis and hindbrain as being key for mediating the expression of salt appetite and show that the neural circuitry subserving performance in different salt appetite paradigms is dissociable (e.g., steroid vs peptide).

CONCLUSION This chapter concentrates on what Burton and Benjamin (1971) called the “elementary problem of localization of primary taste connections”. We know the location and connections of four central gustatory

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areas, but knowledge of their spatial extent, sensory organization, and neurotransmitters remain imprecise. Not surprisingly, we know most about the first central relay in the nucleus of the solitary tract where electrophysiological studies have mapped neural responses to most gustatory receptor subpopulations, permitted intracellular filling of somata and their dendrites, and at least started characterizing the transmitters involved in gustatory processing. One synapse further rostrally in the pontine parabrachial nuclei, however, the extent and organization of the gustatory neurons remain fuzzy. In the thalamus, the same issues are more acute, because there are less data and, in rodents, none in awake, behaving animals. On cortex there are more data, but they are not always coherent. Fitting function to the central gustatory system also presents apparent clarity, but with fuzzy details. The fact that thalamic gustatory lesions fail to interfere with most taste guided behaviors implies a functional difference between the PB gustatory projections to thalamus and those that reach the ventral forebrain. Lesions that include the thalamic gustatory relay have little or no effect on preference-aversion functions, CTA, or sodium appetite. These lesions do affect behavior, however, if appropriate tests are used. Although they do not block sodium appetite per se, they do eliminate the increased intake of strong salt that typically appears with the second and third induction (Scalera et al., 1997). Similarly, thalamic lesions have no effect on some forms of gustatory contrast behavior, but block other forms (Reilly and Pritchard, 1996b, 1997; Reilly and Trifunovic, 1999). The PB taste neurons that project into the ventral forebrain reach more and larger neural structures, but the specific function of these connections remain a cipher. They appear to be critical for CTA or sodium appetite. For CTA, however, there is consistent evidence that disrupting gustatory cortex interferes with this form of learning, even though damage to the thalamic taste relay does not. Two other anatomical connections might account for this apparent disjunction between form and function. First, some neurons in or near the parabrachial gustatory area have axons that reach the vicinity of taste cortex without a synaptic relay in thalamus. Second, taste cortex or contiguous areas have reciprocal connections with the amygdala, which in turn receives gustatory and visceral afferent information directly from the parabrachial nuclei. This is an example of the anomalies that remain in fitting the functions of the gustatory system to its central anatomy. Even when resolved, some of these issues will remain arcane details. Others will turn out to be essential to understanding the perceptual, physiologic, and hedonic consequences of gustatory stimulation.

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Acknowledgments Dr. Loren Evey assisted in the production of Fig. 5. The authors’s research was supported by USPHS Grants DC00240, DC00369, and MH43787.

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C H A P T E R

29 Olfactory System MICHAEL T. SHIPLEY, MATTHEW ENNIS, and ADAM C. PUCHE Department of Anatomy and Neurobiology, University of Maryland School of Medicine, Baltimore, Maryland, USA

The olfactory system is important for reproductive/ maternal functions, neuroendocrine regulation, emotional responses, aggression, and 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 have long been recognized, yet the neural mechanisms that underlie the perception of “smell” are still poorly understood. Since the previous edition of this volume, significant advances have been made in our understanding of the molecular and cellular physiology of olfaction. Odor molecules are transduced by olfactory receptor neurons (ORNs), first-order neurons located in the olfactory epithelium within the nasal cavity (Fig. 1). ORNs are both the receptors that transduce odors and the first-order neurons that convey information to subsequent neurons in the olfactory network. ORN axons project in the olfactory nerve to synaptically terminate in the main olfactory bulb (MOB). MOB output neurons, mitral and tufted cells, convey olfactory information to higher order olfactory structures and to other brain systems. Transmission from the nose to the mitral and tufted cells is strongly regulated by local intrabulbar circuitry and by centrifugal inputs to the MOB 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,

The Rat Nervous System, Third Edition

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. However, this is not the case. 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, somatosensory, and auditory systems. Foremost among these gaps is our limited understanding 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, recognized by different subfamilies of receptors. How selective are each of these receptors to different molecules? Are some ORNs 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? In recent years, the issue of olfactory codes/specificity has been addressed for only a handful of molecules/receptors. Another major gap in our understanding is how the brain analyses the complex activity patterns elicited by a particular odor (e.g., the odor of coffee contains hundreds of different volatile and nonvolatile components). At the cellular level, we are beginning to understand how neurons in the MOB respond to input and how several of the circuits at the level of the bulb function.

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It remains to be determined how the bulb integrates combinatorial odor mixes and how the output of the bulb is processed in higher cortical regions to elicit the perception of smell. This chapter reviews the neurohistology, connections, and chemical anatomy of central olfactory circuitry. Although great progress has been made in understanding ORNs and the olfactory epithelium, this has been the subject of numerous reviews; hence, extensive discussion of these findings beyond the overview presented below is not as germane to a book that serves as a companion to a stereotaxic atlas as is a consideration of central olfactory structures. Thus, the emphasis of this chapter is placed on what is currently known about the circuit organization of the MOB and higher olfactory centers. Although, the discussion of these higher order connections focuses largely on those structures most directly related to the MOB, as a detailed review of circuitry beyond the primary olfactory cortex would make this chapter unwieldy and is covered in part in other chapters in this volume.

THE OLFACTORY EPITHELIUM Organization The sense of smell is mediated through the stimulation of ORNs by volatile chemicals. ORNs are contained in a neuroepithelial sheet that is located at the top of the nasal vault along the upper portion of the nasal septum, the cribriform plate region, and lining the surfaces of endo and ectoturbinates in rat (Fig. 1), and the medial wall of the superior turbinate in human. In several species a small patch of olfactory neuroepithelium is present on the ventral wall of the septum, the septal organ of Masera, although the function of this region of epithelium is unknown. Afferent information from ORNs is transmitted 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 relative partitioning of the odor between air and mucus thus also determines the stimulating effectiveness of an odor. Odorant-binding proteins present in the mucus may function to bind odorants and present them to receptors. Alternatively,

odorant-binding proteins may be required to remove odorants from the receptors and/or chemically inactivate odorants. The olfactory neuroepithelium is a pseudostratified columnar epithelium (Fig. 1A), which is thicker than the surrounding respiratory epithelium of the nasal cavity. This epithelium rests on a highly 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 and extend into the nasal cavity. ORNs are true sensory neurons with both a dendrite and an axon (Cajal, 1911). Their cell bodies lie in the basal twothirds of the epithelium, with apical dendrites that extend to the epithelium surface. The peripheral tip of this dendrite swells slightly to form the olfactory knob, from which six to eight cilia extend into the mucous layer (Menco, 1984; Menco and Farbman, 1985a; Menco and Farbman, 1985b). 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. 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 specialized Schwann cells, the ensheathing cell (DeLorenzo, 1957). These bundles combine to make up the 15 to 20 larger axon fascicles (fila olfactoria) of the olfactory nerve, which pass through the cribriform plate to synapse in the MOB. 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 that project into the mucous layer. These cells appear to regulate serous elements of the mucus composition (Getchell and Getchell, 1992; Getchell et al., 1984) and express molecules of the P450 enzyme systems (Hadley and Dahl, 1982), which suggests a role in detoxification. A third cell type, the microvillar cells, present in humans, rodent, amphibia, and fish at about one-tenth the number of the ORNs, have microvilli on their apical surface, projecting into the mucous layer (Moran et al., 1982; Morrison and Costanzo, 1990; Muller and Marc, 1984; Rowley et al., 1989). Their basal end tapers into a cytoplasmic extension that enters the lamina propria. Retrograde tracing from the olfactory bulb in rat indicates that this extension can project to the olfactory bulb (Rowley et al., 1989). The neuronal ultrastructural appearance of the microvillar cells together with a projection to the olfactory bulb suggests that these cells are a class of bipolar neurons. Deep to the ORNs, sustentacular, and microvillar cells are the globose and horizontal basal cells, which sit on

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FIGURE 1 (A) Scale diagram of the rat nasal cavity and brain shown in a lateral view and cross section diagrams through the nasal cavity at the location of the arrows. The nasal passages of the rat nose contain a series of intricate scroll-like protrusions of bone, collectively termed turbinates. In rat, there are four turbinates, referred to as endoturbinates I to IV, visible on the lateral wall of the nasal cavity with several additional respiratory turbinates in the rostral nose. When these endoturbinates are removed a second set of turbinates, referred to as ectoturbinates 1 to 4, is exposed. Endoturbinate II divides into endoturbinate II (or IIa) and II´ (or IIb). The main olfactory neuroepithelium lines the endo- and ectoturbinates, the roof of the nasal cavity (dorsal recess, DR), and the caudal half of the nasal septum (green outline). The main sensory epithelium is divided into four zones on the basis of the expression of different odorant receptor genes. Dashed lines show these approximate positions of these zones in the adult. Two other smaller regions of sensory neuroepithelium are present within the nasal cavities. The septal organ of Masera is a small oval patch of olfactory neuroepithelium located on the ventral septum rostral to the main olfactory neuroepithelium. Most mammalian species also have a long blind-ended tubular cavity lined by sensory neuroepithelium, called the vomeronasal organ, which is encased by the vomeronasal bone at the base of the septum. (B) Schematic illustration of the olfactory epithelium showing the major cell types present in the rat epithelium; olfactory receptor neuron (ORN), sustentacular cell, microvillar cell, basal cell (globose and horizontal), ensheathing cell, and Bowman’s gland cells.

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a basement membrane just above the lamina propria (Cuschieri and Bannister, 1975a; Cuschieri and Bannister, 1975b; Graziadei and Graziadei, 1979). These 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.

Odorant Recognition and Signal Transduction Electrical signals are generated in ORNs when odorants traverse the mucus layer and bind to specific membrane receptors located on the ORN cilia. Classically, the existence of odorant receptors (OR) as discrete, specific molecules was largely a theoretical construct. However, in 1991, Buck and Axel discovered a large multigene family of odorant receptor genes (ORG). The ORG family consists of approximately 1000 genes that are expressed by ORNs. Our current understanding is that each mammalian ORN expresses only a single unique ORG (the group of ORNs expressing the same ORG can be referred to as an ORN–ORG cohort), although recent intriguing evidence suggests that a subset of ORNs are capable of expressing two separate ORGs (Rawson et al., 2000). ORNs expressing the same odorant receptor gene are widely dispersed in one of four broad expression zones within the epithelium (Fig. 1) (Buck and Axel, 1991; Ressler et al., 1993; Strotmann et al., 1994; Sullivan et al., 1995; Vassar et al., 1993). Although, there are also examples of ORN–ORG cohorts clustering in discrete patches rather than zones (Pyrski et al., 2001; Strotmann et al., 1992; Strotmann et al., 1994). These results indicate that there could be additional patterning for subsets of ORN–ORGs beyond that of the four main expression zones. In the decade since the cloning and characterization of the ORG family our understanding of the molecular biology of olfactory transduction has expanded rapidly. These odorant receptors are members of the seventransmembrane receptor gene superfamily that also includes a family of receptors highly homologous to ORGs isolated from testis (Vanderhaeghen et al., 1993; Vanderhaeghen et al., 1997; Walensky et al., 1998), metabotropic glutamate receptors (mGluR), transforming growth factor receptors, and many other large gene families. In the mammalian genome there are approximately 1000 separate genes encoding the ORG family. It is remarkable that of the 30–35,000 genes present in the mammalian genome fully 1000 genes (~l3% of the genome) code odorant receptors! In humans, approximately two-thirds of these ORGs have become pseudogenes; genes that have undergone evolutionary mutations that insert stop codons into the reading

frame of the gene, rendering them nonfunctional. It is hypothesized that heterogeneity in the human population and the high level of ORG pseudogenes may underlie some instances of specific anosmia. Despite recent advances in the genome project identifying ORG sequences, there are still many unanswered questions about the molecular specificity of odorant receptors. Several efforts have been made to generate three-dimensional models to investigate how putative ligands might “fit” into putative binding pockets on ORGs (Floriano et al., 2000; Sharon et al., 1998; Singer, 2000). A more convincing demonstration of receptor specificity has been provided by heterologous ORG expression (Breer et al., 1998; Krautwurst et al., 1998; Sengupta et al., 1996; Touhara et al., 1999) coupled with measures of physiological responses. The best-characterized ORG to date is the I7 receptor, which has a high affinity for the odorant octanol with weaker responses to related molecules (Krautwurst et al., 1998). The current model for OR specificity is that each OR is “tuned” to recognize a particular chemical moiety that may be present on a number of different odor molecules. The tuning of ORN responses appears to be relatively broad. Receptors responding strongly to one moiety can also respond to other moieties when the odorant concentration is increased. It is clear from these initial studies that Herculean effort may be required to determine receptor-ligand specificity for all 1000 ORGs. Parallel progress has also been made in understanding the transduction events that intervene between the binding of odorants to ORs and the generation of action potentials in ORNs (reviewed in Schild and Restrepo, 1998; Gold, 1999; Zufall and Munger, 2001). Several lines of evidence suggest that a “cAMP” signal transduction pathway predominates in mammalian ORNs. Binding of an odorant to the OR leads to activation of the G-protein Golf which activates an adenylate cyclase type III (ACIII) molecule, leading to a rise in cyclic nucleotides, the opening of olfactory cyclic nucleotide-gated channels (CNGA2/OCNC1), and the entry of calcium into the cilia. The rise in intracellular calcium triggers Na+ and Cl− conductances leading to the generation of action potentials that are propagated down the axon to the MOB. Genetic null mutations for Golf (Belluscio et al., 1998), CNGA2/OCNC1 (Brunet et al., 1996), and ACIII (Wong et al., 2000) firmly establish the essential role for these molecules. Mice null mutant for any of these three transduction elements are functionally anosmic. In other species, notably invertebrate, an IP3 pathway is present (Schild and Restrepo, 1998). In this cascade odorant binding to the OR triggers interactions with G-proteins, which then activates phospholipase C (PLC) to generate a rise in phosphatidal inositol

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(IP3). The IP3 second messenger also opens channels leading to an influx of calcium into the cilia and triggering action potentials within the ORN. However, in mammals the essential requirement of ACIII for olfactory function (Wong et al., 2000) suggests that the IP3 transduction pathway is not a primary cascade. It remains to be tested whether mammalian IP3 transduction is a modulator of the cAMP pathway or mediates inhibitory odor responses that have been reported in ORNs (Morales et al., 1994). ORNs and axons also express high levels of olfactory marker protein (OMP), which is unique to ORNs (Keller and Margolis, 1975; Margolis, 1972). OMP is found in a number of mammalian species, including humans, and appears to be expressed in all ORNs, accounting for l1% of the total protein content of these neurons. The role OMP plays in the function of ORNs is poorly understood; however, recent evidence from mice containing a null mutation for OMP suggests that this protein may play a role in ORN adaptation to odors (Ivic et al., 2000). However, this exciting progress in understanding the fundamental molecular interactions at the level of the olfactory receptor will not bring a complete understanding of the sense of smell 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 first element in a complex neural network involving the MOB and higher cortical areas that leads to the perception of odors. The operations of central olfactory networks and the anatomical organization of the olfactory system appear in some respects to be fundamentally different from the familiar topographically organized circuits of the other major sensory systems. The remainder of this chapter concentrates on the anatomical and neurochemical organization of these brain regions.

The Primary Olfactory Projection In general, any single locus on the surface of the MOB receives innervation from ORNs dispersed widely within the olfactory neuroepithelium (Fig. 2A), and conversely ORNs projecting to an MOB locus are interspersed with other neurons projecting to a different locus. (Astic et al., 1987; Astic and Saucier, 1986; Clancy et al., 1994; Saucier and Astic, 1986; Schoenfeld et al., 1994). However, there is a general tendency for neurons located in the medial, lateral, dorsal, and ventral olfactory neuroepithelium to project to homologous bulbar surfaces (Astic and Saucier, 1986; Saucier and Astic, 1986). A detailed analysis of the primary projection in hamster suggests that patterning in the olfactory neuroepithelium occurs as a series of mediolateral,

dorsoventral areas irrespective of the presence of turbinates (Clancy et al., 1994; Schoenfeld et al., 1994). These epithelium “domains” broadly coincide with the four expression domains for ORGs (for review, see Mori et al., 1999). Subpopulations of ORNs have also been identified by expression of cytoplasmic and membrane associated antigens (Barber, 1989; Carr et al., 1994; Key and Akeson, 1993; Key and Giorgi, 1986; Mori, 1987; Puche and Key, 1996; Riddle et al., 1993; Schwarting et al., 1992a; Schwarting et al., 1992b; Schwarting and Crandall, 1991). Most of these subpopulations of ORN axons “sort” and terminate across wide areas of the MOB in patterns conserved between animals. In contrast to this diffuse topographical organization there is a much more precise receptotopic organization between the MOE and the main olfactory bulb: ORNs expressing the same, single ORG appear to project to the same point on the surface of the MOB (Fig. 2) (Mombaerts et al., 1996; Ressler et al., 1993; Vassar et al., 1994; Wang et al., 1998). In situ hybridization for odorant receptor mRNA suggested that the axons containing mRNA for a specific receptor all terminate within one or a few glomeruli (Ressler et al., 1993; Vassar et al., 1994). More recently, transgenic mice in which ORNs expressing a particular ORG coordinately express β-galactosidase or green fluorescent protein were constructed (Mombaerts et al., 1996; Wang et al., 1998). In these mice ORNs express the same ORG and project to one or two (or a few) glomeruli. Thus, it is hypothesized that specific ORN–ORG cohorts innervate each glomerulus in the MOB and that each glomerulus receives homogenous input from ORNs expressing the same ORG. Thus, glomeruli represent a spatial map of the activity of ORNs to odor stimuli, a conclusion that had been suggested by a variety of experimental approaches for several decades. These functional studies involving 2-deoxyglucose (Astic and Saucier, 1981; Benson et al., 1985; Johnson et al., 1998; Johnson and Leon, 2000), c-fos (Guthrie and Gall, 1995; Onoda, 1992), functional magnetic resonance imaging (Yang et al., 1998), calcium imaging (Wachowiak et al., 2000; Wachowiak and Cohen, 1999), and intrinsic imaging (Rubin and Katz, 1999; Rubin and Katz, 2001; Uchida et al., 2000) all suggested that glomeruli represent functional units. The neural computational problem in the identification of odors thus becomes not how the brain recognizes odors but how the brain recognizes patterns of glomerular activity elicited by different odors.

THE MAIN OLFACTORY BULB Since the brain’s representation of odors reduces to a neural computational problem, knowledge of the

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FIGURE 2 Diagram showing the mosaic projection of ORNs in the (A) main olfactory system and (B) accessory olfactory system. (A) Each ORN has a single axon that passes into the lamina propria underlying the olfactory neuroepithelium (OE), through the cribriform plate (CP), and then into the olfactory bulb. The olfactory bulb is a rostral outcropping of the telencephalon with a highly laminar arrangement shown in more detail in the next figure. ORNs enter the ventral and rostral surfaces of the olfactory bulb and course through the nerve fiber layer (ONL) to their synaptic targets in the glomerular layer (GL). These axons synapse on the dendrites of second-order neurons within globular structures of neuropil called glomeruli. ORNs expressing different odorant receptor genes (shown as purple, green, or blue cells) are interspersed randomly within each zone. Most, if not all, axons from ORNs expressing the same odorant receptor gene converge and project to a defined point on the surface of the olfactory bulb (represented as purple, green, or blue shaded glomeruli). (B) The vomeronasal epithelium can be divided into an apical zone and a basal zone. VRNs in the apical zone express V1R and V3R odorant receptor genes and the G-protein GαI and project to the anterior half of the AOB. VRNs in the basal zone express V2R receptor genes and the G-protein Gαo and project to the posterior half of the AOB. Unlike the main olfactory bulb, ORNs in the vomeronasal organ that express the same ORG project to multiple glomeruli in the AOB. AOB mitral cells also have multiple apical dendrites.

anatomy and physiology of central olfactory circuits is essential to understand olfactory coding. Olfactory circuit organization and development have features in common with other neural systems, especially the cerebral cortex. Improved knowledge of 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 may provide fundamental insights about the underlying computational features that have driven the selective expansion of cortical structures in the evolution of the mammalian brain.

Laminar Organization The MOB is an allocortex that, like other cortical structures, has a characteristic laminar organization

(Figs. 3 and 6). The layers of the MOB and their principal cell types are discussed next. Table 1 lists the neuron types present in the main olfactory bulb, Table 2 summarizes candidate neurotransmitters, and Table 3 summarizes neurotransmitter receptors in the bulb. Olfactory Nerve Layer (ONL) The superficial-most layer of the MOB is the olfactory nerve layer. The ONL contains thin unmyelinated axons from the ORNs and glial cells (Cajal, 1911; Pinching and Powell, 1971b). These glial cells may derive from both the peripheral and the central nervous system. Throughout the ONL, a specialized glial cell, termed the “ensheathing cell” (Doucette, 1989), provides an incomplete wrapping that separates ORN axons and the glomeruli from the periglomerular region (Pinching and Powell, 1971b). The deepest third of the

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FIGURE 3 Schematic representation of neuronal circuitry in the adult rat main olfactory bulb. Olfactory receptor neuron axons course through the olfactory nerve fiber layer (ONL)

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and terminate in the glomerular layer (GL) on dendrites from mitral cells (MC) and tufted cells (external, eTC; middle, mTC; and deep, dTC). These contacts occur in globular neuropil structures termed glomeruli. Glial and neuronal cells surround and delineate each glomerulus. The neurons surrounding these glomeruli form lateral contacts between glomeruli and consist of the periglomerular cells (PGC), short axon cells (SA), and external tufted cells (eTC). 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). The lateral dendrites of mitral and tufted cells form reciprocal synapses with the apical dendrites of granule cells within the external plexiform layer (EPL, lower inset). Mitral and tufted cells send their axons along the lateral olfactory tract (Lot) to the olfactory cortex. Collateral branches of these axons also form within the bulb itself, and a population of tufted cells project solely within the same bulb, the intrabulbar association system (IAS). Granule cells (GC) in the granule cell layer (GCL) form dendrodendritic contacts with mitral and tufted cells in the external plexiform layer. However, some granule cells possess dendrites that do not reach the external plexiform layer, while other granule cell dendrites project toward the center of the bulb. The majority of granule cells are concentrated in the granule cell layer but a few granule cells lie within the mitral cell layer (MCL), superficial to the granule cell layer. A thin neuropil layer termed the internal plexiform layer (IPL) containing the axons of the mitral and tufted cells is located between the mitral and granule cell layers. The olfactory bulb receives both ipsilateral and contralateral central afferent fibers (CFF) from various brain regions that innervate most cell types and layers of the olfactory bulb. The bulb also contains other populations of neurons in various layers with poorly understood function; these are the van Gehuchten cells (vC), horizontal cells (hC), Cajal cells (cC), Golgi cells (gC), and Blanes cells (bC).

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TABLE 1

Neuron Types in the Main Olfactory Bulb

Layer

Neuron type

Size

Density

GL

Periglomerular

5–8 μm

High

Short axon

8–12 μm

Low

External tufted

10–15 μm

Moderate

Middle tufted

15–18 μm

Low

Deep tufted

15–18 μm

Low

Van Gehuchten

12–17 μm

Low

Mitral cell

20–25 μm

High

Granule cell

10–16 μm

Moderate

Cajal cell

Medium

Low

Horizontal cell

Medium

Low

Granule cell

10–16 μm

Very high

Blanes cell

16–23 μm

Low

Golgi cell

12–22 μm

Low

Migratory cell

Small

Low

EPL

GL IPL GCL

SEL

ONL has been found to contain astrocytes (Bailey and Shipley, 1993). These astrocytes express QPRT, the degradative enzyme for quinolinic acid, which is an agonist of the NMDA receptor. Quinolinic acid, like glutamate, may modulate glutamatergic transmission between the ORN axon terminals and the postsynaptic targets. Glomerular Layer (GL) Immediately deep to the ONL is the glomerular layer. 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. The glomeruli are generally ovoid and range from 80 to 160 μm in diameter in rat. Most estimates of glomerular number are similar, around 2000–3000 glomeruli/bulb for rabbits (Allison, 1949) and approximately 1800–2000 in mouse (Allison, 1953; Royet et al., 1988; White, 1972). The number of glomeruli in rats has been estimated at 3000 (Meisami and Safari, 1981). However, recent reestimations using distribution-free stereological methods suggest that there could be as many as 6300 glomeruli in rabbit and 4200 in rat (Royet et al., 1998). It has also been estimated that there are several million ORNs in total projecting to the olfactory glomeruli. Thus there is convergence of several thousand ORN axons in each of the 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. Astrocytes appear to cordon off adjacent glomeruli strengthening the long-standing notion that each glomerulus is a discrete functional unit. The neurons of the glomerular shell are referred to as juxtaglomerular neurons. The majority of the juxtaglomerular neurons can be classified as one of three types—(i) small, periglomerular cells; (ii) slightly larger external tufted cells; and (iii) short axon cells (Pinching and Powell, 1971a). The Golgi studies of Blanes (Blanes, 1898), Golgi (Golgi, 1875), and Cajal (Cajal, 1911), and the Golgi and EM studies of Pinching and Powell (Pinching and Powell, 1971c), provide the classical descriptions of the distribution of juxtaglomerular cell dendrites and axons. The dendrites of the small (5–8 μm) 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 that are the chief morphological feature that distinguishes them from small external tufted cells. The shortaxon cells are somewhat larger than the periglomerular cells (8–12 μm) and are distinguished in Golgi material because their dendrites are confined largely to the periglomerular shell. The axons of periglomerular/ short axon cells usually course along the periphery of up to two to four glomeruli. The external tufted cells, which are 10–15 μm in the long axis, lie in the periglomerular region and usually have a single apical dendrite that arborizes within a single glomerulus, although some external tufted cells have two or three apical dendrites capable of ramifying in several different glomeruli. The apical branching of the external tufted cell fills a larger portion of a glomerulus than does the apical tufts of other classes of tufted cells that are described below or mitral cells (unpublished observations). External tufted cell apical dendrites are varicose but only rarely form spines. In addition to their thick apical dendrite(s), many external tufted cells also have thinner secondary, or lateral, dendrites that extend in the external plexiform layer immediately subjacent to the glomerular layer, although these are less numerous than the lateral dendrites of mitral cells. Some external tufted cells have axons that project to nearby glomeruli; these axons apparently do not enter glomeruli but, rather, terminate between glomeruli, whereas other external tufted cells project an axon out of the olfactory bulb. The presence of an apical tuft, lateral dendrites, and basal axon on external tufted cells is broadly

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TABLE 2 Transmitter

Candidate Transmitters of Main Olfactory Bulb Cells

Cell type/location

Cell type

Frequency

Species

References

Aspartate

Small mitral

Not reported

Few

Rat

(1–3)

CCK

Juxtaglomerular

Not reported

Few

Rat

(4–6)

External tufted

Not reported

Many

Rat

(4–6)

Middle tufted

Not reported

Many

Rat

(4–6)

Deep tufted

Not reported

Few

Rat

(4–6)

Cajal cell

Not reported

Few

Rat

(4–6)

CRF

Tufted cells

Not reported

Few

Rat, Monkey

(7, 8)

Mitral cells

Not reported

Many

Rat, Monkey

(7, 8)

DA

Juxtaglomerular

8–13 μm

Many

Many

(9–15)

ENK

GL—periglomerular

5–8 μm

Many

Hamster, guinea pig, rat

(16–18)

GCL—granule cells

8–16 μm

Few

Hamster, guinea pig, rat

(16–18) (19), and others

GL—periglomerular

5–8 μm

Many

Many

GCL—granule cells

8–16 μm

Many

Many

(19), and others

GABA and DA

GL—periglomerular

5–8 μm

Many

Rat

(20, 21)

GABA and parvalbumin

EPL—External tufted

Small

Few

Rat

(22)

GABA

GABA and ENK

GCL—granule cells

8–16 μm

Rare

Rat

(22)

GABA, SP, and DA

GL—periglomerular

5–8 μm

Few

Hamster

(17)

NAG

MCL—mitral cells

Not reported

Many

Rat

(23, 24)

NADPH diaphorase

GL—short axon

Not reported

Few

Rat

(25, 26)

GL—periglomerular

14–32 μm

Few

Rat, human

(27, 28)

GCL

14–32 μm

Few

Rat, cat, human, marmoset

(27–29)

Somatostatin

GL—juxtaglomerular

Not reported

Few

Rat

(5, 25)

SP

GL—juxtaglomerular

12–17 μm

Many

Hamster, not mouse, rat, cat, guinea pig, rabbit

(10, 17, 30)

EPL—external tufted

12–17 μm

Many

Hamster, not mouse, rat, cat, guinea pig, rabbit

(10, 17, 30)

NPY

TRH

GL—periglomerular

5–8 μm

Many

Rat

(31, 32)

VIP

GL—periglomerular

<10 μm

Few

Cat

(28, 29)

EPL—External tufted

l12 μm

Few

Rat, cat

(28, 29)

EPL—Van Gehuchten

l12 μm

Few

Rat, cat

(28, 29)

References. 1, Fuller and Price (1988); 2, Watanabe and Kawana (1984); 3, Halasz (1987); 4, Seroogy et al. (1985); 5, Matsutani et al. (1988); 6, Bonnemann et al. (1989); 7, Imaki et al. (1989); 8, Bassett et al. (1992); 9, Baker et al. (1988); 10, Baker (1986); 11, Baker et al. (1984); 12, Baker et al. (1983); 13, Gall et al. (1987); 14, Halasz et al. (1981); 15, McLean and Shipley (1988); 16, Bogan et al. (1982); 17, Davis et al. (1982); 18, Kosaka et al. (1995); 19, Mugnaini et al. (1984); 20, Kosaka et al. (1988); 21, Kosaka et al. (1985); 22, Kosaka et al. (1987); 23, Blakely et al. (1987); 24, Ffrench-Mullen et al. (1985); 25, Scott et al. (1987); 26, Davis (1991); 27, Ohm et al. (1988); 28, Gall et al. (1986); 29, Sanides-Kohlrausch and Wahle (1990); 30, Kream et al. (1984); 31, Merchenthaler et al. (1988); 32, Tsuruo et al. (1988).

similarity to the dendritic arrangement of superficial, middle, and deep tufted cells and to that of the mitral cells (to be discussed below). All of these tufted cell types are often grouped with the mitral cell when the output cells of the bulb are discussed; however, there are considerable structural and physiological differences between these classes of cells that suggest different roles in olfactory information processing. 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 (1971b) determined that ORN axons synaptically contact some 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 electron microscopic (EM) sections revealed that the mitral/ tufted dendrites make reciprocal synaptic contacts with the dendrites and gemmules (spine-like processes) of periglomerular cells (Toida et al., 1998; Toida et al., 2000). These specialized reciprocal synapses are often closely associated with each other (Pinching and Powell, 1971b).

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TABLE 3

Transmitter Receptors in the MOB

Receptor

GL

EPL

MCL

IPL

GCL

Cholinergic mAChR1 mAChR2 mAChR3 mAChR4 nAChR

— ++ — — ++

++ ++ ++ ++ ++

— ++ — — ++

++ ++ ++ ++ —

++ ++ — — ++

Noradrenergic α1 α2 β1 β2

— ++ ++ ++

++ ++ — —

++ ++ — —

— — — ++

++ ++ ++ ++

— ++ (ONL) ++ (GL)

— —

— —

— —

— ++

Serotonergic 5-HT1A 5-HT1C 5-HT2A/C

++ — ++

++ — ++

++ — ++

++ — NR

++ ++ NR

Glutamatergic Kainate NMDA AMPA Metabotropic

— ++ ++ ++

++ ++ ++ ++

NR ++ ++ ++

++ ++ ++ ++

— ++ — ++

GABAergic GABAA GABAB

++ ++

++ —

++ ++

— NR

++ ++

Dopaminergic D1 D2

References (1–7)

(1, 8–12)

(13–15)

(13, 16–20)

(21–44)

(3, 30, 31, 45–48) (11, 49–51)

Key. ++, receptors present; —, receptors absent; NR, not reported. References. 1, Booze et al. (1989); 2, Fonseca et al. (1991); 3, Laurie et al. (1992); 4, Rotter et al. (1979); 5, Sahin et al. (1992); 6, Spencer et al. (1986); 7, Le Jeune et al. (1996); 8, Nicholas et al. (1993); 9, Ressler et al. (1994); 10, Wanaka et al. (1989); 11, Young and Kuhar (1980); 12, Woo and Leon (1995); 13, Nickell et al. (1991); 14, Mansour et al. (1990); 15, Koster et al. (1999); 16, McLean et al. (1995); 17, Pompeiano et al. (1992); 18, Pompeiano et al. (1994); 19, Whitaker-Azmitia et al. (1993); 20, Wright et al. (1995); 21, Gall et al. (1990); 22, Martin et al. (1992); 23, Martin et al. (1993); 24, Miller et al. (1990); 25, Molnar et al. (1993); 26, Monaghan and Cotman (1982); 27, Monyer et al. (1994); 28, Ohishi et al. (1993); 29, Petralia and Wenthold (1992); 30, Petralia et al. (1994); 31, Petralia et al. (1994); 32, Ohishi et al. (1993); 33, Shigemoto et al. (1992); 34, Shigemoto et al. (1993); 35, Tanabe et al. (1992); 36, Chen et al. (1996); 37, Watanabe et al. (1993); 38, Wisden and Seeburg (1993); 39, Romano et al. (1995); 40, Kinoshita et al. (1998); 41, Kinzie et al. (1995); 42, Giustetto et al. (1997); 43, Sassoe-Pognetto and Ottersen (2000); 44, Sun et al. (2000); 45, Bowery et al. (1987); 46, Chu et al. (1990); 47, Fritschy et al. (1992); 48, Persohn et al. (1992); 49, Richards et al. (1987); 50, Margeta-Mitrovic et al. (1999); 51, Bonino et al. (1999).

Glomeruli have classically been viewed as relatively homogenous structures consisting of axons, dendrites, synapses, and glial processes. However, recent studies indicate that glomeruli exhibit a degree of subcompartmentalization that is manifested as a segregation of different types of synaptic contacts (Chao et al., 1997; Halasz and Greer, 1993; Kasowski et al., 1999; Toida et al., 2000). ORN axons entering the glomerulus synapse upon target dendrites in subcompartments or islands within the glomerular neuropil, whereas dendrodendritic synapses occur in bundles of 4–100 dendrites that are segregated from the ORN axonal islands by the processes of glia (Chao et al., 1997; Kasowski et al., 1999). 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

portrayed by Golgi and EM studies as neurons with different transmitters may differentially synapse with mitral/tufted cells, olfactory axons, and/or other juxtaglomerular neurons. Indeed, Kosaka and colleagues examination of the synaptic organization of juxtaglomerular neurons expressing tyrosine hydroxylase— dopaminergic neurons—indicates that 80% of the synaptic input to this class of neurons comes from ORN axons, 10% from dendrodendritic contacts with mitral/tufted cells, and the remaining 10% from GABAergic interneurons (Toida et al., 2000). The “synaptology” of dopaminergic neurons is in contrast to juxtaglomerular neurons expressing calbindin D-28K, which receive almost no synapses from ORN axons but instead form dendrodendritic contacts with presumptive mitral/tufted cell dendrites (Toida et al., 1998). It is clear from these studies that populations of juxtaglomerular

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neurons can differ dramatically in their synaptic organization and role in the processing of olfactory input. Transmitters/Peptides of Neurons in the Glomerular Layer ORN axons are a major neural element in the glomerulus. ORN axons utilize glutamate as their primary neurotransmitter which acts on glutamate receptors located on the dendrites of the MOB target cells. Fast synaptic responses in mitral and tufted cells are mediated through AMPA/kainate while slow synaptic responses are mediated via NMDA receptors (Aroniadou-Anderjaska et al., 1997; Ennis et al., 1996). AMPA/kainate and NMDA receptors also appear to mediate fast and slow synaptic responses in juxtaglomerular cells in response to ORN input (Ennis et al., 2001; Keller et al., 1998). Metabotropic glutamate receptors are also present within the glomerular layer, but their role(s) are presently unclear. Many juxtaglomerular cells are dopaminergic (Halasz et al., 1981) or GABAergic (Ribak et al., 1977). In the hamster, about 70% of the dopamine (DA) neurons are reported to colocalize GABA while about 45% of GABAergic cells contain DA (Kosaka et al., 1985). In the rat, virtually all DA periglomerular neurons also contain GABA, whereas DA-expressing tufted cells do not express GABA (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 DA based on the presence of immunocytochemical markers for these transmitters in the same cells (Davis et al., 1982; Kosaka et al., 1988), although juxtaglomerular neurons in the rat do not appear to contain substance P. Thus, some juxtaglomerular cells may contain both a catecholamine (DA) and 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, 1990b). Some periglomerular cells with short axons that project to the deeper granule cell layer contain NADPH diaphorase, neuropeptide Y (NPY), and somatostatin, which Scott and collaborators (1987) suggested may provide a direct route for periglomerular cells to influence granule cells. Davis (1991) concluded that NADPH in the glomerular layer is primarily, if not exclusively, contained in periglomerular cells. Also reportedly present in some juxtaglomerular cells are cholecystokinin (Matsutani et al., 1988; Seroogy et al., 1985), aspartic acid (Fuller and Price, 1988; Halasz, 1987; Watanabe and Kawana, 1984), thyrotropin-releasing hormone (Merchenthaler et al., 1988; Tsuruo et al., 1988), enkephalin (Bogan et al., 1982; Davis et al., 1982; Kosaka

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et al., 1995), and protein kinase C (Saito et al., 1988). In addition, a subpopulation of small juxtaglomerular neurons is positive for acetylcholinesterase; thus, 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 and were identified over a decade ago, little is known of the functional significance of these transmitters/peptides for olfactory signal processing. Because most juxtaglomerular cells are very small (5–12 μm) and because most of them have widely varying local circuit connections and neurochemical content, their physiological characteristics have been difficult to study. As noted above, the glomerular layer contains a large population of dopamine-containing neurons. In the MOB, the predominant DA receptor is of the D2 subtype (Coronas et al., 1997; Guthrie et al., 1991; Mansour et al., 1990a; Nickell et al., 1991; Palacios et al., 1981a). This receptor is localized in the glomerular layer and equally densely in the ONL (Guthrie et al., 1991; Nickell et al., 1991). If ZnSO4 is used to destroy the olfactory epithelium, D2 receptor binding is eliminated in both the olfactory nerve and the glomerular layers. D2 mRNA transcripts are also abundantly expressed by ORNs (Koster et al., 1999). Thus, while it has been presumed for two decades that olfactory nerve terminals do not receive presynaptic contacts, it was hypothesized that juxtaglomerular dendrites release DA, which acts on olfactory terminals but without a classic synaptic specialization. Recently, electrophysiology studies and direct optical imaging of calcium transients in ORN axons demonstrate that DA, acting at the D2 receptor subtype, presynaptically inhibits ORN axons (Berkowicz and Trombley, 2000; Ennis et al., 2001; Wachowiak and Cohen, 1999). Intriguingly, GABA, acting at the GABAB receptor subtype, also exerts a presynaptic inhibitory regulation of ORN terminals in the MOB. GABAB receptors are present at high levels in the glomeruli (Bonino et al., 1999; Margeta-Mitrovic et al., 1999). Recent electrophysiological results demonstrate that GABA released from juxtaglomerular neurons acts on GABAB receptors to presynaptically inhibit ORN terminals (Aroniadou-Anderjaska et al., 2000). External Plexiform Layer (EPL) Immediately deep to the glomeruli is the external plexiform layer, a layer with a relatively low cell density but a very dense neuropil. The predominant neural elements in this layer are the dendrites of mitral/ tufted and granule cells. The principal neuron types in

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EPL are the superficial, middle, and deep tufted cells, named according to their relative depth in EPL, the external tufted cells at the EPL/GL boundary discussed above, and the Van Gehuchten cells, which appear to be local interneurons. There is a gradual increase of tufted cell size from the superficial to the deep parts of the EPL (Pinching and Powell, 1971a; Shipley et al., 1995). 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 miter). The dendritic morphologies of tufted cells vary but usually they have at least one apical dendrite that enters and ramifies within a single glomerulus as do mitral cells. Some tufted cells can have two or three apical dendrites that enter different glomeruli. Tufted cells also have secondary dendrites that run tangentially in the EPL (Shepherd, 1972). These secondary dendrites are thought to form reciprocal synapses with the apical dendrites of granule cells as do mitral cells (Shepherd, 1972). Glutamate released from the secondary dendrites of tufted cells is thought to excite granule cells primarily via AMPA/ kainate and NMDA receptor (Christie et al., 2001). Middle and deep tufted cells have similar, but not identical, axonal projections to the mitral cell (Schoenfeld et al., 1985; Schoenfeld and Macrides, 1984); thus, most tufted and all mitral cells may functionally be considered as the output cells of the olfactory bulb. The axons of many superficial tufted cells project mainly to other sites in the same (ipsilateral) 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 MOB to the anterior olfactory nucleus and other rostral olfactory cortical structures (Schoenfeld et al., 1985; Scott, 1986); few tufted cell axons project into rostral cortical regions. The intrabulbar collaterals of a population of superficial tufted cells form 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 dense tracts that travel within this layer to the opposite side of the same bulb where they terminate as a dense terminal field in the IPL. These tufted cells thus send a discrete, topographically organized projection to the opposite side of the same bulb, a point-to-point reciprocal projection between the lateral and medial bulb. The IAS has the highest degree of point-to-point topographical organization of any known circuit in the olfactory system. Most of the ORN–ORG cohorts studied to date project to one glomerulus on the lateral bulb and a second on the medial bulb (Mombaerts et al., 1996; Ressler et al., 1993; Vassar et al., 1994; Wang et al., 1998). It is appealing

to hypothesize that the IAS could interconnect bulbar regions with the same ORN–ORG axons. The IAS is formed exclusively by cholecystokinin (CCK)-containing tufted cells (Liu and Shipley, 1994) and the terminals of these CCKergic cells terminate preferentially, if not exclusively, onto the apical dendrites of the granule cells (Liu and Shipley, 1994). CCK causes membrane depolarization in all neurons studied to date. Thus, it is likely that when CCKergic tufted cells are active, they cause depolarization of granule cells on the opposite side of the bulb. This may either increase or decrease the release of GABA depending on whether the depolarization invades the dendritic release sites or acts as to shunt currents that would normally cause GABA release. Given the highly topographic organization of the IAS, this may lead either to highly focal inhibition or to excitation on the opposite side of the bulb. A second group 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 in cat (Sanides-Kohlrausch and Wahle, 1990a), and in rat for a variety of calcium-binding proteins (calbindin D-28, calretinin, neurocalcin, and parvalbumin) and NADPH diaphorase (Alonso et al., 1993; Brinon et al., 1999). Tufted cells appear to utilize glutamate as their principle transmitter (Christie et al., 2001). As noted, large populations of tufted cells also contain the neuropeptide cholecystokinin (Seroogy et al., 1985). Using in situ hybridization substance P mRNA transcripts have been detected in some external tufted cells and in up to half of the mitral cells in MOB of rat (Warden and Young, 1988), but to date no studies using immunocytochemistry have detected substance P in mitral cells of any species (Baker, 1986; Inagaki et al., 1982; Shults 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 (Gall et al., 1986), but in the cat, this peptide appears to be in Van Gehuchten cells and not tufted cells (SanidesKohlrausch and Wahle, 1990b). In the hamster, a few middle tufted cells are NADPH diaphorase positive (Davis, 1991). Mitral Cell Layer (MCL) Deep to the external plexiform layer is the mitral cell layer. This is a thin layer that contains the somata of mitral cells (25–35 μm in diameter in rat) arranged in almost a monolayer. These cells are the principal

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output cells of the bulb and, with some minor species variation (cf. Scott, 1986), have one apical dendrite that enters a single glomerulus, where it branches extensively and is synaptically contacted by olfactory axons (Shepherd, 1972). The apical dendrites of approximately 25 mitral cells enter each glomerulus in rat (Cajal, 1911). 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 (unpublished observations). The secondary dendrites of mitral cells may extend up to 2 mm in the EPL and are oriented tangentially, i.e., parallel to the surface of the bulb. These secondary dendrites engage in dendrodendritic synapses with dendrites of granule cells. In addition, they may receive centrifugal and Van Gehuchten cell inputs (Jackowski et al., 1978). It should be noted that while there are approximately 40,000 mitral cells in rat (Meisami, 1989) there are approximately 100,000 granule cells in the mitral cell layer (Frazier and Brunjes, 1988). Thus, while it is referred to as the mitral cell layer, mitral cells make up only approximately 35% of the cells in that layer. Mitral cells are glutamatergic, but have also been hypothesized to utilize N-acetyl-aspartyl-glutamate (NAG) (Blakely et al., 1987; Ffrench-Mullen et al., 1985), which has been observed in mitral cells by immunocytochemistry (Blakely et al., 1987). However, subsequent neurophysiological studies cast doubt on a primary transmitter role for NAG (Whittemore and Koerner, 1989) in mitral cells. Also, a few unusually small, mitral cells appear to contain aspartate and project to the piriform cortex (Fuller and Price, 1988). The neuropeptide, corticotropin-releasing factor (CRF), has been demonstrated in mitral and some tufted cells using both immunocytochemistry and in situ hybridization in the rat (Imaki et al., 1989). CRF fibers were also observed in layer Ia of the piriform cortex. This finding is consistent with CRF being a releasable neuropeptide in mitral cells since mitral cells synaptically terminate in the layer Ia 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 (Imaki et al., 1989). Internal Plexiform Layer (IPL) Immediately subjacent to the mitral cell layer is the internal plexiform layer, a thin layer with many axons and dendrites but a low density of cells. The IPL contains the axons of mitral/tufted cells, dendrites of granule cells, and axons from other centrifugal sources. Some axons in IPL originate from raphe nuclei (5HT; McLean and Shipley, 1987b), the locus coeruleus (NE; McLean et al., 1989), and the nuclei of the diagonal band (ACh;

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Shipley and Adamek, 1984) (see section on “Nucleus of the Diagonal Band” below). As noted earlier, the IPL also contains a very dense plexus of CCK-containing axons and terminals (Liu and Shipley, 1994), which derive from superficial tufted cells. The IPL also contains a few 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 cholinergic inputs from the nucleus of the diagonal band, they may be cholinoceptive. Granule Cell Layer (GCL) The granule cell layer is the deepest neuronal layer in the bulb. It contains many small (8–10 μm in long axis) granule cell neurons. Frequently, three to five 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 are also found mixed with mitral cells in the mitral cell layer. Golgi studies indicate that granule cells lack axons. They have a limited basal dendrite arbor that ramifies in the GCL and a thicker and longer apical dendrite that enters and ramifies extensively in the EPL (Price and Powell, 1970a; Price and Powell, 1970b). Generally, the more superficial the granule cell body is located in the GCL, the more superficially located are its apical dendrite ramifications in the EPL, while granule cells deeper in the GCL have dendrites that ramify in the deeper parts of the EPL (Mori et al., 1983; Orona et al., 1983; Scott, 1986). However, not all granule cells follow this pattern. The exceptions include granule cells whose distal 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 nongranule cells with both dendritic and axonal arbors in the granule cell layer, including the Blanes cell and Golgi cell. There are extensive synapses between the dendrites of the granule cells and the secondary dendrites of mitral/ tufted cells. Mitral/tufted cell dendrites make excitatory glutamatergic synapses onto the dendrites of granule cells and the dendrites of the granule cells make inhibitory synapses onto the dendrites of mitral/tufted cells. The mitral-to-granule cell dendrodendritic excitation that triggers GABA release from granule cells appears to require the NMDA receptor (Chen et al., 2000; Isaacson and Strowbridge, 1998; Schoppa et al., 1998). Most of the granule cells contain GABA (Ribak et al., 1977), which via GABAA receptors inhibits the mitral and tufted cells (Chen et al., 2000; Jahr and Nicoll, 1982; Schoppa et al., 1998; Shepherd, 1972). Some peptides have also been localized in other cells in the granule

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cell layer. 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. Subependymal Layer The deepest layer in the MOB 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 MOB cells derive from this zone (Altman, 1969; Kishi, 1987; Shimada, 1966), although in the adult other MOB interneurons (primarily granule cells with only a few juxtaglomerular cells) are generated in more rostral subventricular forebrain regions then migrate into the bulb along what has been termed “the rostral migratory stream” (Lois and Alvarez-Buylla, 1994; Luskin, 1993; Weiss et al., 1996). The Olfactory Nerve Regulates Transmitters in Some Bulb Neurons Input from the olfactory epithelium is important for the normal expression of transmitters in many types of MOB neurons. For example, the olfactory nerve influences the expression of the DA phenotype in juxtaglomerular neurons. If the olfactory epithelium is destroyed by detergent or 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. This phenomenon of transneuronal regulation of transmitter phenotype has been demonstrated in adult rats (Baker et al., 1983, 1984; Kawano and Margolis, 1982), mice (Baker et al., 1983; Nadi et al., 1981), dogs (Nadi et al., 1981), hamster (Kream et al., 1984), and developing rats (McLean and Shipley, 1988). TH expression is also downregulated in mice homozygous for a null mutation in CNGA2.OCNC1, which renders the mice functionally anosmic (Baker et al., 1999). Thus, these cells appear to require the 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, GABA continues to be expressed in juxtaglomerular neurons in which TH is downregulated following deafferentation (Stone et al., 1990). The mechanism underlying the regulation of TH has been the focus of recent studies. In vitro, depolarizing stimuli induction of TH expression is dependent upon calcium influx into the bulb neurons (Cigola et al., 1998). In an olfactory bulb neuron/olfactory epithelium coculture, transneuronal odor-dependent

regulation of TH expression could be abolished by blockade of glutamatergic signaling through the NMDA receptor (Puche and Shipley, 1999). This suggests a mechanism by which glutamate release by ORNs stimulates dopaminergic neurons leading to the influx of calcium and expression regulation of TH. Epithelial input is also required in order to trigger the initial developmental expression of TH (Baker and Farbman, 1993; McLean and Shipley, 1988). Thus, both the developmental induction and maintenance of the DA phenotype depends on the presence and normal function of the olfactory nerve. Deafferentation does not influence the expression of GABA in periglomerular neurons indicating that odor-evoked activity selectively regulates the expression of different transmitters. DA is not the only transmitter that is 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 (unpublished observations). The influence of the olfactory nerve on MOB cells has been studied in other ways. Laing and colleagues (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. Light and dark granule cells have also been described (Struble and Walters, 1982) and these two subpopulations differ in their reaction to unilateral nares occlusion (Frazier and Brunjes, 1988; Skeen et al., 1985): 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 are recent arrivals in the GCL from the rostral migratory stream. 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 (Frazier and Brunjes, 1988; Kaplan et al., 1985) since the granule cells continue to be born in the proliferative subependymal zone (Frazier and Brunjes, 1988).

Outputs of the Main Olfactory Bulb 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 approximately the level of the AOB. These caudally directed axons give off collaterals in the anterior olfactory

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nucleus (AON) and other regions of olfactory cortex (Figs. 3 and 6). Tufted cells collateralize to an even greater extent in the bulb than mitral cells. The intrabulbar association pathway formed by CCKergic tufted cells was discussed earlier. Mitral/Tufted Cell Axons The primary output of the 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 the MOB project to the dorsal part of the external subdivision of the AON while output cells of the ventral half of the MOB project to the lateral subdivision (Schoenfeld and Macrides, 1984). Output neurons of the MOB that project to more caudal regions (e.g., piriform cortex) are evenly distributed in the MOB. Intracellular injections of horseradish peroxidase (HRP) into mitral cells have been used to show the arborizations of individual axons. The axons often have many collateral terminal arbors in the 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 the AON and piriform cortex in rabbit (Ojima et al., 1984), which is reminiscent of the patchy distribution of thalamic input to the visual cortex. Some mitral cells branch to project to both the 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 a high degree of point-to-point topographical projection to their target structures, as is characteristic of other sensory systems. Projections to Olfactory Cortex Mitral and tufted cells of the MOB project to several structures of the ipsilateral hemisphere, including the superficial plexiform layer of the anterior olfactory nucleus, the piriform, periamygdaloid, and lateral entorhinal cortices, the taenia tecta, the anterior hippocampal continuation, the indusium griseum, and the olfactory tubercle (Figs. 4, 5, and 6). Collectively, the regions directly innervated by the output of the MOB have been referred to as the 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 direct MOB projection to the supraoptic nucleus has also been reported (Smithson et al., 1989). The organization of these projections is discussed below in the section on the primary olfactory cortex.

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Centrifugal Afferents to the MOB Centrifugal afferent inputs to the MOB are very dense, arise from multiple sources (Fig. 4), and play important roles in regulating neural processing in the MOB. 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) modulatory afferents from nonolfactory subcortical structures. These two groups are distinguished because the olfactoryrelated projections mediate specific olfactory sensory and association functions whereas the modulatory afferents have widespread projections that influence CNS functions across other neural systems. 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 brain stem and basal forebrain. Olfactory-related centrifugal afferents to the MOB are discussed below in page 948. The organization of subcortical “modulatory” inputs to the bulb is discussed below in section on “Nonolfactory Modulatory inputs to the Olfaction System.”

PRIMARY OLFACTORY CORTEX The MOB projects to a collection of structures referred to collectively as the primary olfactory cortex (de Olmos et al., 1978). These structures may be usefully divided into three groups: (A) the anterior olfactory nucleus; (B) the medial olfactory cortex comprising the indusium griseum, anterior hippocampal continuation, taenia tecta, infralimbic cortex, and olfactory tubercle; and (C) the lateral olfactory cortex comprising, from rostral to caudal, piriform, periamygdaloid, transitional, and entorhinal cortices.

Anterior Olfactory Nucleus (AON) Architecture of AON Interposed between the MOB and the piriform cortex is a distinctive structure, the anterior olfactory nucleus (Figs. 5 and 6). Herrick (1924), who first defined the AON, 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 for the central region (Haberly and Price, 1978b). Many of the output cells in the different subdivisions of the AON are pyramidal, save for neurons in the external subdivision (Haberly and Price, 1978b). Based on these architectonic features, AON is considered to be a cortical structure. The AON has been divided into

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FIGURE 4 Diagrammatic representation of the connections of the main (MOB) and accessory (AOB) olfactory bulb with cortical (gray panels) and subcortical structures (circles). The outputs of the olfactory bulbs along the lateral olfactory tract are shown at the upper (MOB) and lower (AOB) part of the diagram. Centrifugal inputs from the cortical regions back to the olfactory bulb are shown centrally. Abbreviations: ACo, anterior cortical amygdaloid nucleus; AON, anterior olfactory nucleus; AONm, anterior olfactory nucleus, medial division; BAOT, bed nucleus of the lateral olfactory tract; BST, bed nucleus of the stria terminalis; DB, nucleus of the diagonal band; DHR, dorsal hippocampal rudiment; DP, dorsal peduncular cortex; DR, dorsal raphe nucleus; Ent, entorhinal cortex; LC, locus coeruleus; LPO, lateral preoptic area; Me, medial amygdaloid nucleus; MnR, median raphe; NLOT, nucleus of the lateral olfactory tract; PaCo, periamygdaloid cortex; Pir, piriform cortex; TT, taenia tecta; Tu, olfactory tubercle.

several subdivisions based on cytoarchitecture and connections (de Olmos et al., 1978; Haberly and Price, 1978b). In this chapter we do not review the cytoarchitectural criteria used to distinguish subdivisions of the AON but they are illustrated in Fig. 6; readers seeking further detail can consult previous publications (de Olmos et al., 1978; Haberly and Price, 1978b; Shipley et al., 1996; Shipley and Adamek, 1984). Inputs to AON As noted above, the AON is heavily targeted by mitral and tufted cells. In addition, inputs to the AON arise from other subdivisions of the ipsilateral AON and from the contralateral AON (de Carlos et al., 1989; Luskin and Price, 1983), from the piriform cortex and the entorhinal cortex (Luskin and Price, 1983), from the

anterior amygdaloid area (de Carlos et al., 1989; Luskin and Price, 1983), from the posterolateral cortical amygdala nucleus (de Carlos et al., 1989; Luskin and Price, 1983), from the olfactory tubercle (Luskin and Price, 1983), from the nucleus of the lateral olfactory tract (de Carlos et al., 1989), and from the temporal portion of CA1 division of the hippocampus (van Groen and Wyss, 1990). “Modulatory” inputs arise from the raphe nuclei (McLean and Shipley, 1987a), the locus coeruleus (McLean et al., 1989), and the nucleus of the diagonal band (de Carlos et al., 1989). Outputs of AON All subdivisions of the ipsilateral AON project to both the ipsilateral and the contralateral MOB except the external division (AON pars externa, AONe), which

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FIGURE 5 (A–E) Connections of the main olfactory bulb in rostral to caudal coronal sections of the brain showing both anterograde and retrograde transport of wheat germ agglutinin–horse radish peroxidase following injection into the main olfactory bulb. Abbreviations: AHC, anterior hippocampal continuation; AONd, anterior olfactory nucleus, dorsal division; AONe anterior olfactory nucleus, pars externa; AONl, anterior olfactory nucleus, lateral division; Entl, entorhinal cortex, lateral division; Entm, entorhinal cortex, medial division; HDB, horizontal limb of the diagonal band; lot, lateral olfactory tract; NLOT, nucleus of the lateral olfactory tract; Pir, piriform cortex; TTvs, taenia tecta, ventralsuperior; Tu, olfactory tubercle; VDB, ventral limb of the diagonal band.

projects only to the contralateral MOB (de Olmos et al., 1978). The AON contains the largest number of neurons projecting to the bulb from any one source (Carson, 1984). Thus, there is extensive, bilateral representation of olfactory information at the level of the AON. Based on lesion experiments, Kucharski and Hall (1987) showed that the AON can access and recall existing olfactory memories stored in the contralateral AON or MOB. There is some degree of laminar topography of the terminal projections to the MOB from different AON subdivisions in rat (Luskin and Price, 1983; Reyher et al., 1988) and hamster (Davis and Macrides, 1981). In

both species, the ventral and posterior subdivisions of the 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 the 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 in 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 (1983) described additional outputs of the AON including projections to the ventral taenia tecta, the

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FIGURE 6 Cytoarchitecture (Nissl stain) of the olfactory bulb and primary olfactory cortices. (A) Cross-section of the main olfactory bulb at low magnification. (B) Higher magnification photograph of the lamina organization of the main olfactory bulb. (C–N) Cross-sections through the rat brain at the different rostral to caudal levels of the primary olfactory cortex and olfactory-related cortical structures. Abbreviations: ac, anterior commissure; Acb, nucleus accumbens; ACo, anterior cortical amygdaloid nucleus; AI, anterior insular cortex; AONd, anterior olfactory nucleus, dorsal division; AONe, anterior olfactory nucleus, pars externa; AONl, anterior olfactory nucleus, lateral division; AONm, anterior olfactory nucleus, medial division; AONvp, anterior olfactory nucleus, ventroposterior division; APir, amygdalopiriform transition area; BL, basolateral amygdaloid nucleus; BLP, basolateral amygdaloid nucleus, posterior part; BM, basomedial amygdaloid nucleus; BMP, basomedial amygdaloid nucleus, posterior part; CA1, CA1 region of the hippocampus; CA3, CA3 region of the hippocampus; Ce, central amygdaloid nucleus; CG, central (periaqueductal) gray; CPu, caudate–putamen (striatum); DP, dorsal peduncular cortex; En, endopiriform nucleus; Ent, entorhinal cortex; Entl, entorhinal cortex, lateral division; Entm, entorhinal cortex, medial division; EPL, external plexiform layer; EPLA, external plexiform layer of the accessory olfactory bulb; GCL, granule cell layer; GCLA, granule cell layer of the accessory olfactory bulb; GL, glomerular layer; GLA, glomerular layer of the accessory olfactory bulb; HDB, horizontal limb of the diagonal band; ICj, islands of Calleja; IL, infralimbic cortex; IPL, internal plexiform layer; IPLA, internal plexiform layer of the accessory olfactory bulb; La, lateral amygdaloid nucleus; LH, lateral hippocampal area; lot, lateral olfactory tract; MCL, mitral cell layer; MCLA, mitral cell layer of the accessory olfactory bulb; MePD, medial amygdaloid nucleus, posterodorsal part; MePV, medial amygdaloid nucleus, posteroventral part.

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FIGURE 6, cont’d Abbreviations: NLOT, nucleus of the lateral olfactory tract; ONL, olfactory nerve layer; ONLA, olfactory nerve layer of the accessory olfactory bulb; opt, optic tract; PA, posterior amygdaloid nucleus; Pir, piriform cortex; PLCo, posterolateral cortical amygdaloid nucleus; PMCo, posteromedial cortical amygdaloid nucleus; PrS, presubiculum; RF, rhinal fissure; S, subiculum; SEL, subependymal layer; SO, supraoptic nucleus; St, stria terminalis; Tr, transitional area; TTd, taenia tecta, dorsal; TTis, taenia tecta, inferior-superior; TTvi, taenia tecta, ventral-inferior; TTvs, taenia tecta, ventral-superior; Tu, olfactory tubercle; VP, ventral pallidum. Scale bar in A and C–N = 1 mm.

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piriform cortex and olfactory tubercle, the endopiriform cortex, the ventral agranular insular cortex, and the nucleus accumbens (from the ventroposterior portion of the 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.

direct MOB inputs and, in turn, projects to the dentate gyrus of the hippocampus, it has been suggested that the 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).

Transmitters of AON

Anterior Hippocampal Continuation

Candidate transmitters in the AON and other olfactory cortical areas are summarized in Table 4 and neurotransmitter receptors in Table 5. Aspartate has been proposed as a transmitter of AON neurons (mainly the dorsal and external divisions of the AON) based on selective retrograde transport of [3H]aspartate (Fuller and Price, 1988; Watanabe and Kawana, 1984). Fewer neurons in other subdivisions of the 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 the AON and some of these appear to project to the MOB (Davis et al., 1982). It appears that all neurons in the AONe contain the neuropeptide CRF (Bassett et al., 1992).

The anterior hippocampal continuation (AHC) has been described in detail elsewhere (Adamek et al., 1984; Wyss and Sripanidkulchai, 1983). It is located just ventral to the rostrum of the corpus callosum and dorsal to the taenia tecta. The inputs to the AHC are similar to those of the IG as are its efferent connections with the outstanding difference that IG does not project to the MOB while there is a modest projection from the AHC to the MOB (de Olmos et al., 1978; Scheibel and Scheibel, 1978; Wyss and Sripanidkulchai, 1983). Other major efferent projections of the IG and the AHC are to the mammillary bodies and the anterior thalamic nuclei.

Medial Olfactory Cortex Several cortical areas located on the medial wall of the rostral hemisphere compose part of the olfactory pathways 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, and the taenia tecta (Fig. 6). 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. Indusium Griseum The indusium griseum (IG) or dorsal hippocampal continuation receives input from, but does not project to, the MOB. It is a thin layer of cortex that runs parasagittally just dorsal to the corpus callosum. The IG has been the subject of debate as to whether it is more related to the hippocampus or the MOB (cf. Adamek et al., 1984; Smith, 1897; Wyss and Sripanidkulchai, 1983, for further discussion). It now seems clear that the IG receives direct inputs to its tiny molecular layer from the MOB (Adamek et al., 1984; de Olmos et al., 1978; Wyss and Sripanidkulchai, 1983). This input is mainly to the rostral IG with fewer fibers running more caudally. The molecular layer of the IG also receives input from the lateral and medial entorhinal cortex (Luskin and Price, 1983). Since the entorhinal area receives

Taenia Tecta The taenia tecta proper projects strongly to the MOB (de Olmos et al., 1978; Shipley and Adamek, 1984). The pyramidal neurons of this cortical structure are relatively tightly packed and are located dorsal to the anterior olfactory nuclei on the medial wall of the hemisphere anterior to the rostrum of the corpus callosum. Infralimbic Cortex This cortical region is found slightly rostral to but in the same general region as the AHC. Although most studies indicated a lack of direct connections between the infralimbic cortex and the MOB (de Carlos et al., 1989; de Olmos et al., 1978; Shipley and Adamek, 1984), at least one study has reported a weak projection (Neafsey et al., 1986). The cells in the infralimbic cortex that are said 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 the rostral piriform cortex and, possibly, the endopiriform cortex (Neafsey et al., 1986). Because of its extensive connections with autonomic brain centers infralimbic cortex may be a linkage between the olfaction and the autonomic function. Olfactory Tubercle The olfactory tubercle in rodents, rabbits, and other macrosmatic mammals is a prominent bulge on the base of the hemisphere just caudal to the olfactory peduncle. In such species, axons of mitral and tufted cells (de Olmos et al., 1978; Heimer, 1968; Price, 1973) terminate

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TABLE 4 Neuropeptide Calbindin

CCK

Cell location

Neuropeptides in Olfactory Cortical Structures Cell type

Frequency

Species

References

AON—anterior, medial, lateral

Bi/multipolar

4–8%

Rat, human

(1, 2)

TT

Bi/multipolar

4–8%

Rat

(3)

Pir—mainly layer II, none in layer I

Bi/multipolar

4–8%

Rat

(3)

Ent

Bi/multipolar

4–8%

Rat

(3)

AON—anterior, dorsal, lateral, medial

No info

Rare

Rat

(4)

Pir—mainly layer II

Pyramidal, multipolar

12–17/section

Rat

(5)

Ent—layers II and III

No info

No info

Rat

(6)

CRF

AON

Small

Medium

Rat, monkey

(7, 8)

Dyn B

TT

Nonpyramidal fusiform, multipolar

Scattered

Rat

(9)

AON—medial, ventral, posterior

Nonpyramidal, fusiform, multipolar

Scattered

Rat

(9)

OT

Nonpyramidal, fusiform, multipolar

Scattered

Rat

(9)

Pir—layers II and III

Nonpyramidal, fusiform, multipolar

Scattered

Rat

(9)

Ent—lateral

Nonpyramidal, fusiform, multipolar

Scattered

Rat

(9)

ENK

AON

Fusiform, pyramidal, multipolar

Few

Rat

(10)

Pir—mainly layer II

Fusiform, pyramidal, multipolar

Few

Rat

(10)

TT

Fusiform, pyramidal, multipolar

Few

Rat

(10)

OT

Fusiform, pyramidal, multipolar

Few

Rat

(10)

GABA/GAD

All olfactory regions

Small

Varying density

Many

(11, 12)

LHRH

OT

10–15 μm

Scattered

Rat, others

(13)

Medial olfactory tract

10–15 μm

Scattered

Rat, others

(13)

Pir

No info

Decrease with age

Rat

(14)

Neurotensin NPY

Parvalbumin

SP VIP

Ent

No info

Decrease with age

Rat

(14)

AON—all divisions

15–25 μm

1–5/section

Rat

(15, 16)

OT

15–25 μm

10/section

Rat

(15, 16)

Islands of Calleja

>25 μm

10/section

Rat, Cat

(15–17)

AON—complementary to calbindin

Multipolar

Dense

Rat

(3)

Pir—mainly layer II

Multipolar

Dense

Rat

(3)

AON—all divisions

Small/medium

Many

Rat

(18)

OT

Small/medium

Few

Rat, Cat

(18–20)

Pir—mainly layers II and III

Bipolar (12μm)

Scattered

Rat, mouse, cat

(21, 22)

Ent—mainly layers II and III

Bipolar (12μm)

Scattered

Rat, mouse, cat

(21, 22)

References. 1, Garcia-Segura et al. (1984); 2, Ohm et al. (1991); 3, Celio (1990); 4, Vanderhaeghen et al. (1985); 5, Westenbroek et al. (1987); 6, Greenwood et al. (1981); 7, Bassett et al. (1992); 8, Sakanaka et al. (1987); 9, Fallon and Leslie (1986); 10, Harlan et al. (1987); 11, Mugnaini et al. (1984); 12, Haberly et al. (1987); 13, Zheng et al. (1988); 14, Hara et al. (1982); 15, de Quidt and Emson (1986); 16, de Quidt and Emson (1986); 17, Sanides-Kohlrausch and Wahle (1990); 18, Warden and Young, (1988); 19, Ljungdahl et al. (1978); 20, Ljungdahl et al. (1978); 21, Roberts et al. (1982); 22, Sanides-Kohlrausch and Wahle (1990).

in the superficial layer of the tubercle as in the AON and primary olfactory cortex. The tubercle has a superficial plexiform layer like the 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 the

primary olfactory cortex but rather are polymorphic, and their dendrites do not appear to preferentially extend into the plexiform layer as do those of the pyramidal cells of layer III in the olfactory cortex. These polymorphic neurons appear to be more akin to neurons of the striatum and indeed 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).

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TABLE 5

Receptor Subtypes of the AON

Receptor

AON

Cholinergic mAChR1 mAChR2 mAChR3 mAChR4 nAChR

++ ++ ++ ++ ++

Noradrenergic α1 α2 β1 β2

++ ++ ++ ++

Dopaminergic D1 D2

++ —

Serotonergic 5-HT1A 5-HT1C 5-HT2A/C

++ ++ ++

Glutamatergic Kainate NMDA AMPA Metabotropic

++ ++ ++ ++

GABAergic GABAA GABAB

++ ++

References (1–4)

(5–9; 9; 10)

(11)

(12–14)

(15–26)

(27–31)

Key. ++, receptors present; —, receptors absent; NR, not reported. References. 1, Fonseca et al. (1991); 2, Hunt and Schmidt (1978); 3, Rotter et al. (1979); 4, Spencer et al. (1986); 5, Nicholas et al. (1993); 6, Palacios and Kuhar (1980); 7, Wanaka et al. (1989); 8, Day et al. (1997); 9, Rosin et al. (1996); 10, Woo and Leon (1995); 11, Mansour et al. (1990); 12, McLean et al. (1995); 13, Pompeiano et al. (1992); 14, Wright et al. (1995); 15, Gall et al. (1990); 16, Martin et al. (1993); 17, Monaghan et al. (1985); 18, Ohishi et al. (1993); 19, Shigemoto et al. (1993); 20, Petralia and Wenthold (1992); 21, Petralia et al. (1994); 22, Petralia et al. (1994); 23, Shigemoto et al. (1992); 24, Romano et al. (1995); 25, Kinoshita et al. (1998); 26, Kinzie et al. (1995); 27, Bowery et al. (1987); 28, Palacios et al. (1981); 29, Richards et al. (1987); 30, Zhang et al. (1991); 31, Margeta-Mitrovic et al. (1999).

Lateral Olfactory Cortex Architecture of 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 to the lateral entorhinal cortex (Fig. 6). Collectively, these cortical structures compose the entire temporal cortical mantle ventral to the rhinal sulcus. Haberly and Price (1978a) divided the piriform cortex into three layers that are further subdivided on the basis of cytoarchitecture and afferent connections. Layer I, the superficial plexiform layer, is divided into Ia and Ib which receive different afferents: layer Ia receives afferents from the ipsilateral MOB while layer Ib receives association fibers from the AON and from

other parts of the primary olfactory cortex. Layer II, the superficial compact cell layer is also divided into two sublayers; the more superficial zone has lower cell density and the deeper division has a higher cell density. Layer III is the widest cell layer. Deep to the piriform cortex is the endopiriform nucleus that was recognized by Loo (1931) in the opossum and was further described by Haberly and Price (1978a). Behan and Haberly (1999) reported that the endopiriform nucleus, while similar in many regards to the piriform cortex proper, has unique intrinsic and extrinsic connections that differ from the piriform cortex. In the piriform cortex, the pyramidal cells in layers IIb and III have extensive basal dendrites while some smaller cells in layer IIa lack basal dendrites (Haberly and Price, 1978a) and are reminiscent of dentate granule cells in the hippocampus. Reconstructions of individually intracellularly filled pyramidal cells in the piriform cortex show that pyramidal cells can have extensive axonal projections covering almost an entire cerebral hemisphere, including local connections to anterior and posterior piriform cortex as well as arborizations in orbital cortex, insular cortex, olfactory tubercle, perirhinal cortex, entorhinal cortex, and amygdaloid cortex (Johnson et al., 2000). Many of these cortical areas are implicated in behavior, cognition, emotion, and memory. Piriform cortex also contains numerous interneurons that are distributed throughout all layers and regions of this structure in both opossum and rat (Ekstrand et al., 2001; Haberly et al., 1987; Haberly and Presto, 1986). Some interneurons are found in layer I, where they may function in feedforward inhibitory systems (Ekstrand et al., 2001). Many of these GABAergic interneurons, described as “basket cells,” colocalize with calcium-binding proteins (parvalbumin, calbindin), VIP, or CCK (Ekstrand et al., 2001; Kubota and Jones, 1993). These basket interneurons are primarily distributed in layers II/III and exhibit diverse molecular markers and morphological characteristics. These cells are thought to predominantly form axosomatic or proximal axodendritic synapses with neurons in layers II and III and participate both in feedback and feedforward inhibitory circuits. Several informationprocessing models have been proposed for the function of piriform cortex, but it is beyond the scope of this chapter to present all the models and evidence (for review, Haberly, 2001). As noted by de Olmos and colleagues (1978), there are several transitional regions (including periamygdaloid cortex) between the piriform cortex and the entorhinal cortex and between olfactory cortices and the 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

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29. OLFACTORY SYSTEM

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. The MOB sends a projection to the entire extent of the 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 the AON and piriform cortex while the projection to the more caudal parts of olfactory cortex becomes progressively dominated by mitral cells (Schoenfeld and Macrides, 1984). Candidate transmitters in the piriform cortex are summarized in Table 6.

TABLE 6

Receptor Subtypes of the Piriform Cortex

Receptor

Piriform

Cholinergic mAChR1 mAChR2 mAChR3 mAChR4 nAChR

++(I,II) ++ ++ ++(II) ++(II,III)

Noradrenergic α1 α2 β1 β2

++(II) ++(II/III) ++(II) ++(II)

Dopaminergic D1 D2

++(II) ++

Serotonergic 5-HT1A 5-HT1C 5-HT2A/C

++(II) ++(II) ++/++(II)

Glutamatergic Kainate NMDA AMPA

++(II) ++(Ia,II) ++(II)

Metabotropic GABAergic GABAA GABAB

References (1,2)

(3–10)

(11–14)

945

Connections of Olfactory Cortex Feedback to the Olfactory Bulb The piriform cortex, the lateral entorhinal cortex, and the transitional cortical areas in between all project heavily back to the MOB (Figs. 4 and 6). The projections are heavier from the rostral than from the caudal parts of the 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 MOB arise mainly from pyramidal neurons in layer II and to a lesser extent layer III in the primary olfactory cortex. The transmitters of these olfactory cortical projections to the bulb are not known, although glutamate is suspected. These feedback projections from the olfactory cortex to the MOB are believed to primarily excite the GABAergic granule cells in the MOB which in turn inhibit firing of mitral cells (Nicoll, 1971) via dendrodendritic synapses between granule and mitral cell dendrites (Halasz and Shepherd, 1983). In the hamster, neurons in the rostral to caudal levels of the piriform cortex terminate from superficial to deep in the GCL of the 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 projections from the lateral olfactory cortex to the MOB terminate most heavily in the ipsilateral GCL. Intrinsic and Association Connections

(15–21)

In addition to the feedback projections to the MOB, the olfactory cortex has other extensive connections which can be discussed as four classes: intrinsic or local—short connections between neurons in different layers of the POC; associative—connections with different parts of the POC; extrinsic—connections with other structures; and modulatory inputs—afferents that terminate in the POC as part of a broader innervation of other cortical and subcortical neural systems.

(3,9,20,22–33)

NR (1,7,34,35) ++(I,III) ++

Key. ++, receptors present; —, receptors absent; NR, not reported; I, II, III, piriform cortex layer. References. 1, Bowery et al. (1987); 2, Seguela et al. (1993); 3, Nicholas et al. (1993); 4, Palacios and Kuhar (1980); 5, Unnerstall et al. (1984); 6, Wanaka et al. (1989); 7, Young and Kuhar (1980); 8, Day et al. (1997); 9, Rosin et al. (1996); 10, Jones et al. (1985); 11, Fremeau et al. (1991); 12, Huang et al. (1992); 13, Mansour et al. (1990); 14, Mansour et al. (1990); 15, Hoffman and Mezey (1989); 16, McLean and Darby-King (1994); 17, Mengod et al. (1990); 18, Mengod et al. (1990); 19, Pompeiano et al. (1992); 20, Pompeiano et al. (1994); 21, Wright et al. (1995); 22, Gall et al. (1990); 23, Monaghan (1985); 24, Petralia and Wenthold (1992); 25, Shigemoto et al. (1992); 26, Wisden and Seeburg (1993); 27, Romano et al. (1995); 28, Ohishi et al. (1993); 29, Ohishi et al. (1993); 30, Wada et al. (1998); 31, Kinoshita et al. (1998); 32, Kinzie et al. (1995); 33, Sun et al. (2000); 34, Palacios et al. (1981); 35, Margeta-Mitrovic et al. (1999).

Intrinsic or local connections The POC has two principal layers of pyramidal neurons, layers II and III, which comprise several morphological classes and also several classes of nonpyramidal neurons. There are extensive translaminar or local connections among POC neurons. Layer II neurons give off axon collaterals to deeper layer III pyramidal cells and there are local inhibitory interneurons in layers I and II that are contacted by MOB terminals and by local collaterals of pyramidal cells. Deeper pyramidal cells also give rise to extensive local collaterals that may synapse with

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local interneurons or with more superficial pyramidal cells. Thus, there are extensive translaminar connections both from superficial to deeper layers and vice versa. As noted, there are several classes of GABAergic and neuropeptide-containing neurons in the POC which have the appearance of local interneurons (Ekstrand et al., 2001; Haberly et al., 1987; Haberly and Presto, 1986). GABAergic neurons play an important role in regulating olfactory cortical functions through feedback and feedforward inhibition (Ekstrand et al., 2001). Studies by Haberly’s group indicate that intrinsic GABAergic cells may regulate long-term potentiation (LTP) and the expression of epilepsy; the piriform cortex (Pir) is one of the most seizure-prone structures in the brain. GABAergic neurons are scattered throughout layers II and III of the Pir 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 the Pir. These cells may be excited directly by MOB inputs and possibly by association projections (see below) and function to regulate olfactory cortical excitability by feedforward inhibition. Association connections Corticocortical projections within the POC are extensive and exhibit laminar and regional organization. Axons from pyramidal cells of layer IIb are primarily directed at more caudal sites in the POC; cells in layer III project predominantly to rostral parts of the POC. Commissural fibers to the contralateral POC arise from layer IIb of the anterior parts of the POC. The ipsilateral and commissural association projections of the POC terminate in a highly laminar fashion in layer Ib, immediately below the zone that contains the afferent input from the MOB; a lighter projection terminates in layer III. POC projections back to the AON also terminate in layer Ib, below the MOB recipient zone in layer Ia. Neurons in layers IIb and III send a dense projection back to the MOB; as noted previously, this feedback pathway terminates primarily in the GCL. Extrinsic outputs of olfactory cortex Two classes of POC outputs are discussed above—the feedback projection back to the MOB and the association connections between rostral and caudal olfactory cortex. A third class of outputs is treated separately because it represents the projections of the POC to brain regions not generally included in the olfactory system per se, although their receipt of inputs from the POC obviously implicates these POC targets in olfactory function. The extrinsic outputs of the POC are to both cortical and subcortical structures (Fig. 7). Neocortical projections (Fig. 7) The MOB projection to the POC extends dorsally beyond the cytoarchitectural

limits of the POC into the ventral parts of the granular insular and perirhinal cortices. There are also direct projections from the POC to insular and orbital cortices. Insular and orbital cortices are also the primary cortical targets of ascending pathways arising in the nucleus of the solitary tract (Sol) in the medulla and appear to contain the primary cortical representations for both gustatory and visceral sensation. Thus, olfactory projections to insular and orbital cortex may form part of the circuitry that integrates olfactory and gustatory signals to generate the integrated perception of flavor. These same cortical areas also have descending projections to the hypothalamus and back to the Sol that may influence visceral-autonomic and possibly gustatory functions. Neurons in these cortical areas in primates respond to odors with a higher degree of selectivity than neurons in either the MOB or the POC. Thus these neocortical sites may play a role in the discrimination of different odors. Subcortical projections (Fig. 7) (a) Hypothalamus— The heaviest and most direct olfactory projections to the hypothalamus arise from neurons in the deepest layers of the piriform cortex and the anterior olfactory nucleus. These projections terminate most heavily in the lateral hypothalamic area. Olfactory-recipient parts of the cortical and medial amygdaloid nuclei also project to medial and anterior parts of the hypothalamus. (b) Thalamus—There is a strong projection from the POC to the magnocellular, medial part of the mediodorsal thalamic nucleus and the submedial nucleus (nucleus gelatinosus) (Benjamin et al., 1982; Price and Slotnick, 1983). These thalamic nuclei project to the orbital cortex and the frontal lobes.

Odors and Cognition Physiological studies in monkeys suggest that some degree of odor discrimination may take place in 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 olfactory–neocortical circuit via the thalamus. For example, the olfactory tubercle, insular cortex, and piriform cortex receive input from mitral cell axons (Broadwell, 1975; de Olmos et al., 1978; Ojima et al., 1984; Shipley and Adamek, 1984). The olfactory tubercle and piriform cortex project to the dorsomedial thalamic nucleus (Benjamin et al., 1982; Powell et al., 1965) and the submedial thalamic nucleus (Price and Slotnick, 1983), although the projection from the piriform cortex (but not insular cortex) has been questioned (Motokizawa et al., 1988). The dorsomedial thalamus projects to the posterior orbitofrontal cortex

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29. OLFACTORY SYSTEM

947

FIGURE 7 Diagrammatic representation of some of the higher order connections of the main olfactory system. Emphasis is placed on possible circuits that mediate output responses such as autonomic or hormonal changes. Output projections of the MOB are shown as thick lines; higher order connections are shown as thin lines. Cortical structures are depicted as boxes; subcortical structures as ellipses. Abbreviations: 10, dorsal motor nucleus of the vagus; ACo, anterior cortical amygdaloid nucleus; AON, anterior olfactory nucleus; CE, central amygdala nucleus; DHR, dorsal hippocampal rudiment; DP, dorsal peduncular cortex; Ent, entorhinal cortex; HC, hippocampus; IC, insular cortex; IML, intermediolateral cell column; MD, mediodorsal nucleus; NLOT, nucleus of the lateral olfactory tract; OFC, orbital frontal cortex; P/Amb, nucleus ambiguous and periambigual area; PaCo, periamygdaloid cortex; CG, midbrain periaqueductal gray; Pir, piriform cortex; Pit, pituitary; RVL, rostroventrolateral medulla; SO, supraoptic nucleus; TT, taenia tecta; Tu, olfactory tubercle.

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 the orbitofrontal cortex that, according to Takagi (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 the lateral posterior orbitofrontal cortex more commonly respond to a single odor. Thus, corticocortical 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 the piriform cortex (Kaneko and Mizuno, 1988).

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 poorly understood. Lateral olfactory cortex projects heavily to the lateral hypothalamus which is known to be involved in visceroautonomic functions. Another possibility is that MOB

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and piriform cortex connections with the insular cortex (Fig. 7) might be involved in these functions (Ruggiero et al., 1987; Saper, 1982; Shipley, 1982; Shipley and Geinisman, 1984; Shipley and Sanders, 1982). A portion of the granular insular cortex is a site of significant overlap between olfactory and visceral information in mouse (Shipley and Geinisman, 1984) and rat (Krushel and Van Der Kooy, 1988). In addition, the medial olfactory cortex may be an area efferent control of visceral activity (Neafsey et al., 1986). There are direct projections from the MOB to the ventral part of the medial frontal cortex and there are reciprocal connections between the insular and the medial frontal cortices. Olfactory inputs to the insular cortex and the medial frontal cortex may influence autonomic and visceral function via direct cortical projections to cardiovascular regions of the ventral medulla and the solitary nucleus (Ruggiero et al., 1987) or 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 brain stem autonomic centers such as the periaqueductal gray and the dorsal vagal complex (Hopkins et al., 1981; Hopkins and Holstege, 1978; Rizvi et al., 1992). Portions of the periaqueductal gray project to the ventral lateral medulla (Ennis et al., 1997; Van Bockstaele et al., 1989) and may be involved in pressor and depressor responses of the cardiovascular system (Carrive et al., 1989). 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 and many neurons in the orbitofrontal cortex of the primate react to both taste and smell 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.

Olfaction and Motor Activity As noted earlier, several pathways link olfactoryrelated 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 is integrated with the motor control regions of the striatum so that visceral and somatic effectors may be controlled by these pathways (Newman and Winans, 1980). Olfactory linkages to the ventral striatum are mediated by parallel projections from the MOB, the AON, and the piriform cortex to the olfactory tubercle. Neurons in the olfactory tubercle and some in the 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, 1980).

Olfaction and Memory The entorhinal cortex receives a substantial input from the MOB (Broadwell, 1975; de Olmos et al., 1978; Shipley and Adamek, 1984). In turn, the medial and lateral entorhinal cortices project to the dentate gyrus and CA fields of the hippocampus. MOB projections to the 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, the piriform cortex has direct connections to the entorhinal cortex. Because the hippocampus is important in memory function, these olfactory– entorhinal–hippocampal circuits may be important for the establishment or recall of olfactory memories formed by or associated with other events.

THE ACCESSORY OLFACTORY SYSTEM In macrosmatic mammals such as the rat, two components of the olfactory system are recognized: the main and the accessory olfactory systems. These two components are parallel, but are, for the most part, anatomically and functionally separate.

Vomeronasal Organ (VNO) ORNs in the olfactory epithelium transduce mainly volatile odors and transmit this information to the MOB. By contrast, ORNs located in the vomeronasal organ (vomeronasal receptor neurons, VRNs; Figs. 1A and 2B) are exposed to nonvolatile odors by the engagement of a physiologically regulated pump mechanism. The vomeronasal epithelium can be divided into an apical zone and a basal zone (Fig. 2B). VRNs in the apical zone express V1R and V3R odorant receptor genes and the G-protein Gαi. VRNs in the basal zone express V2R receptor genes and the G-protein Gαo (Halpern et al., 1995; Herrada and Dulac, 1997; Matsunami and Buck, 1997; Pantages and Dulac, 2000). In the main olfactory system, our current understanding is that ORNs respond to chemical epitopes that may be present on different odorant molecules and that the tuning of main olfactory neurons is broad; i.e., a single ORN responds to related chemical epitopes with different affinities. However, VRNs show a narrower tuning in that single VRNs respond specifically to only one putative pheromone molecule. This response takes place at very low concentrations (e.g., 10−11M), which is several orders of magnitude lower than that observed in the main olfactory epithelium (LeindersZufall et al., 2000). This response profile to only one putative pheromone molecule does not broaden with

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29. OLFACTORY SYSTEM

increasing concentration, suggesting that VRNs are very narrowly tuned for putative pheromones (LeindersZufall et al., 2000). Populations of VRNs also exhibit strong selectivity for either male or female urine, although the molecules underlying this response are unknown (Holy et al., 2000). Thus, if ORNs in the main olfactory epithelium are hypothesized to be epitope sensors, VRNs might be detectors for single putative pheromone molecules. Axons of vomeronasal neurons project exclusively to the AOB located at the dorsocaudal limit of the MOB. VRNs from the apical zone project to the anterior half of the AOB, whereas VRNs in the basal zone project to the posterior half of the AOB. 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; Keverne, 1999; Macchi, 1951; Meredith, 2001). Some mammals are anosmic (e.g., porpoises) and lack an olfactory bulb (Breathnach, 1961; 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.

Accessory Olfactory Bulb The AOB is located at the caudal–dorsal end of the MOB. The AOB has some cytoarchitectural features similar to those of the MOB, but is much smaller. The vomeronasal nerve transmits information from the VNO to the glomeruli of the AOB. The glomerular layer in the AOB (GLA) is less distinct than that 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. The term “periglomerular” is thus less appropriate in the AOB than the 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 the AOB (EPLA) and the mitral cell layer (MCLA) are also less distinct than the corresponding layers of the MOB. The AOB internal plexiform layer (IPLA) is unremarkable and is situated between the mitral cell layer and the lateral olfactory tract. The granule cell layer (GCLA) of the 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 the AOB are much more

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polymorphic than their counterparts in the MOB (Takami and Graziadei, 1990, 1991). Indeed, there are five main categories of mitral cells based upon the dendritic arrangements of the cells in Golgi impregnated material. These mitral cells can have from one to five apical dendrites that ramify within different glomeruli (Takami and Graziadei, 1990, 1991). Vomeronasal receptor neurons (VRNs) expressing the same ORG also project to multiple glomeruli in the AOB (6 to 20) glomeruli per gene have been reported; (Belluscio et al., 1999; Rodriguez et al., 1999). Interestingly, the dendrites of at least some AOB mitral cells specifically project to different glomeruli innervated by VRNs expressing the same ORG (Del Punta et al., 2002). Neurotransmitters Electrophysiological studies demonstrate that glutamate is the vomeronasal receptor cell transmitter (Dudley and Moss, 1995; Jia et al., 1999). 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). Electrophysiological studies also indicate that mitral-to-granule cell dendrodendritic transmission is mediated by glutamate or aspartate, as in the MOB (Jia et al., 1999). 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-immunoreactive granule cells increase in number at the time when mitral cell expression is decreasing (Matsutani et al., 1989). The few “periglomerular cells” in the AOB are neurochemically different from those in the MOB. The most obvious difference is the lack of dopaminergic periglomerular cells in the AOB. Also lacking are the substance P-containing external tufted cells that are abundant in the MOB of some species (Baker, 1986). GABAergic periglomerular and granule cells are present in the AOB (Baker et al., 1988). Substance P-containing cells are most prominent in the GCLA of rats. In contrast, there are fewer substance P-immunoreactive cells in the GCLA in rabbit, guinea pig, cat, and hamster, and in mice these cells appear to be absent (Baker, 1986). Candidate transmitters in the accessory olfactory bulb are summarized in Table 7 and neurotransmitter receptors in Table 8.

Outputs of AOB The central connections of the AOB and the MOB to higher order olfactory structures are essentially nonoverlapping. The AOB has direct projections to the

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TABLE 7

Candidate Transmitters of Accessory Olfactory Bulb Cells

Transmitter

Cell type/location

Cell type

Frequency

Species

References

Aspartate

Output

Medium

Many

Rat

(1)

DA

Juxtaglomerular

Small

Rare

Many

(2)

GABA

Periglomerular

Small

Many

Many

Granule cells

Small

Many

Many

ENK

GL

Small

Few

Rat

Neurotensin

Output

Medium

Many

Guinea pig

(5)

SP

Output

Medium

Many

Rat, guinea pig

(3, 5)

GCL—granule cells

9 μm

Many

Rat, hamster

(2, 3)

VIP

EPL—mitral or Van Gehuchten

12 μm

Few

Cat

(6)

(3, 4)

References. 1, Fuller and Price (1988); 2, Baker (1986); 3, Matsutani et al. (1988); 4, Gouda et al. (1990); 5, Matsutani et al. (1989); 6, Sanides-Kohlrausch and Wahle (1990).

TABLE 8

Receptor Subtypes of the AOB

Receptor

AOB

Cholinergic mAChR nAChR

NR NR

receptors and thus may be modulated directly by circulating hormones. The efferent connections of the accessory olfactory system are summarized in Fig. 4.

References

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 size similar to that of females. These findings correlate with sexual dimorphisms in other structures (e.g. preoptic area and hypothalamic nuclei) that are known to influence sexual behavior (for review, Arnold and Gorski, 1984) and receive direct or indirect projections from the AOB (Scalia and Winans, 1975; Simerly et al., 1989).

(1–4)

Noradrenergic α1 α2 β1 β2

++ ++ ++ —

Dopaminergic D1 D2

NR NR

Serotonergic 5-HT1A 5-HT2A/C

NR —

Glutamatergic Kainate NMDA AMPA Metabotropic

NR ++ ++ ++

GABAergic GABAA GABAB

++ ++

(5)

(6–14)

Centrifugal Afferents to AOB

(15–20)

Key. ++, receptors present; —, receptors absent; NR, not reported. References. 1, Day et al. (1997); 2, Domyancic and Morilak (1997); 3, Rosin et al. (1996); 4, Woo and Leon (1995); 5, McLean et al. (1995); 6, Ohishi et al. (1993); 7, Ohishi et al. (1993); 8, Petralia and Wenthold (1992); 9, Petralia et al. (1994); 10, Petralia et al. (1994); 11, van den Pol (1995); 12, Shigemoto et al. (1992); 13, Romano et al. (1995); 14, Kinoshita et al. (1998); 15, Laurie et al. (1992); 16, Persohn et al. (1992); 17, Richards et al. (1987); 18, Young and Kuhar (1980); 19, Zhang et al. (1996); 20, Margeta-Mitrovic et al. (1999).

amygdala, specifically to the medial and posterior cortical nuclei, the bed nucleus of the stria terminalis, and the nucleus of the accessory olfactory tract (see also de Olmos et al., 1978). These pathways may be involved in the processing of pheromonal information. Neurons in the AOB targets express gonadal steroid

There are major differences between centrifugal inputs to the MOB and AOB (Fig. 6). First, centrifugal inputs to the AOB arise from far fewer brain regions than inputs to the MOB. The major afferents to the 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 the AON sends a dense projection to the granule cell layer of the AOB (Rizvi et al., 1992), but all other divisions of the AON lack connections with the AOB.

Higher Order Connections of the Accessory Olfactory System and Reproductive Functions Olfaction plays an important role in sexual behavior in many animals. Macrosmatic animals have a highly developed ability to use olfaction for identifying sexual partners, enemies, and food; i.e., these animals use

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29. OLFACTORY SYSTEM

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 macrosmatic animals, the AOB is involved in processing pheromones that are initially transduced by vomeronasal neurons, which project to the AOB. The AOB projects to the anteromedial (Me) and posterior cortical (PCo) nuclei of the amygdala, the bed nucleus of the stria terminalis, and the nucleus of the accessory olfactory tract (Figs. 4 and 8) (de Olmos et al., 1978; Scalia and Winans, 1975; Shipley and Adamek, 1984). Me and PCo have projections to other amygdaloid nuclei, notably the posterior amygdaloid nucleus (Canteras et al., 1992), and to the preoptic area and the hypothalamus (for review, Shiosaka et al., 1982). The posterior

951

amygdaloid nucleus (Po) appears to receive convergent input from both the Me and PCo and projects heavily upon some of the same structures targeted by the Me and PCo, 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 Me (MePD) 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 the MePD and lesions of the MPO decrease male copulatory behavior (for review, Simerly et al., 1989). Estrogen regulates the expression of CCK at the mRNA level in cells of the MePD; many cells containing CCK in the MePD project to the MPO. In the female rat, CCK injection in the medial preoptic

FIGURE 8 Diagrammatic representation of 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: 10, dorsal motor nucleus of the vagus; AONm, anterior olfactory nucleus, medial division; AP, posterior amygdaloid nucleus; APVH, anterior periventricular hypothalamus; BAOT, bed nucleus of the lateral olfactory tract; Bar, Barrington’s nucleus; BST, bed nucleus of the stria terminalis; IML, intermediolateral cell column; Me, medial amygdaloid nucleus; MPO, medial preoptic area; P/Amb, nucleus ambiguous and periambigual area; PAG, midbrain periaqueductal gray, Pit, pituitary; RVL, rostroventrolateral medulla; SCis, lumbosacral spinal cord; SO, supraoptic nucleus; VMH, ventromedial hypothalamic nucleus.

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region enhances luteinizing hormone secretion (Kimura et al., 1987), although its action on male sexual responses is not known.

“NONOLFACTORY” MODULATORY INPUTS TO THE OLFACTORY SYSTEM The olfactory system is heavily targeted by inputs from nonolfactory subcortical modulatory systems. These inputs arise from three principal sources: the nucleus of the diagonal band, dorsal and median raphe nuclei, and the locus coeruleus. The nucleus of the diagonal band (DB) is a component of the basal forebrain magnocellular system including the DB, the 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.

Nucleus of the Diagonal Band (DB) In the mouse about 3.5% of all neurons that project to the bulb originate in the horizontal limb of the DB (Carson, 1984); far fewer originate in the vertical limb of the DB (Carson, 1984; Shipley and Adamek, 1984). At least two distinct transmitter-specific populations of DB neurons project to the MOB (Zaborszky et al., 1986). 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 the 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 the DB (Zaborszky et al., 1986). Acetylcholinesterase (AChE) is one marker for the cholinergic axons. AChE staining in the bulb is preferentially concentrated in the IPL, the GCL, the 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 the vertical limb of the 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 (Greer et al., 1982; Teicher et al., 1980). It has been suggested that these glomeruli may be areas of specialized olfactory processing during development (Teicher et al., 1980). There are also AChE-positive neurons in the bulb (Nickell and Shipley, 1988). A more specific marker for

cholinergic axons is choline acetyltransferase (ChAT), the requisite enzyme for acetylcholine synthesis. ChATstained axons are located in layers similar to those described for AChE and are very fine in diameter (unpublished observations). The glomeruli of the AOB lack both ChAT and AChE staining. There are no ChATpositive neurons in the bulb. The GABAergic projection from the DB (Zaborszky 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.

Raphe Nuclei The midbrain dorsal and median raphe provides strong inputs to the MOB. In the rat, about 1000 dorsal and 300 median raphe neurons project to the bulb. These neurons are serotonergic and do not contain tyrosine hydroxylase (McLean and Shipley, 1987b) or substance P (Zaborszky et al., 1986). Thick serotonergic fibers preferentially innervate the glomeruli of the MOB while thinner serotonergic axons preferentially innervate inframitral layers (McLean and Shipley, 1987b). In neocortex, thick axons arise from the median raphe and thin axons arise from the dorsal raphe (McLean and Shipley, 1987b) and the same segregation occurs in the MOB. Serotonergic axons do not innervate the glomeruli of the AOB just as cholinergic axons avoid this layer. Since the AOB has far fewer PG cells than the MOB, the paucity of 5HT and ACh innervation of the AOB glomerular layer suggests that serotonergic and cholinergic inputs target PG cells in the MOB. The relative absence of PG neurons in the AOB, thus, might account for the lack of serotonergic/cholinergic input to that layer in the AOB.

Locus Coeruleus A significant modulatory input to the bulb is from the pontine nucleus, the locus coeruleus (LC). In the rat all LC neurons contain the neurotransmitter norepinephrine (NE); the LC contains the largest population of NE neurons in the brain. It has been estimated that up to 40% of LC neurons (400–600 of a total of 1600 LC neurons) project to the bulb in the rat (Shipley et al., 1985). LC axons project mainly to the infraglomerular layers of the bulb, particularly the internal plexiform and granule cell layers (McLean et al., 1989). 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 pattern, unusual for LC terminal fields, is observed in both the MOB and the AOB. In the AOB, the internal plexiform layer is, in fact, sharply demarcated by the

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29. OLFACTORY SYSTEM

dense NE fibers running through it just deep to the multicellular output cell layer (mitral cell layer) (McLean et al., 1989). Based on these light microscopic studies it was suggested that the major target of the NE input is granule cells (McLean et al., 1989). The nature of the physiological actions of NE in the bulb has been a matter of controversy. Salmoiraghi et al. (1964) and McLennan (1971) reported that iontophoretic applications of NE and NE agonists decreased mitral cell firing rates. McLennan further showed that NE failed to decrease mitral cell firing when bicuculline, a specific antagonist of the GABAA receptor, was coapplied with NE. This was interpreted to mean that NE causes granule cells to increase GABA release, thus inhibiting mitral cells. However, using an in vitro whole turtle bulb preparation, Jahr and Nicoll (1982) found that NE application caused increased mitral cell firing. Other studies indicate that activation of the LC, and hence synaptic release of NE, causes a twofold increase in the response of mitral cells to weak but not strong shocks applied to the olfactory nerve (Jiang et al., 1996). NE application in vitro also produced a similar enhancement of mitral cell responses to weak olfactory nerve input (Ciombor et al., 1999; Hayar et al., 2001). This is consistent with the idea that NE preferentially enhances responses to weak stimuli. The functional significance of this action might be to increase the sensitivity of mitral cells to weak olfactory inputs. Thus, when the 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 weak 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. Recently mated female mice abort if presented with the odors of a strange male mouse that is not the mate, a phenomena termed the Bruce effect after the discoverer. 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 AOB memory formation is important in the context of pregnancy block (Kaba et al., 1989). Keverne and colleagues suggest that the dendrodendritic synapse between granule cells and mitral cells in the AOB may be critical for 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. 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 MOB circuits are still being established in the bulb. The timing of noradrenergic axon arrival in the MOB correlates with pharmacological evidence of noradrenergic influence on mitral cell excitability in the immature bulb (Wilson and Leon, 1988).

Differential Innervation of MOB and AOB Modulatory centrifugal afferents from the diagonal band, the dorsal and median raphe nuclei, and the locus coeruleus are common to both the MOB and the AOB. The terminal distribution of these common inputs differs in the AOB and the MOB, especially with respect to the cholinergic and serotonergic inputs. In the AOB, both the cholinergic and serotonergic inputs avoid the glomeruli whereas they heavily innervate the glomeruli of the MOB. The cholinergic–serotonergic inputs to the AOB are mainly to the granule cell layer and internal plexiform layer (Le Jeune and Jourdan, 1991; McLean and Shipley, 1987a). The NE input appears to have similar laminar termination patterns in both the MOB and the AOB.

Olfactory Cortex Modulatory inputs to the olfactory cortex have not been studied in as much detail as those to the MOB. A recent preliminary report, however, indicated that ACh, 5HT, and NE all heavily innervate the olfactory cortex and, in addition, there is a heavy extrinsic DAergic input. Similarly, the functions of theses modulatory inputs to the olfactory cortex are poorly understood. There is evidence that ACh may be involved in associative learning and that excessive ACh release, as during anti-AChE intoxication, causes the generation of epileptic seizures. The functions of 5HT, NE, and DA are unknown.

Abbreviations Alternate abbreviations used in “The Rat Brain in Stereotaxic Coordinates” by G. Paxinos for some structures are listed in parentheses. 10 5HT

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Dorsal motor nucleus of the vagus 5-hydroxytryptophan (serotonin)

954 5-HT1A 5-HT1C 5-HT2A/C ac Acb ACh ACH AChE ACIII ACo AHC AI AMPA AOB AON AONd AONe AONl AONm AONvp APir APVH BAOT Bar BL BLP BM BMP BST CA1 CA3 CCK Ce CG ChAT CPu CRF D1 D2 DA DB DG DHR DP Dyn B En ENK Ent Entl Entm EPL EPLA GABA GABAA GABAB GCL GCLA GL GLA HC HDB IAS IC

MICHAEL T. SHIPLEY ET AL.

Serotonin receptor subtype 1A Serotonin receptor subtype 1C Serotonin receptor subtype 2A/C Anterior commissure Nucleus accumbens Acetylcholine Anterior hippocampal continuation Acetylcholinesterase Adenylate cyclase type III Anterior cortical amygdaloid nucleus Anterior hippocampal continuation Anterior insular cortex Amino-3-hydroxy-5-methyl-4-isoxazole propionic acid receptor Accessory olfactory bulb Anterior olfactory nucleus Anterior olfactory nucleus, dorsal division (AOD) Anterior olfactory nucleus, pars externa (AOE) Anterior olfactory nucleus, lateral division (AOL) Anterior olfactory nucleus, medial division (AOM) Anterior olfactory nucleus, ventroposterior division Amygdalopiriform transition area Anterior periventricular hypothalamus Bed nucleus of the lateral olfactory tract Barrington’s nucleus Basolateral amygdaloid nucleus Basolateral amygdaloid nucleus, posterior part Basomedial amygdaloid nucleus Basomedial amygdaloid nucleus, posterior part Bed nucleus of the stria terminalis CA1 region of the hippocampus CA3 region of the hippocampus Cholecystokinin Central amygdaloid nucleus Central (periaqueductal) gray Choline acetyltransferase Caudate–putamen (striatum) Corticotrophin-releasing hormone Dopamine receptor subtype 1 Dopamine receptor subtype 2 Dopamine Nucleus of the diagonal band Dentate gyrus Dorsal hippocampal rudiment Dorsal peduncular cortex Dynorphin B Endopiriform nucleus Enkephalin Entorhinal cortex Entorhinal cortex, lateral division Entorhinal cortex, medial division External plexiform layer (EPl) External plexiform layer of the accessory olfactory bulb (EPlA) γ-Aminobutyric acid γ-Aminobutyric acid receptor subtype A γ-Aminobutyric acid receptor subtype B Granule cell layer (GrO) Granule cell layer of the accessory olfactory bulb (GrA) Glomerular layer (Gl) Glomerular layer of the accessory olfactory bulb GlA) Hippocampus Horizontal limb of the diagonal band Intrabulbar association system Insular cortex

ICj IG IL IML IPL IPLA La LC LH LHRH Lot LPO mAChR MCL MCLA MD Me MePD MePV MnR MOB MPO nAChR NAG NE NLOT NMDA NPY OCNC OFC OMP ONL ONLA opt OR ORG ORN P/Amb PA PaCo PCo PG Pir Pit PLCo PMCo POC PrS RF RVL S SCis SEL SO SP St TH Tr TRH TT TTd TTis

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Islands of Calleja Indusium griseum Infralimbic cortex Intermediolateral cell column Internal plexiform layer (Ipl) Internal plexiform layer of the accessory olfactory bulb (IPlA) Lateral amygdaloid nucleus Locus coeruleus Lateral hippocampal area Leutinizing hormone releasing hormone Lateral olfactory tract (lo) Lateral preoptic area Muscarinic cholinergic receptor Mitral cell layer (Mi) Mitral cell layer of the accessory olfactory bulb (MiA) Mediodorsal thalamic nucleus Medial amygdaloid nucleus Medial amygdaloid nucleus, posterodorsal part Medial amygdaloid nucleus, posteroventral part Median raphe nucleus Main olfactory bulb (OB) Medial preoptic area Nicotinic cholinergic receptor; alpha and beta adrenergic receptors N-Acetylaspartylglutamine Norepinephrine Nucleus of the lateral olfactory tract (LOT) N-Methyl-D-aspartate receptor Neuropeptide Y Olfactory cyclic nucleotide gated channel Orbital frontal cortex Olfactory marker protein Olfactory nerve layer (ON) Olfactory nerve layer of the accessory olfactory bulb (VN) Optic tract Odorant receptor Odorant receptor gene Olfactory receptor neuron Nucleus ambiguous and periambigual area Posterior amygdaloid nucleus Periamygdaloid cortex Posterior cortical nucleus of the amygdala Periglomerular cells Piriform cortex Pituitary Posterolateral cortical amygdaloid nucleus Posteromedial cortical amygdaloid nucleus Primary olfactory cortex Presubiculum Rhinal fissure Rostroventrolateral medulla Subiculum Lumbosacral spinal cord Subependymal layer Supraoptic nucleus Substance P Stria terminalis Tyrosine hydroxylase Transitional area Thyrotropin releasing hormone Taenia tecta Taenia tecta, dorsal Taenia tecta, inferior-superior

29. OLFACTORY SYSTEM

TTvi TTvs Tu VDB VIP VMH VNO VP VRN

Taenia tecta, ventral-inferior Taenia tecta, ventral-superior Olfactory tubercle Ventral limb of the diagonal band Vasoactive intestinal peptide Ventromedial hypothalamic nucleus Vomeronasal organ Ventral pallidum Vomeronasal olfactory receptor neuron

Acknowledgments We would like to acknowledge grant support for portions of this chapter that refer to research of the authors: PHS Grants DC00347, DC03195, DC02588, DC05676, and DC005739.

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Wong, S. T., Trinh, K., Hacker, B., Chan, G. C., Lowe, G., Gaggar, A., Xia, Z., Gold, G. H., and Storm, D. R. (2000). Disruption of the type III adenylyl cyclase gene leads to peripheral and behavioral anosmia in transgenic mice. Neuron 27, 487–497. Woo, C. C., and Leon, M. (1995). Distribution and development of beta-adrenergic receptors in the rat olfactory bulb. J. Comp. Neurol. 352, 1–10. Wright, D. E., Seroogy, K. B., Lundgren, K. H., Davis, B. M., and Jennes, L. (1995). Comparative localization of serotonin1A, 1C, and 2 receptor subtype mRNAs in rat brain. J. Comp. Neurol. 351, 357–373. Wyss, J. M., and Sripanidkulchai, K. (1983). The indusium griseum and anterior hippocampal continuation in the rat. J. Comp. Neurol. 219, 251–272. Yang, X., Renken, R., Hyder, F., Siddeek, M., Greer, C. A., Shepherd, G. M., and Shulman, R. G. (1998). Dynamic mapping at the laminar level of odor-elicited responses in rat olfactory bulb by functional MRI. Proc. Natl. Acad. Sci. USA 95, 7715–7720. Young, W. S., III, and Kuhar, M. J. (1980). Noradrenergic α1 and α2 receptors: Light microscopic autoradiographic localization. Proc. Natl. Acad. Sci. USA 77, 1696–1700. Zaborszky, L., Carlsen, J., Brashear, H. R., and Heimer, L. (1986). Cholinergic and GABAergic afferents to the olfactory bulb in the rat with special emphasis on the projection neurons in the nucleus of the horizontal limb of the diagonal band. J. Comp. Neurol. 243, 488–509. Zhang, J. H., Cerretti, D. P., Yu, T., Flanagan, J. G., and Zhou, R. (1996). Detection of ligands in regions anatomically connected to neurons expressing the Eph receptor Bsk: Potential roles in neuron–target interaction. J. Neurosci. 16, 7182–7192. Zhang, J. H., Sato, M., and Tohyama, M. (1991). Different postnatal development profiles of neurons containing distinct GABAA receptor beta subunit mRNAs (β1, β2, and β3) in the rat forebrain. J. Comp. Neurol. 308, 586–613. Zheng, L. M., Caldani, M., and Jourdan, F. (1988). Immunocytochemical identification of luteinizing hormone-releasing hormonepositive fibres and terminals in the olfactory system of the rat. Neuroscience 24, 567–578. Zufall, F., and Munger, S. D. (2001). From odor and pheromone transduction to the organization of the sense of smell. Trends Neurosci. 24, 191–193.

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30 Vestibular System PIERRE-PAUL VIDAL Laboratoire de Neurobiologie des Réseaux Sensori-moteurs (LNRS) CNRS ESA 7060, Université Paris V, Paris, France

ALAIN SANS Neurobiologie et Développement du Système Vestibulaire, INSERM Unité 432 Université Montpellier II, Place Eugène Bataillon, Montpellier, France

For vertebrates, maintaining their body equilibrium in the gravitational field and being capable of orienting themselves in their environment are fundamental aspects of survival. These constraints imply permanent control of the head and trunk position in space but also control of the head in relation to the trunk. Three sensory modalities are strongly implicated: visual, proprioceptive, and vestibular. Gaze and postural stabilization are, therefore, results of a complex multisensory integration. This integration can be defined as the process of matching multiple internal representations of an external event (head and/or trunk rotation), obtained from different sensory modalities, into a unique intrinsic frame of reference in which appropriate motor commands can be coded (Gdowski et al., 1999, 2000; Roy and Cullen, 1998, 2001). Past studies have demonstrated that well-defined neuronal networks in the central nervous system implement these complex sensorimotor transformations, known as the vestibular, cervical, and optokinetic reflexes. In rat, vestibulospinal reflexes have not been studied in detail while a considerable amount of data is available in other species (for a review, see Wilson et al., 1995, 1999). However, eye–head coordination (Fuller, 1985; Dieringer and Meier, 1993; Meier and Dieringer suppress, 1993), the angular vestibuloocular reflex (Lannou et al., 1982; Hess et al., 1989; Reber et al., 1996), and the maculoocular reflex (Hess and Dieringer, 1991a,b) had been well described in rat, as has the effect of gravity on the horizontal and vertical vestibuloocular reflex (Brettler et al., 2000; Plotnik et al., 1999). The optokinetic reflex (Hess et al., 1985, 1989;

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Sirkin et al., 1985; Niklasson et al., 1990) and the cervicoocular reflex (Niklasson et al., 1988, 1990) have also been well described in that species. In addition, the functional plasticity of the rat vestibuloocular reflexes has been investigated following adaptation to visuovestibular conflicts (Tempia et al., 1991; Gauthier et al., 1995; for a review across species, see du Lac and Lisberger, 1995c) and habituation (Tempia et al., 1991; for a review across species, see Kawato and Gomi 1992). The postlesional plasticity of the rat vestibular system has been investigated in a variety of models: following unilateral ablation of the labyrinth [Hamman et al., 1998 in the rat; see Curthoys and Halmagyi (1995) and Dieringer (1995) for reviews across species), in a strain of mutant rat (Rabbath et al., 2001, Kaiser et al., 2001), following lesions of the commissural vestibular fibers (Tham et al., 1989), the frontal eye field (Bahring et al., 1994), the nucleus reticularis tegmenti pontis (Hess et al., 1989), and after exposition to toxic agents (Tham et al., 1982, 1984; Larsby et al., 1986; Niklasson et al., 1993; Magnusson et al., 1998). Finally, the vestibular reflexes have been also investigated following exposure to hypergravity (Wubbels and de Jong, 2001) and microgravity (Shipov and Aizikov, 1992). All these behavioral studies are not summarized here due to the limited scope of this review. The interested reader should refer to the papers quoted in this paragraph. The following sections consider the main characteristics of the vestibular system, including the physiologic properties related to the anatomic features of the peripheral and central vestibular system. In this case, we have insisted on the electrophysiological aspects

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and the neurotransmitters implicated in the vestibular nuclei, with preponderance of the medial vestibular nucleus, which has been mainly studied in these studies. Although an attempt has been made to focus the review on rat data, results obtained in other species have also been included when required.

THE VESTIBULAR MESSAGE: FROM THE PERIPHERY TO THE CENTER Vestibular Receptors The vestibular labyrinth includes five sensory organs: three cristae ampullares (horizontal, anterior, and posterior cristae) associated with a semicircular canal and two otolith organs (the utricular and saccular maculae). The horizontal canal is situated in a plane constituting a 30° angle with the horizontal plane, whereas anterior and posterior canals are in a vertical position, almost orthogonal to each other. Utricular and saccular maculae are opposed at approximately right angles to each other, with the utricle lying in a horizontal position. In the rodent the anterior part of the utricular maculae is tilted at an angle of 30° from the posterior horizontal part (Curthoys, 1982a). Cristae ampullares are sensitive to angular accelerations; each canal is preferentially stimulated by accelerations acting within its plane, but as the canals are not totally orthogonal, angular movements more or less affect the other canals. The otolith organs—utricles and saccules—specifically transduce the linear accelerations induced by the head movements and by the gravity. They do not respond to angular accelerations (Goldberg, 1979; Baloh and Honrubia, 1990). The sensory epithelium presents the same features in the five vestibular organs in mammals and birds and has been described extensively. Briefly, it consists of two types of sensory hair cell: the flask-shaped type I hair cell surrounded by an afferent nerve calyx and the cylindrical-shaped type II hair cell contacted by afferent buttons issuing from different nerve fibers (Fig. 1). Glutamate is the neuromediator of sensory cells. The presence of small clear vesicles and dense core vesicles has been shown at the apex of the afferent calyx, suggesting a release of neuromediators (Scarfone et al., 1988; Demêmes et al., 2000). Cell bodies of the efferent endings, which control vestibular messages, are located in the brain stem, below the fourth ventricle (for a review in fishes, see Highstein, 1991). These efferent fibers directly contact the membrane of type II hair cell or membranes of calyx and afferent fibers. Neuromediators of the efferent vestibular fibers—acetylcholine (Schwartz et al., 1986; Kong et al., 1997) and the calcitonin gene-related peptide—are colocalized in the

FIGURE 1 Transmission electron micrograph of a transverse section of a guinea pig utricular epithelium. Type I hair cells (1) are surrounded by afferent nerve calyces containing numerous mitochondria. Type II hair cells (2) are contacted at their bases by afferent endings and by efferent boutons (white arrows). Nuclei of the supporting cells (SC) line the lower part of epithelium, and these cells have numerous intracytoplasmic secretory granules at their apices. St, stereocilia.

efferent buttons (Tamaka et al., 1988; Ohno et al., 1991; Scarfone et al., 1996; Demêmes and Broca, 1998). Acetylcholine could exert an inhibitory control on afferent vestibular messages through nicotinic receptors (Ohno et al., 1993) while CGRP is thought to be excitatory. The electrophysiological characteristics of the two types of vestibular hair cells are varied (for developmental aspects, see Eatock and Hurley, 2003; Chabbert et al., 2003). These cells present several types of various currents. Nevertheless, type I hair cells are clearly identified by large stereotypical delayed outward potassium currents with voltage-dependent activation kinetics (IKI) (in the rat, Lennan et al., 1999; in the pigeon, Ricci et al., 1996). In rat utricles it has been shown that this current is activated at low voltages and has been named gKL (L for low-voltage activation; Rüsch and Eatock, 1996; Rüsch et al., 1998). Delayed outward potassium currents could be responsible for the irregular activity recorded in fibers of vestibular nerve (see following section). Type II hair cells possess a mixture of different currents: a delayed rectified potassium current (IKII), a potassium calcium current IKCa, a fast-inactivating current IA, and calcium currents (Rennie and Correia, 1994; Masetto et al., 1994; Rüsch and Eatock, 1996; Norris et al., 1992; Prigioni et al., 1992). Type II hair cells are contacted by small and medium afferent fibers presenting regular activity. The vestibular receptors with their two types of sensory cells are not organized in a simple network. In fact the five vestibular end organs present regional structural differences with complex organizations at the root of various physiological messages, not yet totally

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elucidated. For example, Lim (1984–1976) indicated that, in the utricle of the rat, the striolar zone presents short hair bundles not attached to the otoconial membrane, whereas the extrastriolar zone has long stereocilia, which partly penetrate the otoconial membrane. He suggested that hair cells in striolar regions could be more sensitive to velocity than to displacement. A correlation has also been found between afferent discharge properties and innervation patterns on the utricular macula (Golberg et al., 1990). The cristae ampullaris present, in their apical parts, short stereocilia not included in the cupula, whereas the stereocilia of medial and basal parts are long and included in the cupula (Mbiene and Sans, 1986). Numerous other features distinguish the central parts (striola area of macula; apical area of cristae) and the peripheral parts in the rat. Central parts mainly contain type I hair cells, presenting particular calcium-binding proteins as parvalbumin, enclosed in afferent multicalices immunoreactive for calretinin and calbindin. Peripheral parts present type I and type II hair cells contacted by medium fibers (dimorphic units), whereas the more peripheral parts contain numerous type II hair cells contacted by buttons of small fibers. It should be noted that medium and small fibers, as opposed to multicalyx, are not immunoreactive to calretinin and calbindin but contain substance P (Demêmes and Rhyzhova, 1996; Sans et al., 2001). There are also regional labyrinth specificities concerning the control of peripheral messages afferent to vestibular nuclei. The efferent terminals are highly concentrated at the peripheral parts of the receptors and rare in the central parts (Raymond and Demêmes, 1983). The transduction of the message in type I hair cells is possibly controlled by neurotransmitters included in the microvesicles located at the apex of the calyxes (Sans and Scarfone, 1996), as these afferent endings possess presynaptic Ca2+ channels and components of the presynaptic soluble N-ethylmaleimide-sensitive fusion factor attachment protein receptor SNARE proteins involved in synaptic vesicle docking and calcium-dependent exocytosis (Demêmes et al., 2000). In addition, a local control could be present at the base of the sensory cells by afferent vesiculated terminals, as shown in rat vestibular maculae (Ross, 1997). In summary, morphological features indicate that the vestibular receptors could address the central nervous system via the vestibular fibers and the primary neurons, elaborate coded messages whose meaning could depend on the receptor type (cristae or macula) but also on the receptor regions origin (for a review, Sans et al., 2001). However, recordings of the secondary neurons do not show the high specificity suggested by the morphological organization at the peripheral level,

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as it is not possible to realize physiological (natural) stimulations of only part of the vestibular receptors. However, some neurons of vestibular nuclei receive selective inputs of the various vestibular end organs (see sections on vestibular complex and second-order vestibular neurons).

First-Order Vestibular Neurons Regular and Irregular Neurons The first-order vestibular neurons are bipolar cells, which contact the hair cells of the labyrinth and the second-order vestibular neurons. Their soma is localized in the Scarpa’s ganglion (Curthoys, 1981), which is divided into two parts: the superior and the inferior. In rat and mice, anterior and lateral cristae and utricule and saccule maculae are in relation with neurons largely overlapping in the superior ganglion. Neurons localized in the inferior ganglion and connected to the posterior crista and saccule macula show tighter clustering but incomplete segregation (Maklad and Fritzch, 1999). Ultrastructural characteristics lead to classify vestibular ganglion neurons in two main types: large and small neurons (Rosenbluth, 1962). On the basis of neurochemical components, i.e., calbindin-D28K (Ca BP), calretinin (Ca R), and neurofilaments (NF) proteins, three subpopulations were identified: (1) CaBP- and CaR-positive neurons corresponding to the largest neurons (16%), (2) exclusively NF-positive neurons, and (3) unlabeled by Ca BP, Ca R, and NF antibody neurons (70%) (Demêmes et al., 1992; Dechesne et al., 1993). These different types of neurons express glutamate (Harper et al., 1995), acetylcholine receptor inducing activity ARIA (Morley, 1998), glycine (Baurle et al., 1997) choline acetyltransferase, and different receptors as glutamate receptors (Fugita, 1994; Doi et al.,1995; Usami et al., 1995; Niedzielski and Wenthold, 1995) and purinergic (P2X) receptors (Xiang et al., 1999). Most vestibular afferents are characterized by a resting discharge permitting the afferents to respond bidirectionally (for reviews, see Goldberg et al., 1991, 2000). The resting discharges can be regular or irregular. Regular first-order vestibular neurons are sensitive to the mechanical displacement of the canalar cupula and to the mechanical displacement of the otolithic membrane. They encode angular head velocity and the combination of linear forces applied to the otoliths (linear acceleration of the head combined with gravitoinertial forces). Regular first-order vestibular neurons are commonly referred to as tonic neurons. Tonic afferents include button and dimorphic units, which synapse in the extrastriolar zone of the otolithic maculae and in the peripheral zone of the cristae. They are characterized

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by thin- or medium-sized slow conducting axons and by a low sensitivity to head rotations and electrical stimulation. Irregular afferents are sensitive to the mechanical displacement of the mechanoreceptors and also to the velocity of their displacement. They are often referred to as phasic neurons. They include calyx and dimorphic units, which innervate the central cristae and the striolar zones. They are characterized by large- or medium-sized fast conducting axons and by a large excitatory response to efferent system stimulation (Baird et al., 1988; Goldberg et al., 1990 Brichta and Goldberg, 2000). Their sensitivity to linear or angular head rotations and electrical stimulations is on average six times higher than regular afferents (Goldberg et al., 1984). First-order vestibular neurons activated from the horizontal semicircular canal have been recorded in anesthetized rats by Curthoys (1982a). Regular firstorder vestibular neurons (43% of the cells) displayed higher spontaneous activity (63 spikes/s vs 18 spike/s), lower sensitivity (0.79 spike/s/s−2 vs 1.1 spike/s/s−2), and longer time constants (3–4 s vs 2–3 s) than irregular cells when tested with constant angular acceleration. Some irregular neurons were more sensitive to increasing accelerations than to decreasing ones, which was not the case for regular neurons. When challenged with sinusoidal angular acceleration (0.01–1.5 Hz), regular first-order vestibular neurons have a lower gain (0.83 vs 1 spike/s/s−2 at 0.5 Hz), a smaller phase lead with respect to velocity, and a longer time constant (4.36 s vs 4.03 s) than irregular cells. However, for both types of stimulations, there was considerable overlap between the characteristics of the two populations of cells. Clearly, there is a continuum of regularity spiking in first-order neurons (as is the case for second-order vestibular neurons, see later). Ionic Currents in First-Order Vestibular Neurons Analysis of voltage-activated conductances recorded in first-order vestibular neurons revealed the presence of a single type of sodium current in all vestibular neurons (Chabbert et al., 1997), five calcium currents (Desmadryl et al., 1997; Chambard et al., 1999), one type of hyperpolarization-activated inward current (IH) (Chabbert et al., 2001a), and a tetraethylammonion TEA-sensitive potassium current (IK) (Chabbert et al., 2001b). However, the properties of these currents and their distributions cannot account for the various electrical activities recorded in primary afferents. More interestingly, considerable variability in the expression of two fast activating potassium currents (IA and ID) (Chabbert et al., 2001b) suggests that, if they are involved in shaping action potential, a variation in their relative expression could explain the heterogeneity of the firstorder vestibular neurons.

Postnatal Maturation of First-Order Vestibular Neurons In the rat, head accelerations can be transduced by the first-order vestibular neurons (Curthoys, 1978, 1979, 1982b) and encoded by the second-order vestibular neurons at birth (Lannou et al., 1979; Rüsch et al., 1998 in mouse). The semicircular canals reach their adult size by P20 and vestibular hair cells appear to be mature at 2 weeks following birth. During the first 2 weeks, first-order vestibular neurons (Desmadryl, 1991; Curthoys, 1978, 1979, 1982a) and second-order vestibular neurons (Lannou et al., 1979) also progressively mature. As a result, the vestibuloocular reflexes, which are already present at birth, increase in sensitivity during the first postnatal month (Curthoys, 1979). Irregular first-order vestibular neurons have been recorded at birth. They respond to head accelerations with a gain similar to the adult rat at P6, while their phase lead with respect to velocity continues to decrease up to P30. Regular afferents are absent at birth and begin to appear at P5. Then, their number and their resting discharge rise steadily to reach adult values around P30. In contrast to irregular cells, regular first-order vestibular neurons display low sensitivity and a long phase lag in the adult as soon as they appear. These data can be compared with the results of previous studies in mice concerning the maturation of the first-order vestibular neurons (Dememes and Sans, 1985; Desmadryl et al., 1986, 1997, 1998; Desmadryl, 1991; Chambard et al., 1999; Dutia et al., 1995, 1998).

THE VESTIBULAR NUCLEAR COMPLEX: MORPHOFUNCTIONAL PROPERTIES Anatomy of Vestibular Nuclei The vestibular primary fibers, which connect to the vestibular receptors at the periphery, arrive in the central nervous system in a nuclear complex, which forms the vestibular nuclei. In fact, this denomination is partially incorrect, as (1) the greater part of these nuclei is totally free of direct vestibular terminal fibers, (2) they receive other sensory modalities and specially proprioceptive influx, and (3) the main tract projecting to vestibular nuclei originates in the cerebellum (Brodal, 1974). Hence, the vestibular complex must be understood as an important junction, which receives various information (visual, proprioceptive, vestibular), leading nuclei to assume a major role in the maintenance of balance and equilibrium. The vestibular complex is classically divided into superior, lateral, medial, and spinal vestibular nuclei

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and smaller cell groups, the interstitial nucleus of the vestibular nerve and groups F, L, X, Y, and Z. These small groups of cells are topographically associated with the four main vestibular nuclei. The central trunk of the vestibular nerve leaves the inner auditive meatus and penetrates orthogonally into the brain stem in front of the external cochlear nucleus. It then splits, in the lateral vestibular nucleus, into two secondary sections; the ascending and descending branches. The ascending branches project into superior, medial, and lateral nuclei; the descending branches project into lateral and spinal vestibular nuclei. Each vestibular fiber provides numerous collaterals, which project into the four vestibular nuclei (Fig. 2). They form an original pattern on the second-order vestibular neurons: some of these cells are specifically connected to the periphery and receive fibers of only one kind of receptor (vertical or horizontal cristae, maculae), whereas others are multiconvergent and receive fibers of different kinds of receptors [in the cat, Graf and Ezure (1986); in the rat, Kubo et al. (1977); Lannou et al. (1980); and Angelaki and Perachio (1993)].

FIGURE 2 Transversal brain stem section through the medial portion of a postnatal 3-day mouse vestibular complex. Parvalbumin immunocytochemical labeling (green) shows the distribution of vestibular nerve fibers (arrow) and their branching in vestibular nuclei. MVe, medial vestibular nucleus; LVeD, dorsal part of the lateral vestibular nucleus; LVeV, ventral part of the lateral vestibular nucleus; 4V, fourth ventricle; Co, cochlear nucleus; 8vn, vestibular nerve; PrH, prepositus hypoglossi nucleus. Courtesy of J. Puyal.

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Brodal and Pompeiano (1957) have studied extensively the anatomy and the afferent and efferent connections of the vestibular nuclei in the cat (for reviews, see Brodal 1972–1974). In the rat, the intranuclear organization of the vestibular nuclei and their relationship with other structures has been investigated by Rubertone and co-workers (Rubertone and Mehler, 1981; Rubertone and Haroian, 1982; Rubertone et al., 1983, 1995). This section only summarizes the anatomy of the main vestibular nuclei to the extent that they are useful for understanding some important morphofunctional correlations. Vestibular nuclei are limited rostrally by the brachium conjunctivum, laterally by the inferior cerebella peduncle (restiform body), ventrally by the nucleus and spinal tract of the trigeminal nerve, and medially by the fourth ventricle and the reticular formation. Superior Vestibular Nucleus (SuVe) This nucleus extends from the level of the motor nucleus of the trigeminal nerve to the anterior portion of the lateral vestibular nucleus. The SuVe is composed of medium-sized cells and small cells. The medium-sized cells are predominant in the central part of the nucleus, forming a core. This arrangement, which can be found in most mammals, including humans (Sadjadpour and Brodal, 1968), is less evident in rodents, especially in rats (Rubertone et al., 1995). The five vestibular receptors provide terminals in the SuVe (Korte and Friedrich, 1979), the major input coming from the ampullae. In fact, a selective projection of each receptor in a specific area has been reported. A massive fiber input issuing from the cristae arrives on the central part of the nucleus on large- and medium-sized cells, whereas fibers of the utricle and the saccule only innervate the periphery of the nucleus (Baloh and Honrubia, 1990). Such a selective projection also exists for the important contingent of fibers arriving from the cerebellum: inputs from the flocculus arrive in the central part and inputs from nodulus, uvula, and the medial nucleus arrive in the marginal regions of the SuVe [in the rat, Rubertone and Mehler (1981); Xiong and Matsushita (2000); in other species, Brodal (1974)]. All parts of the lateral cerebellar nucleus or dentate nucleus send fibers onto the ipsilateral vestibular nuclear complex with preferential projections of the dentate nucleus on the SuVe and spinal nucleus (Delfini et al., 2000). The SuVe projects, via the ipsilateral and contralateral medial longitudinal fasciculus in oculomotor nuclei (Graybiel and Hartwieg, 1974), but in the rat the ipsilateral projections predominate (Rubertone et al., 1995). The SuVe send specific projections to the thalamus, especially in the lateral part of the parafascicular nucleus and the dorsal part of the caudal ventrolateral

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nucleus (Shiroyama et al., 1999). The SuVe also projects to the oral pontine reticular nucleus (Hoddevik et al., 1975) and to the cerebellum. In this case the projections are bilateral and reciprocal with a heavy projection on the flocculus (Rubertone and Mehler, 1981). Commissural connections exist with other contralateral vestibular nuclei, with point-to-point connections between the two SuVe (Ketterer et al., 1990). Considering the massive inputs of the cristae ampullae on the SuVe and their principal efferent projections on oculomotor nuclei, the SuVe appears to play a major role in the control of eye movements. Lateral Vestibular Nucleus (LVe) Rostrally, the LVe is bounded by the caudal pole of the SuVe, caudoventrally by the spinal vestibular nucleus, and medially by the medial vestibular nucleus. Classically, a rostroventral part and a caudodorsal part have been distinguished on the basis of cytoarchitectural differences. In particular, giant cells, typical of the LVe (60 μm in rats), are mostly located in the caudodorsal part of this nucleus. Rubertone et al. (1995) proposed the incorporation of the ventral part of the LVe of the rat in the Mve as a magnocellular part of the latter. Nevertheless, the previous description of LVe is functionally coherent, as the two parts of the latter are the source of the main vestibular projection to the spinal cord via the lateral vestibular tract. The rostroventral part projects into the cervical cord, the caudodorsal part to the lumbosacral cord, and the intermediate part into the thoracic cord. Lateral vestibular spinal fibers terminate in the VIII and VII lamina. Hence, these projections “form” a somatotopic pattern with a clear-cut correspondence between each part of the LVe and the spinal cord (Brodal, 1967; Shamboul, 1980). The two parts of LVe also present other specific and complementary organization patterns : the ventral part is in relation with the labyrinth and the dorsal part with the cerebellum. Indeed the rostroventral part receives vestibular primary fibers originating from the labyrinth end organs, with a main contingent coming from the otolith receptor (utricle and saccule). The dorsocaudal part receives afferent fibers from the cerebellar cortex (vermis) and medial nucleus (Brodal, 1974) with clear somatotopic organization. In the rat projections coming from the lateral part of lobule VII and the adjacent copula pyramis have been described with autoradiographic techniques (Umetani et al., 1986; Umetani and Tabuchi, 1988). The lateral vestibular nucleus also receives a contingent of spinal fibers from cervical (Jones et al., 1986) to lumbar segments in the rostrodorsal part and from cervical and upper thoracic segments in its ventral part (Matsushita et al., 1995). The rostroventral part of the LVe also sends numerous commissural fibers onto the

lateral nucleus and the caudal superior vestibular nucleus (gerbil, Newlands et al., 1989) and sends efferent fibers unilaterally to the medial longitudinal fascicle. The major projections of the maculae on the ventral part of the lateral vestibular nuclei and the massive efferent fibers leaving the two parts of this nucleus, via the spinal cord, indicate that the LVe is mainly implicated in the control of posture. Medial Vestibular Nucleus (Mve) The fourth ventricle limits the dorsomedial border of the Mve over its entire length. The rostral part extends up to the posterior ventral zone of the superior vestibular nucleus and the caudal part ends at the level of the prepositus hypoglossal nucleus bounded ventrally by the nucleus of the solitary tract. Laterally the Mve is limited by the LVe and then by the spinal vestibular nucleus. The cytostructure of this nucleus is relatively homogeneous with small and medium cells densely packed together, giving a dark staining aspect to the Mve, which contrasts with the neighboring nuclei. Small cells are more numerous in the rostral part and thin fibers cross the nucleus in all directions. In the cat, the cristae ampullaris project in the dorsorostral part and the otolith organs in the Mve ventromedial part (Gacek, 1969). The selective projection of these sensors has so far not been documented in the rat. The rat Mve receives a contingent of cerebellar fibers from the caudal vermis (Bernard, 1987; Umetami, 1992; Xiong and Matsushita, 2000) and bilaterally from the medial nucleus (Rubertone et al., 1995). Medial nucleus GABAergic neurons project via axon collaterals to both the vestibular nuclei and the inferior olive (Diagne et al., 2001). A zonal organization of flocculovestibular connections has been established in rats with a specific pattern for dorsal and ventral surfaces of the flocculus. This pattern organization in relation with climbing fiber zones could be a specialization for controlling the activity of the extraocular muscles (Balaban et al., 2000). Other afferents concern a spinovestibular projection through the medial reticular formation in the caudal part of the nucleus with inputs originating from nerve endings in the ligaments and capsule of the upper cervical joints. Afferents from rostral cervical segments (C2–C3) mainly project in the caudal part of the Mve (Bankoul and Neuhuber, 1990). A more extensive projection of cervical afferents in the rat has also been described in the spinal vestibular nucleus and throughout the LVe (Matsushita et al., 1995, Xiong and Matsushita, 2001). Each Mve also receives an important projection from the controlateral medial nucleus through commissural fibers. The caudal part of Mve also receives (as spinal vestibular nucleus) inputs from the cerebral cortex. Cortical projections are bilateral

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predominantly and concern somatosensory areas and the frontal cortex. The rat corticofugal projections to the caudal vestibular nuclei could participate in modulation of the vestibuloocular and vestibulospinal reflex during locomotion (Nishiike et al., 2000). Efferent connections of the Mve concern ascending and descending projections in the medial longitudinal fasciculus. Ascending connections are bilateral and mainly project onto the oculomotor nuclei. The ventral Mve sends fibers in the thalamus, especially in the lateral parafascicular nucleus and the dorsal part of the ventrobasal thalamus nucleus, with an ipsilateral predominance (Shiroyama et al., 1999). The descending connections form the medial vestibulospinal tract with the majority of fibers ending on the cervical anterior cord, essentially in laminae III to V, with some scattered fibers in laminae I and VI (Bankoul and Neuhuler, 1992). Efferent bilateral connections to the cerebellum with caudal vermis and medial nucleus corresponding to reciprocal projections between medial vestibular nucleus and cerebellum have also been demonstrated in the rat (Rubertone and Mehler, 1981; Haroian, 1984; Barmak et al., 1992; Voogd et al., 1996). The convergence of inputs coming from vestibular receptors and neck in the Mve and the major output, which reach the oculomotor nuclei and upper cervical cord, indicate that the Mve is in stabilizing gaze and posture in the horizontal plane. Spinal Vestibular Nucleus (Spinal Nucleus) (SpVe) The rostral part of this nucleus is situated under the posterior section of the lateral vestibular nucleus and its caudal part is situated at the level of the parasolitary nucleus. Laterally it is limited by the external cuneatus nucleus and the restiform body and medially by the Mve. Small- and medium-sized cells constitute the majority of the cell population of the SpVe, but larger cells are numerous in the more anterior part. Longitudinally oriented fibers coming from the vestibular root and from the cerebellum interlace the cells throughout the nucleus; this feature is characteristic of the SpVe. Like the other vestibular nuclei, the SpVe receives primary vestibular fibers with fibers originating from the cristae projecting to the center of the nucleus and those of the maculae projecting to the periphery. The major projections to the spinal vestibular nucleus originate in the cerebellar cortex (flocculus, nodulus, and uvula) and the ipsilateral and contrateral medial cerebellar nuclei (Rubertone et al., 1995; De Zeeuw and Berrebi, 1995; Mouginot and Gahwiler, 1995). Fibers from the magnocellular region and the dorsolateral humps of the dentate nucleus reach the spinal nucleus (Delfini et al., 2000). Spinal fibers issued from the dorsal

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horn of cervical and lumbar levels run with the dorsal spinocerebellar tract and also terminate in the SpVe. The SpVe sends important vestibulocerebellar projections to the cerebellar cortex and to the deep cerebellar nuclei, in particular the medial nucleus, constituting to reciprocal connections (Brodal, 1972; Haroian, 1984). The SpVe also projects in the thalamus: the lateral part of the parafascicular nucleus, the transitional zone between ventrolateral and ventral posterolateral nuclei, and the caudal part of the ventrobasal complex (Shiroyama et al., 1999). The SpVe also sends fibers in the spinal cord, as spinal vestibular nucleus neurons are labeled retrogradly by choleratoxin injected at the c2–c3 level in rats (Matsushita et al., 1995). The reticular formation (paragigantocellular reticular nucleus) also receives afferent input from the spinal vestibular nucleus. It also sends numerous commissural fibers to the contralateral vestibular nuclei. The SpVe seems to be mainly implicated in the integration of vestibular spinal and cerebellar messages implicated in the control of body posture.

Second-Order Vestibular Neurons The following section focuses on how the nonlinear properties of the second-order vestibular neurons combine in vivo with the emerging properties of these networks to achieve gaze and postural control. The internal representations of head and trunk movements (Gdowski et al., 1999, 2000; Roy and Cullen, 1998, 2001) processed by the vestibular nuclei also influence various cortical areas at the origin of the perception of egomotion. This point will not be further developed in the absence of data on that topic in rat (apart from the nice study of Bahring et al., 1994). The interested reader should refer to the review of Fukushima (1997) for further information on that topic. In Vivo Studies Second-order vestibular neurons are activated monosynaptically by first-order ones. They are localized in the vestibular nuclei and project to a variety of targets, including the prepositus hypoglossi, reticular and thalamic neurons, the cerebellar Purkinje cells, and the extraocular motor neurons. Until now the main results have been obtained on the medial vestibular nucleus neurons (MveN), which mediate the horizontal vestibuloocular reflex. Therefore, this review essentially focuses on these cells. When used alone, the term MveN will refer to neurons localized in the medial vestibular nuclei, which have not been specifically identified as secondorder cells. Some MveN increase their discharge rate during horizontal head rotations directed toward the side of the recordings: they are designated type I MveN

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(Duensing and Shaeffer, 1959). Other MVe cells, known as type II MveN, increase their discharge rate during horizontal head rotations directed away from the recording side. They are inhibitory interneurons responsible for the commissural inhibition linking the two vestibular complexes (Shimazu and Precht, 1966). Response to angular accelerations MveN responding to horizontal head rotations have been investigated in the anesthetized rat (Kubo et al., 1975, 1977; Hamman and Lannou, 1987; de’Sperati et al., 1993). Type I and type II neurons represented, respectively, 58 and 42% of the population of neurons recorded by Hamman and Lannou (1987). Their spontaneous activity amounted to 22.7 and 17.4 sp/s, respectively. During sinusoidal horizontal rotations, their gain varied from 0.61 to 1.48 spike/s/s−2 for the tested frequencies. One group of cells discharged in phase with head velocity (lead of 2.8° revelocity at 0.3 Hz in the study of de’Sperati et al., 1993), whereas two other smaller groups of MveN led and lagged head velocity, respectively. Due to the absence of a detailed description of identified second-order vestibular neurons in the alert rat, we briefly summarize here their characteristics in the medial vestibular nucleus in the alert guinea pig (Serafin et al., 1995, 1999). Guinea pig type I MveN were characterized by (a) their monosynaptic activation (0.93 ± 0.11 ms, range: 0.85 to 1.15 ms) following single-shock stimulations of the ipsilateral vestibular nerve and (b) an increasing activity during horizontal head accelerations toward the recording side. These cells exhibited a resting discharge of 24.8 ± 12.9 spikes/s−1 (range: 3.4 to 54 spikes/s−1). MveN could be described as a continuum of cells extending from regular to irregular units with an average coefficient of variation of 0.52 ± 0.18 (range: 0.22 to 1.0, n=51). About one-third of the cells exhibited an eye position sensitivity. In addition, half of them decreased their firing or became silent during ipsilaterally directed rapid eye movements (fast phases or saccades) and exhibited a burst of action potential during the contralaterally directed ones. When challenged with horizontal sinusoidal head rotations in darkness, second-order MveN discharged in phase with head velocity over a large frequency range (0.1 to 3 Hz). Their gain increased from 0.65 ± 0.21 spikes/s−1/deg/s−1 at 0.1 Hz to 1.15 ± 0.24 spikes/s−1/deg/s−1 at 3 Hz. The phase lead of their discharge re head velocity decreased from 23.5 ± 15.1° at 0.1 Hz to 2.9 ± 6.9° at 0.5 Hz and remained in phase with head velocity up to 3 Hz. Altogether, the characteristics of type I MveN in the guinea pig are, therefore, qualitatively similar to those obtained in cats and monkeys (Gdowski and McCrea, 1999; Roy and Cullen, 1998, 2001).

Response to linear accelerations Several groups (Lannou et al., 1980; Angelaki et al., 1993) have investigated the response of rat central vestibular neurons to horizontal linear acceleration. They recorded neurons in and around the medial vestibular nuclei in decerebrated rats. Neurons were functionally identified as either horizontal semicircular canal-related (HC) neurons or vertical semicircular canal-related (VC) ones on the basis of their responses to angular rotations. In addition, some cells were classified as purely otolith organ-related (OTO) neurons. These different types of neurons were challenged with sinusoidal linear translation in the horizontal head plane. Convergence of macular and canal afferents (Kubo et al., 1977) takes place in 73% of type I HC, in 79% of type II HC, and in 86% of VC neurons. In all the recorded neurons, a direction of maximum sensitivity to the linear stimulations was determined together with a “null” direction perpendicular to it. Most of the cells exhibited a significative response along the null direction and their response phases varied as a function of stimulus direction; therefore, two response vectors were used to describe the spatiotemporal response properties of these cells : the largest response vector Smax and the smallest response vector Smin, which was perpendicular to Smax and led Smax by 90°. The tuning ratio (Smin gain/Smax gain), a measure of the two-dimensional spatial sensitivity of the recorded neurons, varied with the stimulus frequency. While these neurons encoded linear acceleration when stimulated along their maximum sensitivity direction, they discharged proportionally to the change of linear acceleration (jerk) when stimulated along their minimum sensitivity direction. Smax vectors were distributed throughout the horizontal plane, and Smin vectors were clustered around the nasoocciput, ipsilaterally with respect to the head acceleration head axis. HC and VC neurons display different spatial and temporal properties, which fit with the spatiotemporal organization of the ocular responses recorded during linear translation in the horizontal head plane. In addition, Bush et al. (1993) proposed that the central otolithic system was organized in canal coordinates, i.e., there is a close alignment between the plane of angular acceleration (canal) sensitivity and the plane of linear acceleration (otolith) sensitivity in otolith/canal-convergent vestibular nuclei neurons. The processing of otolith information has also been investigated in lateral and spinal vestibular nuclear neurons during off-vertical axis rotation (Lai and Chan, 1995, 1996). This protocol is convenient for testing the sensitivity of the vestibular neurons to their otolith afferences, independently of their canalar inputs.

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Response to visual and proprioceptive stimulations As for other vertebrate species, it was shown in alert rat (Cazin et al., 1980a, 1980b; Lannou et al., 1982) that MveN responding to horizontal head acceleration also responded to rotation of a large field visual pattern directed away from the recording side (optokinetic stimulus). During head rotation in the light, the recorded MveN increased activity during horizontal head accelerations toward the recording side, whereas the retinal slip induced by the head movement was directed away from that side. That is to say the MveN visual response was synergistic with their vestibular sensitivity. All type I and type II neurons responded to optokinetic stimuli. There was often a delay of 1 to 3 s following the onset of visual stimuli and firing changes. The gain of the response to optokinetic sinusoidal stimuli dropped steeply above 0.5 Hz. For low stimuli, the optokinetic response was linear and in phase with surround velocity, whereas with higher frequencies a progressive phase lag occurred. Two visual pathways were involved (Cazin et al., 1984). Vestibular neurons were driven optokinetically from the contralateral eye via (a) the pretectum and the prepositus hypoglossal complex and (b) the pretectum, the reticular tegmental nucleus of the pons, and the prepositus hypoglossal complex. Interestingly, the authors also described a transcerebellar route from the pretectum via the nucleus reticularis tegmenti pontis to the ipsi or contralateral flocculi. These results show that with combined vestibular–visual stimuli, i.e., head rotation in the light, the optokinetic system improves the poor low-frequency response of the semicircular canal. In summary, the combination of the two inputs expands the bandwidth of the vestibular reflexes. Finally, anatomical evidence (Neuhuber and Zenker, 1989; Arvidsson and Pfaller, 1990; Bankoul and Neuhuber, 1992; Matsushita et al., 1995) strongly suggests those rat central vestibular neurons process proprioceptive afferences, as is the case in other vertebrate species. The proprioceptive signals are generated by the numerous muscular and/or articular receptors activated during head, trunk, and leg movements. Postnatal maturation Semicircular canal-related vestibular neurons As is the case for vestibular afferents, the second-order vestibular neurons resting discharge is low and irregular during the first postnatal days. Then their resting discharge increases gradually and becomes more regular to reach adult values by the end of the first month. The first low threshold units appear at P4 at a time when regular firing begins in first- and second-order vestibular neurons. Their threshold continues to decrease to become five times smaller at P14, where it reaches adult values.

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The sensitivity of the central vestibular neurons is low at birth and increases steadily to reach adult values by P30. Their time constants and their phase lead tend to diminish slightly during that period. Despite the fact that the eyes open around P13, central vestibular neurons only encode the visual signal at P20 (Lannou et al., 1979, 1980; Reber-Pelle, 1984), at a time when spike generation becomes mature in MveN (see later). Otolith-related vestibular neurons Lai and Chan (2001) recorded the spontaneous activities and response dynamics of otolith-related vestibular neurons in decerebrated rats at P7, P14, P21, and P84 (adult). Cells were recorded extracellularly in the lateral and spinal vestibular nucleus during constant velocity off-vertical axis rotation, which selectively stimulates the otolith sensors. They displayed sinusoidal position-dependent modulation and could be either modulated during the full cycle or silenced during parts of each rotary cycle (clipped). From P7 to P84, the proportion of clipped cells decreased progressively from about 75% to less than 25%. The sensitivity of the otolith-related vestibular neurons increased with age to reach a plateau at P21 for clipped cells and P14 for nonclipped cells. At P7, both types of cells had irregular spontaneous activity, which became more regular as the rats matured. Beyond P14, while spontaneous activity increased in all cells, clipped neurons tended to have significantly lower resting rates and higher gains than nonclipped ones. Irregular neurons could display phase-stable and phase-shift responses, whereas regular ones always displayed a phase-stable response. In Vitro Studies As for second-order vestibular neurons, only cells located in the medial vestibular nucleus, which mediate gaze stabilization in the horizontal plane, have been investigated so far. Intracellular studies in rat slices (Gallagher et al., 1985; Lewis et al., 1989; Johnston et al., 1994) led to the conclusion that medial vestibular nuclei neurons (MveN) can be segregated into two main classes according to their intrinsic membrane properties: type A and B cells. Nevertheless, before describing these two types of neurons, two points should be stressed. First, membrane properties of neurons located in other vestibular nuclei may be different than those in the medial vestibular nucleus. Second, the classification of the MveN on the basis of the profile of their conductances is still a subject of debate. On the one hand, MveN could be distributed as a continuum between two stereotyped schemes corresponding to the canonical type A and B cells (du Lac and Lisberger, 1995a). On the other hand, it could be that once the inherent variability of intracellular recordings is taken into account

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by averaging several successive action potentials, MveN could be categorized unambiguously as type A or B cells (Johnston et al., 1994). The following description is mainly based on the results of Gallagher et al. (1985) Johnston et al. (1994 and Genlain et al., 2003) in rat and is compared to results recorded in guinea pig (Serafin et al., 1991a, 1991b, 1992a, 1992b) and fowl (du Lac and Lisberger, 1995a, 1995b). MveN identification MveN on slice exhibit a regular tonic discharge. It ranges from 4.3 to 36.4 spikes/s (mean 17.1 spikes/s) when recorded extracellularly in the rat (Dutia et al., 1992). Type A MveN constitute 33% of the 123 cells recorded by Johnston et al. (1994) in the rat, a percentage similar to previous findings in the guinea pig (Serafin et al., 1991a). They are characterized by wide action potentials (0.84 ms at firing threshold) followed by a single deep afterhyperpolarization (AHP). Type A MveN also exhibit a transient rectification of the membrane potential, which delays the firing of these cells at the break of hyperpolarizing current steps. This rectification is likely due to activation of an A-like current. In rats, this current is present in both type A and B MveN and is larger in type A cells. In guinea pig, an A-like current is only present in type A MveN. Type A MveN also display small, high-threshold calcium spikes and modest persistent subthreshold sodium plateau potentials (these plateau potentials are not present in guinea pig type A MveN). Type B MveN constitute 67% of the 123 cells recorded by Johnston et al. (1994) in the rat. This percentage was inferior in the guinea pig. This is probably due to the fact that the spike-shape averaging method has led to the identification of cells, described previously as having intermediate characteristics in the guinea pig (type C cell), as type B neurons in the rat. Type B MveN have thinner action potentials (0.63 ms at firing threshold) followed by a biphasic AHP with an early fast and small component followed by a delayed and slower one. As a rule, the AHP is clearly smaller than in type A MveN. Type B MveN also display persistent subthreshold sodium plateau potentials, high-threshold calcium spikes, and prolonged calcium-dependent plateau potentials [in guinea pig, about one-quarter of B MveN, namely type B+LTS MveN, display low-threshold calcium spikes (LTS) that grant them bursting properties]. The membrane resistance (61.5 MΩ) and the membrane time constant (11.65 ms) are not significantly different in type A and B MveN in the rat. The two types of neurons do not appear to be spatially segregated within rat medial vestibular nuclei. Ionic currents in MveN Sodium currents Both types of MveN display the

usual TTX-sensitive sodium conductances that generate the action potential (Patko et al., 2003, Vassias et al., 2003). More interestingly, type A and B MveN are endowed with a long-lasting subthreshold plateau. When IK is blocked by TEA, these plateau can be observed in response to depolarizing pulses from a hyperpolarized membrane potential. In rat, while subthreshold plateau are short lasting in type A MveN, they are larger and longer in type B cells. In both cell types, subthreshold plateau are suppressed by tetrodotoxin (TTX) but persist when calcium is replaced by cadmium or in Ca2+-free medium, which indicates that they are provoked by a sodium persistent current (INaP). Similar results were obtained previously in guinea pig (Serafin et al., 1991a, 1991b), although INaP was never recorded in type A MveN. Interestingly, in guinea pig in vitro whole brain preparation, these plateau potentials can be triggered in some type B second-order vestibular neurons by monosynaptic EPSPs evoked by single-pulse stimulation of the ipsilateral vestibular nerve (Babalian et al., 1997). A tetrodotoxin-sensitive sodium persistent current (INaP) is present in many types of CNS neurons, notably in the cortex, the hippocampus, and the cerebellum (for a review, see Crill 1996). It comes into play at a level of membrane potential where few voltage-gated channels are activated and where input resistance is high. Its impact is, therefore, likely to be important in the MveN. In particular, in type B MveN, the sodiumpersistent current should drive the neurons toward the threshold for fast sodium spikes and could play a major role in the spontaneous activity of these cells. Moreover, preliminary data in guinea pig suggest that type B cells correspond to phasic neurons in vivo and it could facilitate a synchronous discharge of these cells. Calcium currents Calcium currents can be subdivided into low- and high-voltage-activated (LVA and HVA) currents based on their threshold for activation (Jones, 1998). In the guinea pig, it is quite difficult to show calcium conductances in type A MveN. A combination of TTX, TEA, and 4-AP is required and only small TTXresistant action potentials are displayed that are suppressed by cadmium. In contrast, some B cells display calcium-dependent low-threshold spikes (LTS; Serafin et al., 1990a, 1991a, 1991b). The LTS appears as a rebound following large hyperpolarizing current pulses applied at rest or as a burst of action potentials in response to depolarizing current pulses in hyperpolarized cells. LTS was first described in olivary cells (Llinás and Yarom, 1981) and was later shown to underlie the burst firing of a number of CNS neurons (Llinás, 1988; Huguenard, 1996). In B+LTS MveN, the LVA calcium current could confer a high sensitivity to synaptic inputs.

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In B cells, depolarizing current pulses applied in the presence of TTX, TEA, and 4-AP also reveal calciumdependent high-threshold spikes (HTS) and plateau potentials (HTP) that were both eliminated by cadmium (Serafin et al., 1991b). The functional roles of highthreshold calcium conductances, which are believed to play a role in the control of the repetitive firing of neurons, are still unknown in vestibular neurons. Potassium currents The potassium channels expressed in vestibular and auditory neurons have been reviewed by Peusner et al. (1998). In MveN, as in other neurons, spike repolarization and the AHP result from the activation of voltage-dependent (for a review, see Halliwell 1990) and calcium-dependent (for reviews, see Sah 1996; Vergara et al., 1998) potassium conductances. In the rat (Johnston et al., 1994; Dutia and Johnston, 1998, Smith et al., 2002; Patko et al., 2003) and in other species investigated (Smith and Goldberg, 1986), potassium currents appear to differ among the type A and B MveN. In type A MveN, TEA slows down repolarization, and Ca2+-free medium decreases the AHP amplitude considerably. These results suggest that the voltagegated IK conductance and the AHP presumably mediate the action potential repolarization by the calciumactivated conductance IC. In contrast, 4-AP decreases the rate of repolarization slightly and leads to a small decrease in the amplitude of the AHP. The A-like outwardly rectifying current, therefore, appears to play a minor role in cell repolarization. This contrasts with previous results in guinea pig type A neurons (Serafin et al., 1991a, 1991b) showing a more important role of this conductance. The small effect of apamin and carbachol perfusion also indicates the minor role of the small conductance (SK) type of calcium-activated potassium channel and of the IM current in cell repolarization. In type B MveN, the fast AHP is insensitive to apamin but is blocked by TEA and 4-AP. This shows that both a TEA-sensitive potassium current (IK) and a transient A-type current (IA) were involved. While an IA current was observed in rat type B MveN (Johnston et al., 1994), this was not the case in guinea pig. The delayed slower AHP was insensitive to TEA but was abolished either in a free-calcium medium or following bath application of apamin, which indicated a major involvement of SK Ca2+-activated K+ channels, as is the case in guinea pig (de Waele et al., 1993). In contrast to type A cells, when apamin is perfused in the bath, type B cells exhibit very irregular firing, which suggests that calcium-dependent potassium conductances constitute a major determinant of the regularity of their discharge in the slice. Like many other CNS neurons, type A and B MveN types exhibit a sag in membrane potential during longlasting hyperpolarizing current steps. Being sensitive

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to cesium, this rectification is likely due to a mixed sodium and potassium Ih current (Pape, 1996). In summary, rat MveN are endowed with at least five different types of potassium channels: the delayed outward rectifier (IK), a transient outward rectifier (IA), an anomalous inward rectifier (Ih), and calcium-activated potassium channels BK (IC) and SK (IAHP). The potassium channels referred to probably play key functional roles. These ionic conductances contribute to spike repolarization and afterpotentials in MVe neurons and determine the excitability of these cells following the emission of an action potential. This is, therefore, a major determinant of the sensitivity of the MveN to synaptic inputs. In vivo, a deep long-lasting AHP of type A MveN is likely to (a) slow down their activity, (b) promote a regular resting discharge, (c) decrease their sensitivity to excitatory inputs, and (d) augment the duration of their recovery period. This is very reminiscent of the characteristics of tonic second-order vestibular neurons (Precht and Shimazu, 1965; Shimazu, and Precht, 1965). In contrast, B MveN display—with a noninactivating sodium conductance maintaining them near threshold—a steeper slope in the frequency/current curve and a smaller diphasic AHP than A MveN. In vivo, these neurons could (a) be sensitive to excitatory inputs, (b) display an irregular firing resting discharge due to the synaptic noise, and (c) have a lower threshold than the A MveN. This is reminiscent of the properties of kinetic vestibular second-order neurons. In support of this hypothesis, type B neurons, when recorded in the in vitro whole brain preparation, display an irregular resting discharge (Serafin et al., 1992b; Babalian et al., 1997). Rhythmic activities Data are lacking in the rat. However, in the guinea pig, type B MveN can display spontaneous oscillatory discharges. Various pharmacological manipulations induce voltage-dependent rhythmic activities in these cells once they are slightly hyperpolarized. First, N-methyl-D-aspartate (NMDA) triggers rhythmic activity in B but not in type B+LTS MveN (Serafin et al., 1992a). These oscillations are TTX resistant and are abolished by APV, a specific NMDA antagonist, or by replacing sodium by choline. Second, perfusion with a low Ca2+/high Mg2+ medium, which interrupts synaptic transmission, also induces rhythmic activities in B but not in B+LTS MveN. These APVresistant oscillations are abolished by TTX. Finally, perfusion of apamin, a selective blocker of SK channels, induces rhythmic activities in both B and B+LTS MveN (de Waele et al., 1993). This oscillatory behavior is APV resistant and is abolished by TTX or with ouabain, a specific antagonist of the active Na+/K+ exchanger. Oscillations could be a side effect of biologically active drugs applied in a nonspecific way (perfusion through

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a cannula in vivo or in the bath in vitro). Nevertheless, rhythmic activities occur because a particular set of conductances is activated in B cells. At the very least, this pattern of activation is likely to be important for properly processing the multimodal sensory inputs converging on these cells. A working hypothesis would be that it could promote the synchronization of the discharges of the type B MveN. In addition, vestibular neurons have been shown to discharge rhythmically in phase with the limb extensors during locomotion in the guinea pig (Marlinsky, 1992). Gap junctions Electrical coupling has been demonstrated between primary vestibular afferents and second-order vestibular neurons in the adult rat (Korn et al., 1973; Wylie, 1973) and in the adult pigeon (Wilson and Wylie, 1970). The raison d’être of this coupling remains to be elucidated. In vitro, it plays a minor role, if any, at the synapses between first- and second-order vestibular neurons in the chicken (Arabshahi et al., 1997). In addition, short-latency EPSPs (<0.6 m) compatible with an electrical transmission were rarely recorded in second-order vestibular neurons following stimulation of the vestibular nerve in the in vitro whole brain of the guinea pig (Babalian et al., 1997) and were never noticed in the alert guinea pig (Serafin et al., 1999). Nevertheless, numerous neuromodulators modulate electrical coupling. From a functional point of view, gap junctions could play a major role in synchronization of the discharge MveN or in the postlesional plasticity of these neurons. Postnatal maturation Murphy and Du Lac (2001) studied in vitro the maturation of the spike generation in rat MveN. While neurons of rats younger than P12 had a mean spontaneous firing rate of 5.2 spikes/s (maximum 10 spikes/s), they fired with a mean of 9.8 spikes/s (max 25 spikes/s) between P13 and P30. During the first 2 postnatal weeks, they displayed strong adaptation to sustained steps of current. They had a limited range of input amplitude, presumably due to cumulative sodium inactivation due to improper membrane repolarization following each action potential. These cells never discharged faster than 60 spikes/s and exhibited saturation when challenged with increasing input. By P17, MveN discharged up to 200 spikes and responded linearly to a broad range of input. Before P15, the low gain of the primary vestibular afferents may result in the fact that the immature MveN remained in their narrow linear range of response. However, adaptation clearly favors a preferential detection of transient high-frequency head movements. The increasing sensitivity of the MveN during the first 6 days does not result from an increase in spike generation but

rather follows the increasing gain of the first-order vestibular neurons response. In a study in the mouse, Dutia and Johnston (1998) have showed that type A and B MveN could be individualized as early as P5. Immature type A MveN had broad spontaneous action potentials and small AHP. Immature type B MveN had thinner spontaneous action potentials and displayed a delayed, apamin-sensitive AHP. The fast AHP of the mature cell was replaced by a period of isopotentiality. Interestingly, the proportion of the two types of cells was similar to that in the adult rat and the guinea pig (about one-third of type A MveN and two-thirds of type B MveN, if one includes type C cells as type B MveN). Between P10 and P15, the AHP amplitude rose in type A cells, whereas the fast AHP appeared in type B ones. The negativity of the resting potential increased, while the rise-and-fall time of the action potentials shortened until P15. In particular, the apamin-sensitive I (AHP) appeared very early in type B cells and was a major determinant of its intrinsic rhythmicity and excitability. At P30, they were of the adult type. Clearly then, variations in the density and the kinetic of potassium conductances play a major role in MveN maturation, as demonstrated previously by Peusner et al. (1998) in chick central vestibular neurons. Functional Considerations In alert-behaving vertebrates, first-order vestibular neurons (Goldberg, 1991, 2000) and second-order neurons (Chen-Huang et al., 1997; Serafin et al., 1999) can be considered as a broad continuum of cells extending from regular tonic neurons to irregular, phasic ones. The terms phasic and tonic refer to the dynamics of the responses of these cells to the large frequency spectrum of natural head accelerations (DC to 20 Hz). In vitro recordings in a guinea pig isolated whole brain preparation (Babalian et al., 1997) revealed that central vestibular neurons formed a heterogeneous population of cells, whose intrinsic membrane properties were distributed in between two canonical types. These two types of cells, type A and type B MveN, have been described previously in slice preparation in rat and guinea pig. Type B neurons were found to be moderately regular to very irregular in the guinea pig in vitro whole brain, whereas type A cells were always regular. As stated earlier, these results, therefore, suggest that phasic irregular, second-order neurons (identified in vivo) correspond to the irregular type B MveN (identified in vitro), whereas the tonic, regular second-order MveN (identified in vivo) included both the regular type A and a minority of type B cells (identified in vitro). It was, therefore, proposed that the membrane properties of type A and B MveN contribute to determine the dynamics of these neurons in vivo (du Lac and

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Lisberger, 1995a, 1996b; Babalian et al., 1997), as suggested previously for first-order vestibular neurons. In particular, they could contribute to synchronize the discharge of type B MveN. Theoretical evidence, based on realistic biophysical models of type A and B MveN (Av-Ron and Vidal, 1999) and in vitro studies on the cellular processing of temporal information in MveN in fowl (du Lac and Lisberger, 1995a, 1995b) and guinea pig (Ris and Godaux, 2001), supports this hypothesis. However, while it is plausible that membrane properties could characterize the dynamics of the first- and second-order vestibular neurons, the exact contribution of the various subtypes of these neurons during a given behavior remains to be elucidated.

NEUROTRANSMITTERS AND NEUROMODULATORS OF CENTRAL VESTIBULAR NEURONS The pharmacological properties of vestibular neurons have been the subject of several studies over the past years. They have been the topic of various reviews (Raymond et al., 1988; Darlington et al., 1995, 1996; Smith and Darlington, 1994a, 1994b; de Waele et al., 1995; Takahashi and Kubo, 1997; Vibert et al., 1994, 1997, 1999, 2000; Vidal et al., 1996a, 1996b, 1998, 1999) and of a book (Anderson, 2000), which can be consulted to complement the present chapter. The neurotransmitters involved in the neurotransmission and neuromodulation of the central vestibular neurons are conveniently classified into three groups. The excitatory and inhibitory amino acids, aspartate, glutamate, GABA, and glycine, mediate fast synaptic events through their actions on pre- and postsynaptic ionotropic receptors. The five monoamines (histamine, dopamine, serotonin, noradrenaline, and adrenaline) and acetylcholine modulate with a slower time constant, discharge of vestibular neurons by activating metabotropic receptors linked to second messenger systems (Hille, 1992). A third group encompasses several neuroactive peptides. This section stresses that in vivo microiontophoretic applications, in vivo perfusion through chronically implanted cannula, and in vitro bath application on slice have in common that they test the effect of a drug at a population level. The changes of neural activities are the summation of its effect both on the recorded neurons and on the neighboring excitatory and inhibitory cells, which project to them. The results must, therefore, be interpreted with caution. One way of solving this problem on in vitro preparation is to reversibly block synaptic transmission with high Mg2+–low Ca2+ solution or tetrodotoxin. In these con-

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ditions, the postsynaptic effect of a given compound can be studied on recorded neurons, independently of its impact on the interneurons, which synapse the cell under scrutiny. The vast majority of the studies quoted in this section were performed in the rat. Nevertheless, studies performed in the cat and the guinea pig will be included to present a comprehensive view of the neuropharmacology of the vestibular neurons. Finally, it should be mentioned that, apart from subtle differences, type A and B MveN could not be differentiated through their sensitivity to any of the drugs tested so far.

Excitatory and Inhibitory Amino Acids The amino acids can be subdivided into excitatory amino acids aspartate and glutamate and into inhibitory amino acids GABA and glycine. The ionotropic and metabotropic receptors of the amino acids are known under the name of their main specific agonists (Nakanishi, 1992; Pin and Duvoisin, 1995). Excitatory Amino Acid Receptors All types of excitatory amino acid receptors are expressed by vestibular nuclei neurons, including the mGluR1, mGluR2 (Fig. 4), mGluR5, and mGluR7 subtypes of the metabotropic receptors (in the rat, see Shigemoto et al., 1992; Ohishi et al., 1995; Neki et al., 1996; Rabbath et al., 2002; Devau et al., 2003; Horii et al., 2001). In situ hybridization studies have been used to investigate the various subunits of the ionotropic receptors. Rat vestibular nuclei displayed a high density of GluR1, GluR2/3, and GluR4 subunits of the AMPA receptor (Fig. 3), a high density of the R1 and R2C

FIGURE 3 Double GFAP and GluR2 immunoflourescence fluorescence staining of the lateral vestibular nucleus. Neuron of the adult rat lateral vestibular nucleus exhibiting GluR2-immunopositive labeling (soma colored in red). Note that this neuron is surrounded by numerous astrocytic processes (colored in yellow). Zoom : ×1000.

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subunits, and a lower density of the R2B and R2D subunits of the NMDA receptors (Petralia and Wenthold, 1992; de Waele et al., 1994; Watanabe et al., 1994; King et al, 2002; Chen et al., 2003). NMDA receptors are colocalized with AMPA receptors in most rat vestibular neurons (Chen et al., 2000). Hence crossmodulation between NMDA and AMPA receptors is likely to occur during glutamate-mediated excitatory synaptic transmission and could play a role in synaptic plasticity (see later). In good agreement with these morphological data, several in vitro electrophysiological studies reported that vestibular neurons were responsive to the various agonists and antagonists of the AMPA/kainate, NMDA, and trans-ACPD receptors (for reviews, see Gallagher et al., 1992; de Waele et al., 1995; Vidal et al., 1996a). The effects of AMPA, kainate, NMDA, and trans-ACPD are largely mediated by postsynaptic receptors, as shown by their persistence during bath application of TTX and in a high Mg2+/low Ca2+ solution (in guinea pig, see also Darlington et al., 1995). Pharmacological Analysis of Excitatory Amino Acid-Mediated Synaptic Transmission in the MVe An excitatory amino acid (glutamate and/or aspartate) most probably mediates synaptic transmission between (a) first-order and second-order vestibular neurons (for reviews, see Raymond et al., 1988; Gallagher et al., 1992; Yamanaka et al., 1997), (b) second-order vestibular neurons and contralateral abducens motor neurons, and (c) excitatory second-order vestibular neurons and some of the spinal motor neurons (Dieringer, 1995). As demonstrated in the frog (Cochran et al., 1987; Dieringer, 1995), it is also likely that several afferents of the vestibular nuclei neurons, including the spinal fibers and the excitatory commissural pathways, use glutamate and/or aspartate mediated transmission (Chun et al., 2003). The contribution of NMDA receptors to the synaptic transmission between first- and second-order vestibular neurons is still under scrutiny (in the rat, Lewis et al., 1989; Doi et al., 1990; Kinney et al., 1994; Takahashi et al., 1994; in the frog, Cochran et al., 1987; Straka et al., 1995b). In this regard, the isolated in vitro whole brain of the guinea pig (Babalian et al., 1997) is useful as it allows stimulation of first-order vestibular neurons without the problem of current spread to adjacent groups of cells. In this preparation, CNQX (a selective antagonist of AMPA/kainate receptors) suppressed a major part of the field potentials and the EPSPs evoked following stimulation of the eighth nerve. In contrast, APV (an antagonist of NMDA receptors) abolished in about half of the recorded cells a small and variable portion of field potentials or EPSPs, which persisted

following CNQX perfusion. In addition, a previous study by Straka et al. (1995b) in isolated frog’s brain stem indicated that the large vestibular afferents (presumably the kinetic irregular ones) activate NMDA receptors in second-order vestibular neurons. Clearly then, NMDA receptors are involved in the monosynaptic transmission between first- and secondorder vestibular neurons. However, their exact contribution remains to be determined. Indeed, NMDA receptors are subjected to a voltage-dependent block by extracellular Mg2+ (Ascher and Nowak, 1988). Hence, the minor contribution of NMDA receptors recorded in vitro on preparations could become a major one in vivo when MveN are more depolarized (see later). Finally, NMDA and trans-ACPD receptors are present at the presynaptic level in the vestibular nucleus (in rat, Gallagher et al., 1992; in guinea pig, Darlington and Smith, 1995). They could be expressed at the terminal arborization of first-order neurons, as several NMDA and trans-ACPD receptor subunits are expressed by rat vestibular ganglion cells (Doi et al., 1995; Safieddine and Wenthold, 1997). The functional contributions of these receptors remain to be determined. Functional Roles of NMDA Receptors in Vestibular Nuclei: Correlation with in Vivo Data In vivo, NMDA receptors of the central vestibular neuron receptors play a major role in the maintenance of the resting discharge of cells. Indeed, unilateral perfusion of APV in vestibular nuclei in alert unrestrained guinea pigs induces a massive postural and oculomotor syndrome, mimicking unilateral labyrinthectomy (de Waele et al., 1990). It also disables the neural integrator, which processes the velocity signal encoded by the firstorder vestibular neurons into a position signal necessary for stabilizing the eyes. Finally, NMDA antagonists impair the cat vestibuloocular reflex triggered by lowfrequency head rotations (Priesol et al., 2000). These data show that the voltage-dependent block of NMDA receptors of the central vestibular neurons by Mg2+ is compensated by the alert-behaving preparation. Different factors could be at play. First, the high resting discharge of the first-order neurons could sufficiently depolarize the central vestibular neurons to prevent the Mg2+. Second, glycine, a coagonist of the strychnineinsensitive site of the NMDA receptors (for a review, see Wood, 1995), is colocalized in frog and rat with glutamate in the largest afferent fibers (Reichenberger and Dieringer, 1994; Reichenberger et al., 1997). As shown earlier, these fibers underlie the NMDA-mediated response in vestibular neurons in the frog (Straka et al., 1995a, 1995b), the guinea pig (Babalian et al., 1997), and, most likely, the rat (Doi et al., 1990; Kinney et al., 1994). Hence, corelease of glutamate and glycine by large

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irregular fibers could potentiate postsynaptic NMDA receptors. This is a plausible scenario as the strychnineinsensitive site is probably not saturated in vivo (Wood, 1995). In addition, a specific agonist (D-serine) and antagonist (7-chlorokynurenate) of the strychnineinsensitive binding site of NMDA receptors, when perfused chronically in a vestibular complex of alert, unrestrained guinea pigs (Bénazet et al., 1993), induced opposite asymmetries of the vestibuloocular reflex and a reversible postural syndrome. Excitatory Amino Acid Receptors and Vestibular Plasticity Postnatal development Changes in the expression of the AMPA receptor subunits GluR1–4 and of the NMDA receptor subunits NR1 and NR2A–D were investigated in the developing rat medial and lateral vestibular nuclei. During postnatal development, glutamatergic receptor subunits were expressed differentially in the vestibular nuclei. The level of expression of GluR1, GuR4, and NR1 subunits was higher in the developing brain as compared to the adult. A gradual increase in GluR2/3, NR2A, NR2B, and NR2C levels of expression in the medial and lateral vestibular nuclei during the first 3 weeks of postnatal development has been observed. The differential expression of AMPA and NMDA receptor subunits in immature vestibular neurons is consistent with changes in glutamate receptor properties. This may be related to the postsynaptic regulation of receptor subunits associated with the synaptic plasticity of the vestibular neuron connections during specific sequences of postnatal development (Sans et al., 2000). Postlesional plasticity NMDA receptors play a key role in various types of synaptic plasticity (for a review, see, Nakanishi, 1992), which raises the question of their role in vestibular plasticity. Vestibular compensation is a remarkable model of plasticity of the adult central nervous system. Indeed, the postural and oculomotor syndromes observed after unilateral labyrinthectomy largely recover in about a week in rodents [Hamman et al. (1998) in the rat and Smith and Curthoys (1989) and Dieringer (1995) for reviews across species]. At the acute stage, these syndromes result from an asymmetry of the mass discharge between intact and deafferented vestibular nuclei: the deafferented neurons are silent, whereas the spontaneous activity of contralateral medial ones is increased by 15%. At the compensated stage, deafferented vestibular neurons recover a quasinormal resting activity (Ris et al., 1995) and this new balance plays a key role in the compensation process. As NMDA receptors are instrumental in the maintenance of the resting dis-

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charge of central vestibular neurons, they could be strongly involved in the recovery of their resting discharge after unilateral labyrinthectomy. Several experimental studies support this hypothesis [Smith and Darlington (1988) and de Waele et al. (1990) in the guinea pig; de Waele et al. (1994) and Kitahara et al. (1995) in the rat], while in the frog they did not reveal the expected NMDA receptor supersensitivity (Dieringer, 1995). Functional plasticity Habituation and adaptation of the vestibuloocular and vestibulospinal reflexes are believed to be mediated by long-term modifications of synaptic efficacy occurring at various levels of the vestibuloocular and/or vestibulospinal pathways (for reviews, see Kawato and Gomi, 1992, Cohen et al., 1992; du Lac et al., 1995c). Extracellular field recordings suggest that LTP and LTD occur in rat vestibular nuclei (Racine et al., 1986; Capocchi et al., 1992; Grassi et al., 1996, 1999; Grassi and Pettorossi, 2000, 2001; Pett Orossi et al., 2003; Puyal et al., 2003) and involvement of the NMDA receptors is likely. Interestingly, intracellular recordings of guinea pig MveN, either in vivo or in the in vitro whole brain, tend to indicate that these modifications do not take place at the synapse between first-and second-order vestibular neurons. Inhibitory Amino Acid Receptors of Vestibular Nuclei Neurons Anatomical studies GABA and glycine are the most prevalent inhibitory transmitters in the CNS (in the rat, Sivilotti and Nistri, 1991; Sato et al., 1991). GABA receptors are subdivided into two groups: ionotropic GABAA receptors include chloride ion channels and GABAB metabotropic receptors are associated with second messenger systems (for review, see Misgeld et al., 1995). Glycine receptors are ionotropic receptors quite similar to GABAA ones (for review, see Betz et al., 1994). Vestibular nuclei are densely innervated by GABAergic and glycinergic afferent fibers’ (in rat, Rampon et al., 1996). Not surprisingly, vestibular neurons are also endowed with GABAA (Fig. 4A), pre- and postsynaptic GABAB (Fig. 4B) (in the rat, Holstein et al., 1992), and glycinergic receptors (Eleore et al., 2003). Furthermore, about a third of rat MveN are GABAergic neurons (de Waele et al., 1994). These inhibitory cells encompass the inhibitory type II interneurons (Shimazu and Precht, 1966) involved in commissural inhibition and the inhibitory secondorder vestibuloocular and vestibulospinal neurons (Graf et al., 1997). Electrophysiological studies In vitro studies in slices confirm these morphological findings. Extracellular recordings in MveN reveal inhibition through GABAB

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A

B

FIGURE 4 Bright-field photomicrograph showing (A) positive GABAA mRNA labeling in the lateral vestibular nucleus (zoom : ×400) and (B) GABAB immunoreactivity (R1 subunit) in the lateral vestibular nuclear complex of a control adult rat (zoom : ×200).

and GABAB receptors (Dutia et al., 1992; Him et al., 2001). As expected, intracellular recordings in guinea pig (Vibert et al., 1995b, 1995c) in a high Mg2+/low Ca2+ solution or in the presence of TTX reveal that both type A and B MveN are inhibited by GABA, muscimol (a specific GABAA agonist), and baclofen (a specific GABAB agonist). However, in normal saline, while GABA hyperpolarizes several MveN, others are depolarized. Once TTX is added to the bath, these depolarized cells become hyperpolarized. These results in normal saline can be explained as follows. (a) Inhibitory interneurons are tonically active in the slice and inhibit some of the recorded MveN. (b) GABA and muscimol perfusion interrupt their spontaneous discharge and consequently depress the inhibition they exert on the recorded cell. This leads to a disinhibition of the MveN. (c) This disinhibition is large enough in some of the MveN to overcome the direct inhibitory effect of GABA and muscimol. Consequently, GABA application results in a net depolarizing effect. Once TTX is added to the bath, the disinhibition is suppressed and the direct inhibition is revealed. Altogether, these results show that both MveN and the local type II inhibitory interneurons express postsynaptic GABAA receptors. A bath application of glycine also inhibits MveN [Lapeyre and de Waele (1995) in the guinea pig]. This inhibition is suppressed by strychnine but persists in a high Mg2+/low Ca2+ solution. This implies that MveN carry postsynaptic, strychnine-sensitive glycinergic receptors. Glycine, therefore, has two opposite effects in MveN: (1) it augments the depolarizing effect of glutamate through the glycinergic, strychnine-insensitive modulatory site of the NMDA receptors (see preceding paragraph) and (2) glycine hyperpolarizes these cells through their strychnine-sensitive receptors.

Functional Considerations Glycine and GABA mediate the effect of at least three inhibitory afferents of the vestibular neurons: • Cerebellar Purkinje cells use GABA as their main neuromediator (Sato and Kawasaki, 1991; de Zeeuw and Berrebi, 1996). • A commissural pathway links the two medial vestibular nuclei in mammals through local inhibitory interneurons (type II neurons). Type II interneurons are activated by contralateral excitatory MveN (Shimazu and Precht, 1966). Type II interneurons are both GABAergic and glycinergic (Precht et al., 1973; Furuya et al., 1991). • Some GABAergic inferior olive neurons project to the contralateral vestibular complex (Matsuoka et al., 1983). In addition, as shown earlier, glycine could be colocalized with glutamate and/or aspartate in some of the large-diameter first-order vestibular neurons (in frog, Straka et al., 1995a, 1995b). GABAA and GABAB receptors of the vestibular neurons play a key role in the control of gaze in the rat (Reber et al., 1996; Kato et al., 2003; Sun et al. 2002). For instance, commissural inhibition mediated by GABAergic interneurons is a key component of the velocity storage integrator included in vestibulooculomotor pathways and is a determinant in the regulation of HVOR gain (Galiana and Outerbridge, 1984; Katz et al., 1991). Perfusions of vestibular nuclei with agonists or antagonists of the GABAA or GABAB receptors provoke a postural and oculomotor syndrome and alter the gain of the horizontal vestibuloocular reflex. Systemic injections of baclofen, a GABAB agonist, disable the velocity storage integrator (in rat, Cohen et al., 1987; Niklasson et al., 1994). Finally, GABAergic cerebellar

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Purkinje cells are known to be instrumental in the adaptation and habituation of the horizontal vestibuloocular reflex.

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inferior olive to the lateral vestibular nuclei, is depressed following a systemic injection of atropine, a muscarinic antagonist (Matsuoka et al., 1985). Behavioral Evidence

Cholinergic Neurotransmission Two types of cholinergic receptors have been described in the central nervous system: nicotinic and muscarinic. Nicotinic receptors are ionotropic receptors, which include a cation channel. Metabotropic, muscarinic receptors act through G proteins and second messenger systems. Several different subtypes of both nicotinic and muscarinic receptors had been individualized (for reviews, see Hosey, 1992). Anatomical Evidence Only a few vestibular neurons appear to be cholinergic in rats (for reviews, see Zanni et al., 1995; Fukushima et al., 2001; de Waele et al., 1995). In rabbits, cholinergic second-order vestibular neurons project to the flocculus, the nodulus, and the dorsal cap of the inferior olive. Cholinergic vestibulospinal neurons have been detected in the rat, mainly in the lateral vestibular nuclei (Jones et al., 1986). In contrast, nicotinic and muscarinic receptors are numerous in all vestibular nuclei, particularly in the medial vestibular nucleus (in rat, Zanni et al., 1995; Dominguez del Toro, 1994). Moreover, vestibular nuclei display choline acetyltransferase (ChAT) activity, especially in the rat medial vestibular nucleus (Burke and Fahn, 1985). The origin of the cholinergic innervation of vestibular nuclei remains to be determined. Afferent cholinergic neurons could be located within vestibular nuclei or in pedunculopontine formation, tegmental dorsal nuclei neurons, and/or the contralateral inferior olive. Electrophysiological Evidence Nicotinic and muscarinic agonists depolarize MveN. Nicotinic or muscarinic antagonists reversibly suppress this depolarization. These effects are mediated by postsynaptic receptors, as they persist in the presence of tetrodotoxin or during perfusion of a low Ca2+/high Mg2+-containing solution. Both muscarinic and nicotinic receptors regulate the rat MveN spontaneous activity on slice (in rat, Ujihara et al., 1989; Phelan and Gallagher, 1992). In vivo, systemic and microiontophoretic injections of acetylcholine, physostigmine (an inhibitor of acetylcholine esterase), and muscarinic agonists result in an excitation of the lateral and medial vestibular neurons. In addition, synaptic transmission between first-order and second-order vestibular neurons was facilitated by cholinergic agonists and not facilitated by muscarinic antagonists. Finally, an excitatory pathway, linking the

Unilateral perfusion of the vestibular complex with muscarinic agonists in alert rodents induces a postural and oculomotor syndrome, which is the mirror image of the syndrome induced by unilateral labyrinthectomy (i.e., it results in a net excitatory effect of the vestibular neurons). Several studies also suggest that acetylcholine plays a key role in the compensation of vestibular deficits (Kitahara et al., 2001). Belladonna alkaloids, which are known to have anticholinergic properties, are the oldest agents used for the prophylaxis of motion sickness (de Waele et al., 1995). Muscarinic receptors are also involved in the cerebellar control of the cat vestibulospinal reflex gain (Andre et al., 1995). In summary, several lines of evidence suggest that cholinergic innervation of the vestibular nuclei has high behavioral consequences for the stabilization of gaze and posture. Why some second-order vestibular neurons are glutamatergic whereas others are cholinergic remains to be determined.

Modulation of Central Vestibular Neurons by Monoamines The three catecholamines (dopamine, noradrenaline, and adrenaline), serotonin, and histamine are key modulators of the activities of large assemblies of neurons, particularly in relation with the different states of vigilance. Furthermore, dysfunctions of the aminergic systems are involved in major neurological disorders, such as schizophrenia or Parkinson’s disease. The monoaminergic modulation of the central vestibular system is likely to be important from a functional point of view. Two arguments support this hypothesis: (1) the turnover rates of monoaminergic metabolites in the rat vestibular nuclei are important (Cransac et al., 1996) and (2) several agonists and antagonists of the monoaminergic receptors are used successfully in clinical medicine to treat vertigo, motion sickness (for review, see Rascol et al., 1995), and to improve recovery following vestibular deficits (for review, see Smith and Darlington, 1994b). The Histaminergic System Central histaminergic pathways In mammals, histaminergic neurons are located in the tuberomammillary nucleus of the posterior hypothalamus. They play an essential role in the control of circadian rhythms. Three types of metabotropic receptors of histamine have been indentified (Arrang et al., 1995). Postsynaptic H1 and H2 receptors are coupled positively to phospholipase C

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and adenylate cyclase, respectively. These two receptors, when activated, have excitatory effects. H3 receptors are often but not exclusively presynaptic. In this situation, they are part of a negative feedback loop, which controls histamine synthesis and release. They could also modulate neurotransmitter release in nonhistaminergic axon terminals. Histaminergic modulation of the vestibular system Morphological studies show that histaminergic neurons project to all rat vestibular nuclei (Takeda et al., 1987; Pillot et al., 2002), with the largest projection in medial and superior nuclei (in cat, Tighilet and Lacour, 1996). Guinea pig vestibular neurons also express H1 and H2 binding sites (Vizuete et al., 1997). In vitro recordings on slice demonstrate that histamine mainly depolarizes MveN (in the rat, Phelan et al., 1990; Wang and Dutia, 1995), whatever their type [A, B, and B+LTS neurons, Serafin et al. (1993) in the guinea pig]. In the rat, depolarization is mediated by H1 and H2 receptors, while it is only mediated by H2 receptors in the guinea pig (Serafin et al., 1993). In vivo studies indicate that medial and lateral vestibular nuclei neurons are both inhibited or excited by histamine or histaminergic agonists. In particular, the oculomotor and postural syndromes induced, in guinea pig, by unilateral perfusion of the H3 agonist strongly suggest that (a) histaminergic fibers projecting to the vestibular nuclei carry presynaptic H3 receptors (Yabe et al., 1993) and (b) vestibular nuclei neurons are tonically excited, in the awake state, by the histaminergic input. In rat, histaminergic modulation appears to play an important role during compensation for a unilateral labyrinthectomy (Horii et al., 1993) or in response to multisensory conflicts inducing motion sickness (Takeda et al., 1993). It is, therefore, not surprising that histaminergic ligands are widely used in patients for the symptomatic treatment of vertigo and motion sickness (Rascol et al., 1995). In this regard, H3 antagonists are particularly promising. In contrast to standard histaminergic agents, they do not induce drowsiness, an unwanted side effect of histaminergic drugs (Lin et al., 1990). The Serotoninergic System Central serotoninergic pathways Serotoninergic cells form eight separate clusters within brain stem reticular formation, which project diffusely over the entire CNS. This explains the variety of behavior they modulate. Up until now, 10 serotoninergic receptor subtypes have been described (for review, see Zifa and Filion, 1992). They are subdivided into four groups: three groups of metabotropic receptors (5-HT1, 5-HT2, and 5-HT4) and one group of ionotropic receptors (5HT3). Three subtypes of 5-HT1 receptors (5-HT1A, 5-HT1B,

and 5-HT1D) have very high affinity for serotonin and are coupled negatively with adenylate cyclase. The three subtypes of 5-HT2 receptors (HT2A, 5-HT2B, and 5-HT2C) are coupled positively with phospholipase C. Their activation increases the intracellular calcium concentration. The 5-HT1 and 5-HT2 receptors can be localized both pre- and postsynaptically. The 5-HT4 receptors are coupled positively to adenylate cyclase and are mostly localized postsynaptically. The 5-HT3 receptors include a cation-selective channel, induce short-lasting depolarization, and could be mainly localized presynaptically. They are believed to facilitate the release of various neurotransmitters, including serotonin itself. Serotoninergic modulation of the vestibular syste All vestibular nuclei are innervated by serotoninergic fibers, which probably originate in the dorsal raphe nucleus (Giuffrida et al., 1991). 5-HT1A, 5-HT1B, and 5HT2 receptors have been shown to be expressed by vestibular neurons in the rat (Pazos and Palacios, 1985; Wright et al., 1995; Kia et al., 1996). Serotonin can both accelerate and decrease the MveN spontaneous activity in rat slices (Johnston et al., 1993), with a predominance of excitatory effects. Intracellular recordings in guinea pig brain stem slices (Vibert et al., 1994) show that 80% of the MveN are depolarized by serotonin. Interestingly, serotonin directly activates postsynaptic receptors in type B MveN, whereas the excitation of type A MveN is mostly indirect. The depolarizing effect of serotonin can be reproduced (Johnston et al., 1993) by α-methyl-serotonin (a specific agonist of 5-HT2 receptors). However, it is only partly blocked by ketanserin (an antagonist of 5-HT2 receptors). Moreover, the depolarization is accompanied by a decrease of membrane resistance, as reported previously in the rat CNS (Andrade and Chaput, 1990). The serotoninergic binding site remains to be determined. In 15% of type A and type B MveN, bath application of serotonin induces a hyperpolarization, which is likely mediated through 5-HT1A receptors. In vivo, a microiontophoretic injection of 5-HT induces a short hyperpolarization, followed by a large depolarization, in rat lateral vestibular nucleus neurons. Medial and superior vestibular nuclei neurons exhibit (a) excitatory responses likely mediated through 5-HT2 receptors (Jeong et al., 2003) (b) inhibitory responses probably mediated through 5-HT1A receptors, and (c) biphasic responses. Finally, an intracerebroventricular injection of serotonin increases the gain of the rat horizontal vestibuloocular reflex (Ternaux and Gambarelli, 1987). The functional raison d’être of serotoninergic modulation remains to be determined. In vivo studies suggest that serotonin is more likely involved in setting the dynamic properties of the

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vestibular system (Li Volsi et al., 2001; Åmano et al., 2001) rather than in controlling the resting discharge of the vestibular neurons (i.e., the static reflexes). The Dopaminergic System Central dopaminergic pathways Three dopaminergic pathways are well described in the central nervous system (for review, see Civelli et al., 1993). The nigrostriatal pathway is involved in locomotion and movement. The mesocorticolimbic pathway originates in the ventral tegmental area and projects to limbic structures involved in the regulation of emotional states. The tuberoinfundibular pathway originates in the hypothalamus and controls hypophyseal activities. Smaller dopaminergic cell groups have also been localized in the olfactory bulb, the retina, the thalamus, and the dorsal motor nucleus of the vagus nerve. Five distinct subtypes of metabotropic receptors have been identified. They are segregated into two classes on the basis of their pharmacological and structural homologies with the D1 and D2 binding sites. The “D1like” receptors include the D1 and D5 subtypes. They are coupled positively with adenylate cyclase and can be localized both pre- and postsynaptically. Presynaptic receptors usually stimulate the release of various neurotransmitters. The “D2-like” receptors include the D2, D3, and D4 subtypes. In most cases, they seem to be coupled negatively with adenylate cyclase and can be localized both pre- and postsynaptically. The presynaptic receptors inhibit the release of several neurotransmitters, whereas postsynaptic “D2-like” receptors generally hyperpolarize neurons by acting on potassium conductances (Vallar and Meldolesi, 1989). Dopaminergic modulation of the vestibular system Dopaminergic innervation of vestibular nuclei was not observed with anatomical methods (Kohl and Lewis, 1987). However, morphological studies detected D2 receptors in the rat vestibular complex, mostly in the MVE (Yokoyama et al., 1994). This discrepancy remains to be clarified. Dopamine depolarizes MveN in the rat (Gallagher et al., 1992) and the guinea pig (Vibert et al., 1995a), whatever the type (type A, B, or B+LTS MveN). In the guinea pig, depolarization is accompanied by an increase in membrane resistance and is mediated through “D2like” receptors. This atypical depolarization is the result of an indirect effect on the inhibitory interneurons contacting the recorded cells (as for the depolarization induced by GABA described earlier). Indeed, once synaptic transmission is blocked, dopamine has a weak postsynaptic, hyperpolarizing action on all types of MveN. The interpretation is that dopaminergic agonists, by acting on presynaptic “D2-like” receptors, inhibit in

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normal medium a spontaneous, tetrodotoxin-resistant release of GABA. In support of this hypothesis, following continuous perfusion of bicuculline in the bath, the depolarizing effects of dopamine are replaced by hyperpolarizing effects, as in synaptic uncoupling conditions. In vivo, systemic injections of L-DOPA increase the resting activity of vestibular neurons, and microiontophoretic application of dopamine modulates the discharge of these cells. Dihydroergocristine, a nonspecific dopaminergic agonist, reduces the nystagmus induced by unilateral labyrinthectomy in the guinea pig. This could explain why vestibular compensation is shortened by systemic injections of “D2-like” dopaminergic agonists (for review, see Vibert et al., 1995a). The Noradrenergic System Central noradrenergic pathways A large part of the noradrenergic neurons are localized in the locus coeruleus. They project to the forebrain, the cerebellum, and the dorsal half of the brain stem. More ventral groups of noradrenergic neurons project to the ventral brain stem and the hypothalamus. Noradrenaline acts at both pre- and postsynaptic levels through metabotropic receptors. Noradrenaline modulation is thought to increase the signal-to-noise ratio of amino acidmediated synaptic transmission in rat (Woodward et al., 1991) and to regulate vigilance together with acetylcholine and serotonin (McCormick and Wang, 1991). Ten types of adrenergic receptors have been defined, which can mostly be localized at both pre- and postsynaptic levels. They are subdivided in three groups, i.e., α1, α2, and β receptors (Bylund et al., 1994; Nicholas et al., 1996). The α1 receptors encompass four subtypes of receptors (α1A to α1D); they increase intracellular calcium concentration through activation of a phospholipase C. Postsynaptic α1 receptors also inactivate voltage-dependent potassium channels (McCormick and Wang, 1991), which result in increasing neuronal excitability. The α2 receptors encompass three subtypes of receptors coupled negatively with adenylate cyclase. Presynaptic α2 receptors decrease transmitter release; the postsynaptic ones activate potassium channels, which decrease neuronal excitability. Finally, the β receptors encompass three subtypes of receptors (β1 to β3) coupled positively with adenylate cyclase. Presynaptic β receptors stimulate transmitter release and postsynaptic ones usually depolarize the neurons (for review, see Bylund et al., 1994). Noradrenergic modulation of the vestibular system Immunohistochemical tracing has revealed that noradrenergic neurons of the locus coeruleus project over all the vestibular complex, with a predominance in the rat superior and lateral vestibular nuclei (Schuerger

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and Balaban, 1993, 1999). Morphological studies have also revealed that vestibular neurons were endowed with β and α2 receptors. β receptors are abundant in the lateral and superior subnuclei and α2 receptors in the MVE (Rosin et al., 1996; Talley et al., 1996). In situ hybridization methods have also demonstrated α1, α2A, and α2C receptor subtypes in all vestibular nuclei (for review, see de Waele et al., 1995). In vitro recordings in guinea pig brain stem slices confirm these anatomical data (Vibert et al., 1994). Noradrenaline depolarizes half of MveN and decreases their membrane resistance. Type A MveN are less sensitive than type B or B+LTS MveN. However, noradrenaline hyperpolarizes one-fifth of the MveN. Isoproterenol (a β receptor agonist) depolarizes about 60% of MveN and decreases their membrane resistance. L-Phenylephrine (a α1 receptor agonist) also depolarizes about 60% of MveN but increases their membrane resistance. Finally, clonidine (a α2 receptor agonist) hyperpolarizes most MveN and decreases their membrane resistance. While the clonidine effect is direct (it persisted in conditions of synaptic uncoupling), most of the depolarizing responses are indirect (it was modified when synaptic transmission is blocked). For instance, α1-mediated effects could not be shown in synaptic uncoupling conditions, indicating that these receptor are probably presynaptic in the vestibular nuclei. In vivo, rat lateral and superior vestibular nuclei neurons are inhibited by noradrenaline through the activation of α2 receptors (Licata et al., 1993). However, microiontophoretic injections of noradrenaline in the vestibular nuclei of cerebellectomized cat increase the lateral vestibular nucleus neurons firing rate (see de Waele et al., 1995). It is also interesting that vestibular and cervical proprioceptive stimulations modulate locus coeruleus neuron activity (Pompeiano et al., 1990). Furthermore, noradrenergic agonists and antagonists have been shown to modify the dynamic properties of vestibulospinal and vestibuloocular reflexes (for review, see Pompeiano, 1989; Pompeiano et al., 1994). The noradrenergic system is, therefore, likely to regulate the adaptive capabilities of these reflexes (McElligott and Freedman, 1988; Nishii Ke et al., 2001). Functional Speculations Histamine and dopamine modulate the excitability of both type A and B MveN. Histamine is likely to modulate the activity of vestibular neurons according to the state of vigilance. Dopamine could control tonic inhibition, which controls the vestibular nuclei activity. In particular, dopamine could be crucial in regulating the commissural pathways linking the two vestibular nuclei and, in turn, the processing of ves-

tibular information and the resting discharge of the vestibular neurons (static reflexes). Serotonin and noradrenaline are more potent on type B and B+LTS neurons. Hence, they possibly modulate the dynamic properties of the vestibular system. Serotonin may increase the responsiveness of the vestibular system to external stimulations and could be linked to arousal Finally, noradrenaline would be fitted to tune the vestibuloocular and vestibulospinal reflexes according to various task requirements.

Neuropeptides in Central Vestibular Networks Somatostatin (or SRIF), the opioid peptides, adrenocorticotropin (ACTH), and substance P modulate the activity of central vestibular neurons through specific, metabotropic receptors (for reviews, see Balaban et al., 1989; de Waele et al., 1995). Vestibular neurons have also been shown to be sensitive to specific growth factors, including adult animals. However, large species differences are observed among mammals concerning the neuropeptides, their receptors, and their effect (see Gehlert and Gackenheimer, 1997). Somatostatin Somatostatin acts as a neurotransmitter in the central nervous system. Five types of somatostatin receptors, SST1–5, have been identified. They modulate neuronal activity through cAMP, G protein, Ca2+, and/or K3pl channels (for review, see Reisine and Bell, 1995). Somatostatin-immunoreactive cell bodies and fibers are present in all vestibular nuclei, with predominance in periventricular medial vestibular nuclei. High-affinity somatostatin SST3 (Thoss et al., 1995) and SST4 (Selmer et al., 2000) receptors have also been detected in vestibular nuclei of the rat. Intraventricular injection of somatostatin induces body rotations around the longitudinal axis (barrel rotations) that is blocked by intrasystemic injection of antimuscarinic drugs (a suggestion of its indirect effect). An in vivo microiontophoretic injection of somatostatin depresses the resting discharge of rabbit lateral vestibular neurons. In addition, Purkinje cells, which inhibit vestibular neurons, are somatostatin immunoreactive and may release somatostatin together with GABA (for reviews, see Balaban et al., 1989). Finally, somatostatin-immunoreactive neurons can be found in the vestibular ganglion cells of rabbits (Won et al., 1996), which suggests that somatostatin might regulate transmission between first- and second-order vestibular neurons. Opioid Peptides Three classes of endogenous opioid peptides have been individualized: enkephalins, β-endorphins, and

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dynorphins. The δ (OP1), κ (OP2), and μ (OP3) receptors are activated preferentially by enkephalins, dynorphins, and β-endorphins, respectively. In the brain, opiate receptors often activate inwardly rectifying potassium channels through adenylate cyclase inhibition or block voltage-dependent calcium channels. As a rule they are mostly inhibitory (for review, see Dhawan et al., 1997). Enkephalin-immunoreactive cell bodies and enkephalin terminal endings have been described in the vestibular nuclei (in rat, Zanni et al., 1995). Preproenkephalin (the precursor of Met- and Leuenkephalin) mRNA-positive cells were also reported in medial and lateral vestibular nuclei. Interestingly, the medial vestibular nucleus contains the highest density of enkephalinergic neurons of all the central nervous system. Dynorphin-immunoreactive sites have also been reported in lateral and medial vestibular nuclei. Finally, rat vestibular neurons expressed both μ and δ receptors, but only few κ receptors (Mansour et al., 1994; Zastawny et al., 1994). The newly described, endogenous opioid peptide orphanin FQ (or nociceptin) has also been detected in central vestibular nuclei (Neal et al., 1999). In vitro, morphine (a selective agonist of μ receptors), Met-enkephaline, and [D-Ala2] Leu-enkephalin (a selective agonist of δ receptors) increase the activity of about a third of the medial vestibular neurons (Carpenter and Hori, 1992; Lin and Carpenter, 1994) by activation of postsynaptic receptors. Naloxone (a specific antagonist of μ and δ receptors) suppresses these excitatory effects and the activity increase resulting from bath application of acetylcholine. It was, therefore, hypothesized that cholinergic and opioid receptors could interact in vestibular neurons. One study demonstrated that orphanin FQ (nociceptin) was able to modulate vestibular function on both in vitro and in vivo preparations (Sulaiman et al., 1999). In vivo, microiontophoretic injections of morphine and enkephalins mainly increase the discharge of medial vestibular neurons (Iasnetsov and Pravdivtsev, 1986), whereas Leu- and Met-enkephalin decrease the resting discharge of lateral vestibular nuclei neurons. Met-enkephalin is believed to be a neurotransmitter in the cerebellovestibular pathways like somatostatine. In contrast, Leu-enkephalin is usually considered a neuromodulator of vestibular function. The very high density of enkephalinergic neurons in vestibular nuclei could be explain by the existence of a built-in defense system against motion sickness and vertigo: naloxone (an opiate antagonist) enhances the incidence of motion sickness (for review, see de Waele et al., 1995).

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Substance P and Tachykinins Three types of endogenous tachykinins have been identified in the central nervous system: substance P, neurokinin A, and neurokinin B., which preferentially activate the NK1, NK2 and NK3 receptors, respectively. All these receptors are coupled to G proteins and their effects are likely mediated through activation of the phophatidylinositol–Ca2+ second-messenger system (for review, see Regoli et al., 1994). Substance P-immunoreactive fibers and terminal endings (as well as a few immunoreactive neurons) have been detected within the vestibular nuclei, predominantly in the caudal medial and inferior vestibular nuclei (in the guinea pig, Vibert et al., 1996). These fibers could originate from the brain stem reticular formation and from the vestibular nerve itself. Numerous first-order vestibular neurons are substance P immunoreactive in the frog, rabbit, guinea pig, cat, squirrel monkey, and human (Felix et al., 1996). Because these neurons innervate the utricle and the saccule in rabbits and the base of the ampullar crests and the peripheral part of the otolithic maculae in guinea pigs, substance P is probably colocalized with glutamate in part of the thinnest (tonic, regular) vestibular afferents (Usami et al., 1993). In this context, it is curious that in situ hybridization studies have demonstrated only a few NK-1 receptors in the medial vestibular nucleus of rats (Maeno et al., 1993). However, unidentified “substance P receptors” were detected in all rat vestibular nuclei (Nakaya et al., 1994). In guinea pig brain stem slices, substance P depolarized about two-thirds of medial vestibular neurons by activating atypical postsynaptic substance P receptors (Vibert et al., 1996). In vivo, an intrasystemic injection of substance P accelerates the recovery from postural deficits following unilateral labyrinthectomy. Adrenocorticotropin (ACTH), Growth Factors, and Other Neuropeptides An in vitro slice study has shown that ACTH depresses the resting discharge of medial vestibular neurons. In vivo, intrasystemic injections of ACTH improve the compensation of the postural and oculomotor syndromes following unilateral labyrinthectomy (for review, see Darlington et al., 1996). The fragment 4–9 appears to be crucial for this effect (Gilchrist et al., 1996). NGF receptors have been detected in medial and spinal vestibular nuclei, mostly at the borders of the prepositus hypoglossi nucleus (Sukhov et al, 1997). This is interesting because there is increasing evidence that neurotrophic peptides are involved in neuronal plasticity (for review, see Thoenen, 1995). Finally, other types of neuropeptides and neuropeptide receptors have been detected in neurons and/or

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FIGURE 5 Summary diagram. Excitatory synapses are in white, inhibitory synapses are in black. Ach, acetylcholine; GLU, glutamate; GLY, glycine; H VOR, horizontal vestibuloocular reflex; 5-HT, serotonin; IO, inferior olive; LDT, laterodorsal tegmentum; locus C, locus coeruleus; NA, noradrenaline; NGF, nerve growth factor; NO, nitric oxide; PPT, pediculopontine tegmentum; Prep. H, prepositus hypoglossi nucleus; SP, substance P; t. m. neurons, tuberomamillary neurons; V VOR, vertical vestibuloocular reflex. Other neuropeptides than those mentioned on the diagram have been detected, namely neuropeptide Y, neurotensin, vasoactive intestinal peptide (VIP), cholecystokinin, and thyrotropinreleasing hormone (TRH). Dopamine and adenosine triphosphate (ATP) have also been detected at the level of vestibular nuclei, but are not mentioned in the diagram (see text). The vestibulocollic reflex is not represented.

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terminals within the vestibular nuclei. Fibers positive for neuropeptide Y, neurotensin, vasoactive intestinal peptide (VIP), and cholecystokinin most probably innervate vestibular nuclei. Thyrotropin-releasing hormone (TRH) and neurotensin receptors have also been described in the vestibular nuclei (Zanni et al., 1995).

cations. Clearly, vestibular compensation and vestibular adaptation are valuable models used to study the postlesional plasticity and adaptive capabilities of the central nervous system. As a reminder, LTD had been postulated and demonstrated in the flocculus in an effort to undercover the neural basis of the vestibuloocular adaptation.

Presence of Purine Receptors in Vestibular Nuclei

Acknowledgments

Adenosine 5´ triphosphate (ATP)n is a provider of energy to neurons. ATP also interacts with several membrane receptors and modulates neuronal activity. Seven types of ionotropic receptors (the P2x receptors) and six types of metabotropic receptors (the P2Y receptors) activated by ATP have been described. In addition, several subtypes of P1 receptors sensitive to adenosine have been identified. ATP usually has excitatory effects on rat neurons (Gu and MacDermott, 1997). Rat MveN are sensitive to ATP and are endowed with P2X and P2Y purine receptors (for review, see Chessell et al., 1997). P2 receptor agonists increase in a dosedependent manner the spontaneous discharge of onethird of the MveN tested in rat brain stem slices and P2 antagonists suppress this effect.

CONCLUSION Stabilizing gaze and posture require complex multisensory integration, which must show a high degree of plasticity. As stated in the first section of this chapter, this integration can be defined as the process of matching multiple internal representations of an external event (head and/or trunk rotation), obtained from different sensory modalities, into a unique intrinsic frame of reference in which appropriate motor commands can be coded. We have summarized here the neuronal networks, which implement these complex sensorimotor transformations in the rat brain stem. The underlying neuronal computations are the by-product of both the emerging properties of the vestibular-related networks and the individual properties of each of their components, i.e., the neurons. It explains why the rat is a model of choice to study gaze and postural control. In that species, in vivo and in vitro preparations can be combined to study these two complementary aspects. This is necessary because data brought by the molecular biology methods and in vitro recordings have to be interpreted in a functional frame to ultimately figure out how the CNS works. It is also clear that investigating gaze and postural control paves the way for new treatments of vestibular syndromes and leads to new clinical appli-

We thank N. Vibert and C. de Waele for their excellent assistance and for critical reading of the manuscript and V. Chazaly and D. Orcel for their technical help.

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31 Auditory System MANUEL S. MALMIERCA and MIGUEL A. MERCHÁN Laboratory for the Neurobiology of Hearing, Department of Cellular Biology and Pathology Faculty of Medicine, University of Salamanca and Institute for Neuroscience of Castilla y Léon, Spain

Prior to the publication of the first and second editions of The Rat Nervous System in the 20th century, the most commonly used experimental animal for auditory research was the cat and, to a lesser extent, the guinea pig, the chinchilla, and the mouse. The only studies on the rat auditory system were the pioneering work by Harrison and associates in the 1960s in the cochlear nuclear complex and superior olivary complex. Over the past 20 years, however, the rat has become a popular research animal for anatomical,1 physiological,2 and behavioral3 studies of the auditory system, and a wealth of complementary data now exits. To mention one

example, the rat brain, because of its small size, has become a specimen of choice for “brain slice preparations,” e.g., Smith, Wu, and their associates. In this context, the rat auditory system has proved to be useful also to explore some general features of the central nervous system. The large nerve terminals (calyces of Held) in the medial nucleus of the trapezoid body offer a unique opportunity to explore the modulation of synaptic channels and mechanisms at a mammalian central synapse (e.g., Forsythe, 1994; Kungel and Friauf, 1997; Löhrke et al., 1998; Borst and Sakmann, 1999; Bollmann et al., 2000; Cuttle et al., 2000; Smith et al., 2000; Sahara and Takahashi, 2001). A major aim in hearing research is to understand the physiology of the human auditory system and to identify the causes and treatments for hearing impairment. Comparative hearing research is important because animal models can be developed, evaluated, and eventually applied to clinical problems (Fay, 1994). The purpose of the current review is to highlight the functional organization of the rat auditory system and to focus on the rat as a useful animal model for hearing research. Thus, this review is based largely on studies of the rat auditory system, and citations refer to the rat unless stated otherwise. Other species are referred to only if particular data are not available in rat, or for comparative purposes. It should be noticed that both the peripheral and central auditory system vary greatly between fish, reptilians, amphibians, and mammals. The auditory system is also highly specialized in certain mammals, especially echolocating mammals such as cetaceans and bats. When complementing the

1

E.g., Faye-Lund and Osen, 1985; Coleman and Clerici, 1987; Merchán et al., 1988, 1994; Clerici and Coleman, 1990; Osen et al., 1991; Saldaña and Merchán, 1992; Bajo et al., 1993; Malmierca et al., 1993, 1995a, 2002; Feliciano et al., 1995; Lee et al., 1996; Merchán and Berbel, 1996; Saldaña et al., 1996; López et al., 1999; Saint-Marie et al., 1999a, 1999b; Winer et al., 1999b, 1999c; Alibardi, 2000, 2001, 2002, 2003; Gabriele et al., 2000a, 2000b; Saldaña and Berrebi, 2000; Kulesza and Berrebi, 2000; Campos et al., 2001; Riquelme et al., 2001; Kulesza et al., 2002. 2 E.g., Rees and Møller 1983, 1987; Kelly and Sally, 1988; Sally and Kelly, 1988; Kelly, 1990; Kelly et al., 1991; Pierson and Snyder-Keller, 1994; Hu et al., 1994; Hu, 1995; Irvine et al., 1995; Wu and Kelly, 1995a, 1995b, 1995c, 1996; Fu et al., 1996, 1997a, 1997b; Wu and Fu, 1998; Peruzzi et al., 1997, 2000; Bajo et al., 1998; Kelly et al., 1998a; Kilgard and Merzenich, 1998, 1999; Wu, 1998, 1999; Barlett and Smith, 1999, 2002; Luo et al., 1999; Barlett et al., 2000; Hefti and Smith; 2000, 2003; Lim et al., 2000; Sivaramakrishnan and Oliver 2001; Zhao and Wu, 2001; Doron et al., 2002; Ma et al., 2002; Malmierca et al., 2003. 3 E.g., Kelly and Masterton, 1977; Kelly, 1980; Kelly and Glazier, 1978; LeDoux et al., 1984; Kelly and Judge, 1985; Kelly and Kavanagh, 1986; Romanski and LeDoux, 1992; Heffner et al., 1994; Ito et al., 1996; Kelly et al., 1996; van Adel and Kelly, 1998, 1999; Syka et al., 2002.

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description with data from other species, therefore, we have chosen mostly mouse, guinea pig, and cat (small and relatively unspecialized). The specific stimulus for the ear is pressure waves within a certain range of frequencies. The range of frequencies to which the ear responds varies between species. Within their characteristic range, many species, like echolocating bats, are particularly tuned to certain frequencies of special importance to their behavior (reviewed in Fay and Popper, 1994) and are considered to be “auditory specialists” (Echteler et al., 1994). In rat the frequency range is from 0.25 to 70 kHz (Kelly and Masterton, 1977; Ryan et al., 1988; Heffner et al., 1994), while it is about 2–70 kHz in mouse, 0.2–45 kHz in guinea pig, and 0.125–60 kHz in cat. In humans the range is 0.02–20 kHz. Since the representation of tonal frequencies, i.e., tonotopic organization, is the main organizing principle in the auditory system, the frequency range influences the anatomy of the auditory structures. Other aspects of sound like intensity, localization, and temporal (including the spectral) patterns are recoded by central neuronal networks. Sound waves are transmitted mechanically through the outer and middle ear to the sensory hair cells of the organ of Corti (Fig. 1), in the cochlear partition of the inner ear. Receptor potentials set up in the sensory hair cells are transmitted to the brain stem by the cochlear nerve (Figs. 2 and 3). In the first relay center, i.e., the cochlear nuclear complex, the signals of the cochlear nerve are shunted into a number of parallel ascending tracts, each with a particular course and destination (conduction velocities and relays). The ascending auditory tracts converge toward the auditory midbrain (Fig. 3), the inferior colliculus, which in contrast to the role of the superior colliculus within the visual system is an obligatory relay in the route to the auditory cortex (but see Malmierca et al., 2002). From the inferior colliculus and upward, the auditory pathway can be divided into a “core projection” where the tonotopic organization is very sharp and a “belt projection” where it is less sharp (e.g., Andersen et al., 1980a). In contrast to the minimum of two relay stations between the periphery and cerebral cortex in the visual and somatosensory cortices, there is a minimum of three relays in the auditory system (Fig. 3), with several stages of convergence and divergence, and at least seven crossing levels (Fig. 3) that make the auditory system unique and poorly suited for topographical clinical diagnostics.

THE ORGAN OF CORTI In terrestrial mammals, the pinna funnels the sound waves, via the external acoustic meatus, to the tympanic

membrane. The chain of three small ossicles—the malleus, incus, and stapes—transfers the sound vibrations of the tympanic membrane through the aircontaining middle ear to the footplate of the stapes which inserts in the oval window and serves to match impedance of air to fluid. The footplate is smaller than the tympanic membrane and the chain and therefore acts as a transformer, the effect of which can be reflexively regulated by means of the two middle ear muscles (m. stapedius and m. tensor tympani). In the inner ear, the endolymph-filled membranous labyrinth is suspended in the perilymph-containing osseous labyrinth of about the same shape. The auditory part of the mammalian osseous labyrinth is the cochlea, and the membranous labyrinth consists of the cochlear duct. Situated in the middle is an elastic membrane, the basilar membrane, which acts as a hydromechanical frequency analyzer. The outer form of the rat cochlea is cylindrical and possesses 2.2 turns (Burda et al., 1988). The structure of the rat cochlea corresponds to the basic mammalian plan. Figure 1A shows a midmodiolar section through the left cochlea in the wild Rattus norvegicus. The cochlear duct divides the bony cochlea into three compartments, so-called scalae, that are fluid-filled. The turns of the cochlea are seen in detail in Figs. 1A and 1C and in particular the longitudinal division into three scalae: media, vestibuli, and tympani. The scalae coil together, keeping their mutual relationships along the length of the cochlea (Figs. 1A and 1C). Near the modiolus, the scala tympani is separated from the scala vestibuli by the osseous spiral lamina. The scala media is separated from the scalae vestibuli and tympani by the vestibular (Reissner’s) membrane above and the basilar membrane below, respectively (Fig. 1A). At the lateral wall, the scala media is limited by the stria vascularis (Fig. 1A). The outer scalae tympani and vestibuli are joined at the apex by an opening called the helicotrema (Fig. 1A). These outer scalae are filled with a fluid known as perilymph that has a high concentration of Na+ (140 mM) and a low concentration of K+ (4 mM), (Johnstone and Sellick, 1972), similar to the extracellular ionic composition. By contrast, the scala media contains endolymph, which is similar to the intracellular fluid as it has a high concentration of K+ (120 mM) and a low concentration of Na+ (1 mM), (Bosher and Warren, 1968). The scala vestibuli is sealed off from the middle ear by the oval window into which the footplate of the stapes inserts. The scala tympani is sealed by the membrane of the round window. The latter membrane prevents pressure buildup and allows movements of the basilar membrane to occur upon vibrations of the stapes. The vestibular membrane is a thin sheet of connective tissue covered on both sides by epithelial cells

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FIGURE 1 (A) Midmodilolar section through the cochlea (Rattus norvergicus) showing the scalae media, tympani, and vestibuli (redrawn from Burda et al., 1988). (B) Scheme of the organ of Corti (redrawn from Nobili et al., 1998). Asterisks, stereocilia. Scanning electron micrographs of the rat cochlea were kindly provided by Dr. David Furness (C–G). (C) The whole cochlea, dissected to reveal the internal spiral structure. (D) Detail of the surface of the organ of Corti, showing the hair bundles of the single row of inner hair cells and three rows of outer hair cells. (E) A single hair bundle of an inner hair cell, showing the straight rows of stereocilia. (F) A hair bundle from an outer hair cell showing the W shape of the bundle and the three rows of stereocilia of increasing height. (G) Detail of stereocilia, showing the tiny tip link (e.g., arrow) connecting each shorter stereocilium to the taller one behind. For abbreviations see list at end of chapter.

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FIGURE 2 Afferent and efferent innervation of the cochlear epithelium. Note that several type I afferent fibers converge onto single IHCs while a single type II afferent fiber terminates onto several OHCs. Type I fibers terminate in the VCA, VCP, and DC. Type II fibers terminate on the GrC and marginal shell areas of the VC and DC. The efferent MOC innervates the OHCs and the efferent LOC innervates the IHCs. (Modified after Brown et al., 1988a.) For abbreviations see list at end of chapter.

joined by tight junctions. This structure prevents the perilymph from mixing with the endolymph. Detailed measurements of the basilar membrane are given by Burda et al. (1988) in three different rat species. The basilar membrane is 9.5–12 mm long, depending on the rat species (Burda et al., 1988); supports the organ of Corti; and is composed of extracellular material rich in filaments that stretch across its width (Fig. 1B) (Iurato, 1962). These filaments provide the basilar membrane with stiffness. The width of the basilar membrane increases gradually from the base (56.9–71.9 μm) to the apex (189.2–200.7 μm; Burda et al., 1988). By contrast, the basilar membrane thickness gradually decreases from the base (15–19.3 μm) to the apex (1.7–2 μm; Burda et al., 1988). The stria vascularis is a three-layer epithelium surrounding a dense network of capillaries (Smith, 1978). The functional role of the stria vascularis is twofold: it regulates the ionic composition of the endolymph and it provides a physical seal between the bony and membranous labyrinths. The cells within the inner margin are in direct contact with the endolymph and have been suggested to be involved in removing Na+ from and transporting K+ into the scala media, a process that results in a high positive endocochlear potential (60–100 mV, discussed in Offner et al., 1987). The organ of Corti is the sensory epithelium and rests upon the basilar membrane (Fig. 1B). It is made up of several types of supporting cells and two types of sensory cells (Fig. 1B): inner hair cells (IHCs, Figs. 1B, 1D, and 1E) and outer hair cells (OHCs, Figs. 1B, 1D,

and 1F). The IHCs are arranged in a single row, the OHCs in three parallel rows (Fig. 1B and 1D). At their free, cuticular end, the hair cells are provided with stereocilia in a typical “W” pattern (for a review, see Lim, 1986; Slepecky, 1996). The organ of Corti is overlain by the gel-like tectorial membrane (Fig. 1B), which is indirectly connected to the osseus spiral lamina through the spiral limbus. Only the stereocilia of the OHCs appear to be in contact with the tectorial membrane (Fig. 1B). Shearing movements between the basilar membrane with the sensory epithelium, on the one hand, and the tectorial membrane, on the other hand, cause receptor potentials to be produced in the hair cells by means of deflections of their stereocilia (reviewed in Lim, 1986; Russell et al., 1986; Nobili et al., 1998). Mechanical vibrations, derived from the incoming sound, are thus transduced into action potentials, the language of the nervous system. The two types of hair cells each receive specific types of afferent and efferent fibers. In rat, the cochlear hair cells together with the supporting cells are arranged in a geometrically regular pattern that is similar to that of all mammalian species studied thus far (Keithley and Feldman, 1979, 1982; Burda et al., 1988; Keithley et al., 1992). The rat (R. rattus) possesses about 3800–4000 OHCs (364, average total density of OHC/mm of the length of the organ of Corti) and 1000–1300 IHCs (121, average total density of IHC/mm of the length of the organ of Corti). However, the density of hair cells is not uniform. The lowest density for the OHCs is in the base of the cochlea with a trend of gradually increasing density toward

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FIGURE 3 Ascending auditory pathaway of the rat (modified after Brodal, 1981; AC is from Herbert et al., 1991). For abbreviations see list at end of chapter.

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the apex. The maximum density for the IHCs is found at about 25% of the basilar membrane length from the base. Both the OHCs and IHCs do not rest on the basilar membrane itself but on specialized supporting cells. The IHCs form a single row oriented parallel to the cochlear spiral. They are flask-shaped with a globular cell soma tapering into a thinner elongated neck (Fig. 1B). Their nucleus is rounded and located halfway along the length of the cells and divides the cells into two topographic domains. At the basal end are found several synaptic contacts from about 20 afferent cochlear nerve fibers; hence this pole is also referred to as the neural pole (Fig. 2). The neural pole receives about 90–95% of all afferent contacts with cochlear nerve fibers (reviewed in Ryugo, 1992). The apical pole is characterized by a bundle of stereocilia in nearly straight rows and is synapse free (Fig. 1E). There are no quantitative studies on the height of the IHC in rat, but in other species it varies (30–40 μm, Lim, 1980; Nadol, 1988). The number and size of stereocilia on each IHC varies along the length of the cochlear duct. As in other species, the IHCs stereocilia increase gradually from the base (32 per IHCs and 2.3 μm long) to the apex (41 per IHC and 4.3 μm long). The OHCs rest on the supporting cells called Deiter’s cells and make up 75–80% of all hair cells (Fig. 1B). OHCs are organized in three parallel rows (Fig. 1D). The OHCs are cylindrically shaped and possess a large spherical nucleus located at the neural pole. OHCs are characterized by having several cisterns of endoplasmic reticulum distinctly located under the cellular membrane in a laminar fashion that extends from the nucleus up to the apical pole (Lim, 1986; Slepecky, 1996). At the cuticular end, three rows of stereocilia arise forming a typical W-shaped configuration (Fig. 1D). The size and number of OHCs and stereocilia gradually vary along the basilar membrane. Thus, OHCs are 15 μm tall at the base and 40 μm at the apex. The cilia are 2.7 μm long at the base and 4.4 μm at the apex. The number of stereocilia per OHCs is larger at the base (75 vs 62; Burda et al., 1988). As opposed to the IHCs, OHCs only receive 5–10% of the afferent innervation from the cochlear nerve but are contacted by a large number of efferent nerve terminals originating in the olivocochlear bundle discussed below (Warr, 1992). The most conspicuous supporting cells in the organ of Corti are inner and outer pillar cells (Fig. 1B). They form the tunnel of Corti between the IHCs and OHCs (Fig. 1B). These cells rest upon the basilar membrane, close to the junction of this membrane to the spiral osseous lamina, and contain thick bundles of microtubules that confer on them a great rigidity. Medially, the IHCs are wedged in a bundle of supporting cells and, laterally, the OHCs are maintained by the Deiter’s cells. The latter sit on

the basilar membrane (Fig. 1B). The tectorial membrane is a large acellular flap of tissue located on the top of the organ of Corti, laterally, and is attached to the tallest stereocilia of the OHCs (Fig. 1B). Medially, the tectorial membrane is only attached to the IHCs in the basal turn of the cochlea in some species (monkey, Hoshino, 1977) which leads to the general idea that the IHCs’ stereocilia are freestanding and the OHCs are firmly anchored to the tectorial membrane (Lim, 1986). Sensory transduction in the cochlea has been studied for many years. It has been known for several years that the stereocilia of the hair cells in the lateral line organ of fish are sensitive to bending (Flock, 1965) and more recent studies on mammalian hair cells confirmed the early studies of hair cells of the lateral line organ (Russell and Sellick, 1978; Russell, 1983; Russell et al., 1986). The hair-cell stereocilia (Figs. 1D–1G) are connected together through tip-links (Fig. 1G, arrow) and displacement of the cilia toward the higher stereocilia allows the opening of ion channels that are located near or at the tips of the stereocilia (guinea pig, Comis et al., 1985; Hudspeth, 1982, 1989; Hudspeth et al., 2000). The inflow of K+ ions results in depolarization of the hair cells and release of a neurotransmitter. The chemical nature of this neurotransmitter is still unknown, but it is likely to be glutamate (Usami et al., 1992; Matsubara et al., 1996; Wang et al., 1998; for review, see Ottersen et al., 1998). It is therefore clear that the hair cells in the inner ear transform vibrations into action potentials when their stereocilia are deflected but the IHCs and OHCs differ in function. Thus, it has been recognized over the past 25 years that the IHCs act as the primary receptor cell (Russell, 1983; Markin and Hudspeth, 1995) while the OHCs also act as motor cells that can convert membrane potential into a mechanical force (reviewed in Nobili et al., 1998). The frequency components of a sound are mapped along the length of the basilar membrane and its overlying organ of Corti (von Bèkèsy, 1960). The cochlear frequency map is determined by the progressive increase in the stiffness of the basilar membrane that is correlated with the decreasing basilar membrane width and increasing thickness from the apex to the base (e.g., von Bèkèsy, 1960; Dallos, 1992; Echteler et al., 1994). But the frequency selectivity and sensitivity depend on the physiological integrity of the cochlea (Rhode, 1984). Over the past 20 years, a fundamentally new concept arose in hearing research, namely, that the cochlea is active (Brownell et al., 1985; guinea pig: Ashmore, 1987). OHCs appear able to contract and elongate and in so doing generate mechanical forces capable of altering the delicate mechanics of the cochlear duct, increasing frequency selectivity and sensitivity (Neely and Kim, 1983; Dallos and Evans,

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1995a, 1995b; Dallos, 1997; Nobili et al., 1998). The discovery of OHC motility has modified the old view that the cochlea is a simple passive frequency analyzer. In this new view, the cochlea has been described as an active nonlinear filter that allows transmission of the auditory signals to the auditory nerve by the IHCs. Furthermore, this frequency selectivity is possible because of the suppression of adjacent frequencies, a mechanical effect equivalent to the concept of lateral inhibition in the central nervous system (Nobili et al., 1998). The pioneering studies of Rasmussen (1940, 1953), Engström (1958), and others (for an extensive review, see Slepecky, 1996) on the innervation of the cochlea identified two types of nerve fibers to the organ of Corti: afferent and efferent fibers (Figs. 2 and 35). The afferent fibers convey impulses to the cochlear nuclear complex while the efferent fibers convey impulses from the superior olivary complex to the organ of Corti (v.i.). More recent studies have detailed the fine structure and organization of the innervation in several species (Spoendlin, 1967, 1968; Iurato, 1967; Kimura, 1975; Warr and Guinan, 1979; Nadol, 1988; Warr, 1992) including rat (Rosenbluth, 1962; Ross and Burkel, 1973; Lenoir et al., 1987; Romand et al., 1988; Warr et al., 1997). There are two subtypes of afferent fibers (Figs. 2 and 35): the thick myelinated fibers arising from the bipolar type I spiral ganglion cells innervating the IHCs and the thin unmyelinated fibers arising from monopolar type II spiral ganglion cells innervating the OHCs (Rosenbluth, 1962). There are about 15,800–19,900 neurons in the spiral ganglion depending on the rat strain and rat age studied (Keithley and Feldman, 1979; Burda et al., 1988; Hoeffding and Feldman, 1988a, 1988b, 1988c; Hall and Massengill, 1997). The density of neurons varies along the cochlear spiral (Keithley and Feldman, 1979; Burda et al., 1988). Thus the maximum innervation density is 4.7 (Wistar rat), 5.3 (R. norvergicus) and 6 mm (R. rattus) from the base. The early studies on the cat demonstrated that 90–95% of type I fibers contact only IHCs and are unbranched and each fiber terminates on a single IHC. Each IHC contacts about 20 different fibers (Figs. 2 and 35) (Spoendlin, 1972). The number of nerve fibers that synapse onto each IHC is uneven but seems to be larger in frequency regions that are functionally important. The synapses between IHCs and afferent fibers are characterized at the presynaptic zone by the existence of a presynaptic bar surrounded by a moderate number of synaptic vesicles (Sobkowicz et al., 1986; Matsubara et al., 1996). The shape and size of these synaptic bars vary among the IHCs and it has been suggested that this change is due to the physiological characteristics of the postsynaptic fibers and the functional state of

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the IHCs (Jorgensen and Flock, 1976; Liberman, 1980; Merchán-Pérez and Liberman, 1996; reviewed in Slepecky, 1996). In the cat, three types of type I fiber have been characterized based on morphological features that correlate with their spontaneous activity (low, medium, and high spontaneous rate) and threshold sensitivity (Liberman and Oliver, 1984; Liberman et al., 1990). Low threshold and high spontaneous rate fibers have the larger diameter and are located at the pillar side of the IHC while high threshold and low spontaneous rate fibers are thinner and located on the modiolar side (Kawase and Liberman, 1992). The unmyelinated type II fibers arise from small unipolar ganglion cells and constitute about 5% of all ganglion cells (Figs. 2 and 35). In contrast to type I, type II fibers are highly branched and a single fiber synapses onto several OHCs (6–100). There are also two systems of efferent fibers (Figs. 2 and 35): the lateral efferent system (lateral olivocochlear system) that innervates the IHCs and the medial efferent system (medial olivocochlear system) that innervates the OHCs (Warr and Guinan, 1979; Warr, 1992; Warr et al., 1997). They form part of the olivocochlear system that belongs to the descending auditory pathway and are treated in detail in a later section.

THE COCHLEAR NUCLEAR COMPLEX In mammals the cochlear nuclear complex (CNC) provides the first relay center in the ascending auditory pathway (Figs. 3–8). Here the signals of the auditory nerve are transmitted to a number of different types of second-order neurons, each apparently recoding and refining specific aspects of the sound stimulus (e.g., frequencies, temporal pattern, intensity). Each cell type conveys the modified signal to selected higher order centers through specific fiber tracts. The transformation and transmission of signals are influenced by the organization of the presynaptic elements of the cochlear nerve fibers, membrane properties of the second-order neurons, intrinsic fiber systems, commissural connections, and descending fibers from higher auditory centers (for review, see Osen, 1988; Young et al., 1988a; Cant, 1992, 1993; Rhode and Greenbergs, 1992; Romand and Avan, 1997). The rat CNC corresponds to the general mammalian pattern with some minor modifications. As in other mammals, the CNC in rat is situated laterally and superficially in the brain stem (Fig. 5). There are some interspecies variations in location and spatial orientation presumably secondary to differences in the shape of the brain stem (Fig. 4) (Osen, 1988). The CNC (Figs. 4 and 5) consists of a dorsal cochlear nucleus (DC) and a ventral cochlear nucleus (VC). The latter is sub-

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FIGURE 4 Diagrams of the CNC in the rat (A, redrawn after Harrison and Irvine, 1966a) and cat (B, C, redrawn after Osen, 1988). Note the locations of different cell types in the DC (B) and the VCA and VCP (C). For abbreviations see list at end of chapter.

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FIGURE 5 Transverse (A–C) and sagittal (D–F) diagrams of the CNC in rat with special emphasis on the granule cell domains (redrawn after Mugnaini et al., 1980b) For abbreviations see list at end of chapter.

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divided by the cochlear nerve root into an anteroventral (VCA) nucleus and a posteroventral (VCP) nucleus. The rat VC is flattened mediolaterally, measuring about 1.5 mm rostrocaudally, 0.5 mm mediolaterally, and 1.5 mm dorsoventrally, with the dorsal end tilted about 45° medially with respect to the horizontal plane (rough measurements based on Mugnaini et al., 1980a, 1980b, and Paxinos and Watson, 1998) (Fig. 5). Thus only a tilted parasagittal section includes the entire dorsoventral axis of the VC, which is the axis along which the frequency gradient lies (see below). The DC curves around the restiform body4 in the floor of the lateral recess of the fourth ventricle (Fig. 5). It measures 0.4 mm mediolaterally, 1 mm rostrocaudally, and 2 mm along its curved dorsoventral (strial) axis.

Primary Afferents As in other mammals (Ramón y Cajal, 1904, 1909; Lorente de Náo, 1933, 1981) the cochlear nerve fibers terminate within the rat CNC in a tonotopical pattern. Seen in Golgi sections, each fiber bifurcates into an ascending branch, which supplies the VCA, and a descending branch, which supplies the VCP and DC (Harrison and Feldman, 1970). More detailed studies in cat show (Sando, 1965) that fibers from the apical, low frequency part of the cochlea divide ventrally and terminate within laminar fields in the ventral part of the CNC, while those from more basal, high frequency parts of the cochlea divide progressively more dorsally and supply laminar fields in the dorsal part of the CNC. The anatomical distribution of the primary fibers forms the basis for the laminar tonotopic organization of the three subnuclei (Fig. 3). This tonotopic organization has been demonstrated in rat by c-fos immunocytochemistry (Friauf, 1992; Rouiller et al., 1992; Luo et al., 1999; Saint-Marie et al., 1999) and also electrophysiologically (Kaltenbach and Lazor, 1991). Axon collaterals arise from the main branches of the cochlear nerve all along their trajectory (Figs. 4 and 5). In the rat, they also arise from the main stem axon (Osen et al., 1991). Myelinated type I and unmyelinated type II axons follow similar bifurcation patterns, but the total terminal area and mode of termination differ. The mode of termination of the myelinated type I fibers has been described in rat by Feldman and Harrison (1969) in protargol-stained sections and in greater detail in intracellular HRP-labeling experiments in cat and mouse (Ryugo and Fekete, 1982; Rhode et al., 1983a, 1983b; Fekete et al., 1984; Liberman and Oliver, 1984; Rouiller and Ryugo, 1984; Rouiller et al., 1986; Brown

4

Also referred to as the inferior cerebellar peduncle, icp.

et al., 1988a; Ryugo, 1992). Two basic types of terminals are found: large, axosomatic endings called “bulbs of Held” (Held, 1893) and small boutons. The bulbs of Held arise mainly from the ascending branches, while the small boutons arise from loosely ramifying collaterals of both ascending and descending branches (Ramón y Cajal, 1904, 1909; Lorente de Nó, 1933; rat, Feldman and Harrison, 1969; mouse, Willard and Ryugo, 1983). Type I fibers supply all parts of the CNC except the superficial granule cell areas, which are devoid of such input (cat, Osen, 1969). The mode of termination of the unmyelinated type II fibers has been studied after HRP injections into the spiral ganglion of gerbil, mouse, and cat (Brown et al., 1988a) and the labeled fibers have been reconstructed in three dimensions (Brown and Ledwith, 1990). The type II fibers innervate areas rich in granule cells and appear to supply the marginal shell of the VC; see below (Brown et al., 1988b; Brown and Ledwith, 1990; Ryugo, 1992; Ye et al., 2000).

Ventral Cochlear Nucleus The cytoarchitecture of the VC shows only minor interspecies variations, although certain morphological characteristics of the neurons may be more or less distinct. In rat several morphologically different cell types were described by Harrison and his colleagues in protargol-stained sections (Harrison and Irving, 1965, 1966a; Feldman and Harrison, 1969; Harrison and Feldman, 1970). Corresponding cell types were later defined in cat and in rat on the basis of Nissl and Golgi studies (rat, Saldaña et al., 1987; Merchán et al., 1988; cat, Osen, 1969; Brawer et al., 1974). Five main types are generally recognized (Figs. 4 and 6): spherical bushy, globular bushy, octopus, multipolar, and small cells (Osen, 1969; Brawer et al., 1974; for discussion of analogies see Osen, 1969). The five main types may be collected into two groups with respect to dendritic patterns and main axonal targets. The first three types are segregated in distinct portions of the nucleus. The spherical bushy cells are found rostrally in the VCA, the globular bushy cells lie centrally on both sides of the nerve root in the caudal VCA and the rostral VCP, and the octopus cells are found caudally in the VCP. The rat VC is characterized by a relatively small spherical bushy cell area and a relatively large globular bushy cell area, in agreement with the relative size of their target areas (see below). In contrast to the first three types, the multipolar and small cells are present throughout the VC. The small cells are most abundant around the peripheral margins of the nucleus deep to the superficial granule cell layer. In the cat, a large collection of small cells located dorsolaterally in a superficial location forms

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FIGURE 6 Main projecting cell types of the CNC in rat with their corresponding physiological responses (modified after Moore and Osen, 1979b). For abbreviations see list at end of chapter.

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FIGURE 7 Diagram of the T- and D-stellate cells in the VC and their projections to the DC. T-stellate (planar) cells have frequency-specific projections while D-stellate (radiate) cells have across frequency projections (redrawn from Doucet and Ryugo, 1997). For abbreviations see list at end of chapter.

the small cell cap of the VC (Fig. 4) (Osen 1969). A distinct cap area is not easily distinguished in rat. Spherical Bushy, Globular Bushy, and Octopus Neurons The spherical bushy, globular bushy, and octopus neurons (Figs. 4 and 6) have nontapering dendrites ending in bushy-like formations. They differ with regard to the number of root segments and the relative length of the stem dendrites and the terminal bush. In addition each cell type receives different numbers of afferent cochlear fibers and projects to different targets. The bushy cells receive a small number of large axosomatic terminals (cat, Sento and Ryugo, 1989), the bulbs of Held, and have so-called primary-like responses to pure tone stimulation, similar to those of the auditory nerve fibers (Young et al., 1988b). The specific central projections of these neurons were first described by Harrison and Warr (1962) on the basis of silver degeneration studies. Later, Friauf and Ostwald (1988) confirmed by intraaxonal injections in rat that the spherical bushy cells project bilaterally to the

medial superior olive and to the ipsilateral lateral superior olive (Fig. 6). Like the somatic surface of the spherical bushy cells, that of the globular cells is almost completely covered by synaptic terminals and the fine structrure of the terminals is very similar to that described for spherical cells. Although the cochlear terminals are not as large as the endbulbs that contact spherical cells (Fig. 6). The globular bushy cells project to the contralateral medial nucleus of the trapezoid body (Fig. 6). Both spherical and globular bushy cell types seem to be specialized for transmitting precise temporal information necessary for sound localization (Young et al., 1988a). The octopus cells receive small boutons from a number of collaterals of primary fibers (Fig. 6). They respond to a tone burst with a single spike and so have been called onset units (cat, Godfrey et al., 1975; Rhode et al., 1983b). Despite the apparent inhibition of firing after the onset of the stimulus, they receive very few GABAergic and glycinergic inhibitory afferents (mouse, Oertel et al., 1990; cat, Osen et al., 1990). Their main projection is to the superior paraolivary nucleus on

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FIGURE 8 (A) Parasagittal section of a Golgi-impregnated section showing cochlear root neurons. (B) Parasagittal sections of the cochlear nuclear complex showing the primary afferents labeled after HRP injections in the basal regions of the spiral ganglion. (Inset) Unstained cochlear root neuron intermingled among labeled fibers after HRP injection in the basal part of the cochlea. Those fibers send collaterals that terminate on the somata of a cochlear root neuron. (C) Frequency response area of a typical root neuron. Observe, the high best frequency and high threshold at the low frequency tail. (D) PSTH of a root neuron (primary-like with notch type). Figures kindly provided by Dr. López and adapted (C, D) from Sinex et al. (2001). For abbreviations see list at end of chapter.

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both sides and to the contralateral ventral complex of the lateral lemniscus (Fig. 6) (guinea pig, Schofield, 1995; Thompson and Thompson, 1991; Thompson and Schofield, 2000). Their function is still unclear. It has been speculated that these cells, which are the only VC cells not receiving inhibitory input from the DC (mouse, Golding et al., 1995), are involved in encoding short latency echos giving spatial depth to the sound (Wickesberg and Oertel, 1990). It has also been suggested that they encode the pitch period in their temporal firing patterns (mouse, Golding et al., 1995; Oertel, 1997, 1999). While the globular and spherical bushy cell axons course ventrally in the trapezoid body, the octopus cell axons course dorsally, above the restiform body in the intermediate acoustic stria (Fig. 6).

1987; rat, Doucet and Ryugo, 1997) (Fig. 6). The axons apparently give off widely dispersed collaterals to the ipsilateral VC and DC as they course in the dorsal acoustic stria (Fig. 7) (cat, Smith and Rhode, 1989; mouse, Oertel et al., 1990; rat, Doucet and Ryugo, 1997). The Dstellate cells show glycine-like immunoreactivity (guinea pig, Wenthold, 1987; cat, Osen et al., 1990; rat, Doucet et al., 1999) and are probably the only large projection neurons of the CNC that are inhibitory. They are “onchop” responders to pure tone stimulation (guinea pig, Winter and Palmer, 1995). Judged by their projections, they may afford a broad non-frequency-specific bilateral inhibition in the CNC bilaterally (Figs. 6 and 7).

Multipolar Cells

The small cells are abundant in the marginal shell of the VC (Fig. 4) which is composed of the “granule cell layer” and the subjacent “cap area.” The granule cell layer is continuous over the free surface of the rat CNC (Fig. 5) and forms a lamina partly separating the VC and DC (Mugnaini et al., 1980a, 1980b). In the DC the granule cell layer is covered superficially by a molecular layer. The granule cell axons project as parallel fibers (Fig. 6) to the molecular layer (Mugnaini et al., 1980a, 1980b, their Fig. 11) and are described in more detail in connection with the DC. The cap area is small in rat but it still distinguishable due to its contingent of small cells (reviewed in Cant, 1993), many of which show glycine- and/or GABA-like immunoreactivity (cat, Osen et al., 1990). The cap is supplied by both type I and type II fibers. In cat, nearly all type I auditory nerve fibers that innervate the cap have low spontaneous rates in contrast to those terminating in the core (cat, Leake and Snyder, 1989; Kawase and Liberman, 1992). The marginal shell also receives descending input (guinea pig, Shore and Moore, 1998, cat, Ye et al., 2000) and its cells show a wide dynamic range (as wide as 98 dB SPL, cat, Ghoshal and Kim, 1996). Some of the cells in the marginal shell project to the inferior colliculus (cat, Adams, 1979); others project to the medial olivocochlear system bilaterally, to the lateral olivocochlear system ipsilaterally (cat, Ye et al., 2000), and to the medial geniculate body (Malmierca el at., 2002). Altogether, these data suggest that the marginal shell (including the cap area) provides information about stimulus intensity as a part of a feedback gain control system made up of the cochlea, the cochlear nuclear complex, the medial olivocochlear system, and the outer hair cells (cat, Ye et al., 2000).

The multipolar cells (Figs. 4 and 6) have tapering and moderately branched dendrites. They receive primary afferents by means of small boutons from many fibers, mainly on their dendrites (Fig. 6). Two types of multipolar cell, T- and D-stellate cells (Fig. 7), have been defined in mice on the basis of intracellular recording and labeling in in vitro slice experiments (Wickesberg and Oertel, 1988). These cells have also been described in the cat (Rhode et al., 1983a, 1983b). T-stellate cells T-stellate cells seem to correspond to the planar neurons described in rat (Figs. 6 and 7) (Doucet and Ryugo, 1997). They project via the trapezoid body to the periolivary region of the superior olivary complex, the nuclei of the lateral lemniscus, and the central nucleus of the inferior colliculus (cat, Adams, 1979, 1983b; rat, Malmierca et al., 1999a, 1999b) (Fig. 6). They are also said to supply motoneurons of the middle ear muscles (cat, Itoh et al., 1986). Frequency-specific collaterals are given off to both the VC and the DC (Fig. 7) (Lorente de Nó, 1981; cat, Smith and Rhode, 1989; mouse, Oertel and Wu, 1989; Oertel et al., 1990; rat, Doucet and Ryugo, 1997). They show “chopper” responses to tone bursts with a regularly repeated firing pattern5 and may be specialized for conveying frequency-specific excitatory information about stimulus level. D-stellate cells D-stellate cells may correspond to the large multipolar cells with nonoriented dendrites observed in Golgi preparations of rat (Figs. 6 and 7) (Harrison and Feldman, 1970, their Fig. 2) and to the commissural neurons which project to the contralateral CNC (cat, Cant and Gaston, 1982; guinea pig, Wenthold, 5

In a recent study, Ebert and Ostwald (1995) have shown that GABA alters the regular firing pattern for transient choppers units in the rat VCA.

Small Cells

Cochlear Root Neurons In apparent contrast to the situation in other mammals, the CNC of rat and some other rodents contains

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a population of large cells scattered in the cochlear nerve root (Figs. 4, 6, and 8), between the main body of the VC and the glial Schwann-cell border of the cochlear nerve in the so-called acoustic nerve nucleus or intersticial nucleus of the vestibulocochlear nerve (mouse Lorente de Nó, 1926; rat: Harrison and Warr, 1962; Harrison and Feldman, 1970; Merchán et al., 1988; Osen et al., 1991; López et al., 1993, 1999). The cochlear root neurons are calbindin-immunoreactive (López et al., 1993), have dendrites oriented across the direction of the cochlear fibers, and receive axon collaterals terminating in small boutons arising from the main stem axon of the primary afferents (Figs. 6 and 8B). The cochlear root neurons possess an exceptionally thick axon (5–7 μm) that projects, not to the brain stem auditory nuclei,6 but mainly to the contralateral reticular pontine nucleus (Fig. 6) (López et al., 1999; Nodal and López, 2003). A recent electrophysiological study in rat has shown that the cochlear root neurons have a latency of 2–3 ms, their best frequency is about 30 kHz, and they respond with a “primary-like with notch” pattern to pure tone stimulation (Figs. 8C and 8D) (Sinex et al., 2001). These data are consistent with the hypothesis that they participate in the acoustic startle reflex (Lingenhöhl and Friauf, 1992, 1994; Lee et al., 1996).

Dorsal Cochlear Nucleus This part of the CNC shows large interspecies variations, from being distinctly laminated (Figs. 4 and 5) and resembling the cerebellar cortex in rodents and carnivores to being nonlaminated in human (Moore and Osen, 1979a, 1979b) and virtually absent in some cetacea (Osen and Jansen, 1965). In rat, three layers and a central nucleus are discernible (Fig. 5) (Mugnaini et al., 1980a, 1980b), while in cat four layers and a central nucleus can be defined (Lorente de Nó, 1981; Blackstad et al., 1984). In contrast to the VC, the DC contains a large number of GABA- and/or glycine-positive, presumably inhibitory interneurons (rat, Ottersen et al., 1995; cat, Osen et al., 1990; guinea pig, Kolston et al., 1992). Pyramidal Cells The three superficial layers of the DC are related to the bipolar pyramidal (fusiform) cells (Figs. 4–6). The

6

In a detailed study López et al. (1999) have shown that although the axons from the cochlear root neurons pass through several brain stem auditory nuclei, they seem not to innervate them. Weaker projections to other nonauditory nuclei include the ventrolateral tegmental area, the oral pontine reticular nucleus, the rostral and medial paralemniscal regions, the lateral paragigantocellular nucleus, the facial motor nucleus, the intercolliculuar tegmentum, and the superior colliculus.

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apical arbor occupies layer 1, the cell body layer 2, and the basal arbor layer 3 (Figs. 4 and 5). These are the main projection neurons and supply fibers to the contralateral inferior colliculus via the dorsal acoustic stria (cat, Osen, 1972; Oliver, 1984a; rat, Malmierca et al., 1999a, 1999b) (Fig. 6). In addition, they have a direct projection to the medial division of the medial geniculate body (Malmierca et al., 2002). As shown by computer-assisted 3-D reconstructions of Golgi-impregnated material in cat, both dendritic arbors of the pyramidal cells are flattened across the long, frequency gradient axis (Fig. 3) of the DC (Blackstad et al., 1984). The highest degree of flatness and mutually parallel orientation is found in the basal arbor, which is supplied by primary afferents in a strict tonotopical manner (Figs. 3, 4, and 6) (cat, Smith and Rhode, 1985). (Pyramidal cells belong to the type IV frequency response map, Young, 1984; see below.) Interneurons of the DC may be divided into two groups related, respectively, to the apical and basal dendritic arbors of the pyramidal cells. Granule Cell System The granule cell system has been extensively studied in rat (Figs. 3, 4, and 5) (Mugnaini et al., 1980a, 1980b; Wouterlood and Mugnaini, 1984; Wouterlood et al., 1984; Ohlrogge et al., 2001). It is related to the apical arbor and cell body of the pyramidal cells and includes the excitatory granule (Fig. 6) cells and three types of presumably inhibitory cells: the Golgi cells, the molecular stellate cells, and the cartwheel cells (cat, Osen et al., 1990). The granule cells receive direct input from many sources including the somatosensory system (Fig. 6) (cat, Itoh et al., 1987; rat, Weinberg and Rustioni, 1987; Wright and Ryugo, 1996) and pontine nuclei (rat, Ohlrogge et al., 2001). Inputs from these same sources also reach the granule cells indirectly via the GABAimmunoreactive cochlear Golgi cells. The granule cells, which are glutamatergic and excitatory (cat, Godfrey et al., 1975; Osen et al., 1995), contribute parallel fibers to layer I where they form asymmetric contacts en passant with the dendritic spines of both pyramidal cells and cartwheel cells and the smooth dendrites of the stellate cells (Wouterlood and Mugnaini, 1984) (Fig. 6). The stellate cells and cartwheel cells, which are GABA-positive (Mugnaini, 1985) and glycine-positive, respectively, provide a feed-forward inhibition to the pyramidal cells (Wouterlood et al., 1984; guinea pig, Manis, 1989; cat, Osen et al., 1995; Davis et al., 1996; Davis and Young, 1997). Tuberculoventral System The tuberculoventral system interconnects the VC and the DC reciprocally (Fig. 4). It contains both frequency-

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specific and diffuse projections (Lorente de Nó, 1981; Doucet and Ryugo, 1997; Friedland et al., 2003). This system has been extensively studied in Oertel’s group in mouse slice preparations (e.g., Wickesberg and Oertel, 1988, 1993; Wickesberg et al., 1991). The frequency specific projection from the DC to the VC consists of axons of small GABA- and glycinepositive interneurons found between the basal pyramidal cell dendrites in layer 3, the so-called tuberculoventral (or vertical) cells. The relative GABA and glycine contents of these cells vary between species (Moore et al., 1996). The dendritic arbors of these cells are flattened in parallel with the pyramidal cell basal dendrites in the isofrequency planes (cat, Osen, 1983). They receive primary afferents and project their axon to the VC by way of the tuberculoventral tract after giving off recurrent collaterals to the DC, which terminate on pyramidal cells (Lorente de Nó, 1933, 1981; mouse, Oertel and Wu, 1989). Thus, these cells provide a tonotopically organized inhibition in both the DC and the VC. The frequency-specific projection in the opposite direction (from the VC to the DC) (Figs. 4 and 7) is made up of the collaterals of the presumably excitatory Tstellate (multipolar or planar) cells described above (see Doucet and Ryugo, 1997). Perhaps very precise information of intensity level is required by the DC circuitry in order to produce the exquisite sensitivity of its (type IV, see below) projection (pyramidal) neurons to spectral patterns (cat, Nelken and Young, 1994). The diffuse projection from the VC to DC is composed of axons of the glycinergic commissural D-stellate cells (or radiate, see above). The functional implications of this projection need further study, but it has been speculated that off-frequency or wideband inhibition from these cells might act as “spectral contrast detectors” (Doucet et al., 1999). In addition some small cells of the marginal shell project to the DC. They receive input from the type II auditory fibers and descending inputs and thus emerge as very interesting players in the integration of neural activity in ascending and descending systems (Doucet and Ryugo, 1997). Finally. A new type of neuron, referred to as adendric neuron, has been found to participate in the VC to DC projection (Friedland et al., 2003). The deepest layer of the DC contains two size categories of non-frequency-specific cells: the giant cells which project to the contralateral inferior colliculus (Oliver, 1984a) and smaller interneurons. In rat, however, this layer is poorly developed. Functional Significance of the DC The inhibitory interneurons are probably important in generating the “pauser” and “built-up” responses of

the pyramidal cells. The principal DC cells (pyramidal and giant) have a so-called type IV response based on their frequency response area characterized by inhibition at high levels of their best frequency and by levels of spontaneous activity (cat, Evans and Nelson, 1973; Young, 1984; Nelken and Young, 1996). These type IV units are inhibited by two mechanisms. One of them is sensitive to wide band stimuli and the other to narrow band stimuli (Nelken and Young, 1996). Most probably the first one originates in the radiate D-stellate cells in the VC and the second in the type II (tuberculoventral, vertical cells) units of the DC. Behavioral studies in cat, following surgical lesions of the dorsal and intermediate acoustic striae, speak in favor of some role in attention to sound (Masterton and Granger, 1988). The type IV units have been found to be sensitive to spectral notches that the pinna produces. Therefore it has been proposed that they may participate in sound localization, presumably in the elevation component (Young et al., 1992). But the DC may perform other sophisticated tasks. As mentioned, the rat DC projects not only to the inferior colliculus but also to the medial geniculate body (Malmierca et al., 2002). The latter in turn projects to the caudate–putamen and amygdala (LeDoux et al., 1985a, 1985b). Behavioral studies indicate that these pathways mediate the conditioned coupling of emotional responses to an acoustic stimulus (LeDoux et al., 1984). The DC is also related to multisensory integration through its connections to the medial geniculate body (Malmierca et al., 2002).

Connections with Higher Centers The ascending projections of the CNC (Fig. 3) have already been briefly mentioned and are commented on in detail when the target nuclei are described. Here it suffices to point out that the projections are largely tonotopically organized, so that the isofrequency laminae of the CNC are connected with the corresponding isofrequency laminae of the higher order centers. The right and left CNC are interconnected by the fibers of the presumably inhibitory glycinergic commissural neurons as mentioned earlier (cat, Cant and Gaston, 1982; Osen et al., 1990; guinea pig, Wenthold, 1987; rat, Doucet et al., 1999). The CNC also receives descending projections (Fig. 34) from the auditory cortex (Feliciano et al., 1995; Weedman et al., 1996a, 1996b), the inferior colliculus (cat, van Noort, 1969; rat, Caicedo and Herbert, 1993; Saldaña, 1993; guinea pig, Malmierca et al., 1996), the ventral complex of the lateral lemniscus (cat, Whitley and Henkel, 1984), and the superior olivary complex (rat, White and Warr, 1983; Horváth et al., 2000; guinea pig, Schofield, 1994). A large proportion of the latter fibers may also be inhibitory, glycine and/or GABA

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being the transmitters (guinea pig, Ostapoff et al., 1990). But there are also excitatory descending fibers, e.g., collaterals of the cholinergic olivocochlear bundle (discussed below) (Osen et al., 1984).

THE SUPERIOR OLIVARY COMPLEX The mammalian superior olivary complex (SOC, Figs. 3 and 9–13) comprises a number of closely grouped nuclei in the caudal pons. This complex shows considerable variation among species (Irving and Harrison, 1967; Harrison and Feldman, 1970; Osen et al., 1984; primates, Moore and Moore, 1971; reviewed in Heffner and Masterton, 1974, and Schwartz, 1992). The three nuclei most consistently identified across species are the lateral superior olive (LSO), the medial superior olive (MSO), and the medial nucleus of the trapezoid body (MTz7) (Fig. 9), which are also clearly defined in Ramón y Cajal’s (1904, 1909) classical drawings. The three nuclei are surrounded by diffuse cellular areas, collectively referred to as the periolivary region (PO). It may be divided into subnuclei (Morest, 1973), zones, or cell types named according to their position relative to the LSO–MSO–MTz complex (cat, Adams, 1983a; rat, Osen et al., 1984). In rat, as in other rodents, there is a fourth, welldefined nucleus, the superior paraolivary nucleus (SPO; Figs. 9–11), found in the dorsomedial part of the complex (Harrison and Feldman, 1970; Osen et al., 1984; Paxinos and Watson, 1998; Saldaña and Berrebi, 2000; Kulesza and Berrebi, 2000; guinea pig, Schofield, 1991, 1995). It projects to the ipsilateral inferior colliculus (Fig. 3) (Friauf and Kandler, 1990; Saldaña and Berrebi, 2000; guinea pig, Schofield, 1995) and may represent a hyperdevelopment of periolivary cells with a similar projection, present in smaller numbers in other mammals (Fig. 10) cat, Adams, 1983a). Within the PO, perhaps the most conspicuous nucleus in the rat is the medioventral periolivary nucleus (Osen et al., 1984), also referred to as the ventral

7 The nomenclature used for the MTz and periolivary regions surrounding the LSO–MSO in the 4th edition of The Rat Brain in Stereotaxic Coordinates (based on that of Osen et al., 1984) is somewhat different but consistent with that used here. The “medial nucleus of the trapezoid body” (MTz) is referred to as the “nucleus of the trapezoid body” (Tz), the “ventral nucleus of the trapezoid body” (VTz) as the “ventromedial periolivary region” (MVPO); and the “lateral nucleus of the trapezoid body” (LTz) as the “lateroventral periolivary region” (LVPO). We have chosen the terms MTz, VTz, and LTz, because most recent studies related to the rat SOC use them (e.g.; Smith, 1992; Friauf, 1993; Warr and Beck, 1906; Warr et al., 1997; Saldaña and Berrebi, 1990; Kulesza and Berrebi, 2000; Kulesza et al., 2002).

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nucleus of the trapezoid body (VTz; Warr and Beck, 1996) (Figs. 9–11). Another feature of the SOC is the variable, but consistently parallel, development of the LSO and MTz and the independent development of the MSO (Moore and Moore, 1971). The LSO and MTz are well developed in both the rat and the cat, while they are quite small in human (Moore and Moore, 1971). The MSO is diminutive8 in rat, while it is well developed in both the cat and human (Fig. 9). The differences may be related to the range of frequency in hearing. The MSO responds mainly to low frequencies while the LSO responds to all tonal frequencies. Both systems may be related to directional hearing. Kainic acid lesions of the SOC result in an elevation in minimum audible angles for sound localization (van Adel and Kelly, 1998). The MSO is probably more important for localization of low frequency sounds, while the LSO–MTz may code all frequencies. In the following we describe the anatomy and physiology of the LSO, MTz, MSO, SPO, and periolivary regions.

Lateral Superior Olive In transverse sections, the LSO (Figs. 9–11) appears as an S-shaped row of bipolar neurons, with dendrites oriented approximately perpendicular to the long axis (Lavilla, 1898; Ramón y Cajal, 1909; cat, Helfert and Schwartz, 1989; mice, Ollo and Schwartz, 1979; rat, Rietzel and Friauf, 1998), which is also the tonotopic axis of the nucleus: low frequencies are represented laterally and high frequencies are represented medially (cat, Tsuchitani and Boudreau, 1966; rat, Friauf, 1992). When sectioned in other planes, the cells prove to be multipolar with flattened dendritic arbors forming rostrocaudally oriented laminae which match the orientation of the afferent fiber plexus (Ramón y Cajal, 1909; cat, Scheibel and Scheibel, 1974; Cant, 1984; Friauf, 1993; rat, Rietzel and Friauf, 1998). Computer-assisted 3-D reconstructions have indicated that the thickness of the dendritic arbors varies along the tonotopic axis with the thickest arbors in the lateral low frequency areas (gerbil, Sanes et al., 1990). Intracellular injections of Lucifer yellow or neurobiotin into brain slices allow identification of seven classes of LSO neurons in the rat (Rietzel and Friauf, 1998). Bipolar and multipolar are the major cell types. The other five, less abundant,

8 Using an unbiased stereological method to estimate the total number of neurons, Kulesza et al. (2002) found that the rat MSO possesses the least number of neurons within the principal nuclei of the SOC, although the MSO is much longer in the rostrocaudal extent than previously appreciated.

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FIGURE 9 A, Comparison of the superior olivary complex in the rat and cat (redrawn after Osen et al., 1984). Note the relative size of of the LSO–MSO in the two species and the existance of a distinct SPO in the rat. (B) Camera lucida drawing of a section showing calbindin-positive neurons in the MTz and processes in the rat SOC (redrawn after Friauf, 1993). For abbreviations see list at end of chapter.

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FIGURE 10 Camera lucida drawing of a principal cell of the MTz labeled intracellularly and its projections to different nuclei of the LSO. Note the widespread projection that is frequency specific (redrawn from Banks and Smith, 1992). For abbreviations see list at end of chapter.

cell types are small multipolar, banana-like, bushy, unipolar, and marginal cells. The LSO receives input from the VCA on both sides (Figs. 3 and 6) (Harrison and Irving, 1966b). The ipsilateral input derives from spherical bushy cells and is direct and excitatory (Fig. 6) (rat, Warr, 1966; cat, Cant, 1984; Glendenning et al., 1985; Spangler et al., 1985; Brugge, 1988; Osen, 1988). Multipolar cells from the VCA whose axons travel in the trapezoid body or in the intermediate acoustic stria (Friauf and Ostwald, 1988; reviewed in Helfert and Aschoff, 1997; Thompson, 1998; Thompson and Schofield, 2000) probably also innervate the LSO. Input from the contralateral VCA originates from globular bushy cells (Henkel and Gabriele, 1999). These project across the midline to the MTz, which, in turn, has a glycinergic inhibitory (strychnine-sensitive) projection to the LSO on the same side (cat, Moore and Caspary, 1983; Saint-Marie et al., 1989; Adams and Mugnaini, 1990). Due to the thick axons of the globular cells and their large calycine synapses on the MTz cells (see below), the inputs from the two sides reach the LSO neurons simultaneously (for review, see Irvine, 1986, 1992). Consequently, LSO neurons are excited by ipsilateral sounds and inhibited by contralateral sounds

(EI units) and faithfully encode interaural intensity differences in the high frequency range of audition. The LSO projects bilaterally to the central nucleus of the inferior colliculus (Fig. 3) (cat, Shneiderman and Henkel, 1987). The ipsi- and contralateral projections are provided by different cells. Most of the ipsilaterally projecting cells are glycinergic and probably inhibitory, as shown by immunocytochemistry (cat, Glendenning and Baker, 1988; Saint-Marie et al., 1989) and retrograde transport of [3H]glycine (guinea pig, Saint-Marie and Baker, 1990). The contralaterally projecting cells are glycine-negative and probably excitatory (Saint-Marie et al., 1989; Saint-Marie and Baker, 1990). The LSO also innervates the dorsal nucleus of the lateral lemniscus bilaterally (Fig. 3). In addition, the LSO is related to the lateral olivocochlear system. In the rat, as in other rodents, cells of the lateral olivocochlear system are found in the interior of the LSO (White and Warr, 1983; Robertson, 1985; Robertson and Gummer, 1985; Aschoff and Ostwald, 1987, 1988; Helfert et al., 1988) and are either GABAergic or cholinergic (Vetter et al., 1991; Vetter and Mugnaini, 1992). In the cat, they are situated in the dorsal hilus of the LSO and may all be cholinergic (Osen and Roth, 1969; Osen et al., 1984; Warr et al., 1986).

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FIGURE 11 Glutamic acid decarboxylase (GAD) and glycine immunoreactivity in the SOC. Transverse sections through the SOC, immunoreacting against (A) GAD, the rate-limiting enzyme in the synthesis of GABA (γ-aminobutyric acid), and (B) glycine. Note the numerous GAD-immunolabeled cell bodies in the LSO, SPO, and LTz and the absence of immunolabeled cells in the MSO and MTz. The VTz and the dorsal SPO display a high degree of GAD-immunolabeled punctate profiles, interpreted as axon terminals. Glycine-immunoreactive somata are found mainly in the MTz; however, some cell body labeling is also found in the LTz. Nearly all the SOC nuclei, especially the LSO and SPO, display high densities of glycine-immunoreactive punctate profiles. Many of the cell bodies located in the LSO and SPO appear as immunonegative profiles, completely surrounded by glycine immunoreactive profiles. Figure reproduced courtesy of Dr. Randy Kulesza and Dr. Albert Berrebi. For abbreviations see list at end of chapter.

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Medial Nucleus of the Trapezoid Body The MTz constitutes the most medial part of the SOC (Figs. 9–11 and 13A). The principal cells (Fig. 10) resemble the globular bushy cells of the VCA and are situated in between fascicles of fibers in the trapezoid body. The MTz is substantial both in the cat and the rat. While in the rat it constitutes primarily a pure population of principal cells (Casey and Feldman, 1985; Banks and Smith, 1992), in the cat it contains a variety of other neurons that may belong to the PO (Morest, 1968).

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As mentioned, the MTz receives input from the VCA (Figs. 3 and 6) (Warr, 1966; van Noort, 1969; Harrison and Feldman, 1970; cat: Tolbert et al., 1982; Glendenning et al., 1985; Brugge, 1988). The fibers arise from globular bushy cells (Figs. 3 and 6) which have thick axons that terminate as large axosomatic calyces of Held (1893) (Fig. 13A) in a one-to-one relationship. The calyces constitute the largest synaptic terminals in the mammalian brain (Morest, 1973) (Fig. 13A) and provide a fast and secure relay of information from the globular bushy cells to the LSO. However, responses to acoustic stimuli of MTz neurons in vivo are not

FIGURE 12 Scheme of the different projections from the VTz (kindly provided by Dr. Bruce Warr; redrawn fom Warr and Beck, 1996). For abbreviations see list at end of chapter.

FIGURE 13 Comparison of the endbulbs of Held labeled in the MTz (A) and in the VLL (B) after injections of BDA in the rat VCA (A) and VCP (B) (Malmierca, Oliver, and Merchán, unpublished material). For abbreviations see list at end of chapter.

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always the same as those of globular bushy cells. Occasionally, they fail to show the sharp onset that is characteristic of the globular bushy cells. The fact that these cells do not always behave as simple relays may be explained by the presence of somatic synapses of noncalyceal origin (Jean-Baptiste and Morest, 1975). The source of these inputs is not known. The first description of the rat’s MTz neurons, including cytological features at EM level, was provided by Casey and Feldman (1982, 1985). More rencently, Banks and Smith (1992) and Sommer et al. (1993) have correlated the anatomy and physiology of the rat’s MTz in vitro. These authors have shown that there are principal and nonprincipal cells. The principal cells have spherical or ellipsoid somata that give rise to a single large-diameter dendrite that branches extensively, often extending beyond the MTz borders. These cells show several nonlinearities in response to current injection, including a “sag” in the membrane potential for hyperpolarizing currents and rectification for depolarizing currents (Banks and Smith, 1992). The nonprincipal cells are elongated and have very few spines. They may be the source of projections to the dorsal portion of the ventral complex of the lateral lemniscus (Banks and Smith, 1992). The MTz has a wide projection within the ipsilateral SOC (Fig. 10), including the LSO, MSO, SPO, and VTz and lateral nucleus of the trapezoid body (LTz) as well as the dorsal portion of the ventral complex of the lateral lemniscus (cat, Adams, 1979; Glendenning et al., 1981, 1985; Henkel and Spangler, 1983; Spangler et al., 1985; Adams and Mugnaini, 1990, rat, Banks and Smith, 1992; Sommer et al., 1993; reviewed in Thompson and Schofield, 2000). The projections of individual MTz neurons show an orderly topography that is consistent with the known tonotopic maps of their target nuclei (rat, Banks and Smith, 1992; Friauf, 1992). The MTz cells, their fibers, and their fiber terminals show a strong glycine immunoreactivity (cat, Saint-Marie and Baker, 1990; rat, Kulesza and Berrebi, 2000) (Fig. 11). The MTz has been shown to have a strong strychnine-sensitive, inhibitory effect on the LSO (Moore and Caspary, 1983) where strychnine-sensitive receptors have been shown (Friauf et al., 1997). Thus, the principal cells convert excitatory inputs arriving from the contralateral cochlear nucleus to inhibitory projections onto principal cells in the ipsilateral LSO. There is a wealth of experiments in vitro (in rat and mouse) that have confirmed that MTz cells are specialized to convey signals with synaptic delays that vary minutely (Sommer et al., 1993; Wu and Kelly, 1991, 1993, 1995a; Banks and Smith, 1992; Forsythe and Barnes-Davies, 1993; Forsythe, 1994; Kungel and Friauf, 1997; Löhrke et al., 1998; Borst and Sakmann, 1999; Oertel, 1999; Smith et al., 2000). In view of its many collateral projections, the MTz appears to exert an

extensive damping (tonotopic) effect on the entire SOC in response to stimulation of the contralateral ear.

Medial Superior Olive The medial superior olive (MSO) is situated between the LSO and the MTz (Figs. 3, 9–11). In transverse sections it appears as a transversely oriented row of “principal cells” with dendrites extending in the medial and lateral directions (Ramón y Cajal, 1909; Stotler, 1953). Golgi studies carried out in different species have shown that the MSO principal cells are multipolar rather than bipolar (cat, Morest, 1973; Scheibel and Scheibel, 1974; Schwartz, 1977; Henkel and Brunsø-Betchtold, 1990) and extend rostrocaudally in a series of horizontal laminae (guinea pig, Smith, 1995, see his Fig. 1). It is tonotopically organized with low frequency tones represented dorsally and high frequency tones ventrally, with most of the nucleus devoted to low frequency tones (dog, Goldberg and Brown, 1968). As mentioned, the MSO is diminutive in rat which has predominantly high frequency hearing (Fig. 9–11). The MSO also contains a small population of nonprincipal cells (marginal and multipolar). The marginal cells are oriented vertically along the margins of the nucleus. A higher proportion of terminals on the marginal cell bodies are inhibitory (cat, Lindsey, 1975) and probably GABAergic as compared to principal cells (cat, Adams and Mugnaini, 1990). These two cell types project to the inferior colliculus (Fig. 3) (cat, Adams, 1979; Henkel and Spangler, 1983). Finally, the nonprincipal multipolar cells are in the core of the MSO (cat, Morest, 1973; Scheibel and Scheibel, 1974; Kiss and Majorossy, 1983; guinea pig, Smith, 1995), but they do not project to the inferior colliculus (Adams, 1979). The MSO receives input from the VCA bilaterally (Figs. 3 and 6). The fibers arise from low frequency spherical bushy cells (rat, Harrison and Irving, 1965; Warr, 1966, 1982; Friauf and Ostwald, 1988; cat, Goldberg and Brown, 1968; van Noort, 1969; Osen, 1969; Brugge, 1988 monkey, Strominger and Strominger, 1971) (Fig. 3). Their terminal arborizations climb the horizontally oriented dendrites of the MSO cells. In contrast to the situation in the LSO, the MSO receives direct and excitatory input from both sides but the inputs remain segregated (cat, Lindsey, 1975; guinea pig, Smith, 1995). The lateral dendrites receive input from the ipsilateral side; the medial dendrites from the contralateral side (Stotler, 1953; guinea pig, Smith, 1995). The collaterals of individual axons from the spherical bushy cells travel different distances in the rostrocaudal axis so that the collaterals innervating the rostral MSO are shorter than those innervating the caudal MSO in the cat (Smith et al., 1993; Beckius et al., 1999), This organization is

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reminiscent of the required input configuration in the Jeffress model for sound localization (Jeffress, 1948). Innervation by the axons in the ipsilateral MSO do not conform to a delay line (cat, Smith et al., 1993). The direct bilateral input suggests that the MSO neurons, at least in the cat, are ideally suited to measure interaural phase or time differences (Yin and Chan, 1988; Joris et al., 1998). The marginal cells may receive CNC afferents predominantly from one side (cat, Lindsey, 1975). The MSO has also been found to receive inhibitory inputs, primarily glycinergic, from the MTz and the LTz on the same side (Figs. 3 and 6). The latter may also provide GABAergic inhibitory input (cat, Adams and Mugnaini, 1990; rat, Bank and Smith, 1992; bat, Kuwabara and Zook, 1992; gerbil, Cant and Hyson, 1992; guinea pig, Smith, 1995). The MSO projects mainly to the ipsilateral dorsal nucleus of the lateral lemniscus (cat, Oliver and Shneiderman, 1989) and the central nucleus of the inferior colliculus (Fig. 3) (cat, Henkel and Spangler, 1983). The neurons are GABA- and glycine-immunonegative (cat, Helfert et al., 1989; rat, Kulesza and Berrebi, 2000) (Fig. 11) and their projection, therefore, is probably excitatory. The concept that the function of the MSO is to analyze interaural time differences is based on the coincidence detection model of Jeffress (see above). In theory, such coincidence detection would not require inhibitory inputs, but the MTz and LTz provide MSO cells with inhibitory inputs (gerbil, Cant and Hyson, 1992). In bats, inhibition shapes MSO single unit responses (n.b., the bat’s head size may be comparable to that of the rat; Covey et al., 1991; Grothe et al., 1992, 1997; Grothe and Neuweiler, 2000; reviewed in Grothe, 2000). Thus the latter authors have proposed that interaural time difference sensitivity is an epiphenomenon and that MSO responses are too crude to serve a purpose in sound localization neurons in small animals. Instead, Grothe and Neuweiler (2000) suggest that, due to the lagging inhibitory input that MSO cells receive, its original function is to supress reverberations and echoes from the acoustic background and that the MSO performs a fundamental task of auditory analysis, i.e., the MSO creates binaural and temporal receptive fields for time contrast.

Superior Paraolivary Nucleus The SPO is a conspicuous nucleus in rodents like the rat (Fig. 9–11), gerbil, and guinea pig. It consists of multipolar cells which are the largest in the SOC. Based on studies of Golgi-impregnated material (Berrebi et al., 1997) and track-tracing methods (Saldaña and Berrebi, 2000), it has been suggested that these cells

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have an anisotropic orientation similar to the laminar organization seen, e.g., in the inferior colliculus. It has also been argued that the SPO contains multiple cells types (Harrison and Feldman, 1970; mouse, Willard and Ryugo, 1983; guinea pig, Schofield, 1991; Schofield and Cant, 1991; Kiss and Majorossy, 1996). The SPO receives inputs from octopus and multipolar cells in the contralateral VC, inputs from multipolar cells in the ipsilateral VCP (Fig. 6) (Friauf and Ostwald, 1988; guinea pig, Schofield, 1995), and a substantial glycinergic input from the MTz on the same side (Fig. 10) (Banks and Smith, 1992; Sommer et al., 1993; bat, Kuwabara and Zook, 1999). The SPO projects to the ipsilateral inferior colliculus (Fig. 3) Faye-Lund, 1986; Friauf and Kandler, 1990; Saldaña and Berrebi, 2000). It has been claimed that this projection is heterogenous as many neurons projecting to the inferior colliculus appear to use GABA, while others use glycine or excitatory neurotransmitters (guinea pig, Saint-Marie and Baker, 1990; rat, GonzálezHernández et al., 1996). But a recent study in rats using antisera directed against the 65- or 67-kDa isoform of GAD concludes that the rat SPO contains a homogeneous population of multipolar GABAergic neurons (Fig. 11) (Kulesza and Berrebi, 2000). Saldaña and Berrebi (2000) have described a topographic projection to the inferior colliculus, which strongly suggests that the SPO is tonotopic. The exact function of the SPO is still unknown, but the electrophysiological properties of these neurons are starting to emerge (Kulesza et al., 2003) and suggest that the SPO may serve to encode temporal features of complex sounds.

Periolivary Nuclei The PO (Fig. 9–12) contains several distinct types of neurons with different projection patterns (cat, Adams, 1983a). Although there is some interspecies variation, the various cell types all appear to have specific sites of location within the superior olivary complex (cat, Adams, 1983a; rat, Osen et al., 1984; Faye-Lund, 1986; and others). The PO receives input from the VC bilaterally, the lateral part from the ipsilateral side, and the medial part from both sides. These afferents arise mainly from Tstellate cells and perhaps from octopus cells (Warr, 1969) and are probably excitatory. Certain parts of the PO also receive input from the ipsilateral MTz (probably inhibitory) and from the ipsilateral inferior colliculus (probably excitatory) (cat, Andersen et al., 1980a; rat, Faye-Lund, 1986; Huffman and Henson, 1990; Saldaña, 1993) and dorsal nucleus of the lateral lemniscus (cat, Kane and Conlee, 1979; Kudo, 1981; rat, Bajo et al., 1993). PO cells project either to the cochlea, the CNC, or the inferior colliculus (cat, Adams, 1983a; rat, Warr and

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Beck, 1996). The neurons of the medial olivocochlear system, which innervate the OHCs ipsi- and contralaterally, are located medial to the MSO. In the rat they are all situated ventrally in the VTz (Fig. 12) (Warr and Beck, 1996; see below). In contrast, in the cat they are found dorsally in the dorsomedial periolivary region. Collaterals of ascending fibers from the cochlear nucleus to the inferior colliculus establish direct synaptic contact with the medial olivocochlear cells and can thus influence the OHCs via a two-neuron pathway. Of the cells projecting to the CNC, those projecting to the ipsilateral CNC are situated lateral to the MSO in the lateroventral periolivary region, while those projecting to the CNC on both sides are located medial to the MSO in the medioventral periolivary region (cat, Adams, 1983a; rat, Warr, 1969, 1972, 1982; Faye-Lund, 1986; Warr and Beck, 1996; Horváth et al., 2000). The lateral part of the PO is reciprocally connected with the ipsilateral CNC, while the medial part is reciprocally connected with the CNC on both sides. Of the cells projecting to the inferior colliculus, cells situated medial to the MSO are connected to the ipsilateral inferior colliculus, whereas those situated lateral to the MSO and predominantly ventral to the LSO project to the inferior colliculus bilaterally (Adams, 1983a). The LTz is situated ventral to the LSO. Cell types in the CNC that provide the LTz with projections include spherical- and globular-bushy cells, small cells from the marginal shell, and multipolar cells (Warr, 1966; Friauf and Ostwald, 1988; Henkel and Gabriele, 1999; cat, Ye et al., 2000). The LTz projects to the MSO (gerbil, Cant and Hyson, 1992) and is a source of inhibitory input. [The LTz also projects back to the cochlear nucleus (cat, Spangler et al., 1987) and to the inferior colliculus (guinea pig, Schofield and Cant 1991; gerbil, Cant, personal communication).] Special mention should be made of the VTz (Fig. 12), which is situated ventral to the MTz (Fig. 9). The VTz is a heterogeneous group of cells situated strategically at the intersection of ascending projections from the CNC and descending projections from the inferior colliculus (Huffman and Henson, 1990; cat, Spangler and Warr, 1991; rat, Warr and Beck, 1996). The VTz receives major afferent projections from the globular bushy cells, the octopus cells, and the multipolar cells in contralateral VC (Warr, 1972; Friauf and Ostwald, 1988; cat, Thompson, 1998). The input originating from the globular bushy cells arises from precalyceal collaterals (Friauf and Ostwald, 1988; bat, Kuwabara et al., 1991; cat, Smith et al., 1991). The VTz also receives projections from the multipolar cells in the ipsilateral VCP (cat, Thompson, 1998) and the marginal shell in the VCA from both sides (cat, Ye et al., 2000). Finally, the VTz is the major target in the SOC for the descending

projections from the inferior colliculus (Warr and Beck, 1996). Neuroanatomical and physiological evidence suggest that this descending projection may be involved in the activation of the olivocochlear neurons (cat, Rajan, 1990; rat, Vetter et al., 1993). In a comprehensive study carried out in the rat, Warr and Beck (1996) demonstrated that, in addition to projecting to the cochlea on both sides via the olivocohlear bundle, VTz neurons project to the contralateral DC (molecular layer) and LSO, and to the ipsilateral inferior colliculus, deep DC, VC, and nuclei of the lateral lemniscus including the VTz itself (Fig. 12). The VTz has also been shown to have a tonotopic organization (Friauf, 1992; Warr and Beck, 1996).

THE NUCLEI OF THE LATERAL LEMNISCUS Remarkable progress has been achieved in understanding the anatomy and physiology of the rat nuclei of the lateral lemniscus (NLL, Figs. 13–19) over the past decade (Glenn and Kelly, 1992; Li and Kelly, 1992; Bajo et al., 1993, 1998; Merchán et al., 1994; Wu and Kelly, 1995b, 1995c, 1996; Kidd and Kelly, 1996; Fu et al., 1996, 1997a, 1997b; Merchán and Berbel, 1996; Kelly et al., 1998a, 1998b; Wu, 1998, 1999; van Adel et al., 1999; Kelly and Kidd, 2000; Riquelme et al., 2001; Zhao and Wu, 2001). Unfortunately, however, the terminology used to refer to subdivisions of the NLL is problematic. Several cytoarchitectonic schemes have been proposed for different species by different authors (e.g., cat, Adams, 1979; Glendenning et al., 1981; Malmierca et al., 1998; Bajo et al., 1999; bat, Covey and Casseday, 1986, 1991; Covey 1993a, 1993b; Yang et al., 1996; rat, Paxinos and Watson, 1998; Bajo et al., 1993; Caicedo and Herbert, 1993; Merchán et al., 1994; Merchán and Berbel, 1996; Riquelme et al., 2001). Most authors agree on the boundaries of the dorsal nucleus of the lateral lemniscus (DLL; Fig. 3), but terminology referring to the ventral cells has been used inconsistently. In cat, the ventral complex of the lateral lemniscus (VLL) has been considered to be a single nucleus, called the ventral nucleus (Adams, 1979) made up of three subdivisions (ventral, middle and dorsal), each one having a distinct pattern of Nissl-stained cells. Glendenning et al. (1981), on the other hand, considered the dorsal zone to be a distinct nucleus, the intermendiate nucleus which is characterized by a projection from the MTz (Warr and Beck, 1996). This projection has been confirmed after injections of tritiated leucine in the MTz (cat, Spangler et al., 1985) and intracellular injection of single cells (rat, Sommer et al., 1993). In the present review we refer collectively to the cell groups ventral to the DLL as the ventral

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complex of the lateral lemniscus (Fig. 3). The division into VLL and DLL is consistent with the presence of two distinct functional systems: a monaural ventral and a binaural dorsal system (cat, Aitkin et al., 1970; Brugge et al., 1970; Guinan et al., 1972a, 1972b). There are some connectional, neurochemical, and physiological properties that are unique to each system. In the following, they are treated separately.

The Ventral Complex of the Lateral Lemniscus: The Monaural System The VLL consists of groups of neurons embedded within the lateral lemniscus, located between the SOC and the DLL (Fig. 3). It receives inputs mainly from the contralateral ear, as opposed to the DLL which receives inputs from both ears mostly through its connections with the superior olive. Recently, four detailed studies on the rat VLL have disclosed the anatomy and physiology of its neurons (Merchán and Berbel, 1996; Wu, 1999; Riquelme et al., 2001; Zhao and Wu, 2001). Merchán and Berbel (1996) showed that the cells exhibit a variety of shapes and sizes in Nissl-stained sections in accordance with previous studies in cat (Adams, 1979; Glendenning et al., 1981). The same authors, using tract-tracing methods combined with computer-assisted 3-D reconstructions, have shown that these cells are organized in laminae (Fig. 14) (Merchán and Berbel, 1996). After injections in the central nucleus of the inferior colliculus of the tracer biotinylated dextran amine (BDA), which is transported both anterogradely and retrogradely, Merchán and Berbel (1996) showed that the labeled cells form clusters along the dorsoventral and rostrocaudal extent of the VLL without any apparent orientation of the population of cells. Previous studies in cat (e.g., Glendenning et al., 1981; Glendening and Hutson, 1998) also described a lack of orientation. However, 3-D reconstructions, based on every section through the VLL, demonstrate that the cells do form a partly continuous structure (Fig. 14), i.e., a lamina. Furthermore, they also demonstrated that the VLL is topographically organized with respect to the frequency representation in central nucleus of the inferior colliculus (Fig. 14). Thus, Merchán and Berbel (1996) concluded that the rat VLL is a single nucleus, composed of isofrequency laminae extending through the whole dorsoventral and rostrocaudal length of the complex. It is interesting to mention in this context that injections made with biocytin and BDA in the central nucleus of the inferior colliculus of the guinea pig and cat (Malmierca et al., 1996, 1998) resulted in a similar pattern of labeled patches in the VLL, suggesting that the complex in guinea pig and cat may be organized as in the rat. Despite previous

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anatomical and physiological studies (Aitkin et al., 1970; Guinan et al., 1972a, 1972b; Whitley and Henkel, 1984) that argued against a tonotopic organization, it seems that the VLL may indeed be tonotopically organized (Merchán and Berbel, 1996; Malmierca et al., 1998). From the study of Merchán and Berbel (1996) it is also clear that virtually all cells in the rat VLL project to the central nucleus of the inferior colliculus as in the cat (Whitley and Henkel, 1984). Wu (1999) and Zhao and Wu (2001) have used wholecell patch-clamp recordings combined with intracellular labeling of cells to correlate the morphological cell types (Fig. 15), intrinsic membrane properties, and postsynaptic responses (Fig. 16) in the rat VLL neurons. Based on the morphology of the dendritic arbors, they described two main types of VLL neurons, bushy and stellate cells (Fig. 15). The bushy cells (Fig. 15D) show an onset firing pattern and nonlinear current–voltage relationship (Fig. 16). In contrast, the stellate cells (Figs. 15A and 15B) show a linear current–voltage relationship, but exhibit different firing patterns from each other (Fig. 16; v.i.). In addition, the stellate cells show differences in the shape of the soma and the dendritic branching pattern and orientation (Fig. 15). Thus, Zhao and Wu (2001) suggest that the stellate cells constitute a heterogeneous group made up of three subtypes, stellate I (Fig. 15A), stellate II (Fig. 15B), and elongate (Fig. 15C) cells. Stellate I cells show regular or onset–pause firing (Figs. 15A and 16), stellate II cells show adapting firing (Figs. 15B and 16), and elongated cells show burst firing (Figs. 15C and 16). These firing patterns have also been found in in vivo studies (cat, Aitkin et al., 1970; Guinan et al., 1972a, Adams, 1997; bat, Covey and Casseday, 1991; Covey, 1993a, 1993b; rabbit, Batra and Fitzpatrick, 1999). Finally it is worth mentioning that stimulation of the lateral lemniscus ventral to the VLL evoked EPSPs and/or IPSPs in all four cell types (Zhao and Wu, 2001). Connections Afferent projections to the VLL arise mainly from the contralateral VC (Figs. 3 and 6) and ipsilateral MTz (cat, van Noort, 1969; Glendenning et al., 1981; Spangler et al., 1985; reviewed in Merchán et al., 1997). Specific cell types from the VC project to the VLL (cat, Adams, 1979, 1997; Glendenning et al., 1981, bat, Covey, 1993a, guinea pig, Schofield and Cant, 1997; rat, Malmierca et al., 1999a, 1999b). The octopus cells project only to the ventral part of the complex, while the stellate (Fig. 6) and globular cells project to the whole complex (cat, Adams, 1979; Glendenning et al., 1981; Smith et al., 1991; rat, Friauf and Ostwald, 1988). Large terminals resembling the calyces of Held of the MTz have been shown in the ventral portion of the complex (Fig. 13B) (Malmierca et al., 1999a, 1999b) and, most probably, originate from

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the octopus cells (cat, Adams, 1997; rat, Malmierca et al., 1999a, 1999b). Thus, it seems that these cells would be suited to convey precise and secure temporal information. Both primary-like and chopper responses have been observed (bat, Covey, 1993a) after stimulation with pure tones, consistent with the idea that globular and multipolar cells take part in this projection (cat, Smith et al., 1991). Because of these connections, the VLL may be acting as a relay station in the projection to the inferior colliculus, but these responses may also be generated de novo within the VLL as suggested by the elegant study of Zhao and Wu (2001). The MTz projects to the dorsal portion of the complex (cat, Glendenning et al., 1981; Spangler et al., 1985). Others sources of projections to the VLL are periolivary nuclei on the ipsilateral side (cat, Glendenning et al., 1981; guinea pig, Schofield and Cant, 1997). It has been shown that many auditory neurons have extensive local connections in addition to their main projection (e.g., cochlear nuclear complex, Wickesberg et al. (1991, mouse); Doucet and Ryugo (1997, rat); inferior colliculus, Oliver et al. (1991, cat); Malmierca (1991, rat); Malmierca et al. (1995b, guinea pig); Reetz and Ehret (1999, mouse); medial geniculate body: Morest and Winer (1986, cat), and the same seems to hold true for the VLL neurons in rat (Zhao and Wu, 2001). Neurochemistry and Functional Significance The most detailed study of the neurochemistry of the rat VLL is that of Riquelme et al. (2001). This study supports previous findings in the cat (Saint-Marie et al., 1997) and rat (González-Hernández et al., 1996) that the majority of cells in the ventral part of the complex are glycine and/or GABA immunoreactive, although the incidence of colocalization of the two transmitters seems considerably higher than previously estimated. Indeed, Riquelme et al. (2001) have demonstrated that virtually all cells in the ventral two-thirds of the VLL colocalize9 GABA and glycine and that the dorsal part of the complex is composed exclusively of glycine- and GABA-negative cells, although the excitatory nature of these cells remains to be demonstrated (cat, SaintMarie, 1996). The larger number of glycine-positive pericellular puncta in the dorsal part of the complex is consistent with the denser input from the MTz. The afferents and intrinsic connections of the VLL may influence the discharge properties of the VLL neurons.

FIGURE 14 Computer assisted 3-D reconstruction of neurons in the rat VLL after injections of HRP and BDA into the low and high frequency regions of the IC. Note the segregation of retrogradely labeled neurons in the VLL. These findings suggest that the VLL projection to the IC is topographic. (Redrawn from Merchán and Berbel, 1996.) For abbreviations see list at end of chapter.

9 In a recent study, Lim et al. (2000) studied the functional significance of the colocalization of these two inhibitory neurontransmitters on the bushy cells in the rat VCA. They demonstrated that glycine evoked inhibitory postsynaptic currents, while activation of GABAB receptors with baclofen revealed a significant attenuation of this glycinergic inhibitory postsynaptic current. Furthermore, they stated that the effect of baclofen was presynaptic.

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FIGURE 15 Camera lucida drawings of neurobiotin-labeled cells in the VLL. Four cells types have been described: stellate I, stellate II, elongated, and bushy. (Redrawn from Zhao and Wu, 2001.) For abbreviations see list at end of chapter.

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FIGURE 16 VLL neuronal responses and current–voltage relation to injection of depolarizing (A,B1–B3) and hyperpolarizing (B4–B5) currents. Onset response is from the bushy neuron shown in Fig. 15D. Regular response is from the stellate I neuron shown in Fig. 15A. Adapting response is from the stellate II neuron shown in Fig. 15B. (Redrawn from Zhao and Wu, 2001.)

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The heavy bias of projections from the contralateral VC is reflected in the response properties of the VLL neurons, i.e., most of them respond only to contralateral stimulation, and the VLL is primarily devoted to the monaural analysis (cat, Aitkin et al., 1970; Brugge et al., 1970; Guinan et al., 1972a, 1972b). A weak projection from the ipsilateral CNC has been described (cat, Glendenning et al., 1981) and recently Batra and Fitzpatrick (1997) have shown preliminary data of binaural responses in some neurons located in the medial margin of the complex in the rabbit. But it is uncertain whether or not their recordings are outside the VLL as defined in the rat. All this together implies that the VLL neurons are suitable for encoding temporal events. Because of the ability of its cells to detect variations in the temporal features of the auditory stimulus, the VLL may be a fundamental component of the neural circuitry involved in the perception of vocalizations and speech-like communications (reviewed in Merchán et al., 1997).

The Dorsal Nucleus of the Lateral Lemniscus: The Binaural System The DLL is a distinctive group of neurons embedded within the dorsal part of the lateral lemniscus (Fig. 3) that projects bilaterally to the inferior colliculus (Fig. 17). Dorsally, it is separated from the inferior colliculus by a region devoid of neurons. Ventrally, it is delimited from the VLL by a narrow region of flat neurons. Laterally, the DLL is separated from the pial surface by the nucleus sagulum, which is populated by small neurons (cat, Henkel and Shneiderman, 1988; Shneiderman et al., 1988; Bajo et al., 1999; rat, Bajo et al., 1993; Merchán et al., 1994). Because the DLL also contains some small cells, the lateral limit cannot be determined precisely at every rostrocaudal level. In contrast to the VLL, the DLL receives input from both ears, and it projects to both inferior colliculus (Figs. 3 and 17) and also to its homolog on the opposite side (Fig. 3) through the commissure of lateral lemniscus (cll) or Probst’s commissure (Bajo et al., 1993; Zhang et al., 1998; Chen et al., 1999). Therefore, DLL cells are influenced binaurally (cat, Aitkin et al., 1970; Brugge et al., 1970; rat, Bajo et al., 1998; Kelly et al., 1998a). The DLL in the rat is cube-shaped (Bajo et al., 1993; Merchán et al., 1994). In contrast to the VLL, a similarity in structure among species is apparent, with only minor variations (Adams, 1979; Kane and Barone, 1980; Glendenning et al., 1981; Covey and Casseday, 1986; Iwahori, 1986; Shneiderman et al., 1988; Hutson et al., 1991; Wu and Kelly, 1995b; Yang et al., 1996). Several neuronal types have been proposed, depending on the species and the criteria used for cell classification. In

FIGURE 17 Laminar organization and onion-like model of the DLL. Neurons located in the center of the DLL project to the low frequency region of the CNIC while DLL neurons located in the periphery project to the high frequency region. (Redrawn from Merchán et al., 1994.) For abbreviations see list at end of chapter.

the cat, Kane and Barone (1980) described as many as nine types, while Adams and Mugnaini (1984) described only two types. Based on Nissl-stained material, Bajo et al. (1993) described four types in the rat: Type I are large and multipolar, type II are large and bipolar, type III are medium-sized with round cell bodies, and type IV are small round neurons. Similar classes have been described in the mouse (Willard and Ryugo, 1983) and opossum (Willard and Martin, 1983, 1984). More recently, using intracellular injection of biocytin in brain slice preparations of rat DLL, Wu and Kelly (1985b) have identified five types of cells based on the size and shape of both soma and dendritic arbors (Fig. 18). These include multipolar; elongate type I, type II, and type III; and round. The correspondence between the study by Bajo et al. (1993) and Wu and Kelly (1995b) is very close except for the medium-sized and elongate type III as described previously. In addition, the study by Wu and Kelly (1995b) provides a wealth of data concerning the membrane properties of the DLL cells in young rats. Despite their morphological diversity, all injected cells had similar membrane properties with a sustained

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FIGURE 18 Camera lucida drawings of neurobiotin-labeled cells in the DLL. Five cells types have been described: multipolar, round, elongate I, elongate II, and elongate III. (Redrawn from Wu and Kelly, 1995b.)

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FIGURE 19 Intracellular recording from DLL neurons of different morphological types in response to intracellular injection of positive and negative current. (Top left quadrant) Three examples of multipolar, (top right quadrant) three examples of round, (bottom left quadrant) three examples of elongate I, (bottom right) one example of elongate I and elongate II. (Redrawn from Wu and Kelly, 1995b.) (A single calibration bar provides the scale for both large and small reconstructions of each neuron.)

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series of regular action potentials produced by the injection of positive current (Fig. 19). The injection of negative current led to a hyperpolarization proportional to the current amplitude. Wu and Kelly (1995b) also describe two types of after-hyperpolarization. Some neurons have a single undershoot and others have a double undershoot after the occurrence of a spike. Merchán et al. (1994) have revealed a novel principle of organization of the DLL after injections of BDA into the rat inferior colliculus. They showed that a single, small injection of tracer into the inferior colliculus produced a distinct pattern of somatic and axonal labeling in the ipsiand contralateral DLL They also found more neurons in the contralateral DLL and more fibers in the ipsilateral DLL. The labeled structures formed an annular band in both transverse (Fig. 17) and sagittal sections. This band extends over serial sections and so they concluded that the DLL in the rat has a concentric organization of layers like that of an onion (Fig. 17). Merchán et al.’s finding strongly suggests that the DLL is primarily composed of neurons with flattened dendritic arbors as also depicted from Golgi studies (cat, Kane and Barone, 1980; Morest and Oliver, 1984; Iwahori, 1986). This concentric organization may be fundamental for the tonotopic representation in the DLL. Although a concentric tonotopic model has been suggested on the basis of functional mapping studies with c-fos (Friauf, 1992; SaintMarie et al., 1999b), it is not clear what the tonotopic organization is as derived from single-unit recording (Bajo et al., 1998; Kelly et al., 1998a). A similar concentric organization has also been recently described in the gerbil (Zook et al., 1996). In the cat (Aitkin et al., 1970; Shneiderman et al., 1988; Bajo et al., 1999) and the guinea pig (Malmierca et al., 1996) the tonotopic sequence differs slightly from the rodent model. Nevertheless, these minor variations may simply reflect evolutionary changes across species reflecting their audible range (Heffner and Masterton, 1990) rather than a genuine change in the intrinsic organization of the DLL.

bats (following injections of two different tracers into the same frequency regions of MSO and LSO in the same animal) show that the output of these olivary nuclei overlap extensively in the DLL as well as in the inferior colliculus. This opportunity for convergence at the DLL, in addition to the fact that the DLL projects to the contralateral DLL and bilaterally to the inferior colliculus, suggests that binaural processing takes place in multiple iterative steps. The studies by Kelly’s group have shown how the DLL refines the binaural response properties of inferior colliculus neurons (Liy and Kelly, 1992; Kidd and Kelly, 1996; van Adel et al., 1999; Kelly and Kidd, 2000) and supports accurate sound localization as demonstrated by behavioral studies (Ito et al., 1996 and Kelly et al., 1996). The known chemical nature of the nuclei that project to the DLL conforms with the GABA- and glycineimmunoreactive perisomatic boutons observed around the DLL neurons (cat, Saint-Marie et al., 1997; bat, Winer et al., 1995; rat, Riquelme et al., 2001). The excitatory transmitter seems to be glutamate as demonstrated in in vitro studies (v.i., Wu and Kelly, 1995b, 1995c, 1996; Wu, 1998). The neurochemistry of the DLL cells is relatively clear. Adams and Mugnaini (1984) showed that most, if not all, DLL cells use GABA as their transmitter. Since their pioneering study, several reports have confirmed the GABAergic nature of the DLL cells and their inhibitory influence on inferior colliculus responses in different animal species (Hutson et al., 1991; Shneiderman et al., 1993; Yang et al., 1992; Faingold et al., 1993; Yang and Pollak, 1994a, 1994b; 1997; Winer et al., 1995) including the rat (Zhang et al., 1998; Chen et al., 1999; Riquelme et al., 2001). The DLL projection to the inferior colliculus is laminar and bilateral, with a predominant projection to the contralateral inferior colliculus (cat, Shneiderman et al., 1988; rat, Bajo et al., 1993; Merchán et al., 1994; Zhang et al., 1998; Chen et al., 1999).

Connections and Neurochemistry

Three types of synaptic responses have been shown in the rat DLL: EPSPs only (67%), IPSPs only (6%), and combined EPSPs and IPSPs (27%) (Wu and Kelly, 1995b, 1995c; 1996; Wu, 1998). The majority of cells receive excitatory input from the lateral lemniscus. The intracellular recordings in the elegant studies by Wu and Kelly show that both EPSPs and IPSPs could be observed after pharmacological manipulation, which suggests a high degree of convergence on the same DLL cells of synaptic excitation and inhibition. This is not surprising given the large variety of input that the DLL receives from lower auditory nuclei (v.s.). Furthermore, up to 75% of the DLL neurons respond to both lemniscal and commissural inputs from the contralateral DLL.

The DLL receives input from both ears, contralaterally from the VC and DLL, ipsilaterally from the MSO, SPO, and VLL, and bilaterally from the LSO (Fig. 3) (cat, van Noort, 1969; Glendenning and Masterton, 1983; Glendenning et al., 1981; rat: van Adel and Kelly, 1998; Chen et al., 1999). Generally speaking, the DLL receives collaterals from afferents that also innervate the inferior colliculus. Careful inspection of the drawings from cat studies that are concerned with connections of the VC (Oliver, 1987), MSO, and LSO (Henkel and Spangler, 1983; Shneiderman et al., 1988) show that the projection from these nuclei to the DLL is tonotopic. Furthermore, the elegant study by Vater et al. (1995) in

Synaptic Responses

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As mentioned above, the neurotransmitter involved in the excitatory input may be glutamate (cat, Glendenning et al., 1992; rat, Madl et al., 1986; Wu, 1998). Wu and Kelly (1996) and Fu et al. (1997a, 1997b) have shown that there are two separate excitatory mechanisms operating on the DLL neurons. One is mediated by αamino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptors; the other by N-methyl-D-aspartate (NMDA) receptors. The AMPA receptors mediate a quick excitatory response; while the NMDA receptors mediate a late long-latency synaptic transmission. Thus, the NMDA synapses may result in a long-lasting inhibition both in the contralateral DLL and inferior colliculus (Kelly and Kidd, 2000). This extended inhibition in the inferior colliculus may subserve a neural mechanism for the suppression of echoes (cat, Yin, 1994; bat, Fitzpatrick et al., 1995; rat, Kelly and Kidd, 2000). Functional Significance An essential issue in the central auditory system concerns how the origin of a sound is localized. Whereas functions such as pitch or intensity coding can be performed by one ear alone, the accurate localization of a sound requires the use of both ears and depends on the comparison of the sounds they receive (for review, see Yin and Chan, 1988). The afferent and efferent connections of the DLL suggest that this nucleus plays an important role in binaural processing, i.e., sound localization. The early studies of Aitkin et al. (1970) and Brugge et al. (1970) in cat demonstrated that most DLL neurons were sensitive to binaural stimulation. These studies from Aitkin, Brugge, and co-workers showed that many DLL neurons had non-monotonic rate intensity curves, indicating that inhibitory processes were present in the DLL. More recently, in the bat (Covey, 1993b; Markovitz and Pollak, 1993, 1994; Yang et al., 1996) and in the rat (Bajo et al., 1998; Kelly et al., 1998a), studies have confirmed these earlier findings and have extended knowledge about the response properties of single neurons in the DLL. There is also pharmacological evidence on the role of the DLL in binaural processing for the DLL. The blockade of excitatory responses in DLL with kynurenic acid or lesioning the DLL with kainic acid demonstrated that there are changes in the response of neurons in the inferior colliculus to interaural time and intensity differences (rat, Li and Kelly, 1992; bat: Burger and Pollak, 2001; Pollak et al., 2002, 2003) and reduced the binaural suppression in the inferior colliculus (Kidd and Kelly, 1996; van Adel et al., 1999; Kelly and Kidd, 2000) and in the auditory cortex (Glenn and Kelly, 1992). Also, behavioral studies have shown deficits in sound localization, a reduction in spatial acuity, after lesions of DLL or after the transection of the commissure of

Probst, which results in bilateral degeneration of contralaterally projecting neurons in the DLL (Ito et al., 1996 and Kelly et al., 1996). Last, we should mention that the DLL may possess an interesting physiological feature. When the ipsilateral ear is stimulated, the inhibition imposed upon the DLL cells is relatively long-lasting. This effect extends far beyond the acoustic stimulation period and has been described both in the mustache bat (Yang and Pollak, 1994a, 1994b; Pollak et al., 2002, 2003) and in the rat (Kelly et al., 1998a). These authors suggest that this sustained inhibition may be important for localizing multiple sound sources.

THE INFERIOR COLLICULUS The inferior colliculus (IC; Figs. 20–26) is visible on the dorsal surface of the midbrain immediately caudal to the superior colliculus (Figs. 3 and 20). The IC has a key position in the auditory system as an obligatory relay center for most ascending (Fig. 3 and 6) auditory tracts (for review, see van Noort, 1969; Aitkin and Phillips, 1984a, Irvine, 1986, 1992; Malmierca, 1991; Oliver and Huerta, 1992; Ehret, 1997a; Waitzman and Oliver, 2002). In rat, it is ellipsoid with diameters 3.5 and 2 mm (Fig. 20). The long diameter (or axis) is oriented from ventral, lateral, and rostral to dorsal, medial, and caudal forming an angle of about 15° with the frontal plane and 45° with the sagittal plane (FayeLund and Osen, 1985). (In cat, it is nearly spherical with a diameter of 5.5 mm.) Ramón y Cajal (1902, 1904, 1909) identified three major subdivisions in Golgi-impregnated material from a number of mammals: the nucleus, the internuclear cortex, and the lateral cortex. Some modern authors (opposum, Willard and Martin, 1983; mouse, Willard and Ryugo, 1983; rat, Faye-Lund and Osen, 1985) have retained this simple scheme in the form of a central nucleus (CIC), a laterally and rostrally placed external cortex10 (ECIC), and a dorsal cortex (DCIC) (Fig. 20). Morest and Oliver (1984) introduced further subdivisions on the basis of their unique neuropil and neuronal populations’ in Golgi-impregnated material from cat. They subdivided the central nucleus into four subdivisions and observed differences in the dorsal and caudal cortex. Five pericentral nuclei surrounding the 10

As described below, the ECIC is made of a lateral part and a rostral part. Both have different cell types (Fig. 26) and for this reason, Malmierca (1991) defined them as two separate and distinct cortices, the lateral cortex of the IC (cf., Waitzman and Oliver, 2002) and the rostral cortex of the IC (also referred to as the intercollicular zone, cf., Herbert et al., 1991, v.s.).

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FIGURE 20 Semidiagramatic drawings of serial sections of the rat IC with the approximated Stereotaxic coordinates of the Paxinos and Watson’s atlas (1998). (Adapted from Faye-Lund and Osen, 1985). Observe the CIC covered by the DCIC dorsally and caudally, and the ECIC laterally and rostrally. Arrows in the CIC indicate the orientation of the fibrodendritic laminae. Arrow in the DCIC indicates the orientation of dendritic arbors from transitional cells. Arrow in the ECIC indicates the orientation of the dendritic arbors of the pyramidal-like, bitufted, and chandelier cells. Note that the arrow outside the CIC in section Br.- 9 is in the DCIC, but actually this region corresponds to the dorcolateralmost corner of the ECIC (cf. Fig. 21 and 22). For abbreviations see list. VI. SYSTEMS

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central nucleus and cortex included the lateral cortex of Ramón y Cajal, the commissure of the inferior colliculus, and the rostral pole. (The ECIC as it is used in the rat would include their lateral nucleus, ventrolateral nucleus, and rostral pole nucleus.) The subdivisions of the IC may not be the same size in all species, and some may be absent. For example, in the rat, the ECIC appears to occupy a relatively larger portion of the IC than in the cat (Faye-Lund and Osen, 1985; Smith, 1992; Malmierca et al., 1993, 1995a). In the bat, the “dorsoposterior division” of the CIC has a hypertrophic representation for the high tonal frequencies used for echolocation (Zook et al., 1985; Zook and Casseday, 1987). Multiple subdivisions in the CIC of the rat are unclear, but there are differences in the laminar organization in low and high frequency areas (Malmierca et al., 1993). The IC receives fibers from lower and higher auditory centers as well as from nonauditory structures (Fig. 3, 6, and 34). For a long time, it was believed that fibers from lower centers terminated in the CIC, while the fibers from the auditory cortex and nonauditory cortex sources supplied the ECIC and the DCIC (cat, Massopust and Ordy, 1962; van Noort, 1969; Andersen et al., 1980b; Wiberg, 1986; rat, Beyerl, 1978; Faye-Lund, 1985; Coleman and Clerici, 1987; LeDoux et al., 1987; Yasui et al., 1990; Herbert et al., 1991; Smith, 1992). Thus, generally speaking, ascending inputs originate in the lower auditory centers and tend to terminate more densely in the ventral portions of the IC, while the descending input from the auditory cortex and the commissural input terminate more densely in the dorsal portions of the IC. More recently however, Saldaña et al. (1996) have described in the rat a direct excitatory projection from the auditory cortex to the CIC (Figs. 21 and 34). A similar, but sparser, projection has also been described in the cat (Winer et al., 1998). Altogether, these findings suggest the presence of complementary gradients of innervation by the ascending and descending pathways to the IC. Neurons in the ventral IC, i.e., the CIC would be mainly under the influence of the lower centers, while neurons in the dorsal IC, i.e., the DCIC would be mainly under the influence of the descending pathways. Since an area of overlap between the ascending and descending inputs occurs (Saldaña et al., 1996), cells at the border of the CIC and DCIC, in particular, may be under the influence of a combination of ascending, descending, intrinsic, and commissural inputs (Figs. 3, 21, and 34). The extent of such overlapping gradients of ascending and descending inputs may differ from species to species. The IC has ascending projections to the medial geniculate body (Fig. 3) (cat, Oliver, 1984b; guinea pig, Malmierca et al., 1997; rat, González-Hernández et al.,

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1991; Peruzzi et al., 1997; Oliver et al., 1999) and descending projections to the superior olivary complex and the cochlear nuclear complex (Fig. 34) (Warr, 1992; Caicedo and Herbert, 1993; Saldaña, 1993; Vetter et al., 1993). In addition, the IC possesses well-developed commissural (cat, Aitkin and Phillips, 1984b; guinea pig, Malmierca et al., 1995b; rat, Coleman and Clerici, 1987; Saldaña and Merchán, 1992; Malmierca et al., 2001, 2003) and intrinsic (rat, Malmierca, 1991; González-Hernández et al., 1986; Saldaña and Merchán, 1992; cat, Oliver et al., 1991; guinea pig, Malmierca et al., 1995b) fiber systems (Fig. 21).

The Central Nucleus The CIC is defined by the presence of laminae distinguishable in Golgi material as a parallel orientation of afferent lemniscal fibers and neurons with flattened dendritic arbors (Figs. 22 and 23), usually referred to as “fibrodendritic laminae” (cat, Morest, 1964a). This characteristic laminar organization of the CIC has been observed in all species studied (Morest, 1964a; Geniec and Morest, 1971; Rockel and Jones, 1973a, 1973b; FitzPatrick, 1975; Willard and Martin, 1983; Morest and Oliver, 1984; Oliver and Morest, 1984 Zook et al., 1985; Ross et al., 1988; Meininger et al., 1986; Malmierca et al., 1995b) including rat (Faye-lund and Osen, 1985; Malmierca, 1991; Malmierca et al., 1993, 1995a). It constitutes the structural basis for the tonotopic organization (Fig. 3) (cat, Rose et al., 1963; Merzenich and Reid, 1974; rat, Ryan et al., 1988; Kelly et al., 1991; Friauf, 1992; Pierson and Snyder-Keller, 1995). Two main types of neurons have been defined in cat: “disk-shaped” neurons which have flattened dendritic arbors and “stellate cells” which have dendritic arbors that often transverse the laminae (Rockel and Jones, 1973a, 1973b; Morest and Oliver, 1984; Oliver and Morest, 1984; Oliver et al., 1991). In the rat, Malmierca et al. (1993, 1995a) have defined two types of neuron: F (flat) and LF (less flat) neurons by means of computerassisted 3-D reconstructions of Golgi-impregnated material (Figs. 2 and 23). The F neurons clearly conform with the definition of disk-shaped described in cat (Oliver and Morest, 1984), but the correspondence between the LF and stellate is less clear. The F and LF neurons differ in several respects, including dendritic arbor thickness, dendritic branching pattern, orientation, and their location with regard to the laminae (Figs. 22 and 23). The dendritic arbor of the F neurons is 50 μm thick and denser than that of the LF neurons. The F neurons are strictly parallel and form laminae mostly one cell thick, about 40–70 μm (FayeLund and Osen, 1985). The laminae are separated by interlaminar compartments that are populated by the

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FIGURE 21 (A) Composite diagram (adapted from Saldaña and Merchán, 1992) showing the intrinsic (right IC) and commissural (left IC) projections after three separate injections of PHA-L into the right IC. Note that the intrinsic and commissural axons originating in the CIC form distinct laminae that extend into the DCIC and ECIC. (B) Composite diagram (adapted from Saldaña et al., 1996) showing the descending projection from the primary auditory cortex to the IC. Note that these descending projections to the ipsilateral IC mimic the intrinsic ones, but those to the contralateral IC terminate only in the DCIC, the rostral part of the ECIC, and the dorsal portion of the CIC. They do not extend into the lateral part of the ECIC and the ventral part of the CIC. For abbreviations see list at end of chapter.

LF neurons (Fig. 23). The dendritic arbor of these LF neurons is 100 μm thick, and less dense than that of the F neurons. They are roughly parallel to the F neurons (Malmierca et al., 1993). The dendritic arbors of most F and LF neurons are elongated and oriented in parallel with the ventrolaterally to dorsomedially oriented long axis of the laminae. A few F neurons are oriented

rostrocaudally (Fig. 23). A gradual shift takes place so that the orientation is more horizontal in the dorsolateral part of the nucleus and more vertical in the medial part (Malmierca et al., 1993). A gradient in cell size and packing density of cell bodies also prevails with the smallest cell bodies and highest packing density in the low frequency area (Faye-Lund and Osen, 1985,

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FIGURE 22 (A) Computer assisted 3-D reconstruction of 35 neurons from the low and high frequency regions of the CIC maintaining their mutual relationship within the CIC. An arrow points to a perpendicular F neuron, an asterisk to the longest cell, and triangles to large cells in the outskirts of the CIC. Camera lucida drawings of two flat neurons (B) and less flat neurons (C). (Redrawn from Malmierca et al., 1993.) For abbreviations see list at end of chapter.

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FIGURE 23 Diagram of the laminae and interlaminar compartments of the CIC seen en face and on edge (redrawn from Malmierca et al., 1993). F neurons are confined to the laminae (50 μm thick) while LF neurons are segregated and located to the interlaminar compartments (100 μm thick). Lemniscal afferents may be centered on a lamina (a) or on a laminar compartment (b). Note F neurons with their main axis oriented either rostrocaudally or perpedicular to this axis. For abbreviations see list at end of chapter.

their Figs. 3E–3G). The interlaminar compartments were less distinct in the dorsomedial (low frequency) than in the ventromedial (high frequency) region (Fig. 22). The latter region also differed from the former by having F neurons with a higher number of intermediate and terminal dendritic segments and a denser dendritic arbor (Malmierca et al., 1993, 1995a). Using whole-cell patch-clamp techniques in CIC neurons in brain slices of rats, Sivaramakrishnan and Oliver (2001) have characterized the potassium currents

present and correlated them with the firing patterns observed by Peruzzi et al. (2000). This study demonstrated the presence of six distinct physiological cell types (Fig. 24): sustained–regular, onset, pause–build, rebound–regular, rebound–adapting, and rebound– transient. Each of these six types possess a firing pattern caused by a unique potassium current and a set of other parameters. Because of differences in ionic currents, some neurons in the CIC may act as simple relays whereas others are likely to transform their temporal

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FIGURE 24 The six firing patterns found in the IC after injecting depolarizing and hyperpolarizing current pulses. In each panel the top two traces are the voltage response to each current pulse (bottom traces). (A) Sustained-regular firing and (B) onset firing to depolarization with an anode-break spike and no calcium rebound following hyperpolarization. (C) Pause–build response to depolarization after prehyperpolarization and no anode-break spike after hyperopolarization. The cell shows a pause in firing (p) after the first spike. The response to a hyperpolarizing curent shows a buildup response (b). (D) Sustained regular firing to depolarization with an anode-break sodium spike and a calcium rebound following the hyperpolarization. (E) Sustained firing with adaptation and calcium–sodium rebound activity. (F) Transient response to depolarization with calcium–sodium rebound activity. (Figure kindly provided by Dr. Oliver; redrawn from Sivaramakrishnan and Oliver, 2001).

and intensity input. There is apparently no simple correlation between the anatomy and physiology of the F and LF cells. Peruzzi et al. (2000) showed that in 26-well filled cells, both F (23) and LF (3) neurons may have similar firing patterns. Thus, while the F and LF morphology is clearly related to the maintenance of tonotopic organization, there may be several types of F and LF cells with complex functional roles. LeBeau et al. (1995, 1996, 2001) have demonstrated in guinea pig that response types can change by modifying inhibitory input.

Afferent (Ascending) Projections The CIC receives input from several lower auditory centers (Fig. 6) (reviewed in Covey and Casseday, 1999). As demonstrated by retrograde axonal transport of tracers injected into the IC, the fibers originate in the DLL and LSO bilaterally, the VLL and MSO ipsilaterally, and the VCA, VCP, and DC contralaterally (cat, Adams, 1979, 1983b; Brunsø-Betchtold et al., 1981; Aitkin and Phillips, 1984a, 1984b; Oliver, 1984a, 1987; mouse, Ryugo et al., 1981; ferret, Henkel et al., 2003; rat, Beyerl, 1978;

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Coleman and Clerici, 1987; Bajo et al., 1993; Merchán et al., 1994; González-Hernández et al., 1996; Merchán and Berbel, 1996; Kelly et al., 1998b; Oliver et al., 1999; Malmierca et al., 1999a, 1999b). The cat has an ipsilateral projection from the VCA to the low frequency part of the CIC. It also has a bilateral projection from the higest frequencies parts of the MSO (Oliver, 1987; Oliver et al., 1997). Experiments carried out in cat using anterograde tracers, like WGA–HRP or [3H]leucine, injected into the lower centers have shown that the afferents are tonotopically organized. Besides being tonotopically organized, these experiments have shown that many of the ascending systems show a nonuniform distribution in the CIC, defined as a “banded” pattern with the dense bands parallel to, but thicker (about 200 μm, as seen with the 2-deoxiglucose, v.s.) than, the isofrequency fibrodendritic laminae (Oliver, 1984a, 1987; Shneiderman and Henkel, 1987; Shneiderman et al., 1988). According to Oliver and Shneiderman (1991) the banded pattern of bilateral projections to the CIC from nuclei like the LSO and the DLL may be more distinct on one side than on the other side. In the case of the input from the LSO, the distinct bands formed by the ipsilateral projection are intercalated between, rather than overlapping, the more diffuse bands formed by the contralateral projections. The terminal fields of the various ascending projections may also vary in extent along the main axis of the IC. While the terminal fibers from the SOC are confined to the ventral part of the CIC, those from the CNC and the DLL extend more dorsally, including also the deep region of the DCIC. Recent studies in the cat making injections physiologically characterized in the CNC and SOC in the same animal compared the distribution of afferent axons from the dorsal cochlear nucleus and the lateral superior olive to the contralateral inferior colliculus (Oliver et al., 1997). Their results show that layered axons from the dorsal cochlear nucleus and lateral superior olive are superimposed in part of the contralateral central nucleus. However, in the dorsal part of the central nucleus, the same layer of axons from the dorsal cochlear nucleus did not terminate with afferents from the lateral superior olive. In this context, preliminary results carried out in rat combining the injection of two different tracers with recordings of the neuronal activity to pure tone in the VC and DCN (Malmierca et al., 1999a, 1999b) suggest that different parts of the laminae in the CIC receive input from these nuclei that remain segregated. These data suggest that the layers may create specific functional zones in the central nucleus of the inferior colliculus. The development of some afferents to the CIC has been studied in great detail in rat. Thus, Kandler and Friauf (1993) and Niblock et al. (1995) studied the

projections from the CNC, and Gabriele et al. (2000a, 2000b) those from the DLL. The development of the CNC and DNL projections to the IC can be divided into several periods (Kandler and Friauf, 1993; Gabriele et al., 2000a, 2000b). Over the first period this development is charaterized by fiber outgrowth and commissural crossing. It starts about the same embryonic day (day 15). However during the subsequent periods characterized by axonal growth into target, collateralization in target, etc., the development of the DLL projections to the IC appears to lag 2–4 days behind the CNC projections (Gabriele et al., 2000a, 2000b). The results as discussed in Gabriele et al. (2000) indicate that some mature projection patterns are in place prior to the onset of hearing. Such findings suggest that evoked activity may not be required for the initial organization of patterned projections in the ascending auditory pathway. Efferent (Ascending) Projections The CIC projects in a strictly tonotopic manner to the laminated ventral division of the medial geniculate body (Fig. 3), largely to the ipsilateral side but also with a crossed component (tree shrew, Oliver and Hall, 1978a; cat, Oliver, 1984b; guinea pig, Malmierca et al., 1997). The CIC also has a weaker projection to the medial and dorsal divisions. The CIC projections originate from both the F and LF neurons (rat, Peruzzi et al., 1997; Oliver et al., 1999). The majority of neurons that project from the CIC to the medial geniculate body probably use glutamate as the neurotransmitter. However, recent studies show that a significant proportion of this projection uses GABA as the transmitter. Although there is little evidence for a GABAergic projection in other sensory thalamic nuclei, an ascending auditory projection from the IC to the medial geniculate body was reported recently in the cat (Winer et al., 1996). In the rat, Peruzzi et al. (1997) used immunohistochemical and tract-tracing methods to identify neurons in the IC that contain GABA and project to the medial geniculate body. They found GABA-positive projection neurons were found in the CIC and a lower proportion in the DCIC and ECIC. Peruzzi et al. (1997) also used brain slices to stimulate the brachium of the IC. Their results showed monosynaptic inhibitory postsynaptic potentials in morphologically identified thalamocortical relay neurons. Thus, these authors conclude that GABA-containing neurons in the IC make short-latency, monosynaptic inputs to the thalamocortical projection neurons in the medial geniculate body (Fig. 30). As we describe below, no GABAergic cells are present in the rat medial geniculate body (Winer and Larue, 1988) and such monosynaptic inhibitory inputs to the medial geniculate body may be important for the regulation of firing patterns

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in thalamocortical neurons in conjuntion with the known GABAergic projection from the auditory sector of the reticular thalamic nucleus (Guillery et al., 1998).

The External Cortex The definition of the ECIC varies among species and the homology in rat and cat is unclear. In rat, the ECIC covers the CIC laterally, ventrally (lateral cortex), and rostrally (rostral cortex11; Malmierca, 1991) (Figs. 3, 20, 26). Three layers are defined in the lateral part (Faye-Lund and Osen, 1985; Malmierca, 1991). Layer 1 is a continuation of the fibrodendritic capsule of the DCIC (see below). Layer 2 is composed of small- and medium-sized neurons partly aggregated in dense clusters in myelinrich neuropil. The aggregates are also rich in acetylcholinesterase (Paxinos et al., 1999a) and GABA (Ottersen and Størm-Mathisen, 1984). Layer 3 constitutes the largest part and appears to continue into the nonstratified rostral part (rostral cortex) that is topographically related to the fascicles of the commissural fibers. In addition to small- and medium-sized cells, layer 3 contains large multipolar cells especially ventromedially and rostrally. The border toward the CIC is blurred ventrally and rostrally, while dorsolaterally it is indicated by a distinct shift in dendritic orientation, particularly conspicuous in caudal transverse sections (Faye-Lund and Osen, 1985; Malmierca, 1991) where three morphologically distinct neuron types (bitufted, pyramidal-like, and chandelier; Figs. 25A–25C). have been described (Malmierca and Karagülle, 1990; Malmierca, 1991). Computer-assisted 3-D reconstruction of neurons in this region have demonstrated that their dendritic arbors are different from those of the F neurons in the CIC in several respects including their thickness and orientation (Malmierca, 1991). Interestingly, Willard and Ryugo (1983) have described that “large stellate cells with elongated dendritic arbors whose axis tends to align perpendicular to the pial surface” are present in the mouse. The presence of these types of neurons have lead some authors to conclude that, at least in rat, the lateral part of the ECIC is compatible with a cortical-like neuronal architecture in conformity with the original description made by Ramón y Cajal in 1902. In support of this idea, it has been proposed that in neonatal rats, prior to the maduration of the cerebral cortex, the modulation of sensorymotor function may be modulated by the cortices of the IC (McCown and Breese, 1991). The regions homologous

11

Also referred to as the intercollicular zone by Herbert et al. (1991, v.s.).

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to the ECIC in the rat may include the lateral ventrolateral nucleus and the rostral pole nucleus in the cat (Morest and Oliver, 1984). The ventrolateral nucleus may be relatively expanded in the rat and represent the largest component of the ECIC, in contrast to the cat where it is restricted to the bottom third of the ECIC. The ventrolateral nucleus in cat and the comparable part of the ECIC in rat both receive inputs from the lateral lemniscus (Malmierca et al., 1999a, 1999b). These parts of the ECIC have been thought to be parts of the CIC, but the details below will show that the ECIC has many differences from the CIC that justify it being identified as a separate structure. The neurons in the rostral part of the ECIC (rostral cortex) also differ from those in the CIC and the lateral part of the ECIC by being very large multipolar cells (Fig. 27E) (FayeLund and Osen, 1985; Malmierca, 1991; Malmierca et al., 1993). Besides, small- and medium-sized multipollar neurons are present in this rostral cortex (Fig. 25D). Smith (1992) and Li et al. (1998, 1999) have characterized the intracellular responses in the dorsal aspects of the ECIC and DCIC in the rat. They have shown that these cells usually responded to depolarizing current pulses of increasing intensity with phasic spiking and then, as levels increased, with tonic spiking that showed rate adaptation. Spiking also occurred at the offset after hyperpolarizing pluses. These patterns were due primarily to the activation of calcium conductances (Smith, 1992; Li et al., 1998). More recently, Hosomi et al. (1995, 1997) have reported the presence of voltage-activated Ca+2 currents in neonatal IC neurons. N’Gouemo and Rittenhouse (2000) have shown that IC neurons express three types of Ca+2 currents, namely, low-threshold, midthreshold, and high-threshold voltage-activated, on the basis of their biophysical characteristics and pharmacological sensitivities. Besides, using an in vitro study in rat, Li et al. (1998) have shown that ECIC neurons exhibit a greater degree of synaptic excitability than neurons in the DCIC and CIC. The external cortex receives input from the cerebral cortex originating immediately rostral to the primary auditory cortex (Faye-Lund, 1985; Hufman and Henson, 1990; Herbert et al., 1991; Saldaña et al., 1996). It is ipsilateral and terminates in layer 3. The ECIC also receives fibers from many nonauditory structures, including the cuneate (Coleman and Clerici, 1987) and trigeminal (cat, Wiberg, 1986) nuclei, the lateral nucleus of the subtantia nigra (Olazabal and Moore, 1989), the parabrachial region, the midbrain central gray, the periventricular nucleus (Coleman and Clerici, 1987), and the globus pallidus (Yasui et al., 1990). In agreement with the multitude of input the neurons of the ECIC have been shown electrophysiologically to repond not only to auditory, but also to somatosensory input (cat, Aitkin

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FIGURE 25 Camera lucida drawings of neurons from the ECIC (A–E) and DCIC (F–I). (A) Pyramidal-like, (B) chandelier, and (C) bi-tufted neurons from the lateral cortex. Arrow indicates main orientation perpendicular to the surface (cf. Fig. 20, Br. 9). Small (D) and very large (E) multipolar cells from the rostral cortex. (F, G) Spiny and aspinous neurons from the DCIC. (H, I) Transitional cells from the DCIC near the CIC border. The arrows indicate the orientation almost parallel to the CIC laminae (cf. Fig. 20, Lat 2 mm). Redrawn from Malmierca (1991). For abbreviations see list at end of chapter.

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et al., 1978, 1981). The ECIC in rat may also receive input from the ipsilateral medial geniculate body (Senatorov and Hu, 1999), a finding also reported in mouse (Willard and Ryugo, 1983), cat (Paydar et al., 1993), and gerbil (Kuwabara and Zook, 2000; v.i.). The ECIC projects to the dorsal and medial divisions of the medial geniculate body (Fig. 3). These projections from the three subdivisions of the IC overlap, especially in the medial divison.

The Dorsal Cortex The DCIC (Figs. 3, 20, and 25) covers the dorsomedial and caudal aspects of the CIC. In the cat it has been divided in a dorsal part with four layers and the thinner unlayered caudal part (Morest and Oliver, 1984). In rat three layers have been defined (Faye-Lund and Osen, 1985). The most superficial layer (layer 1) is the thin fibrocellular capsule that is continuous with that over the ECIC. It contains scattered, small, flattened neurons. The deeper slightly thicker layer 2 consists of small- and medium-sized, mostly multipolar neurons (Fig. 25F). These two layers together constitute about one-third the maximum thickness of the DCIC. Layer 3 contains small- and medium-sized cells (Fig. 25G). Large multipolar neurons are found at the border toward the CIC. Computer-assisted 3-D reconstructions of Golgi-impregnated neurons in the rat DCIC have demonstrated that the DCIC neurons as a group differ from the F neurons that belong to the CIC in several respects (Malmierca, 1991). There are, however, some neurons with an elongated shape located at the border between the CIC and the DCIC whose dendritic arbors parallel the orientation of the laminae. These cells have been referred to as transitional neurons (Malmierca, 1991) and may be modified disk-shaped cells (Figs. 25H and 25I). A distinct feature of neurons in the DCIC (and also the ECIC) is the presence of nitric oxide. This has been revealed by means of NADPH diaphorase staining for nitric oxide synthase in rat (Herbert et al., 1991; Druga and Syka, 1993) and immunocytochemistry in guinea pig (Coote and Rees, 1999). In guinea pig, staining for nitric oxide synthase was seen in some flat cells in the dorsal part of the central nucleus as well as in the cortices, consistent with a gradient of distribution running from the dorsal cortex toward the ventral part of the CIC. Nitric oxide is a neuromodulator that can interact with other systems such as NMDA. For this reason it has been implicated in long-term potentiation and neuronal plasticity (v.i.), a phenomenon seen in the rat IC (Zhang and Wu, 2000). However, in the guinea pig IC there was no correlation between the distribution of nitric oxide synthase and NMDA receptor subtypes (Coote and Rees, 2000). Lesioning the auditory

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cortex alters the pattern of staining of NADPH diaphorase in the rat IC (Druga and Syka, 2001). The input to the rat DCIC from the cerebral cortex (Saldaña et al., 1996) may originate largely from the primary auditory cortex bilaterally, with a small component to layer 1 from the area ventrocaudal to the primary cortex. The neocortical terminals make a tonotopic banded pattern like that of the ascending projections to the CIC, and they are tonotopically organized with the isofrequency contours continuous with and overlapping those of the CIC (monkey, Fitzpatrick and Imig, 1978; Luethke et al., 1989; cat, Andersen et al., 1980b; Winer et al., 1998; rat, Faye-Lund, 1985; Herbert et al., 1991; Saldaña et al., 1996; Druga et al., 1997). The DCIC also receives input from the sagulum (cat, Shneiderman and Henkel, 1988) and, as mentioned previously, some of the ascending input from lower auditory centers to the CIC encroaches upon the DCIC, as do the intrinsic projections (rat, Saldaña and Merchán, 1992; guinea pig, Malmierca et al., 1995b), a finding that has lead to the hypothesis that the CIC and the DCIC may form a single structure (as discussed in Irvine, 1992), although the same argument could be applied to the ECIC (v.i.). The DCIC projects to the dorsal division of the medial geniculate body.

Commissural and Intrinsic Connections Fibers that interconnect the two sides are defined as commissural while fibers that interconnect the three subdivisions of the IC on one side are defined here as intrinsic (Fig. 21). Both types of fibers may represent collaterals of axons with long projections to the thalamus or lower brain stem or, alternatively, they may represent the sole projection of a neuron that is truly an interneuron restricted to the IC (cat, van Noort, 1969; Adams, 1979; Aitkin and Phillips, 1984b; Oliver et al., 1991; mouse: González-Hernández et al., 1986; rat, Saldaña and Merchán, 1992; González-Hernández et al., 1991, 1996; Malmierca et al., 2001). Important information about the two sets of connections have derived from recent studies with anterograde axonal tracers. Small injections of Phaseolus vulgaris-leucoaggutinin (PHA-L) into the CIC of the rat have shown that the terminal territories of the intrinsic fibers form “sheets” that are parallel to the isofrequency contours of the CIC. The sheets extend into the DCIC and via a sharp bend into the ECIC (mouse, González-Hernández et al., 1986; rat, Saldaña and Merchán, 1992). The thickness varies with the injection site, apparently without any banded variation in the density of terminals as seen in some of the ascending and descending afferent systems. Preliminary results show that the majority of these projections may be excitatory (Yang et al., 2000). Labeled fibers extend over the midline

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forming a mirror-like sheet on the contralateral side, thus indicating connections between the homotopic isofrequency lamina on the two sides. GonzálezHernández et al. (1986) have also shown that the majority of cells that project to the ipsilateral medial geniculate body also send collaterals to the contralateral IC. Retrograde transport of HRP has also shown that the CIC receives input from the DCIC bilaterally and from the ECIC ipsilaterally. The DCIC and ECIC on the same side are also mutually interconnected (Coleman and Clerici, 1987; Malmierca et al., 2001). In guinea pig, it has been shown that the commissural projection may be glutamatergic (guinea pig, Saint-Marie, 1996). Recent preliminary studies on rat show that at least 25% of the commissural neurons may be GABAergic (Yang et al., 2000). Physiological evidence in vivo (rat, Malmierca et al., 2001) and in vitro (rat, Smith, 1992; Li et al., 1998, 1999; gerbil, Moore et al., 1998) shows that the commissural inputs can have either an excitatory or an inhibitory influence on the contralateral IC. In a preliminay study, Viñuela et al. (2000, 2001) have described in rat a group of neurons that project to the SOC. This group of neurons is located near the midline; its shape is cylindrical with a diameter of about 250 μm and it spans from the caudal pole of the commissure of the IC to the rostral pole of the commissure of the superior colliculus. Such neurons may correspond to those first reported by Vetter and Mugnaini (1985) being GABAergic and Faye-Lund (1986) projecting to the SOC and referred to as the “tectal commissural column” by Viñuela et al. (2000). Their afferences have been reported in Viñuela et al. (2001) and may be a source of inhibitory inputs (Mugnaini and Oertel, 1985; Vetter and Mugnaini, 1985). A preliminary report on the electrophysiological features of a few neurons (Pearson et al., 2003) suggests that they are related with frequency rather than with stimulus timing or sound localization.

Neurochemistry and Functional Significance The neurochemistry of the F and LF neurons has been studied in the rat (Roberts and Ribak, 1987; Merchán et al., 2001) and cat (Oliver et al., 1994). In cat about 20% of the cells are GABAergic while in rat the proportion may be slightly larger (up to 25%) according to Merchán et al. (2001). It seems that both F and LF may be GABAergic (Roberts and Ribak, 1987). The rat IC neurons possess GABAA, GABAB, and glycine receptors (Bowery et al., 1987; Edgar and Schwartz, 1990; Vaughn et al., 1996; Campos et al., 2001). The GABAA receptors seem to be more abundant in the DCIC than in the CIC. Studies using microiontophoresis in vivo have demonstrated that both GABA and glycine inhibit IC neurons in several species (LeBeau et al., 1995, 1996,

2001; Palombi and Caspary, 1996a; Vaughn et al., 1996), including rat (Faingold et al., 1989, 1991). In a recent study, Ma et al. (2002) have shown a role for presynaptic GABAB receptors. The rat IC neurons also possess NMDA and AMPA receptors (Petralia and Wenthold, 1992; Ishii et al., 1993; Petralia et al., 1994; Gaza and Ribak, 1997; Caicedo and Eybalin, 1999; Schmid et al., 2001). The NMDA receptors are more abundant in the cortices than in the CIC (Ohishi et al., 1993), i.e., their distribution pattern match the denser projection of the descending projections from the auditory cortex (v.i.). In this context, it is worth mentioning that it has been shown in bats (Suga et al., 2000) that this projection causes long-lasting changes in the neuronal responses of the IC, suggesting that the NMDA receptors may play an important role in neuronal plasticity. Further evidence on this role has been shown in an in vitro study in the rat by Hosomi et al. (1995) and Zhang and Wu (2000) who demonstrate that some IC neurons exhibit long-term potentation. Zhang and Kelly (2001) have studied the different physiological roles of the NMDA and AMPA receptors in the rat using the microiontophoresis application of NMDA and AMPA antagonists in vivo. Both AMPA and NMDA receptors contribute to excitatory responses at all levels of acoustic stimulation that elicit action potentials. Their results confirm the early studies by Faingold et al. (1989, 1991) related to the NMDA receptors. The NMDA and AMPA receptors have a selective influence on early and late components of tone-evoked responses (Zhang and Kelly, 2001). Thus, both AMPA and NMDA are involved in the maintenance of the response for the duration of the stimulus while it is only the AMPA receptors that seem to be important at the onset of the neuronal responses in the IC. Serotonin terminals and receptors as well as noradrenergic fibers have been reported in the rat IC (Klepper and Herbert, 1991; Thompson et al., 1994). They seem to be more abundant in the cortical regions and their origin is from the locus coeruleus and the dorsal raphe nucleus. Their functional role is unknown, although a recent study has been reported in the bat (Hurley and Pollak, 2001). Single-unit recording to pure tone stimulation (Clopton and Winfield, 1973; Kelly et al., 1991; Palombi and Caspari, 1996b, 1996c; Hernández et al., 2001) and functional mapping studies with c-fos (Friauf, 1992; Luo et al., 1999; Saint-Marie et al., 1999a) have shown that a fundamental physiological feature of the CIC is its tonotopic organization (Fig. 3). The dynamic range covers 50–60 dB and the most sensitive thresholds match the behavioral data for the rat (Kelly and Masterton, 1977). A narrow range of best frequencies is represented within each isofrequency lamina. The laminae, in addi-

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tion, are said to show a systematic topographic representation of neurons with respect to temporal response characteristics, with the frequency gradient axis of amplitude modulated tones being oriented orthogonal to the frequency gradient axis of the carrier tone, in the plane of the laminae (for review, see Schreiner and Langner, 1988, 1997). The laminae of the CIC, thus, show a highly organized representation of acoustic signals based on both spectral and temporal signals. In a preliminary study, Hernández et al. (2001) have shown that after stimulation with pure tone, neurons in the rat IC may have different types of PSTHs including onset, on-sustained, pauser and sustained, and regular responses. Similar responses have been described in the guinea pig (Rees et al., 1997). Kelly et al. (1991) and Palombi and Caspary (1996b) showed that the majority of neurons in the rat CIC have V-shaped tuning curves. More recently, Malmierca et al. (2003) have shown a larger variety of frequency response areas that include both V-shaped and non-V-shaped as described, e.g., in the guinea pig (LeBeau et al., 1995, 2001), cat (Ramachandran et al., 1999), and mouse (Egorova et al., 2001). The non-V-shaped curves form a heterogeneous group that includes the types: closed, narrow, low- and high-tilt, and multipeaked. Kelly et al. (1991) studies the sharpness of the tuning curves by means of the Q10 value. They found that maximum Q10 values were larger than previously reported and increased monotonically with the best frequency. The maximum Q10 values were larger than those previously reported for the auditory cortex at the same best frequency. Kelly et al. (1991) and Palombi and Caspary (1996c) also studied the binaural responses and classified them as either suppression, summation, or mixed. Binaural suppression responses were more numerous at high frequencies and summation at low frequencies. Studies based on the iontophoretic application of GABA and glycine antagonists have shown that neural inhibition contributes to the binaural response of neurons in the IC (Faingold et al., 1989, 1991). But only intracellular recordings can demonstrate directly that binaural responses in the IC result from the interaction between excitation and inhibition (cat, Nelson and Erulkar, 1963; Kuwada et al., 1997; guinea pig, Torterolo et al., 1995; bat, Covey et al., 1996). An important issue is why further binaural interaction needs to occur at the level of the IC given that the basic binaural comparison may have already occurred at the SOC. Comparing the comparisons may be an important component of binaural processing in some IC neurons (Casseday et al., 2002). It may also be that the convergence of binaural pathways and spectral information at the IC could produce a topographic map of the auditory space as is the case in the barn owl’s midbrain (Konishi, 1993). Such an auditory map is not clear

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in the mammalian IC (guinea pig, Withington-Wray et al., 1990; Binns et al., 1992), although recently King and associates have shown in the ferret (Doubell et al., 2000) that the nucleus of the brachium of the inferior colliculus may deliver and calibrate the representation of the auditory space in the superior colliculus. Responses in rat to amplitude modulation (AM) and frequency modulation (FM) as a function of the modulation frequency and other parameters were reported by Rees and Møller (Rees and Møller, 1983, 1987; Møller and Rees, 1986). With AM depths of only a few percent, period histograms of the spike discharge synchronized to the modulation waveform of the sound showed a marked modulation of the spike discharge. Responses became progressively less sinusoidal as modulation depth was increased. At the highest depths, the histograms from some neurons contained two peaks, consistent with the neuron showing sensitivity to the rate of change of amplitude. Modulation transfer functions based on synchronized firing to AM or FM (Rees and Møller, 1983) are often bandpass, and in some neurons a similar-shaped modulation transfer function is obtained when the response is calculated from the average firing rate. For AM the distribution of best modulation frequencies peaks between 80 and 120 Hz, but it is important to appreciate that the shape of the modulation transfer function and the best modulation frequency is influenced by the mean level of the stimulus and the presence of background noise (Rees and Møller, 1987). Transfer functions are usually low-pass at low sound levels and become bandpass as levels are increased. The best modulation frequencies of IC neurons are lower than those reported in the cochlear nuclei (Møller, 1972) where best modulation frequencies are typically greater than 250 Hz. Responses obtained in the IC to tones amplitude modulated with pseudorandom noise show that nonlinearities are an important feature in the processing of AM in this nucleus (Møller and Rees, 1986). Zhang and Kelly (2001, 2003) have demonstrated that both AMPA and NMDA receptors are involved in maintaining firing of CIC neurons to dynamically changing acoustic stimuli as well as tone bursts, but no compelling evidence was apparent for the involvement of AMPA and NMDA receptors in mediating to different rates of modulation.

THE MEDIAL GENICULATE BODY, NOT HE MEDIAL GENICULATE BODY Although the auditory thalamus is often referred to as a sensory “relay” center, suggesting a passive role limited to sensory processing, it is actually implicated in a variety of physiological processes (discussed in

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Winer et al., 1999a, 1999b). These include pure auditory processing (e.g., reviewed in Clarey et al., 1992; rat, Hu et al., 1994; Hu, 1995; Senatorov et al., 1997; Senatorov and Hu, 1997; Bartlett and Smith, 1999; guinea pig, He, 2001) and associated changes involved in learning and memory (e.g., Edeline, 1990; McIntosh and GonzálezLima, 1995, 1998). These complementary roles underscore the thalamus in the hierarchy of sensory processing (Winer et al., 1999b). The medial geniculate body (MG; Figs. 3, 26–31) lies on the posterolateral surface of the thalamus (see Groenewegen and Witter, Chapter 17, this volume) as a rounded eminence, lateral and ventral to the superior colliculus, marking the rostral pole of the brachium of the IC. It represents the main auditory center of the thalamus and is the last center for auditory processing before inputs reach the auditory cortex. In rat, the MG is 1.2 mm long in the rostrocaudal dimension, 1.5 mm wide, and 1 mm high (Fig. 26) (Winer et al., 1999b). Many ascending afferents arise in the IC (Figs. 3 and 30) (tree shrew, Oliver and Hall, 1978, cat, Andersen et al., 1980a; Oliver, 1984b; guinea pig, Malmierca et al., 1997; rat, LeDoux et al., 1985a; 1987; Oliver et al., 1999; Bartlett et al., 2000) and others originate in the CNC (Malmierca et al., 2002), SOC, and NLL (cat, Papez, 1929; Henkel, 1983; Winer, 1985, 1991; rat, LeDoux et al., 1985a, 1987). These fibers enter the MG via the brachium of the inferior colliculus (Figs. 3 and 26). The MG also receives descending fibers from the auditory cortex (v.i., for reviews, see Spangler and Warr, 1991; Winer, 1991, 1992) and the reticular thalamic nucleus (Rt; Shosaku and Sumitomo, 1983; cat, Crabtree, 1998) (Figs. 30 and 34). The rat MG differs from the IC in several respects (Winer, 1991, 1992; Winer et al., 1996). Its afferent projections are primarily ipsilateral. Its targets include the auditory cortex as well as subcortical limbic forebrain structures (e.g., amygdala and caudate nucleus). Likewise, there are no parallels for these limbic-related pathways in either the somatosensory (LeDoux et al., 1987) or the visual (Linke et al., 1999) thalamus. A further significant feature of the medial geniculate body is that only 1% of its neurons are GABAergic (Golgi type II, Winer and Larue, 1988), a feature which aligns it with the rodent ventrobasal complex and which distinguishes it from the lateral geniculate body. There are no intrinsic connections known between the different subnuclei or commissural projections interconnecting the MGs on the two sides. The MG contains three main subdivisions (Figs. 3 and 26): ventral (MGV), dorsal (MGD), and medial (MGM), each defined on the basis of cytoarchitecture and fiber connections (cat, Morest, 1964b, 1965; tree shrew, Oliver and Hall, 1975, 1978a, 1978b; Winer, 1985, 1991, 1992; Morest and Winer, 1986; rat, LeDoux et al., 1985a, 1987;

Clerici and Coleman, 1990; Clerici et al., 1990; Winer et al., 1999b, 1999c). Physiologcal studies have shown that the caudal part of the Rt (Fig. 3) is also part of the auditory pathway (cat, Simm et al., 1990; rat, Shosaku and Shumimoto, 1983; Cotillon et al., 1999; Falconi and Malmierca, 2000) and provides the MG with an inhibitory GABAergic projection (Montero, 1983; Bartlett and Smith, 1999) (Fig. 30). The thalamic nuclei surrounding the MG in the adjacent posterior, medial, and rostral parts to the MG proper constitute the caudal paralaminar nuclei (Fig. 3) (Herkenham, 1980; Linke, 1999a, 1999b), which play a key role in the acquisition of a conditioned emotional response to an acoustic stimulus (LeDoux et al., 1983; LeDoux, 1993, 1995). Finally, it is worth mentioning that recently Mitrofanis (2002) discovered an auditory area in the zona incerta of the thalamus.

Ventral Division The ventral division (MGV, Figs. 3 and 26) is the main part in the ascending auditory core projection. It is tonotopically organized (Bordi and LeDoux, 1994a,b; cat, Aitkin and Webster, 1971, 1972; rabbit, Cetas et al., 2001) and shows a laminar arrangement of afferent fibers and principal neurons (Figs. 3 and 26) with tufted dendritic arbors (Figs. 27A and 27B; cat: Morest, 1964b, 1965; Winer, 1991, 1992; rat: Clerici et al., 1990; Winer et al., 1999a,b,c). Three subdivisions with different laminar patterns occur in the rat (Fig. 26): the ventral nucleus, the ovoid nucleus, and the marginal zone (Clerici et al., 1990; Winer et al., 1999b). The dendritic bushy arbors of the tufted neurons, together with the ascending afferent axons from the IC, form fibrodendritic laminae that have a long and gently curved dorsoventral axis inclined about 40° from the stereotaxic vertical plane (Winer et al., 1999b). These laminae are the hallmark of the ventral division (cat: Morest, 1946b; rat: Winer et al., 1999b). The laminae are longer and less curved in the ventral nucleus and shorter and more coiled in the ovoid nucleus (Clerici et al., 1990; Winer et al., 1999b). This suggests that the tonotopic map may be less clear in this subdivision. Golgi type II interneurons with elaborated dendritic arborizations are abundant in the cat (Winer, 1985) and virtually absent in rat (Clerici et al., 1990; Winer and Larue, 1996; Winer et al., 1996, 1999b). This is in agreement with the rarity of GABAergic cells in the rat MG (Winer and Larue, 1988). In cat, the afferent axons and dendrites take part in the formation of complicated glomerular synaptic arrays together with the dendrites of the principal neurons and the afferent fibers (Jones and Rockel, 1971; Morest, 1971, 1975). In rat, these glomeruli are almost absent. In fact very few if any have been observed in rat main sensory thalamic nuclei of the

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FIGURE 26 (A) Nuclear boundaries of subdivisions of the rat MG. (B) Composite drawing made from a Golgi–Cox impregnation in an adult rat. Note the tufted neurons in the MGV, the stellate cells in the MGD, and magnocellular cells in the MGM. (Adapted from Winer et al., 1999b.) For abbreviations see list at end of chapter.

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FIGURE 27 Camera lucida drawings of Neurobiotin-labeled cells of the rat MG. Tufted cells populate the MGV. Cells A, B, and C are typical examples of MGV tufted cells. Cells with two distinct morphologies, tufted and stellate, populate the MGD. Cells D and E and cells F and G are typical examples of MGD tufted and stellate cells, respectively. The dendritic trees of these cells appear similar regardless of the plane of section. Besides stellate cells, cells with very long, thin, sparcely branching dendrites are seen in the MGB and SG nuclei in the coronal plane (cells H, I, and J). The dendrites extend over much greater distances dorsoventrally than those of tufted or stellate cells. Cells H and I were in the MGM; cell J was in the SG. When viewed in the horizontal plane (cell K) these cells do not show as great a dendritic extent. Cell K was in the SG. These cells may either have numerous (see enlarged portion of cell I dendrite) or few (see enlarged portion of cell J dendrite) dendritic spines. Although cells in the MGV and MGD never had axon collaterals within the MGB some cells in the MGM/SG/PIL had local collaterals (curved arrow, cell J). Scale bar applies to all drawings. Figure kindly provided by Dr. Philip Smith (some parts reproduced from Bartlett and Smith, 1999). For abbreviations see list at end of chapter.

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(Spacek and Liberman, 1974; Bartlett and Smith, 2000). Thus, the network of intrinsic neurons and the laminar arrangement may be less developed in rat than in cat (Clerici et al., 1990; Clerici and Coleman, 1990; Winer et al., 1999b). Unfortunately, there are no physiological studies that could help us to understand the functional significance of these glomeruli. Thus, any functional inference should be made with great caution. As discussed below, the Rt takes part in a feedback system that controls transmission of information from the thalamus to the cortex and may also paly a role in selective attention (Arcelli et al., 1997). One can speculate that given the lack of GABAergic cells in the rat MG, the Rt may play an important role in modulating the MG auditory signal processing in addition to its role in the regulation of the sleep–waking cycle or acting as an internal thalamic pacemaker (Arcelli et al., 1997). Another view, as discussed by Winer and Larue (1996), is that the dramatic species-specific differences in the number of GABAergic neurons (mustached bat, less than 1%; rat, about 1%; cat and monkey, 25–35%) may be related to the animal’s need to process more stereotyped (fewer interneurons) versus more complicated (more) auditory stimuli. Winer and Morest have also speculated that the synaptic glomeruli act as dynamic frequency-specific or aurally specific temporal filters adapted to the analysis of the fine structure of the stimulus, such as onset and offset behavior or frequency or AM (Winer and Morest, 1983, 1984; Winer, 1985; Huang et al., 1999). A recent in vitro study with intracellular recording and filling of cells in brain slices (Bartlett and Smith, 1999) has confirmed that the main cell type in the rat MGV is the tufted neuron (Figs. 27A and 27B). Bartlett and Smith (1999) found that these tufted neurons resemble those described in Golgi studies except that the dendritic polarization was weaker (Clerici et al., 1990; Clerici and Coleman, 1990; Winer et al., 1999b). Physiologically, tufted cells in the MGV and MGD have similar intrinsic membrane properties (Fig. 28). A unique feature of the MGV tufted neurons (and some in MGD) is the prevalence of a depolarizing sag potential (Fig. 29) as previously described (Hu, 1995). This sag is thought to reflect activation of a Cs+-sensitive mixed cation current (Ih) (Hu, 1995; Tennigkeit et al., 1996, 1998). The blockade of Ih by Cs+ hyperpolarizes the membrane potential in MGV cells and converts incoming single-spike synaptic responses into a burst (Hu, 1995). This suggests that Cs+-induced membrane potential variations play a dominant role in the regulation of the cell’s firing pattern (Hu, 1995). The main input to the MGV is from the ipsilateral CIC (Fig. 3) (tree shrew, Oliver and Hall, 1978a; cat, Andersen et al., 1980a; Oliver, 1984b; Rouiller and

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Ribaupierre, 1985; guinea pig, Malmierca et al., 1997; rat, González-Hernández et al., 1991; LeDoux et al., 1987; Peruzzi et al., 1997); although a small crossed projection from the contralateral CIC is also present (Fig. 3) (Chernock and Winer, 2001). The ipsilateral input has excitatory and inhibitory components in both cat (Winer et al., 1996) and rat (Peruzzi et al., 1997) (Fig. 30). Many of the tufted neurons from the MGV receive the convergence of excitatory and inhibitory input from the IC but a significant number of neurons receive only excitatory inputs and a few only inhibitory (Figs. 29 and 30) (Bartlett and Smith, 1999). When both excitatory and inhibitory inputs are activated simultaneously by shocking the brachium, the inhibitory inputs often reach the medial geniculate body cells before the excitatory inputs. One study in the bat (Suga et al., 1997) showed that blocking GABA inhibition in the mustached bat medial geniculate body removed the inhibitory side bands and broadened the excitatory tuning curves of cells there. Given the relative lack of GABAergic interneurons in bat medial geniculate body this would indicate that the ascending inhibition might serve to shape/narrow the frequency responses. The functional significance of more rapid conduction time of this inhibitory input is at present unknown. The axons from the IC and the auditory cortex are glutamatergic and activate both AMPA and NMDA receptors on the tufted neurons (Bartlett and Smith, 1999). The MGV also receives GABAergic input from the Rt (Montero, 1983). The stimulation of these Rt axons activates both GABAA and GABAB receptors on tufted neurons in the MGV and MGD as well as on stellate neurons in the MGD (Fig. 30) (Bartlett and Smith, 1999). Bartlett et al. (2000) found that most IC terminals make axodendritic synapses (Fig. 30) and are slightly larger in the MGV than in the MGD. Almost all cortical terminals are small, presumably arising from pyramidal cells in cortical layer VI, and end on higher order dendrites of thalamocortical cells in both the MGV and the MGD (Fig. 30). Occasional large cortical terminals, presumably arising from cortical layer V pyramidal cells, are seen but they are never located in the MGV (Fig. 30). Tennigkeit et al. (1999) found that metabotropic glutamate receptors in the MGV are expressed predominantly in the dendrites. These may be activated by repetitive stimulation of the cortical, but not the collicular, inputs (Bartlett and Smith, personal communication). The MGV is reciprocally connected with the primary auditory cortex Te1 (Figs. 3 and 31) (Ryugo and Killackey, 1974; Willard and Ryugo, 1983; Winer, 1985; Winer and Larue, 1987; Games and Winer, 1988; Roger and Arnault, 1989; Arnault and Roger 1990; Clerici and Coleman, 1990; Shi and Cassell, 1997; Winer et al., 1999c), which is tonotopically organized (Figs. 3 and 31) (Sally

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FIGURE 28 Intrinsic firing properties. Tufted (top left cell) and stellate (top right cell) cells in both the MGV and the MGD all show two firing modes that are membrane potential dependent (right column). When depolarized (top trace) a train of single spikes is evoked (tonic firing mode). As the cell is hyperpolarized the same depolarization evokes a calcium conductance (arrow in third trace) that elicits a burst of spikes (burst firing mode). When a hyperpolarizing current pulse is presented (bottom trace) a rebound burst occurs at current offset (arrow) that consists of a large calcium conductance and a high frequency spike burst. In contrast, many of the cells (left cell) in nuclei that are medial to the MGV and MGD (MGM/SG/PIL) fire tonically and do not show a burst depolarization at any membrane potential (top three traces in left column). When a hyperpolarizing current is applied these cells often do not show a rebound burst (arrow in bottom panel). An additional difference is noted in the spike waveforms. The tufted and stellate cells have monophasic after hyperpolerizations (AHPs) while many cells in medial nuclei display biphasic AHPs (Arrows in panel 3 of left column). Scale bars in third trace apply to all records in that column. The voltage next to each trace represents the cell membrane potential during that trial. Figure kindly provided by Dr. Philip Smith (some parts reproduced from Bartlett and Smith, 1999). For abbreviations see list at end of chapter.

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FIGURE 29 Synaptic inputs from the IC. Tufted cells (top two cells) in the MGV and MGD can receive either a large excitatory (designated EX/O, top trace) IC input or both excitatory and inhibitory (IN/EX or EX/IN, second trace) IC inputs. Stellate cells in the MGD (cell below MGD label) typically receive both excitation and inhibition. The inhibition can precede (IN/EX) or follow (EX/IN) the excitation. The EPSPS are glutamatergic and contain both a NMDA and a non-NMDA component. The IPSPS are GABAergic and contain both GABAA and GABAB components. Increasing the shock strength either in the presence of GABA blockers (third trace) or glutamate blockers (bottom trade) shows that the cells appear to receive several small excitatory and inhibitory inputs. Data on synaptic inputs to cells (large cell at left) in nuclei medial to the MGV and MGD are not available yet. Figure kindly provided by Dr. Philip Smith (some parts reproduced from Bartlett and Smith, 1999). For abbreviations see list at end of chapter.

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FIGURE 30 Scheme of the afferent input to MG neurons (adapted and modified from Winer and Larue, 1996, and Barlett et al., 2000). Small excitatory terminals from the IC contact high order dendrites, while medium and large IC terminals contact secondary dendritic branches. Small inhibitory IC terminals contact the soma sparsely and high order dendrites. Inhibitory input from the Rt terminates on the soma and primary dendrites. Small corticothalamic terminals contact high order dendrites and large corticothalamic terminals contact secondary dendrites. For abbreviations see list at end of chapter.

and Kelly, 1988). The MGV also receives descending projections from the secondary auditory cortex Te3 (Shi and Cassell, 1997). The role of these descending inputs, including those to the MGD and MGM, are discussed in the final section of this account dealing with the descending auditory pathway.

Dorsal Division The dorsal division occupies the caudal and dorsal part of the MG. It represents a mosaic of five regions or subnuclei (Fig. 26) whose functional roles in hearing are uncertain. These include the dorsal superficial, dorsal, deep dorsal, suprageniculate, and ventrolateral divisions (LeDoux et al., 1987; Clerici and Coleman, 1990; Clerici et al., 1990; Winer et al., 1999b). The major source of inputs to the MGD is from nonlemniscal parts of the inferior colliculus (DCIC and ECIC), and its main target is the nonprimary auditory cortex (Fig. 3). Two types of neurons are present (Figs. 27–29). Tufted neurons (Figs. 27D and 27E), with a structure similar, although not identical, to those of the MGV, and stellate neurons with radiate dendrites (Figs. 27F and 27G) (Clerici et al., 1990; Bartlett and Smith, 1999; Winer et al., 1999b). These five subdivisions differ in the concentration of their cell types and in packing density (Winer et al., 1999b) and may differ as well in their midbrain

afferents and cortical targets. The stellate cells have larger dendritic arbors than tufted cells, less branched dendrites, and less dendritic polarization (Fig. 27). The biophysical properties of the MGD neurons resemble those of the MGV (Fig. 28) (except for the prevalence of a sag in MGV cells, v.s.). Both cell types receive excitatory and inhibitory inputs from the IC, excitatory inputs from the cortex, and inhibitory inputs from the Rt (Figs. 29 and 30) (Bartlett and Smith, 1999; Bartlett et al., 2000). An additional difference between MGV and MGD neurons found by Senatorov and Hu (1997) is an enhanced activity of Na+–K+–ATPase in MGD neurons relative to MGV in a whole cell recording study. Neither the cell bodies nor the dendrites appear to be oriented in any particular fashion except in the dorsal superficial nucleus (Winer et al., 1999b). The MGD is not tonotopically organized, with the possible exception of its deepest part that abuts the MGV and shares some of its inputs from the CIC (cat, Calford and Webster, 1981). The MGD receives ascending inputs mainly from the ipsilateral DCIC and descending inputs from the Rt and cortical areas Te1, Te2, and Te3 (Figs. 3, 26, and 34) (Vaudano et al., 1990; Shi and Cassell, 1997). It projects to the second auditory cortical areas (Te2 and Te3; Fig. 3), to the insular cortex (cat, Winer et al., 1977; mouse, Willard and Ryugo, 1983; rat, Roger and Arnault, 1989; Clerici and Coleman, 1990; Arnault and Roger,

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FIGURE 31 (A) Lateral view of the auditory cortex with three main divisions based on Zilles et al. (1980, 1990). (B) Isofrequency representation in Te1 derived from the mapping studies by Sally and Kelly (1988). (C, and D) Hypothetical tonotopic representation of the rat MGV derived from the representation of unit best frequency in the albino rat (adapted from Winer et al., 1999c). For abbreviations see list at end of chapter.

1990; Winer et al., 1999c), and to the lateral nucleus of the amygdala (Doron and LeDoux, 1999). Paydar et al. (1993), Kuwabara and Zook (2000), and Senatorov and Hu (2002) have reported a descending projection from the MGD and MGM to the ECIC, but the functional role of this thalamotectal feedback is unknown.

Medial Division The medial division is the smallest of the three MG regions (Fig. 26). The MGM extends the full length of the MG as a flat lentiform nucleus about 200 μm wide and contains a diverse population of neurons, the most unusual ones being the “magnocellular” neurons (Figs. 27–29) (cat, Morest, 1964b; Winer, 1985; rat, Clerici et al., 1990; Winer et al., 1999c). A clue to the possible functions of the MGM is the diversity of its inputs. These MGM include ascending

projections from the multimodal ECIC (Fig. 3), the CNC (Malmierca et al., 2002), the SOC and VLL (cat, Henkel, 1983; rat, Bajo et al., 1993), the vestibular nuclei (cat, Roucoux-Hanus and Boisacq-Schepens, 1977), the superior colliculus (cat, Morest and Winer, 1986), and the spinal cord (cat and monkey, Berkley, 1980; mouse, Willard and Ryugo, 1983: rat, LeDoux et al., 1987). The descending input arise in the Rt and Te1 and Te3 (Fig. 34) (Rouiller and Welker, 1990; Shi and Cassell, 1997). The MGM projects to all areas of the auditory cortex (Fig. 3), but primarily to Te2 and Te3 and to nonauditory regions (Winer, 1985; Roger and Arnault, 1989; Clerici et al., 1990; Clerici and Coleman, 1990; Arnault and Roger, 1990; Winer et al., 1999c) including the somatosensory cortex (cat, Spreafico et al., 1981), the caudate–putamen, and the amygdala (Ottersen and Ben-Ari, 1979; Ottersen, 1981, 1982; LeDoux, 1985b, 1987; Doron and LeDoux, 1999). The suprageniculate nucleus

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(Figs. 26, 27J, and 27K) also projects to these areas, and its singular pattern of afferent input distinguishes it from the remainder of the dorsal divisions (Winer, 1992). For this reason it is thought by some to be part of the MGM rather than the MGD (LeDoux et al., 1987; Clerici and Coleman, 1990). The MGM and suprageniculate also participate in a descending projection to the IC (Senatorov and Hu, 2002; Winer et al., 2002).

Auditory Sector of the Reticular Thalamic Nucleus The thalamic reticular nucleus (Fig. 3) is a sheet of GABAergic cells situated along the rostral and lateral surface of the dorsal thalamus (reviewed in Guillery, 1998). The auditory sector of the Rt is located in the caudoventral region of the nucleus (Jones, 1983; Shosaku and Sumitomo, 1983). Cells in the auditory sector are topographically segregated according to the region/ nuclear subdivision of the MG they innervate (cat, Rouiller et al., 1985; Conley et al., 1991; Crabtree, 1998). The auditory sector of the Rt projects to the MG (Montero, 1983) and acts via GABAA and GABA B receptor mechanisms on MGV, MGD, and MGM cells (Fig. 30) (Bartlett and Smith, 1999). Recordings from auditory Rt cells have found that they are responsive primarily to auditory stimuli or to shock stimuli applied to the inferior colliculus or the auditory cortex (Shosaku and Sumitomo, 1983; Cotillon et al., 1999; Cotillon and Edeline, 2000; Falconi and Malmierca, 2000; cat, Simm et al., 1990). Some neurons respond with reverberating spikes at 7–10 Hz (Cotillon et al., 1999; Cotillon and Edeline, 2000; Falconi and Malmierca, 2000). Stimulation of the auditory sector of the Rt produces, as expected, suppression of the spontaneous activity of MG cells as well as their response to auditory stimuli or shock stimuli applied to the IC (Shosaku and Sumitomo, 1983). Since the MG in rat lacks local inhibitory interneurons, the role of the auditory Rt in hearing may be more important than in other species as it provides a direct inhibitory input that could modulate neuronal responses in the MG. This is supported by the preliminary results of Falconi and Malmierca (2000) that have shown some temporal, spectral, and binaural properties in singleunit responses to pure tone stimulation. The inhibitory projection from the IC to the MG is also more prominent in rat (about 40% of the cells projecting to the MG are GABAergic) than in other species (Peruzzi et al., 1997). These two inhibitory sources of projections to the rat MG might parallel the physiological role of the Golgi type II cell in other species (Figs. 30 and 34) (Huang et al., 1999; Winer et al., 1999a). The Rt feedback has been implicated in the global synchronization of thalamic neurons during certain

slow-wave phases of sleep and in certain forms of epilepsy (Steriade et al., 1986, 1993; Buzsaki et al., 1990; Steriade, 1998). Thus, Rt cells may also play a role in the synchronized activity of MG cells or during the waking state to selectively enhance the transfer of sensory information through certain thalamocortical cells (cat, Warren and Jones, 1994) and also play a role in cortical arousal during the attentive state (McDonald et al., 1998). McDonald et al. (1998) have noted that electrical stimulation of the auditory sector of the rat Rt evoke focal gamma band oscillations (40 Hz) in auditory cortex. Barth and McDonald (1996) demonstrated that gamma oscillations could occur spontaneously or be evoked in a very specific region of auditory cortex by auditory stimuli. They further showed that the oscillations could be modulated by the auditory thalamus. The finding that stimulation of the auditory sector of the Rt can elicit, presumably through its inhibitory connection with MG, oscillations in the same area of auditory cortex suggests a functional role for Rt in cognition.

Posterior Paralaminar Thalamic Nuclei This group of nuclei includes the posterior intralaminar nucleus (PIL), peripeduncular nucleus (PP), suprageniculate nucleus (considered as a distinct nucleus within the MGD subdivision by some authors) (Fig. 3) and the MGM. The PIL is a part of the intralaminar and midline thalamic nuclei that provide the cortex with a nonspecific input because of their widespread projections (Herkenham, 1980; Jones, 1983; Macchi and Bentivoglio, 1999). Recent studies suggest that the PIL and PP belong to the ascending auditory pathway in rat (Linke, 1999a) as they receive inputs from the IC (LeDoux et al., 1987; Linke, 1999a) and project into layer I of the auditory cortex (Linke, 1999b; Linke and Schwegler, 2000). These paralaminar nuclei are a source of direct sensory inputs to the amygdala. Thus, these nuclei play an important role in the processing of auditory stimuli during various emotional states (Linke et al., 1999, 2000; Linke and Schwegler, 2000; Kimura et al., 2003). Furthermore, the cortical regions that receive projections from these paralaminar thalamic nuclei give rise to amygdalopetal projections (Ottersen, 1981, 1982; Arnault and Roger, 1990; Romanski and LeDoux, 1993a; McIntyre et al., 1996; Shi and Cassell, 1997; Linke and Schwegler, 2000). Thus, two pathways may be implicated in the processing of the emotional significance of sensory stimuli, a “slow” thalamocorticoamygdaloid pathway and a “fast” thalamoamydaloid pathway (Romanski and LeDoux, 1992; Linke et al., 1999, 2000). Linke (1999a) has shown experimental evidence to suggest that these paralaminar thalamic nuclei receive

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input from the superior colliculus. Thus, these connections may represent a subcortical, “fast,” pathway for visual information transfer to the amygdala. Recent electrophysiological studies have shown a further important role for the PIL. The epipial response to electrical stimulation of the PIL evokes localized responses in the primary and secondary auditory cortices (Barth and McDonald, 1996; Brett and Barth, 1997; Sukov and Barth, 1998, 2001). PIL stimulation also produces oscillation of extracellular membrane potential in the gamma frequency band (Sukov and Barth, 2001), suggesting a role for the thalamus in their neurogenesis. These gamma rhythms are not unique to the cortex and may occur in, e.g., the Rt (v.s., Pinault and Deschênes, 1992; Steriade, 1996). Such oscillations persist after destruction of the PIL, so this nucleus may modulate the excitation driving the intrinsic cortical oscillatory mechanism by tonically depolarizing pyramidal cells rather than acting as a pacemaker for cortical gamma oscillation (Sukov and Barth, 2001). Gamma oscillations appear to reflect transient synchronization to neural populations during information processing.

Functional Significance On the basis of cytoarchitectonic, connectional, and functional features, it has been suggested (LeDoux et al., 1987; Bordi and LeDoux, 1994a, 1994b) that the MGV has a purely acoustic function, while the MGD is involved in acoustic attention, and the MGM and remaining posterior paralaminar thalamic nuclei are involved in multisensory arousal and emotional auditory learning (LeDoux and Bordi, 1994a, 1994b; Linke et al., 2000). Single-unit studies in the cat have also shown the existence of a tonotopic organization in the MGV (Imig and Morel, 1985) and the presence of both monotonic and nonmonotonic responses suggesting the presence of inhibitory mechanisms operating in the MGV (Aitkin and Dunlop, 1968, 1969). Neurons in the cat MGV respond to binaural stimuli (Aitkin and Webster, 1971, 1972; Calford, 1983; Calford and Webster, 1981). There are few physiological studies on the rat MG (e.g., LeDoux and Bordi, 1994a, 1994b; Falconi and Malmierca, 2000). The MGV possesses a tonotopic organization with low frequency tones represented dorsally and high frequency tones ventrally (LeDoux and Bordi, 1994a). About 90% of the units recorded in the rat MGV exhibit spontaneous activity (LeDoux and Bordi, 1994a, 1994b). White noise bursts produced a variety of PSTH responses that include on, sustained, reverberating, off, and on–off types. The on response was reported as the most common, both in the MGV and in other auditory thalamic regions (LeDoux and Bordi, 1994a, 1994b). Frequency tuning curves and Q10 values have also been

evaluated in the rat MGV (LeDoux and Bordi, 1994a). MGV neurons have short latencies and show a clear best frequency. For cells with best frequencies above 9 kHz, the Q10 values were significantly larger in the MGV than in other MG subdivisions. Neurons in the cat MGV have been shown to respond to binaural stimuli (Aitkin and Webster, 1971, 1972; Calford, 1983). Behavioral studies in rat (Kelly and Judge, 1985) found that lesions in the MGV have little or no effect on sound localization to noise bursts. This is consistent with the limited effect of cortical lesions on sound localization by rats but contrasts with the severe effects produced by auditory cortical lesions that resulted in MGV degeneration in other species (e.g., cat, Jenkins and Masterton, 1982; ferret, Kavanagh and Kelly, 1987). The MGD lacks a tonotopic organization. Its neurons are less responsive to acoustic stimuli, and they have latencies longer than those in the MGV (Bordi and LeDoux, 1994a, 1994b). The MGD neurons prefer complex sounds to pure tone stimuli (cat, Aitkin et al., 1966; rat, Bordi and LeDoux, 1994a, 1994b). These properties suggest that the MGD may have a role in the discrimination of sound patterns. Rat MGM cells have latencies similar to those of the MGV (Falconi and Malmierca, 2000) but the neurons prefer frequencies above 16 kHz (Bordi and LeDoux, 1994a, 1994b). Although there is no tonotopic map, cells with higher best frequencies tend to be located more ventrally in the MGM. These neurons also are tuned more broadly than those with similar best frequency in the MGV and their complex tuning curves have multipeaked responses (Bordi and LeDoux, 1994a, 1994b). MGM neurons may also respond to somatosensory or to combined auditory and somatosensory stimuli (Bordi and LeDoux, 1994b). Activation of the amygdala can antidromically activate neurons in the MGM but not in the MGV (Bordi and LeDoux, 1994b), confirming that some MGM cells project to the amygdala (LeDoux et al., 1990a, 1990b). Based on this pathway to the amygdala it is evident that the MGM is implicated in the processing of the emotional significance of acoustic stimuli (LeDoux et al., 1984, 1985b, 1986).

THE AUDITORY CORTEX The auditory cortex is part of the temporal cortex (see Palomero-Gallagher and Zilles, Chapter 23, this volume) and represents the site of termination of fibers ascending from the MG (Figs. 3 and 31–33). The auditory cortex shows large variations between species. It has been most extensively studied in the cat. In all mammals studied so far, a core koniocortex (granular cortex) with one or more complete and orderly representations of audible

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FIGURE 32 Neuronal types and laminar boundaries in the primary auditory cortex (Te1; Au1) seen in Nissl-stained sections (A), HRP-labeled cells after injections into the contralateral AC and ipsilateral IC (B), and Golgi-impregnated material (C). Note different cell types, neuronal density, and dendritic branching patterns in each of the six layers. (D–F) Insets showing from which panels B, A, and C were drawn, respectively. (Redrawn from Games and Winer, 1988.)

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FIGURE 33 Spiny cells and intrinsic firing properties of the rat auditory cortex. (Top) Camera lucida drawings of cells in layers 2–6 whose dendrites displayed spines. Pyramidal cells were observed in layers 2–6. A few spiny stellate cells were labeled in layer 4 but it is not yet clear whether these or pyramidal cells predominate in this layer. Labels next to cells indicate the response type each cell displayed to suprathreshold depolarizing current pulses (see lower part of figure). (Bottom) Responses of cells to square current pulses. (A) Regular spiking type 1 (RS1) responses had fast adaptation over the first 50–100 ms of the response and then fired spikes at regular intervals for the remainder of the current pulse. Pyramidal cells in more superficial layers and the small sample of spiny stellate cells in layer 4 tended to show this response. (B) Regular spiking type 2 (RS2) responses had spike adaptation throughout the current pulse which often led to cessation of spike firing. This response was seen in pyramidal cells in deeper layers. (C) Intrinsically bursting (IB) responses had a burst of action potentials at the beginning of the current pulse followed by single, regular spikes for the duration of the current pulse. This response was elicited by large pyramidal cells in layer 5 and deep layer 4. (D) Fast spiking (FS) responses had a rapid spike rate with little or no spike frequency adaptation for the duration of the stimulus. This response was not seen in cells with dendritic spines. Figure kindly provided by Dr. Philip Smith (some parts reproduced from Hefti and Smith, 2000).

frequencies are found to be surrounded by belt areas with less sharp frequency representation (for review, see Kelly, 1990; Winer, 1992; De Ribaupierre, 1997). In the cat, the auditory cortex comprises part of the anterior, medial and posterior ectosylvian gyri, below the suprasylvian sulcus (Winer, 1992; De Ribaupierre, 1997). Four tonotopically organized regions have been defined on the basis of responses to pure tone stimulation (Rose, 1949): the primary auditory cortex, the

anterior auditory field, the posterior auditory field, and the ventroposterior auditory field. The secondary auditory cortex and the temporal auditory area are situated ventral to the primary auditory area and show a less distinct frequency representation (Merzenich et al., 1975; Reale and Imig, 1980). In the rat, several cortical maps have been proposed (e.g., Krieg, 1946a, 1946b, 1947; Ryugo, 1976; Paterson, 1976; Cipolloni and Peter, 1979; Zilles and Wree, 1985,

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1995) but because of the lack of gross anatomical landmarks, like the neocortical sulci, a comparison of the various maps is difficult. A new system has recently been introduced by Zilles and coworkers (e.g., Zilles et al., 1980; Zilles and Wree, 1985) and is followed in principle by Schober (1986), Roger and Arnault (1989), Arnault and Roger (1990), Herbert et al. (1991), and Romanski and LeDoux (1993a, 1993b). Three temporal areas: Te1, Te2, and Te3 (Figs. 3, 31A, and 34) are defined on the basis of the “gray level index” measured in Nissl serial sections by means of computer-controlled image analysis (Zilles et al., 1980; Zilles, 1985).

Neuronal Architecture and Connections of the Cortical Auditory Fields On the basis of the patterns of thalamocortical, corticothalamic, and callosal input as well as physiological responses, the rat auditory cortex can be divided into a central core (Te1 or Au1) and a surrounding belt region that includes secondary auditory areas Te2 and Te3, (Figs. 3, 31a, and 34) (reviewed in Kelly, 1990). Temporal area 1 (Te1; Zilles and Wree, 1985; Zilles, 1990; Zilles et al., 1990) corresponds to parts of the cortical areas 41 and 39 of Krieg and it is considered to be the primary auditory cortex (Figs. 31A and 32) (Games and Winer, 1987; Roger and Arnault, 1989; Herbert et al., 1991; Romanski and LeDoux, 1993a, 1993b; Saldaña et al., 1996; Shi and Cassell, 1997). Temporal area 2 (Te2; Zilles and Wree, 1985) corresponds to parts of areas 36 and 20 of Krieg’s (1946) and temporal area 3 (Te3; Zilles and Wree, 1985) corresponds to parts of areas 20 and 14 of Krieg’s (1946a, 1946b, 1947). Te2 and Te3 constitute the secondary auditory cortices (Arnault and Roger, 1990). However, in a recent study based on the projections of these two areas, questions arose as to whether Te2 is actually a part of the auditory system or is instead related to the visual pathway (Shi and Cassell, 1997; Kimura et al., 2003). In the rat stereotaxic atlases of Swanson (1992), Paxinos and Watson (1998), and Paxinos et al. (1999b), the auditory cortex has been divided into a primary auditory cortex and a surrounding belt located dorsally, caudally, and ventrally. These belt areas compose the secondary auditory cortex and are referred to as the ventral secondary auditory cortex and the dorsal secondary auditory cortex based on the presence of nonphosphorylated neurofilament protein (recognized by the antibody SMI32) positive cells in the superficial layers. The schemes of Zilles and associates differ significantly from those of Swanson and Paxinos and Watson. We use this simple division into Te1, Te2, and Te3 by Zilles’ here for descriptive purposes and because most studies thus far related to the auditory cortex

follow it (e.g., Shi and Cassell, 1997; Winer et al., 1999c; Hefti and Smith, 2000; Doron et al., 2002; Kimura et al., 2003). But the reader is referred to Fig. 3 in PalomeroGallagher and Zilles, Chapter 23, and the atlases of Swanson (1992), Paxinos and Watson (1998), and Paxinos et al. (1999b) for an in-depth comparison of the areal structure. Based on Nissl-stained and Golgi-impregnated material obtained from adult rats, Games and Winer (1988) have analyzed the neuronal architecture of area Te1 (Fig. 32). They distinguish six distinct layers extending over about 1.1 to 1.2 mm (Fig. 32). Layer I extends about 140 μm from the pial surface and forms 13% of the total thickness of the cortex. It has very few neurons (Fig. 32A). Layer II is densely packed with many small neurons and is 125 μm thick (11% of the cortical thickness; Figs. 32A–32C). Layer III is populated by both pyramidal and nonpyramidal neurons whose dendritic arbors are heterogeneous in orientation and it is 190 μm thick (17% of the cortical thickness; Figs. 32A–32C). In layer IV neurons are smaller and slightly more densely packed than in layer III. The main cell type in layer IV is the small stellate. The majority have smooth aspinous dendrites and spiny stellates are much less common (Winer, 1992), and layer IV is only about 100 μm thick (a bit more than 10% of the cortical thickness; Figs. 32A–32C). Layer V is the thickest layer at 270 μm (26% of the cortical thickness). It begins approximately midway through the cortical depth and possesses a lower neuronal packing density and larger cells than layer IV. The main cell type is the pyramidal cell, which are larger and more numerous in the lower than in the upper half (Figs. 32A–32C). Finally, layer VI contains both pyramidal (including inverted pyramidal neurons whose “basal” dendritic arbor is directed toward the white matter) and nonpyramidal cell (bitufted and multipolar neurons) types that are closely packed. It is 245 μm thick and represents about 22% of the cortical depth (Games and Winer, 1988) (Figs. 32A and 32C). Te2 is characterized by a lack of specific differentiation of layers and Te3 exhibits the smallest cortical thickness (about 1000 μm; Roger and Arnault, 1989). Layer V is of particular interest because its cells form part of the projection to the thalamus, subthalamic nuclei, and contralateral cortex through the corpus callosum (Games and Winer, 1988; Moriizumi and Hattori, 1991; Feliciano et al., 1995; Saldaña et al., 1996; Weedman and Ryugo, 1996a, 1996b; Hefti and Smith, 2000; cat, Ojima et al., 1994). In other cortical regions two distinct types of pyramidal cells, “intrinsically bursting” and “regular spiking” can be distinguished in vitro on the basis of their correlated morphology and physiology (McCorwick et al., 1985; Kawaguchi, 1993; Kasper et al.,

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1994). These two types of pyramidal cells are also present in the primary auditory cortex in the rat (Fig. 33) and the role of inhibition in shaping their synaptic responses has been addressed by Hefti and Smith (2000, 2003). The intrinsically bursting pyramidal cells (Fig. 33) have large cell bodies and long, thick apical dendrites that branch extensively in layer I. Their axons project into subcortical white matter and arborize locally in the infragranular cortical layers. Current injections into these intrinsically bursting neurons in in vitro slice preparations has revealed a characteristic fire pattern with a burst of action potential followed by either additional bursts or single spikes (Fig. 33C) (Hefti and Smith, 2000, 2003). In contrast, the regular spiking pyramidal cells (Fig. 33) have smaller cell bodies and a thinner apical dendrite that seldom extends to layer I. Their axons also project to the white matter and arborize locally in the supragranular cortex. Current injections cause these cells to fire single spikes with a variable degree of adaptation (Figs. 33A and 33B) (Hefti and Smith, 2000, 2003). The evidence suggests that the intrinsic bursting cells receive less GABAA-mediated inhibitory input and are able to burst in response to thalamocortical synaptic stimulation far more readily than the regular spiking cells. Intrinsically bursting cells make up the majority of layer V’s input to subcortical targets (Games and Winer, 1988; Kelly, 1990; Winer, 1992; Weedman and Ryugo, 1996a, 1996b; Saldaña et al., 1996) and are capable of providing a robust input to postsynaptic neurons (Hefti and Smith, 2000). In contrast, most regular spiking neurons are strongly inhibited and may provide less robust, but perhaps more specific, information to their inputs. They may participate in a feed-forward pathway from primary to secondary and contralateral cortices (Games and Winer, 1988). The source of inhibition to these pyramidal cells must be local GABAergic neurons. To date, there are no detailed studies on the morphology and spatial distribution of GABAergic neurons in the rat auditory cortex except for a general description of the neurons containing glutamic acid decarboxylase made by Winer and Larue (1989), Ottersen and Størm-Mathisen (1984), Mugnaini and Oertel (1985), and Winer (1992, his Fig. 6.17A). Most probably, however, all cell types except for the pyramidal neurons are GABAergic in the rat primary auditory cortex (Prieto, personal communication, Ottersen and Størm-Mathisen, 1984; Winer and Larue, 1989; Winer, 1992) as is the case in the cat primary auditory cortex (Prieto et al., 1994a, 1994b). The auditory cortex is reciprocally connected with the MG, although in rat the reciprocity is not absolute (Winer and Larue, 1987). Te1 is topographically connected with the laminated MGV in a tonotopic fashion (Figs. 3 and 34) (Scheel, 1988; Roger and Arnault, 1989;

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Clerici and Coleman, 1990; Romanski and LeDoux, 1993a, Winer et al., 1999c; Kimura et al., 2003). It also receives a weak projection from the caudal parts of the MGD and the MGM (Fig. 3) (Roger and Arnault, 1989; Kimura et al., 2003). The caudally situated Te2 is connected with the MGD and MGM, and the ventrorostrally situated Te3 is connected with the MGM (Arnault and Roger, 1990; Clerici and Coleman, 1990; Winer et al., 1999c; Kimura et al., 2003). These thalamocortical fibers terminate in layers III and IV and at the junction of layers V and VI (Ryugo and Killackey, 1974; Ryugo, 1976; Vaughan, 1983; Mitani et al., 1985; Romanski and LeDoux, 1993a). The callosal fibers (Figs. 3 and 34) constitute another important fiber system which interconnects homotopic and heterotopic areas of the left and right auditory cortex (Ruttgers et al., 1990). These fibers terminate in all layers (Cipolloni and Peters, 1979, 1983; Vaughan, 1983; Mitani et al., 1985; Roger and Arnault, 1989) and originate from all layers with a preponderance of neurons lying in the infragranular layers (Granger et al., 1985; Ruttgers et al., 1990). Iontophoretic injections of PHA-L in the rat auditory cortex have revealed a system of intrinsic fibers that run from rostral to caudal (Collía et al., 1990; Lanciego, 1994). These corticocortical projections form columns, perpendicular to the tangential surface of the cortex, that range from 200 to 800 μm in width and are made up of terminal axons that extend through all layers from I to VI (Romanski and LeDoux, 1993b). Te1 projects to the secondary auditory Te2 and Te3 cortical areas (Romanski and LeDoux, 1993b). The auditory cortex projects to nonauditory areas too, like the cingulate gyrus and the perirhinal cortex (Rouiller et al., 1990, cat). Te2 and Te3 also project to the amygdala (Romanski and LeDoux, 1993a; Shi and Cassell, 1997). Studies have also shown that the auditory cortex is connected to the cerebellar cortex. Electrical stimulation of regions corresponding to parts of Te1 generates neuronal responses in the parafloculus of the cerebellum that are similar to those produced by acoustical stimulation (reviewed in Kelly, 1990). The parafloculus also receives input from the auditory cortex through a corticopontine cerebellar pathway (Azizi et al., 1985). The auditory cortex also gives rise to a descending pathway to the thalamus, inferior colliculus, and pons (Fig. 34). These corticothalamic, corticocollicular, and corticopontine systems are commented on in the next section.

Physiological Studies A cochleotopic representation in the primary auditory cortex was first shown in the dog by Tunturi (1952)

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FIGURE 34 Descending auditory pathway of the rat (modified after Brodal, 1981; AC is from Herbert et al., 1991). For abbreviations see list at end of chapter.

and in the rat by Horikawa et al. (1988) and Sally and Kelly (1988) (Figs. 3 and 31B) using single-unit recordings to pure tone stimulation. In rat the representation of high frequencies is located rostrally in Te1 and low frequencies caudally (Fig. 3) (Sally and Kelly, 1988; Kilgard

and Merzenich, 1998, 1999). Neurons at different dorsoventral positions within Te1 have similar best frequencies; consequently, for each rostrocaudal location there is a dorsoventrally oriented isofrequency contour (Fig. 3). These contours extend about 1 mm across the

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cortical surface. Up to 80% of Te1 is devoted to frequencies above 8 kHz. This disproportion is clearly seen when the best frequency of single cells is plotted as a function of the location with the Te1 (Kelly and Sally, 1988). The majority of neurons show short latency responses (8–14 ms) and monotonic rate-intensity functions. Their dynamic range is usually from 5 to 35 dB although it can be as great as 60 dB. Neurons with nonmonotonic rate-intensity functions are also present and their dynamic range is larger, from 30 to 40 dB, but sometimes reaching up to 80 dB (Phillips and Kelly, 1989). The poststimulus responses to pure tone stimulation are characterized by a weak period of suppression followed by a series of bursts with a lower firing rate that appear after a short latency response (Sally and Kelly, 1988). At the borders of Te1, responses have longer latencies. Sally and Kelly have also reported V-shaped tuning curves with Q10 values ranging from 0.97 to 28.4 that increase as a function of the unit best frequency. The lowest thresholds at best frequency usually matched the behavioral audiogram of the rat for many neurons but, interestingly, Sally and Kelly (1988) also found many neurons whose thresholds were above the behavioral limit. Dorsal and ventral to the primary auditory cortex best frequencies are discontinuous with the neighboring isofrequency contours (Sally and Kelly, 1988), suggesting the existence of secondary auditory regions. This conforms to the anatomical mapping of the auditory cortex. In a recent study in the rat, Gaese and Ostwald (2001) have pointed out that the anesthesia can affect responses in auditory cortex. They found that anesthesia reduced spontaneous activity, suppressed sustained responses, and narrowed frequency tuning curves presumably through enhanced inhibitory mechanisms. As demonstrated by Kelly and Sally (1988), over 95% of the units in the auditory cortex exhibit summation, suppression, or mixed binaural interactions (as defined by Goldberg and Brown, 1969) similar to those found in other species (e.g., Brugge et al., 1969; Brugge and Merzenich, 1973; Imig and Adrian, 1977; Kitzes et al., 1980; Middlebrooks et al., 1980; reviewed in Kelly, 1990, and Clarey et al., 1992). The summation type neurons, i.e., those excited by either ear alone or facilitated by stimulation of both ears together, represent about 35% of all units recorded in the study of Kelly and Sally (1988). The suppression type neurons, i.e., those excited by contralateral stimulation and inhibited under binaural stimulation, represent about 42%. Mixed neurons, i.e., those facilitated by binaural stimulation near threshold, but inhibited at higher sound pressure levels, represent about 20%. Although all binaural types are found over a wide range of sound frequencies, the summation type prevails at low fre-

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quencies and the suppression type at high frequencies (Kelly and Sally, 1988). Another interesting feature about the binaural interaction is that cells of the same binaural response type tend to be grouped together and form aggregates of summation, suppression, or mixed patterns (Kelly and Sally, 1988). Cortical areas with similar binaural response properties appear to extend across isofrequency contours to form rostrocaudally oriented bands as found in other species (e.g., Imig and Adrian, 1977; Middlebrooks et al., 1980; Middlebrook and Zook, 1983, reviewed in Clarey et al., 1992; v.i.). The early studies of Evans and Whitfield (1964) showed that units in the cat auditory cortex respond better to FM than to pure tone stimulation. Gaese and Ostwald (1995) have studied the neuronal responses to sinusoidally amplitude-modulated and frequency-modulated stimuli in the rat auditory cortex. They have shown that the tuning characteristics to the modulation frequency are mainly bandpass with best modulation frequencies between 4 and 15 Hz. They also showed that the vast majority of the units are influenced by varying the modulation frequency whereas varying the rate of frequency change had little effect. More recently, Ricketts et al. (1998) have shown that the majority of neurons prefer fast FM sweeps. Orduña et al. (2001) have confirmed that the response patterns exhibited by rat auditory cortical neurons to FM sounds are similar to those reported by Gaese and Ostwald and those observed in the auditory cortex of other species. Since the trains of up-sweeps and down-sweeps used by Orduña et al. (2001) were spectrally equivalent, they suggest that the different responses produced by these stimuli demonstrate that a pure spectral analysis is not enough to predict how cortical neurons will respond to complex sound. They further show that neuronal selectivity for repetition rate is an important factor in determining how cells will respond. Thus, Orduña et al. (2001) conclude that what determines the response properties of auditory cortical neurons is a dynamic interaction between the temporal and spectral features of the sound.

Modular Organization The columnar organization of the sensory cortices has attracted considerable attention over the last 40 years. There is some experimental evidence of a columnar organization in the auditory cortex. According to Merzenich et al. (1975), each of the isofrequency strips of the cat primary auditory cortex represents a set of isofrequency columns with the same characteristic frequency for all units throughout the entire depth of the cortex. Binaural interaction columns have also been described, exhibiting either binaural summation or

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inhibition (cat, Imig and Adrian, 1977; Middlebrooks et al., 1980). These columns are organized in strips running roughly at right angles to the isofrequency contours (cat, Middlebrooks et al., 1980). Nevertheless, the existence of a binaural functional organization orthogonal to the isofrequency contours has been questioned by some authors and a detailed discussion on the issue is provided by Clarey et al. (1992). In addition to the best frequency and binaurality, there are other features of the sound that are represented in the primary auditory cortex, at least in cat. These include threshold, monotonicity, dynamic range, latency, FM speed, and width of tuning curves (Ehret, 1997b; rat: Kilgard and Merzenich, 1999). Furthermore, Read et al. (2001, 2002) have demonstrated that, in cat, neurons with narrow band frequency response areas within an isofrequency contour form patches and constitute a functionally and anatomically segregated system. Likewise, other patches contain neurons suited for broadband spectral processing. Whether or not this modular organization of intrinsic connections associated with spectral tuning is a general feature of mammalian primary auditory cortex, and hence is present in the rat, is unclear. Physiological studies in primates have suggested the existence of multiple spectral tuning modules (Recanzone et al., 1999; Recanzone, 2001; Cheung et al., 2001) but in rat (Kilgard and Merzenich, 1998, 1999) and other small mammals they may be absent or too fine to be disclosed with the available methods given that the isofrequency axis is <2 mm (Sally and Kelly, 1988; Read et al., 2001). Studies in bats (Fitzpatrick et al., 1998), birds (Cohen and Knudsen, 1999), and rats (Collía et al., 1990; Lanciego, 1994) have demonstrated intrinsic connections across isofrequency contours.

Behavioral Studies and Functional Significance As demonstrated by Kelly and colleagues (Kelly and Glazier, 1978; Kavanagh and Kelly, 1988) bilateral ablation of the auditory cortex in the rat and ferret results in very little impairment in sound localization. Using tests with two sound sources positioned symmetrically right and left of the midline, rats with complete bilateral destruction of the auditory cortex maintain their ability to localize sound accurately (Kelly, 1980; Kelly and Glazier, 1978; Kelly and Kavanagh, 1986; Jenkins and Masterton, 1982). In contrast, ablation of the auditory cortex in other species results in severe sound localization deficits (e.g., Neff, 1968; Neff et al., 1975; Heffner and Masterton, 1975; Kavanagh and Kelly, 1987). Complete destruction of the rat auditory cortex also produces no obvious abnormalities in conditioned autonomic responses or conditioned emotional responses to auditory stimuli. These deficits are pre-

sent, however, when the lesions destroy the MG bilaterally (LeDoux et al., 1984; Romanski and LeDoux, 1992). This may be due to the projections originating in the MGM and MGD that target the amygdala (LeDoux et al., 1984, 1985b, 1986; Doron and LeDoux, 1999; Linke, 1999a, 1999b; Linke et al., 1999, 2000). Thus, the behavioral studies in rat suggest that their auditory cortex is not essential for sound localization or for conditioned emotional responses to sounds. What then is functional significance of the auditory cortex in rat? Although there are no behavioral studies on temporal pattern discriminations carried out in rats, Tees (1967) and Patchett (1977) have shown that rats are capable of temporal pattern discriminations and their ability is disrupted if auditory stimulation during development is altered (for review, see Kelly, 1990. The majority of studies, however, have used pure tone stimuli, which are not typical of the spectrally and temporally complex, that have biological significance for the animal. Furthermore, the neurophysiological results of Gaese and Ostwald (1995) mentioned above suggest that the auditory cortex may have a mechanism for periodicity detection based on a temporal code that could be important for the recognition of complex acoustic signals. Other recent behavioral studies (Wan et al., 2001) using Fos imaging have shown that stimulation with novel complex sounds produces significantly greater neuronal activation in Te3 than familiar sounds. This finding suggests that Te3 has an important role in recognition memory process of familiarity discrimination. Although a detailed analysis of plasticity in the auditory system is out of the scope of this review the issue merits some discussion in the present context. Sensory receptive fields and corresponding sensorytopic maps can be modified throughout adulthood. This plasticity occurs because neuronal resources devoted to sensory processing can be reallocated to those particular features of the environment, determined by past experience, to be most behaviorally relevant. Such learning changes have been observed in the auditory system as well as in the somatosensory and visual systems (Cahill et al., 1996; Buonomano and Merzenich, 1998; Edeline, 1999; Kisley and Gerstein, 2001). Studies in rat have shown that frequency receptive fields can be modified by learning paradigms (Maldonado and Gerstein, 1996; Talwar and Gerstein, 2001) or chronic manipulation such as paring specific tonal stimuli with nucleus basalis stimulation (Kilgard and Merzenich, 1998). Adult cortical plasticity appears to be involved in the improvement of a variety of behavioral skills including compensation for damage of sensory systems, functional recovery from central nervous system damage, or maintenance of precise sensory representation (Merzenich et al., 1990).

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31. AUDITORY SYSTEM

THE DESCENDING AUDITORY PATHWAY Parallel to the ascending pathway, there is a stepwise descending projection from the auditory cortex to the organ of Corti (Fig. 34). The centrifugal pathways have been known since the end of the 19th century (Held, 1893), but interest in their study was triggered by the description of the olivocochlear bundle in 1946 by Rasmussen (1946, 1953) (Figs. 2, 34, and 35). The relays of this projection are also influenced by ascending fibers, whereby feedback loops of various sizes and complexities are established. Despite some physiological evidence of inhibitory and facilitatory actions (e.g., Watanabe, 1966; Zhang and Suga, 1997), the role of these loops in audition is still poorly understood (for review, see Harrison and Howe, 1974; Warr et al., 1986; Pickles, 1988; Huffman and Henson, 1990; Spangler and Warr, 1991; Warr, 1992) but recent studies using bats have started to disclose some of the possible functional roles. Suga and associates have shown that the excitatory and inhibitory descending projections serve to sharpen and amplify ascending inputs of the same best frequency as the site of cortical activation (e.g., Gao and Suga, 1998; Yan and Suga, 1998). A recent study in mice confirms a role in plasticity for this corticofugal pro-

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jection (Yan and Ehret, 2001). It is still not clear whether or not the descending auditory system should be considered a series of regional feedback loops or a descending chain, although there seems to be evidence now that both of these arrangements may coexist (Spangler and Warr, 1991). The first step of the chain is the auditory cortex that gives off three descending tracts (Fig. 34): the corticogeniculate, the corticocollicular, and the corticopontine projections. The corticogeniculate projection is part of the reciprocal connection between the thalamus and the cortex (Winer and Larue, 1987). The corticocollicular projection bypasses the MG and terminates in the IC (Faye-Lund, 1985; Herbert et al., 1991; Saldaña et al., 1996). In cat, the cortical belt area has been shown to supply the superior colliculus (Paula-Barbosa and Sousa-Pinto, 1973). More recently, Feliciano et al. (1995) have shown projections from the auditory cortex to subcollicular nuclei. The second step consists of fibers arising from the IC (Fig. 34). These fibers constitute a colliculoolivary and colliculocochleonuclear projection. The colliculoolivary fibers arise from the ECIC and the ventral part of the CIC (Faye-Lund, 1986; Caicedo and Herbert, 1993; Vetter et al., 1993; Saldaña, 1993). The fibers terminate on the periolivary medial olivocochlear cells (MOC)

FIGURE 35 Scheme of the LOC and MOC neurons and their projections to the IHCs and OHCs, respectively. Two types of LOC cells occur: intrinsic located inside the LSO and shell located in the margins of the LSO. Afferent fiber type I and II projections to the CNC are also represented. Note the convergence of type I onto single IHCs. (Figure kindly provided by Dr. Bruce Warr; adapted from Warr, 1992.)

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which supply the OHCs. Thus, these may constitute a three-neuron pathway from the auditory cortex to the sensory cells. The lateral olivocochlear cells (LOC) which supply the ICHs may not be directly influenced from higher auditory centers, although Felciano et al. (1995) have shown a direct corticoolivary projection that may indeed have some influence upon LOC neurons (v.i.). These sensory cells, therefore, may be controlled by a shorter neuronal loop involving only the CNC and the SOC. The third step in the chain is the olivocochlear system (Figs. 34 and 35) that constitutes the efferent innervation of the cochlea which consists of the MOC and the LOC already mentioned. These three main parts are treated subsequently, separately.

The Corticofugal Pathways The rat auditory cortex may modulate the processing of ascending auditory information via descending projections to the MG (Winer and Larue, 1987; Roger and Arnault, 1989; Arnault and Roger, 1990; Shi and Cassell, 1997; Barlett et al., 2000), the RTN (Rouiller and Welker, 1991; Barlett et al., 2000), the IC (Faye-Lund, 1995; Coleman and Clerici, 1987; Games and Winer, 1988; Roger and Arnault, 1989; Herbert et al., 1991; Vaudano et al., 1990; Saldaña et al., 1996; Druga et al., 1997), the nucleus sagulum, the paraleminical regions, and the SOC (Feliciano et al., 1995), as well as the cochlear nuclear complex (Felciano et al., 1995; Weedman et al., 1996a, 1996b) as seen in Fig. 34. Winer and Larue (1987) have demonstrated that the MGV receives the heaviest cortical input, while the MGM receives the least and the MGD receives an intermediate amount. They also showed that despite a largescale topographical overlap in the spatial territories of thalamocortical cells and corticothalamic axonal terminals, there are nonoverlapping zones. There seems to be general agreement that Te1 projects to the MGV and MGD, Te2 to the MGD and sparsely to the MGM, and Te3 to the MGD, the MGV, and sparsely to the MGM (Winer and Larue, 1987; Roger and Arnault, 1989; Arnault and Roger, 1990; Rouiller and Welker, 1991; Shi and Cassell, 1997; Barlett et al., 2000). Barlett et al. (2000) have also found that most terminal boutons arising from the auditory cortex are small (0.57 μm2) and end in small caliber dendrites in the MGV and MGD (Fig. 30). Most probably these small boutons arise from the pyramidal cells of layer VI (Romanski and LeDoux, 1993a, 1993b; Barlett et al., 2000). These small terminals have round vesicles and thick postsynaptic densities (Barlett et al., 2000). The major corticofugal projections appear to be glutamatergic (Potashner et al., 1988), suggesting an excitatory function. In addition, a few very large boutons (>2 μm2) have been described in

the MGD and marginal zone of the MG (Fig. 30) (Rouiller and Welker, 1991; Barlett et al., 2000). These have also been described in cat (Ojima, 1994; Bajo et al., 1995; Winer et al., 1999a; 2001). They terminate on large dendrites and they make multiple contacts on the postsynaptic dendrites. Usually, these large corticothalamic terminals form complexes with the dendrites partially surrounded by astrocytic processes (Barlett et al., 2000). These endings are thought to originate from neurons in layer V (Rouiller and Welker, 1991; Barlett et al., 2000). Electrical stimulation of the auditory cortex results in both excitatory and inhibitory effects on MG neurons (Watanabe et al., 1966; mouse: Ryugo and Weinberger, 1976; bat: Zhang and Suga, 1997). The auditory cortex also projects to the auditory sector of the reticular thalamic nucleus which in turns projects to the MG (Fig. 34) (Rouiller and Welker, 1991; Barlett et al., 2000), thus providing the MG with an inhibitory influence (Montero, 1983; Barlett et al., 2000). The corticofugal projection, therefore, can modulate the MG responses to sound through a direct excitatory pathway and/or an indirect inhibitory pathway. In 1900, Thompson reported a projection from the temporal cortex to the corpora quadrigemina in the primate. Since then, a number of studies have confirmed and extended the knowledge of this pathway in several species (e.g., Massopust and Ordy, 1962; van Noort, 1969; Casseday et al., 1979; Andersen et al., 1980b, Morest and Oliver, 1984; Feliciano and Potashner, 1995; Winer et al., 1998) including rat (Krieg, 1947; Beyerl, 1978; FayeLund, 1985; Coleman and Clerici, 1987; Games and Winer, 1988; Roger and Arnault, 1989; Herbert et al., 1991; Saldaña et al., 1996; Druga et al., 1997) (Figs. 21B and 34). Most of these studies have emphasized the traditional view in which the auditory cortex innervates the collicular cortices only. Furthermore, they have shown a topographic organization of these projections from the primary auditory cortex, such that the caudal low frequency regions of Te1 project to the dorsolateral region of the IC and the rostral high frequency region of Te1 projects to the ventromedial region of the IC. However, Saldaña et al. (1996), based on a study using PHA-L as an anterograde tracer, have demonstrated that Te1 indeed projects into the CIC as already reported by Beyerl (1978) and the topography of corticocollicular projections mimics that of the intrinsic connections in the ipsilateral IC (Figs. 21B and 34) (Saldaña and Merchán 1992). Herbert et al. (1991), in addition, has shown the projection to the IC originating from the secondary auditory cortices, i.e., Te2 and Te3. They have shown that Te2 projects primarily to the superficial layers of the DCIC and ECIC, while Te3 innervates primarily the rostral part of the ECIC (Fig. 34). Winer and Games (1988) demonstrated that the cellular origin

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31. AUDITORY SYSTEM

of these projections is the pyramidal cells in layer V. Feliciano and Potashner (1995) have demonstrated the glutamatergic nature of this projection in guinea pig. This is in agreement with the study of Saldaña et al. (1996), who showed at the electron microscopic level that in all subdivisions of the rat IC these corticocollicular fibers terminate in small boutons with round synaptic vesicles and form asymmetric synapses with thin dendritic shafts and spines. Although these studies suggest a purely excitatory corticollicular projection, electrical stimulation of the auditory cortex in cat elicits not only excitatory but also inhibitory and complex interactions in the IC neurons (cat, Mitani et al., 1983). In a recent study on the functional role of this corticofugal projection in rat using tetrodotoxin to block the effect of the auditory cortex upon the IC, Nwabueze-Ogbo et al. (2002) have shown an enhancement or suppression of the neuronal activity in the IC and prolongation of the first spike latency. Thus, the auditory cortex may modulate the processing of sounds in the IC through the activation of local inhibitory connections within the IC. Electrical stimulation of the auditory cortex in bats causes a downward shift in the preferred frequencies of collicular neurons toward that of the stimulated cortical neurons. This results in a change in the frequency map within the colliculus. Similar changes can be induced by repeated bursts of sound at moderate intensities. Thus, one role of the mammalian corticofugal system may be to modify subcortical sensory maps in response to sensory experience (Yan and Suga, 1998). Several studies have suggested a direct neocortical projection to auditory brain stem nuclei since 1935 (e.g., Mettler, 1935; Kuypers and Lawrence, 1967), nevertheless it has been a general dogma that projections from the auditory cortex do not extend beyond the IC. In 1995, using PHA-L, Feliciano et al. reexamined this issue and demonstrated that indeed Te1 sends direct projections to regions surrounding the lateral lemniscus, including the nucleus sagulum ipsilaterally and bilaterally to SOC and CNC, namely, the VTz, to LSO, a narrow region that overlies the dorsal aspect of the SOC; to the DC; and to the granule cells domain in the VC (Fig. 34). Since this descending input terminates on the LSO, perhaps the LOC neurons (at least the shell neurons) are probably involved in local short distance feedback loops. These latter projections have also been confirmed by Weedman et al. (1996a, 1996b). Felciano et al. (1995) have speculated that the functional role of these corticosubcollicular projections is related to the control of acousticomotor reflexes or perception of sound, but future studies are needed to confirm their hypothesis. Before we turn our attention on the descending projections arising from the IC, we should mention a few recent reports on a new thalamotectal loop (Fig. 34).

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The descending projections from the MG to the IC have been viewed with some scepticism, but there is some evidence in cat (Paydar, 1993), gerbil (Kuwabara and Zook, 2000), and rat (Senatorov and Hu, 2002; Winer et al., 2002) for this projection. The recent study of Kuwabara and Zook (2000) shows that the MGM and MGD project to the ECIC (Fig. 34). These authors speculate that this thalamotectal feedback circuit may be involved in some multisensory task.

The Colliculofugal Pathways Here we focus on the descending projections from the IC to the brain stem nuclei (Fig. 34). The IC has been shown to originate colliculolemniscal, colliculoolivary, and colliculocochlear projections (Faye-Lund, 1986; Caicedo and Herbert, 1993; Saldaña, 1993; Vetter et al., 1993). Similar projections have been described in guinea pig (Malmierca et al., 1996). Caicedo and Herbert (1993) have reported that colliculolemniscal projections are largerly confined to the dorsal nucleus, the sagulum, the horizontal cell group, and the perilemniscal zone. The projections to the DLL arise from the CIC, are topographic, and follows the pattern described by Merchán et al. (1994). The sagulum and perilemnical zones receive the collicular projections from the ECIC and DCIC. The study of Caicedo and Herbert (1993) is based on injections using the tracer PHA-L, however, they showed some retrogradely labeled cells in the DLL and DC; so, owing to the bidirectional nature of the transport, it remains open to question whether the descending projection to the DLL is genunine (Merchán et al., 1994). The colliculoolivary projections originate in the CIC and ECIC and terminate in a band of terminals that extends from the RPO to the VTz (Fig. 34) (FayeLund, 1986; Caicedo and Herbert, 1993; Saldaña, 1993; Vetter et al., 1993). This projection is also topographic, such that the dorsolateral low frequency region in the IC projects to the lateral part of the VTz and the ventromedial high frequency region of the IC projects to the medial portion of the VTz. The terminal fibers in this area appear to overlap the site of origin of the MOC (White and Warr, 1983). Vetter et al. (1993) have provided evidence of termination of IC fibers on these MOC neurons using a double-labeling technique in which PHA-L-labeled fibers from the IC are found in close apposition to retrogradely labeled neurons of the MOC. Although it remains to be demonstrated whether these fibers make synaptic contact on the MOC neurons by electron microscopy, the study of Vetter et al. (1993) strongly suggests that the IC may modulate cochlear responses. This idea is supported by the electrophysiological studies of Dolan and Nuttall (1988) and Rajan (1990) in cat. Electrical stimulation of the IC

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produces an increase in the latency and a reduction in the amplitude of the auditory whole-nerve response. These effects are similar to those elicited by electrical stimulation of the MOC (Rajan, 1990). Electrical stimulation of the IC also reduces the temporal threshold shift that appears after the exposure to a loud noise (Rajan, 1990). The colliculocochlear projection originates in the CIC and ECIC and targets the DC and granule cell domain of the VC (Faye-Lund, 1986; Caicedo and Herbert, 1993; Saldaña, 1993). Currently, the functional role of this projection it unknown. Finally, it should be mentioned that the rat IC also projects to the pontine and mesencephalic reticular nuclei (Caicedo and Herbert, 1993). Projections from the IC to these nonauditory nuclei include the pontine nuclei, the lateral paragigantocellular nucleus, gigantocellular reticular nucleus, the ventrolateral tegmental nucleus, and the caudal pontine reticular nucleus. Such projections have been previously reported in other species (reviewed in Caicedo and Herbert, 1993). Interestingly, the same nuclei receive prominent projections from the cochlear nucleus (Kandler and Herbert, 1991; López et al., 1999), which led researchers to suggest that these nonauditory areas may play a role long- and short-latency auditory-motor behaviors or in acoustically elicited autonomic responses (Kandler and Herbert, 1991; Caicedo and Herbert, 1993).

The Olivocochlear System It has been known for more than 50 years that the olivocochlear bundle (Figs. 2, 34, and 35) provides the organ of Corti with the efferent innervation (Rasmussen, 1946, 1953). There is now substantial evidence that justifies the recognition of two systems of olivocochlear neurons, medial (MOC) and lateral (LOC) (Figs. 34 and 35) (White and Warr, 1983; Warr et al., 1986; Spangler and Warr, 1991; Warr, 1992; Vetter et al., 1991; Vetter and Mugnaini, 1992; Warr et al., 1997; Cantos et al., 2000). As their names imply, the location of their respective cell bodies is in the medial and lateral regions of the SOC. The MOC neurons project mainly to the contralateral cochlea, whereas the LOC neurons project to the ipsilateral cochlea (Warr, 1992). A few MOC neurons (<5%) in rat and other species have been found to project bilaterally (Aschoff and Ostwald, 1988). This is in agreement with physiological studies in which the majority of units respond to ipsilateral tone stimulation and a few MOC neurons are driven binaurally, at least in cat and guinea pig (Liberman and Brown, 1986; Brown, 1989; reviewed in Guinan, 1996). The MOC neurons innervate the OHCs (Figs. 2, 34, and 35), about 55% (rat, Robertson et al., 1989; Horváth et al., 2000) of them originate in the contralateral side

and the reminder in the ipsilateral side, and are made of thick myelinated axons that terminate directly on the OHCs (White and Warr, 1983; Warr et al., 1997; Horváth et al., 2000). MOC neurons (38) have generally large cell bodies, stellate- or triangular-shaped, and possess dendritic arbors that radiate and branch profusely and taper (Osen et al., 1984; Vetter and Mugnaini, 1992). They are located underneath the MSO, lateral to the MTz, in the VTz (Fig. 34) (Vetter and Mugnaini, 1992), and constitute a neurochemically homogeneous population of cholinergic cells (Osen et al., 1984; Vetter et al., 1991). They may receive descending input from the ipsilateral IC (see above, Vetter et al., 1993) and ascending input bilaterally from the VCP (Thompson and Thompson, 1991), and possibly from the globular bushy cells in the caudal VC as White (1986) has found terminal boutons similar to the calyces of Held terminating upon MOC cells. MOC neurons innervate VC neurons (Fig. 34) (Hováth et al., 2000) and clusters of OHCs spanning as much as an octave of cochlear length, but the terminal axonal arbor is often centered basal to the corresponding location of radial afferent fibers of similar best frequency. MOC neurons are sharply tuned and the MOC efferent innervation is tonotopically organized (reviewed in Warr, 1992; Guinan, 1996; Helfert and Aschoff, 1997). The LOC neurons innervate the regions beneath the IHCs and originate predominantly in the ipsilateral side (Fig. 34 and 35) (rat, White and Warr, 1983; Warr et al., 1997; Horváth et al., 2000; cat: Arnesen and Osen, 1984; Liberman and Brown, 1986; mouse, Brown, 1987; Wilson et al., 1991; guinea pig, Brown, 1993). In rat, two distinct types of neurons form the LOC, intrinsic and shell neurons (Fig. 34) (Vetter and Mugnaini, 1992; Warr et al., 1997). All intrinsic neurons (Figs. 34 and 35) are small cells confined within the limits of the ipsilateral LSO. They possess a disk-shaped dendritic arbor as the principal LSO neurons with thin untapered dendrites and constitute about 85´ of all LOC neurons (Vetter and Mugnaini, 1992; Warr et al., 1997; Horváth et al., 2000). The intrinsic neurons probably they receive the same input as the principal cells in the LSO. About 50% of them are GABAergic and the remaining 50% that are cholinergic colocalize calcitonin gene-related peptide (Vetter et al., 1991). Their axons are thin unmyelinated (being 0.77 μm thick in the organ of Corti), travel relatively short distances in the inner spiral bundle, and terminate forming discrete (less than 0.2 octave in extent) terminal arborization with a focal, tonotopic organization (Warr et al., 1997). They do not innervate the VC (Horváth et al., 2000). Most shell neurons (Figs. 34 and 35; about 95%) are located at the ipsilateral areas surrounding the LSO. They are large multipolar cells

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with thick and tapered dendrites and constitute about 15% (Vetter and Mugnaini, 1992; Warr et al., 1997). Little is known about the source of the inputs they receive. Only one third of these shell neurons is GABAergic while the other two-thirds are cholinergic but do not colocalize calcitonin gene-related peptide (Vetter et al., 1991). Their axons are thin, unmyelinated (being 0.37 μm thick in the organ of Corti), travel long distances (greater than 1mm) in the inner spiral bundle, and terminate forming diffuse (greater than 1 octave in extent) terminal arborizations, being probably not tonotopic (Warr et al., 1997). In contrast to the intrinsic neurons, but similar to the MOC, they innervate the VC (Fig. 34) (Horváth et al., 2000). Physiological studies in several species have shown that MOC neurons are sharply tuned with a wide dynamic range. Most of them respond to binaural stimuli, two-thirds to contralateral stimuli and one-third to ipsilateral stimuli (Roberston and Gummer, 1985; Liberman, 1988; reviewed in Guinan, 1996). Stimulation of the MOC neurons is thought to raise the threshold of the primary afferents through the modulation of the biomechanical properties of the basilar and tectorial membranes (Dallos, 1985a, 1985b; Flock, 1988). Thus, MOC neurons may enhance transduction or signal detection through an unmasking effect, thus regulating the slow motility of the OHCs and thereby the stiffness of the basilar membrane (reviewed in Helfert and Aschoff, 1997; Eggermont, 2001). Previous studies have also demonstrated that the MOC can somewhat reduce the temporary threshold shift that occurs as an early manifestation of the damaging effects of loud sounds (guinea pig, Roberston and Johnstone, 1980; Cody, 1992; reviewed in Guinan, 1996), protecting the cochlea. The functional role of the LOC is still unclear. However several immunocytochemical studies have shown that these neurons contain neuroactive substances such as dopamine, serotonin, and opioid peptides (e.g., Safieddine and Eybalin, 1992; Eybalin, 1993; Safieddine et al., 1997), which speaks in favor of a modulatory effect on the IHCs afferents. More recently, Rajan (2000) has shown in cat that even moderate noise backgrounds can significantly exacerbate the cochlear temporary threshold shift induced by loud tones, but this is prevented because under such conditions there is additional activation of the LOC, enhancing protection of cochlear hearing sensitivity. Thus, in background noise there is a conjoint activation of the MOC and LOC to powerfully protect (by almost 30 dB) from loud sound that otherwise would be exacerbated by the noise (Rajan, 2000). Clearly, the olivocochlear system plays a critical role in maintaining the normal operation of the cochlea and the presence of efferent connections with feedback

loops may introduce nonlinear dynamics (Eggermont, 2001) into the auditory system.

Abbreviations Aq AC AM AMPA BDA bic BM C Cb cc CIC cic cll CNC das D DC DD DCIC DLL DO DPO DS EPSP ECIC EE EI Ep F FM FS GABA GAD GrC h I8 IB IC icp IHC IPSP l L LF ll LPO LVPO LOC LR4V LSO LTz M MG MGD MGM MGV MOC MSO MTz

VI. SYSTEMS

Aqueduct (Sylvius) Auditory cortex Amplitude modulation α-Amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid Biotinylated dextran amine Brachium of the inferior colliculus Basilar membrane Caudal Cerebellum Corpus callosum Central nucleus of the inferior colliculus Commissure of the inferior colliculus Commissure of the lateral lemniscus (Prosbt) Cochlear nuclear complex Dorsal acoustic stria Dorsal Dorsal cochlear complex Deep dorsal nucleus of the MGD Dorsal cortex of the inferior colliculus Dorsal nucleus of the lateral lemniscus Dorsal nucleus of the MGD Dorsal periolivary region Dorsal superficial nucleus of the MGD Excitatory postsynaptic potential External cortex of the inferior colliculus Excitatory–excitatory Excitatory–inhibitory Ependimum Flat neurons Frequency modulation Fast spiking γ-Aminobutyric acid Glutamic acid decarboxylase Granule cell layer high frequency region Intersticial nucleus of the vestibulocochlear nerve Intrinsically bursing Inferior colliculus inferior cerebellar peduncle Inner hair cell Inhibitory postsynaptic potential Low-frequency region Lateral Less flat neurons Lateral lemniscus Lateral periolivary zone Lateroventral periolivary zone Lateral olivocochlear system Lateral recess of the fouth ventricle Lateral superior olive Lateral nucleus of the trapezoid body Medial Medial geniculate body Dorsal division of the medial geniculate body Medial division of the medial geniculate body Ventral division of the medial geniculate body Medial olivocochlear system Medial superior olive Medial nucleus of the trapezoid body

1064 MVPO MZ NLL NMDA OC ocb OHC OV PAG PC PHA-L PIL PnC PO PP PSL PSTH R ReIC RM RS Rt SC SG SL SOC SpG sp5 SPO SSL SV T1-T5 Te1 Te2 Te3 TM Tz tz V1 V VC VCA VCP VLL VTT VTz 5 7 8cn 8vn

MANUEL S. MALMIERCA and MIGUEL A. MERCHÁN

Medioventral periolivary zone Marginal zone of the MG Nuclei of the lateral lemniscus N-Methyl-D-aspartate Organ of Corti Olivocochlear bundle Outer hair cells Ovoid nucleus of the MGV Periaqueductal gray matter Pilar cells Phaseolus vulgaris-leucoagglutinin Posterior intralaminar nucleus Pontine reticular nucleus, caudalis Periolivary regions Peripeduncular nucleus Primary spiral lamina Peristimulus histogram Rostral Recess of the inferior colliculus Reinser’s membrane Regular spiking Auditory sector of the reticular thalamic nucleus Superior colliculus Suprageniculate nucleus Spiral ligament Superior olivary complex Spiral ganglion Spinal trigeminal tract Superior paraolivary nucleus Secondary spiral lamina Stria vascularis Scalae tympani (first to fifth turns) Temporal area 1 Temporal area 2 Temporal area 3 Tectorial membrane Nucleus of the trapezoid body Trapezoid body (or ventral acoustic stria) Scalae vestibuli Ventral Ventral cochlear complex Anteroventral cochlear complex Posteroventral cochlear complex Ventral complex of the lateral lemniscus Ventrotubercular tract Ventral nucleus of the trapezoid body 4V 4th ventricle Trigeminal ganglion Facial nucleus Cochlear root of the vestibulocochlear nerve Vestibular root of the vestibulocochlear nerve

Financial support was provided by Spanish Junta de Castilla y León, Fondo Social Europeo’ (Grant SA084/01) and Dirección General de Enseñanza Superior (Grant FI-2000-1396). M.S.M. dedicates this chapter to his parents, Martha, Marco, and Pablo for their continuous and invisible support.

References

Acknowledgments Many people have contributed generously to the ideas and concepts that we present in this account. It is our pleasure to acknowledge their encouragement and constructive criticism at different stages as well as their intense and extensive comments, corrections, and suggestions. We are most grateful to Nell Cant, David Furness, Craig Henkel, Jack Kelly, Douglas Oliver, Kirsten Osen, Adrian Rees, Philip Smith, Bruce Warr, and Jeffery Winer. We would also like to thank Albert Berrebi, David Furness, Randy Kulesza, Dolores López, Douglas Oliver, Philip Smith, Bruce Warr for providing us with beautiful and original pictures.

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C H A P T E R

32 Visual System ANN JERVIE SEFTON1 and BOGDAN DREHER2 1

Departments of Physiology and 2Anatomy and Histology University of SydneyNew South Wales, Australia

ALAN HARVEY Department of Anatomy and Human Biology University of Western Australia, Australia

We dedicate this chapter to the memory of Istvan Törk (1939–1992), one of the pioneers of research into monoaminergic systems of the brain and a valued friend.

The rat, a nocturnal murid rodent, has sophisticated and effectively functioning visual system. Its laterally placed eyes provide it with a panoramic view but there is a binocular overlap, estimated to be 40–60º in front of the animal, above its snout, so that the image of the binocular field falls on the lower temporal retinae (see Fig. 2). Even though the rat’s eye may not accommodate, with its pupil constricted it has a considerable depth of focus (Hughes, 1977b). Although rods are the predominant photoreceptors, cones, although relatively rare (about 0.85% of photoreceptors), are also present and the retina is therefore capable of functioning in both scotopic and photopic conditions (Cicerone, 1976; Szél and Röhlich, 1992). Of the two types of cones identified, the majority (93%) contain a photopigment with a peak sensitivity at about 500–520 nm (Deegan and Jacobs, 1993). Unlike other dichromatic animals, but like some other nocturnal rodents (Jacobs, 1992), the rat appears to have photoreceptors sensitive to ultraviolet light. Indeed, the second, albeit rare, cone type (7% of cones, i.e., about 0.05% of all photoreceptors; Szél and Röhlich, 1992) contains a photopigment with a peak sensitivity at about 370 nm, in the ultraviolet range (Deegan and Jacobs, 1993; Szél and Röhlich, 1992). As in the mouse and gerbil, the rat has only one type of horizontal cell—the axon-bearing B type; the axon-less A type present in

The Rat Nervous System, Third Edition

virtually all other mammals, including cavid rodents, is absent (Peichl and González-Soriano, 1994). There has been controversy concerning reported differences in the sensitivity to light of albino and pigmented strains (cf. Balkema and Dräger, 1991; Creel et al., 1970; Dodt and Echte, 1961; Dyer and Schwartzwelder, 1978; Green and Powers, 1982; Green et al., 1991; Ralto et al., 1991). Herreros de Tejada and colleagues (1992), however, suggested that the dark-adapted thresholds for visually evoked cortical and collicular potentials— at about −5.2 to −5.8 log cd/m2—are virtually identical in both strains. Furthermore, the thresholds are not significantly different from the average absolute threshold for dark-adapted humans. The rat’s eye is capable of optically resolving 12 min of visual angle (Hughes, 1979b; Hughes and Wässle, 1979). The maximal spatial resolution of dark- and lightadapted retinal ganglion cells recorded from optic tract of the hooded rat is, however, only about 1.2 cycles/ degree or about 25 min of visual angle (Friedman and Green, 1982). Similarly, visual acuity estimated on the basis of the sampling theorem from the peak density of ganglion cells is about 1.3 cycles/degree (Pettigrew et al., 1988). Consistent with this, the upper limit of spatial resolution of single neurons recorded from the primary visual cortices of Lister hooded rats is 1.2 cycles/degree

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(Girman et al., 1999). Finally, behaviorally measured visual acuity or acuity extrapolated on the basis of evoked potentials in the visual cortex of awake rats is also about 1.2 cycles per degree (Birch and Jacobs, 1979; Boyes and Dyer, 1983; Dean, 1978, 1981a, 1981b; Fagiolini et al., 1994; Harnois et al., 1984; Lashley, 1938; Seymoure and Juraska, 1997; Silviera et al., 1987; Wiesenfeld and Branchek, 1976). Consistent with these data, the upper limit of spatial resolution of single neurons recorded from the primary visual cortices of Lister hooded rats is 1.2 cycles/degree (Girman et al., 1999). In hooded rats in which one eye has been enucleated at birth, however, behaviorally measured visuospatial resolution reaches 1.6 cycles/degee (Yagi et al., 1995). The better spatial resolution in those animals is likely to be related to the fact that the ipsilateral retinofugal pathway is larger than normal and might correlate with larger than normal numbers of retinal ganglion cells in the area centralis (Chan et al., 1989; Chan and Jen, 1988; Jen et al., 1990; Lund et al., 1980; Lund and Lund, 1971a, 1971b). Visual acuity develops gradually in the first month after natural eye opening for the first time [about 14 days after birth (Dreher and Robinson, 1988)]. Thus, on postnatal day 19, when the optical media became clear, the visual acuity extrapolated on the basis of evoked potentials in the visual cortex of awake hooded rat pups is only 0.51 cycles per degree and the adult values are reached only by postnatal days 40–45 (Fagiolini et al., 1994). The retina is developmentally part of the central nervous system (diencephalon). Processing occurs in layers of the retina antecedent to the ganglion cell layer, but in this chapter, intraretinal processing is not included (see, for a review, Rodieck, 1998). The visual system has been investigated extensively in the adult and developing rat using anatomical, biochemical, molecular, physiological, and behavioral techniques, but space precludes detailed considerations of development in this chapter. Ganglion cells are relatively evenly distributed across the rat retina, with the variation from the highest to the lowest density being 5:1—from only 3000 to 600 cells per mm.2 Dendritic trees and therefore the sizes of the receptive field centers of ganglion cells located in the area of highest density (area centralis) are not significantly different from their counterparts located peripherally, in the areas of lowest density (Dreher et al., 1984; Huxlin and Goodchild, 1997; Perry, 1979). Therefore, exploratory fixation—eye movement bringing the image of an object of interest to a region of high resolution—is not likely to occur. In the rat, however, as in mammals with better developed exploratory eye movements, such as cats or virtually all primates, stimulation of the superior colliculus (SC) elicits conjugate saccadic eye movements, implying that there is a clear-cut relationship between

the retinotopic map and the topographic representation of eye movement fields in the SC (McHaffie and Stein, 1982).

VISUAL PATHWAYS The concept of the mammalian visual system operating as a set of pathways working largely in parallel has proved particularly fruitful (see, for reviews, Stone, 1983; Stone et al., 1979). To a large extent, especially within the retinogeniculocortical pathway, separate classes of retinal ganglion cells send specific information through separate classes of relay cells within one particular visual nucleus. However, we sound a note of caution here: results and interpretations of visual studies in the widely used cat and primate should not be uncritically extrapolated to the rat. For example, there is a prominent class of X or X-like cells in the visual systems of cats and primates, but in the rat the X-like system is poorly developed. However, observations made in the rat, as a representative of the large and successful order of Rodentia, test the general validity of hypotheses generated on the basis of studies of other species and may offer insights into the way the organization of the visual system has developed under the influence of particular environmental factors. The visual pathways of the rat are illustrated schematically in Fig. 1. Retino-recipient nuclei within the primary and accessory visual pathways are considered in the section on “Major Retino-recipient Nuclei.” The suprachiasmatic nucleus is discussed in Chapter 15. Retinal projections of dystrophic rats are similar to normals, albeit somewhat reduced (Decker et al., 1995). It should be noted that in addition to the nuclei illustrated in Fig. 1, there are small, “nonimage-forming” (Pickard, 1982) retinal projections, which are not considered further in this chapter. Retinal ganglion cells lying in the upper temporal quadrant (Leak and Moore, 1997) project to the lateral hypothalamus (Kita and Oomura, 1982; Leak and Moore, 1997; Riley et al., 1981; Sousa-Pinto and Castro-Correia, 1970). They also innervate hypothalamic paraventricular (Johnson et al., 1988 ) and supraoptic (Levine et al., 1991) nuclei; inferior colliculus (Itaya and van Hoesen, 1982a); subthalamus (Yamadori and Yamauchi, 1983); basal telencephalon (Cooper et al., 1989); lateral habenular (Qu et al., 1996) and dorsal raphe (Fite et al., 1999) nuclei; and anterodorsal and anteroventral thalamic nuclei (Ahmed et al., 1996b; Itaya et al., 1981; Repérent et al., 1987). These latter do not represent branches of fibers projecting to visual centers (Ahmed et al., 1995). There is extensive literature on the small ipsilateral pathway in the rat that contributes to the binocular

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SuG OP InG

SC

PPT OPT OT PBG

APT

LP

Rt

IGL VLGMC

sao

DT

MT

DLG LT

iao

SCh VLGPC

eye FIGURE 1 Semidiagrammatic representation of visual pathways originating in the retina of the right eye as if viewed from an elevation of about 45º facing the front of the rat. Each nucleus is represented by a typical coronal section through it. The rostrocaudal sequence is chosen but the dorsoventral locations are only approximate. On the left-hand side of the diagram, visual nuclei are labeled. Note that the contralateral pathway, depicted on the right-hand side by the thick unbroken line, is more extensive than the ipsilateral, shown on the left-hand side(fine unbroken line). Shaded nuclei (on the right) are those reported to receive an input from visual cortical areas. For simplicity, each projection is shown as a branch of a single fiber, but this is not meant to imply that all nuclei are necessarily innervated by branches of retinocollicular axons. Included are two visual nuclei, Rt and PBG, which do not receive a direct retinal input, but which are connected reciprocally to DLG and SC, respectively. The pathways interconnecting PBG and SC ipsilaterally and contralaterally are illustrated with a dashed line; PBG also projects to DLG (not shown). LP receives only a small contralateral retinal input but a substantial input from SC (not shown). The extensive interconnections between major retino-recipient nuclei are not indicated. Accessory optic nuclei (DT, LT, MT) are innervated by the accessory optic tracts contralaterally (sao and iao, shaded lines); the trajectory of the pathways is schematic. Note that although accessory nuclei receive an ipsilateral input (illustrated), the precise pathway has not been determined.

representation of the visual field in target nuclei. Although Polyak’s (1957) estimate that 5–10% of axons in the optic nerve project ipsilaterally is widely quoted, Jeffrey (1984) demonstrated that only about 3% of all ganglion cells project ipsilaterally. In albinos, the ipsilateral projection is even smaller (about 1.5%)1 (Ahmed et al., 1996b; Dreher et al., 1985a; Lund, 1965). Individual ganglion cells may project to more than one nucleus 1

The difference between albinos and other strains is correlated with a difference in the amount of melanin in the retina and retinal derivatives of the optic cup rather than with the amount of melanin in the choroid and iris (Creel and Giolli, 1976; Wise and Lund, 1976).

ipsilaterally (Ahmed et al., 1995, 1996a, 1998). Despite the small size of the ipsilateral pathway, activation of one eye in hooded rats induces a considerable response in ipsilateral retino-recipient regions—the dorsal lateral geniculate nucleus (DLG) and superior colliculus (SC) (Isseroff and Madar, 1983)—as well as in the primary and “higher” visual cortices (Silviera et al., 1989; Thurlow and Cooper, 1988), although responses to patterned visual stimuli are exclusively contralateral (Montero and Jian, 1995). In pigmented rats, the bilateral response is probably mainly due to substantial inputs to both DLG and SC from the contralateral parabigeminal nucleus (Harting et al., 1991b; Sefton and Martin, 1984; Turlesjski

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et al., 1993). In albinos, at the cortical level, information from the ipsilateral eye is, to a large extent, relayed via commissural (callosal) projections from the contralateral visual cortex (Diao et al., 1983a; Silviera et al., 1989). In turn, feedback projections from layers 5 and 6 of visual cortical areas are probably responsible for the bulk of the input, albeit indirectly, from the ipsilateral eye to SC and DLG (see page 1120). Visual centers in the brain receive inputs from a range of ipsi- and contralateral brain stem nuclei (for specifics, see for each nucleus). The location of the cell bodies in locus coeruleus and dorsal raphe projecting to the different functional regions is, to some extent, topographically ordered (Waterhouse et al., 1993). Many studies show that in rats the ipsilateral visual pathway is larger than normal following unilateral neonatal optic tract lesions (Jen and Lund, 1981), tectal lesions (Perry and Cowey, 1979), thalamic lesions (Chan et al., 1989; Chan and Jen, 1988; Jen et al., or 1990; Jen and Lund, 1981), or removal of one eye during fetal (Lund and Miller, 1975) or early postnatal life (Chan et al., 1989; Chan and Jen, 1988; Jen et al., 1990; Lund et al., 1980; Lund and Lund, 1971a, 1971b). The expanded pathway in DLG is functional, but the properties of the cells are not entirely normal (Fukuda et al., 1983). After prenatal or early postnatal enucleation, the expansion usually takes the form of an increase in the area within retino-recipient nuclei occupied by the ipsilateral projection. Neonatal enucleation appears to stabilize the exuberant ipsilateral projection present prenatally and at birth in SC (Laemle and Labriola, 1982; Land and Lund, 1979; Martin et al., 1983). Indeed, unilateral thalamic lesions combined with ipsilateral monocular eye removal in neonatal animals result in the preservation of the entire population of normally transient ipsilaterally projecting retinal ganglion cells (Jen et al., 1989). In adult rats, following cortical ablation and eye removal, there is a substantial sprouting of retinal axons from the remaining eye into the ventral VLG and pretectum (Goodman and Horel, 1966). Because the neural retina is derived embryologically from the central nervous system, the retina and optic nerve of the rat have provided widely used models for studies of central neural damage and regeneration. Space precludes any detailed consideration here; reviews include Dezawa and Adachi-Usami (2000), Harvey et al. (1995), Heiduschka and Thanos (2000), So and Yip (1998), and von Bartheld (1998).

RETINAL OUTPUT Retinal Ganglion Cells Because the axons of ganglion cells are the only retinal output, they determine the nature of the information supplied to each of the retino-recipient nuclei. Repre-

sentation of the visual field on the rat’s retina in relation to various landmarks is illustrated in Fig. 2B. Danias et al. (2002) counted 100,000 ganglion cells in the albino retina; earlier authors reported 110,000 in different strains (Perry et al., 1983; Potts et al., 1982; Schober and Gruschka, 1977). In early postnatal life the number of cells and optic axons is much greater (Perry et al., 1983; Potts et al., 1982; Sefton and Lam, 1984), and in a period of naturally occurring cell death during the first 5–10 postnatal days it is reduced to the stable adult number (Dreher et al., 1983). The majority of ganglion cells have their somata in the ganglion cell layer, but, as in other species, a small proportion—about 2.1% of ipsilaterally projecting and about 0.9% of contralaterally projecting cells (Liu and Jen, 1986)—have somata displaced into the inner plexiform layer (Buhl and Dann, 1988; Dreher et al., 1985a; Linden, 1987). Peichl (1989, see later) identified some as “outer” α cells, but others remain unclassified (Buhl and Dann, 1988). Amacrine cells are not restricted to the amacrine layer of the retina, but are also found in the ganglion layer as well as in the inner nuclear layer (Bunt et al., 1974; Perry, 1981); as in other species, they exhibit a range of morphologies and transmitters (see Vaney, 1990; Rodieck, 1998). It is apparent that there is a morphological diversity of retinal ganglion cells in the rat, but because of technical difficulties of recording from the small eye, there are few correlations among morphology of the retinal ganglion cells, their targets, and functions. Three morphological groups have been distinguished (Dreher et al., 1985; Fukuda, 1977; Huxlin and Goodchild, 1997; Ni and Dreher, 1981; Perry, 1979), perhaps reflecting the three conduction velocity peaks recorded in optic nerve (Sefton and Swinburne,1964). Ni and Dreher (1981) and Dreher et al. (1985) added a subclass to Perry’s (1979) types II and III. Later, Huxlin and Goodchild (1997) added an additional subclass to each of types I and II and suggested a new nomenclature (RGA,B,C); many of their conclusions were verified by Sun et al. (2002), who developed a four group classification. Type I, RGA, or RGA1 cells, also identified using neurofibrillar stains, have large somata (16–32 μm), thick axons (mean diameter 0.9 μm), three to seven large gauge primary dendrites, and large dendritic trees (220 to 790 μm). They are distributed across the retina with a peak density in the temporal region, where receptive fields are smallest (Dreher et al., 1985; Reese and Cowey, 1986; Schall et al., 1987). A weak orientation bias is apparent in the dendritic field (Dreher et al., 1985a; Schall et al., 1987). Sun et al. (2002) correlated their RgA2 cells with the outer ramifying cells of Tauchi et al. (2002) and Peichl’s (1989) subclass (∂), which Huxlin and Goodchild (1997) classified within type III (RGC2). Dendrites ramify in either the inner or the outer

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A. RIGHT VISUAL FIELD upper vertical meridian nasal

OD ut

horizontal meridian

un x ln AC

temporal lt o

30 azimuth lower B. RIGHT RETINA

C. LEFT SUPERIOR COLLICULUS anterior

dorsal lt

un

ln

AC

ln un posterior

x

ut OD

x

lateral lt

1.0 mm

medial

ut

ventral posterior D. LEFT DLG

E. LEFT VISUAL CORTEX

anterior

lateral

anterior

medial

lateral

Oc2l

Oc2m

medial

RL

dorsal LLA

AM AL l u ln lu LM lt l lu PM u un lu

LL LI

ventral

PL

ut Oc1

posterior

1.0 mm

posterior

FIGURE 2 Schematic diagram of representation of the visual field of the right eye on the retina, superior colliculus, dorsal lateral geniculate nucleus, and visual cortex (Oc). The location of the optic disk is indicated (*), although no projection is present. Open circles, the vertical meridian; dotted line, the horizontal meridian; dashed line, the 30º azimuth. ln, lower nasal; lt, lower temporal; un, upper nasal; ut, upper temporal; u, upper; l, lower. (A) Visual field as seen by the right eye of a rat, which can be imagined to be lying parallel to and above the page, nose to the left. (B) Representation of the field on the right retina viewed from behind the eye, animal’s nose to the left. The region of greatest density of ganglion cells (area centralis, after Dreher et al., 1985) is indicated (x). The shaded area (un) represents the binocular field and is the region of the retina from which some ganglion cells project ipsilaterally (Cowey and Franzini, 1979; Cowey and Perry, 1979). (C) Representation of the visual field of the right eye on the left superior colliculus (SC) viewed from above, rat facing the top of the page; prepared from data of Siminoff et al. (1966). Note that ipsilateral axons terminate in the anteromedial region (un). (D) Representation of the visual field of the right eye on the left dorsal lateral geniculate nucleus (DLG) based on data of Montero et al. (1968) and Reese and Jeffery (1983). Four coronal sections through the nucleus, approximately 0.3 mm apart, are shown. Note that not all parts of the visual field are represented in each section. If the sections were aligned, a line joining the location of the optic disk (*) would indicate the direction of the lines of projection in the nucleus. (E) Representation of the visual field of the right eye on to the left visual cortical areas, redrawn from Dreher et al. (1990) and Thomas and Espinoza (1987). The cytoarchitectonic boundaries are indicated (Oc1 or area 17, Oc2L or area 18a, Oc2M or area 18b). Within each cytoarchitectonic area lie visuotopically organized areas; note that the cytoarchitectonic and functional areas are not exactly coextensive. Subdivisions AM (anteromedial) and PM (posteromedial) lie within Oc2M; AL (anterolateral), LI (laterointermediate), LL (laterolateral), LLA (laterolateral anterior), LM (lateromedial), and RL lie rostrolaterally within Oc2L. Each subdivision contains a representation of the visual field. Callosally connected areas are indicated with shading; note that they are generally prominent around the representation of the vertical meridians. Callosal connections are more prominent at the border between Oc1 and Oc2L and in Oc2M; those medial to Oc1 are less dense (from Olavarria and Montero, 1984).

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sublaminae of the inner plexiform layer; these are likely to represent morphological equivalents of functional on- and off-center cells, respectively (Peichl, 1989; Tauchi et al., 1992). Peichl (1989, 1991) estimated that type I cells in the rat represent 2–4% of all retinal ganglion cells with a preponderance of outer α cells; both innerand outer-ramifying mosaics of type I cells cover the retina with a factor of 3.0 to 3.6. Ipsilaterally projecting cells are among the largest and lie in ventrolateral retina (Ahmed et al., 1996a). It is difficult to align the different classificatory systems. Perry’s type II cells resemble Huxlin and Goodchild’s RGB2 and B3, but the other reported subclasses are not correlated so easily. Although some resemble β ganglion cells (presumed counterparts of X cells) in the cat’s retina (Boycott and Wässle, 1974), type II cells are probably not the morphological counterparts of X-like ganglion cells. In view of the paucity of X-like cells in the rat’s visual pathways (Fukuda et al., 1979; Hale et al., 1979), type II and III cells are likely to correspond to subtypes of W-like cells (for review, see Stone, 1983). Of the two subgroups of type III or RGC cells, Huxlin and Goodchild (1997) suggested that their class RGC1 cells correlate with those projecting to the medial terminal nucleus (MT) of the accessory optic tract (Dann and Buhl, 1987). Sun et al. (2002) confirmed Huxlin and Goodchild’s observations but added subclasses RGC3 and C4, as well as adding additional subclasses within a new class RG (RGD1 and D2). The latter bistratified class strongly resemble the directionselective cells of the rabbit retina. In sections of Golgi- or methylene blue-stained retinae, neurons in the ganglion layer have been classified on the basis of their morphology and the vertical extent of their dendritic spread in the inner plexiform layer (Brown, 1965; Bunt, 1976). These classes cannot be correlated readily with the types identified in whole mounts, although some of Brown’s “loose” ganglion cells probably correspond to type I, whereas Brown’s “tight” cells may correspond to Ni and Dreher’s type IIa cells. The somal sizes of cells projecting exclusively to the SC are smaller than those projecting to DLG (Dreher et al., 1985a; Moriya and Yamadori, 1993; Ni and Dreher, 1981; Schober and Gruschka, 1978), although some project to both nuclei (see discussion in following section). Monoamine oxidase type A has been demonstrated in somata and terminals: in smaller cells located centrally in the retina projecting to SC and larger cells distributed more evenly across the retina projecting to DLG (Nakajima et al., 1998). Cells projecting to the MT appear to be distinctive (Dann and Buhl, 1987; Leak and Moore, 1997) and resemble class RGC (Huxlin and Goodchild, 1997) (see Section III,E). Most of the 7000 retinal ganglion cells projecting to the olivary pretectal

nucleus (OPT) are small (type III), but some appear to be type I or RGA. The majority project contralaterally and, interestingly, are largely distributed within the lower retina (Young and Lund, 1998). Some contain substance P (Miguel-Hidalgo et al., 1991). A subpopulation of small type III or RGC ganglion cells with sparse dendrites, located across the retina, projects to the hypothalamus (Moore et al., 1995). The cell bodies, dendrites and proximal segments of the retinal ganglion cells projecting to the hypothalamic suprachiasmatic nucleus and other retino-recipirnt nuclei involved in circadian photoentrinment or the pupilary light reflex (e.g. olivary pretectal nucleus OPT, see further) contain melanopsin which acts as a phototrnasducer (Hattar et al., 2002). Those ganglion cells which are intrinsically photosensitive depolarize in response to light even when their synaptic inputs from photoreceptors are blocked; they respondi very sluggishly to tonic ambient light (Berson et al., 2002). The somata of the great majority (95%) of these cells are located in the ganglion cell layer, the remainder are displaced to the inner nuclear layer. There are only about 2300–2600 intrinsically photosensitive ganglion cells in rat’s retina (about 2.3% of all ganglion cells), their somata are in the range 12–25 μm (mean 16.3 μm). Although they are distributed throughout the entire retina their density is higher in the superior and temporal quadrants of the retina (Hatter et al., 2002). Morphologically similar (Type III) retinohypothalamic cells contain colocalized glutamate and pituitary adenylate cyclase-activating polypeptide (Hannibal et al., 2000). Other retinohypothalamic cells contain substance P (Takatsuji and Tohyama, 1989). At least some retinohypothalamic cells presumably also project to other visual targets from branches observed in the optic chiasm (Millhouse, 1977). A small number of ganglion cells in inferior retina project to dorsal raphe nucleus; they are skewed to larger somal diameters but have not otherwise been characterized (Fite et al., 1999). About 3000 presumed ganglion cells contain substantial amounts of the enzyme cytochrome oxidase; although cells of all sizes strongly express cytochrome oxidase, many appear to be type I (Land, 1987). Of the ganglion cells projecting to SC, 6% contain γaminobutyric acid (GABA), the principal inhibitory neurotransmitter of the mammalian central nervous system (Caruso et al., 1989); about 3% have also been reported to contain neuropeptide substance P along with GABA (Caruso et al., 1990). Because of technical difficulties in recording single neuron activity from the small eye, virtually the only information on the properties of retinal ganglion cells in the rat comes from recordings made from their axons in optic nerve or tract. Most exhibit classical receptive field organization (Kuffler, 1953): on- or off-discharge

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center and antagonistic off- or on-surrounds (Brown and Rojas, 1965). However, the antagonistic concentric center-surround organization is apparent only under conditions of light adaptation (Cicerone and Green, 1980). In the hooded rat at least, some retinal ganglion cells have on/off-receptive field centers and are directionally selective (Hughes, 1980).

Optic Nerve, Chiasm, and Tract The optic nerve in the adult albino rat contains about 100,000 axons; virtually all are myelinated (Fukuda et al., 1982; Lam et al., 1982; Sefton and Lam, 1984). Fully pigmented and hooded rats are estimated to have higher numbers, 120,000 and 114,000 axons, respectively (Hughes, 1977a; Perry et al., 1983). Although three groups of axons with different conduction velocities are apparent electrophysiologically (Sefton and Swinburn, 1964), myelinated axonal diameters distribute around a single peak (Fukuda et al., 1982; Sefton and Lam, 1984). Intraretinal diameters of nonmyelinated axons of HRP-labeled ganglion cells, however, scatter around three peaks (Ni and Dreher, 1981). A nonactivating Na+ conductance (Stys et al., 1993) is present, and patterns of recovery of K+ after activity have been studied (Ransom et al., 2000). Pathways that project ipsilaterally arise from lower temporal retina (Cowey and Franzini, 1979; Kondo et al., 1993); fibers with fast conduction velocities predominate (Fukuda et al., 1981a; Hale, 1980). Indeed, the largest ganglion cells in the lower temporal retina form about 10–14% of the ipsilaterally projecting cells (Dreher et al., 1985a), but only about 2–4% of the total population (Peichl, 1989). Within the optic nerve immediately behind the eye, the relative positions of axons arising in different retinal quadrants are maintained (Baker and Jeffery, 1989; Yamadori, 1981). Topographic order, however, is lost caudally in the nerve (Baker and Jeffery, 1989) and optic tract (Reese, 1987b), where the location of axons appears to reflect a temporal developmental gradient with the largest axons lying most deeply (Reese, 1987a). The number of fibers in the optic tract, which exceeds that in the nerve (K. Lam and A. Sefton, unpublished), presumably includes any branches of optic axons arising at the chiasm (Millhouse, 1977), as well as fibers that pass from one parabigeminal nucleus (PBG) through the optic chiasm to contralateral visual nuclei (Stevenson and Lund, 1982a, 1982b; Watanabe and Kawana, 1979). These additional fibers likely account for most of the presumed centrifugal axons described in the tract by Schober (1974); only a few would represent truly centrifugal axons, including the serotonergic projections from the dorsal raphe nucleus (Hoogland et al., 1985;

Itaya, 1980; Itaya and Itaya, 1985; Labandiera-Garcia, 1988; Uchiyama, 1989; Villar et al., 1987). Optic tract fibers of various diameters are strongly immunoreactive to an antibody against the calcium-binding protein parvalbumin, but another calcium-binding protein, calbindin D-28k, appears to be present only in smalldiameter fibers (Schmidt-Kastner et al., 1992). Branching is seen in the primary visual pathway (Jeffrey and Kuypers, 1981). Although virtually all ganglion cells project to SC (Linden and Perry, 1983a; Dreher et al., 1985a), fast-conducting axons (Sefton, 1968) and large ganglion cells (Ahmed et al., 1996; Kondo et al., 1993; Moriya and Yamadori, 1993) branch to innervate the contralateral DLG and SC. Large cells in the ventrotemporal crescent also branch to innervate ipsilateral DLG and SC (Ahmed et al., 1996a; Kondo et al., 1993; SC Yamadori et al., 1989); some innervate both ventral geniculate (VLG) and lateral posterior (LP) nuclei (Ahmed et al., 1995) or VLG and pretectum (see Giolli and Towns, 1980). Around 0.4% of retinal ganglion cells branch to innervate both VLG and SC ipsilaterally (Ahmed et al., 1998). Bilateral projections from large type I or RGA cells have been reported (Kondo et al., 1993); only a small minority of fibers (Jeffery et al., 1981) branch at the chiasm to project into both optic tracts (Cowey and Perry, 1979; Cunningham and Freeman, 1977). Glutamate appears to be the major transmitter used by optic nerve fibers (Li et al., 1996). N-Acetylaspartylglutamate (NAAG), a neuron-specific dipeptide that activates glutamic receptors, has been identified in cell bodies and axons of retinal ganglion cells, as well as in retino-recipient nuclei (Anderson et al., 1987; Guarda et al., 1988). When ganglion cells are depolarized, NAAG is released in a calcium-dependent manner in synapses between optic tract terminals and their target neurons in the retino-recipient nuclei (Tsai et al., 1988, 1990). Furthermore, following transection of the optic nerve, there is a dramatic loss of NAAG immunoreactivity in the retino-recipient nuclei (Li et al., 1996; Moffett et al., 1990, 1991). However, NAAG may be a neuromodulator rather than a classical transmitter (Jones and Sillito, 1992), perhaps acting via metabotropic glutamate receptors (Binns, 1999). A small fraction (3%) of retinal ganglion cells are immunoreactive for substance P; they represent a subpopulation of ganglion cells projecting to the SC, which contain GABA (Caruso et al., 1990).

RETINO-RECIPIENT NUCLEI Dorsal Lateral Geniculate Nucleus Introduction The dorsal division of the DLG occupies the dorsolateral part of the dorsal thalamus and relays information

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to the visual cortex. As seen in Atlas, plates 32–39, DLG is bounded dorsally and laterally by retinogeniculate and retinocollicular axons (Paxinos and Watson, 1986). Retinogeniculate axons of various diameters stain intensely with an antibody against parvalbumin, but only fine axons traversing DLG are immunoreactive to calbindin D-28k (Schmidt-Kastner et al., 1992). Axons The DLG has a “striated” appearance imparted by the bundles of axons that traverse it (Cajal, 1911). One set

of axons runs in the direction of the “lines of projection” observed electrophysiologically (Montero and Guillery, 1968; Reese, 1988; Reese and Cowey, 1983; Reese and Jeffery, 1983) (see Fig. 2D). This set incorporates geniculocortical axons that leave the nucleus from its anterior pole, together with corticogeniculate axons and retinogeniculate terminals as they ramify around their targets. A second group of axonal bundles, prominent in coronal sections, runs transversely to the first, parallel to the thalamic border, and includes retinogeniculate axons and other retinofugal fibers destined to terminate

FIGURE 3 Cells and axons in DLG. (A) Golgi-impregnated neurons in DLG. Three examples of TCR (relay) cells and one PSD cell (interneuron). PSD cells contain GABA and calbindin D-28k (see text for references). From Parnavelas et al. (1977). (B) Axon (Golgi material) of the larger type 2a bears, on average, 23 large terminal boutons, shown here drawn from an electron micrograph. From Brauer et al. (1979). (C) The smaller type 2b axon (Golgi preparation) and terminal (electron micrograph). Note that the 2b axon ramifies more extensively than the 2a and has more terminal boutons (on average, 160). From Brauer et al. (1979).

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32. VISUAL SYSTEM

elsewhere, as well as corticogeniculate and corticofugal axons terminating in other structures (Brauer et al., 1979; Cajal, 1911; Lashley, 1934b; Lund, 1966; Montero and Guillery, 1968; Nauta and Bucher, 1954). Three populations of retinofugal axons innervate DLG (Brauer et al., 1979). Only two types have, however, been distinguished in Golgi material (see Fig. 3B) (Brauer et al., 1979,1988; Brauer and Winkelmann, 1974). Collaterals of retinocollicular fibers, particularly in rostral DLG, bear the larger (type 2a) terminals, which can be correlated with simple encapsulated synaptic endings (Brauer et al., 1979, 1988; Lund and Cunningham, 1972). Because virtually all the fast-conducting (presumably largest) axons in optic pathways branch to supply both DLG and superior colliculus (SC) (Sefton, 1968), 2a axons probably include the fast-conducting axons, which synapse with Y-like cells in DLG (Hale et al., 1979). The finer type 2b axons (see Fig. 3B) are found throughout DLG, but concentrate predominantly in its caudal third. Type 2b axons are unlikely to be homogeneous; they probably synapse with cells with fairly diverse, W-like properties (Fukuda et al., 1979; Hale et al., 1979). Fine corticogeniculate axons have numerous short terminal branches, bearing about 400 small boutons that make contact with distal dendrites of relay cells in DLG (Brauer et al., 1974; Grossman et al., 1973). One subset arises in upper cortical layer 6 and a second (destined also for LP) from lower lamina 6. Both arborize in parallel with the lines of projection of the retinogeniculate axons and synapse en passant; their collaterals also innervate the thalamic reticular nucleus (Rt). A third subset, arising as branches of corticocollicular fibers arising in layer 5, terminate in varicose endings (Bourassa and Deschenes, 1995). Myelinated axons probably originate in the visual thalamic reticular nucleus (Rt), whereas rarer, unbranched axons make contacts en passant and could be catecholaminergic (Brauer et al., 1974; Lüth et al., 1977; see, however, see page 1093). Cells Generally, in thalamic nuclei, Cajal (1911) observed two classes of cells: the larger project beyond the given nucleus (Golgi type I) whereas the smaller have axonal terminals confined to the parent nucleus (Golgi type II). It is now, however, apparent that the smaller cells lack an axon. The description here has been largely derived from the papers of Brauer and Schober (1973); Brauer et al. (1974); Brauer and Winkelmann (1974); Grossman et al. (1973); Kriebel (1975); Parnavelas et al. (1977a, 1977b); Webster and Rowe (1984); Werner and Brauer (1984); Werner and Kruger (1973); and Winkelmann et al. (1979). As seen in Fig. 3A, the predominant class of multipolar neurons, class A neurons, have round or oval perikarya 15–25 μm in diameter in Golgi material,

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averaging 13.5 μm in Nissl-stained sections. Distributed throughout DLG, they project to the occipital cortex (Lieberman and Webster, 1974). Class A neurons have multipolar dendritic arbors characterized by three to eight primary dendrites of roughly equal size that branch 6–12 gmm from the cell body to form two or three secondary dendrites. Both secondary and tertiary dendrites of class A neurons have irregularly spaced appendages (Webster and Rowe, 1984). Their dendrites extend for up to 150 μm, bearing scattered, or occasionally clustered, spines. Axons of class A cells do not give rise to recurrent collaterals within DLG. Although morphological differences exist among class A cells, there is no agreement on the basis for any subclassification into Y- or W-like types and none are immunoreactive for a range of neuropeptides (Takatsuji and Tohyama, 1989). In the DLG, smaller relay neurons tend to accumulate laterally, close to the surface of the nucleus in the “outer shell,” whereas the larger class A cells concentrate medially (Grossman et al., 1973; Webster and Rowe, 1984; Werner and Kruger, 1973, see following section). In contrast, the smaller class B (Golgi type II) interneurons in DLG, which represent about 20% of the total number of neurons (Gabbott and Bacon, 1994a), tend to be oriented dorsoventrally parallel to the lateral border of the nucleus (see Fig. 3A). They have ovoid or spindle-shaped cell bodies about 10 μm in diameter (Gabbott et al., 1985, 1986a, 1986b, 1988; Gabbott and Bacon, 1994a) and are most common in the lateral part of DLG (Webster and Rowe, 1984). Lacking an axon, they bear only two to four elongated, relatively thick primary dendrites that branch infrequently; some have spine-like as well as more complex appendages (Brauer and Schober, 1973; Grossman et al., 1973; Kriebel, 1975; Parnavelas et al., 1977; Webster and Rowe, 1984; Werner and Brauer, 1984). These interneurons do not represent the I cells of Burke and Sefton (1966a), although they contain the enzyme glutamic acid decarboxylase (GAD), which synthesizes the inhibitory transmitter GABA as well as GABA itself (Gabbott et al., 1985, 1986a, 1986b, 1988; Gabbott and Bacon, 1994a; Mitrofanis, 1992; Ohara et al., 1983; Ottersen and Storm-Mathisen, 1984; Takatsuji and Tohyama, 1989). They probably correspond to interneurons identified physiologically by Sumitomo and Iwama (1977). Ultrastructurally, interneurons in DLG have discoidal vesicles in their dendrites or appendages and have been described as cells with presynaptic dendrites (PSD; Lieberman and Webster, 1972). The presence of discoidal vesicles within their dendrites or appendages distinguishes them unequivocally from class A cells, which contain round vesicles. About 40% of GABAergic intrageniculate interneurons (class B2) also contain nicotinamide adenine

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dinucleotide phosphate diaphorase (NADPHdiaphorase) (Gabbott and Bacon, 1994a; Mitrofanis, 1992). Diaphorase-positive fibers, presumed extrinsic, are also present (Gabbott and Bacon, 1994a). Furthermore, unlike GABAergic interneurons, which are distributed fairly evenly throughout the DLG, the subpopulation containing both GABA and NADPH-diaphorase is restricted almost exclusively to the dorsolateral part of the nucleus corresponding to the “outer shell” discussed in the following section (Gabbott and Bacon, 1994a; Mitrofanis, 1992). At least some intrageniculate interneurons are also strongly stained with antibodies against calbindin D-28k (Schmidt-Kastner et al., 1992). Interneurons are also involved in mediating cholecystokinin-induced changes in firing rates of geniculate relay neurones and may enhance antagonistic receptive field organization (Albrecht et al., 1995). Regional Organization in DLG Despite the lack of lamination in DLG of the rat, several authors have discerned regional variations in the distribution of cells of different sizes, in the composition of afferent axons, and in patterns of retrograde degeneration after cortical lesions, although accounts differ in detail. Reese (1988) emphasized that the nucleus consists of an “outer shell” located caudodorsally and an “inner core” constituting the ventromedial part. Regions of DLG, which reveal different levels of cytochrome oxidase activity (Land, 1987), appear to be innervated by different morphological classes of retinal ganglion cells (Martin, 1986) characterized by different conduction velocities (Gabriel et al., 1985). Because the conduction velocities of axons in optic nerve and relay cells in DLG are closely correlated, differences in the regional distribution of inputs imply that different cell types within DLG are located in different broad regions of the nucleus (Fukuda et al., 1979; Hale et al., 1979; Noda and Iwama, 1967; Reese, 1988). The outer shell receives afferents from retina that are almost exclusively of the small 2b class (Brauer et al., 1979), as well as from SC (Harting et al., 1991a; Reese, 1984; Taylor et al., 1986) and the parabigeminal nucleus (Harting et al., 1991b). Ultrastructurally, complex endings are most evident (Lund and Cunningham, 1972). Cells in the region project to cortical area Oc2L (cytoarchitectonic area 18a), not only to Oc1 (primary visual or striate cortex, cytoarchitectonic area 17 or area V1; Hughes, 1977). Caudally, small to medium-sized cells with small dendritic fields predominate (Gabbott et al., 1986a; Werner and Kruger, 1973) and the density of interneurons is reduced (Gabbott et al., 1986a). Cells recorded in this region are more likely than cells lying rostromedially to change their responses to photic stimuli when paired with conditioning shocks applied

to the tail (Albrecht et al., 1990, 1991). More rostrally, the organization is more complex, with large and small optic tract terminals overlapping completely (Brauer et al., 1979). Ipsilateral axons with fast and medium conduction velocities terminate medially, in the inner core (Fukuda et al., 1981a; Hale, 1980; Hayhow et al., 1962; Reese, 1988; Reese and Cowey, 1983), where larger cells lie (Werner and Kruger, 1973). After eye removal, axons terminating medially in DLG (including the large fibers originating from the ipsilateral eye) degenerate more rapidly than axons terminating laterally; this latter group exhibit neurofibrillar changes (Cunningham and Lund, 1971; Lund et al., 1976). We conclude that neurofibrillar degeneration after eye removal is a feature of the smaller axons, which degenerate more slowly. A close relationship exists between neurons and glia in DLG; glial fibrillary acidic protein is increased in astrocytes following enucleation of the eye or inactivation of retinal cells (Canady et al., 1994). Synaptic Organization In this description we use the terminology of Lieberman and Webster (1974); for a summary of other nomenclature, see Winkelmann et al. (1976). Retinal terminals within DLG (R boutons), containing round synaptic vesicles and pale mitochondria, make Gray type 1 (asymmetrical) presumably excitatory synapses with the dendrites of both PSD (interneurons with presynaptic dendrites) and TCR (thalamocortical relay) cells (Bruckner et al., 1972; Lieberman and Webster, 1972, 1974) (see Figs. 4A and 4B). R boutons, which include the largest terminals in DLG, are found within (1) simple unencapsulated, (2) simple, and (3) complex encapsulated synaptic regions. They are never postsynaptic to other terminals (Lund and Cunningham, 1972). By analogy with terminals identified in Golgi material, they have been divided into two classes (see Figs. 3B and 3C) (Brauer et al., 1979; Lenkov et al., 1978; Winkelmann et al., 1976). As seen in Fig. 4C, the small terminals of cortical origin (SR) also contain round synaptic vesicles but have dark mitochondria (Bruckner et al., 1972; Lund and Cunningham, 1972) They make Gray type 1, simple unencapsulated (extraglomerular), presumably excitatory synapses with distal TCR cell dendrites but not with PSD cells (Lieberman and Webster, 1974). Additional synaptic terminals in DLG containing round or dense-cored vesicles also make Gray type 1 synapses (Bruckner et al., 1972; Lund and Cunningham, 1972). Two classes of terminals containing pleomorphic or discoidal vesicles (P and F boutons) were first distinguished by Lieberman and Webster (1972). P boutons (Fig. 4A, B) arise from dendrites of the intrinsic PSD cells, occurring singly or in interconnected chains,

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FIGURE 4 Synaptic relationships in DLG. Prepared by A.R.Lieberman from previously unpublished micrographs of P.T.Ohara (A), A.R. Lieberman (B), and A.Sefton (C). (A) Examples of a TCR cell dendrite (D) with a large retinal terminal containing round vesicles (R) and a presynaptic dendrite containing discoidal vesicles (P) are enclosed in glial lamellae (a glomerulus). Filamentous contacts (nonsynaptic) can be seen between R and D profiles. Above is an F terminal containing flattened vesicles. A smaller extraglomerular terminal (SR), which is probably of cortical origin, also contains round vesicles but its mitochondrion is different from those in the R bouton. (B) Both an R and a P bouton are making synaptic contact with the spine of a TCR dendrite. (C) An F terminal and an SR bouton (possibly cortical) are shown at synapses. Note the differences in their synaptic specializations.

corresponding to the complex appendages recognized in Golgi impregnations (Grossman et al., 1973). P boutons make Gray type 2 (symmetrical), presumably inhibitory synaptic contacts with dendrites (shafts and spines) of relay cells and with other P boutons. They are postsynaptic to F, R, and other P boutons and are found

commonly in complex zones encapsulated by glia (glomeruli; see Szentagothai, 1973). F boutons (Figs. 4A and 4C) have a dark cytoplasmic matrix containing densely packed cylindrical vesicles and often several mitochondria. Some have been traced to myelinated axons. Never postsynaptic to other

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profiles and found infrequently in glomeruli, F boutons make Gray type 2 synapses with TCR and PSD cells (Lieberman and Webster, 1974). They include terminals of axons arising in the adjacent visual Rt, which synapse with relay cells (Montero and Scott, 1981; Ohara et al., 1980); those F boutons that synapse with PSD cells must arise elsewhere. Although synaptic relationships may be simple, occasional reciprocal synapses are seen. In glomeruli, a P bouton is often both postsynaptic to an R bouton and presynaptic to a TCR dendrite with which the R bouton also synapses directly (a triad or triplet synapse): R ⎯⎯⎯⎯→ P → TCR → ⎯

Additional complex arrays all have a central P terminal, with R, F, or another P bouton presynaptically and a PSD or a TCR dendrite postsynaptically (Lieberman and Webster, 1974). Physiology of DLG and Rt Physiology and function of DLG Burke and Sefton (1966a, 1966b, 1966c) classified cells recorded in DLG as P (principal or relay) and I (interneuronal) on the basis of electrical properties. P cells respond to a shock applied to the optic nerve or tract with a single, short latency action potential followed intermittently by a burst of three to five spikes for periods of 1 s or more; they can be activated antidromically from Oc1 (Burke and Sefton, 1966) and from Oc2L (Sumitomo et al., 1988). They possess intrinsic low-frequency activity; two calcium currents—low and high voltage activated—have been suggested to underlie their oscillatory behavior (Leresche et al., 1991). A differential distribution of L-type calcium channels may affect the transition from bursting to single spike activity. They are located at the bases of dendrites in relay cells, in contrast to their distribution across the somata of interneurons and their centralized localization in Rt cells (Budde et al., 1998). The Na+ current exhibits both inactivating and noninactivating characteristics; the latter may also contribute to both tonic and burst firing of neurons (Parri and Crunelli, 1998). Within the DLG of the cat, inputs from retina and layer 6 of the ipsilateral visual cortex are associated with two different metabotropic glutamate receptors: mGluR1 is located in terminals postsynaptic to corticogeniculate axons and mGluR5 in terminal profiles associated with inhibitory F2 terminals associated with retinal input to relay cells (Godwin et al., 1996). About two-thirds of the cells in DLG of the rat have properties reminiscent of the heterogeneous W class of cells in the cat, whereas most of the remainder have Y-

like properties. Very few resemble the X cells of cat (Fukuda et al., 1979; Hale et al., 1979). When Hale et al. (1979) correlated conduction velocities of afferents with receptive field properties, the fastest conducting group of optic tract fibers (t1) innervates Y-like cells and a few tonic cells with X-like properties; the second conduction velocity group (t2) innervates phasic W-like cells, including all the direction-selective cells (Montero et al., 1968; Sefton and Bruce, 1971); the third group of optic axons (t3) innervates nonconcentric cells. In the small ipsilateral pathway, conduction velocities of the retinal afferents fall into the t1 or t2 groups, and cells in DLG innervated from the ipsilateral eye respond to light either on or off, with tonic responses predominating (Fukuda et al., 1981a, 1981b; Hale, 1980). Attempts have been made to align aspects of the DLG in the rat with that of the cat. Although Fukuda (1973) had suggested that the rat’s DLG contains a region homologous or even analogous to the cat’s A laminae, such homology is doubtful. Indeed, the overlap of latencies (both orthodromic and antidromic) and the range of properties of P cells do not support this idea (Hale et al., 1979). Reese (1984), on the basis of the location of colliculogeniculate terminals in the rat’s DLG, suggested that the caudal and outer parts correspond to the C laminae of the cat’s DLG. Although Lennie and Perry (1981) classified P cells in the DLG of the rat as “X” (with linear spatial summation of visual signals) or “Y” (nonlinear spatial summation), these two groups no doubt include concentric W-like cells defined using other criteria, as concentric W cells in the cat reveal either linear (X-like) or nonlinear (Y-like) summation (Sur and Sherman, 1982). Distinguishing intrageniculate interneurons from P cells on the basis of their receptive field properties is difficult unless antidromic activation from cortex can firmly be excluded (Dubin and Cleland, 1977). Nevertheless, Sumitomo and Iwama (1977) recorded from a small population of cells that they regarded as intrageniculate interneurons Otherwise resembling P cells, they lack the postexcitatory inhibition that is a feature of most P cells (Anderson et al., 1977; Burke and Sefton, 1966b). The electrical characteristics of P and geniculate interneurons in slices are distinctive, with the interneurons displaying a longer time constant and higher input resistance and oscillatory responses, regardless of polarization (Zhu et al., 1999a, 1999b). In interneurons, both EPSPs and IPSPs were generated following low-frequency stimulation of the optic tract, indicating that the interneurons are subject to both excitatory and inhibitory influences. The interneurons are themselves inhibited by GABAA but not GABAB, and their responses are marked by a slow spike–frequency adaptation (Williams et al., 1996). Both a short-acting GABAA-

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32. VISUAL SYSTEM

mediated IPSP and a long-acting GABAB-mediated IPSP, possibly arising from separate populations of intrageniculate interneurons, had been observed previously (Crunelli et al., 1988). It seems likely that geniculate interneurons take part in a feedforward inhibition of P cells, presumably altering their transfer characteristics (Zhu and Lo, 1999). If they represent the functional counterparts of PSD cells, their reported activation from the cortex must be indirect, via TCR (P) cells. Indeed, it is likely that some of the relay cells in DLG in the rat, as in the cat, could be classified as “lagged cells.” Such cells are characterized by a prolonged latency to the onset of a stimulus and their ongoing discharges are initially suppressed before increasing to a relatively steady maintained level. In the cat, geniculate interneurons with receptive fields congruent with those of relay cells innervated by them are responsible for the inhibition observed in the lagged relay neurons (Humphrey and Weller, 1988; Mastronarde, 1987a, 1987b). In cat, the responses of nonlagged cells are mediated via both Nmethylaspartate (NMDA) and non-NMDA receptors (Jones and Sillito, 1992). Physiology and function of visual thalamic reticular nucleus (Rt) I cells respond disynaptically with a burst of up to 12 spikes to an optic nerve stimulus and transynaptically to cortical stimulation; late bursts also occur. I cells are responsible for the powerful postsynaptic inhibition that follows the initial activity of most P cells (Burke and Sefton, 1966a, 1966b, 1966c), probably mediated by GABAB receptors (Crunelli et al., 1988; Hirsch and Burnod, 1987). The somata of I cells lie outside DLG, in the visual sector of the Rt (Hale et al., 1982; Sumitomo et al., 1976) (see Section IV,A). These reticular I (rI) cells, which inhibit P cells in DLG, appear to be functionally identical with the original I cells of Burke and Sefton (1966a, 1966c). First, the inhibitory effects on P cells disappear after specific damage to visual Rt (French et al., 1985; Sumitomo et al., 1976). Second, inhibitory activity is altered in slices of DLG separated from visual Rt (Godfraind and Kelly, 1981). Third, cells in visual Rt contain the GABA-synthesizing enzyme, GAD (Ohara et al., 1983), and thus GABA (Ottersen and StormMathisen, 1984), which inhibits cells in DLG (Kayama, 1985; Kayama et al., 1981). In contrast to P cells in DLG, rI cells have large receptive fields and transiently increase their discharges when photic stimuli are switched both on and off. They also respond to fast-moving stimuli (Sefton and Bruce, 1971; Sumitomo et al., 1977) and are innervated indirectly, via P cells in DLG, by the fastestconducting retinal axons (Hale et al., 1982). Cells in Rt fire in two modes—tonic and oscillatory—as a result of

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strong afterhyperpolarizing potentials that follow bursts of activity (Avanzini et al., 1989). Cells in Rt generally, and presumably also within the visual sector, have a similar orientation and disklike dendritic field (Ohara and Havton, 1996). They are innervated by terminals with asymmetrical synapses (presumably excitatory) that outnumber presumed inhibitory GABAergic terminals. Small corticothalamic terminals are more numerous in Rt than those of thalamic origin, presumed to be branches of thalamocortical fibers (Liu and Jones, 1999). Terminals from each sector of Rt display circumscribed terminal patterns with about 4000 boutons in the related primary thalamic nucleus, including DLG (Pinault and Deschênes, 1998). Dendrodendritic synapses are present in Rt (Pinault et al., 1997), and inhibitory interactions have been reported within Rt (Huguenard and Prince, 1994; Ulrich and Hugeuenard, 1995, 1996). Consistent with these observations, activation of one subtype of metabotropic glutamate receptors excited cells in Rt in slices, whereas activation of another led to a long-lasting hyperpolarization, potentially providing a mechanism for disinhibition (Cox and Sherman, 1999). Cells in Rt are depolarized by cholecystokinin (Cox et al., 1995). The other major input to the visual Rt arises in visual cortical areas Oc1, 2L, and 2M (Ohara and Lieberman, 1981a, 1981b; Sefton et al., 1981), overlapping with the projections from DLG (Coleman and Mitrofanis, 1996). Cortical projections arise in layer 6 (Bourassa and Deschênes, 1995; Montero, 1999) as branches of fibers projecting to DLG and lateral posterior nuclei (LP) (Bourassa and Deschênes, 1995). These projections are mapped topographically and overlap with the projections from DLG (Coleman and Mitrofanis, 1996) (see Table 1). Visual regions of the Rt also receive at least partly segregated projections from brain stem regions: the sector related to DLG from midbrain reticular nucleus, deep SC, and peripeduncular nucleus; those projections related to LP arise particularly from deep SC (Kolmac et al., 2000). Damage to the visual sector of Rt is associated with deficits on an orienting task (Weese et al., 1999). Rt has been implicated in focal attentional mechanisms (Montero, 1999); for a general review, see Guillery et al. (1998). The most complete description of the fine structure and organization of visual Rt is that of Ohara and Lieberman (1985). Dendrites within it, either smooth or varicose, are oriented like a grid across the bundles of axons passing from the thalamus to the cortex (Requena et al., 1991; Scheibel and Scheibel, 1966). In the visual sector, two morphological types of neurons have been described. Bitufted neurons have two to four primary dendrites and somata of variable sizes and shapes; their dendrites are spinous, with spines being

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up to 4 μm in length. The multipolar cells have stellate or ovoid somata, up to 30–40 μm in diameter and four to seven primary dendrites with hair-like protrusions (Requena et al., 1991). Within each sector of the Rt, axons arise from either somata or the proximal segments of primary dendrites (Requena et al., 1991) and project exclusively subcortically, back into the thalamus, particularly to the nucleus supplying the reticular input (Jones, 1975). In this way, cells in visual Rt project back to the topographically related parts of DLG (Hale et al., 1982; Montero and Scott, 1981; Ohara and Lieberman, 1981a, 1981b; 1985; Ohara et al., 1980). Despite some earlier suggestions, the thalamic reticular nucleus does not appear to project to SC (Taylor et al., 1986) or to VLG (Ohara et al., 1980). Visual Rt also receives a specific

TABLE 1 Afferentsa Retina (c) Retina (c,i) Rt(i) Oc1(i) Oc1(i) layer 6b Oc2L(i) Layer 6b Oc2M layer 6 SC(i) SuG Pretectum OPT(i) OPT(c) OT(i) PBG(c) PBG(i) DR(i,c) LC(i,c) LDTg PAG(i,c)b PPTg(i,c)b Efferents Rt Oc1 Layer 4 Lower layer 3 Layer 6 Layer 1 Oc2L Oc2Mb

projection from visual cortical areas 17 and 18 (Coleman and Mitrofanis, 1996) themselves innervated from DLG (see Table 1). Rt also receives inputs from the brain stem (MackaySim et al., 1983): the sector related to DLG from the midbrain reticular nucleus, deep SC, and peripeduncular nucleus; that related to LP particularly from deep SC (Kolmac et al., 2000). Locus coeruleus provides a particularly dense norepinephrinergic innervation (Lindvall et al., 1974; Swanson and Hartman, 1975), with en passant boutons making asymmetrical synapses with dendrites (Asanuma, 1992). Iontophoretic application of norepinephrine or electrical stimulation of locus coeruleus results in the development of a slow depolarization of GABAergic cells in Rt and DLG mediated

Connections of Dorsal Lateral Geniculate Nucleus

Lashley, 1934a Nauta and Straaten, 1947; Hayhow et al., 1962; Schober, 1975; Hickey and Spear, 1976; Perry and Cowey, 1979; Kondo et al., 1993; Moriya and Yamadori, 1993; Ahmed et al., 1995, 1996a; Nakajima et al., 1998 Ohara et al., 1980; Montero and Scott, 1981; Ohara and Lieberman, 1981a; Hale et al., 1982; Pinault et al., 1995; Pinault and Deschênes, 1998 Nauta and Bucher, 1954; Goodman and Horel, 1966; Montero and Guillery, 1968; Jacobson and Trojanowski, 1975; Sefton and Martin, 1984; Al-Abdulla et al., 1998; Mason and Groos, 1981; Sefton et al., 1981; Bourassa and Deschênes, 1995 Takahashi, 1985 Mason and Groos, 1981; Sefton et al., 1981 Sefton et al., 1981 Perry, 1980; Pasquier and Villar, 1982b; Takahashi, 1985 Mason and Groos, 1981; Mackay-Sim et al., 1983; Lane et al., 1997 Pasquier and Villar, 1982a Mackay-Sim et al., 1983 Turlejski et al., 1993 Mackay-Sim et al., 1983 Mackay-Sim et al., 1983; Sefton and Martin, 1984; Harting et al., 19991b; Turlesjski et al., 1993 Harting et al., 1991a; Turlesjski et al., 1993 Lüth et al., 1977; Moore et al., 1978; Pasquier and Villar, 1982a; Kromer and Moore, 1980; Pasquier and Villar, 1982b; Mackay-Sim et al., 1983; Turlejski et al., 1994 Swanson and Hartman, 1975; Jones and Moore, 1977; Kromer and Moore, 1980; Pasquier and Villar, 1982b; Mackay-Sim et al., 1983; Turlejski et al., 1994 Kayama et al., 1986a; Kayama et al., 1986b; Satoh and Fibiger, 1986; Woolf and Butcher, 1986; Cornwell et al., 1990; Shiromani et al., 1990; Turlejski et al., 1994 Mackay-Sim et al., 1983; Turlejski et al., 1994 Hoover and Jacobowitz, 1979; Pasquier and Villar, 1982a; Mackay-Sim et al., 1983; Mesulam et al., 1983; Jones and Yang, 1985; Sofroniew et al., 1985; Hallanger et al., 1987; Rye et al., 1987; Shiromani et al., 1990; Turlejski et al., 1994 Coleman and Clerici, 1980; Sefton et al., 1981; Hale et al., 1982; Itaya and van Hoesen, 1982b Clarke, 1932; Lashley, 1934b; Waller, 1934; Ribak and Peters, 1975; Hughes, 1977; Coleman and Clerici, 1980; Perry, 1980; Cornwell et al., 1990 Peters and Feldman, 1976; Lüth et al., 1977; Johnson and Burkhalter, 1992 Peters and Feldman, 1976 Peters and Saldanha, 1975; Peters and Feldman, 1976 Peters and Feldman, 1976; Lüth et al., 1977 Ribak and Peters, 1975; H. Hughes, 1977; Coleman and Clerici, 1980; Cusick and Lund, 1981; Cornwell et al., 1990 Perry, 1980

a

c, contralateral; i, ipsilateral; c,i, bilateral. Also projects to visual Rt (Sefton et al., Mackay-Sim et al., 1983; Bourassa and Deschênes, 1995).

b

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through the activation of α1-adrenoreceptors (Kayama et al., 1982; McCormick and Wang, 1991). Cells containing choline acetyltransferase and parvalbumin (Jourdain et al., 1989) in the caudal basal forebrain provide another source of inputs to Rt (Asanuma and Porter, 1990; Hallanger et al., 1987; Levey et al., 1987b) and dense plexuses of immunoreactive fibers positive for choline acetyltransferase are present (Levey et al., 1987a). Weak stimulation of the cholinergic projection arising in the laterodorsal tegmental nucleus (Cornwell et al., 1990; Cornwell and Phillipson, 1988; Hallanger et al., 1987) depresses spontaneous and evoked activity in visual Rt cells, whereas stronger stimulation results in a return of spontaneous activity, presumably by activating a cortical loop (Kayama et al., 1986a, 1986b). Iontophoretically applied acetylcholine, while not generally altering the ongoing activity of cells in Rt, reduces their excitatory responses to glutamate (Marks and Roffwarg, 1991). Other immunoreactive plexuses are positive for serotonin (Cropper et al., 1984). Afferents to DLG Apart from the retinal input, afferents to DLG arise in diverse sources: Oc, visual Rt, and various nuclei of the brain stem. These are listed in Table 1, but some specific issues are discussed here. For the locations of sources of the chemical pathways described, see Paxinos et al. (1999a, 1999b). Retinal projection to DLG One of the major inputs to DLG arises from the contralateral eye. In albino animals, within the region of the restricted ipsilateral input, a concealed lamination has been discerned in DLG, such that alternate laminae receive their predominant input from alternate eyes (Hayhow et al., 1962). In contrast, fully pigmented strains with dark eyes have one lamina within the region of the ipsilateral input (Giolli and Creel, 1974; Guillery et al., 1971; Lund et al., 1974). As first demonstrated by (Lashley, 1934a), the retina is mapped topographically in the rat’s DLG (see Fig. 2D). A point in the retina is represented by a “line of projection” through DLG (Montero et al., 1968; Montero and Guillery, 1968; Reese and Cowey, 1983). Because each temporal retina projects to the DLG on both sides, the vertical meridian is not represented in a sharp line as a column of cells (Lund et al., 1974). At least in hooded rats, the ipsilateral projection, arising from the far temporal retinal crescent, is binocularly conjugate with an adjacent crescent displaced toward the optic disc (Reese and Jeffery, 1983) and terminates in DLG in such a way that corresponding retinal points are represented along the lines of projection (Reese and Cowey, 1983). As noted in the section on “Retinal Output”, evidence suggests that retinogeniculate transmitters include

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acetylated dipeptides related to NAAG, which is present in retinal ganglion cells and their projections (Anderson et al., 1987; Moffett et al., 1991; for review, see Pasik et al., 1990). Glutamate is an important transmitter in SC (Binns, 1999). NAAG acts on glutamate receptors either directly or through acidic amino acids derived from the dipeptide by endopeptidase-mediated cleavage. Because relay neurons possess NAAG receptors permeable to Ca2+ and other less permeable N-methyl-d-aspartate (AMPA) receptors, it is possible that NAAG acts on the latter receptors with a weak kainate-like action (Harata et al., 1999). Synchronous activity in DLG results in an extracellular Ca2+-dependent H+ sink (Tong and Chesler, 1999), perhaps modulating the role of NMDA receptors present in DLG (Turner et al., 1994; Williams et al., 1996). Following enucleation, a fiber plexus immunoreactive to calretinin is reduced, suggesting that it is present in retinal axonal terminals (Arai et al., 1992). GABAA and GABAB receptor-mediated responses are recorded in DLG neurons (Soltesz and Crunelli, 1992). Three types of GABA receptors are expressed in local interneurons of rat’s DLG: GABAA bicuculline-sensitive receptors, GABAA bicuculline-insensitive receptors, and GABAB receptors (Zhu and Lo, 1999). Consistent with mediation via Cl− channels, GABAA receptor-mediated responses have a reversal potential of approximately − 82 mV. In contrast, GABAB receptor-mediated responses have a reversal potential of approximately −97 mV, which is consistent with mediation via the K+ channel. Stimulation of the Rt evokes inhibitory postsynaptic potentials (IPSP) in DLG interneurons; these IPSP are mediated exclusively by GABAA bicuculline-sensitive receptors. They exhibit a prominent hyperpolarizationactivated cation conductance, likely to modulate firing patterns and thalamic oscillations. Interneurons in DLG are electrotonically compact and frequently fire in bursts in response to a small depolarizing current, perhaps mediating the feedforward inhibition on relay cells discussed earlier (Zhu et al., 1999c). Thus, DLG interneurons are likely to be involved in complex neuronal circuitry involving geniculocortical relay cells as well as Rt cells (Zhu and Lo, 1999) (see page 1088). Cortical projections to DLG In all eutherian mammals, the numerically largest input to DLG originates from the ipsilateral cortex (for review, see Garey et al., 1991). Axons arising in the occipital cortex (Oc) terminate on relay cells in the retinotopically related part of the ipsilateral DLG (Jacobson and Trojanowski, 1975; Montero and Guillery, 1968), making Gray type I synapses (Lieberman and Webster, 1974). Those projecting to DLG arise in upper layer 6, branching also to supply Rt but not other thalamic visual centers. Within

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DLG they terminate in rostrocaudal “rods,” emitting synapse en passant. Cells in lower layer 6 of Oc projecting to LP emit branches to DLG (Bourassa and Deschênes, 1995). In DLG, activation of corticogeniculate axons enhances the responses of receptive field centers and reduces the strength of responses and antagonism of the surrounds of the majority of off-center, but only about half of the on-center, cells (Molotchnikoff et al., 1984). The most likely neurotransmitters released by corticogeniculate axons are excitatory amino acids: glutamate (Baughman and Gilbert, 1981; Kvale et al., 1983; Lund et al, 1978) and/or D-aspartate (Johnson and Burkhalter, 1992). In experiments involving either stimulation or cooling of the visual cortex, neurons in occipital cortical area 1 (Oc1) appear both to excite and to inhibit cells in DLG (Kayama et al., 1984; Sefton and Bruce, 1971), presumably via GABAergic interneurons within DLG or visual Rt. Indeed, axons arising in Oc also innervate the visual Rt (Bourassa and Deschênes, 1995; Leong, 1980; Ohara and Lieberman, 1981a, 1981b). The projection arises in small pyramidal cells of layer 6 of areas Oc1, Oc2L, and Oc2M (Sefton et al., 1981; Bourassa and Deschênes, 1995). Visual Rt is supplied by branches of corticogeniculate axons (Bourassa and Deschênes, 1995), and terminals of cortical origin within it are similar to SR terminals in DLG (Ohara and Lieberman, 1981b). The corticoreticular projection modulates the activity of relay cells in DLG in a nonspecific way (Molotchnikoff et al., 1984), and it is suggested that neurons of the visual Rt are dependent on a tonic cortical input (Kayama et al., 1984). Subcortical projections to DLG So far, no specific role has been proposed for the projections to DLG from retino-recipient regions: SC, nucleus of the optic tract (OT), olivary pretectal (OPT), parabigminal (PBG), and lateral hypothalamus (LH). The tectogeniculate projection, however, originates exclusively from the caudal two-thirds of the superficial retino-recipient laminae of SC where the contralateral visual field is represented (see Harting et al., 1991a; Hilbig et al., 2000). In pigmented rats the parabigeminogeniculate projection originates mainly from the rostral third of the contralateral nucleus in which the temporal retina and vertical meridian are represented. In albino rats, the projection is significantly smaller, originating mainly from the caudal half of the contralateral parabigeminal nucleus where the nasal retina is represented. In both strains a very small projection from the ipsilateral parabigeminal nucleus constitutes about 2.5% of the cells projecting to DLG in pigmented, but only 1.5% in albino rats (Turlejski et al., 1993). The terminal fields of the tectogeniculate projection overlap with those of the parabigeminogeniculate

projection in the lateral portion of the DLG (Harting et al., 1991b), adjacent to the optic tract (Reese, 1984; see Section IV,B,6). Thus, the tectogeniculate and parabigeminogeniculate projections to the DLG terminate in that portion of the nucleus containing W-like, but not Y-like, cells (Brauer et al., 1988; Fukuda et al., 1979; Gabriel et al., 1985; Hale et al., 1979; Martin, 1986). In turn, the tectogeniculate outflow from the superficial gray (Mackay-Sim et al., 1983; Sugita et al., 1983) is likely to be driven by W-like retinal afferents (Fukuda, 1977). Evidence shows that substance P and/or cholecystokinin containing neurons are involved in these SC projections to DLG (Harvey et al., 2001; OgawaMeguro et al., 1992) and many projection neurons are immunoreactive for calbindin (Lane et al., 1993, 1997). The role of a small projection arising in zona incerta (Power et al., 1999) is not known. Early functional studies of the “nonvisual” projections to the DLG involved placing stimulating electrodes in the brain stem in close proximity to the dorsal tegmental bundle in which axons from several brain stem centers travel to DLG and Rt (Mackay-Sim et al., 1983). Not surprisingly, electrical stimulation at these sites resulted in mixed facilitatory and inhibitory effects on neurons in DLG (Fukuda and Iwama, 1971; Theil et al., 1972). Disinhibition does not account for the facilitation observed in such studies (Waring, 1979). Cells in DLG exhibit oscillatory rhythms, thought to be under the influence of aminergic inputs (Zhu and Ulrich, 1998). Diffuse catecholaminergic fibers observed within DLG (Andén et al., 1966; Fuxe, 1965; Papadopoulos et al., 1989b) arise from norepinephrinergic cells in locus coeruleus (LC, Jones and Moore, 1977; Lüth et al., 1978; Jones and Yang, 1985; Kromer and Moore, 1980; Lindvall et al., 1974; Swanson and Hartman, 1975; Turlejski et al., 1994). Of the neurons in locus coeruleus that innervate DLG, about 60–65% project ipsilaterally, about 35–40% contralaterally (Kromer and Moore, 1980), with about 25% of the latter projecting bihemispherically to innervate the ipsilateral nucleus as well (Turlejski et al., 1994). Fibers containing norepinephrine form highly branched networks of unmyelinated varicosities. The varicosities have diameters ranging from 0.4 to 1.5 μm and contain a few mitochondria among densely packed vesicles. They usually form symmetrical synapses with dendritic shafts and spines, outside the synaptic glomeruli (Papadopoulos and Parnavelas, 1990b). Both electrical stimulation of locus coeruleus and local iontophoretic release of norepinephrine produce a slow membrane depolarization of cells in DLG (Kayama et al., 1982; Rogawski and Aghajanian, 1980a, 1980b; 1982). The effect is mediated by α1 receptors (Kayama, 1985; Rogawski and Aghajanian, 1982), which are present in large numbers in DLG (Jones et al., 1985;

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Palacios et al., 1987; Palacios and Kuhar, 1982; Unnerstall et al., 1984). At the same time, electrical stimulation of locus coeruleus or iontophoretic application of norepinephrine reduces the excitability of intrageniculate interneurons (Kayama, 1985). The norepinephrinergic innervation of Rt is even more prominent (see later); activity in locus coeruleus has been suggested to mediate a tonic activation of the thalamus (Asanuma, 1992; McCormick and Wang, 1991). Although locus coeruleus has been implicated in the generation of pontogeniculooccipital waves, which correlate with features of rapid eye movement sleep in the rat (Marks et al., 1980), depletion of norepinephrine and serotonin (see later) does not alter the sleep state-related patterns of activity in neurons of DLG (Marks et al., 1989). There is a smaller dopaminergic input to DLG (Papadopoulos and Parnavelas, 1990a) arising bilaterally (Turlejski et all., 1994). Moderate numbers of D2 dopaminergic receptors have been found throughout DLG (Bouthenet et al., 1987). The fine dopaminergic fibers, which might originate in mesencephalic reticular formation, as well as from dopaminergic neurons in the dorsal raphe nucleus (DR) (Steinbusch et al., 1980), give rise to irregularly spaced varicosities. The varicosities, typically containing one to three mitochondria and numerous ovoid synaptic vesicles, usually form asymmetrical contacts on dendritic shafts or spines, some of which were identified as those of GABAergic interneurons (Papadopoulos and Parnavelas, 1990a). DLG also contains fine, varicose, immunoreactive serotonergic (5-HT) fibers (Cropper et al., 1984; Fuxe, 1965; Lüth and Seidel, 1987; Mantyh and Kemp, 1983; Papadopoulos et al., 1987; Parent et al., 1981; Steinbuch, 1981; Ueda and Sano, 1986; Wadhwa et al., 1990). They appear to originate from serotonergic cell bodies that constitute 40–50% of all neurons in dorsal raphe (Azmitia and Segal, 1978; Bobillier et al., 1975; Lüth et al., 1977; Moore et al., 1978; Pasquier and Villar, 1982a; Turlejski et al., 1994; Villar et al., 1988). Of dorsal raphe neurons projecting to DLG, about 60% project ipsilaterally and 40% contralaterally; of the latter, about 25% also project ipsilaterally (Turlejski et al., 1994). In addition, about 20–25% of both serotonergic and nonserotonergic neurons in the ipsilateral dorsal raphe that innervate DLG send collaterals to the SC while few in the contralateral dorsal raphe innervate both DLG and SC. About 20% of dorsal raphe neurons projecting to DLG are not serotonergic (Villar et al., 1988). Serotonergic varicosities form synapses with dendritic spines and shafts and only occasionally with somata; axodendritic synapses are asymmetrical (presumably excitatory), whereas axosomatic synapses are symmetrical (Papadopoulos and Parnavelas, 1990b). These synapses change postnatally and may contribute to morphogenesis (Dinopoulos et al.,

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1995). The local iontophoretic release of 5-HT results in a slow onset prolonged inhibition (Yoshida et al., 1984) of the spontaneous firing of P cells within DLG (Marks et al., 1989; Rogawski and Aghajanian, 1980b), as well as thalamic reticular cells (Yoshida et al., 1984). These effects are reproduced by the electrical stimulation of dorsal raphe (Kayama et al., 1984; Yoshida et al., 1984). The inhibitory effects of 5-HT on P cells is probably indirect, as, at least in cat and guinea pig, 5-HT exerts a potent excitatory action on the GABAergic neurons of the Rt (McCormick and Wang, 1991). The primary action of 5-HT in DLG is likely to involve 5-HT1 rather than the other three groups of 5-HT receptors, as 5-HT1 receptors are present in greater concentrations (Pazos et al., 1985; Pazos and Palacios, 1985). Cholinergic terminals in DLG (Hoover and Jacobowitz, 1979; Sofroniew et al., 1985) most likely arise from the cholinergic pedunculopontine nucleus (PPTg; Rye et al., 1987), parabrachial, and/or laterodorsal tegmental nuclei (LDTg; Cornwell et al., 1990; Hallanger et al., 1987; Mackay-Sim et al., 1983; Mesulam et al., 1983; Satoh and Fibiger, 1986; Shiromani et al., 1990; Sofroniew et al., 1985; Turlejski et al., 1994; Woolf and Butcher, 1986) and project via the dorsal tegmental pathway (Shute and Lewis, 1966) or a more lateral ascending pathway (Rye et al., 1987). The tegmental projection to DLG includes both cholinergic and noncholinergic neurons (Shiromani et al., 1990; see Chapter 35 for maps of the distribution of cholinergic cell bodies in the tegmentum). In both tegmental nuclei, atriopeptide and substance P are colocalized with ChAT (Standaert et al., 1986) and about two-thirds of their constituent neurons, which innervate DLG, are located ipsilaterally and about one-third contralaterally (Shiromani et al., 1990; Turlejski et al., 1994). About 20% of tegmental neurons projecting contralaterally branch to the ipsilateral DLG (Turlejski et al., 1994), whereas about 5% of those that innervate DLG also project to the central lateral intralaminar nucleus (Shiromani et al., 1990). Another source of cholinergic input to DLG is the parabigeminal nucleus (see page 1090), which at least in the mouse (Mufson et al., 1986), tree shrew, ferret, and cat contains cells virtually all of which are immunoreactive for choline acetyltransferase (for review, see Fitzpatrick and Raczkowski, 1991). In the rat, neurons in the dorsal and ventral subnuclei contain ChAT, whereas the middle subnucleus shows reduced immunoreactivity (Tan and Harvey, 1989; Wang et al., 1988). It has been suggested that activation of the cholinergic system enhances inhibitory tuning in the DLG. Two types of nicotinic receptors are present in both relay cells and interneurons in the DLG and appear to mediate depolarization (Zhu and Uhlrich, 1997). Two types of muscarinic receptors are also present and appear to be

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associated with a decrease in an inward K current and an increase in the cation current (Zhu and Ulrich, 1998). Stimulation of the laterodorsal tegmentum facilitates P cells in DLG and weakly inhibits intrinsic interneurons (Kayama et al., 1986b); acetylcholine facilitates presumed P neurons in DLG in rats during both arousal and rapid eye movement sleep (Marks and Roffwarg, 1989). Acetylcholine in the rat DLG binds to muscarinic receptors and produces a slow depolarization due to a reduced K+ conductance (McCormick and Prince, 1987). In the presence of atropine, which blocks muscarinic receptors, acetylcholine appears to have a secondary action via nicotinic receptors (Clarke et al., 1985). Cholinergic terminals contain densely packed round vesicles and form synaptic contacts on dendrites of DLG neurons (Hallanger and Wainer, 1988). Nicotinic receptors are present in both relay cells and interneurons. Interestingly, the cholinergic input to Rt arises not only from the tegmental nuclei, but also from the telencephalic basal nucleus of Meynert (Asanuma and Porter, 1990; Hallanger et al., 1987; Levey et al., 1987b). There is strong acetylcholinesterase activity within the DLG of rat pups, especially during the second postnatal week; a transient expression appears to be induced by retinal afferents, as it is reduced dramatically by neonatal enucleations (Robertson et al., 1989). Histamine-immunoreactive fibers, although low in density, are present throughout DLG (Inagaki et al., 1988; Panula et al., 1989), but their source and functional significance are unknown. In the guinea pig, histaminergic fibers innervating DLG appear to arise from the tuberomammillary nucleus of the hypothalamus (Airaksinen and Panula, 1988). In the rat, histaminergic neurons of the hypothalamus form groups E4 and E5, which largely correspond to the dorsomedial hypothalamic nucleus (Wada et al., 1991). There are high densities of H1 and H2 histaminergic receptors in the rat’s DLG (Ruat et al., 1990). Iontophoretic application of histamine to the relay cells in DLG in the cat and guinea pig results in a slow depolarization (McCormick and Williamson, 1991). Substance P-immunoreactive profiles are present in the adult rat, but those containing Leu-enkephalin are virtually absent (Wadhwa et al., 1990). SC lesions result in loss of substance P, cholecystokinin, and vasoactive intestinal polypeptide immunoreactivity in the ipsilateral DLG (MiguelHidalgo et al., 1991; Ogawa-Meguro et al., 1992). The neurochemical nature of two small additional projections is unknown. One originates in the caudal part of the lateral hypothalamus (Turlejski et al., 1993), which receives direct retinal inputs (Kita and Oomura, 1982; Riley et al., 1981; Sousa-Pinto and Castro-Carreia, 1970). The second originates in the dorsolateral neuronal column of the periaqueductal gray (Turlejski et al., 1993).

Efferent Projections from DLG Projections from DLG, summarized in Table 1, are restricted to the ipsilateral Oc and visual Rt. After ablation of the visual cortex, the bulk of cells in DLG degenerates rapidly (Matthews, 1973) by a process resembling apoptosis (Al-Abdulla et al., 1998). The geniculocortical projection fibers, which possibly use excitatory amino acids (most likely glutamate) as transmitters (Johnson and Burkhalter, 1992; Saéz et al., 1998), terminate in layers 4, lower 3, and, to a lesser extent, in layers 6 and 1 (see page 1125). Thalamocortical axons branch and project to different parts of Rt as they pass through it (Scheibel and Scheibel, 1966). The projection from DLG is restricted to the most dorsal, topographically organized part (visual Rt; Hale et al., 1982; Itaya and van Hoesen, 1982b; Ohara and Lieberman, 1981a; Webster et al., 1981). The upper temporal visual field is mapped dorsolaterally and nasal fields more deeply (Hale et al., 1982); dorsal and rostral visual Rt relate to dorsal and rostral DLG (Ohara and Lieberman, 1985). Other nuclei (lateral posterior, posterior thalamic group) project to nonoverlapping sectors of Rt (Coleman and Mitrofanis, 1996; Pinault et al., 1995), which are visually responsive (Sumitomo et al., 1988). DLG as a Visual Relay Nucleus In DLG, the number of relay neurons has been estimated to be about 16,000–18,000 (Satorre et al., 1986; Villena et al., 1989; P.R. Martin, personal communication), representing about 78% of the neurons in the DLG (Gabbott et al., 1986b). All the fast-conducting optic axons (with Y-like properties) appear to project to the DLG (Dreher et al., 1985a; Sefton, 1968). In turn, relay cells receiving fast inputs have fast-conducting axons projecting to occipital cortical areas (Hale et al., 1979; Noda and Iwama, 1967). About 2000–4000 type I retinal ganglion cells, which are the presumed counterparts of Y-like cells (Dreher et al., 1985a; Peichl, 1989), project to DLG. Cells with Y-like properties represent 27% (Hale et al., 1979) to 48% (calculated from Fukuda et al., 1979) of all neurons recorded in DLG. Sampling bias of the electrode does not seem to be an important factor in DLG of the cat (Friedlander et al., 1981). If this assumption is valid for the rat, a single incoming Y-like optic axon is likely to diverge to innervate a minimum of about one, and possibly up to four, relay cells in DLG. Indeed, an even larger amplification is observed for the Y system in DLG of the cat (Friedlander et al., 1981). Because each 2a (presumed Y-like) axon has on average 23 terminals (Brauer et al., 1975), between 6 and 23 2a terminals would excite a fast-conducting relay cell in DLG.

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How many of the remaining optic tract axons project to DLG has not been established. However, we assume that the relative amount of the radioactive material deposited in a given retinorecipient nucleus (about 14% in the DLG) corresponds to the relative strength of the retinofugal projection to the nucleus. Virtually all ganglion cells project to SC (Linden and Perry, 1983a; Dreher et al., 1985a), with 64% of radioactive material being found there (Toga and Collins, 1981). On that basis, it appears that no more than 25,000 ganglion cells project to DLG, a figure in good agreement with our own estimates of about 20–25,000, based on the fraction of retinal ganglion cells labeled after small injections into DLG or those surviving after complete neonatal lesions of SC (Carpenter et al., 1986; Dreher et al., 1985a; Martin, 1986). Therefore, there would be about one retinal ganglion cell for each relay cell in DLG. As noted earlier, the terminals of type I cells must diverge within DLG. Conversely, there must be some convergence among the other classes, consistent with the report of Brauer et al. (1979) of overlap of large and small terminals at least in rostral DLG. In macaque monkeys, there is a good correspondence between the number of axons in the optic nerve (1.2 million; Rakic and Riley, 1983) and the number of cells in DLG (approximately 1 million; Beaulieu and Colonnier, 1983). However, in the cat, there are about four times as many relay cells in DLG as there are optic axons projecting there (Friedlander et al., 1981; Madarász et al., 1978; Wässle, 1982; Weber and Kalil, 1983).

Superior Colliculus Introduction In the rat, virtually all ganglion cells project to the superior (anterior) colliculus (SC) a mammalian homologue of optic tectum of other vertebrates (Linden and Perry, 1983a; Dreher et al., 1985a). Most optic axons arrive prenatally, but axons of late-born ganglion cells are still growing into the rat SC in the first few days after birth (Dallimore et al., 2002). The horizontally laminated organization of SC is apparent in Atlas plates 40–50. The layers, from the surface, are zonal or stratum zonale; superficial gray or stratum griseum superficiale; optic or stratum opticum; intermediate gray or stratum griseum intermediale; intermediate white or stratum album intermediale; deep gray or stratum griseum profundum; and deep white or stratum album profundum; (Huber and Crosby, 1943). The superficial gray and upper optic strata are innervated by retinal axons and thus process visual information. Cells in these layers in turn project to intermediate and deep layers, which also receive inputs from other sensory systems—audi-

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tory and somatosensory (including nociceptive). This organization is apparently a general feature of all mammals (Harting et al., 1973; Stein, 1981) and, indeed, of all vertebrates (Stein, 1981; Stein and Gaither, 1983). In several mammals, including nocturnal murid rodents, such as the rat (Beckstead and Frankfurter, 1983), however, a direct retinal projection also terminates in the intermediate gray layer.. It has been claimed that in rodents, the SC is involved in the spatial localization of a biologically significant stimulus rather than its recognition (“where is it” rather than “what is it?”) (Schneider, 1969). The position of the stimulus may be just as important as the modality of the sensory cue, and different types of inputs can influence head/eye movements and guidance toward or away from a stimulus. To summarize, in the rat the functional roles of the SC include orientation to visual stimuli (Goodale et al., 1978; Sahibzada et al., 1986); influence on navigation and spatially guided movements (Cooper et al., 1998); the direction and speed of eye movements (McHaffie and Stein, 1982; Okada, 1992); avoidance, defensive, or escape reactions (Olds and Olds, 1962; Redgrave et al., 1981; King, 1999), perhaps reflecting the representation of the upper visual field (Sahibzada et al., 1986; Westby et al., 1990); sensorimotor gating (Fendt et al., 1994; Fendt, 1999; Meloni and Davis, 2000); turning (DiChiara and Morelli, 1982; Hebb and Robertson, 1999); and locomotor exploration (Dean et al., 1982), including approach and orienting behaviors (Dean et al., 1986; Westby et al., 1990). Damage to deeper layers leads to sensory neglect (Kirvel et al., 1974). The SC also appears to mediate the effects of light on sleep/wakefulness (Miller et al., 1998) and may be involved in circuits that control circadian rhythms (Moore et al., 2000). Evidence shows that uncrossed (avoidance and defense) and crossed (orienting and approach) output pathways from SC arise from distinct and segregated groups of cells (Dean et al., 1986, 1989; Redgrave et al., 1987a, 1987b; Sahibzada et al., 1986; Westby et al., 1990). The SC appears to be involved in the initiation of defensive or escape responses such as micturition and defecation behaviors (Vargas et al., 2000). The SC also exerts an influence on the control of blood pressure and peripheral blood flow (D’Amico et al., 1996, 1998a, 1998b; Merabet et al., 1997). Cells in intermediate layers of rat SC project contralaterally via the predorsal bundle to initiate shifts in gaze, intrinsic to orienting reflexes (Bickford and Hall, 1989; Redgrave et al., 1986). Interestingly, many nociceptive neurons in the intermediate SC layers project axons contralaterally; behavioral studies support the idea that these projections may be involved in orientation toward a source of discomfort to locate and remove the noxious stimulus (Redgrave et al., 1996a, 1996b; Telford et al., 1996; Wang et al., 2000). Complex circuitries linking

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the multimodal intermediate and deep tectal layers with cerebellar nuclei, substantia nigra, and basal ganglia enable the SC to participate in the selection of appropriate (and suppression of competing or inappropriate) motor programs in response to novel stimuli (NiemiJunkola and Westby, 1998, 2000). Neurons in the deep SC layers are also involved in circuitries linked to audiogenic seizures (Fairgold and Randall, 1999). Lesions in SC produce visual and other sensory neglect (Krauthamer et al., 1992) and deficits in visual search (Heywood and Cowey, 1987). Dean and Redgrave (1984) suggested that visual neglect could be explained by a deficit of a specific response to visual stimuli, an attentional deficit, and a more general lack of visual responsiveness; for example, neglect of stimuli in the far peripheral visual field following collicular lesions may contribute to visual attentional deficits (Overton et al., 1985). SC does not, however, appear to play a significant role in brightness or pattern discrimination (Dean, 1978) or optokinetic nystagmus (Simpson, 1984). Structure Cells Cells of the rat’s SC were described in detail by Cajal (1911), and there is no difficulty in identifying many of the cells he illustrated in Golgi-stained material. More recently, neurons in the upper layers, based on the orientation of their dendrites, have been differentiated as horizontal, vertical, and stellate (Labriola and Laemle, 1977; Langer and Lund, 1974; Tokunaga and

Otani, 1976). Several types of cells appear to be interneurons, with their axons confined to the upper layers. These include horizontal and marginal cells (Fig. 5), lying parallel to the surface in the narrow, poorly differentiated zonal layer. Both horizontal and marginal cells stain strongly for nicotinamide phosphate (NADPH)diaphorase, a synthetic enzyme for the putative cotransmitter nitric oxide (González-Hernandez et al., 1982). These cell types can also contain substance P (Behan et al., 1993). In addition, some stellate cells seem to be interneurons. Consistent with an intrinsic inhibitory role, horizontal, piriform, and small stellate-like cells in the superficial SC have been identified using GABA immunohistochemistry (Mize, 1992). The superficial layers contain a particularly high density of cells expressing GAD mRNA (Harvey et al., 2001). Vertically oriented cells in the superficial gray and optic layers, which appear to be relay cells projecting to deeper layers of SC, include wide-field vertical cells (Isa et al., 1998), piriform cells (Fig. 5), located at the junction between the zonal layer and superficial gray, and some narrowfield cells. These latter cells (Fig. 5) usually have two ascending and two descending primary dendrites; two subtypes, vertical fusiform and pyramidal cells, have been distinguished. Pyramidal cells do not contain NADPH-diaphorase (González-Hernandez et al., 1982). Superficial to deep projecting neurons are often found rostromedially in the rat SC (Hilbig, 1994; Hilbig et al., 2000), and many of the connections are excitatory and

FIGURE 5 Summary diagram from Langer and Lund (1974). Typical examples of Golgi-impregnated neurons in the upper (retino-recipient) layers of SC. A, axon. Marginal (M) and horizontal (H) cells, with horizontally oriented dendrites, have axons that ramify locally. Narrow-field vertical (V), wide-field (W), and pyriform (P) cells have dendrites projecting vertically and axons that usually project to deeper layers but may also ramify in the vicinity of the dendrites.

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appear to use glutamate as the neurotransmitter (Lee et al., 1997; Isa et al., 1998). Note that studies in hamster indicate that neurons in deeper tectal layers can also access information entering the visual layers of the SC via ascending superficially located dendrites (Mooney et al., 1994). Some vertically oriented fusiform cells in the superficial gray are weakly immunoreactive for Met-enkephalin-Arg-Gly-leucine; after enucleation, a different population of small cells expresses the peptide (Okamoto et al., 1990). Wide-field vertical cells (Fig. 5), with up to seven primary dendrites, lie at the junction of the superficial gray and optic layers (Labriola and Laemle, 1977; Langer and Lund, 1974). Narrow- and wide-field cells appear to project to other visual nuclei (DLG, VLG, and LP; Mason and Groos, 1981; MackaySim et al., 1983; Okoyama and Kudo, 1997), and those projecting to DLG may contain substance P, cholecystokyinin (CCK), or VIP (Miguel-Hidalgo et al., 1991; Ogawa-Meguro et al., 1992). Medium-sized neurons in superficial gray and large neurons ventral to the optic layer are strongly reactive for calbindin D-28k (SchmidtKastner et al., 1992). Analysis of neuropeptide mRNA expression in rat SC has revealed a neurochemical parcellation of the superficial layers into three distinct sublaminae (Harvey et al., 2001). Cells expressing preprotachykinin (PPT, substance P) are mostly found in the zonal and in the upper two-thirds of the superficial gray layers. At least some of these cells appear to project to the ipsilateral parabigeminal nucleus (Bennett-Clarke et al., 1989). Seventy-five percent of CCK mRNA expressing cells are located in the upper SC layers, forming a prominent band in the middle third of the superficial gray, whereas somatostatin-expressing neurons form a dense band in its lower third, extending into the upper part of the optic layer. Cells expressing preproenkephalin (PENK) mRNA are also found in lower superficial gray and upper optic layers. In intermediate and deep layers of SC, multipolar cells (including stellate cells) may have axons descending to the deep white layer. Vertical cells have one or two oblique primary dendrites and an axon that either enters intermediate white or, less commonly, ascends for some distance (Tokanuga and Otani, 1976). Only small cells in the intermediate and deep layers stain positively for NADPH-diaphorase (González-Hernandez et al., 1982). Cells expressing mRNAs for PPT, somatostatin, or PENK are distributed differentially in the intermediate and deep layers (Harvey et al., 2001). PPT mRNA-expressing cells are especially prominent caudomedially, and cells expressing somatostatin mRNA can often be seen to form two tiers of label in the intermediate gray and white layers. A cluster of PENK mRNA-expressing neurons is located medially in the deep tectal layers.

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Regional differences in the distribution of other histochemical and immunological stains and markers are also apparent (Illing, 1996). Thus, acetylcholinesterase activity is high in the superficial gray and an extensive network of cholinergic fibers is present in the intermediate gray (Tan and Harvey, 1989) in which there is a clustered or patchy distribution of choline acetyltransferase and acetylcholinesterase activity (Harvey and MacDonald, 1985; Tan and Harvey, 1989; Schnurr et al., 1992; Illing, 1996). No cells in rat SC display immunoreactivity or mRNA for choline acetyltransferase (Tan and Harvey, 1989; Schnurr et al., 1992; Harvey et al., 2001). As in the mouse, one significant source of cholinergic projections to the SC (Illing et al., 1990) is likely to be the parabigeminal nucleus (Mufson et al., 1986), although in the rat perhaps mainly from the dorsal and ventral subnuclei, which project to the ipsilateral tectum (Tan and Harvey, 1989; Wang et al., 1988). Application of anticholinergic agents has been reported to increase 2-deoxyglucose activity, mediated by nicotinic receptors (Pazdernik et al., 1982). More recent data have shown that activation of nicotinic receptors, perhaps located presynaptically on retinal terminals (Prusky and Cynader, 1988), depresses visual responses in the superficial SC and responses can be potentiated with nicotinic antagonists (Binns, 1999; Binns and Salt, 2000; Lee et al., 2001). While patches of acetylcholinesterase are not aligned with regions of high oxidative metabolism (Wallace, 1986a, 1986b; Wiener, 1986), they do correlate with patches of NADPH-diaphorase activity in the intermediate and deep layers. The latter distribution reflects afferents arising in tegmental nuclei PPTg and LDTg (Wallace, 1986b) but not other afferents (Wallace and Fredens, 1989), although, as noted earlier, NADPH-diaphorase activity is concentrated in zonal and superficial gray layers (González-Hernandez et al., 1982). Parvalbumin-positive neurons are rarely found in the zonal layer (13–16% of all parvalbumin-positive neurons in the retinorecipient layers; Barker and Dreher, 1998) but are quite numerous throughout the superficial gray forming complementary sublaminae with calbindin D-28kD (Barker and Dreher, 1998; Cork et al., 1998); vertically oriented parvalbumin-positive cells are also seen in the optic layer (Barker and Dreher, 1998) with a periodicity complementary to that of acetylcholinesterase (Barker and Dreher, 1998; Illing et al., 1990; Schmidt-Kastner et al., 1992). Parvalbumin-positive neurons are present throughout the mediolateral and rostrocaudal extent of the superficial gray with peak density in the region where area centralis is represented (Barker and Dreher, 1998). Almost all morphological classes of collicular cells distinguished in Golgi studies (Labriola and Laemle, 1977; Langer and Lund, 1974;

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Tokunaga and Otani, 1976) express parvalbumin (Barker and Dreher, 1998). Only about one in six parvalbuminimmunoreactive neurons send axons to the DLG and LP (Lane et al., 1993, 1997). Calbindin D-28kD-positive neurons and plexi are located predominantly in the deep part of the superficial gray and in the ventral part of the optic layer (Cello, 1990; Lane et al., 1993, 1997; Dreher et al., 1996) although some (10–15%) calbindin D-28kD-expressing cells are present in the zonal region (Dreher et al., 1996). Again, although these latter neurons are present throughout the mediolateral and rostrocaudal extent of superficial gray, there is a density gradient with its peak in the region where area centralis is represented (Dreher et al., 1996). Several morphological classes of the SC neurons express calbindin D28kD (Dreher et al., 1996). These include “marginal cells” of Langer and Lund (1974) and Labriola and Laemle (1977) “narrow-field vertical cells” of Langer and Lund (1974; cf. also Labriola and Laemle, 1977); “cylindrical type” of Tokunaga and Otani (1976); “type III ganglion cells” of Langer and Lund (1974) and “wide-field vertical cells of Labriola and Laemle (1977); “stellate cells” of Langer and Lund (1974; cf. also Labriola and Laemle, 1977). About 90% of calbindin D-28kD-positive neurons in the superficial gray and optic layers project to the DLG and LP, respectively (Lane et al., 1993, 1997). Some rat SC neurons that project to LP contain the enzyme adenosine deaminase (Miguel-Hidalgo et al., 1989). In intermediate layers, patches of acetylcholinesterase, cytochrome oxidase, and parvalbumin are aligned medially but not laterally, suggesting that the latter do not reflect the pattern of afferents (Illing et al., 1990; Schmidt-Kastner et al., 1992). More recent studies have shown that the distributions of enkephalin and substance P, as well as choline acetyltransferase and parvalbumin, are spatially related to the network of acetylcholinesterase-positive patches in the intermediate layers (Illing, 1996). This lattice-like organization has now been shown to be aligned with a number of afferent and efferent pathways (Mana and Chevalier, 2001). There is also a dense network of serotonergic fibers throughout the SC (see Chapter 34), densest in the deeper part of the superficial gray and upper optic layers, forming fine varicosities (Harvey and MacDonald, 1987). The superficial gray contains a high density of serotonin receptors; 5-HT 1B receptors in particular appear to be mostly located on retinal terminals, thus serotonergic inputs can modulate retinotectal signaling (Boulenguez et al., 1996; Mooney et al., 1996; Sari et al., 1999). In deeper layers, the serotonergic fibers are mostly oriented vertically to the superficial layers (Ueda et al., 1985). Fibers originate from the ipsilateral and, to a lesser extent, contralateral dorsal raphe nuclei (Villar et al., 1988).

Axons and synaptic relations Axons of retinal origin enter SC through the optic layer. They turn vertically and terminate most frequently in the upper part of superficial gray, sometimes synapsing en passant with vertical cells. Ultrastructural studies reveal that small optic terminals, located superficially, contain round synaptic vesicles and make Gray type I (asymmetric) excitatory synapses. Retinocollicular axons fall into three conduction velocity groups (Fukuda, 1977; Sefton, 1968) and their terminals are segregated within the superficial layers. The slower-conducting axons (presumed W-like) project to upper parts of the superficial gray, whereas axons with faster conduction velocities (presumed Y-like), which degenerate more rapidly after eye removal, terminate more deeply in superficial gray and optic layers (Lund et al., 1976; Sefton, 1969). Most retinal ganglion cells are glutamatergic (Binns, 1999) and activate tectal neurons via both NMDA and nonNMDA (AMPA/kainate) ionotropic receptors (Binns, 1999; Lo and Mize, 1999; Kondo et al., 2000; Petralia et al., 1994). AMPA receptor subtypes on putative GABAergic neurons may be involved in remote inhibitory interactions within the superficial SC (Endo and Isa, 2001). The rat SC also contains different types of metabotropic glutamate receptors that influence visual responses in superficial layers (Binns, 1999; Cirone and Salt, 2000; Cirone et al., 2002a; Hudtloff and Thomsen, 1998). Some of these receptors may be associated with retinal axons (Cirone et al., 2002b) or possibly cortical afferents and are likely to be involved in the presynaptic inhibition of glutamate release in superficial gray (Cirone et al., 2002a). A dipeptide related to Nacetylaspartyl-glutamate (NAAG) may also be a neuromodulator of retinocollicular interactions, perhaps acting through metabotropic glutamate receptors (Binns, 1999) (see page 1085). NAAG-reactive terminals and neuropil are present in SC, and following section of the optic nerve, a decrease of 50–60% in levels of NAAG in the contralateral SC is observed (Moffett et al., 1991). Corticotectal axons arise from large pyramidal cells located in the upper two-thirds of layer 5 of topographically related regions of the visual cortex (Oc; Bolz et al., 1991; Bourassa and Deschênes, 1995; Hallman et al., 1988; Hübener and Bolz, 1988; Olavarria and Van Sluyters, 1982; Schofield et al., 1987; Sefton et al., 1981; Thong and Dreher, 1986, 1987). Their terminals contain round vesicles, make Gray type 1 synapses (Lund, 1969; Lund and Lund, 1971a, 1971b), and most appear to be glutamatergic (Binns, 1999; Ortega et al., 1995). They branch to innervate LP, LD, and VLG (Bourassa and Deschênes, 1995). Corticotectal axons reach SC on the third postnatal day (24th postconceptional day) and even at this early stage, all cells of origin are pyramidal cells located in layer 5 (Hübener and Bolz, 1988; Thong

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and Dreher, 1986, 1987). Axons arising in different visuotopically organized cortical areas terminate at different depths within SC: Oc (area 17) deeply in superficial gray and optic layers; Oc2L (especially LM, LI and LL) in the intermediate gray as a pattern of periodic “puffs,” deep in the optic and less densely in the superficial gray layers; and Oc2M in two tiers, one in the intermediate gray and one between the deep white and gray (Harvey and Worthington, 1990). The intermediate layers also receive inputs from the frontal eye fields (Guandalini, 2001; Tsumori et al., 1997), pupillary constriction area (Guandalini, 2003), and retrosplenial cortex (Garcia del Cano et al., 2000). Most of the terminals in the upper layers of SC in normal adult rats contain round vesicles (Lund, 1969) and the majority of them, particularly in the upper superficial gray, arise in the retina and synapse on dendrites or their spines. Round terminals (probably retinal) are occasionally observed in serial arrays, being presynaptic to pleomorphic terminals, which in turn are presynaptic to conventional dendrites. Terminals of corticotectal axons do not take part in serial arrays (Lund and Lund, 1971a). As yet, the ultrastructural characteristics of the different classes observed in Golgi material have not been identified. GABA-immunoreactive terminals generally make Gray type II synapses with both GABA-positive and GABA-negative dendrites and are the presynaptic elements in dendrodendritic synapses, implying that some arise from interneurons. Some GABA-containing and retinal terminals converge on to the same dendrite (Pinard et al., 1991), and evidence shows that retinotectal and corticotectal terminals make direct contact with GABAergic interneurons superficially (Hirai and Okada, 1993). Deep to the optic layer, the reticular substantia nigra is one source of myelinated axons with terminals containing pleomorphic vesicles, making Gray type II synapses with dendritic shafts and they appear to be GABAergic (Vincent et al., 1978). Many different cell types in the rat superficial SC are responsive to GABA (Edwards et al., 2002), and many types of GABAA, GABAB, and GABAC receptors are found in the rat SC, some at particularly high density in superficial gray (Binns, 1999; Fritschy and Mohler, 1995; Pinard et al., 1991; Pirker et al., 2000; Wegelius et al., 1998). Evidence shows that different types of GABA receptors are involved in different inhibitory processes; thus GABAA receptors on horizontal cells mediate surround inhibition and spatial sharpening, whereas GABAB receptors, most likely on piriform and stellate cells, are involved in response habituation (Binns, 1999). These receptors also appear to be involved in the inhibitory modulation of collicular projection neurons (Lee et al., 2001). Activation of GABAC receptors on superficial gray interneurons

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may have a disinhibitory role (Pasternack et al., 1999), increasing tectal output levels to the thalamus and brain stem (Lee et al., 2001; Schmidt et al., 2001). There is increasing evidence for cholinergic modulation of at least some of these efferent pathways (Lee et al., 2001). Retinotopic Organization In the rat, as in all vertebrates, axons arising in the contralateral eye provide the principal input to the upper players of SC (Stein and Gaither, 1983; Toga and Collins, 1981). As seen in Fig. 2, the projection is organized topographically in a manner very similar to that for all vertebrate species studied so far (Lashley, 1934; Siminoff et al., 1966). In the rat, unlike some other mammalian species, however, representation of the central visual field is not substantially magnified in comparison with the peripheral field (Siminoff et al., 1966; Stein, 1981; Toga and Collins, 1981). Retinal terminals are in register with corticotectal terminals and activate cells in superficial gray and optic layers (Lund, 1972; Takeuchi et al., 1982). In normal adult rats, the ipsilateral retinal projection is restricted to rostral SC (Hayhow et al., 1962; Martin et al., 1983), within which monocular and binocular neurons with receptive fields on either side of the vertical meridian have been recorded (Diao et al., 1983b). Extensive developmental studies have revealed many of the mechanisms that underlie the formation of these ordered retinotectal maps in rat and other mammals (e.g., see O’Leary and Wilkinson, 1998). Physiology Cells with relatively small receptive fields lying dorsally in the superficial gray have been correlated with the morphological class of “narrow-field vertical cells” distinguished in Golgi studies (Labriola and Laemle, 1977; Langer and Lund, 1974) and further subdivided into three subtypes. Cells that are located more deeply, in superficial gray and optic layers, have larger receptive fields. They receive a convergent input, perhaps from both cortex and retina (Humphrey, 1968), and have been correlated with the more deeply located morphological class of wide-field vertical neurons (Fukuda and Iwama, 1978; Lo et al., 1998). Collicular cells are characterized by a high-contrast sensitivity and narrow dynamic range (Gonzalez et al., 1991) and some (9%) are orientation selective (Gonzalez et al., 1992). The receptive field sizes of cells in the upper layers in another rodent, the hamster, like those of retinal ganglion cells, do not increase with retinal eccentricity (Chalupa and Rhoades, 1977; Tiao and Blakemore, 1976). The majority of collicular cells in all murid rodents respond optimally to relatively slow motion, and few units can be activated through either eye (for review, see Stein, 1981).

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Several receptive field types such as on, off, on–off and motion-sensitive types have been distinguished on the basis of the responsiveness (or lack thereof) of a cell to stationary flashing spots or bars presented in the receptive fields (Fortin et al., 1999; Fukuda and Iwama, 1978; Gonzalez et al., 1992). On-type receptive fields are not present in young pups (postnatal days 13–14) before the time of natural eye opening (Fortin et al., 1999). Relatively few direction-selective cells have been recorded in rat (10%, Fortin et al., 1999; 14%, Fukuda and Iwama, 1978; 16%, Gonzalez et al., 1992) with the majority lying deeply (Gonzalez et al., 1992). Two-thirds of these respond to upward movement of the stimulus (Fortin et al., 1999; Fukuda and Iwama, 1978; Gonzalez et al., 1992), a characteristic also noted in cells of the SC of other nocturnal rodents such as mouse (Dräger and Hubel, 1975) and hamster (Chalupa and Rhoades, 1977; Tiao and Blakemore, 1976). In addition to direction-selective cells, about a quarter of collicular neurons in the rat are directionally biased, again preferring usually upward movement (Fortin et al., 1999). In the hamster, decortication reduces the number of directionally selective cells in the upper layers of the ipsilateral SC (Chalupa and Rhoades, 1977). Direction-selective and directionally biased neurons are present in the SC of the rat before the time of natural eye opening and before corticotectal inputs are functionally effective (Fortin et al., 1999). Thus the role of corticotectal projections in direction selectivity of collicular neurons in this species is less clear. However, unilateral ablation of the visual cortex produces a reduction in the magnitude of the light-evoked potential in the ipsilateral SC (Goodale, 1973). Apart from the aforementioned paucity of on-type collicular neurons before eye opening, the main developmental change in properties of collicular neurons is a dramatic reduction in the latency of the responses to bright, full-field flash stimuli. Thus, according to Fortin and collaborators (1999), the mean latency of collicular neurons recorded from the superficial gray layer at postnatal days 13/14 is 60 ms (± 5.7 ms), whereas that of adult animals is 21.4 ms (+ 0.8 ms). The reduction in latency is most likely due to the onset of myelination in the retinocollicular pathway at around postnatal day 15 (Warton and Jones, 1985). Many superficial tectal cells receive convergent retinal and visual cortical (area 17) input (Binns, 1999). At least in cat, NMDA receptors are found opposite both retinotectal and corticotectal terminals (Mize and Butler, 2000), and it has been proposed that activation of these receptors is important in the integration and temporal matching of primary sensory and cortical inputs (Binns, 1999). The corticotectal input is functionally dependent on NMDA receptor activation. In young rats, the establishment of excitatory corticotectal inputs appears to be

important in the maturation of intrinsic GABAergic networks after eye opening (Aamodt et al., 2000). Development of LTP effects involving both glutamatergic and GABAergic transmission also occurs during this period (White and Platt, 2000). Later on, cortical afferents may have conditioning or instructional influences on collicular activity in the superficial layers as well as in deeper multimodal laminae, perhaps controlling experience-dependent changes in tectally mediated sensorimotor behaviors (Wallace and Stein, 2000). Inhibition in the SC of the rat appears to be different from that in DLG; in particular the postexcitatory inhibition is weaker (Fukuda et al., 1978; Sefton, 1969). Rt does not project there (Taylor et al., 1986). Inhibitory regions within receptive fields of cells in the superficial layers are superimposed spatially on the excitatory regions, thus the optimal size of a stimulus is smaller than the size of the discharge center (for review, see Stein, 1981) and some cells are end stopped (Gonzalez et al., 1992). Long-term potentiation in the superficial layers of SC can be revealed in rats only after removal of the visual cortex (Shibata et al., 1990), suggesting that cortical inputs may tonically inhibit long-term potentiation in upper collicular layers, probably via GABAergic interneurons (Hirai and Okada, 1993). Cells in deeper laminae of SC can be activated transynaptically (via cells in upper layers) by stimulating the optic chiasm, optic tract, or occipital cortex (Takemoto et al., 1978; Isa et al., 1998), although some retinofugal axons terminate directly in the intermediate gray (Beckstead and Frankfurter, 1983). Electrical stimulation of the optic tract evokes monosynaptic excitatory postsynaptic potentials (EPSPs) in the superficial gray and optic layers and disynaptic EPSPs in the intermediate layers (Isa et al., 1998). Projection neurons in the optic layer respond transiently to retinal input (Lo et al., 1998), but a proportion of neurons in the superficial gray appears to generate prolonged bursts of excitatory postsynaptic currents in cells in the intermediate gray (Ozen et al., 2000). Excitatory superficial to deep projections are mediated by AMPA and NMDAtype glutamate receptors (Isa et al., 1998). Neurons in the intermediate gray exhibit a range of electrophysiological properties, although wide-field vertical cells generally express time-dependent delayed rectification (Saito and Isa, 1998). Inhibition of GABA using bicucculine enhances excitatory synaptic potentials and bursting activity in the intermediate gray layer, suggesting that this intratectal pathway is normally suppressed by GABAergic systems (Isa et al., 1998); GABA is implicated in various behaviors (Adachi et al., 2003; Dean and Redgrave, 1992; Saito and Isa, 2003). In the rat, as in many other mammals, cells deep to the optic layer can be activated by visual or somatosensory

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or, less frequently, auditory stimuli, whereas some respond to two or three modalities (Vidyasagar, 1978). Auditory-responsive cells in the intermediate and deep layers exhibit spatial tuning to sound, a property that enables the encoding of auditory space in the rat SC (Gaese and Johnen, 2000). In other rodents, visually responsive cells with large receptive fields are recorded particularly in the intermediate gray, whereas multimodal, auditory, and somatosensory cells are found more deeply. Topographically organized maps of the different sensory systems are in register (Chalupa and Rhoades, 1977; Dräger and Hubel, 1976; Stein, 1981). During development, alignment of the retinotopic map with other sensory modalities in deeper laminae requires visual signals relayed via the superficial SC (King et al., 1998). Habituation is a common feature of cells in deep collicular layers of rodents (Chalupa and Rhoades, 1977; Dräger and Hubel, 1975; Tiao and Blakemore, 1976). It is generated by intrinsic tectal circuitries involving NDMA and non-NMDA receptors, as well as by feedforward GABAergic inhibition mediated by GABAB receptors (Binns, 1999). Electrical stimulation of SC in the rat produces saccadic eye movements, which are roughly horizontal. Although both direction and magnitude vary with the location of the stimulating electrode across SC, the direction remains constant at different depths; saccades generated rostrally in SC (where nasal visual field is represented) are smaller. Medially (upper visual field), stimulation generates contralateral saccades with an upward component; laterally (lower visual field), saccades with a downward component. Stimulation in, and deep to, the intermediate gray is most effective in eliciting movements, not only of eyes, but also of pinnae and vibrissae (McHaffie and Stein, 1982). Intercollicular connections are prominent; some terminate in the rostral half of the superficial layers and the caudal half of the intermediate and deep layers in the rostral half of the SC (Jen and Au, 1986; Hilbig, 1994; Hilbig et al., 2000; Yamasaki et al., 1984). Intercollicular fibers contain GABA (Binns, 1999) and appear to exert a suppressive effect on visual responses of contralateral collicular neurons (Goodale, 1973). Removal of the SC contralateral to the ablated visual cortex (or sectioning of the tectal commissure) results in amelioration of visual (as well as other sensory) neglect in the visual field contralateral to the ablated visual cortex (Kirval et al., 1974). Connections The afferent and efferent connections of superficial and intermediate/deep layers are summarized separately in Table 2, and the sources of chemical pathways are illustrated in Paxinos et al. (1999a, 1999b). The list

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is undoubtedly incomplete. Neurons in the superficial layers send axons to a variety of visually related nuclei in the midbrain and diencephalon; it has been estimated that about two-thirds of neurons in the superficial SC give rise to ascending projections (Hilbig et al., 2000). The projection to the DLG arises from cells in the ventral twothirds of superficial gray (Lane et al., 1997; Mackay-Sim et al., 1983; Sugita et al., 1983; Ogawa-Meguro et al., 1992). The projection to the anterior pretectal nucleus also originates from superficial gray. The tectal projection to the parabigeminal nucleus originates in superficial gray, but mostly from rostral SC. Neurons efferent to the ventral lateral geniculate tend to be located more ventrally, in lower superficial gray and upper optic layer (MackaySim et al., 1983). Projections to the lateral posterior nucleus mostly arise from cells in the optic layer (Lane et al., 1993, 1997; Miguel-Hidalgo et al., 1989; Sugita et al., 1983) as do the projections to suprageniculate (Linke, 1999; Linke et al., 1999). Stratification in the efferent tectal outflow to DLG and LP is mirrored by the depth segregation of retinal afferents (Fukuda, 1977) and may indicate parallel processing of W-like and Y-like streams through the midbrain and thalamus and thence to different visual cortical regions. Two major descending output pathways arise from largely independent populations of cells in SC. One, in deeper layers, sends a descending projection to the ipsilateral spinal cord and also innervates targets medially in the pons and medulla. It has been suggested that the uncrossed descending pathway to the cuneiform nucleus may contain substance P, whereas some projections to the pons may be enkephalinergic (Harvey et al., 2001). The other pathway (the predorsal bundle) arises mainly from neurons in lateral intermediate gray and white laminae (Bickford and Hall, 1989; Dean et al., 1989; Sahabzida et al., 1987) and innervates predominantly contralateral gaze centers in the midbrain and pons concerned with orienting reactions (Redgrave et al., 1986, 1987b), as well as the inferior olive, reticular formation, and cervical regions of the spinal cord (Dean et al., 1989; Herbert et al., 1997; Murray and Coulter, 1982; Sahabzida et al., 1987; Yasui et al., 1995). Predorsal cells, restricted to intermediate layers, receive inputs from GABAergic fibers arising in substantia nigra (Bickford and Hall, 1992). A monosynaptic GABAergic input from the ventral part of the zona incerta in the ventral thalamus to the cells of origin of the predorsal bundle in intermediate gray has been described (Kim et al., 1992; Romanowski et al., 1985; Watanabe and Kawana, 1982). Because the same ventral region of zona incerta also receives a reciprocal input from SC (Kim et al., 1992; Roger and Cadusseau, 1985; Shammah-Lagnado et al., 1985), it may contribute to the generation of orienting movements

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TABLE 2 Upper layersa Afferents Retina(c,i) Oc1(i)

to SuG,ZS to Op Oc2L(i) to SuG,Op Area LM to SuG,Op Oc2M from layer 5 SC(c) SuG, Op VLGMC(i) to Op,SuG IGL Pretectum(i) APT OP(i) OT(c,i) PPT(c,i) PBG(c,i) PBG(c) PBG(i) LP(c,i) BIN DR LDTg PCom PPTg PrH(c) Efferents SC SuG, Op (c) Dp and In (i) Dp and In (c) DLG(i) from narrow-field vertical cells in lower two-thirds LP(c,i) (from SO)

VLG(i) from wide-field cells in SuG and Op IGL APT OPT, OT PPT, from SuG PBG(i) PBG(c,i) Hab Pcom SG ZI

Connections of Superior Colliculus

Cajal, 1911; Huber and Crosby, 1943; Nauta and van Straaten, 1947; Lund and Lund, 1971a; Ahmed et al., 1998 Huber and Crosby, 1943; Nauta and Bucher, 1954; Goodman and Horel, 1966; Lund, 1966; Hopkins and Niessen, 1976; Jen et al., 1978; Olavarria and Van Sluyters, 1982; Bourassa and Deschênes, 1995 Lund and Lund, 1971a; Takahashi, 1985; Harvey and Worthington, 1990 Takahashi, 1985; Harvey and Worthington, 1990 Takahashi, 1985; Harvey and Worthington, 1990 Harvey and Worthington, 1990 Mason and Groos, 1981; Sefton et al., 1981; Olavarria and Van Sluyters, 1982 Sahibzada et al., 1987; Bickford and Hall, 1989 Hilbig et al., 2000 Brauer and Schober, 1982; Taylor et al., 1986 Graybiel, 1974; Swanson et al., 1974; Ribak and Peters, 1975; Legg, 1979a Taylor et al., 1986; Moore et al., 2000 Huber and Crosby, 1943 Foster et al 1989; Cadusseau and Roger, 1991 Taylor et al., 1986 Taylor et al., 1986 Taylor et al., 1986 Watanabe and Kawana, 1979; Stevenson and Lund, 1982a, 1982b; Taylor et al., 1986 Pasquier and Tramezzani, 1979; Linden and Perry, 1983b; Sefton and Martin, 1984 Hartiing et al., 1991b; Turlejski et al., 1993 Taylor et al., 1986 Lee et al., 1989 Villar et al., 1988; Harvey and MacDonald, 1987 Wallace and Fredens, 1989 Taylor et al., 1986 Wallace and Fredens, 1989; Krauthamer et al., 1995 Ohtsuki et al., 1992 Taylor et al., 1986; Hilbig et al., 2000 Cajal, 1911; Rhoades et al., 1989b; Hilbig and Schierwagen, 1994; Isa et al., 1998; Hilbig et al., 2000 Sahibzada et al., 1987; Bickford and Hall, 1989; Pasquier and Villar, 1982 Perry, 1980; Pasquier and Villar, 1982; Sugita et al., 1983; Reese, 1984; Taylor et al., 1986; OgawaMeguro et al., 1992; Reese, 1988; Lane et al., 1993,1997; Hilbig et al., 2000 SuG Mason and Groos, 1981; Mackay-Sim et al., 1983 Perry, 1980; Mason and Groos, 1981; Schober, 1981; Donnelly et al., 1983; Sugita et al., 1983; Takahashi, 1985; Taylor et al., 1986; Miguel-Hidalgo et al., 1989; Lane et al., 1993,1997; Linke et al., 1999 Pasquier and Villar, 1982; Takahashi, 1985; Taylor et al., 1986; Linke et al., 1999 Mackay-Sim et al., 1983 Taylor et al., 1986 Taylor et al., 1986, Cadusseau and Roger, 1991 Takahashi, 1985; Taylor et al., 1986 Takahashi, 1985; Taylor et al., 1986 Martin and Sefton, 1981; Takahashi, 1985 Stevenson and Lund, 1982a,b; Linden and Perry, 1983b; Sefton and Martin, 1984; BennettClarke et al., 1989; Hilbig et al., 2000 Taylor et al., 1986 Taylor et al., 1986 Taylor et al., 1986 Linke, 1999; Linke et al., 1999 Taylor et al., 1986

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TABLE 2 Intermediate and deep layersa Afferents SC(i) SuG and Op SC(c) InG, DpG; Opb Retina(c) Oc2M Cortical area 4 Cortical area 29c,d RSA, RSG Somatosensory cortex Orofacial motor cortex(i) Vibrissal motor cortex AGm – frontal eye fields(i>c) VLGMC to InG and InW Caudal LP APT (i) OPT InG(c,i) OT Deep cerebellar n(c) DR LC LC(norepinephrinergic) Me5(c,i) Medullary dorsal horn (InG)(c) Parabrachial RF PCom PF Pr5(c,i) PrH(c) Rostral ventral resp gp (InW,DpG)(c,i) SG SN SN(c) GABAergic GABAergic to *DpG SNR Sp5O,I Sp5I(c) Sp5C Spinal cord – all segments(c) to InW; DpW and DpG VLTg ZI Efferents VLGPC LD APT(Anterior) CL(c,i) CnF(i) Hypothalamus IO(c) Lamina V, VII(c>i) Lamina VIII

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Connections of Superior Colliculus—cont’d

Cajal, 1911; Hilbig and Schierwagen 1994; Isa et al., 1998; Rhoades et al., 1989a; Linke et al., 1999; Hilbig et al., 2000 Hilbig et al., 2000 Fish et al., 1982; Rhoades and Fish, 1982 Beckstead and Frankfurter, 1983 Benzinger and Massopust, 1983; Takahashi, 1985; Cadusseau and Roger, 1985; Redgrave et al., 1987a Lee et al., 1989 Benzinger and Massopust, 1983 Garcia Del Cano et al., 2000 Veinante et al., 2000 Tsumori et al., 1997 Miyashita and Mori 1995 Tsumori et al., 2001; Guandalini, 2001 Brauer and Schober, 1982 Graybiel, 1974; Swanson et al., 1974; Ribak and Peters, 1975; Legg, 1979a; Brauer and Schober, 1982; Beitz, 1989 Moore et al, 2000 Beitz, 1989 Foster et al., 1989, Terenzi et al., 1995 Beitz, 1989 Klooster et al., 1995 Beitz, 1989 Lee et al., 1989; Kurimoto et al., 1995 Villar et al., 1988 Pasquier and Tramezzani, 1979 Swanson and Hartman, 1975 Ndiaye et al., 2000 Iwata et al., 1998 Billet et al., 1999 Pasquier and Tramezzani, 1979 Marini et al., 1999 Rhoades et al., 1989a Ohtsuki et al., 1992 Gayton and Pesaro, 1998 Ohtsuki et al., 1992 Pasquier and Tramezzani, 1979 Hopkins and Niessen, 1976 Vincent et al., 1978; Bickford and Hall, 1992 Rhoades et al., 1982 Yasui et al., 1995 Veniente et al., 2000 Rhoades et al., 1989a Rhoades et al., 1989a Antonetty and Webster, 1975 Herbert et al., 1997 Watanabe and Kawana, 1982; Romanowski et al., 1985; Kim et al., 1992 Kolmac and Mitrofanis 2000 Thompson and Robertson, 1987 Bickford and Hall, 1989 Yamasaki et al., 1986; Krauthamer et al., 1992 Redgrave et al., 1987; Redgrave et al., 1988; Dean et al., 1989 Fallon and Moore, 1979 Redgrave et al., 1987a Yasui et al., 1998 Waldron and Gwyn, 1969; Yasui et al., 1998 Continued

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TABLE 2 Efferents—cont’d Lamina VIII from DpG to rostral cord and cervical enlargement Me5 medial to PBg(i) medulla(c,i) MGM Midline and intralaminar n: Pa PAG in midbrain PC(c,i) Periabducens raphe PF(c,i) Pn(i) Po PP, postcomplex PPRFmedial to bic(i) PPTg(c) reticular midbrain(i) pons(c) reticular pontomedulla(i) Rostral ventral resp gp (c,i) RtTg(c) sp cord cervical(c) SN(c) STh VLTg ZI 7 In rabbit:

Connections of Superior Colliculus—cont’d

Murray and Coulter, 1982 Ndiaye, 2000 Redgrave et al., 1987 Waldron and Gwyn, 1969 Linke et al., 1999 Linke et al., 1999; Krout et al., 2001 Krout et al., 2001 Reese, 1988 Yamasaki et al., 1986; Krauthamer et al., 1992 Dean et al., 1989 Yamasaki et al., 1986; Krauthamer et al., 1992 Waldron and Gwyn, 1969; Petrovicky, 1975; Chevalier and Deniau, 1984 Krauthamer et al., 1992 Linke et al., 1999 Redgrave et al., 1987a Redgrave et al., 1987a Petrovicky, 1975; Redgrave et al., 1987a; Bickford and Hall, 1989 Chevalier and Deniau, 1984; Redgrave et al., 1987a Gayton and Pasaro,1998 Petrovicky, 1975; Burne et al., 1981; Torigoe et al., 1986; Redgrave et al., 1987a Murray and Coulter, 1982; Redgrave et al., 1987a; Sahibzada et al., 1987; Bickford and Hall, 1989 Cornoli et al., 2003 Tokuno et al., 1994 Herbert et al., 1997; Yasui et al., 1998 Cadusseau and Roger, 1985; Shammah-Lagnado et al., 1985; Kim et al., 1992 Miyashita and Mori, 1995 thalamus: CM, LD, MD, MGM, PF, Re, SG, ZI; pretectum; midbrain: IC, PAG, PPTg, SC; pons: C, RtTg, Pn, PnO; medulla: IO, Md,; motor nuclei 7, 13; hypothalamus (Holstege and Collewijn, 1982)

a

c, contralateral; i, ipsilateral; c,i, bilateral. In hamster.

b

through the SC (Kim et al., 1992). A small proportion of incertocollicular neurons also projects to nuclei of the dorsal thalamus (Romanowski et al., 1985). Note also that the zona incerta, as well as the central laminar and paracentral intralaminar nuclei of the dorsal thalamus, projects to Oc1, (see Section V). Deeper laminae of the SC also project rostrally to the intralaminar nuclei—central laminar, paracentral, and parafascicular nuclei (Linke, 1999; Krout et al., 2001; Yamasaki et al., 1986)—which also receive projections from a variety of other visually related regions, including visual cortical areas, frontal eye fields (caudal part of cytoarchitectonic area 8), and pretectum. There is evidence, at least in the cat, that intralaminar nuclei are involved in eye movements, orienting, and defense reactions (for review, see Garey et al., 1991; Macchi and Bentivoglio, 1986; Nakamura and Kawamura, 1988), a role that is consistent with the presence of a collicular input. In the rat, the SC forms part of a subcortical pathway for the transfer of sensory information to the limbic system via projections from superficial and deep SC to the suprapeduncular, peripeduncular, and medial division of the medial geniculate nuclei, all of

which in turn project to the lateral nucleus of the amygdala (Linke, 1999; Linke et al., 1999).

Ventral Lateral Geniculate Nucleus and Intergeniculate Leaflet Organization and Subdivisions The ventral lateral geniculate nucleus (VLG), subjacent to DLG, constitutes a part of ventral rather than dorsal thalamus (Jones, 1985). It is bordered laterally by the optic tract and its substance is dissected by bundles of retinal axons (Atlas plates 33–36). Between VLG and DLG lies a small lamina, the intergeniculate leaflet (IGL, Atlas plates 34–37; Hickey and Spear, 1976; Perry and Cowey, 1979; Moore et al., 2000). Two divisions can be clearly recognized within the VLG itself (Niimi et al., 1963). Only the external, lateral, or magnocellular (VLGMC) division receives a significant retinal input, which arises from both eyes (Cajal, 1911; Hickey and Spear, 1976). It contains neurons responsive to visual stimuli (Sumitomo et al., 1979) and projects to visual nuclei (Brauer and Schober, 1982; Kolmac et al., 2000;

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32. VISUAL SYSTEM

Moore et al., 2000). A range of nonvisual thalamic nuclei projects to the medial or parvocellular region (VLGPC) (Kolmac et al., 2000). Due to their close apposition, projections from the two divisions of VLG have not always been distinguished in the rat, although recent studies are more precise (see Table 3). Selective damage to VLG or to adjacent zona incerta (ZI) results in a substantial impairment in the discrimination of light intensity (simultaneous black vs white) but not in pattern discrimination (simultaneous horizontal vs vertical) (Legg, 1979b), suggesting that ZI lies on the output pathway from VLG. There are reciprocal connections between VLG and ZI (Moore et al., 2000; Roger and Cadusseau, 1985; Watanabe and Kawana, 1982). Cells in VLG and IGL project via four pathways. Overall, the efferent projections of VLG terminate predominantly in visual regions and those of the IGL in hypothalamus, as well as some visual regions (including suprachiasmatic nucleus) (Moore et al., 2000). Anatomically, two classes of neurons have been distinguished in VLG (Brauer and Schober, 1973; Mounty et al., 1977; Stelzner et al., 1976; Werner and Brauer, 1984). Two-thirds of the cells are multipolar, probably projection neurons, and these can be further subdivided into three classes on the basis of their size, morphology, and location in VLG. One class, lying medially, may represent the neurons of VLGPC. The remaining cells in VLG are usually bipolar and resemble closely the interneurons with presynaptic dendrites found in DLG (Mounty et al., 1977). Although somata containing GABA are distributed throughout, they are found predominantly in VLGMC (Ohara et al., 1983). Clusters of cells immunoreactive for calretinin are found in the VLGPC and IGL (Arai et al., 1992). IGL Consistent with its role as an integrator of visual inputs to feedback to the SCN (Moore and Card, 1994), cells in the IGL projecting to the suprachiasmatic nucleus respond to light with tonic increases or decreases in firing rates (Zhang and Rusak, 1989). Some are “suppressed-by-contrast” cells, but the majority have properties of the luminance units described in the VLG (Hale and Sefton, 1978; Sumitomo et al., 1979). Cells in IGL contain GABA (Ohara et al., 1983) colocalized with neuropeptide Y (NPY) or enkephalin (Moore and Card, 1994). In addition, estrogen receptor β and progesterone receptor mRNA are expressed in cells in IGL and VLG, suggesting that these visual regions may be involved in the integration of hormonal activity (Horvath et al., 1999). In the IGL of rats (but not in hamsters), enkephalincontaining neurons (Mantyh and Kemp, 1983; Wadhwa et al., 1990), which also contain GABA (Moore and Card, 1994), project to the opposite IGL (Card and Moore, 1982, 1989; Moore and Card, 1994) via optic chiasm

1111

and posterior commissure (Mikkelsen, 1992). Stimulation of one IGL diminishes activity in its contralateral counterpart, perhaps representing a mechanism to reduce the responsiveness of this tonic system to transient changes in illumination (Zhang and Rusak, 1989). A projection from the IGL to the deep pineal gland in rats (Mikkelson, 1994; Mikkelsen and Møller, 1990) and gerbils (Mikkelsen et al., 1991) provides the means by which the secretion of melatonin may be influenced by retinal activity. Other projections from IGL are to visual areas: pretectum, SC, and accessory optic nuclei (Moore et al., 2000). The geniculohypothalamic tract, arising in the IGL and the immediately adjacent VLG, terminates in the ventral part of the suprachiasmatic nucleus (Card et al., 1981; Mikkelson, 1994; Moore et al., 1984; Ribak and Peters, 1975; Swanson et al., 1974). At least in hamsters, it modifies the photic responsiveness of circadian rhythms (Harrington and Rusak, 1988; Rusak et al., 1989). The projection arises from a population of IGL neurons positive for NPY (colocalized with GABA) (Moore and Card, 1994) and its C-flanking peptide (CPON) or avian pancreatic polypeptide (Card and Moore, 1982, 1989; Harrington et al., 1987; Hendrickson et al., 1972; Mantyh and Kemp, 1983; Moore and Lenn, 1972; Moore et al., 1984; Shinohara et al., 1993).. The proportion of NPY cells changes with the length of the photoperiod (Jacob et al., 1998). IGL is thought to modulate suprachiasmatic pacemaker cells (Zhang and Rusak, 1989) by facilitating the difference between light and darkness (Shinohara et al., 1993). IGL also projects to the subcommissioural organ, suggesting that it may be under circadian regulation (Mikkelsen and Vrang, 1994). There is a reciprocal projection from the dorsal part of the suprachiasmatic nucleus to the IGL and VLG (Watts et al., 1987; Watts and Swanson, 1987); the suprachiasmatic nucleus also receives an input directly from retinal ganglion cells that contain substance P (Takatsuji and Tohyama, 1989). IGL receives projections from anterior hypothalamic and retrochiasmatic regions (Moore and Card, 1994). In response to light, both cFos and Fos-B are expressed; c-Fos is present in 15% of cells projecting to the suprachiasmatic nucleus and 34% of those projecting to the contralateral IGL (Peters et al., 1996). The IGL receives substantial serotonergic and dopaminergic inputs, which make fine varicose contacts with dendritic shafts and spines. Serotonergic varicosities establish asymmetrical (presumably inhibitory) synaptic contacts with dendritic shafts and spines. Dopaminergic fibers, which usually establish asymmetrical synapses, occasionally form (presumably inhibitory) symmetrical synapses on dendritic spines and shafts (Papadopoulos and Parnavelas, 1990a, 1990b). Sparse cholinergic inputs have also been reported (Moore and Card, 1994).

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VLGMC (lateral division of VLG) The VLG and its connections with the pretectum contribute to the light reflex (Legg, 1975). VLG has been also strongly implicated in the neuronal circuit underlying brightness discrimination (Legg and Cowey, 1977). There are two morphological and hodological divisions (Kolmac et al., 2000) that are also functionally distinct: the laterally located magnocellular part (VLGMC) and the medially located parvocellular part (VLGPC). About 8% of all retinofugal axons appear to terminate in VLGMC (calculated from Toga and Collins, 1981). VLGMC is organized retinotopically, with a representation of the part of the contralateral visual field extending 20º above and 20º below the horizontal meridian (Nagata and Hayashi, 1984); it receives its substantial input from contralateral retina predominantly via slowly conducting fibers (Hale and Sefton, 1978; Montero et al., 1968; Nagata and Hayashi, 1984). Cells in the VLGM appear to form two distinct functional classes: the so-called “luminance units” and on-center cells with a silent suppressive surround (Hale and Sefton, 1978; Sumitomo et al., 1979). It is likely therefore that VLGMC transmits information about light intensity. Nearly three-quarters of the VLGMC cells are diaphorase positive but GABA negative (Gabbott and Bacon, 1994b; Mitrofanis, 1992) and some cells are positive for enkephalin (Mantyh and Kemp, 1983). Diaphorase-positive cells in VLGMC project to SC (González-Hernández et al., 1994). The ultrastructure of VLG resembles that of DLG, except that the largest somata are the targets of many synaptic terminals containing flattened synaptic vesicles (F1; Stelzner et al., 1976). Both encapsulated and unencapsulated simple and complex serial synapses are observed. Retinogeniculate and corticogeniculate terminals, similar to those in DLG, as well as presynaptic dendrites, have been identified (Lieberman, 1973). The origin and the role of F terminals in VLGMC are unknown; visual Rt does not have a reciprocal relationship with VLG (Hale et al., 1982; Mackay-Sim et al., 1983) and no degenerating F terminals are observed in VLG after damage to it (A.Sefton, unpublished results). The long-lasting postsynaptic inhibition and late bursts observed in DLG do not occur in VLG (Hale and Sefton, 1978; Soltesz et al., 1989; Sumitomo et al., 1979). Nearly three-quarters of cells in the division are NADPH positive. Two subclasses of NADPH cells have been identified; class A is more numerous (14:1). GABA is colocalised in class B cells; they are considered to be interneurons (Gabbot and Bacon, 1994b). Class A neurons containing NADPH project to SC (Gabbott and Bacon, 1994b; González-Hernandez et al., 1994). Cells containing nitric oxide synthase (NOS) project to ipsilateral pretectum, their principal or only target (Meng et al., 1998).

VLGMC also receives inputs from a large variety of subcortical regions, including DLG and lateral posterior, as well as layer 5 of cortical areas Oc1, Oc2L, and Oc2M (Bourassa and Deschênes, 1995; Kolmac et al., 2000; Leong, 1980; Nauta and Bucher, 1954; Pasquier and Villar, 1982b; Sefton et al., 1981; Takahashi, 1985, see Table 3). The cortical innervation of VLG arises as branches of corticotectal and/or corticopontine projections, which may also innervate lateral posterior and lateral dorsal nuclei (Bourassa and Deschênes, 1995). The projection from SC to VLGMC is characterized by the presence of uniform terminals, which contain spherical synaptic vesicles and small dark mitochondria, making Gray type 1 synapses with projection neurons (Taylor and Lieberman, 1987). They form a contingent of R boutons (Lieberman and Webster, 1974) and would thus appear to be providing a facilitatory input to geniculocollicular relay cells, perhaps triggering orienting movements (Dean and Redgrave, 1984; Goodale et al., 1978; Sahibzada et al., 1986) mediated through the projections of VLG to lateral posterior and intrageniculate leaflet (IGL) in the dorsal thalamus. Immunocytochemical studies of VLGMC indicate the presence of terminals containing neurotransmitters and cotransmitters 5-HT, NPY, enkephalin, SP, VIP, dopamine, and, to a lesser extent, norepinephrine (Mantyh and Kemp, 1983; Papadopoulos and Parnavelas, 1989a, 1989b; 1990a, 1990b; Villar et al., 1988). Staining for NADPH activity reveals a striated appearance, with cells and processes distributed among the fiber bundles coursing through the nucleus (Gabbott and Bacon, 1994b). The serotonergic innervation of VLGMC is much denser than that of DLG and consists of fine varicose fibers, which establish asymmetrical contacts with dendritic profiles and symmetrical contacts with somata. In contrast, the norepinephric innervation of VLGMC is much less dense than that of DLG. Norepinephric fibers are fine (0.4–1.5 μm in diameter) and usually form symmetrical synapses with dendritic spines and shafts (Papadopoulos and Parnavelas, 1990b). Dopaminergic fibers mostly form asymmetrical synapses with dendritic spines and shafts, although a few symmetrical synapses with these elements are also present (Papadopoulos and Parnavelas, 1990a). VLGPC (medial division) Less is known of the adjacent small-celled division of VLG. One-third of the neurons are diaphorase positive; the small fraction that are also GABA positive are assumed to be interneurons (Gabbott and Bacon, 1994b). The most caudal regions have been reported to receive two very sparse projections from the visual cortex (Swanson et al., 1974), specifically from areas Oc2L and 2M (Takahashi, 1985), and retina (Hickey and Spear, 1976). This region of VLGPC

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32. VISUAL SYSTEM

1113

and the adjacent ZI, however, merge imperceptibly into the lateral terminal nucleus of the accessory optic system, which also receives a retinal input (Hayhow et al., 1960). Serotonergic networks within VLGPC are less dense than those within VLGMC (Papadopoulos and Parnavelas, 1990b). In contrast, dopaminergic innervation of the medial VLGPC is as dense as that of the MC division (Papadopoulos and Parnavelas, 1990a).

between probably segregating cells that process information from lateral and medial visual fields. In PPT, the dorsal peripheral retina is represented caudally, the nasal retina along the boundary with the OPT so there is a mirror-image representation of the visual field in these nuclei. In contrast in OT, the temporal retina is mapped dorsally, with the nasal retina represented ventrally and the dorsal retina caudally (Scalia and Arango, 1979).

Connections of VLG

Extraretinal Connections

Striking features of VLG are the richness and variety of its connections and the reciprocal nature of many of them. The patterns of connections for VLG and IGL are summarized in Table 3. Wherever possible, projections to and from PC and MC divisions of VLG have been indicated separately. Although interconnections reported between VLGs of both sides have been suggested to be restricted to IGL (Mikkelsen, 1992), the IGL and VLG from each side interconnect, albeit rather differently (Moore et al., 2000).

Connections of the pretectal complex are summarized in Table 4. Whenever it has proved possible to distinguish the connections of individual nuclei, they are indicated. Separate populations of cells in pretectum project to the lateral dorsal nucleus and to the inferior olive, supporting the contention that pretectal nuclei contain functionally distinct cells (Robertson, 1983; see later).

Pretectum General: Cytoarchitecture and Subdivisions The pretectum is derived from the epithalamus and lies at the most rostral pole of the midbrain bordering the thalamus (see Atlas Plates 23–26; also see Chapter 17). Although often referred to as a single entity, the pretectum contains a number of distinct nuclear groups. The most satisfactory description of this area of the rat’s brain is probably that of Scalia (1972), who reviewed earlier nomenclature and defined four major nuclei (see later). Although some of these can be identified in Nissl-stained material, they can be better recognized by Golgistaining (Gregory, 1985) and by their connectivity. Scalia (1972) described some optic axons terminating medially in an area that has been reported to project to retina and that comprises separate populations of cells projecting to the thalamus and to inferior olive (Itaya, 1980; Robertson, 1983). General: Topographical Organization In the rat, only about 13% of retinofugal axons terminate in the contralateral pretectum (estimated from data of Toga and Collins, 1981). Retinal ganglion cells project to both ipsi- and contralateral olivary and posterior pretectal nuclei (OPT and PPT), but only to the contralateral nucleus of the optic tract (OT). In OPT, the vertical meridian of the visual field (temporal retina) is represented anterolaterally, with dorsal peripheral retina (lower visual field) caudally; the lines of projection run from dorsal to ventromedial. Retinal terminals cluster into two separate regions caudally, with the zone

General: Physiology and Behavior Pretectal units responding to visual stimuli can be classified into groups: tonic on center; tonic off center; phasic, some of which are direction selective (Robertson, 1983); and suppressed by light. Tonic off center, concentric, and suppressed by light cells are more sensitive to cortical inactivation (Molotchnikoff et al., 1988). Following lesions of pretectum, sensitivity to both high and low spatial frequencies was reduced; additional involvement of the adjacent medial thalamus (including parafascicular) specifically impaired the detection of high frequencies (Legg and Turkish, 1983). Substantial damage to pretectum and SC alters light–dark sleep patterns (Miller et al., 1998), perhaps mediated by direct pathways from OPT and PPT to suprachiasmatic nuclei (Mikkelsen and Vrang, 1994). Direct pathways from OPT and PPT that innervate suprachiasmatic nucleus bilaterally may contribute to phase-shifting mechanisms (Mikkelsen and Vrang, 1994). Specific Pretectal Nuclei: Structure and Function Olivary pretectal nucleus (OPT) The olivary nucleus is central to the pupillary light reflex (Chan et al., 1995; Young and Lund, 1994) and receives an input largely (but not exclusively) from small retinal ganglion cells (Young and Lund, 1998). On the basis of its rich interconnectivity, the nucleus may also contribute to other visual functions, including coordinating head and eye movements (Klooster et al., 1995a, 1995b). Lying ventral to the brachium of the SC, it is the most readily identifiable pretectal nucleus, particularly in fiber-stained sections. Rostrally, where it lies superficially, the nucleus forms an olive-shaped core region, but caudally, where it lies more deeply, it forms a shell region that merges with the nucleus of the optic tract. In the core region, only a few crossing axons and a few

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TABLE 3 VLGa Afferents Retina(c,i) Oc1 to VLGMC Oc1,2L.2M, layer 5 Oc2L to VLGMC Oc2M to VLGPC to VLGMC(c) VLG(c) DLG to MC,PC SC(i) Pretectum APT(i) OPT(c,i) OT(i) CL to VLGPC DR In LC LD to MC,PC LP to MC,PC LT MD to MC,PC MnR PAG(i) Perirubral retic PF to PC Po to PC PPTg(c,i) Efferents VLG(c) MC Pretectum APT(i) NOT OPT(i) OPT(c,i); OT(i) OT(c,i) SC(i) from cells in MC NADPH-d cells to SuG, Op, InG DTN LT(i) LT(c,i) MT SCN(c,i) LP LD PBG HT (lat, dorsal) IO access PAG (i) Pn RF pons, medulla RF perirubral Rh

Connections of Ventral Lateral Geniculate Nucleus and Intrageniculate Leaflet

Hayhow et al., 1962; Hickey and Spear, 1976; Perry and Cowey, 1979; Ahmed et al., 1995, 1998 Nauta and Bucher, 1954; Leong, 1980; Takahashi, 1985 Nauta and Bucher, 1954; Leong, 1980; Sefton et al., 1981; Pasquier and Villar, 1982; Takahashi, 1985; Bourassa and Deschênes, 1995; Kolmac et al., 2000 Takahashi, 1985 Takahashi, 1985 Mackay-Sim et al., 1983 Moore et al., 2000 Kolmac et al., 2000 Perry, 1980; Pasquier and Villar, 1982a; Takahashi, 1985; Taylor et al., 1986 from wide-field cells in Op and deep Mackay-Sim et al., 1983 Pasquier and Villar, 1982 Mackay-Sim et al., 1983 Mackay-Sim et al., 1983 Mackay-Sim et al., 1983 Kolmac et al., 2000 Pasquier and Villar, 1982a, 1982b; Mackay-Sim et al., 1983; Waterhouse et al., 1993 Mikkelsen, 1992 Mackay-Sim et al., 1983 Kolmac et al., 2000 Kolmac et al., 2000 Moore et al., 2000 Kolmac et al., 2000 Pasquier and Villar, 1982a Mackay-Sim et al., 1983 Mackay-Sim et al., 1983 Kolmac et al., 2000 Kolmac et al., 2000 Mackay-Sim et al., 1983 Graybiel, 1974; Swanson et al., 1974; Legg, 1979a; Mikkelsen and Møller, 1990; Moore et al., 2000 Mackay-Sim et al., 1983 Meng et al., 1998 Graybiel, 1974; Swanson et al., 1974; Ribak and Peters, 1975; Legg, 1979a; Foster et al., 1989; Moore et al., 2000 Moore et al., 2000 Graybiel, 1974 Swanson et al., 1974; Ribak and Peters, 1975; Legg, 1979a Graybiel, 1974 Swanson et al., 1974; Ribak and Peters, 1975 Perry 1980 Gonzalez-Hernández et al., 1994 Graybiel, 1974; Swanson et al., 1974; Ribak and Peters, 1975; Legg, 1979a; Brauer and Schober, 1982; Taylor et al., 1986; Beitz, 1989; Moore et al., 2000 Moore et al., 2000 Graybiel, 1974 Swanson et al., 1974; Ribak and Peters, 1975; Legg, 1979a Graybiel, 1974; Swanson et al., 1974; Beitz, 1989 Swanson et al., 1974; Ribak and Peters, 1975; Legg, 1979a; Card and Moore, 1982 Moore et al., 2000 Thompson and Robertson, 1987; Moore et al., 2000 Moore et al., 2000 Moore et al., 2000 Moore et al., 2000 Graybiel, 1974; Legg, 1979a Graybiel, 1974; Ribak and Peters, 1975; Legg, 1979a; Moore et al., 2000 Moore et al., 2000 Graybiel, 1974; Legg, 1979a Moore et al., 2000

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32. VISUAL SYSTEM

TABLE 3 Efferents—cont’d VRe ZI(i)

1115

Connections of Ventral Lateral Geniculate Nucleus and Intrageniculate Leaflet—cont’d

Moore et al., 2000 Graybiel, 1974; Swanson et al., 1974; Ribak and Peters, 1975; Legg, 1979a

IGL (when identified separately)a Afferents Retina Hickey and Spear, 1976; Moore and Card, 1994 IGL(c) Watts et al., 1987; Watts and Swanson, 1987; Giolli et al., 1988; Zhang and Rusak, 1989; Mikkelsen and Møller, 1990; Moore and Card, 1994 SC Taylor et al., 1986 SCN Watts et al., 1987; Watts and Swanson, 1987; Card and Moore, 1989; Morin et al., 1994 Efferents SC SuG SCN(c,i) IGL(c) VLGPC and ZI(c) OPT OT DTN LTN AH, DH Pa Pineal SCO Rh VRe ZI

Taylor et al., 1986 Moore et al., 2000 Watts et al., 1987; Watts and Swanson, 1987; Card and Moore, 1989; Mikkelson and Vrang, 1994; Moga and Moore 1997 Card and Moore, 1989; Zhang and Rusak, 1989; Mikkelsen and Møller, 1990; Mikkelsen and Møller, 1990 Moga and Moore, 1997; Moore et al., 2000 Moore et al., 2000 Moore et al., 2000 Moore et al., 2000 Moore et al., 2000 Moore et al., 2000 Mikkelsen and Møller, 1990 Mikkelson, 1994 Moore et al., 2000 Moore et al., 2000 Moore et al., 2000

a

c, contralateral; i, ipsilateral; c,i, bilateral.

neurons are immunoreactive to the calcium-binding proteins parvalbumin or calbindin D-28kD. In contrast, a network of densely intertwined neurons in the shell region is strongly reactive for parvalbumin (SchmidtKastner et al., 1992). OPT, which receives a strong bilateral retinal input, contains distinctive, large, wellbranched cells that have tonic on-center receptive fields (Clarke and Ikeda, 1981) and that mediate the pupilloconstrictor light reflex (Clarke and Ikeda, 1985; Trejo and Cicerone, 1984; Chan et al., 1995). Ultrastructurally in OPT, the two major types of cells are presumed to be projection cells and interneurons. Terminals of retinal axons comprise about two-thirds of all terminals present and resemble those in DLG; they contain round vesicles and make Gray type I synaptic contacts. About onequarter of terminals are P boutons, presynaptic dendrites of the smaller neurons containing pleomorphic vesicles. Of the remaining terminals, about half are small with round vesicles probably arising in SC, whereas the other half contain flattened vesicles. The surprisingly complex organization, including serial synapses, is similar to that described for the DLG (Campbell and Lieberman, 1985). Some somata and cell processes in OPT contain GABA (Ohara et al., 1983). In the dark, fibers immunoreactive to substance P increase, but are unaffected by constant light (Takatsuji and Tohyama, 1993).

Tonic on-center units, found only in OPT, have large (mean size 31º) receptive fields. It is likely that they receive a convergent retinal input, as afferent retinal axons have substantially smaller receptive fields (about 10º; Trejo and Cicerone, 1984). Additional inputs might arise from the cortex, although their influence appears to be small (Molotchnikoff et al., 1988), and from cells in VLG with similar receptive fields (Hale and Sefton, 1978; Ribak and Peters, 1975). Electrical microstimulation of OPT induces pupillary constriction (Trejo and Cicerone, 1984). OPT projects to the suprachiasmatic nucleus (Mikkelsen and Vrang, 1994; Moga and Moore, 1997). Unilateral lesions of OPT diminish but do not abolish the light reflexes. The nucleus projects bilaterally to the Edinger–Westphal nucleus (EW) and to the nucleus of the posterior commissure, which also projects bilaterally to EW, providing pathways for the consensual light reflex (Klooster et al., 1995a; Young and Lund, 1994). Integration occurs not only in OPT, but also in EW. In albino rats, with a reduced ipsilateral pathway, the consensual reflex is reduced (Chan et al., 1995). The reflex is preserved following the transplantation of fetal retinae (Radel et al., 1995), suggesting not only that precise topography is not required, but also that a transplant interacts dynamically with the host (Young and Lund, 1997).

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TABLE 4

Connections of the Pretectum

Afferentsa Pretectum (not differentiated) Oc1(i) Nauta and Bucher, 1954; Leong, 1980; Benzinger and Massopust, 1983; Takahashi, 1985 Oc1 layer 5 Martin and Sefton, 1981 Oc2L Takahashi, 1985 Oc2M Benzinger and Massopust, 1983; Takahashi, 1985 SC(i) Perry, 1980 from SuG Martin and Sefton, 1981 VLG Meng et al., 1998 PrH(c,i) Ohtsuki et al., 1992 APT VLG(i) Graybiel, 1974; Swanson et al., 1974; Ribak and Peters, 1975; Legg, 1979 MC Foster et al., 1989 SC Foster et al., 1989 OPT(i) Klooster et al., 1995 PPT Foster et al., 1989 PBG(c) Foster et al., 1989 DCN Yoshida et al., 1992 Dk Foster et al., 1989 DpMe Foster et al., 1989 FL Foster et al., 1989 HL Foster et al., 1989 LC Foster et al., 1989 PAG Foster et al., 1989 Pcom Foster et al., 1989 PP Foster et al., 1989 PPTg Foster et al., 1989 VMH Foster et al., 1989 ZI Foster et al., 1989 5 Yoshida et al., 1992 OPT Retina(c,i) Scalia, 1972; Perry and Cowey, 1979; Scalia and Arango, 1979; Campbell and Lieberman, 1985; Miguel-Hidalgo et al., 1991 (contain substance P); Radel et al., 1995; Young and Lund, 1998 VLG(c,i) Swanson et al., 1974; Ribak and Peters, 1975; Legg, 1979a VLG(i) Graybiel, 1974 OPT(c) Radel et al., 1995 Oc1, Oc2L, Oc2M Takahashi, 1985 OT Retina(c,i) Scalia, 1972; Scalia and Arango, 1979 Retina(i) Perry and Cowey, 1979; Kato et al., 1992 Oc1, Oc2L Takahashi, 1985; Molotchnikoff et al., 1988 Oc2M Benzinger and Massopust, 198 VLG(c,i) Swanson et al., 1974; Ribak and Peters, 1975 VLG(i) Graybiel, 1974 SC(i) Takahashi, 1985 MT Giolli et al., 1988; van der Togt and van der Want, 1992; van der Togt and Schmidt, 1994 Three paths Blanks et al., 1982 GABAergic Giolli et al., 1992 DT Giolli et al., 1992 LT Giolli et al., 1992 PPT Retina(c) Scalia, 1972; Scalia and Arango, 1979 Retina(c,i) Perry and Cowey, 1979 Oc1, Oc2L, Oc2M Takahashi, 1985 SC(c,i) Takahashi, 1985 Efferentsa Pretectum (not differentiated) SC(i) Watanabe and Kawana, 1979 LG Pasquier and Villar, 1982 LP Schober, 1981

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TABLE 4 Efferentsa—cont’d APT Visual Rt(i) VLG(i) LD(i) IO(i) PAG OPT VLG(c,i) DLG(i) IGL(c,i) LD(i) OPT(c) OT(i) SC SCN Dk(c,i) EW(c,i) F(c) Interst Cajal PAG(c,i) Pcom Pn(i) ZI(c) 3(c) OT VLG(c,i) DLG(i) SCdeep OPT(i) OT(c) DT(c) MT LP(i) LD(i) IO(i) Pn(i) PrH RtTg(i) PPT APT LD(i) SCN IO(i)

1117

Connections of the Pretectum—cont’d

Hale et al., 1982 Mackay-Sim et al., 1983 Robertson, 1983; Thompson and Robertson, 1987 Robertson, 1983 Beitz, 1989 Mackay-Sim et al., 1983; Klooster et al., 1995; Young and Lund, 1998; Klooster et al., 1995b Mackay-Sim et al., 1983 Klooster et al., 1995a Robertson, 1983; Thompson and Robertson, 1987 Klooster et al., 1995a Klooster et al., 1995a Beitz, 1989 (deep); Klooster et al., 1995a (in) Mikkelsen and Vrang, 1994; Moga and Moore, 1997 Klooster et al., 1995a, 1995b Campbell and Lieberman, 1985; Klooster et al., 1995a,b; Young and Lund, 1998 Klooster et al., 1995a Klooster et al., 1995b Klooster et al., 1995a, 1995b Young and Lund, 1998 Klooster et al., 1995a, 1995b Klooster et al., 1995a Klooster et al., 1995a Mackay-Sim et al., 1983 Mackay-Sim et al., 1983 Beitz, 1989 Terasawa et al., 1979 Terasawa et al., 1979 Terasawa et al., 1979 Terasawa et al., 1979; Blanks et al., 1982; Natal and Britto, 1987; Giolli et al., 1988; van der Togt and Schmidt, 1994 Mason and Groos, 1981 Mason and Groos, 1981; Robertson, 1983; Thompson and Robertson, 1987 Robertson, 1983 Terasawa et al., 1979 Korp et al., 1989 Terasawa et al., 1979; Cazin et al., 1980a, 1980b Foster et al., 1983 Robertson, 1983 Mikkelson and Vrang, 1994; Moga and Moore, 1997 Robertson, 1983

a

c, contralateral; i, ipsilateral; c,i, bilateral.

Anterior pretectal nucleus (APT) This, the most rostral part of pretectum, comprises cells with small to medium somata. It has been divided into two regions (see Scalia, 1972; Scalia and Arango, 1979), which may represent separate nuclei: the compact (lying posterior, medial to lateral posterior nucleus and ventral to OPT) and the reticular (ventrally), although other possible functional subdivisions may be present (Foster et al., 1989). Although APT appears to receive inputs from other pretectal nuclei, which mediate visual reflexes, it does not receive a direct retinal input (Scalia and Arango, 1979). Its rostral part, which receives inputs from the deep layers of the SC and the contralateral

parabigeminal nucleus, may be more related to nociception than to vision (Foster et al., 1989). Posterior pretectal nucleus (PPT) This nucleus, also containing a variety of characteristic small to medium cells (Gregory, 1985), lies medial to OT and OPT, just rostral to the SC. Although in caudal sections it moves to a more lateral position, it always remains medial to OP. It has been reported to receive a projection from the visual cortex (Nauta and Bucher, 1954) and to project extensively, including the suprachiasmatic nucleus (Mikkelsen and Vrang, 1994). As yet the functional properties of its constituent cells have not been

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elucidated, although P.R. Martin and A.J. Sefton (personal communication) recorded “suppressed by contrast” activity in the region of the nucleus. Nucleus of optic tract (OT) The comparatively large multipolar cells of OT lie among axons of the brachium of SC (Gregory, 1985); it lies adjacent to the accessory dorsal terminal nucleus with which it has a very close relationship (Schmidt et al., 1995) (see following section). In several species, OT cells are innervated by the contralateral retina and have been reported to have large receptive fields. Their responses are strongly directionally selective, with the neurons responding particularly well to large targets moving forward along a horizontal axis at about 1º per second (Cazin et al., 1980a). The responses are mediated by both NMDA and non-NMDA receptor mechanisms (Schmidt, 1991). GABA-mediated inhibition is present, but it appears to be tonic and independent of direction (Schmidt et al., 1994). Stimulation of the medial terminal nucleus inhibits responses in OT (van der Togt and Schmidt, 1994). Three independent efferent populations of cells have been described. Those projecting to the ipsilateral inferior olivary nucleus, which bifurcate to the prepositus hypoglossal nucleus, reveal direction-selective responses, consistent with transferring information about retinal slip (Schmidt et al., 1995). OT has been implicated in the generation of horizontal optokinetic nystagmus in the rat (Cazin et al., 1980b; Reber et al., 1991), which, like another afoveate lateral-eyed species, the rabbit (Hess et al., 1985), has characteristically a stronger response to temporonasal rather than to nasotemporal movement. The strong asymmetry of the horizontal optokinetic nystagmus is reduced by monocular the enucleation (Reber et al., 1991). Projections from the OT mediating the optokinetic nystagmus are to the pontine reticulotegmental nucleus (Cazin et al., 1980b; Terasawa et al., 1979), which is also responsive to vestibular stimulation (Cazin et al., 1980b) and, to a lesser extent, to the prepositus hypoglossal nucleus (Korp et al., 1989). Cells in OT with different response properties also project to the ipsilateral DLG and contralateral dorsal terminal nucleus, suggesting that OT might serve additional functions (Schmidt et al., 1995).

Accessory Optic System Subdivisions The accessory optic system, recognized by many early anatomists, is present in all mammalian species studied so far (Marg, 1973). Accessory nuclei are related to vestibuloocular and optokinetic nystagmus and may signal self-motion (see Giolli et al., 1992; Simpson, 1984).

In the rat, Hayhow et al. (1960) named the terminal nuclei dorsal (DT), lateral (LT), and medial (MT). An additional area, the visual tegmental relay zone (Giolli et al., 1992; Atlas Figs. 38–40), has also been related functionally to the accessory optic system. The pretectal OT has been associated with the accessory system (see Cazin et al., 1980; van der Togt and Schmidt, 1994). Accessory nuclei are interconnected reciprocally (Blanks et al., 1982; Ding et al., 1995; Giolli et al., 1988) (see Table 5). Two crossed accessory fasciculi, superior and inferior, leave the main optic tract behind the chiasm; an alternative nomenclature suggested by Yamadori and Yamauchi (1983) has not been adopted. The inferior fasciculus can initially be identified as a small separate bundle that lies medial to the main optic tract. Subsequently, its fibers intermingle with the median forebrain bundle as they run close to the base of the peduncle to innervate the MT, the principal nucleus for the accessory system. The superior fascicle is a diffuse collection of fibers leaving the posterior edge of optic tract and inferior margin of the brachium of the SC. One branch travels downward, joins the inferior fasciculus, and terminates in the MT, whereas some axons travel over the lateral surface of the medial geniculate nucleus to terminate in the LT. Interspersed among the fibers of the superior fasciculus are neurons that constitute the interstitial nucleus of the superior fasciculus (Giolli et al., 1984). The most caudal axons travel only a short distance to terminate in the DT (Hayhow et al., 1960). Retinal innervation of the DT and LT is sparse (Kostovic, 1971; Yamauchi et al., 1983); the input from the ipsilateral retina is particularly small (Wree and Zilles, 1983). In terminal nuclei, GABAergic neurons postsynaptic to optic afferents contain a calcitonin gene-related peptide, expressing different calcium-binding proteins. They make symmetric synapses with somata, dendritic shafts, and spines (Zhou et al., 1999). Opiate receptors are located in nuclei of the accessory system (Atweh et al., 1978), and μ-opioid receptor-like activity is found in terminals of retinal axons within accessory nuclei; they are associated with asymmetrical contacts and round synaptic vesicles (Ding et al., 1995). Medial terminal nucleus (MT) MT is the most prominent of the nuclei, as is particularly evident in horizontal acetylcholinesterase-stained sections (see Atlas plate 93). The nucleus receives an input from about 1750 bistratified retinal ganglion cells distributed across the retina; the cells presumably have on–off responses and direction-selective properties (Dann and Buhl, 1987). It has been suggested that within it, retinal terminals contact dendrites of cells in the adjacent substantia nigra, which in turn project to the striatum as well as to cingulate and prefrontal cortex (Giolli et al., 1985b).

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TABLE 5 Afferentsa MT Retina(c) Retina(c,i) DT LT VLG OT DpMe IMLF PAG RPn RtTg DT Retina(c) Oc MT GABAergic Interstitial nucleus of superior fascicle Ventral tegmental relay zone LT Retina(c) Oc MT GABAergic VLG Efferentsa MT DT LT OT(i) GABAergic Cajal, Dk DpMe PnO PAG PB(Tsai) LVe, SVe IO Pons DT MT LT MT OT

1119

Connections of the Accessory Optic Nuclei

Hayhow et al., 1960; Lenn, 1972; Dann and Buhl, 1987 Wree and Zilles, 1983 Blanks et al., 1982; Giolli et al., 1988; Ding et al., 1995 Blanks et al., 1982; Giolli et al., 1988; Ding et al., 1995 Graybiel, 1974; Swanson et al., 1974; Swanson and Hartman, 1975; Giolli et al., 1988 Terasawa et al., 1979; Blanks et al., 1982; Natal and Britto, 1987; Giolli et al., 1988; van der Togt and Schmidt, 1994 Giolli et al., 1988 Giolli et al., 1988 Giolli et al., 1988 Giolli et al., 1988 Giolli et al., 1988 Hayhow et al., 1960; Kostovic, 1971; Yamauchi et al., 1983 Leong, 1980 Blanks et al., 1982;. Giolli et al., 1984, 1985a, 1988; Ding et al., 1995 Giolli et al., 1992 Giolli et al., 1992 Giolli et al., 1992 Hayhow et al., 1960; Kostovic, 1971; Yamauchi et al., 1983 Leong, 1980 Blanks et al., 1982; Giolli et al., 1984, 1985a, 1988; Ding et al., 1995 Giolli et al., 1992 Graybiel, 1974; Ribak and Peters, 1975; Legg, 1979a; Mackay-Sim et al., 1983

Blanks et al., 1982;. Giolli et al., 1984, 1985a, 1988; Ding et al., 1995 Blanks et al., 1982; Giolli et al., 1984, 1985a, 1988; Ding et al., 1995 Giolli et al., 1984, 1985a; Clarke et al., 1989; Giolli et al., 1992 van der Togt and Schmidt, 1994 Giolli et al., 1984 Giolli et al., 1984 Giolli et al., 1984 Giolli et al., 1984; Clarke et al., 1989 Giolli et al., 1984, 1985a Giolli et al., 1984; Clarke et al., 1989 Giolli et al., 1984; Clarke et al., 1989 Giolli et al., 1984; Clarke et al., 1989 Giolli et al., 1988; Ding et al., 1995 Blanks et al., 1982; Giolli et al., 1984, 1985a, 1988, 1992; Ding et al., 1995 van der Togt and van der Want, 1992

a

c, contralateral; i, ipsilateral; c,i, bilateral.

The nucleus can be subdivided into two cytologically distinct regions: the dorsal (MTd), in which the inferior retinofugal fascicle terminates, contains large, slender fusiform cells, whereas the ventral (MTv), innervated by the superior fascicle, contains round or oval cells. In both subdivisions, GABA-containing cells are present (Giolli et al., 1985b; van der Togt et al., 1991). Following transection of the contralateral optic nerve, there is a complete loss of immunoreactivity to NAAG in the MT, thus it appears that retinal axons terminating there probably

release an acetylated dipeptide such as NAAG (Moffett et al., 1991). Virtually all cells in the MT project with little collateralization, suggesting that the different morphological types of cells mediate different aspects of the optokinetic nystagmus (Clarke et al., 1989). Through the nucleus run many myelinated axons, only some of which are of retinal origin (Hayhow et al., 1960); other afferents and efferents are listed in Table 6. Only about half of the spherical synaptic contacts were shown to degenerate after eye removal (Lenn, 1972).

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TABLE 6 Afferentsa Retina Oc1 Layer 5 Oc2L Layer 6 Layer 5 Oc2M Cortex area 36 Cortex area 4 Rt SC(c,i) SuG Op Pretectum OT Cu DR LC v(to LD) ZI

Connections of Lateral Posterior Nucleus

Goodman et al., 1973; Yamadori, 1977; Perry and Cowey, 1979 Nauta and Bucher, 1954; Leong, 1980; Benzinger and Massopust, 1983; Takahashi, 1985; Beitz, 1989 Mason and Groos, 1981; Schober, 1981; Bourassa and Deschênes, 1995 Takahashi, 1985 Johnson and Burkhalter, 1992 Mason and Groos, 1981 Benzinger and Massopust, 1983; Robertson, 1983 Hughes, 1977; Olavarria, 1979; Mason and Groos, 1981; Schober, 1981 Schober, 1981 Schober, 1981; Robertson, 1983; Sumitomo et al., 1988; Pinault and Deschênes, 1998; Kolman et al., 2000 Perry, 1980; Takahashi, 1985 Mason and Groos, 1981 Schober, 1981; Lane et al., 1993; 1997 Schober, 1981; Robertson, 1983 Mason and Groos, 1981 Lund and Webster, 1967 Mason and Groos, 1981 Swanson and Hartman, 1975 Takahashi, 1985 Schober, 1981

Efferents Oc2L Layers 4,5,?6 Oc1 Oc2M Cortex area 7 Cortex area 20 Cortex area 36 Rt

Lashley, 1941; Hughes, 1977; Olavarria, 1979; Coleman and Clerici, 1980; Perry, 1980; Sumitomo et al., 1988 Schober, 1981 Waller, 1934; Schober et al., 1976; Hughes, 1977; Perry, 1980; Olavarria and Montero, 1981; Sumitomo et al., 1988; Johnson and Burkhalter, 1992 Hughes, 1977; Perry, 1980 Hughes, 1977; Hughes, 1977; Mason and Groos, 1981 Coleman and Clerici, 1980; Leong, 1980 Sumitomo et al., 1988

SC deep

Beitz, 1989

a

c, contralateral; i, ipsilateral; c,i, bilateral.

Consistent with its postulated role, MT is related reciprocally to other visual nuclei involved in optokinetic nystagmus (OT, DT, and LT) and projects to the pons, vestibular complex, inferior olive, interstitial nucleus of the medial longitudinal fasciculus, and nuclei of Cajal and Darkschewitsch (Blanks et al., 1982; Giolli et al., 1985a; Giolli and Creel, 1974; Giolli et al., 1988). The projection from the MT to the ipsilateral DT and OT consists of GABAergic and non-GABAergic neurons, with GABAergic neurons constituting 72% of the projection. A substantial fraction (50–58%) of GABAergic neurons in both subdivisions, however, project elsewhere (Giolli et al., 1992). The nonreciprocal projection from VLG (Giolli et al., 1988; Graybiel, 1974; Swanson et al., 1974) is likely to arise in VLGPC and to mediate vestibuloocular interactions (see page 1106). In most species, including rat, there appears to be no evidence of a cortical input to MT (see Giolli et al., 1988). Cells in MT in several species have directional properties broadly similar to those in OT (see Simpson, 1984). In MT of the rat, for the majority of cells, the

preferred direction of a slowly moving stimulus is uptemporally, for the minority, down-nasally, although some activity is elicited by horizontally directed stimuli (Biral et al., 1987). Similarly, in guinea pig, some neurons in the dorsal region are strongly activated by vertical and horizontal movements of large parts of the visual field (Lui et al., 1990). The projection from OT appears to modulate the selectivity of cells in MT, as lesions of OT abolished their preference for movements in the nasal direction and increased responses to stimuli moving temporally (Natal and Britto, 1987). Cells in MT inhibit activity in the OT, possibly mediated through GABAB receptors (van der Togt and Schmidt, 1994). Lateral terminal nucleus (LT) This group of cells, poorly distinguished in Nissl-stained sections (see Atlas, Fig. 40), lies below the MG and rostral to SN, merging with ZI and VLG. Medially, it borders the interstitial nucleus of the superior fasciculus, and its oval or fusiform cells, which include a contingent of GABApositive cells (Giolli et al., 1985b, 1992), are surrounded

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32. VISUAL SYSTEM

by a plexus of fibers. Some of the axons are of retinal origin, traveling in the main optic tract or the superior fascicle of the accessory system, while others are derived from the commissures of Meynert and Gudden (Hayhow et al., 1960). The GABAergic output neurons of LT do not project to OT or DT (Giolli et al., 1992). Because of its location, the connections of LT are difficult to distinguish from those of VLG, which itself projects to LT (Mackay-Sim et al., 1983; Ribak and Peters, 1975; Swanson et al., 1974). It is likely that the latter connection is likely to be restricted to VLGPC (see earlier discussion and see page 1106). LT has, however, been reported to be reciprocally connected with MT (Giolli et al., 1988) and to project to DT and OT (Giolli et al., 1992) as well as to receive projections from the Oc (Leong, 1980) and locus coeruleus(Swanson et al., 1974). Cells recorded in LT of species with frontally positioned eyes, such as cats or macaques, have been reported to be vertically direction selective, with a preference for posterior (temporal) motion downward (for reviews, see Grasse and Cynader, 1991; Simpson, 1984). Selective damage results in impairment of simultaneous horizontal/vertical (pattern) and black/white (intensity) visual discrimination (Legg, 1979b). Dorsal terminal nucleus (DT) This small nucleus is not readily identified in Nissl-stained sections; it lies superficially above the brachium of SC, sharing a common border with OT between the medial geniculate nucleus and SC (Atlas Fig. 42). It receives a retinal input directly from the superior retinofugal fascicle (Hayhow et al., 1960; Yamadori and Yamauchi, 1983) and indirectly via the visual cortex (Leong, 1980); it is reciprocally connected with MT (Blanks et al., 1982; Giolli et al., 1984, 1985a, 1988, 1992). In species with frontally placed eyes, cells in DT have properties similar to those of OT and respond to horizontal movement of the target, with the temporonasal direction being preferred (for reviews, see Giolli et al., 1992; Grasse and Cynader, 1991; Simpson, 1984). Functional Considerations The similarity in preferred directionality of DT and OT (horizontal), MT and LT (vertical) suggest that the first pair is related to horizontal optokinetic nystagmus and the second to vertical. Through their connections with the brain stem, they appear to be involved in stabilization of the drift of image position on the retina. Thus the predominance of GABAergic interconnections between some of the nuclei of the accessory optic tract (Giolli et al., 1992) may contribute to the finetuning of responses by reducing activity orthogonal to the plane of retinal slip, as well as that in the same plane but the opposite direction.

ASSOCIATED VISUAL NUCLEI Parabigeminal Nucleus (PBG) The mammalian PBG is considered to be the homologue of the isthmooptic nucleus of lower vertebrates although it lacks a projection to the eye. In the rat, it is located laterally in the brain stem (Atlas plates 29 and 30) adjacent to the brachium of the inferior colliculus and is divided into dorsal, middle, and ventral groups of cells (Tokanuga and Otani, 1978). The superficial retino-recipient laminae of SC project topographically to the ipsilateral PBG particularly to its ventral and dorsal divisions (Linden and Perry, 1983b; Sefton and Martin, 1984). At least some of the tectal cells contributing to this projection contain substance P (BennetClark et al., 1989), an observation consistent with the presence of substance P-immunoreactive fibres (Wang et al., 1988) and receptors (Nakaya et al., 1994) in this nucleus. PBG also contains many fibers immunoreactive for cholecystokinin (Wang et al., 1988), also possibly of tectal origin (Harvey et al., 2001). PBG sends a small projection back to the ipsilateral SC (Stevenson and Lund, 1982a, 1982b; Watanabe and Kawana, 1979); this projection derives in part from cholinergic cells in the dorsal and ventral subnuclei (Tan and Harvey, 1989). In contrast, there is a strong projection, via the optic tract and chiasm (Pasquier and Tramezzani, 1979; Sefton and Martin, 1984; Stevenson and Lund, 1982a, 1982b; Watanabe and Kawana, 1979), from the middle subgroup of the PBG to the rostral part of contralateral SC. In the cat, the PBG contains an orderly map of the visual field, and the properties of its cells resemble those of cells recorded in the superficial gray and optic layers of the SC, except that virtually all cells in PBG are dominated by the contralateral eye (Graybiel, 1978; Sherk, 1979). It seems therefore in the rat that cells in PBG mainly relay information derived from the contralateral eye (via SC) to the opposite SC, as in other mammals with only a small input from ipsilateral temporal retina (cf. Harting et al., 1991a; Stevenson and Lund, 1982a, 1982b; Watanabe and Kawana, 1979). In rats, many PBG neurons are cholinergic (see Mufson et al., 1986; Fitzpatrick and Raczkowski, 1991; Tan and Harvey, 1989; Wang et al., 1988; see also Chapter 35). ChAT immunoreactivity is heaviest in the dorsal and ventral subnuclei/divisions, although occasional ChAT-positive cells (Wang et al., 1988) and some ChAT mRNA expression (A.R. Harvey et al., unpublished) are seen in the middle subnucleus. Rat PBG appears to contain very few interneurons, although it contains a high density of four different GABAA receptor subtypes (Fritschy and Mohler, 1995). There is little or no glutamic acid decarboxylase (GAD)67

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mRNA expression in any of the subnuclei (A.R. Harvey, unpublished observations). Many GAD-expressing cells are, however, found in the area surrounding PBG itself. Neurons in the rostral half of PBG project to the lateral region of the contralateral DLG (Sefton and Martin, 1984; Harting et al., 1991a), a proportion innervating both DLG and SC (Sefton and Martin, 1984). Some PBG fibers terminate in the caudal part of the external medullary lamina of the thalamus. In both albino and pigmented rats, a small projection to the ipsilateral DLG arises in caudal PBG (Harting et al., 1991a; Turlejski et al., 1993).

Lateral Posterior Nucleus (LP) LP, located in lateral dorsal thalamus, lies medial and caudal to DLG (see Atlas Plates 23–41 and Chapter 17) lateral to intralaminar nuclei and habenula, caudal to anterior and ventral nuclei, and rostral to MG and pretectum. LP receives a small retinal input and a significant projection from cells in layer V of the occipital cortex via branches of corticopontine and corticocollicular axons that terminate in restricted varicose endings (Bourassa and Deschênes, 1995). It is also innervated from a distinct sector of Rt, in large, low-density patches (Pinault and Deschênes, 1998). A significant projection is seen from SC (see Table 6). In hamster, terminals arising from the three primary projection sites are distinctive, even when overlapping (Ling et al., 1997). LP is also interconnected with VLGMC (Kolmac et al., 2000; Moore et al., 2000). In rodents, LP is considered to be the homologue of the primate pulvinar (Harting et al., 1972) and of the LP-pulvinar complex in cats (cf. Berson and Graybiel, 1983; Updyke, 1983). It represents a “higher order” nucleus that interconnects cortical areas (Guillery, 1995). In rodents (rat and hamster), as in other mammals, the LP has been subdivided cytoarchitectonically (Lund and Webster, 1967) and on the basis of its afferent input (Crain and Hall, 1980; Ling et al., 1997). Caudal tectorecipient and rostral cortico-recipient regions have been identified in rat (Mason and Groos, 1981; Perry, 1980; Schober, 1981). LP projects cortically to visuotopically organized visual areas in Oc, including Oc1, as well as visual association areas in temporal regions (Coleman and Clerici, 1980; Dreher et al., 1990; Hughes, 1977; Johnson and Burkhalter, 1992; Mason and Groos, 1981; Olavarria, 1979; Perry, 1980; Sanderson et al., 1991; Schober, 1981). On the basis of its cytoarchitecture and connectivity, LP in the rat has been divided into six subregions while the adjacent lateral dorsal nucleus contains two; the following description relies on Takahashi’s (1985) observations. Caudally, the suprageniculate region of the LP receives inputs from the

superficial layers of the ipsilateral SC and from the intermediate layers of the contralateral SC Similarly, the caudomedial region receives a substantial bilateral projection from the superficial layers of the SC (although not from Oc). The caudal sector of the lateral part is innervated ipsilaterally by both SC and area Oc2L (cytoarchitectonic area 18a), while its rostral region and the rostromedial subdivision are innervated not by SC but by ipsilateral areas Oc1, Oc2L, and Oc2M (area 18b). Cells in the caudal regions fire in a mode described as “regularly spiking” with a steep linear increase in firing, whereas those located more rostrally reveal a more clustered mode of spiking. The differences may reflect the dominance of collicular or cortical inputs (Li et al., 2003). The ipsilateral tectal projections derive mainly from cells in the optic layer (Sugita et al., 1983) and many contain calbindin (Lane et al., 1993, 1997). The intramedullary region is innervated from Oc2L and appears to include the retinorecipient zone of LP (Goodman et al., 1973; Perry and Cowey, 1982; Yamadori, 1977). The adjacent laterodorsal nucleus is also related to the visual system, with the ventrolateral region receiving an input from ipsilateral areas Oc1, Oc2L, and Oc2M (Takahashi, 1985), as well as from the pretectum (Robertson, 1983). Its dorsomedial part is innervated from cortical area 4, rostral to area Oc2M. The connections of the LP are summarized in Table 6. D-Aspartate is a possible candidate for the neurotransmitter involved in the reciprocal connections between LP and Oc (Johnson and Burkhalter, 1992; see also Chapter 22 for a discussion on the amino acid transmitter that may be involved in corticothalamic actions). The ultrastructure of LP has been investigated thoroughly in the hamster by Crain and Hall (1980). Two different types of terminals originate in SC, suggesting that they may arise from different cell types. In the rat, both wide- and narrow-field vertical cells deep in superficial gray have been reported to project to the LP (Mason and Groos, 1981), although Lane et al. (1997) located SC-LP cells in the optic layer, noting that the majority contained calbindin with only a minority reacting for parvalbumin. In the LP of the hamster, terminals from Oc have also been subdivided into two classes, suggesting that they arise in different types of cells (Crain and Hall, 1980); in the rat, cells in layer 6 of Oc2L, as well as in layer 5 of Oc1 and Oc2L, project to LP (Mason and Groos, 1981; Schober, 1981). In the hamster, despite earlier suggestions of dense terminal foci for retinal axons, Ling et al. (1997) demonstrated that preterminal and terminal specializations extend throughout LP. F terminals represent a mixed group and may perhaps include presynaptic dendrites (Crain and Hall, 1980). In the rat, some F terminals appear to

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originate in the most posterior region of the Rt (Sumitomo et al., 1988), which has reciprocal relations with LP (Kolmac et al., 2000; Schober, 1981). Like P cells in DLG, cells in LP respond with a single spike to an electrical stimulus to the optic nerve, followed by 200–250 ms of intense inhibition, terminating in an oscillatory recovery, as is consistent with the relationships with the Rt referred to earlier. The initial latency is, however, longer than that of cells in DLG, suggesting perhaps indirect activation via SC in some cases. Cells in LP can be activated antidromically from area Oc1 as well as Oc2L (Sumitomo et al., 1988).

VISUAL CORTEX Cytoarchitectonic and Visuotopic Organization In a series of electrophysiological studies, Montero et al. (1973a, 1973b; for review, see Montero, 1981) distinguished seven visuotopically organized areas in the occipital cortex of the rat. The largest of these areas— their primary visual area—includes area 17 (striate area) distinguished by Krieg (1946a, 1946b) on the basis of qualitative cytoarchitectonic criteria. On the one hand, the primary visual area of Montero and co-workers is larger than Krieg’s area 17 and the primary visual cortex determined electrophysiologically by others (Adams and Forrester, 1968; La Messurier, 1948). On the other hand, it corresponds well to area 17 distinguished on the basis of cytoarchitectonic criteria (Schober, 1986; Schober and Winkelmann, 1975), as well as to the cortical region receiving a direct input from DLG (Ribak and Peters, 1975; Schober and Winkelmann, 1977) and to Oc1 distinguished on the basis of quantitative cytoarchitectonics (Reid and Juraska, 1991; Zilles et al., 1980, 1984; see also Chapter 23). In an aldehyde-perfused brain, the primary visual cortex (area Oc1, area 17, striate cortex, area V1) of the rat constitutes one part of a typical granular cortex (koniocortex) with a prominent layer 4, about 1490 μm in thickness (Peters et al., 1985). In Nissl-stained sections, Oc1 appears striated: the greater the cell density, the darker its appearance and the striation is mainly due to a high density of cells in layers 4 (granular) and 6 (infragranular) and a low density of cells in layers 5 (infragranular) and 2/3 (supragranular). On its lateral and medial aspects, area 17 (Oc1) is bounded, respectively, by cytoarchitectonic areas 18a (Oc2L) and 18b (Oc2M). In contrast with area 17, the density of neurons in layers 2 to 6 of areas 18a and 18b is fairly even. In area 18a, but not in 18b, supragranular layer 2 can be distinguished from layer 3, and the packing density of cells in layer 5 of area 18a is lower than that in 18b

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(Krieg, 1946b; Miller and Vogt, 1984a, 1984b; Schober and Winkelmann, 1975). Finally, infragranular layers 5 and 6 occupy the lower half of the cortical thickness in area 18a but the lower two-thirds in area 18b (Miller and Vogt, 1984a, 1984b; Schober and Winkelmann, 1975). As shown in Fig. 2E, the topographic organization of the primary visual area (Oc1, striate cortex, area 17 or V1) is such that the superior visual field (lower retina) is represented caudally, the nasal visual field (temporal retina) is represented laterally, and the vertical meridian is at the lateral border of Oc1. The lateral part of Oc1 constitutes the binocular field, whereas in its medial part, only the monocular (contralateral eye) field is represented (Adams and Forrester, 1968; Montero, 1973; Thurlow and Cooper, 1988). A potential elicited by stimulation of the ipsilateral eye can be recorded in a restricted region lying laterocaudally in Oc1 (Sakai et al., 1983). The surface area of Oc1 varies from about 7.1 mm2 (Espinoza and Thomas, 1983) to about 9.4 mm2 (Peters et al., 1985; Schober and Winkelmann, 1975), constituting about 50–60% of the combined area of all visual cortical regions and about 10–12% of the entire neocortex (Espinoza and Thomas, 1983). Two groups of investigators, Montero and co-workers and Espinoza and Thomas (1983; cf. also Thomas and Espinoza, 1987), distinguished six other visuotopically organized areas, located in cytoarchitectonic areas 18 (18b) and 18a of Krieg (1946a, 1946b) and Schober and Winkelmann (1975) but including some of Krieg’s area 7. The map by Espinoza and Thomas is illustrated in Fig. 2; it differs in several important details from that of Montero and co-workers. Although Espinoza and Thomas confirmed the presence of four visuotopic areas in Oc2L (cytoarchitectonic area 18a of Krieg), they did not confirm the existence of the fifth area reported by Montero and co-workers in Oc2L. However, medial to the primary visual area (within Oc2M or cytoarchitectonic area 18b), Espinoza and Thomas reported that an additional area is located posteriorly, behind the area described by Montero and co-workers. There is a problem in correlating closely the cytoarchitecture of the visual cortex with its functional subdivisions. In particular, both cytoarchitectonic areas 18a (Oc2L in the combined cytoarchitectonic and myeloarchitectonic nomenclature of Zilles and colleagues) and 18b (Oc2M) are substantially larger than the regions containing visuotopically organized areas within them. Several authors, using techniques of degeneration, autoradiography, and retrograde labeling with HRP, have demonstrated that all the visuotopically organized cortical areas located outside the primary visual cortex are connected reciprocally with visuotopically (retinotopically) corresponding regions of the primary visual area (Coogan and Burkhalter, 1990; Montero

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et al., 1973; Olavarria and Montero, 1981; Sanderson et al., 1991). In addition, each of the regions distinguished by Montero and co-workers and by Espinoza and Thomas (1983) projects directly to the retinotopically corresponding regions of the ipsilateral SC (Harvey and Worthington, 1990; Olavarria and Van Sluyters, 1982; Thong and Dreher, 1986).

Intrinsic Organization of Primary Visual Cortex (Oc1, Striate Cortex, Area 17, Area V1) Cellular Organization In area Oc2L, there is an abundance of both pyramidal cells with their characteristic apical dendrites extending to layer 1 and nonpyramidal (stellate) neurons that are functionally and morphologically heterogeneous. About 80% of the population, are pyramidal cells (see Peters and Kara, 1984; Peters and Kara, 1984a, 1984b; Peters et al., 1985; see also Chapter 32. Although nonpyramidal neurons are present in all layers (Johnson et al., 1988; Peters et al., 1985), they clearly concentrate in layers 2/3 and 4 (Parnavelas et al., 1977; Peters et al., 1985; Werner et al., 1982). Over 90% of neurons in area 17 are rich in cytoplasm. The proportion of neurons with a small amount of cytoplasm, presumed to be predominantly interneurons, is much higher in superficial layers (Werner et al., 1982, 1985). The medial (monocular) part of Oc1 (Oc1m) differs cytoarchitectonically and myeloarchitectonically, as well as in the distribution of acetylcholinesterase from the lateral binocular subfield (Oc1L) (Zilles et al., 1984). In comparison, layer 4 in Oc1M is thicker and the density of granule cells is greater; unlike in Oc1L, layer 6 in Oc1M is thicker than the combined layers 2 and 3 (Reid and Juraska, 1991; Zilles et al., 1984). Several groups of investigators, especially Peters and collaborators, using autoradiography, degeneration, and electron microscopy of neurons identified by gold-toned Golgi impregnation, have provided much detailed information on the cortical inputs and synaptic connectivity of the primary visual area of the rat (Feldman and Peters, 1978; Peters and Feldman, 1976, 1977; Peters et al., 1976, 1979, 1982; Peters and Kimerer, 1981; Peters and Proskauer, 1980; Peters and Saldanha, 1975). The great majority of thalamic afferents (principally arising in DLG) terminates in layer 4 and lower layer 3, although some of the geniculocortical afferents reach layers 1 and 6. Current source density analysis reveals that layers 4 and 6 receive monosynaptic inputs possibly via collaterals of the same geniculocortical axons (Bode-Greuel et al., 1987). It appears that geniculocortical axons form asymmetrical synapses on all neural elements in layer 4 and the lower part of layer 3. In layer 4 and in the lower part of layer 3, 83% of the

thalamic terminals synapse with (1) dendritic spines of apical dendrites and collaterals of layer 5 and 6 pyramidal neurons; (2) dendritic spines of basal dendrites of pyramidal cells located in layer 3; (3) sparsely spined stellate cells; (4) spiny nonpyramidal cells with perikarya are located in layers adjacent to layer 4 but with dendrites extending into layer 4; and (5) dendritic spines of pyramidal cells located in layer 4. About 15% of thalamic terminals synapse on the shafts of apical dendrites of layer 3 and 5 pyramids or on the dendritic shafts of smooth stellate cells. Finally, 2% of the thalamic terminals in layers 4 and lower 3 terminate on the perikarya of sparsely spined or smooth stellate cells. The transmitter(s) used by geniculocortical axons may be excitatory amino acids, such as glutamate and aspartate (Johnson and Burkhalter, 1992; Kaneko and Mizuno, 1988; see Chapter 36). In particular, relay cells in DLG are retrograde labeled with D-aspartate injected into the middle layers of Oc1 where DLG axons terminate (Johnson and Burkhalter, 1992). Furthermore, relay cells in DLG are immunoreactive to phosphate-activated glutaminase, a major synthetic enzyme of transmitterassociated glutamate (Kaneko and Mizuno, 1988). Staining for zinc in terminal fields reveals two heavy bands in the rat neocortex (including visual cortex); one in deep layer 1 and supragranular layers 2 and 3 and the other in upper layer 5. A moderate to heavy reaction is found in layer 6, whereas lighter bands of zinc-rich terminal fields are apparent in upper layers 1, 4, and deep 5 (Pérez- Clausell, 1996). Overall, the zincrich terminal fields are associated with pyramidal neurons, and pyramidal neurons appear to give rise to zinc-rich projections (Pérez-Clausell, 1996; CasanovasAguilar et al., 1998). Based on the measurements of area by Espinoza and Thomas (1983) and Schober and Winkelmann (1975) and the neuronal counts of Rockel et al. (1980), as well as their own neuronal counts and estimates of area, Peters and colleagues (1985) calculated that there are about 850,000–1,128,000 neurons in the primary visual area (Oc1) of the rat. Because there are about 16,000–18,000 relay cells in the DLG (see page 1085), it appears that there are about 50–60 cells in the primary visual cortex of the rat for every relay cell in DLG. Interestingly, in two mammalian species with visual systems more highly developed than that in the rat, the ratio of cells in the primary visual cortex to relay cells in DLG is either slightly (cat about 65:1) or substantially greater (macaque, about 160:1) (Beaulieu and Colonnier, 1983). There are clear sex differences in the overall size of the rat visual cortex (Reid and Juraska,1992a). In particular, the binocular region of Oc1 adult Long–Evans hooded male rats has 19% more neurons than the corresponding region in females (Reid and Juraska, 1992b; Nuñez et al., 2002). A similar difference in the number

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of neurons between adult Long–Evans hooded males and females has been reported for the monocular region of Oc1 (Nuñez et al., 2002). The differences in numbers are apparently related to the activity of peripubertal ovarian hormones (Nuñez et al., 2002). It is not known to what extent, if any, the numerical differences apply to all strains and if the differences are largely specific to particular neuronal classes. Furthermore, it is not known if these differences are reflected in the functional properties of Oc1 neurons and/or gender-related visual capabilities. While all pyramidal neurons have numerous spines on their apical dendrites, only some of the morphologically more diverse groups of nonpyramidal cells have spines (Fig. 6). Three or more dendrites of large and, to a lesser extent, medium pyramidal cells of lamina 5 form oriented clusters. The average center-to-center spacing between the clusters is about 50 μm. In the supragranular laminae, the apical dendrites of pyramidal cells located there are added to the clusters. In contrast, the apical dendrites of small pyramidal cells located in aminae 4 and 6a do not contribute to the clusters (Peters and Kara, 1987). Spinous stellate cells (SPS in Fig. 6) are usually located in layer 4 and seem to receive afferents from DLG onto their spines (Feldman

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and Peters, 1978). Their axons form asymmetric (presumably excitatory) synapses and some of the spinous stellate cells, such as pyramidal cells, send their axons outside Oc1. There is strong evidence indicating that many cortical projection neurons (pyramidal and at least some spinous stellate cells) use glutamate and/or aspartate as excitatory transmitters (Dinopoulos et al., 1989; Dori et al., 1989; Emson and Hunt, 1981; Fosse and Fonnum, 1987; Johnsonand Burkhalter, 1992). In particular, antibodies staining the transmitter-related (rather than the metabolic-related) pool of glutamate and aspartate label pyramidal neurons in layers 2–6. While glutamatelabeled neurons are mainly concentrated in layers 2 and 3, aspartate-immunoreactive neurons are mainly pyramidal neurons in layers 2–6. In the visual cortex of the cat, zinc-rich boutons are associated with pyramidal cells and exhibit much higher glutamate-like immunoreactivity than the background levels (Beaulieu et al., 1992). In the visual cortex of the rat, about 10% of aspartateimmunoreactive cells are nonpyramidal neurons scattered through cortex. Layer 5 neurons immunoreactive to glutamate or aspartate tend to have larger somata than those in other layers (especially those in layers 4 and 6). Furthermore, aspartate-immunoreactive cells in

FIGURE 6 Summary diagram drawn from the papers of Peters and collaborators cited in Section V, B, indicating some of the features of thalamocortical and intracortical connections in area Oc1. Cortical layers are indicated by Roman numerals on the left. B, bipolar cell; C, chandelier cell; M, multipolar cell; NSS, nonspiny (smooth) stellate cell; P, pyramidal cell; Pr, bitufted projection cell with a myelinated axon; SpS, spiny stellate cell; SSM, sparsely spinous multipolar cell. Filled terminals, excitatory; open terminals, inhibitory. Thal. aff., thalamic afferents, mainly geniculocortical. Efferents (eff.) from layer 5 pyramidal cells project to SC, LP, and VLG, as well as to ipsilateral areas Oc2L and Oc2M and contralateral homotopical and heterotopical visual cortices. Pyramidal cells in layer 6 (not shown) project to DLG and Rt, as well as ipsilateral and contralateral visual cortices; pyramidal cells in layers 2 and 3 project to other visual cortical areas both ipsilaterally and contralaterally. Commissural neurons in layers 5 and 6 do not send collaterals to subcortical structures such as SC, VLG, LP, DLG, or pontine nuclei (Hallman et al., 1988, cf. Section VI,D,1). Pyramidal cells use glutamate and/or aspartate as neurotransmitters. Bipolar, chandelier, multipolar, and smooth stellate cells are mostly GABAergic; a substantial proportion of those GABAergic neurons are also immunoreactive to antibodies against choline acetyltransferase, as well as neuropeptides such as somatostatin and vasoactive intestinal polypeptide (VIP). For references, see text.

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Oc1 tend to have larger somata than those immunoreactive to glutamate. The axons of both glutamate- and aspartate-immunoreactive cells form asymmetrical (presumed excitatory) synapses on dendritic spines (Dori et al., 1989). A “honeycomb-like” mosaic of modules less then 100 μm wide at the border of layers 1 and 2 of the visual cortex has been described (Ichinohe et al., 2003). The walls and hollows of the “ honeycomb” consist, respectively, of zinc-enriched corticocortical terminals (and parvalbumin-dense neuropil) and thalamocortical terminations labeled by cytochrome oxidase and antibody against vescicular glutamate transporter 2. The corticocortical and thalamocortical terminations might constitute parallel circuits at the level of layer 2 associated with different dendritic systems (Ichinohe et al., 2003). The majority of nonpyramidal neurons, which represent about 15% of the entire neuronal population of visual cortex, are likely to be GABAergic, as they are GAD immunoreactive (Lin et al., 1986; Ribak, 1978) and GABA immunoreactive (Meinecke and Peters, 1987). Although GAD-immunoreactive neurons are most numerous in lamina 4, many of them are located in supragranular layers, as well as in the ventral half of layer 6, layer 5, and in white matter below layer 6. Indeed, although the GAD activity is highest in layer 4, both supragranular and infragranular layers display quite substantial GAD activity (McDonald et al., 1987). Somal sizes of GAD-immunoreactive cells vary from 50 to 240 mm2. Cells in layers 2 and 3 are larger than cells in other layers and those in layer 6 tend to have the smallest somata (Lin et al., 1986). The majority of GAD-immunoreactive neurons are multipolar or bipolar, with smooth or sparsely spinous dendrites (Lin et al., 1986; Ribak, 1978). The density of GAD-immunoreactive puncta (presumed synapses of GABAergic neurons) is highest around the dendrites of pyramidal cells in layer 4 and is less in the superficial half of layer 1 and the border between layer 6 and the white matter. Some less dense GADimmunoreactive terminals are seen around the somata and basal dendrites of pyramidal cells in layer 5. In addition, a substantial proportion (15–20%) of presumed GABAergic neurons is also immunoreactive to antibodies against the neuropeptide somatostatin. Some neurons immunoreactive to both GAD and somatostatin are also found in the deep part of layer 6 and in the adjoining white matter (Lin et al., 1986). GAD- and somatostatin-immunoreactive neurons are usually multipolar neurons located in layers 2 and 3 (Schmechel et al., 1984; Lin et al., 1986). Gonchar and Burkhalter (1997) studied directly the distribution and morphology of cells immunoreactive

to GABA in the primary visual cortex in the rat. They concluded that there are at least three distinct families of GABAergic neurons. One family is constituted by nonpyramidal cells that are not only immunoreactive to GABA, but also to the calcium-binding protein parvalbumin (PV). A small proportion (5.3%) of GABAergic neurons immunoreactive to PV are also immunoreactive to calbindin (CB), Furthermore, all PV-immunoreactive neurons are also GABAergic and are present in all layers but layer 1. Cortical cells immunoreactive for both GABA and PV have multipolar somata and account for almost 51% of GABAergic neurons. The second group of GABAergic neurons are nonpyramdial, expressing somatostatin (SOM). The great majority (over 86%) of SOM-expressing GABAergic neurons coexpresses CB and a small proportion (2%) coexpresses nitric oxide synthase (NOS). Virtually all NOS-expressing GABAergic cells coexpressed SOM and almost 90% of cells expressing SOM also express GABA. SOMimmunoreactive GABAergic neurons have usually bitufted or multipolar morphologies and are located mainly in infragranular layers 5 and 6, being completely absent from layer 1. Finally, SOM-immunoreactive GABAergic neurons are among the largest GABAergic neurons, and these cells constitute about 17% of GABAergic neurons. The third family of GABAergic neurons in rat’s area Oc1 coexpresses calretinin (CR). About 94% of CR expressing neurons is GABAergic, and these cells constitute about 17% of GABAergic neurons in the striate cortex of the rat. The CR expressing neurons are multipolar, bipolar, or bitufted, but not pyramidal; they do not coexpress PV, SOM, CB, or NOS. Unlike PV expressing axons and dendrites, the axons and dendrites expressing CR are numerous in layer 1 and generally the CR-expressing GABAergic cells are mainly found in the supragranular layers. In Golgi material, several types of nonpyramidal neurons are fairly frequently impregnated. Multipolar cells (M, Fig. 6) have smooth or sparsely spinous dendrites and are located mainly in layer 4, receiving excitatory inputs from geniculocortical axons. There is evidence that they use GABA as their neurotransmitter (Peters and Fairén, 1978; Ribak, 1978) and their axon terminals form symmetrical (presumed inhibitory) synapses on the cell bodies of some pyramidal and nonpyramidal neurons (Peters and Proskauer, 1980). Bipolar cells (B in Fig. 6) are encountered throughout layers 2 to 5 and have vertically elongated, narrow dendritic trees that may traverse the cortex from layers 2 to 5. They receive an excitatory input from DLG, and some of the axons of these neurons in the visual cortices of cat and rat appear to form asymmetric (presumed excitatory) synapses with the dendritic spines of apical dendrites of pyramidal neurons (Fairén et al., 1984;

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Peters and Fairén, 1978; Peters and Kimerer, 1981). Substantial evidence indicates that bipolar cells might use vasoactive intestinal polypeptide (VIP) as a cotransmitter (Connor and Peters, 1984; Emson and Hunt, 1981; Hajos et al., 1988a, 1988b; Loren et al., 1979; McDonald et al., 1982; Morrison et al., 1984; Peters, 1990; Peters and Harriman, 1988; Peters and Kimerer, 1981; Sims et al., 1980). It turns out, however, that apart from bipolar cells that form asymmetrical synapses, up to 50% of bipolar cells in the visual cortex of the rat form symmetrical synapses, either on the shafts of the smooth dendrites of nonpyramidal cells and the spiny dendrites or on cell bodies of pyramidal neurons (Peters and Harriman, 1988, 1990). Furthermore, terminals labeled with VIP contain closely packed pleomorphic vesicles, a feature characteristic of axon terminals in which the peptides coexist with GABA (Peters, 1990). Thus it appears that many VIP-positive bipolar neurons have an inhibitory function. They could play a role in inhibiting narrow modules of pyramidal cells (see later). It is likely that vertically oriented clusters constitute functional modules made up of the apical dendrites of pyramidal neurons, together with local associational neurons such as bipolar and multipolar neurons. Different combinations of such neuronal modules are excited by various inputs, e.g., arising from the DLG or the contralateral cortex (Peters and Kara, 1987). A proportion of bipolar cells mainly in layers 2 to 4, apart from being immunoreactive to antibody specific to VIP, are also immunoreactive to antibody against ChAT—the enzyme synthesizing acetylcholine (Eckenstein and Thoenen, 1983; Eckenstein et al., 1988; Parnavelas et al., 1986). ChAT-labeled intrinsic neurons in the visual cortex usually form symmetrical (presumed inhibitory) synapses on the dendrites of ChATnegative neurons and some are multipolar cells located in layers 5 and 6 (Parnavelas et al., 1986). Chandelier cells (C, Fig. 6) have smooth or sparsely spinous dendrites and cell bodies within layers 2 and 3; they tend to congregate at the borders of Oc1 and Oc2L and, to a lesser extent, between Oc2L and Oc2M. Chandelier cells are probably inhibitory (GABAergic) and their axonal terminals form symmetrical synapses with the initial segments of axons of pyramidal cells in layers 2 and 3 (Peters et al., 1982). Intrinsic Intralaminar and Interlaminar Connections within Area Oc1 There is a well-developed network of clustered intrinsic lateral interconnections within Oc1 (Burkhalter, 1989; Burkhalter and Charles, 1990; Casanovas-Aguilar et al., 1998; Divac et al., 1987; Miller and Vogt, 1984a; Rumberger et al., 2001). Intrinsic interconnections in the

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rat’s visual cortex, like those in other mammals (see, for review, LeVay, 1988), appear to be excitatory, as they employ excitatory amino acids as neurotransmitters (Johnson and Burkhalter, 1992). In contrast to primary visual cortices of cats and primates (Hubel, 1982; LeVay, 1988), there is no evidence of orientation columns in primary visual cortices of nocturnal rodents such as mice (Métin et al., 1988), hamster (Tiao and Blakemore, 1976a), or rat (Girman et al., 1999). It is unlikely therefore that intrinsic clustered interconnections within area Oc1 of the rat (unlike these in the cat and macaque monkey; for review, see Gilbert, 1998) involve visuotopically noncorresponding regions of Oc1, which contain neurons with similar orientation selectivity. Indeed, the parts of area Oc1 that are interconnected intrinsically contain the neurons whose receptive fields are at least partially overlapping and therefore retinotopic (visuotopic) space is represented without a break across the entire field of connected patches within area Oc1 (Rumberger et al., 2001). However, it is likely that the intrinsic clustered interconnections within area Oc1 of the rat, like those in primary visual cortices of cat and macaque monkey, underlie the dynamic nature of cortical receptive fields (see Gilbert, 1998) and the remarkable ability of the mammalian primary visual cortex to reorganize itself following retinal and/or cortical damage (for reviews, see Eysel et al., 1999; Dreher et al., 2001). Layer 4 cells project strongly and in a topographically precise manner to lower layers 2/3 as well as weakly and in a somewhat more diffusely horizontally extended manner to upper layers 2/3 (Burkhalter, 1989). There are also narrow, vertically oriented projections from layer 4 to infragranular layers 5 and 6 (Burkhalter, 1989; Burkhalter and Charles, 1990). In contrast, cells in layers 2/3, 5, and 6 tend to make fairly wide, horizontally extended connections. Cells in layers 2/3 make widespread connections within the same layer and project strongly to lamina 5 (Burkhalter, 1989), with many being constituted by collaterals of fibers projecting to other visual cortical areas in the same hemisphere or to the contralateral Oc1 (Burkhalter and Charles, 1990). Pyramidal neurons in layer 5 receive monosynaptic excitatory input from layers 2/3 and from the lower part of layer 6 (Kenan-Vaknin et al., 1992). The most prominent projections from the upper part of layer 5 are to layers 2/3 and 6, whereas cells in lower layer 5 make wide-ranging clustered projections to layer 1, the bottom of layers 2/3, and the top of layers 4 and 5. The axonal terminals of neurons in layer 5 form patches that are 130–160 μm wide, separated by 230–260 μm. Cells in lower layer 6 make clustered projections to the layer 3/4 border and to layers 1 and 2, as well as to the layer 5/6 border. Their

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terminals form patches 190–220 μm wide, separated by 320–390 μm (Burkhalter, 1989). The associational neurons with long-ranging intrinsic projections within Oc1 are located in layers 2/3, 5, and 6 (Burkhalter, 1989; Burkhalter and Charles, 1990; Divac et al., 1987; Miller and Vogt, 1984a). In contrast, subcortically projecting neurons in layers 5 and 6 (see later) do not contribute to long-range projections and their local axonal collaterals appear to be confined to single functional cortical columns (Burkhalter and Charles, 1990; Hübener and Bolz, 1988).

Receptive Field Properties of Neurons in Visual Cortex There are relatively few published studies on the receptive field properties of neurons in the rat’s Oc1 (Burne et al., 1984; Fagiolini et al., 1994; Girman et al., 1999; Montero, 1981; Parnavelas et al., 1981, 1983; Shaw et al., 1975; Wiesenfeld and Kornel, 1975). About half of the population of cells responds well to flashing uniform stationary on–off stimuli, the other half responds poorly or not at all to such stimuli. The latter cells respond best to optimally oriented moving gratings of an appropriate spatial frequency. The upper limit of spatial resolution of single neurons in Lister hooded rats is 1.2 cycles per degree (Girman et al., 1999). Around 75–80% of cells in the primary visual cortices of hooded rats are orientation selective (Montero 1981; Parnavelas et al., 1981; Fagiolini et al., 1994; Girman et al., 1999). Fagiolini and colleagues (1994) reported that at postnatal days 19–21 the average size of the receptive fields of neurons in the primary visual cortices of hooded Long Evans rat pups is much greater than these of cells recorded on postnatal day 45 or in adulthood (34º with SD-6º vs 6.7º with SD-2.5º). Furthermore, the reduction in the average size of the receptive field of cortical neurons correlates well with the improvements in visual acuity observed in the same period (0.51 cycles per degree vs 1.1 cycles per degree). Around 75–80% of cells in the primary visual cortices of hooded rats are orientation selective (Montero 1981; Parnavelas et al.. 1981; Fagiolini et al., 1994; Girman et al., 1999). As mentioned in the previous section, there is no evidence of orientation columns in rat’s primary visual cortices (Girman et al., 1999). Girman and colleagues (1999) reported that almost 63% of the orientation-selective cells have an orientation tuning width at half-height of 60º or less. Furthermore, an additional 16% of cortical neurons were orientation biased. Thus, at least in hooded rats, almost 95% of cells in the primary visual cortices appear to exhibit some orientation selectivity. About 35% of orientationselective or orientation biased cells show preference

for bars and/or gratings oriented horizontally. As mentioned in the previous section, there is no evidence of orientation columns in rat’s primary visual cortices (Girman et al., 1999). Cells in supragranular layers II and III tend to exhibit little background (“spontaneous”) activity, tend to be sharply orientation selective, and respond to stimuli of relatively low temporal frequencies (low velocities) and relatively high spatial frequencies. Cells in the granular layer IV tend to exhibit high temporal resolution and respond well to relatively high stimulus velocities (over 500º/s) (Girman et al., 1999; see also honeycomb-like mosaic in layers 1 and 2; Ichinohe et al., 2003). Cells in granular layer IV tend to exhibit high temporal resolution and respond well to relatively high stimulus velocities (over 500º/s; Girman et al., 1999). A similar relation between laminar location and receptive field properties has been reported for visual cortical cells recorded in another species of murid rodents, the mouse (Dräger, 1975; Mangini and Pearlman, 1980; Métin et al., 1988), and in lagomorphs rabbit; Murphy and Berman, 1979). On postnatal day 17 (i.e., about 3 days after natural eye opening for the first time), virtually all cells in primary visual cortices of hooded Long–Evans pups, which respond to visual stimuli, are not orientation selective. The proportion of orientation-selective cells and their orientation tuning width became adult-like only by postnatal day 45. Furthermore, dark rearing of pups (until postnatal day 60) results not only in poor responsiveness, a tendency to rapid habituation, and low spontaneous activity of cortical neurons, but also in a paucity of orientation and direction-selective cells in area Oc1. The proportion of orientation biased cells in area Oc1 is, however, not significantly reduced by dark rearing in the first 5–6 weeks of postnatal life (Fagiolini et al., 1994). Despite the fact that the ipsilateral retinofugal projection in the rat is rather small (see “Introduction” I), the percentage of binocular cells in the rat primary visual cortex is high (about 80%; Fagiolini et al., 1994), i.e., quite comparable with that in striate cortices of cats (e.g., Hubel and Wiesel, 1962; Burke et al., 1992) or macaque monkeys (Baker et al., 1974). It is likely that the high percentage of binocular neurons is related to numerous intrinsic short-range associational connections between cortical cells whose geniculate input relays information from corresponding regions of contralateral and ipsilateral eyes. There appears to be no dramatic “magnification” of the representation of the ipsilateral retina in the geniculate. Indeed, unlike in striate cortices of cats or macaque monkeys, most binocular cells in primary visual cortices of the rats are dominated by the contralateral eye. On postnatal day 17, binocular cells

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dominated by the contralateral eye constitute the great majority of visually responsive neurons recorded from area Oc1 of Long–Evans hooded pups, and the proportion of monocular cells driven exclusively via the contralateral eye increases slighthy with age (up to postnatal day 45; Fagiolini et al., 1994). In rats, as in cats and macaque monkeys (for reviews, see Movshon and Van Sluyters, 1981; Sherman and Spear, 1982, Wiesel, 1982), there is a clear critical period in which the neurons in primary visual cortices are highly susceptible to monocular visual pattern deprivation (Fagiolini et al., 1994). In the primary visual cortices of hooded Long–Evans rats deprived monocularly of patterned visual stimulation in the period from postnatal day 14 to postnatal day 45, the proportion of binocular cells dropped to about 45% (vs about 80% in normal animals; Fagiolini et al., 1994). Furthermore, the deprived contralateral eye dominated only less than 2% of cortical neurons, whereas the nondeprived ipsilateral eye dominated over 90% of cortical neurons. Cells in the primary visual cortex of the rat are maximally susceptible to monocular visual deprivation in the fourth postnatal week (Fagiolini et al., 1994). Cells in the primary visual cortex of the rat (e.g., Bourne et al., 1984; Parnavelas et al., 1981, 1983; Shaw et al., 1975; Wiesenfeld and Kornel, 1975) can be classified as simple (S), complex (C), or end stopped (hypercomplex) using criteria developed in cats and monkeys (Bishop and Henry, 1972; Dreher, 1972; Henry, 1977; Hubel and Wiesel, 1962,1968; Shottun et al., 1991). Some cells of each type exhibit direction selectivity (Parnavelas et al., 1981, 1983; Girman et al., 1999). Overall, almost 60% of visual cortical cells recorded in rats exhibit either clear directional selectivity or directional bias (Girman et al., 1999). For many cortical neurons, especially those recorded in supragranular layers in the upper third of the cortex, the magnitude of responses is reduced substantially when stimuli extend beyond the discharge field (so-called classical receptive field). In some cases, the reduction in the magnitude of responses is independent of the orientation of the stimuli extending beyond the classical receptive fields. In most cases, however, the level of suppression of the responses is dependent on the orientation of the stimuli outside the classical receptive field: the suppression is usually maximal when the orientation of the stimuli covering both fields is the same (Girman et al., 1999). Only pyramidal cells (identified morphologically) have complex or end-stopped receptive fields, with complex cells being located in layers 2 to 4 and end-stopped cells in layers 2, 3, and 5. In contrast, simple cells may have either pyramidal or nonpyramidal morphology and are found in layers 2, 3, and 4. Of those cells that do not exhibit orientation selectivity, some have on- or

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off-discharge centers with antagonistic surrounds, whereas others have on- or off-discharge centers with silent suppressive surrounds; some do not respond to stationary flashing stimuli, although their responses to moving stimuli are quite vigorous. Nonoriented cells in the rat do not appear to concentrate in any particular layer of the cortex (Parnavelas et al., 1983). In contrast, in two other species of murid rodents (hamster and mouse), cells that are not orientation selective or only weakly so tend to be located in layer 4 and in infragranular layers 5 and 6; in the latter case they tend to project to subcortical nuclei (Klein et al., 1986; Mangini and Pearlman, 1980; Métin et al., 1988; Simmons and Pearlman, 1983). In view of the laminar distribution of geniculocortical terminals and the fact that they terminate on all neural elements in layers 4 and lower 3, which can form asymmetric synapses (see previous section), it appears that not only cells with simple receptive field properties, but also cells with complex and end-stopped properties receive a direct input from the DLG. This argues against a strictly hierarchical model of processing information in the visual cortex (Hubel and Wiesel, 1962, 1968), which assumes that only cells with simple receptive fields receive a direct input from DLG and that they in turn drive complex and end-stopped cells (for reviews, see Stone, 1983; Stone et al., 1979). Furthermore, cells with simple- and complex-like receptive fields have also been recorded from area Oc2L (area 18a; Molotchnikoff and Hubert, 1990). Again, the presence of cells with simple-like properties in area Oc2L, which does not receive a significant thalamic input from DLG but does receive a substantial “feedforward” associational input originating from supragranular layers of area Oc1 (see page 1134), argues against a strictly hierarchical model of processing of information in the rat visual cortex.

Extrinsic Connections of Areas Oc1, Oc2L, and Oc2M Diencephalic, Mesencephalic, and Pontine Connections The connections of the various cortical areas are summarized in Chapter 23, but some interrelationships of particular interest are discussed here. Locations of the cells of origin of the chemical pathways are illustrated in Paxinos et al. (1999a, 1999b). As mentioned earlier, geniculocortical fibers form Gray type I (asymmetric) synapses, presumably excitatory, in layer 4 as well as in 6, the lower part of 3, and layer 1 (Peters and Feldman, 1976; Peters and Saldanha, 1975). In both carnivores and primates, the axons of different functional classes of relay cells in DLG terminate to a large extent in different sublayers of layer 4 of Oc1, and the information is

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relayed largely in parallel to different visuotopically organized cortical areas (for reviews, see Garey et al., 1991; Stone and Dreher, 1982; Stone et al., 1979). It is not known in the rat, however, if there is a similar segregation of functionally distinct geniculocortical terminals into different layers and/or visual cortical areas. The projection from Oc1 to DLG (Coleman and Clerici, 1980; Cusick and Lund, 1981, 1982; Hughes, 1977; Jacobson and Trojanowski, 1975; Nauta and Bucher, 1954; Ribak and Peters, 1975) arises from small pyramidal cells in layer 6 (Mason and Groos, 1981; Sefton et al., 1981). Evidence suggests the presence of a monosynaptic loop through layer 6 from and to DLG (Peters and Saldanha, 1975). Oc1 also receives a small thalamic input from the lateral posterior nucleus (LP) and from more rostrally located intralaminar nuclei—centromedial (CM), paracentral (PC), and centrolateral (CL) (Dreher et al., 1990; Herkenham, 1986; Sanderson et al., 1991) and the reuniens nucleus of the midline (Dreher et al., 1990). The principal thalamic input to areas LM in caudal Oc2L (cytoarchitectonic area 18a) and PM in caudal Oc2M (cytoarchitectonic area 18b) originates in the LP; both areas also receive a substantial thalamic input from the posterior nuclear complex (Sanderson et al., 1991). The LP projection to cytoarchitectonic area 18a terminates mainly in layer 4 with a small projection to layer 1 (Schober, 1981; cf. also Coleman and Clerici, 1980; Johnson and Burkhalter, 1992; Olavarria, 1979; Olavarria and Torrealba, 1978). Area Oc2M, but not Oc2L, also receives a smaller thalamic input from the caudalmost part of the laterodorsal nucleus (Van Groen and Wyss, 1992a, 1992b). In addition, cortical area LM but not area PM receives a small thalamic input from DLG, rostral intralaminar nuclei, and the ventromedial nucleus (Sanderson et al., 1991). Projections from the posterior complex and the ventromedial nucleus terminate mainly in layer 1, those from the ventral anterolateral nucleus terminate mainly in layers 1 and 6, whereas those from the centromedial nucleus terminate mainly in layer 6 (Herkenham, 1979, 1980; for review, see Parnavelas and McDonald, 1983). It has been claimed that there is a small input to Oc1, as well as to the other visuotopically organized areas in the rat’s occipital cortex, from the ipsilateral zona incerta of the ventral thalamus (Dreher et al., 1990; Lin et al., 1990). Many of the incertal neurons projecting to Oc1 or other cortical areas, such as those projecting to the intermediate layers of SC, appear to be GABAergic (Lin et al., 1990; Nikolelis et al., 1992, 1997). More recent studies suggest, however, that the incertocortical projections to Oc1 (Mitrofanis and Mikuletic, 1999; Power and Mitrofanis, 1999), as well as these from frontal (motor), parietal (somatosensory), and cingulate (limbic) neocortices (Lin et al., 1990, 1997; Nicolelis

et al., 1992, 1995; Mitrofanis and Mikuletic, 1999; Power and Mitrofanis, 1999), most likely originate from displaced hypothalamic cells scattered among incertal neurons. The support for such interpretations of incertocortical projections stems from the fact that incertal cells projecting to the neocortex (unlike other cells in the given segment of zona incerta) have large multipolar morphologies and express the same type of antigens as cells in the adjacent hypothalamus. Apart from the projection from ectopic hypothalamic cells in the ipsilateral zona incerta, cells in Oc1 and Oc2M also receive inputs from the ipsilateral hypothalamus and dorsomedial nucleus as well as tuberal and posterior lateral areas (Divac, 1975, 1981; Henderson, 1981; Bigl et al., 1982; Lamour et al., 1982; McKinney et al., 1983; Rieck and Carey, 1984; Sloniewski et al., 1984; Tago et al., 1984; Saper, 1985; Sloniewski et al., 1986; Carey and Rieck, 1987; Dreher et al., 1990; Sanderson et al., 1991). Projections from the dorsomedial nucleus and tuberal area might be histaminergic (see page 1093). Cells in the upper and lower visual cortical layer 6 of Oc1, Oc2L, and Oc 2M project to DLG and Rt (Bourassa and Deschênes, 1995). Cells in layer 5 of Oc1, Oc2L, and Oc2M project to SC and to VLG (Olavarria and Van Sluyters, 1982; Sefton et al., 1981); some corticocollicular fibers branch to innervate DLG (Bourassa and Deschênes, 1995). Similar cells, at least in areas Oc1 and Oc2L, project to the LP (Mason and Groos, 1981; Olavarria and Van Sluyters, 1982; Schober, 1981; Sefton et al., 1981). In particular, a substantial proportion of neurons in layer 5 of areas Oc1, Oc2L, and Oc2M project to lateral parts of ipsilateral pontine nuclei (Hallman et al., 1988; Thong, 1991; Wiesendanger and Wiesendanger, 1982a, 1982b) and many corticopontine neurons located in visual cortices also send collaterals to the SC (Hallman et al., 1988; Thong, 1991). Projections from Oc1 to pontine nuclei provide one of several possible routes for visual information to reach the cerebellum. A few cells in Oc2M and Oc1 project to the caudal sector of zona incerta (Benzinger and Massopust, 1983; Nauta and Bucher, 1954; Roger and Cadusseau, 1985; Mitrofanis and Mikuletic, 1999). The projection from Oc1 to zona incerta is much smaller then those from cingulate, frontal, or parietal cortices. (Mitrofanis and Mikuletic, 1999). Occipitoincertal projections, like incertal projections from other cortical areas, originate in lamina 5 (Mitrofanis and Mikuletic, 1999). There is a direct norepinephrinergic input to the visual cortex from the dorsal division of locus coeruleus. In particular, the input to the visual cortex originates from the part of dorsal division of the locus coeruleus, which is located caudally to the part from which the projection to sensorimotor or frontal cortices arises. While the bulk (95%) of the projection originates in the

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ipsilateral locus, a small contralateral projection is also apparent (Sloniewski et al., 1986; Waterhouse et al., 1983). Overall, the activity of neurons in the locus coeruleus and hence the release of norepinephrine in the visual cortex appear to be regulated by the level of attention (Sara and Segel, 1991). Indeed, repeated exposures to the same stimulus result in reduced activations of cells in locus coeruleus (Sara and Segel, 1991). In the adult rat, norepinephrinergic fibers are present in all cortical layers, although innervation of the supragranular layers (especially layer 1) appears to be denser than that of infragranular layers (Latsari et al., 2002, Morrison et al., 1978; Papadopoulos et al., 1989b; Parnavelas et al., 1985). In the visual cortex of newborn rats, norepinephric fibers are present above and below the cortical plate and during subsequent weeks they gradually arborize and innervate all cortical layers (Latsari et al., 2002). Substantial evidence indicates that norepinephrinergic cortical terminals communicate with cortical cells via a nonsynaptic mode (see Beaudet and Descarries, 1978; Séguéla et al., 1990; for review, see Descarries et al., 1991). Consistent with the volume transmission means of neuronal communication (Agnati et al., 1992), the majority of α,2A-adrenergic receptors are located at nonjunctional sites of axons, dendrites, perikarya, and glial processes (Venkatesan et al., 1996). Furthermore, unlike norepinephric terminals, α1-adrenergic receptors concentrate in deep layers (Jones et al., 1985). However, a number of electron microscopic data indicate that a proportion of norepinephrinergic varicosities (0.45 to 2.2 μm in diameter) and terminals in the rat’s visual cortex form conventional symmetrical or asymmetrical synapses with dendritic spines, dendritic shafts (most commonly), or somata of both pyramical and nonpyramidal neurons (Latsari et al., 2002; Papadopoulos et al., 1989b; Parnavelas and Papadopoulos, 1989; Paspalas and Papadopoulos, 1996, 1998; Séguéla et al., 1990). Norepinephric fibers form symmetrical synapses on the perikarya and proximal dendrites of peptidergic interneurons containing somatostatin (SRIF), neuropeptide Y (NPY), or vasoactive intestinal polypeptide (VIP). About 27% of SRIF-immunoreactive neurons, distributed in layers II–VI (but mostly in the infragranular layers), are contacted by norepinephric fibers. The synaptic varicosities on perikarya of SRIF neurons are 0.5–1.6 μm in diameter, round or oval in shape, and contain clear round vesicles. About 25% of NPYimmunoreactive neurons, distributed in layers II–VI (again mostly in the infragranular layers) are contacted by norepinephric fibers. NPY cells are also contacted by multiple varicosities formed by norepinephric fibers en passant. The synaptic varicosities on perikarya of NPY neurons are 0.55–1.75 μm in diameter, round or

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oval in shape, and contain clear round vesicles. Finally, about 20% of VIP-immunoreactive neurons, distributed in layers II–VI, are contacted by norepinephric fibers. The synaptic varicosities on perikarya of VIP neurons are 0.5–1.7 μm in diameter, round or oval in shape, and contain clear round vesicles (Paspalas and Papadopoulos, 1999). Only about 15% of fibers terminate in close proximity to blood vessels (Papadopoulos et al., 1989b; Paspalas and Papadopoulos, 1996, 1998). Norepinephrine appears to suppress horizontal propagation of excitatory transmission and an enhancement of inhibitory transmission to pyramidal cells in supragranular layers II and III of the rat visual cortex (Kobayashi et al., 2000). Iontophoretic application of norepinephrine, at least in some neurons of primary visual cortices of cats (Kasamatsu and Heggelund, 1982; Videen et al., 1984) and rats (Waterhouse et al., 1990), produces an enhancement of visually evoked responses and, at the same time, reduces the background (spontaneous) activity, thus generally increasing “signal-tonoise” ratios. In the superficial layers of sliced preparations of the rat visual cortex, paired-pulse stimulation combined with an application of norepinephrine elicits long-term homosynaptic depression. Thus, it has been proposed that norepinephrine might play an important role in naturally occurring receptive field plasticity (Kirkwood et al., 1999). Both D1 and D2 dopamine receptors are present in the rat’s visual cortex (Boyson et al., 1986). Immunohistochemical studies indicate that there are also dopaminergic terminals in the rat’s Oc1 and Oc2L; in Oc1, however, they are relatively rare and concentrate in layer 6 (Berger et al., 1985; Descarries et al., 1987; Papadopoulos et al., 1989a; Phillipson et al., 1987). The source of these fibers appears to be the ipsilateral and, to a lesser extent, the contralateral mesencephalic ventral tegmental area (Dreher et al., 1990; Phillipson et al., 1987; Saper, 1985; cf. also presumed dopaminergic projections from the ventral tegmental area to visual cortex of the cat, Törk and Turner, 1981). Dopamine-containing varicosities and terminals in the visual cortex form synaptic contacts with dendritic shafts and spines; the synapses are usually asymmetrical (Papadopoulos et al., 1989a). The iontophoretic release of dopamine in Oc1 reduces the “spontaneous” firing rate, as well as responses to full field flashes (Kolta and Reader, 1989). Serotonergic (5-HT) projections to the visual cortex originate from dorsal and median raphe nuclei in the midbrain as well as from B6 and B9 clusters, respectively, in the pons and in the ventrolateral tegmentum of the midbrain (Dahlström and Fuxe, 1964; Törk, 1985). While many of these projections to areas Oc1, Oc2L, and Oc2M are serotonergic (Jacobs et al., 1978; Koh et al., 1991; Kosofsky and Molliver, 1987; Mamanous and

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Molliver, 1988; Newman and Liu, 1987; O’Hearn and Molliver, 1984; Sloniewski et al., 1986; Waterhouse et al., 1986), a substantial proportion is not; most dorsal raphe neurons projecting to areas Oc1 and Oc2L are located close to the midline in the ipsilateral ventromedial caudal segment of the nucleus (Koh et al., 1991; O’Hearn and Molliver, 1984; Waterhouse et al., 1986). A proportion (20–30%) of dorsal raphe neurons projecting to the visual cortex also projects to the ipsilateral or contralateral parafloccular cerebellar cortex (Waterhouse et al., 1986). There is also a smaller projection from the contralateral dorsal raphe, whereas similar numbers of serotonergic projections to the visual cortex from the medial raphe, B6, and B9 clusters originate from ipsilateral and contralateral nuclei (Koh et al., 1991; O’Hearn and Molliver, 1984). Two morphologically distinct networks of serotonergic fibers are apparent in the visual cortex of rat (Kosofsky and Molliver, 1987; Lidov et al., 1980; Papadopoulos et al., 1987) and cat (Mulligan and Törk, 1988). The great majority are fine fibers characterized by small varicosities (0.5–0.8 μm in diameter), whereas the minority are thick fibers with large spherical or elongated varicosities; both types of fibers form conventional symmetrical and asymmetrical synapses with dendritic spines, shafts, and apical dendrites of pyramidal cells (Papadopoulos et al., 1987). Dense plexuses of immunocytochemically identified serotonergic fibers are present in all layers of area Oc1 (Lidov et al., 1980; Papadopoulos et al., 1987). The density of serotonergic fibers, especially those running parallel to the pia, is greatest in the outer part of layer 1 (Lidov et al., 1980; Papadopoulos et al., 1987). However, layers 2 and 3 contain a high density network of mainly radially oriented serotonergic fibers. The density of serotonergic fibers is somewhat lower in layers 4 and 5, whereas in lower layer 6 there is a dense network of serotonergic fibers oriented parallel to the white matter (Papadopoulos et al., 1987). The serotonergic innervation of area Oc2L is less dense than that of area Oc1 (D’Amato et al., 1987). Serotonergic fibers in the rat visual cortex preferentially innervate perikarya, and proximal dendrites of peptidergic interneurons containing SOM or NPY, as well as distal dendrites of interneurons containing SOM but neither synaptic relationship nor close appositions between 5-HT fibers and interneurons containing VIP, were observed (Paspalas and Papadopoulos, 2001). The iontophoretic application of 5-HT at least in area Oc1 usually results in net suppression of maintained activity as well as a reduction in both excitatory and inhibitory responses to visual stimuli and spatial reorganization of the receptive fields of the cortical neurons (Waterhouse et al., 1990). Furthermore, stimulation of 5-HT1A receptors in

slices of the rat visual cortex inhibits the induction of long-term potentiation by suppressing NMDA receptormediated synaptic excitation (Edgawa et al., 1999). Subcortical Telencephalic Connections of Areas Oc1, Oc2L, and Oc2M A small direct cholinergic projection arises from several magnocellular nuclei constituting mainly but not exclusively the Ch4 subgroup of cholinergic neurons (Mesulam et al., 1983) in the basal forebrain (medial septal nucleus, nuclei of the vertical and horizontal limbs of the diagonal band, lateral and medial preoptic area, and substantia innominata). It terminates in Oc1 (Bigl et al., 1982; Divac, 1975; Henderson, 1981; Lamour et al., 1982; McKinney et al., 1983; Price and Stern, 1983; Rieck and Carey, 1984; Saper, 1984; Sloniewski et al., 1986; Tago et al., 1984) and, to a lesser extent, several visuotopically distinct regionss within Oc2L and Oc2M (Carey and Rieck, 1987; Dreher et al., 1990; Sanderson et al., 1991). Contrary to earlier claims, neurons of the basal forebrain are not highly collateralized. Each visual cortical area or subregion receives an input from distinct and separate subpopulations of neurons in the basal forebrain (Carey and Rieck, 1987). The cholinergic fibers form a dense diffuse network that terminates mainly in layers 1,2/3, and 5; the density of cholinergic fibers in layers 4 and 6 is low (Eckenstein et al., 1988; Lysakowski et al., 1986, 1989). However, Parnavelas and colleagues (1983) reported that layers 4, 5, and 6 receive the major cholinergic input. Acetylcholinesterase activity is found primarily in layer 5 and the deep part of layer 4 (Hanes et al., 1992; Robertson et al., 1990; Zilles et al., 1984). Choline Oacetyltransferase (ChAT) activity is highest in layers 5 and 2/3, with lower activity in layers 6,4, and 1 (Walch and Schliebs, 1989). ACh in the rat’s visual cortex appears to act via muscarinic receptors, as M1 and M2 receptors are prominent there (Mash and Potter, 1986). The density of muscarinic receptors is highest in layer 1, followed by layers 2/3 and 6 with low densities of receptors in layers 4 and 5, whereas the density of sodiumdependent, high-affinity choline uptake sites is highest in layer 1, with a more uniform distribution in lower layers (Walch and Schliebs, 1989). Cholinergic terminals stained for ChAT form mainly symmetrical (presumably inhibitory) synapses on dendrites and occasionally on perikarya of noncholinergic, nonpyramidal cells as well as on dendrites or perikarya of ChAT-labeled neurons. However, a small proportion of pyramidal neurons is immunoreactive to antibodies specific to nicotinic ACh receptors. About 15% of layer 5 pyramidal cells projecting to SC display immunoreactivity to nicotinic ACh receptors (Bravo and Karten, 1992). Finally, some cholinergic terminals

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form asymmetrical (presumably excitatory) synapses (Parnavelas et al., 1986). The preponderance of symmetrical (presumed inhibitory) synapses formed by cholinergic terminals is rather surprising in view of the fact that the iontophoretic application of ACh, at least in the cat’s primary visual cortex, usually produces prolonged and marked excitation (McCormick, 1989; Sillito and Kemp, 1983). Overall, the effects of ACh on neurons in the visual cortex are quite specific. Thus, following the iontophoretic application of ACh to the visual cortex of the cat, there is a substantial increase in the degree of direction selectivity and the sharpness of orientation-tuning curves of many neurons without any substantial accompanying increase in their background firing rates (Murphy and Sillito, 1991; Sillito and Kemp, 1983). Therefore, the application of ACh appears to result in a better “signal-tonoise” ratio when neurons are activated by visual stimuli. In the rat, iontophoretic application of ACh to layer 6 cells of the rat primary visual cortex produces a large depolarization accompanied by a train of action potentials (Kenan-Vaknin et al., 1992). In contrast, some neurons in the superficial layers of the cat’s visual cortex are strongly inhibited following local application of ACh (Sillito and Kemp, 1983). Similarly, in the rat, application to cells in layers 2/3 causes hyperpolarization followed by a large depolarization and a train of action potentials (Kenan-Vaknin et al., 1992). Inhibition following the ACh application, however, is not likely to represent a primary inhibitory effect, as in vitro it results from the excitation of local inhibitory interneurons (McCormick and Prince, 1986). In the superficial layers of sliced preparations of the rat visual cortex, paired-pulse stimulation combined with an application of carbachole (activating cholinergic receptors) elicits long-term homosynaptic depression (Kirkwood et al., 1999). Thus, it has been proposed that ACh-like norepinephrine might play an important role in naturally occurring receptive field plasticity (Kirkwood et al., 1999). Local pressure application of the ACh agonist, carbachol, in slice preparations of rat visual cortex reduces intracellularly recorded fast excitatory postsynaptic potentials, as well as both fast and slow inhibitory postsynaptic potentials evoked by the electrical stimulation of white matter (Murakoshi, 1994). The effect of carbachol is antagonized by atropine and is probably presynaptic, as carbachol does not reduce glutamate or γ-aminobutyric acid-induced depolarizations. Overall, ACh appears to modulate neuronal activity in the visual cortex by inducing a long-lasting increase in neuronal excitability in the late phase of synaptic transmissions by a mechanism of reduced inhibitory transmissions (Murakoshi, 1994).

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As in other mammalian species studied so far (for review, see LeVay and Sherk, 1981), Oc1 of the rat receives a substantial input mainly from the caudal part of the ipsilateral claustrum and a smaller input from the caudal part of the contralateral claustrum (Dreher et al., 1990; Jakubowska-Sadowska et al., 1998; Li et al., 1986; Minciacchii et al., 1985; Sadowski et al., 1997; Shameem et al., 1984; Sloniewski et al., 1986; cf., however, Carey and Rieck, 1987; Miller and Vogt, 1984a). The claustral projection to the visual cortex has been reported to originate mainly in the posteroventral part of the claustrum (Jakubowska-Sadowska et al., 1998; Li et al., 1986; Minciacchii et al., 1985; Sadowski et al., 1997). However, Shameem and her colleagues (1984) indicated that ipsilateral projections originate mainly from the dorsoventral claustrum, whereas Carey and Neal (1985) and Sloniewski et al. (1986) did not see any dorsoventral gradient among claustral neurons projecting to the ipsilateral visual cortex. Oc 2M has also been reported to receive a substantial claustral input, and this projection seems to be numerically even larger than that to Oc1 (Carey and Neal, 1985; Dreher et al., 1990; Miller and Vogt, 1984a). More recently, however, claustral input to area Oc 2L has also been noted; furthermore, virtually all claustral neurons projecting to Oc1 also project to areas Oc2M and Oc 2L (Jakubowska-Sadowska et al., 1998). In contrast, only about 1% of claustral cells projecting to visual cortex send axonal collaterals to the ipsilateral cingulate cortices, whereas about 10% send axonal collaterals ipsilaterally to the frontal cortices and to both frontal and cingulate cortices (Li et al., 1986). Finally, claustral neurons projecting to visual cortical areas do not send collaterals to the primary somatosensory cortical areas (Jakubowska-Sadowska et al., 1998). However, after injections of selenite salts (allowing tracing of specific zinc-rich presumably glutaminergic pathways) into the visual cortices, only very few labeled neurons (one or two per section) were present in the ipsilateral claustrum (Casanovas-Aguilar et al., 1998). The Oc layer in which the claustral afferents terminate is not known, although in other mammalian orders, they terminate in layer 4. A corticoclaustral projection has not been demonstrated in the rat, but in view of the fact that corticoclaustral connections are reciprocal in all mammalian species studied so far, it seems very likely that Oc1 also projects to the caudal claustrum. In other species, this projection arises in layer 6 (LeVay and Sherk, 1981). While the functional role of these interconnections remains obscure, some evidence indicates that claustral cells projecting to area 17 of the visual cortex in the cat play a significant role in regulating the length sensitivity of end-stopped cells in area 17 (Sherk and LeVay, 1983). However, because claustrocortical

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connections in the rat are quite divergent, they might be involved in the integration of visual, motor, and limbic information (cf. Li et al., 1986). Associational Interareal Connections of Visuotopically Organized Cortices Using degenerative techniques, Nauta and Bucher (1954) concluded that there are projections from Oc1 (cytoarchitectonic area 17) to both Oc2L and Oc2M, with the projection from Oc1 to Oc2L being reciprocal. Since then, area Oc1 has been shown to have reciprocal associational connections with virtually all visuotopically organized areas (cf. Espinoza and Thomas, 1983; Thomas and Espinoza, 1987) within areas Oc2L and Oc2M (respectively, cytoarchitectonic areas 18a and 18b, Miller and Vogt, 1984a; Montero, 1981, 1993; Montero et al., 1973a; Montero and Guillery, 1968; Olavarria and Montero, 1981, 1984; Sanderson et al., 1991; Schober et al., 1976). Furthermore, projections from area Oc1 to extrastriate visual areas are organized visuotopically (Montero, 1993). Rumberger and colleagues (2001), however, argued that although the “feedforward” (see further) projections from the single region in the primary visual cortex to areas Oc2l and Oc2M are patchy, this “patchiness” does not necessarily indicate the existence of a number of distinct visuotopically organized areas within areas Oc2L and Oc2M but might be related to the presence of functional subunits within these areas. The pattern of associational interconnections of visual cortical areas is summarized in Fig. 7A1. The associational projection from area Oc1 to area LM in the caudal part of Oc2L is numerically the largest (Sanderson et al., 1991). Virtually all associational cells in Oc1, including those projecting from area Oc1 to area Oc2L (Coogan and Burkhalter, 1990), are pyramidal neurons whose somata are located mainly in supragranular layer 2/3 with a smaller proportion located in infragranular layers 5 and 6 and a few in layer 4 (Miller and Vogt, 1984a; Sanderson et al., 1991). This tendency of associational neurons projecting from Oc1 to be located in supragranular layers (“feedforward” projections) rather than in infragranular layers (“feedback” projections, see following section) is particularly apparent in the projection from area Oc1 to PM (within Oc2M) in which about 85% of associational neurons are located in supragranular layer 2/3. In the case of projections from area Oc1 to area AM (within Oc2M) and PL (within area Oc2L), only about 70% of associational neurons are located in layer 2/3 (Sanderson et al., 1991). Associational fibers projecting from area Oc1 to areas Oc2L and Oc2M terminate most heavily in layers 2/3 and 4, with fewer in the deep part of layers 1, 5, and the upper part of 6 (a feedforward projection; Coogan and Burkhalter, 1990, 1993).

Area Oc1 receives its major associational “feedback” inputs from visuotopically organized area LM (within Oc2L) with smaller inputs from visuotopically organized area PM in Oc2M, area AL in Oc2L, and the smallest input from area AM in Oc2M (Dreher et al., 1985b; Miller and Vogt, 1984a; Montero, 1981; Olavarria and Montero, 1981, 1984; Sanderson et al., 1991; Schober et al., 1976; Torrealba et al., 1984). It is worth noting that a substantial proportion (about 30%) of associational neurons in area Oc2L projecting to ipsilateral Oc1 also projects to the contralateral areas Oc1, Oc2L, and Oc2M; thus they represent associational–commissural neurons that are located in all layers except layer 1 and lower layer 6 (Dreher et al., 1990). Projections to Oc1 from all visuotopically organized areas, however, originate predominantly in infragranular layers 5 and 6 (feedback projections, see following section) rather than from supragranular layers 2/3 (Miller and Vogt, 1984a; Sanderson et al., 1991). The trend is most apparent in the projection from area LM (within Oc2L) to Oc1 in which 65% of associational neurons are pyramidal cells located in infragranular layers 5 and 6 while the remainder are pyramidal cells in supragranular layer 2/3 (Sanderson et al., 1991). Like other feedback projections, they terminate mainly in layer 1 and, to a lesser extent, in layers 2 and 5 of Oc1; associational terminals are virtually absent from layers 3 and 4 of area Oc1 (Coogan and Burkhalter, 1990, 1993). Associational Connections of Visuotopically Organized Cortices with Nonvisuotopically Organized Cortices The associational connections of the visuotopically organized areas in Oc with other regions of the neocortex are summarized in Fig. 7A2. Note first that, with the exception of area LM within Oc2L, each area receives inputs from the caudal third of the frontal eye field of motor cortex (cytoarchitectonic area 8; Dreher et al., 1990; Miller and Vogt, 1984a; Sanderson et al., 1991; Sefton et al., 1991; Sukekawa, 1988; Torrealba et al., 1984). The frontooccipital fibers arise mainly from pyramidal cells in layers 3 and 5; in area Oc1 they terminate in layers 5 and 6, in Oc2L in layers 1, 5, and 6, and in Oc2M in all layers except layer 2. Furthermore, connections between area 8 and areas Oc1, Oc2L, and Oc2M are reciprocal. The projection from area Oc1 to area 8 originates mainly from pyramidal cells in layer 5 at the border between Oc1 and Oc2L. The occipitofrontal projection from area Oc2L arises mainly from pyramidal cells in layer 5 and, to a lesser extent, from layers 2, 3, and 6; that from Oc2M arises mainly from pyramidal cells in layers 2 and 3 and, to a lesser extent, layers 5, 6, and 4 (Sukekawa, 1988). After injections of selenite salts (allowing tracing of specific zinc-rich presumably glutaminergic pathways)

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FIGURE 7 Diagrammatic representation of cortical connections of the visual cortical areas. (A) Associational connections of visuotopic areas of Oc: Oc1 (primary visual cortex, cytoarchitectonic area 17, area V1, striate cortex), AL and LM in Oc2L (cytoarchitectonic area 18a), and AM and PM in Oc2M (cytoarchitectonic area 18b). Viewed as if from above, right cortex; rostral upward. (A.1) Associational interconnections between visuotopically organized areas; note that all are reciprocal and that all areas in Oc2L and Oc2M are interconnected with Oc1. Area LM within area Oc2L is also connected reciprocally with all other designated regions. For references from which data are summarized, see Section V,D,3. (A.2) Associational connections between visuotopically organized areas within Oc1, Oc2L, and Oc2M and several nonvisuotopically organized regions of the cerebral cortex. Reciprocal connections are indicated with double-headed arrows. Areas Oc1, AL in Oc2L, and AM in Oc2M receive projections from five areas (SS: somatosensory cytoarchitectonic area 3; FEF: frontal eye field, area 8; retrosplenial cingulate area 29d; perirhinal areas 13/35; “paravisual” temporal area 36), area PM in Oc2M receives inputs from all but one (area 3) of the above cortical regions as well as a projection from area 11 in the prefrontal cortex, and area LM in Oc2L is innervated by three of the nonvisuotopically organized cortical regions (areas 13/35, 29d, and 36). For references, see Section V,E,3. (B) Commissural interconnections between visual cortical areas (Oc1, Oc2L, and Oc2M) and commissural connections with other regions of the cortex. Viewed from above, rostral upward. Note that Oc1, Oc2L, and Oc2M are interconnected reciprocally both homotopically and heterotopically (bottom arrows). In addition, each area receives a callosal input from the contralateral frontal eye field (area 8) and perirhinal area 13/35. Furthermore, areas Oc1 and Oc2M receive commissural inputs from retrosplenial cingulate area 29d and, in the case of area Oc2M, the connection is reciprocal. For references, see Section V,D,4.

into the frontal cortices, a substantial number of retrogradelly labeled neurons was present in supragranular layers 2 and 3 of ipsilateral areas Oc2M and Oc2L (Casanovas-Aguilar et al., 1998). However, selenite salt injections into frontal cortices reveal only few retrogradelly labeled neurons in infragranular layers 5 and 6 of ipsilateral areas Oc2M and Oc2L and in layer 6 of area Oc1 (Casanovas-Aguilar et al., 1998). Second, areas Oc1, Oc2L, and Oc2M all receive inputs from the retrosplenial cingulate cortex (cytoarchitectonic area 29d;

Dreher et al., 1990; Miller and Vogt, 1984a; Nauta and Bucher, 1954; Sanderson et al., 1991; Vogt and Miller, 1983); these connections between area 29d and Oc1, Oc2L, and Oc2M, especially the latter, appear to be reciprocal (Vogt and Miller, 1983). Third, all visuotopically organized areas, especially area AL (within Oc2L), receive inputs from paralimbic perirhinal areas 13/35 and, to a much lesser extent, “paravisual” associational area 36 (area Te2; see Chapter 35) in the temporal lobe. Fourth, areas Oc1 and AM (within Oc2M) and AL

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(within the Oc2L) receive inputs from the somatosensory cortex (cytoarchitectonic area 3), and the connections between Oc1 and the somatosensory cortex appear to be reciprocal (Olavarria and Montero, 1984; Sanderson et al., 1991; cf. also Casanovas-Aguilar et al., 1998). Finally, area PM (within cytoarchitectonic area 18b) receives associational input from cytoarchitectonic area 11 in the prefrontal cortex (Miller and Vogt, 1984a; Sanderson et al., 1991). In a study of bihemispherically projecting cortical neurons, Dreher and colleagues (1990) demonstrated that about 20% of associational neurons projecting to Oc1 from perirhinal area 13/35 are associational– commissural neurons that also contribute to the heterotopic callosal projection to contralateral areas Oc1, Oc2L, and Oc2M; all are pyramidal cells located in layer 5. Similarly, in limbic area 29d, about 10% of associational neurons projecting to Oc1 also project to contralateral Oc1, Oc2L, and Oc2M. In area 29d, both associational and associational–commissural neurons are located in layers 4, 5, and the upper and lower (but not the middle) parts of layer 6 (Dreher et al., 1990; Sanderson et al., 1991; Vogt and Miller, 1983). Finally, in oculomotor area 8, associational neurons projecting to the ipsilateral Oc1 are located in layer 4 and the upper half of layer 5; about 10% of them, found in layer 5, also project to contralateral Oc1, Oc2L, and Oc2M (Dreher et al., 1990). The interconnections between areas AM and PM within area Oc2M originate mainly in the supragranular layers 2/3 (feedforward projections; Sanderson et al., 1991) and those from Oc2L to areas in Oc2M terminate mainly in layers 2/3 and 4 (Coogan and Burkhalter, 1990). In contrast, 75% or more of the associational projections from area LM and areas 13/35 and 36 in the temporal lobe to area AL originate from infragranular layers, whereas other associational projections from areas AL and LM within Oc2L originate fairly evenly in both infragranular and supragranular layers (Sanderson et al., 1991). The responsiveness to photic stimuli and the visuotopic organization within area Oc2L survive acute ablation of Oc1, probably being sustained by projections from extrageniculate visual structures, particularly those from LP (Johnson and Burkhalter, 1992; Olavarria and Torrealba, 1978; Sanderson et al., 1991). Evidence indicates that Oc1 (area 17) exerts mainly an excitatory influence on cells in visuotopically organized areas within Oc2L and Oc2M. First, associational projections from area Oc1 use excitatory amino acids as transmitters (Johnson and Burkhalter, 1992). Second, at any visual eccentricity, the receptive fields in visuotopically organized areas within Oc2L and Oc2M are substantially larger than those in the visuotopically

corresponding regions of Oc1 (Espinoza and Thomas, 1983). Third, reversible inactivation of parts of Oc1 results in a substantial reduction in the firing rate of neurons in corresponding parts of Oc2L. While the responses of most cells to stationary visual stimuli are hardly affected, responses to moving stimuli, especially those of strongly orientation-selective cells, are reduced substantially (Molotchnikoff and Hubert, 1990). Commissural Connections of Visual Cortex The occipital cortices are interconnected commissurally via the caudal section of the corpus callosum (Olavarria and Van Sluyters, 1986). Figure 7B illustrates the connections diagrammatically. The bulk of Oc1, however, does not receive an input from the opposite cortex; the commissural neurons and terminals are concentrated in the lateral third at the border of areas LM (Oc2L) and AL where the vertical meridian is represented (Cipolloni and Peters, 1979; Cusick and Lund, 1981; Gravel and Hawkes, 1990; Jacobson, 1970; Lewis and Olavarria, 1995; Lund and Lund, 1970; Lund et al., 1984; Malach, 1988; Miller and Vogt, 1984b; Nauta and Bucher, 1954; Olavarria et al., 1987; Olavarria and Van Sluyters, 1985; Ribak, 1977; Schober et al., 1976; Sefton et al., 1991; Záborszky and Wolff, 1982). The medial twothirds of Oc1 are relatively free of commissural connections (Gravel and Hawkes, 1990; Miller and Vogt, 1984b; Olavarria and Van Sluyters, 1983, 1985; Sefton et al., 1991). Homotopic callosal connections of Oc1 originate mainly from pyramidal neurons located in layers 2/3 and in layers 5 and 6 (Hallman et al., 1988; Miller and Vogt, 1984b; Olavarria and Van Sluyters, 1983, 1985; Sefton et al., 1991). They terminate throughout all layers, although they concentrate in layers 1, 2/3, and 5 (Jacobson and Trojanowski, 1974; Miller and Vogt, 1984b; Schober et al., 1976). In addition, some callosally projecting neurons and terminals are scattered in the infragranular layers (especially layer 5) throughout the entire mediolateral extent of areas Oc1 and Oc2L (Granger et al., 1985; Gravel and Hawkes, 1990; Jacobson and Trojanowski, 1974; Miller and Vogt, 1984b; Olavarria and Van Sluyters, 1983, 1985; Sefton et al., 1991). The lateral part of area Oc1 also projects heterotopically to the medial part of the contralateral area Oc2L to terminate mainly in the supragranular layers (Miller and Vogt, 1984b; Schober et al., 1976). While the medial two-thirds of area Oc1 project to the heterotopic lateral part of the contralateral area Oc1 and to the medial part of area Oc2L, there is also a very small callosal projection from Oc1 to area Oc2M. Neurons projecting to the homotopic areas of the opposite hemisphere are distributed fairly evenly in layers 2/3, 4, and 5, whereas the heterotopically projecting callosal neurons are, to a large extent, concentrated in layer 5 (Miller and Vogt, 1984b).

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Commissural neurons in layers 5 and 6 do not send subcortical collaterals (Hallman et al., 1988). Apart from the callosally connected narrow region at the border with Oc1, area Oc2L is largely free from callosal neurons and terminals. There is, however, a rostrocaudal chain of five rings enclosing acallosal islands of visuotopically organized areas (Olavarria and Montero, 1984; Thomas and Espinoza, 1987). There are at least four callosally connected patches of cortex within area Oc2M: a small rostrolateral patch, a larger rostromedial patch, and two small patches along the rostrocaudal axis of Oc2M; the patches more or less merge (Malach, 1988; Miller and Vogt, 1984b; Olavarria and Van Sluyters, 1985). Callosal neurons in area Oc2L project mainly to the homotopic region of Oc2L in the opposite hemisphere; their terminals concentrate in supragranular layers 1,2,3, and 5 (Miller and Vogt, 1984b; Sanderson et al., 1991). There is also a small heterotopic projection from area Oc2L to Oc2M. The homotopically projecting neurons in area Oc2L are distributed throughout layers 2 to 5, whereas the heterotopically projecting neurons are virtually restricted to layer 5 (Miller and Vogt, 1984b). Area Oc2M (cytoarchitectonic area 18b) is interconnected with homotopic regions of area Oc2M as well as areas Oc1 and Oc2L of the opposite hemisphere. In area Oc2M, homotopically projecting callosal neurons are located mainly in layers 2/3, 4, and 5, whereas heterotopically projecting neurons are found exclusively in layer 5. Numerous neurons (presumably glutaminergic) are heavily labeled by injections of selenite salts into occipital cortices (zinc-rich projections). They lie in supragranular layers 2 and 3 and in inragranular layer 6 (less heavily labeled neurons were present in layer 5 and none in layer 4) of both homotopic and heterotopic contralateral visual areas (CasanovasAguilar et al., 1998). Finally, area 29d projects heterotopically to contralateral areas Oc1 and Oc2M, whereas callosal neurons in area Oc2M project to layers 2, 3, 4, and 5 of the contralateral retrosplenial area 29d (Miller and Vogt, 1984b). Heterotopically projecting neurons in area 29d are distributed throughout layers 2 to 5 (Dreher et al., 1990; Miller and Vogt, 1984b; Sefton et al., 1991). Visuotopically organized areas in Oc1, Oc2L, and Oc2M also receive commissural projections from the contralateral caudal third of cytoarchitectonic area 8 (frontal eye field) and contralateral perirhinal areas 13/35 in the temporal lobe (Dreher et al., 1990; Sefton et al., 1991). About 10% of the associational neurons projecting from area 8 to the ipsilateral area Oc1 also project contralaterally to the visual cortex (Dreher et al., 1990). Similarly, cells in perirhinal areas 13/35 that project to the contralateral visual cortex are located in layer 5 (Dreher et al., 1990; Sefton et al.,

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1991). They constitute about 20% of the associational neurons projecting from areas 13/35 to the ipsilateral Oc1 (Dreher et al., 1990). The perirhinal temporal cortex also receives a commissural input from the contralateral visual cortex (Olavarria and Montero, 1984). At certain stages of development (during the first 10–12 postnatal days), callosally interconnected regions apparently involve the entire mediolateral extent of the visual cortex, including regions in which noncorresponding parts of the visual field are represented (Bullier et al., 1990; Gravel and Hawkes, 1990; Lund et al., 1984; Miller and Vogt, 1984c; Olavarria and Van Sluyters, 1985). Furthermore, at that period more than half of the associational neurons projecting to area Oc1 also send an axonal branch to the contralateral visual cortical areas (Bullier et al., 1990). Neonatal or early postnatal eye enucleations (Cusick and Lund, 1982; Lund et al., 1984; Olavarria et al., 1987; Rothblatt and Hayes, 1982) result in a stabilization of the exuberant neonatal areal distribution of callosal neurons and terminals in the cortex contralateral to the enucleated eye. A similar stabilization of exuberant commissural connections is seen ipsilaterally following neonatal or early postnatal surgical damage to the dorsal thalamus (Cusick and Lund, 1982) or after kainate-induced cellular damage restricted to DLG (Sefton et al., 1991). Generally, regions of the visual cortex that are strongly callosally interconnected are complementarily interleaved with regions that have strong associational interconnections with visuotopically organized areas (Olavarria and Montero, 1984; Olavarria and Van Sluyters, 1985; Thomas and Espinoza, 1987). It is likely that heterotopic callosal connections between different hierarchical levels of cortical areas (see later) are functionally similar to the associational connections and could be classified as feedforward and feedback projections (see Dreher, 1986; Kennedy et al., 1991). Indeed, as mentioned earlier, about 30% of associational neurons projecting from Oc2L to Oc1 also project via the corpus callosum to the contralateral visuotopically organized areas Oc1, Oc2L, and Oc2M (Dreher et al., 1990). In albino rats, commissural cortical connections play a crucial role in generating the binocular excitatory responses characteristic of many cortical neurons located in the vicinity of the border between areas Oc1 and Oc2L (Diao et al., 1983a) and, in both albino and hooded rats, visual cortical potentials evoked by stimulation of the ipsilateral eye (Silviera et al., 1989). Consistent with the relative paucity of ipsilateral retinofugal projections, the posterior portion of the corpus callosum in the rat plays a significant role not only in the interhemispheric, but also in the interocular transfer of brightness and visual pattern discrimination (cf. Mohn and Russell, 1981).

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Feedforward and Feedback Associational Pathways: Homologies, Feedforward, and Feedback-Based Hierarchies of Cortical Visual Areas Despite the existence of a number of visuotopically organized representations of the contralateral visual field in the rat’s occipital cortex (see page 1119), there is controversy concerning the number of functionally distinct areas and possible homologies between the visuotopically organized areas of murid rodents and those of carnivores and primates. Thus, Malach (1989), while confirming numerous previous reports of associational connections between Oc1 and a number of visuotopically organized regions within Oc2L (area 18a) and Oc2M (area 18b), concluded that areas 18a and 18b together constitute a single visual area, homologous to area V2 of carnivores and primates and suggested that visuotopically organized patches of cortex within Oc2L and Oc2M represent interconnected but different functional modules within one global area [cf. Mitchison and Crick (1982) for a general exposition of the concept]. In contrast, Rumberger and colleagues (2001) argued on the basis of both qualitative and quantitative analyses of the anatomical connections between the rat’s primary visual cortex and the region of the visual cortex surrounding it that there are two distinct visual areas in the region of the cortex surrounding area V1; one of these areas appears to correspond to area Oc2L and the other to area Oc2M (see Zilles, 1990, Zilles and Wree, 1995). Sanderson and colleagues (1991; cf. also Coogan and Burkhalter, 1993; Montero, 1993) go even further and argue that only the lateromedial area LM within caudal Oc2L could be considered a homologue of areas V2 of carnivores and primates and that at least some of the patches of visuotopically organized cortex constitute distinct functional areas within Oc2L and Oc2M of the rat visual cortex. Thus, area LM of the rat, like areas V2 of cats and primates, receives its principal associational input from area 17 (Oc1). Second, the border region between areas LM and Oc1 is callosally connected and contains representation of the vertical meridian (Espinoza and Thomas, 1983; Thomas and Espinoza, 1987), as is the border between area V1 (area 17) and area V2 of carnivores and primates (Kaas, 1980; Kennedy et al., 1991; Tusa et al., 1981). Third, while in the rat pyramidal cells in layer 5 of areas V2 of cats and primates send only one visuotopically organized projection to the ipsilateral SC (Kawamura et al., 1974; Tsumoto et al., 1983), pyramidal cells in layer 5 of all visuotopically organized areas within Oc2L and Oc2M send separate visuotopically organized projections to the ipsilateral SC (Harvey and Worthington, 1990; Olavarria and Van Sluyters, 1982; Thong and Dreher, 1986).

Fourth, the laminar patterns of termination of corticotectal projections from area 17 [Oc1, area AL (Oc2L)] and from area Oc2M are distinctly different from each other. Thus, the corticotectal projection from Oc1 terminates in the superficial gray and upper part of optic layers, while the projection from the caudal part of area 18a (Oc2L), which includes area LM (the presumed homologue of V2 in carnivores and primates), terminates in the lower part of superficial gray, optic, and intermediate gray layers. Furthermore, the projection from area AL (in the rostral part of area 18a or Oc2L) terminates almost exclusively in optic and superficial gray layers. Finally, the corticotectal projection from area Oc2M (cytoarchitectonic area 18b) terminates almost exclusively in the deeper collicular layers (Coogan and Burkhalter, 1993; Harvey and Worthington, 1990). Fifth, while areas Oc1 (17) and Oc2M (18b) receive afferents from the ipsilateral and, to a lesser extent, the contralateral dorsomedial claustrum, area Oc2L (18a) does not receive claustral afferents (Carey and Neal, 1985; Dreher et al., 1990; Sanderson et al., 1991). In both primates and carnivores (for reviews, see Dreher, 1986; Kaas and Krubitzer, 1991; Van Essen and Mounsell, 1983), there is a “hierarchy” of visual cortical areas. The primary visual cortex that receives the bulk of its thalamic input from the DLG constitutes the base of the hierarchy. Associational projections from the primary visual cortex, which originate predominantly from supragranular layers 2 and 3 and terminate mainly in the granular layers of the “higher” cortical areas, are considered to constitute feedforward projections. However, associational projections from “higher order” areas to “lower order” areas, which originate mainly from the infragranular layers 5 and 6 and terminate in infragranular layers rather than layer 4, are considered to represent feedback projections. Applying the same principles to the rat’s visual cortex, the following hierarchy can be constructed. Area Oc1 (area 17, area V1, striate cortex), which receives most, if not all, of the cortical projection of the DLG (Olavarria, 1979; Ribak and Peters, 1975; Sanderson et al., 1991; Schober and Winkelmann, 1977), constitutes the base of the hierarchy. It sends feedforward projections to all visuotopically organized areas within areas Oc2L and Oc2M (Coogan and Burkhalter, 1990; Miller and Vogt, 1984a; Montero, 1993; Sanderson et al., 1991). Area LM within area Oc2L is probably homologous to area V2 of carnivores and primates and constitutes a second level of hierarchy. It receives its major associational feedforward input from area Oc1 (Sanderson et al., 1991) and, like other parts of area Oc2L, sends mainly a feedback type of projection to area Oc1 (Coogan and Burkhalter, 1990). Area AL in rostral area Oc2L, which receives feedforward input from area Oc1 and area LM

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(Sanderson et al., 1991), might constitute the third level of hierarchy. Area Oc2M, with its feedforward input from Oc1 and visuotopically organized areas within Oc2L (Coogan and Burkhalter, 1990; Miller and Vogt, 1984a; Sanderson et al., 1991) and feedback projections to areas Oc1 and Oc2L, would constitute an additional step in the hierarchy (Coogan and Burkhalter, 1990; Sanderson et al., 1991). A word of caution is necessary here. As mentioned in the previous section, Rumberger and colleagues (2001) argued that although the feedforward projections from a single region in the primary visual cortex to areas Oc2l and Oc2M are patchy, this “patchiness” does not necessarily indicate the existence of a number of distinct visuotopically organized areas within areas Oc2L and Oc2M but might be related to the presence of functional subunits within these areas. Nevertheless, despite these arguments, Schuett and colleagues (2002), mapping retinotopic organization of the mouse visual cortex using an optical imaging technique, concluded that there are at least four distinct extrastriate areas in the mouse visual cortex. These results support the view that the visual cortex of murine rodents consists of a multiple number of distinct areas. Another set of data that supports the aforementioned concept of a “hierarchy” of areas in the rat’s visual cortex is the laminar pattern of distribution of corticotectal terminals. Thus, in both carnivores and primates (for reviews, see Cusick, 1988; Dreher, 1986; Illing and Graybiel, 1986; Harting et al., 1992), corticotectal projections from the primary visual cortex terminate almost exclusively in the retino-recipient superficial layers of the ipsilateral SC, whereas the projections from progressively “higher” cortical areas terminate in progressively deeper layers in SC. As mentioned earlier, in the rat, the corticotectal terminals originating in area 17 (Oc1), the caudal part of Oc2L (LM), the rostral part of area Oc2L (AL), and in area Oc2M terminate in progressively deeper layers of the ipsilateral SC (Harvey and Worthington, 1990). Both feedforward and feedback corticocortical interactions are mediated via excitatory amino acids (Johnson and Burkhalter, 1994). The feedforward projections from the primary visual cortex to area LM within area Oc2L, as well as feedback projections from area LM to the primary visual cortex, terminate mainly on dendritic spines, rather than shafts, of pyramidal neurons (Johnson and Burkhalter, 1996). However, while 90 and 10% of feedforward projections terminate on processes of pyramidal cells and GABAergic interneurons, respectively, 98% of feedback terminates on pyramidal cells and only 2% on GABAergic interneurons (Johnson and Burkhalter, 1996). Both feedforward and feedback interactions between Oc1 and area LM (within area Oc2L) are mainly exci-

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tatory and are mediated via glutamate receptors, which are blocked by kynurenic acid. Electrical stimulation of feedforward projections from Oc1 to area LM (within Oc2L) results in synaptic activity in all layers (including strong activity in layer 4) of area LM (Domenici et al., 1995). Furthermore, the spatiotemporal pattern of feedforward synaptic activation in area LM following electrical activation of area 17 strongly resembles the spatiotemporal pattern of synaptic activation in Oc1 following activation of geniculocortical input (KenanVaknin and Teyler, 1994). In contrast, the synaptic activity in Oc1 following electrical activation of area LM (feedback input from area LM) is prominent in superficial layers but weak in layer 4 (Domenici et al., 1995). The spatiotemporal pattern of synaptic activity in Oc1 following electrical activation of area LM strongly resembles the spatiotemporal pattern of synaptic activation in Oc1 following activation of another part of Oc1 (intraareal connections within area 17; Domenici et al., 1995). Electrical stimulation of feedforward pathway from Oc1 evokes monosynaptic depolarizing (excitatory) postsynaptic potentials in almost 90% of cells in area LM. The EPSPs are followed by disynaptic hypolarizing (inhibitory) postsynaptic potentials (IPSP) (Shao and Burkhalter, 1996). Furthermore, in most cells in supragranular layers 2/3 in area LM, the EPSPs evoked by electrical stimulation of striate cortex are followed first by fast IPSP, which are mediated by GABAA receptors, which in turn are followed by slow IPSPs mediated by GABAB receptors (Shao and Burkhalter, 1999). In contrast, although electrical activation of area LM evokes monosynaptic EPSP in most neurons in Oc1, the inhibitory feedback input is mainly apparent in a slight acceleration in the decay of EPSPs (Shao and Burkhalter, 1996). Indeed, disynaptic fast and slow IPSPs (both mediated by GABAB receptors) are very rare following activation of the feedback pathway. Interestingly, activation of the feedback pathway by stimulation of area LM results in an increased rate of ongoing neuronal firing in Oc1 (Shao and Burkhalter, 1999). Morphological analysis also reveals that feedback projections from area LM provide strong monosynaptic excitatory input to forward projecting pyramidal neurons in the Oc1, which in turn form preferentially excitatory synapses on other local pyramidal cells (Johnson and Burkhalter, 1997). Thus excitatory networks within Oc1 appear to amplify presumably “modulatory” feedback activity from the “higher” visual areas (Johnson and Burkhalter, 1997).

Some Functional Considerations Ablation of Oc1 (primary visual cortex) results in a relatively small reduction in behavioral measured

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spatial acuity from about 1.0 to about 0.7 cycles per degree (Dean, 1981b). However, contrary to some previous claims (see Lashley, 1931; Lashley and Frank, 1934), it appears that ablation of Oc1 does not seriously impair the rat’s ability to discriminate patterns (Dean, 1981a; Hughes, 1977). Oc1 of the rat, therefore, as in many other mammals, seems to be mainly concerned with the discrimination of local features. Neurons of the primary visual cortex generally seem to be involved in acuity mechanisms rather than primary pattern and form analysis (for reviews, see Sprague et al., 1981; Stone and Dreher, 1982). The apparent discrepancy between these recent studies and the classical studies of Lashley seems to be related to the involvement of the lateral visual cortical area (Oc2L) in addition to Oc1, at least in some animals in Lashley’s series. Indeed, lesions involving both Oc1 and laterally located area Oc2L (which might contain several retinotopically organized distinct areas; Espinoza and Thomas, 1983; Montero, 1981; Thomas and Espinoza, 1987; see Fig. 2E) produce dramatic impairments in the rat’s ability to discriminate visual patterns and reduce the behaviorally measured acuity to 0.3 cycles per degree (Dean, 1981a). Furthermore, it appears that similar impairments in visual pattern discrimination can be produced by combined lesions of Oc1 and the lateral posterior nucleus (LP), which provides the principal thalamic input to areas LM in the caudal part of Oc2L and PM in caudal Oc2M. In contrast, lesions of LP without accompanying damage to Oc2L do not produce any significant impairment in pattern vision (Hughes, 1977). Evidence indicates that the pathway from retina via SC and LP nucleus to the visual cortex contributes to visual spatial resolution and pattern discrimination. First, if ablation of Oc1 is followed by ablation of SC, pattern vision and visual acuity are impaired to the same extent as after lesions involving both Oc1 and Oc2L (Dean, 1981a). Second, ablations of Oc2L with minimal involvement of Oc1 result in severe impairments in visual pattern discrimination and reversal visual learning tasks (Dean, 1981a; Gallardo et al., 1979; McDaniel et al., 1982). It can therefore be concluded that in the rat, as in other mammals, visual areas located outside the primary visual cortex play a crucial role in visual pattern discrimination (for reviews, see Sprague, 1981; Stone, 1983; Stone and Dreher, 1982; Stone et al., 1979). In the rat, these areas are located within Oc2L. However, selective ablations of medial extrastriate areas (e.g., the anteromedial area within Oc2M) result in the severe impairment of visuosomatic conditional responses (PintoHarnuy et al., 1987). On that basis, Montero (1993) suggested that in the rat, as in primates (cf. Mishkin et al., 1983; Goodale and Milner, 1992), there might be

separate cortical “streams” of processing of visual information: one involved in pattern recognition (areas within Oc2L) and the other in spatially oriented actions (areas within Oc2M). Feedforward and feedback corticocortical interactions are both mediated via excitatory amino acids (Johnson and Burkhalter, 1994). Feedforward projections from the primary visual cortex to area LM within area Oc2L, as well as feedback projections from area LM to the primary visual cortex, terminate mainly on the dendritic spines, rather then shafts, of pyramidal neurons. However, while 90 and 10% of feedforward projections terminate on processes of pyramidal cells and GABAergic interneurons, respectively, 98% of feedback terminates on pyramidal cells with only 2% on GABAergic interneurons (Johnson and Burkhalter, 1996). Both feedforward and feedback interactions between the striate cortex and area LM (within area Oc2L) are mainly excitatory and are mediated via glutamate receptors, which are blocked by kynurenic acid. Electrical stimulation of feedforward projections from area 17 to area LM results in synaptic activity in all layers (including layer 4) of LM (Domenici et al., 1995). Furthermore, the spatiotemporal pattern of synaptic activity resembles strongly the spatiotemporal pattern of synaptic activation in the striate cortex following the activation of geniculocortical input (Kenan-Vaknin and Teyler, 1994). In contrast, the synaptic activity in the striate cortex following electrical activation of area LM (feedback) is prominent in superficial layers but weak in layer 4. Furthermore, its spatiotemporal pattern resembles strongly the pattern of synaptic activation in the striate cortex following activation of another part of the striate cortex (intraareal connections within area 17; Domenici et al., 1995) or that following intraareal stimulation within the extrastriate cortex. Unlike thalamocortical projection fibers, tangential fibers within the striate area and extrastriate area Oc2L (18a) conduct extremely slowly (0.28 m/s; Lohmann and Rörig, 1994; Domenici et al., 1995)) Electrical stimulation of the feedforward pathway from the striate cortex evokes monosynaptic depolarizing (excitatory) postsynaptic potentials in almost 90% of cells in area LM, and the EPSPs are followed by disynaptic (inhibitory) postsynaptic potentials. Furthermore, in most cells in supragranular layers 2/3 in area LM, EPSPs evoked by electrical stimulation of the striate cortex are followed by fast IPSPs mediated by GABAA receptors and later by slow IPSPs mediated by GABAB receptors. In contrast, although electrical activation of area LM evokes monosynaptic EPSPs in most neurons in the striate cortex, the inhibitory feedback is mainly apparent in a slight acceleration in the decay of EPSPs. Indeed, disynaptic fast and slow IPSPs (both mediated

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by GABAB receptors) are very rare following activation of the feedback pathway. Interestingly, activation of the feedback pathway by stimulation of area LM results in an increased rate of ongoing neuronal firing in area 17 (Shao and Burkhalter, 1999). Morphological analysis also reveals that feedback projections from area LM provide strong monosynaptic excitatory input to forward-projecting pyramidal neurons in the striate cortex, which in turn form preferentially excitatory synapses on other local pyramidal cells. Thus excitatory networks within the striate cortex appear to amplify presumably “modulatory” feedback activity from the “higher” visual areas (Johnson and Burkhalter, 1997). Indeed, it is likely that in the rat, as in the cat (Alonso et al., 1993a, 1993b; Marinez-Conde et al., 1999; Wang et al., 2000) and macaque monkeys (Hupé et al., 1998, 2001a, 2001b), the feedback activity from the higherorder visual areas modulates not only the magnitude of responses of neurons in the striate cortex, but also their receptive field properties. Although rats have severely impaired visual capabilities after the removal of all visuotopically organized cortical areas, they are still able to perform some visual pattern discriminations (Spear and Barbas, 1975). Their remaining capabilities are presumably related to cortical areas outside the visuotopically organized regions and/or to subcortical visual nuclei. The cortical areas most likely to contribute to those remaining visual capabilities are perirhinal area 13/35—the “higher order” visual associational temporal area (Te2; see Chapter 19) within area 36 of Krieg (1946a, 1946b) and cingulate retrosplenial area 29d. Thus area Te2 within area 36 receives a direct input from the caudal part of the LP (Coleman and Clerici, 1980). In turn, LP receives inputs from several visual structures, such as the retino-recipient layers of SC, pretectum, and even some directly from the retina (see Table 3). The connections between area 36 and LP are reciprocal (Coleman and Clerici, 1980; Mason and Groos, 1981). Apart from its weak direct projection to visuotopically organized areas in the occipital cortex (see page 1130), area 36 projects to perirhinal area 13/35 (Deacon et al., 1983), which has substantial associational– commissural interconnections with Oc1, Oc2L, and Oc2M (see pages 1130–1131). Similarly, area 29d in the retrosplenial cingulate cortex receives its principal input from the anterior and lateral dorsal thalamic nuclei (Sripanidkulchai and Wyss, 1986; Van Groen and Wyss, 1992b), which in turn receive some input not only directly from the retina (Itaya et al., 1981; Repérent et al., 1987), but also retino-recipient regions of the pretectum (Thompson and Robertson, 1987). As discussed in pages 1130–1131, area 29d has substantial associational and commis-

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sural interconnections with area Oc1 and visuotopically organized areas within areas Oc2L (18a) and Oc2M (18b). The substantial interconnections between perirhinal area 13/35, paravisual area 36, and cingulate retrosplenial area 29d with area Oc1 and visuotopically organized areas within Oc2L and Oc2M suggest that those areas are involved in the integration of visual information. In particular, area 29d has bihemispheric interconnections with visuotopically organized areas, with the frontal eye field (Vogt and Miller, 1983) and the dorsocaudal claustrum (Divac et al., 1978). Thus, as for the presumably homologous areas in carnivores (Olson and Musil, 1992) and primates (Musil et al., 1992), it is probably involved in the analysis of the size and brightness of visual stimuli and might carry the signals related to the position of the eye in the orbit and/or the direction of head movements (Chen et al., 1990).

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33 Cerebral Vascular System OSCAR U. SCREMIN VA Greater Los Angeles Healthcare System and Department of Physiology UCLA School of Medicine, Los Angeles, California, USA

The vascular system is, in many aspects, unique and more heterogeneous in the brain than in most other organs. Cerebral capillary density ranges in the cerebrum from 260 mm/mm3 (average of white matter) to 2000 mm/mm3 in the paraventricular and supraoptic nuclei of the hypothalamus (Zeman and Innes, 1963). In addition to these quantitative differences, which reflect variations in metabolic rate, as well as the existence of specialized endocrine secretory mechanisms, there are also wide variations in capillary structure. Most vessels in the nervous system possess endothelial cell tight junctions that provide a seal and prevent or considerably hinder the passage of water-soluble molecules unless they interact with specialized transport systems. Capillaries in certain areas, however, are considerably more permeable and, thus, create microenvironments within brain tissue where many more blood constituents are accessible to nerve cells (Pardridge, 1988; Davson et al., 1987; Gross et al., 1986). Another unique aspect of brain circulation is the absence of a lymphatic system as known in most other organs. Although materials deposited in brain tissue or subarachnoid space can be recovered in the lymph nodes, this phenomenon is believed to be due to transfer from cerebrospinal fluid to extracranial extracellular fluid through the cribriform plate and the spaces around the trunks of the emerging cranial and spinal nerves (Davson et al., 1987). Wide variations in cerebral vascular anatomy exist among species. An intricate system of branching arteries, usually intermingled with veins or sinuses, the carotid rete mirabilis, exists in birds, cats, sheep,

The Rat Nervous System, Third Edition

swine, and, to a lesser extent, dogs (Daniel et al., 1953; Baker, 1982). Rats, however, like rabbits and primates, lack this system. In rats, as in all Placentalia, cerebral arteries and veins almost never run in pairs, although they connect through a complex network of capillaries. In contrast, in the brains of Marsupialia, arteries and veins meet at the surface of the brain, penetrate the brain tissue together, and remain paired throughout to the end of the terminal loop (Wislocki and Campbell, 1937; Scharrer, 1962). Other important characteristics of the rat cerebral vascular system, shared by most species, are the lack of direct communications between arteries and veins (A–V shunts) and the existence of numerous arterial and venous anastomoses that create considerable redundancy and make it almost impossible to produce complete localized ischemia of the brain by the occlusion of blood vessels. In rats, arterial anastomoses are found at all levels of branching, from the best-known arterial circle to the capillaries themselves, which form extensive uninterrupted meshes. The same is true of the rat venous system.

METHODOLOGY The description of the rat vascular system that follows is based on vascular casts obtained by injections of Batson’s No. 17 anatomical corrosion compound (Polysciences Inc., Warrington, PA) and cerebral sections of animals injected with a mixture of nine parts Neoprene Latex 571 (Dupont De Nemours & Co, Inc.,

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Wilmington, DE) and one part waterproof Black Drawing Ink A (Pelikan AG, Hannover, Germany). Animals were of the Sprague–Dawley strain, 250–350 g body mass. In preparation for intravascular injections, animals were anesthetized with halothane and an intravenous injection of 400 units of heparin sodium (The Upjohn Co., Kalamazoo, MI) was given. In order to study the arterial system, the chest was opened, and a stainless-steel cannula was inserted into the ascending aorta through the left ventricle and secured with a ligature at the root of the aorta. The right auricle was then opened and an infusion of 100 ml of heparinized saline (10 units heparin sodium/ml) was performed, followed by the Batson compound or the neoprene latex–ink mixture. In the case of the Batson compound, brains were left in place and, after 14 h, the entire head was immersed in 40% KOH until soft tissues were completely digested. The bone was then removed manually. Brains of animals injected with the neoprene latex–ink mixture were removed, frozen in methylbutane cooled to dry ice temperature, and later sectioned serially in a cryostat (Microm, Waldorf, Germany) at − 20ºC. The sections were dehydrated and mounted with Permount (Fisher Scientific, Pittsburgh, PA). The venous system was studied by retrograde injections of Batson compound into the external jugular vein with a technique similar to that used for the arterial system.

CEREBRAL BLOOD VESSELS Arteries Extracranial Origins Four arteries, two common carotids and two vertebrals, supply the cerebrum, brain stem, cerebellum, and cervical spinal cord (Fig. 1). Vascular diameters given in the following description were obtained from a total cast of the head arterial system (Batson compound). They should be taken as representative of the relative size of the blood vessels described and only as an approximation to the diameters in vivo. The common carotid arteries (0.90 mm in diameter) originate from the aortic arch on the left and the brachiocephalic trunk on the right. They divide at the level of the inferior border of the thyroid gland into external (0.77 mm in diameter) and internal (0.71 mm in diameter) carotid arteries (Fig. 1) [Paxinos et al., 1994 (Fig. 140)]. The second one gives origin, approximately 2 mm distal to the carotid bifurcation, to the pterygopalatine artery (0.53 mm in diameter), after which its diameter decreases to 0.56 mm. The vertebral arteries (0.34 mm in diameter) originate from the subclavian arteries, enter the vertebral

foramen of the sixth cervical vertebra, and then course inside the transverse canal, formed by the superposition of the transverse foraminae of the cervical vertebrae, until they reach the atlas bone (Fig. 1). Inside the canal, the vertebral arteries give off radicular arteries that feed into the ventral and dorsal spinal arteries. Pterygopalatine Artery On account of the notable arrangement of being encircled by the stapes, the pterygopalatine artery is also known as the stapedial artery (Tandler, 1899). Tandler has studied the comparative anatomy of this vessel in numerous species and has defined it as “. . . that artery which arises from the internal carotid, penetrating the stapes anlage in the embryo and is later located between the crurae of the stapes.” It is present as a fully developed artery in some animals or as a rudimentary vessel in others. Even within a single order, there are extreme variations in the relative size of the pterygopalatine and internal carotid arteries. Within rodents, for instance, Arctomys marmota (marmot) shows a large pterygopalatine and a rudimentary internal carotid, whereas the reverse is true of Pedetes caffer (springhare). In rats, however, both vessels are of a comparable size. The pterygopalatine artery is an inconstant, rudimentary vessel in humans. In Tandler’s systematic study, two main divisions of this vessel are described, the ramus superior and the ramus inferior, the first related to the middle meningeal and the orbital arteries (external ophthalmic artery in the rat) and the second one to the internal maxillary artery (pterygoid, descending palatine, sphenopalatine, and infraorbital arteries in the rat). The pterygopalatine artery (Fig. 1), the equivalent of the pterygopalatine portion of the internal maxillary artery, a branch of the external carotid artery of humans, supplies mostly extracranial structures, with the notable exception of the middle meningeal artery. It does not give off any branches on its extracranial course between its origin and its entrance to the cranium through the posterior lacerated foramen. After it enters the tympanic bulla, it travels along the medial wall of the tympanic cavity, crossing through the space between the crurae and the base of the stapes. It then emerges intracranially at the angle between the tympanic bulla and the petrous bone. Throughout its intracranial course, it remains in the subdural space and courses on an arched path around the bulla and slightly downward. It is at the outermost portion of this arched trajectory, and just underneath the lateral end of the transverse sinus, that the pterygopalatine artery gives origin to the middle meningeal artery (0.16 mm diameter), which divides into anterior, middle, and posterior branches to supply the duramater of the cerebrum (Fig. 2A). This is the

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FIGURE 1 Representation of the origins and distribution of the main cephalic arteries of the rat in sagittal (top) and horizontal (bottom) projections. These vascular outlines were drawn by tracing photographs of a complete cast (Batson compound) of the head arteries. Contours of the olfactory bulb, cerebrum, and cerebellum are indicated in the top projection. Parent vessels and main branches of the vertebral and external carotid arteries are blackened in the top projection, whereas parent vessels and main branches of the arterial circle are blackened in the bottom projection.

only intracranial branch of the pterygopalatine artery, which exits the cranium through the petrotympanic fissure and turns medially, terminating in a dorsal and a ventral group of vessels. The former includes the external ophthalmic artery, an anastomotic branch to the angular artery, the ethmoidal artery, and an artery to the pterygoid fossa (pterygoid artery). The arterial supply of the orbit and contents originates mostly from the two terminal branches of the external ophthalmic artery (Figs. 1 and 2A). Other vessels that contribute to the vascular supply of the orbit are branches from the angular artery and two intracranial branches of the internal carotid artery, the trigeminal

artery, and the internal ophthalmic artery. The ethmoidal artery supplies a few small branches to the posterior portion of the medial wall of the orbit and leaves the orbit through the ethmoidal foramen to distribute over the ethmoidal region of the nasal cavity. The diameters of these arteries at their entrance to the orbit are external ophthalmic, ramus superior, 0.21 mm; external ophthalmic, ramus inferior, 0.13 mm; internal ophthalmic, 0.03 mm; trigeminal artery, 0.04 mm; angular artery, 0.15 mm; and ethmoidal artery, 0.15 mm. The pterygoid artery, after supplying numerous small arteries to the tissues of the pterygopalatine fossa, continues ventrally and laterally, crossing over the mandible

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FIGURE 2 (Left) Shallow depth of focus photographs of a total arterial cast (Batson compound) of the rat head obtained with an AF Nikkor 50-mm lens and a +6 close-up lens, f/1.8, on a Nikon N8008 35-mm camera with APX 25 Agfapan film. The focal plane was at 6 mm from the midline in the top photograph and at the midline in the bottom photograph. (Right) Outlines of the main arteries present within the field of each photograph.

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between the base of the coronoid process and the last molar. This vessel then runs lateral to the last three molars and curves downward to anastomose with the facial artery (Figs. 1 and 2A). A similar anastomosis is provided in humans by the buccal artery (Platzer, 1989), a branch of the internal maxillary artery. The ventral group of vessels at the termination of the pterygopalatine artery includes the descending palatine, sphenopalatine, and infraorbital arteries (Figs. 1 and 2). The descending palatine, after running rostrally on the roof of the hard palate [Paxinos et al., 1994 (Fig. 136)] joins the contralateral homologous vessel. The infraorbital artery terminates by branching into six vibrissal arteries and additional branches for the dorsal portion of the nose, after its exit through the infraorbital foramen (Figs. 1 and 2A). Internal Carotid Artery After it has given off the pterygopalatine artery, the internal carotid artery continues in a dorsal and medial direction to enter the cranium through the carotid foramen, situated between the tympanic bulla and the basal plate of the occipital bone [Paxinos et al., 1994 (Fig. 41)], midway between the posterior lacerated foramen and the symphysis between occipital and basisphenoid bones. It emerges inside the skull at the level of the caudal border of the pituitary gland and then runs under the lateral border of the gland, where it leaves a depression. The first intracranial branch of the internal carotid artery, given off its ventral wall, is the trigeminal artery. This vessel can be followed on the surface of the ophthalmic nerve to the entrance of this nerve into the orbit. The trigeminal artery anastomoses with branches of the external ophthalmic artery and the termination of the internal ophthalmic artery at the caudal pole of the eye. The internal carotid artery then gives off the posterior communicating artery and from that point incorporates itself into the arterial circle, described in detail later (Fig. 1). Extracranial Anastomotic Circles Three wide anastomotic circles interconnect the internal and external carotid territories at several levels (Figs. 1 and 2). The three circles start at the origin of the external carotid bifurcation and then take one of three courses to join the pterygopalatine artery: (1) the pterygoid artery; (2) the angular artery, which is the termination of the trunk of the facial artery; or (3) the multiple anastomoses between the nasal branches of the facial artery (external carotid) and the vibrissal arteries, which are terminal branches of the infraorbital artery (pterygopalatine). In addition, a long anastomotic arch can be visualized on a horizontal projection (Fig. 1A). It starts at the point where the pterygopalatine artery

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branches from the internal carotid artery and continues with the descending palatine artery; the latter runs ventrally and rostrally and joins the contralateral homologous artery at the frontal end of the palate. These anastomotic systems are connected to the arterial circle, caudally through the internal carotid arteries at the point of origin of the pterygopalatine arteries and rostrally through the anastomoses of the olfactory arteries with the angular artery. Vertebral Arteries After they have reached the atlas, the vertebral arteries curve laterally and dorsally, giving dorsal muscular branches that exit through the alar foraminae of the dorsal arch of the atlas, at the point where they bend sharply to proceed medially and forward, while entering the foramen magnum until they finally fuse to form the origin of the basilar artery (Figs. 1 and 3A) [Paxinos et al., 1994 (Figs. 130–141)]. At the beginning of this segment, between the bend and the fusion, one dorsal spinal artery (0.10 mm in diameter) originates on each side. From points close to the fusion, and through a short but complex anastomotic network, a single ventral spinal artery (0.13 mm diameter) originates (Figs. 1 and 3A) [Paxinos et al., 1994 (Figs. 131–132)]. These three vessels descend into the cervical spinal canal and receive ventral and dorsal radicular arteries from the vertebrals and also from branches of the subclavian artery and aorta at more caudal levels. At the point where they bend sharply, the vertebral arteries can be found in close proximity to the surface of the posterior arch of the atlas and can be approached through the alar foramen (Pulsinelli and Brierley, 1979) (Fig. 1). Occlusion at this level leaves the origin of the ventral and dorsal spinal arteries intact and blood can then bypass the occlusion through these vessels that are fed, as described earlier, by the radicular arteries proximal to the occlusion. When both common carotid and vertebral arteries are occluded simultaneously, blood flow in the forebrain decreases to values low enough to abolish EEG activity (Pulsinelli et al., 1982). Spontaneous respiration and control of blood pressure are still maintained, as the ventral and dorsolateral spinal arteries are sufficient to maintain blood flow in the basilar territory at levels compatible with neural function. The posterior inferior cerebellar artery originates at the same level as the dorsal spinal artery, which occasionally is a branch of the former. At a level approximately midway between the emergence of the dorsal and ventral spinal arteries, the vertebral artery gives off the paraolivary artery (Figs. 3A and 13F), which runs lateral to the inferior olive, giving off numerous laterally directed branches that supply perforating vessels to the subjacent medullary structures.

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FIGURE 3 Ventral (A), dorsal (B), and lateral (C) views of the superficial cerebral arteries of a rat brain treated by selective arterial injection with a neoprene latex–black ink mixture.

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Basilar Artery The basilar artery, a median vessel (0.36 mm in diameter) resulting from the fusion of the two vertebral arteries (0.34 mm in diameter), runs over the ventral surface of the brain stem and supplies the brain stem and cerebellum (Figs. 3 and 4). Over its entire trajectory, this vessel gives off a palisade of dorsal branches (median medullary, pontine, and mesencephalic arteries) that penetrate the brain stem until they reach the floor of the fourth ventricle or the periaqueductal gray matter (Figs. 4, 8, and 11). At the medullary level, these median arteries terminate at the floor of the fourth ventricle in two vessels directed laterally and also give off laterally directed branches throughout their trajectory, which spans a distance about 0.5 mm from the midline (Figs. 11J, 13F, and 13G). The dorsal terminal branches frequently anastomose end to end with terminal, medially directed branches of the inferior anterior cerebellar artery, thus forming a complete circle around the brain stem (Fig. 3A). Numerous branches can be observed stemming off the basilar artery on the ventral surface of the brain stem. Four to six small arteries encircle the bulbar pyramids and penetrate the brain tissue just lateral to these structures. Immediately rostral to these vessels arise two arteries that course laterally for about 2 mm, giving off medial and lateral, as well as penetrating, branches. These vessels frequently anastomose end to end with the paraolivary arteries (branches of the vertebral arteries), thus forming two ventral medullary anastomotic circles (Fig. 3A). The inferior anterior cerebellar arteries (0.28 mm in diameter) originate at the junction between the medulla oblongata and the pons (Figs. 3A, 3C, 8A, and 11J). These vessels encircle the brain stem, giving off numerous penetrating arteries, and then terminate dorsally, providing the main arterial supply to the choroid plexus of the fourth ventricle, as well as branches to the cerebellum. The internal auditory arteries originate twothirds of the way between the emergence of the anterior inferior cerebellar arteries and that of the superior cerebellar arteries, [Paxinos et al., 1994 (Figs. 132–137)] These arteries course toward the internal auditory meatus, which they enter alongside the seventh and eighth nerves. Above and below these vessels, the basilar artery gives off a number of laterally directed vessels, the pontine ventral arteries, that supply penetrating vessels and terminate at the roots of the fifth, seventh, and eighth nerves (Fig. 3A). The superior cerebellar arteries (0.28 mm in diameter) are part of the terminal arborization of the basilar artery. As can be observed in Fig. 5, this terminal arborization can take the form of a quadrifurcation (Figs. 5A and 5D), but generally the basilar artery ends in two superior

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cerebellar arteries that give off, at a variable distance from their origin, the posterior cerebral arteries. Not infrequently, both posterior cerebral arteries originate from the same superior cerebellar artery (Fig. 5B and 5C). After their origin at the midline, the superior cerebellar arteries run laterally, and dorsally, along the rostral border of the pons. They then divide into a medial and a lateral branch. The medial superior cerebellar arteries supply vessels to the dorsal aspect of the cerebellum (dorsal cerebellar arteries) (Figs. 3B, 8B, and 12A) and to the caudal portion of the inferior colliculus (Fig. 12B). The lateral superior cerebellar arteries follow the outer border of the cerebellum (Figs. 1, 3C, 8A, 8B, and 13E), and, at the level of the uvula (lobule 9), they turn rostrally and divide into numerous medially directed branches, which almost close a circle around the cerebellum (Fig. 12D). The cerebellar tissue is supplied by arteries that run in the interfolial space and give off perforating vessels that run normal to the surface (Fig. 4). The perforating arteries destined for the granular layer run through the molecular layer, giving a few or no collaterals and they start branching at the limit between molecular and granular layers. Some capillary loops find their way back to the surface through the molecular layer, but the capillary plexus of this layer is mainly supplied from arteries on the surface of the cerebellar cortex. The capillary plexus of the granular layer is significantly more dense than that of the molecular layer (Fig. 4) (Zeman and Innes, 1963), a phenomenon in keeping with the higher cellular density of the granular layer. The posterior cerebral artery (0.23 mm diameter) commonly originates, as stated earlier, from the initial portion of the superior cerebellar artery (Fig. 5). The first branches from this vessel are an anastomosis with its contralateral homologous vessel and a variable number of thalamo-perforating arteries, usually three on each side, that course rostrally and dorsally to reach the ventral posterior region of the thalamus (Figs. 7, 8C, 10E, 10F, 10G, 13F, and 13G). The next branch, given off just before or after the junction with the posterior communicating artery, is the transverse collicular artery (0.15 mm in diameter) (Fig. 11I) that courses over the surface of the brachium of the inferior colliculus to be distributed mainly over the external cortex of the inferior colliculus (Coyle, 1975), contributing some anastomotic branches to the supracollicular network described later. This vessel also gives off perforating branches that supply the substantia nigra. Close to the origin of the transverse collicular artery, the posterior cerebral artery gives origin to the longitudinal hippocampal artery (0.25 mm in diameter) (Fig. 10G), which runs initially in the same general direction as its parent vessel and then follows the longitudinal axis of the hippocampus.

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FIGURE 4 (Bottom) A photograph of a 1-mm-thick midsagittal section of a specimen with a selective arterial injection (neoprene latex–black ink mixture). The palisade of arteries stemming from the basilar artery and supplying the medial portion of the brain stem is clearly shown. The location of the main anatomical structures is indicated. (Top) A portion of the cerebellum, identified with a rectangular insert, is shown at higher magnification. The top left portion of the top cerebellum image has been replaced by a photograph of a 0.1-mm-thick midsagittal section through the same location in order to highlight the variation in capillary density between molecular (MoCb) and granular (GrCb) layers of the cerebellar cortex.

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FIGURE 5 Six variations of the arterial circle anatomy (ventral view) are shown as described by Brown (1966). A symmetrical arrangement is represented in A, where the individual components have been identified. Both posterior cerebral arteries originate from the right superior cerebellar artery in B and from the left in C. An anterior communicating artery is shown in D. Double artery variations (buttonhole formations) are shown in the anterior cerebral arteries (A, B, E, and F) extending into the azygos anterior cerebral artery in C and also in the middle cerebral artery in E. A double origin of the middle cerebral artery is shown in B and F. Modified from Brown (1966) and reprinted by permission of WileyLiss, a division of John Wiley & Sons, Inc.

The posterior lateral choroidal artery (Figs. 10F and 12D) stems from the longitudinal hippocampal artery close to its origin or from the posterior cerebral artery and courses in an anterior, dorsal, and medial direction to join the distal portion of the anterior choroidal artery forming the common choroidal artery. These vessels supply the choroid plexus of the lateral ventricle and the anterior portion of the choroid plexus of the third ventricle. The posterior medial choroidal artery (Fig. 10E) originates from the supracollicular network and runs medially and dorsally to supply the caudal portion of the choroid plexus of the third ventricle. This vessel usually anastomoses rostrally with the commom choroidal artery. The terminal branches of the posterior lateral choroidal artery supply some of the dorsal thalamic arteries. The longitudinal (with respect to the axis of the hippocampus) hippocampal artery gives origin, at nearly regular intervals, to perpendicular short transverse

arteries (transverse hippocampal arteries) that course in the hippocampal fissure (Figs. 8B, 9D, 10E, 10F, 10G, 12B, 12C, 12D, 13E, and 13F). Numerous anastomoses can be found between this artery and the anterior boundary of the supracollicular network (Figs. 10G and 10H). Occasionally, the longitudinal hippocampal artery originates from the posterior communicating artery. Further details on the vascular anatomy of the hippocampus can be obtained from the excellent descriptions provided by Coyle (1975, 1976, 1978). Beyond the origin of the longitudinal hippocampal artery, the posterior cerebral artery gives off three or four main cortical branches that run in a dorsolateral direction over the surface of the occipital pole and reflect over the posterior border of the hemisphere to reach the dorsal aspect of the occipital cortex where they anastomose end to end with the occipital terminal branches of the middle cerebral artery (Figs. 11I, 12C, 12D, and 12E).

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FIGURE 6 Dorsal view of cerebral arteries injected with latex 20 days after occlusion of the right middle cerebral artery in a Wistar rat. Considerable enlargement and tortuosity of end-to-end anastomoses among branches of azygos anterior cerebral, azygos pericallosal, and posterior cerebral arteries, with those of the occluded middle cerebral artery, are clearly shown. Reproduced from Coyle (1984) and reprinted by permission of Wiley-Liss, a division of John Wiley & Sons, Inc.

The posterior cerebral artery ends in a variable number of branches that feed into an anastomotic network, which spreads over the dorsal surface of the superior and inferior colliculi (Figs. 4, 7, 10G, 10H, 11I, and 12B). This supracollicular network gives numerous perforating vessels that supply the superior and inferior colliculi. On its anterior border, this network also gives origin to arteries that supply the dorsal hippocampus and dorsal thalamus. On its posterior border, the supracollicular network anastomoses freely with the cortical pial network over the occipital cortex. On its anteromedial portion, the supracollicular network anastomoses with the terminal branches of the azygos pericallosal artery. Arterial Circle The anastomoses of arteries at the base of the brain are a remarkable feature of cerebral vascular anatomy

that was described in the 16th century. Although this structure is associated with the name of Thomas Willis, who described it in his Cerebri Anatome, published in 1664, it was in fact first mentioned by Gabriel Fallopius in 1561 and first illustrated by Giulio Casserio, one of William Harvey’s teachers in 1632, and by Johann Vesling in 1647, all from the School of Padua (McHenry, 1969). The anatomy of the arterial circle in rats is well known (Greene, 1963; Farris and Griffith, 1962; Zeman and Innes, 1963; Brown, 1966). Although there is a general resemblance to the analogous structure in humans, there are notable differences. The human posterior communicating artery is a slender blood vessel, not infrequently absent on one side, that joins the internal carotid artery and the terminal branching of the basilar artery. In rats, however, this vessel is of a size comparable to that of the middle and anterior cerebral arteries and anastomoses with the posterior cerebral artery at a point relatively more distant from the origin of these vessels from the basilar artery than that in humans. The anterior communicating artery is absent in rats, except as an anomaly (Fig. 5D). Instead, the anterior cerebral arteries fuse to form the azygos anterior cerebral artery at the point where the anterior communicating artery is found in humans (Fig. 5A). The rest of the arterial circle of the rat includes the same blood vessels as its human counterpart but the relative lengths of these are quite different. Whereas in humans the internal carotid gives rise to the posterior communicating and anterior cerebral arteries at the angle between the optic tract and the optic nerve, in rats it gives off the posterior communicating artery behind the posterior border of the optic chiasm, at the level of the median eminence. In consequence, the segment of the arterial circle occupied by the internal carotid is longer, and that occupied by the posterior communicating is shorter in rats than in humans. Moreover, the contribution of the posterior cerebral artery to the arterial circle in rats is greater than in humans. It is tempting to speculate that this may relate to the greater relative size of the rat hippocampus, which derives its arterial supply from this artery, precisely at the point of its connection with the posterior communicating artery. The courses and relative sizes of the arteries in the rat that are termed “posterior communicating” and “posterior cerebral” in the present work suggest that in this species the vessel distal to the junction between posterior communicating and posterior cerebral arteries is a continuation of the former (Fig. 5). In that case, the vessel that originates in the internal carotid should be named posterior cerebral, and the name “posterior communicating artery” should be applied to the connection between the posterior cerebral (originating in

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FIGURE 7 Photograph of a 1-mm-thick midsagittal section of a specimen with a selective arterial injection (neoprene latex–black ink mixture). Location of main structures and arteries are shown. In this and following figures, the approximate equivalence of planes of cerebral sections with those of the stereotaxic atlas of Paxinos and Watson (1998) is indicated. This section corresponds approximately to the Atlas, lateral, 0.4 mm, plate 80. The fan-like arrangement of vessels originating from the posterior cerebral arteries (pcer) in the posterior perforated space is evident on the bottom right-hand portion of the figure. The corpus callosum (cc) shapes the course of the azygos pericallosal artery (azp) and of the subfornical artery (sfa), which, after originating in the dorsum of the corpus callosum, follows a long course to reach its final destination, the subfornical organ (SFO). All the vessels supplying the medial septum (MS) are shown, as well as the origin of three branches of the azygos pericallosal artery: anterior (aif), middle (mif), and posterior (pif) internal frontal arteries.

the internal carotid) and the basilar arteries. Such an arrangement, in which the posterior cerebral artery originates from the internal carotid artery and the posterior communicating artery runs between the posterior cerebral and basilar arteries, has been described in humans as an anomaly (Polyak, 1957). An alternative approach is that of Greene (1963) in which the posterior communicating artery is seen running between internal carotid and basilar arteries, with the posterior cerebral artery emerging as a branch of the posterior communicating artery. However, there are ontogenetic arguments (Brown, 1966) that support the nomenclature used in this work (Fig. 5A) over these alternative concepts. More importantly, the present nomenclature maintains the conventions adopted almost universally for the description of the human arterial circle.

Although the general pattern of the arterial circle is maintained in most rats, significant variations can be observed. (Brown, 1966) has studied this phenomenon, and the main anomalies found by this author are presented in Fig. 5. The branches that originate from the arterial circle show important differences with regard to their human counterparts. A large olfactory artery arises from the anterior cerebral artery, approximately midway between its origin and the fusion with the contralateral artery to form the azygos anterior cerebral artery (Figs. 1, 5, and 8C). A vessel of similar origin and destination can be found in human embryos (Padget, 1944) but it does not persist into adulthood. The ethmoidal artery takes the place of the olfactory artery in mammals lacking this vessel.

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FIGURE 8 Shallow depth of focus photographs of a total arterial cast (Batson compound) of the rat brain, cerebellum, and brain stem obtained with an AF Micro-Nikkor 60-mm lens, f/2.8, on a Nikon N8008 35-mm camera with APX 25 Agfapan film. The focal plane was at 4 mm from the midline in A, 2.5 mm from the midline in B, and at the midline in C. The middle cerebral artery cortical distribution can be observed in A, with the cortical penetrating arteries coming into focus on the dorsal aspect of the brain. The midportion of the anterior choroidal artery (ach) can be appreciated in A, and its proximal portion in B, which also shows the general orientation of the transverse hippocampal arteries (trhi). The anterior striate artery (astr) and vessels approaching the amygdala from its ventral surface can be appreciated in A. The course of the anterior cerebral artery can be followed from B, after the emergence of the olfactory artery (olfa), to C, where the azygos anterior cerebral artery and its branches can be identified. The parallel course of the superior (scba) and anterior inferior (aica) cerebellar arteries can be appreciated in A.

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The hypothalamus is supplied by dorsomedially directed perforating vessels that originate from the posterior cerebral, internal carotid, and anterior cerebral arteries, either directly or from branches of these vessels that run medially over the ventral surface of the mamillary body, median eminence, and anterior hypothalamic area (Scremin, 1970; Ambach and Palkovits, 1975, 1977) (Figs 3A, 9B, 9C, 9D, 10E, and 13G). The infundibulum is irrigated by branches from the internal carotid artery (Scremin, 1970). This is at variance from humans in which there is an important contribution of the posterior communicating artery to this region. A prominent infundibular artery originates from the ventromedial wall of the internal carotid artery (Figs. 9C and 9D), just proximal to the origin of the posterior communicating artery (Figs. 1 and 3A). Midway between its origin and the midline, the infundibular artery divides into several branches that supply the median eminence (Fig. 9D), providing the arterial input of the hypothalamohypophyseal portal system. The infundibular arteries of the two sides are connected with each other through relatively large anastomotic vessels (0.03 to 0.06 mm in diameter). The posterior hypothalamus and mammillary bodies are irrigated by branches of the posterior communicating artery, although occasionally the branches belong to the internal carotid artery just opposite the infundibular artery. The anterior choroidal artery arises from the internal carotid artery, 0.3 to 0.6 mm rostral to the emergence of the posterior communicating artery (Fig. 1, 8A, and 8B). It gives off the posterior amygdaloid arteries and a small branch to the posteromedial portion of the piriform cortex. It then courses in a dorsolateral direction (Figs. 13G, 13E, and 13D) and divides into a lateral branch that supplies the choroid plexus of the lateral ventricle (Fig. 10E) and a dorsomedial branch that ascends to supply two to three dorsolateral thalamic arteries (Fig. 13E) and branches to the rostral portion of the choroid plexus of the third ventricle. The anterior choroidal artery anastomoses with the posterior lateral choroidal artery to form the common choroidal artery and also with the longitudinal hippocampal artery and the dorsal thalamic arteries. An internal ophthalmic artery (0.09 mm in diameter) originates from the medial wall of the internal carotid artery, 0.2 to 0.7 mm proximal to the origin of the middle cerebral artery (Figs. 1 and 3A). This vessel courses in an anterior and medial direction under the optic tract, gives penetrating branches to the lateral hypothalamus, and then joins the optic nerve, continuing on its surface until it enters the orbit. Inside the orbit the artery joins an anastomotic network that surrounds the caudal pole of the eye at the point of entrance of the optic nerve, which is also fed by branches from the

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external ophthalmic artery (from the pterygopalatine artery) and the trigeminal artery (from the internal carotid artery). The ophthalmic artery of humans originates from the internal carotid artery at the angle between the optic nerve and the optic tract, providing the bulk of the blood supply to the orbit and its contents. In rats, the contribution of this vessel to the vascular supply of the eye and orbit is minimal, and this territory is instead supplied, as described in detail earlier, largely by the external ophthalmic artery. Opposite the origin of the internal ophthalmic artery and at a level just rostral to the posterior border of the optic chiasm, an artery of 0.13 mm in diameter (corticoamygdaloid artery) originates from the lateral wall of the internal carotid artery (Figs. 1 and 3A). This vessel is distributed caudally over the piriform cortex (Figs. 9C and 9D), and its lateral branches anastomose with branches of the rhinal artery (Fig. 3C). The anterior amygdaloid arteries originate from the corticoamygdaloid artery and supply the amygdaloid complex (Fig. 9C). The middle cerebral artery (0.24 mm diameter) is one of the two terminal branches of the internal carotid artery. It originates from the arterial circle at a point about 2 mm caudal to bregma (“Atlas,” bregma, −2 mm) on the outer border of the optic tract (Fig. 3A). This vessel courses laterally and rostrally over the olfactory cortex and gives off several branches to the piriform cortex; at the level of the lateral olfactory tract, it yields a forwardly directed vessel, the corticostriate artery (Fig. 3A). The latter vessel supplies both the anterior portion of the piriform cortex and the lateral olfactory tract. It then gives off the anterior striate arteries, which course dorsally following the medial edge of the external capsule to supply the lateral and dorsal portions of the caudate-putamen (Figs. 8A, 9A, 12D, 13E, and 13F). A variable number of arteries (posterior striate arteries) that supply more caudal areas of the striatum (Figs. 9B, 9C, 12D, and 13E) originate from the middle cerebral artery, around the origin of the corticostriate arteries. These vessels are the equivalent of the lenticulate-striate arteries of humans. After giving off the corticostriate artery, the middle cerebral artery curves over the lateral surface of the cerebral hemisphere and branches in a variable pattern that, in general, is represented by groups of rostral, medial, and caudal vessels (Fig. 3). The second terminal branch of the internal carotid artery is the anterior cerebral artery (0.28 mm in diameter). This vessel courses in a forward and medial direction immediately ventral to the outer border of the optic chiasm (Figs. 1, 3A, and 5A). At a point approximately corresponding to the bregma (“Atlas,” bregma, −0.3 mm), it gives off the olfactory artery (0.20 mm in diameter) (Figs. 1, 2B, 5A, 7, and 8C). It then moves

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medially and dorsally, crossing the outer border of the optic chiasm at its origin and finally fusing with its contralateral homologous artery to form the azygos anterior cerebral artery (0.25 mm in diameter) (Fig. 5A). Generally, after the emergence of the olfactory artery, the anterior cerebral artery gives off the lateral orbitofrontal artery (Fig. 3A), which supplies the olfactory tubercle, the ventral surface of the olfactory bulb, and the rostral portion of the nucleus accumbens. The azygos anterior cerebral artery gives off one medial orbitofrontal artery to each hemisphere from its ventral wall (Fig. 7). This vessel divides into two terminal branches: (1) a cortical branch that supplies the medial and ventral orbital cortex, cingulate cortex, and frontal cortex and (2) an olfactory branch that irrigates the dorsal aspect of the olfactory bulb (Fig. 3B). The azygos anterior cerebral artery also gives off the ascending septal artery, which supplies the vertical limb of the diagonal band and the medial septum (Figs. 7, 8C, and 13G). The rostral portions of the septum are supplied by two to four smaller branches (rostral septal arteries) that stem off the posterior wall of the azygos anterior cerebral artery between the emergence of the ascending septal artery and the genu of the corpus callosum (Figs. 7 and 13E). The azygos anterior cerebral artery ascends in a dorsal and slightly caudal direction to bend over the genu of the corpus callosum, becoming the azygos pericallosal artery. At this transition, cortical branches emerge (anterior and middle internal frontal arteries) and course over the cingulate cortex and medial portions of the frontal cortex of both hemispheres to finally anastomose end to end with termination of the medial branches of the middle cerebral artery (Figs. 7, 12A, and 12B). The azygos pericallosal artery proceeds caudally (Figs. 2B, 7, and 8C), giving off the posterior internal frontal arteries, the retrosplenial artery, and terminal branches that supply the retrosplenial and occipital cortex (Figs. 10F, 10G, 10H, 11I, 12A, and 12B). Prominent end-to-end anastomoses can be seen between these branches and the caudal branches of the middle cerebral artery (Fig. 12A). The olfactory artery courses parallel to the external border of the optic chiasm, continues under the olfactory bulbs, and finally divides into two to four terminal branches that pass through the cribriform plate of the ethmoid bone to supply the nasal cavity (Fig. 1). This vessel gives off only a few intracranial branches, such as the rostral basal forebrain arteries that supply the horizontal limb of the diagonal band and the ventral pallidum, although these forebrain arteries may originate from the anterior cerebral artery itself or the proximal portion of the lateral orbitofrontal artery, a branch of the former.

Pial Arterial Network The pial arteries form a complex anastomotic network over the cortical surface. The middle cerebral, anterior cerebral, posterior cerebral, and internal carotid arteries all contribute to it. Inspection of Fig. 3 will reveal numerous end-to-end anastomoses among vessels originating in all these tributaries. Predominant in the dorsal view of the brain are anastomoses between branches from the azygos anterior cerebral, azygos pericallosal, and middle cerebral arteries in the paramedian region and among branches from the azygos pericallosal, middle cerebral, and posterior cerebral in the caudal region (Fig. 12A). In the lateral views, the rhinal artery, a branch from the middle cerebral artery, running almost horizontally in the caudal direction, receives numerous anastomoses from the most ventral rami of the terminal arborization of the middle cerebral artery and usually joins branches of the posterior cerebral artery with large end-to-end anastomoses (Fig. 3C). The rhinal artery lies deep in the rhinal fissure, usually under the caudal rhinal vein. Frequently this artery is represented by two or more smaller arteries that exchange anastomoses and surround the rhinal vein. These communications between territories are of crucial importance in the incidence of infarction following the partial occlusion of cortical vessels. There is a species variation as well as an effect of maturation on the frequency, tortuosity, and size of the anastomotic channels. This issue has been studied in great detail by Coyle and collaborators, who have produced convincing evidence of the plasticity of the cortical vascular architecture (Coyle, 1984, 1986; Coyle and Jokelainen, 1982; Coyle and Heistad, 1986, 1987; Coyle and Panzenbeck, 1990). Occlusion of the middle cerebral artery, for instance, is followed by an increase in diameter and tortuosity of these collateral channels, which explain the partial recovery of blood flow levels observed over time after such a lesion (Coyle, 1984) (Fig. 6). Another characteristic of the pial arterial network is the existence of anastomosis between adjacent arteries, which take the form of polygons of variable shape and size. In our material, the mean greater diameter of the polygons was 0.70 mm (SD 0.26); n= 115; and range, 0.13–1.58 mm. These small anastomotic circles were present on all areas of the cortex. Penetrating arteries originate from the sides of the polygons and enter the cortical tissue perpendicularly to the pial surface (Figs. 9–13). These anastomotic systems, placed at the site of emergence of the cortical penetrating arteries, are present in all mammals, including primates (Mchedlishvili and Kuridze, 1984). Although their functional significance has not been established, it is tempting to speculate that they may provide uniform

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perfusion pressure in all penetrating arteries derived from a single anastomotic polygon, perhaps ensuring uniform blood flow within the tissue polyhedron delimited by the penetrating arteries derived from the anastomotic polygon. This idea is coherent with a columnar organization of cortical neurons.

Veins Three main venous outflow systems can be outlined in rats: dorsal, ventral, and caudal. The dorsal system is represented by the retroglenoid vein, which does not exist in humans except as a rare anomaly, and is the main cerebral vein of rats (Figs. 14–16 and 18). This vessel can be considered as the extracranial continuation of the transverse sinus. The ventral system is represented by the cavernous sinus, which receives the basal vein, the anterior rhinal vein, and tributaries from the inferior and superior olfactory sinuses (Figs. 15 and 16). The caudal system is composed of the vertebral vein, vertebral canal sinus, and internal jugular vein. The three cerebral venous outflow systems are interconnected by a large number of intra- and extracranial anastomoses (Figs. 14–17). Retroglenoid Vein The diameter of the retroglenoid vein as it leaves the cranium through the retroglenoid foramen is 1.3 mm. This vessel drains the caudal and dorsal aspects of the brain and the anterior cerebellum (Figs. 14–16 and 18). It is the continuation, extracranially, of the transverse sinus that originates at the confluence of the superior sagittal and straight sinuses (torcular or caudal confluence of sinuses). The transverse sinus receives the dorsal cerebellar veins, lateral collicular vein, superior petrosal sinus, caudal rhinal vein, sigmoid sinus, and parafloccular vein (Figs. 14–17). As the retroglenoid vein exits the cranium, it is covered by a fascia for 1 mm of its trajectory. Below the inferior border of this fascia, and above the insertion of the tragicus muscle, it anastomoses with the internal maxillary vein through a 0.6-mm-wide, 0.6-mm-long vessel. It then courses outward and ventrally to anastomose with the superficial temporal vein (Figs. 14–16 and 18). Between these two anastomoses it receives tributaries on its anterior border from the adjacent temporal region, masseter muscle, and the temporomandibular joint and one tributary on its posterior border from the external auditory canal (Fig. 18). The internal maxillary vein is formed by the union of a large vein from the pterygoid plexus and the interpterygoid emissary vein; the latter drains the intercavernous sinus through which it communicates widely with its homologue on the opposite side. The internal maxillary and superficial temporal veins join

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to form the posterior facial vein, which then joins with the anterior facial vein to form the external jugular vein. Internal Jugular Vein The internal jugular vein originates at the junction of the sigmoid sinus; the inferior petrosal sinus, which is the caudal extension of the cavernous sinus; and an anastomosis with the pterygoid plexus (Figs. 14–16). The vertebral vein and the vertebral canal sinus originate from the same point. At variance with humans, the internal jugular vein of rats is a small, almost vestigial vessel. Its diameter at the posterior lacerated foramen, through which it leaves the skull, is 0.25 mm. Superficial Venous Systems The olfactory bulb is drained by a prominent superior olfactory sinus and by two inferior olfactory sinuses that drain into the rostral confluence of sinuses. From this point emerge the nasal emissary vein, which traverses the nasal bone and communicates with the supraorbital vein, and the olfactory emissary vein, which exits the cranium ventrally and joins the anterior communication between the two cavernous sinuses (Figs. 14–16). The ventral aspect of the cerebral cortex is drained, on its anterior portion, by the rostral rhinal vein, which drains into the rostral confluence of sinuses and also communicates by way of two branches with the cavernous sinus and the olfactory emissary vein. The posterior ventral aspect of the cerebral cortex is drained by the caudal rhinal vein, which empties in the transverse sinus. Both rhinal veins course on the rhinal fissure. The continuity between these two vessels is interrupted at the point where the middle cerebral artery crosses over the rhinal fissure (Figs. 15 and 16). At this level, branches of both rhinal veins flank the course of the artery for a short distance. Occasionally, a small end-to-end anastomosis that joins both veins is found under the middle cerebral artery. The posteromedial portion of the base of the brain is drained by the basal vein, which receives veins that run parallel to the ventral segment of the middle cerebral artery (middle cerebral vein) and to the initial segment of the anterior cerebral artery (anterior cerebral vein) (Figs. 15 and 16). It also receives veins from the anterior hypothalamic area. The basal vein drains caudally into the dorsal portion of the cavernous sinus, at the point of origin of the inferior and superior petrosal sinuses (Fig. 15). A similar vein exists in humans but, at variance with its homolog in rats, it continues dorsally to end on the great cerebral vein of Galen (Johanson, 1954; Duvernoy, 1975). The dorsal (superior sagittal and transverse sinuses and their tributaries) and ventral (cavernous sinus and its tributaries) cerebral venous systems are connected at numerous points. Intracranially, this communication is

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effected through a main channel, the superior petrosal sinus, which runs between the caudal termination of the cavernous sinus and the transverse sinus. The two systems also communicate by numerous anastomoses found over the surface of the cerebral hemispheres between tributaries of the basal vein and those of the anterior and posterior rhinal veins. The inferior petrosal sinus and the sigmoid sinus also connect both systems at the point of origin of the internal jugular vein and the vertebral vein and sinus (Figs. 15 and 16). Extracranial Venous Anastomoses Extracranially, the ventral and dorsal venous systems are connected through the interpterygoid emissary vein, which runs through the interpterygoid foramen and connects the intercavernous sinus with the internal maxillary veins and pterygoid plexi of both sides (Fig. 15). These in turn receive an anastomosis from the retroglenoid vein, the extracranial termination of the transverse sinus. The interpterygoid emissary vein may be

considered analogous to the emissary vein of the sphenoidal foramen of Vesalius, which has been described in humans. This emissary vein connects the cavernous sinus and the pterygoid plexus (Mettler, 1948). The analogy is suggested by the similar location of the interpterygoid foramen of rats and the foramen of Vesalius in humans because both are situated medial to the foramen ovale. The emissary vein of the foramen of Vesalius is a small vessel, which may not be always present because it has been omitted in the majority of descriptions of the human cranial venous system. In contrast, the interpterygoid vein of rats is a prominent channel (1 to 1.6 mm in diameter). Deep Venous Systems The superior and inferior colliculi, as well as dorsal portions of the hippocampus, thalamus, septum, and caudal striatum, are drained by a system composed of the azygos internal cerebral vein, the vein of Galen, and the straight sinus. At variance with humans, in

FIGURE 9 Coronal sections, 1 mm thick, of a brain injected intraarterially with a neoprene latex–black ink mixture. The approximate correspondence between these sections and the “Atlas” figures are as follows: A, interaural, 9.2 mm, plate 17; B, interaural, 8.2 mm, plate 21; C, interaural, 7.2 mm, plate 26; D, interaural, 6.2 mm, plate 31. The vascular supply of the anterior portion of the caudate putamen (Cpu) can be appreciated in A and that of its posterior portion in B and C. The majority of cortical penetrating arteries stay within the cortex, although vessels that are distributed to the subcortical white matter (scop) are found at regular intervals. The entire trajectory of the ventral thalamic arteries (vth) that supply the paraventricular nuclei of the hypothalamus and the reuniens thalamic nucleus (Re) can be observed in C and D.

FIGURE 10 Coronal sections, 1 mm thick, of a brain injected intraarterially with a neoprene latex–black ink mixture. The approximate correspondence between these sections and the Atlas figures are as follows: E, interaural, 5.2 mm, plate 35; F, interaural, 4.2 mm, plate 39; G, interaural, 3.4 mm, plate 42; H, interaural, 2.7 mm, plate 45. The anterior choroidal artery (ach) can be observed best on the left side of E, showing the choroid plexus (ChP). The thalamo-perforating arteries (thp), which first appear in E, can be followed in F, and their origin from the posterior cerebral arteries is revealed in G. The supracollicular arterial network can be appreciated best in G. The radial orientation of mesencephalic arteries is apparent in H.

FIGURE 11 Coronal sections, 1 mm thick, of a brain injected intraarterially with a neoprene latex–black ink mixture. The approximate correspondence between these sections and the “Atlas” figures are as follows: I, interaural, 1.7 mm, plate 49; J, interaural, −1.8 mm, plate 63. Numerous anastomoses are seen on I, between cortical branches of the posterior cerebral arteries (copc) and the occipital terminal branches of the middle cerebral arteries. Arteries converge on the periaqueductal gray (PAG) from the supracollicular network (scol), the surface of the pons, and the basilar artery. J shows the entire trajectory of a median medullary artery (mmd).

FIGURE 12 Horizontal sections, 1 mm thick, of a brain injected intraarterially with a neoprene latex–black ink mixture. Because of a slightly different orientation of the sections, the sections do not correspond exactly with single figures of the “Atlas.” The approximate correspondence is as follows: C (rostral), interaural, 6.90 mm, plate 116; C (caudal), interaural, 6.62 mm, plate 115; D (rostral), interaural, 5.90 mm, plate 112; D (caudal), interaural, 5.72 mm, plate 111. The cerebral cortex of the vertex is shown in A. The cortex is intact on the right side, showing large anastomoses on its surface between branches of the middle cerebral (mcer) and the middle (mif), and posterior (pif) internal frontal arteries, as well as the terminal branches of the azygos pericallosal artery (tep). Because the section plane is not perfectly horizontal, the surface of the cortex is in part absent on the left side of A, showing the penetrating cortical arteries (cop). This asymmetry is also present in B, where, on the left side, the transverse hippocampal arteries (trhi) can be seen. Most of the azygos pericallosal artery (azp) is shown in B, where the supracollicular arterial network is well developed. The vascular supply of the hippocampus can be appreciated best in C and D. FIGURE 13 Horizontal sections, 1 mm thick, of a brain injected intraarterially with a neoprene latex–black ink mixture. Due to a slightly different orientation of the sections, the sections do not correspond exactly with single figures of the “Atlas.” The approximate correspondence is as follows: E (rostral), interaural, 4.90 mm, plate 105; E (caudal), interaural, 4.18 mm, plate 105; F (rostral), interaural, 3.40 mm, plate 102; F (caudal), interaural, 2.40 mm, plate 98; G (rostral), interaural, 2.20 mm, plate 97; G (caudal), interaural, 1.40 mm, plate 94. The cortical branches (cof) of the medial orbitofrontal artery (mofr) are clearly seen in E. This figure also shows the anterior (astr) and posterior (pstr) striate arteries. The thalamo-perforating arteries (thp) are observed best in F and their origin is found on G, which shows best the general organization of the brain stem arteries in the horizontal plane.

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FIGURE 14 Dorsal view of the brain venous system. These vascular outlines, as well as those of Figs. 15, 16, and 17, were drawn by tracing photographs of casts (Batson compound) of the cerebral and cerebellar veins. Of the veins that drain into the transverse sinus, only the superficial ones are shown.

FIGURE 15 Ventral view of the rat brain venous system. Some of the veins presented in the dorsal view (Fig. 14) are shown with dotted lines for reference. The short anastomosis between the retroglenoid vein (rglv) and the internal maxillary vein (imaxv) is perpendicular to the plane of the figure and is shown as a shaded area (rg).

FIGURE 16 Lateral view of the rat brain venous system. The continuity between the caudal and the rostral rhinal veins is interrupted at the point where the middle cerebral artery crosses over the rhinal fissure. Branches from the two veins flank the trajectory of the artery. The interpterygoid emissary vein, which communicates the cavernous sinus (cav) with the internal maxillary vein (imaxv), is almost perpendicular to the plane of this figure (see Fig. 15) and thus appears short in this projection.

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FIGURE 17 Dorsal view of the deep dorsal veins of the rat brain. The superficial tributaries of the transverse sinus are not shown. The dorsal wall of the superior sagittal sinus has been removed to show the opening of the straight sinus into the caudal confluence of sinuses. The entrance of the inferior sagittal sinus (iss) into the straight sinus is shown as a shaded area.

which two internal cerebral veins are usually found, an unpaired (azygos) internal cerebral vein is characteristic of rats (Fig. 17). This vessel receives the dorsal septal veins anteriorly and two lateral thalamostriate veins. In commonly observed variations, one or both of the longitudinal hippocampal veins may run alongside the azygos internal cerebral vein before joining it. The great cerebral vein of Galen can be recognized by its position below the splenium of the corpus callosum. It is formed by the confluence of the longitudinal hippocampal veins and the azygos internal cerebral vein. From that point, this vessel turns slightly dorsally and joins the inferior sagittal sinus to form the straight sinus, which joins the superior sagittal and transverse sinuses at the caudal confluence of sinuses

(torcular). Just before this point, the straight sinus receives the medial collicular veins (Fig. 17). The longitudinal hippocampal veins continue ventrally and caudally and then ventrally and rostrally, following the longitudinal axis of the hippocampus. These vessels connect with the straight and transverse sinuses via the medial and lateral collicular veins, respectively, and also with the superior petrosal sinus, through several small channels. The hypothalamus and the ventral portions of the septum, striatum, and thalamus are drained by a system composed of the basal vein and the cavernous and inferior petrosal sinuses. Finally, the rostral striatum and claustrum are drained by deep branches of the cortical afferents to the sagittal sinus and rostral rhinal veins.

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FIGURE 18 Semischematic representation of the retroglenoid vein (rglv) as it exits the cranium. This vessel has been retracted forwardly to show its anastomosis (rg) with the internal maxillary vein. This anastomosis cannot be seen on a lateral view, as it is hidden by the retroglenoid vein. The insertion of the tragicus muscle (trm) is a good reference for the localization of the anastomosis. The temporalis muscle and the exorbital lacrimal gland are shown retracted dorsally and ventrally, respectively, to expose the veins.

SPINAL CORD BLOOD VESSELS Arteries The rat spinal cord arterial supply bears an almost exact resemblance to that of humans. It originates from three longitudinal systems that extend throughout the entire cord. These are the ventral spinal artery, which can be found at the entrance of the ventral median fissure, and two dorsal spinal arteries located just ventral to the entrance of the dorsal roots. These vessels are fed by the ventral and dorsal radicular arteries. The ventral spinal artery is a continuous channel that extends from vessels stemming from the vertebral arteries rostrally and ends at the filum terminale. Two additional longitudinal arterial channels are found on the surface of the cord with less frequency, except for the cervical segments, in which they are an almost constant feature.

These are a median dorsal spinal artery, situated at or close to the dorsal septum, and two lateral spinal arteries, situated about midway between the attachment of the dorsal and that of the ventral roots. These arteries may occasionally be replaced by two or more parallel vessels. The longitudinal channels described earlier are joined by transverse anastomoses that may take the form of a well-developed artery (transverse anastomotic circle) or an irregular anastomotic network (Fig. 19). The total number and precise location of the ventral and dorsal radicular arteries show some variability. Tveten (1976a) studied this issue in a series of 115 rats and found that dorsal radicular arteries were more numerous and evenly distributed, but of a smaller size, than the ventral radicular arteries. This author also found that the total number of ventral radicular arteries ranged from 3 to 14, with an average of 7. They were more frequent at C5 and C6 and from T11 to L1. The lowest

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FIGURE 19 Coronal section, 1 mm thick, of the spinal cord (C5) injected intraarterially with a neoprene latex–black ink mixture. Because it is common in the cervical spinal cord, this specimen shows the ventral (vsp), dorsal (dsp), median dorsal (mdosa), and lateral (lsp) spinal arteries. The remarkable difference in vascular density of gray and white matter is apparent in this unstained section. Contrast is provided only by the presence of injection material in small arteries and precapillary vessels.

frequencies of occurrence of these vessels were at C1–C3, T1–T3, and caudal to L2. Dorsal radicular arteries ranged from 16 to 35, with an average of 25, and were more abundant at the level of the cervical and lumbar enlargements. One radicular artery is considerably larger than any of the others and, in most cases, constitutes the main arterial supply for the lower thoracic and lumbosacral cord. This is the equivalent of the great ventral radicular artery of Adamkiewicz of humans (Adamkiewicz, 1881, 1882; Lazorthes et al., 1971). Its most frequent location in the study of Tveten was at T13, ranging from T11 to L2. The ventral spinal artery gives off, at regular intervals, the ventromedian or sulcal arteries (Fig. 19). These vessels ascend in the ventromedian sulcus, usually in pairs destined one to a side, and reach the medial junction between the gray commissure and the ventral horn. They are then distributed widely within the gray matter, giving one or more branches to the ventral horn with collaterals to the commissure, lateral gray column, and base of the dorsal horn. The capillary loops that

originate in these distribution vessels travel beyond the boundary between gray and white matter and supply the anterior and ventral portion of the lateral white columns. The rest of the spinal cord is supplied by perforating rami from the pial arterial network that interconnects the dorsal, dorsolateral, and lateral longitudinal arterial channels described earlier. These perforating arteries are arranged in a radial orientation and travel through the white matter, giving occasional small branches, to a final distribution in the gray matter. They are designated ventral paramedian, ventrolateral, mediolateral, dorsolateral, dorsal paramedian, and dorsomedian arteries (Fig. 19).

Veins The veins of the spinal cord may be divided, according to (Tveten, 1976b), into a central and a peripheral system. The former drains the ventral gray matter into the ventral median or sulcal veins, which run parallel to the arteries of the same name. The peripheral system

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comprises veins that reach the surface of the cord and drain into a pial plexus that eventually empties into the dorsomedian and ventral spinal and radicular veins. The ventromedian veins drain smaller vessels from both sides of the ventral gray matter, in contrast with the ventromedian spinal arteries, which, as described earlier, alternate between both sides. These veins are also more numerous and smaller in size than the corresponding arteries; they drain into a ventral median spinal vein that runs dorsal to the ventral spinal artery. This vein extends along the entire cord, with infrequent discontinuities. It anastomoses rostrally with an irregular venous plexus on the ventral surface of the brain stem and ends caudally at the conus medullaris. Radicular veins of variable size and position originate from the ventral spinal vein and follow the ventral and dorsal roots, but do not correspond exactly to the radicular arteries. A prominent dorsal median vein receives veins from the dorsal white matter columns, dorsal gray matter, and dorsal gray commissure, as well as veins from the pial venous plexus. Numerous dorsal radicular veins, usually larger than the ventral radicular veins, drain this vessel and in turn empty into the segmental vertebral veins and the epidural venous plexus (Tveten, 1976b).

VASCULAR INNERVATION Adrenergic Innervation The rat cerebral blood vessels are richly innervated. The course of sympathetic nerve fibers can be traced from the superior sympathetic ganglion that gives off the internal carotid nerve at its rostral end. This branch, the largest efferent from this ganglion, joins the internal carotid artery and contributes to the network of nerve fibers that surrounds the artery. The supply of noradrenergic nerve fibers to the vessels of the arterial circle, including the basilar artery, is provided by the superior cervical sympathetic ganglion because specific fluorescence and noradrenaline contents disappear after superior cervical ganglionectomy (Iwayama, 1970; Edvinsson and MacKenzie, 1977). The contribution of the stellate ganglion may be limited to the more caudal portion of the arterial circle (Edvinsson et al., 1972; Purves, 1972; Arbab et al., 1988). The density of noradrenergic innervation is greatest in the arteries of the base of the brain and it decreases more distally, although it is still present in intracerebral arterioles (MacKenzie and Scatton, 1987). These fibers have been shown to contain norepinephrine (NE), neuropeptide Y (Edvinsson et al., 1984), and 5-hydroxytryptamine (5-HT) (Bonvento and Lacombe, 1993). Unilateral stimulation of the supe-

rior cervical sympathetic ganglion in rats induces a modest reduction of cerebral blood flow (CBF) in the territories of the stimulated side supplied by the middle cerebral, posterior cerebral, and posterior communicating arteries, whereas CBF is preserved in the anterior cerebral and basilar territories (Tuor, 1990). Activation of the cerebrovascular sympathetic innervation may limit CBF in hypertension. This phenomenon, first demonstrated in cats (Bill and Linder, 1976), is also present in rats (Kobayashi et al., 1991).

Cholinergic Innervation Although, as discussed earlier, superior cervical ganglionectomy causes loss of the noradrenergic nerve fibers in cerebral vessels, considerable numbers of nonadrenergic nerve fibers still remain after this intervention (Iwayama, 1970; Edvinsson and MacKenzie, 1977). Many of these fibers have been identified as cholinergic on the basis of morphological (Iwayama, 1970) or histochemical (Lavrentieva et al., 1968; Miao and Lee, 1990) criteria. Tracing the origin of these fibers has proved more difficult than in the case of their adrenergic counterparts. All members of the parasympathetic outflow to the head and neck [preganglionic fibers from nuclei of the third, seventh, and ninth cranial nerves and postganglionic fibers from the ciliary (third), sphenopalatine (seventh), submandibular (seventh), and otic (ninth) ganglia, as well as from several microscopic ganglia and scattered cells] supply fibers to cerebral blood vessels or contribute to the internal carotid nervous plexus (Chorobski and Penfield, 1932; Mitchell, 1953; Suzuki, 1989; Hara et al., 1985). These components may merely make a transit along cerebral blood vessels on route to their final destinations or they may instead innervate the smooth muscle of these vessels with a potential for control of CBF. Evidence for a role of these systems in cerebrovascular control remains controversial because some authors have reported an increase in CBF following stimulation of the seventh nerve, superficial petrosal nerve, or sphenopalatine ganglion (James et al., 1969; D’Alecy and Rose, 1977; Pinard et al., 1979; Goadsby, 1989; Seylaz et al., 1988), whereas others have failed to detect any changes (Meyer et al., 1971; Busija and Heistad, 1981; Linder, 1981; Scremin et al., 1983). However, a central origin of cholinergic nerve fibers to cerebral blood vessels has been proposed (Scremin et al., 1980; Dauphin et al., 1991; Biesold et al., 1989), although evidence against the nucleus basalis being the sole source of this innervation has been adduced (Scremin et al., 1991; Galea et al., 1991; Santos-Benito et al., 1988; Waite et al., 1999). A physiological role for acetylcholine (ACh) in the control of CBF is indicated by several experimental

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findings. The slope of the regression of CBF on CGU, which defines the gain of the mechanism that adjusts blood flow to metabolic demand, is enhanced after pharmacologic inhibition of the enzymatic breakdown of ACh and is decreased by the cholinergic muscarinic blocker scopolamine (Scremin et al., 1993; Scremin and Jenden, 1996). Moreover, the increase in CBF observed during spontaneous cortical arousal or somatosensory stimulation is reduced markedly by cholinergic blockade with atropine or scopolamine and is enhanced by the inhibition of acetylcholinesterase with physostigmine (Scremin et al., 1973; Pearce et al., 1981; Tsukada et al., 1997).

Noncholinergic, Nonadrenergic Innervation Evidence for a nonadrenergic, noncholinergic innervation of cerebral blood vessels has been growing in recent years. An important supply of nerve fibers containing neuropeptide Y is present in the arteries of the arterial circle, and most of these fibers disappear after superior cervical ganglionectomy. Vasoactive intestinal peptide immunoreactive fibers are present in rat cerebral arteries (Kobayashi et al., 1983; Miao and Lee, 1991), and it has been proposed that these fibers originate in the sphenopalatine ganglia in this species (Hara et al., 1989). Substance P (Cuello et al., 1978) and calcitonin gene-related peptide (CGRP) (Rosenfeld et al., 1983) are known to exist in neurons of the trigeminal ganglion. Lesions of this ganglion cause the disappearance of these peptides from cerebral vessels (Edvinsson et al., 1988). Although both peptides are potent cerebral vasodilators, trigeminal ganglionectomy does not affect CBF (Edvinsson et al., 1988). It is then conceivable that substance P- and CGRP-containing nerve fibers of cerebral vessels may serve a sensory function or induce vascular relaxation in emergency situations (Edvinsson, 1988). A sensory role of dynorphin B-containing nerve fibers found in brain blood vessels has been proposed (Moskowitz et al., 1986). The presence of nerve fibers containing 5-HT has been well demonstrated for pial vessels and arteries of the arterial circle. Concentration of 5-HT in rat cerebral vessels is reduced by lesions of the medial and dorsal raphe and by superior cervical ganglionectomy (Reinhard et al., 1979; Marco et al., 1985; Bonvento et al., 1991; Chang et al., 1989). A potent constrictor effect of 5-HT on cerebral vessels in situ, blocked by ketanserin or methysergide, is well documented, although arterioles of diameter less than 0.07 mm show a propranololsensitive dilatation (Edvinsson, 1988). Several authors have provided evidence for a role of nitric oxide (NO) in cerebral vascular tone in vitro (Gonzalez and Estrada, 1991; Lee and Sarwinski, 1991;

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Toda and Okamura, 1991). NO is synthesized by NO synthase (NOS) from L-arginine in the presence of nicotinamide adenine dinucleotide hydrogen phosphate (NADPH) (Knowles et al., 1990). NO synthase (Nozaki et al., 1993) and NADPH (Tsuchida et al., 2001) have been detected in perivascular nerves of the arterial circle and proximal vessels stemming from it. NADPH colocalizes with tyrosine hydroxylase in some fibers, suggesting a sympathetic origin (Tsuchida et al., 2001). Others appear to originate from the sphenopalatine ganglion (Nozaki et al., 1993). NO may also originate from endothelial cells, macrophages, neurons, or glia (Groves and Wang, 2000); consequently, the presence of NOS or NADPH in perivascular nerves does not necessarily imply that they are involved in the responses of the cerebral circulation to NOS inhibition, such as attenuation of hypercapnic vasodilatation (Harada et al., 1997; Heinert et al., 1998).

FUNCTIONAL LOCALIZATION WITH BLOOD FLOW Experimenters in the first half of the last century already provided evidence for the usefulness of local blood flow in functional localization (Gerard, 1938; Schmidt and Hendrix, 1938). An early description of enhancement of blood flow in the visual pathways by photic stimulation was provided by Kety and collaborators using tissue autoradiographic detection of a radioactive blood flow tracer gas in cats (Landau et al., 1955). This methodology was later adapted to rats with the use of Iodo-[14C]-antipyrine as a blood flow tracer (Sakurada et al., 1978). The avoidance of anesthesia during the data acquisition phase with this technique considerably enhanced the usefulness of blood flow in functional localization within the brain of conscious animals. However, the need to inject the blood flow tracer intravenously in this approach requires restraint or tethering of the subject. This limitation has been recently overcome with the use of an implantable, remotely activated, infusion pump that permits tracer injection without manipulating the subject (Holschneider et al., 2002). The temporal resolution of the autoradiographic measurement of blood flow for functional localization can be reduced to 10 s, a significant advantage over metabolic measurements with 2-deoxyglucose, which requires a minimum of 30 min, thus impeding the study of rapidly changing sensory, motor, or behavioral states. Examples of localization of function with autoradiography in freely moving, nontethered rats are shown in Fig. 20. It is generally assumed that blood flow changes follow, and are a consequence of, the increase in

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FIGURE 20 Maps of the “flattened” cerebral cortex in which the x axis represents distance from the midline along the cortex and the y axis represents distance from bregma, positive if rostral and negative if caudal to this landmark. Iodo-[14C]-antipyrine, a blood flow tracer, was injected by remote activation of an implanted minipump to freely moving rats walking on a rotarod treadmill (TOP) or while exposed to 1- to 8-KHz alternating tones (bottom). Mean differences in Z scores of tracer activity between locomotor activity (n= 8) and controls at rest (n= 7) or tone exposure (n= 9) and nonstimulated controls (n= 12) are shown in a color-coded scale. These maps allow visualization of functional activation of perfusion (yellow to red color) on the dorsal, lateral, and basal cortical surfaces of the cerebral cortex in a single plane. Abbreviations of cortical regions: A, amygdala; FrA, frontal association cortex; I, insular cortex; M1, primary motor cortex; M2, secondary motor cortex; S1, primary somatosensory cortex; S2, secondary somatosensory cortex; Pir, piriform cortex; PRh, perirhinal cortex; RSA, retrosplenial agranular cortex; Te, temporal cortex; Tu, olfactory tubercle; V1, primary visual cortex; V2, secondary visual cortex. Graphs courtesy of Dr. Daniel Holschneider.

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33. CEREBRAL VASCULAR SYSTEM

metabolic activity. Although undoubtedly this is a primary function of the blood flow increase associated with enhanced activity, many experimental facts indicate that other processes are also at work. Transsynaptic (and not antidromic) activation appears to be required to elicit functional hyperemia in the spinal cord ventral horn (Scremin and Decima, 1983), implying that local blood flow may be controlled by a feedforward mechanism involving synaptically released neurotransmitters rather than a feedback mechanism based on a neuronal metabolic error signal. Such an “anticipatory,” primarily vascular mechanism could explain why the increase in CBF observed in functional activation studies is often in excess of the metabolic requirements. It has been shown, for instance, that during the initial phase of convulsive seizures, CBF increases severalfold, and simultaneous records of tissue pH not only fail to show a metabolic error signal (acidification), but indicate tissue alkalosis instead due to washout of CO2 by enhanced circulatory convection (Astrup et al., 1978). CO2 partial pressure is reciprocally related to synaptic efficacy (Esplin et al., 1973), and thus enhanced circulatory convection associated with activity may help preserve or enhance synaptic function. The same excess perfusion occurs in nearly all forms of physiological focal brain activation, a phenomenon that is exploited by the blood oxygen level dependent (BOLD) functional MRI protocol, based on the enhanced oxygenation of cerebral venous blood associated with activity. The local increase in blood flow associated with activity also participates in the clearance of metabolically generated heat and released neurotransmitters. Thus, the broad spectrum of functions served by local blood flow makes methods based on high-resolution detection of this variable useful nonspecific probes in the exploration of functional localization.

Abbreviations Arteries ang ama acer ach acom aif astr aorta asa azac azp bas bcph cab caud

Angular artery (facial) Anterior amygdaloid arteries (corticoamygdaloid) Anterior cerebral artery (internal carotid) Anterior choroidal artery (internal carotid) Anterior communicating artery (anterior cerebral) Anterior internal frontal artery (azygos pericallosal) Anterior striate arteries (corticostriate) Aorta (left ventricle) Ascending septal artery (azygos anterior cerebral) Azygos anterior cerebral artery (anterior cerebral) Azygos pericallosal artery (azygos anterior cerebral) Basilar artery (vertebral) Brachiocephalic trunk (aorta) Caudal ascending basal forebrain arteries (anterior cerebral) Caudal branches (middle cerebral)

com cctd cch cof copc cop coamg costr dpal dcb dop dpaq dsp dth dol dlth dmcb dom eth ectd eoph faci aica pica iorb infa ifl iaud ictd ioph lhy lofr lpaq lsp lscb ling lhia med mofr mstr mscb mdosa mmd mmes mpn mel mdac mcer mif mma nas

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1197 Commisural artery (ventromedian spinal) Common carotid artery (right: brachiocefalic, left: aorta) Common choroidal artery (posterior lateral choroidal, anterior choroidal) Cortical branch of orbitofrontal artery (lateral, medial orbitofrontal) Cortical branches of posterior cerebal artery (posterior cerebral) Cortical penetrating arteries (middle, anterior, posterior cerebral) Corticoamygdaloid artery (internal carotid) Corticostriate artery (middle cerebral) Descending palatine artery (pterygopalatine) Dorsal cerebellar arteries (medial superior cerebellar) Dorsal paramedian spinal artery (dorsal spinal) Dorsal periaqueductal arteries (supracollicular network) Dorsal spinal artery (vertebral, radicular) Dorsal thalamic arteries (posterior lateral choroidal, supracollicular network) Dorsolateral spinal artery (spinal transverse anastomoses) Dorsolateral thalamic arteries (anterior choroidal) Dorsomedial cerebellar arteries (superior cerebellar) Dorsomedian spinal artery (median dorsal) Ethmoidal artery (pterygopalatine) External carotid artery (common carotid) External ophthalmic artery (pterygopalatine) Facial artery (external carotid) Inferior anterior cerebellar artery (basilar) Inferior posterior cerebellar artery (vertebral) Infraorbital artery (pterygopalatine) Infundibular artery (internal carotid) Interfolial artery (superior, anterior inferior, posterior inferior cerebellar) Internal auditory artery (basilar) Internal carotid artery (common carotid) Internal ophthalmic artery (internal carotid) Lateral hypothalamic arteries (anterior cerebral, internal carotid, posterior communicating) Lateral orbitofrontal artery (anterior cerebral artery) Lateral periaqueductal arteries (supracollicular network) Lateral spinal artery (vertebral, radicular) Lateral superior cerebellar artery (superior cerebellar) Lingual artery (external carotid) Longitudinal hippocampal artery (posterior cerebral) Medial branches (middle cerebral) Medial orbitofrontal artery (azygos anterior cerebral) Medial striate arteries (anterior cerebral) Medial superior cerebellar artery (superior cerebellar) Median dorsal spinal artery (radicular) Median medullary arteries (basilar) Median mesencephalic arteries (basilar) Median pontine arteries (basilar) Mediolateral spinal artery (spinal transverse anastomoses) Medullary arterial circle (basilar) Middle cerebral artery (internal carotid) Middle internal frontal artery (azygos pericallosal) Middle meningeal artery (pterygopalatine) Nasal arteries (facial)

1198 occ olfa olo orb pol pa pira pnr pva pamy pcer pcom pif plch pmch pstr ptg ptgpal radar rha rabfa ros rsa spa tra sbcl scop sfa scba sopha scol tep thp trac trcol trhi trig vep vsp vth vlmd vlsp vmsp vert vbr Sinuses ccs cav ios ipets iss icav

OSCAR U. SCREMIN

Occipital artery (external carotid) Olfactory artery (anterior cerebral) Olfactory branch of medial orbitofrontal artery (medial orbitofrontal) Orbital artery (pterygopalatine) Paraolivary artery (vertebral) Perforating arteries (interfolial) Piriform arteries (middle cerebral) Pontine rami of basilar artery (basilar) Pontine ventral arteries (basilar) Posterior amygdaloid arteries (anterior choroidal) Posterior cerebral artery (superior cerebellar) Posterior communicating artery (internal carotid) Posterior internal frontal artery (azygos pericallosal) Posterior lateral choroidal artery (longitudinal hippocampal) Posterior medial choroidal artery (supracollicular network) Posterior striate arteries (middle cerebral) Pterygoid artery (pterygopalatine) Pterygopalatine artery (internal carotid) Radicular artery (lateral spinal, vertebral) Rhinal artery (middle cerebral) Rostral ascending basal forebrain arteries (olfactory, anterior cerebral) Rostral branches (middle cerebral) Rostral septal arteries (azygos anterior cerebral) Sphenopalatine artery (pterygopalatine) Spinal arterial transverse anastomoses (ventral, dorsal spinal) Subclavian artery (right: brachiocephalic, left: aorta) Subcortical penetrating arteries (middle, anterior, posterior cerebral) Subfornical artery (azygos pericallosal) Superior cerebellar artery (basilar) Superior ophthalmic artery (external ophthalmic) Supracollicular arterial network (posterior cerebral) Terminal pericallosal branch (azygos pericallosal) Thalamo-perforating arteries (posterior cerebral) Transverse anastomotic arterial circle (ventral and dorsal spinal) Transverse collicular artery (posterior cerebral) Transverse hippocampal arteries (longitudinal hippocampal) Trigeminal artery (internal carotid) Ventral paramedian spinal artery (spinal transverse anastomoses) Ventral spinal artery (vertebral, radicular) Ventral thalamic arteries (internal carotid, anterior cerebral) Ventrolateral medullary arteries (anterior inferior cerebellar) Ventrolateral spinal artery (spinal transverse anastomoses) Ventromedian spinal artery (ventral spinal) Vertebral artery (subclavian) Vibrissal arteries (infraorbital) Caudal confluence of sinuses (transverse sinus) Cavernous sinus (inferior petrosal sinus, interpterygoid emissary vein) Inferior olfactory sinus (rostral confluence of sinuses) Inferior petrosal sinus (internal jugular vein) Inferior sagittal sinus (straight sinus) Intercavernous sinus (interpterygoid emissary)

occs rcs sigs sts sos spets sss trs vertcs Veins acerv afv azicv azyv basv crhv dcbv dsv dmspv auv exjug gcv ivc itcv ifv ijug imxv iptgv lchv lcolv lhiv

luv masv mcolv mcerv nasem olfev ophv pflv pfv ptgpl rav rg rglv rrhv sav segv stemv scv sorb tmjv thsv trhiv vspv vmspv

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Occipital sinus (internal jugular vein) Rostral confluence of sinuses (nasal, olfactory emissary veins) Sigmoid sinus (internal jugular vein) Straight sinus (caudal confluence of sinuses) Superior olfactory sinus (rostral confluence of sinuses) Superior petrosal sinus (transverse sinus) Superior sagittal sinus (caudal confluence of sinuses) Transverse sinus (retroglenoid vein) Vertebral canal sinus (vertebral vein, vertebral epidural plexus) Anterior cerebral vein (basal) Anterior facial vein (external jugular) Azygos internal cerebral vein (great cerebral vein of Galen) Azygos vein (left superior vena cava) Basal vein (cavernous sinus) Caudal rhinal vein (transverse sinus) Dorsal cerebellar vein (transverse sinus) Dorsal septal veins (azygos internal cerebral) Dorsomedian spinal vein (vertebral canal sinus) External auditory canal vein (retroglenoid) External jugular vein (superior vena cava) Great cerebral vein of Galen (straight sinus) Inferior vena cava (right atrium) Intercostal vein (superior vena cava, azygos) Interfolial vein (dorsal cerebellar vein or sigmoid sinus) Internal jugular vein (superior vena cava) Internal maxillary vein (posterior facial) Interpterygoid emissary vein (internal maxillary) Lateral choroidal vein (thalamostriate) Lateral collicular vein (transverse sinus) Longitudinal hippocampal vein (medial, lateral collicular, superior petrosal sinus, great cerebral vein) Lumbar vein (azygos, inferior vena cava) Masseter vein (retroglenoid) Medial collicular vein (straight sinus) Middle cerebral vein (basal) Nasal emissary vein (supraorbital) Olfactory emissary vein (cavernous sinus) Ophthalmic vein (cavernous sinus) Parafloccular vein (transverse sinus) Posterior facial vein (external jugular) Pterygoid venous plexus (internal maxillary) Radicular vein (epidural plexus, segmental vertebral) Retroglenoid vein to internal maxillary vein anastomosis Retroglenoid vein (internal maxillary, superficial temporal) Rostral rhinal vein (rostral confluence of sinuses) Sacral vein (inferior vena cava) Segmental vertebral vein (vertebral, intercostal, lumbar, sacral) Superficial temporal vein (posterior facial) Superior vena cava (right atrium) Supraorbital vein (superficial temporal) Temporomandibular join vein (retroglenoid) Thalamostriate vein (azygos internal cerebral) Transverse hippocampal vein (longitudinal hippocampal) Ventral spinal vein (radicular, vertebral) Ventromedian spinal veins (ventral spinal)

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vep vertv Structures Al AFor Amyg AH AM BL BMP BST CPu CM GrCb MoCb 1–10 Cp Cb ChP Cg DG DC DCIC DLG DR dr DM DI Ent 7n CA1–3 FrA fi FL G g7 Gi GP GI Hb HL Hi HDB 12 Hy I IC IAM IP LH LSD LV M1 M2 MB Me MG MS MVe ME Md Dk LL Oc

Vertebral epidural venous plexus (vertebral vein, segmental vertebral) Vertebral vein (superior vena cava) Agranular insular cortex Alar foramen Amygdala Anterior hypothalamic area Anteromedial thalamic nucleus Basolateral amygdaloid nucleus Basomedial amygdaloid nucleus, posterior part Bed nucleus of the stria terminalis Caudate putamen Central medial thalamic nucleus Cerebellar cortex, granular layer Cerebellar cortex, molecular layer Cerebellar lobules Cerebellar peduncle, basal part Cerebellum Choroid plexus Cingulate cortex Dentate gyrus Dorsal cochlear nucleus Dorsal cortex of the inferior colliculus Dorsal lateral geniculate nucleus Dorsal raphe nucleus Dorsal root Dorsomedial hypothalamic nucleus Dysgranular insular cortex Entorhinal cortex Facial nerve Fields CA1–3 of Ammon’s horn Frontal association cortex Fimbria Forelimb area of the cortex Gelatinous thalamic nucleus Genu of the facial nerve Gigantocellular reticular nucleus Globus pallidus Granular insular cortex Habenula Hindlimb area of cortex Hippocampus Horizontal limb of the diagonal band Hypoglossal nucleus Hypothalamus Insular cortex Inferior colliculus Interanteromedial thalamic nucleus Interpeduncular nucleus Lateral hypothalamic area Lateral septal nucleus, dorsal part Lateral ventricle Primary motor cortex Secondary motor cortex Mammillary body Medial amygdaloid nucleus Medial geniculate Medial septum Medial vestibular nucleus Median eminence Medulla Nucleus of Darkschewitsch Nucleus of the lateral lemniscus Occipital cortex

ox Pa PV Par Par2 PAG PRh Pir Pons Pn PnO PH Prh py RCh RS Re Spt sol SpC Sp5C S1 S2 SI SN SC Te Te1 Te2 Th trm TS TU TyBu V1 V2 VLG VP VPL VPM vr

Optic chiasm Paraventricular nucleus of hypothalamus Paraventricular thalamic nucleus Parietal cortex Parietal cortex, area 2 Periaqueductal gray (central gray) Perirhinal cortex Piriform cortex Pons Pontine nuclei Pontine reticular nucleus, oral part Posterior hypothalamic area Prepositus hipoglossal nucleus Pyramidal tract Retrochiasmatic area Retrosplenial cortex Reuniens thalamic nucleus Septum Solitary tract Spinal cord Spinal trigeminal nucleus, caudal part Primary somatosensory cortex Secondary somatosensory cortex Substantia innominata Substantia nigra Superior colliculus Temporal cortex Temporal cortex, area 1 (primary auditory cortex) Temporal cortex, area 2 Thalamus Tragicus muscle Triangular septal nucleus Olfactory tubercle Tympanic bulla Primary visual cortex Secondary visual cortex Ventral lateral geniculate nucleus Ventral pallidum Ventral posterolateral thalamic nucleus Ventral posteromedial thalamic nucleus Ventral root

Acknowledgments This work was supported by a Senior Research Career Scientist Award from the Rehabilitation Research and Development Service, United States Department of Veterans Affairs. I thank Dr. George Paxinos for identification of structures in photographs of brain sections, Dr. Ralph R. Sonneschein for a review of the manuscript and many helpful suggestions, Ms. Mingen G. Li for assistance with brain sections, and Ms. Maria A. Scremin for help with photographic reproductions.

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C H A P T E R

34 The Serotonin and Tachykinin Systems ANTONY HARDING, GEORGE PAXINOS, and GLENDA HALLIDAY Prince of Wales Medical Research Institute, Sydney, New South Wales, Australia and The University of New South Wales, Sydney, New South Wales, Australia

The serotonin [5-hydroxytryptamine (5-HT)] system consists of clusters of neurons containing 5-HT within the raphe nuclei, the periaqueductal gray, and the surrounding reticular formation of the brain stem and has diverse projections throughout the central nervous system. These projections are organized topographically with parallel innervation from different neuronal clusters being a common feature. This divergent system contrasts markedly with the tachykinin system, where tachykinin-containing neurons are more ubiquitous in the central nervous system. Tachykinins are found both as long projection neurons and as interneurons with a restricted terminal field. In many instances, several tachykinin inputs may influence a single neuron. In addition, many neurons that contain tachykinins also contain other neurotransmitter substances, including 5-HT (Hökfelt et al., 2000). The possible involvement of 5-HT in affective behaviors, and both 5-HT and the tachykinins in pain perception, has stimulated research on the anatomy and function of neurons utilizing these neuromodulators.

SEROTONIN SYSTEM 5-HT is one of a class of chemicals called indolamines synthesized from tryptophan, which contains an indole ring and a carboxyl-amide side chain (Azmitia, 2001). Serotonin was researched extensively in the 1930s and 1940s under the name “enteramine” (Whitaker-Azmitia, 1999). Although the importance of 5-HT for its many different functions in the brain is now well established,

The Rat Nervous System, Third Edition

the presence of 5-HT in the rat central nervous system was only first demonstrated biochemically in the mid 20th century (Twarog and Page, 1953), but it was not until formaldehyde-induced fluorescence techniques were developed that the visualization of 5-HTcontaining structures was achieved (Dahlström and Fuxe, 1964). 5-HT is the best known and most studied of the indolamines, although several other indolamines have now been located in separate and more limited neuronal systems, including 5-methoxytryptamine, tryptamine, and 6-hydroxytryptamine (Dabadie et al., 1992; Wallace et al., 1982). The distribution of 5-HT-containing neurons was confirmed through specific immunohistochemical techniques to localize 5-HT (Steinbusch, 1981), its precursor, 5-hydroxytryptophan (Touret et al., 1987), and its synthesizing enzyme, tryptophan hydroxylase (Weissmann et al., 1987). The introduction of these immunohistochemical techniques overcame the major problem of the fluorescence technique, i.e., fluorophore instability under ultraviolet and white light. However, immunohistochemistry of serotonin has its own difficulties, with the majority of the serotonin chemical being lost during fixation and possibly also during immunohistochemistry incubations (Hökfelt et al., 2000). Fortunately, enough detectable 5-HT remains within the tissue to enable the location of 5-HT by immunohistochemistry, particularly within small diameter fibers, so that 5-HT local circuit relationships can be identified. It is interesting to note that the precursor to 5HT is found in fewer brain stem neurons than 5-HT itself (Touret et al., 1987). It has been shown that the

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release of 5-HT is proportional to the levels of intracellular 5-HT and that the dynamics of 5-HT release appear to differ from the classic quantal model of neurosecretion (Schaechter and Wurtman, 1990). More recently, it has been found that, as a backup to neurotransmitter reuptake failure, a cross-clearance of 5-HT into dopamine neurons can occur to clear the extracellular space (Zhou et al., 2002). This leads to the possibility that dopamine neurons can store and release 5HT in limited situations as a “false neurotransmitter” (Zhou et al., 2002). Studies on the development of the serotonin system reveal that 5-HT neurons develop early in fetal life as two separate clusters within the brain stem (Fig. 1): a rostral group just caudal to the mesencephalic flexure and a caudal group in the medulla oblongata (Aitken and Törk, 1988). The rostral group gives rise to almost all ascending 5-HT fibers, whereas the caudal group gives rise to the majority of descending fibers (Aitken and Törk, 1988). The early development of these neurons and their extensive brain connections prompted speculation that 5-HT may play an important role in regulating development of the brain (Azmitia, 2001; Dooley et al., 1997; Lauder and Krebs, 1978; Lauder et al., 1983).

Location of 5-HT Containing Cell Bodies The majority of serotonergic neurons are located in the midline raphe nuclei of the brain stem, which in Nissl-stained preparations appear as unpaired aggregations of neurons (Törk, 1985). However, not all raphe neurons contain 5-HT, and the proportion of 5-HT neurons within each nucleus varies considerably (Törk, 1985). In many instances, boundaries of the raphe nuclei merge with the surrounding reticular formation, making it difficult to specify the precise anatomical location of 5-HT-containing neurons. For this reason, the clusters of 5-HT neurons in the brain stem have been classified alphanumerically into nine groups, with group B1 found in the caudal medulla oblongata and B9 within the midbrain (Dahlström and Fuxe, 1964; Steinbusch, 1981; Törk, 1985, 1990; Weissmann et al., 1987). The location of these 5-HT neuronal clusters within particular cytoarchitectural regions is listed in Table 1 and shown in Figs. 16–27. B1, B2/4, and B3 groups develop from 5-HT cells born in the caudal group, whereas B5/8, B6/7, and B9 groups develop from 5-HT cells born in the rostral group (Törk, 1990). Group B1 in the Raphe Pallidus Nucleus (RPa) and Surrounding Reticular Formation 5-HT neurons in RPa are the most ventrally placed 5-HT neurons in the medulla oblongata. The majority

FIGURE 1 Camera lucinda drawing of a whole mount of an embryo at embryonic day 14 (E14). At E14, the rostral cluster has increased in its extent and the ascending fiber bundle reaches the rostral mesencephalon. A second cluster of 5-HT-immunoreactive neurons, together with prominent descending fiber bundles, appears in the rostral medulla. Some additional 5-HT-immunoreactive fibers appear close to the midline, but the cells of origin have not yet been demonstrated. Reproduced with permission from Aitken and Törk (1988).

of neurons are located between the pyramids in RPa, although occasional 5-HT neurons are found within the pyramids, on the ventral surface of the pyramids, and laterally around the exiting hypoglossal fibers. These laterally placed neurons form a separate and extensive rostrocaudal group. 5-HT neurons are found throughout the rostrocaudal extent of RPa, from the level of the rostral inferior olive to the level of the pyramidal decussation. The most rostral 5-HT neurons in RPa are contiguous with 5-HT neurons in the raphe magnus nucleus. The laterally placed 5-HT neurons designated as B1 cells are contiguous with laterally placed 5-HT

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TABLE 1 Alphanumeric classification

Nomenclature Assigned to 5-HT Neurons Cytoarchitectural regions

B1

Raphe pallidus nucleus (RPa) Caudal ventrolateral medulla (CVL)

B2

Raphe obscurus nucleus (ROb)

B3

Raphe magnus nucleus (RMg) Rostral ventrolateral medulla (RVL) Lateral paragigantocellular reticular nucleus (LPGi)

B4

Central gray of the medulla oblongata

B5

Pontine median raphe nucleus (MnR)

B6

Pontine dorsal raphe nucleus (DR)

B7

Midbrain dorsal raphe nucleus (DR)

B8

Midbrain median raphe nucleus (MnR) Caudal linear nucleus (CLi)

B9

Medial lemniscus (ml)

neurons in the rostral paramedian reticular formation designated as B3 cells. The location of these two groups of B1 cells can be seen in Figs. 23–26. Group B2/4 in the Raphe Obscurus Nucleus (ROb), Surrounding Reticular Formation, and Periaqueductal Gray Structures 5-HT neurons in ROb are found along the midline throughout the medulla oblongata as two vertical parallel sheets of neurons dorsal to the inferior olive and pyramids. The majority of neurons are located in the ventral half of the medulla oblongata, although occasional 5-HT neurons are located within periaqueductal gray structures (B4 cells) and more laterally in the reticular formation. The dendrites of many midline B2 cells have a distinctive dorsoventral orientation. 5-HT neurons are found throughout the rostrocaudal extent of ROb, from the level of the rostral inferior olive to the level of the pyramidal decussation. The most rostral 5HT neurons in ROb are contiguous with 5-HT neurons in the raphe magnus nucleus. The location of B2 and B4 cells can be seen in Figs. 24–27. Group B3 in the Raphe Magnus Nucleus (RMg) and Surrounding Reticular Formation 5-HT neurons in RMg are arranged more loosely than 5-HT neurons in RPa or ROb. The majority of neurons are located dorsal to the pyramids in the ventral half of the tegmentum of the rostral medulla oblongata and caudal pons, extending over a similar distance within the brain stem as the facial nucleus. The distribution of B3 neurons resembles an equilateral triangle in transverse section. The 5-HT neurons are not found directly on the midline, but extend

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laterally over several cytoarchitectural regions with a small number lying close to the ventromedial limits of the facial nucleus. The most caudal 5-HT neurons in RMg are contiguous with the midline 5-HT neurons in RPa and ROb, and the laterally placed 5-HT cells of the B3 group are contiguous with the more caudally and laterally located B1 group. The location of B3 cells can be seen in Figs. 21–23. Group B5/8 in the Median Raphe Nucleus (MnR) and Caudal Linear Nucleus (CLi) 5-HT neurons in the B5 group are located in the pontine raphe nucleus, although many studies have noted the variable distribution of 5-HT neurons within this structure (Steinbusch, 1981; Törk, 1985, 1990). It has been suggested that B5 neurons are the caudal remnant of 5-HT neurons located in the MnR designated as B8 cells (Törk, 1990). The larger B8 group of 5-HT neurons is found throughout the midline raphe of the rostral pons (MnR) and caudal linear nucleus of the midbrain (CLi). The 5-HT cells in these nuclei are separated artificially by the decussating fibers of the superior cerebellar peduncles. Both raphe nuclei are considered to have several subdivisions. Occasional 5-HT neurons are located in the surrounding reticular formation. Dorsal 5-HT neurons are contiguous with 5-HT cells in the dorsal raphe nucleus, whereas ventrally placed 5-HT neurons are contiguous with the B9 group. The location of B5 and B8 cells can be seen in Figs. 16–21. The number of 5-HT neurons in the B8 group declines significantly with age (Lolova and Davidoff, 1991). Group B6/7 in the Dorsal Raphe Nucleus (DR) and the Surrounding Periaqueductal Gray (PAG) 5-HT neurons in the B6/7 group are located largely within DR, although occasional 5-HT neurons are located more dorsally and laterally within the PAG. This cell group is the largest and most dense 5-HT aggregate in the rat brain. The B6 group is the caudal extension of the DR in the pons, whereas the B7 group is larger and located within the midbrain periaqueductal gray. The B7 group has a characteristic fountain shape in transverse sections with several obvious subregions. The location of B6/7 cells can be seen in Figs. 16–22. In contrast to the B8 group, there is no age-related loss of 5-HT neurons within DR, although 5-HT cell size and dendritic length declines significantly with age (Lolova and Davidoff, 1991). Group B9 in the Medial Lemniscus (ml) and Surrounding Reticular Formation 5-HT neurons in the B9 group are associated with the ml, although occasional 5-HT cells are located within the pontine and mesencephalic reticular formation. Cells

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of the B9 group are medium sized (15–25 μm), are mainly fusiform in shape, and are orientated predominantly in the mediolateral direction (Vertes and Crane, 1997). This group of 5-HT neurons is not as compactly organized as some of the midline (raphe) groups, but extends from the rostral midbrain to midpontine levels, although the 5-HT cells concentrate in the midbrain (Vertes and Crane, 1997). The B9 group has been reported to contain more 5-HT neurons than any other serotonergic group with the exception of the dorsal raphe group (Vertes

and Crane, 1997). The B9 group appears crescent shaped in transverse section and their location can be seen in Figs. 16–19.

Main 5-HT Pathways The relatively primitive and basic distribution of 5HT cell groups is conserved across vertebrate species, with almost all neuronal 5-HT cell bodies located in the brain stem on or near the midline (Jacobs and Fornal, 1999). From the brain stem, 5-HT neurons have extensive projections to virtually all areas of the brain and spinal cord (Steinbusch, 1981; Törk, 1985, 1990). The rostral groups of 5-HT neurons (B5–B9) give rise to almost all ascending fibers, whereas the caudal groups (B1–B4) give rise to the majority of descending fibers (Aitken and Törk, 1988; Jacobs and Fornal, 1999). In rats, the majority of 5-HT fibers are unmyelinated, although some myelinated fibers can also be found (Azmitia and Gannon, 1983). Studies on the development of these fiber systems have shown that the 5-HT innervation of cortical structures changes with postnatal development (Gould, 1999). Paradoxically, this innervation occurs irrespective of whether primary sensory input is present in the sensory cortical areas (Nakazawa et al., 1992), suggesting that intrinsic cortical factors predominate in the development of this system. Over the first 8 postnatal weeks there is a substantial decrease (37%) of 5-HT neurons in the B5–B9 cell groups (Koh et al., 1991). Results of these studies strengthen the hypothesis that the serotonin system functions during the critical period of neural development. Further evidence suggests that 5-HT fibers degenerate with age (van Luijtelaar et al., 1989). Thus, the serotonergic system appears to be capable of substantial changes over the life span of a rat. 5-HT Projections to the Spinal Cord

FIGURE 2 Computer-generated three-dimensional reconstructions of serotonin-immunopositive cells in sections represented on the left side of Fig. 16–27. Tracings were made around each section. Sections are spaced 500 μm apart and are aligned to the ventral aspect of the aqueduct and the fourth ventricle. The cerebellum has been removed. Rostral is to the left of the figure and caudal is to the right. Each dot represents 10 serotonin-immunopositive cells. (A) Ventral view displaying the midline nature of serotonin cells in the raphe nuclei. The lateral groups of serotonin cells (B3 and B9) are also represented. (B) Lateral view displaying two distinct groups of serotoninimmunopositive cells. The more caudal groups of cells are B1, B2, and B3 serotonin cells. The more rostral groups of cells are B5, B6, B7, B8, and B9 serotonin cells.

The spinal cord receives strong and highly organized 5-HT projections. 5-HT fibers are found throughout the gray matter of most spinal cord segments, with several regions containing a higher density of 5-HT terminals. These include layers 1 and 2 of the dorsal horn, the intermediolateral column, layer 10 surrounding the central canal, and ventral horn motor nuclei (laminae VIII and IX) (Anderson et al., 1989). There is still some controversy over whether 5-HT varicosities form substantial numbers of classical synaptic contacts with postsynaptic neural elements, with studies hypothesizing that 5-HT regulates astrocytic glial elements and plays a significant role in volume transmission, a more diffuse mode of neurotransmission (Ridet et al., 1993). In addition, 5-HT varicosities have extensive nonneuronal contacts, including central nervous system microvasculature

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(Cohen et al., 1995), and ependymal cells lining the ventricles and spinal central canal (Azmitia, 1999; Brusco et al., 1998). A study using retrograde tracers has localized 5-HT projections from ependymal cells and the subcommisural organ to a specific population of cells at the midline of dorsal raphe (B6/7) and median raphe (B5/8), respectively (Mikkelsen et al., 1997). The major source of 5-HT innervation to the spinal cord arises from the caudal groups of 5-HT cells (B1–B3) with predominance from B3 (RMg) cells (Araneda et al., 1989; Bowker et al., 1981b; Gao and Mason, 2001; Hökfelt et al., 1978; Johansson et al., 1981; Jones et al., 1991; Kwiat and Basbaum, 1990; Magoul et al., 1988; Mason, 1997; Millhorn et al., 1987, 1989; Skagerberg and Björklund, 1985; Zemlan et al., 1984). The B3 projection ends in the dorsal horn traveling in the dorsolateral fasciculus. The location of this fiber pathway within the cord has made it accessible and relatively easy to study. In contrast, the innervation of other spinal structures is mediated through deeper fiber bundles located in the ventral funiculus. 5-HT cells in ROb and RPa project to the ventral horn (and possibly the intermediolateral column), whereas lateral B1 cells (and some B3 cells) project to the intermediolateral column (Bacon et al., 1990; Bowker and Abbott, 1990; Gao and Mason, 1997; Helke et al., 1986; Sasek et al., 1990; Skagerberg and Björklund, 1985; Strack et al., 1989; Törk, 1990; Zagon and Bacon, 1991; Zemlan et al., 1984). The majority of 5-HT neurons projecting to the spinal cord have extensive collaterals over many segments. However, cervical spinal cord segments receive additional input largely from the B3 cell group, but there is some input from the B6, B7, B8, and B9 groups (Bowker and Abbott, 1990; Bowker et al., 1981a; Li et al., 2001; Skagerberg and Björklund, 1985; Zemlan et al., 1984). Physiologic evidence is now being presented to identify subclasses of B3 groups in their spinal projections with differing functions (Gao and Mason, 2001). Brain Stem and Cerebellar 5-HT Projections Many brain stem and cerebellar structures receive dense 5-HT projections, although in many instances the exact location of the cells of origin remains uncertain as different 5-HT cell groups often innervate the same structures. In addition, certain 5-HT cell groups have extremely diverse projections to several structures throughout the brain and spinal cord. The extent of collateralization of single 5-HT neurons has not been well studied, although attempts are now being made using both anatomical (Gao and Mason, 2001; Li et al., 2001) and physiological (Gao and Mason, 2001; Hentall et al., 2000; Mason, 1997; McQuade and Sharp, 1997) techniques. In the medulla oblongata (Figs. 19–27), 5-HT fibers are found in the reticular formation, the solitary complex,

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the cranial nerve nuclei, and the superior and inferior olive (Kolta et al., 1993; Steinbusch, 1981; Takeuchi et al., 1983; Törk, 1985; Woods and Azeredo, 1999). The dense projection to the solitary complex (Figs. 24–26) arises from a variety of cell groups, although the B3 and B7 groups predominate (Schaffar et al., 1988; Thor and Helke, 1987; Zemlan et al., 1984). 5-HT cells from at least the B3 group also innervate the dorsal vagal and hypoglossal nuclei (Zemlan et al., 1984) (Figs. 26 and 27). In addition, 5-HT fibers from peripheral ganglia (nodose and petrosal ganglia) innervate the solitary complex (Nosjean et al., 1990; Thor et al., 1988), and B6 and B7 cells innervate the rostral ventrolateral medulla of the B3 group (Underwood et al., 1999). Interestingly, only a small proportion of rostral ventromedial medulla neurons have serotonin innervation (Odeh and Antal, 2001). B3 cells have also been shown to innervate catecholamine cells located in the reticular formation (A1/C1) and solitary complex (A2/C2), which have projections to the spinal cord or diencephalon (Nicholas and Hancock, 1988, 1989, 1990). The rostral part of the ventral respiratory column, located in the reticular formation, receives a dense 5-HT innervation from all caudal raphe groups (Connelly et al., 1989; Holtman et al., 1990). There is also a dense 5-HT innervation of spinally projecting B3 cells (Figs. 16–22) from B6, B7, B8, and B9 (Beitz, 1982b; Lakos and Basbaum, 1988; Zeng et al., 1991). In addition, lateral B3 cells innervate more medial B3 cells (Beitz, 1982b). In the pons (Figs. 19–22), 5-HT fibers are found in periaqueductal gray structures, the parabrachial nucleus, and the cranial nerve nuclei (except those supplying the extraocular muscles) (Kolta et al., 1993; Odeh and Antal, 2001; Steinbusch, 1981; Takeuchi et al., 1983; Törk, 1985). The innervation of pontine structures is largely by B3, B6, and B8 cells (Clark and Proudfit, 1991; Imai et al., 1986; Törk, 1985; Zemlan et al., 1984). In the cerebellum, the highest density of 5-HT fibers is found in the granule cell layer, although most of the cortex and deep nuclei receive 5-HT input (Steinbusch, 1981; Törk, 1985). This input is also largely from B3 and B6 cell groups (Waterhouse et al., 1986). In the midbrain (Figs. 16–18), 5-HT fibers are also found in periaqueductal gray structures, as well as in the superficial layers of the superior colliculus, the cranial nerve nuclei (except those supplying the extraocular muscles), the interpeduncular complex, and the dopaminergic cell groups (Kolta et al., 1993; Steinbusch, 1981; Takeuchi et al., 1983; Törk, 1985). The majority of midbrain structures are innervated by B7 cells. Thus B7 cells innervate dopaminergic nuclei (Imai et al., 1986; Vertes, 1991), although B8 cells also contribute to this pathway (Vertes and Martin, 1988). B7 cells also innervate the periaqueductal gray and surrounding structures

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in the reticular formation (Vertes, 1991), most notably the cholinergic pedunculopontine tegmental nucleus (Figs. 17 and 18). B7 cells also innervate neurons in the MnR and CLi (Vertes, 1991) (Figs. 16–21). An interesting feature of B7 cells is that they send collaterals to regions of similar function, for example, B7 cells innervate the inferior colliculus and the cochlear nuclei (Klepper and Herbert, 1991). In contrast, B9 cells innervate the superior colliculus and B8 cells innervate the interpeduncular complex (Vertes and Crane, 1997; Vertes and Martin, 1988). 5-HT Fibers Innervating Forebrain Structures The ascending projections initially congregate in the medial forebrain bundle before diverging into different fiber pathways to innervate their target structures. The main target structures of these fibers are the olfactory bulb, hypothalamus, thalamus, septal area, striatum, hippocampus, and cerebral cortex (Araneda et al., 1989; Cumming et al., 1997; Ljubic-Thibal et al., 1999; Magoul et al., 1988; Stamford et al., 2000; Steinbusch, 1981; Törk, 1985; Vertes, 1991). Most forebrain structures receive parallel innervation from B7 and B8 cells, and within these cell groups the projections are organized topographically (Imai et al., 1986; Törk, 1990). In addition, collaterals from single 5-HT neurons are common with many neurons projecting to functionally related structures (Imai et al., 1986; Törk, 1990). One study suggests that some forebrain pathways are more vulnerable to age than others. For example, degenerating fibers were found in the basal ganglia and frontoparietal cortices (although, strangely, there was little change in 5-HT levels) compared with little pathology found in the septum or amygdala (van Luijtelaar et al., 1992). One interesting effect reported as a result of experimental transection of the medial forebrain bundle is an increase in the levels of 5-HT in the ventral mesencephalon, especially the substantia nigra and ventral tegmental area, which, associated with astroglial and microglial activation, may play a role in directing the regeneration of the 5-HT axons (Revuelta et al., 1999). Diencephalic structures receive heavy projections from B5 (MnR), B6 (DR), and especially B7 (DR), B8 (MnR and CLi), and B9 (ml) cells (Vertes, 1991; Vertes and Crane, 1997; Vertes and Martin, 1988). The most dense 5-HT innervation of the hypothalamus is seen in the suprachiasmatic nucleus, the lateral part of the medial preoptic nucleus, and dorsal and ventral premamillary nuclei (Törk, 1985). The dorsolateral hypothalamus receives projections predominantly from the midline portions of the dorsal raphe (Ljubic-Thibal et al., 1999), in particular involving the regulation of renin and prolactin (Van de Kar et al., 1996). Hypothalamic neurons containing histamine receive a specific

5-HT input (Ericson et al., 1989). The subfornical organ and other circumventricular structures are densely innervated by 5-HT fibers (Törk, 1985). The DR has a heavy projection to midline and intralaminar thalamic nuclei (Vertes, 1991), whereas B9 cells project to the intralaminar, central medial, lateral geniculate, reticular, paraventricular, reuniens, and rhomboid thalamic nuclei, as well as to the subthalamus nucleus (Vertes and Martin, 1988). In contrast, the MnR projects to the lateral habenula nucleus (Vertes and Martin, 1988). The basal forebrain and septal regions also receive a dense 5-HT input. 5-HT fibers in the septum arise from B7 and B8, as well as from some B3 cells, and these cells often send collaterals to the entorhinal cortex (Köhler et al., 1982). In the dorsolateral septum, many of the innervated neurons contain tachykinins (Gall and Moore, 1984). The cholinergic basal forebrain receives projections from B7, B8, and B9 cells (Jones and Cuello, 1989; Vertes, 1991; Vertes and Crane, 1997; Vertes and Martin, 1988). Furthermore, the DR projects to the bed nucleus of the stria terminalis, claustrum, and accumbens nucleus (Vertes, 1991), whereas the MnR projects to the accumbens nucleus (Vertes and Martin, 1988). The basal ganglia also receives a dense 5-HT innervation almost exclusively from the B7 cell group, although 5-HT cells within CLi also innervate these structures (Imai et al., 1986; Vertes, 1991). The basal ganglia 5-HT projection is concentrated in the posteromedial striatum and globus pallidus (McQuade and Sharp, 1997; Soghomonian et al., 1989; Vertes, 1991). Many neurons projecting to the striatum also send axon collaterals to the globus pallidus and substantia nigra (Imai et al., 1986; Törk, 1985). In addition, the amygdala receives a substantial 5-HT innervation mainly to its central, lateral, and basolateral nuclei (Imai et al., 1986; Vertes, 1991). The source of this projection is from 5-HT cells located in the B6, B7, and B8 cell groups (Imai et al., 1986; Vertes, 1991). There is an early significant contribution of 5-HT to growth and proliferation of hippocampal granule cells during neurogenesis (Gould, 1999). Consequently, the 5-HT innervation of the hippocampus in the adult is highly organized. Both B6 (DR) and B5/8 (MnR) cell groups contribute to the hippocampal 5-HT projection (Imai et al., 1986; McQuade and Sharp, 1997; Patel et al., 1996; Törk, 1985; Vertes, 1991; Vertes and Martin, 1988). Within the hippocampal formation, dense bands of 5-HT fibers can be seen within the strata lacunosum moleculare and oriens, with CA1 and CA2 regions receiving the most dense innervation compared with CA3 and CA4 (Ihara et al., 1988; Törk, 1985). The 5-HT terminals within the hippocampus have been shown to preferentially target neurons containing the inhibitory neurotransmitter GABA and the calcium-binding protein calbindin D-28K (Freund et al., 1990), although it is still

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questionable whether many anatomical synapses are formed by these terminals (Oleskevich et al., 1991). Most cytoarchitectural areas of the cortex receive an overlapping innervation from the B7, B8, and B9 cell groups, although the B7 and B9 cell groups project topographically to the cortex whereas the B8 group does not (O’Hearn and Molliver, 1984). It is currently thought that two basic systems innervate the cortex differentially (Kosofsky and Molliver, 1987; Mamounas et al., 1992). The B7 and B9 cell groups innervate the entire cortex, with particular subgroups of neurons innervating related cortical and subcortical fields. For example, 5-HT cells in the lateral wing of DR influence the primary visual cortex as well as the superior colliculus and the lateral geniculate thalamic nucleus (Villar et al., 1988). The laminar distribution of 5-HT fibers arising from these neurons differs between various cortical regions. In contrast, B8 cells seem to innervate the outer cortical layers of most cortical regions. Several cortical regions are more heavily innervated than others, including the piriform, insular, and frontal cortices, with the occipital, entorhinal, perirhinal, frontal orbital, anterior cingulate, and infralimbic cortices receiving a moderate innervation (Vertes, 1991). There has been considerable controversy over whether 5-HT axon terminals in the cortex form conventional synapses. Initial studies using serial section analysis suggested that the majority of terminals have a small synaptic specialization (Papadopoulos et al., 1987), but serial section computer reconstructions have questioned this finding (Séguéla et al., 1989). This discrepancy may reflect cortical specialization rather than true anatomical differences. Nevertheless, the anatomical distribution of 5-HT terminals reflects a highly ordered projection system. In addition to these central nervous system 5-HT projections, several studies have suggested that cerebral blood vessels are innervated by brain stem 5-HT neurons located in the DR (Cohen et al., 1995). This has led to the theory that brain stem 5-HT neurons may play an important part in the regulation of cerebral blood flow under physiological conditions and may also have a role in the pathogenesis of cerebrovascular disorders. However, there is some suggestion that this projection does not exist (Mathiau et al., 1993a, 1993b, 1993c). The identification of serotonergic neurons in the ventrolateral medulla as being central chemoreceptors that react when stimulated by acidosis in the blood is a more recent development (Bradley et al., 2002; Richerson et al., 2001). The neurons involved are tightly apposed to the large arteries that run through this medullary region (Bradley et al., 2002) and have projections that run to all of the major respiratory nuclei in the brain (Richerson et al., 2001). It is suggested that these neu-

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rons make a significant contribution to the sleep–wake cycle and respiratory control (Richerson et al., 2001).

5-HT Receptor Types and Location In addition to the variety of structures innervated by 5-HT, various types of postsynaptic receptors determine the final response to 5-HT release. Rapid advances in molecular biological techniques over the past decade have resulted in a recent update of the classification of 5-HT receptors (Barnes and Sharp, 1999; Hoyer and Martin, 1996, 1997). There are now seven different subclasses of 5-HT receptors (numbered consecutively), with at least 14 distinct subtypes recognized (Barnes and Sharp, 1999). These receptors have distinct distributions and are thought to mediate the different cellular responses to 5-HT. There are five subtypes of 5-HT1 receptors, labeled 5-HT1A, 5-HT1B, 5-HT1D, 5-HT1E, and 5-HT1F (Barnes and Sharp, 1999). 5-HT1A receptors are found within the raphe nuclei and hippocampus of the brain as well as in the enteric nervous system. This receptor hypopolarizes the membrane by opening K+ channels and also modulates the adenylate cyclase second messenger system. 5-HT1B receptors were originally identified as being localized in rodent brain only. More recently, however, species homologues have been identified and are one of the factors driving reclassification of the 5-HT receptor nomenclature (see Barnes and Sharp, 1999; Hoyer and Martin, 1996, 1997). 5-HT1B receptors are found in high density in the rat basal ganglia (including substantia nigra and globus pallidus) and also in many other regions, including deep cerebellar nuclei (Barnes and Sharp, 1999; Ramboz et al., 1996). These receptors act by inhibiting adenylate cyclase. 5-HT1C receptors have been reclassified as 5-HT2C receptors based on its operational and transductional similarities with the 5-HT2 receptors (now 5-HT2A) (Hoyer and Martin, 1996, 1997). 5-HT1D receptors are found in the basal ganglia, hippocampus, and cortex and act in a similar fashion to 5-HT1B receptors. 5-HT1D receptors are located predominantly on axon terminals of both 5-HT and non-5-HT neurons (Barnes and Sharp, 1999). 5-HT1E receptors are present mainly in the cortex (particularly entorhinal cortex), caudate, putamen, and claustrum, with detectable levels also found in the subiculum of the hippocampus and the amygdala (Barnes and Sharp, 1999). 5-HT1F receptors are located in hippocampal CA1–3, cortex (particularly cingulate and entorhinal cortices) dorsal raphe, claustrum, and caudate (Barnes and Sharp, 1999). There is currently no experimental evidence identifying a functional role for 5-HT1E and 5-HT1F receptors (Barnes and Sharp, 1999; Hoyer and Martin, 1996, 1997; Stamford et al., 2000).

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There are three subtypes of 5-HT2 receptors, labeled 5-HT2A, 5-HT2B, and 5-HT2C. These are similar in terms of their molecular structure, pharmacology, and signal transduction pathways (Barnes and Sharp, 1999). All subtypes of 5-HT2 receptors are coupled to phospholipase C and mobilize intracellular calcium (Barnes and Sharp, 1999). 5-HT2A receptors (previously 5-HT2 receptors) are localized mainly in the cerebral cortex, largely in layer IV (Barnes and Sharp, 1999; Hoyer and Martin, 1997; Kalkman and Fozard, 1991; Peroutka, 1988), but this receptor expression has been identified in the hypoglossal and vestibular nuclei, the inferior olive, and lateral reticular nucleus (Fonseca et al., 2001). These receptors depolarize membrane potential and stimulate phosphatidylinositol hydrolysis. These receptors are downregulated following chronic administration of antidepressant and antipsychotic medications (Eison, 1996). The 5-HT2B receptor is the newest member of the 5-HT2 family. For many years this receptor was unable to be located in the rat brain (unlike mouse and human); however, its presence has been identified and is restricted to just a few brain regions, including the cerebellum, lateral septum, dorsal hypothalamus, and medial amygdala (Barnes and Sharp, 1999). Few specific effects of the 5-HT2B receptor have been identified. 5-HT2C receptors (previously 5-HT1C receptors) are located in very high concentrations in the choroid plexus and are also located in areas of the cortex (olfactory nucleus, pyriform, cingulate, and retrosplenial cortices) and limbic system (nucleus accumbens, amygdala, and hippocampus), as well as the basal ganglia (Barnes and Sharp, 1999). In the medulla, 5-HT2C receptor mRNA is located at high levels in many nuclei, whereas in the spinal cord, 5-HT2C receptor mRNA is located at most levels of the spinal cord except for lamina II (for details, see Fonseca et al., 2001). One of the actions that occurs following activation of 5-HT2C receptors is depolarizing of the pyramidal neurons of the pyriform cortex (Barnes and Sharp, 1999). In the choroid plexus, the resulting action of these receptors occurs by stimulating phosphatidylinositol hydrolysis. There is currently only one receptor gene in the rat that encodes 5-HT3 receptors, although 5-HT3A and 5-HT3B subtypes are reported in the human brain (Barnes and Sharp, 1999). The 5-HT3 receptor is unique in that it is the only 5-HT receptor subtype known to be a ligandgated ion channel (Bloom and Morales, 1998). 5-HT3 receptors are located mainly on peripheral neurons that act like a cation channel by depolarizing the membrane potential (Barnes and Sharp, 1999; Kalkman and Fozard, 1991; Peroutka, 1988). These receptors are located primarily within the dorsal vagal complex in the brain stem, with comparatively low levels elsewhere, of which the highest levels are in mesolimbic regions (Barnes and

Sharp, 1999; Kilpatrick et al., 1996). Associations with GABA-containing cells by 5-HT3 receptors in the neocortex, olfactory cortex, hippocampus, and amygdala have been reported (Bloom and Morales, 1998). Although usually identified as a single receptor type, 5-HT4 receptors are sometimes subdivided into two subtypes, being designated as 5-HT4S and 5-HT4L or, alternatively, 5-HT4(a) and 5-HT4(b), respectively, for the short and long forms of this receptor (Barnes and Sharp, 1999; Hoyer and Martin, 1997). High levels of this receptor are located in the nigrostriatal and mesolimbic systems of the brain and their action is via inhibition of adenylate cyclase (Barnes and Sharp, 1999; Kalkman and Fozard, 1991) and release of acetyl choline in the rat frontal cortex (Eglen et al., 1995). There are putative roles for this receptor in cognition and anxiety (Barnes and Sharp, 1999; Eglen et al., 1995). There are two subtypes of 5-HT5 receptors: 5-HT5A and 5-HT5B. 5-HT5A receptors have a wide distribution in the rat brain, whereas 5-HT5B has a more limited distribution in the supraoptic nucleus of the hypothalamus and some other brain regions, including the subiculum, hippocampal CA1, habenula, dorsal raphe nucleus, olfactory bulb, and the entorhinal and pyriform cortices (Barnes and Sharp, 1999). Like some of the other 5-HT receptors, 5-HT5 receptors may act by inhibition of adenylate cyclase (Hoyer and Martin, 1997). 5-HT6 receptors are a single class of receptors located within the rat brain in the caudate nucleus, olfactory tubercles, nucleus accumbens, and hippocampus (Barnes and Sharp, 1999). This receptor has also been shown to couple positively with adenylate cyclase (Barnes and Sharp, 1999; Hoyer and Martin, 1997). The 5-HT7 receptor is the most recently identified class of 5-HT receptor in the rat. Three isoforms [5-HT7(a), 5-HT7(b), and 5-HT7(c)] are expressed in rats. Receptor expression is high within regions of the thalamus, hypothalamus, and hippocampus (Barnes and Sharp, 1999). The 5-HT7 receptor acts by stimulation of adenylate cyclase (Barnes and Sharp, 1999; Gannon, 2001; Vanhoenacker et al., 2000). The identification of this receptor in the suprachiasmatic nucleus of the hypothalamus suggests a functional role in the serotonergic modulation of circadian rhythms, although proof is still somewhat lacking (Gannon, 2001; Vanhoenacker et al., 2000). In addition to selective membrane receptors for serotonin, selective membrane transporters of serotonin are found on serotonin neurons and glia within the central nervous system (Adell et al., 1991; Gershon and Jonakait, 1979). Such transporters regulate extracellular serotonin concentrations by uptake mechanisms, which may be directed toward both serotonergic and nonserotonergic neurons (Zhou et al., 2002). In the

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future it is likely that several new 5-HT receptors will be found as research continues.

Functions of 5-HT Systems 5-HT is known to have diverse effects in a variety of systems, and this feature has made the identification of separate postsynaptic actions and their integrative effects difficult. 5-HT has been implicated in the central regulation of many autonomic functions (blood pressure, sodium, and glucose balance, body fluid homeostasis), motor behaviors (sex, feeding), nociception, cognition, arousal, and affective behaviors (depression, anxiety). In many instances, 5-HT acts at several neural levels in the normal performance of these functions. It has been shown that different types of ascending 5-HT fibers are functionally distinct as shown by their vulnerability to a variety of 5-HT neurotoxic amphetamines (Battaglia et al., 1991; Mamounas and Molliver, 1988; Mamounas et al., 1991, 1992; Tohyama et al., 1988). B7/9 axons are more susceptible to neurotoxic damage than B5/8 axons, and this preferential damage is important for the psychotropic action of amphetamines and, therefore, the control of affective states. The ablation of B7/9 axons is long lasting and occurs after a single, low dosage of amphetamine (Battaglia et al., 1991; Mamounas et al., 1991) by affecting 5-HT terminal uptake mechanisms (Battaglia et al., 1991). The control of affective states has been emphasized in the hippocampus where 5-HT modulates local inhibitory circuits in a statedependent fashion through a feedforward system (Freund et al., 1990). In contrast to changing affective state, damage to this system does not hamper adequate spatial learning (van Luijtelaar et al., 1991). The 5-HT hippocampal system differs significantly from the feedback septohippocampal system (Freund et al., 1990). In humans, many affective disorders can be exacerbated by the administration of 5-HT2C (formerly 5-HT1C; see earlier discussion) receptor agonists (Kalkman and Fozard, 1991). 5-HT2C receptor agonists also induce a reduction in feeding in rats (Barnes and Sharp, 1999; Kennett and Curzon, 1988), and it has been shown that appetite suppressants damage B7/9 axons in the same manner as amphetamines (Molliver and Molliver, 1990). Reports of stimulus-dependent increases in sympathetic discharge and hyperglycemia after administration of 5HT to the hypothalamus (Sakaguchi and Bray, 1989) or stimulation of descending 5-HT projections (Shian and Lin, 1990) suggest that 5-HT may act at several sites to modulate food intake. This notion is enhanced by reports that the destruction of 5-HT nerves by 5,7dihydroxytryptamine reduces stimulus-induced hyperglycemia (Shian and Lin, 1990). In a similar fashion,

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treatments that increase central extracellular 5-HT increase sodium excretion, and renal denervation attenuates the natriuresis produced by centrally administered 5-HT (Montes and Johnson, 1990). It is likely that these systems are integrated with the hypothalamic regulation of these behaviors. The role of 5-HT in motor activity is controversial, although there is ample evidence for a direct role in modulating activities necessary for many behaviors. In addition to the feeding behaviors associated with food and fluid intake, a discrete 5-HT projection from lateral B3 neurons to pudendal motoneurons inhibits sexual reflexes in rats (Marson and McKenna, 1992). This inhibition is enhanced by 5-HT injections into both B7 and B8 cell groups (Hillegaart et al., 1989), suggesting that 5-HT may act at several neural levels to produce such a response. These 5-HT injections into the B7 cell group also produce a dose-dependent decrease in motor activity, whereas the opposite effect was elicited with 5-HT injections into the B8 cell group (Hillegaart et al., 1989). It is currently speculative to say that these motoric affects can be directly attributed to these 5-HT cell groups. Activation of 5HT1A receptors on 5-HT neurons at the ventral surface of the medulla induces hemodynamic effects, in particular a fall in blood pressure and heart rate due to the inhibition of central sympathetic outflow and/or increase in vagal tone (Kalkman and Fozard, 1991; Ramage, 2001). This effect is limited to the medial B3 cells in the medulla, as 5-HT cells in the lateral B3 group appear to raise blood pressure via a parallel pathway (Minson et al., 1987; Pilowsky et al., 1986). The central receptor involved in this opposing action is the 5-HT2A receptor (Ramage, 2001), with evidence suggesting that perturbations of the 5HT1A and 5HT2A receptors are important in generating and controlling hypertension (Kalkman and Fozard, 1991; Ramage, 2001). The ability of 5-HT to regulate the transmission of nociceptive information at various levels of the nervous system has been appreciated for some time. It is now clear that 5-HT neurons not only modify nociceptive input at spinal and supraspinal levels, but are also involved in mediating the antinociceptive action of morphine and the analgesia produced by the electrical stimulation of certain brain stem regions (Fields et al., 1991). 5-HT B3 neurons are considered to be off cells (suppress nociception) because supraspinal injections of opioids release 5-HT in the spinal cord (Fields et al., 1991). B3 cells are excited by local 5-HT (5-HT2 receptor mediated), whereas nearby neurons are inhibited by 5-HT (5-HT1 receptor mediated). These nearby cells are thought to be on cells (Fields et al., 1991). Thus 5-HT exerts the similar effect of nociception

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suppression. The spinal release of 5-HT produces antinociception via 5-HT1A and 5-HT3 receptors (Fields et al., 1991). These receptors are found on primary afferent terminals, with 5HT1A receptors inhibiting nociceptive dorsal horn neurons. The blockade of 5-HT3 receptors causes hyperalgesia and reverses 5-HT antinociception (Fields et al., 1991). The anatomical diversity within the serotonergic system is reflected by the multiple physiological, biochemical, and behavioral effects produced by central 5-HT neurons. Future studies are likely to identify more specific anatomical circuits coupled to particular 5-HTmediated postsynaptic mechanisms underlying characteristic behaviors.

TACHYKININ SYSTEM The undecapeptide substance P (SP) was first detected in crude brain extract by von Euler and Gaddum (1931) and was found to cause a wide spectrum of physiological effects. It was only much later in the 20th century that it was generally recognized that SP is just one member of a structurally related family of bioactive peptides: the tachykinins (see Hökfelt et al., 2001). The term “tachykinin” literally means fast acting, and its compounds contain the COOH-terminal sequence of Phe-X-Gly-Leu-Met-NH2 (Helke et al., 1990; Maggio, 1988) (Table 2). Tachykinins are widely distributed and active in both the central and the peripheral nervous systems, evoking a variety of responses in a variety of tissues. Unlike classical neurotransmitters, tachykinins are synthesized ribosomally as larger protein precursors in the neuronal cell body and shipped to the terminals. Enzymatic processing within the axon converts them to mature forms. Mammalian tachykinins are the products of three genes. The SP/NKA gene is the preproachykinin I (PPT1 or PPTa) gene. The preprotachykinin II (PPTII or PPTb) gene is the gene for neurokinin B (NKB) (Helke et al., 1990; Hökfelt et al., 2001; Maggio, 1988). A gene labeled as the preprotachykinin III (PPTIII or PPTc) gene for a novel tachykinin, hemokinin 1 (HEK-1), has been reported (Bellucci et al., 2002; Zhang et al., 2000). This tachykinin has not had a proven presence in the rat nervous system, being identified

originally in hemopoietic tissue. However, its actions do not appear to be mediated by a unique receptor but through the same receptor (NK1; see later) as that used by SP (Bellucci et al., 2002; Camarda et al., 2002). Due to the early stage of investigation of this tachykinin, further discussion is currently premature. Four precursors for tachykinins are produced by the SP/NKA gene, α-, β-, γ-, and δ-preprotachykinins (Table 2), as a result of differential RNA splicing and differential posttranslational processing (Helke et al., 1990; Hökfelt et al., 2001; Maggio, 1988). The structure of preprotachykinins suggests but does not require a similar regional distribution for SP and NKA, and in several regions the tachykinins are colocalized, cosynthesized, and coreleased (Helke et al., 1990; Maggio, 1988). Despite this, SP is found in higher concentrations than NKA in most locations (Kanazawa et al., 1984). Although tachykinins can be found in most neuroanatomical regions (Harlan et al., 1989; Marksteiner et al., 1992; Warden and Young, 1988), the distribution of tachykinin precursors reveals that brain regions usually contain only one tachykinin type. For example, the nucleus of the lateral olfactory tract contains only NKB cells, whereas raphe nuclei contain only SP cells (Warden and Young, 1988). Unfortunately, many antisera used in tachykinin research have been directed against the common tachykinin C terminus (with exceptions) (Brodin et al., 1986; Kanazawa et al., 1984; Marksteiner et al., 1992) that are therefore unable to discriminate between different tachykinins. Our understanding of the tachykinin system and specific tachykinin pathways and function is therefore still rudimentary. In the following text, the distribution of various tachykinins is noted from the literature where possible, although the material presented in the figures has been stained using a monoclonal antibody raised against the C terminus of SP (Seralab) and therefore cannot discriminate between the tachykinins.

Location of Tachykinin-Containing Cell Bodies Because the processing of tachykinins to their mature form occurs within axons, few studies have reported the distribution of tachykinin-containing cells in the central nervous system without the use of colchicine to

FIGURES 3–27 Photomicrographs of serial sections every 500 μm through one rat brain. Sections are stained immunohistochemically for serotonin [(Incstar, 1:10,000 dilution) (left side of photomicrographs in Figs. 16–27)] and substance P [(Sera-lab, 1:1000 dilution) (right side of photomicrographs in Figs. 16–27 and 3–15)]. The scale bar in Fig. 27 represents 500 μm.

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FIGURE 3

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FIGURE 4

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FIGURE 5

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FIGURE 6

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FIGURE 7

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FIGURE 8

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FIGURE 9

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FIGURE 10

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FIGURE 11

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FIGURE 12

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FIGURE 13

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FIGURE 14

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FIGURE 15

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FIGURE 16

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FIGURE 17

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FIGURE 18

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FIGURE 19

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FIGURE 20

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FIGURE 21

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FIGURE 22

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FIGURE 23

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FIGURE 24

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FIGURE 25

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FIGURE 26

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FIGURE 27

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TABLE 2 Gene

Tachykinin Peptide Family

Preprotachykinin mRNA

SP/NKA (PPT I; PPTa)

Peptide

α,β,γ,δ

Substance P (SP)

β,γ

Neurokinin A (NKA)

β

Neuropeptide K (NPK)

γ

Neuropeptide γ (NPγ)

β,γ

Neurokinin A3–10 (NKA3–10)

NKB (PPT II: PPTb)

—

Neurokinin B (NKB)

PPT III (PPTc)a

—

Hemokinin 1 (HK-1)a

a

Requires further studies to confirm presence in rat nervous system.

eliminate axoplasmic transport (Ljungdahl et al., 1978; Shults et al., 1984). Most studies utilizing this technique have limited their study to discrete regions. The ontogeny of tachykinin-containing neurons shows that many neurons contain mature tachykinins during development (Inagaki et al., 1982; Sakanaka et al., 1982a). Other studies visualizing specific tachykinin mRNA have determined the distribution of SP/NKA- and NKB-containing neurons in adult rats (Harlan et al., 1989; Marksteiner et al., 1992; Warden and Young, 1988). Tachykinin-containing neurons appear on gestational day 14 in the central trigeminal system (Inagaki et al., 1982; Sakanaka et al., 1982a). The early appearance of mature tachykinins in the nervous system has stimulated speculation that tachykinins may have an additional role in neural development (Inagaki et al., 1982; Sakanaka et al., 1982a). Several tachykinin-containing cell groups develop after birth and tend to retain mature tachykinin within their neuronal somata throughout adulthood. The remaining tachykinin-containing neurons progressively decrease in number with maturity (Inagaki et al., 1982; Sakanaka et al., 1982a). SP/NKA neurons are concentrated in spinal cord laminae 1 and 2 with occasional cells in lamina 5 (Ljungdahl et al., 1978; McGonigle et al., 1996; Warden and Young, 1988). The lateral cervical nucleus in the cervical cord and lateral spinal nucleus elsewhere contains significant numbers of SP/NKA neurons (Ljungdahl et al., 1978; Warden and Young, 1988). In addition, SP/NKA neurons can be seen throughout the intermediolateral cell column. In contrast, NKB neurons are most prominent in lamina 3 and are also found in dorsal root ganglia cells (Ljungdahl et al., 1978; Warden and Young, 1988). The medulla oblongata is rich in tachykinin cell groups. Most striking is the number of SP/NKA neurons within the medullary raphe (magnus, pallidus, and obscurus) (Harlan et al., 1989; Ljungdahl et al., 1978; Shults et al., 1984; Thor et al., 1988; Warden and Young, 1988). SP/NKA neurons are prominent in the

solitary complex, whereas scattered SP/NKA neurons are found in trigeminal and vestibular complexes and areas of the reticular formation (Harlan et al., 1989; Leibstein et al., 1985; Ljungdahl et al., 1978; Warden and Young, 1988). In contrast, NKB neurons are found in substantial numbers in the external cuneate nucleus and occasionally in the spinal trigeminal nucleus (Marksteiner et al., 1992; Warden and Young, 1988). The cerebellum contains few tachykinin-containing neurons and only occasional NKB neurons are found in the pons within the reticular formation (Marksteiner et al., 1992; Warden and Young, 1988). SP/NKA neurons are prominent in the dorsal tegmental nucleus, lateral dorsal tegmental nucleus, locus coeruleus, parabrachial nuclei, raphe cell groups (B5/8 and B6), and the pontine reticular formation (Harlan et al., 1989; Ljungdahl et al., 1978; Shults et al., 1984; Warden and Young, 1988). In the midbrain, occasional SP/NKA neurons are seen in the ventral tegmental area and dorsal regions of the mesencephalic tegmentum, although many areas contain large numbers of tachykinin-containing neurons (Harlan et al., 1989; Ljungdahl et al., 1978; Shults et al., 1984; Warden and Young, 1988). The interpeduncular nucleus contains numerous SP/NKA neurons (Harlan et al., 1989; Ljungdahl et al., 1978; Shults et al., 1984; Warden and Young, 1988), which have mature tachykinin within their somata throughout adulthood (Sakanaka et al., 1982a) (Figs. 15 and 16). The superficial layers of the superior colliculus contains numerous SP/NKA neurons as well as NKB neurons (Warden and Young, 1988), which also have mature tachykinin throughout adulthood (Inagaki et al., 1982; Sakanaka et al., 1982a) (Figs. 14–17). In contrast, only SP/NKA neurons are found in the deeper layers of the superior colliculus and throughout the inferior colliculus (Harlan et al., 1989; Ljungdahl et al., 1978; Shults et al., 1984; Warden and Young, 1988). Both types of tachykinin-containing neurons are found in discrete regions of the periaqueductal gray; SP/NKA neurons are located in central regions whereas NKB neurons are seen in dorsal regions (Warden and Young,

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1988). Many large SP/NKA neurons are found in the lateral dorsal tegmental nucleus and the pedunculopontine tegmental nucleus (Harlan et al., 1989; Ljungdahl et al., 1978; Shults et al., 1984; Warden and Young, 1988). In addition, the Edinger–Westphal nucleus contains a large number of SP/NKA neurons (Harlan et al., 1989; Ljungdahl et al., 1978; Shults et al., 1984; Warden and Young, 1988). Interestingly, the dorsal raphe B7 group contains SP/NKA neurons (Harlan et al., 1989), whereas all tachykinins are found within the same neurons of the B9 cell group (Warden and Young, 1988). The hypothalamus is rich in both SP/NKA and NKB neurons (Nussdorfer and Malendowicz, 1998). SP/NKA neurons are especially abundant in dorsomedial, ventromedial, and premamillary nuclei (Harlan et al., 1989; Ljungdahl et al., 1978; Warden and Young, 1988) with many of these neurons containing mature tachykinin throughout adulthood (Inagaki et al., 1982). Lower densities of SP/NKA neurons are seen in most other regions, including preoptic, dorsolateral, lateral, posterior, suprachiasmatic, retrochiasmatic, septohippocampal, arcuate, and supramamillary nuclei (Harlan et al., 1989; Kawano et al., 1989; Larsen, 1992; Ljungdahl et al., 1978; Shults et al., 1984; Warden and Young, 1988). NKB neurons are numerous in the anterior hypothalamic area and lateral mamillary nuclei (Marksteiner et al., 1992; Warden and Young, 1988) and have been reported in the periventricular nucleus and the medial magnocellular paraventricular subnucleus (Koutcherov et al., 2000). Fewer NKB neurons are found in medial preoptic, dorsal, lateral, posterior, suprachiasmatic, retrochiasmatic, septohippocampal, and arcuate nuclei (Marksteiner et al., 1992; Warden and Young, 1988). The habenula has a high concentration of SP/NKA neurons (Harlan et al., 1989; Ljungdahl et al., 1978; Marksteiner et al., 1992; Shults et al., 1984; Warden and Young, 1988) with most neurons in the medial habenula containing NKB as well (Marksteiner et al., 1992; Warden and Young, 1988). These neurons contain visible tachykinin throughout adulthood (Inagaki et al., 1982). Some SP/NKA neurons are found in precommissural and lateral geniculate nuclei of the thalamus (Takatsuji and Tohyama, 1989; Warden and Young, 1988). In the olfactory bulb, SP/NKA mitral and granule neurons are found in main and anterior olfactory nuclei (Gouda et al., 1990; Harlan et al., 1989; Warden and Young, 1988). In contrast, NKB periglomerular neurons are seen throughout the anterior olfactory nucleus (Marksteiner et al., 1992; Warden and Young, 1988). In the basal forebrain, SP/NKA neurons were abundant in the islands of Calleja and the olfactory tubercle (Harlan et al., 1989; Warden and Young, 1988). NKB neurons are present in the dorsolateral olfactory tubercle descending from the tip of the corpus callosum, in the

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striatal cell bridges descending from the ventral striatum, and in the basal nucleus (Marksteiner et al., 1992; Warden and Young, 1988). All divisions of the septum contain SP/NKA neurons, whereas NKB neurons are found only in medial and lateral septal nuclei (Harlan et al., 1989; Marksteiner et al., 1992; Warden and Young, 1988). Occasional tachykinin-containing neurons are found throughout both limbs of the diagonal band of Broca (Warden and Young, 1988). The caudate-putamen and accumbens nuclei contain numerous tachykinin neurons in a fairly homogeneous pattern (Furuta et al., 2000; Harlan et al., 1989; Ljungdahl et al., 1978; Marksteiner et al., 1992; Shults et al., 1984; Warden and Young, 1988), which are visible throughout adulthood (Inagaki et al., 1982). Similarly, tachykinin-containing neurons are found throughout the entopeduncular nucleus (Murakami et al., 1989). Bed nuclei of the stria terminalis contain large numbers of both SP/NKA and NKB neurons (Harlan et al., 1989; Ju et al., 1989; Marksteiner et al., 1992; Shimada et al., 1989), suggesting some colocalization of these tachykinins in this region. In contrast, all neurons in nuclei of the olfactory tract contain NKB only (Marksteiner et al., 1992; Warden and Young, 1988). Most amygdoid nuclei contain SP/NKA neurons, especially the anterior cortical, central, and medial nuclei (Cassell and Gray, 1989; Harlan et al., 1989; Ljungdahl et al., 1978; Shimada et al., 1989; Shults et al., 1984; Warden and Young, 1988). Scattered NKB neurons are found throughout the amygdala with large numbers present in central and basolateral nuclei (Marksteiner et al., 1992; Warden and Young, 1988). Tachykinin-containing neurons were scattered through all regions of the hippocampus and neocortex (Borhegyi and Leranth, 1997; Harlan et al., 1989; Marksteiner et al., 1992; Warden and Young, 1988). NKB neurons are concentrated along the hilar surface of the dentate gyrus, but are found throughout the hippocampus (Marksteiner et al., 1992; Warden and Young, 1988). Fewer SP/NKA neurons are seen in the hippocampus, but many SP/NKA neurons are present in the subiculum and entorhinal cortex (Borhegyi and Leranth, 1997; Harlan et al., 1989; Warden and Young, 1988). In the neocortex, SP/NKA neurons concentrate in layer 4, whereas NKB neurons concentrate in layers 2–4 (Harlan et al., 1989; Marksteiner et al., 1992; Warden and Young, 1988). There appears to be a differential distribution of these neurons throughout the neocortex, although this is not well documented.

Main Tachykinin Pathways Although there are extensive numbers of tachykinin neurons throughout the central nervous system,

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tachykinin fibers innervate discrete regions. Most previous studies have not been able to distinguish between specific tachykinin innervation (Ljungdahl et al., 1978; Shults et al., 1984), although studies have published the distribution of NKB fibers in the central nervous system (Koutcherov et al., 2000; Marksteiner et al., 1992) and SP innervation of the rat hippocampal formation (Borhegyi and Leranth, 1997). The following description of tachykinin fiber innervation concentrates on extrinsic projections. It should be noted that in many instances tachykinin neurons also act as local interneurons. Tachykinin Projections to the Spinal Cord Very high densities of tachykinin fibers are found in discrete regions of the spinal cord (Ljungdahl et al., 1978; Sakamoto et al., 1999; Shults et al., 1984; Todd, 2002), particularly laminae 1 and 2. Throughout the thoracic cord, a dense tachykinin network is found in the intermediolateral cell column. Tachykinin fibers also run from the lateral cell column toward the central canal. The remaining laminae contain only scattered tachykinin fibers. Many tachykinin fibers terminating in the dorsal horn are thought to be the central axons of sensory dorsal horn neurons. It has also been known for some time that tachykinin-producing neurons in the ventral medulla project to the spinal cord, particularly those in the raphe nuclei (Araneda et al., 1989; Bowker et al., 1981a; Johansson et al., 1981; Magoul et al., 1988). It appears that only tachykinin neurons in the ROb and RPa, as well as tachykinin neurons in the lateral B1 area, project to the spinal cord. In contrast, tachykinin neurons in the RMg and surrounding tegmentum do not project to spinal regions (Menetrey and Basbaum, 1987; Strack et al., 1989). It has been shown that many tachykinin neurons in these regions project to the intermediolateral cell column (Charlton and Helke, 1987; Sasek and Helke, 1989), as well as dorsal horn regions. In the dorsal horn, tachykinins are found in type I synaptic glomeruli (Ribeiro-da-Silva et al., 1989). Brain Stem and Cerebellar Tachykinin Projections Within the medulla oblongata, dense tachykinin fibers are detected in many reticular nuclei (Ljungdahl et al., 1978; Shults et al., 1984), in particular the parvocellular, intermediate and lateral paragigantocellular reticular nuclei (Figs. 23–27). Raphe nuclei receive tachykinin innervation from medullary neurons in the lateral paragigantocellular reticular nucleus, the trigeminal division of the lateral reticular nucleus, and the solitary complex (Beitz, 1982a), as well as neurons in the cuneate nucleus, the cholinergic pedunculopontine tegmental nucleus, the B5–B8 groups, and the periaqueductal gray of the midbrain (Beitz, 1982a; Zeng et al., 1991). In the dorsomedial medulla (Figs. 23–27), the

solitary complex and dorsal motor vagal nucleus are very densely innervated (Ljungdahl et al., 1978; Shults et al., 1984), with the solitary complex containing NKBimmunoreactive fibers (Marksteiner et al., 1992). Parvicellular adrenergic neurons within the solitary complex receive a dense innervation (Kawai and Takagi, 1989), possibly from tachykinin cells located in the caudal raphe nuclei (Thor and Helke, 1987; Thor et al., 1988). Tachykinin fibers from peripheral ganglia (nodose and petrosal ganglia) innervate the solitary complex (Thor et al., 1988). In the dorsal motor vagal nucleus, tachykinin innervation is restricted to the region containing neurons projecting to the stomach (Buchan et al., 1991). The tachykinin innervation to these cells is from diverse sources, including the caudal raphe nuclei (Thor and Helke, 1987; Thor et al., 1988), the ventral division of the lateral bed nucleus of the stria terminalis, and the medial division of the central amygdaloid nucleus (Shimada et al., 1989). The spinal trigeminal nucleus is also densely innervated (Figs. 23–27), with a similar distribution to that observed in the dorsal horn of the spinal cord (Ribeiro-da-Silva et al., 1989). NKB-immunoreactive fibers have been identified in this nucleus (Marksteiner et al., 1992). There is a discrete tachykinin innervation of specific regions of the inferior olivary complex that has not been well described in past studies (Ljungdahl et al., 1978; Shults et al., 1984). In the pons, dorsal and medial parabrachial nuclei (Fig. 20) contain a high density of tachykinin fibers (Ljungdahl et al., 1978; Shults et al., 1984), including fibers containing NKB (Marksteiner et al., 1992). This innervation arises from a number of diverse sources, including the caudal, medial solitary complex (Mantyh and Hunt, 1984; Milner et al., 1984; Milner and Pickel, 1986; Riche et al., 1990), the caudal ventrolateral reticular formation, the lateral dorsal tegmental nucleus, the paraventricular, dorsomedial, and lateral hypothalamic nuclei (Milner and Pickel, 1986), the ventral division of the lateral bed nucleus of the stria terminalis, and the medial division of the central nucleus of the amygdaloid complex (Shimada et al., 1989). The catecholaminergic A7 group also receives tachykinin innervation from tachykinin cells located in the raphe magnus nucleus and surrounding reticular formation (Kohlmeier et al., 2002; Yeomans and Proudfit, 1990). Most periaqueductal gray structures receive a moderate innervation of tachykinin fibers (Figs. 19–22), including the caudal DR and locus coeruleus (Ljungdahl et al., 1978; Shults et al., 1984). A few NKB fibers are found in the MnR and surrounding reticular formation (Marksteiner et al., 1992), while very few tachykinin fibers are observed in the cerebellum (Ljungdahl et al., 1978; Marksteiner et al., 1992; Shults et al., 1984).

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The highest density of tachykinin fibers is found in midbrain structures (Figs. 13–15), particularly the substantia nigra and interpeduncular complex. In the substantia nigra, tachykinin fibers from the striatal neurons (Anderson and Reiner, 1990; Bolam and Smith, 1990) innervate the pars reticulata, pars compacta, and pars lateralis (Lee et al., 1997; Ljungdahl et al., 1978; Shults et al., 1984), with NKB-immunoreactive fibers present only in the pars compacta and pars lateralis (Marksteiner et al., 1992). Most tachykinin fibers in the substantia nigra innervate dopaminergic neurons (Bolam and Smith, 1990). The more medially situated ventromedial tegmentum also contains high densities of tachykinin fibers (Ljungdahl et al., 1978; Shults et al., 1984), in particular the ventral tegmental nuclei and interfascicular nucleus (Halliday and Törk, 1988). The interpeduncular nucleus is innervated by tachykinin fibers in a discrete manner, with fibers concentrated in the rostral ventrolateral, caudal intermediate, and lateral nuclei (Ljungdahl et al., 1978; Shults et al., 1984) and are NKB positive (Marksteiner et al., 1992). This innervation arises solely from habenular nuclei via the fasciculus retroflexus (Marksteiner et al., 1992) (Fig. 12). The periaqueductal gray, particularly dorsally, contains a high density of tachykinin fibers (Ljungdahl et al., 1978; Shults et al., 1984), as does the nearby cuneiform nucleus (Figs. 17–20). In contrast, NKB-immunoreactive fibers concentrate in the more ventral DR (Marksteiner et al., 1992). The superior colliculus contains a high density of tachykinin fibers in specific regions: zona layer, superficial gray, intermediate gray, and deep gray (Ljungdahl et al., 1978; Shults et al., 1984) (Figs. 14–17). Tachykinin fibers are also found in the area containing B9 neurons (Ljungdahl et al., 1978; Shults et al., 1984) (Figs. 16–18). Tachykinin Fibers Innervating Forebrain Structures Many ascending tachykinin fibers projecting to the forebrain congregate in the medial forebrain bundle before diverging into different fiber pathways to innervate their target structures. The main target structures of these fibers are the cortex, amygdala, hypothalamus, and septum (Ljungdahl et al., 1978; Marksteiner et al., 1992; Shults et al., 1984). In the diencephalon, high densities of tachykinin fibers are found in most of the hypothalamic nuclei and in midline nuclei of the thalamus (Ljungdahl et al., 1978; Nussdorfer and Malendowicz, 1998; Shults et al., 1984) (Figs. 8–12). In the preoptic hypothalamic area (Figs. 7 and 8), high fiber densities are found in medial, lateral, anterior suprachiasmatic, and periventricular nuclei (Larsen, 1992; Ljungdahl et al., 1978; Shults et al., 1984). Tachykinin neurons in the arcuate and the ventrolateral portion of the anterior hypothalamic nucleus project to neurons containing luteinizing hormorne-

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releasing hormone in the septopreoptic area (Tsuruo et al., 1991) (Figs. 7 and 8). Innervation to the anterior suprachiasmatic nucleus is directly from tachykinin containing retinal ganglia cells (Takatsuji et al., 1991). The anterior medial preoptic area and periventricular nucleus contain many NKB-immunoreactive fibers (Marksteiner et al., 1992). The anterior hypothalamic nucleus, the dorsal part of the supraoptic nucleus and the paraventricular nucleus contain moderate numbers of tachykinin fibers (Larsen, 1992; Ljungdahl et al., 1978; Shults et al., 1984) (Figs. 7 and 8), with a proportion of these containing NKB (Marksteiner et al., 1992). In the caudal magnocellular nucleus, tachykinin fibers target histaminergic neurons (Tamiya et al., 1990). Once again, tachykinin innervation of magnocellular hypothalamic nuclei (Fig. 11) comes from more than one source. These include the bed nucleus of the stria terminalis, the anterior periventricular nucleus, the dorsomedial hypothalamic nucleus, the ventromedial hypothalamic nucleus, the lateral hypothalamic area, the pedunculopontine tegmental nucleus (cholinergic), the laterodorsal tegmental nucleus (also cholinergic), and tachykinin neurons located in medullary catecholamine groups C1, C2, C3, A1, and A2 (Bittencourt et al., 1991). High densities of tachykinin fibers are found in the lateral hypothalamic area (Larsen, 1992; Ljungdahl et al., 1978; Shults et al., 1984) (Fig. 11). In the medial hypothalamus, high densities of tachykinin fibers are found in the arcuate nucleus, the dorsomedial nucleus, the periformical area, and the median eminence (Larsen, 1992; Ljungdahl et al., 1978; Shults et al., 1984) (Figs. 9–12). Most of these areas contain NKB-immunoreactive fibers, particularly the median eminence (Marksteiner et al., 1992). Tachykinin neurons in the ventromedial arcuate nucleus are the source of innervation for the median eminence (Palkovits et al., 1989) (Figs. 9–11). In the posterior hypothalamus, high densities of tachykinin fibers are found in posterior premamillary nuclei (Ljungdahl et al., 1978; Shults et al., 1984), and high densities of NKB fibers are seen in the lateral mamillary bodies (Marksteiner et al., 1992). Large regions of the thalamus contain relatively few tachykinin fibers (Ljungdahl et al., 1978; Marksteiner et al., 1992; Shults et al., 1984). Despite this, tachykinin spinothalamic tract neurons in spinal cord laminae 1 and 5 project to somatosensory thalamic nuclei (Battaglia and Rustioni, 1992). High densities of tachykinin fibers are found in the lateral habenular nucleus (Ljungdahl et al., 1978; Shults et al., 1984) (Figs. 10–12). These fibers originate in the entopeduncular nucleus (Murakami et al., 1989). Other thalamic nuclei containing tachykinin fibers include paraventricular, centromedial, mediodorsal, and ethomoid nuclei (Ljungdahl et al., 1978; Shults et al., 1984) (Figs. 9–13). Tachykinin cells in the

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parabrachial nucleus project to the medial part of the posterior thalamic nuclear group (Mantyh and Hunt, 1984), and tachykinin neurons in the superficial layers of the superior colliculus project to the lateral geniculate nucleus (Ogawa-Meguro et al., 1992). The basal forebrain and septal regions also contain tachykinin fibers (Ljungdahl et al., 1978; Marksteiner et al., 1992; Shults et al., 1984). In the lateral septum, there are dense zones of tachykinin fibers, with the tachykinin fibers often basketing septal neurons (Ljungdahl et al., 1978). Some NKB-immunoreactive fibers are found in both lateral and medial septal nuclei (Marksteiner et al., 1992). Tachykinin innervation to the spetum arises from neurons located between the anterior hypothalamic nucleus and the lateral hypothalamus (Sakanaka et al., 1982b), as well as from neurons in the lateral dorsal tegmental nucleus (Sakanaka et al., 1981, 1982b). The cholinergic basal forebrain also contains a high density of tachykinin fibers (Ljungdahl et al., 1978; Shults et al., 1984), including NKB-immunoreactive fibers (Marksteiner et al., 1992). The olfactory tubercle (Figs. 3–6) contains a dense tachykinin fiber network, particularly in the plexiform layer (Ljungdahl et al., 1978; Shults et al., 1984) where NKB fibers congregate (Marksteiner et al., 1992). The basal ganglia also contains a dense network of tachykinin fibers (Ljungdahl et al., 1978; Marksteiner et al., 1992; Shults et al., 1984). The distribution of tachykinin fibers in the caudate-putamen features a number of high density patches, particularly in the ventral anterior part with a decreasing gradient in the dorsal and posterior direction (Furuta et al., 2000; Ljungdahl et al., 1978; Shults et al., 1984) (Figs. 3–9). There are high-density bands along the ventrolateral border and abutting the globus pallidus. The globus pallidus contains a high density of tachykinin fibers (Ljungdahl et al., 1978; Shults et al., 1984) (Figs 7–11), with many fibers originating from striatal tachykinin neurons (Anderson and Reiner, 1990). Another group of principally NKB-immunoreactive fibers originating from striatal tachykinin neurons has been investigated with projections to the substantia innominata (Furuta et al., 2000). The anterior and medial parts of the accumbens nucleus contain high densities of tachykinin fibers (Ljungdahl et al., 1978; Shults et al., 1984) (Figs. 3 and 4) that originate from at least two sources: medial and commissural nuclei of the solitary complex (Li et al., 1990b) and the ventral and ventrolateral periaqueductal gray (Li et al., 1990a). Some tachykinin fibers in the accumbens nucleus contain NKB (Marksteiner et al., 1992). The amygdala receives some tachykinin innervation, mainly to its central and medial nuclei (Ljungdahl et al., 1978; Shults et al., 1984). The central amygdaloid nucleus (Fig. 9) contains many NKB-immunoreactive

fibers, as basolateral and intercalated amygdaloid nuclei (Marksteiner et al., 1992) (Fig. 10). The posterior division of the medial amygdaloid nucleus (Figs. 10 and 11) is sexually dimorphic with twice the area innervated by tachykinins in male rats (Malsbury and McKay, 1989). The high density of tachykinin fibers throughout bed nuclei of the stria terminalis (Figs. 5–11) includes many NKB-immunoreactive fibers (Ljungdahl et al., 1978; Marksteiner et al., 1992; Shults et al., 1984). This region contains one of the most dense NKB fiber plexuses (Marksteiner et al., 1992). Many of the tachykinin fibers within bed nuclei of the stria terminalis originate in the amygdala. In concert with the enlargement of the medial amygdaloid nucleus in male rats, the posterior medial division of the bed nucleus of stria terminalis (Fig. 7) is also twice as large in male rats (Malsbury and McKay, 1989). There is only a sparse tachykinin innervation of the hippocampus (Borhegyi and Leranth, 1997; Ljungdahl et al., 1978; Shults et al., 1984) (Figs. 9–12), with the most dense innervation in the molecular layer of the dentate gyrus, in the stratum moleculare of CA1, and in the pyramidal layer of CA2 (Iritani et al., 1989). Intrinsic SP innervation from within the hippocampus is suggested as arising from the interneurons of CA1–3 and subiculum and granule cells (Borhegyi and Leranth, 1997). NKB-immunoreactive terminals concentrate in the stratum lucidum of CA3 where mossy fibers terminate (Marksteiner et al., 1992). There is an increased density of tachykinin fibers in the subiculum and entorhinal cortex (Iritani et al., 1989). Tachykinin fibers, including those containing NKB, are found in all regions of the cortex (Iritani et al., 1989; Ljungdahl et al., 1978; Marksteiner et al., 1992; Shults et al., 1984). These fine varicose fibers are concentrated in layers 2 and 4 (Iritani et al., 1989). An increased concentration of fine tachykinin fibers and the addition of thick varicose tachykinin fibers are found in medial prefrontal cortical areas (Iritani et al., 1989; Ljungdahl et al., 1978; Shults et al., 1984). Tachykinin neurons in the lateral dorsal tegmental nucleus are known to project to the medial prefrontal cortex (Sakanaka et al., 1983; Vincent et al., 1983). In addition to these central nervous system tachykinin projections, several studies have shown that cerebral blood vessels are densely innervated by tachykinin fibers (Reuss et al., 1992; Suzuki et al., 1989). These tachykinin fibers are sensory in nature and are the peripheral axon of tachykinin neurons in the trigeminal ganglia (opthalmic division) and other cranial and upper cervical dorsal root ganglia (Suzuki et al., 1989). Approximately one-third of trigeminal ganglia neurons contain tachykinins and innervate the meninges and meningeal arteries (Reuss et al., 1992).

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Tachykinin Receptor Types and Location In addition to the multiplicity of tachykinin peptides throughout the central nervous system, at least three pharmacologically distinct postsynaptic receptors have been defined largely on the rank order of potency and binding of tachykinins in bioassays. Thus the receptors, neurokinin-1 (NK-1), NK-2, and NK-3, each display a moderate degree of selectivity for SP, NKA, and NKB, respectively (Helke et al., 1990; Nakanishi, 1991). Early reports of a putative fourth tachykinin receptor (NK-4) have been published (Donaldson et al., 2001). Each of the tachykinins is capable, if present in sufficiently high concentrations, of stimulating all these receptors, presumably because of the similarity in their carboxy terminals. All of the three well-established receptor subtypes contain the seven transmembrane-spanning domains characteristic of members of the G-proteincoupled receptor family (Nakanishi, 1991). However, the three well-recognized receptor subtypes have different distributions in the central nervous system. The tachykinin NK-1 receptor is widely distributed in both central and peripheral nervous systems (Quartara and Maggi, 1998; Saria, 1999). In the central nervous system, the NK-1 receptor is found in high concentration in the dorsal horn of the spinal cord, particularly in layers 1 and 2 where they are located postsynaptic to sensory nerve terminals (Helke et al., 1990; Todd, 2002). There is also a population of NK-1 receptor-immunoreactive neurons in layers 3 and 4, whose dendrites arborize with those in layers 1 and 2 (Sakamoto et al., 1999). In the ventral horn and intermediolateral cell column, NK-1 receptors are located on autonomic and motor neurons (Helke et al., 1990). High densities of NK-1 receptor-binding sites are found in the striatum, stria terminalis, septohippocampal nucleus, and the accumbens nucleus, with moderate densities in the amygdala, habenula, periventricular nucleus, and the olfactory bulb (Beaujouan et al., 2000; Helke et al., 1990; Mazzone and Geraghty, 2000; Nakanishi, 1991; Quartara and Maggi, 1998; Saria, 1999). NK-1 receptors are also found on glial cells. Differences in the reaction of NK-1 receptors to different agonists and antagonists have led to the suggestion that different subtypes of NK-1 receptor exist (Beaujouan et al., 2000; Glowinski, 1995). NK-2 receptors are mainly located in peripheral tissues and for some time there was some controversy over whether they were present at all in the central nervous system (Helke et al., 1990; Nakanishi, 1991). Highly selective radioligands now available show highest NK-2 receptor densities in the hippocampus, septum, and thalamus with much lower densities elsewhere in the brain (Helke et al., 1990; Saffroy et al., 2001).

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NK-2 receptors in the spinal cord are only associated with the dorsal horn, presumably the sensory afferent system (Helke et al., 1990). NK-3 receptors are found primarily in the central nervous system, most notably in the neocortex, septum, diagonal band of Broca, hypothalamus, zona incerta, amygdala, substantia nigra, and raphe nuclei (Ding et al., 1996; Koutcherov et al., 2000; Massi et al., 2000; Mileusnic et al., 1999; Shughrue et al., 1996). In contrast to the NK-1 receptor, NK-3 receptors are dense in the deep layers of the cerebral cortex (Helke et al., 1990; Nakanishi, 1991). Because NK-3-binding sites are found in a similar distribution to central NK-2 receptors, there is some suggestion that NK-2 receptors are really NK-3 receptors (Helke et al., 1990; Nakanishi, 1991). A putative new receptor, NK-4, has been reported that is highly homologous to the NK-3 receptor and may be also referred to as the NK-3B receptor (Donaldson et al., 2001). This receptor is reported as being widely expressed in the rat central nervous system, including the cerebral cortex, hippocampus, hypothalamus, and dorsal horn of the spinal cord. This receptor may also be present in tissues other than the central nervous system (Donaldson et al., 2001).

Function of Tachykinin Peptides The functions of tachykinins have mainly been determined by their location and distribution. Thus tachykinin peptides have been implicated in motor, sensory, cardiovascular, respiratory, and gastrointestinal functions. In many instances, tachykinins are thought to act at several neural levels in the normal performance of these functions. Since the discovery that the highest concentration of SP occurred in the substantia nigra, the effects of SP on dopaminergic neurons have been widely studied. It has been shown that SP and NKA both cause release of dopamine in the striatum (Reid et al., 1990), although there has been some controversy over the most potent tachykinin in this pathway (Deutch et al., 1985; Helke et al., 1990; Tremblay et al., 1992). Disruption of this pathway produces the parkinsonian features of rigidity and bradykinesia. Consistent with this is the report that striatal tachykinin neurons are selectively activated when challenged after dopamine denervation (Zhang et al., 1992). Tachykinins are important peptides in the transmission of sensory signals, particularly nociception (Doak and Sawynok, 1997; Hökfelt et al., 2001; Saria, 1999). All three tachykinins are found at multiple sites in areas processing pain signals (Battaglia and Rustioni, 1992), including input to parallel processing pathways (Yeomans and Proudfit, 1990). Intrathecal administration

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of selective tachykinin receptor agonists has revealed that NK-1 receptors participate in nociception, whereas NK-3 receptors participate in antinociception (Helke et al., 1990). Anxiolytic-like and antidepressant-like effects have also been reported following administration of NK-1 receptor antagonists and NK-3 receptor agonists in rats (Massi et al., 2000). It has also been noted that tachykinins are associated with the peripheral projections of sensory nerves. Tachykinins are also involved in the neural control of cardiovascular events. Tachykinin antagonists decrease mean arterial pressure by 33% but leave heart rate unaffected when placed intrathecally in the spinal cord (Keeler and Helke, 1985), with this response mediated through NK-1 receptors (Helke et al., 1990; Quartara and Maggi, 1998). In addition, tachykinins directly influence medullary relay neurons in cardiovascular reflex pathways. Tachykinins microinjected into the caudal third of the solitary complex decrease mean arterial pressure and heart rate in a dose responsive manner (Kubo and Kihara, 1987; Lukovic et al., 1987), whereas tachykinin antagonists increase blood pressure and heart rate (Kubo and Kihara, 1987). Changes in respiration were also noted in these experiments with tachykinins producing a dose-dependent apnea when injected into the caudal solitary complex (Kubo and Kihara, 1987). A greater variety of responses is found when tachykinins and their antagonists are applied to the ventral surface of the medulla oblongata or injected into the medullary reticular formation. Thus tachykinins affect different neural pools to change ventilation in response to hypoxia, hypercapnea, and somatosensory stimuli (Chen et al., 1990). Another autonomic function affected by the administration of central tachykinins is the control of gastric motility and acid secretion through innervation of the dorsal motor vagal nucleus (Buchan et al., 1991). The high concentrations of tachykinins in hypothalamic regions suggest that tachykinins may be important for many other autonomic and behavioral functions. In particular, the direct retinal innervation of the suprachiasmatic nucleus suggests that tachykinins establish the light–dark cycle (Takatsuji et al., 1991). The heavy innervation of sexually dimorphic brain regions suggests that tachykinins may be important in sexual behavior, whereas tachykinins in other hypothalamic regions are likely to be involved in the release of pituitary hormones (Tsuruo et al., 1991). A role of SP in the initiation and maintenance of sleep has been proposed (Kohlmeier et al., 2002). It is also clear that tachykinins are involved in maintaining water homeostasis (Larsen, 1992; Lecci and Maggi, 2001). Further studies are required to establish the exact role of tachykinins in these and other brain regions.

COEXISTENCE OF SEROTONIN AND TACHYKININS The colocalization of classical neurotransmitters and peptides within the same neuron was first demonstrated in endocrine cells, but has since been noted as a more general phenomenon in the central nervous system (Hökfelt et al., 1987, 2000). The functional significance of the coexistence of multiple messenger molecules is not well understood, although advances are being made (Hökfelt et al., 2000; Merighi, 2002). Past studies have postulated that peptides colocalized with classical neurotransmitters are likely to be responsible for functions other than fast ionic exchanges, e.g., long-term trophic effects (Hökfelt et al., 1987). Several studies have concentrated on the colocalization of 5-HT and tachykinins (Léger et al., 2002; Sergeyev et al., 1999).

Location of Cell Bodies Both 5-HT and tachykinins are found within neurons in the midline and lateral B1 and B3 cell groups as well as within the midline B2 (ROb) cell group (Bowker et al., 1981a; Chan-Palay et al., 1978; Hökfelt et al., 1978; Johansson et al., 1981; Kachidian et al., 1991; Léger et al., 2002). It should be noted that within these cell groups twice as many 5-HT neurons are found compared to tachykinin-containing neurons (Johansson et al., 1981). It is thought that the majority of these neurons project to the spinal cord (Araneda et al., 1989; Bowker and Abbott, 1990; Johansson et al., 1981; Magoul et al., 1988), although it has been reported that neurons containing both neurotransmitters project to the solitary complex (Thor and Helke, 1987; Thor et al., 1988). Neurons containing both neurotransmitters predominate in the B1/2 cell groups. Although tachykinins are found throughout all medullary 5-HT cell groups, the caudal B1/2 cell groups can be distinguished from the rostral B3 cell group by the combination of other peptide neurotransmitters (Kachidian et al., 1991). Such neurotransmitter distinctions probably underlie functional differences within these pathways. In contrast to the B1, B2, and B3 cell groups, tachykinins are not found within any other 5-HT cell groups in the rat brain stem (Araneda et al., 1989; Magoul et al., 1988; Towle et al., 1986).

Main Pathways It has been known for some time that 5-HT and tachykinins are located within the same terminals and within the same dense-cored vesicles (Pelletier et al., 1981). In the spinal cord, both neurotransmitters are located in fibers within the ventral horn, whereas few fibers in the superficial layers of the dorsal horn

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contain both 5-HT and tachykinins (Wessendorf and Elde, 1987). This is despite the high density of both fiber types in this region (see earlier discussion). In the intermediolateral cell column and around the central canal over half of the fibers containing 5-HT and tachykinins contain both neurotransmitters (Appel et al., 1986, 1987; Wessendorf and Elde, 1987). A similar density of fibers containing both neurotransmitters is found in many dorsal medullary regions, including the solitary complex, dorsal motor vagal nucleus, area postrema, and the hypoglossal nucleus (Thor et al., 1988). However, the vast majority of fibers in these regions display immunoreactivity only for either 5-HT or tachykinin (Tallaksen-Greene et al., 1993; Thor et al., 1988). In a study of the distribution of these compounds throughout the brain stem, it was found that the highest density of colocalized fibers occurred in cranial nerve motor nuclei and in reticular regions of the ventral medulla (Tallaksen-Greene et al., 1993). Double-labeled fibers are uncommon in regions involved in processing special sensory information (Tallaksen-Greene et al., 1993). Because of the distribution of fibers containing both neurotransmitters, neurons containing 5-HT and tachykinins are thought to modulate somatic and sympathetic autonomic motoneurons (Tallaksen-Greene et al., 1993; Wessendorf and Elde, 1987). Pharmacological evidence in the spinal cord has shown that descending 5-HT neurons innervating the intermediolateral cell column release tachykinins as well as 5-HT to increase blood pressure (Chalmers et al., 1987; Hökfelt et al., 2000).

FUNCTIONAL INTERACTION BETWEEN SEROTONIN AND TACHYKININS Although the colocalization of 5-HT and tachykinin is quite restricted in the rat central nervous system, 5HT and tachykinin fibers often innervate the same structures, including brain stem regions containing monoamine neurons, the periaqueductal gray, the superior colliculus, and cuneiform nucleus (Figs. 16 and 17). Several studies have shown that these neurotransmitters interact throughout the neuroaxis (see below). In the dorsal horn of the spinal cord, both neurotransmitters are thought to modulate behavioral responses to painful stimuli (see earlier discussion). Intrathecal injections of 5-HT or 5-HT1 receptor agonists inhibit the response to painful stimuli in contrast to intrathecal injections of SP (Eide and Hole, 1991). When SP is injected prior to 5-HT, this inhibition is reduced without a reduction in skin temperature (Eide and Hole, 1991). These responses appear to be dose

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dependent, with similar nociception stimulation when both compounds are injected in very low doses, although these responses can be blocked by receptor antagonists (Eide and Hole, 1991). Intracellular recording studies in the solitary complex have shown that 5-HT and SP are both excitatory to solitary neurons and reduce the duration of prolonged action potentials when applied alone, but this action can be reversed under certain conditions (Jacquin et al., 1989). 5-HT inhibits solitary neurons after they have been exposed to prolonged conditioning by SP (Jacquin et al., 1989). The same reversal is seen for SP when 5-HT has been used for preconditioning. This suggests that the underlying mechanism for this action takes place at a common level, possibly in relation to second messenger processes. The extracellular concentration of 5-HT is increased by all three tachykinins (Iverfeldt et al., 1990; Solti and Bartfai, 1987). This response appears to be mediated through separate receptor mechanisms (Iverfeldt et al., 1990; Solti and Bartfai, 1987), suggesting that specific tachykinin pathways can independently modify the extracellular concentration of 5-HT. In addition to these direct postsynaptic actions, these compounds can modify biosynthetic pathways. In the rat neostriatum, depletion of 5-HT by amphetamine treatment decreases the SP/NKA precursor (Walker et al., 1991). Conversely, increasing extracellular 5-HT levels increases the SP/NKA precursor. These changes can be mimicked by 5-HT2 receptor agonists (Walker et al., 1991).

Abbreviations 3N 3V 4N 5HT 7N 7n 10N 12N A5 AC aca AcbC AcbSh acp AD AHiAL AHP AM Amb AP APT Aq Arc AV

VII. NEUROTRANSMITTERS

Oculomotor nucleus Third ventricle Trochlear nucleus 5-Hydroxytryptamine, serotonin Facial nucleus Facial nerve Dorsal motor nucleus, vagus Hypoglossal nucleus A5 noradrenaline cells Anterior commissural nucleus Anterior commissure, anterior Accumbens nucleus, core Accumbens nucleus, shell Anterior commissure, posterior Anterodorsal thalamic nucleus Amygdalohippocampal area, anterolateral Anterior hypothalamic area, posterior Anteromedial thalamic nucleus Ambiguus nucleus Area postrema Anterior pretectal nucleus Aqueduct Arcuate hypothalamic nucleus Anteroventral thalamic nucleus

1248 AVPe B1–B9 BAC Bic BM BMP BST BSTIA BSTLD BSTLV BSTMA BSTMP BSTV CA3 cc CeC CeL CeM cg Cg CIC CLi CM CnF cp CPu Cu CVL DA DCIC DG DLG DLL DMC DMD DMSp5 DpG DPGi DpMe DpWh DR DTg dtg ECIC ECu EGP EP Eth EW f F fmi fr FStr G g7 gcc Gi GiA GiV GP Gr gr HDB I

ANTONY HARDING ET AL.

Anteroventral periventricular nucleus B1 to B9 serotonin cell groups Bed nucleus, anterior commissure Brachium of the inferior colliculus Basomedial amygdaloid nucleus Basomedial amygdaloid nucleus, posterior Bed nucleus, stria terminalis Bed nucleus, stria terminalis, intraamygdaloid division Bed nucleus, stria terminalis, lateral division, dorsal Bed nucleus, stria terminalis, lateral division, ventral Bed nucleus, stria terminalis, medial division, anterior Bed nucleus, stria terminalis, medial division, posterior Bed nucleus, stria terminalis, ventral division Area CA3 of Ammon’s horn Corpus callosum Central amygdaloid nucleus, capsular Central amygdaloid nucleus, lateral Central amygdaloid nucleus, medial Cingulum Cingulate cortex Central nucleus of the inferior colliculus Caudal linear nucleus of the raphe Central median thalamic nucleus Cuneiform nucleus Cerebral peduncle, basal Caudate putamen Cuneate nucleus Caudal ventrolateral reticular nucleus Dorsal hypothalamic area Dorsal cortex of the inferior colliculus Dentate gyrus Dorsal lateral geniculate nucleus Dorsal nucleus of the lateral lemniscus Dorsomedial hypothalamic nucleus, compact Dorsomedial hypothalamic nucleus, diffuse Dorsomedial spinal trigeminal nucleus Deep gray layer of superior colliculus Dorsal paragigantocellular nucleus Deep mesencephalic nucleus Deep white layer of superior colliculus Dorsal raphe nucleus Dorsal tegmental nucleus Dorsal tegmental bundle External cortex inferior colliculus External cuneate nucleus External globus pallidus Entopeduncular nucleus Ethmoid thalamic nucleus Edinger–Westphal nucleus Fornix Nucleus of the fields of Forel Forceps minor, corpus callosum Fasciculus retroflexus Fundus striati Gelatinosus thalamic nucleus Genu facial nerve Genu corpus callosum Gigantocellular reticular nucleus Gigantocellular reticular nucleus, alpha Gigantocellular reticular nucleus, ventral Globus pallidus Gracile nucleus Gracile fasciculus Nucleus horizontal limb diagonal band Intercalated nuclei, amygdala

ic ICjM icp IF IGL IGP IMD InCo InG InWh IO IP IPC IPI IPL IPR IPAC IPRL IRt KF LA LC LDTg LH LHb LHbL LHbM ll LM LPB LPGi LPMC LPMR LPO LRt LSD LSI LSO LSS LSV LT LV LVe MCLH MCPO MePD MD MDC MdD MDM ME MeAD MePD MePV MG MHb MiTg ml MMn MnPo MnR Mo5 MPA MPB MPOC

VII. NEUROTRANSMITTERS

Internal capsule Islands of Calleja, major island Inferior cerebellar peduncle Interfascicular nucleus Intergeniculate leaf Internal globus pallidus Intermediodorsal thalamic nucleus Intercollicular nucleus Intermediate gray layer, superior colliculus Intermediate white layer, superior colliculus Inferior olive Interpeduncular nucleus Interpeduncular nucleus, caudal subnucleus Interpeduncular nucleus, intermediate subnucleus Interpeduncular nucleus, lateral subnucleus Interpeduncular nucleus, rostral Interstitial nucleus, posterior limb, anterior commissure Interpeduncular nucleus, rostrolateral subnucleus Intermediate reticular nucleus Kölliker–Fuse nucleus Lateroanterior hypothalamic area Locus coeruleus Laterodorsal tegmental nucleus Lateral hypothalamic area Lateral habenular nucleus Lateral habenular nucleus, lateral Lateral habenular nucleus, medial Lateral lemniscus Lateral mamillary nucleus Lateral parabrachial nucleus Lateral paragigantocellular nucleus Lateral posterior thalamic nucleus, mediocaudal Lateral posterior thalamic nucleus, mediorostral Lateral preoptic area Lateral reticular nucleus Lateral septal nucleus, dorsal Lateral septal nucleus, intermediate Lateral superior olivary nucleus Lateral striatal stripe Lateral septal nucleus, ventral Lateral terminal nucleus, accessory optic tract Lateral ventricle Lateral vestibular nucleus Magnocellular nucleus lateral hypothalamus Magnocellular preoptic nucleus Medial amygdaloid nucleus, posterodorsal Mediodorsal thalamic nucleus Mediodorsal thalamic nucleus, central Medullary reticular field, dorsal Mediodorsal thalamic nucleus, medial Median eminence Medial amygdaloid nucleus, anterodorsal Medial amygdaloid nucleus, posterodorsal Medial amygdaloid nucleus, posteroventral Medial geniculate nucleus Medial habenular nucleus Microcellular tegmental nucleus Medial lemniscus Medial mamillary nucleus, median Median preoptic nucleus Median raphe nucleus Motor trigeminal nerve Medial preoptic area Medial parabrachial nucleus Medial preoptic nucleus, central

34. THE SEROTONIN AND TACHYKININ SYSTEMS

MS mt MT Mtu MVe ω Op OPT OT ox Pa4 PaAP PAG PaPo PBP pc PCRt PCRtA PDTg Pe PeF PH PIL PLi PMD PMR PMV Pn PnC PnO Po PoT PP PPT PPTg PR Pr5 PrH PSol PT PV py pyx R RAmb Rbd Re Rh RI RLi RMg ROb RPa RPO RRF rs Rt RtTg RVL SCh SCO scp SFO

Medial septal nucleus Mamillothalamic tract Medial terminal nucleus of the accessory optic tract Medial tuberal nucleus Medial vestibular nucleus Omega Optic nerve layer superior colliculus Olivary pretectal nucleus Nucleus optic tract Optic chiasm Paratrochlear nucleus Paraventricular hypothalamic nucleus, anterior parvocellular Periaqueductal gray Paraventricular hypothalamic nucleus, posterior Parabrachial pigmented nucleus Posterior commissure Parvocellular reticular nucleus Parvocellular reticular nucleus, alpha Posterodorsal tegmental nucleus Periventricular hypothalamic nucleus Perifornical nucleus Posterior hypothalamic area Posterior intralaminar thalamic nucleus Posterior limitans thalamic nucleus Premamillary nucleus, dorsal Paramedian raphe nucleus Premamillary nucleus, ventral Pontine nucleus Pontine reticular nucleus, caudal Pontine reticular nucleus, oral Posterior thalamic nuclear group Posterior thalamic nuclear group, triangular Peripeduncular nucleus Posterior pretectal nucleus Pedunculopontine tegmental nucleus Prerubral field Principal sensory trigeminal nucleus Prepositus hypoglossal nucleus Parasolitary nucleus Paratenial nucleus Paraventricular nucleus Pyramidal tract Pyramidal decussation Red nucleus Retroambiguus nucleus Rhabdoid nucleus Reuniens thalamic nucleus Rhomboid thalamic nucleus Rostral interstitial nucleus, medial longitudinal fasciculus Rostral linear nucleus raphe Raphe magnus nucleus Raphe obscurus nucleus Raphe pallidus nucleus Rostral periolivary region Retrorubrual field Rubrospinal tract Reticular thalamic nucleus Reticulotegmental nucleus, pons Rostral ventrolateral reticular nucleus Suprachiasmatic nucleus Subcommissural organ Superior cerebellar peduncle Subfornical organ

SG SGe SHi SHy SI sm SNC SNL SNR SO Sol sol SolC SolM Sp5C Sp5I Sp5O SPF SP SPFPC Sph SPTg SpVe st STh Su3 Su3C Sub SubC subI SuG SuM TC Te TS TT Tu Tz VDB VLGPC VLL VMH VOLT VP VPPC VRe VTA VTg xcsp ZI ZL Zo

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Suprageniculate thalamic nucleus Supragenual nucleus Septohippocampal nucleus Septohypothalamic nucleus Substantia innominata Stria medullaris thalami Substantia nigra, compact Substantia nigra, lateral Substantia nigra, reticular Supraoptic nucleus Nucleus solitary tract Solitary tract Nucleus solitary tract, commissural Nucleus solitary tract, medial Spinal trigeminal nucleus, caudal Spinal trigeminal nucleus, interpolar Spinal trigeminal nucleus, oral Subparafascicular thalamic nucleus Substance P Subparafascicular thalamic nucleus, parvocellular Sphenoid nucleus Subpeduncular tegmental nucleus Spinal vestibular nucleus Stria terminalis Subthalamic nucleus Supraoculomotor central gray Supraoculomotor central gray, cap Submedius thalamic nucleus Subcoeruleus nucleus Subincertal nucleus Superficial gray layer superior colliculus Supramamillary nucleus Tuber cinereum Terete hypothalamic nucleus Triangular septal nucleus Tenia tecta Olfactory tubercle Nucleus trapezoid body Nucleus vertical limb diagonal band Ventral lateral geniculate nucleus, parvocellular Ventral nucleus lateral lemniscus Ventromedial hypothalamic nucleus Vascular organ of the laminar terminalis Ventral pallidum Ventroposterior thalamic nucleus, parvocellular Ventral reuniens nucleus Ventral tegmental area Ventral tegmental nucleus Decussation superior cerebellar peduncle Zona incerta Zona limitans Zonal layer superior colliculus

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C H A P T E R

35 Cholinergic Neurons and Networks Revisited LARRY L. BUTCHER1 and NANCY J. WOOLF2 1

Laboratory of Chemical Neuroanatomy and 2Laboratory of Nanoneuroscience Department of Psychology, University of California Los Angeles, California, USA

process of extraction. If acetylcholine, however, or any other substance of comparable activity, existed in quantities sufficient for chemical detection, its action would inevitably overpower that of adrenine in a gland extract. (pp. 188–189)

Approximately a century ago, Reid Hunt, in a paper authored with Traveau (Hunt and Traveau, 1906), made the prophetic commentary that I frequently obtained extracts of the suprarenal (and also of the brain) which caused a fall of blood pressure . . . and which were also more powerful than cholin. . . . I also got results . . . which led me to think that at least some of these results were to be attributed to a precursor of cholin or to some compound of cholin. . . . From these observations it seemed not impossible that . . . cholin compounds . . . might arise in the body; and, further, that such compounds may have some importance in certain pathological conditions. . . . Acetylcholin, the first of this series, is a substance of extraordinary physiological activity. In fact, I think it safe to state that, as regards its effect upon the circulation, it is the most powerful substance known. . . . We have not determined the cause of the fall of blood pressure from acetylcholine, but from the fact that it can be prevented entirely by atropine, I am inclined to think that it is due to an effect upon the terminations of the vagus in the heart. (p. 1789)

Armed with more data, as well as the wisdom of hindsight, Dale adopted a less equivocal stand 24 years later (Dale, 1938): If Reid Hunt . . . had examined in more detail the action of acetylcholine, the intense depressor activity of which he described . . . I think it must have been realized that acetylcholine would be a more suitable and likely parasympathetic transmitter than muscarine. (p. 416)

The significant contributions that Hunt made to early work on acetylcholine were acknowledged by Dale 8 years later but with qualifications (Dale, 1914): Reid Hunt found evidence of the existence of a substance in the suprarenal gland, which was not choline itself, but easily yielded that base in the

The Rat Nervous System, Third Edition

Since the time of Hunt’s seminal observations and conjectures, considerable experimental evidence has accumulated indicating that acetylcholine is involved importantly in intercellular communication processes in both peripheral and central nervous systems (for a recent treatment of topic, see Bartus, 2000). The nicotinic actions of acetylcholine at the neuromuscular junction and its muscarinic effects on the postganglionic targets of the parasympathetic autonomic nervous system, for example, are well understood and are now considered common knowledge in standard textbooks on pharmacology, physiology, pathology, and biochemistry, among others. Despite the long history of the study of acetylcholine as a physiologically significant chemical messenger, valid histochemical methods for its demonstration

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have only appeared since the 1980s. The first of these, the immunocytochemical demonstration of choline O-acetyltransferse (ChAT), the synthetic enzyme of acetylcholine, by use of polyclonal and monoclonal antibodies was employed by many laboratories with generally converging results (e.g., see Butcher, 1995). These techniques were followed by in situ hybridization methods for the ChAT mRNA (Oh et al., 1992). Most recently, immunocytochemical (Roghani et al., 1997) and in situ hybridization (Roghani et al., 1996) procedures have been used to map the distribution of the vesicular acetylcholine transporter (VAChT, see Figs. 1 and 2). All of these methods have yielded a remarkably similar picture of the organiza-

tion and distribution of cholinergic neurons in the central and peripheral nervous systems, a picture that has remained essentially unchanged since the second edition of “The Rat Nervous System” was published in 1995 (Butcher, 1995). In fact, the interested reader is referred to that chapter for a discussion of the properties and principles of organization of central cholinergic neurons and networks that remain as valid now as then. This chapter, therefore, confines itself to a consideration of cholinergic neuroanatomy in the context of possible functions, both normal and pathologic. It concludes with consideration of a novel conjecture concerning the role of forebrain cholinergic neurons in the genesis of Alzheimer’s disease.

FIGURE 1 Localization of the vesicular acetylcholine transporter in the rat anterior basal forebrain visualized by use of digoxigenin-labeled riboprobes and in situ hybridization. Reformatted, with permission, from Roghani et al. (1996). Scale: 200 μm.

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CHOLINERGIC NEUROANATOMY IN THE CONTEXT OF FUNCTION Cholinergic systems of the mammalian brain appear to play important roles in learning, memory, and consciousness. Acetylcholine is one of the key chemical messengers in brain and is found in significantly higher concentrations in the rat cortex than classic monoamine transmitters (approximately 10 times higher than levels of dopamine, serotonin, and norepinephrine; see Molinengo and Ghi, 1997). Still, our understanding is incomplete regarding how acetylcholine, along with other messengers, takes primary sensory input to the next level of processing, for example, nor do we have a complete picture of how acetylcholine and other transmitters might interact with fast-acting neurotransmission at ionotropic glutamate and GABA receptors. In the central nervous system, acetylcholine and the monoamine transmitters act predominately at the slower responding metabotropic receptors, which are coupled to G-proteins. Ionotrophic receptor actions of acetylcholine, however, occur in many parts of the central nervous system (e.g., see Levin et al., 1990).

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Overview of the Anatomy of Cholinergic Systems The cholinergic anatomy of rat brain is organized into several major cell groups and pathways (Figs. 3 and 4; for reviews, see Wainer et al., 1993; Butcher, 1995). Cholinergic neurons form a relatively continuous chain of somata from the caudal to rostral extent of the rat brain (Fig. 3). Regions of sparse cholinergic cells are filled in by monoaminergic neurons, including serotonergic neurons in the raphe nuclei, noradrenergic neurons in the locus ceruleus and hypothalamus, and dopamine neurons in the substantia nigra and ventral tegmental area. The significance of this continuum is not known. During development, it may reflect patterns of cell migration. In the adult, it may enable multiple neurotransmitters to participate effectively in a variety of functions ranging from learning to consciousness. The somatic motor neurons, including gamma motoneurons, and autonomic preganglionic and parasympathetic postganglionic neurons are all cholinergic, regardless of whether they innervate skeletal musculature, smooth muscle, cardiac muscle, or glands. Mesopontine cholinergic neurons in the laterodorsal

FIGURE 2 Localization of the vesicular acetylcholine transporter in the rat spinal cord visualized by use of polyclonal antibodies and immunocytochemistry. Arrows point to puncta putatively representing regions of afferent synaptic contact. Reformatted, with permission, from Roghani et al. (1997). Scale: 20 μm.

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FIGURE 3 Schematic representation of the rostral–caudal organization of cholinergic somata in the rat brain.

tegmental nucleus and pedunculopontine tegmental nucleus innervate extensive portions of the brain, including the brain stem, spinal cord, thalamus, hypothalamus, basal forebrain, and medial limbic cortex. These cholinergic cells are probably a major component of the so-called ascending reticular activating system, known for its ability to activate the cerebrum. The medial habenula of the epithalamus, specifically its ventral two-thirds, contains a collection of densely packed, relatively small cholinergic somata. The basal forebrain is composed of a complex of cholinergic cells, including the medial septal nucleus, vertral diagonal band nucleus, horizontal diagonal band nucleus, substantia innominata, magnocellular preoptic nucleus, and basal nucleus. These cells project to the entirety of the cerebral cortex, as well as to the hippocampus and amygdala (Fig. 4). The dorsal and ventral striata, composed of the caudate putamen complex, accumbens nucleus, olfactory tubercle, and islands of Calleja complex, contain cholinergic interneurons (Fig. 4). Cholinergic and monoaminergic neurons appear to play pivotal roles in learning. Studies that simultaneously assessed these modulators in aged rats found that learning deficits are associated with an immunohistochemical loss of markers for acetylcholine, norepi-

nephrine, and serotonin (Stemmelin et al., 2000). This is in partial agreement with earlier studies suggesting that learning and memory can be totally blocked by combined cholinergic and serotonergic suppression (Vanderwolf, 1987). Research in this area is still vigorous, and many questions remain unanswered. Interplay among cholinergic and monoaminergic neurons is evident in different states of wakefulness or rapid eye movement sleep. Mesopontine tegmental cholinergic neurons are active during rapid eye movement sleep, whereas noradrenergic and serotonergic neurons are inactive (Maloney et al., 1999). The effects of these messengers differ between waking and dreaming to the extent that cortical “binding” (a mechanism that determines coherent activity among neurons) is tuned to either sensory stimuli or internally perceived events (Kahn et al., 1997). The complexity of this interplay is not completely understood.

Efferent Cholinergic Neurons in Spinal Cord and Brain Stem Spinal and brain stem cholinergic efferent neurons provide fibers to different somatic, smooth, and cardiac muscles, as well as to glands. Before these efferent fibers leave the spinal cord, they frequently emit

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FIGURE 4 Schematic representation in the rat brain of telencephalic local-circuit cholinergic neurons and projections of the basal forebrain and mesopontine cholinergic systems. n, nuclei.

collateral axon fibers. Terminals from these cholinergic collateral fibers make contact with other cholinergic neurons (Li et al., 1995). The connectivity between cholinergic neurons and interneurons may enable them to act as an integrated ensemble. Efferent spinal and brain stem cholinergic neurons appear exquisitely plastic, especially during development. The glial-derived neurotrophic factor has been discovered to promote axonal growth of rat cholinergic motoneurons (Blesch and Tuszynski, 2001). The brainderived growth factor also stimulates the axonal growth of motoneurons in adult rats (Kishino et al., 1997). Reelin, an extracellular matrix protein, may be essential in the proper migration of sympathetic and parasympathetic cholinergic motoneurons (Phelps et al., 2002). These factors, among others, contribute to the topography of cholinergic motoneurons and to the arrangement of their connections.

Mesopontine Cholinergic Neurons Mesopontine cholinergic neurons have attracted considerable interest because they give rise to ascending projections to the thalamus and basal forebrain. The pedunculopontine tegmental nucleus is located lateral to the superior cerebellar peduncle in rat. The ascending projections from the mesopontine tegmentum to the thalamus and basal forebrain are implicated in

the overall regulation of wakefulness and rapid eye movement sleep (Jones, 1993). Lesions in mesopontine areas that contain cholinergic cells can produce coma (Stewart-Amidei, 1991); less complete lesions produce a syndrome called peduncular hallucinosis (Manford and Andermann, 1998). This latter atypical state of consciousness often includes vivid visual hallucinations of geometric figures. Related to a role in consciousness, mesopontine cells also appear to mediate attention. Lesions of the pedunculopontine tegmental nucleus produce deficits (increased errors and latency times) on a five-choice serial reaction time task constructed to test attention (Inglis et al., 2001). Mesopontine neurons putatively filter sensory information and select salient stimuli upon which the animal then focuses attention. The anatomical localization of mesopontine cholinergic cells, specifically their intermingling with ascending sensory fibers, may well underlie this role as a sensory filter. Autoregulation of mesopontine cholinergic neurons may have an important role in their physiology and function. Electron microscopic studies on mesopontine cholinergic cells find the VAChT protein on membranous organelles highly localized to dendrites; this suggests that the dendritic release of acetylcholine may be prominent among mesopontine cells (Garzon and Pickel, 2000). The dendritic release of acetylcholine could be one more way that mesopontine cholinergic

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cells act as an integrated ensemble. In general, the diffuse release of acetylcholine acting extrasynaptically could be of physiological relevance in the mammalian central nervous system for a variety of functions (Descarries et al., 1997).

Medial Habenula The medial habenula contains a dense aggregate of cholinergic somata. Due to the atypical morphology of these neurons (compared to other large multipolar neurons), it was not clear to us at first if these neurons were indeed cholinergic (Woolf and Butcher, 1985; cf. Houser et al., 1983). Subsequent studies unequivocally revealed that cells in the medial habenula were cholinergic by use of ChAT mRNA hybridocytochemistry (Oh et al., 1992), VAChT immunohistochemistry (Roghani et al., 1996; Arvidsson et al., 1997), and methods for the high-affinity choline transporter (Misawa et al., 2001). The medial habenula provides cholinergic axons to the fasciculus retroflexus, many of which terminate in the interpeduncular nucleus. Some cholinergic axons from the laterodorsal tegmental and the horizontal diagonal band nuclei also join the fasciculus retroflexus, entering through the lateral habenula. Thus, the fasciculus retroflexus is unique among cholinergic pathways insofar as it combines mesopontine, thalamic, and basal forebrain cholinergic fibers into one bundle. The function of the cholinergic component of this system may include regulation of EEG patterns and rapid eye movement sleep, as evidenced by reciprocal connections between the habenular complex and the laterodorsal tegmental nucleus (Semba and Fibiger, 1992). Lesions of the fasciculus retroflexus decrease time spent in hippocampal theta and rapid eye movement sleep (Valjakka et al., 1998).

Cholinergic Interneurons in the Dorsal Striatum and Ventral Striatum Cholinergic interneurons of the caudate-putamen complex, accumbens nucleus, and olfactory tubercle are spaced fairly regularly apart. Their somata are medium to large in size and multipolar. Although the function of these cells is, in part, extrapyramidal in nature, at least one study reported a role for both dorsal and ventral striatum in active avoidance in the T maze (Shapalova et al., 1996). The cholinergic neurons of the islands of Calleja are smaller than those in the striatum. It is conceivable that they are functionally related to the cholinergic neurons of the ventral striatum. No definite behavioral role has been established for cholingeric neurons of the islands of Calleja complex.

Basal Forebrain Cholinergic Neurons A rostral-to-caudal gradient exists among cholinergic projections emanating from cholinergic basal forebrain cells. Medial septal and ventral diagonal band nuclei constitute the rostral extension. These cholinergic neurons project mainly to the hippocampus (CA1–CA4 and dentate gyrus) and subiculum but also provide part of the cholinergic innervation of the entorhinal, perirhinal, and retrosplenial cortex. A major component of these projections is the septohippocampal pathway, which has been studied extensively in rats (see Chapter 20). A role in spatial memory has been established for this pathway, although controversies remain. Spatial learning performance during the acquisition of a radial arm maze task correlates with nearly doubled levels of acetylcholine release (Fadda et al., 2000), suggesting an important relationship between septohippocampal cholinergic activity and learning. Controversies emerge when one tries to relate cholinergic neuron damage to the presence or absence of behavioral deficits. It is not clear if cholinergic deafferentation alone is sufficient to produce behavioral deficits in all cases. It has been shown, for example, that damage to both cholinergic and GABAergic septohippocampal pathways is necessary for spatial memory deficits (Pang et al., 2001). The selective lesion of septal cholinergic neurons with 192-IgG saporin has been reported to impair delayed matching to position in a T-maze test (Johnson et al., 2002). The sensitivity of this task to selective deficits is cited as resolving the issue of whether selective versus nonselective damage to the cholinergic system is needed to produce behavioral deficits (see also discussion of the basal nucleus later). In addition to spatial memory, contextual memory appears to depend on the integrity of the cholinergic septohippocampal pathway. Intraseptally infused antisense DNA phosphothioate oligonucleotides directed at TrkA, the high-affinity uptake receptor of nerve growth factor, impair contextual memory (Woolf et al., 2001). Cholinergic basal forebrain neurons are not poisoned by the antisense oligonucleotides; rather, the synthesis of ChAT and VAChT is decreased dramatically. Contextual fear conditioning is also accompanied by an increase in acetylcholine release during acquisition and a fourfold increase during retention (Nail-Boucherie et al., 2000). Rewiring of cholinergic circuits appears likely during the acquisition phase to enable the increase in acetylcholine release upon reexposure to the training chamber. Of possible relevance to its role in memory, the septohippocampal pathway is responsible for generating a 4- to 10-Hz oscillation pattern called

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hippocampal theta. Hippocampal theta correlates with the presence or absence of particular behaviors and is implicated in spatial memory, although others have argued that the rhythm is related to gross movement (Paxinos and Bindra, 1970). Selective destruction of cholinergic septohippocampal cells with 192-IgG saporin diminishes hippocampal theta greatly (Apartis et al., 1998). Acetylcholine release is positively correlated with frequency at the peak power of the theta band, and hippocampal acetylcholine outflow covaries with relative power of the theta band (Keita et al., 2000). Although noncholinergic cells in the septal area or hippocampus may additionally contribute to these oscillations, acetylcholine appears to be involved pivotally. The middle territories of the cholinergic basal forebrain include the horizontal diagonal band nucleus and magnocellular preoptic nucleus. These cholinergic neurons innervate olfactory bulb, amygdale, and the cingulate, retrosplenial, entorhinal, perirhinal, insular cortices, as well as parts of the frontal cortex. Cholinergic terminals are especially dense in the basolateral and lateral amygdala. These medial cholinergic pathways exhibit considerable plasticity following axotomy, even in the adult organism, that can be modified by trophic factors (e.g., see Farris et al., 1993, 1995). Although behavioral studies on the medial cholinergic pathway are fewer in number than studies on the septohippocampal and basalocortical systems, it has been reported that removal of the olfactory bulbs produces spatial memory deficits and disruption of cholinergic indexes in the horizontal diagonal band and magnocellular preoptic nuclei (Bobkova et al., 2001). The caudal part of the cholinergic basal forebrain pathway is composed of the basal nucleus and substantia inominata. These structures contain neurons that project cholinergic axons to all of the neocortex: frontal, parietal, temporal, and visual. Two exceptions to this rule in rat brain are that the medial part of the visual cortex receives input from the ventral diagonal band nucleus (see Carey and Rieck, 1987), similar to that of the retrosplenial cortex, which lies adjacent, and the limbic regions of frontal cortex that receive afferents from the horizontal diagonal band and magnocellular preoptic nuclei. The role of the basal nucleus in behavior has been widely studied. Nearly 10,000 citations are listed on Medline searching these key words, and two-thirds of the studies are done on rats. The cholinergic basal forebrain appears to be involved in mediating attention, memory, learning, perception, and consciousness. Infusions of 192-IgG saporin into the basal nucleus, for example, produce deficits in a five-choice serial

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reaction time task designed to measure attention (McGaughy et al., 2002). These deficits are correlated with cell loss in the basal nucleus and decreased acetylcholine release. The use of 192-IgG saporin to demonstrate the role of the basal nucleus in memory has been less straightforward. To produce clear memory deficits on working memory in the Morris water maze, a 90% decrease of ChAT is needed (Waite et al., 1995). Some Purkinje cells are also damaged by that dose of 192IgG saporin. When lesser doses of 192-IgG are applied, memory deficits are not apparent but attention is impaired (Baxter et al., 1995). An important consideration is whether cholinergic systems must be compromised appreciably (>90% decrease) to produce memory deficits or if nonselective damage is necessary to produce those deficits. It is likely that cholinergic neurons must be compromised greatly or that another transmitter plus acetylcholine must be reduced. Residual cholinergic terminals are able to compensate by upregulating acetylcholine synthesis (Waite and Chen, 2001). Serotonin systems can also compensate for cholinergic loss. Combined application of the selective cholinergic toxin, 192-IgG saporin, and the serotonin toxin, 5,7-dihydroxytryptamine, impair performances in the T-maze alternation test, the water maze working memory test, and the radial arm maze, even when ChAT is only reduced to 40% of normal (Lehmann et al., 2000). This could be due to the fact that acetylcholine and serotonin stimulate some of the same signal transduction cascades inside their targeted neurons. Perception appears, in part, to involve cholinergic basal nucleus projections to the neocortex. This is indicated by physiological studies done in the sensory cortex. Cholinergic deafferentation during postnatal development alters ocular dominance columns, alone and in combination with noradrenergic deafferentation (Siciliano et al., 1999). A muscarinic receptordependent alteration is produced for receptive fields in the rat auditory cortex in response to basal nucleus stimulation (Miasnikov et al., 2001). When cholinergic nicotinic and muscarinic agonists are applied to the somatosensory cortex, receptive fields are reorganized within about 1 h (Penschuck et al., 2002). Across sensory modalities, acetylcholine appears to have potent effects on receptive field characteristics, parameters that doubtless affect perception. The cholinergic basal forebrain has also been suggested to play a role in the mediation of consciousness (Woolf, 1996; Smythies, 1997). The rationale behind this suggestion derives from the widespread cortical projections of the basal nucleus and the effects of anticholinergic drugs acting at muscarinic receptors

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to produce delirium. That the nicotinic receptor is antagonized by inhalational anesthetics, that there are visual hallucinations in patients with dementia with Lewy bodies, and that Parkinson’s disease patients with an additional loss of pedunclopontine tegmental neurons have rapid eye movement sleep abnormalities further suggest that cholinergic systems are involved in consciousness (Perry et al., 1999).

CENTRAL CHOLINERGIC NEURONS: MODES OF OPERATION Cholinergic neurons in the central nervous system, along with monoaminergic neurons, appear to mediate both overt and covert types of behavior. The efferent cholinergic fibers of the spinal cord and brain stem mediate covert or observable behavior through their direct stimulation of muscle. Commands upon skeletal muscle produce observable movements. Effects on smooth muscle, cardiac muscle, and glands produce different observable behaviors (e.g., sweating, lacrimation). Alternatively, autonomic states are relayed back to the central nervous system via autonomic afferents to produce observable emotional behaviors mediated through commands over skeletal muscle (i.e., facial expression, instrumental responses). Muscle contraction is mediated through motor proteins, such as myosin and kinesin, which attach to actin filaments and microtubules, respectively. These motor proteins use a similar type of conformation shift to convert ATP to ADP and affect muscle contraction (Vale and Milligan, 2000). Acetylcholine actions at nicotinic receptors produce Ca2+ currents that are coupled to skeletal muscle contraction (Ashcroft, 1991). Acetylcholine affects smooth muscle via the second messenger, phosphoinositide-specific phospholipase C (PI-PLC), which in turn releases internal stores of Ca2+ and causes contraction (Eglin et al., 1994). Mesopontine and basal forebrain cholinergic neurons affect covert behavior at virtually every level. These cholinergic groups, along with monoamine cells, appear to play a role in cognition, learning, memory, attention, and consciousness. How this is accomplished undoubtedly involves cholinergic receptors and the signal transduction cascades initiated by these receptors. Central acetylcholine is known to activate postsynaptic muscarinic receptors M1, M3, and M5 (for review, see Taylor and Brown, 1999). M1 is found in higher concentrations in the cerebral cortex and its localization to cortical pyramidal cells is certain. Postsynaptic M1 receptors (and M3/M5) activate

PI-PLC. Through mediation of the α subunit of a GTP-binding protein, PI-PLC activation leads to the subsequent activation of inositol 1,4,5-triphosphate (IP3) and diacylglycerol. In turn, diacylglycerol activates Ca2+/phospholipid-dependent protein kinase (PKC). IP3 increases the release of calcium from internal stores, which activates Ca2+/calmodulindependent kinase II (C/CMK). In dendrites, these kinases affect the cytoskeleton via the phosphorylation of microtubule-associated proteins. The mechanism by which acetylcholine mediates learning and memory may involve effects on dendrite reorganization (for review, see Woolf, 1998). In addition to changes in synaptic efficacy, long-lasting storage may be possible with the reorganization of dendrites. Protein kinases affected by acetylcholine (e.g., PKC and C/CMK) phosphorylate the microtubuleassociated protein, MAP-2 (for review, see Johnson and Jope, 1992). Studies in rat hippocampal cells show that PKC specifically increases branching of dendrites, whereas other kinases affect branching in both dendrites and axons (Audesirk et al., 1997). The mechanism by which acetylcholine mediates consciousness can be considered in light of the signal transduction cascades it initiates. Cholinergic muscarinic actions on PKC and C/CMK, taken with the known roles of PKC and C/CMK in the phosphorylation of MAP-2, suggest that one postsynaptic effect of acetylcholine is to phosphorylate MAP-2. In addition to M1 effects, 5-HT-2, α1-adrenergic, and histaminergic receptors activate PI-PLC. Hence activation of these receptors will also increase the phosphorylation of MAP-2 via PI-PLC. Dopamine D1 receptors, 5-HT-3 receptors, and β-adrenergic are postsynaptic receptors that stimulate adenylyl cyclase, which in turn activates cAMP-dependent kinase (PKA), which is also known to phosphorylate MAP-2. Dopamine D2 receptors, among others, inhibit adenylyl cyclase and thereby inhibit PKA. MAP-2 has been shown to be an anchoring protein for PKA (Harada et al., 2002). Thus, our notion of MAP-2 as a strictly structural protein has to be revised. Acetylcholine and monoamine transmitters exert effects on different receptors, which in turn activate second messenger pathways; nonetheless, all converge on the cytoskeleton, especially the cytoskeletal-binding protein/signal transduction molecule, MAP-2. In this regard, the hypothetical model advanced by Hameroff and Penorse, called orchestrated objective reduction (Orch OR), may have relevance for the cholinergic mediation of consciousness (see Woolf and Hameroff, 2001). Orch OR refers to the collapse of superpositioned states held momentarily by the electrons in tubulin, the molecule that, paired in dimers, forms micro-

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tubules. MAP-2 bound to tubulins of the microtubules orchestrates this objective reduction, or collapse, because it physically constrains the molecule (Hameroff and Penrose, 1996). Mental processes come fully to mind or to the level of consciousness when a sufficient number of tubulins are coherently superpositioned and then collapse. This coherence requires a confluence between neurons and glial cells that could be provided by gap junctions. Isolation from membrane events is also required. Phosphorylation of MAP-2 would provide such isolation by decoupling the membrane from the microtubule and by increasing the repulsive force of the MAP-2 surrounding the microtubule. Although the Orch OR model is speculative, it is noteworthy that, similar to the case with muscle contraction, acetylcholine could affect a behavioral outcome, in this case covert behavior, via its action on microtubules.

GENESIS OF ALZHEIMER’S DISEASE: A HYPOTHESIS Two major, fundamentally different, types of cell death have been recognized (e.g., Wyllie et al., 1980): necrosis and apoptosis. The former term connotes death induced by pathologic insults deriving from sources ostensibly independent of normal biologic processes, as exemplified by injury, complement attack, lytic viral infection, hyperthermia, prolonged hypoxia, and exposure to diverse toxins. The latter term denotes a process by which cells die in a systematic manner according to an intrinisic biologic program(s) initiated by specific stimuli. Apoptosis has been observed to occur in neutrophil polymorph senescence, developmentally regulated cell death during embryogenesis and metamorphosis, endocrinedependent tissue atrophy (e.g., lymphocytes deprived of interleukin-2 or exposed to excess amounts of glucocorticoids), cell death following trophic factor withdrawal, and normal tissue turnover (e.g., Wyllie et al., 1980). It has been referred to variously as “natural cell death,” “cell self-destruction or suicide,” and “programmed cell death” and is believed to involve highly orchestrated biochemical and morphologic sequelae distinct from those operating during necrosis (e.g., see Wyllie et al., 1980). Evidence has been obtained in tissue culture that the p75 neurotrophin receptor, known as p75NTR and having as one of its naturally occurring ligands nerve growth factor (NGF), is involved in programmed cell death in specific populations of neurons (Rabizadeh et al., 1993). In these experiments, p75NTR was expressed in temperature-sensitive immortalized

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CSM 4.1 neural cells by means of a retroviral vector, pBabe-puro-p75NTR; these cells did not express TrkA, the high-affinity receptor for NGF. Control cells transfected with pBabe-puro alone expressed neither p75NTR nor TrkA. In cells cultured in medium containing serum, expression of p75NTR had no effect on cell death, but when serum was withdrawn to induce apoptosis, expression of p75NTR led to an increase in neural cell death. If NGF was added, however, not only was the negative effect on cell survival suppressed, but the cells had a death rate less than that of control cells transfected with the identical vector lacking the p75NTR sequence. Binding of the p75NTR by a monoclonal antibody against p75NTR also suppressed the enhancement of neuronal cell death by the p75NTR. Addition of a control monoclonal antibody did not affect cell survival. Neither NGF nor monoclonal antibody against p75NTR affected the survival of control cells. The picture emerging from this research is that if the p75NTR is expressed in neurons, then it must be bound by appropriate concentrations of NGF or some other ligand, not necessarily an agonist (e.g., the 192 IgG antibody against the p75NTR), or else an apoptotic death program will be initiated. Concentrations of NGF below the Kd for the p75NTR (10−9M) do not rescue neurons from apoptosis, but those at and above that value do (Rabizadeh et al., 1993). Furthermore, p75NTR -mediated apoptosis in cells in culture can be potentiated by β-amyloid (Rabizadeh et al., 1994), excesses of which may contribute significantly to the pathogenesis of Alzheimer’s disease (e.g., see Friedlich and Butcher, 1994). Although the role of the p75NTR in mediating apoptosis in tissue culture seems certain, little has been established unequivocally about its function in vivo. Nonetheless, on the basis of the aforementioned tissue culture results and the distribution of the p75NTR in the intact mammalian brain, as well as other lines of experimental evidence, the following working hypothesis can be advanced: p75NTR is the arbiter of programmed cell death in basal forebrain cholinergic neurons and plays a role in the pathologic sequela of Alzheimer’s disease. First, the p75NTR is colocalized virtually exclusively with cholinergic neurons in the mature basal forebrain but not with the morphologically similar cholinergic cells of the mesopontine complex (Woolf et al., 1989a). Among cholinergic neurons, only those in the forebrain degenerate in Alzheimer’s disease (Woolf et al., 1989b) or show dystrophic changes in normal aging. Cholinergic neurons in the basal forebrain are also among the systems affected earliest and most severely in Alzheimer’s disease, and degeneration of that

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system correlates well with the severity of dementia, including prominently memory impairment (e.g., see Butcher and Woolf, 1989; Bartus, 2000). Second, chronic hyperthyroidism during early postnatal development produces preferential degeneration of cholinergic neurons in the basal forebrain beginning approximately 60 days following T3 administration (Gould and Butcher, 1989). Neuronal loss is preceded by a steady increase in intraneuronal p75NTR immunoreactivity (unpublished observations, this laboratory). Chronic hypothyroidism (i.e., propylthiouracil treatment), however, markedly decreases p75NTR immunopositivity in basal forebrain cholinergic neurons (unpublished observations, this laboratory). Surprisingly, none of these latter neurons die, even though they display morphologic features consistent with stunted growth, including shrunken somata and reduced dendritic number, lengths, and branch points (Gould and Butcher, 1989 and unpublished observations, this laboratory). Third, the p75NTR is increased in degenerating cholinergic neurons of the basal nuclear complex in Alzheimer’s disease while levels of NGF are slightly reduced or unchanged (Kordower et al., 1989; Goedert et al., 1989). Levels of the mRNA for NGF are also unchanged in the basal nucleus in Alzheimer’s disease patients, but there is a threefold increase in the mRNA for the p75NTR (Goedert et al., 1989; Persson and Ernfors, 1990). Thus, even though levels of NGF and the mRNA for NGF are normal in Alzheimer’s disease, the ratio of NGF to the p75NTR is decidedly abnormal in a direction favoring increased amounts of unbound p75NTR, which, in turn, could initiate programmed death in affected cells. [Additional data demonstrating a functional relationship between basal forebrain cholinergic neurons and p75NTR can be found in Yeo et al. (1997).] In operational terms, foregoing data suggest that if the ratio of unbound to bound p75NTR is greater than one, then the cell containing that receptor becomes increasingly at risk for apoptosis. If the ratio is less than one, then the risk of programmed cell death decreases. In Alzheimer’s disease, it is proposed that the ratio of unbound to bound p75NTR becomes greater than one in an increasing number of neurons in the cholinergic basal forebrain, eventually reaching apoptosis-inducing values, leading to cell death. It is not known how much greater than one this ratio must become in individual neurons to trigger apoptosis and whether that value is the same for all cells. Unanswered questions for future research include these issues, as well as determining the mechanisms or conditions altering the balance between bound and unbound p75NTR and the precise pathologic sequence that ensues in Alzheimer’s disease.

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Wainer, B. H., Steininger, T. L., Roback, J. D., Burke-Watson, M. A., Mufson, E. J., and Kordower, J. (1993). Ascending cholinergic pathways: Functional organization and implications for disease models. Prog. Brain Res. 98, 9–30. Waite, J. J., and Chen, A. D. (2001). Differential changes in rat cholinergic parameters subsequent to immunotoxic lesion of the basal forebrain nuclei. Brain Res. 918, 113–120. Waite, J. J., Chen, A. D., Wardlow, M. L., Wiley, R. G., Lappi, D. A., and Thal, L. J. (1995). 192 immunoglobulin G-saporin produces graded behavioral and biochemical changes accompanying the loss of cholinergic neurons of the basal forebrain and cerebellar Purkinje cells. Neuroscience 65, 463–476. Woolf, N. J. (1996). Global and serial neurons form a hierarchically arranged interface proposed to underlie memory and cognition. Neuroscience 74, 625–651. Woolf, N. J. (1998). A structural basis for memory storage in mammals. Prog. Neurobiol. 55, 59–77. Woolf, N. J., Butcher, L. L. (1985). Cholinergic systems in the rat brain. II. Projections to the interpeduncular nucleus. Brain Res. Bull. 14, 63–83. Woolf, N. J., Gould, E., and Butcher, L. L. (1989a). Nerve growth factor receptor is associated with cholinergic neurons of the basal forebrain but not the pontomesencephalon. Neuroscience 30, 143–152. Woolf, N. J., and Hameroff, S. R. (2001). A quantum approach to visual consciousness. Trends Cogn. Sci. 5, 472–478. Woolf, N. J., Jacobs, R. W., and Butcher, L. L. (1989b). The pontomesencephalic cholinergic system does not degenerate in Alzheimer’s disease. Neurosci. Lett. 96, 277–282. Woolf, N. J., Milov, A. M., Schweitzer, E. S., and Roghani, A. (2001). Elevation of nerve growth factor and antisense knockdown of TrkA receptor during contextual memory consolidation. J. Neurosci. 21, 1047–1055. Wyllie, A. H., Kerr, J. F. R., and Currie, A. R. (1980). Cell death: The significance of apoptosis. Int. Rev. Cytol. 68, 251–306. Yeo, T. T., Chua-Couzens, J., Butcher, L. L., Bredesen, D. E., Cooper, J. D., Valletta, J. S., Mobley, W. C., and Longo, F. M. (1997). Absence of p75NTR causes increased basal forebrain cholinergic neuron size, choline acetyltransferase activity, and target innervation. J. Neurosci. 17, 7594–7605.

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C H A P T E R

36 Glutamate JONAS BROMAN Department of Physiological Sciences, Lund University Lund, Sweden

ERIC RINVIK, MARCO SASSOE-POGNETTO, HOSSEIN KHALKHALI SHANDIZ and OLE PETTER OTTERSEN Centre for Molecular Biology and Neuroscience, Institute of Basic Medical Sciences University of Oslo, Blindern, Oslo, Norway Dipartimento di Anatomia, Farmacologia e Medicina Legale University of Turin, Italy

Glutamate (Glu) is undoubtedly the most prevalent transmitter in the brain. This amino acid is probably being used as a signaling substance in a majority of synapses, alone or along with peptides or other neuroactive compounds that colocalize with Glu. The excitatory effect of Glu was recognized in the early 1950s (Hayashi, 1954; Curtis and Watkins, 1960), but it took a long time until Glu was generally accepted as a neurotransmitter (Krnjevic, 1986; Watkins, 1986). Notably, the high concentration and relatively even distribution of Glu among brain regions were difficult to reconcile with a transmitter role. By the mid-1980s (Fonnum, 1984), Glu largely fulfilled the four main criteria for classification as a neurotransmitter: presynaptic localization, release by physiological stimuli, identical action with naturally occurring transmitter, and mechanism for rapid termination of transmitter action. Later investigations have strengthened a neurotransmitter role for Glu by demonstrating an ATPdependent selective transport of Glu into purified synaptic vesicles (Naito and Ueda, 1985; Maycox et al., 1990; Fykse et al., 1989; Winther and Ueda, 1993), the presence of high concentrations of Glu in synaptic vesicles isolated from the brain (Riveros et al., 1986; Burger et al., 1989; Orrego and Villanueva, 1993), and Ca2+-dependent exocytotic release of Glu from isolated nerve terminals (Nicholls, 1995). However, the

The Rat Nervous System, Third Edition

molecular basis for vesicular accumulation of Glu was long unknown. This has changed with the discovery of a family of vesicular Glu transporters (VGLUT1-3; Bellocchio et al., 2000; Takamori et al., 2000, 2001, 2002; Fremeau et al., 2001; Gras et al., 2002). As to the postsynaptic effect of Glu, rapid application of Glu to neuronal membrane patches at a concentration similar to that estimated to be present in the synaptic cleft following exocytotic release mimics the response that is obtained following the activation of excitatory synapses (Colquhoun et al., 1992; Clements et al., 1992; Bergles et al., 1999). Extensive molecular studies during the recent decade have provided detailed knowledge on the subunit proteins and gene families of Glu receptors (for reviews, see Blackstone and Huganir, 1995; Scannevin and Huganir, 2000). This chapter deals with anatomical aspects of transmitter Glu and provides an overview of the neuronal populations that use Glu as a neurotransmitter. The wide distribution of Glu precludes a comprehensive analysis of the topic within the available space and, in many cases, we have had to cite reviews rather than the original publications. Before describing the putative Glu-ergic projections in the brain, it is necessary to discuss the techniques that are available for the identification of neurons that use Glu as a transmitter. Biochemical procedures,

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detecting reduced content or uptake of Glu or Glu analogues following lesions, have proved useful in investigations of major projections (e.g., corticofugal fiber tracts; Fonnum, 1984; Storm-Mathisen and Ottersen, 1988), but poor sensitivity hampers analysis of smaller pathways. Identification of many minor Glu-ergic projections was made possible by the use of the metabolically inert Glu analogue D-[3H]aspartate as a transmitter-specific retrograde tracer (Baughman and Gilbert, 1980; Streit, 1980). It is a problem that 3 D-[ H]aspartate does not differentiate between putative Glu-ergic and aspartergic projections. Further, a number of fiber tracts likely to use Glu as a neurotransmitter are poorly labeled or unlabeled by D-[3H]aspartate, possibly due to the low presynaptic Glu uptake capacity of the terminals of such pathways (Ottersen, 1991). Analyses of the detailed anatomical distribution of Glu became possible with the development of antibodies to aldehyde-fixed amino acids (Storm-Mathisen et al., 1983). Early studies based on amino acid immunocytochemistry (Ottersen and Storm-Mathisen, 1984a, 1984b; Somogyi et al., 1986; Wanaka et al., 1987; Yoshida et al., 1987; Hepler et al., 1988; Chagnaud et al., 1989; Liu et al., 1989; Pow and Crook, 1993) demonstrated that Glu is widely distributed in the brain and localized not only in presumed Glu-ergic neurons, but also in neurons with other transmitter signatures. This is in line with biochemical data pointing to the involvement of Glu in several metabolic functions (protein synthesis, intermediary metabolism, and as a precursor for GABA). Introduction of the postembedding immunogold technique to amino acid immunocytochemistry (Somogyi and Hodgson, 1985) made it possible to analyze the distribution of Glu at a quantitative level and at higher anatomical resolution. This helped distinguish transmitter Glu from other pools of Glu. Using the immunogold approach, Somogyi et al. (1986) demonstrated enrichment of Glu immunoreactivity in parallel and mossy fiber terminals in the cerebellum, and later studies showed that the strength of the immunogold signal in these terminals was strongly correlated to the density of synaptic vesicles (Ji et al., 1991). Based on the assumption that a vesicular enrichment of Glu is a hallmark of Glu-ergic synapses, the quantitative immunogold approach has been used extensively to identify such synapses in the mammalian brain. The usefulness of this approach has been further improved by the development of combinations of anterograde tracing and immunogold labeling (De Biasi and Rustioni, 1988; Broman et al., 1990). As many of the data reviewed here are based on immunogold labeling, a critical evaluation of this technique appears relevant.

The ubiquitous presence of Glu in the central nervous system (CNS) sets hurdles for the analysis of Glu immunogold-labeled preparations and calls for quantitative analyses. Thus, the mere presence of Glu does not necessarily indicate a transmitter role. It serves to illustrate this that low levels of Glu occur in terminals rich in GABA or glycine (e.g., Somogyi et al., 1986; Bramham et al., 1990; Broman et al., 1990, 1993; Todd et al., 1994; Örnung et al., 1998). As biochemical studies have demonstrated high levels of Glu in synaptic vesicles (see earlier discussion), Glu-ergic terminals should be rich in Glu. Data from immunogold studies support this notion. However, demonstration of an enrichment of Glu fulfills only the first of the four main criteria of a neurotransmitter. A critical question is whether Glu may be present in high concentrations also in terminals not using Glu for synaptic transmission. The levels of Glu in terminals with other transmitter signatures than GABA or glycine (e.g., monoaminergic fibers) are largely unknown (but see Torrealba and Müller, 1999). It is noteworthy that strong immunogold signals for Glu were detected in motor nerve terminals innervating fast-twitch (but not slow-twitch) muscle fibers in rats (Waerhaug and Ottersen, 1993), although it remains to be shown whether these terminals release Glu in addition to acetylcholine. More recently Clarke et al. (1997) reported that cholinergic terminals in the basal ganglia contained levels of Glu that were intermediate to those in terminals with asymmetric and symmetric synapses, respectively. A corelease of acetylcholine and Glu has been demonstrated from presumed cholinergic synaptosomes and from cholinergic terminals of the Torpedo electric organ (Docherty et al., 1987; Vyas and Bradford, 1987). Although a colocalization of Glu with another neuroactive compound may point to transmitter roles for both substances, one cannot rule out that significant levels of “metabolic” Glu are present in certain populations of terminals. We must conclude that an enrichment of Glu within nerve terminals speaks strongly in favor of a transmitter role for Glu, but that immunogold data, like data obtained with other techniques, must be interpreted with due caution and with reference to alternative techniques addressing different features of Glu-ergic synapses. One such feature is vesicular Glu uptake. The discovery of a family of vesicular Glu transporters (VGLUT1-3; Bellocchio et al., 2000; Takamori et al., 2000, 2001, 2002; Fremeau et al., 2001; Gras et al., 2002) has opened up new possibilities for the identification of putative Glu-ergic neurons. Antibodies to these transporters have provided selective labeling of vesicle clusters in well-characterized Glu-ergic path-

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36. GLUTAMATE

ways (e.g., Fremeau et al., 2001). However, with this approach, any negative results must be interpreted with caution as there might be VGLUT isoforms that remain to be discovered. For references to early studies of Glu pathways, the reader may consult previous reviews (e.g., Ottersen and Storm-Mathisen, 1984b; Fonnum, 1984; Ottersen, 1991; Storm-Mathisen et al., 1995; Broman et al., 2000). In keeping with the scope of the present volume, we have largely ignored analyses in species other than rat.

ANATOMICAL SYSTEMS Neocortex Being easily amenable to biochemical analysis, the massive corticofugal projections, especially corticostriatal projections, were among the first for which Glu was assigned a transmitter role (Divac et al., 1977; McGeer et al., 1977). Since then, biochemical, pharmacological, and immunocytochemical studies have implicated Glu as a neurotransmitter in a large number of neocortical output systems (for reviews, see Fonnum, 1984; Storm-Mathisen and Ottersen, 1988; Tsumoto, 1990; Ottersen, 1991; McCormick and von Krosigk, 1992; Broman et al., 2000). In addition to the corticostriatal path (Fonnum et al., 1981; Girault et al., 1986; Gundersen et al., 1996), putative Glu-ergic pathways include cortical projections to the thalamus (Lund-Karlsen and Fonnum, 1978; Baughman and Gilbert, 1980, 1981; Fonnum et al., 1981; Young et al., 1981; Montero and Wenthold, 1989; Montero, 1990; Broman and Ottersen, 1992; McCormick and von Krosigk, 1992; De Biasi et al., 1994a; Ericson et al., 1995; Blomqvist et al., 1996; Eaton and Salt, 1996), to several loci in the brain stem (Young et al., 1981; Rustioni and Cuenod, 1982; Matute and Streit, 1985; Azkue et al., 1995; Ortega et al., 1995; Mize and Butler, 1996; Torrealba and Müller, 1996, 1999), and to the spinal cord (Young et al., 1981; Rustioni and Cuenod, 1982; Potashner et al., 1988; Valtschanoff et al., 1993). Retrograde tracing with D-[3H]aspartate also supports Glu as a neurotransmitter in pyramidal neurons projecting to the ipsilateral or contralateral cortex (local, associational, and commissural connections; Streit, 1980; Barbaresi et al., 1987; Elberger, 1989; Kisvarday et al., 1989; Johnson and Burkhalter, 1992). Although the majority of cortical interneurons are inhibitory and immunoreactive for GABA (Somogyi et al., 1998), a special type of local circuit neuron in layer IV, the spiny stellate neuron, is excitatory and assumed to use Glu as a neurotransmitter (Saint Marie and Peters, 1985; Tsumoto, 1990; Anderson et al., 1994).

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Details are yet sparse as to what populations of cortical neurons express vesicular Glu transporters. VGLUT1 mRNA is expressed at high levels in neurons of all layers (except layer I), whereas VGLUT2 mRNA shows a more restricted distribution with a preference for small neurons of layer IV (Ni et al., 1994; Hisano et al., 2000; Fremeau et al., 2001; Herzog et al., 2001; Fremeau et al., 2002). Although further analyses are required, distribution patterns suggest that pyramidal projection neurons primarily use VGLUT1 to accumulate Glu into synaptic vesicles (this is also supported by the distribution and appearance of VGLUT1-immunoreactive terminals in subcortical areas; Bellocchio et al., 1998; Sakata-Haga et al., 2001; Kaneko et al., 2002; Varoqui et al., 2002), whereas excitatory cortical local circuit neurons may use VGLUT2 for this purpose. With respect to the recently described VGLUT3, data on cortical expression are sparse and partly conflicting (Fremeau et al., 2002; Gras et al., 2002; Schäfer et al., 2002; Takamori et al., 2002). In conclusion, there is strong and overwhelming evidence that Glu acts as a neurotransmitter in most, if not all, projection neurons of the cerebral cortex and presumably also in excitatory cortical local circuit neurons (Fig. 1). Although there is now strong evidence in support of Glu as a neurotransmitter in the thalamic inputs to the cerebral cortex, the evidence in favor of such a role has been less straightforward than for corticofugal projections. In some studies, cortical injections of 3 D-[ H]aspartate have resulted in no or only few retrogradely labeled neurons in the thalamus (Streit, 1980; Baughman and Gilbert, 1981; Barbaresi et al., 1987). Others have demonstrated retrograde D-[3H]aspartate transport from the cortex to a large number of neurons in the nonspecific groups of nuclei (e.g., midline and intralaminar nuclei; Ottersen et al., 1983), in the lateral geniculate nucleus (Johnson and Burkhalter, 1992), and in the mediodorsal and other medial and intralaminar nuclei (Pirot et al., 1994). In Glu immunogold studies, high levels of Glu have been detected in collateral terminals of geniculocortical neurons in cats (Montero, 1990) and in anterogradely labeled thalamocortical terminals in the somatic sensory, auditory, and visual cortices of rats (Kharazia and Weinberg, 1993, 1994). The latter authors also noted a significant positive correlation between the densities of Glu immunogold labeling and synaptic vesicles in thalamocortical terminals. Terminals of thalamocortical axons projecting from the anterior thalamic nuclei to the retrosplenial granular cortex are similarly rich in Glu (Wang et al., 2001). In situ hybridization reveals strong expression of VGLUT2 mRNA in numerous neurons throughout the

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thalamocortical input, i.e., most dense in layer IV but also evident in layers I and VI (Bellocchio et al., 1998; Fremeau et al., 2001, Fujiyama et al., 2001; Herzog et al., 2001; Sakata-Haga et al., 2001; Kaneko and Fujiyama,, 2002; Kaneko et al., 2002; Varoqui et al., 2002; Minelli et al., 2003). Further, kainic acid lesions of the thalamic ventrobasal complex result in almost complete disappearance of VGLUT2 immunoreactivity in the somatosensory cortices, with no apparent reduction of VGLUT1 immunoreactivity (Fujiyama et al., 2001). Thus, available data from D-[3H]aspartate tracing, Glu immunogold labeling, and detection of vesicular Glu transporters and their mRNAs concur with physiological and pharmacological observations (see, e.g., Tsumoto, 1990; Hicks et al., 1991; McCormick and von Krosigk, 1992) in providing strong support for Glu as a transmitter in most, if not all, thalamocortical neurons. The literature on excitatory connections in the hippocampus has been reviewed elsewhere (Ottersen and Storm-Mathisen, 2000) and is not dealt with here.

Sensory Systems Somatosensory Pathways

FIGURE 1 Glutamatergic projections originating in the neocortex are shown. The neocortex gives rise to glutamatergic projections to the ipsilateral (1) and contralateral (2) neocortices, as well as to a large number of subcortical structures (some target structures have been left out for the sake of clarity): ACb, accumbens nucleus; Amg, amygdala; CG, central gray; CPu, caudate putamen; Cu, cuneate nucleus; Gr, gracile nucleus; IC, inferior colliculus; Pn, pontine nuclei; R, nucleus ruber; SC, superior colliculus; SN, substantia nigra; Th, thalamus; Tu, olfactory tubercle; VTA, ventral tegmental area.

thalamus, whereas VGLUT1 mRNA is expressed at low levels in many nuclei, except in the medial habenula where VGLUT1 mRNA expression is high (Ni et al., 1994; Hisano et al., 2000; Fremeau et al., 2001; Herzog et al., 2001). While VGLUT1-immunoreactive terminals are distributed fairly homogeneously throughout all cortical layers, immunocytochemical detection of VGLUT2 reveals high densities of terminal staining primarily in cortical layers receiving

Glu has been regarded as a strong transmitter candidate in primary afferent neurons ever since its excitatory effect on spinal neurons was detected (Curtis and Watkins, 1960; Rustioni and Weinberg, 1989; Broman, 1994; Broman et al., 2000), although there have been uncertainties regarding the proportion and types of primary afferent fibers using Glu as a neurotransmitter (Salt and Hill, 1983; Schneider and Perl, 1988). Investigations during the recent decade have provided strong support for Glu as a neurotransmitter in all categories of primary afferent fibers terminating in the dorsal horn and in dorsal column nuclei. Primary afferent terminals in all laminae of the spinal cord dorsal horn are rich in Glu (Broman et al., 1993; Valtschanoff et al., 1994), as are those in the cuneate nucleus (De Biasi et al., 1994b). Enrichment of Glu has also been described in select populations of primary afferent terminals in the spinal or trigeminal dorsal horns (De Biasi and Rustioni, 1988; Maxwell et al., 1990b, 1993; Merighi et al., 1991; Rousselot et al., 1994; Iliakis et al., 1996) and in vagal afferents to the solitary tract nucleus (Saha et al., 1995; Sykes et al., 1997). There are also positive correlations between the density of synaptic vesicles and the Glu immunogold labeling density in different populations of dorsal horn primary afferent terminals, further supporting a vesicular localization and thus a transmitter role of Glu in these terminals (Broman and Ådahl, 1994;

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Larsson et al., 2001). In contrast to Glu, the levels of aspartate in lamina I–IV primary afferent terminals are low and not associated with synaptic vesicles (Larsson et al., 2001). Investigations on the localization of vesicular Glu transporters demonstrate that VGLUT1 is present in relatively large terminals located in the deep laminae of the dorsal horn (lamina III–VI) and less densely in parts of the ventral horn, including the motor nuclei. VGLUT2-immunoreactive terminals are, on average, smaller than VGLUT1-immunoreactive terminals and are distributed more evenly throughout the spinal gray matter with the highest density in laminae I and II (Kaneko et al., 2002; Varoqui et al., 2002; Li et al., 2003; Todd et al., 2003). Transganglionic labeling with cholera toxin subunit B (CTb, which selectively labels myelinated primary afferent fibers) demonstrates that all CTb-labeled primary afferent terminals in the deep dorsal horn contain VGLUT1 and that some also contain VGLUT2 (Todd et al., 2003). Most CTb-labeled terminals in lamina I (presumably A∂ nociceptor terminals) contain VGLUT2 but none contain VGLUT1. Of the examined primary afferent C-fiber terminals in lamina II (defined by isolectin B4 staining or immunolabeling for substance P + CGRP or somatostatin + CGRP), some displayed weak staining and others no staining for VGLUT2, whereas none were stained for VGLUT1 (however, see Li et al., 2003). The latter finding is somewhat surprising considering the high levels of Glu in primary afferent C-fiber terminals (Broman et al., 1993; Broman and Ådahl, 1994; Valtschanoff et al., 1994). A possible explanation is that vesicular Glu uptake in these terminals depends on VGLUT3 (the expression of which has been detected in dorsal root ganglia; Gras et al., 2002) or on a mechanism that remains to be characterized (Todd et al., 2003). Glu has been detected in sizable proportions of terminals contacting the cell bodies or dendrites of neurons that give rise to ascending somatosensory pathways, including the spinothalamic tract (Westlund et al., 1992; Lekan and Carlton, 1995), the spinocervical tract (Maxwell et al., 1992), and the postsynaptic dorsal column pathway (Maxwell et al., 1995). Such terminals may originate from primary afferents, from intrinsic spinal neurons, or from descending pathways (e.g., the corticospinal tract; Valtschanoff et al., 1993). Although their anatomic organization remains to be defined, there is considerable evidence for the presence of intrinsic excitatory circuits in the dorsal horn (Willis and Coggeshall, 1991). In the superficial dorsal horn, high levels of Glu have been detected in neurotensinimmunoreactive terminals, presumed to have an

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exclusive intraspinal origin (Todd et al., 1994). The majority of such terminals also label for VGLUT2, as do most enkephalin-immunoreactive terminals in the superficial dorsal horn (Todd et al., 2003). Further, virtually all substance P and somatostatinimmunoreactive terminals that lack CGRP (i.e., terminals that originate from other sources than the primary afferents) are immunopositive for VGLUT2 (Todd et al., 2003). Thus, current evidence supports the presence of several populations of excitatory local circuit neurons in the dorsal horn that influence the activity of ascending projection neurons through synaptic release of Glu. Negative findings following D-[3H]aspartate tracing from the thalamus initially argued against a role for excitatory amino acids as neurotransmitters in the ascending somatosensory pathways (Rustioni et al., 1983), although findings in electrophysiological/ pharmacological studies did support such a role (Salt, 1986; Klockgether, 1987; Salt and Eaton, 1996). However, since 1990, a series of studies using Glu immunogold labeling have provided strong evidence in support of Glu as a neurotransmitter in a number of ascending somatosensory pathways. The most comprehensive studies have been made in cats or primates, but available data from rats and mice (De Biasi and Rustioni, 1990; Hamori et al., 1990; De Biasi et al., 1994a; Hamlin et al., 1996; Azkue et al., 1998) are entirely consistent with findings in other species. Thus, enrichment of Glu has been detected in spinocervical tract terminals in the lateral cervical nucleus (Broman et al., 1990; Kechagias and Broman, 1994, 1995), in terminals from the lateral cervical and dorsal column nuclei in the thalamic ventral posterior lateral nucleus (VPL; Broman and Ottersen, 1992; De Biasi et al., 1994a; Kechagias and Broman, 1995), in spinothalamic tract terminals in the nucleus submedius and posterior region of the thalamus (Ericson et al., 1995; Blomqvist et al., 1996), and in spinomesencephalic terminals in the periaqueductal gray (Azkue et al., 1998). Further, in several of these terminal populations, significant positive correlations between synaptic vesicle and Glu immunogold labeling densities were evident, thus supporting a vesicular accumulation of Glu. A notable exception to the aforementioned findings is the report of relatively low levels of Glu in terminals of the postsynaptic dorsal column pathway (PSDC; De Biasi et al., 1995). Glu levels in PSDC terminals in the cuneate nucleus are significantly lower than those detected in primary afferent terminals and are about the same as those in inhibitory terminals. Thus, the transmitter of neurons projecting from the spinal cord to dorsal column nuclei remains to be identified.

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Little is known about the expression of vesicular Glu transporters in ascending somatosensory pathways. However, VGLUT2-immunoreactive terminals in the ventral posterior lateral nucleus of the thalamus display light and electron microscopic features typical of terminals derived from ascending somatosensory fibers (Sakata-Haga et al., 2001; Kaneko and Fujiyama, 2002; Kaneko et al., 2002). In conclusion, there is strong and, in some cases, overwhelming evidence that Glu serves a neurotransmitter role in perhaps all somatosensory primary afferent fibers and in at least most central somatosensory pathways, including the somatosensory thalamocortical connections (see Anatomical Systems, Neocortex; Fig. 2). Visual Pathways The initial part of visual pathways is situated in the retina, where photoreceptors form synaptic connections with bipolar cells, which in turn synapse with ganglion neurons in the inner plexiform layer. These connections are referred to as the “vertical” or “through” pathway of the retina, whereas horizontal cells and amacrine cells form the intraretinal lateral connections. There is strong support for Glu as a neurotransmitter in the “vertical” pathway (Ehinger and Dowling, 1987; Massey and Redburn, 1987; Daw et al., 1989). Glu is released from photoreceptors (Copenhagen and Jahr, 1989), and several immunocytochemical studies have reported high levels of Glu in photoreceptor cells, especially in their terminals (Davanger et al., 1991; Kalloniatis and Fletcher, 1993; Jojich and Pourcho, 1996; Huster et al., 1998; Davanger et al., 1994b). Also, bipolar cells are rich in Glu and their terminals contain higher levels of Glu than their parent cell bodies (Ehinger et al., 1988; Davanger et al., 1991; Martin and Grünert, 1992; Kalloniatis and Fletcher, 1993; Davanger et al., 1994a; Jojich and Pourcho, 1996). Although Glu is generally considered an excitatory transmitter, in the synapses between photoreceptors and on-center bipolar cells, Glu exerts an inhibitory action through metabotropic receptors (Copenhagen, 1991; Nakajima et al., 1993; Euler et al., 1996; Sasaki and Kaneko, 1996; Vardi and Morigiwa, 1997; Brandstatter et al., 1997; De Vries and Schwartz, 1999; Morigiwa and Vardi, 1999). Thus, light-induced hyperpolarization of photoreceptors, leading to a diminished release of Glu from their terminals, results in depolarization of on-center bipolar cells (reduced inhibition) and a hyperpolarization of off-center bipolar cells (reduced excitation). The patterns of mRNA expression and immunolabeling for vesicular Glu transporters in the rat retina suggest that VGLUT1 is used for vesicular accumulation of Glu in both photoreceptors and bipolar cells (Mimura et al., 2002).

FIGURE 2 Schematic drawing of somatosensory and visual glutamatergic fiber systems. CTT, cervicothalamic tract; DRG, dorsal root ganglion primary afferent fibers; Hy, retinal projection to the hypothalamus; LG, retinal projection to the lateral geniculate nucleus; ML, fibers in the medial lemniscus from cuneate and gracile nuclei; Ret, retinal photoreceptors and bipolar cells; SC, retinal projection to the superior colliculus; SCT, spinocervical tract; SpPAG, spinomesencephalic input to periaqueductal gray; STT, spinothalamic tract; TCss, somatosensory thalamocortical projections; TCv, visual thalamocortical projections.

Signals generated by photoreceptors are communicated to the brain through the axons of ganglion cells projecting through the optic nerve and tract. The main termination of the optic tract is located in the thalamic lateral geniculate nucleus, which relays the signals to the visual cortices. Pharmacological data support Glu as a transmitter of retinal terminals in the lateral geniculate nucleus of rats (Crunelli et al., 1987), and enucleation in rats results in a loss of Glu in this nucleus on the contralateral side (Sakurai and Okada, 1992). Further, retinal terminals in the lateral geniculate of both cats and monkeys are rich in Glu immunoreactivity (Montero and Wenthold, 1989;

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Montero, 1990). Ganglion neurons in the rat retina also express VGLUT2 mRNA and display VGLUT2 immunoreactivity (Mimura et al., 2002). VGLUT2immunoreactive terminals in the rat lateral geniculate correspond morphologically to retinal terminals, and contralateral enucleation results in a large decrease of VGLUT2 immunoreactivity in the lateral geniculate nucleus (Sakata-Haga et al., 2001; Kaneko and Fujiyama, 2002; Kaneko et al., 2002). Thus, several lines of evidence support a transmitter role for Glu in ganglion cell terminals in the latter nucleus. Studies using 3 D-[ H]aspartate tracing or Glu immunogold labeling also point to a neurotransmitter role of Glu in optic tract terminals in other regions, including the superior colliculus (Matute and Streit, 1985; Ortega et al., 1995; Mize and Butler, 1996), the pretectum (Nunes-Cardoso et al., 1991), and the hypothalamus (Castel et al., 1993; De Vries et al., 1993; Chen and Pourcho, 1995). As stated in the section on the neocortex, both 3 D-[ H]aspartate tracing and Glu immunogold labeling suggest that Glu acts as a transmitter in the thalamocortical connection of the lateral geniculate nucleus. Thus, Glu appears to mediate signal transfer in each step of the visual pathway, from photoreceptor synapses in the retina to geniculocortical synapses in the visual cortex (Fig. 2). Auditory Pathways Considerable evidence supports an excitatory amino acid as a neurotransmitter in the synapses between inner hair cells and cochlear afferent nerve fibers (reviewed by Usami et al., 2000). Hair cells are rich in Glu in comparison to most other cellular elements in the cochlea (Usami et al., 1992), and GluR2/3 and GluR4 AMPA receptor subunits have been detected in inner, but not outer, hair cell synapses (Matsubara et al., 1996). However, although immunogold particles signaling Glu are associated with synaptic vesicles in hair cells, it remains to be shown that hair cell synaptic vesicles are indeed rich in Glu (Usami et al., 2000). Thus, although Glu must be considered a very strong cochlear hair cell transmitter candidate, definitive evidence is pending. Several lines of evidence, including pharmacological data, also support a transmitter role for Glu in cochlear nerve terminals in the cochlear nuclei (reviewed by Parks, 2000). Further, quantitative immunogold studies have shown that type I cochlear afferent terminals are rich in Glu, display high Glu/glutamine ratios, and are depleted in Glu following K+-induced depolarization (Hackney et al., 1996; see also Alibardi, 2003). Among the other terminal populations in the auditory system that have been subjected to Glu immunogold analysis are the calyces of Held in the

FIGURE 3 Schematic drawing of auditory glutamatergic fiber systems. cH, calyces of Held in the medial nucleus of the trapezoid body; CN-LSO, cochlear nucleus inputs to the lateral superior olive; HC, cochlear hair cells; Pf, granule cell/parallel fibers in the dorsal cochlear nucleus; TCa, auditory thalamocortical projections; 8cn, cochlear primary afferent fibers.

medial nucleus of the trapezoid body. These large terminals, which originate in the ventral cochlear nucleus on the contralateral side, exhibit a strong Glu immunogold labeling that is concentrated over vesicle clusters and mitochondria (Grandes and Streit, 1989). Helfert et al. (1992) detected high levels of Glu in round vesicle-containing terminals, presumably originating from the ipsilateral cochlear nucleus, in the lateral superior olive. In the dorsal cochlear nucleus, parallel fiber terminals originating from granule cells are rich in Glu, which is depleted by depolarization with high [K+] (Osen et al., 1995). High levels of Glu have also been recorded in auditory nerve terminals and granule cell terminals, as well as in large “mossy” terminals in the dorsal cochlear nucleus (Rubio and Juiz, 1998). As indicated in the section on the neocortex, enrichment of Glu has also been detected in auditory thalamocortical axon terminals (Kharazia and Weinberg, 1994; Fig. 3).

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Additional excitatory projections in the brain stem auditory system include inputs to the nuclei of the lateral lemniscus from the cochlear nucleus and lateral superior olive contralaterally and from the medial superior olive ipsilaterally. The list of excitatory connections also comprises cochlear nuclei efferents to the bilateral medial superior olive, input through the lateral lemniscus to the inferior colliculus (including fibers from lateral lemniscal nuclei), commissural connections between the inferior colliculi, and the projection from the inferior colliculus to the medial geniculate body (the latter projection also includes an inhibitory component). That these fiber systems use an excitatory amino acid as a neurotransmitter receives support from pharmacological–physiological studies and studies of retrograde transport or uptake/release of D-[3H]aspartate (Schwarz and Schwarz, 1992; Suneja et al., 1995; Saint Marie, 1996; Moore et al., 1998; Wu, 1998; Parks, 2000; Bartlett and Smith, 2002). Glu is the most likely transmitter candidate of the aforementioned fiber systems. In agreement, auditory relay stations in the brain stem and thalamus contain an abundance of nerve terminals immunoreactive for VGLUT1 or 2 (Sakata-Haga et al., 2001; Kaneko et al., 2002; Varoqui et al., 2002). Olfactory Pathways The olfactory system comprises three hierarchically ordered subdivisions: the olfactory epithelium located in the nasal cavity, the olfactory bulb, and the olfactory cortex, an array of cortical areas that receive direct input from the olfactory bulb (Shipley and Ennis, 1996; Chapter 29). Evidence shows that Glu acts as a neurotransmitter in both primary sensory afferents to the olfactory bulb and in bulbar projections to higher order olfactory areas. The axons of sensory neurons that reach the olfactory bulb terminate in spheroid structures of neuropil called glomeruli, where they establish synapses with the dendrites of the output neurons (mitral and tufted cells) and one type of interneuron, the periglomerular cell. The Glu-ergic nature of these afferents is supported by immunocytochemical studies, showing that Glu occurs in high levels in axon terminals of olfactory sensory neurons (Liu et al., 1989; Sassoè-Pognetto et al., 1993). Significantly, Glu immunoreactivity is more elevated in nerve terminals compared with axons and with postsynaptic dendrites (Didier et al., 1994). Electrophysiological recordings also support this conclusion, as stimulation of the olfactory nerve evokes AMPA and NMDA responses in mitral cells (Berkowicz et al., 1994; Ennis et al., 1996). Olfactory neurons also contain taurine and carnosine, a dipeptide that has been proposed to have modu-

latory or neuroprotective functions (Margolis, 1974; Sassoè-Pognetto et al., 1993; Didier et al., 1994; Horning et al., 2000). There is extensive immunocytochemical and pharmacological evidence that the output neurons of the olfactory bulb release Glu (reviewed in Shepherd and Greer, 1998; Haberly, 1998). Mitral and tufted cells are strongly labeled with antibodies against Glu (Ottersen and Storm-Mathisen, 1984a, 1984b; Liu et al., 1989). These neurons also show immunoreactivity for N-acetyl-L-aspartyl-L-glutamic acid (NAAG) and aspartate (Anderson et al., 1986; Saito et al., 1986; Blakely et al., 1987), but a transmitter role of these substances is not supported by functional analyses (Whittemore and Koerner, 1989; Trombley and Shepherd, 1993). Substantial evidence shows that mitral and tufted cells release Glu both from their dendrites in the glomerular layer and external plexiform layer (where they establish dendrodendritic synapses with periglomerular cells and with granule cells, respectively) and from their axons in the olfactory cortex (Hennequet et al., 1998). Dendrodendritic synapses between mitral/tufted cells and granule cells are reciprocal pairs consisting of an asymmetric and a symmetric junction (Rall et al., 1966; Price and Powell, 1970a; Fig. 4). In these reciprocal connections, release of Glu from the principal neurons activates AMPA and NMDA receptors and triggers the release of GABA from granule cell spines (Nicoll, 1971a; Nowycky et al., 1981; Jahr and Nicoll, 1982; Trombley and Shepherd, 1992; Wellis and Kauer, 1993, 1994; Sassoè-Pognetto and Ottersen, 2000). In addition to activating postsynaptic receptors, Glu released by mitral cell dendrites can spread out of the synaptic cleft and activate receptors on the parent dendrite as well as on neighboring cells (Nicoll, 1971b; Aroniadou-Anderjaska et al., 1999; Isaacson, 1999; Friedman and Strowbridge, 2000; Salin et al., 2001). Immunogold and electrophysiological investigations indicate that some granule cell spines may also release Glu, although it is presently unknown whether individual granule cells can release both Glu and GABA (Didier et al., 2001). In conclusion, there is convincing evidence that Glu serves as a neurotransmitter in primary afferents and in local dendrodendritic circuits of the olfactory bulb. Other synapses that presumably use Glu are those formed by axon collaterals of mitral and tufted cells in the internal plexiform layer and granule cell layer. Afferent fibers from cerebral hemispheres also establish asymmetric synapses at various levels in the olfactory bulb (Price and Powell, 1970b; Pinching and Powell, 1972). The Glu-ergic nature of these synapses is supported indirectly by immunolabeling of axodendritic junctions with antibodies directed against Glu

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FIGURE 4 Dendrodendritic reciprocal synapses between a mitral cell (mc) and a granule cell spine (gc) in the rat olfactory bulb. The reciprocal synaptic arrangement consists of an asymmetric, mitral-to-granule synapse (thick arrow) and a symmetric, granule-to-mitral synapse (empty arrow), located side by side. This section was labeled with an antiserum against the NR1 subunit of NMDA receptors, and strong immunolabeling is visible over the asymmetric junction. Another asymmetric synapse with a different granule cell spine (lower left) is also labeled. Adapted from Sassoè-Pognetto and Ottersen (2000).

receptor subunits (Sassoè-Pognetto and Ottersen, 2000). Finally, external tufted cells and possibly other types of Glu-positive juxtaglomerular neurons may establish Glu-ergic synapses in the periglomerular neuropil (Pinching and Powell, 1971; Liu et al., 1989). Vomeronasal System The vomeronasal system is a chemosensory pathway that has evolved in many terrestrial vertebrates to detect nonvolatile pheromones associated primarily with social and reproductive behaviors. Pheromonal

information detected by the vomeronasal sensory organ is conveyed through the accessory olfactory bulb to the amygdala and then to the hypothalamus (Halpern, 1987; Mori, 1987; Liman, 1996; Bargmann, 1997). A convergence of immunocytochemical and electrophysiological data indicates that the basic synaptic organization of the accessory olfactory bulb is similar to that of the main olfactory bulb (Dudley and Moss, 1995; Jia et al., 1999; Quaglino et al., 1999). Thus, Glu appears to be the neurotransmitter used both by vomeronasal afferents and by output neurons. As in

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the main olfactory bulb, a punctate immunoreactivity for Glu is present in the periglomerular region and in the granule cell layer (Quaglino et al., 1999), suggesting that other types of synapse are also Glu-ergic.

Motor Pathways Motoneurons in the spinal cord and brain stem constitute the final common pathway for motor commands. The synaptic inputs to motoneuronal cell bodies and dendrites at different levels of the neuroaxis have been examined extensively in regard to neuroactive amino acid contents (Shupliakov et al., 1993; Murphy et al., 1996, Tai and Goshgarian, 1996; Yang et al., 1997; Örnung et al., 1998; Bae et al., 1999; Lindå et al., 2000; Somogyi, 2002). The proportion of premotoneuron terminals defined as Glu-ergic in immunogold studies varies from about 35% to over 50%. Thus, Glu seems to be a predominant excitatory transmitter in nerve terminals synapsing on motoneurons. The pattern of immunostaining for the vesicular Glu transporters VGLUT1 and VGLUT2 further underscores an essential role for transmitter Glu in the excitation of motoneurons and interneurons in the ventral horn (Kaneko et al., 2002; Varoqui et al., 2002; Todd et al., 2003). In the ventral horn of the rat spinal cord, a moderate density of relatively large VGLUT1-immunoreactive terminals is evident in lamina VII and in motor nuclei. The smaller VGLUT2immunoreactive terminals occur at moderate to high densities throughout the ventral horn. Glu-ergic terminals in the ventral horn may originate from fiber tracts descending from the cortex or the brain stem, from intraspinal neurons, or from primary afferent fibers. The only type of primary afferent fiber connecting directly with motoneurons are Ia fibers from muscle spindles. As for other primary afferent fibers (see section on somatosensory pathways), there is overwhelming evidence in favor of Glu as a transmitter in Ia afferent boutons. Transmission between Ia fibers and motoneurons is blocked by excitatory amino acid receptor antagonists (Jessel et al., 1986). Transganglionically labeled Ia afferent terminals in contact with motoneurons and neurons in the central cervical nucleus are also rich in Glu (Örnung et al., 1995), as are giant boutons (likely to originate from Ia fibers) in Clarke’s column (Maxwell et al., 1990a). Further, all primary afferent terminals (labeled by the transganglionic transport of choleragenoid) in the ventral horn contain VGLUT1 but not VGLUT2 immunoreactivity (Todd et al., 2003). Thus, all types of primary afferent-relayed reflex activity likely depend on Glu-ergic neurotransmission. In contrast to the well-established neurotransmitter role for Glu in primary afferent fibers contacting

motoneurons and ventral horn local circuit neurons, there is weak evidence for Glu neurotransmission in other types of inputs. A notable exception is the corticospinal tract, which is widely recognized to use Glu as a transmitter (Storm-Mathisen and Ottersen, 1988; Rustioni and Weinberg, 1989; Valtschanoff et al., 1993). However, because direct corticospinal input to motoneurons is sparse in rats (Terashima, 1995), most Glu-ergic terminals in this species on motoneurons (except those of Ia afferent origin) must originate from intraspinal neurons or descending tracts. Considerable evidence from physiological– pharmacological studies shows that excitatory amino acid receptors mediate several different inputs to motoneurons (e.g. McCrimmon et al., 1989; Floeter and Lev-Tov, 1993; Pinco and Lev-Tov, 1994; Chitravanshi and Sapru, 1996; Hori et al., 2002). Glu is likely to be the transmitter in these inputs, although conclusive evidence is lacking. Several studies have detected the presence of Glu in cell bodies of brain stem neurons projecting to the spinal cord (Beitz and Ecklund, 1988; Mooney et al., 1990; Nicholas et al., 1992; Liu et al., 1995), but cell body labeling for Glu is unreliable as a marker for Glu-ergic neurons (Broman et al., 2000). However, Stornetta et al. (2003) reported that bulbospinal neurons in the rostral ventral respiratory group express VGLUT2 mRNA and that their terminals in the cervical ventral horn are immunoreactive for the same transporter. These findings indicate that this bulbospinal projection is Glu-ergic.

Basal Ganglia The basal ganglia comprise the striatum (caudate nucleus and putamen), the nucleus accumbens, the globus pallidus, the entopeduncular nucleus, the subthalamic nucleus, and the substantia nigra. These structures have profuse and complex fiber connections with each other, as well as with several other regions of the CNS. In only a minority of the many fiber connections of the basal ganglia in the rat has the transmitter substance been established by means of combined tracing and immunocytochemical studies at the ultrastructural level. However, information gained from various types of investigations has led to the identification of strong transmitter candidates for a number of the basal ganglia connections. The massive corticostriatal projection serves to illustrate this point. Early electrophysiological (Kitai et al., 1976; Wilson, 1986) and neurochemical (Spencer, 1976; Divac et al., 1977; Kim et al., 1977; Streit, 1980; Fonnum et al., 1981) investigations were suggestive of a Glu-ergic excitatory input to the striatum from the cerebral cortex. A subsequent light microscopical immunohistochemical study documented

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that a very large number of fibers and bouton-like structures in striatum of the rat display Glu immunoreactivity (Ottersen and Storm-Mathisen, 1984a, 1984b). It was later shown that striatal boutons with the ultrastructural characteristics of cortical afferents were rich in Glu and that they also sustained a high-affinity uptake of aspartate (Gundersen et al., 1996). Thus, even though immunocytochemical analyses of anterogradely labeled corticostriatal boutons are pending, available data support the notion that cortocostriatal fibers use Glu as a transmitter. This would be in line with the many reports on the distribution in the striatum of the rat of various types of Glu receptors (Bernard et al., 1996; Wullner et al., 1997; Petralia et al., 2000; Shigemoto and Mizuno, 2000; Wisden et al., 2000). It should be noted, however, that organization of the corticostriatal projection is very complex, and it remains an open question whether Glu is a transmitter in all corticostriatal fibers or only in a subpopulation of them. A light microscopic investigation showed that 52–61% of retrogradely labeled corticostriatal neurons in the rat displayed Glu immunoreactivity (Bellomo et al., 1998). Up to 96% of these neurons were immunopositive when antisera against Glu and aspartate were used simultaneously, and Glu- and aspartate-immunopositive cortical neurons appeared to be largely segregated (Bellomo et al., 1998). These data should be interpreted with caution as the level of Glu (and aspartate) in cell bodies may reflect the size of the metabolic pools rather than the transmitter pools of the respective amino acids. The cerebral cortex also sends fibers to other basal ganglia than the striatum, although on a smaller scale. Thus, the subthalamic nucleus (STN) of the rat receives fibers from wide cortical areas (Afsharpour, 1985; Canteras et al., 1988). Electrophysiological investigations suggested that this input was excitatory (Kitai and Deniau, 1981; Rouzaire-Dubois and Scarnati, 1987; Feger and Mouroux, 1991; Fujimoto and Kita, 1993). In a combined tracing and immunocytochemical study in the rat, it was indeed shown that a considerable number of the corticosubthalamic boutons are rich in Glu (Bevan et al., 1995). In the rat the corticosubthalamic projection is accompanied by a more modest corticopallidal pathway (Naito and Kita, 1994). The corticopallidal boutons have an ultrastructural appearance similar to that of cortical efferents in other basal ganglia, suggesting that these afferents are Glu-ergic. Immunocytochemical evidence of this is pending. The thalamus represents the second largest source of afferents to the basal ganglia. In a combined tracing and immunocytochemical study in the rat, it was shown that axon terminals in the subthalamic nucleus

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that originate in the parafascicular nucleus of the thalamus are very rich in Glu (Bevan et al., 1995). This finding lends support to earlier physiological and pharmacological investigations (Mouroux and Féger, 1993; Féger et al., 1997). As far as the massive thalamostriatal projection is concerned, the identity of the transmitter substance remains to be determined (De las Heras et al., 1997), although electrophysiological studies point to an excitatory signal substance (Purpura and Malliani, 1967; Buchwald et al., 1973; Kitai et al., 1976). It is likely that at least part of the complex thalamostriatal projection is Glu-ergic. A light microscopical study has suggested that many afferents to the ventral striatum from the amygdala in the rat use Glu as a transmitter (McDonald, 1996). In recent years the subthalamic nucleus has taken central stage in attempts to explain the pathophysiology of Parkinson’s disease (Albin et al., 1989b). Although several observations clearly indicate that the original model was too simplified (Marsden and Obeso, 1994; Levy et al., 1997; Obeso et al., 1997, 2000; Wichmann and DeLong, 1998; Bar-Gad and Bergman, 2001), it remains unquestionable that hyperactivity of the subthalamic neurons is a prominent feature in Parkinson’s disease. The main efferent projections of the subthalamic nucleus terminate in the two segments of the globus pallidus (primate), the globus pallidus and entopeduncular nucleus (rodents), and in the pars compacta and pars reticulata of the substantia nigra. A smaller contingent of fibers extends to the tegmental pedunculopontine nucleus (PPN) (Parent and Hazrati, 1993). Light microscopical immunohistological studies have shown that practically all cells in STN display strong Glu immunoreactivity in the rat (Ottersen and Storm-Mathisen, 1984a, 1984b) as in other species (Smith and Parent, 1988; Albin et al., 1989a). Because the presence of Glu in neuronal cell bodies is not necessarily indicative of a transmitter role, these findings are not decisive. However, a transmitter role of Glu is supported by electrophysiological observations in the rat (Kitai and Kita, 1987) and by combined tracing and immunocytochemical studies in the cat (Rinvik and Ottersen, 1993). The latter immunogold analysis showed that boutons of subthalamonigral fibers are rich in Glu. Similar investigations have not been undertaken for the subthalamopallidal or subthalamoentopeduncular projections. However, in the rat there is ample evidence that the subthalamofugal fibers send branches to both the substantia nigra and the globus pallidus (van der Kooy and Hattori, 1980). When correlated with immunohistochemical and tracing studies in other species, as well as with electrophysiological investigations in the rat (Robledo and

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Féger, 1990), it appears highly likely that the subthalamic projections to the globus pallidus and entopeduncular nucleus are Glu-ergic. The nature of the transmitter substance in the projection from the subthalamic nucleus to the pedunculopontine nucleus (PPN) remains to be determined. However, some data are available on the reciprocal connection. In combined tracing and immunocytochemical studies in the rat it was shown that PPN sends Glu-enriched fibers to the STN (Bevan and Bolam, 1995) and entopeduncular nucleus (Clarke et al., 1997). The latter authors demonstrated that a significant portion of labeled axon terminals from PPN displayed high levels of immunoreactivity against both Glu and choline acetyltransferase, suggestive of a colocalization of Glu and acetylcholine.

Cerebellum The transmitter systems of the cerebellum have been reviewed by Voogd et al. (1996) and by Ottersen and Walberg (2000). These reviews should be consulted for a complete bibliography. Mossy and climbing fibers constitute the major afferent pathways to the cerebellar cortex (for references, see Palay and Chan-Palay, 1974). It has long been known that these pathways are excitatory and that they have separate origins; the most important ones being the pontine nuclei and the inferior olive, respectively. The mossy fiber system is by far the more massive and establishes contacts with dendritic digits of granule cells. The latter cells are themselves excitatory (see later) and constitute the second leg in a disynaptic excitatory input to Purkinje cells. In contrast, climbing fibers excite Purkinje cells directly through their synapses on Purkinje cell dendritic thorns. It is now well established that both of the major afferent pathways mediate fast excitation (Ito, 1984), and several lines of evidence point to Glu as the likely transmitter. Mossy fibers display a strong immunogold signal for Glu (Somogyi et al., 1986; Fig. 5B). The intensity of this signal is correlated positively to the packing density of

synaptic vesicles (Ji et al., 1991) and is abolished following depolarization of cerebellar slices with high [K+] (Ottersen et al., 1990). This implies that the immunolabeling is likely to represent a transmitter pool. The postsynaptic elements of mossy fibers express several types of Glu receptor (Cox et al., 1990; Gallo et al., 1992; Petralia and Wenthold, 1992), and pharmacological data are consistent with the idea that Glu acts as their endogenous ligand (Garthwaite and Brodbelt, 1990). Mossy fibers are also very rich in phosphateactivated glutaminase (Laake et al., 1999), which is a key enzyme in Glu synthesis. VGLUT1 and VGLUT2 have both been identified in mossy fibers (Fremeau et al., 2001), but there is still some uncertainty as to the degree of colocalization of these two vesicular Glu transporters. It must be emphasized that some mossy fibers may use other signal substances, instead of or in addition to Glu. Subpopulations of mossy fibers express cholinergic markers and neuroactive peptides with presumed modulatory functions. These data are reviewed by Voogd et al. (1996) and by Ottersen and Walberg (2000). Climbing fibers are now believed to use Glu as a transmitter, like the majority of mossy fibers. Climbing fibers are very rich in Glu (Ottersen et al., 1992), contain the vesicular Glu transporter VGLUT2 (Fremeau et al., 2001; Pahner et al., 2003), and face thorns that express high concentrations of AMPA receptors (Landsend et al., 1997). Early studies showed that climbing fibers take up and retrogradely transport 3 D-[ H]aspartate to their perikarya in the inferior olive (Wiklund et al., 1984). In regard to the latter finding, it should be noted that the tracer D-[3H]aspartate does not differentiate between transport of the endogenous substrates L-aspartate and L-Glu (Danbolt et al., 1994). In fact, L-aspartate was long held to be the most likely climbing fiber transmitter. Supporting this view were slice experiments showing that evoked release of endogenous aspartate from the cerebellar cortex could be reduced by lesions of the inferior olive by 3-acetylpyridine (Toggenburger et al., 1983; Vollenweider et al., 1990). However, quantitative immunogold

FIGURE 5 Electron micrographs showing the distribution of glutamate-like immunoreactivity (small gold particles) in the rat cerebellar cortex. The section was also labeled with antibodies to glutamine (an important glutamate precursor), which were visualized by the use of large gold particles. (A) Molecular layer. Parallel fiber terminals (pf) are labeled strongly for glutamate. Some glutamate immunoreactivity is also found in spines (s), reflecting the presence of a metabolic pool. Glial processes (g) contain little glutamate immunoreactivity but display significant glutamine immunolabeling. Some large particles (indicating glutamine) overlie the intercellular space (arrows). (B) Granule cell layer. A mossy fiber terminal (mf) is strongly glutamate immunoreactive and also appears to contain a sizable pool of glutamine. The Golgi (Go) cell terminal (probably GABAergic) is comparatively weakly labeled. Note the flat vesicles (arrows). Asterisks denote granule cell dendrites (adapted from Ottersen et al., 1992). Scale bars: 0.4 μm.

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analyses with specific antibodies demonstrated that the level of L-aspartate in climbing fiber terminals was low compared with the average tissue level and with the level of this amino acid in the parent cell bodies in the inferior olive (Zhang et al., 1990). It is possible that a relative shortage of oxygen and energy substrates during the preparation and incubation of brain slices leads to a buildup of L-aspartate in nerve terminals that contain only sparse amounts of this amino acid under physiological conditions (Gundersen et al., 1998). One must also consider the possibility that immunogold analyses fail to reveal the entire endogenous pool of transmitter L-aspartate, either because this pool is released during the preparation of the tissue or because it is inaccessible to immunogold detection. The latter explanations are unlikely but cannot be ruled out entirely. Another excitatory amino acid that has been implicated in climbing fiber neurotransmission is homocysteic acid (HCA; Cuénod et al., 1989). Like L-aspartate, this sulfur-containing amino acid is released in smaller quantities than normal following lesions of the inferior olive (Vollenweider et al., 1990). However, with the advent of specific antibodies it was shown that HCA-like immunoreactivity is largely confined to glial elements, including Bergmann fibers (Grandes et al., 1991; Zhang and Ottersen, 1992, 1993). This rules out a transmitter role of HCA in climbing fibers. The possibility remains that HCA is engaged in an unorthodox signaling process involving release from glial cells (Do et al., 1997). If a substrate of plasma membrane Glu transporters is responsible for signal transfer in climbing fiber–Purkinje cell synapses, one would expect an interference with Glu transport to affect the postsynaptic response to climbing fiber activation. To test this, Takahashi et al. (1996) injected D-aspartate into Purkinje cells and indeed demonstrated that this led to a prolonged excitatory postsynaptic current at the climbing fiber synapses. The most likely explanation of this finding is that the injected D-aspartate inhibits an excitatory amino acid transporter that normally contributes to the removal of transmitter from the synaptic cleft (also see Otis et al., 1997). Candidate transporters are EAAT4, which is concentrated at specific membrane domains in Purkinje cell spines (Dehnes et al., 1998), and EAAT3 (formerly EAAC1), which is distributed more generally in neuronal plasma membranes (Rothstein et al., 1994). Similar to mossy fibers, climbing fibers contain a number of neuroactive substances in addition to Glu, including peptides with possible modulatory functions (Voogd et al., 1996). Thus data reviewed earlier

must not be taken to indicate that Glu is the sole neuroactive compound released from climbing fibers. Parallel fibers are the axons of cerebellar granule cells and serve as the second leg of a disynaptic excitatory input to Purkinje cells. Parallel fibers also establish synapses with dendritic stems of interneurons (Palay and Chan-Palay, 1974). Compelling evidence points to Glu as a parallel fiber transmitter. One of the first pieces of evidence came with the study of Young et al. (1974), who observed that a granule cell loss (caused by virus infection) was accompanied by a decreased content of Glu and Glu/aspartate uptake in the cerebellar cortex. Subsequent investigations showed that the content and uptake, as well as the release of Glu, depend on intact granule cells (for reviews, see Ito, 1984; Ottersen and Storm-Mathisen, 1984b). Parallel fiber terminals display a strong immunogold signal for Glu (Somogyi et al., 1986; Ottersen, 1989; Fig. 5A) and this signal depends on a Glu pool that can be depleted by high [K+] (Ottersen et al., 1990). Immunogold analyses have also shown that the postsynaptic specializations of parallel fiber synapses express AMPA and δ2 receptors (Baude et al., 1994; Nusser et al., 1994; Landsend et al., 1997). Parallel fiber terminals contain the vesicular Glu transporter VGLUT1 (Fremeau et al., 2001; Pahner et al., 2003). Data supporting a transmitter role of Glu in parallel fibers are indeed overwhelming, but it remains to clarify how their transmitter pool is maintained. Whereas the major Glu-synthesizing enzyme phosphate-activated glutaminase (PAG) is abundant in mossy fiber terminals, it occurs at very low levels in terminals of parallel fibers (Laake et al., 1999). Thus the latter fibers probably depend on alternative sources for transmitter replenishment. Glu is also a strong transmitter candidate for one type of interneuron in the cerebellar cortex: the unipolar brush cell (Mugnaini et al., 1997). This cell is exceptional among cerebellar interneurons by showing an enrichment in Glu and being presynaptic to Glu receptors (Nunzi and Mugnaini, 1999). The unipolar brush cell contacts granule cells and other unipolar brush cells, and presumably also Golgi cells (Nunzi and Mugnaini, 1999).

CONCLUSION The present survey of putative Glu-ergic fiber systems emphasizes the predominant role of Glu as a transmitter of projection neurons in the central nervous system. The evidence is now compelling that Glu is involved in signal transfer in major sensory and

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motor pathways and in associational and commissural connections of the neocortex. Glu also appears to serve a transmitter role in all of the major fiber pathways in the cerebellum with the exception of the GABA-ergic Purkinje cell projection. In contrast, few types of local circuit neurons exhibit a Glu-ergic phenotype. The latter class of neurons largely depends on GABA or glycine for fast synaptic transmission. Glu is certainly a prevalent transmitter, but this must not be equated with uniformity at the level of synaptic transmission. As the present review has been focused on Glu, we have not elaborated on the issue of transmitter colocalization. The fact is that many of the fiber systems that contain Glu also contain other neuroactive compounds. These may be colocalized with Glu or occur in separate fibers. It is also clear that L-aspartate may rival Glu as a transmitter in certain fiber systems (Gundersen and Storm-Mathisen, 2000). Another level of complexity is added by the structural and molecular heterogeneity among Glu synapses. Some Glu synapses are ensheathed by glial processes, whereas other synapses lack glial investment (Chaudry et al., 1995). Such differences obviously affect transmitter diffusion and hence synaptic function, and the complement of glutamate receptors varies widely between synapses even within individual fiber pathways (e.g., Nusser et al., 1998; Takumi et al., 1999). As a result, Glu synapses exhibit a functional diversity that may be underestimated easily when the focus is restricted to the issue of transmitter phenotype. In fact, unraveling the heterogeneity of Glu synapses will be the next major challenge once the general map of Glu pathways has been established.

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VII. NEUROTRANSMITTERS

Index

A A5 group, autonomic control, 772, 774 A5 neuron, features, 284 A7 group, autonomic control, 772 A7 neuron, features, 284 AA, see Anterior aygdaloid area Accessory olfactory bulb (AOB) connections afferents, 950 efferents, 949–950 high-order connections and reproductive function, 950–952 vomeronasal organ, 949 modulatory inputs differential innervation, 953 locus coeruleus, 952–953 nucleus of the diagonal band, 952 raphe nuclei, 952–953 neuron types, 949 neurotransmitters, 949 sexual dimorphism, 950 Accessory optic system connections, 1119 dorsal terminal nucleus, 1121 functions, 1121 lateral terminal nucleus, 1120–1121 medial terminal nucleus, 1118–1120 subdivisions, 1118 Acetylcholine Alzheimer's disease involvement of neurons, 1265–1266 anatomy of cholinergic systems brain stem projections, 1260–1261 hippocampus projections, 1262–1263 medial habenula projections, 1262 mesopontine projections, 1261–1262 overview, 1259–1260 septum projections, 1262–1263 spinal cord projections, 1260–1261 striatum projections, 1262 facial nucleus neurotransmission in medulla, 305 history of study, 1257–1258 receptors and signaling, 1264–1265 striatum medium spiny projection neuron receptors, 478

Acetylcholine Continued vascular innervation, 1194–1195 vesicular transporter localization, 1258 Acetylcholinesterase (AChE) central division of the extended amygdala staining central division of the supracapsular bed nucleus of the stria terminalis, 567 interstitial nucleus of the posterior limb of the anterior commissure, 568 lateral bed nucleus of the stria terminalis, 564 lateral part of the central amygdaloid nucleus, 565–566 laterobasal nuclear complex staining basolateral amygdaloid nucleus, 584 basomedial amygdaloid nucleus, 586 lateral amygadaloid nucleus, 583 ventral basolateral amygdaloid nucleus, 585 medial division of the extended amygdala staining intraamygdaloid bed nucleus of the stria terminalis, 560 medial amygdaloid nucleus, 559–560 medial bed nucleus of the stria terminalis, 558 medial sublenticular extended amygdala, 561 olfactory amygdala staining amygdalopiriform transition area, 533–534 anterior aygdaloid area, 517 anterior cortical amygdaloid nucleus, 524 nucleus of the lateral olfactory tract, 520 posterolateral cortical amygdaloid nucleus, 527 vomeronasal amygdala staining amygdalohippocampal transition area, 543 bed nucleus of the accessory olfactory tract, 535 posteromedial cortical amygdaloid nucleus, 535

1293

AChE, see Acetylcholinesterase ACO, see Anterior cortical amygdaloid nucleus ACTH, see Adrenocorticotropin AD, see Alzheimer's disease α2-Adenosine receptors, striatum medium spiny projection neurons, 477–478 Adrenocorticotropin (ACTH), vestibular neuron modulation, 985 Agranular insular cortex areas, 750 cytoarchitectonics, 750 local cerebral glucose utilization studies, 750 neurotransmitter receptors, 750–751 AHI, see Amygdalohippocampal transition area Alar plate, gene expression in development, 16–17 Alzheimer's disease (AD) cingulate cortex pathology, 705, 721–724 p75 neurotrophin receptor role, 1265–1266 γ-Aminobutyric acid (GABA) dorsal horn neurochemistry, 140 locus coeruleus inputs, 271–273 motor trigeminal nucleus neurotransmission in medulla, 301 striatum medium spiny projection neuron receptors, 478 Amygdala, see also Extended amygdala; Laterobasal nuclear complex amygdalostriatal transition zone acetylcholinesterase staining, 591 cytoarchitectonics, 591 heavy metal staining, 591 topographic landmarks, 591 autonomic control basolateral complex, 783 central nucleus, 782–783 extended amygdala, 783–784 bed nucleus of the anterior commissure, 592 functional overview, 509–510 fusiform nucleus, 593 gestational time of origin, 29–31

1294 Amygdala Continued histochemical staining, 514 intramedullary gray acetylcholinesterase staining, 592 cytoarchitectonics, 591 heavy metal staining, 591–592 topographic landmarks, 591 locus coeruleus connections, 280 neuron origins, 62–63 nucleus of the commissural component of the stria terminalis acetylcholinesterase staining, 593 cytoarchitectonics, 592 heavy metal staining, 592 topographic landmarks, 592 olfactory amygdala amygdalopiriform transition area acetylcholinesterase staining, 533–534 choline acetyltransferase staining, 534 cytoarchitectonics, 530 heavy metal staining, 533 topographic landmarks, 527, 530 anterior aygdaloid area acetylcholinesterase staining, 517 choline acetyltransferase staining, 517 cytoarchitectonics, 515 fibroarchitectonics, 515 heavy metal staining, 515, 517 projections, 514 topographic landmarks, 515 anterior cortical amygdaloid nucleus acetylcholinesterase staining, 524 choline acetyltransferase staining, 524 cytoarchitectonics, 522–523 fibroarchitectonics, 523–524 heavy metal staining, 524 topographic landmarks, 521–522 connections, 543, 547, 551–556 neuropeptides, 543 neurotransmitters, 543 nucleus of the lateral olfactory tract acetylcholinesterase staining, 520 choline acetyltransferase staining, 520–521 cytoarchitectonics, 517 fibroarchitectonics, 517, 520 heavy metal staining, 520 topographic landmarks, 517 posterolateral cortical amygdaloid nucleus acetylcholinesterase staining, 527 choline acetyltransferase staining, 527 cytoarchitectonics, 525 fibroarchitectonics, 525 heavy metal staining, 525 topographic landmarks, 524–525 parastrial nucleus, 592 subdivisions, 510, 512–513 subventricular nucleus, 592 tachykinin projections, 1244

INDEX

Amygdala Continued topography, 509–510, 512–513 unclassified nucleus connections, 593–594 vomeronasal amygdala amygdalohippocampal transition area topographic landmarks, 535, 537 acetylcholinesterase staining, 543 choline acetyltransferase staining, 543 cytoarchitectonics, 537 fibroarchitectonics, 537, 541 heavy metal staining, 541, 543 bed nucleus of the accessory olfactory tract acetylcholinesterase staining, 535 cytoarchitectonics, 534 fibroarchitectonics, 534 heavy metal staining, 534–535 topographic landmarks, 534 connections, 543, 547, 551–556 neuropeptides, 543 neurotransmitters, 543 posteromedial cortical amygdaloid nucleus acetylcholinesterase staining, 535 cytoarchitectonics, 535 heavy metal staining, 535 topographic landmarks, 535 Amygdalohippocampal transition area (AHI) acetylcholinesterase staining, 543 choline acetyltransferase staining, 543 cytoarchitectonics, 537 fibroarchitectonics, 537, 541 heavy metal staining, 541, 543 topographic landmarks, 535, 537 Amygdalopiriform transition area (APIR) acetylcholinesterase staining, 533–534 choline acetyltransferase staining, 534 cytoarchitectonics, 530 heavy metal staining, 533 topographic landmarks, 527, 530 Amygdalostriatal transition zone (Astr) acetylcholinesterase staining, 591 cytoarchitectonics, 591 heavy metal staining, 591 topographic landmarks, 591 Angiotensin II, subfornical organ actions, 394 Anterior aygdaloid area (AA) acetylcholinesterase staining, 517 choline acetyltransferase staining, 517 cytoarchitectonics, 515 fibroarchitectonics, 515 heavy metal staining, 515, 517 projections, 514 topographic landmarks, 515 Anterior cerebral artery, anatomy, 1179–1180 Anterior choroidal artery, anatomy, 1179 Anterior cortical amygdaloid nucleus (ACO) acetylcholinesterase staining, 524 choline acetyltransferase staining, 524 cytoarchitectonics, 522–523 fibroarchitectonics, 523–524 heavy metal staining, 524 topographic landmarks, 521–522

Anterior olfactory nucleus (AON) architecture, 937–938 inputs, 938 neurotransmitters, 942 outputs, 938–939, 942 Anterior periventricular nucleus (PeA), functions, 339 Anterior thalamic nuclei connections, 430–431 functions, 431–432 types, 430 Anterodorsal preoptic nucleus, functions, 344 Anteroventral periventricular nucleus (AVPV), functions, 338–339 AOB, see Accessory olfactory bulb AON, see Anterior olfactory nucleus AOT, see Bed nucleus of the accessory olfactory tract APIR, see Amygdalopiriform transition area Arcuate nucleus autonomic control, 782 functions, 340 Area 24b, role in movement, vision, and pain behaviors, 712–713 Area 25 cytology, 707 infra limbic area IL, 707 Area 30, N-methyl-D-aspartate receptor antagonist-induced deafferentiation, 718–719 Area prostrema afferents, 397 efferents, 397–398 gross anatomy, 397 hormone secretion and receptors, 398 Area X afferent fibers, 114 cytoarchitecture, 125–126 Arterial circle, anatomy, 1176–1177, 1179–1180 Arteries, see Spinal cord; Vasculature, cerebral; Vascular innervation Astr, see Amygdalostriatal transition zone Auditory cortex areas, 1051, 1053–1054 behavioral studies, 1058 descending pathways corticofugal pathways, 1059–1061 overview, 1058–1059 fiber systems, 1055 layers, 1054–1055 mapping, 1051 modular organization, 1057–1058 plasticity, 1058 stimulus response studies, 1055, 1057 Auditory system, see also Auditory cortex; Cochlea; Cochlear nuclear complex; Inferior colliculus; Medial geniculate body; Nuclei of the lateral lemniscus; Superior olivary complex glutaminergic pathways, 1275–1276 organ of Corti, 998–1003 peripheral system, 997–998

INDEX

Autonomic control system forebrain level, behavioral and metabolic integration amygdala basolateral complex, 783 central nucleus, 782–783 extended amygdala, 783–784 cerebral cortex infralimbic cortex, 785 insular cortex, 784–785 motor cortex, 786 perirhinal cortex, 785 prelimbic cortex, 785 somatosensory cortex, 786 hypothalamus anteroventral third ventricular area, 778 arcuate nucleus, 782 dorsal hypothalamic area, 780 dorsomedial nucleus, 780 lateral hypothalamic area, 780, 782 paraventricular nucleus, 778–780 posterior lateral hypothalamic area, 782 retrochiasmatic area, 782 tuberomammillary nucleus, 782 thalamus intralaminar nuclei, 777–778 mediodorsal nucleus, 777 paraventricular nucleus, 777 ventroposterior parvocellular nucleus, 776–777 hierarchical model, 761 medullospinal level and reflex control caudal ventrolateral medulla, 767 nucleus of the solitary tract, 763, 765 raphe nuclei, 767 rostral ventrolateral reticular nucleus, 765, 767 ventromedial medulla, 767 mesopontine level, reflex control and arousal modulation and integration A5 group, 772, 774 A7 group, 772 cerebellum, 776 locus coeruleus, 775 midbrain raphe nuclei, 775–776 parabrachial nucleus, 768, 771–772 pedunculopontine and laterodorsal tegmental nuclei, 776 periaqueductal gray, 772 network model, 761 Autonomic ganglia cardiac plexus, 94 general organization, 77–79 groups, 77 head ganglia, 83–84, 94 intramural ganglia of the gut, 94–97 neuromuscular junctions, 97–99 neurotransmitters, 90–91 pelvic ganglia, 91–94 pelvic plexus, 82–83 preganglionic neuron and fiber structure, 84–85 prevertebral ganglia, 81–82, 91

Autonomic ganglia Continued principal ganglion neuron features, 86–87 rami communicantes, 80 sensory fibers, 99, 101, 103 small intensely fluorescent cells, 87–90 splanchic nerves, 81 sympathetic chains, 79–80 sympathetic ganglia, 85–86 tracheal ganglia, 94 AVPV, see Anteroventral periventricular nucleus

B BAC, see Bed nucleus of the anterior commissure Basal ganglia, see also Striatum; Substantia nigra functional overview, 455 glutaminergic pathways, 1278–1280 organization, 455–458 outputs medial globus pallidus, 482 substantia nigra, 482–484 tachykinin projections, 1244 Basal plate, gene expression in development, 14–16 Basilar artery, anatomy, 1173 Basilar pontine nuclei afferents cerebellum, 169 cerebral cortex, 168–169 Edinger–Westphal nucleus, 171 hypothalamus, 171 locus coeruleus, 171 nucleus of Darschewitsch, 171 raphe nuclei, 171 spinal cord, 171 tectum, 171 cytoarchitecture, 168 efferents, 171–172 functions, 172–173 neurons, 167 Basolateral amygdaloid nucleus (BL) acetylcholinesterase staining, 584 choline acetyltransferase staining, 584 cytoarchitectonics, 583–584 fibroarchitectonics, 584 heavy metal staining, 584 topographic landmarks, 583 Basomedial amygdaloid nucleus (BM) acetylcholinesterase staining, 585–586 choline acetyltransferase staining, 586 cytoarchitectonics, 585 fibroarchitectonics, 585 heavy metal staining, 585 topographic landmarks, 585 Bauplan anteo-posterior patterning, 9–10 differential aspects of neurogenesis, 6–7 dorsoventral patterning, 7–9 gene expressoin, 10 tagma, 10

1295 Bed nucleus of the accessory olfactory tract (AOT) acetylcholinesterase staining, 535 cytoarchitectonics, 534 fibroarchitectonics, 534 heavy metal staining, 534–535 topographic landmarks, 534 Bed nucleus of the anterior commissure (BAC), features, 592 Bed nucleus of the stria terminalis (BST), locus coeruleus connections, 279–280 BL, see Basolateral amygdaloid nucleus BLV, see Ventral basolateral amygdaloid nucleus BM, see Basomedial amygdaloid nucleus Brain stem analgesia system, 868–870 cholinergic projections, 1260–1261 dentate gyrus connections, 647–648 paraventricular nucleus connections, 380 periaqueductal gray connections, 247 serotonergic projections, 1209 spinal cord pathways in micturition coordination abdominal pressure control systems, 323 diffuse descending systems, 323 micturition reflex, 325 overview, 323–324 tachykinin projections, 1242–1243 trigeminal sensory system sensory nuclei afferent organization, 825–826 caudal subnucleus chemoarchitecture, 829 cytoarchitecture, 829 projections, 830–831 responses, 829–830 cytoarchitectonics, 826 groups, 824–825 interpolar subnucleus chemoarchitecture, 828–829 cytoarchitecture, 828–829 projections, 829 responses, 829 mesencephalic trigeminal nucleus chemoarchitecture, 827 cytoarchitecture, 827 projections and function, 827 oral subnucleus chemoarchitecture, 828 cytoarchitecture, 828 projections, 828 responses, 828 paratrigeminal nucleus chemoarchitecture, 831 cytoarchitecture, 831 projections, 831 responses, 831 principal sensory nucleus chemoarchitecture, 827–828 cytoarchitecture, 827–828 projections, 828 responses, 828 somatotopy, 826–827 BST, see Bed nucleus of the stria terminalis

1296

INDEX

BSTIA, see Intraamygdaloid bed nucleus of the stria terminalis BSTL, see Lateral bed nucleus of the stria terminalis BSTM, see Medial bed nucleus of the stria terminalis BSTSc, see Central division of the supracapsular bed nucleus of the stria terminalis BSTSm, see Medial division of the supracapsular bed nucleus of the stria terminalis

C Cajal–Retzius neuron, migration, 66 Calcitonin gene-related peptide (CGRP), dorsal horn neurochemistry, 137 Cardiac plexus, anatomy, 94 CeL, see Lateral part of the central amygdaloid nucleus CeM, see Medial part of the central amygdaloid nucleus Central division of the supracapsular bed nucleus of the stria terminalis (BSTSc) acetylcholinesterase staining, 567 cytoarchitectonics, 566 heavy metal staining, 566–567 topographic landmarks, 566 Central lateral nucleus (CL) afferents, 436 efferents, 437 functions, 440 structure, 436 Central medial nucleus (CM) afferents, 436 efferents, 437 functions, 440 structure, 435 Central sublenticular extended amygdala (SLEAc) cytoarchitectonics, 566 heavy metal staining, 566 topographic landmarks, 566 Cerebellum afferent mossy fiber systems, 229, 231 autonomic control, 776 basilar pontine nuclei connections, 169 glutaminergic pathways, 1280, 1282 gross anatomy, 205, 207–208 lateral reticular nucleus connections, 177–178 locus coeruleus connections, 280–281 nuclei anterior interposed nucleus, 215–216 lateral nucleus, 216 medial nucleus, 210–211 neurons, 208–209 posterior interposed nucleus, 211, 214–215 subdivision, 208 Purkinje cell organization and connections corticonuclear projection zones, 216–219 longitudinal zone chemoarchitecture, 224–226

Cerebellum Continued olivocerebellar projection, 219–222 paraflocculus, 223–224 vestibulocerebellum, 223–224 zebrin staining, 226, 228–229 serotonergic projections, 1209 spinal pathways ascending pathways dorsal spinocerebellar tract, 152 spinoolivary tract, 152 ventral spinocerebellar tract, 152 descending pathways, 157 tachykinin projections, 1242–1243 termination of mossy fiber systems lateral reticular nucleus, 233 pons, 234 spinal cord, 231–233 trigeminal nucleus, 233–234 vestibular nucleus, 234–235 Cerebral cortex auditory system, see Auditory cortex autonomic control infralimbic cortex, 785 insular cortex, 784–785 motor cortex, 786 perirhinal cortex, 785 prelimbic cortex, 785 somatosensory cortex, 786 basilar pontine nuclei connections, 168–169 descending spinal cord pathways, 155–156 lateral migratory stream Cajal–Retzius neuron migration, 66 layer VI–VII neuron migration stages, 66–67 piriform cortex, 67 subplate neuron migration, 66 locus coeruleus connections, 278 serotonergic projections, 1210 somatosensory system, see Somatosensory cortex striatum connections corticostriatal neuron subtypes, 458, 460 medium spiny projection neuron connections, 465 organization of corticostriatal afferents convergent corticostriatal organization, 462–463 general organization, 463 quantitative data, 463–464 topographical organization, 460–462 overview, 498 patch/matrix compartment connections, 494–497 tachykinin projections, 1244 visual system, see Visual cortex CGRP, see Calcitonin gene-related peptide ChAT, see Choline acetyltransferase Choline acetyltransferase (ChAT) amygdalohippocampal transition area staining, 543 dorsal horn neurochemistry, 140 intraamygdaloid bed nucleus of the stria terminalis staining, 560

Choline acetyltransferase (ChAT) Continued laterobasal nuclear complex staining basolateral amygdaloid nucleus, 584 basomedial amygdaloid nucleus, 586 lateral amygadaloid nucleus, 583 ventral basolateral amygdaloid nucleus, 585 localization, 1258 olfactory amygdala staining anterior aygdaloid area, 517 anterior cortical amygdaloid nucleus, 524 mygdalopiriform transition area, 534 nucleus of the lateral olfactory tract, 520–521 posterolateral cortical amygdaloid nucleus, 527 Choroid plexus cerebrospinal fluid synthesis, 400–401 innervation, 401 Ciliac plexus, anatomy, 81–82 Ciliary ganglion, neurons, 94 Cingulate cortex area 24b in movement, vision, and pain behaviors, 712–713 area 25 cytology, 707 infra limbic area IL, 707 areas, 752 Brodmann nomenclature, 707–709 cortical connections of retrosplenial cortex and visuospatial function, 713–714 cytology midcingulate cortex, 709–710 perigenual anterior cingulate cortex, 709–710 retrosplenial cortex, 710–711 divisions, 705–707 functional overview, 751–752 layers, 752–753 local cerebral glucose utilization studies, 752 N-methyl-D-aspartate receptor antagonist-induced neurotoxicity in retrosplenial cortex Alzheimer's disease neurodegeneration relevance, 721–724 area 30 deafferentiation, 718–719 overview, 716 pathomorphological response, 717–718 polysynaptic circuit disinhibition adrenergic system, 719 cholinergic system, 719 glutamatergic system, 719–720 markers, 721 metabolic derangements, 720 psychosis induction, 721 neurotransmitter receptors, 752–753 opioid receptors, 711–712 pathology Alzheimer's disease, 705, 721–724 schizophrenia, 705

1297

INDEX

Cingulate cortex Continued primate comparison to rat medial cortex, 722–723 thalamic afferents area 29, 715 axon terminal morphology and multiple heteroreceptor regulation, 715–716 regional differentiation, 714–715 Circumventricular organs (CVOs), see also Area prostrema; Choroid plexus; Median eminence; Pineal gland; Subcommissural organ; Subfornical organ; Vascular organ of the lamina terminalis blood–brain barrier permeability, 389 ependymal cells, 389 CL, see Central lateral nucleus Claustrocortex, see Agranular insular cortex CM, see Central medial nucleus CNC, see Cochlear nuclear complex Cochlea anatomy, 998 basilar membrane, 1000 Deiter's cells, 1002 inner hair cells, 1000, 1002 olivocochlear system, 1062–1063 organ of Corti nerve fiber types, 1003 outer hair cells, 1000, 1002 pillar cells, 1002 sensory transduction, 1002–1003 vestibular membrane, 998, 1000 Cochlear nuclear complex (CNC) afferents, 1007, 1009 anatomy, 1003, 1007 ascending projections, 1012–1013 cochlear root neurons, 1010–1011 dorsal cochlear nucleus functional significance, 1012 granule cell system, 1011 pyramidal cells, 1011 tuberculoventral system, 1011–1012 small cells, 1010–1011 ventral cochlear nucleus cytoarchitecture, 1009 globular bushy neuron, 1009 multipolar cells D-stellate cells, 1010 T-stellate cells, 1010 octopus neuron, 1009–1010 spherical bushy neuron, 1009 Conditioned taste aversion (CTA), circuitry, 911–912 Corticotropin-releasing hormone (CRH), locus coeruleus inputs, 273 CREB, see Cyclic AMP response elementbinding protein CRH, see Corticotropin-releasing hormone CST, see Nucleus of the commissural component of the stria terminalis CTA, see Conditioned taste aversion CVOs, see Circumventricular organs Cyclic AMP response element-binding protein (CREB), hypothalamic integration, 359–360

D DB, see Nucleus of the diagonal band Dentate gyrus connections basket cell projections, 643–644 brain stem, 647–648 granule cell projections, 643 ipsilateral associational/commissural projection, 644–645 mossy fiber intrahippocampal connections, 648–649 septum, 645–647 supramammillary area, 647 cytoarchitectonics, 639 granule cell layer, 639, 641 molecular layer neurons, 641, 643 neuron migration, 68 neuron origins, 65 polymorphic cell layer, 643 Diencephalon components, 407 periaqueductal gray connections, 247 spinal pathways ascending pathways, 153–154 descending pathways, 156 DLG, see Dorsal lateral geniculate nucleus DMH, see Dorsomedial hypothalamic nucleus Dopamine nigrostriatal dopamine system dorsal tier dopamine neurons, 489–490 input to pars comapcta neurons, 490 overview, 488–489 ventral tier dopamine neurons, 490 striatum medium spiny projection neuron receptor subtypes and mediation of gene regulation, 475–477 vestibular neuron modulation, 982–984 Dorsal horn interneurons excitatory circuits, 860 excitatory neurotransmitters aspartate, 858 glutamate, 858 peptides, 858, 860 inhibitory circuits, 863–864 inhibitory neurotransmitters amino acids, 861–862 peptides, 862–863 Dorsal lateral geniculate nucleus (DLG) axons, 1090–1091 connections afferents cerebral cortex, 1097–1098 retina, 1097 subcortical projections, 1098–1100 efferents, 1100 table, 1096 fiber proteins, 411–412 neuron types, 1091 phyisology and function, 1094–1095 projections, 412 regional organization, 1092 relay neurons, 1010–1101 structure, 411 synaptic organization, 1092

Dorsal premammillary nucleus (PMD), functions, 347–348 Dorsomedial hypothalamic nucleus (DMH), functions, 345 Dynorphin, dorsal horn neurochemistry, 139

E Edinger–Westphal nucleus, basilar pontine nuclei connections, 171 EF, see Epifasicular nucleus Emotion, periaqueductal gray coping circuits anatomy, 250 escapable versus inescapable stressors, 253 immediate early gene expression studies, 250–251, 253 Endomorphin, dorsal horn neurochemistry, 139 Enkephalin dorsal horn neurochemistry, 137, 139 locus coeruleus inputs, 273 Entorhinal cortex connections associational projections, 681 commissural projections, 681 dentate gyrus, 677–678 extrinsic connections, 682–686 hippocampal formation, 677 hippocampus CA1, 679 CA2, 679 CA3, 679 crossed connections, 681 topography of pathways, 680–681 parahippocampus, 681–682 subiculum, 679–680 gestational time of origin, 34 layers, 672, 674 neuron origins, 64 neuron types, 674–676 subdivisions, 676–677 Ependymal stem cell, dynamics in cortical germinal zones, 68, 71–72 Epifasicular nucleus (EF), locus coeruleus connections, 266, 271, 285 Extended amygdala central division central division of the supracapsular bed nucleus of the stria terminalis acetylcholinesterase staining, 567 cytoarchitectonics, 566 heavy metal staining, 566–567 topographic landmarks, 566 central sublenticular extended amygdala cytoarchitectonics, 566 heavy metal staining, 566 topographic landmarks, 566 connections, 577–582 interstitial nucleus of the posterior limb of the anterior commissure acetylcholinesterase staining, 568

1298 Extended amygdala Continued cytoarchitectonics, 567–568 heavy metal staining, 568 topographic landmarks, 567–568 lateral bed nucleus of the stria terminalis acetylcholinesterase staining, 564 cytoarchitectonics, 562–563 heavy metal staining, 563 topographic landmarks, 562 lateral part of the central amygdaloid nucleus acetylcholinesterase staining, 565–566 cytoarchitectonics, 565 fibroarchitectonics, 565 heavy metal staining, 565 topographic landmarks, 565 medial part of the central amygdaloid nucleus cytoarchitectonics, 564 fibroarchitectonics, 564 heavy metal staining, 564–565 topographic landmarks, 564 neurotransmitters and neuropeptides cells, 568–570 fibers, 570–572 divisions, overview, 552 history of study, 547, 551 intercalated cell masses acetylcholinesterase staining, 591 cytoarchitectonics, 589 fibroarchitectonics, 589–590 heavy metal staining, 59 topographic landmarks, 588–589 medial division connections, 572–576 intraamygdaloid bed nucleus of the stria terminalis acetylcholinesterase staining, 560 choline acetyltransferase staining, 560 cytoarchitectonics, 560 fibroarchitectonics, 560 heavy metal staining, 560 topographic landmarks, 560 medial amygdaloid nucleus acetylcholinesterase staining, 559–560 choline acetyltransferase staining, 560 cytoarchitectonics, 559 fibroarchitectonics, 559 heavy metal staining, 559 topographic landmarks, 558–559 medial bed nucleus of the stria terminalis acetylcholinesterase staining, 558 cytoarchitectonics, 557–558 fibroarchitectonics, 558 heavy metal staining, 558 topographic landmarks, 557 medial division of the supracapsular bed nucleus of the stria terminalis cytoarchitectonics, 562 topographic landmarks, 561–562

INDEX

Extended amygdala Continued medial sublenticular extended amygdala acetylcholinesterase staining, 561 cytoarchitectonics, 561 fibroarchitectonics, 561 heavy metal staining, 561 topographic landmarks, 560–561 neurotransmitters and neuropeptides cells, 568–570 fibers, 570–572 Eye, see Retina; Visual system

F Facial nucleus afferents medulla, 304–305 midbrain pathways, 303–304 pons, 304 cytoarchitectonics, 303 dendritic architecture, 303 myotopic organization, 301–303 Frontal cortex areas, 730, 732 cytoarchitectonics, 730 layers, 730, 733, 738 local cerebral glucose utilization studies, 734 neurotransmitter receptors, 734, 738 Fu, see Fusiform nucleus Fusiform nucleus (Fu), features, 593

G GABA, see γ-Aminobutyric acid Glial stem cell, dynamics in cortical germinal zones, 68, 71–72 Globus pallidus lateral globus pallidus in indirect striatal output morphology, 479–480 neuron types, 480 neurotransmission, 480 output, 481 synaptic input, 480–481 medial globus pallidus in basal ganglia output, 482 Glutaminergic pathways auditory pathways, 1275–1276 basal ganglia, 1278–1280 cerebellum, 1280, 1282 detection, 1269–1270, 1283 dorsal horn neurochemistry, 139–140 motor pathways, 1278 neocortex, 1271–1272 olfactory pathways, 1276–1277 somatosensory pathways, 1272–1274 vesicular glutamate transporters, 1270–1273 visual pathways, 1274–1275 vomeronasal system, 1277–1278 Glycine dorsal horn neurochemistry, 140 motor trigeminal nucleus transmission in medulla, 301

Golgi tendon organ, somatosensory receptor, 799 Gustatory system conditioned taste aversion circuitry, 911–912 nucleus of the solitary tract connections, 893–896 cytoarchitecture, 905–907 neurochemistry, 908 parabrachial nuclei connections, 896–900 cytoarchitecture, 907 neurochemistry, 908–909 preference–aversion behavior circuitry, 910 reciprocal projections between taste relays, 903 salt appetite circuitry, 912–913 somatosensory cortex connections, 902–903 neurochemistry, 909 taste buds distribution, 892–893 neurochemistry, 907–908 thalamus neurochemistry, 909 relay nuclei, 900, 902 trigeminal innervation, 822, 893

H Hippocampal region, see also Dentate gyrus; Entorhinal cortex; Parasubiculum; Perirhinal cortex; Postrhinal cortex; Presubiculum; Subiculum components, 33–34 definition, 636–637 fiber bundles, 637, 639 gestational time of origin entorhinal cortex, 34 hippocampus, 35 subiculum, 34–35 history of study, 635–636 position in brain, 637 Hippocampus cholinergic projections, 1262–1263 connections CA1 associational connections, 656 commissural projections, 656 extrinsic connections, 656–657, 660 intrahippocampal connections, 656 CA2, 655–656 CA3 associational connections, 654 commissural projections, 654 extrinsic connections, 654–655 intrahippocampal connections, 653–654 fields, 649 gestational time of origin, 35 information flow lamellar concept, 689–690 septotemporal topography of perforant path projections, functional implications, 690–692

1299

INDEX

Hippocampus Continued transverse topography and functional implications, 692–693 trisynaptic circuit and serial/parallel information processing, 690 laminar organization, 649 locus coeruleus connections, 278–279 neuron types and local connections, 649–650, 652 serotonergic projections, 1210 tachykinin projections, 1244 Histamine, vestibular neuron modulation, 981–982, 984 Hypocretin/orexin, locus coeruleus inputs, 271 Hypoglossal nucleus afferents forebrain pathways, 308 lingual proprioceptors, 311 medulla dorsal medulla, 309 reticular formation, 309–310 ventral medulla, 309–310 midbrain projections, 308–309 neuropeptide input, 311 raphe nuclei, 310 trigeminal and solitary nucleus input, 310–311 cytoarchitectonics, 307 dendritic architecture, 307–308 interneurons, 307 motoneurons, 307 myotopic organization, 305–307 Hypothalamus accessory magnocellular neurosecretory neurons, 382 autonomic control anteroventral third ventricular area, 778 arcuate nucleus, 782 dorsal hypothalamic area, 780 dorsomedial nucleus, 780 lateral hypothalamic area, 780, 782 paraventricular nucleus, 778–780 posterior lateral hypothalamic area, 782 retrochiasmatic area, 782 tuberomammillary nucleus, 782 basilar pontine nuclei connections, 171 functional overview, 335, 352 gross anatomy, 336 integrative mechanisms cellular level, 357–358 molecular level, 358–360 neural systems level, 352–354 nuclei level, 354–357 locus coeruleus connections, 280 magnocellular neurosecretory system, 369 morphological organization areas and nuclei, 337 lateral zone lateral hypothalamic area, 351–352 lateral preoptic area, 350–351 overview, 349 medial zone anterior region, 345 mammillary region, 347–349

Hypothalamus Continued overview, 340–342 preoptic region, 342–345 tuberal region, 345–347 periventricular zone anterior region, 339–340 cytoarchitectonics, 336 mammillary region, 340 preoptic region, 338–339 tuberal region, 340 zones and regions, 337 olfactory cortex connections, 946 parvocellular neurosecretory system, 369 serotonergic projections, 1210

I IC, see Inferior colliculus IGL, see Intergeniculate leaflet IMD, see Intermediodorsal nucleus IMG, see Intramedullary gray Inferior colliculus (IC) auditory function, 1029 central nucleus afferent projections, 1035–1036 efferent projections, 1036–1037 electrophysiology, 1034–1035 neuron types, 1031–1032 colliculofugal pathways, 1061–1062 commissural connections, 1039–1040 components, 1029, 1031 dorsal cortex, 1039 external cortex, 1037, 1039 intrinsic connections, 1039–1040 neurochemistry and functional significance, 1040–1041 Inferior mesenteric plexus, anatomy, 81–82 Inferior olivary nucleus afferents, 182–184 cytoarchitecture, 180–182 efferents, 184–185 functions, 185–187 Insular cortex, autonomic control, 784–785 Intergeniculate leaflet (IGL) functions, 413–414 projections, 413 structure, 413 Intermediodorsal nucleus (IMD) afferents, 435 efferents, 437 structure, 434 Internal carotid ganglion, anatomy, 83 Internal carotod artery, anatomy, 1171 Internal jugular vein, anatomy, 1181 Internal ophthalmic artery, anatomy, 1179 Interneurons hypoglossal nucleus, 307 paraventricular nucleus, 379–380 striatum abundance, 471 classification calretinin/γ-aminobutyric acid interneurons, 474 large aspiny cholinergic neurons, 473

Interneurons Continued medium aspiny GABAergic neurons, 473 overview, 471 parvalbumin interneurons, 473 somatostatin/neuropeptide Y interneurons, 473–474 supraoptic nucleus, 372–373 trigeminal nucleus, 297 Interstitial nucleus of the posterior limb of the anterior commissure (IPAC) acetylcholinesterase staining, 568 cytoarchitectonics, 567–568 heavy metal staining, 568 topographic landmarks, 567–568 Intraamygdaloid bed nucleus of the stria terminalis (BSTIA) acetylcholinesterase staining, 560 choline acetyltransferase staining, 560 cytoarchitectonics, 560 fibroarchitectonics, 560 heavy metal staining, 560 topographic landmarks, 560 Intramedullary gray (IMG) acetylcholinesterase staining, 592 cytoarchitectonics, 591 heavy metal staining, 591–592 topographic landmarks, 591 Intramural ganglia glial processes, 97 myenteric ganglia, 94–95 structure, 94–97 IPAC, see Interstitial nucleus of the posterior limb of the anterior commissure Isocortex, see Frontal cortex; Occipital cortex; Parietal cortex; Temporal cortex

L La, see Lateral amygadaloid nucleus Lanceolate ending, somatosensory receptor, 799 Lateral amygadaloid nucleus (La) acetylcholinesterase staining, 583 choline acetyltransferase staining, 583 cytoarchitectonics, 573, 583 fibroarchitectonics, 583 heavy metal staining, 583 topographic landmarks, 573 Lateral bed nucleus of the stria terminalis (BSTL) acetylcholinesterase staining, 564 cytoarchitectonics, 562–563 heavy metal staining, 563 topographic landmarks, 562 Lateral cervical nucleus, cytoarchitecture, 126 Lateral geniculate nucleus, see Dorsal lateral geniculate nucleus; Ventral lateral geniculate nucleus Lateral part of the central amygdaloid nucleus (CeL) acetylcholinesterase staining, 565–566 cytoarchitectonics, 565 fibroarchitectonics, 565

1300 Lateral part of the central amygdaloid nucleus (CeL) Continued heavy metal staining, 565 topographic landmarks, 565 Lateral posterior nucleus (LP) connections, 1120, 1122 cytoarchitectonics, 1122 stimulus response studies, 1122–1123 ultrastructure, 1122 Lateral reticular nucleus (LRt) afferents cerebellum, 177–178 red nucleus, 177 spinal cord, 176–177 cerebellar termination of mossy fiber systems, 233 cytoarchitecture, 174, 176 efferents cerebellar cortex, 178 spinal cord, 178–179 functions, 179–180 localization, 174 Lateral spinal nucleus afferent fibers, 113 cytoarchitecture, 126 Lateral thalamic nuclei connections, 432–433 functions, 433 subdivisions, 432 types, 432 Laterobasal nuclear complex (LBNC) basolateral amygdaloid nucleus acetylcholinesterase staining, 584 choline acetyltransferase staining, 584 cytoarchitectonics, 583–584 fibroarchitectonics, 584 heavy metal staining, 584 topographic landmarks, 583 basomedial amygdaloid nucleus acetylcholinesterase staining, 585–586 choline acetyltransferase staining, 586 cytoarchitectonics, 585 fibroarchitectonics, 585 heavy metal staining, 585 topographic landmarks, 585 connections, 588–590 divisions, 572–573 hodological relationships, 586–587 lateral amygadaloid nucleus acetylcholinesterase staining, 583 choline acetyltransferase staining, 583 cytoarchitectonics, 573, 583 fibroarchitectonics, 583 heavy metal staining, 583 topographic landmarks, 573 neurotransmitters and neuropeptides, 587 ventral basolateral amygdaloid nucleus acetylcholinesterase staining, 585 choline acetyltransferase staining, 585 cytoarchitectonics, 584 fibroarchitectonics, 584–585 heavy metal staining, 585 topographic landmarks, 584 LBNC, see Laterobasal nuclear complex

INDEX

LC, see Locus coeruleus Limbic cortex gestational time of origin, 32 neuron origins, 63–64 Locus coeruleus (LC) afferents indirect afferents, 268 microphysiology studies, 267–269 neurotransmitter inputs adrenergic input, 271–272 γ-aminobutyric acid, 271–273 corticotropin-releasing hormone, 273 double-labeling studies, 271–273 enkephalin, 273 epifasicular nucleus, 271 excitatory amino acids, 270–271 hypocretin/orexin, 271 nucleus paragigantocellularis lateralis, 270–271 overview, 263–265 serotonin, 272 ultrastructural studies, 273–274 retrograde and anterograde tract tracing, 265–269 analgesia system, 869–870 autonomic control, 775 basilar pontine nuclei connections, 171 cytoarchitecture cell types and subnuclei, 259, 261 electrotonic coupling, 262–263 neurotransmitters, 261–262 efferents ascending projections amygdala, 280 bed nucleus of the stria terminalis, 279–280 cerebral cortex, 278 hippocampus, 278–279 hypothalamus, 280 neocortex, 278 olfactory bulb, 277–278 olfactory cortex, 277–278 preoptic area, 280 striatum, 279 thalamus, 279 descending projections cerebellum, 280–281 medulla, 281 pons, 281 spinal cord, 281 glia, 274 olfactory bulb inputs and modulation, 952–953 pericoerulear region, see Pericoerulear region projection properties antidromic activation of neurons from multiple sites, 282–283 collateralization of efferent neurons, 281–282 retrograde labeling of neurons from multiple sites, 282 topography of neurons, 283 ultrastructure of terminals, 283–284

Longitudinal hippocampal artery, anatomy, 1175 LOT, see Nucleus of the lateral olfactory tract LP, see Lateral posterior nucleus LPGi, see Nucleus paragigantocellularis lateralis LRt, see Lateral reticular nucleus

M Magnocellular preoptic nucleus (MCPO), functions, 350–351 Main olfactory bulb (MOB) afferents, 937 efferents intrabulbar collaterals, 936–937 mitral/tufted cell axons, 937 olfactory cortex, 937, 945 layers external plexiform layer, 933–934 glomerular layer neurons, 930–933 neurotransmitters and peptides, 933 granule cell layer, 935–936 internal plexiform layer, 935 mitral cell layer, 934–935 olfactory nerve layer, 928, 930 subependymal layer, 936 locus coeruleus connections, 277–278 modulatory inputs differential innervation, 953 locus coeruleus, 952–953 nucleus of the diagonal band, 952 raphe nuclei, 952–953 neuron migration, 67–68 neuron origins, 65 odor memory circuitry, 948 olfactory nerve regulation of neurotransmitters in bulb neurons, 936 MCPO, see Magnocellular preoptic nucleus MD, see Mediodorsal nucleus Me, see Medial amygdaloid nucleus Medial amygdaloid nucleus (Me) acetylcholinesterase staining, 559–560 choline acetyltransferase staining, 560 cytoarchitectonics, 559 fibroarchitectonics, 559 heavy metal staining, 559 topographic landmarks, 558–559 Medial bed nucleus of the stria terminalis (BSTM) acetylcholinesterase staining, 558 cytoarchitectonics, 557–558 fibroarchitectonics, 558 heavy metal staining, 558 topographic landmarks, 557 Medial division of the supracapsular bed nucleus of the stria terminalis (BSTSm) cytoarchitectonics, 562 topographic landmarks, 561–562 Medial geniculate body (MG) auditory processing, 1041–1042, 1050–1051 components, 1042

1301

INDEX

Medial geniculate body (MG) Continued dorsal division, 1045, 1048 functions, 422 medial division, 1048–1049 posterior paralaminar thalamic nuclei, 1050 projections, 421–422 structure, 420–421 thalamic reticular nucleus auditory sector, 1049–1050 ventral division, 1042, 1045 Medial habenula, cholinergic projections, 1262 Medial part of the central amygdaloid nucleus (CeM) cytoarchitectonics, 564 fibroarchitectonics, 564 heavy metal staining, 564–565 topographic landmarks, 564 Medial preoptic nucleus (MPO), functions, 343–345, 354–356 Medial sublenticular extended amygdala (SLEAm) acetylcholinesterase staining, 561 cytoarchitectonics, 561 fibroarchitectonics, 561 heavy metal staining, 561 topographic landmarks, 560–561 Medial vestibular nucleus anatomy, 970–971 second-order vestibular neurons calcium currents, 974–975 electrophysiological identification, 974 gap junctions, 976 postnatal maturation, 976 potassium currents, 975 rhythmic activities, 975–976 sodium currents, 974 types and functions, 976–977 Median eminence, structure and function, 398–399 Median preoptic nucleus (MnPO), functions, 338 Mediodorsal nucleus (MD) functions, 428–429 projections, 425–428 subdivisions, 425 Medium spiny projection neuron, see Striatum Medulla autonomic control caudal ventrolateral medulla, 767 raphe nuclei, 767 ventromedial medulla, 767 facial nucleus connections, 304–305 hypoglossal nucleus connections dorsal medulla, 309 reticular formation, 309–310 ventral medulla, 309–310 locus coeruleus connections, 281 periaqueductal gray connections, 247–249 serotonergic projections, 1210 somatosensory relay nuclei afferents, 802–803

Medulla Continued cytoarchitecture, 801 efferents, 803–804 plasticity, 801–802 somatotopic organization, 801 spinal pathways ascending pathways direct dorsal column pathway, 149–150 nuclei afferents, 151 postsynaptic dorsal column pathway, 150 spinocervical tract, 151 spinoreticular tracts, 150–151 descending pathways proproispinal connections, 158–159 raphe nuclei, 157–158 reticular formation, 157 trigeminal nucleus, 157 vestibular nuclei, 158 trigeminal nucleus connections, 300–301 Merkel ending, somatosensory receptor, 797 N-Methyl-D-aspartate (NMDA) receptor antagonist-induced neurotoxicity in retrosplenial cortex Alzheimer's disease neurodegeneration relevance, 721–724 area 30 deafferentiation, 718–719 overview, 716 pathomorphological response, 717–718 polysynaptic circuit disinhibition adrenergic system, 719 cholinergic system, 719 glutamatergic system, 719–720 markers, 721 metabolic derangements, 720 psychosis induction, 721 vestibular neuron modulation, 978–979 MG, see Medial geniculate body Micturition brain stem–spinal cord pathways in coordination abdominal pressure control systems, 323 diffuse descending systems, 323 overview, 323–324 human functional neuroimaging studies, 326–327 motoneurons bladder, 321–322 pelvic floor, 322–323 periaqueductal gray in control, 325–327 peripheral afferent nerves, 324–325 physiology, 321 sacral cord reflexes, 323 spinal cord–brain stem pathways in micturition reflex, 325 Midbrain reticular formation, ascending spinal cord pathways, 153 Middle cerebral artery, anatomy, 1179 MnPO, see Median preoptic nucleus MOB, see Main olfactory bulb Molecular specification Bauplan, 7–11 differential aspects of histoogenesis, 6–7

Molecular specification Continued mechanisms, 20–21 neural plate subdivisions, 11–13 neural tube alar plate, 16–17 basal plate, 14–16 overview, 13–14 sharing of molecularly distinct brain domains among vertebrates, 5 telencephalon, 17–20 Motoneurons bladder, 321–322 hypoglossal nucleus, 307 pelvic floor, 322–323 MPO, see Medial preoptic nucleus Muscle spindle, somatosensory receptor, 799

N Neocortex, see also Isocortex gestational time of origin, 32 glutaminergic pathways, 1271–1272 locus coeruleus connections, 278 neuron origins, 63 Nerve growth factor (NGF), vestibular neuron modulation, 985 Neural plate, gene expression in subdivisions, 11–12 Neural stem cell dynamics in cortical germinal zones, 68, 71 maps of mosaics in telencephalic neuroepithelium over time, 38–61 primary cells, 68 secondary cells, 68, 71 Neural tube, gene expression in development alar plate, 16–17 basal plate, 14–16 overview, 13–14 Neuroepithelium, neuron origins and gene expression, 37 Neurokinins amygdala projections, 1244 basal ganglia projections, 1244 brain stem projections, 1242–1243 cerebellum projections, 1242–1243 cerebral cortex projections, 1244 dorsal horn neurochemistry, 136–137 functions, 1245–1246 genes, 1214 hippocampus projections, 1244 immunohistochemical staining, 1214 neuron development, 1240 receptor types and distribution, 1245 septum projections, 1244 serotonin and tachykinin coexistence cell body locations, 1246 functional interactions, 1247 pathways, 1246–1247 spinal cord projections, 1242 thalamus projections, 1243–1244 vestibular neuron modulation, 985 Neuromuscular junction, ganglion neurons, 97–99

1302 Neuron migration dentate migratory stream in dentate gyrus, 68 lateral migratory stream in cerebral cortex Cajal–Retzius neuron migration, 66 layer VI–VII neuron migration stages, 66–67 piriform cortex, 67 subplate neuron migration, 66 rostral migratory stream in olfactory bulb, 67–68 Neurulation, see also Bauplan; Molecular specification differential aspects of histoogenesis, 6–7 fate mapping, 4, 8 field homology, 5 molecular specification state, 4 planar neural patterning, 4 vertical patterning, 4 NGF, see Nerve growth factor Nitric oxide (NO), vascular innervation, 1195 NLL, see Nuclei of the lateral lemniscus NMDA receptor, see N-Methyl-D-aspartate receptor NO, see Nitric oxide Nociception ascending pathways postsynaptic dorsal column pathway, 865–866 spinocervical pathway, 867 spinohypothalamic tract, 867–868 spinomesecephalic tract, 866–867 spinoreticular tract, 867 spinothalamic tract, 864–865 brain stem analgesia system locus coeruleus, 869–870 periaqueductal gray, 868–869 raphe nuclei, 869 dorsal horn interneurons excitatory circuits, 860 excitatory neurotransmitters aspartate, 858 glutamate, 858 peptides, 858, 860 inhibitory circuits, 863–864 inhibitory neurotransmitters amino acids, 861–862 peptides, 862–863 nociceptors, 855–856, 858 overview of pathways, 853–854 plasticity in pathological conditions inflammatory pain, 871–872 mechanisms, 870–871 neuropathic pain dorsal horn plasticity, 872 neuroanatomical plasticity, 872–873 neurochemical plasticity, 873–875 neurophysiological plasticity, 873 somatosensory cortex nociceptive neurons, 868 thalamus nociceptive neurons, 868 Nodose ganglion, anatomy, 83–84 Norepinephrine locus coeruleus neurotransmission, 261–262, 271, 284

INDEX

Norepinephrine Continued vestibular neuron modulation, 983–984, 1194 NTS, see Nucleus of the solitary tract Nuclei of the lateral lemniscus (NLL) cytoarchitectonic schemes, 1020–1021 dorsal nucleus binaural input, 1025 connections, 1028 functional significance, 1029 neurochemistry, 1028 neuron characterization, 1025 organization, 1025, 1028 synaptic responses, 1028–1029 ventral complex connections, 1021–1022 monoaural input, 1021 neurochemistry, 1022, 1025 neuron characterization, 1021 Nucleus accumbens, neuron origins, 62 Nucleus of the commissural component of the stria terminalis (CST) acetylcholinesterase staining, 593 cytoarchitectonics, 592 heavy metal staining, 592 topographic landmarks, 592 Nucleus of Darschewitsch, basilar pontine nuclei connections, 171 Nucleus of the diagonal band (DB), olfactory bulb inputs and modulation, 952 Nucleus of the lateral olfactory tract (LOT) acetylcholinesterase staining, 520 choline acetyltransferase staining, 520–521 cytoarchitectonics, 517 fibroarchitectonics, 517, 520 heavy metal staining, 520 topographic landmarks, 517 Nucleus paragigantocellularis lateralis (LPGi), locus coeruleus connections, 266, 270–271, 285 Nucleus reuniens thalami (Re) afferents, 435 efferents, 438 structure, 434 Nucleus of the solitary tract (NTS) autonomic control, 763, 765 gustatory system connections, 893–896 cytoarchitecture, 905–907 neurochemistry, 908 projections, 763, 765 Nucleus submedius (Sub) connections, 429 functions, 429–320 structure, 429

O Occipital cortex, see also Visual cortex areas, 748 cytoarchitectonics, 747–748 local cerebral glucose utilization studies, 748 neurotransmitter receptors, 748–749 Olfactory artery, anatomy, 1180

Olfactory bulbs, see Accessory olfactory bulb; Main olfactory bulb Olfactory cortex anterior olfactory nucleus architecture, 937–938 inputs, 938 neurotransmitters, 942 outputs, 938–939, 942 connections associative connections, 946 extrinsic outputs, 946 intrinsic connections, 944–946 olfactory bulb, 937, 945 glutaminergic pathways, 1276–1277 integration of taste and visceral response, 947–948 lateral olfactory cortex architecture, 944–945 locus coeruleus connections, 277–278 medial olfactory cortex anterior hippocampal continuation, 942 indusium griseum, 942 infralimbic cortex, 942 olfactory tubercle, 942–943 taenia tecta, 942 modulatory inputs, 953 motor activity linkage, 948 odor cognition circuitry, 946–947 Olfactory epithelium odorant receptor genes, 926 olfactory bulb connections, 927 olfactory receptor neurons, 923–924, 926–927 organization, 924, 926 signal transduction, 926–927 Olfactory peduncle, neuron origins, 65 Olfactory tubercle, neuron origins, 62 Olive, cerebellum projections, 219–222 Opioid receptors cingulate cortex, 711–712 dorsal horn neurochemistry, 139 striatum medium spiny projection neurons, 478 vestibular neuron modulation, 984–985 Optic ganglion, anatomy, 83 Optic nerve anatomy, 1089 neurotransmission, 1089 Orbitofrontal cortex areas, 749 local cerebral glucose utilization studies, 749 neurotransmitter receptors, 749–750 topography, 749 Orexin, see Hypocretin/orexin Oromotor nuclei, see Facial nucleus; Hypoglossal nucleus; Trigeminal nucleus OVLT, see Vascular organ of the lamina terminalis

P p75 neurotrophin receptor, Alzheimer's disease role, 1265–1266

INDEX

Pa, see Paraventricular nucleus Pacinian corpuscle, somatosensory receptor, 798 PAG, see Periaqueductal gray Pain, see Nociception Pallidum gestational time of origin, 28–29 neuron origins, 37, 62 Parabigeminal nucleus connections, 1121 neurotransmission, 1121–1122 Parabrachial nuclei autonomic control, 768, 771–772 gustatory system connections, 896–900 cytoarchitecture, 907 neurochemistry, 908–909 spinal afferents, 151 Paracentral nucleus (PC) afferents, 436 efferents, 437 functions, 440 structure, 435–436 Parafascicular nucleus (PF) afferents, 436–437 efferents, 437 functions, 441 structure, 436 Parastrial nucleus (PS) features, 592 functions, 344 Parasubiculum anatomy, 661 connections associational projections, 671 commissural projections, 671 extrinsic connections, 669–672 hippocampus, 671 parahippocampus, 671 cytoarchitectonics, 671 neuron types, 671 subicular complex concept, 660–662 Parataenial nucleus (PT) afferents, 435 efferents, 438 structure, 434 Paraventricular nucleus (Pa) afferents brain stem, 380 forebrain, 380–381 hypothalamus, 381 functions, 339–340, 354 interneurons, 379–380 magnocellular neurosecretory component magnocellular neuron morphology and efferent path, 377 principal neuron distribution, 375, 377 nonendocrine projections morphology and efferent path, 378–379 principal neuron distribution, 378 parvocellular neurosecretory component morphology and efferent path, 377–378 principal neuron distribution, 377

Paraventricular nucleus (PV) afferents, 434 efferents, 437 functions, 439–440 structure, 434 Parietal cortex anterior parietal cortex areas, 738–739 cytoarchitectonics, 739 layers, 741 local cerebral glucose utilization studies, 739 neurotransmitter receptors, 739–741 topography, 738 posterior parietal cortex areas, 743 layers, 743 neurotransmitter receptors, 743 ventral parietal cortex layers, 742–743 neurotransmitter receptors, 741–743 nomenclature, 741 Parvicellular ventral posterior nucleus (VPPC) connections, 420 functions, 420 structure, 420 Patch/matrix compartments, see Striatum PC, see Paracentral nucleus PeA, see Anterior periventricular nucleus Pelvic ganglion neurotransmitters, 91–93 preganglionic inputs, 93–94 pricipal neurons, 91 Pelvic plexus, anatomy, 82–83 PeP, see Posterior periventricular nucleus Periaqueductal gray (PAG) afferents medulla, 247–249 prefrontal cortex, 249 spinal cord, 247–249 analgesia system, 868–869 autonomic control, 770 columnar organization functional studies, 244–245 neurochemical studies, 245–247 cytoarchitecture, 243–244 efferents brain stem, 247 diencephalon, 247 emotional coping circuits anatomy, 250 escapable versus inescapable stressors, 253 immediate early gene expression studies, 250–251, 253 micturition control, 325–327 spinal pathways ascending pathways, 153 descending pathways, 156 Pericoerulear region architecture, 275 extranuclear dendrites, 275, 277 inputs, 275 locus coeruleus connections, 277

1303 Perirhinal cortex autonomic control, 785 connections associational projections, 688 commissural projections, 688 extrinsic connections, 688–689 hippocampal formation, 687–688 parahippocampus, 687–688 cytoarchitectonics, 751 layers, 686–687, 751 neuron types, 687 neurotransmitter receptors, 751 topology, 686 PF, see Parafascicular nucleus PFC, see Prefrontal cortex Pial arterial network, anatomy, 1180–1181 Pineal gland cell types, 400 structure, 400 Piriform cortex gestational time of origin, 32–33 neuron migration, 67 neuron origins, 64 Pituitary gland gross anatomy, 369–370 hypothalamic connections, 370 median eminence as circumventricular organ, 398–399 Plasticity auditory cortex, 1058 medulla somatosensory relay nuclei, 801–802 nociception in pathological conditions inflammatory pain, 871–872 mechanisms, 870–871 neuropathic pain dorsal horn plasticity, 872 neuroanatomical plasticity, 872–873 neurochemical plasticity, 873–875 neurophysiological plasticity, 873 somatosensory cortex, 806–807 thalamus somatosensory system, 804 trigeminal sensory system, 801–802, 804, 838 PLCO, see Posterolateral cortical amygdaloid nucleus PMCO, see Posteromedial cortical amygdaloid nucleus PMD, see Dorsal premammillary nucleus PMV, see Ventral premammillary nucleus Po, see Posterior thalamic nucleus Pons cerebellar termination of mossy fiber systems, 234 facial nucleus connections, 304 locus coeruleus connections, 281 molecular specification, 5 nuclei, see Basilar pontine nuclei; Reticulotegmental nucleus of the pons spinal pathways ascending pathways, 151–152 descending pathways, 156–157 trigeminal nucleus connections, 299–300

1304 Posterior cerebral artery, anatomy, 1173, 1176 Posterior choroidal arteries, anatomy, 1175 Posterior periventricular nucleus (PeP), functions, 340 Posterior thalamic nucleus (Po) connections, 418–419 functions, 419 localization, 417–418 staining, 418 Posterolateral cortical amygdaloid nucleus (PLCO) acetylcholinesterase staining, 527 choline acetyltransferase staining, 527 cytoarchitectonics, 525 fibroarchitectonics, 525 heavy metal staining, 525 topographic landmarks, 524–525 Posteromedial cortical amygdaloid nucleus (PMCO) acetylcholinesterase staining, 535 cytoarchitectonics, 535 heavy metal staining, 535 topographic landmarks, 535 Postrhinal cortex connections associational projections, 688 commissural projections, 688 extrinsic connections, 688–689 hippocampal formation, 687–688 parahippocampus, 687–688 layers, 686–687 neuron types, 687 topology, 686 Preference–aversion behavior, circuitry, 910 Prefrontal cortex (PFC), periaqueductal gray connections, 249 Preoptic area, locus coeruleus connections, 280 Presubiculum anatomy, 661 connections associational projections, 668–669 commissural projections, 669 extrinsic connections, 669–671 hippocampus, 669 parahippocampus, 669 cytoarchitectonics, 668 neuron types, 668 subicular complex concept, 660–662 Pretectum anterior pretectal nucleus, 1116–1117 connections, 1113, 1116–1117 cytoarchitecture, 1113 nucleus of optic tract, 1118 olivary pretectal nucleus, 1113, 1115 posterior pretectal nucleus, 1117 stimulus response studies, 1113 topography, 1113 Primary olfactory cortex, see Piriform cortex Principal ganglion neurons, features, 86–87 PS, see Parastrial nucleus PSCh, see Suprachiasmic preoptic nucleus PT, see Parataenial nucleus

INDEX

Pterygopalatine artery, anatomy, anatomy, 1168–1169, 1171 Purine receptor, vestibular neuron modulation, 985, 987 Purkinje cell, see Cerebellum PV, see Paraventricular nucleus

R Rami communicantes, anatomy, 80 Raphe nuclei analgesia system, 869 autonomic control medullary nuclei, 767 midbrain nuclei, 775–776 basilar pontine nuclei connections, 171 descending spinal cord pathways, 157–158 hypoglossal nucleus connections, 310 olfactory bulb inputs and modulation, 952–953 serotonergic nerve clusters group B1 in raphe pallidus nucleus, 1206–1207 group B2/4 in raphe obscurus nucleus, 1207 group B3 in raphe magnus nucleus, 1207 group B5/8 in median raphe nucleus, 1207 group B6/7 in dorsal raphe nucleus, 1207 Re, see Nucleus reuniens thalami Receptor autoradiography, isocortex mapping, 729 Red nucleus afferents, 188–190 anatomy, 187 cytoarchitecture, 187–188 descending spinal cord pathways, 156 efferents, 190–191 functions, 192–193 lateral reticular nucleus connections, 177 Relaxin subfornical organ actions, 394 vascular organ of the lamina terminalis actions, 396 Reticular thalamic nucleus (Rt) functions, 442–444 projections, 441–442 structure, 441 Reticulotegmental nucleus of the pons (RtTg) afferents, 173 cytoarchitecture, 173 efferents, 173–174 functions, 173–174 Retina ganglion cells distribution, 1084 neurotransmission, 1088 types, 1086, 1088 recipient nuclei, see Dorsal lateral geniculate nucleus; Superior colliculus; Ventral lateral geniculate nucleus

Retroglenoid vein, anatomy, 1181 Retrosplenial cortex areas, 753 cortical connections and visuospatial function, 713–714, 753 cytoarchitectonics, 753 N-methyl-D-aspartate receptor antagonist-induced neurotoxicity Alzheimer's disease neurodegeneration relevance, 721–724 area 30 deafferentiation, 718–719 overview, 716 pathomorphological response, 717–718 polysynaptic circuit disinhibition adrenergic system, 719 cholinergic system, 719 glutamatergic system, 719–720 markers, 721 metabolic derangements, 720 psychosis induction, 721 neurotransmitter receptors, 753 Rh, see Rhomboid nucleus Rhomboid nucleus (Rh) afferents, 435 efferents, 438 structure, 323 Rostral ventrolateral reticular nucleus (RVL), autonomic control, 765, 767 Rt, see Reticular thalamic nucleus RtTg, see Reticulotegmental nucleus of the pons Ruffini ending, somatosensory receptor, 797–798 RVL, see Rostral ventrolateral reticular nucleus

S Salt appetite, circuitry, 912–913 SI, see Somatosensory cortex SII, see Somatosensory cortex SC, see Superior colliculus SCh, see Suprachiasmatic nucleus Schizophrenia, cingulate cortex pathology, 705 Sensory fibers, autonomic ganglia, 99, 101, 103 Septum chemoarchitecture lateral group, 606–607, 614 medial group, 614 posterior group, 614–615 ventral group, 615 cholinergic projections, 1262–1263 connections lateral group hippocampal formation, 616 hypothalamus, 616 midbrain, 616–617, 619 medial group, 619–620 posterior group, 620–621 ventral group, 621 dentate gyrus connections, 645–647 development, 602–603 functional organization, 621–623

INDEX

Septum Continued gestational time of origin, 31–32 history of study, 601 morphology and cytoarchitecture lateral group, 605 medial group, 605 overview, 603–605 posterior group, 605–606 ventral group, 606 neuron origins, 63 serotonergic projections, 1210 tachykinin projections, 1244 Serotonin brain stem projections, 1209 cerebellum projections, 1209 cerebral cortex projections, 1210 development of neurons, 1206 facial nucleus neurotransmission in medulla, 305 functions of systems appetite, 1213 hemodynamic regulation, 1213 motor activity, 1213 nociception, 1213–1214 hippocampus projections, 1210 hypothalamus projections, 1210 immunohistochemical staining, 1205 locus coeruleus inputs, 272 medulla projections, 1210 motor trigeminal nucleus neurotransmission in medulla, 300–301 neuronal clusters group B1 in raphe pallidus nucleus, 1206–1207 group B2/4 in raphe obscurus nucleus, 1207 group B3 in raphe magnus nucleus, 1207 group B5/8 in median raphe nucleus, 1207 group B6/7 in dorsal raphe nucleus, 1207 group B9 in medial lemniscus, 1207–1208 receptor subtypes and distribution, 1211–1213 septum projections, 1210 spinal cord projections, 1208–1209 tachykinin and serotonin coexistence cell body locations, 1246 functional interactions, 1247 pathways, 1246–1247 vascular innervation, 1195 vestibular neuron modulation, 982, 984 SIF cells, see Small intensely fluorescent cells SLEAc, see Central sublenticular extended amygdala SLEAm, see Medial sublenticular extended amygdala Small intensely fluorescent (SIF) cells cell junctions, 90 differentiation, 88 electron microscopy, 87–88

Small intensely fluorescent (SIF) cells Continued number in ganglions, 88–89 processes b89 synapses, 89–90 types, 89 SO, see Supraoptic nucleus SOC, see Superior olivary complex Somatosensory cortex afferents, 807–808 cytoarchitecture, 805–806 efferents, 808 gustatory system connections, 902–903 neurochemistry, 909 intracortical connections, 808–809 nociceptive neurons, 868 plasticity, 806–807 somatosensory areas, 805 somatotopic organization, 806 trigeminal sensory system SI barrel field injury models, 835 behavioral importance, 836 chemoarchitecture, 833 columnar organization, 833–834 cytoarchitecture, 833 imaging, 835 projections, 835 responses, 834–835 SII, 836 Somatosensory receptors cell bodies and central processes, 799–800 cutaneous receptors free nerve endings, 799 lanceolate endings, 799 Merkel endings, 797 Pacinian corpuscles, 798 Ruffini endings, 797–798 small lamellated corpuscles, 798–799 joint receptors, 799 muscle receptors free nerve endings, 799 Golgi tendon organs, 799 muscle spindles, 799 Somatosensory system glutaminergic pathways, 1272–1274 trigeminal sensory system, see Trigeminal sensory system Somatostatin dorsal horn neurochemistry, 137 vestibular neuron modulation, 984 SP, see Substance P SPF, see Subparafascicular nucleus Spinal cord afferent fibers area X, 114 lamina I, 112 lamina II, 112–113 lamina III–VI, 113–114 lamina VII, 114 lateral spinal nucleus, 113 overview, 111–112 somatotopic organization, 114–115 ventral horn, 114

1305 Spinal cord Continued ascending somatosensory pathways dorsal column pathways, 800 spinothalamic tract, 800–901 basilar pontine nuclei connections, 171 brain stem pathways in micturition coordination abdominal pressure control systems, 323 diffuse descending systems, 323 micturition reflex, 325 overview, 323–324 cerebellar termination of mossy fiber systems, 231–233 cholinergic projections, 1260–1261 cytoarchitecture area X, 125–126 lamina I, 121–122 lamina II, 122 lamina III, 122 lamina IV, 122–123 lamina V, 123 lamina VI, 123 lamina VII, 123 lamina VIII, 124 lamina IX, 124–125 lateral cervical nucleus, 126 lateral spinal nucleus, 126 lateral reticular nucleus connections, 176–179 locus coeruleus connections, 281 periaqueductal gray connections, 247–249 serotonergic projections, 1208–1209 substantia gelatinosa, see Substantia gelatinosa of the spinal cord tachykinin projections, 1242 vasculature arteries, 1192–1193 veins, 1193–192 Spinal dorsal horn, see Dorsal horn interneurons; Substantia gelatinosa of the spinal cord Spinocerebellar tract dorsal spinocerebellar tract, 152 ventral spinocerebellar tract, 152 Spinocervical tract nociception pathway, 867 overview, 151 Spinohypothalamic tract, nociception pathway, 867–868 Spinomesecephalic tract, nociception pathway, 866–867 Spinoreticular tract nociception pathway, 867 overview, 150–151 Spinothalamic tract nociception pathway, 864–865 somatosensory system, 800–901 Splanchic nerves, anatomy, 81 Striatum cholinergic projections, 1262 cortical input corticostriatal neuron subtypes, 458, 460

1306 Striatum Continued organization of corticostriatal afferents convergent corticostriatal organization, 462–463 general organization, 463 quantitative data, 463–464 topographical organization, 460–462 overview, 498 dual output systems, 484–485, 487–488, 500 gestational time of origin, 29 indirect output pathway lateral globus pallidus morphology, 479–480 neuron types, 480 neurotransmission, 480 output, 481 synaptic input, 480–481 overview, 478–479 subthalamic nucleus morphology, 481 output, 482 synaptic input, 481 interneurons abundance, 471 classification calretinin/γ-aminobutyric acid interneurons, 474 large aspiny cholinergic neurons, 473 medium aspiny GABAergic neurons, 473 overview, 471 parvalbumin interneurons, 473 somatostatin/neuropeptide Y interneurons, 473–474 locus coeruleus connections, 279 medium spiny projection neurons inputs axon collateral inputs, 468–470 cerebral cortex, 465 dopaminergic neurons, 468 interneurons, 469 miscellaneous inputs, 469 thalamus, 465, 467–468 morphology, 464–465 neurotransmitters, 465 outputs α2-adenosine receptors, 477–478 acetylcholine receptors, 478 γ-aminobutyric acid receptors, 478 connectional basis, 474–475 direct versus indirect, 474, 498, 500 dopamine receptor subtypes and mediation of gene regulation, 475–477 neurochemical basis, 475 opioid receptors, 478 neuron origins, 62 nigrostriatal dopamine system dorsal tier dopamine neurons, 489–490 input to pars comapcta neurons, 490 overview, 488–489 ventral tier dopamine neurons, 490

INDEX

Striatum Continued patch/matrix compartments cortical input, 494–497 organization, 495–497 overview, 490, 492, 500 striatal outputs, 492–494 thalamic input, 495 Sub, see Nucleus submedius Subcommissural organ function, 399–400 localization, 399 Subfornical organ afferents, 391–393 efferents, 393 hormone receptors, 393–394 vascularization, 391 Subiculum anatomy, 660–661 connections afferents, 664–665 associational projections, 663 efferents, 665–667 hippocampus, 663 parahippocampus, 663–664 topographical organization, 667–668 cytoarchitectonics, 662 gestational time of origin, 34–35 history of study, 660 neuron origins, 65 neuron types, 662 subicular complex concept, 660–662 Submandibular ganglion, anatomy, 83 Subparafascicular nucleus (SPF) afferents, 437 functions, 441 structure, 436 Subplate, neuron migration, 66 Substance P (SP) dorsal horn neurochemistry, 136 functions, 1245–1246 gene, 1214 history of study, 1214 immunohistochemical staining, 1214 neuron development, 1240 neuron distribution, 1240–1241 prcessing, 1214 receptor types and distribution, 1245 serotonin and tachykinin coexistence cell body locations, 1246 functional interactions, 1247 pathways, 1246–1247 vestibular neuron modulation, 985 Substantia gelatinosa of the spinal cord ascending pathways cerebellum dorsal spinocerebellar tract, 152 spinoolivary tract, 152 ventral spinocerebellar tract, 152 diencephalon, 153–154 medulla direct dorsal column pathway, 149–150 nuclei afferents, 151 postsynaptic dorsal column pathway, 150

Substantia gelatinosa of the spinal cord Continued spinocervical tract, 151 spinoreticular tracts, 150–151 midbrain reticular formation, 153 periaqueductal gray, 153 pons, 151–152 superior colliculus, 152–153 telencephalon, 154 definition, 129–130 descending pathways cerebral cortex, 155–156 diencephalon, 156 red nucleus, 156 telencephalon, 154–156 dorsal horn neurochemistry γ-aminobutyric acid, 140 calcitonin gene-related peptide, 137 choline acetyltransferase, 140 dynorphin, 139 endomorphin, 139 enkephalin, 137, 139 glutamate, 139–140 glycine, 140 neurokinin A, 136 neurokinin B, 136–137 neurokinin receptors, 137 nonpeptididergic sensory fiber markers, 142–143 opioid receptors, 139 somatostatin, 137 substance P, 136 neuronal characteristics lamina I, 130–131 lamina II, 131–133, 136 lamina III, 133 synaptic glomeruli functions, 134, 136 neurochemistry, 143 types, 133–134 ultrastructure, 133 Substantia nigra dendrite morophology, 483–484 neuron types, 482–483 pars reticula neurons input, 484 output, 484 Subthalamic nucleus morphology, 481 output, 482 synaptic input, 481 Subventricular nucleus (SV), features, 592 Subventricular zone, neuron sources, 37 Sulci, molecular specification, 5 SuM, see Supramammillary nucleus Superior cervical ganglion anatomy, 86–88 neuron number, 85 neurotransmitters, 90 synapses, 90 Superior colliculus (SC) axons and synaptic relations, 1104–1105 cell types, 1102–1104 connections, 1107–1110 functions, 1101–1102

INDEX

Superior colliculus (SC) Continued layers, 1101 retinotopic organization, 1105 spinal ascending pathways, 152–153 stimulus response studies, 1105–1107 Superior olivary complex (SOC) components, 1013 lateral superior olive, 1013, 1015 medial nucleus of the trapezoid body, 1017–1018 medial superior olive, 1018–1019 periolivary nuclei, 1019–1020 superior paraolivary nucleus, 1019 Suprachiasmatic nucleus (SCh), functions, 339 Suprachiasmic preoptic nucleus (PSCh), functions, 338 Supramammillary area, dentate gyrus connections, 647 Supramammillary nucleus (SuM), functions, 348 Supraoptic nucleus (SO) afferents, 374–375 interneurons, 372–373 magnocellular neuron morphology and efferent path, 371–372 principal neuron distribution and neuropeptides, 370–371 SV, see Subventricular nucleus Synaptic glomeruli, dorsal horn functions, 134, 136 neurochemistry, 143 types, 133–134 ultrastructure, 133

T Tachykinins, see Neurokinins; Substance P Taste, see Gustatory system Tectum, basilar pontine nuclei connections, 171 Telencephalon ependymal stem cells, 68, 71–72 gene expression in development, 17–20 gestational time of origin amygdala, 29–31 hippocampal region entorhinal cortex, 34 hippocampus, 35 subiculum, 34–35 limbic cortex, 32 neocortex, 32 olfactory bulb, 35 olfactory peduncle, 35–36 overview, 27–28 pallidum, 28–29 piriform cortex, 32–33 septum, 31–32 striatum, 29 glial stem cells, 68, 71–72 neural stem cells dynamics in cortical germinal zones, 68, 71 maps of mosaics in telencephalic neuroepithelium over time, 38–61

Telencephalon Continued neuron migration dentate migratory stream in dentate gyrus, 68 lateral migratory stream in cerebral cortex Cajal–Retzius neuron migration, 66 layer VI–VII neuron migration stages, 66–67 piriform cortex, 67 subplate neuron migration, 66 rostral migratory stream in olfactory bulb, 67–68 neuron origins germinal sources of telencephalic neurons amygdala, 62–63 dentate gyrus, 65 entorhinal cortex, 64 limbic cortex, 63–64 neocortex, 63 nucleus accumbens, 62 olfactory bulb, 65 olfactory peduncle, 65 olfactory tubercle, 62 pallidum, 37, 62 piriform cortex, 64 septum, 63 striatum, 62 subiculum, 65 neuroepithelium sources and gene expression, 37 subventricular zone sources, 37 spinal pathways ascending pathways, 154 descending pathways, 154–156 Temporal cortex areas, 745–746 cytoarchitectonics, 746 layers, 746 local cerebral glucose utilization studies, 746 neurotransmitter receptors, 746–747 parcellation schemes, 744–746 Thalamic reticular nucleus auditory sector, 1049–1050 visual sector, 1095–1096 Thalamus afferent classification as drivers or modulators, 409–410 ascending spinal cord pathways cerebellum, 157 medulla proproispinal connections, 158–159 raphe nuclei, 157–158 reticular formation, 157 trigeminal nucleus, 157 vestibular nuclei, 158 overview, 153 periaqueductal gray, 156 pons, 156–157 association thalamic nuclei anterior nuclei, 430–432 lateral nuclei, 432–433

1307 Thalamus Continued mediodorsal nucleus, 425–429 nucleus submedius, 429–430 auditory processing, see Medial geniculate body autonomic control intralaminar nuclei, 777–778 mediodorsal nucleus, 777 paraventricular nucleus, 777 ventroposterior parvocellular nucleus, 776–777 cingulate cortex connections area 29, 715 axon terminal morphology and multiple heteroreceptor regulation, 715–716 regional differentiation, 714–715 dorsal versus ventral thalamus distinctions, 407–408 gustatory system neurochemistry, 909 relay nuclei, 900, 902 intralaminar nuclei afferents, 436–437 central lateral nucleus, 436 central medial nucleus, 435 efferents, 437–439 functions, 439–441 paracentral nucleus, 435–436 parafascicular nucleus, 436 subparafascicular nucleus, 436 locus coeruleus connections, 279 midline nuclei afferents, 434–435 efferents, 437–439 functions, 439–441 intermediodorsal nucleus, 434 nucleus reuniens thalami, 434 parataenial nucleus, 434 paraventricular nucleus, 434 rhomboid nucleus, 323 motor nuclei ventral lateral/ventral anterior complex, 422–424 ventromedial thalamic nucleus, 424–425 nociceptive neurons, 868 nuclei, classical categorization, 408–409 olfactory cortex connections, 946 reciprocity of thalamocorticothalamic relationships, 410–411 reticular thalamic nucleus functions, 442–444 projections, 441–442 structure, 441 sensory nuclei and projections dorsal lateral geniculate nucleus, 411–412 intergeniculate leaflet, 413 medial geniculate nucleus, 420–421 parvicellular ventral posterior nucleus, 420 posterior thalamic nucleus, 417–419 ventral lateral geniculate nucleus, 412–413

1308 Thalamus Continued ventroposterolateral nucleus, 414–417 ventroposteromedial nucleus, 414–417 somatosensory system afferents, 805 cytoarchitecture, 804 efferents, 805 plasticity, 804 somatotopic organization, 804 striatum connections medium spiny projection neuron connections, 465, 467–468 patch/matrix compartment connections, 495 tachykinin projections, 1243–1244 trigeminal sensory system sensory nuclei inputs posterior nucleus chemoarchitecture, 832 cytoarchitecture, 832 projections, 832–833 responses, 832 ventroposteromedial nucleus chemoarchitecture, 832 cytoarchitecture, 832 projections, 832 responses, 832 TM, see Tuberomammillary nucleus Trigeminal nucleus afferents forebrain pathways, 298 medulla, 300–301 midbrain pathways, 298–299 pons, 299–300 cerebellar termination of mossy fiber systems, 233–234 cytoarchitectonics, 297 dendritic architecture, 297 descending spinal cord pathways, 157 interneurons, 297 myotopic organization, 295–297 Trigeminal sensory system ascending spinal pathways dorsal column pathways, 800 spinothalamic tract, 800–901 brain stem sensory nuclei afferent organization, 825–826 caudal subnucleus chemoarchitecture, 829 cytoarchitecture, 829 projections, 830–831 responses, 829–830 cytoarchitectonics, 826 groups, 824–825 interpolar subnucleus chemoarchitecture, 828–829 cytoarchitecture, 828–829 projections, 829 responses, 829 mesencephalic trigeminal nucleus chemoarchitecture, 827 cytoarchitecture, 827 projections and function, 827 oral subnucleus chemoarchitecture, 828

INDEX

Trigeminal sensory system Continued cytoarchitecture, 828 projections, 828 responses, 828 paratrigeminal nucleus chemoarchitecture, 831 cytoarchitecture, 831 projections, 831 responses, 831 principal sensory nucleus chemoarchitecture, 827–828 cytoarchitecture, 827–828 projections, 828 responses, 828 somatotopy, 826–827 cerebral cortex, see Somatosensory cortex cornea and conjuctiva innervation, 819, 822 development central vibrissal pathway, 837–838 ganglion cells and periphery, 836–837 plasticity, 838 ganglion cell fiber neurotransmission, 823–824 numbers, 823 response properties, 824 somatotopy, 824 medullary relay nuclei afferents, 802–803 cytoarchitecture, 801 efferents, 803–804 plasticity, 801–802 somatotopic organization, 801 meninges and cranial vessel innervation, 818–819 peripheral nerves and receptors, 817–818 receptors, see Somatosensory receptors somatosensory cortex SI barrel field injury models, 835 behavioral importance, 836 chemoarchitecture, 833 columnar organization, 833–834 cytoarchitecture, 833 imaging, 835 projections, 835 responses, 834–835 SII, 836 temporomandibular joint innervation, 822 thalamic nuclei inputs posterior nucleus chemoarchitecture, 832 cytoarchitecture, 832 projections, 832–833 responses, 832 ventroposteromedial nucleus chemoarchitecture, 832 cytoarchitecture, 832 projections, 832 responses, 832 thalamus afferents, 805 cytoarchitecture, 804 efferents, 805

Trigeminal sensory system Continued plasticity, 804 somatotopic organization, 804 tongue innervation, 822, 893 tooth and periodontal ligament innervation, 822–823 vibrissal follicle innervation, 822 Tuberal nucleus, functions, 346–347 Tuberomammillary nucleus (TM) autonomic control, 782 functions, 349

V Vagus nerve, anatomy, 83–84 Vascular innervation acetylcholine, 1194–1195 neuropeptides, 1195 nitric oxide, 1195 noradrenaline, 1194 serotonin, 1195 Vascular organ of the lamina terminalis (OVLT) afferents, 395–396 efferents, 396 ependymal cells, 394 hormone secretion and receptors, 396 localization, 394 vascularization, 395 Vasculature, cerebral arteries anterior cerebral artery, 1179–1180 anterior choroidal artery, 1179 arterial circle, 1176–1177, 1179–1180 basilar artery, 1173 extracranial anastomotic circles, 1171 extracranial origins, 1168 internal carotod artery, 1171 internal ophthalmic artery, 1179 longitudinal hippocampal artery, 1175 middle cerebral artery, 1179 olfactory artery, 1180 pial arterial network, 1180–1181 posterior cerebral artery, 1173, 1176 posterior choroidal arteries, 1175 pterygopalatine artery, 1168–1169, 1171 vertebral arteries, 1171 cast preparation, 1167–1168 functional imaging of blood flow, 1195, 1197 innervation, see Vascular innervation mammalian comparison, 1167 veins deep venous systems, 1182, 1191 extracranial anastomoses, 1182 internal jugular vein, 1181 retroglenoid vein, 1181 superficial venous systems, 1181–1182 Veins, see Spinal cord; Vasculature, cerebral Ventral basolateral amygdaloid nucleus (BLV) acetylcholinesterase staining, 585 choline acetyltransferase staining, 585 cytoarchitectonics, 584 fibroarchitectonics, 584–585

INDEX

Ventral basolateral amygdaloid nucleus (BLV) Continued heavy metal staining, 585 topographic landmarks, 584 Ventral horn, afferent fibers, 114 Ventral lateral geniculate nucleus (VLG) connections, 1113–1115 functions, 413 intergeniculate leaflet, 1111 lateral division, 1112 medial division, 1112 organization, 1110–1111 projections, 412–413 structure, 412 Ventral lateral/ventral anterior complex connections, 423 functions, 423–424 structure, 422–423 Ventral premammillary nucleus (PMV), functions, 348 Ventromedial hypothalamic nucleus (VMH), functions, 345–346 Ventromedial thalamic nucleus (VM) connections, 424 functions, 424–425 structure, 424 Ventroposterolateral nucleus (VPL) chemoarchitecture, 414 connections, 414–416 cytoarchitecture, 414 functions, 416–417 Ventroposteromedial nucleus (VPM) chemoarchitecture, 414 connections, 414–416 cytoarchitecture, 414 functions, 416–417 Vertebral arteries, anatomy, 1171 Vestibular nuclei, see also Vestibular system cerebellar termination of mossy fiber systems, 234–235 descending spinal cord pathways, 158 Vestibular system first-order vestibular neurons ionic currents, 968 postnatal maturation, 968 regular neurons, 967–968 ultrastructure, 967 hair cell receptors, 966–967 neurotransmitters and neuromodulators of central neurons adrenocorticotropin, 985 cholinergic receptors anatomy, 981 behavior studies, 981 electrophysiology, 981 dopaminergic modulation, 982–984

Vestibular system Continued excitatory amino acid receptors N-methyl-D-aspartate receptors, 978–979 pharmacological analysis, 978 plasticity role, 979 subunits, 977–978 histaminergic modulation, 981–982, 984 inhibitory amino acid receptors anatomical studies, 979 electrophysiological studies, 979–980 functional considerations, 980 nerve growth factor, 985 neurokinins, 985 noradrenergic modulation, 983–984 opioid receptors, 984–985 purine receptors, 985, 987 serotonergic modulation, 982, 984 somatostatin, 984 substance P, 985 second-order vestibular neurons activation, 971–972 angular acceleration response, 972 linear acceleration response, 972 medial vestibular nucleus neurons calcium currents, 974–975 electrophysiological identification, 974 gap junctions, 976 postnatal maturation, 976 potassium currents, 975 rhythmic activities, 975–976 sodium currents, 974 types and functions, 976–977 postnatal maturation otolith neurons, 973 semicircular canal neurons, 973 visual and proprioceptive stimulation response, 973 vestibular nuclear complex lateral vestibular nucleus, 970 medial vestibular nucleus, 970–971 spinal vestibular nucleus, 971 structure, 968–969 superior vestibular nucleus, 969–970 Visual cortex cytoarchitectonics, 1123 extrinsic connections of areas Oc1, Oc2L, and Oc2M associational connections interareal connections of visuotopically organized cortices, 1134 visuotopically organized cortices with nonvisuotopically organized cortices, 1134–1136

1309 Visual cortex Continued commissural connections, 1136–1137 diencephalon, 1129–1132 mesencephalon, 1129–1132 pons, 1129–1132 subcortical telencephalic connections, 1132–1133 feedforward and feedback associational pathways, 1138–1141 functions, 1139–1141 lesion studies, 1139–1140 primary visual cortex area Oc1 intrinsic connections, 1127–1128 cellular organization, 1124–1127 honeycomb mosaic of modules, 1125–1126 layers, 1125 neuron number, 1124 neurotransmission, 1126–1127 sex differences, 1124–1125 receptive field properties of neurons, 1128–1129 visuotopic organization, 1123 Visual system, see also Accessory optic system; Dorsal lateral geniculate nucleus; Pretectum; Retina; Superior colliculus; Ventral lateral geniculate nucleus; Visual cortex eye photoreceptors, 1083 glutaminergic pathways, 1274–1275 lateral posterior nucleus, 1122–1123 optic nerve anatomy, 1089 neurotransmission, 1089 parabigeminal nucleus, 1121–1122 pathways, overview, 1084–1086 spatial resolution, 1083–1084 thalamic reticular nucleus, 1095–1096 VLG, see Ventral lateral geniculate nucleus VM, see Ventromedial thalamic nucleus VMH, see Ventromedial hypothalamic nucleus VNO, see Vomeronasal organ Vomeronasal organ (VNO) accessory olfactory bulb connections, 949 epithelium structure, 948–949 glutaminergic pathways, 1277–1278 receptor neurons, 948 VPL, see Ventroposterolateral nucleus VPM, see Ventroposteromedial nucleus VPPC, see Parvicellular ventral posterior nucleus Zebrin, staining in cerebellum, 226, 228–229

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