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 (<20 μm) were usually achromatic and were frequently seen in association with neuroglial cells. Medium and small cells made up most of the rostral two thirds (i.e., the parvicellular part). Strominger et al. (1987) only recognized three neuronal populations: neurons with coarse Nissl bodies, neurons with a fine granular Nissl substance, and achromatic neurons. The soma sizes of neurons with coarse and fine Nissl substance overlap considerably, whereas the achromatic neurons are clearly smaller than the other two cell types. “Coarse” neurons were found exclusively in the caudal tip, while the rostral tip harbored predominantly “fine” neurons. In the intermediate red nucleus both cell groups were found. The small neurons were found throughout the nucleus. Finally, studies by Tucker et al. (1989) differentiated between neurons with a relatively small, oval, and eccentrically placed nucleus and displaying a large cytoplasmic–nuclear ratio (magnocellular neurons) and neurons with a relatively large rounded and a centrally placed nucleus displaying a
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small cytoplasmic to nuclear ratio (parvicellular neurons). Approximately 3000 magnocellular neurons were counted which could be divided into a largesized (>25 μm) and a small-sized (<25 μm) population and which dominated the caudal and ventral part of the nucleus. Parvicellular neurons (±20,000) were classified as large-sized (30–45 μm), medium-sized (15–30 μm), or small-sized (>15 μm) cells and may all be found throughout the nucleus. Although there is no consensus between different authors, it is agreed that the caudal pole harbors the largest neurons. Between 250 and 400 μm from the caudal pole, this large-celled part can be subdivided into a rather compact ventrolateral part and a more loosely arranged dorsomedial part (Reid et al., 1975b; see also Figs. 8A and 8B). The ventrolateral part is continuous with the so-called lateral horn of the rat red nucleus (Reid et al., 1975b) and is somewhat separated from the dorsomedial part, which appears to divide into a dorsal and a dorsomedial grouping of neurons. Rostral to this level no subdivisions are apparent. At least a part of the small neuronal population may represent local GABAergic interneurons, which have been demonstrated in turtle, cat, and monkey (VuillonCacciuttolo et al., 1984; Keifer et al., 1992; Ralston, 1994). Large neurons in the caudal two-thirds of the rat red nucleus frequently stain positive for calbinding D28k, whereas medium to large neurons in the rostral two-thirds frequently contain parvalbumin. Both populations, however, were strongly intermingled and double-labeled cells were only seldomly encountered (Hontanilla et al., 1995). The nucleus minimus deserves some comment. This aggregation of small cells lateral to the parvicellular part of the red nucleus was first described by Von Monakov (1909) in the rabbit and later identified in the cat by others (Brodal and Gogstad, 1954; Taber, 1961). In the rat, data are controversial. Reid et al. (1975b) did not find a nucleus minimus. However, they described the lateral horn as a subdivision of the red nucleus and as consisting of predominantly small- and mediumsized neurons. Faull and Carman (1978), who based their opinion on the pattern of terminal degeneration after superior cerebellar peduncle lesions, concluded that a nucleus minimus complying to Von Monakov’s description may be found intercalated between fascicles of the medial lemniscus. However, the position of this nucleus is rather similar to that of the lateral horn of Reid. Finally, Paxinos and Watson’s atlas of the rat brain (1986), based on Faull and Carman’s paper, delineates a nucleus minimus lateral to the dorsal part of the red nucleus, where it appears to be dorsal or dorsolateral from the lateral horn of Reid et al. Based on our own material of cerebellar nuclear projections to this area
(see below), we propose that the term nucleus minimus should be abandoned. Rather, we would advocate the term pararubral area, which forms an integral part of the parvicellular part of the red nucleus rostrally but becomes discernible as a more or less separated cluster of neurons located dorsolateral to its magnocellular part (Figs. 8H and 9: also see Fig. 6 of Ruigrok and Cella, 1995). As such, the pararubral area is located within the medial part of the nucleus reticularis subcuneiformis of Newman (1985). As mentioned earlier, the rostral boundary of the parvicellular red nucleus is difficult to establish. Based on the efferent connections of this area, it has been suggested by Kennedy (1987; Tucker et al., 1989) that the area directly surrounding the fasciculus retroflexus should be considered a part of the red nucleus. In rat, the area dorsomedial to this fiber bundle is recognized as (the rostral part of) the nucleus of Darkschewitsch and its ventrocaudal continuation as the nucleus accessorius medialis of Bechterew (?edial accessory oculomotor nucleus, MA3, of Paxinos and Watson, 1986). The rostral interstitial nucleus of the medial longitudinal fascicle and the prerubral field are situated ventral and ventrolateral to the fasciculus retroflexus, respectively (Paxinos and Watson, 1986). Carlton et al. (1982) designated the whole area surrounding the retroflex bundle as the nucleus parafascicularis prerubralis on the basis of its efferent connections. In this respect, it is noteworthy that in primates the parvicellular red nucleus is indented rostromedially by the retroflex bundle and extends rostrally to it. In human, the fasciculus retroflexus even traverses the parvicellular red nucleus completely (Paxinos and Huang, 1995), thus separating a dorsomedial part, which may be homologous to (part of) the nucleus parafascicularis prerubralis of the rat. It will be evident that extensive and detailed studies on the afferent as well as the efferent connections of these areas in the various animals are needed in order to establish whether areas with similar names indeed subserve similar functions.
Afferents to the Red Nucleus The red nucleus receives input from the cerebellar nuclei as well as from the cerebral cortex. Projections arising from the posterior thalamic nucleus (Roger and Cadusseau, 1987), zona incerta (Ricardo, 1981), hypothalamic areas, central pontine gray, nuclei raphe dorsalis and magnus, gigantoreticular nucleus, parvicellular reticular nucleus, parabrachial nuclei, and locus coeruleus (Bernays et al., 1988) have also been described. In cat, projections from the dorsal column nuclei and the spinal cord have been demonstrated to reach and terminate within the red nucleus (Boivie, 1988).
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FIGURE 8 Relation between the origin of the rubrospinal tract and the cerebellar nucleorubral connections. (A) Retrograde labeling of rubral neurons after WGA–HRP injection into the contralateral cervical spinal cord. (B) Retrograde labeling after WGA–HRP injection into the contralateral lumbar spinal cord. Note that the labeled cells are positioned ventral to those labeled in panel A. (C,E,G) Injection sites of the anterograde tracer PHA-L into the lateral part of the IntA, the medial part of the IntA, and the ventral, parvicellular part of the Lat, respectively. (D,F,H) Corresponding terminal labeling in the contralateral red nucleus. Note the the resultant labeling after a medial IntA injection (D) overlaps with the location of rubrospinal neurons that project to the lumbar cord, whereas the lateral IntA projects to the dorsal red nucleus (F) where the rubrocervical neurons are found. The parvicellular Lat gives rise to a conspicuous terminal labeling in the pararubral area (small arrows in panel H, also noted labeled, nonterminal, fibers within confines of the red nucleus). Open arrow in panels A, B, D, F, and H, idicates the lateral horn. Bar equals 200 μm (Ruigrok, unpublished results).
Various studies employing degeneration (Caughell and Flumerfelt, 1977) and retrograde and/or anterograde techniques (Daniel et al., 1987; Angaut and Cicirata, 1988) have recognized that the projections from the contralateral cerebellar nuclei to the red nucleus in the rat are organized in a somatotopical fashion. Our
material of small injections into the cerebellar nuclei with the anterograde tracer PHA-L (Ruigrok and Voogd, 1990) confirms and extends these observations (Figs. 8C–8H). Massive projections arise from the anterior interposed nucleus (IntA); its lateral part projects to the dorsomedial part of the magnocellular
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red nucleus, whereas its medial part provides heavy terminal arborizations in its ventrolateral part and lateral horn. The projection from the IntA stems from coarse axons and terminates on somata and proximal dendrites of the magnocellular neurons (Flumerfelt, 1978, 1980; Dekker, 1981). The Lat provides a major input to especially the parvicellular part of the nucleus. Here, a topographical organization also can be recognized. The rostral part of the Lat projects rather ventromedially, whereas more dorsolateral areas of the parvicellular part are supplied by the caudal part of the Lat (Angaut and Cicirata, 1988). A conspicuous projection to the cell aggregate located lateral to the magnocellular red nucleus (pararubral area) was found to arise specifically from the parvicellular subdivision of the Lat (Ruigrok et al., 1993; Teune et al., 2000: see also Figs. 8G and 8H). Red nuclear projections from the posterior interposed nucleus are sparse and are found along its dorsomedial aspect. This cerebellar nucleus distributes terminals mainly to the medial accessory oculomotor nucleus and to the medial part of the nucleus parafascicularis prerubralis (Teune et al., 2000, however, cf. Daniel et al., 1987). A small but definite projection is found from the medial cerebellar nucleus to the medial part of the base of the lateral horn. It should be recognized that the projections from the medial, posterior interposed as well as from the lateral cerebellar nuclei consist of rather fine caliber fibers that terminate on intermediate and distal dendrites and are not as massive when compared to the projections from the anterior interposed nucleus (Caughell and Flumerfelt, 1977; see also Figs. 8C and 8F). It is suggested that all rubral projections from the cerebellum are collaterals from ongoing fibers that eventually terminate in the thalamus (Shinoda et al., 1988) and are excitatory (Toyama et al., 1970; Oka, 1988). Most likely they make use of an excitatory amino acid as neurotransmitter (Bernays et al., 1988; Giuffrida et al., 1993; Schwarz and Schmitz, 1997). In the cat, using physiological techniques, it was found that at least some of the nucleoolivary projection neurons (thought to be all GABAergic: De Zeeuw et al., 1989b, Fredette and Mugnaini, 1991) collateralize to the red nucleus and/or thalamus (McCrea et al., 1978; Andersson and Hesslow, 1987). However, so far this notion could not be corroborated by anatomical studies (Teune et al., 1994; Schwarz and Schmitz, 1997). The projections from the rat sensorimotor cortex are not as well documented as those arising from the cerebellar nuclei. Gwyn et al. (1974) using degeneration techniques, reported that the sensorimotor cortex provides projections to the parvicellular part of the red nucleus only. This was confirmed by Brown (1974a) who also noticed that the prerubral field and an area dorsolateral
to the red nucleus (i.e., the pararubral area) receive a particularly heavy cortical input (cf. projections from the sensorimotor cortex to the subcuneiform reticular nucleus, Newman et al., 1989). Physiological studies by Giuffrida et al. (1988a, 1988b), however, claim that both magno- and parvicellular parts of the red nucleus of the rat are controlled by cerebral as well as cerebellar influences. This agrees well with a study using retrograde transport of small injections of lectin coupled to colloidal gold in different parts of the red nucleus in the guinea pig (Giuffrida et al., 1991). Retrogradely labeled neurons in layer V of the agranular frontal cortex were found after injections into the parvi- and into the magnocellular part of the red nucleus. In a study employing the use of selective retrogradely transported tracers, Bernays et al. (1988) suggested that the cerebral afferents to the red nucleus make use of an excitatory amino acid as transmitter (also see Giuffrida et al., 1993). Cadusseau and Roger (1987, 1988) have described a conspicuous and essentially reciprocal projection from the posterior thalamic nucleus to the parvicellular part of the red nucleus in the rat. The posterior thalamic nucleus is positioned within the sub-pretectal area at the mesodiencephalic junction and appears to be involved in the assimilation of somatosensory and/or nociceptive information. In this regard it is interesting that the zona incerta, considered to be a highly integrative somatosensory center, also provides input to both the posterior thalamic nucleus and the parvicellular red nucleus (Ricardo, 1981; Watanabe and Kawana, 1982; Roger and Cadusseau, 1985; Roger and Cadusseau, 1987). A somatosensory input from the spinal cord to the red nucleus that bypasses the cerebellum and cerebral cortex has been reported in the cat (Wiberg and Blomqvist, 1984a, 1984b; Padel et al., 1986; Boivie, 1988; Rathelot and Padel, 1997; Steffens et al., 2000) and in the monkey (Wiberg et al., 1987; Kerr and Bishop, 1991). In the rat, evidence has been obtained that direct reciprocal connections between the red nucleus and the trigeminal complex may function in a potentially similar way (Godefroy et al., 1998). Finally, a serotonergic projection arising from the raphe magnus and the raphe dorsalis nuclei to the red nucleus as well as a noradrenergic projection has been demonstrated in rat and cat (Bosler et al., 1983; André et al., 1987; Bernays et al., 1988) and has been shown to exert a powerful modulatory action on the activity of rubral neurons (Schmied et al., 1991; Ciranna et al., 1996; Faherty et al., 1997; Licata et al., 1998, 2001).
Efferents of the Red Nucleus When examining the efferent connections of the red nucleus a distinction should be made in the efferents
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of its parvicellular part and in those of its magnocellular part. However, it has been pointed out that, contrary to the situation in primates, in the rat this distinction is not easily made. Various authors have pointed out that large neurons may be found well into the parvicellular region. Yet, in monkeys, available evidence indicates that a rather strict separation of the efferent connections of both subdivisions of the red nucleus exists. The magnocellular part gives rise to the crossed rubrospinal and rubrobulbar projection, whereas the parvicellular part gives rise to the central tegmental tract which descends to the ipsilateral inferior olivary complex. In the cat, a similar organization has been suggested, although here, as in the rat, a division of the red nucleus in a parvi- and magnocellular part is less clear. Obviously, the efferent connections of the rat red nucleus may give insight into its subdivisions. However, data on the projections arising from the red nucleus are confusing. Originally, only the caudal third of the nucleus with its giant- and large-sized neurons was thought to give rise to the rubrospinal tract (Gwyn, 1971; Flumerfelt and Gwyn, 1974; Murray and Gurule, 1979). Its dorsomedial part was found to project to the cervical enlargement, whereas its ventrolateral part connected with the lumbosacral cord (Figs. 8A and 8B). However, it has become clear that at least some parvicellular neurons also project to the spinal cord (Huisman et al., 1981, 1983; Shieh et al., 1983; Tucker et al., 1989; Kennedy, 1990). The rubrospinal tract descends dorsally within the contralateral lateral funiculus where it is separated from the substantia gelatinosa by a small spinocervical tract. It terminates at the base of the dorsal horn and intermediate regions of the ventral horn (Brown, 1974b). Rubrospinal terminals may synapse with both excitatory and inhibitory interneurons and a considerable number of fibers may send projections to the ipsilateral side of the cord (Antal et al., 1992). Rubrospinal projections are considered to be excitatory and most likely make use of either glutamate or aspartate as neurotransmitter (Benson et al., 1991) The parvicellular red nucleus is generally thought to provide a projection to the inferior olive. In the rat, using anterograde as well as retrograde techniques, such a projection has been described by Swenson and Castro (1983a, 1983b). However, Rutherford et al. (1984) and Carlton et al. (1982) specifically denied this projection and claimed that most olivary projections stem from the area surrounding the fasciculus retroflexus (i.e., their nucleus parafascicularis prerubralis: also see Figs. 5, 6, and and 9). Finally, Kennedy (1987), using a modified HRP visualization technique, claimed that many neurons not only in the parvicellular part but also in magnocellular part were (faintly) retrogradely
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labeled after inferior olivary injections. Initial studies by Kennedy (Tucker et al., 1989; Kennedy, 1990; Tucker and Kennedy, 1990), employing double retrograde labeling techniques with fluorescent tracers, suggested that the rubroolivary pathway, at least in part, may exist of collaterals of the rubrospinal pathway but these ideas could not be corroborated in more recent years. In analogy with the situation in monkey, Kennedy furthermore proposed that the nucleus parafascicularis prerubralis should be redefined as being a part of the parvicellular red nucleus. Although we can agree with Kennedy on this point, it should be appreciated that (A) the retrograde labeling as the result of inferior olive injections is considerably more abundant and intense in the area parafascicularis prerubralis as compared to the labeling of cells in the parvicellular part of the red nucleus, suggesting the origin of a dense projection to the inferior olive from the former area, and (B) the pararubral area, with its input from the ventral part of the Lat, should also be incorparated in the rubral complex (Figs. 8 and 9). No specific data on the efferent projections of this area have been published as yet. Although Newman (1985) claims that the subcuneiform reticular nucleus projects to the spinal cord, it can be seen from Fig. 8 that the large, retrogradely labeled cells located dorsolateral to the red nucleus do not seem to correspond to the area of anterogradely labeled fibers derived from the parvicellular part of the lateral cerebellar nucleus (compare Figs. 8A and 8B with Fig. 8H). In addition to the rubrospinal and rubroolivary pathways, the red nucleus also provides a crossed rubrobulbar projection, presumably via collaterals of the rubrospinal pathway, which terminate in various brain stem centers like the lateral part of the facial nuclei, the parvicellular reticular formation, the rostral part of the lateral reticular nucleus, the oral part of the spinal trigeminal nucleus, and the principal sensory trigeminal nucleus, the descending vestibular nucleus, and the dorsal column nuclei (Flumerfelt and Gwyn, 1974; Hinrichsen and Watson, 1983; Godefroy et al., 1998). As a fourth major terminal source, the red nucleus has been shown to give rise to a rubrocerebellar pathway (Huisman et al., 1983; Yarom et al., 1991). Presently no information is available on the special characteristics of the rubronuclear and/or rubrocortical projection patterns in the rat, but in the cat these projections have been reported to specifically target the IntA (Nakamura et al., 1987). Finally, rubrothalamic projections, in particular arising from the parvicellular part of the nucleus, have been described to terminate in the posterior thalamic nucleus (Roger and Cadusseau, 1987) and, in the cat, in the ventrolateral thalamic nucleus (Condé and Condé, 1980).
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Functional Considerations Obviously, many questions still remain unanswered concerning the function of the red nucleus. The, previously made, distinction between magno- and parvicellular subdivisions that serve different functions, may still be useful, but appears to be rather naive. Present knowledge suggests that the red nuclear area may comprise at least three populations of neurons. A caudally located, predominantly large-celled, group participates in the execution of motor behavior, it receives its main input from the anterior interposed nucleus of the cerebellum and gives rise to the rubrospinal tract. A more rostrally located cell group consisting of slightly smaller cells, possibly assisted by a group of small cells located lateral to the magnocellular subdivision, receives its main input from the cerebellar dentate, the sensorimotor cortex, and the posterior thalamic nucleus. In the rat, many of these parvicellular neurons appear to project to the spinal cord but may also project to the inferior olive. Finally, a rostrally located group of small neurons surrounding the fasciculus retroflexus (area
parafascicularis prerubralis) receives a massive input from the sensorimotor cortex, but also from the dentate and the posterior interposed nuclei and appears to project mainly to the infe
