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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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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This book is dedicated to: Babis and Kalliopi
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Contents
II
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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CONTENTS
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
2
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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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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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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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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SHIRLEY A. BAYER and JOSEPH ALTMAN
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.
Beal, J. A., Russel, C. T., and Knight, D. S. (1988). Morphological and developmental characterization of local-circuit neurons in lamina III of rat spinal cord. Neurosci. Lett. 86, 1–5. Bester, H., Chapman, V., Besson, J. M., and Bernard, J. F. (2000). Physiological properties of the lamina I spinobrachial neurons in the rat. J. Neurophysiol. 83, 2239–2259. Brown, A. G. (1981). “Organization in the Spinal Cord: The Anatomy and Physiology of Identified Neurons.” Springer-Verlag, Berlin. Burstein, R., Cliffer, K. D., and Giesler, G. J., Jr. (1987). Direct somatosensory projections from the spinal cord to the hypothalamus and telencephalon. J. Neurosci. 7, 4159–4164. Burstein, R., Cliffer, K. D., and Giesler, G. J., Jr. (1990a). Cells of origin of the spinothalamic tract in the rat. J. Comp. Neurol. 291, 329–344. Burstein, R., Dado, R. J., and Giesler, G. J., Jr. (1990b). The cells of origin of the spinothalamic tract of the rat: A quantitative reexamination. Brain Res. 511, 329–337. Burstein, R., and Potrebic, S. (1993). Retrograde labeling of neurons in the spinal cord that project directly to the amygdala or the orbital cortex in the rat. J. Comp. Neurol. 335, 469–485. Cechetto, D. F., Standaert, D. G., and Saper, C. B. (1985). Spinal and trigeminal dorsal horn projections to the parabrachial nucleus in the rat. J. Comp. Neurol. 240, 153–160. Cervero, F., Handwerker, H. O., and Laird, J. M. A. (1988). Prolonged noxious mechanical stimulation of the rat’s tail: Responses and encoding properties of dorsal horn neurones. J. Physiol. (London) 404, 419–436. Chaouch, A., Menétrey, D., Binder, D., and Besson, J. M. (1983). Neurons at the origin of the medial component of the bulbopontine spinoreticular tract in the rat: An anatomical study using horseradish peroxidase retrograde transport. J. Comp. Neurol. 214, 309–320. Cook, A. J., and Woolf, C. J. (1985). Cutaneous receptive field and morphological properties of hamstring flexor α-motoneurones in the rat. J. Physiol. (London) 364, 249–263. Esteves, F., Lima, D., and Coimbra, A. (1993). Structural types of spinal cord marginal (lamina-I) neurons projecting to the nucleus of the tractus solitarius in the rat. Somatosens. Mot. Res. 10, 203–216. Feil, K., and Herbert, H. (1995). Topographic organization of spinal and trigeminal somatosensory pathways to the rat parabrachial and Kolliker–Fuse nuclei. J. Comp. Neurol. 353, 506–528. Fukuyama, U. (1955). On cytoarchitectural lamination of the spinal cord in the albino rat. Anat. Rec. 121, 396. Giesler, G. J., Björkeland, M., Xu, Q., and Grant, G. (1988). Organization of the spinocervicothalamic pathway in the rat. J. Comp. Neurol. 268, 223–233. Giesler, G. J., Cannon, J. T., Urca, G., and Liebeskind, J. C. (1978). Long ascending projections from substantia gelatinosa Rolandi and the subjacent dorsal horn in the rat. Science 202, 984–986. Giesler, G. J., Nahin, R. L., and Madsen, A. M. (1984). Postsynaptic dorsal column pathway of the rat. I. Anatomical studies. J. Neurophysiol. 51, 276–291. Giesler, G. J., Urca, G., Cannon, J. T., and Liebeskind, J. C. (1979). Response properties of neurons of the lateral cervical nucleus in the rat. J. Comp. Neurol. 186, 65–78. Granum, S. L. (1986). The spinothalamic system of the rat. I. Locations of cells of origin. J. Comp. Neurol. 247, 159–180. Han, Z. S., Zhang, E. T., and Craig, A. D. (1998). Nociceptive and thermoreceptive lamina I neurons are anatomically distinct. Nat. Neurosci. 1, 177–178. Hongo, T., Kitazawa, S., Ohki, Y., Sasaki, M., and Xi, M. C. (1989). A physiological and morphological study of premotor interneurones in the cutaneous reflex pathways in cats. Brain Res. 505, 163–166. Hylden, J. L., Anton, F., and Nahin, R. L. (1989). Spinal lamina I projection neurons in the rat: Collateral innervation of parabrachial area and thalamus. Neuroscience 28, 27–37.
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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
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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
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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 (25 μm) and a 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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C H A P T E R
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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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
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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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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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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
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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
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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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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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Neurotransmitters in LC and peri-LC
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; P96%) 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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Kosel, K. C., Van Hoesen, G. W., and Rosene, D. L. (1983). A direct projection from the perirhinal cortex (area 35) to the subiculum in the rat. Brain Res. 269, 347–351. Krettek, J. E., and Price, J. L. (1977a). The cortical projections of the mediodorsal nucleus and adjacent thalamic nuclei in the rat. J. Comp. Neurol. 171, 157–192. Krettek, J. E., and Price, J. L. (1977b). Projection from the amygdaloid complex to the cerebral cortex and thalamus in the rat and cat. J. Comp. Neurol. 172, 687–722. Krieg, W. J. S. (1946a). Connections of the cerebral cortex. I. The albino rat. A. Topography of the cortical areas. J. Comp. Neurol. 84, 221–275. Krieg, W. J. S. (1946b). Accurate placement of minute lesions in the brain of the albino rat. Q. Bull. Northwestern Univ. Med. Sch. 20, 199–208. Krieg, W. J. S. (1947). Connections of the cerebral cortex. I. The albino rat. C. Extrinsic connections. J. Comp. Neurol. 86, 267–394. Lasiter, P. S., and Glanzmann, D. L. (1983). Axon collaterals of pontine taste area neurons project to the posterior ventromedial thalamic nucleus and to the gustatory neocortex. Brain Res. 258, 299–304. Lasiter, P. S., Glanzmann, D. L., and Mensah, P. A. (1982). Direct connectivity between pontine tatste areas and gustatory neocortex in rat. Brain Res. 234, 111–121. Lassen, N. A. (1959). Cerebral blood flow and oxygen consumption in man. Physiol. Rev. 39, 183–238. LeMessurier, D. H. (1948). Auditory and visual areas of the cerebral cortex of the rat. Fed. Proc. 7, 10–71. Leong, S. K. (1983). Localizing the corticospinal neurons in neonatal, developing and mature albino rat. Brain Res. 265, 1–9. Lin, C.-S., Lu, S. M., and Schmechel, D. E. (1986). Glutamic acid decarboxylase and somatostatin immunoreactivities in rat visual cortex. J. Comp. Neurol. 244, 369–383. Markowitsch, H. J., and Guldin, W. O. (1983). Heterotopic interhemispheric cortical connections in the rat. Brain Res. Bull. 10, 805–810. Mason, R., and Gross, G. A. (1981). Cortico-recipient and tectorecipient visual zones in the rat’s posterior (pulvinar) nucleus: An anatomical study. Neurosci. Lett. 25, 107–112. Mata, M., Fink, D. J., Gainer, H., Smith, C. B., Davidsen, L., Savaki, H., Schwartz, W. J., and Sokoloff, L. (1980). Activity dependent energy metabolism in rat posterior pituitary reflects sodium pump activity. J. Neurochem. 34, 213–215. McDaniel, W. F., McDaniel, S. E., and Thomas, R. K. (1978). Thalamocortical projections to the temporal and parietal association cortices in the rat. Neurosci. Lett. 7, 121–125. Miller, M. W., and Vogt, B. A. (1984). Direct connections of rat visual cortex with sensory, motor and association cortices. J. Comp. Neurol. 226, 184–202. Montero, V. M. (1973). Evoked responses in the rat’s visual cortex to contralateral, ipsilateral and restricted photic stimulation. Brain Res. 53, 192–196. Montero, V. M. (1981). Comparative studies on the visual cortex. In “Cortical Sensory Organization” (Woolsey, C. N., Ed.), pp. 33–81. Humana Press, Clifton, NJ. Montero, V. M., Bravo, H., and Fernández, V. (1973a). Striate-peristriate cortico-cortical connections in the albino and gray rat. Brain Res. 53, 202–207. Montero, V. M., Rojas, A., and Torrealba, F. (1973b). Retinotopic organization of striate and peristriate visual cortex in the albino rat. Brain Res. 53, 197–201. Musil, S. Y., and Olson, L. J. (1990). The role of cat cingulate cortex in sensorimotor integration. In “The Cerebral Cortex of the Rat” (Kolb, B., and Tees, R. C., Eds.), pp. 345–365. MIT, Cambridge, MA. Neafsey, E. J. (1990). The complete ratunculus: Output organization of layer V of the cerebral cortex. In “The Cerebral Cortex of the
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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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regulation of blood pressure in the rat. J. Pharmacol. Exp. Ther. 226, 893–899. Witkin, J. W., Paden, C. M., and Silverman, A. J. (1982). The luteinizing hormone releasing hormone (LHRH) systems in the rat brain. Neuroendocrinology 35, 429–238. Woulfe, J. M., Flumerfelt, B. A., and Hrycyshyn, A. W. (1990). Efferent connections of the A1 noradrenergic cell group: A DBH immunohistochemical and PHA-L anterograde tracing study. Exp. Neurol. 109, 308–322. Yamada, K. A., McAllen, R. M., and Loewy, A. D. (1984). GABA antagonists aplied to the ventral surface of the medulla oblongata block the baraoreceptor reflex. Brain Res. 297, 175–180. Yamano, M., Hillyard, C. J., Girgis, S., Emson, P. C., MacIntyre, I., and Tohyama, M. (1988). 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. Exp. Brain Res. 71, 603–610. 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., Cechetto, D. F., and Saper, C. B. (1990). Evidence for a cholinergic projection from the pedunculopontine tegmental nucleus to the rostral ventrolateral medulla of the rat. Brain Res. 517, 19–24. Yasui, Y., Saper, C. B., and Cechetto, D. F. (1989). Calcitonin generelated peptide immunoreactivity in the visceral sensory cortex, thalamus, and related pathways in the rat. J. Comp. Neurol. 290, 487–501. 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. Zaborszky, L., Beinfeld, M. C., Palkovits, M., and Heimer, L. (1984). Brainstem projection to the hypothalamic ventromedial nucleus in the rat: A CCK-containing long ascending pathway. Brain Res. 303, 225–231. Zardetto-Smith, A. M., and Gray, T. S. (1987). A direct neural projection from the nucleus of the solitary tract to the subfornical organ in the rat. Neurosci. Lett. 80, 163–166. Zaretskaia, M. V., Zaretsky, D. V., Shekhar, A., and Dimicco, J. A. (2002). Chemical stimulation of the dorsomedial hypothalamus evokes non-shivering thermogenesis in anesthetized rats. Brain Res. 928, 113–125. Zhang, Y. H., Lu, J., Elmquist, J. K., and Saper, C. B. (2000). Lipopolysaccharide activates specific populations of hypothalamic and brainstem neurons that project to the spinal cord. J. Neurosci. 20, 6578–6586.
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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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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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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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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
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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;
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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
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Right
AGm
AGl
2,7
AGl 1,6
,7
2,
SI
SI
7
1,3
,5
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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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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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D
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 (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.
References Apartis, E., Poindessous-Jazat, F. R., Lamour, Y. A., and Bassant, M. H. (1998). Loss of rhythmically bursting neurons in rat medial septum following selective lesion of septohippocampal cholinergic system. J. Neurophysiol. 79, 1633–1642. Arvidsson, U., Riedl, M., Elde, R., and Meister, B. (1997). Vesicular acetylcholine transporter (VAcetylcholineT) protein: A novel and unique marker for cholinergic neurons in the central and peripheral nervous systems. J. Comp. Neurol. 378, 454–467. Ashcroft, F. M. (1991). Ca2+ channels and excitation–contraction coupling. Curr. Opin. Cell Biol. 3, 671–675. Audesirk, G., Cabell, L., and Kern, M. (1997). Modulation of neurite branching by protein phosphorylation in cultured rat hippocampal neurons. Dev. Brain Res. 102, 247–260. Bartus, R. T. (2000). On neurodegenerative diseases, models, and treatment strategies: Lessons learned and lessons forgotten following a generation following the cholinergic hypothesis. Exp. Neurol. 163, 495–529. Baxter, M. G., Bucci, D. J., Gorman, L. K., Wiley, R. G., and Gallagher, M. (1995). Selective immunotoxic lesions of basal forebrain cholinergic cells: Effects on learning and memory in rats. Behav. Neurosci. 109, 714–722. Blesch, A., and Tuszynski, M. H. (2001). GDNF gene delivery to injured adult CNS motor neurons promotes axonal growth, expression of the trophic neuropeptide CGRP, and cellular protection. J. Comp. Neurol. 436, 399–410. Bobkova, N. V., Nesterova, I. V., and Nesterov, V. V. (2001). The state of cholinergic structures in forebrain of bulbectomized mice. Bull. Exp. Biol. Med. 131, 427–431. Butcher, L. L. (1995). Cholinergic neurons and networks. In “The Rat Nervous System” (G. Paxinos, Ed.), 2nd ed., pp. 1003–1015. Academic Press, San Diego. Butcher, L. L., and Woolf, N. J. (1989). Neurotrophic neurons may exacerbate the pathologic cascade of Alzheimer’s disease. Neurobiol. Aging 10, 557–570. Carey, R. G., and Rieck, R. W. (1987). Topographic projections to the visual cortex from the basal forebrain in the rat. Brain Res. 424, 205–215. Dale, H. H. (1914). The action of certain esters and ethers of choline, and their relation to muscarine. J. Pharmacol. Exp. Ther. 6, 147–190. Dale, H. H. (1938). Acetylcholine as a chemical transmitter of the effects of nerve impulses. II. Chemical transmission at ganglionic synapses and voluntary motor nerve endings: Some general considerations. J. Mt. Sinai Hosp. 4, 416–429. Descarries, L., Gisiger, V., and Steriade, M. (1997). Diffuse transmission by acetylcholine in the CNS. Prog. Neurobiol. 53, 603–625. Eglen, R. M., Reddy, H., Watson, N., and Challiss, R. A. (1994). Muscarinic acetylcholine receptor subtypes in smooth muscle. Trends Pharmacol. Sci. 15, 114–119. Fadda, F., Cocco, S., and Stancampiano, R. (2000). Hippocampal acetylcholine release correlates with spatial learning performance in freely moving rats. Neuroreport 11, 2265–2269. Farris, T., Butcher, L,. L., Oh, J. T., and Woolf, N. J. (1995). Trophic factor modulation of cortical acetylcholinesterase reappearance following transaction of the medial cholinergic pathway in the adult rat. Exp. Neurol. 131, 180–192. Farris, T., Woolf, N. J., Oh, J. T., and Butcher, L. L. (1993). Reestablishment of laminar patterns of cortical acetylcholinesterase activity following axotomy of the medial cholinergic pathway in the adult rat. Exp. Neurol. 121, 77–92. Friedlich, A., and Butcher, L. L. (1994). Involvement of free oxygen radicals in β-amyloidosis: An hypothesis. Neurobiol. Aging 15, 443–455.
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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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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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