Handbook of the Cerebellum and Cerebellar Disorders [1 ed.] 9789400713321, 9789400713338, 9789400714045

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Handbook of the Cerebellum and Cerebellar Disorders

Mario Manto • Donna L. Gruol Jeremy D. Schmahmann Noriyuki Koibuchi • Ferdinando Rossi Editors

Handbook of the Cerebellum and Cerebellar Disorders

With 545 Figures and 69 Tables

Editors Mario Manto Unite´ d’Etude du Mouvement (UEM) FNRS, Neurologie ULB Erasme Bruxelles, Belgium Jeremy D. Schmahmann Ataxia Unit, Cognitive and Behavioral Neurology Unit Department of Neurology Massachusetts General Hospital Harvard Medical School Boston, MA, USA

Donna L. Gruol Molecular and Integrative Neuroscience Department (MIND) The Scripps Research Institute California, CA, USA Noriyuki Koibuchi Department of Integrative Physiolgy Gunma University Graduate School of Medicine Maebashi, Gunma, Japan

Ferdinando Rossi Neuroscience Institute of the Cavalieri-Ottolenghi Foundation (NICO) University of Turin Orbassano, Turin, Italy

ISBN 978-94-007-1332-1 ISBN 978-94-007-1333-8 (eBook) ISBN 978-94-007-1404-5 (print and electronic bundle) DOI 10.1007/978-94-007-1333-8 Springer New York Heidelberg Dordrecht London Library of Congress Control Number: 2012942646 # Springer Science+Business Media Dordrecht 2013 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. Exempted from this legal reservation are brief excerpts in connection with reviews or scholarly analysis or material supplied specifically for the purpose of being entered and executed on a computer system, for exclusive use by the purchaser of the work. Duplication of this publication or parts thereof is permitted only under the provisions of the Copyright Law of the Publisher’s location, in its current version, and permission for use must always be obtained from Springer. Permissions for use may be obtained through RightsLink at the Copyright Clearance Center. Violations are liable to prosecution under the respective Copyright Law. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. While the advice and information in this book are believed to be true and accurate at the date of publication, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Printed on acid-free paper Springer is part of Springer Science+Business Media (www.springer.com)

Foreword

Although research on the cerebellum has a long history of over two centuries, its advancement during the past five decades has been particularly rapid. An enormous amount of knowledge has been accumulated, forming a rich wealth of technological innovations, diverse refined data, novel concepts, and challenging hypotheses. Clearly, it is a timely endeavor to broadly review and reorganize the accumulated knowledge on a commonly understandable basis. This should be a very necessary step toward the full utilization of available outcomes of rigorous research thus far performed on the cerebellum and toward the effective focusing of our future research. It is my pleasure to welcome this great Handbook of the Cerebellum and Cerebellar Disorders as a compilation with such an overall aim. I am certain that it will play a pivotal role in promoting the entire research fields on the cerebellum. This handbook provides an authoritative survey of the experimental and theoretical studies performed in two core areas of cerebellar research. One core covers fundamental knowledge of the cerebellum at the molecular, cellular, neuronal circuit, developmental, and behavioral levels. It includes not only biological and experimental approaches but also modeling and computational approaches to the study of the cerebellum. The other core covers knowledge of disorders involving the cerebellum. This area will be applied in the near future to the development of breakthroughs in the so-far-difficult medical treatment of cerebellar diseases. The handbook embodies the current situation in which significant disparities between these two core areas of research on the cerebellum, which hampered their merging, have been diminished considerably. This handbook will no doubt facilitate the further merging of fundamental and medical knowledge of the cerebellum. The five editors (Mario Manto, Donna L. Gruol, Jeremy D. Schmahmann, Ferdinando Rossi, and Noriyuki Koibuchi) have masterly identified major phenomena, issues, and concepts of central importance in normal and diseased cerebella. They have chosen 106 topics to fill four volumes. Two thirds of these topics are on the fundamental knowledge and the other one third on knowledge of cerebellar disorders. Each of these topics is assigned to a qualified author(s) and is explained in terms of basic components such as genes, messengers, electrical/chemical signals, cellular processes, neuronal circuits, systems functions, theoretical models, mutations, animal models, and evolution.

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Foreword

Among such diverse topics, the degree of establishment and the depth of refinement of concepts could vary, and some might be debated among contemporaries. I take such a variety as a feature of a rapidly expanding research field, in which new research technologies are developed to enable novel observations and in which hypothesis-guided approaches play leading roles. Hence, in this handbook, the readers will find not only an impressive array of new knowledge but also dynamic perspectives of ever-advancing research fields on the cerebellum. Masao Ito Riken Institute Japan

Preface

The cerebellum has long attracted a core group of scientists intrigued by the sophistication of its circuitry, its unique geometric arrangement and developmental biology, and its characteristic clinical manifestations. With the advances in genetic studies, the rising awareness of the roles of the cerebellum in the nonmotor domain, and the profusion of brain imaging techniques that have generated a vast amount of new knowledge revealing novel aspects of cerebellar function, the field of cerebellar neurobiology has expanded rapidly. Large communities of scholars now setting out on their own paths of scientific enquiry are keenly interested in the cerebellum and its multiple roles in nervous system function. The evolution, and in some instances revolution, in knowledge of the cerebellum has sparked new fields of enquiry and attracted new schools of thought and legions of new investigators. The motivating goal of this comprehensive text therefore was to assemble an international panel of experts who could summarize the state of the art of the many facets of cerebellar clinical and basic neuroscience, and incorporate the most recent developments in the field. There are several excellent books on the neurobiology and clinical neurology of the cerebellum, but until the present volume there has been no single comprehensive work that can serve as an in-depth authoritative resource for the international community of scientists, clinicians, and other professionals interested in the science of the cerebellum. The Handbook of the Cerebellum and Cerebellar Disorders has been in preparation for over 2 years. This detailed work required the contributions of an international panel of renowned scientists and clinicians with experience in a diverse array of fields of neuroscience who were invited to write chapters that provide synthesis, analysis, and interpretation of both the historical and contemporary literature. This handbook could not have been completed without their considerable efforts, and we gratefully acknowledge their commitment to the project. We would like to recognize the staff at Springer who provided excellent service throughout this project. We particularly wish to acknowledge Ann Avouris, Martijn Roelandse, Somodatta Roy, Namita Mathur, Mansi Seth, and Vasuki Ravichandran for their input, assistance, constant support, and high degree of professionalism. They have been invaluable in helping to bring this work to completion. In addition to the printed version, we have arranged with Springer that the handbook be made available electronically on the Springer website. The reader may find that the ebook format is more accessible and that it facilitates searches more readily. vii

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Preface

The editors have attempted to cover what we regard as essential material, while striving to avoid redundancy. In the belief that this volume may be useful to the scientific and clinical communities, we plan to produce future editions of this work, and we therefore invite suggestions and critique in order to further strengthen this compilation, and perhaps include other authors and material that could serve to enhance the handbook and draw attention to the increasingly vibrant field of the basic science and clinical neurology of the cerebellum. Mario Manto, Brussels, Belgium Donna L. Gruol, La Jolla, USA Jeremy D. Schmahmann, Boston, USA Noriyuki Koibuchi, Gunma, Japan Ferdinando Rossi, Turin, Italy

Biographical Sketch of the Editors

Mario Manto, M.D. (1992), Ph.D. (1996), is a neurologist. He is Researcher at the FNRS (ULB)-Belgium. He is the founding and current Editor of the international journal The Cerebellum (Springer). He has also founded the Society for Research on the Cerebellum (www.socrecer.org). His research studies are focused on the pathogenesis of cerebellar disorders and have been funded by national and international research organizations: FNRS (Belgium), European Commission (FP5, FP6), NIH (USA). Works carried out by him have been published in peerreviewed journals. He serves as reviewer for more than 30 international journals. Donna L. Gruol, Ph.D., is Associate Professor in the Department of Molecular and Integrative Neuroscience at the Scripps Research Institute and adjunct Associate Professor of the Neuroscience Department at the University of California at San Diego. She obtained a Ph.D. from the Illinois Institute of Technology and did postdoctoral training at the University of Maryland Medical School, The National Institutes of Health, and The Salk Institute. She has been a member of several NIH grant review panels and has served on journal editorial boards and advisory committees. Her current research focuses on neuroadaptive changes in CNS neurophysiology produced by neuroinflammation. Jeremy D. Schmahmann is Professor of Neurology at Harvard Medical School and Massachusetts General Hospital. He is Director of the Ataxia Unit and the Laboratory for Neuroanatomy and Cerebellar Neurobiology, and a member of the Cognitive and Behavioral Neurology Unit at Massachusetts General Hospital. He trained at the University of Cape Town Medical School, the Neurological Unit of Boston City Hospital, and the Department of Anatomy and Neurobiology at Boston University. He received the Norman Geschwind Prize from the American Academy of Neurology and the Behavioral Neurology Society, and the Distinguished Neurology Teacher Award from the American Neurological Association. He is a Fellow of the American Academy of Neurology and the American Neuropsychiatric Association, on the scientific advisory board of the National Ataxia Foundation, and is cited in The Best Doctors in America since 1996. His research and clinical efforts are focused on the neuroanatomical substrates of cognition, and the role of

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Biographical Sketch of the Editors

the cerebellum in intellect and emotion. His other books include The Cerebellum and Cognition (Academic Press), MRI Atlas of the Human Cerebellum (Academic Press), Fiber Pathways of the Brain (Oxford University Press), and Cerebellar Disorders in Children (MacKeith Press). Noriyuki Koibuchi, M.D., Ph.D., obtained an M.D. degree from Gunma University School of Medicine in 1985 and a Ph.D. degree from Institute of Endocrinology, Gunma University, in 1989. Then he had a postdoctoral training at the Rockefeller University, New York. After serving as an Assistant Professor of Physiology at Dokkyo University School of Medicine and a Visiting Assistant Professor of Medicine at Harvard Medical School, he became a Professor of Integrative Physiology, Gunma University Graduate School of Medicine, Japan, in 2001. His major research interest is hormonal regulation of cerebellar development and plasticity. Ferdinando Rossi obtained M.D. (1985) and Ph.D. degrees in Neuroscience (1990) at the University of Turin, Italy. He has been Assistant Professor of Human Physiology (1990–1998), Associate Professor of Neurobiology (1998–1998), and Full Professor of Neuroscience (1999-today) at the Department of Neuroscience, University of Turin. He spent 2 years on sabbatical at the INSERM U-106 (Paris). He is now Director of the Neuroscience Institute of Turin and Dean of the Faculty of Psychology. He is Associate Editor of the European Journal of Neuroscience; member of the editorial board of Neuroscience, The Cerebellum, Neurobiology of Disease, and Frontiers in Neurosciences. His main research interests are focused on the mechanisms of cerebellar development, axonal regeneration, and cell replacement following CNS injury, activity/experience-dependent plasticity, and repair in the CNS.

Contents

Volume 1 Section 1

Cerebellar Development . . . . . . . . . . . . . . . . . . . . . . . . . .

1

1

Specification of the Cerebellar Territory . . . . . . . . . . . . . . . . . . . Marion Wassef

3

2

Proneural Genes and Cerebellar Neurogenesis in the Ventricular Zone and Upper Rhombic Lip . . . . . . . . . . . . . . . . . G. Giacomo Consalez, Marta Florio, Luca Massimino, and Laura Croci

23

3

Zones and Stripes: Development of Cerebellar Topography . . . . Roy V. Sillitoe and Richard Hawkes

43

4

Roof Plate in Cerebellar Neurogenesis . . . . . . . . . . . . . . . . . . . . . Victor V. Chizhikov

61

5

Specification of Cerebellar and Precerebellar Neurons . . . . . . . . Mikio Hoshino, Yusuke Seto, and Mayumi Yamada

75

6

Specification of Granule Cells and Purkinje Cells . . . . . . . . . . . . Thomas Butts, Leigh Wilson, and Richard J. T. Wingate

89

7

Granule Cell Migration and Differentiation . . . . . . . . . . . . . . . . Yutaro Komuro, Jennifer K. Fahrion, Kathryn D. Foote, Kathleen B. Fenner, Tatsuro Kumada, Nobuhiko Ohno, and Hitoshi Komuro

107

8

Analysis of Gene Networks in Cerebellar Development John Oberdick

.......

127

9

Purkinje Cell Migration and Differentiation . . . . . . . . . . . . . . . . Constantino Sotelo and Ferdinando Rossi

147

10

Development of Cerebellar Nuclei . . . . . . . . . . . . . . . . . . . . . . . . Gina E. Elsen, Gordana Juric-Sekhar, Ray A. M. Daza, and Robert F. Hevner

179

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Contents

11

Specification and Development of GABAergic Interneurons Karl Schilling

...

207

12

Development of Glutamatergic and GABAergic Synapses . . . . . Marco Sassoe`-Pognetto and Annarita Patrizi

237

13

Synaptic Remodeling and Neosynaptogenesis . . . . . . . . . . . . . . . Ann M. Lohof, Mathieu Letellier, Jean Mariani, and Rachel M. Sherrard

257

14

Synaptogenesis and Synapse Elimination . . . . . . . . . . . . . . . . . . . Masanobu Kano and Masahiko Watanabe

281

15

Genes and Cell Type Specification in Cerebellar Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Matt Larouche and Daniel Goldowitz

301

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

319

16

Hormones and Cerebellar Development Noriyuki Koibuchi and Yayoi Ikeda

Section 2

Anatomy, Connections and Neuroimaging of the Cerebellum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

341

17

Vascular Supply and Territories of the Cerebellum . . . . . . . . . . Louis Caplan

343

18

Vestibulocerebellar Connections . . . . . . . . . . . . . . . . . . . . . . . . . Neal H. Barmack and Vadim Yakhnitsa

357

19

Cerebellar Nuclei and the Inferior Olivary Nuclei: Organization and Connections . . . . . . . . . . . . . . . . . . . . . . . . . . . Jan Voogd, Yoshikazu Shinoda, Tom J. H. Ruigrok, and Izumi Sugihara

20

21

22

23

377

Axonal Trajectories of Single Climbing and Mossy Fiber Neurons in the Cerebellar Cortex and Nucleus . . . . . . . . . . . . . . Yoshikazu Shinoda and Izumi Sugihara

437

Visual Circuits from Cerebral Cortex to Cerebellum; The Link Through Pons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Mitchell Glickstein

469

Cerebellar Connections with Limbic Circuits: Anatomy and Functional Implications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gene J. Blatt, Adrian L. Oblak, and Jeremy D. Schmahmann

479

Cerebellar Influences on Descending Spinal Motor Systems . . . . Tom J. H. Ruigrok

497

Contents

xiii

24

Cerebellar Thalamic and Thalamocortical Projections . . . . . . . . Sharleen T. Sakai

25

Cerebellar Outputs in Non-human Primates: An Anatomical Perspective Using Transsynaptic Tracers . . . . . . . . . . . . . . . . . . Andreea C. Bostan and Peter L. Strick

549

Delineation of Cerebrocerebellar Networks with MRI Measures of Functional and Structural Connectivity . . . . . . . . . . . . . . . . . . Christophe Habas, William R. Shirer, and Michael D. Greicius

571

Radiographic Features of Cerebellar Disease: Imaging Approach to Differential Diagnosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . O. Rapalino, Robert Chen, and R. G. Gonzalez

587

26

27

28

Imaging Vascular Anatomy and Pathology of The Posterior Fossa . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Zeshan A. Chaudhry, Ronil V. Chandra, R. Gilberto Gonza´lez, and Albert J. Yoo

29

MR Spectroscopy in Health and Disease . . . . . . . . . . . . . . . . . . . ¨z G€ ulin O

30

Functional Topography of the Human Cerebellum Revealed by Functional Neuroimaging Studies . . . . . . . . . . . . . . . . . . . . . . Catherine J. Stoodley, John E. Desmond, and Jeremy D. Schmahmann

529

679

713

735

Volume 2 Section 3

Neurotransmission, Neuromodulation, Physiology

...

765

31

Cerebellar Granule Cell . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Egidio D’Angelo

767

32

Purkinje Neurons: Synaptic Plasticy . . . . . . . . . . . . . . . . . . . . . . Herve´ Daniel and F. Crepel

793

33

Stellate Cells: Synaptic Processing and Plasticity . . . . . . . . . . . . Siqiong June Liu

809

34

Golgi Neurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Katarzyna Pietrajtis and Ste´phane Dieudonne´

829

35

Glutamate Receptor Auxiliary Subunits and Interacting Protein Partners in the Cerebellum . . . . . . . . . . . . . . . . . . . . . . . Ian D. Coombs and Stuart G. Cull-Candy

36

GABA and Synaptic Transmission in the Cerebellum . . . . . . . . . Tomoo Hirano

853 881

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Contents

37

Norepinephrine and Synaptic Transmission in the Cerebellum . . . Daniel J. Chandler, Shevon E. Nicholson, Gerard Zitnik, and Barry D. Waterhouse

895

38

Serotonin and Synaptic Transmission in the Cerebellum . . . . . . Fumihito Saitow, Moritoshi Hirono, and Hidenori Suzuki

915

39

Cannabinoids and Synaptic Transmission in the Cerebellum . . . Michael H. Myoga and Wade G. Regehr

927

40

Purinergic Signaling in the Cerebellum . . . . . . . . . . . . . . . . . . . . Mark J. Wall and Boris P. Klyuch

947

41

Modulatory Role of Neuropeptides in the Cerebellum Georgia A. Bishop and James S. King

........

971

42

Neurosteroids and Synaptic Formation in the Cerebellum . . . . . Kazuyoshi Tsutsui

993

43

Inferior Olive: All Ins and Outs . . . . . . . . . . . . . . . . . . . . . . . . . . 1013 J. R. De Gruijl, L. W. J. Bosman, Chris I. De Zeeuw, and M. T. G. De Jeu

44

Dynamics of the Inferior Olive Oscillator and Cerebellar Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1059 Alexandre Mathy and Beverley A. Clark

45

Feedback Control in the Olivo-Cerebellar Loop . . . . . . . . . . . . . 1079 Fredrik Bengtsson and Germund Hesslow

46

Neurons of the Deep Cerebellar Nuclei . . . . . . . . . . . . . . . . . . . . 1101 Marylka Yoe Uusisaari and Thomas Kn€ opfel

47

Cerebellar Nuclei and Cerebellar Learning . . . . . . . . . . . . . . . . . 1111 Dieter Jaeger

48

Cerebro-Cerebellar Connections . . . . . . . . . . . . . . . . . . . . . . . . . 1131 Richard Apps and Thomas C. Watson

49

Cerebellar Control of Eye Movements . . . . . . . . . . . . . . . . . . . . . 1155 Pablo M. Bla´zquez and Angel M. Pastor

50

Cerebellum and Eyeblink Conditioning . . . . . . . . . . . . . . . . . . . . 1175 Derick H. Lindquist, Joseph E. Steinmetz, and Richard F. Thompson

51

Cerebellar Control of Speech and Song . . . . . . . . . . . . . . . . . . . . 1191 Daniel E. Callan and Mario Manto

52

Cerebellum and Timing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1201 Rebecca M. C. Spencer and Richard B. Ivry

Contents

xv

. . . . . . . . . . . . . . . . . . . . . . . . . . . 1221

53

Cerebellar Control of Posture M. E. Ioffe

54

Cerebellum and Gravity: Altered Earth’s Gravity Perception Under Pathological Conditions and Response to Altered Gravity in Space . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1241 Elizabeth M. Sajdel-Sulkowska

55

Cerebellum-Like Structures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1257 Nathaniel B. Sawtell and Curtis C. Bell

Volume 3 Section 4

Computational Models of Cerebellar Function . . . . . . .

1279

56

Cerebellum and Internal Models . . . . . . . . . . . . . . . . . . . . . . . . . 1281 Timothy J. Ebner

57

State Estimation and the Cerebellum . . . . . . . . . . . . . . . . . . . . . . 1297 Robert M. Hardwick, Maria Dagioglou, and R. Chris Miall

58

Adaptive Filter Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1315 Paul Dean, Henrik J€ orntell, and John Porrill

59

Cerebellum and Human Evolution: A Comparative and Information Theory Perspective . . . . . . . . . . . . . . . . . . . . . . 1337 C. Huang and Robert E. Ricklefs

60

Computational Structure of the Cerebellar Molecular Layer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1359 James M. Bower

61

Recursive Genome Function of the Cerebellum: Geometric Unification of Neuroscience and Genomics . . . . . . . . . . . . . . . . . 1381 Andras J. Pellionisz, Roy Graham, Peter A. Pellionisz, and Jean-Claude Perez

Section 5

Animal Models to Study Cerebellar Function . . . . . . . .

1425

. . . . . . . . . . . . . . . . . . . . . . . . . . . 1427

62

Animal Models: An Overview Noriyuki Koibuchi

63

Cerebellar Development and Neurogenesis in Zebrafish . . . . . . . 1441 Jan Kaslin and Michael Brand

64

Teleost Fish . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1463 Takanori Ikenaga

65

Robotic Mouse . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1481 Emmanuelle Bitoun, Peter L. Oliver, and Kay E. Davies

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Contents

66

Lurcher Mouse . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1499 Jan Cendelı´n and Frantisˇek Vozˇeh

67

Tottering Mouse . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1521 Timothy J. Ebner and Gang Chen

68

Rolling Nagoya Mouse . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1541 Jaap J. Plomp and Arn M. J. M. van den Maagdenberg

69

Ataxic Syrian Hamster . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1563 Kenji Akita

70

Hemicerebellectomy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1579 Marco Molinari, Maria Teresa Viscomi, and Maria G. Leggio

Section 6

Symptoms of Cerebellar Disorders in Human . . . . . . . .

1595

71

Cerebellar Motor Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1597 Giuliana Grimaldi

72

Lesion-Symptom Mapping of the Human Cerebellum Dagmar Timmann, Michael K€ uper, Elke R. Gizewski, Beate Schoch, and Opher Donchin

73

Deficits of Grasping in Cerebellar Disorders . . . . . . . . . . . . . . . . 1657 Dennis A. Nowak, Dagmar Timmann-Braun, and Joachim Hermsd€ orfer

74

Ataxic Hemiparesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1669 Akiyuki Hiraga

75

Cerebellum and Cognition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1687 Maja Steinlin and Kevin Wingeier

76

Cerebellar Sequencing for Cognitive Processing . . . . . . . . . . . . . 1701 M. Molinari and M. G. Leggio

77

Cerebellar Cognitive Affective Syndrome and the Neuropsychiatry of the Cerebellum . . . . . . . . . . . . . . . . . . . . . . . 1717 Jeremy D. Schmahmann

78

Cerebellar Mutism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1753 Peter Marie¨n, Hyo Jung De Smet, Philippe Paquier, Peter P. De Deyn, and Jo Verhoeven

79

Human Cerebellum in Motivation and Emotion . . . . . . . . . . . . . 1771 Dennis J. L. G. Schutter

. . . . . . . . 1627

Contents

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Volume 4 Section 7

Cerebellar Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1783

80

Clinical Scales of Cerebellar Ataxias . . . . . . . . . . . . . . . . . . . . . . 1785 Katrin B€ urk

81

Approach to the Differential Diagnosis of Cerebellar Ataxias Francesc Palau and Carmen Espino´s

82

Cerebellar Malformations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1819 Ozlem Alkan, Osman Kizilkilic, and Tulin Yildirim

83

Consequences for Cerebellar Development of Very Premature Birth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1839 Matthew Allin

84

Cerebellar Agenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1855 Romina Romaniello and Renato Borgatti

85

Chiari Malformations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1873 Mario Manto and Herweh Christian

86

Dandy-Walker Malformations . . . . . . . . . . . . . . . . . . . . . . . . . . . 1887 George A. Alexiou and Neofytos Prodromou

87

Autism Spectrum Disorders and Ataxia . . . . . . . . . . . . . . . . . . . . 1895 S. Hossein Fatemi and Timothy D. Folsom

88

Cerebellum and Schizophrenia – The Cerebellum Volume Reduction Theory of Schizophrenia . . . . . . . . . . . . . . . . . . . . . . . 1907 Gaku Okugawa

89

Progressive Myoclonic Epilepsies . . . . . . . . . . . . . . . . . . . . . . . . . 1923 Benjamin Legros and Mary L. Zupanc

90

Cerebellar Stroke . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1959 Keun-Hwa Jung and Jae-Kyu Roh

91

Immune Diseases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1985 Marios Hadjivassiliou

92

Endocrine Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2009 Mario Manto

93

Infectious Diseases of the Posterior Fossa Mario Manto and Patrice Jissendi

94

Diagnosis of Neoplastic and Paraneoplastic Cerebellar Ataxia . . . 2039 Genevie`ve Demarquay and Je´roˆme Honnorat

. . . 1799

. . . . . . . . . . . . . . . . . . 2027

xviii

Contents

95

Posterior Fossa Trauma . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2055 Matthias Maschke, Maria M€ orsdorf, Dagmar Timmann, and Uwe Dietrich

96

Cerebellotoxic Agents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2079 Mario Manto

97

Multiple System Atrophy (MSA) . . . . . . . . . . . . . . . . . . . . . . . . . 2119 Gregor K. Wenning, Florian Krismer, and Sid Gilman

98

Idiopathic Late Onset Cerebellar Ataxia (ILOCA), and Cerebellar plus Syndrome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2143 Shoji Tsuji

99

Essential Tremor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2151 Elan D. Louis

100

Autosomal Recessive Cerebellar Ataxias . . . . . . . . . . . . . . . . . . . 2177 Anne Noreau, Nicolas Dupre´, Jean-Pierre Bouchard, Patrick A. Dion, and Guy A. Rouleau

101

Autosomal Dominant Spinocerebellar Ataxias and Episodic Ataxias . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2193 Franco Taroni, Luisa Chiapparini, and Caterina Mariotti

102

Mitochondrial Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2269 Stefano Di Donato, Daniele Marmolino, and Franco Taroni

103

X-Linked Ataxias Josef Finsterer

104

Neuropathology of Ataxias . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2327 Mitsunori Yamada

105

General Management of Cerebellar Disorders: An Overview . . . . 2349 Winfried Ilg and Dagmar Timmann

106

Novel Therapeutic Challenges in Cerebellar Diseases . . . . . . . . . 2371 Antoni Matilla-Duen˜as, Carme Serrano, Yerko Iva´novic, Ramiro Alvarez, Pilar Latorre, and David Genı´s

. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2313

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2397

List of Contributors

Kenji Akita Biomedical Institute, Research Center, Hayashibara Biochemical Laboratories, Inc., Naka-ku, Okayama, Japan George A. Alexiou Department of Neurosurgery, Children’s Hospital “Agia Sofia”, Holargos, Athens, Greece Ozlem Alkan Department of Radiology, Baskent University Medical School, Adana, Turkey Matthew Allin Department of Psychosis Studies, Biomedical Research Centre for Mental Health, Institute of Psychiatry and King’s College London, London, United Kingdom Ramiro Alvarez Neurodegeneration Unit, Neurology Service, University Hospital Germans Trias i Pujol (HUGTP), Badalona (Barcelona), Spain Richard Apps School of Physiology and Pharmacology, University of Bristol, Bristol, UK Neal H. Barmack Department of Physiology & Pharmacology, Oregon Health & Science University, Portland, OR, USA Curtis C. Bell Neurological Sciences Institute, Oregon Health and Science University, Beaverton, OR, USA Fredrik Bengtsson Department of Experimental Medical Science, Division for Neuroscience, University of Lund, Lund, Sweden Georgia A. Bishop Department of Neuroscience, The Ohio State University, Columbus, OH, USA Emmanuelle Bitoun Department of Physiology, Anatomy and Genetics, MRC Functional Genomics Unit, University of Oxford, Oxford, UK Gene J. Blatt Anatomy and Neurobiology, Boston University School of Medicine, Boston, MA, USA Pablo M. Bla´zquez Department of Otolaryngology, School of Medicine, Washington University, St. Louis, MO, USA

xix

xx

List of Contributors

Renato Borgatti Department of Child Neuropsychiatry and Neurorehabilitation, Scientific Institute “Eugenio Medea”, Bosisio Parini (LC), Italy L. W. J. Bosman Netherlands Institute for Neuroscience, Royal Dutch Academy of Arts & Sciences (KNAW), Amsterdam, The Netherlands Department of The Netherlands

Neuroscience,

Erasmus

Medical

Center,

Rotterdam,

Andreea C. Bostan Center for the Neural Basis of Cognition, Systems Neuroscience Institute and Department of Neurobiology, School of Medicine, University of Pittsburgh, Pittsburgh, PA, USA Jean-Pierre Bouchard Department of Neurological Sciences, Laval University CHAUQ (Enfant-Je´sus), Que´bec, Que´bec, Canada James M. Bower Barshop Institute to Longevity and Aging Studies, Department of Radiology, University of Texas Health Science Center, San Antonio, TX, USA Department of Biology, Neuroscience Institute, University of Texas, San Antonio, TX, USA Michael Brand Developmental Genetics, Biotechnology Center and Center for Regenerative Therapies Dresden, Dresden University of Technology, Dresden, Germany Katrin B€ urk Department of Neurology, Philipps University of Marburg, Marburg, Germany Thomas Butts MRC Centre for Developmental Neurobiology, King’s College, London, UK Daniel E. Callan Department of Computational Brain Imaging, Neural Information Analysis Laboratories ATR, Soraku-gun, Kyoto, Japan Louis Caplan Department of Neurology, Beth Israel Deaconess Medical Center, Boston, MA, USA Jan Cendelı´n Department of Pathophysiology, Faculty of Medicine in Pilsen, Charles University, Pilsen, Czech Republic Daniel J. Chandler Department of Neurobiology and Anatomy, Drexel University College of Medicine, Philadelphia, PA, USA Ronil V. Chandra Department of Interventional Neuroradiology and Endovascular Neurosurgery, Massachusetts General Hospital, Harvard Medical School, Boston, MA, USA Zeshan A. Chaudhry Department of Diagnostic Neuroradiology, Massachusetts General Hospital, Harvard Medical School, Boston, MA, USA Gang Chen Department of Neuroscience, University of Minnesota, Minneapolis, MN, USA

List of Contributors

xxi

Robert Chen Emergency Imaging Division, Department of Radiology, Massachusetts General Hospital, Boston, MA, USA Luisa Chiapparini Unit of Neuroradiology, Fondazione IRCCS Istituto Neurologico “Carlo Besta”, Milan, Italy Victor V. Chizhikov Center for Integrative Brain Research, Seattle Children’s Hospital Research Institute, Seattle, WA, USA Herweh Christian Department of Neuroradiology, University of Heidelberg, Medical Center, Heidelberg, Germany Beverley A. Clark Wolfson Institute for Biomedical Research, University College London, London, UK G. Giacomo Consalez Division of Neuroscience, San Raffaele Scientific Institute, Milan, Italy Ian D. Coombs Department of Neuroscience, Physiology and Pharmacology, University College London, London, UK F. Crepel Laboratoire de Pharmacologie et Biochimie de la synapse, CNRS UMR 8619, Institut de Biochimie et de Biophysique Mole´culaire et Cellulaire, Universite´ Paris-Sud 12, Orsay Cedex, France Laura Croci Division of Neuroscience, San Raffaele Scientific Institute, Milan, Italy Stuart G. Cull-Candy Department of Neuroscience, Physiology and Pharmacology, University College London, London, UK Maria Dagioglou Behavioural Brain Sciences, School of Psychology, University of Birmingham, Edgbaston, Birmingham, UK Egidio D’Angelo Department of Neuroscience, University of Pavia, Brain Connectivity Center, IRCCS C. Mondino, Pavia, Italy Herve´ Daniel Laboratoire de Pharmacologie et Biochimie de la synapse, CNRS UMR 8619, Institut de Biochimie et de Biophysique Mole´culaire et Cellulaire, Universite´ Paris-Sud 12, Orsay Cedex, France Kay E. Davies Department of Physiology, Anatomy and Genetics, MRC Functional Genomics Unit, University of Oxford, Oxford, UK Ray A. M. Daza Department of Neurological Surgery, Seattle Children’s Research Institute, Center for Integrative Brain Research, M/S C9S-10, Seattle, WA, USA Peter P. De Deyn Department of Neurology, ZNA Middelheim Hospital, Antwerp, Belgium Hyo Jung De Smet Department of Experimental Psychology, University of Ghent, Ghent, Belgium

xxii

List of Contributors

Paul Dean Department of Psychology, University of Sheffield, Sheffield, UK Genevie`ve Demarquay Centre de Re´fe´rence, de Diagnostic et de Traitement des Syndromes Neurologiques Parane´oplasiques, Hospices Civils de Lyon, Lyon, France John E. Desmond Department of Neurology, Johns Hopkins Medical School, Baltimore, MD, USA Stefano Di Donato Fondazione IRCCS Istituto Neurologico C., Milano, Italy Uwe Dietrich Department of Neuroradiology, Evangelisches Krankenhaus Bielefeld, Bielefeld, Germany Ste´phane Dieudonne´ Laboratoire de Neurobiologie, Inhibitory Transmission Team, IBENS, Ecole Normale Supe´rieure (CNRS UMR 8197; INSERM U 1024), Paris, France Patrick A. Dion Centre of Excellence in Neuroscience of Universite´ de Montre´al (CENUM), Centre de Recherche du Centre Hospitalier de l’Universite´ de Montre´al (CRCHUM), Montre´al, Que´bec, Canada Department of Pathology and cellular biology, Universite´ de Montre´al, Montre´al, Que´bec, Canada Opher Donchin Department of Biomedical Engineering and Zlotowski Center for Neuroscience, Ben-Gurion University of the Negev, Be’er Sheva, Israel Nicolas Dupre´ Department of Neurological Sciences, Laval University CHAUQ (Enfant-Je´sus), Que´bec, Que´bec, Canada Timothy J. Ebner Department of Neuroscience, University of Minnesota, Minneapolis, MN, USA Gina E. Elsen Department of Neurological Surgery, Seattle Children’s Research Institute, Center for Integrative Brain Research, M/S C9S-10, Seattle, WA, USA Carmen Espino´s Laboratory of Genetics and Molecular Medicine, Instituto de Biomedicina de Valencia, Consejo Superior de Investigaciones Cientı´ficas (CSIC), and CIBER de Enfermedades Raras (CIBERER), Valencia, Spain Jennifer K. Fahrion Department of Neurosciences, NC30, Lerner Research Institute, Cleveland Clinic Foundation, Cleveland, OH, USA S. Hossein Fatemi Department of Psychiatry, Division of Neuroscience Research, University of Minnesota, Minneapolis, MN, USA Departments of Pharmacology & Neuroscience, University of Minnesota, Minneapolis, MN, USA Kathleen B. Fenner Department of Neurosciences, NC30, Lerner Research Institute, Cleveland Clinic Foundation, Cleveland, OH, USA Josef Finsterer Danube University Krems, Vienna, Austria

List of Contributors

xxiii

Marta Florio Division of Neuroscience, San Raffaele Scientific Institute, Milan, Italy Timothy D. Folsom Department of Psychiatry, Division of Neuroscience Research, University of Minnesota, Minneapolis, MN, USA Kathryn D. Foote Department of Neurosciences, NC30, Lerner Research Institute, Cleveland Clinic Foundation, Cleveland, OH, USA David Genı´s Neurodegenerative Diseases Unit, University Hospital of Girona Dr. Josep Trueta, Girona, Spain Sid Gilman Department of Neurology, University of Michigan, Ann Arbor, MI, USA Elke R. Gizewski Departments of Neuroradiology, University of Duisburg-Essen and Justus-Liebig-Universit€at Gießen, Gießen, Germany Mitchell Glickstein Cell and Developmental Biology, University College London, London, UK Daniel Goldowitz Department of Medical Genetics, Child and Family Research Institute, Centre for Molecular Medicine and Therapeutics, University of British Columbia, Vancouver, BC, Canada R. Gilberto Gonza´lez Department of Diagnostic Neuroradiology, Massachusetts General Hospital, Harvard Medical School, Boston, MA, USA Roy Graham DRC Computer, Sunnyvale, CA, USA Michael D. Greicius Department of Neurology and Neurological Sciences, Functional Imaging in Neuropsychiatric Disorders (FIND) Lab, Stanford University School of Medicine, Stanford, CA, USA Giuliana Grimaldi Unite´ d0 Etude du Mouvement (UEM), Neurologie - ULB Erasme, Bruxelles, Belgium J. R. De Gruijl Netherlands Institute for Neuroscience, Royal Dutch Academy of Arts & Sciences (KNAW), Amsterdam, The Netherlands Christophe Habas Service de NeuroImagerie, CHNO des XV-XX, Universite´ Pierre et Marie Curie Paris 6, Paris, France Marios Hadjivassiliou Department of Neurology, Royal Hallamshire Hospital, Sheffield, UK Robert M. Hardwick Behavioural Brain Sciences, School of Psychology, University of Birmingham, Edgbaston, Birmingham, UK Richard Hawkes Department of Cell Biology and Anatomy Genes and Development Research Group, and Hotchkiss Brain Institute, The University of Calgary, Calgary, AB, Canada

xxiv

List of Contributors

Joachim Hermsd€ orfer Lehrstuhl f€ ur Bewegungswissenschaft, Fakult€at f€ur Sportund Gesundheitswissenschaft, Technische Universit€at M€unchen, Munich, Germany Germund Hesslow Department of Experimental Medical Science, Section for Neuroscience, University of Lund, Lund, Sweden Robert F. Hevner Department of Neurological Surgery, Seattle Children’s Research Institute, Center for Integrative Brain Research, M/S C9S-10, Seattle, WA, USA Akiyuki Hiraga Department of Neurology, Chiba Rosai Hospital, Ichihara-shi, Chiba, Japan Tomoo Hirano Department of Biophysics, Graduate School of Science, Kyoto University, Kyoto, Sakyo-ku, Japan Moritoshi Hirono Laboratory for Motor Learning Control, RIKEN Brain Science Institute, Saitama, Japan Je´roˆme Honnorat Centre de Re´fe´rence, de Diagnostic et de Traitement des Syndromes Neurologiques Parane´oplasiques, Hospices Civils de Lyon, Lyon, France Universite´ Claude Bernard Lyon 1, Lyon, France Neuro–Oncologie, Hoˆpital Neurologique, BRON Cedex, France Mikio Hoshino Department of Biochemistry and Cellular Biology, National Institute of Neuroscience, National Center of Neurology and Psychiatry, Kodaira, Tokyo, Japan C. Huang School of Biological Sciences, University of Missouri-Kansas City, Kansas City, MO, USA Yayoi Ikeda Department of Histology and Cell Biology, Yokohama City University School of Medicine, Yokohama, Japan Takanori Ikenaga Graduate School of Life Science, University of Hyogo, Ako-gun, Hyogo, Japan Winfried Ilg Section Computational Sensomotorics, Department of Cognitive Neurology, Hertie Institute for Clinical Brain Research, Centre for Integrative Neuroscience, University of T€ ubingen, T€ ubingen, Germany M. E. Ioffe Institute of Higher Nervous Activity and Neurophysiology, Russian Academy of Science, Moscow, Russia Yerko Iva´novic Monte Alto Rehabilitation Medical Center, (Madrid), Private Practice, Madrid, Spain National Reference Care Centre for People with Rare Diseases and Their Families– CREER–(Burgos), IMSERSO, Burgos, Spain

List of Contributors

xxv

Richard B. Ivry Department of Psychology, University of California, Berkeley, CA, USA Dieter Jaeger Department of Biology, Emory University, Atlanta, GA, USA M. T. G. De Jeu Department of Neuroscience, Erasmus Medical Center, Rotterdam, The Netherlands Patrice Jissendi Service de Neuroradiologie, ULB Erasme, Bruxelles, Belgium Henrik J€ orntell Section for Neurophysiology, Department of Experimental Medical Sciences, Lund University, Lund, Sweden Keun-Hwa Jung Department of Neurology, Seoul National University, Medical College, Seoul National University Hospital, Seoul, South Korea Gordana Juric-Sekhar Department of Pathology, Harborview Medical Center, Seattle, WA, USA Masanobu Kano Department of Neurophysiology, Graduate School of Medicine, The University of Tokyo, Bunkyo-ku, Tokyo, Japan Jan Kaslin Developmental Genetics, Biotechnology Center and Center for Regenerative Therapies Dresden, Dresden University of Technology, Dresden, Germany Australian Regenerative Medicine Institute (ARMI), Monash University, Melbourne, Australia James S. King Department of Neuroscience, The Ohio State University, Columbus, OH, USA Osman Kizilkilic Department of Radiology, Istanbul University Cerrahpasa Medical School, Istanbul, Turkey Boris P. Klyuch School of Life Sciences, University of Warwick, Coventry, UK Thomas Kn€ opfel Laboratory for Neuronal Circuit Dynamics, RIKEN Brain Science Institute, Saitama, Japan Noriyuki Koibuchi Department of Integrative Physiology, Gunma University Graduate School of Medicine, Maebashi, Gunma, Japan Hitoshi Komuro Department of Neurosciences, NC30, Lerner Research Institute, Cleveland Clinic Foundation, Cleveland, OH, USA Yutaro Komuro Department of Neurosciences, NC30, Lerner Research Institute, Cleveland Clinic Foundation, Cleveland, OH, USA Florian Krismer Division of Clinical Neurobiology, Department of Neurology, Medical University, Innsbruck, Austria Michael K€ uper Department of Neurology, University of Duisburg-Essen, Essen, Germany

xxvi

List of Contributors

Tatsuro Kumada Department of Neurosciences, NC30, Lerner Research Institute, Cleveland Clinic Foundation, Cleveland, OH, USA Matt Larouche Department of Medical Genetics, Child and Family Research Institute, Centre for Molecular Medicine and Therapeutics, University of British Columbia, Vancouver, BC, Canada Pilar Latorre Neurodegeneration Unit, Neurology Service, University Hospital Germans Trias i Pujol (HUGTP), Badalona (Barcelona), Spain Maria G. Leggio Neurorehabilitation Unit A – Ataxia Laboratory, I.R.C.C.S. Santa Lucia Foundation, Rome, Italy Department of Psychology, University of Rome La Sapienza, Rome, Italy Benjamin Legros Department of Neurology; Reference Center for the Treatment of Refractory Epilepsy, Universite´ Libre de Bruxelles- Hoˆpital Erasme, Brussels, Belgium Mathieu Letellier Centre National de la Recherche Scientifique, Universite´ Pierre et Marie Curie–Paris6, Paris, France MRC Laboratory for Molecular Cell Biology and Cell Biology Unit and Department of Neuroscience, Physiology and Pharmacology, University College London, London, United Kingdom Derick H. Lindquist Department of Psychology, The Ohio State University, Columbus, OH, USA Siqiong June Liu Department of Cell Biology and Anatomy, LSU Health Sciences Center Medical Education Building LSUHSC, New Orleans, LA, USA Ann M. Lohof Centre National de la Recherche Scientifique, Universite´ Pierre et Marie Curie-Paris6, Paris, France Elan D. Louis GH Sergievsky Center, College of Physicians and Surgeons, Columbia University, New York, NY, USA Department of Neurology, College of Physicians and Surgeons, Columbia University, New York, NY, USA Taub Institute for Research on Alzheimer’s Disease and the Aging Brain, College of Physicians and Surgeons, Columbia University, New York, NY, USA Department of Epidemiology, Mailman School of Public Health, Columbia University, New York, NY, USA Unit 198, Neurological Institute, New York, NY, USA Arn M. J. M. van den Maagdenberg Departments of Neurology and Human Genetics, Leiden University Medical Centre, Leiden, The Netherlands

List of Contributors

xxvii

Mario Manto Unite´ d’Etude du Mouvement (UEM), FNRS, Neurologie ULB Erasme, Bruxelles, Belgium Jean Mariani Centre National de la Recherche Scientifique, Universite´ Pierre et Marie Curie–Paris6, Paris, France Assistance Publique–Hoˆpitaux de Paris, Hoˆpital Charles Foix, Unite´ d’Exploration Fonctionnelles, Ivry–sur–Seine, France Peter Marie¨n Department of Neurology, ZNA Middelheim Hospital, Antwerp, Belgium Department of Clinical Neurolinguistics, Vrije Universiteit Brussel, Brussels, Belgium Caterina Mariotti Department of Diagnostics and Applied Technology, Unit of Genetics of Neurodegenerative and Metabolic Disease, Fondazione IRCCS Istituto Neurologico “Carlo Besta”, Milan, Italy Daniele Marmolino Laboratoire de Neurologie expe´rimentale, Universite´ Libre de Bruxeles (ULB), Bruxelles, Belgium Matthias Maschke Department of Neurology, Krankenhaus der Barmherzigen Br€ uder, Trier, Germany Luca Massimino Division of Neuroscience, San Raffaele Scientific Institute, Milan, Italy Alexandre Mathy Wolfson Institute for Biomedical Research, University College London, London, UK Antoni Matilla-Duen˜as Department of Neurosciences, Basic, Translational and Molecular Neurogenetics Research Unit, Health Sciences Research Institute Germans Trias I Pujol (IGTP), Universitat Auto`noma de Barcelona, Badalona (Barcelona), Spain R. Chris Miall Behavioural Brain Sciences, School of Psychology, University of Birmingham, Edgbaston, Birmingham, UK Marco Molinari Laboratory of Experimental Neurorehabilitation Unit A – Ataxia Laboratory, I.R.C.C.S. Santa Lucia Foundation, Rome, Italy Maria M€ orsdorf Department of Neuroradiology, Bruederkrankenhaus Trier, Trier, Germany Michael H. Myoga Department of Neurobiology, Harvard Medical School, Boston, MA, USA Shevon E. Nicholson Department of Neurobiology and Anatomy, Drexel University College of Medicine, Philadelphia, PA, USA

xxviii

List of Contributors

Anne Noreau Centre of Excellence in Neuroscience of Universite´ de Montre´al (CENUM), Centre de Recherche du Centre Hospitalier de l’Universite´ de Montre´al (CRCHUM), Montre´al, Que´bec, Canada Dennis A. Nowak Klinik Kipfenberg, Neurologische Fachklinik, Kipfenberg, Germany Neurologische Universit€atsklinik, der Philipps–Universit€at, Marburg John Oberdick Department of Neuroscience & Center for Molecular Neurobiology, The Ohio State University, Columbus, OH, USA Adrian L. Oblak Anatomy and Neurobiology, Boston University School of Medicine, Boston, MA, USA Nobuhiko Ohno Department of Neurosciences, NC30, Lerner Research Institute, Cleveland Clinic Foundation, Cleveland, OH, USA Gaku Okugawa Department of Neuropsychiatry, Kansai Medical University, Hirakata, Osaka, Japan Peter L. Oliver Department of Physiology, Anatomy and Genetics, MRC Functional Genomics Unit, University of Oxford, Oxford, UK ¨ z Center for Magnetic Resonance Research, Department of Radiology, G€ ulin O Medical School, University of Minnesota, Minneapolis, MN, USA Francesc Palau Laboratory of Genetics and Molecular Medicine, Instituto de Biomedicina de Valencia, Consejo Superior de Investigaciones Cientı´ficas (CSIC), and CIBER de Enfermedades Raras (CIBERER), Valencia, Spain Philippe Paquier Department of Clinical Neurolinguistics, Vrije Universiteit Brussel, Brussels, Belgium Department of Neurology and Neuropsychology, University Hospital Erasme, ULB, Brussels, Belgium Unit of Neurosciences, School of Medicine, Antwerp University, Antwerp, Belgium Angel M. Pastor Departamento de Fisiologı´a, Universidad de Sevilla, Sevilla, Spain Annarita Patrizi Department of Anatomy, Pharmacology, and Forensic Medicine, National Institute of Neuroscience–Italy, Turin, Italy F.M. Kirby Neurobiology Center, Children’s Hospital, Harvard Medical School, Boston, MA, USA Peter A. Pellionisz UCLA, Westwood, CA, USA Andras J. Pellionisz HolGenTech, Sunnyvale, CA, USA Jean-Claude Perez IBM Emeritus, Martignas, France

List of Contributors

xxix

Katarzyna Pietrajtis Laboratoire de Neurobiologie, Inhibitory Transmission Team, IBENS, Ecole Normale Supe´rieure (CNRS UMR 8197; INSERM U 1024), Paris, France Jaap J. Plomp Departments of Neurology and Molecular Cell Biology – Group Neurophysiology, Leiden University Medical Centre, Leiden, The Netherlands John Porrill Department of Psychology, University of Sheffield, Sheffield, UK Neofytos Prodromou Department of “Agia Sofia”, Holargos, Athens, Greece

Neurosurgery,

Children’s

Hospital

O. Rapalino Neuroradiology Division, Department of Radiology, Massachusetts General Hospital, Boston, MA, USA Wade G. Regehr Department of Neurobiology, Harvard Medical School, Boston, MA, USA Robert E. Ricklefs University of Missouri-St. Louis, St. Louis, MO, USA Jae-Kyu Roh Department of Neurology, Seoul National University, Medical College, Seoul National University Hospital, Seoul, South Korea Romina Romaniello Department of Child Neuropsychiatry and Neurorehabilitation, Scientific Institute “Eugenio Medea”, Bosisio Parini (LC), Italy Ferdinando Rossi Neuroscience Institute of Turin (NIT), Department of Neuroscience, University of Turin, Turin, Italy Neuroscience Institute of the Cavalieri–Ottolenghi Foundation (NICO), University of Turin, Orbassano, Turin, Italy Guy A. Rouleau Centre of Excellence in Neuroscience of Universite´ de Montre´al (CENUM), Centre de Recherche du Centre Hospitalier de l’Universite´ de Montre´al (CRCHUM), Montre´al, Que´bec, Canada Research Center CHU Ste–Justine, and Department of Pediatrics and Biochemistry, University of Montreal, Montre´al, Que´bec, Canada Research Center CHU Ste–Justine, and Department of Pediatrics and Biochemistry, CHUM Research Centre, Montre´al, Que´bec, Canada Tom J. H. Ruigrok Department of Neuroscience, Erasmus Medical Center Rotterdam, Rotterdam, The Netherlands Fumihito Saitow Department of Pharmacology, Nippon Medical School, Tokyo, Japan Japan Science and Technology Agency, CREST, Tokyo, Japan Elizabeth M. Sajdel-Sulkowska Department of Psychiatry, Harvard Medical School and Brigham and Women’s Hospital, Harvard Institute of Medicine, Rm. 921, Boston, MA, USA

xxx

List of Contributors

Sharleen T. Sakai Department of Psychology and Neuroscience Program, Michigan State University, East Lansing, MI, USA Marco Sassoe`-Pognetto Department of Anatomy, Pharmacology, and Forensic Medicine, National Institute of Neuroscience-Italy, Turin, Italy Nathaniel B. Sawtell Department of Neuroscience and Kavli Institute for Brain Science, Hammer Health Sciences Center, Room 510C Columbia University Medical Center, New York, NY, USA Karl Schilling Anatomisches Institut – Anatomie und Zellbiologie, Rheinische Friedrich-Wilhelms-Universit€at, Bonn, Germany Jeremy D. Schmahmann Ataxia Unit, Cognitive and Behavioral Neurology Unit, Department of Neurology, Massachusetts General Hospital, Harvard Medical School, Boston, MA, USA Beate Schoch Departments of Neurosurgery, University of Duisburg-Essen and Stiftungsklinikum Mittelrhein GmbH, Koblenz, Germany Dennis J. L. G. Schutter Department of Experimental Psychology, Faculty of Social Sciences, Utrecht University, Utrecht, The Netherlands Carme Serrano Neurology Service, Hospital de Martorell, Barcelona, Spain Yusuke Seto Integrative Bioscience and Biomedical Engineering, Graduate School of Science and Engineering, Waseda University, Shinjuku-ku, Tokyo, Japan Rachel M. Sherrard Centre National de la Recherche Scientifique, Universite´ Pierre et Marie Curie-Paris6, Paris, France Yoshikazu Shinoda Department of Systems Neurophysiology, Graduate School of Medicine, Tokyo Medical and Dental University, Bunkyo-ku, Tokyo, Japan William R. Shirer Department of Neurology and Neurological Sciences, Functional Imaging in Neuropsychiatric Disorders (FIND) Lab, Stanford University School of Medicine, Stanford, CA, USA Roy V. Sillitoe Dominick P. Purpura Department of Neuroscience, Albert Einstein College of Medicine 812 Kennedy Center, Bronx, NY, USA Constantino Sotelo Neurociences Institute, Miguel Hernandez University and CSIC, Alicante, Sant Joand’Alacant, Spain INSERM, U968, Paris, France UPMC Univ Paris 06, UMR_S 968, Institut de la Vision, Paris, France CNRS, UMR_7210, Paris, France Rebecca M. C. Spencer Department of Psychology, University of Massachusetts, Amherst, MA, USA

List of Contributors

xxxi

Maja Steinlin Neuropaediatrics, University Children’s Hospital Inselspital, Bern, Switzerland Joseph E. Steinmetz Department of Psychology, The Ohio State University, Columbus, OH, USA Catherine J. Stoodley Department of Psychology, College of Arts and Sciences, American University, Washington, DC, USA Peter L. Strick Pittsburgh Veterans Affairs Medical Center, Pittsburgh, PA, USA Center for the Neural Basis of Cognition, Systems Neuroscience Institute and Department of Neurobiology, School of Medicine, University of Pittsburgh, Pittsburgh, PA, USA Izumi Sugihara Department of Systems Neurophysiology, Graduate School of Medicine, Tokyo Medical and Dental University, Bunkyo-ku, Tokyo, Japan Hidenori Suzuki Department of Pharmacology, Nippon Medical School, Tokyo, Japan Japan Science and Technology Agency, CREST, Tokyo, Japan Franco Taroni Fondazione IRCCS Istituto Neurologico C., Milano, Italy Department of Diagnostics and Applied Technology, Unit of Genetics of Neurodegenerative and Metabolic Diseases Istituto Neurologico “Carlo Besta”, Milan, Italy Richard F. Thompson Department of Psychology, University of Southern California, Los Angeles, CA, USA Dagmar Timmann Department of Neurology, University of Duisburg-Essen, Essen, Germany Shoji Tsuji Department of Neurology, Graduate School of Medicine, The University of Tokyo, Tokyo, Japan Kazuyoshi Tsutsui Laboratory of Integrative Brain Sciences, Department of Biology, Waseda University, and Center for Medical Life Science of Waseda University, Shinjuku-ku, Tokyo, Japan Marylka Yoe Uusisaari Laboratory for Neuronal Circuit Dynamics, RIKEN Brain Science Institute, Saitama, Japan Theoretical and Experimental Neurobiology Unit, Okinawa Institute of Science and Technology (OIST), Onna–Son, Okinawa, Japan Jo Verhoeven Department of Language and Communication Science, City University London, London, UK Maria Teresa Viscomi Laboratory of Experimental Neurorehabilitation, I.R.C.C.S. Santa Lucia Foundation, Rome, Italy

xxxii

List of Contributors

Jan Voogd Department of Neuroscience, Erasmus Medical Center Rotterdam, Rotterdam, The Netherlands Oegstgeest, The Netherlands Frantisˇek Vozˇeh Department of Pathophysiology, Faculty of Medicine in Pilsen, Charles University, Pilsen, Czech Republic Mark J. Wall School of Life Sciences, University of Warwick, Coventry, UK Marion Wassef Institut de Biologie de l’Ecole Normale Supe´rieure (IBENS), Paris, France CNRS UMR 8197, Paris, France INSERM U1024, Paris, France Masahiko Watanabe Department of Anatomy, Hokkaido University Graduate School of Medicine, Sapporo, Japan Barry D. Waterhouse Department of Neurobiology and Anatomy, Drexel University College of Medicine, Philadelphia, PA, USA Thomas C. Watson Department of Pharmacology, University of Cambridge, Cambridge, UK Gregor K. Wenning Division of Clinical Neurobiology, Department of Neurology, Medical University, Innsbruck, Austria Leigh Wilson MRC Centre for Developmental Neurobiology, King’s College, London, UK Richard J. T. Wingate MRC Centre for Developmental Neurobiology, King’s College, London, UK Kevin Wingeier Neuropaediatrics, University Children’s Hospital Inselspital, Bern, Switzerland Vadim Yakhnitsa Department of Physiology & Pharmacology, Oregon Health & Science University, Portland, OR, USA Mitsunori Yamada Department of Clinical Research, National Hospital Organization, Saigata National Hospital, Ohgata-ku Johetsu-city, Niigata, Japan Mayumi Yamada Department of Biochemistry and Cellular Biology, National Institute of Neuroscience, National Center of Neurology and Psychiatry, Kodaira, Tokyo, Japan Tulin Yildirim Department of Radiology, Baskent University Medical School, Adana, Turkey Albert J. Yoo Department of Interventional Neuroradiology and Endovascular Neurosurgery, Massachusetts General Hospital, Harvard Medical School, Boston, MA, USA

List of Contributors

xxxiii

Chris I. De Zeeuw Netherlands Institute for Neuroscience, Royal Dutch Academy of Arts & Sciences (KNAW), Amsterdam, The Netherlands Department of Neuroscience, Erasmus Medical Center, Rotterdam, The Netherlands Gerard Zitnik Department of Neurobiology and Anatomy, Drexel University College of Medicine, Philadelphia, PA, USA Mary L. Zupanc Children Hospital of Orange County, Orange County, CA, USA

Section 1 Cerebellar Development

1

Specification of the Cerebellar Territory Marion Wassef

Abstract

The cerebellar primordium develops dorsally at an intermediate anteroposterior (AP) level of the neural tube. Its size is modulated by the early anteriorizing and posteriorizing signals, which pattern the neural tube. Two important signaling centers, the midbrain–hindbrain organizer and the roof plate, intersect at the level of the cerebellar anlage and control its positioning, differentiation, growth, survival, and patterning. Neural tube bending in the pontine region induces a widening of the fourth ventricle, which is made possible by choroid plexus differentiation and extension. As a consequence of these morphogenetic changes, the AP axis of the cerebellar primordium is rotated by 90 , and the cerebellar vermis and hemispheres derive from the anterior and posterior parts of the early cerebellar plate, respectively. The cerebellar plate is progressively subdivided along its dorsoventral axis into distinct domains, which generate subsets of cerebellar neurons according to their neurotransmitter phenotype. The roof plate marked by Gdf7 expression is at the origin of choroid plexus cells but does not contribute neurons or glia to the cerebellum. The rhombic lip, marked by Atoh1 expression, produces all the glutamatergic neurons of the cerebellum and a large number of non-cerebellar neurons. Finally, the ventral cerebellar neuroepithelium, marked by Ptf1a expression, generates all the GABAergic neurons and can be further subdivided into two progenitor domains, devoted to the production of Purkinje cells and GABAergic projection neurons of the deep

M. Wassef Institut de Biologie de l’Ecole Normale Supe´rieure (IBENS), 46 rue d’Ulm, 75005 Paris, France and CNRS UMR 8197, 46 rue d’Ulm, 75005 Paris, France and INSERM U1024, 46 rue d’Ulm, 75005 Paris, France e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_1, # Springer Science+Business Media Dordrecht 2013

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cerebellar nuclei. The so-called cerebellar primordium is not restricted to the production of cerebellar neurons but contributes to a large number of nuclei in the isthmic region.

Introduction The relative positions of the main brain subdivisions (Fig. 1.1) are established during development and have been conserved in the course of vertebrate evolution. The neural tube progressively differentiates into distinct regional identities resulting in local modifications of the growth and mechanical properties of the neuroepithelium and the formation of bulges called vesicles. In parallel, the different domains of the neuroepithelium evolve distinct competences to respond to adjacent or intrinsic signals. The forebrain, midbrain, and hindbrain vesicles, which form first, are known as the primary brain vesicles and become further subdivided later. The vesicles and intervening constrictions have often been used as stage-specific landmarks of early brain regionalization (but see below). In early studies based on retrospective anatomical observations during development, the cerebellar primordium was identified by its proximity to two landmarks: the midbrain anteriorly and the hindbrain choroid plexus posteriorly. Soon after neural tube closure, the cerebellar plate was identified as a pair of dorsal extensions of the anterior hindbrain adjacent to the midbrain vesicle and limited by the choroidal plate (Fig. 1.2). However, the extent of the cerebellar primordium and the localization of its boundaries could not be determined solely on the basis of anatomical or histological studies. Furthermore, specification of the cerebellar territory is likely to have begun much earlier with neural plate regionalization. Beginning from the 1990s, a series of fate mapping studies were therefore undertaken aiming at mapping early brain subdivisions or, more specifically, at delineating the cerebellar primordium. Several vertebrate species were examined at successive developmental stages in these studies, which used a large variety of lineage tracing methods. The fate maps performed at a given stage provide information about the destiny of a region of the neural tube left under the influence of adjacent structures in its normal context. It is also interesting to determine at what stage the cerebellar primordium becomes specified (or committed) and is able to maintain its cerebellar fate in isolation when cultured in vitro or transplanted to ectopic locations. Finally, the development of sophisticated molecular genetics tools provided new insight into the molecular pathways involved in the control of the size of the cerebellar territory and the positioning of its boundaries.

Delineating the Cerebellar Primordium Obtaining a fate map at a given stage consists in labeling a region of a living embryo in order to follow its fate at later developmental stages when anatomical structures can be unambiguously identified. This requires that the label should not spread to adjacent structures or be diluted by tissue growth. Isotopic transplants are often used

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Fig. 1.1 Relative positions of the main brain subdivisions in adult mice: (a) dorsal and lateral views of a dissected adult mouse brain. The main brain subdivision have been colored, forebrain in brown, midbrain in blue, cerebellum in red, and the rest of the hindbrain in yellow. (b) schematic representation of a medial sagittal section through a similar brain depicting the structures which derive from the r1 (rhombomere 1) developmental compartment. These include the pontine region and the cerebellum which are highlighted in red. (c) higher magnification of the vermal region of the cerebellum illustrating the continuity of neural structures forming the roof of the fourth ventricle. The cerebellum is connected to the midbrain through the vellum medullaris and to the posterior hindbrain through the choroid plexus

to minimize the spread of the label. The donor embryo is labeled by injection, incorporation of a dye, or by genetic tagging. Homologous fragments are excised from the donor and host. The host fragment is discarded and replaced by its labeled donor counterpart. This technique of embryological manipulations is powerful but concerns only few vertebrate species which are accessible to embryological manipulations at early stages. In particular, it is not appropriate for mammalian embryos. Nevertheless, it is possible to extend to other species the fate maps obtained from experimentally amenable species by using the gene expression boundaries as landmarks for performing grafts. The gene map then serves as a common reference. Based on the observation that the pattern of gene expression is globally well conserved across species, it is assumed that the relation between the fate map and the gene map is conserved. The genetic technologies developed in mouse during the

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a midbrain forebrain midbrain hindbrain r1 r2 hindbrain

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Fig. 1.2 Localization of the cerebellar primordium in chick and mouse embryos: (a) schematic representation of the neural tube vesicles and main brain subdivisions in chick embryos at 1.5 days of incubation (E1.5, dorsal view) and E3.5 (lateral view), corresponding to developmental stages 11 and 20 of Hamburger and Hamilton (1951), respectively. The arrowheads mark the limits of the major brain subdivisions. Note the marked bending of the neural tube at 3.5 days of incubation and the appearance of a choroid plexus forming the roof of the hindbrain. The cerebellum (in red) derives from the dorsal part of r1. (b) Lateral (E8.5 and E9.5) and posterior (E11.5) views of three mouse embryos. The cerebellar primordium is colored in pink and the midbrain in green. The mouse neural tube closes between E8.5 and E9.5. Notice that at E11.5 the two cerebellar halves which were previously opposed are separated by the transparent choroid plexus posteriorly and widely diverge

last decades have directly confirmed the suitability of this molecular anatomy. It has become possible to genetically label a specific cell subset at a given embryonic stage in mouse embryos and subsequently trace their lineage (Zinyk et al. 1998; Rodriguez and Dymecki 2000; Sgaier et al. 2005; Dymecki and Kim 2007).

Fate Maps of the Cerebellum The first detailed fate maps of the cerebellar primordium, soon after neural tube closure, were obtained independently by the groups of Alvarado-Mallart

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(Martinez and Alvarado-Mallart 1989, 1990) and Le Douarin (Hallonet et al. 1990) in ten somite-stage avian embryos (stage 10 of Hamburger and Hamilton 1951, noted HH10). The fate maps were obtained by isotopic transplantation of fragments of the neural tube between quail and chick embryos. The quail-chick chimera transplantation method designed by Le Douarin (1982) is based on the similarity between avian species at early developmental stages. It takes advantage of a specific cytological feature to identify quail cells. A clump of nucleolus-associated heterochromatin appears as a central dot within the nucleus of quail cells labeled with DNA stains. In chick cells, the chromatin appears diffusely organized. The QCPN monoclonal antibody which specifically recognizes quail cells was later developed and is now often used instead of DNA stains. The avian fate maps allowed to localize the anterior and posterior limits of the cerebellar primordium soon after neural tube closure. It is also indicated that, unexpectedly, the anterior limit of the cerebellar primordium at the 10 somite stage (HH10) is not marked by the isthmic constriction. Rather, the prospective cerebellum extends anteriorly into the adjacent mesencephalic vesicle. The anterior limit of the cerebellum was not materialized by a morphological landmark but likely corresponded to the MHB boundary as defined by Otx2 expression (Millet et al. 1996). Indeed, the Otx2 boundary lies slightly anterior to the isthmic constriction (Millet et al. 1996). Recent genetic fate maps performed in mouse embryos demonstrated that the anterior brain domain, characterized by Otx2 and Wnt1 expressing cells, are adjacent to the cerebellar primordium (Sgaier et al. 2005; Zervas et al. 2005). Interestingly, at later developmental stages, Otx2 is also expressed at the caudal edge of the cerebellar primordium and is required for its development. The hindbrain is further subdivided into seven developmental units called rhombomeres (r1-r7). The quail-chick fate maps indicated that the cerebellum originates from the most anterior rhombomere, r1. It was unclear if the primordium of the cerebellum is confined to dorsal r1. The auricular part of the cerebellum was identified by Marin and Puelles (1995) as originating from the caudalmost region of the cerebellar primordium and was postulated by these authors to derive from r2. Genetic tracing indicated, however, that dorsal r2 cells do not contribute to the cerebellum in mouse embryos (Awatramani et al. 2003; Farago et al. 2006). Anteroposterior specification along the hindbrain and spinal cord is controlled by a family of widely conserved transcription factors, the Hox genes. The most anteriorly expressed, Hoxa2 reaches r2 but is not expressed in r1. The fate maps thus indicate that the cerebellar primordium lies anterior to the domain of influence of the Hox genes. Accordingly, downregulation of Hoxa2 in mouse (Gavalas et al. 1997) and chick (Irving and Mason 2000) results in a posterior extension of the cerebellar primordium at the expense of r2. As concerns the dorsoventral dimension of the cerebellar primordium, the quail chick fate maps indicated that even if it is restricted to the neural tubular plate, it extends more in ventral r1 than previously suspected, encompassing two thirds of the neural tube dorsoventral axis.

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Fig. 1.3 Differentiation of the hindbrain choroid plexus from the roof plate: Schematic representation of the development of the choroid plexus between E1.5 and E3.5 in chick embryos based on data obtained in mouse and chick. At each age the neural tube is illustrated in toto on the left and the corresponding transverse section through the anterior hindbrain on the right. The choroid plexus derives from epithelial cells which constitute the roof plate at day 1.5. These cells start to proliferate and undergo an epitheliomesenchymal transition in such a way that they form a wide thin sheet of neural cells which constitute the roof of the hindbrain, fourth ventricle. At the same time, anterior bending of the neural tube tends to widen the dorsal hindbrain (forces represented by the divergent arrows at E3.5)

Rotation of the Cerebellar Primordium At the time of neural closure, the cerebellar primordium consists of a pair of dorsal wings which flank the roof plate and extend along the anteroposterior axis of the anterior hindbrain (r1). Fate maps obtained at this stage by transplantation or by genetic cell lineage tracing indicated that the two posterior cerebellar plates will develop as the cerebellar hemispheres. They are pushed apart laterally by the flexure of the neural tube at the level of the pons (pontine flexure) and the constriction of the neural tube at the MHB boundary. Their mediolateral rotation is made possible by the rapid growth of the choroidal plate (Fig. 1.3). The final rotation of the prospective cerebellar vermis takes place slightly later in the anterior cerebellar plate and involves active cell reorganization.

Specification of the Cerebellar Primordium The Midbrain–Hindbrain (MHB) Domain At early developmental stages, under the influence of posteriorizing and anteriorizing signals, the neural plate becomes subdivided into anterior and posterior domains, which express distinct combinations of transcription factors (Wurst and

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Bally-Cuif 2001; Liu and Joyner 2001; Kobayashi et al. 2002; Raible and Brand 2004). The homeodomain transcription factors Otx2 and Gbx2 are expressed, respectively, in the anterior and posterior neural plate. Slightly later, at the end of gastrulation, between E7.5 and E9.5 in the mouse, a new region of the neural tube forms progressively at an intermediate anteroposterior location. Called midbrain– hindbrain (MHB) domain, it expresses a combination of transcription factors and signaling molecules, which endow it with the competence to develop as midbrain and cerebellum. Among transcription factors with conserved function in MHB development are the vertebrate homologs of the drosophila engrailed segmentation genes, En1 and En2 in mouse and chick, eng1, 2 and 3 in zebrafish, the paired homeodomain transcription factors Pax2, Pax5 and Pax8, the Lim homeodomain transcription factors Lim1a and Lim1b (depending on species). Development of the MHB domain is also dependent on the function of two secreted cell–cell signaling molecules Wnt1 (wingless-related) and Fgf8 (fibroblast growth factor 8), which are expressed in the presumptive midbrain and anterior hindbrain, respectively. The major MHB-specific genes are not interdependent at early stages. The onset of their expression in the MHB domain is triggered independently by vertical signals derived from non-neural structures underlying the neural plate (McMahon et al. 1992; Wurst et al. 1994; Ye et al. 2001; Li and Joyner 2001). Closely related genes belonging to the same family often contribute to MHB development displaying small differences in timing or expression patterns between paralogs. These variations may explain why the phenotypes, resulting from inactivating the same paralog, may markedly differ in severity between species. Some of the genes expressed in the early MHB domain will be later maintained in the cerebellum, the midbrain, or both.

Organizing Properties of the Isthmic Neuroepithelium – Fgf8 As a complement of quail-chick fate map studies, ectopic transplantations were performed with the aim of testing the commitment of the neuroepithelium of the posterior midbrain and anterior hindbrain to a cerebellar fate (Alvarado-Mallart et al. 1990; Martinez et al. 1991). The resulting chimeras were analyzed at short and long survival times posttransplantation. Unexpectedly, both anterior and posterior quail cerebellar transplants switched the fate of the adjacent host neuroepithelium. Isthmic transplants, grafted anterior to the MHB, affected the fate or polarity of adjacent structures. The neuroepithelium of the anterior diencephalon (p2) was transformed into optic tectum, a midbrain structure, instead of its normal thalamic fate. More anterior transplants resulted in the formation of one or two ectopic optic tectum structures, which often fused with the endogenous tectum. The optic tectum is a polarized structure marked by the graded expression of several transcripts and proteins including En2, which can be detected by immunocytochemistry. The tectal structures, induced by isthmic transplants, were always polarized with their caudal end close to the grafts. In the hindbrain, the dorsal neuroepithelium of r2, r3, and r4 was recruited to a cerebellar fate by adjacent cerebellar grafts. Thus, pieces of cerebellar primordium

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behaved as long-range signaling centers. When transplanted into a wide competent territory they induced the adjacent neural structures to complete an ectopic MHB domain. These properties are the hallmark of secondary organizers. The neuroepithelium of the isthmic constriction adjacent to the MHB boundary is therefore known as the MHB or isthmic organizer. Wnt1 and Fgf8, two diffusible signaling molecules expressed in the midbrain and anterior hindbrain, respectively, were candidate organizing signals. Loss of function experiments in the mouse indicated that either one is essential for the formation of the cerebellum and midbrain. Gain of function experiments, obtained by implantation of beads soaked in FGF8 in the neural tube of avian embryos, indicated that an ectopic source of FGF8 mimicks the inductive properties of fragments of the MHB organizer (Crossley et al. 1996). FGF8 has several important functions in the specification of the cerebellar territory. First, it prevents the expression of potent repressors of the cerebellar fate, Otx2 anteriorly, and Hoxa2 posteriorly. It also stabilizes the expression of Gbx2, which is required for cerebellum development. FGF8 controls growth and polarity of the MHB domain by maintaining pools of actively proliferating undifferentiated precursors on each side of the MHB boundary and by controlling the gradient of En1/2 expression. FGF8 is a potent morphogen, which serves as a major component of several organizing centers in the brain and body. FGF8 organizing activity relies on the rapid and progressive deployment of a set of target genes with nested expressions many of which act as inhibitors of the FGF8 signaling pathway. Besides, alternative splicing of Fgf8 generates several secreted isoforms differing only at their mature amino terminus (MacArthur et al. 1995). Two of the four FGF8 splice isoforms, FGF8a and FGF8b, are expressed in the mid-hindbrain region during development. Although the only difference between these isoforms is the presence of an additional 11 amino acids at the N terminus of FGF8b, these isoforms possess remarkably different abilities to pattern the midbrain and anterior hindbrain (Olsen et al. 2006). Quantitative analysis showed that Fgf8b signal is 100 times stronger than Fgf8a signal (Sato et al. 2001). The FGF8 subfamily comprises two additional members, FGF17 and FGF18, which are expressed by the MHB organizer. The FGF17b splice variant and FGF18 display intermediate receptor-binding affinities and patterning abilities compared to FGF8a and FGF8b (Olsen et al. 2006).

The MHB Organizer: A Molecular Network Set up at the Otx2-Gbx2 Boundary Otx2 and Gbx2 mutually cross-inhibit each other’s expression resulting in the formation of a sharp boundary between their expression domains. Positioning of the Otx2Gbx2 boundary, also called MHB boundary, depends on the relative “strength” of Otx2 and Gbx2 (Simeone 2000; Wurst and Bally-Cuif 2001). Loss of Gbx2 function or ectopic expression of Otx2 posteriorly results in a posterior displacement of the MHB boundary (Wassarman et al. 1997; Broccoli et al. 1999; Millet et al. 1999). Conversely, reducing the number of Otx2 copies in an Otx1 mutant background shifts the

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Fig. 1.4 Genetic network controlling midbrain–hindbrain organizer positioning and function: At early stages, positioning of the MHB organizer depends on cross interactions between Otx2 and Gbx2, which code for homeodomain transcription factors. Otx2 is a potent repressor of cerebellar fate even at relatively late stages of development. Transcription of the En1/2, Fgf8, and Wnt1 genes is initiated independently in the MHB domain. They soon engage in cross-regulatory interactions constituting a tight genetic network which is deployed in space around the Otx2/ Gbx2 boundary and is completely dependent on the function of each of its components. The potent patterning and polarizing activity of the MHB organizer probably also reflects its capacity to maintain a pool of undifferentiated midbrain and hindbrain cells. Also dysfunction of the MHB organizer results in truncation of adjacent structures

MHB boundary anteriorly up to the p2 prosomere and dramatically increases the size of the cerebellar primordium. Otx2 plays a major role in preventing the midbrain and posterior hindbrain to turn into cerebellum. In contrast, in the absence of Gbx2, the anterior hindbrain fails to develop a coherent midbrain or cerebellum structure (Wassarman et al. 1997). Conversely, anterior ectopic expression of Gbx2 induces a milder and transient enlargement of the cerebellum (Millet et al. 1999). This indicates that other factors (FGF8, Hox. . .) cooperate with Gbx2 to prevent transformation of the anterior hindbrain into midbrain. At the stabilized Otx2-Gbx2 boundary, the Wnt1, En1, FGF8, and Pax2 genes initiate cross-regulatory interactions and soon become interdependent, as schematized in Fig. 1.4. Once established, the function of the MHB organizer relies on each member of this genetic network. Thus, inactivation of a single gene (Wnt1: McMahon et al. 1992, En1: Wurst et al. 1994), or of two members of the same gene family (Pax2 and 5: Urba´nek et al. 1997), or conditional inactivation of Fgf8 to

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bypass its early requirement for embryonic development, all result in the deletion of most of the midbrain and cerebellum, preceded by a downregulation of the expression of the other genes involved in MHB organizer function.

The Otx2-Gbx2 Boundary: A Stable or Drifting Limit? Because Fgf8 is highly expressed adjacent to the Otx2-Gbx2 boundary and ectopic FGF8 induces Gbx2 and represses Otx2 expression the cell lineage restriction and stability of the MHB boundary has been a matter of debate. Distinct methods were used in chick and mouse embryos in order to follow the fate of cells adjacent to the MHB boundary. In chick embryos, the Otx2 boundary at the MHB junction was found to coincide with the posterior end of the early generated neurons of the mesencephalic nucleus of nerve V (Millet et al. 1996). In mouse embryos, a fate map of the MHB boundary was obtained by Wnt1-CreERT conditional labeling at E7.5, E8.5, and E9.5 (Sgaier et al. 2005). A cell lineage restriction was observed in the dorsal posterior midbrain beginning from E8.5 but the lineage restriction was not established ventrally before E9.5. The lineage restriction that prevented Wnt1 expressing cells from contributing to the cerebellar plate was not absolute, as a few cells were observed posteriorly in the anterior part of the cerebellar plate. At later stages, these cells did not contribute to the mature cerebellum. Taken together, the data in chick and mouse indicate that the early MHB boundary is stable in the dorsal neural tube and marks the anterior limit of the mature cerebellum. In contrast, ventral midbrain cells may relocate in the adjacent anterior hindbrain between E8.5 and E9.5, resulting in a discrete anterior shift of the Wnt1 lineage boundary between these two stages.

Genes Regulating the Competence of the Neuroepithelium to Develop a MHB Identity During zebrafish gastrulation, iro1 and iro7 are expressed in a broad neural plate domain that includes the prospective MHB. The iro1 and iro7 expression domain is expanded in headless and masterblind mutants, which are characterized by exaggerated Wnt signaling. Early expansion of iro1 and iro7 expression in these mutants correlates with expansion of the (MHB) domain, raising the possibility that iro1 and iro7 have a role in the determination of the MHB. Knockdown of both iro1 and iro7 genes prevents Pax2.1 expression and the establishment of the MHB organizer. Their ectopic expression is not sufficient to induce a MHB identity (Itoh et al. 2002). These observations suggest that the Iro genes are required for the early acquisition of a MHB identity. The POU domain transcription factor spg/pou2 (Reim and Brand 2002), which is specifically expressed in the MHB primordium of zebrafish is involved in conferring a MHB identity. spg/pou2 is orthologous to mammalian Oct3/Oct4, and is required for the early expression of key molecules that function in the formation of

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the MHB, such as pax2.1, spry4, wnt1, her5, eng2, and eng3. Pou2 mutant embryos are insensitive to FGF8, although FGF receptor expression and activity of the FGF8 signaling pathway appear intact. This indicates that spg/pou2 mediates the regional competence to respond to FGF8 signaling.

Genes Regulating Distinct Neuroepithelium Competences on Either Sides of the MHB Boundary If the MHB boundary remains stable it is because it forms at the interface between two domains, which from the onset have distinct competences. This means that they differ in their responses to the same signals. Several transcription factors expressed on either sides of the MHB boundary have been shown to affect the competence of their expression domains to respond to FGF8 signaling. Chick Irx2, a homeobox gene of the Iroquois family is expressed in the presumptive cerebellum and is a target of the FGF8/MAP kinase cascade (Matsumoto et al. 2004). Chick Irx2 modulates the response of neuroepithelial cells to FGF8 signaling. At the difference of Fgf8b, which when expressed ectopically in the midbrain shifts neuroepithelium identity to a cerebellum fate, Fgf8a stimulates midbrain growth but fails to induce a fate change. Overexpression of Irx2 with Fgf8a turns the entire dorsal midbrain into a cerebellum. Meis2, which is expressed in the chick tectal primordium, is both necessary and sufficient for tectal development. Otx2 transcriptional activity in the tectum depends on the balance between a co-repressor, the Groucho family co-repressor Tle4/Grg4, and a coactivator, the TALE-homeodomain protein Meis2. Meis2 acts by competing with the Grg4 corepressor for binding to Otx2 and thereby restores Otx2 transcriptional activator function in the tectum (Agoston and Schulte 2009).

Subdivisions of the Cerebellar Plate Like other parts of the early neural tube, the neuroepithelium of the cerebellar primordium is progressively subdivided along is anteroposterior and dorsoventral axis. This partition reflects differences in the duration (Sato and Joyner 2009) and intensity (Basson et al. 2008) of exposure to diffusible signaling molecules produced by adjacent signaling centers like FGF8/17/18, WNT1, BMP4/7, GDF7, SHH. . ., and to their various downstream effectors. The identity of cerebellar progenitors thus depends on their distance from the midbrain–hindbrain organizer, the roof plate/choroid plexus, and the floor plate, which contribute to increase their diversity. The current paradigm to understand the successive developmental steps leading to the production of a large diversity of neuronal types derives from the pioneer studies of the T. Jessell group on the developing spinal cord. Neuronal diversity in the spinal cord was shown to result from the subdivision of the neuroepithelium into progenitor domains according to their dorsoventral coordinates. Each progenitor

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Fig. 1.5 Anteroposterior and dorsoventral subdivisions of the cerebellar plate. Schematic representation of anteroposteriorly (AP left) and dorsoventrally (DV right) arranged cerebellar patterns observed on whole-mount stained brains. The dotted line represents a sagittal section. Imagine the pattern changes on sagittal sections when the line is moved mediolaterally and how it is complicated to reconstitute the global pattern from sections

domain expresses a unique combination of transcription factors and generates a characteristic set of neuronal types in a stereotyped succession. The establishment of progenitor domains occurs in two steps. In an early phase, opposed gradients of SHH and BMP signals derived from the floor plate and roof plate, respectively, activate or repress the expression of a first set of transcription factors which provide broad dorsal or ventral identities to epithelial cells. The second step relies on successive and mutual cross-interactions between transcription factors. These interactions subdivide the early broad neural tube domains into dedicated progenitor domains with sharpened boundaries. This second step is less well documented in the cerebellar primordium than in the spinal cord even if mutual crossrepressions between transcription factors expressed in adjacent cerebellar domains have been reported. As illustrated in Fig. 1.5, the rotation of the cerebellar primordium tends to obscure the relationship between the early axes which determine the orientation of gradients of diffusible molecules derived from the midbrain–hindbrain organizer or the roof plate and those of the late embryonic cerebellum.

Anteroposterior Subdivisions of the Cerebellar Plate Cerebellar vermis and hemisphere. Fate map studies in mouse and chick have shown that the cerebellar vermis derives from the anterior cerebellar plate, while the cerebellar hemispheres derive from its posterior part. Several genes involved in the function of the midbrain–hindbrain organizer like Fgf8, En1, and Pax2 are

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known to display differential expression along the anteroposterior axis of the cerebellar plate. This is also the case for several targets or inhibitors of FGF8 signaling including the Fgf8 family members Fgf17 and Fgf18, the Sprouty1/2/4 genes, which together modulate the chronology and spatial deployment of FGF8 signaling. Expression of the basic helix-loop-helix (bHLH) transcription factors Ngn1/2 is also modulated along the anteroposterior axis. Vermis size depends on the duration and intensity of FGF8 signal expressed in the anterior hindbrain (Sato and Joyner 2009). Interestingly, the cerebellar domains from which the vermis and hemispheres derive seem to be distinct and differ in their response to FGF8 signaling. The vermal region is atrophied or deleted in mutants in which the function of the MHB organizer is affected or in which the intensity or duration of FGF8 signaling is decreased. Conversely, FGF8 signaling can be increased in the midbrain–hindbrain domain through the conditional inactivation of two or three of the functionnally redundant Sprouty genes which belong to a family of FGF8 inhibitors. Sprouty1/2 conditional mutation in the MHB results in a marked enlargement of the vermis but does not affect the size of the hemispheres (Yu et al. 2011). The mature cerebellar midline derives from the midbrain-hindbrain boundary. At early developmental stages, the cerebellar vermis is a paired structure. In early embryos, the two adjacent vermal halves are separated by the roof plate. Fate map studies have shown that the midline of the mature cerebellar vermis derives from a narrow group of cells at the midbrain–hindbrain boundary (Kala et al. 2008). This indicates that similar to the cerebellar hemispheres, the vermis also experiences a nearly 90 rotation of its axis from anteroposterior to mediolateral. This complex process also explains the occurrence of incomplete fusions of the vermis in pathological and experimental contexts. An isthmic compartment intervening between the cerebellum and midbrain? In late embryos and pups, the isthmus is an ill-defined region straddling the anterior hindbrain and posterior midbrain. Early embryological studies have described several populations of neurons originating in the cerebellar plate and destined to populate the isthmic nuclei. The Wnt1-CreER fate maps described above indicate that the Wnt1 expression domain contributes cells to a rostral hindbrain subdivision distinct from the cerebellum which could belong to the isthmus. At early stages of cerebellar neurogenesis in mice (E11.5, E12.5), a Ptf1a negative domain intervenes on sagittal sections (see below) between the Ptf1a positive cerebellar neuroepithelium and the ring of Wnt1 positive cells which marks the midbrain–hindbrain boundary. These cells express low level of Pax2. It is unclear if they belong to a portion of the ventral neuroepithelium of the pons repositioned dorsally as a consequence of morphogenetic movements or constitute a bona fide component of the cerebellar primordium. Some authors consider the isthmus as a separate compartment intervening between the midbrain and r1 but the existence of an isthmic neuroepithelium compartment is still questionable. The isthmic compartment is proposed to correspond to the Fgf8 expression domain and would not contribute to the cerebellar primordium located more posteriorly in dorsal r1. Genetic lineage tracing indicates, however, that the progeny of FGF8 expressing cells largely contributes to the mature cerebellum

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(Charles Watson and Tomomi Shimogori, personal communication). It is nevertheless possible that a proportion of the Ptf1a negative cells of the isthmic neuroepithelium turn on Ptf1a expression at some point as is the case for cells adjacent to the RL, which successively turn on Atoh1/Math1 expression (see below).

Dorsoventral Subdivisions of the Cerebellar Plate The rhombic lip/cerebellar neuroepithelium (Atoh1/Ptf1a) subdivision. In the early hindbrain (also called rhombencephalon) as in the rest of the neural tube the roof plate is a row of specialized cells which marks the dorsal midline. The roof plate cells express Gdf7, a member of the BMP family of signaling molecules. Fate mapping of the Gdf7 positive cells using Gdf7-Cre mice indicated that the neural part of the hindbrain choroid plexus or choroidal plate derives from the roof plate. The dorsal most subdivision of the hindbrain neuroepithelium called rhombic lip (RL) lines the roof plate. The RL in r1 is called cerebellar, rostral, anterior, or upper rhombic lip (uRL) whereas the rhombic lip of r2-r7 is called lower rhombic lip (lRL). The uRL has been classically described as the source of cerebellar granule cell precursors and thought to generate only cerebellar granule neurons precursors whereas the mossy fiber precerebellar nuclei is derived from the lRL (Altman and Bayer 1997; Rodriguez and Dymecki 2000). The bHLH transcription factor Atoh1 (for mouse Atonal homolog 1) also known as Math1 is expressed in the RL as early as E9.5. Atoh1 is required for the development of the uRL-derived cerebellar granule neurons and the lRL-derived mossy-fiber precerebellar nuclei (Ben-Arie et al. 1997; Wang and Zoghbi 2001; Wang et al. 2005). Short- and long-term tracing of the progeny of the Atoh1-positive RL progenitors identified a large population of additional derivatives (Wang et al. 2005; Machold and Fishell 2005). Strikingly, the Atoh1+ progenitors of the uRL produce all the glutamatergic neurons in the cerebellum. This includes the granule cells, the scarce population of unipolar brush cells, and all the glutamatergic projection neurons of the deep cerebellar nuclei (Wang et al. 2005; Machold and Fishell. 2005; Fink et al. 2006). The GABAergic neurons of the cerebellum originate from the non-RL cerebellar neuroepithelium, which is characterized by the expression of the Ptf1a bHLH transcription factor (Hoshino 2006). Ptf1a is required for the differentiation of all the GABAergic cell types of the cerebellum (Hoshino et al. 2005). This comprises the two types of projection neurons: Purkinje cells and the GABAergic projection neurons of the deep cerebellar nuclei, and a multitude of Pax2-positive interneuron types, that is, Golgi, basket, and molecular layer interneurons in the cerebellar cortex, and the GABAergic interneurons of the deep cerebellar nuclei (Maricich and Herrup 1999). It is interesting to note that the RL and the cerebellar epithelium cannot be considered as lineage restricted compartments. The fates of successive waves of Atoh1-expressing cells have been traced in Atoh-CreER x R26R mice in which the Cre recombinase is tethered in the cytoplasm of Atoh1 expressing cells (Machold and Fishell 2005). Timed injection of Tamoxifen to Atoh-CreER x R26R

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pregnant mice allows the recombinase to enter the nucleus and recombine the LacZ reporter. The progeny of progenitors, which were expressing Atoh1 at the time of Tamoxifen injection, can be followed. It was observed that the RL progenitors, which express Atoh1 at early stages, give rise to the early generated neurons of the deep cerebellar nuclei and the non-cerebellar populations of isthmic nuclei. They do not contribute to the granule cell population. In contrast, later injections of tamoxifen trace successive waves of granule cells which first populate the anterior cerebellar cortex then progressively its more posterior aspects. These observations indicate that new Atoh-negative “naı¨ve” progenitors in the adjacent cerebellar neuroepithelium are recruited to express Atoh. This suggests that the RL acts as a device in which “naı¨ve” progenitors are transformed into successive RL derivatives, which then migrate out allowing a new population of “naı¨ve” progenitors to enter the RL domain. Ptf1a and Atoh1 repress each other’s expression (Pascual et al. 2007) but beginning from the onset of Purkinje cell generation at E11.5, the pattern of Ptf1a protein expression is “salt and pepper” in the cerebellar neuroepithelium, leaving room for putative “naı¨ve” progenitors. In addition, in Atoh1 mutants, a small population of granule cells develops which is derived from Ptf1a + progenitors (Pascual et al. 2007). Taken together, these findings suggest that GABAergic and glutamatergic cerebellar neurons originate from distinct neuroepithelial regions, the cerebellar neuroepithelium and the RL. Furthermore, normal development of these neurons requires the activity of bHLH transcription factors Ptf1a or Atoh1, respectively (Hoshino et al. 2005; Ben-Arie et al. 1997). A similar subdivision of progenitors related to the neurotransmitter phenotype of their progeny is also observed in the forebrain. The glutamatergic and GABAergic neurons which form the cerebral cortex arise from distinct subdivisions of the forebrain neuroepithelium (Anderson et al. 1997; Wonders and Anderson 2006). Their differentiation is under the control of distinct bHLH transcription factors: the glutamatergic projection neurons of the cerebral cortex are produced in the dorsal pallium under the control of Ngn1/2, whereas generation of the GABAergic interneurons in the subpallial neuroepithelium is dependent on Ascl1/Mash1. Other subdivisions of the cerebellar plate. The cerebellar plate is currently considered to comprise four subdivisions and an intervening non-cerebellar isthmic domain (Fig. 1.6a): from dorsal to ventral: the roof plate, the RL, and two additional domains Ptf1dorsal and Ptf1ventral (Chizhikov et al. 2006; Zordan et al. 2008; Mizuhara et al. 2010). The existence of two subdivisions in the cerebellar neuroepithelium was first suggested by the presence of populations of early postmitotic neurons in the mantle zone of the cerebellar plate, which segregate along the dorsoventral axis and differ in marker expression (Fig. 1.6b). This includes an early ventral expression of Pax2, a marker of cerebellar GABAergic neurons, with the exception of Purkinje cells. Lhx1/5 and Corl2 (Minaki et al. 2008) are both considered to be Purkinje cell markers. More recently, an abrupt change in the level of expression of E-cadherin in the cerebellar neuroepithelium was shown to correspond to fate differences in the overlying neurons (Fig. 1.6c). Nephrocystin3 (Neph3), which encodes a membrane protein and is a target of

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a

b

c

d

e

Is

Ptf1a ventral Ptf1a dorsal RL RP

Fig. 1.6 Dorsoventral subdivisions of the cerebellar primordium: (a) Most studies agree that the cerebellar primordium comprises the following subdivisions from dorsal to ventral. 1, The roof plate does not contribute cells to the cerebellum. 2, The rhombic lip which expresses Atoh1/Math1 and produces all the glutamatergic neurons of the cerebellum and other neurons destined to populate a variety of isthmic nuclei. 3, A Ptf1a expressing domain which can be subdivided into a dorsal and a ventral component on the basis of differences in marker expression in the early neuroepithelium as well as in the overlying neurons. 4, Finally, a ventral domain whose cerebellar fate is uncertain. (b) Several populations of postmitotic neurons overlie different portions of the neuroepithelium of the cerebellar plate (Chizhikov et al. 2006). 1, A stream of migrating deep cerebellar neurons (in green) marked by Atoh1/Math1 and Tbr1 which accumulate ventrally, 2, Purkinje cells (in light purple) marked by Lhx1/5 or Corl2, 3, putative GABAergic neurons of the deep cerebellar nuclei (in orange) marked by Pax2 and Lmx1a. C-E Abrupt changes in the expression of markers in the cerebellar neuroepithelial cells suggest the existence of subdivisions in the cerebellar neuroepithelium. (c) E-Cadherin (Mizuhara et al. 2010), (d) Ngn1 and (e) Ngn2 (Zordan et al. 2008)

Ptf1a is expressed in both compartments. Antibodies to the two surface molecules, Neph3 and E-cadherin, were used for fluorescence activated cell sorting (FACS). The fate of the sorted cells was identified after 2 days of culture in vitro, based on their expression of cell type-specific markers detected by immunocytochemistry. The Neph3+/E-cadherinhigh neuroepithelial cells of the dorsalmost compartment were shown to produce Corl2+ Purkinje cells whereas the Neph3+/E-cadherinlow neuroepithelial cells generated Pax2+ neurons. These neurons, based on their birthdates, were GABAergic projection neurons of the deep cerebellar nuclei (Mizuhara et al. 2010). Subdivisions of the cerebellar neuroepithelium have also been detected on the basis of the expression of the Ngn1 (Fig. 1.6d) and Ngn2 (Fig. 1.6e) neurogenic proteins (Zordan et al. 2008). In summary, like other parts of the brain, the cerebellar plate is subdivided into progenitor domains, which produce different types of neurons. The spatial segregation of progenitors fated to produce specific cell types is, however, much looser than in the rest of the hindbrain or in the spinal cord.

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Conclusions and Future Directions The existence of a neuroepithelial territory specialized in the production of cerebellar neurons could be anticipated based on some of the adult characteristics of the cerebellum. The cerebellum is a well-delineated organ with a characteristic histological organization. If the origin of cerebellar neurons can be traced back to a restricted domain of dorsal r1, the same domain also contributes to a wide range of extracerebellar nuclei: the cerebellar territory is not devoted to the production of cerebellar neurons. Positioning, patterning, and growth of the cerebellar territory depend on two orthogonal organizers, the MHB organizer and the roof plate, which provide specific properties to the adjacent neuroepithelial domains, the isthmic neuroepithelium and the rhombic lip, respectively. Recently obtained fate maps have increased our understanding of the organization of the cerebellar rombic lip and the neurogenic processes taking place in this structure. Future studies should further our understanding of the organization of the rest of the cerebellar plate cell and improve our knowledge of the lineages it produces. Modifications in cerebellar plate patterning are observed in mouse embryos when the late expression of genes involved MHB organizer is modified. It would be interesting to investigate if the diversity in cerebellar shape and organization observed during evolution results from similar molecular variations.

References Agoston Z, Schulte D (2009) Meis2 competes with the Groucho co-repressor Tle4 for binding to Otx2 and specifies tectal fate without induction of a secondary midbrain-hindbrain boundary organizer. Development 136:3311–3322 Altman J, Bayer SA (1997) Development of the cerebellar system: in relation to its evolution, structure, and functions. CRC Press, Boca Raton, FL Alvarado-Mallart RM, Martinez S, Lance-Jones CC (1990) Pluripotentiality of the 2-day-old avian germinative neuroepithelium. Dev Biol 139:75–88 Awatramani R, Soriano P, Rodriguez C et al (2003) Cryptic boundaries in roof plate and choroid plexus identified by intersectional gene activation. Nat Genet 35:70–75 Anderson SA, Eisenstat DD, Shi L, Rubenstein JL (1997) Interneuron migration from basal forebrain to neocortex: dependence on Dlx genes. Science 278:474–476 Basson MA, Echevarria D, Ahn CP et al (2008) Specific regions within the embryonic midbrain and cerebellum require different levels of FGF signaling during development. Development 135:889–898 Ben-Arie N, Bellen HJ, Armstrong DL et al (1997) Math1 is essential for genesis of cerebellar granule neurons. Nature 390:169–172 Broccoli V, Boncinelli E, Wurst W (1999) The caudal limit of Otx2 expression positions the isthmic organizer. Nature 401:164–168 Chizhikov VV, Lindgren AG, Currle DS et al (2006) The roof plate regulates cerebellar cell-type specification and proliferation. Development 133:2793–2804 Crossley PH, Martinez S, Martin GR (1996) Midbrain development induced by FGF8 in the chick embryo. Nature 380:66–68 Dymecki SM, Kim JC (2007) Molecular neuroanatomy’s “Three Gs”: a primer. Neuron 54:17–34 Farago AF, Awatramani RB, Dymecki SM (2006) Assembly of the brainstem cochlear nuclear complex is revealed by intersectional and subtractive genetic fate maps. Neuron 50:205–218

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Fink AJ, Englund C, Daza RA et al (2006) Development of the deep cerebellar nuclei: transcription factors and cell migration from the rhombic lip. J Neurosci 26:3066–3076 Gavalas A, Davenne M, Lumsden A et al (1997) Role of Hoxa-2 in axon pathfinding and rostral hindbrain patterning. Development 124:3693–3702 Hallonet ME, Teillet MA, Le Douarin NM (1990) A new approach to the development of the cerebellum provided by the quail-chick marker system. Development 108:19–31 Hamburger V, Hamilton HL (1951) A series of normal stages in the development of the chick embryo. J Morphol 88:49–92 Hoshino H, Nakamura S, Mori K et al (2005) Ptf1a, a bHLH transcriptional gene, defines GABAergic neuronal fates in cerebellum. Neuron 47:201–213 Hoshino M (2006) Molecular machinery governing GABAergic neuron specification in the cerebellum. Cerebellum 5:193–198 Irving C, Mason I (2000) Signalling by FGF8 from the isthmus patterns anterior hindbrain and establishes the anterior limit of Hox gene expression. Development 127:177–186 Itoh M, Kudoh T, Dedekian M et al (2002) A role for iro1 and iro7 in the establishment of an anteroposterior compartment of the ectoderm adjacent to the midbrain-hindbrain boundary. Development 129:2317–2327 Kala K, Jukkola T, Pata I et al (2008) Analysis of the midbrain-hindbrain boundary cell fate using a boundary cell-specific Cre-mouse strain. Genesis 46:29–36 Kobayashi D, Kobayashi M, Matsumoto K et al (2002) Early subdivisions in the neural plate define distinct competence for inductive signals. Development 129:83–93 Le Douarin N (1982) The neural crest. Cambridge University Press, Cambridge Li JY, Joyner AL (2001) Otx2 and Gbx2 are required for refinement and not induction of midhindbrain gene expression. Development 128:4979–4991 Liu A, Joyner AL (2001) Early anterior/posterior patterning of the midbrain and cerebellum. Annu Rev Neurosci 24:869–896 MacArthur CA, Lawshe´ A, Xu J et al (1995) FGF-8 isoforms activate receptor splice forms that are expressed in mesenchymal regions of mouse development. Development 121:3603–3613 Machold R, Fishell G (2005) Math1 is expressed in temporally discrete pools of cerebellar rhombic-lip neural progenitors. Neuron 48:17–24 Maricich SM, Herrup K (1999) Pax-2 expression defines a subset of GABAergic interneurons and their precursors in the developing murine cerebellum. J Neurobiol 41:281–294 Marin F, Puelles L (1995) Morphological fate of rhombomeres in quail/chick chimeras: a segmental analysis of hindbrain nuclei. Eur J Neurosci 7:1714–1738 Martinez S, Alvarado-Mallart RM (1989) Rostral cerebellum originates from the caudal portion of the so-called “Mesencephalic” vesicle: a study using chick/quail chimeras. Eur J Neurosci 1:549–560 Martinez S, Alvarado-Mallart RM (1990) Expression of the homeobox Chick-en gene in chick/ quail chimeras with inverted mes-metencephalic grafts. Dev Biol 139:432–436 Martinez S, Wassef M, Alvarado-Mallart RM (1991) Induction of a mesencephalic phenotype in the 2-day-old chick prosencephalon is preceded by the early expression of the homeobox gene en. Neuron 6:971–981 Matsumoto K, Nishihara S, Kamimura M et al (2004) The prepattern transcription factor Irx2, a target of the FGF8/MAP kinase cascade, is involved in cerebellum formation. Nat Neurosci 7:605–612 McMahon AP, Joyner AL, Bradley A et al (1992) The midbrain-hindbrain phenotype of Wnt-1-/ Wnt-1- mice results from stepwise deletion of engrailed-expressing cells by 9.5 days postcoitum. Cell 69:581–595 Millet S, Bloch-Gallego E, Simeone A et al (1996) The caudal limit of Otx2 gene expression as a marker of the midbrain/hindbrain boundary: a study using in situ hybridisation and chick/ quail homotopic grafts. Development 122:3785–3797 Millet S, Campbell K, Epstein DJ et al (1999) A role for Gbx2 in repression of Otx2 and positioning the mid/hindbrain organizer. Nature 401:161–164

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Minaki Y, Nakatani T, Mizuhara E et al (2008) Identification of a novel transcriptional corepressor, Corl2, as a cerebellar Purkinje cell-selective marker. Gene Expr Patterns 8:418–423 Mizuhara E, Minaki Y, Nakatani T et al (2010) Purkinje cells originate from cerebellar ventricular zone progenitors positive for Neph3 and E-cadherin. Dev Biol 338:202–214 Olsen SK, Li JY, Bromleigh C et al (2006) Structural basis by which alternative splicing modulates the organizer activity of FGF8 in the brain. Genes Dev 20:185–198 Pascual M, Abasolo I, Mingorance-Le Meur A et al (2007) Cerebellar GABAergic progenitors adopt an external granule cell-like phenotype in the absence of Ptf1a transcription factor expression. Proc Natl Acad Sci USA 104:5193–5198 Raible F, Brand M (2004) Divide et Impera–the midbrain-hindbrain boundary and its organizer. Trends Neurosci 27:727–734 Reim G, Brand M (2002) Spiel-ohne-grenzen/pou2 mediates regional competence to respond to Fgf8 during zebrafish early neural development. Development 129:917–933 Rodriguez CI, Dymecki SM (2000) Origin of the precerebellar system. Neuron 27:475–486 Sato T, Araki I, Nakamura H (2001) Inductive signal and tissue responsiveness defining the tectum and the cerebellum. Development 128:2461–2469 Sato T, Joyner AL (2009) The duration of Fgf8 isthmic organizer expression is key to patterning different tectal-isthmo-cerebellum structures. Development 136:3617–3626 Sgaier SK, Millet S, Villanueva MP et al (2005) Morphogenetic and cellular movements that shape the mouse cerebellum; insights from genetic fate mapping. Neuron 45:27–40 Simeone A (2000) Positioning the isthmic organizer where Otx2 and Gbx2meet. Trends Genet 16:237–240 Urba´nek P, Fetka I, Meisler MH et al (1997) Cooperation of Pax2 and Pax5 in midbrain and cerebellum development. Proc Natl Acad Sci USA 94:5703–5708 Wang VY, Zoghbi HY (2001) Genetic regulation of cerebellar development. Nat Rev Neurosci 2:484–491 Wang VY, Rose MF, Zoghbi HY (2005) Math1 expression redefines the rhombic lip derivatives and reveals novel lineages within the brainstem and cerebellum. Neuron 48:31–43 Wassarman KM, Lewandoski M, Campbell K et al (1997) Specification of the anterior hindbrain and establishment of a normal mid/hindbrain organizer is dependent on Gbx2 gene function. Development 124:2923–2934 Wonders CP, Anderson SA (2006) The origin and specification of cortical interneurons. Nat Rev Neurosci 7:687–696 Wurst W, Auerbach AB, Joyner AL (1994) Multiple developmental defects in Engrailed-1 mutant mice: an early mid-hindbrain deletion and patterning defects in forelimbs and sternum. Development 120:2065–2075 Wurst W, Bally-Cuif L (2001) Neural plate patterning: upstream and downstream of the isthmic organizer. Nat Rev Neurosci 2:99–108 Ye W, Bouchard M, Stone D et al (2001) Distinct regulators control the expression of the midhindbrain organizer signal FGF8. Nat Neurosci 4:1175–1181 Yu T, Yaguchi Y, Echevarria D et al (2011) Sprouty genes prevent excessive FGF signaling in multiple cell types throughout development of the cerebellum. Development 138:2957–2968 Zervas M, Blaess S, Joyner AL (2005) Classical embryological studies and modern genetic analysis of midbrain and cerebellum development. Curr Top Dev Biol 69:101–138 Zinyk DL, Mercer EH, Harris E et al (1998) Fate mapping of the mouse midbrain-hindbrain constriction using a site-specific recombination system. Curr Biol 8:665–668 Zordan P, Croci L, Hawkes R et al (2008) Comparative analysis of proneural gene expression in the embryonic cerebellum. Dev Dyn 237:1726–1735

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Proneural Genes and Cerebellar Neurogenesis in the Ventricular Zone and Upper Rhombic Lip G. Giacomo Consalez, Marta Florio, Luca Massimino, and Laura Croci

Abstract

The cerebellar primordium arises between embryonic days 8.5 and 9.5 from dorsal rhombomere 1, adjacent to the fourth ventricle. Cerebellar patterning requires the concerted action of morphogens secreted by the rhombic lip and roof plate, and leads to the formation of two main neurogenetic centers, the upper rhombic lip and ventricular zone, from which glutamatergic and GABAergic neurons arise, respectively. These territories contain gene expression microdomains that are partially overlapping and, among others, express proneural genes. This gene family is tightly conserved in evolution and encodes basic helix-loop-helix transcription factors implicated in many neurogenetic events, ranging from cell-fate specification to terminal differentiation of a variety of neuronal types across the embryonic nervous system. The present paper deals with the established or suggested roles of proneural genes in cerebellar neurogenesis. Of the proneural genes examined in this chapter, Atoh1 plays a quintessential role in the specification and development of granule cells and other cerebellar glutamatergic neurons. Besides playing key roles at early stages in these early developmental events, Atoh1 is a key player in the clonal expansion of GC progenitors of the external granule layer. NeuroD, formerly regarded as a proneural gene, acts as a master gene in granule cell differentiation, survival, and dendrite formation. Ascl1 participates in GABA interneuron and cerebellar nuclei neurons generation, and suppresses astrogliogenesis. Conversely, little is known to date about the role(s) of Neurog1 and Neurog2 in cerebellar neurogenesis, and a combination of loss- and gain-of-function studies is required to elucidate their role, if any, in cerebellar neurogenesis.

G.G. Consalez (*) • M. Florio • L. Massimino • L. Croci Division of Neuroscience, San Raffaele Scientific Institute, Via Olgettina 58, Milan, 20132, Italy e-mail: [email protected], [email protected], [email protected], [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_2, # Springer Science+Business Media Dordrecht 2013

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Introduction The mature cerebellum represents only 10% of the total brain volume, but contains the majority (80–85%) of human neurons (reviewed in Herrup and Kuemerle 1997). It is the primary center of motor coordination and it is essential for cognitive processing and sensory discrimination (Schmahmann 2004). In humans, the cerebellum achieves its mature configuration only many months after birth and for this reason it is especially vulnerable to developmental abnormalities. Like the cerebrum, the cerebellum comprises an outer cortical structure, a layer of white matter, and a set of cerebellar nuclei (CN) beneath the white matter. The CN project efferent fibers to the thalamus, brainstem, and spinal cord (Paxinos 1995) mediating the fine control of motor movements and balance. During mouse embryonic development, the cerebellum arises between embryonic days (E)8.5 and 9.5 from dorsal rhombomere 1 (r1), adjacent to the fourth ventricle, due to the mutual interactions established by patterning genes, including Gbx2, Otx2, Fgf8, Wnt1, En1, En2, Pax2, Pax5, and others (reviewed in Liu and Joyner 2001). Cerebellar development requires a contribution from the posterior mesencephalon (Martinez and Alvarado-Mallart 1987) and the alar plate of rhombomere 2 (Marin and Puelles 1995). Normal patterning of the cerebellar anlage (Fig. 2.1a, b) depends on the formation and function of the isthmic organizer, a signaling center secreting the morphogens fibroblast growth factor 8 (FGF8) and WNT1, that defines the cerebellar territory along the anterior– posterior axis of the central nervous system (Martinez and Alvarado-Mallart 1987; Marin and Puelles 1994; Crossley et al. 1996; Wassarman et al. 1997; Martinez et al. 1999) and requires a contribution by the roof plate, secreting WNT and bone morphogenetic protein (BMP) ligands (Alder et al. 1999; Chizhikov and Millen 2004; Chizhikov et al. 2006). Cerebellar patterning leads to the formation of two germinal centers that will give rise to the multitude of neuronal types and subtypes observed in the mature cerebellum. These germinal epithelia, called ventricular zone (VZ) and upper rhombic lip (URL), contain gene expression microdomains that are partially overlapping, and that will regulate the genesis of neuronal precursors fated to adopt GABAergic and glutamatergic phenotypes, respectively. Among other genes expressed in these microdomains are four proneural genes, namely neurogenin1 and neurogenin2 (Neurog1/Ngn1, Neurog2/Ngn2) and achaete/scute like1 (Ascl1/Mash1) in the VZ, as well as Atonal homolog1 (Atoh1/Math1) in the URL (Fig. 2.1c). Proneural genes in Drosophila melanogaster development. The defining event of neurogenesis is the switch from uncommitted, cycling progenitor cells to committed neuronal precursors. Nervous system development in Drosophila melanogaster has served as a paradigm for the discovery and dissection of neurogenetic processes and their regulation, providing a conceptual framework for the understanding of mammalian neurogenesis. In Drosophila, the entire nervous system arises from neuroectodermal cells, which delaminate from the surface epithelium and migrate into the interior of the embryo to form neural precursor

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Compartments

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mesencephalon isthmic organizer

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FGF8, WNTs sub-a

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sagittal view anterior to the left

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Fig. 2.1 Schematic of a cerebellar primordium with respect to patterning (a, b), neurogenesis (gene names in red boldface, c) and neuronal migration. See text for details. In d, vertical upward arrows indicate radially migrating ventricular zone progenitors, curved arrows indicate granule cell precursors (purple) and cerebellar neuron precursors (black). Abbreviations: ctz cortical transitory zone, egl external granule layer, ntz nuclear transitory zone, rl upper rhombic lip (abbreviated as URL in the text), rp roof plate, vz ventricular zone

cells, or neuroblasts. The first step of neural fate determination in the Drosophila nervous system is the singling-out of neuroblasts from the neuroectodermal epithelium. Prior to such cell-selection process, all the ectodermal precursors share an equivalent potentiality to become either neuroblasts or epidermoblasts. The choice between these alternative fates was proven to rely on the expression of a small group of transcription factors, belonging to the basic Helix-Loop-Helix (bHLH) family, which instruct neuroectodermal cells to take up the fate of neural precursors – from which they were termed proneural genes (reviewed in Jan and Jan 1994). Several expression studies have shown that a combination of upstream regulatory genes acts on the promoter regions of proneural genes to induce their

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expression within the so-called proneural clusters, i.e., regularly spaced patches of cells, all of which initially share an equivalent potential to become neural progenitors (Skeath and Doe 1996). Progressively, through a cell-cell interaction process called lateral inhibition, clustered progenitors start competing one another. As a result of this competition, the expression of proneural genes becomes restricted to one single prevailing cell, which will delaminate to form the neuroblast and will maintain and further upregulate proneural gene expression. Conversely and simultaneously, the remaining progenitor cells within the cluster receive inhibitory signals from the predominant neural precursor cell, they partially or completely downregulate proneural gene expression, thereby acquiring the alternative fate of epidermoblasts (Artavanis-Tsakonas et al. 1999). Finally, singled-out neural progenitors undergo a stereotyped pattern of cell division producing a fixed number of daughter cells, which will terminally differentiate into the distinct cell types characteristic of the mature fly nervous system: neurons and associated support cells of the sensory organs (Jan and Jan 1994). The final cell type that a given neuron is fated to adopt is decided in a hierarchical, stepwise fashion. At each step, the neuroblast restricts its developmental potential along specific neuronal lineages, undergoing binary or multiple cell-fate choices at determined phenotypic branch-points of the cell-type specification process. As previously mentioned, a striking observation, first made in Drosophila, is the fact that the expression of particular proneural genes is restricted to different neuronal lineages. Indeed, genes belonging to the as-c complex were shown to be specifically involved in the determination of neuroblasts in external sense organs; ato provides the competence to form chordotonal organs (a type of internal sense organs) and photoreceptors; amos is employed in the formation of multidendritic neurons and olfactory sensilla. Such observations raised the possibility that proneural genes might be involved not only in triggering a “generic” program of neural determination (epidermis versus neuroblast) but also in the subsequent specification of a given neural identity (e.g., multidendritic neurons versus chordotonal organs). Support to this hypothesis came from gain-of-function experiments: the ectopic expression of as-c induces the formation of ectopic external sense organs while the forced expression of amos leads to ectopic multidendritic neurons and olfactory sensilla (Rodriguez 1990; Chien 1996; Parras 1996; Goulding et al. 2000). Drosophila proneural genes had been initially subdivided into two classes, based on their function: (a) determination genes, like ac, sc or amos – dominantly expressed before any sign of neuronal differentiation and acting in the sorting-out of neural progenitors from the neuroectoderm; and (b) differentiation genes, like cato and biparous – expressed after the selection of neural precursors and involved in the process of neuronal differentiation per se. The sequential activation of these two classes of proneural genes – in a regulatory cascade in which early expressed determination genes induce downstream effectors of differentiation – was in agreement with the stepwise progression of undifferentiated progenitors toward differentiated neural cells (Campuzano and Modolell 1992). Indeed, a classical concept of experimental embryology is that the transition from an undifferentiated

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to a fully differentiated cell comprises two steps: cell specification/determination and cell differentiation. Nevertheless, such functional categorization of proneural genes turned out not to be completely exhaustive nor completely explanatory, as many proneural genes, like atonal, were proven to exert both functions or either, i.e., to trigger a generic program of neural development and/or to induce neuronal lineage or subtypespecific programs, depending primarily on the temporal- and regional-specific cues active throughout their expression window. Furthermore, an individual bHLH protein could be required for several different cell types at different times or locations during development. Factors affecting the outcome of proneural gene function include the context (cellular and genetic) and timing of their expression. More sophisticated criteria were therefore required to assess proneural function in neural development in order to account for such functional complexity. Placing proneural genes in the context of temporal and, whenever possible, epistatic cascades has been a major goal in the past few years. Correlating these cascades with precise stages and cellular events in neuronal differentiation remains an open challenge. The roles of proneural genes in vertebrate neurogenesis Cell-fate decisions in the development of the vertebrate and invertebrate nervous systems are controlled by remarkably similar mechanisms. Among other ontogenetic programs, the coupling of neural determination and lineage/cell-type specification, by means of the combined action of proneural genes and positional identity determinants, is a mechanism strikingly conserved in evolution. Unsurprisingly, vertebrate genomes were proven to contain several orthologs of Drosophila proneural genes (Ledent and Vervoort 2001; Simionato et al. 2007). Based on the homology to their Drosophila counterparts, such genes have been isolated, cloned, and shown to play a pivotal role in vertebrate neurogenesis (Ghysen and Dambly-Chaudiere 1988; Lee 1997). As in Drosophila vertebrate proneural genes commit cycling progenitor cells to a neuronal fate, which involves activation of Notch signaling and the induction of cell-cycle exit. As in Drosophila development, vertebrate proneural genes act in regulatory cascades, with early expressed genes inducing fate specification of neural progenitors, and later-expressed genes, regulating terminal differentiation (Cau et al. 1997; Lee 1997). In terms of sequence conservation in the bHLH domain, the mouse atonal homologs Atoh1 and Atoh5 are the genes most closely related to Drosophila atonal. Loss-of-function studies in the mouse have shown that Atoh1 is necessary for the generation of cerebellar granule neurons (see further), and for the development of the sensory epithelium of the inner ear (Ben-Arie et al. 1997; Chen et al. 2002), while Atoh5 is essential for retinal ganglion cell production (Wang 2001). Close members of the ato family were also found in Xenopus laevis – Xath1 and Xath5 – where they have been shown to induce a neural fate (Kanekar et al. 1997; Kim 1997). In Xenopus laevis, overexpression of Neurog1 leads to a massive ectopic formation of XneuroD-positive neurons (Ma et al. 1996). Incidentally, NeuroD, a gene initially labeled as a proneural gene by virtue of its homology to other proneural

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factors, and of its ability to convert presumptive dorsal epidermis into neurons in Xenopus laevis (Lee et al. 1995), is in fact involved in later stages of neuronal differentiation in vertebrates. Again, in addition to their generic neurogenetic role, numerous lines of evidence indicate that proneural genes also specify neuronal subtype identity. For instance, in the neocortex, the proneural genes Neurog1 and Neurog2 are absolutely required to specify the identities of early-born, deep layer neurons, whereas they are dispensable for later stages of neuronal fate specification (Schuurmans et al. 2004). In the developing spinal neural tube, Atoh1 on one hand and Neurog1 and Ascl1 on the other are expressed in nonoverlapping patterns, and cross-inhibition occurs between the two sets of genes (Gowan et al. 2001). Furthermore, constitutive or ectopic expression of neurogenins, and other atonal homologs, yields specific neuronal subtypes in vivo (Blader et al. 1997; Kanekar et al. 1997; Olson et al. 1998; Perez et al. 1999; Gowan et al. 2001). Neurog1 or Neurog2 single-mutant mice lack complementary sets of cranial sensory ganglia, and Neurog1/2 double mutants lack, in addition, spinal sensory ganglia and a large fraction of ventral spinal cord neurons (Fode et al. 1998; Ma et al. 1998; Ma et al. 1999). Likewise, loss-of-function and gain-of-function analyses have shown that Ascl1 contributes to the specification of neuronal subtype identity (Fode et al. 2000) and is both required and sufficient to specify ventral telencephalic fates. Further evidence indicates that Ascl1 acts cooperatively in sympathetic ganglia as an instructive determinant of cell fate to induce the noradrenergic phenotype (Goridis and Brunet 1999). Ascl1 has also been implicated in the differentiation of earlyborn (but not late-born) neurons in the striatum, indicating that it regulates both progenitor cell behavior and neuronal fate specification in a temporally defined manner (Casarosa et al. 1999; Yun et al. 2002). The present chapter deals with the established or presumptive roles of proneural genes in the context of neuronal type specification, determination, differentiation, and clonal expansion at various stages in the course of cerebellar neurogenesis. In particular, this chapter will provide a review of the literature as regards the roles played in that context by four established proneural genes and by one of their targets, NeuroD, formerly considered a proneural gene. Emphasis will be placed on the evidence accrued from the analysis of wild-type and transgenic mice.

Atoh1, the Master Gene in Granule Cell Development More than a century ago, the URL was recognized as the origin of the most numerous neuronal population in the CNS, i.e., cerebellar granule cells (GCs), that first migrate tangentially (Fig. 2.1d, purple arrow) and then radially during embryonic and postnatal development, respectively (Cajal 1889; Schaper 1897). During postnatal proliferation in the external granule layer (EGL), GC precursors maintain Atoh1 expression until they start migrating inward to form the internal granule layer (IGL) (Hatten and Heintz 1995; Helms et al. 2001). Atoh1 is the murine homolog of the Drosophila proneural gene atonal, as shown by sequence similarity and functional conservation in evolution (Ben-Arie et al. 2000).

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It encodes the homonymous bHLH transcription factor, and is expressed starting at E9.5 in the dorsal neural tube, from rhombomere 1 to the tail (Akazawa et al. 1995). In the URL, Atoh1 is expressed in progenitors distinct from Lmx1a+ roof plate cells (Chizhikov et al. 2006). Atoh1 plays key roles in the development of all glutamatergic neurons of the cerebellum. The cerebellum of Atoh1 / mice is reduced in size and displays no foliation. In Atoh1 null embryos, unlike at earlier stages (Chizhikov et al. 2006), BrdU incorporation is strikingly reduced in the rhombic lip at E14.5, and the EGL fails to form, resulting in the complete absence of cerebellar GCs (Ben-Arie et al. 1997; Bermingham et al. 2001). Instead, Purkinje cells (PCs), which form normally in the cerebellar VZ, migrate into the outermost part of the cerebellum and form a rudimentary PC layer, although some PCs fail to migrate toward the cortex and are retained in deep regions of the cerebellar primordium. Balanced levels of Atoh1 expression are essential for the correct timing of GC precursor differentiation. As mentioned, Atoh1-null mice have a small, poorly foliated cerebellum devoid of GCs. However, while Atoh1 overexpression leads to the upregulation of early differentiation markers (e.g., NeuroD, Dcx), it is insufficient to promote a complete differentiation of GC precursors (Helms et al. 2001). This paradoxical effect can be explained by the fact that, physiologically, Atoh1 is downregulated by its own protein product through a negative regulatory feedback loop (Gazit et al. 2004), and by other factors such as Notch intracellular domain, and the zinc-finger transcription factor Zic1 at the onset of GC precursor migration from the EGL to the IGL (Ebert et al. 2003). Likewise, in vitro experiments demonstrated that bone morphogenetic protein (BMP) signaling activation leads to the post-translational inactivation of Atoh1, which is targeted to the proteasome for degradation (Zhao et al. 2008). Atoh1 plays a key role in granule cell clonal expansion. Peak cerebellar growth occurs relatively late compared to the rest of the brain, driven primarily by proliferation of EGL cells. In the mouse, starting at the end of embryonic development (around E16.5), GC precursors resident in the EGL start undergoing an impressive wave of clonal expansion, before differentiating and undertaking their radial migration into the IGL. The highest growth rate is recorded during the first 2 weeks of postnatal life, while in humans, the corresponding proliferative peak occurs in utero during the third trimester, although EGL remnants can persist for up to a year after birth (Abraham et al. 2001). This prolonged ontogenetic period makes the cerebellum susceptible to developmental aberrations and tumor formation. Medulloblastoma (MB), the most common brain tumor of childhood, has been studied by histological means and, more recently, by molecular analysis (Eberhart et al. 2000; Thompson et al. 2006). Physiologically, the clonal expansion phase is promoted non-cell-autonomously by PCs, as first shown by John Oberdick and coworkers through transgenic expression of the diphtheria toxin in these neurons, leading to the virtual abolition of GC precursor proliferation in the EGL (Smeyne et al. 1995). This non-cell-autonomous effect is driven by the secreted morphogen and mitogen sonic hedgehog (SHH), as shown by Matthew Scott’s group and by others (Dahmane and Ruiz-i-Altaba 1999; Wallace 1999; Wechsler-Reya and Scott 1999). In fact, conditional knock-out mice in which SHH is specifically

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removed from PCs, MacMahon and coworkers (Lewis et al. 2004) observed a drastic reduction of GC precursors expansion. Accordingly, heterozygous mutants for the inhibitory SHH receptor patched, in which SHH signaling (reviewed in Villavicencio et al. 2000) is upregulated in a ligand-independent fashion, are prone to the development of medulloblastoma (MB)-like tumors. Among the MB subtypes that have been characterized in patients, the desmoplasmic variant seems to be derived from Atoh1+ GC precursors in the EGL (Pomeroy et al. 2002; Salsano et al. 2004). Collectively, this evidence prompted questions as to whether Atoh1 plays any role in GC precursor proliferation. Zoghbi and coworkers addressed this point in a paper published in 2009 (Flora et al. 2009) showing that Atoh1 is required for the peri- and postnatal expansion of GC progenitors. To investigate the molecular effects of Atoh1 deletion, they isolated GC progenitors from Atoh1flox/flox P5 mice and infected them with adenoviruses expressing either the green fluorescent protein (GFP) or the Cre recombinase gene, and cultured transduced cells in the presence of SHH to evaluate their proliferative status. Their results showed that Atoh1 deletion led to a sharp decrease in cell proliferation, suggesting that, in those cells, the response to SHH is dependent on Atoh1. Through various approaches, they showed that Atoh1 mediates this response by transcriptionally upregulating the expression of Gli2, a critical mediator of SHH signaling (reviewed in Villavicencio et al. 2000). Next, they asked whether its expression might be required for the genesis of MBs induced by constitutive activation of the SHH pathway. To address this point they crossed mice harboring a conditional deletion of Atoh1 with tamoxifen-inducible Cre deletors expressing a constitutively active form of the SHH coreceptor smoothened. While, in the presence of Atoh1, smoothened overexpression caused hyperplasia of the EGL, this effect was drastically quenched in mice carrying a homozygous Atoh1 deletion. Different glutamatergic neurons derive from Atoh1+ progenitors. Some Atoh1+ cells migrate from the RL to the nuclear transitory zone (NTZ) (Jensen et al. 2004) (black downturned arrow in Fig. 2.1d), a transitory cell cluster that will give rise to the cerebellar nuclei (CN). A paper published in 2005 by Gord Fishell’s group (Machold and Fishell 2005) clearly revealed that GCs are not the only glutamatergic lineage derived from Atoh1+ progenitors of the URL. This work was conducted taking advantage of a mouse line expressing a tamoxifen-inducible form of Cre recombinase (CreERT2) under transcriptional control of the Atoh1 promoter. This approach, dubbed genetic inducible fate mapping (Joyner and Zervas 2006), allowed the authors to tag several waves of neuronal progenitors born at successive developmental time points, determining their final fate and location in the adult brainstem and cerebellum, thus shedding light on the spatiotemporal regulation of cerebellar glutamatergic neurogenesis. The results of this study clearly demonstrated that, prior to E12.5, Atoh1 is transiently expressed in cohorts of cerebellar rhombic lip neural precursors that populate the anterior hindbrain and deep cerebellar nuclei in the adult. Starting at E12.5, Atoh1+ progenitors start giving rise mostly to granule cells that will populate the anterior

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lobe; finally, Atoh1+ progenitors born at later stages until E16.5 will progressively populate the entire EGL and, after inward migration, the IGL of all cerebellar lobules. A parallel study, conducted by Zoghbi and colleagues (Wang et al. 2005), took advantage of an Atoh1LacZ/+ knock-in line to analyze the migration and fate of Atoh1+ progenitors and corroborated the findings of Machold and Fishell. In addition, this study, using an Atoh1LacZ/ null mutant, demonstrated that Atoh1 is essential for the establishment of the NTZ first, and eventually for the formation of the glutamatergic component of CNs. Thus, Atoh1 is required to support the development of both GCs and the glutamatergic CN neurons. Late Atoh1+ progenitors in the URL give rise to unipolar brush cells. Unipolar brush cells (UBCs) are the other glutamatergic neuron located in the adult IGL. They were first described for their morphology: they feature a single brush-like dendritic ending (Harris et al. 1993; Mugnaini and Floris 1994) and project their axons to GCs and to other UBCs (Nunzi et al. 2001). In 2006, Hevner and colleagues showed that UBCs also originate from the Atoh1+ URL that produces GCs, except they do so later on in development (Englund et al. 2006). UBC precursors are born in the URL between E15.5 and E17.5, and migrate into the prospective white matter during late embryonic and early postnatal development. Noteworthily, while Atoh1 / mice show the complete loss of GC precursors, UBCs were severely reduced but not completely depleted in the same mutants. Conclusions. In the context of cerebellar neurogenesis, Atoh1 expression in the URL plays a quintessential role in the specification and development of GC precursors and of progenitors of other neurons, namely glutamatergic ones populating the CN, besides several ventral hindbrain nuclei. While Atoh1 is expressed in UBC progenitors, it is not strictly required for the determination of this specific cell fate. Besides playing key roles at early stages in these early developmental events, Atoh1 is a key player in the clonal expansion of GC progenitors in the EGL at the end of their tangential migration from the URL (Fig. 2.1d).

NeuroD, an “Almost Proneural” Gene with Key Roles in Cerebellar Development NeuroD is a bHLH transcription factor originally studied in Xenopus laevis for its ability to convert embryonic epithelial cells into differentiated neurons and for its role in promoting cell-cycle exit and neuronal differentiation (Lee et al. 1995). A loss-of-function approach was used to study NeuroD functions during brain development (Miyata et al. 1999). NeuroD null mice died shortly after birth due to diabetes. In the study conducted by Myata et al., the authors rescued this phenotype introducing a transgene that encodes the mouse NeuroD gene under control of the insulin promoter. The rescued null mice featured an ataxic gait, walked around incessantly, and failed to balance themselves. A histological analysis performed at P30 revealed a severe reduction of cerebellar GCs in

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the posterior lobules (VI-X), whereas a small population of GCs survived in the anterior cerebellum. Secondary to GC depletion, PCs of the posterior cerebellum failed to arrange into a proper monolayer. However, PC development appeared normal in terms of PC-specific marker expression. At birth, NeuroD is expressed in the inner layer of the EGL where post-mitotic granule cells reside, and in the postmigratory GCs located in the IGL. In null mice, at birth, Miyata et al. found an increased rate of cell death in the inner half of the posterior EGL, which harbors post-mitotic precursors. GC death continued until P30, indicating a degeneration of the surviving GCs of the IGL. The authors concluded that NeuroD regulates a transcriptional cascade of genes that are essential for the differentiation and survival of post-mitotic cerebellar GCs. Another study, conducted by Azad Bonni’s group at Harvard University (Gaudilliere et al. 2004), revealed a novel role for NeuroD in regulating GC neuron dendritic morphogenesis. They demonstrated that the knockdown of NeuroD expression by RNA interference in both primary cerebellar GC culture and organotypic cerebellar slices resulted in a profound alteration of dendritic morphogenesis, while it had no effect on axonal growth. Moreover, they demonstrated that neuronal activity leads to the phosphorylation of NeuroD by the protein kinase CAMKII. This event activates a downstream intracellular signaling pathway that regulates dendritogenesis in cerebellar granule neurons. Additional studies will be necessary to fully dissect the transcriptional machinery acting downstream of NeuroD and responsible for the growth and maintenance of granule cell neuron dendrites. Conclusions. Taken together, the above findings demonstrate that NeuroD acts as a master gene in the context of cell-intrinsic transcriptional mechanisms that guide GC differentiation, survival, and dendrite formation. It is worth mentioning that NeuroD is not exclusively expressed in the progeny of URL progenitors. Expression of this gene is clearly detectable in the cerebellar cortical transitory zone (CTZ), a post-mitotic compartment hosting PC- and other GABAergic precursors (L. Croci and G.G. Consalez, unpublished observation; Lee et al. 2000).

Ascl1 in Ventricular Zone Neurogenesis Ptf1a is a master gene of cerebellar GABAergic neurogenesis. Although the cerebellum contains a relatively small variety of neurons, the molecular machinery governing neuronal generation and/or subtype specification is still poorly understood. In 2005, Hoshino et al. (2005) published the characterization of a novel mutant mouse, cerebelless, which lacks the entire cerebellar cortex but survives up to adult stages. The analysis of its phenotype, and the characterization of the underlying gene mutation, clarified that Ptf1a (pancreas transcription factor 1a), which encodes a bHLH transcription factor, is required for generating the cerebellar GABAergic compartment. Atoh1 and Ptf1a participate in regionalizing the cerebellar neuroepithelium, and define two distinct areas, the VZ (Ptf1a positive) and the URL (Atoh1 positive), which generate GABAergic and glutamatergic neurons,

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respectively (Hoshino et al. 2005; Pascual et al. 2007). Although Ptf1a is not a proneural gene, the expression of three proneural genes (Ascl1, Neurog1, and Neurog2) overlaps with that of Ptf1 in the VZ, warranting this brief foreword. Ascl1 labels the cerebellar GABAergic lineage. While a large number of studies have investigated the distribution and roles of Atoh1 in the cerebellar glutamatergic lineage, fewer studies have been devoted to analyzing GABAergic precursors born in the cerebellar VZ, and to dissecting the roles of proneural genes in this context. In 2008, Zordan et al. (2008) published a systematic descriptive analysis of proneural gene expression at early stages of mouse cerebellar development. The results of this study established that at the onset of cerebellar neurogenesis, starting at about E11, the Ascl1 transcript becomes detectable by in situ hybridization in the VZ and presumptive NTZ. A similar distribution is observed at later stages, with the Ascl1 transcript occupying the entire thickness of the Ptf1a + VZ all the way to its apical (ventricular) margin. Accordingly, the territories occupied by Ascl1 and Atoh1 are clearly complementary. The Ascl1 transcript remains confined to the VZ until E13.5. An additional study by Jane Johnson and coworkers (Kim et al. 2008), published 1 month later, reported the results of genetic fate mapping performed using two transgenic Ascl1-Cre lines, one of which expressed a tamoxifen-inducible Cre recombinase, CreERTM (Helms et al. 2005; Battiste et al. 2007) and two Creinducible reporter lines (Soriano 1999; Srinivas et al. 2001). The evidence produced in this elegant lineage analysis study was in full agreement with the conclusions drawn by Zordan et al.: in particular, Ascl1+ progenitors are initially (E12.5) restricted to the cerebellar VZ and excluded both from the post-mitotic CTZ and from the rhombic lip migratory stream. The locations of Ascl1+ and Atoh1+ progenitors are mutually exclusive, whereas a high degree of overlap was shown between Ptf1a+ and Ascl1+ progenitors, revealing that Ascl1 labels GABAergic neuronal progenitors. However, at later stages (E17.5), Ascl1+ progenitors were no longer confined to the VZ; instead, they were scattered throughout the cerebellar primordium. Some of these cells coexpressed Ascl1 and the oligodendrocyte marker Olig2. In addition, the fate mapping analysis contained in this study revealed that Ascl1+ progenitors born either before E11.5 or after E13.5 give rise to the GABAergic components of the CN, whereas progenitors born between E11.5 and E13.5 are fated to become PCs. Finally, when Ascl1+ progenitors are tagged by tamoxifen administration after E16, some of them acquire an oligodendroglial identity and localize to the prospective white matter. In the same study, no colocalization was ever scored between Ascl1+ progenitors and astrocyte-specific markers. Finally, a study authored by Marion Wassef and coworkers (Grimaldi et al. 2009) further refined the analysis of the role of Ascl1 in cerebellar neurogenesis, incorporating a description of the effects of Ascl1 gene disruption and overexpression. With regard to the distribution of Ascl1+ progenitors, Grimaldi et al. established that these precursors progressively delaminate out of the VZ to settle into the prospective white matter first, and cerebellar cortex next. By studying an Ascl1-GFP transgenic mouse, they demonstrated that Ascl1+ progenitors give rise to interneurons positive for Pax2 and to oligodendrocyte precursors positive for

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Olig2. Conversely, glutamatergic neurons as well as astrocytes and Bergmann glia cells never expressed GFP. To clarify the role of Ascl1 in the generation of different cerebellar cell types, the authors analyzed Ascl1 null mice at E18.5. The loss of Ascl1 led to a dramatic reduction of Pax2+ and Olig2+ precursors, whereas astrocytic precursors, labeled by Sox9, were moderately increased in number. No change was found as regards PC development. Furthermore, to circumvent the perinatal lethality of Ascl1 null mice and study the effect of Ascl1 at later stages, the authors transplanted solid grafts obtained from E15.5 mutant- and wild-type cerebella into the cerebral cortex of newborn recipients. After 2 months, they analyzed the grafts and, consistent with previous results, they found that they contained a reduced number of parvalbumin + interneurons, but a normal number of PCs. Finally, the authors resorted to a gain-of-function approach (in vivo electroporation of a GFP plasmid, stage E14.5) to determine whether oligodendrocytes and Pax2+ interneurons are lineally related in the cerebellum. After electroporation, cerebella were dissected and cultured for 6 days in vitro. By comparing the results of intraventricular versus parenchymal electroporations, they concluded that most Ascl1+ oligodendrocytes do not originate from the cerebellar VZ. In addition, they electroporated an Ascl1-GFP plasmid into the cerebellar VZ. This led to an increased number of Pax2+ interneurons, to a reduced number of Olig2+ oligodendrocyte precursors, and to a complete absence of astroglial cells. This prompted them to conclude that Ascl1 overexpression pushes progenitors toward the Pax2 interneuron fate and suppresses the astrocytic fate. Conclusions. Taken together, the above evidence suggests that, in cerebellar neurogenesis, Ascl1 contributes to the GABAergic pool. It participates in GABA interneuron and CN neuron generation, and in PC development. However, it is not required for PC specification, suggesting that, in this process, it may either be irrelevant or act redundantly with other VZ-specific proneural genes. When overexpressed at E14.5, Ascl1 promotes the GABA interneuron phenotype, suppressing the astrocytic fate.

Uncertain Roles for Neurogenins in Cerebellar Gabaergic Development Neurog1 and Neurog2 are expressed in the Ptf1a + ventricular neuroepithelium. As shown by Zordan et al. (2008), the Neurog2 transcript is first observed around E11 in CN neuron progenitors of the cerebellar primordium, whereas Neurog1 appears 1 day later, in a rostral region located between the isthmic organizer, labeled by Fgf8, and the territory marked by Ascl1. At E12.5, both Neurog1 (see also Salsano et al. 2007) and Neurog2 are present in the VZ but with a few differences in distribution: in the anterior cerebellum, Neurog1 is expressed at high levels in a region close to the midline, whereas Neurog2 is detected only in the lateral VZ. In posterior territories, the expression patterns of the two proneural genes overlap completely. Neurog1 and Neurog2 are adjacent and partially overlap

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with post-mitotic domains labeled by Lhx1 and Lhx5 two genes that control PC differentiation (Zhao et al. 2007). This suggests that Neurog1 and Neurog2 are expressed in progenitors that are undertaking the last cycle of cell division, to become post-mitotic PC precursors. At E13.5 the differential anterior boundaries of Neurog1 and Neurog2 are maintained, although the transcript levels of both genes are downregulated. The authors conclude that Neurog1 and Neurog2 are mainly expressed in the cerebellar germinal epithelium that gives rise to GABAergic progenitors, while they are completely absent from the URL, the source of all glutamatergic cerebellar progenitors. Moreover, their expression patterns are similar but not totally overlapping, suggesting that they may contribute to the diversity of cerebellar GABA neurons and, possibly, PC subtypes. Neurog1 is expressed in cerebellar GABAergic interneuron progenitors. In 2009, Doughty and coworkers published a lineage analysis in which, by using transgenic fate mapping, they identified the mature cerebellar neurons deriving from Neurog1-positive cell fates in the developing mouse cerebellum (Lundell et al. 2009). They confirmed the findings reported by Zordan et al. and, in addition, extended the Neurog1 expression analysis to late embryonic and postnatal cerebellar development. At E14-E20, Neurog1 is present in Ptf1a + neurons, but it is excluded from the URL and EGL. Moreover, at P7, it colocalizes with Ptf1a and BrdU in the deep white matter. This suggests that Neurog1 is expressed in early GABAergic interneuron precursors that, shortly after birth, migrate from the white matter to reach their final destination in the cortex. To test their hypothesis the authors used two artificial chromosome (BAC)-reporter mice imported from the NIH GENSAT consortium (Rockefeller University, New York) to study short-term and long-term Neurog1-positive cell fates. The first to be characterized was the Neurog1-EGFP transgenic mouse line, previously used to map short-term cell fates in the developing thalamus (Vue et al. 2007). This line acted as a medium/long-term reporter thanks to the persistence of GFP signal. Their results supported the hypothesis that Neurog1 is expressed in Pax2+ interneuron progenitors, whereas it does not contribute to the GABAergic CN neuron lineage. Surprisingly, they did not reveal any fluorescence in PC neurons. Furthermore, the authors bred Neurog1Cre transgenic mice into the double reporter Z/EG line (Novak et al. 2000). Z/EG mice express a LacZ cassette under control of the CMV enhancer/chicken actin promoter (pCAGGS). In the presence of a Cre recombinase the LacZ cassette is excised, leading to the activation of the downstream EGFP gene. Using this approach, they revealed sporadic PC neurons that were GFP+, mostly located in the hemispheres rather than the vermis. Surprisingly though, this reporter failed to detect any GABAergic interneurons. This discrepancy might have been due to epigenetic/positional silencing of the reporter transgene, or to low-level expression of the Neurog1-Cre transgene. A recent study conducted by Jane Johnson and coworkers confirmed the notion that, in the cerebellar primordium, the Neurog1+ lineage gives rise to PCs (Kim et al. 2011). Conclusions. In summary, Neurog1 is expressed in progenitors giving rise to GABAergic interneurons of the cerebellar cortex and at least to some PCs. However, it does not seem to contribute to the development of CN neurons. While the

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role of Atoh1 in cerebellar neurogenesis has been fully elucidated, and that of Ascl1 has been at least partially clarified, nothing much is known about the role(s) of neurogenins in the same context. While both neurogenin genes are expressed in the cerebellar VZ in presumptive GABAergic neurons, nothing can be inferred to date as regards their function(s). Do they affect cell type or subtype specification, or neuronal versus glial commitment? And in either case, do they act redundantly with each other or with Ascl1, whose expression pattern overlaps with that of neurogenins at early stages? There seems to be some degree of selectivity, in that Neurog1 is expressed in only a share of GABAergic progenitors, and, as predicted by Zordan et al., the broader expression domain exhibited by Neurog2 could suggest that this gene plays a unique role in the development of the GABAergic component of CN neurons. A recent study (Dalgard et al. 2011) describes the results of a genome-wide analysis of gene expression in the cerebellar primordium of E11.5 Neurog1 null mice to identify the Neurog1 transcriptome in the developing cerebellum. This screen identified 117 genes differentially enriched in Neurog1 / versus control sample sets with a high presence of gene sets enriched for functions in nervous system development. Furthermore, their findings suggest that Neurog1 and Pax6 interact functionally in the activation of downstream targets. At any rate, a combination of loss- and gain-of-function studies is now required to elucidate the roles of these genes in cerebellar neurogenesis.

Final Remarks Proneural genes are expressed at crucial stages in cerebellar neurogenesis. Their ascertained roles include the regulation of neuronal fate determination, neuronal type specification, terminal differentiation, and GC clonal expansion. Under all those circumstances they may be part of as yet unclarified combinatorial codes that integrate their function with that of many other genes, particularly those encoding other transcription factors. Further studies are required to dissect the molecular machinery in which they function, in cooperation with regulatory cascades controlling positional identity, fate specification, and differentiation. Acknowledgments Giacomo Consalez’ research has been supported by grants from Ataxia UK, Compagnia di San Paolo, the EU (EuroSyStem), and the Berlucchi Foundation. Further support has come from the Stayton family in honor of Dr. Chester A. Stayton of Indianapolis, Indiana, USA. This review is dedicated to his cherished memory.

References Abraham H, Tornoczky T, Kosztolanyi G, Seress L (2001) Cell formation in the cortical layers of the developing human cerebellum. Int J Dev Neurosci 19:53–62 Akazawa C, Ishibashi M, Shimizu C, Nakanishi S, Kageyama R (1995) A mammalian helixloop-helix factor structurally related to the product of Drosophila proneural gene atonal is a positive transcriptional regulator expressed in the developing nervous system. J Biol Chem 270:8730–8738

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and differentiation of progenitor cell types in the subcortical telencephalon. Development 129:5029–5040 Zhao Y, Kwan K-M, Mailloux C, Lee W-K, Grinberg A, Wurst W, Behringer R, Westphal H (2007) LIM-homeodomain proteins Lhx1 and Lhx5, and their cofactors Ldb1, control Purkinje cell differentiation in the developing cerebellum. Proc Natl Acad Sci USA 104:13182–13186 Zhao H, Ayrault O, Zindy F, Kim JH, Roussel MF (2008) Post-transcriptional down-regulation of Atoh1/Math1 by bone morphogenic proteins suppresses medulloblastoma development. Genes Dev 22:722–727 Zordan P, Croci L, Hawkes R, Consalez GG (2008) Comparative analysis of proneural gene expression in the embryonic cerebellum. Dev Dyn 237:1726–1735

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Zones and Stripes: Development of Cerebellar Topography Roy V. Sillitoe and Richard Hawkes

Abstract

Cerebellar architecture is organized around the Purkinje cell. Purkinje cells in the mouse cerebellum come in many different subtypes, organized first into four transverse zones and then further grouped into hundreds of reproducible topographical units – stripes. Stripes are identified by their functional properties, connectivity, and expression profiles. The molecular pattern of stripes is highly reproducible between individuals and conserved through evolution. Pattern formation in the cerebellar cortex is a multistage process that begins with the generation of the Purkinje cells in the ventricular zone (VZ) of the fourth ventricle. During this stage or shortly after, Purkinje cell subtypes are specified toward specific positions. Purkinje cells migrate from the VZ to form an array of clusters that form the framework for cerebellar topography. At around birth these clusters disperse, triggered by a Reelin signaling pathway, to form the adult stripe array. The chapter will begin with a brief overview of adult cerebellar topography, primarily focusing on the mouse cerebellum, and then discuss the cellular and molecular mechanisms that establish these remarkable patterns.

R.V. Sillitoe Dominick P. Purpura Department of Neuroscience, Albert Einstein College of Medicine 812 Kennedy Center, 1410 Pelham Parkway South, Bronx, NY, 10461, USA e-mail: [email protected] R. Hawkes (*) Department of Cell Biology and Anatomy Genes and Development Research Group, and Hotchkiss Brain Institute, The University of Calgary, 3330 Hospital Drive NW, Calgary, AB, T2N 4N1, Canada e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_3, # Springer Science+Business Media Dordrecht 2013

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The Architecture of the Adult Cerebellar Cortex The adult mouse cerebellum is shown in Fig. 3.1, immunoperoxidase stained for the antigen zebrin II (Brochu et al. 1990: zebrin II ¼ aldolase C (AldoC) – Ahn et al. 1994). There are two subsets of Purkinje cells: zebrin II-immunopositive (zebrin II+) and zebrin II-immunonegative (zebrin II-). Purkinje cells in each subset are aligned to form an alternating array of parasagittal stripes (Brochu et al. 1990; Sillitoe and Hawkes 2002). Stripes are reproducible between individuals and symmetrically distributed about the midline (Hawkes et al. 1985; Hawkes and Leclerc 1987; Brochu et al. 1990). Zebrin II+ stripes are numbered as P1+  P7+ from the midline laterally, and the intervening zebrin II- stripes are numbered with reference to the medial zebrin II+ stripe (i.e., P1- lies immediately lateral to P1+, etc.). In the vermis, four transverse domains in the anterior–posterior axis are identified by zebrin II expression: the striped anterior zone (AZ: lobules I–V), the uniformly zebrin II+ central zone (CZ: lobules VI–VII), the striped posterior zone (PZ: lobules VIII–dorsal IX), and the uniformly zebrin II+ nodular zone (NZ: lobules IX ventral and X: Ozol et al. 1999). A similar alternation of zones is seen in the hemispheres (Sarna et al. 2006).

a

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L7/Pcp2-hAP (E15)

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Fig. 3.1 The mouse cerebellum is organized into an array of transverse zones and parasagittal stripes. (a): Adult mouse cerebellum immunoperoxidase stained in whole mount with anti-zebrin II/Aldolase C. (b): Schematic illustrating the pattern of zebrin II in the mouse cerebellum. (c): Embryonic day (E)15 mouse cerebellum stained in whole mount for alkaline phosphatase (hAP) to detect the expression of an L7/Pcp2-hAP transgene (see Sillitoe et al. 2009 for details). (d): Schematic illustrating the pattern of embryonic Purkinje cell clusters as revealed by hAP staining in L7/Pcp2-hAP transgenic mice. Abbreviations: AZ anterior zone, CZ central zone, PZ posterior zone, NZ nodular zone, Sim simplex, Fl/Pfl flocculus/paraflocculus, Pmd paramedian, Cop copula pyramidis. Lobule numbers are indicated by Roman numerals and stripes are labeled with Arabic numerals (Panels C and D were adapted from Sillitoe et al. 2009)

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Numerous molecular markers are co-localized with either the zebrin II+ or zebrin II- Purkinje cells. For example, the GABA-B receptor is expressed in the zebrin II+ population (Chung et al. 2008a) and phospholipase C(PLC)ß4 in the zebrin II- population (Sarna et al. 2006). However, detailed comparisons between zebrin II expression and other antigenic markers reveal that the parasagittal stripes are much more elaborate than the expression of any one antigen indicates. For example, comparisons between zebrin II and the glycoprotein epitope HNK1 reveal that although these two antigens are largely co-localized (Eisenman and Hawkes 1993), in several lobules discrete Purkinje cell populations express both antigens (Marzban et al. 2004). Similarly, expression of the 25 kDa small heat shock protein (HSP)25 reveals parasagittal Purkinje cell heterogeneity in both the CZ and NZ – areas in which zebrin II is homogeneously expressed in all Purkinje cells (Armstrong et al. 2000). As a result, the adult cerebellar cortex of the mouse can reliably and reproducibly be subdivided into several hundred distinct regions, each typically comprising no more than a few hundred Purkinje cells (e.g., reviewed in Hawkes 1997; Sarna and Hawkes 2003; Apps and Hawkes 2009). Stripe and zone compartments influence all aspects of cerebellar biology. They are highly reproducible between individuals, conserved through evolution (AZ – Sillitoe et al. 2005; PZ – Marzban and Hawkes 2010), and insensitive to experimental manipulation (reviewed in Larouche et al. 2006). Afferent topography is also striped. Zone and stripe boundaries restrict afferent terminal fields (e.g., climbing fibers, spinocerebellar mossy fibers, and trigeminocerebellar mossy fibers that relay somatosensory signals terminate mainly in zebrin II- stripes throughout the AZ and into rostral lobule VI, where the AZ interdigitates with the CZ: e.g., reviewed in Voogd and Ruigrok 2004) into compartments that are reflected by functional cerebellar maps (e.g., Chockkan and Hawkes 1994; Hallem et al. 1999). Many cerebellar mutant phenotypes are restricted by zone and stripe boundaries. For example, swaying (Thomas et al. 1991), rostral cerebellar malformation/ Unc5h3 (Napieralski and Eisenman 1996), cerebellar deficient folia (Cook et al. 1997; Beierbach et al. 2001; Park et al. 2002), and meander tail (Ross et al. 1990) all exhibit deficits restricted primarily to the AZ; the gain of function d2 glutamate receptor mutant lurcher (Lc/+) has a zebrin II expression domain during development that is restricted at the CZ/PZ boundary (Tano et al. 1992); and the weaver mouse exhibits a Purkinje cell ectopia that is primarily restricted to the CZ (Eisenman et al. 1998; Armstrong and Hawkes 2001). Finally, most examples of Purkinje cell death due to mutation or insult show restriction to parasagittal stripes (reviewed in Sarna and Hawkes 2003; Duffin et al. 2010; Armstrong et al. 2010). How does this remarkable zone-and-stripe pattern develop?

From Allocation to Rhombomere 1 to Two Germinal Epithelia In mice, the cerebellar primordium arises between E8.5 and E9.5 entirely from within the metencephalon (Wassef and Joyner 1997; Zervas et al. 2004).

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The boundary between Otx2 and Gbx2 expression domains initially demarcates the border between mes- and metencephalon and the location of the isthmic organizer, a tissue patterning structure that promotes interactions between cerebellar patterning genes (reviewed in Zervas et al. 2005). Several studies have examined putative allocation events during this period, which generate the Purkinje cell population: the general conclusion is that the entire Purkinje cell population in the adult arises from 100 to 150 precursors, likely specified at around E7–E8 (Baader et al. 1996; Mathis et al. 1997; Hawkes et al. 1998; Watson et al. 2005), although there is no evidence that these are restricted to a particular Purkinje cell subset. The early stages of cerebellar development are reviewed in detail in ▶ Chap. 6, “Specification of Granule Cells and Purkinje Cells.” This chapter will only consider mechanisms pertinent to the origin of stripe patterning (for other reviews, see Hawkes and Gravel 1991; Hawkes and Eisenman 1997; Herrup and Kuemerle 1997; Oberdick et al. 1998; Armstrong and Hawkes 2000; Larouche and Hawkes 2006; Sillitoe and Joyner 2007). The cerebellum houses two distinct germinal matrices – the dorsal rhombic lip and the ventral ventricular zone (VZ) of the fourth ventricle. Genetic fate mapping studies show that the rhombic lip gives rise to glutamatergic projection neurons of the cerebellar nuclei, cerebellar granule cells, and unipolar brush cells (Wingate 2001; Machold and Fishell 2005; Wang et al. 2005; Englund et al. 2006). The VZ gives rise to GABAergic components of the cerebellum including all GABAergic interneurons, and all Purkinje cells: all cerebellar GABAergic neurons derive from progenitors expressing Ptf1a, which is required for their specification (Hoshino et al. 2005; Pascual et al. 2007). However, the VZ is not homogenous but divided by gene expression into numerous overlapping expression domains (e.g., Chizhikov et al. 2006; Zordan et al. 2008). This issue is discussed in ▶ Chap. 15, “Genes and Cell Type Specification in Cerebellar Development.”

Purkinje Cell Birth Date, Phenotype, and Location Purkinje cells undergo terminal mitosis in the VZ between E10 and E13 in the mouse (Miale and Sidman 1961; Hashimoto and Mikoshiba 2003). Birthdating studies, using incorporation of either adenovirus (Hashimoto and Mikoshiba 2003) or bromodeoxyuridine (e.g., Feirabend et al. 1985; Karam et al. 2000; Larouche and Hawkes 2006), reveal a direct correlation between the birthdate of a Purkinje cell and its final mediolateral location, suggesting that Purkinje cells acquire positional information at or shortly after their terminal differentiation in the VZ. It is not known whether positional information and phenotype are specified at the same time. Postmitotic Purkinje cells migrate dorsally out of the VZ, presumably along radial glia processes (Morales and Hatten 2006), and stack in the cerebellar anlage with the earliest-born Purkinje cells located most dorsally.

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From Ventricular Zone to Clusters After migration from the VZ the Purkinje cells undergo a complex and poorly understood reorganization (E14–E18), possibly involving cell-signaling molecules including cadherin (Redies et al. 2010) and Eph-ephrin (Karam et al. 2000), to yield a stereotyped array of early clusters with a range of molecular phenotypes. The Purkinje cell migration pathways are carefully described in Miyata et al. (2010) (see also ▶ Chap. 9, “Purkinje Cell Migration and Differentiation”). During this same period, Purkinje cell clusters begin to express a variety of early markers that reveal both rostrocaudal and mediolateral compartments (e.g., calbindin – Wassef et al. 1985; cyclic GMP-dependant protein kinase – Wassef and Sotelo 1984; HSP25 – Armstrong et al. 2001; neurogranin – Larouche et al. 2006; cadherins – reviewed in Redies et al. 2010; homeobox genes, including En1, En2, Pax2, and Wnt17b – Bally-Cuif et al. 1992; Millen et al. 1995; L7/pcp2-LacZ – Smeyne et al. 1991; Oberdick et al. 1993; Ozol et al. 1999; OMP-LacZ – Nunzi et al. 1999; inositol 1,4,5-trisphosphate (IP3) receptor (IP3R)nls-LacZ – Furutama et al. 2010, etc.). Detailed comparisons of various early markers are not comprehensive but such data as are available suggest that all fit into a common schema.

Purkinje Cell Subtype Specification When is Purkinje cell phenotype specified? In order to answer this, many attempts have been made to alter Purkinje cell phenotype, which have almost always been ineffective. First, surgical interventions in the neonate have no effect on the expression of compartmentation markers (e.g., zebrin I – Leclerc et al. 1988; L7/pcp2-LacZ – Oberdick et al. 1993; HSP25 – Armstrong et al. 2001). Secondly, in cerebellar explants taken as early as E13 and placed either in slice culture (Oberdick et al. 1993; Seil et al. 1995; Rivkin and Herrup 2003; Furutama et al. 2010) or transplanted (Wassef et al. 1990), Purkinje cell subtypes apparently develop normally. Finally, ectopic Purkinje cells in various mouse mutants develop their normal adult phenotypes (e.g., reeler – Edwards et al. 1994; disabled – Gallagher et al. 1998; weaver – Armstrong and Hawkes 2001). These data suggest that cell autonomous mechanisms early in cerebellar development direct the specification of Purkinje cell phenotypes towards distinct subtypes. The only experimental manipulation that is known to alter Purkinje cell subtype is deletion of Early B-cell Factor 2 (Ebf2: Croci et al. 2006; Chung et al. 2008b). In the adult cerebellum, Ebf2 expression is restricted to the zebrin II- Purkinje cell subset. When Ebf2 is deleted, a complex cerebellar phenotype results, but in particular a prominent subset of zebrin II- Purkinje cells express zebrin II+ markers in addition to the normal zebrin II- ones (Croci et al. 2006; Chung et al. 2008b). This suggests that EBF2 is a repressor of the zebrin II+ phenotype. The role of Ebf2 is discussed in detail in ▶ Chap. 2, “Proneural Genes and Cerebellar Neurogenesis in the Ventricular Zone and Upper Rhombic Lip.”

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From Embryonic Clusters to Adult Stripes Starting at around E18, the embryonic clusters begin to disperse. This occurs at the same time that cerebellar lobules begin to form (Sudarov and Joyner 2007). The two processes are coupled – if Purkinje cell dispersal is blocked then lobulation is prevented and the cerebellum is lissiform – but the mechanistic relationship is unknown. In contrast to the relationship between cluster dispersal and lobules, a recent genetic study demonstrated that Purkinje cell stripe patterning and foliation can be uncoupled and En1/2 controls each process independently (Sillitoe et al. 2008b). Whether cluster dispersal is the passive concomitant of granular layer maturation and lobule formation or requires active Purkinje cell migration is not known. Whatever the case, because cluster dispersal occurs primarily in the rostrocaudal plane – the rostrocaudal length of the mouse vermis increases 25-fold from E17 to P30 while the width of the vermis increases only 1.5x during the same time period (Gallagher et al. 1998) – the clusters elongate into long parasagittal stripes. The transformation of embryonic Purkinje cell clusters into mature stripes is mediated by Reelin signaling (Tissir and Goffinet 2003). The external granular layer (EGL) secretes Reelin starting around E17 (D’Arcangelo et al. 1997; Jensen et al. 2002). Reelin binds two receptors on Purkinje cells – the Apolipoprotein E receptor 2 (Apoer2) and the Very Low Density Lipoprotein Receptor (VLDLR: Trommsdorff et al. 1999). Binding induces receptor clustering (Strasser et al. 2004) and activates an intracellular protein kinase cascade leading to tyrosine phosphorylation of the docking protein Disabled-1 (mdab-1: Howell et al. 1997; Goldowitz et al. 1997; Sheldon et al. 1997). Downstream of Disabled-1 are interactions with Src and Fyn cytoplasmic tyrosine kinases, and with phosphatidylinositol 3-kinase (Bock and Herz 2003; Kuo et al. 2005). The cyclin-dependant kinase (cdk) 5 signaling pathway has also been implicated in Reelin signaling as Purkinje cells in cdk5 pathway-mutants phenocopy reeler (e.g., Ohshima and Mikoshiba 2002). The end result is thought to be a drop in Purkinje cell-cell adhesion, thereby allowing the early clusters to disperse. Accordingly, mutations in the Reelin pathway affect all Purkinje cells and result in the complete failure of cluster dispersal and global Purkinje cell ectopia. However, deletion of either of the Reelin receptors Apoer2 and Vldlr results in selective, specific Purkinje cell ectopias (Larouche et al. 2008): in Apoer2/ mice, ectopic Purkinje cells are largely restricted to the zebrin II- population of the anterior vermis; in contrast, Vldlr/ mice have a much larger population of ectopic Purkinje cells that includes members from both zebrin II+/ phenotypes and HSP25 immunoreactivity reveals that a large portion of ectopic zebrin II+ cells is destined to become stripes in lobules VI–VII. Finally, a small, very specific population of ectopic zebrin II- Purkinje cells is observed in animals heterozygous for both receptors (Apoer2+/: Vldlr+/: no ectopia is present in mice heterozygous for either receptor alone). Despite the known importance of the Reelin pathway in regulating Purkinje cell dispersal, other genetic cues are also likely required. For example, the HSP25+/zebrin II+ Purkinje cell subset in the CZ is selectively ectopic in weaver mutants (Armstrong and Hawkes 2001).

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This model suggests a direct genealogical relationship between embryonic clusters and adult stripes. This is not straightforward to establish because the parasagittal pattern of early antigens tends to disappear perinatally, either because they are downregulated (e.g., neurogranin – Larouche et al. 2006) or because they become expressed uniformly by all Purkinje cells (e.g., calbindin – Wassef et al. 1985), while most adult stripe antigens are not expressed in the mature pattern of stripes until P15 (e.g., zebrin II – Lannoo et al. 1991; HSP25 – Armstrong et al. 2001). While the basic cerebellar architecture seems to be laid down in the embryo, the maturation of stripe phenotypes is not complete until P15 or so. For example, zebrin II is first expressed at around P6, but by P10–P12 all Purkinje cells express zebrin II. From P12 to P15 zebrin II is downregulated in the zebrin II- population to reveal the mature stripe array (Brochu et al. 1990; Lannoo et al. 1991; Rivkin and Herrup 2003). The molecular mechanism that mediates zebrin II downregulation is not known. However, recent studies have identified markers that bridge between clusters to stripes (e.g., Larouche et al. 2006; Marzban et al. 2007; Sillitoe et al. 2009). The current hypothesis is that embryonic clusters are the precursors of the adult stripes. While the hypothesis implies a direct relationship, experimental evidence indicates that it is not at all simple. On the one hand, current maps suggest about 20 different clusters but ten times as many stripes in the adult. Where does the additional complexity come from? While the apparent lack of complexity could merely be a reflection of an underdeveloped toolkit, the internal consistency of the different cluster antigens does not support this view: all known embryonic markers conform to a common schema with 10 clusters on each side of the cerebellum. Therefore, there may be secondary patterning stages, perhaps associated with the transformation of clusters into stripes, which takes the embryonic broad-stroke pattern and elaborates it into a more complex adult form. On the other hand, there is evidence that some stripes in the adult result from the coalescence of multiple clusters (e.g., the P1- stripe in the AZ is the fusion of three distinct clusters in the embryo – Ji and Hawkes 1994; Marzban et al. 2007). Finally, genetic fate mapping using an L7/pcp2-CreER allele supports the hypothesis that at least some embryonic clusters contribute Purkinje cells to multiple stripes in the adult (Sillitoe et al. 2009).

Afferent Topography It is generally believed that the Purkinje cell map serves as a scaffold around which other cerebellar structures are organized – both afferent projections (climbing fibers and mossy fibers: reviewed in Sotelo 2004) and interneurons including granule cells, Golgi cells, and unipolar brush cells (e.g., Chung et al. 2009; reviewed in Apps and Hawkes 2009) are thought to use Purkinje cell cues to establish their own topography.

Climbing Fibers In the adult, climbing fibers project from neurons in the contralateral inferior olivary complex and terminate on Purkinje cell dendrites, with each Purkinje cell

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receiving input from a single climbing fiber. Each subnucleus in the inferior olive projects to a limited number of Purkinje cell stripes (e.g., Voogd and Ruigrok 2004; Sugihara and Quy 2007; Apps and Hawkes 2009). The cerebellar projection neurons of the inferior olive are born in the caudal rhombic lip and migrate ventrally in the submarginal stream (Sotelo and Che´dotal 2005). Similar to their target Purkinje cells the fate, survival, differentiation, and migration of inferior olivary neurons is dependent on the function of Ptf1a (Yamada et al. 2007). Climbing fibers enter the cerebellar anlage at E15 and immediately terminate within specific Purkinje cell clusters (e.g., Che´dotal and Sotelo 1992; Paradies et al. 1996). During Purkinje cell cluster dispersal into stripes the climbing fibers are presumably carried along with them, thereby creating parasagittal terminal fields that align with the Purkinje cell stripes (Gravel et al. 1987; Apps and Hawkes 2009). In the neonatal cerebellum, each Purkinje cell receives input from several climbing fibers. This is converted to the adult mono-innervation by the elimination of all but one (reviewed in Cesa and Strata 2009). However, it appears that the sculpting of climbing fiber innervation does not contribute significantly to the refinement of cerebellar topography (Cre´pel 1982; Sotelo et al. 1984). Sotelo and colleagues have argued that matching gene expression domains between the cerebellum and inferior olive contain cues that guide the formation of a precise topographical projection map (Wassef et al. 1992; Che´dotal et al. 1997; Sotelo and Che´dotal 2005). In support of this model, Nishida et al. (2002) demonstrated that overexpression of Ephrin-A2 by using retroviral vectors disrupts the general topography of the olivocerebellar projection. Moreover, inferior olivary axons expressing high Eph receptor activity are prevented from entering into domains with ectopic Ephrin-A2 ligand expression (Nishida et al. 2002). Although the parasagittal band topography of climbing fibers was never examined, these experiments identify the Eph/Ephrin signaling pathway as likely to provide positional information during afferent/target matching.

Mossy Fibers The other major afferents of the cerebellum are mossy fibers, which arise from multiple sources and terminate in synaptic glomeruli on the dendrites of granule cells. Mossy fibers are also restricted by transverse zone and parasagittal stripe boundaries (e.g., Gravel and Hawkes 1990; Ji and Hawkes 1994; Armstrong et al. 2009; Sillitoe et al. 2010; Ruigrok 2010). In some cases, mossy fiber terminal fields align with specific subsets of stripes (e.g., Armstrong et al. 2009), and in others they split Purkinje cell stripes into smaller units (e.g., cuneocerebellar/ spinocerebellar terminal fields in the P1- stripes of the AZ: Ji and Hawkes 1994). The major features of the development of mossy fiber topography are similar to that for climbing fibers. The earliest mossy fibers enter the cerebellar anlage by around E12 (rat – Ashwell and Zhang 1992, 1998). Mossy fiber topography is established before most granule cells are formed (Arsenio Nunes and Sotelo 1985), and is accompanied by direct contacts between mossy fiber growth cones and

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Purkinje cells in embryonic and early postnatal clusters (Mason and Gregory 1984; Grishkat and Eisenman 1995). This model is consistent with observations from mutant animals with agranular cerebella, in which the spinocerebellar mossy fiber topography is organized into bands despite the absence of a normal mossy fiber-granule cell-Purkinje cell pathway (e.g., Arsenio Nunes and Sotelo 1985; Arsenio Nunes et al. 1988; Eisenman and Arlinghaus 1991) and with the data from neonatal lesion studies demonstrating that there does not seem to be a significant role for competition between mossy fiber sources in sculpting terminal fields (Ji and Hawkes 1995). The molecular basis of mossy fiber terminal field restriction is not well understood but deletion of either the retinoic acid receptor-related orphan receptor alpha (RORalpha: Arsenio Nunes et al. 1988) or En1/2 (Sillitoe et al. 2010), or overexpression of En2 in Purkinje cells (Baader et al. 1999) results in mossy fiber targeting defects. As for climbing fibers, mossy fiber terminals are presumed to disperse along with the Purkinje cells as embryonic clusters transform into stripes. Then postnatally, as granule cells are born in the external granular layer and descend past the Purkinje cells to the granular layer, the mossy fiber terminals displace from the Purkinje cells and synapse with differentiated granule cells. As a result the mossy fiber terminal fields become aligned with the overlying Purkinje cell stripes.

Interneurons Several cerebellar inhibitory interneurons show evidence of restriction by the Purkinje cell scaffold. First, Golgi cell dendrites are restricted by Purkinje cell stripe boundaries (Sillitoe et al. 2008a). Secondly, subsets of unipolar brush cells are associated with particular adult stripes (Chung et al. 2009). Models have been proposed by which both patterns of restriction involve mechanisms similar to those that organize mossy fiber afferent growth cones. Both Golgi cells and unipolar brush cells are thought to intermingle with Purkinje cells at the embryonic cluster stage. Hence, as the Purkinje cell clusters disperse, the interneurons become restricted to a particular stripe. Next, and as the granule cells form, the Golgi cells displace from the Purkinje cells to the neighboring granule cell axons, and the unipolar brush cells displace to the underlying granular layer where they are contacted by mossy fibers and synapse with granule cells and other unipolar brush cells (Mugnaini et al. 2010). Granule cells are born in the external granular layer (EGL), a germinal epithelium that forms from the rhombic lip and spreads to cover the cerebellar surface. Postmitotic granule cells migrate into the cerebellar anlage, following Bergmann glial guides, for 20–30 days, to create the adult granular layer (reviewed in Sillitoe and Joyner 2007). The EGL of the developing cerebellum and the granular layer of the adult cerebellum are subdivided by transverse boundaries revealed by lineage tracing, gene expression patterns, or through the consequences of genetic mutations. Several patterns of granular layer and/or EGL gene expression reveal transverse expression boundaries (e.g., reviewed in Hawkes et al. 1999), one aligned with

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the AZ/CZ boundary (lobule V–VI) and another at the PZ/NZ boundary (in lobule IX: reviewed in Ozol and Hawkes 1997). In addition, mRNA analysis reveals a complex map of Fgf receptor and ligand expression in the EGL and granular layer (Yaguchi et al. 2009). These zonal relationships may reflect either epigenetic interactions with Purkinje cells or distinct cell autonomous effects. It is likely that both occur. From the spatial distribution of genotypes in embryonic stem cell chimeras it was concluded that the cerebellar granular layer derives from two distinct precursor pools on either side of a lineage boundary within the rhombic lip (Hawkes et al. 1999). This is consistent with previous chimera studies, which also suggested that granule cells across the AZ/CZ and PZ/NZ boundaries have separate developmental origins (e.g., Goldowitz 1989). Additional evidence for a multiple origin of the EGL comes from studies of scrambler (Goldowitz et al. 1997) and disabled (Gallagher et al. 1998), mutants in which there is an incomplete fusion of the anterior and posterior granular layers in lobule VI leaving distinct, overlapping anterior and posterior leaflets. Finally, by using a Math1-CreER allele, Machold and Fishell (2005) demonstrated by genetic fate mapping that granule cell progenitors are destined to populate specific anterior–posterior zones. For example, lineages marked at E12.5 selectively populate the AZ whereas those marked at E15.5 populate all but the NZ. Together, these data suggest that the allocation of cells to specific EGL compartments may be dependent on spatial and temporal regulation of cellular movements and gene expression. It is difficult to imagine that the striped expression patterns in the granular layer are generated by the differential migration of EGL lineages. For example, neuronal nitric oxide synthase (nNOS – or its surrogate nicotinamide adenine dinucleotide phosphate (NADPH)-diaphorase) is expressed in the adult granular layer in stripes that align with Purkinje cell stripes (Hawkes and Turner 1994). NADPHd/nNOS activity is first detected at P3. During the first postnatal week of development the granular layer expresses nNOS uniformly (Schilling et al. 1994). Subsequently, clusters of granule cells begin to suppress their expression of nNOS, and from this, a new heterogeneous pattern of nNOS expression emerges that persists into adulthood (Yan et al. 1993; Schilling et al. 1994; Hawkes and Turner 1994). In such cases, it seems more plausible that differential gene expression is induced by the local Purkinje cell environment or by the input from mossy fiber stripes (Schilling et al. 1994). Conclusions

Every facet of cerebellar structure and function is built around the zone-andstripe architecture. While the pattern formation process is complex, a simple theme emerges – Purkinje cells are the scaffold around which other structures organize, and their disruption leads to widespread abnormalities in cerebellar topography. Acknowledgments We thank Dr. Carol Armstrong for her advice. This study was supported by a grant to RH from the Canadian Institutes for Health Research and by new investigator start-up funds to RVS from Albert Einstein College of Medicine of Yeshiva University.

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4

Roof Plate in Cerebellar Neurogenesis Victor V. Chizhikov

Abstract

The roof plate is a distinct group of cells located at the dorsal midline of the developing central nervous system extending along its entire anterior–posterior axis. In the developing hindbrain, the roof plate comprises a simple epithelial layer covering the dorsal opening of the 4th ventricle. As development proceeds, the 4th ventricle roof plate differentiates into choroid plexus epithelium, which produces cerebrospinal fluid and serves as a blood-cerebrospinal fluid barrier. A growing amount of evidence indicates that both the 4th ventricle roof plate and its later derivative, the hindbrain choroid plexus, produce various secreted molecules, which critically regulate development of the adjacent cerebellum. Bone morphogenetic proteins secreted from the roof plate are crucial to the induction of the cerebellar rhombic lip. Signals from the early roof plate and later secretion of Sonic hedgehog from the choroid plexus promote proliferation of progenitors in the cerebellar ventricular zone. This chapter discusses recent studies that established the roles of the 4th ventricle roof plate and the hindbrain choroid plexus in cerebellar neurogenesis and the molecular mechanisms of their action.

Introduction During development, neurogenesis in many regions of the central nervous system is regulated by secreted signals produced by specialized groups of cells called signaling centers (Lee and Jessell 1999; Kiecker and Lumsden 2005; Sillitoe and Joyner 2007; Dessaud et al. 2008). One such signaling center regulating cerebellar development is the isthmic organizer (IsO), located at the mid-hindbrain boundary.

V.V. Chizhikov Center for Integrative Brain Research, Seattle Children’s Hospital Research Institute, 1900, 9th Avenue, Mailstop C9S-10, Seattle, WA 98101, USA e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_4, # Springer Science+Business Media Dordrecht 2013

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As discussed earlier in this book (▶ Chap. 1, “Specification of the Cerebellar Territory”), numerous studies have demonstrated that fibroblast growth factor (FGF) signals secreted from the IsO are important for proper establishment of the cerebellar territory during early developmental stages. More recently, another embryonic structure, the 4th ventricle roof plate and its later derivative, the hindbrain choroid plexus, have emerged as additional signaling centers regulating multiple aspects of cerebellar neurogenesis. The roof plate is a transient embryonic signaling center located at the dorsal midline of the developing central nervous system along its entire anterior-posterior axis (Lee and Jessell 1999; Monuki et al. 2001; Chizhikov and Millen 2005; Wilson and Maden 2005; Cheng et al. 2006; He´bert and Fishell 2008). At most levels of the neural tube, along the anterior–posterior axis, it appears as a dorsal stripe of cells. At the level of hindbrain, however, the roof plate broadens into a single-layer sheet of epithelial cells covering the fourth ventricle (Fig. 4.1a, b). During development, the roof plate epithelium (RPe) derives from the most lateral edges of the neural plate, which occupy the dorsal midline, following transformation of the neural plate into neural tube (Landsberg et al. 2005; Hunter and Dymecki 2007). As the hindbrain flexes (at embryonic day (e) 9.0 in the mouse), the RPe flares out laterally to cover the expansive 4th ventricle (Fig. 4.1a, b). Around e12.5 in the mouse, the RPe differentiates into choroid plexus epithelium (CPe) (Awatramani et al. 2003; Currle et al. 2005; Chizhikov et al. 2006; Hunter and Dymecki 2007), a cuboidal epithelium that produces cerebrospinal fluid and serves as a blood-cerebrospinal fluid barrier (Redzic et al. 2005; Emerich et al. 2005) (Fig. 4.1d). Although the majority of RPe and CPe cells are post-mitotic, both of these structures receive cellular contributions from adjacent tissue. Early during development a restricted dorsal domain of the IsO contributes cells to the anterior part of the 4th ventricle roof plate (Alexandre and Wassef 2003). In addition, cells from the adjacent cerebellar rhombic lip (RL) contribute to both the RPe and CPe, allowing growth of these structures (Hunter and Dymecki 2007; Rose et al. 2009). During development, all cerebellar neurons arise from one of two germinal zones: the cerebellar rhombic lip (RL) and the cerebellar ventricular zone (VZ) (reviewed in Wingate 2005; Hevner et al. 2006; Millen and Gleeson 2008). The cerebellar RL, defined by expression of the basic helix-loop-helix transcription factor Atoh1 (Atonal homolog 1, also known as Math1), is located adjacent to the RPe/CPe (Ben-Arie et al. 1996; Helms and Johnson 1998; Wang et al. 2005) (Fig. 4.1c, d). Recent fate-mapping studies together with previous transplantation experiments have shown that the RL gives rise to all glutamatergic cerebellar neurons, including glutamatergic neurons of the deep cerebellar nuclei (DCN), granule neurons, and unipolar brush cells (reviewed in Wingate 2005; Hevner et al. 2006). During development, these neurons arise in distinct, although partially overlapping, birth cohorts. The first cerebellar neurons generated in the RL are glutamatergic neurons of the DCN, which in mice exit the RL between e10 and e12. They are followed at e13.5 by granule neuron precursors, which continue to be produced until the time of birth, the time of RL regression. Unipolar brush cells begin to exit the RL around e15.5 (Fig. 4.1c, d) (Machold and Fishell 2005;

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Fig. 4.1 Overview of embryonic rhombomere 1 neuroanatomy. (a) Dorsal view of e9.5–12 developing mouse embryo. Top is anterior, bottom is posterior. Midbrain (mid), hindbrain (hind), isthmic organize (IsO), which develops at the mid-hindbrain boundary, rhombomere 1 (rh1), which is the most anterior part of the hindbrain are labeled. 4th ventricle roof plate epithelium (4v RPe) is blue. (b) Sagittal section of the developing mouse neural tube taken at the level of the dashed line in panel “a.” Top is anterior, bottom is posterior, left is ventral, right is dorsal. Midbrain (mid), rhombomere 1 (rh1), RPe, and 4th ventricle (4v) are labeled. (c) A diagram of the developing cerebellar anlage at e9.5–12, corresponding to the region boxed in panel “b.” RPe (blue), cerebellar RL (pink), and cerebellar VZ (green) are shown. At this early developmental stage, RL gives rise to glutamatergic neurons of the DCN (shown as pink circles). These cells migrate tangentially along the dorsal surface of the cerebellar anlage and accumulate at the nuclear transitory zone (NTZ). The early cerebellar VZ gives rise to GABAergic neurons of the DCN and Purkinje cells (PC) (both shown as green circles). Both GABAergic neurons of DCN and Purkinje cells migrate radially from the cerebellar VZ. (d) A diagram of the developing cerebellar anlage at e14.5–18.5. During these stages, RPe transforms into the CPe (blue) and RL gives rise to granule cell precursors (GC) and unipolar brush cells (UBC) (both shown as pink circles). Granule cell precursors migrate tangentially along the dorsal surface of the cerebellar anlage to form the external granule cell layer (EGL). UBCs migrate directly into the cerebellar anlage. The cerebellar VZ gives rise to precursors of GABAergic interneurons (int) (green circles), which migrate radially from the cerebellar VZ

Wang et al. 2005; Fink et al. 2006; Englund et al. 2006). The cerebellar VZ, defined by the basic helix-loop-helix transcription factor Ptf1a (pancreas specific transcription factor 1a), is located ventral to the RL (Hoshino et al. 2005; Glasgow et al. 2005; Hoshino 2006; Pascual et al. 2007) (Fig. 4.1c, d). It gives rise to all GABAergic cerebellar neurons, including GABAergic neurons of the DCN, Purkinje cells,

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granule layer interneurons (such as Golgi cells) and molecular layer interneurons (basket and stellate cells). In the cerebellar VZ, the first born neurons are small GABAergic DCN neurons. In mice, these cells become postmitotic between e10 and e12. Purkinje cells are born from e11 to e13. The last cells arising from the cerebellar VZ are GABAergic interneuron progenitors (which later give rise to Golgi, basket, and stellate cells) (Altman and Bayer 1997; Hoshino et al. 2005; Pascual et al. 2007; Carletti and Rossi 2008; Leto et al. 2008; Schilling et al. 2008) (Fig. 4.1c, d). Once formed, the 4th ventricle roof plate and the choroid plexus produce secreted molecules that critically regulate development of both the adjacent RL and the more distantly located VZ. This chapter discusses the roles of RPe/CPederived signals in cerebellar neurogenesis and the mechanisms of their action.

Roof Plate and Choroid Plexus in Rhombic Lip Development The cerebellar RL arises adjacent to the RPe/CPe, and the RL and RPe/CPe critically depend on each other during development. Bone morphogenetic proteins (Bmp) signals from the RPe/CPe are required for early RL induction, while the RPe/CPe receive cellular contributions from the RL helping their growth.

Secreted Signals from the 4th Ventricle Roof Plate and the Hindbrain Choroid Plexus as Regulators of Rhombic Lip Development Several groups of experiments have shown that signals secreted from the 4th ventricle roof plate are required for the early induction of the cerebellar RL. Genetic ablation of the 4th ventricle roof plate by diphtheria toxin expression in early (e10–12.5) mouse embryos resulted in complete loss of the cerebellar RL. In addition, differentiating glutamatergic DCN neurons, which normally originate from the RL, were absent based on lack of expression of their marker Lhx2 (LIM homeobox protein 2) (Chizhikov et al. 2006). At the same time, coculturing naı¨ve rhombomere 1 (rh1) neural tissue with exogenous 4th ventricle roof plate resulted in the appearance of ectopic Atoh1+ RL cells, as well as Lhx2+ cells, suggesting that roof plate signaling is not only necessary but also sufficient for induction of the RL and production of at least one of its derivatives – Lhx2+ glutamatergic neurons of the DCN (Chizhikov et al. 2006). In vitro tissue culture experiments identified members of the bone morphogenetic proteins (Bmp) family expressed in the 4th ventricle roof plate, including Bmp6, Bmp7, and Gdf7 (growth differentiation factor 7), as major components of roof plate signaling. For example, when added to culture media, Bmps induced numerous Atoh1+ cells in naı¨ve rh1 neural tissue, mimicking coculturing naı¨ve neural tissue with 4th ventricle roof plate. Moreover, this Bmp-treated neural tissue formed mature granule neurons after transplantation into the early postnatal

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Fig. 4.2 A diagram showing a sagittal section of the developing mouse cerebellar anlage. Signals secreted from the RPe/CPe control development of the cerebellar RL and cerebellar VZ. RPe/CPe (blue), cerebellar RL (pink), and cerebellar VZ (green) are shown. Fourth ventricle (4v). Arrows show signals secreted from the RPe/CPe. At early developmental stages (e9.5–12.5 in the mouse), Bmp signals from the RPe/CPe induce Atoh1-positive cerebellar RL adjacent to the RPe/CPe. At the same early stages, RPe/CPe signaling (which likely involves Bmp and Wnt molecules) is also essential for normal proliferation of progenitors in the cerebellar VZ. Beginning from e14.5 in the mouse, CPe produces Shh molecules, which promote proliferation of progenitors in the cerebellar VZ at this and later developmental stages. Shh is secreted into the cereberospinal fluid through which it is likely delivered to the cerebellar VZ progenitors

cerebellum, suggesting that Bmps not only induce RL progenitors but are also sufficient to initiate generation of granule neurons, the most numerous RL-derived cells (Alder et al. 1999). Finally, electroporation of activated Bmp receptors into chick developing rh1 neural tissue induced ectopic Atoh1+ cells (Machold et al. 2007), further confirming ability of Bmps to initiate the RL developmental program (Fig. 4.2). Although the studies described above have clearly demonstrated the role of the 4th ventricle roof plate in RL induction and established Bmps as major components of roof plate signaling, it is important to note that all these studies were performed using very early mouse or chick embryos. It has recently been demonstrated that rather than representing a single population, Atoh1+ cells within the cerebellar RL are a highly dynamic population, with different progenitors expressing Atoh1 at early versus late developmental stages. Indeed, inducible genetic fate mapping in mice reveled that once progenitors in the RL induce Atoh1 expression, they rapidly transit out from the RL leaving no progeny behind. Therefore, Atoh1 expression is induced de novo in naı¨ve progenitors throughout cerebellar neurogenesis (Machold and Fishell 2005). Since Bmps continue to be expressed in the CPe throughout late embryonic development, it is likely that similar to roof plate in early embryos, CPe continues induction of Atoh1+ RL cells at later developmental stages. This hypothesis, however, remains to be experimentally tested. Besides its likely role in RL induction at late embryonic stages, recent in vitro culture experiments identified an additional role for choroid plexus in attenuating

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differentiation of RL progenitors. Coculturing with choroid plexus significantly decreased neuronal differentiation of RL cells. Moreover, whereas addition of purified Bmp7 to culture mimicked this phenotype, using a blocking antibody against Bmp7 abolished the inhibitory effect of the choroid plexus, demonstrating casual involvement of Bmps (Krizhanovsky and Ben-Arie 2006). This attenuating of neuronal differentiation by CPe-derived Bmp signals may be important to maintain a pool of undifferentiated progenitors in the RL during development. In addition to members of the Bmp family, other factors expressed in the 4th ventricle roof plate and choroid plexus have been suggested as regulators of RL development. For example, both the 4th ventricle roof plate and the hindbrain choroid plexus synthesize retinoic acid, while the cerebellar RL is a site of active retinoic acid usage (Yamamoto et al. 1996; Wilson et al. 2007). The role of retinoic acid or other RPe/CPe-derived molecules in RL development has not been experimentally confirmed yet.

Segregation of the RPe/CPe Lineage and Neuronal RL Derivatives Although the majority of RL-derived cells adopt neuronal fates, the RL also contributes to the 4th ventricle roof plate and the hindbrain choroid plexus, allowing growth of these structures during development (Landsberg et al. 2005; Hunter and Dymecki 2007; Rose et al. 2009) (Fig. 4.3a). Since appropriate numbers of neuronal and RPe/CPe cells are generated in each normal embryo, there must be precise molecular mechanisms segregating RPe/CPe from neuronal RL lineages. Loss of function experiments in mouse embryos demonstrated that the LIM homeodomain transcription factor 1a (Lmx1a) plays a unique role in roof plate maintenance, segregating the RPe/CPe lineage from neuronal RL derivatives (Chizhikov et al. 2010). In Lmx1a-/- (dreher) embryos, the 4th ventricle roof plate is normally induced, but as development proceeds, cells belonging to the RPe/CPe lineage migrate into the adjacent cerebellum and adopt fates of neuronal RL derivatives, including glutamatergic DCN neurons, granule cells, and unipolar brush cells (Chizhikov et al. 2010). This cell fate switch results in a severely reduced choroid plexus observed in dreher mice (Millonig et al. 2000; Manzanares et al. 2000) (Fig. 4.3b). In contrast, Atoh1 is required for generation of all classes of neurons originating from the RL. In Atoh1-/- mice, glutamatergic DCN neurons are not present and very few granule cells and unipolar brush cells are produced (Ben-Arie et al. 1997; Jensen et al. 2004; Wang et al. 2005; Machold and Fishell 2005; Englund et al. 2006). Interestingly, in these mice RL cells contribute excessively to the choroid plexus, suggesting that Atoh1 is not only an important activator of the general RL program but that it is also important for segregation of neuronal RL lineages from RPe/CPe cells (Rose et al. 2009) (Fig. 4.3c). Taken together, these data suggest that Lmx1a and Atoh1 antagonize each other’s expression or activity to ensure proper generation of the 4th ventricle roof plate/choroid plexus and cerebellar glutamatergic neurons.

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a

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NTZ EGL

DCN

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NTZ EGL

GC DCN

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GC UBC

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Fig. 4.3 Lmx1a and Atoh1 segregate the RPe/CPe lineage and neuronal RL derivatives. A diagram showing sagittal sections of the developing cerebellar anlage in wild-type (wt) (a), Lmx1a-/-(dreher) (b), and Atoh1-/- (c) mouse embryos. Cerebellar RL is pink. RL neuronal derivatives such as granule cells (GC), glutamatergic neurons of the DCN and UBCs are shown as pink circles. The nuclear transitory zone (NTZ) and external granule cell layer (EGL) are shown. CPe is blue. (a) In wild-type mice, cerebellar RL mostly produces cerebellar neurons, although it also contributes into the adjacent RPe/CPe. RL-derived CPe is shown in red. (b) In Lmx1a-/- (dreher) mutant mice, many cells of the RPe/CPe lineage abnormally adopt fates of glutamatergic neurons of the DCN, granule cells, and unipolar brush cells (UBC), and migrate into the cerebellar anlage. These RPe/CPe-derived neurons are shown as blue circles. This transformation results in a smaller 4th ventricle roof plate and hindbrain choroid plexus in dreher mice. In addition, the entire cerebellum is smaller in dreher mice compared to wild-type littermates. Since signals from the RPe/CPe are essential for both RL induction and proliferation in the cerebellar VZ, reduction of cerebellar size in dreher mice is likely secondary to the roof plate and choroid plexus abnormalities. (c) In Atoh1-/- embryos, glutamatergic neurons of the DCN are not produced and very few granule cells or UBCs (shown as pink circles) are detected in these mutants. Instead, in Atoh1-/- mice, RL cells contribute excessively to adjacent CPe

Roof Plate and Choroid Plexus in Development of the Cerebellar Ventricular Zone In contrast to the cerebellar RL, induction of the cerebellar VZ does not depend on roof plate signaling. Signals secreted from the roof plate and choroid plexus, however, regulate proliferation of progenitors in the cerebellar VZ during both early and late embryonic stages.

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Signals from the 4th Ventricle Roof Plate Regulate Proliferation in the Early Cerebellar Ventricular Zone Analysis of roof plate–ablated mouse embryos was instrumental in uncovering the role of the 4th ventricle roof plate in VZ development. In e12.5 roof plate–ablated embryos, the cerebellar VZ was normally induced based on the presence of an appropriate Ptf1a expression domain in rh1 VZ. The numbers of Ptf1a+ progenitors as well as numbers of their derivative Lhx1/5+ cells, which at this stage likely include differentiating GABAergic neurons of DCN and Purkinje cells (Morales and Hatten 2006; Zhao et al. 2007), were, however, significantly decreased in these roof plate–ablated embryos (Chizhikov et al. 2006). The decreased numbers of Ptf1a+ and Lhx1/5+ cells were associated with reduced proliferation of cells in the earlier cerebellar VZ, which was proposed to explain this phenotype (Chizhikov et al. 2006) (Fig. 4.2). Since roof plate–ablated embryos die shortly after e12.5, analysis of neuronal VZ derivatives at later developmental stages could not be performed. Nevertheless, although this analysis was limited to very early developmental stages, these roof plate–ablated embryos strongly suggested that signals secreted from the 4th ventricle roof plate are critical for normal progenitor proliferation in the cerebellar VZ. Analysis of Lmx1a-/- (dreher) and Lmx1a/Lmx1b double knockout (Lmx1a-/-; Lmx1bcko/-) mice further confirmed the role of the RPe/CPe in cerebellar VZ proliferation (Mishima et al. 2009). As mentioned earlier, inactivation of Lmx1a leads to reduction of the 4th ventricle roof plate and the hindbrain choroid plexus, which are both even smaller in Lmx1a-/-;Lmx1bcko/- mice. Reduction of the RPe/ CPe in Lmx1a-/- or Lmx1a-/-;Lmx1bcko/- embryos was associated with graded decreases in VZ proliferation at e12.5, resulting in a small and mispatterned cerebellum in adult Lmx1a-/- and Lmx1a-/-;Lmx1bcko/- mice (Mishima et al. 2009). Currently, the nature of roof plate signals regulating proliferation of VZ progenitors at early developmental stages is not completely understood. It is likely, however, that Bmp and Wnt (wingless-related MMTV integration site) signals are involved. Potential involvement of Bmp signals is supported by both gene expression and mouse transgenic studies. As discussed above, Bmps are expressed in both RPe and CPe, and strong pSmad staining, a readout of active Bmp signaling, was detected in the embryonic cerebellar VZ (Qin et al. 2006). Furthermore, transgenic mice expressing activated Bmp receptor 1a (Bmpr1a) under the control of the nestin promoter demonstrated overproliferation of neuroepithelium throughout the central nervous system, including the cerebellar VZ (Panchision et al. 2001). Several members of the Wnt family are also well-known mitogens, which regulate proliferation of neuronal progenitors in different areas of the central nervous system (Megason and McMahon 2002; Chenn and Walsh 2002; Chesnutt et al. 2004; Ille et al. 2007). In particular, Wnt1 and Wnt3a promote proliferation of neural progenitors by activating expression of positive cell cycle regulators, including cyclin D1 and D2 (Megason and McMahon 2002). In roof plate–ablated mouse embryos, expression of Wnt1 was lost and cyclin D2 was significantly downregulated throughout the cerebellar VZ, implicating a Wnt1-cyclin D2 molecular pathway as a potential mediator of this VZ phenotype (Chizhikov et al. 2006).

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It is important to note, however, that in contrast to Bmps, Wnt1 is not expressed in rh1 RPe/CPe. Instead it is expressed in the adjacent dorsal domain, which includes the cerebellar RL and the RL–roof plate junction (Landsberg et al. 2005; Chizhikov et al. 2006). Therefore, loss of the 4th ventricle roof plate likely affects Wnt1 expression nonautonomously. Notably, Bmps are potent activators of Wnt1 expression in several experimental systems (Timmer et al. 2002; Zechner et al. 2007; Caronia et al. 2010). Therefore, it is possible that in developing rh1, Bmp signals from the RPe/CPe regulate proliferation in the cerebellar VZ both directly and indirectly, via activation of Wnt1 expression in the adjacent dorsal tissue.

Shh Signals from the Hindbrain Choroid Plexus Regulate Proliferation of Progenitors in the Late Embryonic Cerebellar Ventricular Zone Recent loss and gain of function experiments in mouse embryos have identified Sonic hedgehog (Shh), which is secreted from the hindbrain choroid plexus, as an important and previously unappreciated regulator of VZ development (Huang et al. 2010). In the cerebellum, Shh activity is not detectable at early developmental stages, but this pathway becomes activated in the cerebellar VZ at e14.5 and remains active in this germinal zone throughout late embryonic development. Additionally, the widespread presence of primary cilia, essential sensors and transducers of Shh signals (Huangfu et al. 2003; Berbari et al. 2009; Goetz and Anderson 2010) was detected in VZ progenitors in late embryonic mouse embryos (Huang et al. 2010). To study the role of Shh in VZ development, mouse embryos with decreased or increased levels of Shh signaling were generated and analyzed. Nestincre;Smof/- embryos, in which Shh signaling is conditionally ablated in all neural cells throughout the developing central nervous system, had a remarkably thin cerebellar VZ at e16.5. This was associated with reduced proliferation of VZ progenitors and decreased levels of cell cycle promoting cyclin D1. In addition, a dramatic reduction of the GABAergic interneuronal population, which originates from the cerebellar VZ, was detected in these embryos. In contrast, embryos with ectopic Shh signaling in the central nervous system (Nestin-cre;SmoM2 embryos), revealed enhanced proliferation in the cerebellar VZ, and, importantly, increased numbers of GABAergic interneuron precursors were detected in the cerebella of these mice (Huang et al. 2010). Together, these experiments established that Shh signaling is critical for proper proliferation of progenitors in late embryonic cerebellar VZ. This progenitor pool, in turn, is essential for the generation of the appropriate number of GABAergic interneuronal precursors, which originate from the cerebellar VZ at late embryonic stages. Interestingly, other VZ-derived neurons, such as Purkinje cells and GABAergic DCN neurons were not significantly affected in embryos with abnormal levels of Shh signaling (Huang et al. 2010), suggesting that they are generated from the cerebellar VZ before its development becomes dependent on Shh signaling.

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Interestingly, although the cerebellar VZ is molecularly defined by Ptf1a expression (Hoshino et al. 2005; Glasgow et al. 2005; Wingate 2005; Pascual et al. 2007), Ptf1a+ cells in the cerebellar VZ are largely postmitotic, suggesting that they represent committed neuronal progenitors. In contrast, similar to other CNS regions (Anthony et al. 2004), radial glia represent an uncommitted stem cell population in the cerebellar VZ (Spassky et al. 2008; Sch€ uller et al. 2008; Huang et al. 2010). Analysis of Nestincre;Smof/-, Nestin-cre;SmoM2, and Ptf1a-cre;Smof/- embryos revealed that, in the cerebellar VZ, Shh signaling specifically regulates proliferation of radial glia without affecting Ptf1a+ progenitors (Yang et al. 2008; Huang et al. 2010). Gene expression studies revealed that before e16.5, the mouse cerebellum itself is not a source of Shh signals, while this gene is strongly expressed in the hindbrain CPe beginning from e13.5 (Huang et al. 2009). Specific inactivation of Shh expression in the CPe resulted in strong defects in the cerebellar VZ proliferation and GABAergic interneuron expansion. These defects were comparable to those observed in Nestincre;Smof/- embryos, in which Shh signaling was ablated throughout the cerebellar VZ neuroepithelium, suggesting the hindbrain choroid plexus as a likely source of Shh for cerebellar VZ progenitor proliferation (Huang et al. 2010). Interestingly, a significant level of Shh protein was detected in the cerebrospinal fluid, suggesting a transventricular path for Shh ligand delivery from the CPe to the cerebellar VZ (Huang et al. 2010). Taken together, these data strongly suggest that CPe-derived signals regulate VZ proliferation not only at early but also at late embryonic stages, and introduce Shh as a major component of this late CPe-derived signaling (Fig 4.2).

Conclusions and Future Directions During the last decade, the 4th ventricle roof plate and its later derivative, the hindbrain choroid plexus, have emerged as important signaling centers which nonautonomously regulate cerebellar development by producing various secreted molecules. This knowledge has been derived from a combination of genetic ablation experiments, mouse mutant analyses, gene expression studies, and in vitro explant experiments. Together, these studies have firmly established roles for RPe/ CPe-derived signals in several different steps of cerebellar development. In particular, RPe/CPe-derived signals have been shown to be critical for proper development of two embryonic cerebellar germinal zones: the cerebellar RL and the cerebellar VZ. Bmp signals from the RPe/CPe induce the RL progenitor population during early development and attenuate neuronal differentiation within the RL at later developmental stages, maintaining the RL progenitor pool. Shh, and likely other molecules, expressed in the roof plate and choroid plexus, are also important positive regulators of proliferation in the cerebellar VZ. Cerebellar neurogenesis, however, is a complex multistep process that is not limited to initial specification or proliferation of neuronal progenitors, but also includes differentiation and migration of different types of neurons, and finally, the interconnection of the cerebellar circuitry. The role of roof plate or choroid plexus has not yet been clearly demonstrated in these later aspects of cerebellar

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neurogenesis. Taking into account the large size of the 4th ventricle roof plate and the hindbrain choroid plexus, their location directly adjacent to the developing cerebellum, and the large number of molecules secreted by these tissues, it is likely that the role of RPe/CPe in cerebellar neurogenesis is not limited by regulating development of progenitors in the cerebellar RL or VZ. Indeed, in vitro experiments, in which embryonic rat cerebellar slices were cultured together with exogenous choroid plexus, suggested a possible role for CPe in promoting the differentiation of cerebellar neurons. In these cultures, neurites grew out in a radial fashion from cultured cerebellar slices. Only short neuritic outgrowths were detected when cerebella were cultured alone. When the cerebella were cocultured with choroid plexus, a large increase in the length of neurites was observed, suggesting that signals secreted from the choroid plexus positively regulate neuronal differentiation under these experimental conditions. This effect can be mimicked by retinoic acid, which is normally produced by the choroid plexus (Yamamoto et al. 1996). It is remain unclear, however, if CPe-derived retinoic acid has a comparable function in vivo and if so, which cerebellar neurons primarily respond to this signaling. Additional experiments are clearly required to investigate the role of RPe/CPe in regulation of neuronal differentiation, migration, and establishment of the cerebellar connectivity.

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Specification of Cerebellar and Precerebellar Neurons Mikio Hoshino, Yusuke Seto, and Mayumi Yamada

Abstract

The cerebellum is thought to participate in the regulation of movement and is comprised of various types of neurons in the cerebellar cortex and nuclei. Each type of neurons has morphologically, immunohistochemically, and electrophysiologically distinct characteristics. In addition, there are two precerebellar afferent systems, the mossy fiber (MF) system and the climbing fiber (CF) system. MF neurons are located in various nuclei throughout the brainstem and send their axons to cerebellar granule cells, whereas CF neurons reside exclusively in the inferior olivary nucleus (ION) and project to Purkinje cells. Recently developed genetic lineage-tracing methods as well as gene-transfer technologies have accelerated the studies on the molecular machinery to specify neuronal subtypes in the cerebellum and the precerebellar systems.

Specification of Cerebellar Neurons The cerebellum consists of three parts: cortex, white matter, and nucleus. The cerebellar cortex contains Purkinje, Golgi, Lugaro, stellate, basket, granule, and unipolar brush cells. The latter two cell types are glutamatergic excitatory neurons, while the others are all GABAergic inhibitory neurons. The cerebellar nucleus (CN) includes three types of neurons: large glutamatergic projection neurons (CN-Glu neurons), mid-sized GABAergic inhibitory projection neurons

M. Hoshino (*) • M. Yamada Department of Biochemistry and Cellular Biology, National Institute of Neuroscience, National Center of Neurology and Psychiatry, 4-1-1 Ogawa-Higashi, Kodaira, Tokyo, 187–8502, Japan e-mail: [email protected] Y. Seto Integrative Bioscience and Biomedical Engineering, Graduate School of Science and Engineering, Waseda University, 3-4-1 Okubo, Shinjuku-ku, Tokyo, 169–8555, Japan M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_5, # Springer Science+Business Media Dordrecht 2013

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(CN-GABA-ION neurons), and small GABAergic interneurons (CN-GABA interneurons). CN-GABA-ION neurons extend their axons to the inferior olivary nucleus (ION) (Carletti and Rossi 2008), while CN-Glu neurons send their axons to nuclei outside the cerebellum, including the red nucleus and the thalamus. It is believed that all types of cerebellar neurons are generated from the neuroepithelium of the alar plate of rhombomere 1 (r1) during development (Millet et al. 1996; Wingate and Hatten 1999; Chizhikov and Millen 2003; Zervas et al. 2004). The dorsal-most part of the r1 neuroepithelium, that is, the roof plate, does not produce neurons but cells of the choroid plexus (Chizhikov et al. 2006). Neuroepithelium that produces cerebellar neurons can be divided into two regions: the rhombic lip (RL) and the ventricular zone (VZ). These two regions can be morphologically discriminated by a notch located on the border. Although the history of studies on the cerebellum is very long (Ramo´n y Cajal 1911), the molecular machinery underlying cerebellar neuron development is still unclear. In 1997, Ben-Arie et al. reported that a basic-helix-loop-helix type (bHLH) transcription factor, Atoh1 (also called Math1), is expressed in the RL and involved in producing cerebellar granule cells (Ben-Arie et al. 1997). However, the development of the other types of neurons in the cerebellum remained elusive until three breakthrough papers were published in 2005. While generating certain transgenic lines, Hoshino et al. found a novel mutant mouse line, cerebelless, which lacked the entire cerebellar cortex. In this mutant, all types of GABAergic neurons are not produced in the cerebellum, which leads to the secondary loss of glutamatergic granule cells and eventually, the entire cerebellar cortex (Hoshino et al. 2005). The responsible gene was identified as pancreatic transcription factor 1a (Ptf1a), which was known to participate in pancreatic development and to encode a bHLH transcription factor. This gene is expressed in the neuroepithelium of the VZ but not of the RL and its expression is lost in the cerebelless mutants. Cre-loxP recombination-based lineage tracing analysis revealed that all types of cerebellar GABAergic neurons are derived from Ptf1aexpressing neuroepithelial cells in the VZ, but glutamatergic neurons, such as granule cells and CN-Glu neurons, are not. Loss of Ptf1a expression in cerebelless as well as Ptf1a-knock out mice resulted in inhibition of the production of GABAergic neurons in the cerebellar primordium. Furthermore, ectopic introduction of Ptf1a by means of in utero electroporation resulted in the abnormal production of neurons with GABAergic characteristics from the dorsal telencephalon that should only produce glutamatergic neurons under normal conditions. In addition, Pascual et al. reported that, in the Ptf1a-null mutants, the fate of neurons produced from the VZ is changed to that of granule cells (Pascual et al. 2007). These observations suggested that Ptf1a, expressed in the cerebellar VZ, determines GABAergic neuronal fate in the cerebellum. PTF1A was also identified as a causative gene for a human disease that exhibits permanent neonatal diabetes mellitus and cerebellar agenesis (Sellick et al. 2004). On the other hand, two other groups revealed a molecular fate map of the derivatives of Atoh1-expressing neuroepithelial cells in the cerebellar RL (Machold and Fishell 2005; Wang et al. 2005). They showed that not only granule cells but

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also, at least in part, some neurons in the CN are derived from the RL, although they did not discriminate between neuron types in the CN. In their studies, the development of RL-derived CN neurons was shown to be disrupted in the Atoh1-null mice. Because Hoshino et al. reported that GABAergic but not glutamatergic CN neurons are derived from Ptf1a-expressing neuroepithelial cells in the VZ (Hoshino et al. 2005), their findings suggest that cerebellar glutamatergic neurons such as granule cells and CN-Glu neurons are derived from the RL. Accordingly, unipolar brush cells, which are glutamatergic, were also shown to emerge from the RL (Englund et al. 2006). Together, these studies indicate the presence of two molecularly defined neuroepithelial areas in the cerebellum, the Atoh1-expressing RL and the Ptf1aexpressing VZ, which generate glutamatergic and GABAergic neurons, respectively. Each bHLH transcription factor is involved in producing the corresponding neuronal subtype in the cerebellum. This suggests a model in which the cerebellar neuroepithelium is regionalized into two distinct regions, the VZ and the RL, by the two bHLH transcription factors (Hoshino 2006). During embryonic development, the ventral part of the cerebellar neuroepithelium expresses Ptf1a, leading to the acquirement of cerebellar VZ characteristics to generate GABAergic neurons, while the dorsal part of cerebellar neuroepithelium expresses Atoh1 and becomes the cerebellar RL, producing glutamatergic neurons. In the telencephalon, similar regionalization by bHLH transcription factors takes place. Glutamatergic neurons emerge from dorsal neuroepithelium expressing Neurogenin 1/2 (Ngn 1/2), and GABAergic neurons are produced from ventral neuroepithelium expressing Mash1 (Wilson and Rubenstein 2000). How are these neuroepithelial areas formed? In general, the roof plate can affect the dorsal structure of the neural tube (Lee et al. 2000; Millonig et al. 2000). Chizhikov et al. revealed that the roof plate plays an important role in the formation of the cerebellar dorsoventral domain formation by analyzing cerebellar mutants that lack the roof plate (Chizhikov et al. 2006). Moreover, it has been suggested that bone morphogenetic proteins (BMPs) secreted from the roof plate as well as Notch signaling are involved in the formation of the RL and the VZ (Machold et al. 2007). A recent study that induced Purkinje cells from ES cells suggested that loss of sonic hedgehog signaling may give the dorsoventral spatial information of the cerebellar VZ to the cerebellar neuroepithelium which eventually leads to the expression of Ptf1a (Muguruma et al. 2010). Although the machinery governing GABAergic and glutamatergic neuronal subtype specification by transcription factors has been clarified to some extent, molecular mechanisms to specify each member of GABAergic (e.g., Purkinje, Golgi, basket, stellate cells and CN-ION, CN-interneurons) or glutamatergic (e.g., granule, unipolar brush cells, and CN-Glu neurons) subtype remain unclear. However, birthdating studies using 3H-thymidine and BrdU (Chan-Palay et al. 1977; Batini et al. 1992; De Zeeuw and Berrebi 1995; Sultan et al. 2003; Leto et al. 2006) as well as adenovirus (Hashimoto and Mikoshiba 2003) revealed that each type of neuron is generated at distinct developmental stages. As to GABAergic neurons, Purkinje cells are produced at an early stage (embryonic day (E) 10.5–13.5 in

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c1 . . .Atoh1 + c2 . . .Lhx1/5 + (GABAergic)

c2 (GABAergic) c4

c3

c2v

c2d . . .Corl2 +

c2d c1

Pax2 Lhx1/5

Corl2 Lhx1/5

c3 . . .Lmx1a +

Atoh1

E-cad⫾ Ptf1a pc2v

E-cad+++ Ptf1a pc2d

c2v . . .Pax2 +

RP

c4 . . .Lhx1/5 + pc2 . . .Ptf1a + pc2d . . .E-cad +++ pc2v . . .E-cad⫾

pc2(Ptf1a +)

Fig. 5.1 Domain structure of the cerebellar primordium at an early developmental stage (e.g. E12.5 in mice). The c1 domain, expressing Atoh1, corresponds to the rhombic lip that produces all types of glutamatergic neurons in the cerebellum. The pc2 is the Ptf1a-expressing neuroepithelial domain that generates all types of GABAergic cerebellar neurons. At early neurogenesis stages, such as E12.5, the pc2 domain can be subdivided into pc2d and pc2v subdomains, which expresses E-cadherin strongly and weakly, respectively. The c2 domain, expressing Lhx1/5, consists of immature GABAergic neurons putatively generated from pc2 neuroepithelial domain. This domain can also be subdivided into two subdomains, c2d and c2v. The c2d subdomain consists of corl2expressing neurons or Purkinje cells, whereas the c2v subdomain includes Pax2-positive cerebellar GABAergic interneurons. Although c3 and c4 domains are Lmx1a- and Lhx1/5-positive, respectively, cell types which consist these domains are unknown. The roof plate (RP) is located most dorsally, and plays prominent roles in organizing this cerebellar domain structure

mice), Golgi cells at middle stages (E14.5), and stellate/basket cells at a late stage (Perinatal). Regarding glutamatergic neurons, in addition to the experiment above, molecule-based lineage tracing analyses (Machold and Fishell 2005; Wang et al. 2005; Englund et al. 2006) have clarified that CN-Glu neurons leave the cerebellar RL at early stages (E10.5–12.5) and granule cells and unipolar brush cells at middle to late stages (granule cell:E12.5, ubc: E12.5–E18.5). In addition, somatic recombination-based clonal analyses suggested that Purkinje, Golgi, and basket/stellate cells as well as some CN neurons (probably GABAergic) belong to the same lineage (Mathis et al. 1997; Mathis and Nicolas 2003). These data indicate that some temporal information in the neuroepithelium may be involved in specification of neuronal types in the RL and VZ, respectively. However, the underlying molecular mechanisms have not yet been clarified. Some scientists tried to divide the structure of the cerebellar primordium into several domains (Fig. 5.1). Chizhikov et al. defined four cellular populations (denoted c1–c4 domains) in the cerebellar primordium by the expression of a few transcription factors (Chizhikov et al. 2006). c1 corresponds to the Atoh1expressing RL and c2 is located just above the Ptf1a-expressing VZ (denoted

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pc2), indicating that c2 cells mainly consist of GABAergic inhibitory neurons. Although c3 and c4 express Lmx1a and Lhx1/5 respectively, their neuronal subtypes remain to be determined. This subdomain structure is disrupted when the roof plate was removed (Chizhikov et al. 2006). Furthermore, at the early neurogenesis stage (e.g., E12.5 in mice), Minaki et al. subdivided the c2 domain into dorsally (c2d) and ventrally (c2v) located subdomains that express corl2 and Pax2, respectively (Minaki et al. 2008). While corl2 is exclusively expressed in immature and mature Purkinje cells (Minaki et al. 2008), Pax2 is expressed in GABAergic interneurons (e.g., Golgi, stellate, basket, CN-GABA neurons) in the cerebellum (Maricich and Herrup 1999; Weisheit et al. 2006). They also subdivided the Ptf1aexpressing neuroepithelial domain (pc2) into pc2d and pc2v, which strongly and weakly express E-cadherin, respectively. From the positions of the neuroepithelial and neuronal subdomains, they suggested that the pc2d neuroepithelial subdomain produces cells in the c2d domain which give rise to Purkinje cells and pc2v subdomain generates cells in the c2v that become GABAergic interneurons (Mizuhara et al. 2010). As development proceeds, pc2d and pc2v subdomains become smaller and larger, respectively, and by E14.5 in mice, the Ptf1a-expressing pc2 domain is comprised only by the pc2v subdomain which expresses E-cadherin weakly. This correlates with the fact that, at E14.5 in mice, Ptf1a-expressing neuroepithelium does not produce Purkinje cells but Pax2-positive interneurons (Maricich and Herrup 1999; Hashimoto and Mikoshiba 2003). Expression patterns of several other transcription factors in the cerebellar VZ during development were also reported. For example, some groups reported the expression patterns of proneural bHLH transcription factors, such as Ngn1, Ngn2, and Mash1 in the cerebellar VZ although their function in cerebellar development is still unclear (Zordan et al. 2008; Lundell et al. 2009). However, it has been recently reported that Pax2-positive neurons, but not Purkinje cells, are reduced in the Mash1-null cerebellum (Grimaldi et al. 2009), suggesting that these bHLH transcription factors may play distinct roles in cerebellar development. In addition, several transcription factors have been reported to participate in the development of a certain type of cerebellar neurons. Double knockout of Lhx1 and Lhx5 as well as the targeted disruption of their cofactor Ldb1 resulted in lack of Purkinje cell production in the cerebellum although Pax2-positive interneurons did not seem to be affected. Because Lhx1 and Lhx5 are expressed in post-mitotic cells, this suggests that Lhx1, Lhx5, and Ldb1 are post-mitotically involved in Purkinje cell specification (Zhao et al. 2007). In addition, targeted disruption of cyclin D2 caused loss of stellate cells in the cerebellar molecular layer, suggesting its involvement in the development of stellate cells (Huard et al. 1999). From the RL, several types of glutamatergic neurons, such as CN-Glu neurons, granule cells, and unipolar brush cells, are generated. CN-Glu neurons leave the RL at early neurogenesis stages. Some transcription factors, such as Tbr1, Irx3, Meis2, Lhx2, and Lhx9 have been found to be expressed in post-mitotic progenitors of CN-Glu neurons, but their roles have not been clarified (Morales and Hatten 2006). Other molecules, such as Zic1 (Aruga et al. 1998), have been reported to play important roles in the migration, maturation, and survival of granule cells, but the

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molecular machinery underlying the specification of granule cell identity is unknown. Although unipolar brush cells strongly express Tbr2, its function is also elusive. In addition to genetic analyses, heterotopic and heterochronic transplantation studies have also provided important clues to understanding cerebellar development (Carletti and Rossi 2008). When tissues from embryonic and postnatal cerebella were mixed and transplanted to the fourth ventricle of an adult mouse, the postnatal-derived cells differentiated only into interneurons such as granule, basket, and stellate cells, but not projection neurons, such as Purkinje cells, whereas the embryonic-derived cells were capable of becoming all types of cerebellar neurons (Jankovski et al. 1996). In addition, it was shown that dissociated cells taken from cerebellar primordium at early neurogenesis stages could differentiate into all major types of cerebellar neurons, but those from postnatal cerebellum differentiated only to Pax2-positive interneurons (Carletti et al. 2002). These findings suggest that the differentiation competence of cerebellar progenitors becomes restricted as development proceeds. However, the molecular mechanisms underlying this fate restriction process have not yet been clarified. Interestingly, Leto et al. suggested that pax2-positive interneurons, such as Golgi, stellate, basket cells, and CN-GABA interneurons are derived from same progenitor pool (Leto et al. 2006).

Specification of Precerebellar Neurons There are two types of precerebellar afferent systems: mossy fiber (MF) and climbing fiber (CF) systems. MF neurons are located in several nuclei throughout the brain stem and extend their glutamatergic projections to granule cells conveying peripheral and cortical information to the cerebellum. Four major nuclei containing MF neurons are the pontine gray nucleus (PGN), the reticulotegmental nucleus (RTN), the lateral reticular nucleus (LRN), and the external cuneate nucleus (ECN) in the hindbrain (Altman and Bayer 1987). In addition, some MF neurons are also located in the spinal trigeminal nucleus (Sp5) in the hindbrain and Clarke’s column in the spinal cord. In contrast, CF neurons reside exclusively in the inferior olive nucleus (ION), which receive input from the cerebral cortex, the red nucleus, spinal cord, and other brain stem nuclei and send glutamatergic projections to Purkinje cells (Ruigrok et al. 1995). Both types of precerebellar neurons also send branch axons to the neurons in the CN. These precerebellar systems are thought to transmit the external and internal information to the cerebellar cortex to modulate cerebellar function, including regulation of animal movement. Previous birthdating studies in mice revealed that CF neurons are generated at relatively early neurogenesis stages (E9.5–11.5) and MF neurons are produced at slightly later stages (E10.5–16.5) (Pierce 1973). Along the rostrocaudal axis, both MF and CF neurons in the hindbrain are generated from the caudal hindbrain, around rhombomeres 6–8 (r6–8), as suggested by avian grafting studies as well as mammalian fate map analyses (Ambrosiani et al. 1996; Cambronero and Puelles

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Lmx1a Atoh1 Ngn1 Mash1 Ptf1a Olig3 Pax6 Wnt1

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RP dP1 dP2 dP3 dP4 dP5 dP6

MF neurons

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Fig. 5.2 Neuroepithelial domain structure in the caudal hindbrain. In the caudal hindbrain (r6–8), several transcription factors are expressed within the dorsal neuroepithelium during embryonic development. The dorsal-most part, the roof plate (RP), expresses Lmx1a. Other than the roof plate, the dorsal neuroepithelium can be divided into six domains (dP1–dP6) according to the expression pattern of transcription factors such as Atoh1, Ngn1, Pax6, Mash1, Ptf1a, and Olig3. While mossy fiber (MF) neurons are derived from the dP1 domain expressing Atoh1, climbing fiber (CF) neurons are generated from the dP4 domain expressing Ptf1a and Olig3

2000; Farago et al. 2006; Kawauchi et al. 2006). By contrast, MF neurons in the Clarke’s nucleus are generated in the spinal cord (Bermingham et al. 2001). Classic anatomical and immunohistochemical studies have suggested that these precerebellar nuclei neurons in the hindbrain emerge from the dorsal part of the hindbrain and migrate tangentially or circumferentially to their final loci (BlochGallego et al. 1999; Yee et al. 1999; Kyriakopoulou et al. 2002). However, they take slightly different paths from each other; MF and CF neurons move extramurally and intramurally, respectively. Introduction of a GFP-expressing vector into the embryonic dorsal hindbrain allowed the dramatic visualization of migrating precerebellar nuclei neurons during development (Kawauchi et al. 2006; Okada et al. 2007). Many groups have reported transcription factors that are expressed within the dorsal neuroepithelium of the caudal (r6–8) hindbrain during embryonic development, trying to define domains along the dorsoventral axis. The dorsal-most part expressing Lmx1a corresponds to the roof plate which gives rise to the choroid plexus (Chizhikov et al. 2006). Other than the roof plate, the dorsal neuroepithelium can be divided into six domains (dP1–dP6) according to the expression pattern of the transcription factors, such as Atoh1, Ngn1, Mash1, Ptf1a, and Olig3 (Fig. 5.2). As to the precerebellar nuclei neurons, a series of studies have tried to clarify the precise origins of MF and CF neurons by genetic lineage-tracing methods. By analyzing genetically engineered mice that express lacZ or Cre recombinase under the control of the endogenous or exogenous Atoh1 promoter, MF neurons of PGN, RTN, LRN, and ECN were shown to emerge from the Atoh1-expressing neuroepithelial domain (dP1, Ben-Arie et al. 2000; Rodriguez and Dymecki 2000;

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Landsberg et al. 2005; Wang et al. 2005). Targeted disruption of the Atoh1 gene resulted in loss of these MF neurons, suggesting an involvement of Atoh1 in the MF neuron development. Atoh1 regulates the expression of the transcription factor Barhl1 (Mbh2) that is expressed in MF neurons. Loss of Barhl1 expression resulted in a decrease of MF neurons, leading to a decrease in the size of MF precerebellar nuclei (Li et al. 2004). In addition, Flora et al. reported that one of the E-proteins, Tcf4, interacts with Atoh1 and regulates differentiation of a specific subset (PGN, RTN) of MF neurons (Flora et al. 2007). Landsberg et al. also performed lineage trace analysis by using two variants of FLP (Flipperase recombinase) with different recombinase activities that were expressed under the control of the Wnt-1 promoter whose strength is the highest at the dorsal-most part and gradually decreases ventrally. They demonstrated that CF neurons are derived from the neuroepithelial region where Wnt-1 is very weakly expressed, whereas MF neurons emerge from the strongly Wnt-1-expressing region (Landsberg et al. 2005). In addition, Nichols and Bruce generated transgenic mice carrying a Wnt-1-enhancer/lacZ transgene and observed that MF neurons but not CF neurons were labeled by b-gal in those mice (Nichols and Bruce 2006). These findings suggested that CF neurons are generated from the neuroepithelial region ventral to the Atoh1-expressing domain. By Cre-loxP-based lineage trace analysis, Yamada et al. showed that all CF neurons in the ION are derived from the Ptf1a-expressing neuroepithelial region (Yamada et al. 2007). Loss of the Ptf1a gene resulted in the fate change of some CF neurons to MF neurons, suggesting that Ptf1a plays a critical role in fate determination of CF neurons. They also showed an involvement of Ptf1a in migration, differentiation, and survival of CF neurons. Storm et al. reported that not only MF neurons but also CF neurons are derived from the Olig3-expressing neuroepithelial region that broadly expands within the dorsal hindbrain (Storm et al. 2009) by CreloxP-based linage tracing. Targeted disruption of the Olig3 gene caused the disorganized development of MF neurons and complete loss of CF neurons (Liu et al. 2008; Storm et al. 2009). Moreover, the ectopic co-expression of Olig3 and Ptf1a induced cells expressing a CF neuron marker in chick embryos (Storm et al. 2009). These findings suggest that CF neurons emerge from the Ptf1a/Olig3-expressing neuroepithelial domain (dP4) and that Ptf1a and Olig3 are cooperatively involved in the development of CF neurons. Domain structure of the dorsal neuroepithelium in the caudal hindbrain region is shown in Fig. 5.2.

Conclusions and Future Directions Various types of neurons are generated from the dorsal hindbrain. As described above, the dorsal neuroepithelium of the rostral hindbrain (r1) produces all types of cerebellar neurons, while the dorsal regions of the caudal hindbrain (r6–8) generate neurons that include the precerebellar system neurons, such as MF and CF neurons. In addition, histological observations suggested that the dorsal part of the middle

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hindbrain produces neurons of the cochlear nucleus, where auditory information is processed and relayed to the brain (Pierce 1967; Ivanova and Yuasa 1998). More directly, genetic-fate-mapping studies using transgenic mice confirmed that neurons of the cochlear nucleus are derived from the dorsal part of r2–5 in mice (Farago et al. 2006), although in avians, they were shown to emerge from a broader part (r3–8) by grafting studies (Tan and Le Douarin 1991; Cambronero and Puelles 2000; Cramer et al. 2000). As to neuronal subtypes, Fujiyama et al. identified origins of inhibitory and excitatory neurons of the cochlear nucleus; inhibitory (glycinergic and GABAergic) and excitatory (glutamatergic) neurons are derived from Ptf1a- and Atoh1-expressing neuroepithelial regions, respectively (Fujiyama et al. 2009), and their development is dependent on the corresponding bHLH proteins. In the hindbrain from r1 to r8, there are dorsoventral domain structures defined by several transcription factors, which are longitudinally expressed throughout the hindbrain. Especially, two bHLH transcription factors, Atoh1 and Ptf1a seem to play important roles in specifying distinct neuronal subtypes. These two proteins are expressed in different neuroepithelial regions throughout the hindbrain

Rostral

Caudal

Dorsal

Atoh1 Ptf1a

r1

2

3

4

5

6

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Ventral Cerebellar anlage cerebellum Glutamatergic neurons GABAergic neurons

Middle hindbrain Cochlear nu.

Caudal hindbrain Precerebellar systems

Glutamatergic neurons GABAergic/ glycinergic neurons

Mossy fiber neurons Climbing fiber neurons

Fig. 5.3 Basic HLH proteins and neurons produced from the dorsal hindbrain. Atoh1 and Ptf1a are expressed in distinct neuroepithelial regions throughout the rhombomeres 1–8 (r1–8). Each number represents the rhombomeric number. Upper side is dorsal, lower is ventral. Left side is rostral, right side is caudal. Neuronal subtypes generated from the dorsal neuroepithelium of the rostral, middle, and caudal hindbrain regions are shown

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(Fig. 5.3). In both the rostral (r1) and middle hindbrain (r2–5 in mice), Atoh1 and Ptf1a participate in generating excitatory and inhibitory neurons, respectively. However, this rule is not applicable to the caudal hindbrain. The Ptf1a neuroepithelial domain in the caudal hindbrain (r6–8 in mice) produces not only inhibitory neurons (local circuit neurons) but also glutamatergic neurons (CF neurons, Yamada et al. 2007), while the Atoh1 domain generates glutamatergic MF neurons. This raises the possibility that the rostral/middle (r1–5) and caudal (r6–8) hindbrain subregions have distinct characteristics. Overall, throughout the hindbrain regions, transcription factors, such as Atoh1 and Ptf1a, seem to define neuroepithelial domains along the dorsoventral axis and participate in specifying distinct neuronal subtypes according to the rostrocaudal spatial information (Fig. 5.3).

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Specification of Granule Cells and Purkinje Cells Thomas Butts, Leigh Wilson, and Richard J. T. Wingate

Abstract

Granule cells and Purkinje cells are the major populations of neurons in the cerebellum. Their specification depends on a combination of regional identity and spatiotemporal cues. These are conferred by patterning systems in the early embryo that determine anteroposterior and dorsoventral positional coordinates and an age-dependent signal (or signals) whose nature is obscure. While a number of important questions remain about the nature of cerebellar progenitor pools and their precise boundaries, a variety of fate-mapping and genetic approaches have indicated that both granule cells and Purkinje cells arise from different dorsoventral domains within hindbrain rhombomere 1. Unusually, granule cell precursors undergo a subsequent transit amplification stage regulated by Purkinje cell signals, within a transient superficial germinal layer. Recent evolutionary insights suggest that this phase of Sonic hedgehogdependent transit amplification is only found in amniotes. Evolutionarily, since secondary proliferation arose independently of granule cell specification, it is likely to be an adaptation purely for postspecification regulation of granule cell numbers.

Introduction Over recent years, the understanding of the mechanisms that systematically pattern neurogenesis in the vertebrate CNS has been revolutionized by molecular insights into CNS patterning. A conceptual framework of Cartesian patterning axes in the early neural tube (Lumsden and Krumlauf 1996) has served as a model for

T. Butts • L. Wilson • R.J.T. Wingate (*) MRC Centre for Developmental Neurobiology, King’s College, 4th floor New Hunt’s House Guy’s Campus, SE1 1UL London, UK e-mail: [email protected], [email protected], [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_6, # Springer Science+Business Media Dordrecht 2013

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analyzing the development of a number of more complex structures. Of these, the cortex and the cerebellum provide a significant challenge by virtue of their scale and late development with respect to embryogenesis. Relating early patterning events in these regions to adult structure and function is thus a case of reconciling a number of patterning events that take place within territories that are not only growing but undergoing considerable structural reconfiguration during development. Nevertheless, through a history of fate-mapping studies and more recently developed transgenic techniques, the cerebellum has emerged as a comparatively simple developmental structure comprising two main lineages of GABAergic and glutamatergic cell types (reviewed in (Wingate 2005)). The most populous representatives of each lineage are Purkinje cells and granule cells, respectively, which also form the conserved synaptic partnership in the molecular layer that is the substrate of cerebellar function. This chapter discusses how the cerebellum is regionally allocated in the early neural tube as a molecularly distinct territory. Purkinje and granule cells arise from separate dorsoventrally located domains within this territory as part of a temporally orchestrated pattern of neurogenesis. Finally, the chapter explores the developmental and evolutionary significance of the secondary proliferation of granule cell precursors within a transient superficial layer, the external germinal layer (EGL). In particular, the emergence of the EGL in amniotes represents a marked shift in the potential interactions of neurogenic pools as granule cell precursors and Purkinje cells are brought into spatial alignment and close proximity.

Territorial Allocation Of all the regions of the CNS, the cerebellum has perhaps the longest history of study by fate-mapping techniques. Early attempts to define the territorial boundaries of presumptive cerebellum used microsurgical approaches to generate chimeric embryos from chick and quail tissue (Le Douarin 1993). Interest focused on the structural constrictions that divide the early neural tube into a superficially segmental structure. It was clear that the cerebellum developed from the region at the interface between presumptive midbrain vesicle (mesencephalon) and anterior presumptive hindbrain vesicle (metencephalon). Using a number of different types of the grafting condition, the embryonic constriction corresponding to the division between these vesicles (the midbrain/hindbrain boundary or isthmus) was, surprisingly, found to partition Purkinje cells into an anterior (midbrain derived) and posterior (hindbrain derived) pool (Hallonet et al. 1990). By contrast, the overlying layer granule cell precursors were exclusively of hindbrain origin. These studies suggested that the adult cerebellum is comprised of cells with different territorial origins. With the advent of molecular labeling techniques, the regional origins of the cerebellum were reassessed in terms of gene expression. In particular, the attention

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of various groups turned to the most anterior hindbrain segment (rhombomere 1) that comprises the anterior two thirds of the metencephalon. Rhombomere 1 lies between the nested domains of Otx gene expression that define the head, midbrain, and forebrain (Acampora et al. 1995) and the cluster of Hox genes that are expressed in hindbrain and spinal cord (Lumsden and Krumlauf 1996). In this model, the posterior limit of Otx expression defines the anterior boundary of the cerebellum. Its caudal extent is delimited by the boundary of the most anterior Hox gene, Hoxa2 (Fig. 6.1a, b). Supporting evidence for Hox genes setting a caudal limit to cerebellar neurogenesis came from analysis of the Hoxa2 mutant mouse. Hoxa2 is the most anterior homolog and the only Hox gene to be expressed in rhombomere 2. Its deletion in mice results in a Hox-free territory between rhombomere 1 and rhombomere 3 and a caudal expansion of the cerebellum into the hindbrain (Gavalas et al. 1997), while the complementary overexpression of Hoxa2 within the previously Hox-free rhombomere 1 suppresses the formation of granule cells (Eddison et al. 2004). Molecular evidence for Otx genes comprising an anterior boundary of the cerebellum was initially prompted by an investigation of the expression domain of Otx2 with respect to morphological constriction that had previously been used as fate-mapping landmark in avian chimeras. Careful analysis revealed that the molecular boundary of midbrain territory only converges on the morphological landmark of the isthmic constriction, at stages after fate-mapping had previously been carried out (Millet et al. 1996). This raised the possibility that the dual origin of the Purkinje cell epithelium from midbrain and hindbrain resulted from a reliance on morphological rather than genetic regional boundaries. Subsequent fate-mapping of granule cells with respect to both Hoxa2 expression demonstrated that the precursors of this population are entirely contained within rhombomere 1 (Wingate and Hatten 1999). The designation of rhombomere 1 as the origin of all cerebellar neurons is consistent with a wealth of studies on the molecular patterning of cerebellar territory with respect to the isthmus, an organizer that forms at the midbrain/ hindbrain boundary (Joyner 1996). Firstly, deletions of genes whose expression domains overlap rhombomere 1 at some stage of development, such as Wnt1 (McMahon and Bradley 1990) and Gbx2 (Wassarman et al. 1997) or Engrailed-1 (Wurst et al. 1994) can result in a dramatic loss of the majority of cerebellar neurogenesis. Secondly, disruption of genes expressed specifically at the midbrain/hindbrain boundary such as fibroblast growth factors (FGF) can lead to the loss or severe reduction in size of the adjacent cerebellum in mice (Xu et al. 2000) and zebrafish (Brand et al. 1996; Reifers et al. 1998). These and a number of experiments using protein overexpression have suggested that FGFs may consequently have an instructive role in cerebellar induction (Koster and Fraser 2006; Martinez et al. 1999; Matsumoto et al. 2004). Others have argued for a more indirect role in patterning through, for example, regulating the position of the rostral boundary of Hoxa2 expression (Irving and Mason 2000) and hence the

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Fig. 6.1 The origins of cerebellum territory and Purkinje and granule cells. (a) In situ hybridization for Otx2 and Hoxa2 an early stage vertebrate neural tube (Chick, E2) shows the unlabelled region corresponding to rhombomere (r)1. Fate-mapping and experimental studies indicate that this is the source of granule cells and possibly all other cerebellar neurons. (b) Schematic representation of how r1 territory maps onto a later embryo lying between midbrain (mb) and hindbrain (hb) regions. The posterior boundary of cerebellar territory is defined by widest angles of the roofplate of the fourth ventricle (iv). The diamond-shaped rhombic lip lies at the edge of the fourth ventricle roofplate. (c) Granule cell precursors and Purkinje cells are generated from nonoverlapping territories progressively more distal to the roofplate. (d) The standard model of the relationship between progenitors/precursors (solid circle) and their derivatives (empty circle) is shown for Ptf1a (Purkinje) and Atoh1 (granule) lineages. The molecular signature of progenitors of the Atoh1 migratory precursor pool are as yet unknown (gray). Patterns of cell movement are shown by arrows: Purkinje neurons and granule precursors migrate along radial (i) and tangential (ii) trajectories, respectively. Roofplate cells are generated predominantly adjacent to the rhombic lip and passively progress into this nonneural epithelium (Currle et al. 2005; Nielsen and Dymecki 2010). The ventricular zone (vz), rhombic lip (rl), and roofplate (rp) produce different lineages and are generally believed to represent different developmental compartments

ultimate size of the cerebellum. Of these latter studies, evidence that FGF can induce cerebellar territory indirectly, by down regulating Otx expression (Foucher et al. 2006; Sato and Joyner 2009), offers strongest support for the cerebellum as the derivative of an Otx-negative domain.

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Dorsoventral “Compartments” and the Origin of Cell Types As in the spinal cord, dorsoventral patterning cues play a major part in the allocation of cell types within the presumptive cerebellum. The cerebellum is approximately derived from the dorsal half of the neural tube (Hallonet and Le Douarin 1993) and as might be expected, dorsalizing signals, in particular TGFbs, play an important role in specifying at least some neuron fates (Alder et al. 1996, 1999; Helms and Johnson 2003). The source of dorsalizing signals is the roof plate of the fourth ventricle (Chizhikov et al. 2006; Lee et al. 2000; Millonig et al. 2000), which comprises an expanded nonneural sheet of cells which also secretes a number of potent patterning molecules such as Wnt proteins (Rodriguez and Dymecki 2000). Gene ablation and genetic fate-mapping techniques have revealed that Purkinje cells and granule neuron precursors are born in distinct domains, respectively, defined by the expression of Ptf1a (Hoshino et al. 2005) and Atoh1 (Ben-Arie et al. 1997; Machold and Fishell 2005; Wang et al. 2005) (Fig. 6.1c). The expression of Atoh1 is restricted to the neural margins of roof plate, the embryonic rhombic lip (Wingate 2001). Ptf1a defines broader territory embryonically ventral to the rhombic lip (Hoshino et al. 2005). The expression of Atoh1 can be induced by ectopic TGFbs (Alder et al. 1999) and is lost following either TGFb knockdown (Lee et al. 1998) or following a more radical genetic ablation of roof plate (Lee et al. 2000). However, this latter manipulation does not, however, lead to a complete loss of the Ptf1a domain (Chizhikov et al. 2006). Thus, while there appears to be a conserved function for Atoh1 and Ptf1a between cerebellum and spinal cord in the specification of a glutamatergic or GABAergic neuronal phenotype, respectively (Bermingham et al. 2001; Glasgow et al. 2005; Helms and Johnson 1998; Miesegaes et al. 2009), induction of the Ptf1a domains in the hindbrain might rely on additional, as yet undetermined, patterning molecules. One defining feature of Purkinje and granule cell lineages is their mutual exclusivity. No Mash1 (Kim et al. 2008) or Ptf1a (Hoshino et al. 2005) positive cells give rise to precursors within the rhombic lip (Fig. 6.1c). No Atoh1(Math1) rhombic lip precursors give rise to GABAergic neurons (Machold and Fishell 2005; Rose et al. 2009; Wang et al. 2005). Does this reflect a genuine compartmentation of the dorsoventral axis into lineage restriction units, or the segregation of neuroblasts or committed precursors from an uncommitted progenitor or stem cell pool? Determining the answer to this question depends on when bHLH genes are expressed in progenitors (stem cells), precursors (partially or wholly committed dividing cells), and/or neuroblasts (young neurons). Evidence from tamoxifen induced pulse labeling demonstrates that the expression of at least one bHLH gene, Atoh1(Math1), is confined to young neuroblasts and committed precursors (including the EGL) but not progenitors (Machold and Fishell 2005): no derivatives of Atoh1-positive cells reenter the rhombic lip (Fig. 6.1d). Extensive mixing of proliferative cells along the DV axis (Clarke et al. 1998; Wingate and Lumsden 1996; Wingate and Hatten 1999) suggests that rhombic lip progenitors should

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Fig. 6.2 Alternative models to explain the effects of mutation of bHLH genes in Purkinje and granule cell lineages. The compartmental model where progenitors are lineage restricted (a) is contrasted with a “replenishment” model (b) where progenitors can mix between territories. The location of ectopic cells in bHLH mutant mice is shown on a single composite of cell locations in the Ptf1a (Pascual et al. 2007) and Atoh1(Math1) (Rose et al. 2009) mutant mice. In a compartment model (c) ectopic, ventricular zone-derived Atoh1+ve cells that enter the EGL must be derived by aberrant radial migration (Pascual et al. 2007). Ectopic rhombic lip derived cells must similarly be assumed to enter the roofplate lineage through aberrant tangential migration. In a “replenishment” model where uncommitted progenitors can cross into each of these precursor zones (d) knockout phenotypes interpret the role of bHLHs as simply regulating commitment and differentiation. When Ptf1a is downregulated, precursors can enter the EGL via tangential migration. The final position of ectopic EGL cells may reflect the earlier birth date of GABAergic interneurons with respect to the prolonged assembly of the EGL (cells generated early in the lineage lie most distal to the rhombic lip)

stochastically be drawn from the ventricular zone. However, this would require that both Ascl1(Mash1) and Ptf1a bHLH genes were similarly restricted to committed precursors (but not progenitors) since neither Ptf1a nor Ascl1(Mash1) expressing cell lineages contribute to the rhombic lip. Resolving this discrepancy remains an important question in cerebellar patterning. On the one hand, the observations of Machold and Fishell (2005) could be interpreted as revealing the presence of a spatially compartmentalized population of dedicated rhombic lip stem cells (Fig. 6.2a). On the other hand, if the rhombic lip were continually replenished from the ventricular zone (Fig. 6.2b) there is an as yet unidentified progenitor common to all cerebellar neuronal subtypes that might be free to spread across “compartment” boundaries. Limited evidence in favor of the latter “replenishment” model comes from tamoxifen-mediated, pulse labeling of Ascl1(Mash1) derivatives in mouse. This reveals that few if any Ascl1(Mash1)-positive cells reenter the ventricular zone (Kim et al. 2008) but analysis was only performed after a relatively long postinjection survival of several days. Similarly, in vitro evidence that cerebellar

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stem cell progenitors can generate both GABAergic and glutamatergic lineages (Lee et al. 2005), is countered by evidence that prominin-positive cerebellar stem cells in vivo are restricted to nongranule cell fates, at least in late embryogenesis (Zhang and Goldman 1996). Finally, a replenishment model might explain the unusual lineage “violations” that accompany the knockout of both Ptf1a and Atoh1(Math1) bHLH genes. Following Ptf1a knockout, cells from the Purkinje cell domain enter the EGL and express Atoh1(Math1) (Pascual et al. 2007). Following the knockout of Atoh1 (Math1), cells from the rhombic lip can enter the gdf7 roof plate lineage (Rose et al. 2009). In both cases, normal lineage segregation between progenitor has been disrupted. While this has been interpreted as indicating cross-inhibition of expression between bHLH genes (Pascual et al. 2007) (Fig. 6.2c), it is possible that bHLH genes are upstream of events that segregate precursors and neurons, but that multipotent progenitors themselves are not restricted to lineage compartments (Fig. 10.2d). This interpretation leaves open the possibility that a progenitor is free to contribute to any lineage, in vivo, depending on patterns of stochastic cell mixing between rhombic lip and the rest of the ventricular zone and/or roof plate.

Temporal Patterning and Lineage in the Rhombic Lip Both granule cell precursors and Purkinje cells represent only a subset of the neurons born within their respective dorsoventral domains. Purkinje cells are one of a number of GABAergic interneurons that collectively regulate cerebellar activity. For GABAergic neurons generated from the ventricular zone distal to the rhombic lip, regional differences in gene expression may underlie some of this cell fate diversity (Chizhikov et al. 2006; Mizuhara et al. 2010; Morales and Hatten 2006; Zordan et al. 2008) in addition to a demonstrable sequence in the production of cerebellar nuclei and Purkinje cells from the Ascl1(Mash1)-positive domain (Kim et al. 2008). For Atoh1-positive precursors, temporal patterning plays a pivotal role in generating cell fate diversity. Application of a transgenic inducible reporter in mouse (Machold and Fishell 2005) confirmed avian fate-mapping studies (Gilthorpe et al. 2002) that established a temporal sequence of cell production from the cerebellar rhombic lip that culminates in the production of migratory granule cell precursors. Rhombic lip derivatives include a subset of cerebellar nuclei (Fink et al. 2006; Machold and Fishell 2005), unipolar brush cells (Englund et al. 2006), and a variety of early born, rostral hindbrain nuclei (Gilthorpe et al. 2002; Machold and Fishell 2005; Rose et al. 2009). The factors that determine the temporal allocation of cell fate in the cerebellum remain obscure. Heterochronic transplantation reveals that, in the rhombic lip, precursors are temporally committed to a given fate, but that temporal transition between commitment states requires an exogenous signal (Wilson and Wingate 2006). The nature this exogenous signal is of central importance to understanding the specification of granule cell precursors (the last born lineage). In early

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embryonic stages, local signaling factors released from the roof plate (Chizhikov and Millen 2004), including the nuclear hormone retinoic acid (Wilson et al. 2007; Yamamoto et al. 2003), are well placed to influence temporal patterning. However, it is possible that circulating hormones also influence maturation, similar to the role thyroid hormone plays in regulating later cerebellar maturation (reviewed in (Koibuchi 2008)).

Secondary Proliferation and Neurogenesis A unique characteristic of cerebellar development is the segmentation of granule cell production into two distinct phases. The first is at the rhombic lip while the second takes place within a transient superficial layer of proliferating cells formed by the tangential migration of committed granule cell precursors (Alder et al. 1996; Wingate and Hatten 1999). Precursors in this external germinal layer are characterized by the extended expression of Atoh1(Math1). When this gene is mutated, the EGL fails to form (Ben-Arie et al. 1997). Elegant fate-mapping using the MADM technique in mouse has shown that the proliferation of EGL precursors is characterized, in the main, by symmetrical divisions (Espinosa and Luo 2008). Precursors have moreover been recently shown to give rise exclusively to granule cells (Alder et al. 1996; Espinosa and Luo 2008; Klein et al. 2005), contradicting earlier assumptions (Lin et al. 2001) about the possible derivatives of the EGL. Both mode of division and committed cell fate make the EGL an excellent model for studying the biological significance of transit amplification in CNS development. Transit amplification of granule cell precursors within the EGL is driven by sonic hedeghog (Shh) secreted from the underlying layer of Purkinje cells (Dahmane and Ruiz-i-Altaba 1999; Lewis et al. 2004; Wallace 1999; WechslerReya and Scott 1999). Shh signaling is downstream of the expression of the orphan retinoic receptor, RORalpha in Purkinje cells (Gold et al. 2003) and its transduction in granule cells is modulated by IGF signaling (Fernandez et al. 2010) and mediated by the proto-oncogene, Nmyc (Kenney et al. 2003, 2004). The retention of Atoh1 by granule cells is absolutely required for their proliferative response to Shh (Flora et al. 2009; Zhao et al. 2008) and the amplitude of Shh signaling determines the size of the EGL and hence the surface area and degree of foliation of the cerebellum (Corrales et al. 2004, 2006). The transit amplifying function of the EGL also has important clinical relevance. A significant proportion of medulloblastoma can be attributed to the mutations within the Shh signaling pathway (Ellison et al. 2003; Goodrich et al. 1997; Kool et al. 2008) with respect to granule cell proliferation (Schuller et al. 2008; Yang et al. 2008). In addition to exponentially expanding the population of granule cell precursors, formation of the EGL spreads granule cells throughout the cerebellum to evenly cover the developing Purkinje cell layer. A postmitotic phase of radial migration translates this spatial distribution into the internal granule cell layer that is thus in

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spatial alignment with its target neurons. This correlated development implies interactions between Purkinje cells and granule cells that extend beyond the regulation of the pace of proliferation. Accordingly, Purkinje cell layering is disrupted in the absence of granule cells (Jensen et al. 2004). Similarly, in mice that are chimeric for mutations that disrupt the spatial distribution of granule cells (Unc5H3 (Goldowitz et al. 2000)), their number or maturation (Pax6 (Swanson and Goldowitz in press)), Goldowitz and colleagues have demonstrated how the normal developmental program of granule cells is essential for normal Purkinje cell spatial distribution. Whether reciprocal interactions are required to mediate normal differentiation of either cell type is, however, unclear.

Diversity of Granule and Purkinje Cells The adult cerebellum is notable for a circuitry whose extraordinary uniformity has long been seen as critical for its function (Braitenberg 1961; Cajal 1894, 1911; Dean et al. 2010). Superimposed on this uniform cellular architecture is a sagittal molecular organization, famously revealed in stripes of expression of Zebrin/ aldolase C that divides the Purkinje cells into longitudinal molecular domains (Hawkes and Herrup 1995). This parasagittal organization is reflected by the subdivision of the functional cerebellar circuit in parasagittal microzones (Apps and Garwicz 2005). Running perpendicular to sagittal domains are transverse folia. To some extent folia simply appear to be a structural consequence of increasing surface area through EGL proliferation (Corrales et al. 2006). However, it is clear that the folia lobe imposes a functional organization on certain aspects of interneuron connectivity (Watt et al. 2009). Similarly, the folia may correspond to anteroposterior molecular and lineage landmarks within the cerebellum (Hawkes et al. 1999). Together, these sets of observations suggest a large degree of molecular heterogeneity within the uniform cellular structure of the cerebellum. How much, if any, of these aspects of cellular heterogeneity are determined at neurogenesis? Purkinje cells are born with a relatively discrete temporal window (Kim et al. 2008) and ventricular zone characterized by uniform gene expression (Hoshino et al. 2005). Indirect studies of lineage through x-inactivation model in mouse (Baader et al. 1996) and retroviral lineage dispersal in chick (Lin and Cepko 1999) show no evidence for topographic organization of neurogenesis within the Purkinje cell precursor pool. Nevertheless, birth dating evidence in the rodent suggests an anterior posterior variation in birth dates of Purkinje cells in different regions of the cerebellum (Altman and Bayer 1985). How this relates to patterns of neurogenesis in early development is unclear. The paired plates of the mammalian cerebellar anlage undergo a significant rotation as the midline fuses (Sgaier et al. 2005) reconfiguring the early embryonic axes with respect to adult topography. The significance of maturational gradients is also challenged by the observation that these morphological movements are absent during fusion of the avian cerebellum (Alexandre and Wassef 2003) and yet both bird and mouse have a distinct parasagittal molecular organization.

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For granule cells, timing of migration and division confer different aspects of morphological and spatial ordering. Just as different populations of rhombic lip derivatives terminate their migration at successively more proximal targets (Gilthorpe et al. 2002; Machold and Fishell 2005; Wang et al. 2005; Wingate 2005), temporal cohorts within the granule cell precursor pool appear to be spatially addressed as they leave the rhombic lip (Jensen et al. 2004). The last born granule cell population in posterior cerebellum is also characterized by the expression of Lmx1a (Chizhikov et al. 2010). Once a coherent EGL has formed, tangentially orientated, migratory inner cells (Cajal 1894) disperse over large distances (Ryder and Cepko 1994), obscuring any potential temporal order in initial EGL deposition. Subsequent proliferation results in a single point within the inner granule cell layer being populated by granule cells of a wide range of birthdates. Within each locus, younger granule cells project axons to more superficial strata within the molecular layer and may in turn be exposed to different molecular layer interneuron milieu (Espinosa and Luo 2008). However, there is no evidence, by comparison to the cerebral cortex (McConnell and Kaznowski 1991), that the timing of granule cell birth confers laminar identity on parallel fibers. In summary, it seems unlikely that any of the spatiotemporal patterning that might potentially specify subpopulations of Purkinje cell or granule cell have significance for later functional organization. Although developmental mechanisms result in chronotopic ordering at various stage of granule cell development, the processes involved in successive stages of development erase the order established in the previous phase. The limited degree to which later embryonic pattern is transferred to adulthood is also a characteristic of the ontogeny “late onset” banding patterns (Larouche and Hawkes 2006). Overall, the rewriting of developmental patterns in successive stages of development appears to be a feature of cerebellar development.

An Evolutionary Perspective on Neurogenesis The cerebellum in vertebrates presents an enormous variety of size and form but is characterized by a highly conserved synaptic arrangement between granule cell axons and Purkinje cell dendrites within a conserved molecular layer (Nieuwenhuys et al. 1998; Sultan and Glickstein 2007). From this, the conserved elements of neurogenesis that confer Purkinje cell and granule cell identity might expect to be highly conserved. Rhombomere 1 itself, as defined as an Otx-negative, Hox-negative region, is present in all chordates (Wada et al. 1998) and may represent a fundamental component of the tripartite vertebrate neural tube (Holland and Holland 1999). Although, molecular correlates of the Purkinje cell and granule cell precursor pools have not been studied in basal vertebrates, it seems likely that a rudimentary cerebellum is found in agnathans such as lamprey. Granule cells and Purkinje cells arise in different developmental compartments as part of temporally patterned lineages that give rise to a range of other glutamatergic and GABAergic neurons, respectively. The range of cell types

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produced in these two compartments varies between vertebrates (Butler and Hodos 1996; Nieuwenhuys et al. 1998), suggesting that the temporal sequence of cell fate allocation may be a substrate of evolutionary adaptation. Within this diversity, Purkinje cells and granule cell production is conserved. The increasing recognition of the importance of non-Purkinje GABAergic interneurons in cerebellar function (Dean et al. 2010) make these variations potentially highly significant. While changes in the interneuron complement account for some of the developmental variability between species, changes in the scale and foliation of the cerebellum have been more thoroughly investigated and correlate with behavioral attributes (Lisney et al. 2008; Sultan and Glickstein 2007; Yopak et al. 2007; Yopak and Montgomery 2008). The developmental origins of changes in the gross morphology of cerebellum can be traced to variations in the way that neurogenesis is organized in different species (Butts et al. 2011). In particular, size and number of folia can be related to the presence of an EGL and the degree of proliferation of its component precursors. Molecular analyses have only recently shown that an Atoh1positive EGL only emerges relatively late in evolution as a feature of amniote embryogenesis (Chaplin et al. 2010). Ancestral granule cells can be presumed to have differentiated at the rhombic lip as they do in modern chondrichthyans. Granule cells became tangential migrants in the bony fish lineage (Chaplin et al. 2010; Kani et al. 2010), but only first formed a superficial granule cell layer in tetrapods (Gona 1972). Shh responses in Atoh1-positive precursors are a feature that is likely to first appear in the reptile clade (Cajal 1891) (Fig. 6.3). The absence of an EGL is likely to be significant for both the distribution of granule cells and their interactions with Purkinje cells. For example, formation of an internal granule cell layer in the absence of an EGL engages a different mode of tangential migration. In zebrafish, postmitotic rhombic lip derived granule cells migrate directly to their target position through neuronal layers using a form of chain migration (Kani et al. 2010; Rieger et al. 2009). Correspondingly, Purkinje cells fail to express Shh (Chaplin et al. 2010; Kani et al. 2010), raising the question of how much of the interaction between cell types that characterizes the development of the amniote cerebellum is present in teleosts (Fig. 6.3). What the development of the fish cerebellum also demonstrates is that, perhaps surprisingly, an EGL is neither required to regulate appropriate development of the cerebellum or the organization of the molecular layer. The fish cerebellum reaches its most complex in electrosensory species such as the mormyrid fish where it becomes the dominant CNS structure covering much of the surface of the brain (Nieuwenhuys and Nicholson 1969). What then is the significance of the EGL, and specifically Shh-mediated transit amplification in granule cell neurogenesis? One explanation for the evolution of the EGL may be its ability to rapidly amplify granule cell numbers during the defined period of embryogenesis mandated by land colonization. By contrast, fish grow continuously throughout life and require a continuous addition of new circuit elements to the expanding cerebellum. In amniotes, a defined transient amplification stage may also provide a selective advantage through the fine control of complementary cell numbers that it affords. Even closely related bird and mammal species can show large differences in

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Fig. 6.3 Developmental innovations in the evolution of the cerebellum. The vertebrate radiation (left) is shown with respect to adult cerebellum morphology (right) and the first appearance of developmental motifs (middle). While tangential migration of granule cells is found in fish, a superficial transient granule cell layer first appears only in frog. Shh expression is absent in fish (and possibly amphibians). Shh is, however, expressed in sharks although granule cells do not transduce Shh signals (Chaplin et al. 2010). Purkinje cell/granule cell interactions appear to be a feature that may have first evolved in amniotes

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cerebellar size which are presumably mediated through subtle changes in Shh signaling from Purkinje cells to granule cells (Corrales et al. 2006; Sultan 2005; Sultan and Glickstein 2007). Evolutionary evidence might hence argue that one distinct role of the EGL may be to simply link the neurogenic program of Purkinje and granule cells by bringing displaced populations into close proximity. Conclusions

Neurogenesis in the mammalian cerebellum is circumscribed by anteroposterior, dorsoventral, and temporal patterning constraints. The production of the two major cell types, granule cells and Purkinje cells, is dependent on the expression of the bHLH genes Atoh1 and Ptf1a, respectively. Specification in populations of either cell appears to be homogenous with respect to the later organization of the cerebellum into functional and molecular microdomains. This suggests that cerebellar patterning with respect to function is an ongoing process throughout development. Neurogenesis thus seems mainly to be concerned with making the right number of neurons. To accomplish this, in birds and mammals, a unique phase of granule precursor cell tangential migration brings granule cells into a signaling interaction with Purkinje cells, as the transient EGL. However, the formation of an EGL is not a prerequisite for a large and functionally sophisticated cerebellum and anamniotes (sharks, fish, and amphibians) have evolved alternative solutions to generating the appropriate number of appropriately deployed granule cells.

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Granule Cell Migration and Differentiation Yutaro Komuro, Jennifer K. Fahrion, Kathryn D. Foote, Kathleen B. Fenner, Tatsuro Kumada, Nobuhiko Ohno, and Hitoshi Komuro

Abstract

In the developing cerebellum, granule cells migrate from their birth place to their final destination. The active translocation of granule cells is essential for the formation of cerebellar cortical layers and their proper differentiation. This chapter will review (1) how granule cells migrate from their origin to their resident destinations in the developing cerebellum, (2) the mechanisms involved in normal and abnormal migration of granule cells, and (3) the mechanisms underlying the differentiation of granule cells.

Introduction After final cell division, postmitotic neurons migrate from their sites of origin to their final destinations, where they reside during their entire adult life. This movement of immature neurons is a fundamental cellular event essential for building large neuronal assemblies (Valiente and Marin 2010). Distinct genetic mutations and environmental toxins can affect neuronal migration in humans and result in abnormal development of the brain, leading to neurological disorders (Guerrini and Parrini 2010). During the last five decades, granule cell migration has been extensively studied and used as a model system for neuronal migration. This is because the mechanisms underlying granule cell migration are utilized during the migration of immature neurons in other brain regions (Komuro and Rakic 1998b; Jiang et al. 2008). The role of neuron–glia interaction in neuronal migration was first discovered in the migration of granule cells along the Bergmann glial processes in the

Y. Komuro • J.K. Fahrion • K.D. Foote • K.B. Fenner • T. Kumada • N. Ohno • H. Komuro (*) Department of Neurosciences, NC30, Lerner Research Institute, Cleveland Clinic Foundation, 9500 Euclid Avenue, Cleveland, OH, 44195, USA e-mail: [email protected], [email protected], [email protected], [email protected], [email protected], [email protected], [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 107 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_7, # Springer Science+Business Media Dordrecht 2013

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developing cerebellum (Rakic 1971). This discovery led to the findings that in the developing cerebrum, immature neurons use radial glial processes as a scaffold for their migration. Likewise, the role of cell adhesion molecules in neuronal cell migration was first discovered in granule cells (Rakic et al. 1994). To date, wide varieties of cell adhesion molecules, which play a critical role in neuronal cell migration, have been identified in other regions of the brain. Moreover, the regulation of neuronal cell migration by neurotransmitters was first reported in granule cell migration (Komuro and Rakic 1993), followed by the discovery showing the key role of neurotransmitters in the migration of cerebral neurons. This chapter will first describe the recent findings revealing that granule cells exhibit cortical layer-specific changes in their migration. It will then discuss how the cortical layer-specific migration of granule cells is regulated by intrinsic programs and both extracellular and intracellular signals. Next, it will present recent studies examining how alcohol exposure adversely affects granule cell migration in the developing cerebellum. Finally, it will review the mechanisms involved in the early differentiation of granule cells during the period of their migration.

Granule Cells Exhibit Different Modes, Speeds, and Directions of Migration at Different Cortical Layers Granule cell precursors begin to proliferate in the upper rhombic lip of the mouse embryo by embryonic day 10 (E10). Thereafter, granule cell precursors start to migrate tangentially in a lateromedial and posteroanterior direction to cover the superficial zone of the embryonic cerebellum (Miale and Sidman 1961). By E15, most of the cerebellar surface is covered by granule cell precursors. The cell layer occupied by granule cell precursors is called the external granular layer (EGL). After clonal expansion in the EGL, granule cell precursors begin to produce postmitotic granule cells after birth. Postmitotic granule cells alter their shape concomitantly with changes in the mode and rate of migration as they migrate toward their final destination within the internal granular layer (IGL) (Komuro and Yacubova 2003). This section will describe the translocation and transformation of postmitotic granule cells from the EGL to the IGL. The cortical layer-specific changes in granule cell migration are schematically represented in Fig 7.1.

The External Granular Layer (EGL) In the early postnatal cerebellum of mice, granule cell precursors actively proliferate every 20 h at the top of the EGL. After their final mitosis, granule cells remain in the EGL for 20–48 h before initiating their radial migration across the molecular layer (ML). The significance of this latent period is not well understood. Recent studies revealed that at the middle of the EGL, coincident with the extension of two horizontal processes, postmitotic granule cells start to migrate tangentially in the direction parallel to the longitudinal axis of the folium (Komuro et al. 2001). Their

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Fig. 7.1 The three-dimensional representation of granule cell migration from the EGL to the IGL in the early postnatal mouse cerebellum. 1, Extension of two uneven horizontal processes near the top of the EGL; 2, Tangential migration in the middle of the EGL; 3, Development of a vertical process near the border between the EGL and the ML; 4, initiation of radial migration at the EGL-ML border; 5, Bergmann glia-associated radial migration in the ML; 6, Stationary state in the PCL; 7, Glia-independent radial migration in the IGL; 8, Completion of migration in the middle or the bottom of the IGL. P Purkinje cell, B Bergmann glia, G Golgi cell, g postmigratory granule cell, cf climbing fiber, mft mossy fiber terminal

morphology and the speed of cell movement change systematically with their position within the EGL (Komuro et al. 2001). For example, the rate of tangential cell movement is fastest (14.8 mm/h) in the middle of the EGL, when the cells have two short horizontal processes. As granule cells elongate their soma and extend longer horizontal processes at the bottom of the EGL, they move at a reduced rate (12.6 mm/h). At the interface of the EGL and the ML, granule cells migrate tangentially at the slowest rate (4.1 mm/h). During the tangential migration at the EGL-ML border, granule cells begin to extend descending processes from their somata into the ML. Granule cells retain two elongated horizontal processes while their nucleus and surrounding cytoplasm start to enter into the short vertical process descending into the ML. It takes 30 min for the complete translocation of the nucleus and surrounding cytoplasm from the horizontally extended process to the vertical process. After the completion of nucleus re-orientation, the granule cell somata quickly enter the ML. As a result of the soma’s translocation within the leading process at the EGL-ML border, granule cells develop a thin trailing process connected to two horizontal processes. These

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horizontal processes emitted from each side of the granule cell somata transform into future parallel fibers. Although the majority of parallel fibers develop from the two preexisting horizontal processes of tangentially migrating granule cells, there is another mechanism for the formation of parallel fibers. During the initiation of radial migration at the EGL-ML border, the tip of horizontally extended leading processes turns toward the ML, which is followed by the granule cell somata. As a result, the horizontal trailing process of granule cells becomes one side of the parallel fibers. Subsequently, granule cells develop a new small process at the rear part of the vertically elongated somata. The new process extends toward the opposite direction of the extension of the horizontal trailing process, and becomes the other side of the parallel fibers (Komuro et al. 2001).

The Molecular Layer (ML) In the ML, granule cells have vertically elongated cell bodies, thin trailing processes, more voluminous leading processes, and migrate radially along the Bergmann glial processes. The rates of granule cell migration in the ML depend critically on the age of the cerebellum: the average rate of cell migration in the ML increases systematically as development proceeds (Komuro and Rakic 1995). Consequently, granule cells traverse the developing ML within a relatively constant time period despite the doubling in width of the ML during the second week of postnatal life. Granule cell migration in the ML is characterized by alternations of short stationary phases with movement in a forward or backward direction. The net displacement of the cells depends on the duration and frequency of these phases as well as on the speed of movement (Komuro and Rakic 1995). At the bottom of the ML, the vertically elongated somata of granule cells move toward the Purkinje cell layer (PCL), while the length of their leading process gradually decreases (Komuro and Rakic 1998a). The shortening of the leading process is due to the advance of the granule cell somata within the leading process rather than from active retraction. The distal portion of the leading process positioned in the PCL begins to extend large motile lamellipodia and filopodia. This is not a characteristic of the leading process of granule cells in the ML, which is invariably associated with Bergmann glial fibers and usually tapers without motile lamellipodia.

The Purkinje Cell Layer (PCL) Once the somata of granule cells enter the PCL, the shape abruptly transforms from a vertically elongated spindle to a sphere (Komuro and Rakic 1998a). The rounded somata significantly slow their movement, and stop completely in the PCL. The rounded somata remain stationary in the PCL for an average of 115 min, with time ranging from 30 to 220 min (Komuro and Rakic 1998a). However, highly motile lamellipodia and filopodia develop at the distal portion of the leading process, suggesting that the tips of leading processes actively search for potential guidance cues. After a prolonged stationary period, granule cells in the PCL begin

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to re-extend their somata and leading processes. During this transformation, granule cells gradually accelerate the rate of their migration and cross the PCL-IGL border.

The Internal Granular Layer (IGL) The spindle-shaped granule cells migrate toward the bottom of the IGL at a rate comparable to that recorded for granule cells migrating along Bergmann glial fibers within the ML (Komuro and Rakic 1998a). The long axis of the granule cell somata remains oriented perpendicular to the PCL-IGL boundary line during this radial migration. Once the tip of a leading process approaches the IGL-white matter (WM) border, the granule cell somata become rounded. Granule cells then slow their migration and stop their movement near the IGL-WM border. In the postnatal day 10 (P10) mouse cerebellum, the majority of granule cells complete their migration at the bottom stratum of the IGL, while less than 20% of the cells settle in the middle or top strata (Komuro and Rakic 1998a). Although there are large differences in the total migrating distance of granule cells between different species and between different ages in a given species, in the P10 mouse cerebellum, granule cells first move tangentially 220 mm in the EGL, and then migrate radially 250 mm to attain their final position in the IGL. The average transit time of granule cells is 25.0 h in the EGL, 9.8 h in the ML, 5.2 h in the PCL, and 11.1 h to attain their final position in the IGL. Therefore, granule cells move from the top of the EGL through the ML and the PCL to their final position in the IGL within 2 days (average, 51 h) after the initiation of their tangential migration in the EGL (Yacubova and Komuro 2003).

Mechanisms Involved in Normal and Abnormal Migration of Granule Cells This section will first review the mechanisms which are responsible for the regulation of granule cell migration in a cortical layer-specific manner. It will then discuss how alcohol exposure during the period of cerebellar development adversely affects granule cell migration.

Control of Granule Cell Migration by Intrinsic Programs Although the cortical layer-specific changes in granule cell migration are likely to be induced by responses to local environmental cues, the alterations of migratory behavior may also depend partially on an internal clock or intrinsic programs. This is because in the microexplant cultures of the early postnatal mouse cerebellum, isolated granule cells sequentially go through three characteristic phases of migrating behavior and morphology without contacting other cells and processes (Yacubova and Komuro 2002a). This indicated that inherent (intrinsic) programs

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control the alterations of granule cell migration. Three characteristic phases (phase I, phase II, and phase III) of sequential changes in migratory behavior of isolated granule cells and their morphology are as follows:

Phase I (PI, a Period of 0–20 h In Vitro) During the early stage of PI, granule cells repeatedly change the shape of their somata from spherical to spindle and vice versa. They frequently turn to the left or right without extending leading processes (Yacubova and Komuro 2002a). At the points at which granule cells change their direction of movement, they stop their movement, become round, and then extend their cell bodies in the direction of the upcoming movement. Shortly after the extension, the cells resume their movement parallel to the direction of the longitudinal axis of the cell bodies. During the middle stage of PI, granule cells repeatedly extend and withdraw short leading processes, and move at a fast rate only after the process fully extends. The extension of a new leading process toward a different direction is an essential prerequisite for changing the direction of cell movement. Near the end of PI, granule cells start to develop a new mode of turning behavior; the tip of the leading process turns in a new direction and then the cell body follows the changes. Granule cells exhibit a dynamic cycle of cell advancement and stationary phase every 3 h; the active cell migration lasts for 2 h, and the stationary period is 1 h in length. Phase II (PII, a Period of 20–40 h In Vitro) During the early stage of PII, granule cells develop another mode of turning as follows: (1) the tip of the leading process bifurcates, (2) both branches extend in the opposite direction, (3) one of the branches collapses and retracts, and (4) the cell body follows the direction of extension of the remaining branch (Yacubova and Komuro 2002a). Granule cells exhibit this mode of turning behavior throughout PII. During the late stage of PII, granule cells become stationary for 2–3 h and retract their processes. Phase III (PIII, a Period of 40–60 h In Vitro) During the early stage of PIII, granule cells start to exhibit the initial signs of termination of their migration, which is a morphological change of the leading process (Yacubova and Komuro 2002a). During the late stage of PIII, granule cells slow down their movement, and slightly increase their turnings. At the end of PIII, the cells become permanently stationary, extend a lamellipodium around the soma, and emit several thin processes. The majority of granule cells terminate their migration 50–60 h after the initiation of their movement. Time-Dependent Changes in Granule Cell Migration and Their Morphology by Intrinsic Programs There are distinct relationships between the migratory behavior of granule cells, their morphology, and the elapsed time in vitro (Yacubova and Komuro 2002a). In PI, granule cells migrate at an average rate of 26.0 mm/h and exhibit the highest rate of turning behavior (1.3 turns/h), when the cells have multiple

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(3.7 processes/cell) and short (20.8 mm) processes. The length of the cycle of cell movement and stationary state is shortest (218 min). In PII, granule cells extend a long and thick leading process-like process (55.6 mm), and exhibit an elongated cycle (244 min) of cell movement and stationary state. The rate of cell movement is fastest (33.1 mm/h), while the amount of turning is lowest (0.3 turn/h). In PIII, granule cells slow down their movement (25.2 mm/h), but slightly increase their turning amount (0.5 turn/h). The length of the cycle of cell movement further increases to 297 min. These results indicate the existence of intrinsic (inherent) programs for controlling granule cell migration in a developmental stage-dependent manner.

Possible Roles of Intrinsic Programs in the Regulation of Granule Cell Migration In Vivo Although the question of whether and how intrinsic programs regulate granule cell migration in vivo remains to be determined, the comparison between in vivo and in vitro migration suggests possible roles of intrinsic programs in granule cell migration in vivo (Yacubova and Komuro 2002a; Komuro and Yacubova 2003). First, in PII (20–40 h in vitro), granule cells have two long processes and move at the fastest rate, while in the ML (25–35 h after the initiation of migration) granule cells have a long leading process and a trailing process and move at an increased rate. This similarity suggests that granule cell migration in the ML is partially regulated by intrinsic programs. Second, the 2–3 h stationary state of granule cells at the late stage of PII suggests that the prolonged stationary state (an average of 115 min) of granule cells in the PLC (35–40 h after the initiation of migration) is controlled by intrinsic signals. Third, in PIII (40–60 h in vitro), granule cells terminate their migration without cell–cell contact and start to express the a6 subunit of GABAA receptors, which are expressed only when the cells arrive in the IGL. This suggests that granule cells in PIII are in a similar stage of differentiation with those in the IGL (40–50 h after the initiation of migration). The time schedule for completion of granule cell migration in vitro is quite similar to that for granule cell migration in vivo. This similarity suggests that an internal program (or clock) may be involved in determining the length of granule cell migration.

Extracellular Glutamate Accelerates Granule Cell Migration in the ML Through the Activation of NMDA Receptors The role of extracellular glutamate in the regulation of granule cell migration was discovered in the early 1990s (Komuro and Rakic 1993, 1998b; Rakic and Komuro 1995). The first evidence came from the study examining whether inhibition of glutamate receptors affects the rate of granule cell migration. Blocking NMDAtype glutamate receptors with D-AP5 or MK-801 significantly decreases the rate of granule cell migration in the ML, while blocking kainate- and AMPA-type glutamate receptors with CNQX does not alter the rate of migration in the ML (Komuro and Rakic 1993). The role of the NMDA receptors in granule cell migration is

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further supported by evidence that changes in Mg2+ or glycine concentration affect the rate of granule cell movement. Because extracellular Mg2+ blocks NMDA receptor activity in a voltage-dependent manner and application of glycine potentiates NMDA receptor activity, it is expected that they both would influence granule cell migration. Indeed, the removal of Mg2+ from the medium significantly increases the rate of granule cell migration in the ML, whereas the rate of migration is reduced in a high Mg2+ medium in the ML. Likewise, application of 10 mM glycine significantly increases the rate of granule cell migration in the ML. These results demonstrate that the speed of granule cell migration in the ML is highly sensitive to small fluctuations in extracellular Mg2+ and glycine levels, implying that the activity of NMDA receptors modulates the rate of granule cell migration in the ML. The presence of spontaneously active NMDA receptors on the surface of migrating granule cells has been confirmed by patch-clamp analysis (Komuro and Rakic 1998b). The frequency of spontaneous NMDA receptor-coupled channel activity is low in the EGL, with large increases recorded in migrating neurons of the ML. Furthermore, single-channel recordings reveal developmentally related changes in the biophysical properties of the NMDA receptors during the course of granule cell differentiation, suggesting that migrating granule cells express one or more specific receptor subunits that are distinct from those comprising the receptors present in mature granule cells. It has been shown that migrating granule cells co-express the NR1 and NR2A or NR2B subunits, whereas postmigratory cells in the IGL express the NR1 and NR2C types (Komuro and Rakic 1998b). This progressive alteration in subunit composition could account for a change in NMDA receptor function during development. Moreover, there is evidence that the sensitivity of the NMDA receptors on granule cells in the EGL to glutamate increases during the course of cerebellar development. This increase can account for the acceleration of granule cell migration during the late stages of cerebellar development. The question of how NMDA receptors of migrating granule cells could be activated is intriguing, because the cells do not form synapses before the completion of their translocation in the IGL. One possibility is that endogenous extracellular glutamate may activate the immature form of the NMDA receptor by nonsynaptic means. Interestingly, the elevation of extracellular glutamate concentrations by inhibiting glutamate uptake by astrocytes increases the frequency of spontaneous NMDA receptor-coupled channel activity, and significantly accelerates the rate of granule cell movement in the ML (Komuro and Rakic 1993). These results suggest that endogenous extracellular glutamate is an important signal for the activation of NMDA receptors and that the increase of extracellular glutamate levels could enhance the rate of granule cell migration until the concentration reaches a toxic level. Several possible sources alone or in combination could be sufficient to activate NMDA receptors in migrating granule cells. It has been shown that glutamate released from cultured astrocytes in response to local signals causes an activation of NMDA receptor in neighboring neurons (Komuro and Rakic 1998b). Therefore, it is possible that Bergmann glial cells, which belong to the astrocyte cell class,

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release glutamate in response to local signals and influence the rate of granule cell migration by regulating the activity of NMDA receptors. The intimate association and large mutually shared surface area that exists between Bergmann glial processes and adjacent migrating granule cells could considerably facilitate this process. Furthermore, parallel fibers (the axons of postmigratory granule cells) are the most obvious source of extracellular glutamate in the ML. Because migrating granule cells do not form synapses with other cells, any glutamate released by parallel fibers must activate the NMDA receptor of migrating granule cells in a paracrine manner. Indeed, nonsynaptic activation of the NMDA receptor by extracellular glutamate has been observed in migrating granule cells.

Reciprocal Regulation of Granule Cell Migration in the EGL and the IGL by Somatostatin The role of somatostatin, a neuropeptide, in the control of granule cell migration was discovered in the early 2000s (Yacubova and Komuro 2002b). Somatostatin has two bioactive products, somatostatin-14 (SST-14) and somatostatin-28 (SST-28), which is a congener of SST-14 extended at the N-terminus. Five somatostatin receptors (SSTRs) have been cloned and named SSTR1–5 according to their order of identification. Both SST-14 and SST-28 bind to all five somatostatin receptors. It has been expected that SST may play a critical role in neurogenesis or neural differentiation, because numerous brain regions, including the cerebral cortex and cerebellum, exhibit high levels of SST and its receptor early in development. Postmitotic granule cells express all five types of SSTRs before the initiation of migration, while differentiated granule cells in the adult do not express the receptors (Yacubova and Komuro 2003). High levels of SST are present along the migratory route of granule cells and in their final destination (Yacubova and Komuro 2002b). During periods of granule cell migration, SST-14 is present in Purkinje cells, Golgi cells, and climbing fibers, while SST-28 is present in Golgi cells and mossy fiber terminals. The use of cerebellar slices obtained from P10 mice revealed that the addition of exogenous SST-14 or SST-28 to the medium significantly increases the rate of granule cell migration in the EGL, slightly decreases the rate in the ML, and significantly decreases the rate in the IGL (Yacubova and Komuro 2002b). In contrast, the addition of an SST antagonist (AC-178,335) to the medium significantly decreases the rate of granule cell migration in the EGL, slightly increases the rate in the ML, and significantly increases the rate in the IGL (Yacubova and Komuro 2002b). These results indicate that endogenous SST differentially controls granule cell migration in the EGL, the ML, and the IGL. SST accelerates the tangential migration of granule cells near their birthplace within the EGL, but slows down the radial migration near their final destination within the IGL. The next question is whether SST acts directly on migrating granule cells or acts on other cells, which then indirectly influence granule cell migration. The use of

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microexplant cultures of P0-P2 mouse cerebella demonstrates that the application of exogenous SST-14 or SST-28 significantly increases the rate of granule cell movement at 1 day in vitro, while exogenous SST-14 or SST-28 substantially decreases the rate at 2 days in vitro, suggesting that SST directly acts on migrating granule cells in a developmental stage-specific manner (Yacubova and Komuro 2002b). The use of Ca2+ indicator dyes reveals that SST-14 increases the size and frequency of Ca2+ spikes of granule cells at 1 day in vitro, whereas SST-14 eliminates Ca2+ spikes at 2 days in vitro (Yacubova and Komuro 2002b). An increase in the rate of granule cell movement follows the enlargement of Ca2+ spikes at 1 day in vitro, while the elimination of Ca2+ spikes at 2 days in vitro decreases the rate. These results demonstrate that the differential effects of SST at 1 day in vitro and 2 days in vitro on Ca2+ spikes might explain how SST switches its effect on granule cell migration from acceleration at the early phase of migration to deceleration at the late phase of migration.

Halt of Granule Cell Migration in the PCL by PACAP The role of pituitary adenylate cyclase-activating polypeptide (PACAP) in the control of granule cell migration was discovered in the mid-2000s (Cameron et al. 2007). PACAP, a member of the secretin/glucagon/vasoactive intestinal polypeptide family, is known to control physiological functions of a wide range of cells (Botia et al. 2007; Vaudry et al. 2009). PACAP has two bioactive products, PACAP38 and PACAP27. PACAP27 is the N-terminal 27-amino acid sequence of PACAP38. There are three types of PACAP receptors (PAC1, VPAC1, and VPAC2), which belong to the class-B G protein-coupled receptor superfamily (Vaudry et al. 2009). There is a unique pattern of endogenous PACAP expression in the developing cerebellum: PACAP is present sporadically in the bottom of the ML, expressed intensively in the PCL, and dispersedly throughout the IGL (Cameron et al. 2007, 2009). PACAP is expressed by Purkinje cell dendrites in the ML, Purkinje cell somata in the PCL, and mossy fiber terminals in the IGL. Therefore, endogenous PACAP is highly expressed in the route of granule cell migration. Granule cell precursors and postmitotic granule cells are devoid of PACAP, but they express high levels of PAC1 receptors prior to the initiation of their migration (Cameron et al. 2007). After the completion of migration, granule cells start to lose PAC1 receptors. Granule cells and their precursors in the EGL also express VPAC1 receptors at a much lower level than the PAC1 receptors, but do not express VPAC2 receptors. The use of microexplant cultures of early postnatal mouse cerebellum demonstrates that application of exogenous PACAP38 reduces the rate of granule cell migration, while application of its antagonist (PACAP6-38) does not affect the rate (Cameron et al. 2007). Furthermore, the use of cerebellar tissue slices obtained from the early postnatal mouse also demonstrates that the application of exogenous PACAP38 slows down the radial migration of granule cells in the ML. Collectively, these results indicate that PACAP acts on granule cell migration as a “brake” (stop signal) for cell movement.

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The use of cerebellar tissue slices reveals that the effect of exogenous PACAP on granule cell migration varies within each cortical layer (Cameron et al. 2007). The application of exogenous PACAP38 significantly reduces granule cell motility in the EGL and ML, but fails to alter their movement in the PCL and the IGL. How does this happen? The answer lies in the differing expression of endogenous PACAP in different cortical layers. The use of PACAP6-38 indicates the role of endogenous PACAP38 in granule cell migration. The application of PACAP6-38 significantly increases granule cell motility in the PCL, but does not alter motility in the EGL, ML, and IGL (Cameron et al. 2007), suggesting that the reduction of the rate of granule cell migration in the PCL is caused by endogenous PACAP release. At the top of the IGL, where high levels of endogenous PACAP are present, granule cells migrate at a speed comparable to that observed in the ML. Furthermore, the application of exogenous PACAP or its antagonist does not significantly alter granule cell migration in the IGL. How does this occur? Although the mechanisms underlying the mysterious effects of PACAP on granule cell migration in the IGL remain to be determined, there is a possible explanation. For example, before entering the IGL, granule cells may lose their response to PACAP38 via the desensitization of PACAP receptors. This occurs because PACAP receptors undergo a rapid desensitization after an initial activation, as seen in other G protein-coupled receptors (Vaudry et al. 2009). Although the continuous application of PACAP first reduces the rate of granule cell migration, the cells gradually recover their motility even in the presence of PACAP (Cameron et al. 2007). The average time required for returning the motility of granule cells to control levels under continuous exposure to PACAP is 2.1 h, which is similar to the stationary period of the cells observed in the PCL (1.9 h). Furthermore, the recovery from the PACAP-induced reduction of granule cell motility is delayed by inhibiting protein kinase C (PKC). This provides additional evidence that PACAP receptors undergo desensitization in the PCL and IGL, because the G protein-coupled receptor kinases, which mediate the desensitization of PACAP receptors, are sensitive to changes in the activity of PKC (Vaudry et al. 2009). These results indicate that after an initial response to endogenous PACAP in the PCL, PACAP receptors on granule cells undergo desensitization, which allows the cells to actively migrate within the endogenous PACAP-rich IGL.

Ca2+ Spikes Control Granule Cell Migration and Its Termination In the early 1990s, the combined use of cerebellar slices of the early postnatal mice and pharmacological tools revealed the role of voltage-gated Ca2+ channels, especially the N-type Ca2+ channel, in granule cell migration (Komuro and Rakic 1992). Postmitotic granule cells in the EGL start to express N-type Ca2+ channels prior to the initiation of their migration. The number of N-type Ca2+ channels on the plasmalemmal surface of granule cells rapidly increases during their migration to the IGL. The blockade of N-type Ca2+ channel activity by a specific antagonist significantly reduces the speed of granule cell migration in the ML, suggesting that

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the N-type Ca2+ channels play a key role in controlling the speed of granule cell migration in the ML (Komuro and Rakic 1992; Rakic and Komuro 1995). In the mid-1990s, the role of intracellular Ca2+ levels in granule cell migration was examined because the activation of N-type Ca2+ channels induces the elevation of intracellular Ca2+ levels by increasing the Ca2+ influx. The use of Ca2+ indicator dyes reveals that migrating granule cells exhibit dynamic changes in intracellular Ca2+ levels of their somata (Komuro and Rakic 1996). In microexplant cultures of the early postnatal mouse cerebellum, Ca2+ spikes in the granule cell somata occur 4–24 times per hour, with average frequencies of 13/h. There is a positive correlation between the speed of granule cell migration and both the amplitude and frequency components of Ca2+ spikes (Komuro and Rakic 1996). The experimental reduction of the Ca2+ influx by lowering extracellular Ca2+ concentrations or blocking Ca2+ channels results in a decrease in the amplitude and frequency of Ca2+ spikes in the granule cell somata. This reduction is linearly related to the speed of granule cell movement (Komuro and Rakic 1996). These results suggest that the migration of granule cells in vitro is controlled by the combination of the amplitude and frequency of Ca2+ spikes. During the mid-2000s, the combined use of cerebellar slices of the early postnatal mouse and Ca2+ indicator dye (Oregon Green 488 BAPTA-1) demonstrated that granule cells exhibit a distinct pattern of Ca2+ spikes as they migrate in different cortical layers (Kumada and Komuro 2004; Komuro and Kumada 2005). The changes in the frequency of Ca2+ spikes in the granule cell somata in each cortical layer are as follows:

EGL At the top of the EGL, granule cell precursors exhibit Ca2+ spikes in their somata with a low frequency (average frequency, 8.3/h). The intervals of occurrences are regular and the amplitude is uniform. Concomitant with the initiation of tangential migration at the middle of the EGL, postmitotic granule cells significantly increase the frequency of Ca2+ spikes (20.9/h). The Ca2+ spikes gradually decrease in number at the bottom of the EGL (15.9/h) and the EGL-ML border (12.8/h), and the rhythm becomes irregular, containing short, silent periods. ML Once granule cells enter the ML, the cells slightly increase the number of Ca2+ spikes (15.1/h at the top of the ML, and 17.2/h at the middle). However, at the bottom of the ML, the frequency of Ca2+ spikes gradually decreases to 12.2/h and the amplitudes of Ca2+ spikes become variable. PCL Upon entering the PCL, granule cells significantly reduce the frequency of Ca2+ spikes with long, silent periods, and also decrease the amplitude of Ca2+ spikes. The average frequencies of Ca2+ spikes are 7.3/h at the top of the PCL and 6.9/h at the bottom.

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IGL At the top of the IGL, granule cells significantly increase the frequency of Ca2+ spikes (15.1/h), although the rhythms are irregular and the amplitudes are variable. As granule cells migrate through the middle of the IGL, the frequency of Ca2+ spikes gradually decreases to 9.3/h and the amplitude becomes smaller. At the bottom of the IGL, the Ca2+ spikes disappear, or significantly decrease in frequency (2.4/h). The frequency of Ca2+ spikes in the granule cell somata dynamically changes along the migratory pathway and positively correlates with the rate of cell migration (correlation coefficient, 0.85) (Kumada and Komuro 2004). Granule cells reduce the frequency of Ca2+ spikes and the rate of cell movement at each boundary between cortical layers. These results suggest that the frequency of Ca2+ spikes is one of the factors that control the alterations of granule cell migration in a corticallayer dependent manner. Next, the question of whether the changes in Ca2+ transients directly affect granule cell migration is examined by experimental modifications of the frequency of Ca2+ spikes. Reducing Ca2+ influx and decreasing internal Ca2+ release result in a significant reduction of the frequency of Ca2+ spikes in the granule cell somata and a slowdown of their migration at the top of the IGL (Kumada and Komuro 2004). Furthermore, inhibiting Ca2+ signaling-upstream by blocking phospholipase C (PLC) also decreases the frequency of Ca2+ spikes and slows down granule cell movement. Likewise, the inhibition of Ca2+ signaling-downstream by blocking protein kinase C (PKC) and Ca2+/calmodulin results in a significant reduction of Ca2+ spike frequency and the migration rate (Kumada and Komuro 2004). These results demonstrate that the reduction of Ca2+ spike frequency in the granule cell somata is always accompanied by the slowdown of cell movement, implying that the Ca2+ spike frequency provides an intracellular signal for controlling the rate of granule cell migration. At their final destination within the IGL, granule cells lose Ca2+ spikes, or significantly reduce the frequency. The loss of Ca2+ spikes is not caused by physiological deterioration after a prolonged period of observation, because 2–5 h later postmigratory granule cells resume spontaneous Ca2+ spikes. Time-lapse observation of intracellular Ca2+ levels and cell movement reveals the sequence of the loss of Ca2+ spikes and the completion of granule cell migration (Kumada and Komuro 2004; Komuro and Kumada 2005). At the bottom of the IGL, granule cells initially migrate with variable amplitudes of Ca2+spikes, but completely lose Ca2+ spikes before becoming permanently stationary. The average time lag between the loss of Ca2+ spikes and the cessation of migration is 16.8 min with a range of 5–27 min. These results suggest that the loss of Ca2+ spikes is a prerequisite for completing granule cell migration at their final destination. The role of Ca2+ spike loss in the completion of granule cell migration was determined by experimentally altering Ca2+ spike frequency at their final destination (the bottom of the IGL). Inhibiting the Ca2+ influx or reducing internal Ca2+ release results in a significant reduction of the Ca2+ spike frequency and a slowdown of granule cell movement (Kumada and Komuro 2004). On the other hand, stimulating internal Ca2+ release

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significantly increases the Ca2+ spike frequency and accelerates cell movement (Kumada and Komuro 2004). Furthermore, inhibiting the activity of PLC, PKC, and Ca2+/calmodulin significantly decreases the Ca2+ spike frequency and slows down cell movement. These results indicate that the loss of Ca2+ transients may trigger molecular cascades leading to the completion of granule cell migration at their final destination. The effects of the stimulation of internal Ca2+ release on granule cell migration suggest that the basic levels (or amount) of spontaneous Ca2+ release from the internal stores may be reduced at the bottom of the IGL. Such reduction of internal Ca2+ release may be responsible for the slowdown (or termination) of granule cell migration at their final destination. The loss of Ca2+ spikes may be induced by external stop signals or contact with other cells and processes, but intrinsic programs may also be responsible. The use of microexplant cultures of early postnatal mouse cerebella reveals that during a period of active cell movement, granule cells exhibit spontaneous Ca2+ spikes in their somata, and the Ca2+ spike frequency depends on the elapsed time after plating (Kumada and Komuro 2004). There is a positive correlation between the Ca2+ spike frequency and the migration rate (correlation coefficient, 0.81). Also, the Ca2+ spikes disappear or significantly reduce their occurrences when granule cells stop migrating at 50–60 h in vitro, although 1–3 h later the postmigratory granule cells resume generating the Ca2+ spikes (Kumada and Komuro 2004). The loss of Ca2+ spikes always precedes the completion of migration. The average time lag between the loss of Ca2+ transients and the cessation of migration is 11.6 min with a range of 3–21 min. The reduction of the Ca2+ spike frequency induced by the use of pharmacological tools is always accompanied by the slowdown of granule cell movement. The increasing of Ca2+ spike frequency by stimulating internal Ca2+ release significantly accelerates granule cell migration at the final phase of migration (50–60 h in vitro), leading to a delay in the completion of migration. These results suggest that intrinsic programs may set the timing of the loss of Ca2+ spikes in granule cells at approximately 50–60 h in vitro, and may trigger the completion of migration. It is not well understood how Ca2+ spikes control granule cell motility, but there are several possible scenarios (Kumada and Komuro 2004; Komuro and Kumada 2005). One possibility is that Ca2+ spikes regulate the dynamic assembly and disassembly of cytoskeletal elements required for the operation of a forcegenerating mechanism involved in granule cell movement. Furthermore, Ca2+ spikes may modulate the repetitive formation and elimination of binding sites between granule cells and their migratory substrates. Ca2+ spikes may control conformational changes of cell adhesion molecules, such as integrins, which are expressed on the plasma membrane of granule cells. At present, little is known about how the loss of Ca2+ spikes induces the cessation of granule cell migration. One possibility is that the loss of Ca2+ spikes might cause changes in the Ca2+dependent activation of specific enzymes, which in turn affects the phosphorylation state or extent of proteolysis of large numbers of proteins. These changes could induce the rearrangement of cytoskeletal components that are required for the completion of granule cell migration.

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Impairment of Granule Cell Migration by Alcohol Alcohol is presently the most common chemical teratogen causing malformation and mental deficiency in humans (Kumada et al. 2007, 2010; Jiang et al. 2008). Prolonged exposure to alcohol during gestation and lactation correlates with a pattern of abnormal development in newborns. This developmental disturbance is known as “fetal alcohol spectrum disorder” (FASD). Children with FASD often show neurological signs associated with cerebellar damage such as delayed motor development, problems with fine tasks, and ataxia. Multiple aspects of central nervous system development can be affected by alcohol. Among them, striking abnormalities appear to involve the impairment of neuronal migration. However, until recently, little was known about how alcohol adversely affects neuronal cell migration. Real-time observation of cell movement in cerebellar slices of early postnatal mice demonstrates that application of ethanol immediately slows granule cell migration in a dose-dependent manner (Kumada et al. 2006). Application of 10 mM ethanol (equivalent to blood ethanol level Purkinje cell synapse stabilization during cerebellar development of mutant mice lacking the glutamate receptor delta2 subunit. J Neurosci 17:9613–9623 Lalouette A, Lohof A, Sotelo C et al (2001) Neurobiological effects of a null mutation depend on genetic context: Comparison between two hotfoot alleles of the delta-2 ionotropic glutamate receptor. Neuroscience 105:443–455 Letellier M, Bailly Y, Demais V et al (2007) Reinnervation of late postnatal Purkinje cells by climbing fibers: neosynaptogenesis without transient multi-innervation. J Neurosci 27:5373–5383 Letellier M, Wehrle R, Mariani J et al (2009) Synapse elimination in olivo-cerebellar explants occurs during a critical period and leaves an indelible trace in Purkinje cells. Proc Natl Acad Sci USA 106:14102–14107 Lichtman JW, Colman H (2000) Synapse elimination and indelible memory. Neuron 25:269–278 Lohof AM, Mariani J, Sherrard RM (2005) Afferent-target interactions during olivocerebellar development: Transcommissural reinnervation indicates interdependence of Purkinje cell maturation and climbing fibre synapse elimination. Eur J Neurosci 22:2681–2688 Lomeli H, Sprengel R, Laurie DJ et al (1993) The rat delta-1 and delta-2 subunits extend the excitatory amino acid receptor family. FEBS Lett 315:318–322 Mariani J (1982) Extent of multiple innervation of Purkinje cells by climbing fibers in the olivocerebellar system of weaver, reeler, and staggerer mutant mice. J Neurobiol 13:119–126 Mariani J (1983) Elimination of synapses during the development of the central nervous system. Prog Brain Res 58:383–392 Mariani J, Changeux JP (1981) Ontogenesis of olivocerebellar relationships. I. Studies by intracellular recordings of the multiple innervation of Purkinje cells by climbing fibers in the developing rat cerebellum. J Neurosci 1:696–702 Mariani J, Benoit P, Hoang MD et al (1990) Extent of multiple innervation of cerebellar Purkinje cells by climbing fibers in adult X-irradiated rats. Comparison of different schedules of irradiation during the first postnatal week. Brain Res Dev Brain Res 57:63–70 Mason CA, Christakos S, Catalano SM (1990) Early climbing fiber interactions with Purkinje cells in the postnatal mouse cerebellum. J Comp Neurol 297:77–90

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Sherrard RM, Dixon KJ, Bakouche J et al (2009) Differential expression of TrkB isoforms switches climbing fiber-Purkinje cell synaptogenesis to selective synapse elimination. Dev Neurobiol 69:647–662 Sotelo C, Hillman DE, Zamora AJ et al (1975) Climbing fiber deafferentation: Its action on Purkinje cell dendritic spines. Brain Res 98:574–581 Sugihara I (2005) Microzonal projection and climbing fiber remodeling in single olivocerebellar axons of newborn rats at postnatal days 4–7. J Comp Neurol 487:93–106 Sugihara I, Wu HS, Shinoda Y (2001) The entire trajectories of single olivocerebellar axons in the cerebellar cortex and their contribution to cerebellar compartmentalization. J Neurosci 21:7715–7723 Sugihara I, Lohof AM, Letellier M et al (2003) Post-lesion transcommissural growth of olivary climbing fibres creates functional synaptic microzones. Eur J Neurosci 18:3027–3036 Takeuchi T, Miyazaki T, Watanabe M et al (2005) Control of synaptic connection by glutamate receptor delta2 in the adult cerebellum. J Neurosci 25:2146–2156 Tempia F, Bravin M, Strata P (1996) Postsynaptic currents and short-term synaptic plasticity in Purkinje cells grafted onto an uninjured adult cerebellar cortex. Eur J Neurosci 8:2690–2701 Tohgo A, Eiraku M, Miyazaki T et al (2006) Impaired cerebellar functions in mutant mice lacking DNER. Mol Cell Neurosci 31:326–333 Uemura T, Kakizawa S, Yamasaki M et al (2007) Regulation of long-term depression and climbing fiber territory by glutamate receptor delta2 at parallel fiber synapses through its C-terminal domain in cerebellar Purkinje cells. J Neurosci 27:12096–12108 Usowicz MM, Sugimori M, Cherksey B et al (1992) P-type calcium channels in the somata and dendrites of adult cerebellar Purkinje cells. Neuron 9:1185–1199 Voneida TJ, Christie D, Bogdanski R et al (1990) Changes in instrumentally and classically conditioned limb-flexion responses following inferior olivary lesions and olivocerebellar tractotomy in the cat. J Neurosci 10:3583–3593 Watanabe F, Miyazaki T, Takeuchi T et al (2008) Effects of FAK ablation on cerebellar foliation, Bergmann glia positioning and climbing fiber territory on Purkinje cells. Eur J Neurosci 27:836–854 Willson ML, Bower AJ, Sherrard RM (2007) Developmental neural plasticity and its cognitive benefits: Olivocerebellar reinnervation compensates for spatial function in the cerebellum. Eur J Neurosci 25:1475–1483 Willson ML, McElnea C, Mariani J et al (2008) BDNF increases homotypic olivocerebellar reinnervation and associated fine motor and cognitive skill. Brain 131:1099–1112 Zagrebelsky M, Rossi F, Hawkes R et al (1996) Topographically organized climbing fibre sprouting in the adult rat cerebellum. Eur J Neurosci 8:1051–1054 Zagrebelsky M, Strata P, Hawkes R et al (1997) Reestablishment of the olivocerebellar projection map by compensatory transcommissural reinnervation following unilateral transection of the inferior cerebellar peduncle in the newborn rat. J Comp Neurol 379:283–299 Zuo J, De Jager PL, Takahashi KA et al (1997) Neurodegeneration in Lurcher mice caused by mutation in delta2 glutamate receptor gene. Nature 388:769–773

Synaptogenesis and Synapse Elimination

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Masanobu Kano and Masahiko Watanabe

Abstract

Formation of excess synaptic connections at perinatal stage and subsequent elimination of redundant synapses and strengthening of the surviving ones are crucial steps for functional neural circuit formation in developing nervous system. Shortly after birth, excitatory synapses are formed on the somata of Purkinje cells (PCs) from climbing fibers (CFs) that originate from neurons in the inferior olive of the contralateral medulla oblongata. At this developmental stage, each PC is innervated by multiple (around five) CFs with equal strengths. Subsequently, a single CF is selectively strengthened relative to other CFs during the first postnatal week. Then, around postnatal day 9 (P9), only the strongest CF (“winner” CF) starts to extend its innervation to PC dendrites. On the other hand, synapses of the weaker CFs (“loser” CFs) remain on the soma and the most proximal portion of the dendrite, and they are eliminated progressively during the second and the third postnatal weeks. From P6 to P11, the elimination proceeds independently of the formation of the synapses on PC dendrites from parallel fibers (PFs), the other excitatory inputs to PCs. From P12 and thereafter, the elimination of weaker CFs requires normal PF-PC synapse formation and is presumably dependent on the PF synaptic inputs that activate type 1 metabotropic glutamate receptor (mGluR1) and its downstream signaling in PCs. Most PCs become mono-innervated by single CFs in the third

M. Kano (*) Department of Neurophysiology, Graduate School of Medicine, The University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo, 113-0033, Japan e-mail: [email protected] M. Watanabe Department of Anatomy, Hokkaido University Graduate School of Medicine, Sapporo, 060-8638, Japan e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 281 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_14, # Springer Science+Business Media Dordrecht 2013

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postnatal week. This chapter integrates the current knowledge of synaptogenesis and subsequent synapse elimination at CF to PC connections during postnatal cerebellar development.

Introduction In the process of neural circuit formation during postnatal development, supernumerary synapses are formed transiently around birth and then functionally important synapses are strengthened while unnecessary synapses are weakened and eventually eliminated. This process is known as “synapse elimination” and is widely thought to be an important mechanism to form functional neural circuits. Synapse elimination has been studied extensively in the neuromuscular junction and autonomic ganglia (Gan et al. 2003; Walsh and Lichtman 2003). However, it is difficult to perform such detailed analyses in the central nervous system (CNS), because of small size of synapse, heterogeneity and abundance of synaptic inputs to each neuron, and complexity of synaptic organization. In this respect, the climbing fiber (CF) to Purkinje cell (PC) synapse in the cerebellum is an exceptional case and provides an excellent model to study synapse elimination in the CNS (Crepel 1982; Lohof et al. 1996; Hashimoto and Kano 2005; Kano and Hashimoto 2009). PCs in the adult cerebellum receive two major excitatory inputs, namely parallel fibers (PFs) and CFs (Palay and Chan-Palay 1974; Ito 1984). PFs are bifurcated axons of cerebellar granule cells (GCs) and form synapses on spines of PC’s distal dendrites. Each synaptic input is weak but as many as 100,000 PFs make contacts on dendritic spines of a single PC (Palay and Chan-Palay 1974; Ito 1984). In contrast, the majority of PCs in the adult cerebellum are innervated by single CFs (mono-innervation) but each CF makes strong synaptic contacts on PC’s proximal dendrites (Ito 1984). In early postnatal days, however, all PCs are innervated by multiple CFs (multiple innervation) (Crepel 1982; Lohof et al. 1996; Hashimoto and Kano 2005; Kano and Hashimoto 2009). These surplus CFs are eliminated eventually and mono-innervation is attained in the third postnatal week (Kano et al. 1995, 1997, 1998; Offermanns et al. 1997). The chapter describes how CF synapses are formed, single CFs are selected and their synapses are strengthened, and synapses of redundant CFs are eliminated during postnatal cerebellar development.

Climbing Fiber Synaptogenesis on Immature Purkinje Cells Axons from inferior olivary neurons reach the primitive cerebellum around E18 (Wassef et al. 1992). They ramify and give rise to thick and thin collaterals. This stage is called the “creeper stage” (Chedotal and Sotelo 1993) at which the thick collaterals, the “climbing fibers” creep between the PCs. At this stage, PCs have bipolar shapes [called “simple- and complex-fusiform cells” (Armengol and Sotelo 1991)] and have just completed their migration and are organized in a multilayer. Initially, each olivocerebellar axon forms about 100 “creeper” CFs (Sugihara

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2005). Then, the three stages of CF to PC synapse formation follow, which are described by Ramo´n y Cajal in his pioneering studies (1911): the “pericellular nest” stage, the “capuchon” stage, and the “dendritic stage.” At the “pericellular nest” stage, CFs surround the cell bodies of PCs which undergo explosive outgrowth of perisomatic protrusions in all directions from the cell bodies [called the phase of “stellate cells” (Armengol and Sotelo 1991)]. CFs establish contacts with the abundant pseudopodia stemming from the soma and form a plexus on the lower part of the PC somata. Among the 100 “creeper” CFs of each olivocerebellar axon, only around 10 can develop “pericellular nests.” The “capuchon” stage is characterized by the displacement of the plexus to the apical portion of PC somata and main dendrites. Then, the “dendritic” stage follows which is characterized by the upward spread of CF innervation into dendrites. Electrophysiological evidence indicates that functional CF synapses are formed on immature PCs around P3. In juvenile rats and mice in vivo, stimulation in the inferior olive after P3 elicits CF-mediated responses in PCs (Crepel 1971). However, the responses of juvenile PCs are graded in parallel with the increase in the stimulus strength (Crepel et al. 1976), which indicates that PCs are innervated by multiple CFs in juvenile rodent cerebellum. Later studies in vivo clarified that both the percentage of PCs innervated by multiple CFs and the average number of CFs innervating individual PCs decrease with postnatal development, and that most PCs become innervated by single CFs by the end of the third postnatal week (Crepel et al. 1981; Mariani and Changeux 1981). Thus, these earlier studies performed in mice and rats in vivo have established that elimination of redundant CF inputs occurs during postnatal development of the cerebellum as in the neuromuscular synapses in the periphery.

Functional Differentiation and Selective Strengthening of Single Climbing Fiber Inputs When recorded from PCs in cerebellar slices at P2-3, clearly discernible excitatory postsynaptic currents (EPSCs) can be elicited by stimulating CFs. EPSCs appear with discrete steps by gradually increasing stimulus strength, clearly indicating that PCs at this age are innervated by multiple CFs. The amplitudes of EPSCs elicited by stimulating multiply innervating CFs at this stage are much smaller than those evoked by mature CFs at later developmental stage (Hashimoto and Kano 2003; Hashimoto and Kano 2005; Scelfo and Strata 2005; Bosman et al. 2008; Ohtsuki and Hirano 2008). Therefore, CF inputs become stronger, while redundant CFs are eliminated during postnatal development. In anesthetized animals in vivo, Mariani and Changeux found that CF stimulation elicited in some PCs two CF-mediated responses whose amplitudes were quite different around P10 to P13 (Mariani and Changeux 1981). This result suggests that only one CF is strengthened relative to others before the completion of synapse elimination. Developmental changes in the synaptic strengths of multiple CFs innervating the same PC were systematically investigated in mice aged P2 to P21 using whole-cell

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Fig. 14.1 Postnatal development of CF-PC synapses (a) Diagrams of CF-PC synapses at four representative stages of postnatal development in mice. (b) Four distinct phases in postnatal development of CF synapses

voltage-clamp recordings from PCs in cerebellar slices (Hashimoto and Kano 2003). To search CFs innervating the PC under recording, the stimulation pipette was moved systematically and the stimulus strength was increased gradually at each stimulation site. The number of CFs innervating each PC was judged by the number of discrete CF-EPSCs, and the strengths of individual CF inputs were estimated from the sizes of CF-EPSCs. This quantitative study shows that more than five discrete CF-EPSCs with similar amplitudes are present in PCs from mice around P3 (Fig. 14.1a, P3). In contrast, in the second postnatal week, PCs with multiple CF innervation have one large CF-EPSC and a few small CF-EPSCs (Fig. 14.1a, P7 and P12). These results suggest that synaptic strengths of multiply innervating CFs are relatively uniform in neonatal mice, and one CF is selectively strengthened during postnatal development (Hashimoto and Kano 2003; Hashimoto and Kano 2005; Bosman et al. 2008; Ohtsuki and Hirano 2008). Quantitative assessments of the disparity among the synaptic strengths (i.e., amplitudes of multiple CF-EPSCs) in individual PCs demonstrate that the disparity progressively increases from P3 to P6 and reaches a stable level at P7. This result indicates that one CF is selectively strengthened among multiple CFs innervating the same PC during the first postnatal week (Fig. 14.1b, (1) Functional differentiation) (Hashimoto and Kano 2003). Meanwhile, it was shown that the innervation pattern of CFs over PCs drastically changes during the first postnatal week in rats (Sugihara 2005), which is consistent with the electrophysiological data in mice (Hashimoto and Kano 2003). At P4 in rat, CFs have many creeping terminals in the PC layer and their swellings do not aggregate at particular PC somata

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(creeper type). Then, from P4 to P7, CFs surround several specific PC somata and form aggregated terminals on them (nest type) (Sugihara 2005). There are clear difference in electrophysiological properties between EPSCs elicited by the strongest CF input and those by other weaker inputs. The sizes of glutamate transient in synaptic cleft in response to CF stimulation can be estimated by using nonequilibrium inhibition of AMPA receptors by a low-affinity competitive antagonist (Clements 1996). These values are significantly larger for the strongest CF than for the weaker CFs (Hashimoto and Kano 2003). However, in a low extracellular Ca2+ concentration in which CF-EPSCs result from one-site one vesicle release, the amplitudes of glutamate transients for the strongest and the weaker CFs are not different (Hashimoto and Kano 2003). This result indicates that the sizes of glutamate transients caused by single synaptic vesicles are the same between the two types of CF inputs. Therefore, the larger glutamate transients by stimulating the strongest CF are thought to result from the higher probability of multivesicular release. Further electrophysiological examination suggests that the release probability is not different between the two types of CF inputs. Therefore, it is thought that the number of release site facing a narrow postsynaptic region of PC is larger in the strongest CF than in weaker CFs.

Dendritic Translocation of Single Climbing Fibers Morphological evidence indicates that the sites of CF synapses on PCs change from soma to dendrite during early postnatal development, which is known as “climbing fiber translocation” (Altman and Bayer 1997). A recent study has clarified the relationship between the selective strengthening of single CFs and CF translocation by using electrophysiological and morphological techniques (Hashimoto et al. 2009a). The location of synapses along the somato-dendritic domains of PCs can be estimated by analyzing the kinetics of EPSCs arising from single synaptic vesicles (termed quantal EPSCs) in CF terminals. In a Sr2+-containing external solution, stimulation of presynaptic axons causes asynchronous release of synaptic vesicles. The quantal EPSCs arising from a stimulated CF can be recorded under this experimental condition (Hashimoto and Kano 2003; Hashimoto et al. 2009a). Under whole-cell recordings from the soma, quantal EPSCs originating from PC dendrites are strongly attenuated by dendritic filtering. Because CFs that have undergone translocation to PC dendrites have synapses with different electrotonic lengths from the somatic recording site, individual quantal EPSCs should undergo different degrees of distortion depending on the locations of CF synapses along PC dendrites. Since the rise time of quantal EPSCs is reported to be proportional to the distance from the synaptic sites to the soma (Roth and Hausser 2001), distribution of the rise of quantal EPSCs for a CF reflects the extent of dendritic translocation of that CF. At P7-P8 when the selective strengthening of single CF in each PC has just completed, there is no significant difference in the distribution of qEPSCs rise time for the strongest CF and for the weaker CFs. This result indicates that synapses of

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the strongest CF and weaker CFs are located on the soma at this developmental stage (Fig. 14.1a, P7). At P9-P10, the incidence of qEPSCs with slow rise time is more frequent for the strongest CF than for the weaker CFs, suggesting that CFs begin to expand their innervations territories to dendrites (Fig. 14.1b (3) CF translocation). The difference in the distribution of quantal EPSC rise times for the strongest CF and for the weaker CFs becomes larger from P11 to P14. While the incidence of quantal EPSCs with slow rise time become more frequent for the strongest CF with age, the quantal EPSC rise time for weaker CFs remain almost unchanged from P9 to P14. These electrophysiological data collectively indicate that: (1) synaptic competition among multiple CFs occurs on the soma before P7 (Fig. 14.1a, P3 and  P7, Fig. 14.1b (1) Functional differentiation), (2) only the strongest CF (“winner” CF) starts to translocate to dendrites at P9 and the translocation continues thereafter (Fig. 14.1a, P12, Fig. 14.1b (3) CF translocation), (3) synapses of the weaker CFs (“loser” CFs) remain around the soma (Fig. 14.1a, P12). Morphological data also support these notions of CF synapse development. When subsets of CFs are labeled by an anterograde tracer, BDA, injected into the inferior olive, pericellular nests with extensive branching of CFs are observed at P7, P9, and P12. At P7, in spite of the presence of immature stem dendrite in PCs, CF synapses are confined to the soma and are absent on the dendrites. CF synapses are first found on PC dendrites at P9. At P12 and thereafter, the territory of CF innervations extended progressively along the PC dendrites. It should be noted that the strongest CFs translocating to dendrites keep their synapses on PC somata until around P12. In contrast, synaptic terminals of the weaker CFs are confined to the soma and the basal part of the primary dendrite. These weaker CFs are thought to be collaterals of the strongest CFs innervating adjacent PCs. Thus, pericellular nests represent multiple CF innervations of PCs (Hashimoto et al. 2009a).

Early Phase of Climbing Fiber Synapse Elimination Earlier studies on spontaneous mutant mice (Crepel and Mariani 1976; Mariani et al. 1977; Crepel et al. 1980; Mariani and Changeux 1980) and animals with experimentally induced “hypogranular” cerebella (Woodward et al. 1974; Crepel and Delhaye-Bouchaud 1979; Bravin et al. 1995; Sugihara et al. 2000) have revealed that the presence of intact granule cells and normal formation of PF-PC synapses are prerequisite for CF synapse elimination. Crepel et al. (1981) showed that elimination of surplus CFs consists of two distinct phases, the early phase up to around P8 and the late phase from around P9 to P17 (Crepel et al. 1981). The early phase occurs normally in animals with “hypogranular” cerebella, whereas the late phase is severely impaired by inhibiting granule cell production, indicating that the early phase CF synapse elimination proceeds independently of PF-PC synapse formation whereas the late phase critically dependent on it. However, since the animal models with “hypogranular” or “agranular” cerebella have abnormalities of cerebellar development other than granule cell genesis and PF-PC synapse

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formation, there remains a possibility that CF synapse elimination might be influenced by such developmental defects. Detailed assessment of the postnatal development of CF innervations in mouse cerebellar slices demonstrates that there is no significant reduction but rather a tendency of increase in the average number of CFs per PC from P3 to P6 when functional differentiation of multiple CFs occurs (Hashimoto et al. 2009b). Then, the value decreased progressively from P6 to around P15 (Scelfo and Strata 2005; Hashimoto et al. 2009b). These results indicate that CF synapse elimination does not proceed in parallel with functional differentiation of multiple CFs but starts after the strengthening of single CFs in individual PCs. The analysis of mutant mice deficient in glutamate receptor d2 subunit (GluRd2 or GluD2) indicates two distinct phases of CF synapse elimination (Hashimoto et al. 2009b). GluRd2 is richly expressed in PCs and its deletion causes impairment of PF-PC synapse formation leading to reduction of PF-PC synapse number to about half of the wild-type mice. Nevertheless, GluRd2 deletion does not significantly affect the histoarchitecture of the cerebellum and morphology of PC and its dendritic tree (Kashiwabuchi et al. 1995; Kurihara et al. 1997). In GluRd2 knockout mice, the average number of CFs innervating each PC was similar to that of control mice from P5 to P11. However, the value was significantly larger than that of control mice from P12 to P14 (Hashimoto et al. 2009b). These results collectively indicate that CF synapse elimination in mice can be classified into two distinct phases, namely the “early phase” from P6 to around P11 which is independent of PF-PC synapse formation and the “late phase” from around P12 and thereafter which requires normal PF-PC synapse formation (Fig. 14.1b (2) Early phase and (4) Late phase of CF elimination) (Hashimoto et al. 2009b). Mechanisms of the early phase of CF synapse elimination remain largely unknown. However, patterns of CF activity have been reported to influence the early phase of CF synapse elimination. Andjus et al. disrupted the normal activity pattern of CF in rat at P9-P12 by administration of harmaline, which induced synchronous activation of inferior olive neurons (Andjus et al. 2003). This treatment caused persistent multiple CF innervations of PCs in rats at P15-P87. Furthermore, a recent report strongly suggests that PC activity is crucial for CF synapse elimination (Lorenzetto et al. 2009). Lorenzetto et al. generated transgenic mice that expressed a chloride channel-YFP fusion protein specifically in PCs to suppress their excitabilities. In these mice, the expression of chloride channel was observed in PCs during the “early phase” at P9, and multiple CF innervations persisted up to P90. Therefore, perturbation of PC activity is considered to cause impairment of the “early phase” of CF synapse elimination. As for possible molecular mechanisms, insulin-like growth factor I (IGF-1) is reported to be involved in CF synapse elimination from P8 to P12 (Kakizawa et al. 2003). IGF1 is thought to enhance the strengths of CF synapses and promote their survival, whereas the shortage of IGF-1 appears to impair the development of CF synapses (Kakizawa et al. 2003). In addition, Sherrard et al. reported recently that the activated and full-length forms of TrkB, a receptor for BDNF, fall while the expression of the truncated form, which acts as a negative regulator of TrkB

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signaling, increases around the onset of the early phase of CF synapse elimination (Sherrard et al. 2009). This finding suggests that decrease in Trk B signaling might permit the elimination of surplus CF synapses. Morphological data indicate that CFs that undergo dendritic translocation keep their synapses on the PC soma during the second postnatal week. In contrast, synaptic terminals of the weaker CFs are confined to the soma and the basal part of the primary dendrite. The characteristic pericellular nest consists of somatic synapses originating from collaterals of a single predominant CF and from weaker CFs, and thus represents multiple CF innervation of PCs (Hashimoto et al. 2009a). Therefore, CF synapse elimination is thought to be a process of nonselective pruning of perisomatic synapses, which spares dendritic synapses of a single predominant CF and leads to mono-innervations of that CF (Hashimoto et al. 2009a).

Late Phase of Climbing Fiber Synapse Elimination In the mature cerebellum, PFs form synaptic contacts on spines of PC distal dendrites, whereas CFs innervate their proximal dendrites. In the GluRd2 knockout mice, the density of PF-PC synapses is decreased about half of the wild-type mice, and about 40% of spines are “naked” spines that are not contacted by PF terminals. Consequently, CFs invade the distal dendrites and form ectopic synapses there (Hashimoto et al. 2001; Ichikawa et al. 2002). These ectopic CF synapses appeared around P10 when PF synapse formation and extension of PC dendritic arbor occur most vigorously. Similar abnormality of CF innervation is also found in a mutant mouse deficient in cbln1 in which PF to PC synapse formation is severely impaired (Hirai et al. 2005). These results suggest that PFs compete for postsynaptic sites on PC dendrites with CFs during development, and confine the CF innervation territories to proximal dendrites (Fig. 14.2, “Restriction to proximal dendrites”). PF-PC synapses transmit signals from mossy fibers to PCs. In particular, impulses along PFs activates type 1 metabotropic glutamate receptor (mGluR1) and its downstream signaling cascades in PCs, which has been shown to drive the process of CF synapse elimination (Kano and Hashimoto 2009). Electrophysiological examination demonstrates that the mutant mice deficient in mGluR1 are impaired in CF synapse elimination (Kano et al. 1997; Levenes et al. 1997). Mice deficient in signaling molecules downstream of mGluR1, Gaq, PLCb4, or PKCg, are also impaired in CF synapse elimination (Kano et al. 1995, 1998; Offermanns et al. 1997; Hashimoto et al. 2000). Electrophysiological examination of CF innervation following postnatal development demonstrates that the regression of CF synapse occurs normally during the first and second postnatal weeks in all of the four mouse strains. However, these mice display abnormality in synapse elimination during the third postnatal week. These results suggest that the signaling cascade from mGluR1 to PKCg is essential for the late phase of CF synapse elimination, but it is not required for the early phase of CF synapse elimination. Importantly, formation of PF to PC synapses is not impaired in these mutant mice. Gross anatomy of the cerebellum and morphology of PC are largely normal. PCs

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Fig. 14.2 Mechanisms underlying the late phase of CF synapse elimination. PF-PC synapses play two distinct roles in the late phase. PF synapses are formed and maintained on distal dendrites of PCs through the interaction of Cbln1 and GluRd2. First, PF synapses occupy the post synaptic sites on the distal dendrites and confine the CF innervation sites to the proximal dendrites (Restriction to proximal dendrites). Second, neural activity along the pathway of mossy fiber, GC and PF involving NMDA receptor at MF to GC synapses drives mGluR1 to PKCg signaling cascades in PCs (Activation of mGluR1 signaling cascade), which eventually leads to elimination of somatic CF synapses including those of weak CFs and of somatic collaterals of strong CF (elimination of somatic synapses). TrkB, myosin Va, BSRP and GLAST are also involved in the late phase of CF synapse elimination. (Modified from Kano et al. 2008)

have well-differentiated dendritic arbors with numerous dendritic spines, which is indistinguishable from wild-type PCs. Furthermore, structure and density of PF-PC synapses are normal in electron microscopic examination. These results indicate that the impaired CF synapse elimination is not caused secondarily by the defect in PF-PC synapse formation. Several lines of evidence indicate that mGluR1 signaling within PC is crucial for CF synapse elimination. The defect in the CF synapse elimination in the mGluR1 knockout mice is restored in the mGluR1-rescue mice in which mGluR1a has been introduced specifically into PCs (Ichise et al. 2000). Regression of CF synapses is impaired in mice by PC-specific expression of a PKC inhibitor peptide (De Zeeuw et al. 1998). Furthermore, the distribution of multiply innervated PCs in the cerebellum of PLCb4 knockout mouse exactly matches that of the PCs with

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predominant expression of PLCb4 in the wild-type mouse cerebellum (Kano et al. 1998; Hashimoto et al. 2000). Thus, the signaling from mGluR1 to PKCg in PCs but not other cell types should play a central role in CF synapse elimination. Evidence suggests that mGluR1 activation at PF synapses drives the late-phase of CF synapse elimination. It is known that mGluR1 can readily be activated by PF inputs (Batchelor et al. 1994; Finch and Augustine 1998; Takechi et al. 1998), while hardly by CF inputs without blockade of glutamate transporters (Dzubay and Otis 2002). Furthermore, chronic blockade of NMDA receptors within the cerebellum results in the impairment of CF synapse elimination (Rabacchi et al. 1992) specifically in its later phase (Kakizawa et al. 2000). NMDA receptors are not present at either PF or CF synapses on PCs but they are abundantly expressed at mossy fiber to granule cell synapses. Therefore, the chronic blockade of NMDA receptors within the cerebellum should affect mossy fiber to granule cell transmission. These results indicate that neural activity along mossy fiber–granule cell–PF-PC pathway and subsequent activation of mGluR1 is prerequisite for the late phase of CF synapse elimination (Fig. 14.2, “Activation of mGluR1 signaling cascade”) (Kakizawa et al. 2000). As for other molecules that may be involved in CF synapse elimination, a neurotrophin receptor, TrkB (Bosman et al. 2006; Johnson et al. 2007), a motor protein, myosin Va (Takagishi et al. 2007), a glutamate transporter, GLAST (Watase et al. 1998) and a novel brain-specific receptor-like protein family, BSRP (Miyazaki et al. 2006) have been reported. Since genetic or pharmacological deletion of these molecules in mice impairs CF synapse elimination in the second postnatal week, these signaling cascades are thought to be involved in the “late phase” of CF synapse elimination. Although details of signaling cascades in which these molecules are involved are not known, a neurotrophin receptor, TrkB is especially interesting, because TrkB signaling is required for normal development of GABAergic innervations of PCs. In TrkB knockout mice, the number of GABAergic synapses is reduced (Rico et al. 2002) and the inhibitory postsynaptic currents are prolonged (Bosman et al. 2006), which suggest the GABAA receptors do not undergo the normal a3 to a1 subunit switching (Takayama and Inoue 2004). It is therefore possible that the impaired CF synapse elimination in TrkB knockout mice may be caused secondarily by the insufficient developmental maturation of inhibitory synapses onto PCs. It is reported that null mutant mice deficient in Ca2+/ calmodulin-dependent protein kinase IV (CaMKIV) have persistent multiple CF innervations, but it is unclear at what stage of postnatal development the impairment occurs (Ribar et al. 2000). It is also reported that null mutant mice deficient in aCaMKII display multiple CF innervations at P21-P28, but this phenotype disappears in adulthood (Hansel et al. 2006), suggesting that aCaMKII deficiency delays but not prevent CF synapse elimination. It is reported that Lysotracker positive structures surrounding PCs, which are presumed to be within Bergmann glia, are abundant during the second and third postnatal weeks (Song et al. 2008). Lysotracker Red is a marker for the lysozomes and late endosomes of living cells, which positively stains bulb-shaped tips of retreating motor axons and the axon fragments (“axozomes”) engulfed by Schwann

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cells during synapse elimination of neuromuscular junction (Bishop et al. 2004). Therefore, it is possible that retreating CF axons might be digested by glia in a manner similar to the retreating motor axons at neuromuscular junction. Figure 14.2 describes schematically the current model of the late phase of CF synapse elimination. As mentioned above, PF-PC synapses play two distinct roles. First, PFs occupy the spines of distal dendrites of PCs and confine the CF synapses to the proximal dendrites (Fig. 14.2, “Restriction to proximal dendrites”). Second, PFs convey neural activity from mossy fibers through NMDA receptor activation at mossy fiber to granule cell synapses. This neural activity activates mGluR1 at PF to PC synapses and drives its downstream signaling to PKCg in PCs (Fig. 14.2, “Activation of mGluR1 signaling cascade”). The signaling molecules downstream of PKCg are currently unknown, but these molecules nonselectively eliminate somatic CF synapses from the weaker CFs and from somatic collaterals of the strong CF (Fig. 14.2, “Elimination of somatic CF synapses”). The molecular mechanisms underlying morphological elimination of the weaker CFs are also unknown. Some mechanisms must convey trans-synaptic retrograde signaling from PCs to weaker CFs, either by some diffusible retrograde messengers such as cytokines or by molecules mediating protein–protein interactions that bridge preand postsynaptic membranes.

Heterosynaptic Competition between Parallel Fiber and Climbing Fiber Inputs In PCs, dendritic spines in the proximal dendritic compartment are innervated by single CFs, while those in the distal compartment are innervated by PFs (Fig. 14.3 middle). The construction of the characteristic excitatory wirings stands on competitive equilibrium among afferents promoted by distinct molecular mechanisms. Surgical, pharmacological, and genetic manipulations that shift the equilibrium also alter this territorial innervation. Disruption of GluRd2 gene in mice not only impairs PF synapse formation, but also affects the mode of CF innervations (Ichikawa et al. 2002). In the molecular layer, CF branches are distributed over inner four-fifth portions in control mice (84%), whereas their distribution almost reaches the pial surface in GluRd2 knockout mice (95%). When the tracer-labeled CFs were followed from the soma to the tips of PC dendrites by serial electron microscopy, this expanded distribution represents distal extension of CF branches to take over free spines on the distal dendritic compartment. Such aberrant extension occurs toward not only distal dendrites of the same PCs but also those of the neighboring PCs. The latter type of spine takeover thus causes the innervation of a given PC by multiple CFs of different neuronal origins (Fig. 14.3, left). This anatomical evidence for multiple CF innervation is consistent with electrophysiological recording combined with Ca2+ imaging. In GluRd2 knockout mice, a single strong CF elicits large EPSCs with a fast rise time and high Ca2+ elevation over the entire dendritic tree, whereas weak CFs elicit small EPSCs with a slow rise time and low Ca2+ elevation confined

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PF PF PF+CF CF

CF

PF+CF

CF CF GluRδ2-KO Cbln1-KO

CF Wild type

CF

CF

Cav2.1-KO

Fig. 14.3 Summary diagram of molecular mechanisms for competitive synaptic wiring in PCs. Note that the CF (CF) and PF (PF) territories are reversed in mutant mice defective in GluRd2/ Cbln1 (left) and CaV2.1 (right). With both mechanisms, CF and PF territories are sharply segregated and CF mono-innervation is established in wild-type animals (middle). (Reproduced from Watanabe 2008, with permission)

to some distal dendrites (Hashimoto et al. 2001). These findings indicate that GluRd2 is also essential to restrict CF innervation to the proximal dendritic compartment, which eventually prevents multiple CF innervation at this compartment (Fig. 14.3, left). Interestingly, Cbln1-knockout mice also manifest similar phenotypes, including impaired PF-PC synaptogenesis, persistent multiple CF innervation, impaired long-term depression at PF-PC synapses, and severe ataxia (Hirai et al. 2005). This is because postsynaptic GluRd2 trans-synaptically interacts with presynaptic neurexins through Cbln1 (Matsuda et al. 2010; Uemura et al. 2010). Thus, Cbln1 acts as a bidirectional synaptic organizer at PF-PC synapses on the distal dendritic compartment, which eventually restricts CF territory to the proximal compartment. This mechanism is also active in the adult cerebellum. The ablation of GluRd2 in adulthood also leads to disconnection and mismatching of PF-PC synapses and progressive distal extension of ascending branches of CFs (Takeuchi et al. 2005; Miyazaki et al. 2010). Ascending CF branches aberrantly innervate distal dendrites of not only the target PCs, but also neighboring ones. Furthermore, transverse branches of CFs, which are a short motile collateral forming no synapses in wildtype animals (Rossi et al. 1991; Sugihara et al. 1999; Nishiyama et al. 2007), display aberrant mediolateral extension to innervate distal dendrites of neighboring and remote PCs. Consequently, many PCs are wired by single main CF and surplus CFs innervating small parts of distal dendrites. Surplus CF-EPSCs with slow rise time and small amplitude also emerge progressively after GluRd2 ablation.

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Likewise, when recombinant Cbln1 is applied to the subarachnoid space of adult Cbln1-knociout mice, only a single injection can rapidly restore PF-PC synapse structure and function, and cerebellar ataxia (Ito-Ishida et al. 2008). Therefore, the GluRd2-Cbln1-neurexin system is essential to maintain PF-PC synapses at the distal dendritic compartment and to keep CF mono-innervation in the adult cerebellum by suppressing aberrant invasion of CF branches to the distal dendritic compartment. In contrast, CF innervation is regressed but PF innervation expands to the proximal compartment, when surgical lesion is given to olivocerebellar projections in adult animals or when activities in the cerebellar cortical neurons are blocked with the sodium channel blocker tetrodotoxin or with the AMPA receptor antagonist NBQX (Bravin et al. 1995; Kakizawa et al. 2000; Cesa et al. 2007). Such a change often accompanies hyperspiny transformation at the proximal dendritic compartment (Bravin et al. 1995; Cesa et al. 2007). Similar changes are reproduced in mutant mice defective in the pore-forming a subunit CaV2.1 (a1A) of P/Q-type Ca2+ channels (Miyazaki et al. 2004). This channel is one of the high voltageactivated Ca2+ channels, and constitutes >90% of the total Ca2+ current density in PCs (Mintz et al. 1992; Stea et al. 1994). CaV2.1 is abundantly distributed in PC dendrites and spines (Kulik et al. 2004). In CaV2.1 knockout mice, hyperspiny transformation is induced at proximal dendrites and somata of PCs, and many of these ectopic spines are innervated by PF terminals. Conversely, the distribution of CFs is regressed to lower portions of the molecular layer, and they innervated spines from somata and basal dendrites. Furthermore, in more than 90% of CaV2.1 knockout PCs, their basal dendrites and somata are innervated by CFs of different neuronal origins. As a result, the proximal somatodendritic compartment in CaV2.1 lacking PCs receives chaotic innervation by numerous PFs and multiple CF (Fig. 14.3, right). Considering that PCs lack functional NMDA receptors (Yamada et al. 2001), P/Q-type Ca2+ channels in PCs might substitute for the role of NMDA receptors, and function as a coincidence detector to regulate synaptic strengthening and elimination. Thus, CF activities leading to AMPA receptor activation and subsequent Ca2+ influx through P/Q-type Ca2+ channels are essential for monopolizing the proximal dendritic compartment by a single main CF and for expelling other excitatory inputs from that compartment. Taken altogether, excitatory synaptic wiring in PCs is formed and maintained through homosynaptic competition among CFs and heterosynaptic competition between PFs and CFs. GluRd2 and Cbln1 fuel heterosynaptic competition in favor of PF innervation, whereas P/Q-type Ca2+ channels facilitate both heterosynaptic and homosynaptic competitions in favor of single main CFs. Based on these molecular mechanisms, PCs can establish territorial innervation by PF and CF and mono-innervation by CF.

Conclusions and Future Directions This chapter made an overview of postnatal development of CF-PC synapses, which is one of the best studied examples of synapse elimination in the brain.

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The outline has now become clear from a number of electrophysiological and morphological studies. Shortly after birth, each PC is innervated by multiple CFs with similar synaptic strengths on the soma. Subsequently, a single CF is selectively strengthened during the first postnatal week. Then, around postnatal day 9 (P9), only the strongest CF (“winner” CF) starts to extend its innervation to PC dendrites. In contrast, synapses of the weaker CFs (“loser” CFs) remain around the soma, and they are eliminated progressively during the second and the third postnatal weeks. From P6 to P11, the elimination proceeds independently of PF-PC synapse formation. From P12 and thereafter, the elimination of weaker CFs requires normal PFPC synapse formation. There is a heterosynaptic competition between PFs and CFs for the postsynaptic sites on Purkinje cell dendrites, which is at work not only during postnatal development but also in adulthood. Molecular mechanisms of the late phase of CF synapse elimination and the heterosynaptic competition have been elucidated to some extent in the past 15 years by using a number of knockout mice and pharmacological approaches (see Figs. 14.2 and 14.3). In contrast, little is known about the molecular mechanisms of selective strengthening of single CF, the early phase of CF synapse elimination, and CF translocation. In addition to knockout mice, the RNAi technology to knockdown candidate molecules will be useful. In addition, using an in vitro organotypic culture system is another approach to pursue the molecular mechanisms of CF synapse elimination. Letellier et al. recently reported very interesting results by using an organotypic slice culture that CF synapse elimination occurs only during the critical period that depends on the maturation stage of postsynaptic PCs but not on that of presynaptic olivary neurons (Letellier et al. 2009). The cerebellum has been attracting many neuroscientists who pursue the mechanisms of synaptogenesis and synapse elimination. Continuing research on cerebellar microcircuits will elucidate fundamental mechanisms of the formation, elimination, and maturation of neural circuits. Acknowledgments This work has been supported in part by the Strategic Research Program for Brain Sciences (Development of Biomarker Candidates for Social Behavior), Grants-in-Aid for Scientific Research 21220006 (M.K.) and 19100005 (M.W.), and the Global COE Program (Integrative Life Science Based on the Study of Biosignaling Mechanisms) from MEXT, Japan.

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Genes and Cell Type Specification in Cerebellar Development

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Matt Larouche and Daniel Goldowitz

Abstract

One of the key goals of neural development is to make specific cell types that originate from multipotent progenitor cells. The process of cell specification is only beginning to be understood. Evidence thus far suggests that it occurs in a stepwise fashion, and it is likely that each step requires the coordinated expression of a unique set of genes. The cerebellum is an excellent model system for understanding cell fate questions because it contains only a handful of defined cell types that are each located in a specific lamina and are therefore easily identified. These features have made the cerebellum an essential brain region in the understanding of the gene networks that give rise to specific cell types during development. This chapter will first discuss recent advances in parsing the pathways necessary to produce specific cerebellar cell types. Next, the open-source cerebellar GRiTS (Gene Regulation in Time and Space) project (CBGRiTS.org), which has amassed a microarray-based readout of cerebellar gene expression on a daily basis during embryogenesis and every 3 days postnatally, will be discussed. Finally, efforts to mine this transcriptomic information using novel bioinformatic tools to search for new genes that may confer cell-type specificity during cerebellar development will also be discussed.

M. Larouche (*) • D. Goldowitz Department of Medical Genetics, Child and Family Research Institute, Centre for Molecular Medicine and Therapeutics, University of British Columbia, 950 West 28th Avenue, Vancouver, BC, V5Z 4H4, Canada e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 301 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_15, # Springer Science+Business Media Dordrecht 2013

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Introduction The cerebellum is a well-studied model of neural development largely because it is a simple structure. It contains only a handful of cell types, including Purkinje, granule, stellate, basket, Golgi, unipolar brush, and Lugaro neurons, in addition to glial populations that include astrocytes, oligodendrocytes, and Golgi epithelial cells (also known as Bergmann glia). This region of the nervous system has been an essential contributor to the understanding of a variety of developmental processes, including progenitor proliferation, neuronal specification, differentiation, and migration (Fig. 15.1a). Recent progress in cell fate research has demonstrated that sequential cascades are responsible for defining some cell types. The cerebellar model is consistent with these ideas; however, at present, there is only a rough understanding of the genes necessary to specify particular cell types. For example, while it appears that a common progenitor population gives rise to all glutamatergic cells in the cerebellum, the genetic cascades required to produce each specific

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Fig. 15.1 Cerebellar development. (a) The cerebellum has played an essential role in understanding the genes that underlie key developmental processes. The cerebellum has been critical for elucidating the role of the Sonic hedgehog (Shh) pathway in regulating proliferation. It has also been essential for understanding specification (see this chapter), differentiation (see this chapter), and essential neuronal migratory pathways like Reelin. A partial list of genes and their role in neurodevelopment as studied in the cerebellum is shown. (b) A list of birthdates for the major cell types within the cerebellum is shown. Green bars represent the period when birth dating methods (H3 or BrdU) reveal incorporation into the DNA of that particular cell-type. References: a – (Altman and Bayer 1997); b – (Carletti and Rossi 2008); c – (Englund et al. 2006); d – (Leto et al. 2006); e – (Machold and Fishell 2005); f – (Miale and Sidman 1961); g – (Sekerkova et al. 2004); h – (Yamada and Watanabe 2002); i – (Yuasa 1996); j – (Zhang and Goldman 1996). (c) Cells that populate the cerebellum arise from two germinal zones – the rhombic lip (red – E11), located at the caudal aspect of the cerebellar anlage, and the neuroepithelium lining the roof of fourth ventricle (green – E11, E14). Glutamatergic cells arise from rhombic lip and typically migrate along the dorsal aspect of the cerebellum (red -E14) before finally entering this structure. Granule cells arise from precursors that are the primary constituents of the external germinal layer (EGL). In contrast, GABAergic cells, like Purkinje cells, arise in the neuroepithelium and migrate dorsally into the cerebellar anlage (E14, P30). Granule cells originate from precursors derived from the rhombic lip and surround the cerebellum in a secondary germinal known as the external germinal layer (EGL ¼ dark red, in E14 and P7 views). As granule neurons are born in the EGL, they migrate radially through the nascent molecular layer (ML//orange-shading) until they arrive in the internal granule layer (GL//light-red shaded area). In contrast, GABAergic cells, like Purkinje cells, arise in the neuroepithelium and migrate dorsally into the cerebellar anlage (E14, P30). In the mature cerebellum, Purkinje cells reside in a monolayer (PCL – black line P30) that separates the molecular and granular layers. Table in panel “C” summarizes the major cell types and the respective layer that each cell type is located in the mature cerebellum. Abbreviations: Ascl1 – achaete-scute complex homolog 1; Dab1 – disabled homolog 1; Dll – delta-like; E – embryonic; Gli – Gli-Kruppel family member; Hes – hairy and enhancer of split; Jag – jagged; Lrp8 – low-density lipoprotein receptor-related protein 8; NE – neuroepithelium of the 4th ventricle; P – postnatal; Ptc – patched homolog; RL – rhombic lip; Shh – Sonic hedgehog; Smo – smoothened;. Vldlr – very low-density lipoprotein receptor; Rln – reelin

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subtype are largely unknown (see below). This chapter will first summarize cerebellar development focusing on the various cell types that populate this structure. Next, an examination of the current understanding of the genes involved in conferring cell identity in the cerebellum will be conducted. For the purposes of brevity, the focus of this chapter will be restricted to genes that have been specifically reported to regulate the production of a particular cell type as opposed to influencing the survival/proliferation of cells. Finally, the opportunities that the GRiTS database can provide for discovering novel genes and networks involved in promoting the development of cell types during development will be discussed.

Cerebellar Structure and Development The cerebellum originates from the alar plate of the neural tube, and this region is known to give rise to the sensory structures of the nervous system. Functionally, the cerebellum is at the crossroads between the sensory and motor systems and is essential for coordinating communications between these two systems. (Altman and Bayer 1997). The cerebellar cortex has a trilaminar organization (Fig. 15.1c), and each layer contains a defined set of cell types – the outer most molecular layer contains stellate and basket interneurons in addition to glial cells; located beneath the molecular layer, the Purkinje cell layer contains the cell bodies of the namesake neuron; located immediately beneath the Purkinje layer is the granule layer which contains granule cells, Golgi cells, unipolar brush cells, Golgi epithelial cells, and a few other minor cell types (Altman and Bayer 1997). Internal to the cerebellar cortex is the cerebellar white matter, which contains the majority of the cerebellar astrocytes and oligodendrocytes. The final major cell type in the cerebellum is the cerebellar nuclear neurons, which are restricted to four bilateral pairs of nuclei symmetrically distributed on either side of the midline. During development, the cerebellar territory is established through the actions of the isthmic organizer (IO). The IO sets up the boundary between the mes – and metencephalic vesicles around embryonic day (E) 8.5–9 in the mouse (Wassef and Joyner 1997). The cerebellum arises from rhombomere 1 of the metencephalic vesicle. Mutations in the genes encoding the key morphogens secreted by the isthmic organizer, which include Otx2, Gbx2, Fgf8, and Wnt1, typically result in the absence of a cerebellum and/or midbrain. These mutations demonstrate the essential nature of the IO to the development of the caudal CNS. Because of the essential nature of these genes for the establishment and maintenance of the IO, inducible transgenics, which allow temporal control over gene inactivation, have been invaluable in establishing the role of many of these genes (e.g., Joyner and Zervas 2006). A more detailed discussion of the role of the IO can be found elsewhere (e.g., Wassef and Joyner 1997; Hidalgo-Sanchez et al. 2005). Around the time that the midbrain-hindbrain territory is established, the neural tube closes (Ybot-Gonzalez et al. 2002, 2007). Like most areas of the brain, the ventricular surface of the neural tube in the hindbrain serves as the germinal zone for the

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developing cerebellum. In the cerebellum, there are two germinal zones – the roof of the fourth ventricle, known as the neuroepithelium, and the edges of neural tube surrounding the fourth ventricle, known as the rhombic lip.

Cerebellar Cell Specification Cerebellar cells have well-defined developmental profiles that are characterized by discrete time periods in which each cell type is generated (Fig. 15.1b). Despite that understanding, there is a poor knowledge of the processes that regulate how each specific cell type is produced. Evidence suggests that combinations of transcription factors may work together in cerebellar development to specify various cell types. For example, glutamatergic nuclear neurons and progenitor populations express transcription factors known to influence cell fate such as Meis1 and 2, Irx3, and Lhx2/9 (Morales and Hatten 2006). Components of the notch-signaling cascade also define discrete zones in the roof of the fourth ventricle during periods where cell birth is ongoing in this germinal zone (Zordan et al. 2008). There are mutant alleles in mice that also predict the action of broadly acting regulators of cell specification in the cerebellum. For example, the absence of granule cells in the anterior cerebellum of the meander tail mutant points to the actions of a gene on a subpopulation of granule cell progenitors (Ross et al. 1990). Chimera experiments demonstrate that these effects are mediated in a cell autonomous manner (Goldowitz and Hamre 1998). Currently, the best-understood process in cerebellar cell differentiation is the specification of cells along either the GABAergic or glutamatergic lineages (Fig. 15.2 – green ovals). There is growing evidence indicating that the rhombic lip produces glutamatergic neurons, whereas GABAergic neurons originate from the neuroepithelium (Fig. 15.1c; e.g., Carletti and Rossi 2008). Thus, granule neurons, unipolar brush cells, and the glutamatergic cerebellar nuclear cells arise from the rhombic lip (Wingate 2001; Wang et al. 2005; Englund et al. 2006). By contrast, Purkinje neurons, GABAergic cerebellar nuclear neurons, the interneuron populations, including stellate, basket, Golgi, and Lugaro neurons, are all generated from progenitors that originate in the neuroepithelium of the forth ventricle (Altman and Bayer 1985a).

Glutamatergic Cells The first cerebellar cells are initially generated around the time when the neural tube closes (Fig. 15.1c). The glutamatergic cerebellar nuclear neurons are the earliest cells produced in the cerebellum between E10.5 and 12.5 in the mouse (Miale and Sidman 1961). These cells arise from the rhombic lip and take a tangential course within the cerebellar parenchyma where they migrate near to the dorsal surface of the cerebellum. They arrive at the region termed the nuclear transitory zone before being displaced to their ultimate destination – the cerebellar

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Fig. 15.2 Genes mediating cell-lineage decisions in the developing cerebellum. Unique cell types within the cerebellum depend on the action of various genes, and each necessary gene is listed above the line leading to that cell type. Dotted arrows indicate putative relationships suggested by the literature, black solid arrows indicate published relationships. The least differentiated cell types reside on the left-hand side (green squares), while the most differentiated cells are on the right (red circles/ovals). Abbreviations used: GCP ¼ granule cell precursor; Glu ¼ glutamatergic; IGL -internal granule layer; UBC ¼ unipolar brush cell

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nuclei – situated in the ventral aspect of the mature cerebellum (Altman and Bayer 1985b). Whether this process is by active migration or passive mechanical forces is not currently known. The next glutamatergic cells born in the cerebellum are granule cells, which are born over a protracted period of time beginning with the allocation of the precursor population in the rhombic lip between E12 and 14.5 (Miale and Sidman 1961; Machold and Fishell 2005). Following their generation, these precursors migrate out over the dorsal surface of the cerebellum and form a layer known as the external germinal layer (EGL), which covers the dorsal surface of the cerebellum. Beginning around E15, these precursors proliferate and give rise to the terminally differentiated granule neurons that then migrate inward, radially, from their position in the EGL to the internal granular layer, inside of the cerebellum. Unipolar brush cells (UBCs) are the latest born glutamatergic neuron generated between E14 and 19 in the rhombic lip (Englund et al. 2006; Sekerkova et al. 2004). Once born, UBCs migrate away from the rhombic lip through the inner cerebellar parenchyma finally ending up largely in lobules VI/VII and IX/X in the mature cerebellum (Sekerkova et al. 2004). Recent research indicates that the master transcription factor for the glutamatergic lineage in the cerebellum is Math1 (Fig. 15.2). This transcription factor is essential to promote glutamatergic neuron production both in the cerebellum as well as several nuclei in the neighboring brainstem (Ben-Arie et al. 1997). Math1 is a bHLH transcription factor and the mouse orthologue of the Drosophila Atonal (Ben-Arie et al. 1997). Math1 is expressed in the mouse rhombic lip as early as E9.5 (Akazawa et al. 1995; Gazit et al. 2004). Math1 knockout mice produce very few glutamatergic cells and lack almost all granule neurons, unipolar brush cells, and glutamatergic cerebellar nuclear neurons (Wang et al. 2005; Englund et al. 2006; Ben-Arie et al. 1997). Math1 is the earliest known marker of granule cell precursors, and the first granule cell precursors express this gene as early as E12.5 (Machold and Fishell 2005). Mouse chimeras demonstrate that Math1 acts cell autonomously in granule cells during development, as there are no mutant Math1 granule cells present in cerebella consisting of predominantly wild type cells (Jensen et al. 2004). Granule cells have the best-understood lineage of all glutamatergic cells in the cerebellum, and there are multiple genes involved in specifying these cells. For example, there is evidence that bone morphogenetic proteins (BMPs) may modulate Math1 in order to specify granule cells (Fig. 15.2). BMPs are a group of proteins related to the TGFß transcription factor family, and they are released from the roof plate, including that overlying the cerebellum (Miyazono et al. 2009). Ectopic expression of constitutively activated BMP receptor 1b results in an upregulation of the chick Math1 homologue – Cath1-transcripts (Machold et al. 2007). Conversely, mice with null mutations for both BMP receptors (BMP1a and 1b) fail to induce Math1 expression and have a disorganized cerebellum (Qin et al. 2006). Mice with either BMP receptor singly knocked out have no Math1 phenotype (Qin et al. 2006) and suggest that the two BMP receptors are functionally redundant. Exogenous application of BMPs to cultured granule cells can also induce Math1 expression in culture (Alder et al. 1999). Together, these observations suggest that BMPs can induce Math1 and indicate that Math1 signaling occurs downstream from BMPs.

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In addition to Math1 and BMP, there are several other genes that regulate glutamatergic cell identity in the cerebellum that have been previously characterized as proliferation regulators. Cyclin D2 is one such example. Cyclins, in general, bind and activate cell division kinases (CDKs) and regulate progression through the cell cycle (Kozar and Sicinski 2005). Cyclin D2, in particular, is necessary to produce granule cells. Mice lacking functional cyclin D2 have a reduced number of granule neurons and also lack GABAergic stellate cells (see below for discussion about stellate cells Huard et al. 1999). Currently, it is not clear if Cyclin D2 regulates aspects of the cell cycle in granule or stellate cells, or their precursors. It has been postulated that reductions in the cell-division rate of precursors would have a direct impact on the number of cells produced. Huard and colleagues did observe an increase in the number of granule cell precursors experiencing apoptosis in the EGL of cyclin D2 mutants, suggesting that CyclinD2 could act to maintain the progenitor populations (Huard et al. 1998). Like CyclinD2, members of the Zic family of zinc-finger transcription factors also regulate cell proliferation and glutamatergic identity in the cerebellum. Zics are related to the Drosophila, odd-paired pair rule gene, and the vertebrate family has five homologues in vertebrates (Zic1-5). These transcription factors typically have roles in cell specification but also regulate cell proliferation (Aruga et al. 1996). Zics are importantly involved in cerebellar development; for example, deletion of Zic1 results in severe cerebellar malformations, in addition to axial deficits in other parts of the body (Aruga et al. 1998). The cerebellar phenotype is at least partially manifested as a reduction in the number of granule cell precursors in these mutants as demonstrated through labeling experiments using the thymidine analogue – BrdU (Aruga et al. 1998). Deletion of Zic1 leads to the loss of a lobule located in the anterior cerebellum and could suggest effects of this gene in a localized progenitor subpopulation. Zic2 has also been proposed to regulate precursor proliferation rates in cooperation with Zic1. Compound mutant mice (Zic1+/::Zic2+/kd) also exhibit a reduction in the rates of cell proliferation in the EGL when compared to Zic1/ animals. These compound mutants are deficient in an anterior lobe folia but, in addition, they also lack a lobule in the posterior cerebellum (Aruga et al. 2002). One explanation for these effects is that Zic-regulation of granule cell proliferation could occur through interactions with cyclin D1 and/or D2 (Huard et al. 1999; Aruga et al. 2002). Tandem deletion of both Zic1 and Zic4 genes in mice gives rise to cerebellar abnormalities that are reminiscent of those seen in human Dandy–Walker malformations (Grinberg et al. 2004). The expression domains of different Zics often overlap in the cerebellum and suggest that these transcription factors may interact to exert their effects. Notch signaling is also well known to regulate cell-fate decisions and proliferation rates in both vertebrates and invertebrates. Consistent with these roles, there is growing evidence that this conserved pathway regulates the development of several cell types within the cerebellum. Whole-organism knockouts of notch-pathway members often result in embryonic lethality, and this limits the ability to understand how this pathway affects cell specification. However, conditional approaches have been invaluable for assessing the role of notch signaling in nervous system

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development. For example, knockout of floxed Notch1 mediated by the En1-cre transgene resulted in an almost complete elimination of nuclear neurons and granule cells in the cerebellum (Machold et al. 2007). In addition to Notch1, several other members of the notch-signaling pathway are expressed in granule neurons. Activity mediated by the Notch2 receptor is also necessary for producing postmitotic granule cells (Solecki et al. 2001). Overexpressing Notch2 in the postnatal cerebellum results in an increase in granule cell progenitor proliferation and suggests that this receptor could direct the terminal differentiation of granule cells (Solecki et al. 2001). Finally, NeuroD (also known as NeuroD1) acts later in granule cell development to specify these cells after they have undergone terminal division. NeuroD is a bHLH protein that binds to DNA and probably regulates the production of other neurogenic genes. There are discrete neuronal populations that are absent in NeuroD1 knockout mice, indicating that NeuroD1 is necessary for the development of only certain neuron subtypes. The affected neuronal populations include cells in the telencephalon, olfactory epithelium, and granule cells from the hippocampus and cerebellum (Lee et al. 2000). Within the cerebellum, the neuroD1 transcript is heavily expressed in the cell layer beneath the proliferating layer in the EGL, suggesting a role in promoting terminal differentiation in granule cells (Miyata et al. 1999).

GABAergic Cells The GABAergic cells first generated in the cerebellum are the Purkinje cells, which are born between E11 and 13.5 in the neuroepithelium lining the fourth ventricle (Miale and Sidman 1961). GABAergic nuclear neurons are also generated beginning around E11, and like Purkinje cells, the progenitors for nuclear neurons originate in the neuroepithelium of the fourth ventricle (Miale and Sidman 1961; Taber Pierce 1975; Leto et al. 2006). In contrast to the narrow time window for Purkinje cell genesis, nuclear neurons are generated into postnatal time periods and also arise from progenitor populations that reside in the white matter in addition to the neuroepithelium (Carletti and Rossi 2008; Leto et al. 2006). GABAergic interneuron precursors also originate in the neuroepithelium and begin to produce terminally differentiated neurons as early as E16; however, most interneurons are not born until P0–12 (Leto et al. 2006; Maricich and Herrup 1999). Most interneurons are born from progenitors that arise from the neuroepithelium and then come to reside in the cerebellar white matter. Finally, Lugaro cells are likely born in late embryogenesis or early postnatal stages (Laine et al. 1992). The pancreatic transcription factor 1a (Ptf1a) is essential to produce neurons with a GABAergic phenotype in the cerebellum. As its name suggests, Ptf1a was originally characterized for its essential role in the pancreas, where it functions to specify cells during development (Kawaguchi et al. 2002; Sellick et al. 2004). Interestingly, knockout mice lacking functional Ptf1a do not have Purkinje cells or

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other GABAergic cerebellar interneurons, including basket, stellate, and Golgi neurons (Hoshino et al. 2005; Pascual et al. 2007). These observations suggest that it is a master regulator for the GABAergic phenotype in the cerebellum. During development, Ptf1a is expressed in the neuroepithelium of the fourth ventricle at a time when Purkinje cells and nuclear neurons are normally generated (Hoshino et al. 2005). In the absence of Ptf1a, some of the progenitors adopt a granule cell– like phenotype, suggesting that Ptf1a may function to repress the glutamatergic phenotype (Pascual et al. 2007). The Lim homeodomain proteins have been associated with the GABAergic lineage in the cortex (e.g., Fogarty et al. 2007), and in the cerebellum they have been demonstrated necessary for Purkinje-cell development. In particular, Lhx1 and 5 are two closely related Lim-homeodomain transcription factors that are expressed in cells exiting the neuroepithelium at E12.5 – a time during Purkinjecell genesis (Zhao et al. 2007). Double mutants of these transcription factors exhibit severe reductions in the number of Purkinje cells, as assessed using anti-Calbindin immunostaining (Zhao et al. 2007). Likewise, genetically inactivating the essential cofactor for Lhx1/5 signaling – Ldb1 – results in a phenotype similar to the double knockout (Zhao et al. 2007). It is possible that the Lhx proteins regulate cell survival since they are expressed in post-mitotic cells; however, this hypothesis has not yet been investigated. In addition to Lim proteins, notch signaling also regulates the production of Purkinje cells in the cerebellum. Consistent with this idea is that several notchfamily members are expressed in the neuroepithelium, suggesting that these could regulate the production of other GABAergic cell types as well. For example, there is robust expression of Notch1 and its receptor Dll1 in the neuroepithelium between E10 and 12 – a time when Purkinje cells and nuclear neurons are born from this germinal zone (Fig 15.1b; Machold et al. 2007). Thus far, Notch1 activity is the only member of the notch-signaling pathway demonstrated to be involved in specifying Purkinje cells in the cerebellum. In addition to reduced numbers of granule and nuclear neurons, mice with a conditional deletion of Notch 1 receptor (En1-cre mediated) have reduced numbers of Purkinje neurons (Machold et al. 2007). Interestingly, there was no Purkinje-cell deficit observed in mice with an En2-cre mediated deletion of Notch1 (Lutolf et al. 2002). However, Machold and colleagues argue that since En1 is expressed both earlier and broader in the cerebellar anlagen than En2, there is a temporal window, where Notch1 activity is necessary to promote the Purkinje-cell fate (Machold et al. 2007). One well-characterized downstream effector of the notch pathway is Ascl1, formerly known as Mash1. Ascl1 and its Drosophila orthologues, Achaete and Scute, were originally characterized as neurogenic effectors of the notch pathway (Ishibashi 2004). Ascl1 is a bHLH transcription factor that has well-established roles in regulating fate decisions of GABAergic neurons in the cortex (reviewed in Schuurmans and Guillemot 2002). In the cerebellum of Asc1 knockout mice, Pax2+ interneurons are significantly reduced (Grimaldi et al. 2009). Moreover, transfecting the neuroepithelium of the fourth ventricle of wild type mice with a vector expressing Acsl1 increases the number of Pax2+ cells without affecting the

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rate of proliferation (Grimaldi et al. 2009). This evidence suggests that Ascl1 promotes the Pax2+ interneuron fate at a time after the cells have undergone terminal division. Interestingly, manipulating Ascl1 also has consequences on glial cell numbers in the developing cerebellum (see below) and indicates that interneurons and cerebellar glia potentially arise from similar progenitor pools. However, since Pax2 and Olig2 were never observed in the same cells in the cerebellum, it seems likely that the common progenitor of interneurons and oligodendrocytes occurs prior to the stage when Ascl1 is expressed (Grimaldi et al. 2009). Finally, it appears that, in addition to its role in promoting the granule cell fate, CyclinD2 is also essential for GABAergic stellate cells. These molecular interneurons are severely depleted in Cyclin D2 knockout cerebella (Huard et al. 1999). As with granule cells, it is unclear if this gene functions at the progenitor level or within the differentiated cell types. Most accounts suggest that granule cells and stellate cells are derived from separate progenitor populations (e.g., Zhang and Goldman 1996). However, there is some recent evidence suggesting that cells from both classes could arise from GFAP-expressing progenitor populations (Silbereis et al. 2009, 2010).

Cerebellar Glial Cells Classically, it was found that cerebellar glia have a postnatal birthdate (Miale and Sidman 1961). More recent birthdating evidence suggests that some glial progenitors are detected as early as E13–14 in the mouse, arising from the neuroepithelium of the fourth ventricle (Yamada and Watanabe 2002). There is conflicting evidence that suggests glial progenitors may also reside in the EGL. For example, experiments where replication-deficient retroviruses were applied directly to the EGL resulted in no glial cells becoming labeled (Zhang and Goldman 1996). However, recent experiments using GFAP-cre based lineage tracing methods indicate that both glial and granule cells from the GFAP lineage could arise from the EGL (Silbereis et al. 2009, 2010). There is compelling evidence to suggest that at least some cerebellar oligodendrocytes are derived from extracerebellar sources (Grimaldi et al. 2009). Recent evidence suggests that Pax2-expressing interneurons could share a common progenitor with glial cells in the developing cerebellum. In addition to decreased numbers of Pax2+ interneurons, Ascl1/ knockout cerebella also have reduced numbers of oligodendrocytes and increased numbers of astrocytes (Grimaldi et al. 2009). Transfecting the neuroepithelium of the fourth ventricle with vectors expressing Ascl1 reduces the number of astrocytes (identified by Glast and/or Sox9 expression). Interestingly, these overexpression experiments do not affect the number of oligodendrocytes in the nascent cerebellum. This observation is consistent with the hypothesis that oligodendrocytes at least, partially, originate from extracerebellar sources. In addition to the role of Ascl1 in regulating the production of interneurons in the cerebellum, notch signaling is also involved in regulating the formation of Bergmann glial cells. Cerebella from mice, where Jagged-1 is deleted using an

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En2-cre dependent-approach, have reduced numbers of Bergmann glial in the adult cerebellum. This observation suggests that Jag1 is responsible for specifying this glial subtype (Weller et al. 2006). However, further investigation is needed to determine if Jagged1 is functioning as a survival factor rather than a differentiation factor, and whether this is a cell-intrinsic or cell-extrinsic manner.

Strategies to Identify Novel Factors Necessary for Specifying Cells During Cerebellar Development In an effort to better understand cerebellar development from a molecular perspective, a project aimed at assembling a microarray-based developmental transcriptome for two strains of mice – C57B6/J and DBA across embryonic and postnatal development was recently initiated. RNA from cerebellar tissue was obtained from each strain from each day during embryogenesis (E12-birth) and every third day postnatally, currently up until P9. These RNA isolates were then analyzed using the Illumina microarray platform to assess the expression levels of each gene on each particular day. These data are publicly available on the Internet – www.cbgrits.org. Analysis of these data using novel bioinformatic tools to identify developmentally important genes is currently underway. Two bioinformatic approaches are being employed to identify genes that play candidate roles in cell specification and differentiation in cerebellar development. In the first approach, the birthdates of various cell types in the cerebellum were used to identify genes putatively involved in cell specification (see Fig. 15.1b). A guilt-by-association approach was employed reasoning that genes essential to a particular cell specification event would be up-regulated (or down-regulated) at the time when a particular cell type was born. Differential equation modeling was used to assess the expression of each gene across time and the significance of fit to the differential model was determined (Bao et al. 2007). Using this approach, significant inflection points in expression data were identified, and these inflections represent “on” and “off” points in gene activity. The list of genes significantly fitting the DEM was scanned to identify on-off points that coincide with cell specification events. For example, the literature indicates that Purkinje cells are born between E11 and 13.5 (Miale and Sidman 1961). Using this developmentally discrete time frame to query the list of genes that significantly fits the differential model reveals that Lhx5 is a candidate gene regulating the production of these neurons (Fig. 15.3a). The DEM identifies on and off inflection points at approximately E10.5 and 13.5, respectively, for Lhx5 in both the C57 and DBA strains of mice. Consistent with the hypothesis that Lhx5 is necessary for Purkinje-cell development, these neurons are absent in the E18 cerebellum of animals null for both Lhx1/5 (Fig. 15.3a; Zhao et al. 2007). The second bioinformatic approach to analyze the GRiTS transcriptome data is aimed at identifying gene networks essential for producing a cerebellum. To this end, the GRiTS data was analyzed using a comparative modeling approach capable of revealing putative regulatory relationships within a network. Conceptually, this analysis begins by organizing gene expression data into groups or clusters restricted

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Fig. 15.3 Bioinformatic tools generate testable hypotheses. (a) A differential equation modeling (DEM) approach compares the temporal expression profile for each gene to a differential equation model (46,632 probes measured via microarray for each time point). The DEM analysis identifies significant inflections in gene expression, and the first two inflections (first upward – on – and first downward – off) were identified for each gene significantly fitting the DEM. Two wild type strains were analyzed – DBA2J (D2) and C57BI/6 J (C57) – and about 2,000 genes fit the DEM for each strain. Reasoning that genes essential for cerebellar development would be common between both strains, lists for each strain were cross-referenced and yielded a set of 187 common genes. Finally, this list of 187 was filtered to identify genes having on-off points (+/ 0.5 day) coinciding with the birthdates of specific cerebellar cell types (i.e., Fig. 15.1b). Purkinje cells were selected as a seed for this approach, and their birthdate (E11–13) yielded a single gene – Lhx5 – that coincided with this period. The expression pattern reported in the Allen Brain Atlas (www.brain-map.org) for Lhx5 yielded expression localized to the areas, where differentiated Purkinje cells (P) are found at E11.5 (b, d) and E13.5 (c, e) in the cerebellum. Lhx5 expression is absent in both cerebellar germinal zones – the neuroepithelium (NE) and rhombic lip (RL) – and suggest that it is restricted to post-mitotic cells. (b) The parent–child tool identifies putative gene networks that merit further investigation. With this approach, genes are first organized into clusters, where all genes within a particular cluster share statistically similar temporal gene expression profiles. These temporally related clusters are then compared to each other in order to establish a putative hierarchy between clusters. Members of the notch-signaling pathway were used to seed this parent–child model. Circles represent temporally distinct gene clusters and arrows indicate hierarchical relationships. Notch-related genes appearing in each cluster are listed. Although, most of relationships proposed by the parent–child model are not reported in the literature, their putative arrangement is plausible based on the current understanding of this signaling cascade within the literature

to genes that have significantly similar temporal expression patterns. Once the clusters are established, regulatory relationships are then inferred between clusters based on conserved expression patterns offset by temporal shifts (Song et al. 2009a, b). This approach ultimately organizes the gene expression clusters into

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a hierarchy based on the timing of expression differentials. Arranged in this way, these data are capable of identifying both established relationships and are capable of revealing novel regulatory relationships. For example, one pathway that is well represented by this bioinformatic approach is the notch-signaling pathway (Fig. 15.3b). Specifically, a comparative analysis of the GRiTS data reveals that the transcriptional activator Mastermind-like 1 (Maml1) acts as a parent gene regulating the transcription of Hairy Enhancer of Split 1 (Hes1). This relationship is supported by published findings (Wu et al. 2000). Similarly, the comparative analysis predicts that the Notch RNA-binding protein Musashi will act as a parent putatively regulating Numb1 expression, and these observations are supported by reports in the literature (Okano et al. 2002). Other interactions between notch-pathway members are predicted but not yet confirmed (Fig 15.3b). Thus, analyses like the comparative modeling approach could reveal the nature of networks responsible for shaping cerebellar development by regulating characteristics like cell identity during development. In conclusion, the last decade has proven very fruitful in improving the understanding of the processes of cell specification in brain development. However, only a limited number of genes have been clearly implicated in this process. The cytoarchitectural simplicity of cerebellum that includes a detailed understanding of the birthdate and developmental characteristics of many of its cell types suggests that the cerebellum is poised to make great advances in the understanding of brain development in the coming years. The GRiTS time-series data will be a valuable contribution to this process because of its richness of temporal data and accessibility to the general research community. This is an unparalleled tool, and in combination with the application of algorithms for its analysis, it will help reveal genes and genetic interactions necessary to build a CNS structure.

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Miale IL, Sidman RL (1961) An autoradiographic analysis of histogenesis in the mouse cerebellum. Exp Neurol 4:277–296 Miyata T, Maeda T, Lee JE (1999) NeuroD is required for differentiation of the granule cells in the cerebellum and hippocampus. Genes Dev 13:1647–1652 Miyazono K, Kamiya Y, Morikawa M (2009) Bone morphogenetic protein receptors and signal transduction. J Biochem 147:35–51 Morales D, Hatten ME (2006) Molecular markers of neuronal progenitors in the embryonic cerebellar anlage. J Neurosci 26:12226–12236 Okano H, Imai T, Okabe M (2002) Musashi: a translational regulator of cell fate. J Cell Sci 115:1355–1359 Pascual M, Abasolo I, Mingorance-Le Meur A et al (2007) Cerebellar GABAergic progenitors adopt an external granule cell-like phenotype in the absence of Ptf1a transcription factor expression. Proc Natl Acad Sci USA 104:5193–5198 Qin L, Wine-Lee L, Ahn KJ et al (2006) Genetic analyses demonstrate that bone morphogenetic protein signaling is required for embryonic cerebellar development. J Neurosci 26:1896–1905 Ross ME, Fletcher C, Mason CA et al (1990) Meander tail reveals a discrete developmental unit in the mouse cerebellum. Proc Natl Acad Sci 87:4189–4192 Schuurmans C, Guillemot F (2002) Molecular mechanisms underlying cell fate specification in the developing telencephalon. Curr Opin Neurobiol 12:26–34 Sekerkova G, Ilijic E, Mugnaini E (2004) Time of origin of unipolar brush cells in the rat cerebellum as observed by prenatal bromodeoxyuridine labeling. Neuroscience 127:845–858 Sellick GS, Barker KT, Stolte-Dijkstra I et al (2004) Mutations in PTF1A cause pancreatic and cerebellar agenesis. Nat Genet 36:1301–1305 Silbereis J, Cheng E, Ganat YM et al (2009) Precursors with glial fibrillary acidic protein promoter activity transiently generate GABA interneurons in the postnatal cerebellum. Stem Cells 27:1152–1163 Silbereis J, Heintz T, Taylor MM et al (2010) Astroglial cells in the external granular layer are precursors of cerebellar granule neurons in neonates. Mol Cell Neurosci 44:362–373 Solecki DJ, Liu XL, Tomoda T et al (2001) Activated Notch2 signaling inhibits differentiation of cerebellar granule neuron precursors by maintaining proliferation. Neuron 31:557–568 Song MJ, Lewis CK, Lance ER et al (2009a) Reconstructing generalized logical networks of transcriptional regulation in mouse brain from temporal gene expression data. EURASIP J Bioinform Syst Biol 2009:545176 Song M, Ouyang Z, Liu ZL (2009b) Discrete dynamical system modelling for gene regulatory networks of 5-hydroxymethylfurfural tolerance for ethanologenic yeast. IET Syst Biol 3:203–218 Taber Pierce E (1975) Histogenesis of the deep cerebellar nuclei in the mouse: an autoradiographic study. Brain Res 95:503–518 Wang VY, Rose MF, Zoghbi HY (2005) Math1 expression redefines the rhombic lip derivatives and reveals novel lineages within the brainstem and cerebellum. Neuron 48:31–43 Wassef M, Joyner AL (1997) Early mesencephalon/metencephalon patterning and development of the cerebellum. Perspect Dev Neurobiol 5:3–16 Weller M, Krautler N, Mantei N et al (2006) Jagged1 ablation results in cerebellar granule cell migration defects and depletion of Bergmann glia. Dev Neurosci 28:70–80 Wingate RJ (2001) The rhombic lip and early cerebellar development. Curr Opin Neurobiol 11:82–88 Wu L, Aster JC, Blacklow SC et al (2000) MAML1, a human homologue of Drosophila mastermind, is a transcriptional co-activator for NOTCH receptors. Nat Genet 26:484–489 Yamada K, Watanabe M (2002) Cytodifferentiation of Bergmann glia and its relationship with Purkinje cells. Anat Sci Int 77:94–108 Ybot-Gonzalez P, Cogram P, Gerrelli D et al (2002) Sonic hedgehog and the molecular regulation of mouse neural tube closure. Development 129:2507–2517

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Hormones and Cerebellar Development

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Noriyuki Koibuchi and Yayoi Ikeda

Abstract

Cerebellar development involves various epigenetic processes that activate specific genes at different time points. The epigenetic influences include humoral influences from endocrine cells. Among circulating hormones, a group of small lipophilic hormones such as steroids (corticosteroids, progesterone, androgens, and estrogens) and thyroid hormone may particularly serve an important role in mediating environmental influences to the cerebellum. Receptors for such lipophilic hormones are mainly located in the cell nucleus (nuclear receptor, NR), and represent the largest family of ligand-regulated transcription factors. In the cerebellum, these are expressed in a specific temporal and spatial pattern. Among lipophilic hormones, involvement of thyroid hormone and gonadal steroids on cerebellar development has been well studied. Deficiency of thyroid hormone during postnatal development results in abnormal cerebellar morphogenesis in rodents. Estrogen and progesterone also play an important role in this process. In addition to the supply from circulation, several gonadal steroids are produced locally within the Purkinje cell (neurosteroids). In this chapter, the effect of thyroid and steroid hormones are separately discussed. Neurosteroids that are locally synthesized in the cerebellum are discussed in ▶ Chap. 42, “Neurosteroids and Synaptic Formation in the Cerebellum”.

N. Koibuchi (*) Department of Integrative Physiology, Gunma University Graduate School of Medicine, 3-39-22 Showa-machi, 371-8511 Maebashi, Gunma, Japan e-mail: [email protected] Y. Ikeda Department of Histology and Cell Biology, Yokohama City University School of Medicine, 3-9 Fukuura, 236-0044 Yokohama, Japan e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 319 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_16, # Springer Science+Business Media Dordrecht 2013

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Introduction Development of the brain involves epigenetic processes that activate specific genes at different time points. Epigenetic influences that control neuronal development may originate from the neuronal cell itself, or from outside of the brain. The former includes spatial and temporal patterning of gene expression tightly regulated by their intrinsic molecular programs. The latter includes sensory influence, mediated by peripheral nervous system, and humoral influence from endocrine cells. Environmental influences, such as stressors, social experience, nutrients, drugs, and environmental chemicals, may affect such processes. The cerebellar cortex forms well-organized structures: a highly specific and uniform arrangement of cells and microcircuitry. The cerebellum is one of the few sites in the brain where the pattern of intrinsic connections is known in considerable detail. These features make the cerebellum an ideal system to study the mechanisms of neural development and plasticity. Based on such advantage, many excellent works have been done at various levels from basic science to clinical disorders. On the other hand, although a number of hormone receptors are expressed in the cerebellum and cerebellar development and function are greatly influenced by hormonal status, relatively smaller number of studies may have been done on the role of hormonal signaling on development and plasticity of cerebellum. Among circulating hormones, a group of small lipophilic hormones such as steroids (corticosteroids, progesterone, androgens, and estrogens) and thyroid hormone may serve an important role in mediating environmental influences. Because of their chemical nature, these may be able to cross blood–brain barrier more easily than peptide hormones, although existence of specific transporters has been proposed (Suzuki and Abe 2008). Receptors for such lipophilic hormones are mainly located in the cell nucleus (nuclear receptor, NR), and represent the largest family of ligand-regulated transcription factors (Mangelsdorf et al. 1995). Schematic showing on the molecular mechanisms of NR-mediated transcription is shown in Fig. 16.1. NRs are widely distributed in the central nervous system (CNS) as well as other organs with a specific pattern of expression (Bookout et al. 2006). In the cerebellum, NRs are expressed in a specific temporal and spatial pattern (Qin et al. 2007). However, the role of these NRs on cerebellar function is not fully understood. The action of NRs is exerted by binding to their specific coregulators, such as coactivators and corepressors to regulate transcription of their target genes (Rosenfeld et al. 2006). Cofactors may alter chromatin structure by modulating histone acetylation/methylation or stabilization of basal transcriptional machinery. Genetically modified mouse lacking one of such coactivators shows aberrant cerebellar development (Nishihara 2008), indicating that temporal and spatial expression of these coregulators also play an important role in mediating NR signaling. Among lipophilic hormones, involvement of thyroid hormone and gonadal steroids on cerebellar development and plasticity has been well studied. Deficiency of thyroid hormone during postnatal development results in abnormal cerebellar morphogenesis in mammals including humans and rodents (Koibuchi et al. 2003). Estrogen and progesterone also play an important role in this process (Dean and

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Ligand (hormones)

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Fig. 16.1 Schematic figure showing the mechanisms of steroid/thyroid hormone receptor (nuclear receptor, NR)-mediated transcription. Nuclear receptor (NR) binds to specific nucleotide sequences known as hormone response element (HRE) as a homodimer, or heterodimer with retinoid X receptor. Various coregulators bind to NRs in a ligand dependent manner. Cofactors may alter chromatin structure by modulating histone acetylation/methylation or stabilization of basal transcriptional machinery (basal TFs). Usually, coactivator complex is recruited in the presence of ligand, whereas corepressor complex in the absence of ligand

McCarthy 2008). In addition to the supply from circulation, these gonadal steroids are produced locally within the Purkinje cell (Tsutsui 2006). These steroids may not only act through NRs, but also through membrane-associated receptor (Sakamoto et al. 2008). Although the functional significance of rapid action of estrogen and progestin has not yet been fully understood, these steroids may modulate neurotransmitter action such as GABA and NMDA-receptor mediated signaling (Belcher et al. 2005; Frye 2001). It should be noted that these thyroid/steroid hormonemediated pathway may be disrupted by prescribed drugs and environmental chemicals (Nguon et al. 2005; Darras 2008).

Thyroid Hormone and Cerebellar Development Molecular Mechanisms of Thyroid Hormone Action The thyroid hormone (L-triiodothyronine, T3; L-tetraiodothyronine, thyroxine, T4) binds to the thyroid hormone receptor (TR) and regulates transcription of target genes (Wu and Koenig 2000). TR genes are encoded in two genetic loci, termed as a and b, which are located at chromosomes 17 and 3 in human, and 11 and 14 in mouse, respectively (Lazar 1993). Each locus produces at least two proteins, which are termed as TRa1 and a2 (or c-erbAa2), and TRb1, TRb2, and TRb3. Furthermore, some introns have a weak promoter activity such as intron 7 of TRa gene. Thus, deletion of upstream exon may result in the expression of additional TR-related proteins, whose expression is limited under normal condition (Chassande 2003). So far, at least three additional TR-related proteins may be

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Fig. 16.2 Thyroid hormone receptor and its related proteins generated from a or b gene locus. Numbers indicate the number of amino acid. Hatched region with the same pattern indicates that the amino acid sequence is identical

generated. Such proteins, termed as TRDa1, TRDa2, and TRDb3, lack N-terminus and DNA-binding domain (DBD). TR and its related proteins generated from a or b gene locus are shown in Fig. 16.2. TR forms homodimer or heterodimer with retinoid X receptor (RXR) and binds to thyroid hormone response element (TRE) located at the promoter region of target genes. TR binds to TRE regardless of the presence of T3 and regulates transcription in a ligand dependent manner. In the presence of T3, it recruits protein complexes called coactivators to activate transcription, whereas in the absence of T3 it recruits corepressor complexes to repress transcription. It should be noted that, although TRa2 can bind to TRE, T3 cannot bind to it. TRa2 may act as endogenous inhibitor for other TRs. Because of this bi-directional function of TR, phenotype of TR gene knockout mouse is different from that of hypothyroid (thyroid hormone deficient) animals (Koibuchi 2009). Thus, TR gene knockout and hypothyroid animal models that are induced by thyroid dysgenesis or dyshormonogenesis are equally important to understand the mechanisms of thyroid hormone system in brain. Animal models to study thyroid hormone action in developing cerebellum are discussed more in detail later in this chapter.

The Effect of Thyroid Hormone in Developing Cerebellum As shown in Fig. 16.3, perinatal hypothyroidism dramatically affects cerebellar development. The growth and branching of Purkinje cell dendrites are greatly reduced by perinatal hypothyroidism (Nicholson and Altman 1972a). The number

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Fig. 16.3 The effect of altered thyroid hormone status during rat cerebellar development. Rats were rendered hypothyroid by administering antithyroid drug (propylthiouracil) starting from day 17 after conception. They were sacrificed at day 15 after birth. Compared with control rat (a, c, e), hypothyroid rat cerebellum is smaller (b). Retardation of dendrite arborization is evident in the Purkinje cell, as shown by immunohistochemistry for calbindin (d). Proliferation and migration of granule cell from the external granule cell layer (EGL) to the internal granule cell layer (IGL) is also retarded (f). Also note the decrease in the width of the molecular layer (ML) by perinatal hypothyroidism (arrows in e, f)

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of synapses between Purkinje cell dendrite and granule cell axons is decreased (Nicholson and Altman 1972a, b). The disappearance of the external granule cell layer (EGL) and migration of granule cells into the internal granule cell layer (IGL) are delayed (Nicholson and Altman 1972c). Myelination is also delayed (Bala´zs et al. 1971). Furthermore, synaptic connection among cerebellar neurons and afferent neuronal fibers from other brain region is also affected (Hajo´s et al. 1973). Such abnormal development cannot be rescued unless TH is replaced within first 2 weeks of postnatal life in rodents (Koibuchi et al. 2003). Recently, a dispersed primary culture system from rat newborn cerebellum has been developed (Ibhazehiebo et al. 2011a). By using this system, the effect of T4 treatment on Purkinje cell dendrite arborization was studied. As shown in Fig. 16.4, T4 treatment for 14 days dramatically induces an increase in Purkinje cell dendritic areas. Thyroid hormone can also induce an increase in neurite extension in purified cerebellar granule cell aggregate culture (Ibhazehiebo et al. 2011b). These findings indicate that thyroid hormone acts directly to cerebellar cells to promote development. Thyroid hormone crosses the blood–brain barrier or blood–cerebrospinal fluid barrier through its specific transporters such as organic anion transporter (Oatp) – 1c1 and monocarboxylate transporter (MCT) – 8 (Bernal 2005). T4 seems to be crossing such barriers more easily than T3 (Calvo et al. 1990). Then, T4 is taken up by astrocyte or tanycyte, in which T4 is converted to T3, an active form of thyroid hormone, by type 2 iodothyronine deiodinase, which is predominantly expressed in these cell types (Guadan˜o-Ferraz et al. 1997). T3 is then transferred to neuron or oligodendrocyte and binds to TR. T3 in the neuron/oligodendrocyte is converted to T2 for inactivation by type 3 iodothyronine deiodinase, which is predominantly present in neuronal cells (Tu et al. 1999). In human fetal cerebellum, T3 levels remain low during the first 20 weeks of pregnancy, followed by a gradual increase until birth (Kester et al. 2004). Such change may not because of the change in circulating thyroid hormone levels, but because of the differential expression of type 2 and 3 deiodinases. In rat, type 3 deiodinase expression is relatively high during fetal life, whereas type 2 deiodinase activity keeps increasing until early postnatal life (Bates et al. 1999). Such differential activities in deiodinases are consistent with the finding showing the change in T3 levels in the cerebellum. Such pattern of changes in T3 content and deiodinase activities in the cerebellum is greatly different from other brain regions, that is, the cerebral cortex, in which a striking increase in T3 content is seen during the first trimester with a very low level of type 3 deiodinase in human (Kester et al. 2004). Such difference may reflect the temporal difference in developmental period in each brain region. It has been generally accepted that the TR mediates most thyroid hormone actions in the brain, although non-genomic thyroid hormone actions have also been proposed. TRs are widely expressed in the developing and adult cerebellum (Bradley et al. 1992). TRb1 is predominantly expressed in the Purkinje cell, whereas TRa1 in other subset of cells. C-erbAa2 or TRa2, which is produced from TRa gene by alternative splicing, and which does not bind to thyroid hormone, is also widely expressed, although its physiological role has not yet been clarified.

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Fig. 16.4 The effect of thyroid hormone (T4) on Purkinje cell development in rat primary cerebellar culture. Newborn rats were sacrificed on postnatal day 2 to dissect out the cerebellum. Dispersed cells were plated and cultured for 14 days with indicated amount of T4. Immunocytochemistry was performed using anti-calbindin antibody. (a) Photomicrographs of Purkinje cells in culture; (b) Quantitative analysis for the dendritic area of Purkinje cells in culture

Although TRs are expressed widely in the developing cerebellum, and their levels are not decreased in adult, thyroid hormone regulates many thyroid hormoneresponsive genes only during the first 2 weeks of postnatal life in rodents. For example, the expression of neurotrophin (NT)-3 gene is regulated only during such period (Koibuchi et al. 2001). Not only TR but also cofactors such as steroid receptor coactivator (SRC)-1 is also strongly expressed in the adult and developing cerebellum (Martinez de Arrieta et al. 2000; Yousefi et al. 2005). Disruption of SRC-1 results in abnormal cerebellar development similar to those seen in hypothyroid animals (Nishihara et al. 2003). SRC-1 is predominantly expressed in the

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Purkinje cell and the IGL, whereas only cells located in the premigratory zone express SRC-1 in the EGL (Yousefi et al. 2005). These results are consistent with previous data showing that thyroid hormone is insensitive to proliferating granule cells (Messer et al. 1984). Furthermore, cerebellar SRC-1 protein levels were greatest at P15, when thyroid hormone most strongly affects cerebellar development (Yousefi et al. 2005). These results indicate that the change in expression of coactivators may play an important role in TH sensitivity in the cerebellum. Indeed, not only SRC-1, but also other brain-specific coregulators that are expressed in the developing cerebellum may also be involved. Some of such candidates are hairless (Thompson and Bottcher 1997). Furthermore, developmental alteration of DNA methylation may also play a role in altering thyroid hormone sensitivity. These epigenetic processes may control not only TR action, but also other NR action, which also has a distinct critical period controlling cerebellar development. In addition to coactivators/corepressors, TR may interact with other nuclear receptors and regulate gene expression. One such example is retinoic acid–related orphan receptor (ROR) a, which is also a member of steroid/thyroid receptor superfamily. It is strongly expressed in the Purkinje cell and plays a critical role in its development. Cerebellar phenotype and alteration of neurotrophin expression of natural mutant mouse (staggerer) harboring RORa mutation is similar to that of hypothyroid mouse (Qiu et al. 2007), although its thyroid function is normal, indicating that RORa may be involved in thyroid hormone-regulated gene expression in the developing cerebellum. In fact, thyroid hormone regulates RORa expression during first postnatal 2 weeks (Koibuchi et al. 2001), indicating that thyroid hormone may alter the gene expression critical for cerebellar development through regulation of RORa. Furthermore, RORa augments TR-mediated transcription, whereas staggerer type mutant RORa does not have such action (Qiu et al. 2007). A recent study has shown that RORa may directly interact with TR without binding to TRE. DNA-binding domain of RORa may play a role in such interaction (Qiu et al. 2009). These results indicate that RORa is required for full function of TR in the developing cerebellum.

Animal Models to Study Thyroid Hormone Action in Developing Cerebellum The cerebellum is the most commonly used brain region to study the mechanisms of thyroid hormone action on brain development. Since the development of rodent cerebellum occurs mostly postnatal, perinatal hypothyroidism causes various cerebellar abnormalities, as discussed above. Perinatal hypothyroidism can be easily induced by administering antithyroid drugs such as propylthiouracil and methimazole (methylmercaptoimidazole) (Koibuchi 2009). These drugs inhibit synthesis of thyroid hormone by inhibiting thyroid peroxidase activity. In addition to drug-induced hypothyroid animal models, many mutant or gene-modified animal models showing congenital hypothyroidism have been reported, some of which

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have been used to study the thyroid hormone action in the cerebellum (Koibuchi 2009). One example is the pax8 gene knockout mouse (Poguet et al. 2003). Pax8 is essential for thyroid follicular cell differentiation and thus its knockout mouse shows a severe hypothyroidism. Morphological development and gene expression in cerebellum is greatly affected in this mouse. Regarding the TR gene knockout model, these animal models may not always be suitable to study thyroid hormone action in the cerebellum. As discussed above, TR has bi-directional actions of transcriptional regulation of target genes; without T3 it represses transcription, whereas it activates transcription with T3. Since TR deletion abolishes the repressive action of TR, phenotypes of TR knockout mice are greatly different from those of mice harboring low thyroid hormone level. To study the role of TR on organ development and function, however, TR knockout mice are essential. Another issue that may be considered to generate TR knockout mouse is that some introns have a weak promoter activity such as intron 7 of TRa gene. Thus, deletion of upstream exon may result in the expression of additional TR-related proteins, whose expression may be limited under normal condition (Chassande 2003). As discussed above, at least three additional TR-related proteins, TRDa1, TRDa2, and TRDb3, may be generated. Thus, phenotypes of TR knockout mice may be due to combination of deletion of a specific TR with overexpression of other TR species. Table 16.1 shows the list of TR knockout mice. Possible remaining TR proteins in each animal are also indicated. TRa1 deleted mice are also reported to show a limited alteration of behavior and neural circuit (Guadan˜o-Ferraz et al. 2003). However, their cerebellar phenotype appeared to be normal except for aberrant maturation of astrocytes (Morte et al. 2004). More strikingly, deletion of TRa1 prevents structural alteration of the cerebellum in hypothyroidism that was induced by methimazole and perchlorate treatment (Morte et al. 2002). These results indicate that abnormal cerebellar phenotype in thyroid dyshormonogenesis animal may be due to dominant negative action of unliganded TRa proteins. On the other hand, TRa2 knockout mouse shows both hyper- and hypothyroid phenotype in an organ-specific manner (Salto´ et al. 2001). This may be due to elevated expression of TRa1 in this mouse. TRa1 expression in brain is also elevated, but cerebellar phenotype was not clear. Deletion of both TRa1 and TRa2 also shows only limited phenotype in cerebellum. However, apart from cerebellar phenotype, existence of TRDa1 and/or TRDa2 shows altered phenotype in various organs. When TRa1 and TRa2 are deleted but TRDa1 and TRDa2 expressions are not inhibited (TRa / ) (Fraichard et al. 1997), their phenotype is more severe than those of mice in which all TRa proteins are deleted (TRa0/0) (Gauthier et al. 2001; Macchia et al. 2001). The decrease in plasma thyroid hormone levels is greater, and more severe impairment of bone and intestine development is observed. More limited brain phenotype is observed in TRb knockout mice. While TRb1 is widely expressed including cerebellum, particularly in the Purkinje cell (Bradley et al. 1992), the expression of TRb2 is confined within pituitary, hypothalamus (TRH neuron), retina, and inner ear. TRb2 knockout mice show central resistance to thyroid hormone with elevated T3, T4, and TSH levels in serum (Abel et al. 1999).

Exon 3

TRβ-/-

TRα-/-: see above TRb-/-: exon 4–5

Gauthier 1999

Go¨the 1999

Forrest 1996 Sandhofer 1998

Abel 1999 Ng 2001

Gauthier 2001 Macchia 2001

Fraichard 1997

TRα0/0 TRβ-/- TRα 0/0: see above Gauthier 2001 TRβ-/-: exon 4–5

TRα-/- TRβ-/-

TRα and β TRα1-/- TRβ-/- See above

Exon 2

Exon5–intron7

TRα 0/0

TRβ TRβ2-/-

Exon 2

TRα-/-

Normal T3 with slightly reduced T4 level Prevention of hypothyroid phenotype in the cerebellum Overexpression of TRα1, inducing both hyper- (high body temperature, increased hear rate) and hypothyroid phenotype (increased body fat) Aberrant intestine and bone development

Representative phenotypes

α2, Δα2 All β All β

None

All α All β

β1, (β3, Δβ3) All α All α

High T3 and T4 levels due to high TSH level Growth retardation. Abnormal bone maturation Δα1, Δα2 Aberrant intestine and bone development (more severe than TRα-/-) Eleveted TSH, T3, and T4 levels (more severe than TRβ-/-) Reduced body temperature and bone maturation (more severe than TRα-/-) Aberrant auditory function (more severe than TRβ-/-) Aberrant intestine development (milder than TRα-/-, or TRα-/- TRβ-/-)

Central resistance to thyroid hormone levels Elevated TSH, T3, and T4 levels Selective loss of M-cone in retina Central resistance to thyroid hormone Elevated TSH, T3, and T4 levels Aberrant auditory functional development

Δα1, Δα2, all β All β Aberrant intestine and bone development, but the phenotype is less severe than those in TRα-/-

α1, Δα1 α1, α2

All β

β2

All α

α1, α2

Table 16.1. Thyroid hormone receptor (TR) gene knockout mouse models Deleted Remained Targeted gene Targeted exon References TRs TRs TRα TRα1-/Exon 9 Guadan˜o-Ferraz 2003 α1, α2, Δα2, Morte 2002, 2004 Δα1 all β Exon 10 Salto´ 2001 α2, α1, Δα1, TRα2-/Δα2 all β

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Furthermore, this deletion causes a selective loss of M-cones in retina (Ng et al. 2001). However, abnormal brain phenotype seems to be confined within hypothalamus, and any change in cerebellar phenotype has not been reported. In TRb knockout mouse, on the other hand, it shows aberrant development of auditory function in addition to central hypothyroidism (Forrest et al. 1996). However, although TRb is strongly expressed in the Purkinje cell, its deletion does not induce any alteration of thyroid hormone-responsive genes in cerebellum (Sandhofer et al. 1998). In case of TRa and b double knockout, since one receptor cannot substitute the function for the other, their phenotypes are more severe than those of single gene knockout. In TRa1 / TRb / mice, delayed general growth and aberrant bone maturation, which are not seen in each single knockout mouse, are observed (G€othe et al. 1999). In TRa / TRb / mice, aberrant intestinal development, which is seen in TRa1 / , and high T3, T4, and TSH levels, which is seen in TRb / , are observed, both of which are more severe than those of single knockout mice (Gauthier et al. 1999). However, in TRa0/0TRb / mice, while low body temperature and abnormal auditory function, which are more severe than those of TRa0/0 or TRb / , are seen, aberrant intestinal development is milder than those of TRa / TRb / or TRa1 / (Gauthier et al. 2001). These results indicate the possible contribution of TRa variants (Da1 and/or Da2) in generating differential phenotypes. Alteration of brain development of these double knockout mice has not yet been studied in detail. In addition to TR knockout mice, several knockin mice harboring mutant TRs have been generated (Hashimoto et al. 2001; Itoh et al. 2001). Such animals are considered as a model for human syndrome of resistance to thyroid hormone (RTH), which is characterized as reduced action of thyroid hormone in thyroid hormone target tissue (Refetoff et al. 1993). In addition to elevated serum levels of T3 and T4 with non-suppressed thyrotropin (TSH), many patients show clinical features related to neurological disorders such as hyperactivity and learning disability. The majority of patients harbor a mutation in the TRb gene. Their phenotypes are similar to those seen in human RTH patient such as elevated levels of T3 and T4 with unsuppressed TSH levels in serum, delayed general growth, and goiter. Furthermore, such animals show an aberrant cerebellar development similar to those seen in congenital hypothyroid animals (Hashimoto et al. 2001). This mouse shows decreased arborization of Purkinje cell dendrite with aberrant locomotor activity and decreased expression of thyroid hormone-responsive genes in cerebellum. In addition, although RTH patient harboring mutated TRa gene has not yet been identified, knockin mice harboring mutant TRa have also been generated to examine the involvement of unliganded TRa on development and functional maintenance of target organ including the cerebellum (Itoh et al. 2001). Compared to TRb mutated mice, their neurological phenotype is more severe. This tendency is evident when the phenotypes of mice harboring the same point mutation in TRa and b are compared (Itoh et al. 2001). Mutant TRa knockin mice show various aberrant cerebellar developments similar to those seen in congenital hypothyroid animals. These results indicate that TRa may be more involved in regulating cerebellar gene expression than TRb.

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Steroid Hormones and Cerebellar Development General Overview Adrenal and gonadal steroid hormones are known as stress and sex hormones, respectively, and both play important roles in the development of the CNS. They affect various developmental events of neurons, such as survival, differentiation, and remodeling of axons and dendrites. These effects are associated with brain organization, sexual differentiation, and stress responses. Steroid hormones bind cognate ligand-activated receptors, members of the steroid/thyroid superfamily of nuclear receptors, to modulate the transcription of hormone-responsive genes. This chapter summarizes the published data characterizing the actions of these hormones and the localization of the receptors in the developing cerebellum. Most of the information presented here is from the studies of rodents.

Adrenal Steroid Hormones and Cerebellar Development Mineralocorticoids and glucocorticoids are major adrenal steroid hormones synthesized in the adrenal cortex. Mineralocorticoids play roles in maintaining sodium and potassium levels, and glucocorticoids are involved in the stress response and in regulating carbohydrate metabolism. Their levels are controlled via the hypothalamo-pituitary-adrenal (HPA) axis by pituitary adrenocorticotropic hormone and hypothalamic corticotropin-releasing factor. Most of the effects on the brain are mediated via binding to intracellular receptors, the glucocorticoid receptor (GR) and mineralocorticoid receptor (MR) (Rashid and Lewis 2005). Regulation at the genomic level is thought to be responsible for slow and long-lasting effects, such as the actions of corticosteroid hormones on neurogenesis, neuronal morphology, and function in response to chronic stress, while rapid effects that respond within minutes are regulated by non-genomic action (Evanson et al. 2010). Recently, membraneassociated glucocorticoid and mineralocorticoid receptors have been suggested to be involved in the mediation of non-genomic rapid effects (Prager and Johnson 2009). GRs are expressed widely in the adult brain, with highest levels in the hippocampus and hypothalamic paraventricular nucleus, regions associated with activation of the HPA axis. Expression is first detected in the embryonic rat brain, and levels are high and similar in the developing cerebellum and hippocampus (Lawson et al. 1992). Prenatal glucocorticoids influence the development of Purkinje neurons (Rugerio-Vargas et al. 2007) and a recent study using chick embryos has reported the presence of GR mRNA in the embryonic cerebellum (Yamate et al. 2010). Furthermore, Yamate et al. also showed that effects after treatment with excess glucocorticoids are mediated via GRs and indirectly influence behavioral activity after hatching. In the mouse, GR immunoreactivity is intense in the external granule cell layer, the Purkinje cell layer, and white matter regions, and weak in the molecular layer and internal granule cell layer at postnatal day 7 (P7). These results suggest that glucocorticoids exert actions on cellular differentiation in the developing cerebellum to induce multiple changes in peripheral responses and brain function.

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It has been suggested that steroid hormones can affect neuronal susceptibility to different types of insults, including oxidative stress. In rats, cortisone treatment during prenatal (Velazquez and Romano 1987) and postnatal (Bohn and Lauder 1980) development resulted in a decreased number of cerebellar granule cells. Ahlbom et al. (2000) further showed that cerebellar granule cells exposed to high levels of glucocorticoids during the prenatal period become more sensitive to oxidative stress-induced cell death. A single glucocorticoid injection into the neonatal mouse can also induce apoptotic changes in the cerebellum, which results in permanent reductions in the number of neurons within the internal granule layer, suggesting that there is a limited period of vulnerability to glucocorticoids during development (Noguchi et al. 2008). Stressful experiences, such as maternal deprivation (MD) in the early postnatal period in rats, cause retardation in the development of cerebellar-dependent motor coordination and behavioral abnormalities similar to schizophrenia (Llorente et al. 2009). Rats that are maternally separated during P4-P14, which corresponds to the stress-hyporesponsive period characterized by reduced responsiveness of the HPA axis (Walker et al. 2001), have elevated levels of corticosterone at P13 and display behavioral alterations at adolescence and in adulthood (Viveros et al. 2009). These results support the possibility that abnormally increased levels of glucocorticoids due to neonatal stress during development are associated with structural abnormalities in the cerebellum. In addition, a recent study using chick embryos has suggested that apoptosis induced in immature granule neurons by corticosteroid may be mediated via a non-genomic mechanism (Aden et al. 2008). Based on these animal studies, these actions of glucocorticoids on the developing human cerebellum may be related to reduced birth weight and may also be responsible for emotional and behavioral problems observed in children whose mothers are treated during pregnancy with glucocorticoids for respiratory dysfunction (Noguchi et al. 2008). In addition, structural and functional cerebellar abnormalities, such as Purkinje cell loss, have been detected in many psychiatric disorders, such as autism and schizophrenia (Baldac¸ara et al. 2008; Martin et al. 2010). Interestingly, some effects of neonatal MD are different between the sexes. Effects on cellular degeneration and astrocyte proliferation in the cerebellum of neonatal MD rats were greater in males than in females (Llorente et al. 2009). This is attributed to males being more vulnerable to stress and/or a sex difference in the onset of sensitivity to stress. Furthermore, impairment of eyeblink conditioning was observed only in MD males. This is thought to be associated with the sexually dimorphic pattern of developmental GR expression in the posterior region of the cerebellar interpositus nucleus, a key region for eyeblink conditioning (Wilber and Wellman 2009).

Gonadal Hormones and Cerebellar Development Testosterone and estradiol (E2) are the two major gonadal steroids synthesized in the testis and the ovary. An important factor for the actions of these gonadal hormones is aromatase, an enzyme that is responsible for producing estrogens from androgens.

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In the developing brain, gonadal steroids are well known for their functions in the formation of brain structures that are different between males and females. During a limited perinatal period from late embryonic development through the first few days of postnatal life, testosterone is produced in males by the testis, which is differentiated from the indifferent gonad during early embryonic development directed by the testis-determining gene Sry (Koopman et al. 1991). In the brain, testosterone is converted to E2 by aromatase, whereas female ovaries, whose differentiation occurs postnatally, do not secrete E2 during this period. Thus, during the perinatal critical period, significantly higher levels of E2 in males compared to females are thought to act on male brain development. Regulatory mechanisms of E2 action have been reviewed in detail elsewhere (Wright et al. 2010); however, briefly, E2 regulates apoptosis to produce sexually dimorphic cell numbers, dendritic spine formation, neuronal migration, and synaptic organization in hypothalamic regions, most of which are key regions for regulating male and female sexual functions in the adult brain. In the developing rat hippocampus, gonadal steroids control sexually dimorphic neurogenesis (Zhang et al. 2008). Because of the lack of estrogen exposure during the perinatal period, the female brain was thought to develop without E2. However, studies using knockout mice of the aromatase gene have suggested that E2 produced by the ovary during a prepubertal period plays a role in the differentiation of the female-typical brain (Bakker and Brock 2010). The two major estrogen receptors (ERs), ERa and ERb, are expressed in hypothalamic regions, where development occurs differently between the sexes. A degree of overlapping distribution and colocalization occurs between ERa and ERb in some hypothalamic regions, such as the ventromedial hypothalamus, a key region in female reproduction, suggesting a possible interaction of the two proteins (Ikeda et al. 2003). Phenotypes of ERa and ERb knockout mice are different, suggesting distinctive roles (Kudwa et al. 2006). It has also been suggested that rapid plasma membrane-mediated non-genomic actions of estrogen, which possibly interacts with classical transcriptional regulation (genomic mechanisms), also plays an important physiological role in regulating the neural actions of estrogen in the brain (Raz et al. 2008; Vasudevan and Pfaff 2008; Kelly and Qiu 2010). These results suggest complicated interactions of ERa and ERb by various signaling regulatory mechanisms. Furthermore, it has recently been suggested that epigenetic alterations of DNA methylation patterns on the promoters of ER genes induced by estradiol during development is important for sexually dimorphic brain organization. This may be a mechanism by which estradiol regulates ER gene expression to produce permanent masculinization of the brain (Wilson and Westberry 2009). Androgens, directly acting on the androgen receptor (AR), are also thought to play a role in brain masculinization. This is based on studies of human patients with complete androgen insensitivity syndrome and on patients with mutations in the aromatase gene, as well as on studies of rodents with the testicular feminization mutation, which produces a nonfunctional AR (Zuloaga et al. 2008). Gonadal steroids also play an important role in the development of brain regions that are not significantly different between the sexes, including the cerebellum.

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Estradiol levels in the cerebellum are higher during the first postnatal week compared to later developmental stages (Sakamoto et al. 2003; Biamonte et al. 2009), and treatment of newborn rats with estradiol promotes dendritic growth and spine formation of Purkinje cells (Sakamoto et al. 2003). These studies indicate that estrogens play a role in cerebellar development. Higher expression levels of ERa compared to ERb in the hypothalamus indicate that ERa is predominantly associated with reproduction, whereas ERb is expressed in non-reproductive regions, such as in the cerebral cortex, hippocampus, cerebellum, and the dorsal raphe and is thought to play an important role in brain morphogenesis (Fan et al. 2010). Cerebellar development occurs at late embryonic and early postnatal stages in rodents. Both ERa and ERb are detected in an immature cerebellar granule cell line that was derived from late embryonic mouse cerebellum. Experiments with ER-subtype selective agonists and overexpression of ERa and ERb in this cell line have indicated that ERa, but not ERb, mediates E2 actions in the embryonic cerebellar granule cells (Gottfried-Blackmore et al. 2007). Quantitative RT-PCR studies have shown that both receptors are expressed in the cerebellum from birth through to adulthood but levels of ERb mRNA in neonatal rats are significantly higher than those of ERa (Ikeda and Nagai 2006). These studies have also shown that levels of ERa in the cerebellum during early postnatal development are significantly higher than in the adult; adult levels being only slightly higher than background. In contrast, no significant changes in the level of cerebellar ERb mRNA occurred during development or in adulthood (Ikeda and Nagai 2006). However, this pattern of ERb expression differs somewhat from that previously studied using western blot analysis, in which the level of ERb protein decreased transiently at P5 and P7 and then increased dramatically at P10 followed by a subsequent decrease (Jakab et al. 2001). By in situ hybridization and immunohistochemistry, ERa-positive Purkinje cells are abundant during early postnatal stages, but that these cells are reduced to only a few in adulthood. Since considerable outgrowth and differentiation of Purkinje cell dendrites proceeds during the first 3 postnatal weeks, these results suggest a possible role for ERa in Purkinje cell differentiation (Ikeda and Nagai 2006). ERb immunoreactivity was detected in various neurons, including Golgi cells, Purkinje cells, and basket cells, and the expression in each cell type occurs at different postnatal days. Jakab et al. (2001) detected additional ERbimmunoreactive cells, such as differentiating external granular layer cells and glial cells, although protein or mRNA for ERb in these cells could not be detected. The cellular localization of ERb may reflect its role in cellular differentiation and maturation in cerebellar development. As shown in Fig. 16.5, the different expression profiles of ERa and ERb suggest that E2 exerts its actions in a cell-type specific manner via binding to the two ERs, which play distinctive roles in cerebellar development. The possibility of rapid estrogen signaling mechanisms in the developing cerebellum has been recently discussed elsewhere (Belcher 2008). Although no sex differences in architecture have been reported in the normal development of the cerebellum, there is a clear sex difference of cerebellar pathology in several developmental diseases in humans and in the corresponding animal models. This has been linked to alterations of circulating gonadal steroids during critical

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Fig. 16.5 Localization of ERa and ERb proteins during postnatal cerebellar development. Representative images of immunohistochemistry for ERa (a–c) and ERb (d–f) in the cerebellum at postnatal day (P) 7, P14, and P21. Arrows indicate representative ERa-immunoreactive Purkinje cells. egl External germinal layer, gl granular layer, ml molecular layer, pl Purkinje cell layer. Scale bar, 50 mm

periods of cerebellar development (Dean and McCarthy 2008). Estrogens can protect neurons from oxidative stress-induced death (Dare´ et al. 2000) in several brain regions, including the cerebellum, by modifying neuronal vulnerability (Min˜ano et al. 2007). The antioxidative action of E2 may be relevant to sex differences in negative symptom scores of schizophrenia, scores being higher in males (Goldstein and Link 1988). A recent study of reeler mice, which have a mutation in the reelin gene, a candidate gene in neurodevelopmental disorders such as schizophrenia and autism (Fatemi 2001), indicates that E2, by interacting with reelin, acts on survival and maturation of Purkinje cells in female mice, and that the female-specific regulation of the reelin promoter by E2 might be epigenetic (Biamonte et al. 2009). Thus, estrogens are thought to play important roles in neuronal protection, in addition to their neurotrophic actions in the developing cerebellum. Cerebellar testosterone levels are transiently higher in males compared to females at P5 (Biamonte et al. 2009). An in vitro study showed the presence of AR protein in cultured P7 cerebellar granule cells, and demonstrated that cerebellar

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granule cells obtained from testosterone-treated neonatal rats are protected from cell death induced by oxidative stress via a mechanism mediated by the androgen receptor (Ahlbom et al. 2001). Autism spectrum disorders are more frequent in men than in women, and high fetal testosterone levels have been suggested to be involved (Knickmeyer and Baron-Cohen 2006). Several studies have reported that aromatase mRNA levels in the cerebellum during the early postnatal period are higher than in later stages (Sakamoto et al. 2003; Lavaque et al. 2006; Biamonte et al. 2009). Lavaque et al. (2006) detected a transient and marked elevation in aromatase mRNA levels at P10 in the male, but not female, cerebellum and they suggested a possible contribution of this gene to the local synthesis of estrogen, which plays an important role in cerebellar development, in combination with estradiol derived from the gonads. Higher vulnerability of males to MD stress might be associated with male-specific induction of aromatase, although the mechanisms for this remain to be elucidated.

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Min˜ano A, Cerbo´n MA, Xifro´ X (2007) 17beta-estradiol does not protect cerebellar granule cells from excitotoxicity or apoptosis. J Neurochem 102:354–364 Morte B, Manzano J, Scanlan T et al (2002) Deletion of the thyroid hormone receptor alpha 1 prevents the structural alterations of the cerebellum induced by hypothyroidism. Proc Natl Acad Sci USA 99:3985–3989 Morte B, Manzano J, Scanlan TS et al (2004) Aberrant maturation of astrocytes in thyroid hormone receptor alpha 1 knockout mice reveals an interplay between thyroid hormone receptor isoforms. Endocrinology 145:1386–1391 Ng L, Hurley JB, Dierks B et al (2001) A thyroid hormone receptor that is required for the development of green cone photoreceptors. Nat Genet 27:94–98 Nguon K, Ladd B, Baxter MG et al (2005) Sexual dimorphism in cerebellar structure, function, and response to environmental perturbations. Prog Brain Res 148:199–212 Nicholson JL, Altman J (1972a) The effects of early hypo- and hyperthyroidism on the development of the rat cerebellar cortex. II. Synaptogenesis in the molecular layer. Brain Res 44:25–36 Nicholson JL, Altman J (1972b) Synaptogenesis in the rat cerebellum: effects of early hypo- and hyperthyroidism. Science 176:530–532 Nicholson JL, Altman J (1972c) The effects of early hypo- and hyperthyroidism on development of rat cerebellar cortex. I. Cell proliferation and differentiation. Brain Res 44:13–23 Nishihara E (2008) An overview of nuclear receptor coregulators involved in cerebellar development. Cerebellum 7:48–59 Nishihara E, Yoshida-Komiya H, Chan CS et al (2003) SRC-1 null mice exhibit moderate motor dysfunction and delayed development of cerebellar Purkinje cells. J Neurosci 23:213–222 Noguchi KK, Walls KC, Wozniak DF et al (2008) Acute neonatal glucocorticoid exposure produces selective and rapid cerebellar neural progenitor cell apoptotic death. Cell Death Differ 15:1582–1592 Poguet AL, Legrand C, Feng X et al (2003) Microarray analysis of knockout mice identifies cyclin D2 as a possible mediator for the action of thyroid hormone during the postnatal development of the cerebellum. Dev Biol 254:188–199 Prager EM, Johnson LR (2009) Stress at the synapse: signal transduction mechanisms of adrenal steroids at neuronal membranes. Sci Signal 2:re5 Qin J, Suh JM, Kim BJ et al (2007) The expression pattern of nuclear receptors during cerebellar development. Dev Dyn 236:810–820 Qiu C-H, Shimokawa N, Iwasaki T et al (2007) Alteration of cerebellar neurotrophin messenger ribonucleic acids and the lack of thyroid hormone receptor augmentation by staggerer- type retinoic acid receptor-related orphan receptor-a mutation. Endocrinology 148:1745–1753 Qiu C-H, Miyazaki W, Iwasaki T et al (2009) Retinoic Acid receptor-related orphan receptor alpha-enhanced thyroid hormone receptor-mediated transcription requires its ligand binding domain which is not, by itself, sufficient: possible direct interaction of two receptors. Thyroid 19:893–898 Rashid S, Lewis GF (2005) The mechanisms of differential glucocorticoid and mineralocorticoid action in the brain and peripheral tissues. Clin Biochem 38:401–409 Raz L, Khan MM, Mahesh VB et al (2008) Rapid estrogen signaling in the brain. Neurosignals 16:140–153 Refetoff S, Weiss RE, Usala SJ (1993) The syndromes of resistance to thyroid hormone. Endocr Rev 14:348–399 Rosenfeld MG, Lunyak VV, Glass CK (2006) Sensors and signals: a coactivator/corepressor/ epigenetic code for integrating signal-dependent programs of transcriptional response. Genes Dev 20:1405–1428 Rugerio-Vargas C, Ramı´rez-Escoto M, DelaRosa-Rugerio C et al (2007) Prenatal corticosterone influences the trajectory of neuronal development, delaying or accelerating aspects of the Purkinje cell differentiation. Histol Histopathol 22:963–969 Sakamoto H, Mezaki Y, Shikimi H et al (2003) Dendritic growth and spine formation in response to estrogen in the developing Purkinje cell. Endocrinology 144:4466–4477

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Sakamoto H, Ukena K, Kawata M et al (2008) Expression, localization and possible actions of 25Dx, a membrane-associated putative progesterone-binding protein, in the developing Purkinje cell of the cerebellum: a new insight into the biosynthesis, metabolism and multiple actions of progesterone as a neurosteroid. Cerebellum 7:18–25 Salto´ C, Kindblom JM, Johansson C et al (2001) Ablation of TRa2 and a concomitant overexpression of alpha1 yields a mixed hypo- and hyperthyroid phenotype in mice. Mol Endocrinol 15:2115–2128 Sandhofer C, Schwartz HL, Mariash CN et al (1998) Beta receptor isoforms are not essential for thyroid hormone-dependent acceleration of PCP-2 and myelin basic protein gene expression in the developing brains of neonatal mice. Mol Cell Endocrinol 137:109–115 Suzuki T, Abe T (2008) Thyroid hormone transporters in the brain. Cerebellum 7:75–83 Thompson CC, Bottcher M (1997) The product of a thyroid hormone-responsive gene interacts with thyroid hormone receptors. Proc Natl Acad Sci USA 94:8527–8532 Tsutsui K (2006) Biosynthesis and organizing action of neurosteroids in the developing Purkinje cell. Cerebellum 5:89–96 Tu HM, Legradi G, Bartha T et al (1999) Regional expression of the type 3 iodothyronine deiodinase messenger ribonucleic acid in the rat central nervous system and its regulation by thyroid hormone. Endocrinology 140:784–790 Vasudevan N, Pfaff DW (2008) Non-genomic actions of estrogens and their interaction with genomic actions in the brain. Front Neuroendocrinol 29:238–257 Velazquez PN, Romano MC (1987) Corticosterone therapy during gestation: effects on the development of rat cerebellum. Int J Dev Neurosci 5:189–194 Viveros MP, Llorente R, Lo´pez-Gallardo M et al (2009) Sex-dependent alterations in response to maternal deprivation in rats. Psychoneuroendocrinology 34(Suppl 1):S217–226 Walker CD, Kanand JS, Plotsky PM (2001) Development of the hypothalamic–pituitary–adrenal axis and the stress response. In: McEwen BS (ed) Handbook of physiology: coping with the environment. Oxford University Press, New York Wilber AA, Wellman CL (2009) Neonatal maternal separation alters the development of glucocorticoid receptor expression in the interpositus nucleus of the cerebellum. Int J Dev Neurosci 27:649–654 Wilson ME, Westberry JM (2009) Regulation of oestrogen receptor gene expression: new insights and novel mechanisms. J Neuroendocrinol 21:238–242 Wright CL, Schwarz JS, Dean SL et al (2010) Cellular mechanisms of estradiol-mediated sexual differentiation of the brain. Trends Endocrinol Metab 21:553–561 Wu Y, Koenig RJ (2000) Gene regulation by thyroid hormone. Trends Endocrinol Metab 11:207–211 Yamate S, Nishigori H, Kishimoto S et al (2010) Effects of glucocorticoid on brain acetylcholinesterase of developing chick embryos. J Obstet Gynaecol Res 36:11–18 Yousefi B, Jingu H, Ohta M et al (2005) Postnatal changes of steroid receptor coactivator-1 immunoreactivity in rat cerebellar cortex. Thyroid 15:314–319 Zhang JM, Konkle AT, Zup SL et al (2008) Impact of sex and hormones on new cells in the developing rat hippocampus: a novel source of sex dimorphism? Eur J Neurosci 27:791–800 Zuloaga DG, Puts DA, Jordan CL et al (2008) The role of androgen receptors in the masculinization of brain and behavior: what we’ve learned from the testicular feminization mutation. Horm Behav 53:613–626

Section 2 Anatomy, Connections and Neuroimaging of the Cerebellum

Vascular Supply and Territories of the Cerebellum

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Louis Caplan

Overview Within the posterior circulation, brain and vascular structures are characterized as involving the proximal, middle, and distal posterior circulation territories (Caplan 1996; Caplan 2000; Caplan et al. 2004, 2005; Chaves et al. 1994; Savitz and Caplan 2005). The proximal intracranial posterior circulation territory includes regions supplied by the intracranial vertebral arteries (ICVAs) – the medulla oblongata and the posterior inferior cerebellar artery (PICA)-supplied region of the cerebellum. The ICVAs join at the medullo-pontine junction to form the basilar artery (BA). The middle intracranial posterior circulation territory includes the portion of the brain supplied by the BA up to its superior cerebellar artery (SCA) branches – the pons and the anterior inferior cerebellar artery (AICA)-supplied portions of the cerebellum. The BA divides to form the two posterior cerebral arteries (PCAs) at the junction between the pons and the midbrain, just beyond the origins of the superior cerebellar arteries (SCAs). The distal intracranial posterior circulation territory includes all of the territory supplied by the rostral BA and its SCA, PCA, and their penetrating artery branches – midbrain, thalamus, SCA-supplied cerebellum, and PCA territories. This distribution is shown diagrammatically in Fig. 17.1. The three surfaces of the cerebellum are: tentorial (or superior) facing the tentorium cerebelli, petrosal facing toward the petrous bone, and suboccipital facing the suboccipital bone located between the lateral and sigmoid dural sinuses

L. Caplan Department of Neurology, Beth Israel Deaconess Medical Center, Palmer 127 West Campus, Boston, MA, 02215, USA e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 343 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_17, # Springer Science+Business Media Dordrecht 2013

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Fig. 17.1 Schema of the proximal, middle, and distal intracranial territories of the vertebrobasilar arterial system (Drawn by Laurel Cook-Lowe, modeled after a figure in Duvernoy (1978), p. 41)

(Lister et al. 1982). The PICAs encircle the medulla and supply the suboccipital cerebellar surface; the AICAs course around the pons and supply the petrosal surface of the cerebellum, and the SCAs encircle the midbrain and supply the tentorial, superior surface of the cerebellum (Lister et al. 1982). The arteries to the cerebellum are distributed rostrocaudally so that the posterior inferior cerebellar arteries (PICAs) arise from the ICVAs, the anterior inferior

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cerebellar arteries (AICAs) arise from the BA, and the most rostral arteries, the SCAs, arise near the BA bifurcation (Fig. 17.2). The PICAs and the SCAs, the two largest arterial pairs have medial branches that supply mostly the vermian and paravermian portions of their respective regions of the cerebellum, and lateral branches which supply the cerebellar hemispheres. Infarcts in the cerebellum are often limited to the territory of one of these branches, e.g., medial PICA (mPICA), lateral SCA (lSCA), etc. These cerebellar branch territory infarcts correspond to functional regions such as the inferior vermis or superior lateral neocerebellum. The AICAs, in contrast, supply only a small part of the anterior inferior cerebellum and the flocculus, but their major supply is to the lateral pontine tegmentum and the brachium pontis. The AICAs do not divide into medial and lateral major cerebellar branches but give off twigs to various structures.

Fig. 17.2 Schematic diagram of the cerebellar arteries. 1 Superior cerebellar artery (SCA); 2 medial branch of the SCA; 3 lateral branch of the SCA; 4 anterior inferior cerebellar artery (AICA); 5 posterior inferior cerebellar artery (PICA); 6 medial branch of PICA; 7 lateral branch of PICA; 8 basilar artery; 9 vertebral artery (From Amarenco (1991))

Fig. 17.3 Sketch showing course and branching of the posterior inferior cerebellar artery (PICA). 1 PICA; 2 lateral branch of PICA; 3 medial branch of PICA; 4 cerebellar hemisphere; 5 cerebellar vermis; 6 cerebellar tonsil (Reproduced with permission from Amarenco P, Hauw J-J, Caplan LR. Cerebellar infarctions in Handbook of Cerebellar disease, New York, Marcel Dekker, 1993;251–290)

346 Fig. 17.4 The supply zone of PICA (Reproduced with permission from Amarenco P. The spectrum of Cerebellar infarctions. Neurology 1991;41:973–979)

Fig. 17.5 The supply zone of the medial branch of PICA (Reproduced with permission from Amarenco P. The spectrum of Cerebellar infarctions. Neurology 1991;41:973–979)

L. Caplan

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Posterior Inferior Cerebellar Arteries (PICAs) The PICAs usually originate from the ICVAs about 2 cm below the origin of the basilar artery, and, on average, about 8.6 mm above the foramen magnum (Marinkovic et al. 1995). The site of origin, however, varies from 14 mm below the foramen magnum to 26 mm above the foramen magnum (Marinkovic et al. 1995). About 10% arise from the basilar artery (Amarenco and Hauw 1989). Size varies; the diameters varied between .58 and 2.10 mm in one analysis (Amarenco and Hauw 1989). Some ICVAs end in PICA, and PICA can be absent in which case there usually is a large artery that arises from the proximal basilar artery that supplies both the PICA and AICA territories. Occasionally PICA is duplicated. After coursing laterally and downward to go around the lateral medulla (the lateral medullary segment), the PICAs make a cranially directed loop and ascend between the dorsal portion of the medulla and the caudal part of the cerebellar tonsil on that side (the tonsillo-medullary segment) (Lister et al. 1982; Marinkovic et al. 1995). They then make a second loop above the cranial portion of the tonsil and descend along the inferior vermis coursing between the inferior medullary velum and the rostral portion of the tonsil (the telovelotonsillar segment). Finally the artery becomes superficial and supplies branches to the tonsil, medulla, choroid plexus, and cerebellar cortex. Medial and lateral branches (mPICA and lPICA) arise from the main trunks (Fig. 17.3) at variable locations between the two PICA loops. mPICA supplies the inferior vermis including the nodulus, uvula, pyramis, tuber, and sometimes the declive and the medial portions of the semilunar lobule, gracile lobule, and the tonsil (Chaves et al. 1994; Amarenco and Hauw 1989; Amarenco et al. 1989, 1993; Amarenco 1991; Gilman et al. 1981; Duvernoy 1978). mPICA often sends a supply to the dorsal medulla. lPICA supplies the inferior two thirds of the biventer, most of the inferior portion of the semilunar

Fig. 17.6 The supply zone of the lateral branch of PICA (Reproduced with permission from Amarenco P. The spectrum of Cerebellar infarctions. Neurology 1991;41:973–979)

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Fig. 17.7 Right lateral medullary fossa. 1 Vertebral artery; 2 posterior inferior cerebellar artery (PICA); 3 accessory nerve; 4 lateral medullary fossa; 5 vagus nerve; 6 IV ventricle choroids plexus; 7 glossopharyngeal nerve; 8 vestibulocochlear nerve; 9 facial nerve; 10 lateral pontine vein; 11 pons; 12 abducens nerve; 13 olive; A, A0 rami arising from PICA; B rami arising from the vertebral artery to supply the lateral medulla; C rami arising from the basilar artery; C0 and D rami arising from AICA (From Duvernoy (1978), p. 70)

and the gracile lobules, and the anterolateral portion of the tonsil (Chaves et al. 1994; Amarenco and Hauw 1989; Amarenco et al. 1989, 1993; Amarenco 1991; Gilman et al. 1981; Duvernoy 1978). Figures 17.4, 17.5, and 17.6 show diagrammatically the supply territories of PICA, mPICA, and lPICA. The PICAs sometimes supply the deep cerebellar structures including the fastigial nuclei and may contribute to the supply of the dentate nuclei (Amarenco and Hauw 1989). Dye injections into SCA and PICA postmortem show SCA irrigation of dorsal portion of the dentate nucleus and PICA irrigation of ventral portion of the dentate nucleus (Schmahmann 2000).

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Vascular Supply and Territories of the Cerebellum

Fig. 17.8 MRI sagittal T2-weighted scan showing a PICA territory infarct (From Caplan (1996))

Fig. 17.9 Necropsy specimen showing an infarct in the territory of the medial branch of the posterior inferior cerebellar artery (From Amarenco P, Hauw J-J, Henin D et al. Les infarctus du territoire de l’arte`re ce´re´belleuse poste´roinfe´rieure; e´tude clinicopathologique de 28 cas. Rev Neurol 1989;145:277–286. With permission)

Fig. 17.10 Base of the brain at necropsy showing the origin of the anterior inferior cerebellar arteries

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Although many equate the Wallenberg syndrome with an occlusion of PICA causing infarction in the lateral medulla, PICA does not supply the lateral medullary tegmentum. This region is supplied by a group of parallel small arteries that originate directly from the intracranial vertebral artery and pass through the lateral medullary fossa to supply the lateral medulla (Fig. 17.7) (Duvernoy 1978). The medial branch of PICA supplies a small area in the dorsal medulla that includes vestibular nuclei and the dorsal motor nucleus of the vagus. Figure 17.8 is a sagittal section MRI showing a PICA infarct. Figure 17.9 shows a brain specimen with a medial PICA territory infarct.

Anterior Inferior Cerebellar Arteries (AICAs) The AICAs are nearly constant arteries but their origins, sizes, and supply zones vary greatly. They have the smallest territory of supply of any of the cerebellar arteries. The AICAs usually arise about 1 cm above the vertebrobasilar artery junction (Fig. 17.10), but they can sometimes arise directly from the ICVA, or from a common trunk with PICA. The internal auditory arteries are usually branches of the AICAs but in some individuals they arise directly from the basilar artery. Asymetry and reciprocal size relationship of AICA and PICA are common. In one study the diameters of AICA ranged from 0.38 to 1.8 mm (mean 1.1 mm) (Marinkovic et al. 1995). After arising from the basilar artery the AICAs travel

Fig. 17.11 Blood supply of the caudolateral pons from the Anterior Inferior Cerebellar Artery (AICA). The shaded area to the right is the supply of a lateral branch of AICA. A-basilar artery; B-medial pontine segment of AICA; C-loop segment of AICA around flocculus; D-paramedian basilar artery branches; E-brainstem branches of AICA; F-flocculus; G-posterior fossa cistern; H-IV ventricle; I-brachium pontis; J-medial lemniscus; K-lateral spinothalamic tract; L-motor nucleus of V; M-spinal tract and nucleus of V; N-main sensory nucleus of V; O-Vll and Vlll cranial nerves; P-internal acoustic meatus (From Perneczky A, Perneczky G, Tschabitscher M, Samec P. The relation between the caudolateral pontine syndrome and the anterior inferior cerebellar artery. Acta Neurochirurgica 1981;58:245–257. With permission)

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Fig. 17.12 Diagramatic depiction of the supply zones of the anterior inferior cerebellar arteries. (a) Shows the pontine territory. 1 flocculus, 2 brachium pontis, 3 restiform body, 4 brachium conjunctivum, 5 dentate nucleus, 6 vestibular nuclei, 7 spinothalamic tract, 8 central tegmental tract, 9 medial lemniscus, 10 cerebellar nodulus. (b) Shows the cerebellar supply on a lateral view of the cerebellum and (c) shows the supply on cut sections of the cerebellum and brainstem. The supply zones are shaded (Reproduced with permission from Amarenco P, Hauw J-J, Caplan LR. Cerebellar infarctions in Handbook of Cerebellar disease, New York, Marcel Dekker, 1993;251–290)

toward the cerebellopontine angle, passing below the Vth nerve, crossing the VIth nerve, and meeting the VIIth and VIIIth nerves at the cerebellopontine angle (Marinkovic et al. 1995; Amarenco and Hauw 1989, 1990a; Amarenco et al. 1993; Amarenco 1991; Gilman et al. 1981; Duvernoy 1978; Perneczky et al. 1981). After crossing the VIIIth nerve, the AICAs give rise to the internal auditory arteries and then divide into two branches. One branch courses laterally and inferiorly to supply the anterior inferior portion of the cerebellum on the petrosal surface.

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Fig. 17.13 Necropsy specimen (H and E stained) showing an anterior inferior cerebellar artery territory infarct

Fig. 17.14 Brain at necropsy showing the superior cerebellar arteries circling the midbrain and giving off branches

The other branch loops around the bundle made by the VIIth and VIIIth nerves, and supplies the flocculus, brachium pontis and the lateral part of the pons (Marinkovic et al. 1995; Amarenco and Hauw 1989, 1990a; Amarenco et al. 1993; Amarenco 1991; Perneczky et al. 1981). The internal auditory arteries supply the facial and vestibulocochlear nerves as well as the structures of the inner ear. Figure 17.11 is a schematic drawing of the AICA and its supply (Perneczky et al. 1981). Figure 17.12 shows the brainstem and cerebellar distribution of the AICA supply territory. Figure 17.13 is a necropsy specimen showing an AICA territory infarct at the level of the pons.

Superior Cerebellar Arteries (SCAs) The SCAs arise as the last pair of branches from the basilar artery just before the basilar artery bifurcates into the paired PCAs (Fig. 17.14). The third cranial nerves

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Fig. 17.15 Schematic diagram of the superior cerebellar artery (SCA) and its medial (mSCA) and lateral (lSCA) branches. The top branch is the mSCA and the lower branch is the lSCA (From Amarenco P, Roullet E, Goujon C et al. Infarction in the anterior rostral cerebellum (the territory of the lateral branch of the superior cerebellar artery). Neurology 1991;41:253–258. With permission)

Fig. 17.16 The SCA supply territories are shaded (From Amarenco and Hauw (1989))

run between the SCAs and the PCAs near the posterior communicating arteries. In about 15% of patients there are bifid SCAs. In one series the diameters ranged from 0.7 to 1.93 mm (mean 1.1 mm) (Marinkovic et al. 1995). The SCA encircles the brainstem close to or within the ponto-mesencephalic sulcus, just below the third

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Fig. 17.17 Supply zones of the superior cerebellar arteries. (a) Shows the superior pontine supply. 18 brachium conjunctivum, 20 lateral lemniscus, 21 medial lemniscus, 22 locus coeruleus, 23 mesencephalic tract of V, 24 spinothalamic tract, 25 cortico-tegmental tract, 26 decussation of IV nerve, 28 medial longitudinal fasciculus. (b) Shows an anteroposterior view and (c) a lateral view of the cerebellum (From Amarenco and Hauw (1989)) (Note that the numbers referred to here in the legend are not in the figure, and the numbers in the figure are not in the legend)

Fig. 17.18 Necropsy specimen showing SCA territory infarct in the rostral pons. The pontine tectum and a small part of the dorsolateral pontine tegmentum are involved (From Amarenco P, Hauw JJ. Cerebellar infarction in the territory of the superior cerebellar artery: a clinicopathologic study of 33 cases. Neurology 1990;40:1383–1390. With permission)

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nerve and just above the trigeminal nerve. While coursing around the midbrain, the SCAs give off branches that supply the brainstem including the superior portion of the lateral pontine tegmentum and the pontine and mesencephalic tectum. The SCAs have an early division within the cerebello-mesencephalic cistern where it divides into the mSCA and lSCA branches. Figure 17.15 shows the usual branching of the SCAs and the course of the lateral and medial branches. The mSCA branch extends more laterally than the mPICA. Occasionally these branches arise directly from the basilar artery and the SCAs. Both the major branches of the SCAs course toward the pedunculo-cerebellar sulcus and reach the superior and anterior aspects of the cerebellum above the horizontal fissure. The mSCAs mostly supply the superior portions of the vermis including the central, culmen, declive, and folium lobules; the lSCAs supply mostly the lateral portions of the cerebellar hemispheres including the anterior, simplex, and superior portion of the semilunar lobules. The SCAs also supply the cerebellar nuclei (dentate, fastigial, emboliform, and globose) as well as the bulk of the cerebellar white matter (Amarenco 1991; Amarenco and Hauw 1989, 1990b; Amarenco et al. 1991, 1993). Figures 17.16 and 17.17 show the cerebellar and brainstem supply territories of the SCA. Figure 17.18 is a necropsy specimen showing a large SCA territory infarct. Figure 17.19 shows three MRI scans that

a

b

c

Fig. 17.19 MRI T2-weighted scans (From Caplan (1996)). (a) Sagittal view showing SCA territory infarct. (b) Axial section showing small vermal cerebellar infarct in the territory of the medial branch of the superior cerebellar artery (mSCA). (c) Coronal section showing a bilateral SCA territory infarct appearing like “icing on a cake”

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illustrate the imaging distribution of various SCA territory infarcts. The distribution of the supply territories of the cerebellar arteries as found on CT and MRI scanning has been illustrated and reviewed (Savoiardo et al. 1987; Courchesne et al. 1989; Press et al. 1989; Press et al. 1990).

References Amarenco P (1991) The spectrum of cerebellar infarctions. Neurology 41:973–979 Amarenco P, Hauw J-J (1989) Anatomie des arteres cerebelleuses. Rev Neurol 145:267–276 Amarenco P, Hauw J-J (1990a) Cerebellar infarction in the territory of the anterior and inferior cerebellar artery. Brain 113:139–155 Amarenco P, Hauw JJ (1990b) Cerebellar infarction in the territory of the superior cerebellar artery: a clinicopathologic study of 33 cases. Neurology 40:1383–1390 Amarenco P, Hauw J-J, Henin D et al (1989) Les infarctus du territoire de l’arte`re ce´re´belleuse poste´ro-infe´rieure; e´tude clinico-pathologique de 28 cas. Rev Neurol 145:277–286 Amarenco P, Roullet E, Goujon C et al (1991) Infarction in the anterior rostral cerebellum (the territory of the lateral branch of the superior cerebellar artery). Neurology 41:253–258 Amarenco P, Hauw J-J, Caplan LR (1993) Cerebellar infarctions. In: Lechtenberg R (ed) Handbook of cerebellar diseases. Marcel Dekker, New York, pp 251–290 Caplan LR (1996) Posterior circulation disease: clinical findings, diagnosis, and management. Blackwell Scientific, Boston Caplan LR (2000) Posterior circulation ischemia: then, now, and tomorrow The Thomas Willis Lecture – 2000. Stroke 31:2011–2013 Caplan LR, Wityk RJ, Glass TA et al (2004) New England Medical Center Posterior Circulation Registry. Ann Neurol 56:389–398 Caplan LR, Wityk RJ, Pazdera L et al (2005) New England Medical Center posterior circulation stroke registry: II. Vascular lesions. J Clin Neurol 1:31–49 Chaves CJ, Caplan LR, Chung C-S, Amarenco P (1994) Cerebellar infarcts. In: Appel S (ed) Current neurology, vol 14. Mosby-Year Book, St. Louis, pp 143–177 Courchesne E, Press GA, Murakami J et al (1989) The cerebellum in sagittal plane-anatomic-MR correlation: 1. The vermis. AJNR Am J Neuroradiol 10:659–665 Duvernoy HM (1978) Human brainstem vessels. Springer, Berlin Gilman S, Bloedel J, Lechtenberg R (1981) Disorders of the cerebellum. FA Davis, Philadelphia Lister JR, Rhoton AL, Matsushima T, Peace DA (1982) Microsurgical anatomy of the posterior inferior cerebellar artery. Neurosurgery 10:170–199 Marinkovic S, Kovacevic M, Gibo H, Milisavljevic M, Bumbasirevic L (1995) The anatomical basis for the cerebellar infarcts. Surg Neurol 44:450–461 Perneczky A, Perneczky G, Tschabitscher M, Samec P (1981) The relation between the caudolateral pontine syndrome and the anterior inferior cerebellar artery. Acta Neurochir 58:245–257 Press GA, Murakami J, Courchesne E et al (1989) The cerebellum in sagittal plane-anatomic-MR correlation: 2. The cerebellar hemispheres. AJNR Am J Neuroradiol 10:667–676 Press GA, Murakami JW, Courchesne E et al (1990) The cerebellum: 3. Anatomic-MR correlation in the coronal plane. AJNR Am J Neuroradiol 11:41–50 Savitz SI, Caplan LR (2005) Current concepts: vertebrobasilar disease. N Engl J Med 352:2618–2626 Savoiardo M, Bracchi M, Passerini A, Visciani A (1987) The vascular territories in the cerebellum and brainstem: CT and MR study. AJNR Am J Neuroradiol 8:199–209 Schmahmann JD (2000) Cerebellum and brainstem. In: Toga A, Mazziotta J (eds) Brain mapping, the systems. Academic, San Diego, pp 207–259

Vestibulocerebellar Connections

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Neal H. Barmack and Vadim Yakhnitsa

Abstract

The vestibular system projects onto the cerebellum via three major pathways that are composed of primary and secondary vestibular mossy fiber afferents and tertiary vestibular climbing fibers. Vestibular primary afferent mossy fibers project to the ipsilateral uvula-nodulus (folia 9d and 10). Secondary vestibular mossy fibers originate from three of the five vestibular nuclei: medial, descending, and superior (MVN, DVN, and SVN). These mossy fibers terminate in the uvula-nodulus and flocculus. The MVN, DVN, and SVN receive convergent vestibular, optokinetic, and neck proprioceptive information. Vestibular tertiary afferents originate from two subnuclei of the inferior olive, the b-nucleus and dorsomedial cell column (DMCC), and send climbing fibers to the contralateral uvula-nodulus. The b-nucleus and DMCC receive direct projections from the parasolitary nucleus (Psol). The Psol, b-nucleus, and DMCC convey information to the cerebellum from the vertical semicircular canals and otoliths, but not from the horizontal semicircular canals. Climbing fiber projections are arrayed in sagittal zones, establishing a mediolateral map on the uvula-nodulus of the 180
of possible head angles during roll-tilt. Signals conveyed from climbing fibers are preeminent in modulating the discharge of both complex and simple spikes (CSs and SSs) in cerebellar Purkinje cells. This discharge is fed back onto a fraction of neurons in the dorsal aspect of the descending and medial vestibular nuclei as well as the prepositus hypoglossal nucleus. The vestibulocerebellum imposes a climbing fiber-constructed coordinate system on postural responses and permits adaptive guidance of movement.

N.H. Barmack (*) • V. Yakhnitsa Department of Physiology & Pharmacology, Oregon Health & Science University, 3181 SW Sam Jackson Park Road, 97239 Portland, OR, USA e-mail: [email protected], [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 357 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_18, # Springer Science+Business Media Dordrecht 2013

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Introduction The vestibulocerebellum participates in sensorimotor integration and motor control. While reflexive and reactive movements can be executed with or without an intact cerebellum, the cerebellum adds accuracy and modifiability to movements using sensory feedback and superimposing centrally generated commands (Morton and Bastian 2006). Cerebellar circuitry itself is modifiable (Eccles et al. 1967; Ito 1976, 2002). For this reason, the cerebellum attracts interest from both basic and clinical neurobiologists. Impaired cerebellar function can be the consequence of chronic alcoholism (Schapiro et al. 1984), genetic mutation (Zee et al. 1976; Falk et al. 1999; Koeppen 2005), immunological disorders (Hida et al. 1994; Sakai et al. 1995), and neural trauma. This chapter describes one of the key sensory inputs to the cerebellum, the vestibular system. It characterizes how the vestibular system interacts with the cerebellar circuitry in the uvula-nodulus. This interactive system is used as a model for understanding how cerebellar circuitry functions.

Vestibular Primary Afferent Cerebellum Projections: Mossy Fibers The peripheral vestibular apparatus consists of five end organs; three semicircular canals are orthogonally oriented and sense angular rotation in the horizontal, pitch, and roll planes. Two otoliths sense linear acceleration of gravity during roll-tilt (utricle) and pitch (saccule). Vestibular primary afferents that originate from each of the five ipsilateral vestibular end organs combine in the vestibular nerve. As it approaches the brainstem, the vestibular nerve divides into two fiber bundles containing axons of unequal thickness. The thicker axons enter the medulla passing into the vestibular complex where they terminate on the medial, descending, and superior vestibular nuclei (MVN, DVN, and SVN) as well as the parasolitary nucleus (Psol) (Fig. 18.1a, b) (Cajal 1911). The bundle with thinner axons ascends to the cerebellum after passing through the SVN and LVN. Vestibular primary afferent mossy fibers project to the ipsilateral nodulus (folium 10) and the ventral aspect of the uvula (folium 9d) (Fig. 18.1c). Collectively, vestibular primary afferents comprise the largest mossy fiber projection to the nodulus (Carpenter et al. 1972; Alley et al. 1975; Korte 1979; Carleton and Carpenter 1984; Kevetter and Perachio 1986; Gerrits et al. 1989; Barmack et al. 1993a; Purcell and Perachio 2001). Mossy fiber projections to the uvula-nodulus are diffuse. A single mossy fiber branches several times after it enters the cerebellum, and each branch may yield synaptic terminals over a distance of several mm. This branching pattern can be seen in single mossy fibers whose cell bodies in the lateral reticular nucleus (LRN) have been labeled with injections of biotinylated dextran amine (BDA). Mossy fibers from the LRN project bilaterally to several folia in the anterior lobe vermis (Fig. 18.3a) (Wu et al. 1999). Vestibular primary afferent mossy fibers project unilaterally to a distinct region of the cerebellum, the uvula-nodulus. However, the projection remains

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Fig. 18.1 Projection of vestibular primary afferents to the vestibular complex and to the cerebellum. (a) Vestibular primary afferents bifurcate as they enter the brainstem (modified from Cajal (1911)). (b) The terminal fields within the cat vestibular complex of five horizontal semicircular canal afferents, intra-axonally labeled with HRP, are illustrated. The terminals are mapped onto the vestibular complex (modified from Sato and Sasaki (1993)). (c) Vestibular primary afferents were labeled by a labyrinthine injection of the C-fragment of tetanus toxin. The toxin was transported to mossy fiber terminals in the granule cell layer of folia 9c and 10 in the rabbit. The arrow heads delimit the folia with dense reaction product. (d) Vestibular secondary afferents were labeled with an injection of WGA-HRP into the caudal medial and descending vestibular nuclei. As in the case of the primary afferent projection, secondary vestibular mossy fiber afferents terminate in folia 9d and 10 in the rabbit as indicated by arrow heads. Abbreviations: DVN, LVN, MVN, and SVN descending, lateral, medial, and superior vestibular nuclei, Psol parasolitary nucleus

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topographically diffuse. A single vestibular primary afferent mossy fiber not only projects onto multiple folia in the uvula-nodulus, it also has a wide mediolateral distribution within each folium (Maklad and Fritzsch 2003). Saccular and utricular afferents project mainly, but not exclusively, to the ventral uvula where they synapse on granule cells. Semicircular canal afferents project mainly, but again not exclusively, to the nodulus (Kevetter and Perachio 1986; Purcell and

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Perachio 2001; Newlands et al. 2002; Maklad and Fritzsch 2003; Newlands et al. 2003). Granule cell axons ascend through the Purkinje cell layer and then divide into parallel fibers that extend for distances of 3–6 mm in the molecular layer as they innervate Purkinje cell dendrites. Consequently, the spatial resolution of an already diffuse mossy fiber signal is further degraded as it is distributed by parallel fibers to 100–500 Purkinje cells (Brand et al. 1976; Mugnaini 1983; Pichitpornchai et al. 1994; Barmack and Yakhnitsa 2008). Even though the vestibular primary afferent mossy fiber projection is exclusively ipsilateral, the length of parallel fibers assures that Purkinje cells located within a single folium are likely to receive parallel fiber-mediated mossy fiber signals representing all vestibular end organs from both the ipsilateral and contralateral labyrinths. Ascending axons of granule cells could potentially reduce the dispersion contributed by parallel fibers by making preferential synapses on overlying Purkinje cells (Cohen and Yarom 1998; Gundappa-Sulur et al. 1999). However, synapses made by a single granule cell ascending axon are few (1,000). Current electrophysiological evidence demonstrates that the strength of parallel fiber synapses on Purkinje cells and the strength of synapses made by ascending axons are equivalent (Isope and Barbour 2002). Furthermore, there is no evidence to suggest that the ascending axons reaching a particular Purkinje cell convey signals from only a single modality or a single vestibular end organ. In the classic cerebellar homunculus, the vestibular system is preferentially represented in the uvula-nodulus. However, the homunculus merely characterizes the cerebellar region that receives vestibular signals as opposed to somatosensory or cutaneous signals from the limbs and trunk. It provides no explicit topography of these signals. In other words, vestibular primary afferent mossy fibers project to the uvula-nodulus as opposed to other cerebellar folia. However, this primary afferent mossy fiber projection provides no organizational specialization of vestibular information within the uvula-nodulus. Rather, vestibular mossy fiber signals are mixed to begin with and subsequently dispersed by parallel fiber projections (Isope and Barbour 2002; Sims and Hartell 2006).

Vestibular Secondary Afferent Mossy Fiber Cerebellar Projections from Vestibular Nuclei Secondary vestibular afferents originate from the vestibular complex that consists of five nuclear subgroups: MVN, DVN, LVN (also known as Dieter’s nucleus), SVN (Brodal and Pompeiano 1957; Brodal 1974), and the parasolitary nucleus (Psol) (Barmack et al. 1998). Nuclear subgroups are defined as part of the vestibular complex based on contiguity and sharing vestibular primary afferent projections – although the primary afferent projection to LVN is modest (Fig. 18.1b). Vestibular secondary afferents project bilaterally to the uvula-nodulus and flocculus from the caudal aspects of the DVN, MVN, and SVN (Brodal and Torvik 1957; Kotchabhakdi and Walberg 1978; Yamamoto 1979; Brodal and

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Brodal 1985; Thunnissen et al. 1989). WGA-HRP injections into the caudal MVN and DVN reveal that vestibular secondary afferent mossy fibers terminate in the granule cell layers of folia 9d and 10 in a pattern that is similar to that of vestibular primary afferents (Fig. 18.1c, d). However, the projection of vestibular secondary afferents is not restricted to the uvula-nodulus. The caudal aspects of the DVN and MVN project to several other cerebellar folia, including the anterior vermis, flocculus, and paraflocculus (Thunnissen et al. 1989; Epema et al. 1990; Barmack et al. 1992b). Immunohistochemical and radiochemical evidence shows that most, if not all, of these ascending projections are cholinergic (Tago et al. 1989; Barmack et al. 1992a, c). Two of the five vestibular nuclei, LVN and Psol, do not project to the cerebellum. LVN projects to the spinal cord. Psol projects to the inferior olive.

Tertiary Vestibular Climbing Fiber Projections to the Uvula-Nodulus The vestibular climbing fiber projection to the uvula nodulus originates from the contralateral b-nucleus and dorsomedial cell column (DMCC) of the inferior olive (Barmack et al. 1993b; Barmack 1996; Kaufman et al. 1996) (Fig. 18.2). In contrast to mossy fibers, climbing fibers terminate in narrow, functionally distinct, sagittal zones (Fig. 18.3b) (Barmack et al. 1993b; Voogd et al. 1996; Sugihara et al. 2001). The width of these zones ranges from 400 mm in mice to 800 mm in rabbits. Within these sagittal climbing fiber zones, all the climbing fibers share a common vestibular receptor identity. The b-nucleus and DMCC receive vestibular secondary afferent projections primarily from the ipsilateral Psol which, in turn, receives vestibular primary afferents from the ipsilateral labyrinth. The descending projection from the Psol to the ipsilateral b-nucleus and DMCC is GABAergic. Three interesting consequences emerge from the vestibular climbing fiber projection to the uvula-nodulus. First, the vestibular climbing fiber projection to the one side of the uvula-nodulus is controlled by climbing fibers whose activity is controlled primarily by contralateral labyrinth. Second, the activity of climbing fibers is controlled mostly by an inhibitory signal: the descending GABAergic signal from Psol. Third, it is possible to analyze separately the contributions of vestibular primary and secondary afferent mossy fibers from vestibular climbing fibers because mossy fiber signals arise primarily from the ipsilateral labyrinth, while climbing fiber signals arise from the contralateral labyrinth (Fig. 18.2b). The scheme of separate labyrinthine origins of vestibular mossy and climbing fiber projections is partially violated by the existence of a small descending pathway from the dorsal Y group to the contralateral b-nucleus and DMCC (Fig. 18.2b). This pathway is glutamatergic and excites contralateral olivary neurons (Kevetter and Perachio 1986; De Zeeuw et al. 1993; Wentzel et al. 1995; Blazquez et al. 2000). Neurons in each dorsal Y group receive both ipsilateral primary and bilateral secondary vestibular projections. By any logical standard, the dorsal Y group should be included as part of the vestibular complex. It is excluded

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by its location. Although it receives a vestibular primary afferent projection, it lies at the floor of the cerebellar nuclei, rather within the confines of the dorsal brainstem.

Climbing Fiber Zones Vestibular roll-tilt modulates the activity of Purkinje cell complex spikes (CSs) and simple spikes (SSs) in the uvula-nodulus. Climbing fiber projections are converted from the caudal-rostral organization of the inferior olive to a mediolateral organization in the uvula-nodulus (Sato and Barmack 1985). Pairs of vertical semicircular canals are represented in two distinct sagittal zones. The ipsilateral posteriorcontralateral anterior semicircular canals are represented in a medial zone. The ipsilateral anterior-contralateral posterior semicircular canals are represented in a lateral zone. Interposed between these two zones on the ventral surface of the nodulus is an optokinetic climbing fiber projection from the contralateral dorsal cap. This projection is excited by posterior ! anterior stimulation of the ipsilateral eye (Alley et al. 1975; Graf et al. 1988; Takeda and Maekawa 1989; Barmack and Shojaku 1995). Signals from the horizontal semicircular canals are not represented in the projection of climbing fibers from either the b-nucleus or DMCC to the uvula-nodulus although they are represented in the primary vestibular mossy fiber afferent projection.

Climbing Fiber Evoked Discharge in Purkinje Cells Cerebellar Purkinje cells have two action potentials. CSs are evoked by the multisynaptic contacts of a single climbing fiber as it wraps around the Purkinje cell proximal dendrite (Granit and Phillips 1956; Eccles et al. 1966a). CSs are longduration (2–8 ms), multipeaked action potentials that discharge at 0.1–5.0 imp/s. SSs are short-duration (0.75–1.25 ms), single-peaked action potentials that discharge at 20–60 imp/s. SSs reflect the excitatory drive of the more than 150,000 ä Fig. 18.2 Cerebellar and vestibular circuitry. (a) Diagram of cerebellar folium illustrates seven neuronal types. Inhibitory interneurons are indicated in red. (b) Vestibular primary and secondary afferent mossy fibers (mf) and tertiary climbing fibers (cf) project to uvula-nodulus. The bilateral secondary afferent projections are not illustrated. Solid lines indicate excitatory pathways. Red dashed lines indicate inhibitory pathways. Primary afferent mossy fibers project to the ipsilateral parasolitary nucleus (Psol), medial, descending, and superior vestibular nuclei (MVN, DVN, and SVN), Y group (Y), and granule cell layer of nodulus (green lines). Psol sends an inhibitory GABAergic projection to the ipsilateral b-nucleus and dorsomedial cell column (DMCC) (dashed red lines). Climbing fibers from the b-nucleus and DMCC project to contralateral nodulus (darkblue lines). Y group projects to contralateral dorsal cap (DC), b-nucleus, and DMCC (light-blue lines). Abbreviations: Ba basket cell, Bg Bergmann astrocyte, Fl flocculus, Gc granule cell, Go Golgi cell, IntP interpositus nucleus, LCN and MCN lateral and medial cerebellar nucleus, Lu Lugaro cell, NG2+ glia, Pc Purkinje cell, PFl paraflocculus, pf parallel fiber, Sc stellate cell, UBC unipolar brush cell, and 8n, vestibular nerve

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Fig. 18.3 Comparison of single mossy and climbing fiber axon terminals within the cerebellum. (a) A single mossy fiber in the left lateral reticular nucleus was injected with biotinylated dextran amine (BDA). It projected bilaterally to several folia in the anterior lobe vermis (modified from Wu et al. (1999)). (b) A single neuron in the medial accessory olive was injected with BDA. The labeled climbing fiber axon terminals of this cell projected in a narrow sagittal pattern onto the contralateral uvula-nodulus (modified from Sugihara et al. (2001)). Abbreviations: icp inferior cerebellar peduncle, IP nucleus interpositus, LCN lateral cerebellar nucleus, and MCN medial cerebellar nucleus

parallel fiber synapses on the Purkinje cell dendritic tree. The widely held assumption that mossy fibers modulate the higher discharge frequency of SSs, after a synapse on granule cells and distribution by parallel fibers to Purkinje cell dendrites, is echoed in textbooks (Ghez and Thach 2000), scholarly reviews

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(Apps and Garwicz 2005; Bloedel and Bracha 2009), and research reports (Ebner and Bloedel 1981; Armstrong and Edgley 1988; Nagao 1989; Kano et al. 1991; Lisberger et al. 1994; Walter and Khodakhah 2006). However, this assumption obscures the possible contributions of other factors that contribute to modulated SS discharge. Spontaneous SSs occur in isolated Purkinje cells in the absence of any parallel fiber input (Llina´s and Sugimori 1980; Raman and Bean 1999). Interneurons in the molecular layer – basket and stellate cells – exert inhibitory control of Purkinje cells (Andersen et al. 1964; Midtgaard 1992; Pouzat and Hestrin 1997; Szapiro and Barbour 2007). Golgi cells in the granule cell layer indirectly influence SS discharge by inhibiting the discharge of granule cells (Eccles et al. 1966b) (Fig. 18.2a). During sinusoidal vestibular roll-tilt, it is possible to record the modulated discharge of CSs and SSs. It is also possible to record from and identify mossy fiber afferent terminals (MFTs) (Fig. 18.4c). Interestingly, MFTs increase their rate of discharge during ipsilateral roll-tilt. Purkinje cell CSs also increase their rate of discharge during ipsilateral roll-tilt. However, SSs increase their rate of discharge during contralateral roll-tilt (Fig. 18.4a, c). This leads to the conclusion that the mossy fiber ! granule cell ! parallel fiber pathway cannot account for the modulation of SSs because it has the wrong phase. Although CSs are defined by their sensitivity to stimulation in the planes of vertical semicircular canals, they are also sensitive to static roll-tilt, indicating sensitivity to otolithic inputs. More than 90% of the Purkinje cells recorded in folia 9c, 9d, and 10 are modulated by sinusoidal and static roll-tilt. The antiphasic modulation of SSs and CSs is manifested in other ways. By changing the orientation of the head about the vertical axis while an animal is sinusoidally roll-tilted, it is possible to characterize a “null plane” at which vestibular stimulation no longer modulates the activity of either CSs or SSs (Fig. 18.4a, c, d). On either side of this “null plane,” the phase of modulated activity shifts by 180
. An “optimal plane” occurs at 90
with respect to the null plane (Fig. 18.4a, b). At the “optimal plane,” neuronal activity is maximally modulated. For each Purkinje cell, CSs and SSs share the same “null” and “optimal” planes and maintain their antiphasic discharge (Fig. 18.4a–d). Furthermore, the depth of modulation of both CSs and SSs is correlated. In a Purkinje cell that has a CS with a large depth of modulation, the SS will also be well modulated (Fig. 18.4e). These observations suggest that climbing fibers are likely the source of modulation for both CSs and SSs. However improbable, it remains logically possible that the antiphasic discharge of CSs and SSs is the consequence of two afferent systems, the responses of which are modulated out of phase. This idea can be evaluated experimentally. The influence of climbing fiber activity on SSs can be tested independently of vestibular primary afferent mossy fibers by acutely disrupting the vestibular primary afferent pathway. If vestibular primary afferent mossy fibers convey a signal necessary for the modulation of SSs in the ipsilateral uvula-nodulus, then the modulation should be blocked when the ipsilateral labyrinth is destroyed by a unilateral labyrinthectomy (UL). A UL leaves the climbing fiber input to the

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Fig. 18.4 Antiphasic discharge of Purkinje cell CSs and SSs during vestibular stimulation. Sinusoidal vestibular stimulation in the roll-pitch axes evoked antiphasic discharge of CSs and SSs from a Purkinje cell in the left folium 9d. (a) The cartoon shows the head orientation of the rabbit during sinusoidal roll-tilt. This orientation was optimal for modulating both CSs and SSs. This orientation maximally stimulated the left posterior and right anterior semicircular canals. When the rabbit was rolled onto its left side CSs increased and SSs decreased. (b) A “null plane” for minimal modulation of CSs and SSs occurred when the orientation of the head was changed so that it was 54
CCW with respect to a vertical axis. In this orientation, the left posterior and right anterior semicircular canals were nearly aligned with the longitudinal axis of rotation. (c) The phase of discharge of 123 CSs, 55 SSs and 11 mossy fiber terminals (MFTs) was measured during sinusoidal roll-tilt about the longitudinal axis. CSs discharged in phase with MFTs and 180
out of phase with SSs. (d) Responses of CSs and SSs were compared in 29 Purkinje cells during stimulation in optimal and null planes. Optimal and null modulation of CSs and SSs always corresponded. (e) The peak increase in CS depth of modulation is plotted against the peak decrease of SS depth of modulation for 29 Purkinje cells stimulated in the optimal plane. Increased depth of modulation of CSs was correlated with increased depth of modulation of SSs. Modified from Barmack and Shojaku (1995) and Fushiki and Barmack (1997)

ipsilateral uvula-nodulus substantially intact since it originates primarily from the contralateral labyrinth. In rabbits, a UL does not prevent vestibular modulation of SSs in ipsilateral Purkinje cells. In these cells, the modulation of SSs remains antiphasic to CSs (Barmack and Yakhnitsa 2003).

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Conversely, if the climbing fiber projection to the uvula-nodulus only influences the modulation of CSs, then blocking this projection leaves the primary and secondary mossy fiber projections intact and should not change vestibular modulation of SSs. However, unilateral electrolytic lesions of the b-nucleus of the inferior olive abolish CSs and block vestibular modulation of SSs in Purkinje cells contralateral to the olivary lesion (Barmack and Yakhnitsa 2003, 2011).

Climbing and Mossy Fiber Function Climbing fiber discharge has two consequences. First, it directly evokes CSs from Purkinje cells arrayed in a sagittal zone. Second, it indirectly reduces SSs in Purkinje cells within this zone by exciting inhibitory interneurons, most likely stellate cells. Paradoxically, the influence of inhibitory interneurons appears greater than that conveyed by vestibular primary afferent mossy fiber ! granule cell ! parallel fiber signals for three reasons: (1) mossy fiber ! granule cell ! parallel fiber signals are too diffuse to account for the specificity of SSs, (2) they have the wrong polarity to account for the phase of SSs evoked by vestibular stimulation, and (3) they are too weak to modulate SSs even when climbing fiber signals are blocked by microlesions of the contralateral inferior olive. What do mossy fibers do? Possibly, they convey a regionally selective level of excitability. The net influence of primary and secondary vestibular mossy fiber afferents could indicate “your head is moving.” The details of head movement are encoded in specific climbing fiber vectors: “you are falling forward and to your left.” Increased mossy fiber activity could broadly increase the discharge of SSs. This higher discharge rate could enhance the depth of modulation of SSs by climbing fiber-evoked interneuronal inhibition.

Reciprocal Cerebellar Projections to the Vestibular Complex Cerebellar projections onto the vestibular and cerebellar nuclei can be studied with conventional orthograde and retrograde tracer methods or, alternatively, with Purkinje cell–specific antibodies (Bernard 1987; Shojaku et al. 1987; Walberg and Dietrichs 1988; Tabuchi et al. 1989; Wylie et al. 1994; Barmack et al. 2000; Highstein and Holstein 2005). Antibodies to two protein kinase C isoforms, PKC-g and PKC-d, provide insights into general cerebellar projections to vestibular and cerebellar nuclei. PKC-g labels the cell bodies and axon terminals of all Purkinje cells. PKC-d labels primarily Purkinje cells in the uvula-nodulus and, to a lesser extent, Purkinje cells in the flocculus (Fig. 18.5). These antibodies demonstrate that the LVN, SVN, MVN, DCN, and NPH (nucleus prepositus hypoglossi) receive projections from “nonnodular” Purkinje cells. Virtually, every cell in the LVN and SVN receive such projections. However, the cerebellar projections to the MVN, DVN, and NPH are more restricted. Uvula-nodular Purkinje cells project only

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Fig. 18.5 Purkinje cell projections to vestibular and cerebellar nuclei of the rat. Transverse sections through the vestibular and cerebellar nuclei from the caudal to rostral poles are illustrated. An antiserum to PKC-g labels all Purkinje cells. Regions containing immunolabeled axon are shown in pink. Parts of each cerebellar nucleus receive projections from cerebellar Purkinje cells. All cells in the lateral and superior vestibular nuclei (LVN and SVN) receive Purkinje cell projections, as do cells in parts of the descending and medial vestibular nuclei (DVN and MVN) and nucleus prepositus hypoglossal nucleus (NPH). An antiserum to PKC-d identified the more restricted projection of Purkinje cells from the uvula-nodulus and flocculus to vestibular and

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to the dorsal surfaces of the MVN, DVN, and NPH and the medioventral quadrant of the Interpositus nucleus. Large regions of the MVN, DVN, and NPH receive no cerebellar projections.

Cerebellar Functions The cerebellum is not needed for the execution of postural or oculomotor reflexes. If the cerebellum is removed surgically, these reflexes persist albeit of inappropriate amplitude and phase. Rather, the cerebellum modulates reflexive movements and establishes a sensory hierarchy for the guidance of movements. For example, the flocculus relies on a climbing fiber-mediated retinal slip signal to adjust the gain of vestibuloocular reflexes. Floccular damage or floccular pharmacological inactivation impairs plastic changes in the horizontal vestibuloocular reflex (Ito et al. 1982; Nagao 1983; Katoh et al. 1998; Nagao and Kitazawa 2003). Similarly, the uvula-nodulus encodes information concerning roll-tilt about the longitudinal axis conveyed by signals from the vertical semicircular canals and the utricular otoliths. Damage to the uvula-nodulus impairs remembered postural adjustments to previously maintained head position in space (Barmack et al. 2002). The cerebellum predicts spatial environments. If an animal is oscillated for an extended period of time and then the oscillation is stopped, oscillatory eye movements persist (Kleinschmidt and Collewijn 1975). Information about such oscillations arrives at the cerebellum encoded by vestibular climbing fibers (Barmack and Shojaku 1992). This predictive function involves cerebellar circuitry, but may already be contained in climbing fiber signals and in signals from the Psol. A more intuitive view of this adaptive function applies to the common experience of moving about on land after even brief intervals spent in boat at sea. Otolithic inputs may prove more effective than vision in predicting the spatial environment at sea. Returning to land, these otolithic signals yield to visual and proprioceptive signals. The cerebellum can either recalibrate or suppress inappropriate signals in these instances.

ä Fig. 18.5 (continued) cerebellar nuclei. This antiserum labeled all Purkinje cells in the uvulanodulus and many Purkinje cells in the flocculus. It identified Purkinje cell axon terminals in the DVN, MVN, NPH, and medioventral aspect of the interpositus nucleus (IntP) (black lines). The numbers at the top of each section indicate the distance from the caudal pole of the vestibular complex. Abbreviations: Amb nucleus ambiguus, Cu cuneate nucleus, DCN dorsal cochlear nucleus, Ecu external cuneate nucleus, icp inferior cerebellar peduncle, IO inferior olive, jx juxta restiform body, LCN lateral cerebellar nucleus, LC locus coeruleus, MCN medial cerebellar nucleus, Nsol solitary nucleus, Py pyramidal tract, scp superior cerebellar peduncle, Sg Scarpa’s ganglion, ol solitary tract, SpV and spV spinal trigeminal nucleus and spinal trigeminal tract, VCN ventral cochlear nucleus, VI abducens nucleus, VII facial nucleus, X dorsal motor nucleus of vagus, XII hypoglossal nucleus, 7n facial nerve, and 12n hypoglossal nerve

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In sum, the cerebellum contains a topographic map of vestibular space conveyed by climbing fiber projections. This map provides an anatomic substrate for specific modulation and adaptation of postural reflexes evoked by vestibular and optokinetic stimulation.

Conclusions and Future Directions Abnormal Cerebellar Function Our understanding of the contributions of the cerebellum to normal and abnormal movement will be enhanced by further investigation of how the activity of different types of neurons and glia contribute to the regulation of the Purkinje cell output signal. The importance of climbing fiber activity to the regulation of discharge of both CSs and SSs has only recently been demonstrated, and climbing fibers are not yet generally considered to have a primary role in the regulation of SS discharge. Only future research will determine whether climbing fiber signals are as important in other regions of the cerebellum as they are in the vestibulocerebellum. The role of climbing fibers in the control of cerebellar signaling may be reflected in the etiology and treatment of cerebellar disease. Spinocerebellar ataxia (SCA), a CAG repeat disease, is not always restricted to the cerebellum (Schaefer et al. 2007; Barnes et al. 2011). In SCA-6, degeneration in precerebellar regions also occurs (the vestibular complex, motor cortex, and inferior olive) (Ishikawa et al. 1999). Detailed analysis of cerebellar disease may implicate the inferior olive in other cases, such as in Leigh’s disease (Cavanagh 1994) or in other forms of SCA (Koeppen 2005). Serotoninergic 5-HT1A receptor agonists are often administered to improve ataxia in patients with cerebellar disorders (Takei et al. 2005). It is assumed that the 5-HT1A therapeutic effects are mediated by Purkinje cell 5-HT1A receptors. However, these therapeutic effects could be mediated by 5-HT1A receptors on olivary neurons that receive serotoninergic projections from neurons in the nucleus reticularis paragigantocellularis (Bishop and Ho 1986). It is possible that circumscribed damage to the brainstem (including the inferior olive) could influence the expression of cerebellar symptoms often attributed to cerebellar cortical pathology.

References Alley K, Baker R, Simpson JI (1975) Afferents to the vestibulo-cerebellum and the origin of the visual climbing fibers in the rabbit. Brain Res 98:582–589 Andersen P, Eccles JC, Voorhoeve PE (1964) Postsynaptic inhibition of cerebellar Purkinje cells. J Neurophysiol 27:1138–1153 Apps R, Garwicz M (2005) Anatomical and physiological foundations of cerebellar information processing. Nat Rev Neurosci 6:297–311 Armstrong DM, Edgley SA (1988) Discharges of interpositus and Purkinje cells of the cat cerebellum during locomotion under different conditions. J Physiol 400:425–445

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Cerebellar Nuclei and the Inferior Olivary Nuclei: Organization and Connections

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Jan Voogd, Yoshikazu Shinoda, Tom J. H. Ruigrok, and Izumi Sugihara

Abstract

The cerebellar nuclei, together with certain vestibular nuclei, are the target of the axons of the Purkinje cells of the cerebellar cortex. Each of these nuclei receives a projection from a longitudinal Purkinje cell zone. Climbing fiber projections are organized according to the same zonal pattern. In this chapter, we will review the morphology and the circuitry of the cerebellar nuclei and the inferior olive and the recurrent pathways connecting them.

Introduction The cerebellar nuclei, together with certain vestibular nuclei, are the target of the axons of the Purkinje cells of the cerebellar cortex. The projections of the cerebellar nuclei to the brain stem and the distribution of the cerebello-thalamo-cortical paths determine the sphere of influence of the cerebellum. Jansen and Brodal (1940, 1942) were the first to notice that the topographical organization of the Purkinje cell

J. Voogd (*) Department of Neuroscience, Erasmus Medical Center Rotterdam, P.O. Box 2040, Rotterdam, 3000 CA, The Netherlands and Rhijngeesterstraatweg 1, 2342 AN Oegstgeest, The Netherlands e-mail: [email protected] Y. Shinoda • I. Sugihara Department of Systems Neurophysiology, Graduate School of Medicine, Tokyo Medical and Dental University, 1-5-45 Yushima, Bunkyo-ku, Tokyo, 113-8519, Japan e-mail: [email protected], [email protected] T.J.H. Ruigrok Department of Neuroscience, Erasmus Medical Center Rotterdam, P.O. Box 2040, Rotterdam, 3000 CA, The Netherlands e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 377 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_19, # Springer Science+Business Media Dordrecht 2013

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projections to the cerebellar nuclei and the olivocerebellar climbing fiber system is very similar. Marr (1969) formulated this similarity in his learning theory of the cerebellar cortex as “the olivary cell should respond to a command for the same elemental movement as is initiated by the corresponding Purkinje cell.” In this chapter, we will review the morphology of the cerebellar nuclei and the inferior olive, the afferent connections of the olive, the zonal organization of the corticonuclear and olivocerebellar projection, and the efferent connections of the cerebellar nuclei and the presence of recurrent cerebellar-brain stem circuitry.

The Cerebellar Nuclei Four cerebellar nuclei, known as the fastigial emboliform, globose, and dentate nucleus, were distinguished in the human cerebellum by Stilling (1864). The same four nuclei can be distinguished in different mammalian species, where they are known as the medial, anterior interposed, posterior interposed, and lateral cerebellar nucleus (Ogawa 1935; Weidenreich 1899) (Fig. 19.1). The cerebellar nuclei are arranged in two groups. The rostrolateral group consists of the anterior interposed nucleus (emboliform) that is connected with the lateral (dentate) nucleus. The caudomedial group consists of the medial (fastigial) nucleus with the posterior interposed (globose) nucleus. A fifth cerebellar nucleus, located at the border of the fastigial and posterior interposed nucleus, known as the “interstitial cell groups” was distinguished by Buisseret-Delmas and Angaut (1993) in the rat but appears to be present in other mammals (Fig. 19.1a, b). Three populations of neurons have been distinguished in all cerebellar nuclei. The main population consists of excitatory relay cells with widespread, branching axons, terminating in the brain stem, the spinal cord, and the thalamus. They constitute a mixed population of cells of all shapes and sizes (Courville and Cooper 1970). Most relay cells of the cerebellar nuclei are glutamatergic. Large, glycinergic neurons that give rise to the ipsilateral projections of the fastigial nucleus were identified in mice (Bagnall et al. 2009). Small GABAergic neurons, that project exclusively to the inferior olive constitute a second population, present in all nuclei (Graybiel et al. 1973; Mugnaini and Oertel 1985) (see also section “The Nucleo-olivary Pathway”). A third population of small interneurons was identified by Chan-Palay (1977) on morphological grounds in the monkey, by Chen and Hillman (1993) as glycinergic neurons, and by Leto et al. (2006) as GABAergic neurons, all in rodents. Glycinergic neurons, with projections to the cerebellar cortex, were described in the mouse lateral cerebellar nucleus (Uusisaari and Knopfel 2010). Before, all recurrent projections from the cerebellar nuclei to the cortex were supposed to originate as collaterals from the relay cells (McCrea et al. 1978). The fastigial nucleus is located next to the midline. In rodents, a prominent protrusion, known as the dorsolateral protuberance (Goodman et al. 1963), is present that is lacking in carnivores and primates (Fig. 19.1a). The rodent anterior interposed nucleus includes a lateral portion, indicated as the dorsolateral hump that

Fig. 19.1 The cerebellar nuclei of the rat, Macaca fascicularis, and the human cerebellum: dorsal aspect and selected transverse sections. Abbreviations: AI anterior interposed nucleus, bc brachium conjunctivum, CO cochlear nucleus, D micro microgyric part of the dentate nucleus, D dentate nucleus, DLH dorsolateral hump, DLP dorsolateral protuberance, D macro macrogyric part of the dentate nucleus, DV descending vestibular nucleus, E emboliform nucleus, F fastigial nucleus, G globose nucleus, ICG interstitial cell groups, icp inferior cerebellar peduncle, int.nu. basal interstitial nucleus, L lateral cerebellar nucleus, LV lateral vestibular nucleus, Mcm caudomedial subdivision of the medial nucleus, Mn medial subdivision of the medial nucleus, MV medial vestibular nucleus, PI posterior interposed nucleus, SV superior vestibular nucleus, Y group Y

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sometimes is allocated to the lateral cerebellar nucleus. A parvocellular subnucleus occupies the ventral part of the lateral cerebellar nucleus in these species. The primate dentate nucleus has the shape of a “crumpled purse” (Chan-Palay 1977) with its hilus directed ventromedially and rostrally. In the human cerebellum, two parts of the nucleus can be distinguished, a microgyric and a macrogyric portion (Fig. 19.1c). The dorsomedial, microgyric portion is folded in rather narrow, rostrocaudally directed ridges. The ridges in the ventrolateral, macrogyric portion of the nucleus are broad and subdivided, and the cell band in this part of the nucleus is wider than in the microgyric part. The cells of the microgyric part of the dentate are larger than those in the macrogyric position of the nucleus (Demole´ 1927a, b), but both subdivisions also contain small neurons. This subdivision had already been noticed by Vicq d’Azyr in the first accurate description of the nucleus lateral, dating from the eighteenth century (Glickstein et al. 2009). The ontogenetic development of the macrogyric portion lags far behind the microgyric (Weidenreich 1899). Unfortunately, there are no indications how the two parts of the human dentate are represented in the monkey dentate. A prominent Y nucleus is present, ventral to the dentate, in the monkey cerebellum (Fig. 19.1b). It should be distinguished from Ill-defined groups of small acetylcholinesterase-positive cells dispersed in the white matter of the flocculus and the nodule and ventral to the dentate and the posterior interposed nucleus, in the roof of the fourth ventricle known as the basal interstitial nucleus (Fig. 19.1b). The basal interstitial nucleus is reciprocally connected with the flocculus (Langer et al. 1985) and, possibly, with the nodulus. The group Y is a lateral extension of the superior vestibular nucleus that gives rise to projections to the oculomotor nuclei (Graybiel and Hartwieg 1974; Stanton 1980a; Steiger and B€uttner-Ennever 1979; Yamamoto et al. 1986). Langer’s basal interstitial nucleus and the group Y are welldefined nuclei in the monkey cerebellum but have not yet been recognized in the human cerebellum.

Subdivision of the Inferior Olive The inferior olive derives its name from the olive-shaped prominence on the ventrolateral surface of the medulla. The first description of the nuclei of the inferior olive dates from Stilling (1843). He identified the principal olive, the medial accessory olive (as the “grossen Pyramidenkern”) and the dorsal accessory olive (his “Oliven Nebenkern”) in sections through the human medulla oblongata. Sections through the inferior olive of the human brain and of different species that have been used in experimental studies of the connections of the inferior olive are illustrated in Figs. 19.2 and 19.3. Diagrams of surface-projections of the olivary nuclei, first constructed by Brodal (1940) showing the position of the sections, are included in these figures. Subdivisions of the principal and the accessory olives have been distinguished by inspection of the shape and contour of these nuclei, but their definite identification depends on the recognition of their connections. Moreover, species-specific subdivisions may occur.

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Fig. 19.2 Diagrams of selected transverse sections through the inferior olive and surfaceprojections of its subnuclei of different mammalian species (Redrawn from Brodal (1940), rabbit and cat; Ruigrok and Voogd (1990), rat; and Fujita et al. (2010), marmoset). Abbreviations: a, b, c subnuclei a, b, c of the caudal medial accessory olive, ß group beta, DAOc caudal dorsal accessory olive, DAOdf dorsal fold of the dorsal accessory olive, DAOr rostral dorsal accessory olive, DAOvf ventral fold of the dorsal accessory olive, dlPO dorsal lamina of the principal olive, dm dorsomedial group, dmcc dorsomedial cell column, MAOc caudal medial accessory olive, MAOr rostral medial accessory olive, MDO medial part dorsal accessory olive, vlo ventrolateral outgrowth, vlPO ventral lamina of the principal olive

Fig. 19.3 Diagrams of selected transverse sections through the human inferior olive and of transverse sections and a surface projection of its subnuclei of the macaque monkey (Human inferior olive redrawn from Marechal (1934)). Abbreviations: llPO lateral lamina of the principal olive, mlPO medial lamina of the principal olive. For other abbreviations, see Fig. 19.2

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The medial accessory olive can be subdivided into caudal (MAOc) and rostral (MAOr) parts. The MAOc often has been further subdivided into the mediolaterally disposed subgroups a, b, and c. (Fig. 19.1, rat section 2, marmoset, sections 1–3; Fig. 19.2, macaque monkey, section 3). This subdivision is not distinct at all levels. Moreover, in the rat, subgroups a–c were distinguished, in addition to the group beta, which forms a fourth, medial subdivision of the MAOc, whereas in the macaque monkey, subgroup c is identical to the group beta. The group beta was first distinguished by Brodal (1940). In the cat and the rabbit, it is aligned with the more rostrally located dorsomedial cell column (DMCC). The DMCC was first distinguished by Bertrand and Marechal (1930) in the human inferior olive as a rostrally located cell group, attached to the medial pole of the DAO and, ventrally, to the medial lamella of the principal olive (Fig. 19.3, section 48). This position is even more distinct at late fetal stages (Fig. 19.3, inset). In the cat and the rabbit, Brodal (1940) identified a cell group with a different position, located dorsomedial to the rostral MAO, as the DMCC (Fig. 19.2). In the rat, Brodal’s DMCC is present, but it should be distinguished from another subgroup, the dorsomedial group (Azizi and Woodward 1987). The dorsomedial group in the rat is attached to the medial pole of the ventral lamina of the principal olive, a position it shares with the DMCC from Marechal’s original description. In macaque monkeys, the cell group identified as the DMCC is attached to the medial lamella of the PO and/or the medial pole of the DAO and, therefore, occupies the same position as the human DMCC and the rat dorsomedial group (Bowman and Sladek 1973; Brodal and Brodal 1981; Whitworth and Haines 1986a, b) (Figs. 19.2 and 19.3). In the marmoset, both cell groups can be distinguished (Fujita et al. 2010). The connections of the two cell groups are different: the DMCC as present in cat, rabbit, and rat receives vestibular input, the dorsomedial group and the monkey DMCC (Fig. 19.3, right panel, sections 5,6), are innervated by somatosensory systems. A cell group corresponding to the cat DMCC, innervated by the vestibular system, has not been identified in monkeys. At the level of the MAOc, the dorsal cap (DC) of Kooy (1917) is located dorsal to the group beta. More rostrally, it extends medially as the ventrolateral outgrowth (VLO). The dorsal accessory olive can be divided into caudal (DAOc) and rostral (DAOr) parts. In the rat, the DAOc is folded over the DAOr, and the two divisions of the DAO are known as the dorsal and ventral fold of the DAO (Azizi and Woodward 1987). In surface projections, the DAOc typically forms a caudomedially directed extension of the DAO. The principal olive generally is subdivided into dorsal (dlPO) and ventral (vlPO) laminae. In rat, cat, rabbit, and marmoset, the cell band of the vlPO is thinner than the broad cell band of the dlPO. At some levels, a gap or a constriction separates the vlPO from the dlPO in rat and cat. In artiodactyles, the ventral lamina is always separated from the rest of the PO (Whitworth and Haines 1986b). In the macaque monkey, an additional lateral lamina or “bend” was distinguished. In monkeys, the great apes and in the human inferior olive the dorsal (dlPO), lateral (llPO) and the lateral portion of the ventral lamina, indicated as the vlPO, display the typical convolutions of the inferior olive. In monkeys, the thin, nonvoluted portion of the

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vlPO is indicted as the mlPO (Fig. 19.3). In the human inferior olive, Kooy (1917) and Bertrand and Marechal (1930) described a narrow, medial lamina (mlPO), located medial to the MAO (Fig. 19.3, sections 38–51). In rabbit and rat, the medial pole of the DAOr is connected with the dlPO. In the cat and the marmoset, a similar connection exists with the vlPO. In the human and macaque inferior, the DAOr and the laminae of the principal olive remain isolated from each other. The VLO is continuous with the vlPO in the rat and in the human inferior olive and with the dlPO in the rabbit and the cat. No such continuity appears to exist in the marmoset and the macaque monkey.

Afferent Connections of the Inferior Olive Four main categories of afferents of the inferior olive, each with their own distribution, can be distinguished. The inferior olive processes somatosensory input mostly in the MAOc and DAO, vestibular and optokinetic input in the DC/VLO/ group beta, visual information in the medial part of MAOc. The PO and MAOr process information relayed by nuclei at the mesodiencephalic border from the cerebral cortex. However, overlap between the various modalities within olivary subnuclei has been noted. Below, we provide a more detailed account of these projections.

Projections from Spinal Cord, Trigeminal Nuclei, and Dorsal Column Nuclei Ventral Spino-olivary Pathways Spino-olivary fibers originate mainly from the contralateral cervical and lumbar gray matter, ascend in the ventral funiculus, and terminate in the caudal MAOc and, laterally, in caudal and rostral parts of the DAO (Fig. 19.4) (Boesten and Voogd 1975; Armstrong and Schild 1979; Armstrong et al. 1982). The system has been described in different mammalian species, but not in monkeys (Brodal et al. 1950; Brown et al. 1977; Martin et al. 1980; Mizuno 1966; Swenson and Castro 1983a, b; Whitworth and Haines 1983). Detailed information on the anatomy and the physiology of the ventral funiculus olivocerebellar pathway (vfSOCP) (Oscarsson and Sj€olund 1977a, b, c) is available for the cat (Fig. 19.22a). Spino-olivary fibers originate from the medial ventral horn (Rexed’s (1952) lamina VIII), the nucleus proprius of the dorsal horn (laminae IV/V), cells in the lateral funiculus and the intermediate gray (lamina VII). Except for lower cervical levels, all neurons giving rise to spino-olivary fibers are located contralaterally. The majority of the system takes its origin from the lumbar and sacral cord, the contribution of the thoracic cord is negligible and of the cervical cord rather small. The lateral cervical nucleus does not contribute to the projection (Armstrong and Schild 1979; Armstrong et al. 1982; Buisseret-Delmas 1982; Molinari 1984, 1985; Richmond et al. 1982). Lumbar and sacral spino-olivary fibers terminate ventrolaterally in the MAOc, overlapping with the projection from the cervical cord that extends more medially.

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Fig. 19.4 The ventral funiculus spino-olivary pathway in the cat. The origin of this system is illustrated in diagrams of the lumbosacral cord for cells retrogradely labeled from the entire DAO (L5-S3, Molinari 1984) and from the caudal MAO and DAO (L6-L7, Molinari 1985), the low cervical (Armstrong et al. 1982), and high cervical cord (Richmond et al. 1982). Projections from these levels to the contralateral MAOc and DAOc show much overlap, and a more distinct topical projection is present in the DAOr (Armstrong et al. 1982; Boesten and Voogd 1975). A projection from the contralateral spinal trigeminal nucleus (Trig) occupies the rostromedial MAOc and the medial DAOr (Courville et al. 1983b) (Redrawn from illustrations of the cited papers). For abbreviations, see Fig. 19.1

A similar overlap of sacrolumbar and cervical afferents is present in the caudal DAO. In the rostral DAO, a distinct somatotopical organization is present, with lumbar and cervical fibers terminating in increasingly more medially located lamellae (Armstrong et al. 1982; Boesten and Voogd 1975; Richmond et al. 1982). A very similar somatotopical organization was found in the DAO of the cat for the distribution of evoked potentials on peripheral stimulation (Fig. 19.5) (Gellman et al. 1983). Evoked potentials from light cutaneous stimuli predominate over deep input from muscles and fascia in DAOr, and the latter predominate in the DAOc. These recordings represent the combined projections from the ventral and the dorsal column spino-olivary pathways. Projections to the MAOc and the DAOc originate mainly from the medial ventral horn and projections to the DAOr from the dorsal horn. The intermediate gray projects to both accessory olives (Molinari 1984, 1985). Spino-olivary fibers terminate as boutons containing spherical vesicles, with asymmetrical synapses on dendritic shafts and spines, more inside than outside the glomeruli (King et al. 1975; Molinari and Starr 1989).

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Fig. 19.5 Somatotopical localization in the dorsal accessory olive of the cat (Redrawn from Gellman et al. (1983))

Projections from the Sensory Nuclei of the Trigeminal Nerve The sensory trigeminal nuclei project to the extreme medial part of the contralateral DAOr (Cook and Wiesendanger 1976) and, in addition, to the adjoining vlPO in the cat and the rabbit (Van Ham and Yeo 1992), to the dorsomedial group in the rat (Huerta et al. 1983) and to a separate region in the vlPO in the rabbit (Van Ham and Yeo 1992). No terminations are present in the dorsomedial cell column in any of these species (Fig. 19.6). In the MAOc, the trigeminal nuclei project to its rostromedial region, next to the rostral pole of the group beta in all species where they overlap with the terminals from the cuneate nucleus (see section “Projections from the Dorsal Column Nuclei” and Fig. 19.6c). In the cat, the projection extends more caudally, overlapping with the terminals from the cervical cord (Fig. 19.6). Interestingly, terminations in the rat DAOr were identified as collateral projections of the trigemino-tectal pathway, but the projection of the trigeminal nuclei to the MAOc is an unbranched system (Huerta et al. 1983). No data on the trigeminoolivary projection are available for primates. Most authors agree that the main origin of the trigemino-olivary projection is located in the pars interpolaris of the spinal nucleus (Huerta et al. 1983; Huerta and Harting 1984; Swenson and Castro 1983a; Van Ham and Yeo 1992; Walberg 1982), although some also include the principal sensory nucleus and the pars caudalis.

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Fig. 19.6 Localization of projections of the contralateral spinal trigeminal nucleus illustrated in a rostral and a more caudal section through the inferior olive of the rat (Huerta et al. 1983), the cat (Berkley and Hand 1978), and in transverse sections and a diagram of the flattened inferior olive of the rabbit (Van Ham and Yeo 1992). For abbreviations, see Fig. 19.2

Projections from the Dorsal Column Nuclei Two fiber systems occupy the dorsal columns: the ascending branches of the dorsal root fibers and the postsynaptic dorsal column pathway that contains an extra synapse in the dorsal horn (Fig. 19.7). Both pathways transmit somatosensory information to the dorsal column nuclei, but the latter pathway has been shown to be responsible for the transmission of nociceptive input to the inferior olive in the cat (Ekerot et al. 1991). The connectivity in the dorsal funiculus spinoolivocerebellar pathway (df-SOCP) has been extensively studied by Ekerot and Larson (1979a, b) (Fig. 19.22b). In the dorsal column nuclei of the cat, the fusiform cells that project to the contralateral inferior olive are located outside the cell cluster regions of the internal and gracile nuclei, which give rise to the medial lemniscus (Fig. 19.8). Neurons of the gracile nucleus that project to the rostral and caudal DAO accumulate rostral to the cluster region in the transitional portion of the nucleus. The main projection is to the DAOr. The numbers of the cells labeled from injections of retrograde tracers in the DAOc and the MAOc are small (Molinari 1984, 1985). A wider distribution of neurons with projections to the inferior olive was found for the internal cuneate nucleus (Alonso et al. 1986; Buisseret-Delmas 1982) (Fig. 19.8).

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Fig. 19.7 The dorsal funiculus spino-olivary pathway in the cat. (a) Both ascending collaterals from spinal root fibers and the postsynaptic dorsal column pathway partcipate in the projection to the dorsal column nuclei. Projections to the inferior olive take their origin from regions outside the cell clusters (cl) that give origin to the medial lemniscus. (b) Projections from the contralateral gracile nucleus (Boesten and Voogd 1975). (c) Projections from the contralateral gracile and internal cuneate nuclei (Boesten and Voogd 1975). (d) Projections from the contralateral cuneate nucleus (Groenewegen et al. 1975) (b–d redrawn from illustrations of the cited papers). Abbreviations: CE external cuneate nucleus, Cui, internal cuneate nucleus, GR gracile nucleus. For other abbreviations, see Fig. 19.2

The projections from the gracile and internal cuneate nuclei occupy lateral and more medial bands in the DAOr and mostly spare the DAOc (Fig. 19.7b, d). A minor projection from the gracile nucleus is present in the MAOc (Figs. 19.7b and 19.8). Terminals from the internal cuneate nucleus are found in the rostromedial MAOc at its border with the MAOr (Fig. 19.7c), a region that also receives a projection from the spinal trigeminal nucleus (Fig. 19.5). Projections to the ipsilateral inferior olive from the dorsal column nuclei, described by several authors, may originate from the adjoining reticular formation (Courville et al. 1983b). Few data are available for the projection of the dorsal column nuclei to the inferior olive in monkeys. Sections of cases from Kalil (1979) and Molinari et al. (1996), illustrated in Fig. 19.9, show terminations in the DAOr, the DMCC, and the rostromedial MAOc. These projections are in accordance with the findings in cats, with the exception of the DMCC that receives vestibular, rather than somatosensory input in this species. It may be that the monkey DMCC corresponds to the dorsomedial group, as identified in the rat, rather than with the DMCC of other species.

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Fig. 19.8 Localization of neurons of the dorsal column nuclei projecting to the contralateral inferior olive in the cat. Left panels (blue) show the location of neurons in the gracile nucleus and the rostrocaudal distribution of neurons projecting to the entire DAO, the MAOc and the DAOc and the MAOc only (Molinari 1984, 1985). Right panel shows the distribution of neurons of the internal cuneate nucleus after large injections of a retrograde tracer in the inferior olive (Alonso et al. 1986) (Redrawn from illustrations of the cited papers). Abbreviations: AP area postrema, Cui internal cuneate nucleus, GR gracile nucleus

Fig. 19.9 Projections from the internal cuneate nucleus of the macaque monkey. (a) is the most rostral section (a and d redrawn from Kalil (1979), c and d from Molinari et al. (1996)). Abbreviations see Fig. 19.2

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Electron microscopic studies of the dorsal column-olivary projection documented its terminals as excitatory boutons with spherical vesicles and asymmetrical synaptic contacts (Molinari et al. 1991). GABAergic neurons in the cuneate and less in the gracile nucleus, with projections to the contralateral inferior olive, were observed by Nelson and Mugnaini in the rat (1989). Alternative routes for the transmission of peripheral input to the inferior olive are provided by ascending spinal, trigeminal, and dorsal column pathways terminating in the pretectum that innervates the DAOr (see below), in Darkschewitsch nucleus that innervates the DAOr, or in the parvocellular red nucleus that innervates the principal olive (Wiberg and Blomqvist 1984a, b; Wiberg et al. 1986, 1987).

Optokinetic and Vestibular Projections to the Inferior Olive The optokinetic system with its projections of the nuclei of the accessory optic system and the nucleus of the optic tract in the mesencephalon to the dorsal cap (DC) and the ventrolateral outgrowth (VLO) recently was reviewed by Giolli et al. (2006), Barmack (2006), and Voogd (Voogd and Barmack 2006; Voogd et al. 2011). For complete references, we refer to these reviews. DC and VLO receive information on global movements of the visual surround from large field ganglion cells in the contralateral retina via the optic tract (Fig. 19.10a). These movements generate retinal slip signals, which can serve as an error signal in long-term adaptation of the vestibulo-ocular and optokinetic reflexes. The retinal slip signals around a vertical axis excite neurons of the nucleus of the optic tract and the dorsal nucleus of the accessory system. These nuclei project to the DC of the ipsilateral inferior olive. Global movements of the visual surround around an oblique horizontal axis at 45 azimuth, that is approximately colinear with the axis of the ipsilateral anterior semicircular canal, are transmitted bilaterally by the medial and lateral nuclei of the accessory optic system and contralaterally by the visual tegmental relay zone. These nuclei project to the VLO. The organization of the accessory optic system and the projections of its subnuclei to the inferior olive in primates is very similar to that in lower mammals. The border between the projections of vertical axis and horizontal axis neurons in the rabbit is located halfway within the dorsal cap. Consequently, the two functional subdivisions in this species correspond to the VLO and the rostral dorsal cap that relay information from horizontal axis neurons, and the caudal dorsal cap that serves as the relay for the vertical axis neurons. The VLO merges with the vlPO, and the vertical axis subdivision may spill over in this lamina in species like the rat. The DC and the VLO receive additional input from the paramedian reticular formation (Fig. 19.10b), the dorsal group Y, and the nucleus prepositus hypoglossi (Fig. 19.11a). The paramedian reticular formation is involved in the generation of saccades. Its projection to the dorsal cap has been identified only in the cat (Gerrits and Voogd 1986). The dorsal group Y is located within the floccular peduncle and is the target of Purkinje cell zones influencing eye movements about a similar horizontal axis as the cells of the VLO. In the cat, it projects to the contralateral VLO

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Fig. 19.10 Optokinetic and vestibular afferents of the inferior olive. (a) Projection of the nuclei of the accessory optic system and the nucleus of the optic tract on to the dorsal cap and the ventrolateral outgrowth. (b) Map of the projection of the paramedian pontine reticular formation to the dorsal cap in the cat (Gerrits and Voogd 1986). Abbreviations: bc brachium conjunctivum, cp cerebral peduncle, D dorsal nucleus of the accessory optic system, DC dorsal cap, dlPO dorsal lamina of the principal olive, L lateral nucleus of the accessory optic system, LGB lateral geniculate body, ll lateral lemniscus, M medial nucleus of the accessory optic system, MGB medial geniculate body, NRTP nucleus reticularis tegmenti pontis, NTO nucleus of the optic tract, opt.tr optic tract, Pc posterior commissure, PPRF paramedian pontine reticular formation, transped.tr transpeduncular tract, V trigeminal nerve, VLO ventrolateral outgrowth, vlPO ventral lamina of the principal olive, VTRZ ventral tegmental reflex zone

(Gerrits et al. 1985), and in the rabbit, this projection also includes the rostral DC (De Zeeuw et al. 1994a). This projection is GABAergic. The nucleus prepositus hypoglossi has a role in gaze-holding. Separate neuronal populations connect it with the horizontal gaze center in the paramedian reticular formation, the vestibular nuclei, the cerebellum, and the inferior olive (McCrea and Baker 1985). The projection of the nucleus prepositus hypoglossi to the dorsal cap is contralateral in the cat and rat (Barmack et al. 1993; Gerrits et al. 1985), but bilateral in the rabbit (De Zeeuw et al. 1993). This projection consists of a mixture of cholinergic and GABAergic afferents and of elements expressing both transmitters (Barmack et al. 1993; De Zeeuw et al. 1993). According to Barmack (2006), vestibular input to the group beta and the DMCC is organized as a simple push-pull system, with excitatory input from the group Y and inhibition from the parasolitary nucleus. A projection of the group Y to the contralateral DMCC (but not to the group beta) was traced in the cat (Gerrits et al. 1985) (Fig. 19.11a) but not in the rabbit (De Zeeuw et al. 1994a) where this system, moreover, was found to be GABAergic. The parasolitary nucleus is located lateral to the solitary nucleus, next to the caudal pole of the descending vestibular nucleus

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Fig. 19.11 (a) Injection sites of antegrade tracers in different parts of the vestibular nuclei, the group Y and the nucleus prepositus hypoglossi and their projections to the inferior olive (Redrawn from Gerrits et al. (1985). Additional projections (of the group Y to the contralateral group beta (Barmack 2006) and of the nucleus prepositus hypoglossi to the ipsilateral dorsal cap (De Zeeuw et al. 1993) are hatched. (b) Projection of the parasolitary nucleus to the group beta in the rat (Redrawn from Barmack et al. (1998)). Abbreviations: F fastigial nucleus, IA anterior interposed nucleus, L lateral cerebellar nucleus, Y group Y, FlO flocculus, LV lateral vestibular nucleus, DV descending vestibular nucleus, MV medial vestibular nucleus, PrH nucleus prepositus hypoglossi, S nucleus of the solitary tract, CE external cuneate nucleus, CU internal cuneate nucleus, PO principal olive, vlo ventrolateral outgrowth, dc dorsal cap, dmcc dorsomedial cell column, MAO medial accessory olive, ß group beta

(Fig. 19.11b). It receives root fibers from branches of the vestibular nerve that innervate the ampullae of the vertical canals and the utriculus. Its small, compact neurons provide the ipsilateral group beta and the DMCC with inhibitory synapses (Barmack et al. 1998; Loewy and Burton 1978; Molinari and Starr 1989). Projections of the medial and descending vestibular nuclei to the ipsilateral group beta and the DMCC were found by several authors in cat, rat, and rabbit (Barmack et al. 1993; Brown et al. 1977; Gerrits et al. 1985; Nelson and Mugnaini 1989; Saint-Cyr and Courville 1979; Swenson and Castro 1983a) (Fig. 19.11a). Signals from the ipsilateral anterior semicircular canals, relayed through the parasolitary nucleus, are mapped upon the caudal group beta, signals from the posterior canal onto its rostral part. There are no projections from the horizontal canal. DMCC neurons respond preferentially to otolithic stimulation. The pattern of vestibularly modulated activity in DMCC neurons is consistent with an inhibitory vestibulo-olivary projection, supposedly from the parasolitary nucleus (Barmack 2006).

Afferents from Tectum and Pretectum A contralateral projection of the superior colliculus to the medial MAOc, located next to the group beta, was demonstrated in rat, cat, rabbit, and monkey

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Fig. 19.12 Projections to the inferior olive from the tectum and the pretectum. (a–c) The tectoolivary projection. (a, b) Tecto-olivary neurons in the cat are located in the intermediate gray layer as part of a grid, where they alternate with tectospinal (green) and tectotrigeminal (yellow) neurons (Redrawn from Huerta and Harting (1984) and Weber et al. (1978)). (c) Termination in the rat of tecto-olivary projection in medial MAOc (Akaike 1992). (d, e) The pretecto-olivary projection in the cat. (d) Origin from the anterior pretectal nucleus (Kitao et al. 1989). (e) Termination in the DAOr (Kawamura and Onodera 1984). Abbreviations: APN anterior pretectal nucleus, CG central gray, Cp posterior commissure, DAO dorsal accessory olive, GC central gray, II-V layers II-V of the superior colliculus, MAO medial accessory olive, MAO neurons projecting to MAOc, R red nucleus, THAL thalamus, tsp tectospinal neurons, ttr tectotrigeminal neurons

(Frankfurter et al. 1976; Harting 1977; Hess and Voogd 1986; Holstege and Collewijn 1982; Saint-Cyr and Courville 1982; Weber et al. 1978) (Fig. 19.12c). It takes its origin from the fourth, intermediate gray lamina of the superior colliculus. When Weber’s (Weber et al. 1978) plot of the neurons that were labeled retrogradely from the inferior olive (Fig. 19.12b) is examined, these neurons appear to be located in layers or patches in the superficial and deep parts of the intermediate gray. Huerta and Harting (1984) found similar patches of neurons with projections to the inferior olive to be arranged in a grid in the intermediate gray layer that also contains neurons with projections to the trigeminal nuclei and those giving rise to the predorsal fascicle (Fig. 19.12a). The functional meaning of this organization is not

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known. In the rat MAOc, the tecto-olivary projection remains separated from the more laterally and rostrally terminating trigemino-olivary pathway (Akaike 1989). The ventral anterior pretectal nucleus and scattered neurons in the pretectum, that may belong to the dorsal pretectal nucleus and the nucleus of the posterior commissure, have been found to project to the ipsilateral DAOr in the rabbit and cat (Itoh et al. 1983; Kawamura and Onodera 1984; Kitao et al. 1989) (Fig. 19.12d, e). These nuclei receive afferents from the dorsal column nuclei. In the DAOr, the projections from the pretectum and the dorsal column nuclei overlap (Kawamura et al. 1982; Bull et al. 1990).

Nuclei at the Mesodiencephalic Border: The Central and Medial Tegmental Tracts The origin of the central tegmental tract, one of the main fiber systems of the human brain stem, from the parvocellular red nucleus and its termination in the principal olive was established by Verhaart (1936). The medial tegmental tract was discovered by Ogawa (1939) in the cat. It arises from the region surrounding the fasciculus retroflexus, including Darkschewitsch nucleus in the rostral central gray, descends along the raphe in the ventral part of the medial longitudinal fascicle, and terminates in the MAOr and the vlPO. The presence of the two tegmental tracts, one arising from the parvocellular red nucleus, that innervates the principal olive and the other from different nuclei surrounding the fasciculus retroflexus, innervating MAOr, has been confirmed in different species, including humans (Voogd 2004). The mesodiencephalic projections were analyzed in great detail by Onodera (1984) in the cat (Fig. 19.13). This author distinguished a series of nuclei, surrounding the fasciculus retroflexus with projections to the ipsilateral inferior olive. Darkschewitsch nucleus projects to the MAOr and less intensely to the DMCC. Bechterew’s nucleus connects with the vlPO, and the parvocellular red nucleus with the dlPO. A rather vague projection of Cajal’s interstitial nucleus and/or the prerubral field was found to the lateral MAOc and the VLO. A topographic arrangement was found for the parvocellular red nucleus, with its lateral portion projecting to rostral, and its medial portion to caudal dlPO. Similarly, dorsolateral Darkschewitsch nucleus projects to caudal MAOr and its ventromedial portion to rostral MAOr (Porter et al. 1993). The connections of the parafascicular region with the inferior olive in the rat were reviewed by Ruigrok (2004). Although details on this system in the rat are not known, it seems likely that its organization is very similar to that of the cat. Strominger et al. (1979) found these connections to be very similar in monkey (Fig. 19.14), although they were not able to provide definite proof of the projection of Darkschewitsch nucleus to the MAOr; in Fig. 19.14, this projection is illustrated in analogy with the cat. The dorsomedial subnucleus of the parvocellular red nucleus, located medial to the fasciculus retroflexus, represents Bechterew’s nucleus and projects to the vlPO. The main lateral and caudal parts of the parvocellular red nucleus project topographically to dlPO and llPO, with lateral

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Fig. 19.13 Connections of the parvocellular red nucleus and other nuclei at the mesodiencephalic border to the inferior olive in the cat. Projections of the cerebral cortex to these nuclei are indicated in the upper diagram of rostral view of the cerebral hemisphere (Lower panels redrawn and modified from Onodera (1984)). Abbreviations: aq aqueduct, B Bechterew’s nucleus, D Darkschewitsch nucleus, DAOc/r caudal/rostral dorsal accessory olive, dc dorsal cap, dmcc dorsomedial cell column, FEF frontal eye fields, IC interstitial nucleus Cajal, MAOr/c rostral/caudal part of medial accessory olive, prf prerubral field, r fasciculus retroflexus, Rp parvocellular red nucleus, ß group beta, vl/dlPO ventral/ dorsal lamina of the principal olive; Vlo ventrolateral outgrowth

parts projecting medially in dlPO and medial parts more laterally. Thus, it transmits the somatotopical organization of this nucleus, imposed upon it by the cerebral cortex. The terminations in the PO are distributed in alternating bands of high and low density. The significance of this pattern is not known but may indicate a more precise topical organization of the system. The preolivary nuclei at the mesodiencephalic border receive afferents from the cerebellum and from the cerebral cortex. Direct cortico-olivary connections have been observed but appear to be rather scanty (see Saint-Cyr (1983) for a review and Borra et al. (2010) for a more recent observation). Darkschewitsch’ nucleus receives a cerebellar projection from the posterior interposed nucleus (Fig. 19.32a). The fastigial and dentate nuclei have been mentioned as possible afferent sources. In monkeys, the dentate nucleus projects to the parvocellular red nucleus, with

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Fig. 19.14 Connections of the parvocellular red nucleus and other nuclei at the mesodiencephalic border to the inferior olive in the monkey. Origin of projections to these nuclei is indicated in the upper diagram of the cerebral hemisphere. Stippled pre- and postrolandic areas contain all neurons retrogradely labeled from the parvocellular red nucleus (Humphrey et al. 1984). Antegrade tracing was performed from frontal eye fields, primary ventral premotor, supplementary motor, and posterior parietal areas. Projections from primary motor and supplementary motor areas to the parvocellular red nucleus are somatotopically arranged. Projections to the inferior olive, illustrated in a transverse section at the bottom of the figure, were redrawn from Strominger et al. (1979). Abbreviations: aq aqueduct, Dark Darkschewitsch’ nucleus, dl/ll/vlPO dorsal/lateral/ventral lamina of the principal olive, DM dorsomedial subnucleus of the parvocellular red nucleus (Bechterew’s nucleus), FEF frontal eye field, M1 primary motor cortex, MAOr rostral medial accessory olive, SEF supplementary eye field, SMA supplementary motor area

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a termination from its ventrocaudal part in the dorsomedial nucleus and of its dorsal and rostral parts in its main ventrocaudal portion (Stanton 1980b; Voogd 2004) (Fig. 19.32b, c). In the cat, cortical afferents to the nuclei of the mesodiencephalic border stem from the frontal eye fields and the areas 4, 6, and 3 (Miyashita and Tamai 1989; Nakamura et al. 1983; Saint-Cyr 1987). These areas project to Darkschewitsch’ nucleus and the parvocellular red nucleus (Fig. 19.13). Afferents from posterior parietal cortex terminate in the parvocellular red nucleus (Oka 1988). Bechterew’s nucleus was not considered as a separate nucleus in these studies. Through the topical projection of Darkschewitsch’ nucleus to the MAOr, the rostral pole of this nucleus receives oculomotor, and its caudal part MAOr skeletomotor input (Porter et al. 1993). The corticorubral projection has been studied in greater detail in monkeys (Fig. 19.14). The total cortical area projecting to the parvocellular red nucleus was determined by Humphrey et al. (1984) (Fig. 19.14, stippled). It includes areas 4, 6, and 8 and the posterior parietal cortex area7, extending into the dorsal bank of the intraparietal cortex. It originates from a specific set of pyramidal neurons in upper layer 4 (Catsman-Berrevoets et al. 1979). The projection of the motor cortex to the magnocellular red nucleus is differently organized: it stems from neurons in deep layer 4 and constitutes a collateral projection of the pyramidal tract. The frontal eye field projects to Darkschewitsch’ nucleus and the dorsomedial subnucleus of the parvocellular red nucleus, and the supplementary eye field to the dorsomedial subnucleus only (Burman et al. 2000; Hartmann-von Monakow et al. 1979; Huerta and Kaas 1990; Huerta et al. 1986; Kuypers and Lawrence 1967; Leichnetz 1982; Leichnetz et al. 1984; Shook et al. 1990). The main lateral and caudal part of the parvocellular red nucleus receives overlapping, somatotopically arranged projections from the primary motor cortex, the supplementary motor area, and ventral and dorsal premotor area (Burman et al. 2000; Hartmann- von Monakow et al. 1979; J€ urgens 1984; Kuypers and Lawrence 1967; Leichnetz et al. 1984; Orioli and Strick 1989; Tokuno et al. 1995; Wiesendanger and Wiesendanger 1985). Posterior parietal afferents mainly terminate in Darkschewitsch’ nucleus; the projection to the parvocellular red nucleus is sparse (Burman et al. 2000; Faugier-Grimaud and Ventre 1989; Leichnetz 2001). Prefrontal projections from the prearcuate cortex dorsal to the principal sulcus, specifically from area 9, were noticed by Leichnetz (Leichnetz and Gonzalo-Ruiz 1996; Leichnetz et al. 1984). Because direct connections of the cerebral cortex with the inferior olive are few, most are shunted through the nuclei of the mesodiencephalic border. Alternative pathways use other preolivary nuclei such as ventral regions of the dorsal column nuclei that may serve as a relay between the motor cortex and the DAO and the MAOc in cat and rat (Ackerley et al. 2006; Andersson 1984; McCurdy et al. 1992). This region also receives a projection from the contralateral magnocellular red nucleus. Reports on inhibition of transmission in the inferior olive by rubral or cortical stimulation (Gibson et al. 2002) may be related to the presence of GABAergic neurons in this region that project to the inferior olive (Nelson and Mugnaini 1989). Another alternative pathway for inhibition of the DAOr passes

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through the anterior interposed nucleus, which receives a collateral projection from the rubrospinal tract (Huisman et al. 1983) and the nucleo-olivary pathway from this nucleus.

The Corticonuclear and Olivocerebellar Projections The Corticonuclear Projection The concept of the longitudinal zonal organization of the corticonuclear projection resulted from a combination of Weidenreich’s subdivision of the cerebellar nuclei with observations on the presence of parasagittally oriented white matter compartments in the cerebellum of the ferret and the cat (Voogd 1964, 1969). With the H€aggqvist myelin stain, bundles of medium-sized myelinated Purkinje cell axons could be distinguished in the cerebellar white matter separated by narrow slits that contain small myelinated fibers only (Fig. 19.15). Each compartment channels the Purkinje cell axons from a longitudinal Purkinje cell zone to a particular cerebellar nucleus. These Purkinje cell zones are illustrated for the ferret in Fig. 19.16. The A zone is present in the entire vermis and projects to the fastigial nucleus. The B zone occupies the lateral vermis of the anterior and posterior lobes; Deiters’ lateral vestibular nucleus is its target nucleus. The C1 and C3 zones merge in the ventral anterior lobe. They extend into the simplex lobule. C3 reappears in the Crus II and terminates in the rostral paramedian lobule; C1 is present in the medial paramedian lobule. C1 and C3 project to the anterior interposed nucleus. The C2 zone extends over the entire cerebellum, from lobule II of the anterior lobe to the paraflocculus and the flocculus. It connects with the posterior interposed nucleus. The D zone occupies the lateral hemisphere. In the ansiform lobule and the paraflocculus, it is divided into the medial D1 and the lateral D2 zones that project to ventrocaudal and rotrodorsal parts of the dentate nucleus, respectively. This longitudinal pattern in the corticonuclear projection was confirmed with retrograde axonal transport from the cerebellar nuclei (Fig. 19.17) (Bigare´ 1980; Voogd and Bigare´ 1980) and by Trott et al., using antegrade axonal tracing methods (Trott and Armstrong 1987; Trott et al. 1998), both in the cat. The B zone now was found to be restricted to the anterior vermis. Moreover, an injection of the lateral vestibular nucleus also produced labeled Purkinje cells in the anterior A zone (Fig. 19.17 1b). Exclusive labeling of this subpopulation was obtained from injections of the medial vestibular nucleus (Fig. 19.17 1c). In the dorsal anterior lobe, a gap between the labeled Purkinje cells in the A and B zones contains the X zone (Fig. 19.17 1c, arrow). In the anterior lobe, D1 and D2 zones are restricted to its dorsal folia. A very similar pattern in the corticonuclear projection was described for the anterior lobe of the macaque, the squirrel monkey, and the bush baby, and for the paramedian lobule and the paraflocculus of Tupaia glis using Nauta’s silver impregnation for degenerated axons (Haines et al. 1982). White matter compartments in the macaque monkey, delineated by the accumulation of

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Fig. 19.15 Reconstruction of the white matter compartments in the anterior lobe of the cerebellum of the ferret. Each compartment contains the cerebellar target nucleus of the corresponding Purkinje cell zone, illustrated in the A compartment with the fastigial nucleus. Bottom photograph: border between compartments A and B. H€aggqvist stain

acetylcholinesterase at the borders of the compartments, were found to be arranged like in the cat (Hess and Voogd 1986; Voogd et al. 1987a, b). The zonal organization of the corticonuclear projection was confirmed for the rat by Buisseret-Delmas with antegrade axonal transport of horseradish peroxidase (Buisseret-Delmas and Angaut 1993). In addition to the earlier identified set of zones, they described additional X, A2, and Y zones (Fig. 19.18). The X zone is located between the A and B zones in the dorsal anterior lobe; it was first identified in electrophysiological experiments on the olivocerebellar projection (see below). It projects to cell groups located between the fastigial and posterior interposed nucleus, known as the interstitial cell groups (Trott and Armstrong 1987; Trott et al. 1998). The A2 zone is located in the medial hemisphere of the simplex lobule and the Crus II (Akaike 1992). The dorsolateral protuberance of the fastigial nucleus is the target of its corticonuclear projection. The Y zone (BuisseretDelmas’ D0 zone) projects to the dorsolateral hump of the anterior interposed nucleus. Buisseret-Delmas located the Y zone between C3 and D1. In studies of the olivocerebellar projection, it was found to occupy a more lateral position, between D1 and D2 (Fig. 19.18).

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Fig. 19.16 Reconstructions of the Purkinje cell zones of the cerebellum of the ferret (Redrawn from Voogd (1969)). Abbreviations: ANS ansiform lobule, ANT anterior lobe, PFL paraflocculus, PMD paramedian lobule, SI simplex lobule

Another aspect of the corticonuclear projection was highlighted by the application of immunohistochemical methods, that distinguished between two populations of zonally distributed Purkinje cells, one population immunoreactive for zebrin (zebrin II: aldolase C), and a second population that was zebrin-negative. Alternating zebrin-positive and zebrin-negative bands are arranged in a reproducible pattern (Fig. 19.23). Axons of zebrin-positive Purkinje cells converge upon ventrocaudal and lateral parts of the cerebellar nuclei, and axons from zebrin-negative Purkinje cells terminate more rostrally (Fig. 19.25b). The use of the zebrin pattern as a template for the study of the olivocerebellar projection is considered in section “The Olivocerebellar Projection”.

The Olivocerebellar Projection In early morphological studies of the olivocerebellar projection, the origin of the climbing fibers from the inferior olive was not known. Axonal tracing methods visualizing the climbing fibers only became available in 1974 (Desclin 1974). Their origin from the olive had been established by Eccles et al. (1966), and Oscarsson and his group in Lund, acting on this observation, started their studies on spinoolivocerebellar climbing fiber paths (SOCPs) in the same year (Oscarsson and Uddenberg 1966). The first attempt at a map of the topographical organization of the olivocerebellar projection was by Holmes and Steward (1908) (Fig. 19.19). It

Fig. 19.17 Reconstructions of the anterior lobe (1), the dorsal aspect (2), and the posterior aspect (3) of the cerebellum of the cat, showing labeled Purkinje cells after injections of a retrograde tracer in a particular cerebellar nucleus. For each panel, the injected nucleus (MV medial vestibular nucleus) and the labeled Purkinje cell zone (A zone) is indicated (Redrawn from Bigare´ (Bigare´ 1980; Voogd and Bigare´ 1980)). Abbreviations: AI anterior interposed nucleus, ANS ansiform lobule, ANT anterior lobe, Cr I/II Crus I/II, Dc caudal dentate nucleus, Dr rostral dentate nucleus, F fastigial nucleus, LV lateral vestibular nucleus, MV medial vestibular nucleus, PFL paraflocculus, PI posterior interposed nucleus, PMD paramedian lobule, SI simplex lobule

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Fig. 19.18 Diagram of the zonal organization of the cerebellum of the rat. (a) Diagram of the flattened cerebellar cortex, the Purkinje cell zones are shown in the right half of the diagram. (b) Cerebellar and vestibular target nuclei of the Purkinje cell zones. (c) Diagram of the flattened inferior olive, showing subnuclei with projections to the Purkinje cell zones and their target nuclei. (d) Diagram of the zebrin-positive and zebrin-negative bands. Abbreviations: A-D2 Purkinje cell zones A-D2, 1-7 zebrin-positive bands P + 1-7, AI anterior interposed nucleus, Beta cell group beta, c caudal (MAO or DAO), C subnucleus C of the caudal MAO, DAO dorsal accessory olive, Dc caudal subnucleus of the dentate nucleus, DC dorsal cap and ventrolateral outgrowth, DLP dorsolateral protuberance of the fastigial nucleus, DMCC dorsomedial cell column, DM dorsomedial group, Dr rostral subnucleus of the dentate nucleus, F fastigial nucleus, i intermediate MAO, ICG interstitial cell groups, LV lateral vestibular nucleus, MAO medial accessory olive, PI posterior interposed nucleus, PO principal olive, r rostral (MAO or DAO), vest vestibular nuclei

remains the only diagram of its sort for the human cerebellum. The diagram shares its lobular organization with the diagrams produced for the olivocerebellar projection in rabbit and cat by Brodal 30 years later (Brodal 1940) (Fig. 19.20). Different subdivisions of the olive project to different lobules. The anterior lobe is an exception. In the rabbit, the extreme lateral portion of the anterior lobe, the “hemisphere proper,” receives afferents from the rostral pole of the principal olive. The accessory olives project to the vermis and to a paravermal, intermediate zone. This tripartition was confirmed in later studies of Jansen and Brodal (1940, 1942) of the corticonuclear projection, where the vermis was found to project to the

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Fig. 19.19 Diagram illustrating the topographical relations between the inferior olive and the human cerebellum. The dorsal accessory olive and the medial half of the dorsal leaf of the principal olive are connected with the cortex of the superior surface of the cerebellum. Olivocerebellar fibers from the ventral lamina of the principal olive and the ventral part of the medial accessory olive project to the tonsilla and the caudal vermis (From Holmes and Steward (1908)). Abbreviations: Bi biventral lobule, Ce central lobule, DAO dorsal accessory olive, Fl flocculus, Gr lobulus gracilis, Lg lingula, MAO medial accessory olive, PO principal olive, Qa anterior quadrangular lobule, Qp posterior quadrangular lobule, Si inferior semilunar lobule, Ss superior semilunar lobule, To tonsilla

fastigial and the vestibular nuclei, the intermediate zone to the interposed nucleus, and the hemisphere to the lateral cerebellar nucleus. These observations lead these authors to the conclusion that olivocerebellar and corticonuclear projections are arranged according to the same zonal pattern. The three-zonal arrangement only applies to the anterior lobe and the simplex lobule. Attempts have been made to extrapolate it to the posterior lobe, but these attempts have not met with much success.

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Fig. 19.20 The olivocerebellar projection in the rabbit indicated in flattened maps of the cerebellum and the inferior olive. The construction of the map of the olive is indicated in the bottom figurine. Olivary subnuclei and their projections are indicated with the same color (Redrawn from Brodal (1940)). Abbreviations: ANS ansiform lobule, ANT anterior lobe, DAO dorsal accessory olive, dc dorsal cap, dl dorsal lamina principal olive, dmcc dorsomedial cell column, F fastigial nucleus, FLO flocculus, INT interposed nucleus, LAT lateral cerebellar nucleus, MAO medial accessory olive, PFL paraflocculus, PMD paramedian lobule, PO principal olive, SI simplex lobule, vl ventral lamina principal olive

The conclusions of Voogd (1969) and Groenewegen and Voogd (1977) and Groenewegen et al. (1979) on the longitudinal zonal organization of the olivocerebellar projection were based on the observation that olivocerebellar fibers from subnuclei of the inferior olive use the white matter compartments to terminate on Purkinje cells of the corresponding longitudinal zone and, as collaterals, on its target nucleus. The organization of the olivocerebellar system is not a lobular but a zonal one. Groenewegen and Voogd (1977) and Groenewegen et al. (1979) used autoradiography of [3H]leucine to map the olivocerebellar projection in the cat. Earlier, Courville and colleagues (Courville 1975; Courville and Faraco-Cantin 1978; Courville et al. 1974) had demonstrated the termination of olivocerebellar fibers in longitudinal climbing fiber zones with the same method. However, they concluded that these zones corresponded to Brodal’s lobular pattern of termination. Groenewegen (Groenew(egen and Voogd 1977, Groenewegen et al. (1979) distinguished the same zones recognized earlier from their white matter compartments (Fig. 19.21). The A zone is innervated by the MAOc. It extends over the entire vermis, with the exception of lobule X that receives it climbing fibers, together with the flocculus, from the dorsal cap and the VLO. The DAOc projects to the B zone

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Fig. 19.21 Diagram of the zonal organization of the olivocerebellar and corticonuclear projections in the cat shown as flattened maps of the cerebellar cortex, the cerebellar nuclei and the inferior olive (Redrawn from Groenewegen et al. (1979)). Abbreviations: ANS ansiform lobule, ANT anterior lobe, DAO dorsal accessory olive, dc dorsal cap, Deiters’ Deiters’ lateral vestibular nucleus, dmcc dorsomedial cell column, FLO flocculus, MAO medial accessory olive, PFL paraflocculus, PMD paramedian lobule, PO principal olive, SI simplex lobule, vlo ventrolateral outgrowth

that is limited to the lateral vermis of the anterior lobe and the simplex lobule. C1 and C3 zones are innervated by the DAOr. They are present in the anterior lobe with the simple lobule and in the Crus II of the ansiform lobule and the paramedian lobule. C3 is restricted to the rostral paramedian lobule. Both zones fuse in the rostral anterior lobe, around the rostral extremity of the C2 zone. This zone receives its climbing fibers from the MAOr and extends over the entire cerebellar cortex, including the paraflocculus. The lateral D zone, innervated by the principal olive, also extends over the entire rostrocaudal length of the cerebellar cortex. At the level of the ansiform lobule and the paraflocculus, it is divided into D1 and D2 zones, but a differential origin of these climbing fiber subzones from the principal olive could not be established. Climbing fibers always emit collaterals to the cerebellar or vestibular target nucleus of the Purkinje cell zone they innervate. Such a collateral projection could not be identified for the projections of the VLO and the DC. Oscarsson and his group defined their spino-olivocerebellar climbing fiber paths (SOCPs) by the spinal funiculus they use as their first link. They mapped the terminations of these paths by recording the positive climbing fiber potentials

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Fig. 19.22 Distribution and properties of spino-olivary climbing fiber paths (SOCPs) in the anterior lobe of the cat. (a) Ventral funiculus (vf)-SOCP (Redrawn from Oscarsson and Sj€ olund (1977b)). (b) Dorsal funiculus (df)-SOCP (Redrawn from Ekerot and Larson (1979a)). (c) Somatotopical organization of the forelimb segment of the C3 zone. Color-coded map illustrates partially overlapping, half-moon-shaped terminal fields, activated from subsequently more cranial cutaneous nerves 1–8, indicated in upper figurine. Broken line indicates border of mirrored representations in medial and lateral C3 zone (Redrawn from Ekerot and Larson (1979b)). (d) Transverse branching of climbing fibers innervating dual zones (Redrawn from Ekerot and Larson (1982))

from the cerebellar surface and complex spikes from the underlying Purkinje cells from the anterior lobe and determined their laterality, somatotopical organization and the quality of their peripheral input. The ventral funiculus (vf)-SOCP takes its origin from the contralateral spinal gray, as illustrated in Fig. 19.4. It terminates in the vermis in the A zone, in the B zone, and in the C1 and C3 zones of the rostral anterior lobe (lobules I-IV). Input to the A, C1, and C3 zones stems from the hindlimb and is ipsilateral; the B zone receives a bilateral input from all limbs, with the projection of the forelimb located more laterally, partially overlapping with the projection from the hindlimb (Oscarsson and Sj€olund 1977a, b; Oscarsson and Uddenberg 1966) (Fig. 19.22a). The A and B zones receive their input through the cell groups in the ventral horn and/or the intermediate gray and the MAOc and the DAOc, respectively. The hindlimb input to the C1 and C3 zones is relayed through the cell groups in the dorsal horn that are absent in the cervical enlargement. Because the ventral funiculus was isolated at the C3 segment, they missed the C1 projection to the DAOr (Fig. 19.4) and its contribution to the rostral C1 and C3 zones. Projections to the posterior lobe were rarely studied by the physiologists. The C1 zone in the paramedian lobule was found to share climbing fiber collateral input with the C1 and C3 zones in the anterior lobe (Armstrong et al. 1973; Oscarsson and Sj€ olund 1977a). The presence of a C3 zone in the rostral paramedian lobule was never confirmed.

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The bilateral projection of the four limbs to the B zone was studied in more detail by Andersson and Oscarsson (1978). They described narrow, mediolaterally arranged longitudinal strips of Purkinje cells that receive the same climbing fiber input (i.e., mainly ipsilateral hindlimb, bilateral hindlimb, bilateral hind- and forelimb, etc., with a trigeminal microzone located most medially (Andersson and Eriksson 1981)). These “microzones” are narrow, with a width of a few Purkinje cells, but may extend for tens of millimeters in the rostrocaudal direction. The relationship of the microzones to the morphology of the climbing fiber system is discussed in ▶ Chap. 20 “Axonal Trajectories of Single Climbing and Mossy Fiber Neurons in the Cerebellar Cortex and Nucleus” (Shinoda and Sugihara). The dorsal funiculus (df)-SOCP was studied by Oscarsson (1969) and Ekerot and Larson (1979a, b). Its first link consists of spinal root fibers ascending in the dorsal columns. Its relay in the dorsal column nuclei and its projections to the inferior olive were discussed in section “Projections from the Dorsal Column Nuclei” and illustrated in Figs. 19.7, 19.8, and 19.9. The projection to the anterior lobe is ipsilateral for all the zones. Oscarsson (1969) documented ipsilateral hindand forelimb projections to the B zone and an ipsilateral hindlimb projection to A. These projections presumably overlap with similar projections from the vfSOCP. The relay for the B zone in the DAOc only has been substantiated for the hindlimb (Fig. 19.6). A substantial hindlimb projection from the gracile nucleus to MAOc would provide for the activation of the A zone. Ekerot and Larson (1979a, b) limited their study to the four short-latency zones, X, C1, C3, and D2, relayed monosynaptically by the dorsal column nuclei to the inferior olive, and two longlatency zones, C2 and D1, with an extra synapse, presumably in the mesencephalon (Fig. 19.22b). The C1, C3, and D2 (or Y) zones are innervated by the DAOr. Rostral hindlimb segments of the C1 and C3 zones overlap with the vf-SOCP, and their forelimb segments extend into lobule V. A projection of this subnucleus to the D2 zone was not noticed in earlier morphological studies. Originally, the D2 zone was defined by its lateral location, its projection to the rostral dentate, and its climbing fiber afferents from the dorsal lamina of the principal olive. Although Ekerot’s D2 zone is located in the most lateral part of the anterior lobe of the cat, its connections clearly differ from the original D2 zone. It should be considered as a third zone of the C1-C3 collective, innervated by the rostral DAO and projecting to the anterior interposed nucleus. The true D2 zone may have been inaccessible for the microelectrode, as it is hidden in the lateral pole of the anterior lobe of the cat, Garwicz (1997), in the ferret, proposed the neutral term Y zone for Ekerot’s D2 zone. In the rat, a D0 zone (Buisseret-Delmas 1989) was identified in the lateral anterior lobe and the paramedian lobule, located between the D1 and D2 zones (Voogd et al. 2003; Sugihara and Shinoda 2004). The D0 zone occupies a similar position as the Y zone in the cat. Its connections, however, are with subnuclei that have not been identified in carnivores, receiving climbing fibers from the dorsomedial group of the ventral lamina of the principal olive and projecting to the dorsolateral hump. Moreover, it gives rise to the uncrossed descending limb of the brachium conjunctivum (Mehler 1969) that has not been identified in carnivores either.

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The X zone is located between the A and B. It is a pure forelimb zone located in lobule V. It receives its climbing fibers from intermediate levels of the MAO (Campbell and Armstrong 1985) (illustrated in Fig. 19.18 for the rat). The forelimb projection from the cuneate nucleus to the intermediate MAO is illustrated in Fig. 19.6. A trigeminal input to this region is illustrated in Figs. 19.4 and 19.5. Alternative routes for the transmission of peripheral input to the inferior olive involve the crossed ascending projections of somatosensory relay nuclei to the nuclei of the mesodiencephalic junction, referred to before. For the long-latency projection to D1, the pathway probably involves Bechterew’s nucleus, a subsidiary of the parvocellular red nucleus, and the ventral lamina of the principal olive (Fig. 19.13). The pathway for the C2 zone includes Darkschewitsch’ nucleus (Fig. 19.13) and the MAOr. The D1 zone displays an indistinct somatotopical organization; C2 lacks a somatotopical arrangement. The somatotopical organization of the forelimb region of the C3 zone was studied in more detail. Climbing fibers transmitting information from cutaneous nerves terminate in partially overlapping half-moon-shaped fields in this zone. When stimulating thoracic, ulnar, radial, and neck nerves, the fields shift rostrally in this order (Fig. 19.22c). Apparently, the C3 zone contains two mirror-faced somatotopical maps in its medial and lateral halves. A similar, but single, somatotopical organization is present in the C1 and the Y (D2) zone. Transverse climbing fiber branching was found between medial C3 and C1 and between lateral C3 and Y (D2) (Armstrong et al. 1973; Ekerot and Larson 1982) (Fig. 19.22d). Moreover, climbing fibers branch between the X zone and a new CX zone, located between C2 and C1. The presence of zonal pairs sharing the same topically arranged peripheral climbing fiber input, enclosing a zone innervated by climbing fibers carrying information descending from the mesodiencephalic junction, is a remarkable, but still unexplained feature of the anterior lobe. In rat and mouse transverse branching in the anterior lobe among the C1, C3, and D0 zones in lobules VI and VI between A and A2 and in the copula pyramidis in the C1 zone, was noticed with antegrade labeling of climbing fibers by Sugihara and Shinoda (2004) and Sugihara and Quy (2007). Apart from tactile input, the df-SOCP climbing fibers of the df-SOCP were found to relay nociceptive input. This quality is relayed by the postsynaptic dorsal column pathway, and not by the spinal root fibers of the dorsal funiculus (Ekerot et al. 1991; Uddenberg 1968). Nociceptive input is found for the X, C1, CX, and C3 zones (Garwicz et al. 1992). In the most recent chapter in the study of the olivocerebellar projection, the “zebrin pattern” was used as a template. Antibodies known as the zebrins exclusively stain a subpopulation of Purkinje cells in rodents that are arranged in parallel longitudinal bands, separated by bands of zebrin-negative Purkinje cells (Hawkes and Leclerc 1987) (Fig. 19.23). One of these antibodies, zebrin II, recognizes the enzyme aldolase C (Hawkes and Herrup 1995). Earlier, the same pattern was described by Scott (1964) for the enzyme 5-nucleotidase in the molecular layer of the cerebellum of the mouse. In the nomenclature devised by Hawkes, zebrin-positive bands are numbered from medial to lateral as P1+ to P7+, and the zebrin-negative bands

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Fig. 19.23 Diagram of zebrin-positive (black) and zebrin-negative (white) Purkinje cell bands in the cerebellum of the rat. Zebrinpositive (P+) bands are numbered according to Hawkes and Leclerc (1987). Non-numbered satellite bands are shown in red. Abbreviations: II-IX lobules II-IX, COP copula pyramidis, Cr I/II Crus I/II, PMD paramedian lobule, SI simplex lobule

P1- to P6- are located lateral to the zebrin-positive bands bearing the same number (Fig. 19.23). The “satellite bands” distinguished by these authors (red in Fig. 19.23) were not numbered. Later they were found to be a reproducible feature and indicated with lower case characters (Figs. 19.18, 19.23, and 19.24). The main question that remained was whether the zebrin-positive and zebrinnegative bands were identical to the corticonuclear and olivocerebellar projection zones. The answer to this question was provided by Voogd et al. (2003), using a bidirectional axonal tracer, injected into identified zones of the cerebellar cortex of the rat and, in much greater detail, by Sugihara c.s who mapped the olivocerebellar projection from numerous small injections of an antegrade axonal tracer in subnuclei of the inferior olive in rat (Sugihara and Shinoda 2004) and mouse (Sugihara and Quy 2007) in material counterstained for aldolase C (zebrin II). Additional studies were published by Voogd and Ruigrok (2004) and Pijpers et al. (2005). Earlier, the olivocerebellar projection in the rat had been described by Buisseret-Delmas and Angaut (1993). Their findings largely confirmed the earlier studies in carnivores. Of the additional zones described by them, the X zone receives it climbing fibers from the lateral intermediate MAOc. The A2 zone is innervated by the medial MAOc, and the Y (Delmas’s D0 zone) by the dorsomedial group of the ventral lamina of the principal olive (Fig. 19.18a, c).

Fig. 19.24 (continued)

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In Voogd’s studies, the corticonuclear and olivocerebellar projection zones were found to be congruent with zebrin-positive and zebrin-negative bands. The C2, D1, and D2 zones consist of zebrin-positive Purkinje cells, and the X, B, C1, C3, and Y zones are zebrin-negative (Fig. 19.18.). Because these zebrin-negative zones are absent from the Crus I of the ansiform lobule and the paraflocculus, these lobules are entirely zebrin-positive, and the borders of the constituent C2, D1, and D2 zones cannot be distinguished. This is also the case for the X, B, and C1 zones that occupy the anterior zebrin-negative P2-band. It follows from the analysis of the olivocerebellar projection that Hawkes nomenclature for the zebrin bands in the anterior and posterior cerebellum is not consistent. Anterior bands P2+ to P6+ are continuous with posterior bands P3+ to P7+. The A2 zone occupies a region in the Crus II, containing the zebrin-positive and zebrin-negative P4b and P5a bands. Anteriorly, it corresponds to the c and d bands of the simplex lobule. In the following account, anterior and posterior bands that receive their climbing fibers from the same subdivision of the inferior olive will be indicated with their numbers separated by a dash (i.e., anterior band 2+ and posterior band 3+ are indicated as 2+/3+). Greater detail was obtained in Sugihara and Shinoda’s study (Sugihara and Shinoda 2004) of the olivocerebellar projection in the rat. They distinguished four groups in this projection (Fig. 19.24a, b). Group I (green) includes the lateral subnucleus a of the MAOc, the MAOr, and the principal olive. It projects to zebrinpositive bands P1, with the exception of lobule VII, 2+/3 + and to the 4+/5+, 5+/6+ and 6+/7+. With the exception of subnucleus a of the MAOc that receives somatosensory input, it receives its afferents from the nuclei at the mesodiencephalic junction. Group II (blue) comprises the projections of caudal subnucleus c of the MAOc, group beta and the DMCC and the caudal part of the dorsomedial group to zebrin-positive bands a+/2+, 3+, and 4+ in the uvula and a complicated array of zebrin-positive bands in intermediate regions of lobule VI-VIII (2b+, c+, d+, 4b+, 5a+). Of this group, the beta nucleus and the DMCC receive vestibular input, and lateral MAOc shares the tecto-olivary projection with the group III and receives a spatially segregated somatosensory afferents. Group III (yellow) includes the projections of subnucleus c of the MAOc to zebrin-negative bands in the vermis and in intermediate regions of lobules VI-VIII. In group IV (red), the DAO and the rostral part of the dorsomedial group project to anterior bands 2-, b+ and -, 3+ and -, 3b+ and -, 4- and 5- and posterior 4-, e1 and 2+ and -, 5- and 6-. This region includes ä Fig. 19.24 (a) Olivocerebellar projections to aldolase-C-positive and aldolase-C-negative Purkinje cell zones. The four groups in this projection, their origin from the olive and their termination in the cortex, are indicated in different colors. Group I (green) and group II (blue) project to aldolase-positive bands. Group III (yellow) and group IV (red) project to aldolasenegative bands. (b) Flattened reconstruction of the cerebellar cortex showing aldolase-C-positive and aldolase-C-negative bands. (c) Correspondence of aldolase C banding pattern with A-D zones and their origin from the inferior olive. The A and A2 zones correspond with multiple aldolase-Cpositive (dark blue and dark orange) and aldolase-C-negative (light blue and light orange) bands (a and b, Redrawn from Sugihara and Shinoda (2004))

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both zebrin-negative, and the weakly zebrin-positive bands 3+ and 3b+ in the anterior cerebellum and e1, and e2 in the posterior cerebellum. Group IV is purely somatosensory. The correspondence between the four groups of Sugihara and Shinoda (2004) and the nomenclature of Voogd (Fig. 19.18) and Buisseret-Delmas (1993) is indicated in Fig. 19.24C. The localization of X, B, C1-3, Cx and the D1, Y, and D2 zones is immediately clear. The position of the D1 and D2 zones, flanking the Y (Buiseret-Delmas’s D0 zone), now was definitely established. The possibility still exists that D1 is discontinuous in the Crus I (lobule VIc) because no labeling was observed in this region with injections of the vlPO. The main difference with previous maps is the composition of the A and A2 zones of an array of interdigitating zebrin-positive and zebrin-negative bands. The principle, formulated by Groenewegen et al. (1979) that olivocerebellar fibers emit collaterals that terminate in the target nucleus of the Purkinje cells innervated by these fibers, has not been challenged. The collateral olivocerebellar projection to the cerebellar nuclei was studied in the rat by Ruigrok and Voogd (2000) and Sugihara and Shinoda (2007). The collateral projections of subnuclei of the inferior olive were found to overlap with the corticonuclear projections from the Purkinje cell zones that receive climbing fibers from these subnuclei, as reviewed in the previous paragraphs. Zebrin (aldolase C) immunochemistry cannot be used to map the corticonuclear projection of the individual zones. However, it may serve to visualize the overall topography of the corticonuclear projection of zebrin-(aldolase C)-positive and negative Purkinje cell zones as labeling of fibers and terminals in the cerebellar nuclear neuropil (Hawkes and Leclerc 1986; Sugihara and Shinoda 2007). Zebrinpositive and zebrin-negative zones that interdigitate in the cerebellar cortex, converge upon two separate homogeneously zebrin-positive or zebrin-negative regions of the cerebellar nuclei, that occupy caudoventral and lateral, and rostrodorsal parts of the nuclei, respectively (Fig. 19.25b). As a result, the medial (fastigial) nucleus is divided into the rostrodorsal, zebrin-negative and caudoventral, zebrin-positive parts, which may be related to some functional localization in this nucleus. In the interposed nucleus, this division between the rostrodorsal and caudoventral parts roughly corresponds to the division between the anterior and posterior interposed nuclei. The lateral (dentate) nucleus is entirely aldolase-C-positive, in accordance with its innervation by zebrin-positive bands only. Collateral projections to the fastigial nucleus from groups I and II that innervate zebrin-positive zones include those from subnucleus a of the MAOc (green in Fig. 19.25), group beta and DMCC (light blue), and subnucleus c (dark blue). These collateral projections occupy subsequently more dorsal and rostral laminae (Fig. 19.25a, b, g, h). The collateral projection of subnucleus c to lobules VI and VII, the so-called visual vermis, demarcates the visuomotor subdivision of the fastigial nucleus. Subnucleus b of the MAO that belongs to group III, which innervates zebrin-negative territory, supplies collateral projections to the rostral and dorsal fastigial nucleus. Olivocerebellar fibers from medial subnucleus b that innervate the zebrin-negative bands of the A2 zone emit collaterals to

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Fig. 19.25 (a) Collateral projections of olivocerebellar fibers to the cerebellar nuclei in the rat illustrated in diagrams of parasagittal (a–f) and transverse sections (g, h). Projections from the four groups in the olivocerebellar projection, distinguished in Fig. 19.24a, are shown in the same colors. Within group II, the projection to lobules VI and VII (the “visual vermis”: dark blue) is distinguished from other lobules (light blue). Within group III, the projections of the dorsolateral protuberance and medial posterior interposed nucleus (orange) are distinguished from those to rostral fastigial nucleus and interstitial cell groups (yellow). (b) Caudal view of a 3-D reconstruction of the caudoventral and lateral aldolase-C (zebrin-positive) and rostral aldolase-C-negative compartments of the cerebellar nuclei of the rat (Redrawn from Sugihara and Shinoda (2007)). Abbreviations: AI anterior interposed nucleus, AICG anterior interstitial cell groups, C caudal, CVP caudal pole, D dorsal, DLH dorsolateral hump, DLP dorsolateral protuberance, DMC dorsomedial crest, ICG interstitial cell groups, L lateral cerebellar nucleus, LV lateral vestibular nucleus, M medial cerebellar nucleus, PI posterior interposed nucleus

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Fig. 19.26 Afferent climbing fiber and efferent projections of the flocculus and the nodulus and adjacent lobules. (a) Zonal organization of the olivocerebellar projection to the flocculus and the adjacent ventral paraflocculus and to the nodulus and the uvula in the rat and their origin from the inferior olive, shown in flattened diagrams of these structures. (b) Efferent connections of the flocculus and the nodulus in rabbits. Note different localization of group beta innervated zone in the nodulus of rat and rabbit. Abbreviations: DC dorsal cap, DM dorsomedial group, f0-f4 floccular zones f0-f4, Fast, fastigial nucleus, MAOc/r caudal/rostral medial accessory olive, MV medial vestibular nucleus, n1-n5 nodular zones 1–5, PI posterior interposed nucleus, PO principal olive, ß group beta, SV superior vestibular nucleus, VLO ventrolateral outgrowth, Y group Y

the dorsolateral protuberance and spill over in the adjacent posterior interposed nucleus (orange). Lateral subnucleus b innervates the base of the DLP and the interstitial cell groups (yellow). The latter represents the collateral projection of the X zone. Collateral projections from the MAOr and the ventral and dorsal laminae of the PO, that belong to group I (green), terminate in the zebrinpositive neuropil of the posterior interposed nucleus and in the caudal and rostral portions of the dentate nucleus, respectively. Collateral projections of

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group IV include those of the DAOr that innervate the anterior interposed nucleus (red), and the DAOc that provide collaterals to the lateral vestibular nucleus and to islands of gray matter between the anterior interposed and lateral nuclei, known as the anterior interstitial cell groups (AICG, pink). The neuropil of the target nuclei of group IV is devoid of zebrin-positive elements. Olivocerebellar projections to the flocculus and the nodulus and neighboring lobules have received much attention. In the flocculus of the rabbit, five Purkinje cell zones and corresponding white matter compartments could be distinguished (Tan et al. 1995a, b, c) (Fig. 19.26a). The caudal extension of the C2 zone is located along its lateral border. Actually, this lateral border corresponds to the inner, medial border of the folial chain of the hemisphere that is turned back upon itself as the flocculus. The f1 and f3 zones receive a projection from the rostral dorsal cap (DC) and the ventrolateral outgrowth (VLO), the f2 and f4 zones from the caudal DC. In the flocculus of the rat, the C2 zone and the four floccular zones, with the same dual projection from the DC and the VLO, could be distinguished (Ruigrok et al. 1992). A fifth DC-innervated f0 zone was identified by Sugihara et al. (2004) (Fig. 19.26a). A differential projection of caudal and rostral parts of the VLO was found by these authors, confirming earlier observations by Gerrits (1982) in the cat. A similar zonal organization is present in the flocculus of the mouse (Schonewille et al. 2006). In monkeys, the f4 zone appears to be absent (Voogd et al. 1987a, b). In all species, the floccular zones extend for some distance on the ventral paraflocculus. In the rabbit, this extension is known as folium P (Yamamoto 1978, 1979), in the cat as the medial extension of the ventral paraflocculus (ME: Gerrits and Voogd 1982), and in monkeys, the floccular zones occupy the entire ventral paraflocculus (Voogd et al. 1987b). Collateral projections from the DC terminate in the rostrolateral ventral dentate nucleus, from the VLO in its medial part and in the dorsal group Y (Sugihara et al. 2004). Climbing fibers from the VLO also terminate in the extreme lateral part of the ansiform lobule (Fig. 19.24). Olivocerebellar fibers from DC and VLO do not collateralize to the vestibular nuclei (Ruigrok and Voogd 2000; Sugihara et al. 2004). Collateral projections from the group beta and a lateral part of subnucleus b of the MAOc to parts of the vestibular nuclei were identified by Ruigrok (unpublished) and Sugihara and Shinoda (2007), respectively. The zonal organization of the nodulus and the uvula is fairly complicated. In the uvula, four zebrin-positive bands are present, separated by narrow zebrinnegative slits. These zebrin-negative slits disappear in the ventral uvula. Purkinje cells in the ventral uvula and the nodulus are all zebrin-positive (Fig. 19.26) (Voogd et al. 1996). In the medial and lateral uvula, the n1 and n5 zones are innervated by the DC, and an intermediate N3 band receives climbing fibers from the VLO. Of these zones, only n3 continues into the ventral uvula. In the uvula, the caudal group beta innervates zebrin-positive band P1+ and medial P2+, rostral group beta innervates lateral P2+ and medial

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P3+, and climbing fibers terminating in lateral P3+ and in P4+ are derived from the DMCC. The narrow P1-, P2-, and P3- bands receive climbing fibers from subnucleus b of the MAOc. According to Sugihara and Shinoda (2004), climbing fibers terminating in P4+ are derived from the caudal part of the dorsomedial group and the most caudal and medial part of the vlPO and the dorsomedial group is an alternative source for the P3- projection. The localization of the DC and VLO-innervated zones was confirmed for the rat by Ruigrok (Ruigrok 2003), who also demonstrated collateralization of climbing fibers between the projections from the DC to the rostral DC-innervated zone in the ventral paraflocculus and the n1 and n5 zones in the nodulus and for the VLO between f1 and f3 and n3. The efferent connections of the Purkinje cell zones of the flocculus and the nodulus have been studied in rabbits (Fig. 19.26b). The DC-innervated f2 and f4 zones project to oculomotor relay cells in the medial vestibular nucleus, the VLOinnervated f1 and f3 zones project to oculomotor relay cells in the group Y and the superior vestibular nucleus (De Zeeuw et al. 1994b; Tan et al. 1995a). The zonal organization of the rabbit nodulus differs from the rat in the presence of a medial beta-innervated zone, but DC and VLO-innervated N1, N3, and n4 zones are present in the same configuration as in the rat. All the zones project to the medial vestibular nucleus; additional projections to the superior vestibular nucleus and group Y take their origin from n3 and n4 (Wylie et al. 1994). Terminations in the superior and medial vestibular nuclei are complementary to those from the flocculus (Angaut and Brodal 1967; Haines 1977; Voogd 1964). All or most of the Purkinje cell zones of the flocculus and the nodulus project to the basal interstitial nucleus.

The Nucleo-olivary Pathway The pathway from the cerebellar nuclei to the contralateral inferior olive was discovered by Graybiel et al. (1973) in the cat. It takes its origin from the small, GABAergic neurons in all cerebellar nuclei (Mugnaini and Oertel 1985). The projection is reciprocal with respect to the collateral projections of the subnuclei of the inferior olive to the cerebellar nuclei and has been described in the rat (Ruigrok and Voogd 1990), the cat (Courville et al. 1983a; Legendre and Courville 1986; Tolbert et al. 1976), and the monkey (Beitz 1976; Chan-Palay 1977b; Kalil 1979). Nucleo-olivary fibers from the dentate and interposed nuclei ascend in a separate bundle, ventral to the brachium conjunctivum, decussate caudal to the brachium, and descend in the tegmentum to the olive. Nucleo-olivary fibers from the fastigial nucleus take a more diffuse route to the olive (Legendre and Courville 1986). GABAergic projections from the vestibular nuclei and the group Y to the DC, VLO, the group beta, and the DMCC were considered in section “Optokinetic and Vestibular Projections to the Inferior Olive”. In the rat, small numbers of fibers recross at the level of the olive. Recrossing fibers mainly terminate in MAOr, vlPO,

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Fig. 19.27 White matter compartments in the anterior lobe of Macaca fascicularis. Processed for acetylcholine esterase. Abbreviations: IntA anterior interposed nucleus; bc brachium conjunctivum, cr restiform body

and in the dorsomedial group (Ruigrok and Voogd 1990). In all species, a projection from the ventrocaudal dentate to the vlPO, and from the rostral dentate to the dlPO, was found.

Olivocerebellar and Corticonuclear Projections in Primates The presence of corticonuclear projection zones A-D in prosimians, macaques, and squirrel monkeys was demonstrated by Haines and his colleagues for the anterior lobe of these species (Haines et al. 1982). A division of the D zone into D1 and D2 zones was not observed by these authors. The parasagittal organization of white matter compartments in the macaque cerebellum, visualized by the accumulation of acetylcholinesterase at their borders, was found to be very similar to earlier observations in carnivores (Voogd et al. 1987a) (Fig. 19.27). An X compartment was recognized in the anterior cerebellum; D1 and D2 compartments that issue at the caudal and more rostral portions of the dentate nucleus were recognized in the dorsal paraflocculus and the paramedian lobule. In experiments using autoradiography of antegradely transported tritiated leucine, the A, X, B, and C2 compartments were shown to channel the olivocerebellar fibers to their respective Purkinje cell zones.

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Fig. 19.28 Aldolase-C-positive and aldolase-C-negative bands in a flattened reconstruction of the cerebellar cortex of the marmoset (left). The correspondence with the A, C1, C2, C3, D1, and D2 zones, based on the afferent climbing fiber and efferent nuclear connections of the aldolase-Cpositive and aldolase-C-negative bands, is indicated in the right hand panel. The symmetry axis for the numbering of the bands and the areas without cortex in the center of the ansiform loop are shown in red (Redrawn from Fujita et al. (2010)). Abbreviations: ANT anterior lobe, FLO flocculus, PFL paraflocculus, PMD paramedian lobule, SI simplex lobule, I-X lobules I-X

Knowledge of the corticonuclear and olivocerebellar projections in subhuman primates remains fragmentary, particularly with respect to the connections of the dentate nucleus. A major advance was the publication of a complete map of the aldolase-C (zebrin II)-positive and negative bands in the marmoset, a small primate (Fujita et al. 2010) (Fig. 19.28). For the vermis, the zebrin pattern in primates was analyzed by Sillitoe and colleagues (2005), but the hemisphere remained unexplored. The banding pattern of the marmoset cerebellum is very similar to the rat (Fig. 19.24). A, C1, C2, C3, D1, and D2 zones could be recognized using small injections of bidirectional tracers in specific aldolase-C-positive or aldolaseC-negative bands. D1 receives a projection from the vlPO and D2 from the dlPO. In the posterior lobe, the P5a bands appear to be absent. In the rat, these bands are part of the A2 zone. The target of the P5a- band, the dorsolateral protuberance of the fastigial nucleus, is not present in the marmoset either. A reduction of the A2 zone in this species, therefore, appears likely.

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Fig. 19.29 Diagram of the projections of the fastigial nucleus. Projections of the rostral and ventrocaudal fastigial nucleus are indicated in yellow and those from its visuomotor division in green. The recurrent climbing fiber pathway from the superior colliculus is shown in red. Abbreviations: coll.sup superior colliculus, Fr rostral division fastigial nucleus, Fvc ventrocaudal division fastigial nucleus, I-X vermal lobules I-X, MAOc caudal medial accessory olive, med.s.ret medial reticular formation, no nucleo-olivary pathway, PPRF pontine paramedian reticular formation, riMLF rostral interstitial nucleus of the medial longitudinal fascicle, vest.nu vestibular nuclei, vis.fast visuomotor division fastigial nucleus

The Cerebellar Nuclei: Efferent Connections and Recurrent Climbing Fiber Paths The Fastigial Nucleus The fastigial nucleus receives projections from the anterior vermis and lobule VIII in its rostral portion, from the oculomotor vermis (lobule VII) in a caudal subdivision, and from lobules IX and X in its ventrocaudal part (Fig. 19.29). In rodents, a dorsolateral protuberance is present that receives Purkinje cell axons from the A2 zone (Buisseret-Delmas 1988) (Fig. 19.18). The rostral fastigial nucleus projects bilaterally and symmetrically to the vestibular nuclei (magnocellular medial and descending vestibular nuclei, nucleus parasolitarius) and the medial bulbar and pontine reticular formation (Batton et al. 1977; Homma et al. 1995; Teune et al. 2000). The entire contralateral projection uses the uncinate tract (Thomas 1897) that decussates in the cerebellar commissure and hooks over the brachium conjunctivum to enter the vestibular nuclear complex from laterally. It emits an

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Fig. 19.30 Labeled neurons in the rat fastigial nucleus from an injection of a retrograde tracer affecting the entire left-sided output of the nucleus. Abbreviations: DLP dorsolateral protuberance, Fr rostral fastigial nucleus, Fvc ventrocaudal fastigial nucleus, PI posterior interposed nucleus

ascending branch (Probst 1901) consisting of scattered fibers around the medial pole of the brachium, that does not join it in its decussation. The direct fastigiobulbar tract enters the vestibular nuclei from medially. In the mouse, the ipsilateral pathway was found to originate from glycinergic neurons (Bagnall et al. 2009). The distribution of these glycinergic cells in the ventral and rostral fastigial nucleus in the mouse corresponds closely to the neurons with ipsilateral projections in the rat (Fig. 19.30). The projections of the caudal fastigial nucleus and the dorsolateral protuberance are entirely crossed. The connections of the caudal, oculomotor division of the fastigial nucleus have been studied mainly in monkeys (Noda et al. 1990) (Fig. 19.29). This subdivision appears to be the main source of the crossed ascending branch of the uncinate tract. Its bilateral projection to the vestibular nuclei and the bulbar reticular formation overlaps with the rostral fastigial nucleus. Specific contralateral projections of the oculomotor division include the pontine paramedian reticular formation (PPRF)

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Fig. 19.31 Diagram of the projections of the anterior interposed nucleus. The recurrent climbing fiber pathway from the pretectum is indicated in red. Abbreviations: AI anterior interposed nucleus, Ant.pret. nu anterior pretectal nucleus, DAOr rostral dorsal accessory olive, M primary motor area, no nucleo-olivary pathway, Rmc magnocellular red nucleus, Thal thalamus, VIM ventral intermediate thalamic nucleus

that contains the excitatory and inhibitory burst cells for horizontal eye movements, the rostral interstitial nucleus of the medial longitudinal fascicle (riMLF) with similar neurons for vertical eye movements, the rostral intermediate layer of the superior colliculus, where fibers cross in the tectal commissure to innervate the contralateral side (May et al. 1990), and parts of the suprageniculate, ventral intermediate, ventral lateral and intralaminar thalamic nuclei. A recurrent tecto-olivary pathway takes its origin from the intermediate layer of the superior colliculus to terminate in the contralateral medial MAOc (Figs. 19.12 and 19.30). In all species, this subnucleus projects to the oculomotor vermis. In rodents, the tecto-olivary pathway also innervates a separate neuronal population in subnucleus b of the MAOc that projects to the A2 zone and its target nucleus, the dorsolateral protuberance (Akaike 1992; Ruigrok and Voogd 2000). The contralateral projection of the dorsolateral protuberance to the brain stem is mainly directed at the lateral, parvocellular reticular formation and the adjoining spinal nucleus of the trigeminal nerve, the parabrachial nuclei, the nucleus pedunculopontinus, and the deep mesencephalic nucleus. This system appears to absent or rudimentary in primates and carnivores.

Anterior Interposed Nucleus Projections from the anterior and posterior C1, C3, and Y zones converge on to the anterior interposed nucleus, where they are arranged into a single somatotopical map (Garwicz and Ekerot 1994). The main projections of the anterior interposed nucleus include the magnocellular red nucleus, the nucleus reticularis tegmenti pontis, and, through the ventral intermediate and ventral lateral thalamic nuclei, the primary motor cortex (Fig. 19.31). A recurrent pretecto-olivary pathway arises from the anterior pretectal nucleus and relays in the DAOr (Kawamura and Onodera

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1984; Kawamura et al. 1982; Sugimoto et al. 1982) (see also section “Afferents from Tectum and Pretectum” and Figs. 19.12 and 19.31). In rodents, the dorsolateral hump (Goodman et al. 1963) (Fig. 19.1) may be considered as a subnucleus of the anterior interposed nucleus. It receives a projection of the D0 zone, the presumed equivalent of the Y zone in the cat (Sugihara et al. 2009) (Fig. 19.18). The D0 zone and the dorsolateral hump are innervated by the dorsomedial group of the ventral lamina of the principal olive. The dorsolateral hump gives origin to the uncrossed descending branch of the superior cerebellar peduncle (Cajal 1972; Mehler 1969). It terminates in the lateral parvocellular reticular formation and the adjoining spinal nucleus of the trigeminal nerve (Teune et al. 2000), where it overlaps with terminals of the contralateral dorsolateral protuberance. Both of these systems appear to be absent in non-rodent species.

Posterior Interposed Nucleus and Interstitial Cell Groups The posterior interposed nucleus is the recipient of Purkinje cell axons of the C2 zone that extends over the entire rostrocaudal length of the cerebellar cortex from lobule III into the flocculus. The MAOr provides the C2 zone with its climbing fibers and the posterior interposed nucleus with a collateral projection. The MAOr receives its descending input through the medial tegmental tract from Darkschewitsch nucleus located at the mesodiencephalic junction (Ogawa 1939). This nucleus is a relay in the projection from frontal eye fields to the rostral pole of the MAOr and motor and posterior parietal cortices to its caudal parts (section “Nuclei at the Mesodiencephalic Border: The Central and Medial Tegmental Tracts” and Fig. 19.13). The rostral pole of the MAOr projects to the flocculus and the adjacent ventral paraflocculus. The dorsal paraflocculus and the ansiform lobule receive their climbing fibers from successively more caudal parts of the MAOr. The caudal portion of the MAOr, which receives motor and parietal input, projects to the C2 zone of the anterior lobe and the paramedian lobule (Brodal and Kawamura 1980). Laterocaudal visuomotor and rostromedial skeletomotor divisions also have been recognized in the posterior interposed nucleus (van Kan et al. 1993). The laterocaudal visuomotor division receives a corticonuclear projection from the paraflocculus (Xiong and Nagao 2002). The topical projections of the more rostral segments of the C2 zone to the posterior interposed nucleus have not yet been studied. Efferent connections of the posterior interposed nucleus (Fig. 19.32a) include the contralateral magnocellular red nucleus, Darkschewitsch’ nucleus, the nearresponse region located dorsal to the oculomotor nuclei which is required for vergence eye movements (May et al. 1992), the superior colliculus and, as thalamocortical projections, the primary motor and premotor cortex and the frontal eye fields and posterior parietal areas (Kievit 1979; Lynch and Tian 2006; Matelli

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Fig. 19.32 Diagrams of the projections of the posterior interposed nucleus (a), the caudal dentate nucleus (b), and the rostral dentate nucleus (c). Recurrent climbing fiber pathways are indicated in red. Abbreviations: ctt central tegmental tract, coll.sup superior colliculus, Dark Darkschewitsch’ nucleus, Dc caudal dentate nucleus, Dr rostral dentate nucleus, FA supplementary eye field, FE frontal eye field, MAOr rostral medial accessory olive, mt medial tegmental tract, no nucleo-olivary pathway, Par parietal cortex, PI posterior interposed nucleus, POdl dorsal lamina principal olive, POvl ventral lamina principal olive, Rpc DM dorsomedial subdivision parvocellular red nucleus (Bechterew’s nucleus), Rpc lat lateral subdivision parvocellular red nucleus, Thal thalamus, VIM ventral intermediate thalamic nucleus, VLO ventral lateral thalamic nucleus oral part, VL-X ventral lateral thalamic nucleus, nucleus X

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and Luppino 1996; Matelli et al. 1989). Another link with the frontal eye field and, possibly, parietal areas is provided by the superior colliculus (Lynch et al. 1994; Tian and Lynch 1997). Extensive projections to the medial intraparietal area MIP, from the anterior and paramedian C2 zone, through the laterocaudal half of the posterior interposed nucleus and a more restricted projection of its laterocaudal pole to the ventral lateral intraparietal area LIP were documented by Prevosto et al. (2009). Efferents of the posterior interposed nucleus largely overlap with those from the anterior interposed and dentate nuclei. Darkschewitsch nucleus constitutes the link in the direct and indirect reciprocal climbing fiber paths (Fig. 19.32a). The interstitial cell groups share projections to the vestibular nuclei, the reticular formation, the red nucleus, and the thalamus with neighboring nuclei (Buisseret-Delmas et al. 1998; Teune et al. 2000). A specific feature of this cell group is its collateral projection to the spinal cord with the caudal medial reticular formation and to the thalamus with the superior colliculus (Bentivoglio and Kuypers 1982).

Dentate Nucleus The dentate nucleus can be divided into rostrodorsal and caudoventral parts that receive corticonuclear projections of the D1 and D2 zones, respectively, and collaterals from the olivocerebellar fibers that innervate these two zones (Fig. 19.18). Rostral motor and ventrocaudal non-motor divisions were distinguished by Strick in the cebus monkey (Strick et al. 2009). A visuomotor division was identified in the ventrocaudal pole of the monkey dentate (van Kan et al. 1993). It is not known whether these different modes that partition the dentate nucleus define the same rostrocaudal subdivisions. The caudal visuomotor division of the dentate nucleus gives rise to a component of the brachium conjunctivum that terminates contralaterally in the dorsomedial parvocellular red nucleus, the superior colliculus and medially in the ventrolateral thalamic nucleus including area X, with minor extensions in the adjoining mediodorsal and anterior nuclei (Fig. 19.32b). The rostral dentate projects to the lateral and caudal parvocellular red nucleus and the ventral intermediate and lateral portions of the ventral lateral thalamic nucleus (Chan-Palay 1977; Kievit 1979) (Fig. 19.29c). Nucleo-olivary pathways connect the rostral dentate with the dorsal lamina and the lateral bend of the inferior olive, and the caudal dentate with the ventral lamina (section “The Nucleo-olivary Pathway”). Strick’s distinction in the monkey dentate of rostrodorsal motor and ventrocaudal non-motor divisions is based on retrograde transneuronal labeling experiments. Neurons in the motor division can be labeled from the primary and premotor cortex including the supplementary motor area (SMA). Neurons in the caudal non-motor division are connected with the preSMA and the prefrontal areas 9d and 46 (Fig. 19.33). A projection to the frontal eye field was located in the

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Fig. 19.33 Diagram illustrating transneuronal retrograde labeling from injection sites in different cortical areas in the cebus monkey. (a) Localization of injection sites resulting in labeling within the dentate nucleus (red) and in cases without labeling of this nucleus (blue). (b) Diagram of the unfolded dentate nucleus of the cebus monkey, showing approximate position of the labeling from injections of different cortical areas. Broken line indicates border between motor and non-motor divisions of the dentate nucleus. Question mark indicated the non-explored medial lamina of the dentate nucleus. Red ellipse indicates extent of the labeling from an injection of the anterior intraparietal area that includes both the motor and non-motor divisions of the nucleus (Reproduced and modified from Strick et al. (2009)). Abbreviations: IP anterior intraparietal area, AS arcuate sulcus, CgS cingulate sulcus, CS central sulcus, FEF frontal eye field, PS principal sulcus, IP intraparietal sulcus, LS lateral sulcus, Lu lunate sulcus, M1 primary motor cortex, PMv ventral premotor area, PreSMA rostral division of the supplementary motor area, SMA supplementary motor area (SMA proper), ST superior, TE temporal lobe, ST temporal sulcus

extreme caudal pole of the nucleus (Strick et al. 2009) (Fig. 19.32b, c). Neurons projecting to the anterior intraparietal area (AIP; Clower et al. 2005) are located in both the motor and non-motor divisions, and those projecting to the medial intraparietal (MIP) mainly occupy the rostral dentate (Prevosto et al. 2009). Projections to parietal area 7b (Clower et al. 2001) and the lateral intraparietal area (LIP; Prevosto et al. 2009) are derived from its non-motor division.

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The dorsomedial parvocellular red nucleus, the medial tegmental tract, and the ventral lamina of the principal olive are links in direct and indirect recurrent projections from the frontal eye fields and parietal areas to the caudal dentate and the D1 zone. The lateral and caudal parvocellular red nucleus, the central tegmental tract, and the dorsal lamina of the principal olive are links in the recurrent pathway from motor and premotor cortical areas to the rostral dentate and the D2 zone (Figs. 19.14 and 19.32b, c). The zonal organization of the connections of the inferior olive and the cerebellar nuclei, as described in this chapter, adequately covers the anatomy of the cerebellum of lower mammals. However, our knowledge of the connections of the hemisphere of the great apes and the human cerebellum, which accounts for the bulk of the cerebellar cortex in these species, remains incomplete. The demonstration by Fujita et al. (2010) of the presence of D1 and D2 zones in the marmoset hemisphere does support the idea that the construction of the primate hemisphere is similar to that in rodents and carnivores, but the relative proportions of the marmoset hemisphere are not very different from rodents. The source of the climbing fiber projection to the D1 zone in rodents and primates is the ventral, nonconvoluted lamina of the principal olive (section “The Corticonuclear and Olivocerebellar Projections”). If this subdivision of the principal olive is represented in the human inferior olive by its medial lamina (Fig. 19.3), the contribution of the human D1 zone to the hemisphere would be small indeed. Its major output would be represented by the D2 zone, with its projection to the convoluted dentate nucleus. In this case, the subdivision of the human dentate in rostrodorsal microgyric and caudolateral macrogyric portions would correspond with two major subdivisions of the D2 zone. There is no evidence on the corticonuclear projection of the human cerebellar hemisphere. The only observations on the connections of the two parts of the human dentate stem from papers on crossed cerebro-cerebellar atrophy by Demole´ (1927a, b) and Verhaart and Wieringen-Rauws (1950). Demole´ concluded that degeneration of the rostrodorsal, microgyric dentate occurred with large, chronic lesions involving the pre- and postcentral gyri. Lesions of more posterior parts of the hemisphere, involving parietal, occipital, and temporal areas, resulted in atrophy of the ventrocaudal macrogyric dentate. Prefrontal lesions would spare the dentate nucleus. These observations still need to be confirmed but, at least, emphasize the possibility that the increase in size of the cerebellar hemisphere is also related to the elaboration of its connections with postrolandic areas, in addition to the expansion of the cerebello-prefrontal connectivity, as advocated by many cognitive neuroscientists. Interestingly, Masao Ito (2012) in his recent book “The cerebellum. Brain for an Implicit Self” proposes a neural control system for mental activities that includes both prefrontal and postrolandic areas and the cerebellar hemispheres. In this system, the prefrontal cortex serves as the controller, the representation of the controlled subjects is found in the temporoparietal cortex, and the cerebellar hemispheres provide the internal models of the controlled objects represented in the temporoparietal cortex.

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Axonal Trajectories of Single Climbing and Mossy Fiber Neurons in the Cerebellar Cortex and Nucleus

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Yoshikazu Shinoda and Izumi Sugihara

Abstract

A major factor in determining the function of a particular cerebellar cortical region depends upon its afferent and efferent connections. Two distinct afferent pathways convey information to the cerebellar cortex: climbing fibers and mossy fibers. A large amount of fundamental knowledge about afferent projections to the cerebellum from various precerebellar nuclei has been accumulated by using new anatomical methods, but the axonal trajectories of single climbing fiber neurons in the inferior olive and single mossy fiber neurons of multiple sources have not been well understood. The knowledge about the morphologies of single mossy fiber neurons and climbing fiber neurons in the cerebellum is essential for our understanding of the function of the cerebellum. This article will describe and compare the entire axonal trajectories of single olivocerebellar (OC) neurons and single mossy fiber neurons from the lateral reticular nucleus, pontine nucleus and dorsal column nucleus in the cerebellar cortex and nucleus. Furthermore, this chapter will deal with the relationship between the longitudinal cortical and nuclear compartmentations revealed by aldolase C expression and the longitudinal bands of cortical and nuclear axonal terminals of single climbing fiber neurons and single mossy fiber neurons, and discuss the functional significance of these organizations to generate the final output from the cerebellar nucleus to the targets outside the cerebellum for control of movement and other functions.

Y. Shinoda (*) • I. Sugihara Department of Systems Neurophysiology, Graduate School of Medicine, Tokyo Medical and Dental University, 1-5-45, Yushima, Bunkyo-ku, Tokyo, 113-8519, Japan e-mail: [email protected], [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 437 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_20, # Springer Science+Business Media Dordrecht 2013

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Introduction The Purkinje cell is the only output neuron of the cerebellar cortex (Cx), and the cortical neuronal circuitry in which Purkinje cells (PCs) are embedded remains remarkably constant in different cerebellar regions. Therefore, a major factor in determining the function of a local cerebellar area depends upon its afferent and efferent connections and the relationship between the two (Eccles et al. 1967; Ito 1984; Voogd 1969). Two distinct afferent pathways which arise from many different origins (vestibular, spinal, and cerebral cortical) convey information to the Cx. The afferents that project to the cerebellum via precerebellar nuclei except the inferior olive (IO) take the form of mossy fibers (MFs) in the Cx, while the afferents to the Cx via the IO (olivocerebellar axons) take the form of climbing fibers (CFs). CFs originate only from the contralateral IO, and each PC is directly innervated by a single CF (Cajal 1911; Szentha´gothai and Rajkovits 1959). On the other hand, MFs which originate from multiple sources enter the cerebellum rostrally, and many cross the midline in the cerebellar commissure and project bilaterally. The cerebellar nuclei (CN) comprise the fastigial (medial) (FN), interposed (IP), and dentate (lateral) nuclei (DN), and three subdivisions of the corticonuclear projection are identified: medial (vermis), intermediate, and lateral zones projecting to the FN, IP, and DN, respectively (Jansen and Brodal 1940). In the Cx, longitudinal compartmentation beyond these three subdivisions was revealed by the demonstration of zones A–D based on cholinesterase staining in the cerebellar white matter and the topography of the corticonuclear and olivocortical projections (Voogd 1964, 1969; Groenewegen and Voogd 1977; Kawamura and Hashikawa 1979; Buisseret-Delmas and Angaut 1993). These longitudinal zones extend across one or more lobules, even across the entire rostrocaudal length of the Cx (Voogd and Bigare´ 1980; Voogd et al. 1987). The olivocerebellar (OC) projection is also arranged according to the same principle (Voogd 1969; Oscarsson 1969). For a proper understanding of cerebellar function, it is essential to know how MF and CF afferent systems interact with these cortical longitudinal zones defined both anatomically and electrophysiologically, and emit output signals to the CN via PCs, and how these PC output signals interact with nuclear collateral inputs from MF and CF afferents in nuclear neurons and generate final cerebellar output signals to their targets outside the cerebellum. Neuroanatomists have tried to reveal the unknown neural connections in the mammalian central nervous system (CNS). Along with the development of new anatomical techniques, a large amount of fundamental knowledge about afferent systems to the cerebellum has been accumulated. However, the conclusions reached by neuroanatomists are not necessarily correct, since the following methodological advantages and disadvantages of different anatomical techniques have not been properly assessed. 1. The degeneration method, the most classical for analyzing fiber connections, was used for analysis of cerebellar afferent projections and greatly promoted with the advent of the Nauta method. But this method is capricious and its most serious drawback is that it is very difficult to distinguish degenerated passing fibers from degenerated terminals.

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2. Great technical development in neuroanatomy was achieved in 1970–1990 by the introduction of new retrograde tracers such as horseradish peroxidase (HRP), wheat germ agglutinin-horseradish peroxidase (WGA-HRP), and cholera toxin B subunit. These methods greatly expanded our knowledge about fiber connections in the CNS. However, one serious problem in these retrograde-labeling methods is that these tracers are taken up not only by nerve terminals but also by fibers of passage. The projections from the precerebellar nuclei to the CN was investigated by injecting retrograde traces into the CN, but many studies reported false positive data about collateral projections to the CN. As pointed out by Dietrichs et al. (1983), injection tracks to deliver a tracer solution into the CN inevitably pass through the Cx, or tracers might be taken up by passing fibers in and around the CN. 3. Injection of an orthograde tracer into a certain nucleus is generally useful for detecting projections from the injected nucleus to multiple target regions, although this method cannot differentiate whether single neurons in the nucleus project to two different targets via collaterals, or neurons that project to individual targets coexist in the nucleus. New orthograde tracers were introduced, first WGA-HRP and later Phaseolus vulgaris-leucoagglutinin (PHA-L) and biotinylated dextran amine (BDA). As described for retrograde labeling, tracer uptake by passing fibers may occur for orthograde labeling. For example, in the case of an injection into the pontine nucleus, a tracer is often taken up by passing fibers from the nucleus reticularis tegmenti pontis or contralateral pontine nucleus. PHA-L and BDA were reported to be taken up by passing fibers especially when injected in a large amount, but it does not occur significantly when injected in a small amount. Afferent projections were also investigated by means of anterograde transport of tritiated amino acids such as leucine, since these amino acids are considered to be taken up only by cell bodies. However, termination or fine passing fibers could not be confirmed with certainty from the light microscopical material, and, in addition, this method could not determine whether labeled materials are either axon collaterals from stem axons of MFs or CFs projecting to the Cx, or axon terminals of axons that specifically project to the CN without any projection to the Cx. 4. Despite abundant anatomical studies on the CF and MF systems, there is little information available on the organization of CF and MF projection at the level of single neurons. The Golgi method was the only staining method for observing single cell morphology, but this method could not adequately delineate axonal morphology because it is difficult to impregnate myelinated axons in adult tissue, and the neural processes pass out of the plane of section. Therefore, most of the neurons in the mammalian CNS remain terra incognita as far as their axonal trajectories are concerned. A new intracellular staining method with fluorescent dyes made it possible to study the morphology of physiologically identified neurons (Stretton and Kravitz 1968), but fluorescent dyes were not suitable for staining thin axon collaterals in the mammalian CNS. For visualization of the entire axonal morphology of a single neuron with a long axon, HRP was successfully used for intracellular staining (Jankowska et al. 1976; Snow et al. 1976; Kitai et al. 1976), and Kitai et al. (1976) succeeded in staining single

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electrophysiologically identified PCs with this method. Although all of the processes of a single neuron labeled with a tracer are not contained in a single section, the reconstruction of the axonal trajectory using serial sections makes it possible to reveal the entire axonal trajectory of a single labeled neuron, and, at present, neurons with long axons such as corticospinal and vestibulospinal axons can be visualized with this method over a distance of 10–30 mm (Shinoda et al. 1981, 1986). The single cerebellar afferent fibers first stained with this method were functionally identified MFs that responded to muscle stretch (Krieger et al. 1985). Since then, the branching patterns of cerebellar afferent and efferent systems were investigated with this method (Shinoda et al. 1992). However, intracellular iontophoretic injection of HRP into a cell or an axon requires highly proficient skills and the same result can be achieved by the extracellular injection of BDA. Extracellularly injected BDA is taken up by dendrites and cell bodies in the vicinity of an injection site and carried to terminals arising from stained neurons. By controlling the amount of injection, it is possible to label a limited number of neurons or even a single neuron, which makes it possible to reconstruct the entire trajectories of single cerebellar afferent neurons from serial sections (Sugihara et al. 1999; Wu et al. 1999). This review will summarize and compare the characteristic features of the axonal projection patterns of single CF and MF neurons in the Cx. In addition, the projections of single CF and MF neurons will be correlated to cerebellar cortical longitudinal zones and nuclear zones classified by aldolase C expression. Besides the projection to the Cx, CF and MF projections to the CN have not been fully investigated. The CN gives rise to cerebellar outputs to target structures outside the cerebellum (Toyama et al. 1968; Uno et al. 1970; Allen and Tsukahara 1974; Shinoda et al. 1985a, b; Futami et al. 1986; Sato et al. 1996, 1997), and efferent CN neurons receive inhibitory input from PCs (Eccles et al. 1967; Ito et al. 1970). The presence or absence of excitatory inputs to the CN from the precerebellar nuclei is important for understanding cerebellar function (Shinoda et al. 1993, 1997). It had commonly been assumed that neurons in the precerebellar nuclei project to the Cx and that all have axon collaterals to the CN. This view has been echoed in textbooks, reviews, and even research articles without any experimental evidence. However, cerebellar anatomists could not definitely confirm the existence of afferent axons to the CN with the classical degeneration method. The existence of axon collaterals of OC axons to the CN was first demonstrated by Gerrits and Voogd (1987), using a modern reliable anatomical staining method. The existence of the projections of MFs to the CN had been more controversial (Dietrichs et al. 1983; Chan-Palay 1977; Ito 1984). By introducing an intraaxonal staining method with HRP, electrophysiologically identified MFs were visualized using serial sections, and the existence of nuclear axon collaterals of single MF axons terminating in the Cx was definitely demonstrated (Shinoda et al. 1992). However, in spite of a wealth of anatomical reports on the afferent projections to the Cx, there have been far fewer studies on afferent projections to the CN. Therefore, this review also deals with the presence or absence of cerebellar afferent projections to the CN and discusses the functional significance of these projections for output generation in the cerebellum for motor control.

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Axonal Trajectories of Single Olivocerebellar Axons The entire axonal trajectories of OC neurons were completely reconstructed on serial sections in the rat (Sugihara et al. 1999). For clarification in the following description, the entire axon of an IO neuron is called an OC axon, while a thick branch of the OC axon in the Cx that terminates on dendrites of a PC in the molecular layer is called a CF (Cajal 1911). Stem axons left the IO toward the contralateral side, crossed the midline, and ran transversely above or through the contralateral IO (Fig. 20.1b). After entering the white matter of the inferior cerebellar peduncle (ICP), they ran longitudinally through the dorsolateral ICP and in the white matter rostral and dorsal to the CNs (Voogd 1995), but sometimes they ran through or beneath the CNs (Fig. 20.2c). Axons projecting to the vermis through the rostral ICP entered the deep cerebellar white matter rostral to the FN, whereas those projecting to the lateral and intermediate hemisphere through the caudal ICP entered the cerebellar white matter rostral to the DN. OC stem axons successively ramified into many branches in the deep cerebellar white matter, when the stem axons reached the semi-parasagittal plane in which axonal branches terminated as CFs. These axonal branches entered the folial white matter and while ramifying, ran together in the semi-parasagittal plane to reach the areas for their terminations in the Cx contralateral to the IO of their origin (Fig. 20.2a). The branches of OC axons were grouped into thick branches (0.7–1.4 mm in diameter) and thin collaterals (0.2–0.5 mm), and the diameter of stem axons was within the range of that of thick branches (Fig. 20.1a). A single OC axon generated 2–17 final thick branches (CFs) (6.1
3.7, mean
S.D., n ¼ 16), each of which innervated a single PC as a CF. The number of CFs of a single OC axon is consistent with the average number of CFs per OC axon (about seven in the rat) inferred by counting the total numbers of PCs and IO neurons (Schild 1970).

Distribution of CFs The most distal portion of each thick branch of an OC axon reached a target PC usually at its soma, and a terminal arborization on a single PC was always formed by a single CF. The number of swellings on the terminal arborization of a single CF was 154–321 (249.8
42.2, n ¼ 32). The diameter of swellings ranged from 0.5 to 2.0 mm, mostly 1.2–1.6 mm. Assuming 6.1 CFs per OC axon, 249.8 swellings per CF corresponded to 1,524 swellings on CF terminal arborizations per OC axon, on average. The cortical distribution of CFs of a single OC axon conformed to the pattern predicted from previous mass labeling studies of the OC projection; namely, all CFs of a single OC axon terminated in a single strip-shaped longitudinal band in the Cx. This band spread widely in a rostrocaudal direction, covering single or multiple lobules, but only narrowly in a mediolateral direction (usually about 200–300 mm) (Fig. 20.2). In the example shown in Fig. 20.2, six labeled OC axons in the medial accessory olive (MAO) ran close to each other in the brain stem and the most rostral ICP (Fig. 20.2c). Stem axons curved caudally and ran closely together rostral and dorsal to the FN and started ramifying in the deep white

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a

Crus Ia D M

L V

Crus Ib

0.5 mm

Sim b

b Crus Ia

DN PIN

FN

DN ICP PFL

FL 1 mm

ICP IO

Fig. 20.1 Frontal view of the trajectory and the distribution of climbing fibers (CFs) of a single olivocerebellar (OC) axon innervating crus Ia. Biotinylated dextran amine (BDA) was injected into the dorsal lamella in the principal olive of the inferior olive (IO). A filled arrowhead indicates a thin collateral in the inferior cerebellar peduncle (ICP), an arrow the first bifurcation into two thick branches, and open arrowheads the first two branching points of thin collaterals that terminated in the granular layer. All collaterals of this axon could be traced to their terminals. FL flocculus, FN fastigial nucleus, PFL paraflocculus, PIN posterior interpositus nucleus, Sim b lobule simplex b, D, V, L and M dorsal, ventral, lateral and medial (From Sugihara et al. 1999)

matter in a rostrocaudal direction. The branched main axons ascended in the white matter in the same parasagittal plane toward the vermal cortex of lobules VI and VII, and, ramifying in a rostrocaudal direction in the folial white matter of individual lobules, terminated as CFs in the molecular layers of single (brown axon), multiple separate (light blue axon), or adjacent lobules (pink, green, orange, and purple axons) (Fig. 20.2a). As in this example, IO neurons in a very restricted area of the IO have common features in axonal coursing and terminal distribution in

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a

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VIa VIb VIc 2 mm VII

VIa Rostral Lateral

Medial Caudal

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500 µm 500 µm

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Fig. 20.2 Reconstruction of the entire trajectories of 6 OC axons originating from a centromedial portion of the medial accessory olive (MAO). (a, c) Lateral view of 6 OC axons. CF terminals of each axon are distributed widely in a rostrocaudal direction through lobules VI and VII, but all of the terminals are distributed in a limited mediolateral width of about 0.3 mm. CFs in gray do not belong to the six reconstructed OC axons represented by different colors. Dotted lines, borders of the granular layers. V–VIII and X, lobules V-VIII and X. (b) Longitudinal band of CFs of the 6 OC axons in the cerebellar cortex (Cx). CFs of individual OC axons are plotted on the unfolded parasagittal strip of the cerebellar cortical surface of lobules VI and VII (stippled area in the inset in a). Colors used for CFs in (b) correspond to those used for the OC axons in (a) and (c). Light and dark gray areas in the unfolded scheme represent the Cx exposed on the cerebellar surface and hidden in the sulci, respectively. Note that thin axon collaterals of the OC axons terminate in the FN (From Sugihara et al. 2001)

a particular lobule or lobules. Increasing volumes of BDA injected into a particular portion of the IO subnucleus resulted in the rostrocaudal extension of the projection area in a single longitudinal band, but with only slight extension in the mediolateral

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width. This finding indicates that adjacent IO neurons project to a set of segments in the same longitudinal band rather than to multiple segments in separate longitudinal bands (Sugihara et al. 2001), and that individual longitudinal compartments of single OC axons are much narrower than classical longitudinal zones of A, B, C, and D (Voogd 1969). The width of a longitudinal compartment innervated by a single OC axon almost corresponded to the width of the mediolateral distribution of PCs that show synchronous firing of complex spikes (Sugihara et al. 1995).

Thin Collaterals of OC Axons Thin collaterals, which were given off from stem axons and thick branches of OC axons, were more abundant in terms of the number per OC axon than thick branches. For convenience of description, they were classified into three types according to their main termination sites: (1) white matter in the ICP, (2) the CNs, and (3) the cerebellar granular layer. All of the thin axon collaterals and their terminals in the ICP, the CNs, and the granular layer were derived from OC axons terminating as CFs in the Cx, and none of the OC axons terminated specifically in the CNs without projecting to the Cx. Thin collaterals to the CNs were observed in 20 out of 22 OC axons and each of these 20 axons had one (n ¼ 15), two (n ¼ 3), three (n ¼ 1), or six (n ¼ 1) collaterals (1.4
1.2 collaterals per OC axon) that innervated only a single CN (Sugihara et al. 1999). In the example in Fig. 20.2, all OC axons projected to lobules VI and VII of the vermis, and their nuclear collaterals were given off only to the FN. The diameters of nuclear collaterals were very thin, ranging from 0.2 to 0.3 mm, and their terminal branches bore several swellings of en passant and terminal types. The average number of swellings per nuclear OC collateral was 54.0
66.0 (n ¼ 22). Thin collaterals terminating mainly in the granular layer were given off from stem or thick branches of OC axons in the deep and folial white matter (Fig. 20.1a). Swellings were densest in the upper portion of the granular layer, but were seen at all depths in that layer. Some swellings in the granular layer seemed to make contact with the soma of a presumed Golgi cell, and other swellings were located among a dense aggregation of granule cells. Thin collaterals usually terminated in the same lobule and the same parasagittal zone as CFs of the same OC axons. The number of thin collaterals in the granular layer ranged from 3 to 16 (average, 8.5) per OC axon (see details of thin collaterals in Sugihara et al. 1999).

Relationship Between the Longitudinal Distribution of CFs of Single OC Axons and Aldolase C Bands in the Cx The cortical longitudinal zones are classically divided into A, B, C1, C2, C3, D1, and D2 (Voogd 1969). An electrophysiological study showed that the Cx is composed of fine longitudinal strips with a width of 0.3 mm and length of around 3 mm, and this functional unit is called a “microzone” (Oscarsson 1979). Even finer longitudinal compartmentation in the Cx was defined based on the expression

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pattern of aldolase C (zebrin II). Hawkes and Leclerc (1987) showed that a subset of PCs and their axons that were stained with antizebrins were arranged in longitudinal zones alternating with strips of non-zebrin-immunoreactive PCs, and that most zebrin-positive bands continued uninterruptedly from the anterior to the posterior lobe. Recently, a correspondence between the OC projection pattern and aldolase C compartments of some parts of the Cx have been examined using a combined orthograde labeling of OC axon terminals and aldolase C immunohistochemistry in mass labeling materials (Voogd et al. 2003; Voogd and Ruigrok 2004; Pijpers et al. 2005, 2006). Figure 20.3 shows an example of the relationship between CFs of single OC axons and aldolase C expression pattern in the Cx in a systematic study of the entire Cx and IO (Sugihara and Shinoda 2004). A comprehensive twodimensional map of the aldolase C compartments was first formed in the entire unfolded Cx from serial sections. CFs labeled by a small injection of BDA into the IO were then mapped onto this aldolase C map generated from the same rat. Labeled CFs following injection into the caudolateral MAO (red) were distributed in a longitudinal band of aldolase C-negative band (1-) and spread widely even in the anterior and posterior vermis. Similar distributions of labeled CFs following injections into different parts of the MAO were observed in aldolase C-negative longitudinal bands (green and light blue) and aldolase C-positive band (purple) (Fig. 20.3). However, even though CFs shown in the same color seemed to be located in a single continuous longitudinal band in the anterior and posterior lobes, they belonged to bands of different names in the anterior and posterior lobes (e.g., green CFs in the anterior lobe are located in the (1-) compartment, whereas green CFs in the posterior lobe in the (2-) compartment). Single OC axons often innervate the rostral and caudal cerebellum by their axon collaterals (Sugihara et al. 2001). Taking advantage of this property, a specific pair of rostral and caudal aldolase C compartments were correctly linked based on the common projection of single OC axons. Whereas previous studies presumed that rostral and caudal compartments with the same nomenclature corresponded to each other (Hawkes and Leclerc 1987; Brochu et al. 1990), it turned out that this relationship did not always hold (see the links between rostral and caudal aldolase C compartments in Table I in Sugihara and Shinoda 2004). Individual longitudinal bands of CFs in different colors (Fig. 20.3) were aligned in the new pairs of the linked rostral and caudal compartments determined in relation to the common topographic OC projection to the pair of compartments in the anterior and posterior lobes.

Compartmentation of the CN and Its Relationship to the Cortical Compartments The compartmentation of OC projection in the Cx suggests that an equivalent compartmentation may exist in the CNs, because PCs project topographically onto the CNs (Buisseret-Delmas and Angaut 1993) and topographically projecting OC axons give rise to collaterals to the CN (Sugihara et al. 1999, 2001). Most single OC axons have axon collaterals to the CN on their way to the main cortical

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Fig. 20.3 Reconstruction of aldolase C compartments throughout the Cx and the relationship of CF longitudinal bands to aldolase C (zebrin II) pattern. Continuity of the compartments in the rostrocaudal direction was determined by precise reconstruction of individual compartments on serial sections and then plotted on a map of the unfolded Cx (see Sugihara and Shinoda 2004). The nomenclatures of compartments were adopted according to Hawkes and Leclerc (1987), Voogd et al. (2003), and Voogd and Ruigrok (2004). An aldolase C-positive compartment and its laterally neighboring negative compartment usually have the same name with different suffixes (+, positive compartment; , negative compartment). I–X, Lobules I–X; a–c, sublobules a–c. Distribution of CFs labeled following injection of BDA into the MAO in the four experiments is superimposed on the aldolase C map. The colored dots plotted on an unfolded representation of the Cx indicate locations of individual CFs labeled by injection into four different parts of the MAO. Narrow longitudinal band-shaped OC projection from a single injection site connects corresponding rostral and caudal aldolase C compartments (From Sugihara and Shinoda 2004)

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projection area (Sugihara et al. 1999). In the example in Fig. 20.2, OC axons originating from the MAO projected to lobules VI and VII in the vermis and to the FN by their axon collaterals. Taking advantage of this innervation property of single OC axons, the functional relationship between the OC compartments of the CN and the Cx was systematically investigated (Sugihara and Shinoda 2007). First, aldolase C expression of PC axon terminals was used to map the compartments of the CNs. The localization of PC terminals with or without aldolase C classified the CNs into aldolase C-positive and negative groups, respectively (Fig. 20.4C). The DN and the posterior interpositus (PIN) were aldolase C-positive, whereas the anterior interpositus (AIN) was aldolase C-negative. The FN was divided into the rostral aldolase C-negative and the caudal aldolase C-positive compartments (Fig. 20.4D). Then, a pair of the cortical and the nuclear aldolase C compartments innervated by single OC axons were systematically mapped following injections of a small amount of BDA into various parts of subnuclei of the IO (Fig. 20.4A). In each IO injection case, CFs of labeled OC axons were mapped within cortical aldolase C compartments (Fig. 20.4B), and then terminals of nuclear collaterals of the same OC axons were mapped in aldolase C compartments of the CN (Fig. 20.4C). Based on the results of this analysis, the olivocorticonuclear projection was classified into five “groups” of functional compartments (represented by five different colors in Fig. 20.4A–C) (Sugihara and Shinoda 2007). Each group originated from a subarea within the IO (Fig. 20.4A) and projected to multiple cortical strips of PCs, all of which were either aldolase C-positive (groups I–II and V) or aldolase C-negative (groups III and IV). As mentioned above, the CNs were divided into caudoventral aldolase C-positive (DN, PIN, and caudal FN) and rostrodorsal aldolase C-negative parts (AIN and rostral FN) (Fig. 20.4C, D). The olivonuclear terminations of the five groups projected topographically to five separate compartments within the CNs with the same aldolase C expression (either positive or negative). A PC axon formed a terminal arbor in a specific small area in the CN, and rostrocaudal PCs in a single longitudinal band of aldolase C-positive or C-negative cortical compartment converged to the aldolase C-positive or C-negative nuclear compartment, respectively (Sugihara et al. 2009). Therefore, whether PCs were located in the anterior or posterior lobe, aldolase C-positive PCs projected to the caudoventral aldolase C-positive nuclear compartments, and aldolase C-negative PCs projected to the rostrodorsal aldolase C-negative nuclear compartments. These findings suggest that each nuclear compartment and its corresponding cortical longitudinal strip innervated by the common OC axons are connected to each other by PCs with the same properties in aldolase C expression, and these three form a functional unit in the cerebellum. This structure may be a morphological correlate for a functional unit of a “corticonuclear microcomplex” proposed by Ito (1984).

Morphology of Single Mossy Fibers Quantitatively important sources of MFs are the spinal cord, the pontine nuclei (PNs), the vestibular nuclei, the lateral reticular nucleus (LRN), and the dorsal

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Fig. 20.4 Clustering of termination areas of nuclear collaterals of OC axons in the cerebellar nuclei (CNs) classified into five groups based on their cortical projection patterns. (A) Mapping of 123 injection sites on the dorsal view of individual subnuclei of the right IO. (B) Mapping of labeled CF terminals in aldolase C compartments on the unfolded cortical scheme. Each injection site was classified into one of the five groups based on its projection pattern to aldolase C compartment by referring to the classification of OC projection (Sugihara and Shinoda 2004, their Fig. 20.8a). The injection sites in the IO (A) and their corresponding labeled CFs (B) and olivonuclear termination areas (C) are indicated by different colors, depending on which group they belong to (green, group I; blue, group II; yellow, group III; red, group IV; gray, group V). (C): Rostral (a), dorsal (b), and caudal (c) views of the left CNs showing three-dimensional distribution of nuclear termination areas of 123 olivary injections shown in (A). (D) Three-dimensional reconstruction of the aldolase C-positive (gray) and -negative area in the left CNs shown in a dorsocaudal view for comparison. The wire frames are coronal contours of the cerebellar nuclei. Gray and darker gray areas indicate aldolase C-negative and C-positive compartments, respectively. PO and DAO principal and dorsal accessory olive, CP copula pyramidis in (B), Cr I and Cr II crus I and II of ansiform lobule, Par paramedian lobule, pf primary fissure, VPFL ventral paraflocculus, DC dorsal cap of Kooy, VLO ventrolateral outgrowth, ICG the interstitial cell group, DMC the dorsomedial crest, CP the caudal pole in (C) and (D); DLH the dorsolateral hump, DLP dorsolateral protuberance, AIN the anterior interpositus nucleus, d-Y dorsal group Y nucleus (From Sugihara and Shinoda 2007)

column nuclei (DCNs). However, our knowledge about the precise terminations of most afferent MF systems, especially their single axonal projection patterns in the Cx and CN, is far from complete. Since the cerebellar MF projection from the LRN

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has been investigated well in the rat, this section will describe single axonal morphologies of LRN neurons first and the similarity and difference of the projection pattern between the LRN and other MF systems later.

Lateral Reticular Nucleus Neurons General Features of Axonal Trajectories of Single LRN Neurons Mass labeling studies using autoradiography (K€ unzle 1975: Chan-Palay et al. 1977) and PHA-L (Ruigrok and Cella 1995) clearly showed that MFs of the LRN terminate in longitudinal strips in the granular layer. Data obtained from complete reconstruction of the entire trajectories of single LRN axons confirmed their findings and further extended the information about the projection pattern (Wu et al. 1999). A majority of LRN axons passed through the ipsilateral ICP to the cerebellum (n ¼ 25 out of 29)

Fig. 20.5 Reconstruction of the entire trajectory of an MF axon originating from a lateral reticular nucleus (LRN) neuron in the cerebellum and brain stem. The reconstruction of the axon labeled with extracellular iontophoretic application of BDA was made from 158 serial sections of 50 mm thickness. IP posterior interpositus nucleus, sct spinocerebellar tract, sptV spinal trigeminal tract, SPVI nucleus interpolaris (From Shinoda 1999)

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Fig. 20.6 Frontal (top) and laterall views (bottom) of a completely reconstructed single LRN axon originating from the caudal part of the right LRN. This axon enters the cerebellum through the right icp, projects to the vermis bilaterally (lobules II through VII) and the FN (see the bottom drawing e), and forms a multiple longitudinal zonal projection pattern by its cortical arborescent collaterals. Bottom drawings show lateral views of individual collaterals as indicated in the top drawing with lower case letters (a–g) (Modified from Wu et al. 1999)

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(Fig. 20.5), but some axons (n ¼ 4) ran through or under the contralateral IO and entered the cerebellum through the contralateral ICP. Generally, an arbor of an LRN axon within the cerebellum could be regarded as consisting of a thick stem axon (2.5–3.5 mm in diameter) running transversely, cortical branches of various diameters (around 2 mm), and very thin nuclear branches (0.4–1.0 mm). Most stem axons crossed the midline within the cerebellum to make bilateral projections with ipsilateral preponderance. While running transversely in the cerebellar white matter rostral and dorsal to the CNs toward the contralateral side, stem axons gave rise to several primary collaterals to the Cx and CN. These cortical collaterals arose almost perpendicularly from the stem axons, and ran almost in the parasagittal plane. On their way, individual collaterals widely ramified mainly in a dorsoventral direction, but did not spread so much in a mediolateral direction, so that each collateral terminated as an MF rosette in the granular layer in a relatively narrow longitudinal zone covering one to four lobules (Fig. 20.5). The number of cortical primary collaterals given off from a transverse stem axon was five to nine (7.0
1.0, n ¼ 29). In each bilaterally projecting neuron, the number of ipsilateral and contralateral primary collaterals was three to five (3.8
0.8, n ¼ 25) and two to six (3.7
1.4, n ¼ 25), respectively. In each ipsilaterally projecting neuron, the number of primary collaterals was five to eight (6.0
1.2, n ¼ 4).

Distribution of Terminals of Single LRN Axons in the Cx Each terminal branch of a single LRN axon always terminated as a rosette terminal of terminal type or en passant type in the granular layer. The number of rosettes per axon ranged from 84 to 219 (154.0
37.0, n ¼ 11). In each bilaterally projecting axon, the ratio of axon terminals in the ipsilateral Cx to those in the contralateral Cx ranged from 1.3 to 2.3 (1.7
0.4, n ¼ 7). Axon terminals were mainly seen in lobules III–VI, occasionally lobules II, VII, and VIII in the vermis and paravermal area of the anterior lobe, and sometimes in the hemisphere. Cortical primary collaterals sent terminal branches sometimes to a single lobule but usually to multiple (one to four) lobules and spread rather widely in the semi-parasagittal plane (width, 400–2,000 mm, usually more than 1,000 mm) (lower drawings in Fig. 20.6). In contrast, the spread of the terminal branches of a single primary collateral was relatively restricted in the transverse plane (mediolateral width, usually 300– 650 mm), and the terminal distribution of a single LRN axon roughly showed a pattern of multiple longitudinal bands arranged mediolaterally (upper drawings in Fig. 20.6). Figure 20.7 shows an example of the longitudinal terminal distribution of individual primary collaterals, in which the distances from the midline to individual terminals were plotted on unfolded longitudinal strips of the vermal lobules. In Fig. 20.7a, two adjacent primary collaterals terminate in a longitudinal zone which is labeled zone c in Fig. 20.6. In this zone, terminals of one collateral were distributed in lobule V, and those of the other in lobule VI with two clusters of terminals were slightly separated in lobule VIa and VId. The longitudinal spread of the terminal clusters extended over 6.5 mm (see legend in Fig. 20.7), whereas the mediolateral spread of the clusters was limited between 230 and 400 mm. Even as a whole, these terminals were well aligned in a single longitudinal strip less than 400 mm wide in the

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Fig. 20.7 Longitudinal band distribution of terminals of single primary collaterals of an LRN axon in the unfolded Cx. Primary collaterals in (a) and (b) are the same as shown in Fig. 20.6c and e, respectively. (a) Terminals of two primary collaterals plotted on the unfolded Cx (left). Two

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transverse plane. Similarly, Fig. 20.7b shows the distribution of terminals of another primary collateral of the same axon (labeled zone e in Fig. 20.6) on the unfolded cortical parasagittal strip. This collateral bifurcated into two main branches, one in lobule III and the other in lobules IV and V. Terminals were distributed widely in a parasagittal longitudinal strip (11.6 mm long), but the mediolateral spread was restricted within 600 mm as a whole. As shown in this example, the general feature of the cortical distribution of terminals of a single LRN axon could be summarized as follows. Terminals that belong to one or sometimes two primary collaterals spreading in a few lobules make a longitudinal zone in the parasagittal plane that is usually less than 500 mm wide in a mediolateral direction, and several such longitudinal zones of terminals are arranged almost in parallel mediolaterally.

Morphology of Collaterals Terminating in the Cerebellar and Vestibular Nuclei All reconstructed LRN axons supplied collaterals to the CN. Nuclear collaterals were given off from transversely running stem axons or proximal parts of cortical primary branches and ran in a caudal direction. A single axon had two to three (2.5
0.5, n ¼ 11) primary collaterals to the cerebellar (and/or vestibular) nuclei. Each of these nuclear collaterals was much thinner (diameter, 0.4–1.0 mm) than stem axons (diameter 1.5–2.0 mm) and cortical branches, and had a localized termination area in the CN or CNs. A nuclear collateral ran toward the target nucleus in a relatively straight path without any branching when the branching point was far from the CN. Within the CN, a nuclear collateral bore several en passant swellings and had only one or two ramifications to end in several (usually one to four) terminal branches. Each terminal branch was usually about 200–500 mm long and bore frequent en passant swellings and sometimes short branchlets bearing a terminal swelling. Usually, a single axon innervated only one target nucleus, but sometimes two CNs, or both the cerebellar and vestibular nuclei (most often the FN and IP, and occasionally the DN and IP.) There seemed to be a rough correspondence in the cortical and nuclear projections of single LRN axons, which is compatible with the zonal arrangement in corticonuclear projection (Voogd et al. 1996). For example, when an axon had a collateral in the FN, the same axon always had terminations in the vermis on the same side as the FN. An axon with collaterals in the IP and the DN often innervated the intermediate part including the most lateral vermis and the hemisphere. ä Fig. 20.7 (continued) terminals in lobule VII that were far away from the other terminals in lobule V and VI are not included in the unfolded strip, but they were located in the same parasagittal strip. (b) Terminals borne on a primary collateral plotted on unfolded lobules III–V (right). The rostrocaudal distance regarding distribution of terminals was measured along the surface of the molecular layer on a reconstructed parasagittal section, and the locations of axon terminals in the granular layer were projected to the corresponding sites on the surface of the unfolded Cx. Arabic numbers attached to the portions of the folia correspond to those on the unfolded cortical parasagittal strips. Roman numbers indicate names of lobules. Note the narrow mediolateral and wide longitudinal distributions of terminals belonging to individual primary collaterals (Reproduced from Wu et al. 1999)

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Dorsal Column Nucleus Neurons The connections of the DCNs with the cerebellum have been extensively investigated both anatomically and electrophysiologically. Sagittal organization of MFs from the cuneocerebellar tract was reported by Voogd (1964, 1969) and definitely confirmed by Gerrits et al. (1985) with autoradiography. The axonal trajectories of single dorsal column nucleus (DCN) (gracile (GN), cuneate (CuN) and external cuneate nucleus (ECuN)) neurons were analyzed only recently (Quy et al. 2011). Almost all axons projected only to the ipsilateral cerebellum (Fig. 20.8d), although some axons in the ECuN projected bilaterally (Fig. 20.8a). This finding is consistent with previous reports that the DCN-cerebellar projection is predominantly ipsilateral (Somana and Walberg 1980; Gerrits et al. 1985) with a small but definite projection to the contralateral anterior vermis (Voogd 1964). Stem axons originating from the DCN ran rostrolaterally in the dorsolateral superficial white matter of the medulla without any collaterals, and entered the ipsilateral ICP. While running medially, the stem axons gave rise to several primary collaterals mostly in the ipsilateral (Fig. 20.8d) and occasionally bilateral deep white matter (Fig. 20.8a). These collaterals entered the folial white matter of several lobules which were either adjacent or separate from each other, and further branched before entering the granular layer. Each branch formed a cluster of MF rosettes in a band-shaped small area in a lobule. DCN axons showed trajectories and ramification patterns that were essentially similar to those described above, but the cortical lobules of their termination varied, depending on the locations of their cell bodies in the DCN. In the example in Fig. 20.8a, an ECuN axon projected bilaterally by way of its transcortical transverse stem axon in the deep cerebellar white matter. Several lobular branches were given off perpendicularly from the traversing stem axon, and axon terminals of individual primary collaterals were localized in a mediolaterally narrow-strip zones. This branching pattern is very similar to that of LRN axons (Wu et al. 1999). The number of rosette terminals per axon ranged from 57 to 202 with an average of 123. Nuclear collaterals were relatively unusual in DCNcerebellar axons (1/15 single neurons examined), in contrast to abundant collaterals in the CN of spinocerebellar MF axons (Matsushita and Yaginuma 1995). To examine the relationship between terminal distribution of single DCN axons and the aldolase C expression pattern in the Cx, all terminals of a single DCN axon were mapped on the aldolase C expression map of the same animal (Fig. 20.8b). The GN mainly projected to the copula pyramidis and lobules III–V, the CuN to the paramedian and simple lobules, and the ECuN to lobules I–VI and VIII–IX, although there was some overlap. The majority of terminals of the GN and CuN axons were located within aldolase C-negative or lightly C-positive compartments in different lobules, even though many single DCN axons projected to the anterior and posterior lobes with their axon collaterals. However, terminals of single neurons in the ECuN were located not only in aldolase C-negative but also in aldolase C-positive compartments, since single ECuN neurons often projected to lobule IX, which is mostly aldolase C-positive. One GN neuron that had an axon collateral to the AIP had rosette terminals in aldolase C-negative compartments in the anterior lobe (lobules III–V) and the posterior lobe (lobule VIII) (Fig. 20.8d).

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Fig. 20.8 Morphologies of reconstructed single MF axons of an external cuneate nucleus (ECuN) neuron (a, b) and a gracile nucleus (GN) neuron (d). (a) Frontal view of the axonal trajectory of an ECuN neuron, which projects bilaterally to the Cx, drawn on the montage of rostral and central cerebellar sections and a section of the caudal medulla. (b) Distribution of all the rosette terminals of this axon mapped on the unfolded cortical scheme. (c) Injection site in the ECuN mapped on the horizontal scheme of the dorsal column nucleus. The injection site in the GN applies to the neuron in (d). (d) Lateral view of the axonal trajectory of a single GN neuron on the montage of the cerebellum and the brain stem. Note a nuclear collateral of this neuron in the AIN (arrowhead) (From Quy et al. 2011)

Pontine Nucleus Neurons Inputs from the cerebral cortex to the CN were investigated by stimulating areas 6 and 4, and recording intracellular potentials from CN neurons in the cat (Shinoda et al. 1987). Stimulation of the precruciate cortex (area 6) produced small excitatory postsynaptic potentials (EPSPs) followed by large inhibitory postsynaptic potentials (IPSPs), and these EPSPs were considered to be mediated by the PN and the nucleus reticularis tegmenti pontis (NRTP), since stimulation of the PN and the NRTP produced monosynaptic EPSPs followed by IPSPs in the same DN neurons. To confirm these pathways anatomically, single MF axons originating from the PN were stained with intraaxonal injection of HRP after electrophysiological identification (Fig. 20.9b), and thin tiny axon collaterals of MF axons projecting to the Cx were identified in the DN (Fig. 20.9a),

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Fig. 20.9 Photomicrographs of a nuclear collateral (a) and MF rosettes (b) of an intraaxonally stained pontocerebellar axon in the cat. Scale in B, 50 mm. (c) Axonal trajectory of a reconstructed single pontocerebellar axon superimposed on a montage of several coronal sections of the rat in which many terminals were located. A nuclear collateral was not found in this axon. All MF terminals of this axon are mapped on the scheme of aldolase C compartments of the Cx. BPN basilar pontine nucleus (Reproduced from Shinoda et al. 1992)

(Shinoda et al. 1992). However, nuclear collaterals were found only in about 20% of the PN axons well stained. Since the entire axonal trajectories of single PN axons could not be stained well in the cat, the axonal morphologies of single PN neurons were recently analyzed in the rat (Fig. 20.9c) (Na, Sugihara, and Shinoda unpublished). Typically, single pontocerebellar axons crossed the midline in the pons, and after entering the cerebellum through the contralateral middle cerebellar peduncle, ran transversely in the deep cerebellar white matter toward and often across the midline. From the transversely running stem axons, multiple primary collaterals were successively given off almost perpendicularly in the Cx only contralaterally or bilaterally with a contralateral predominance. Each primary collateral further branched in a parasagittal plane to form a strip-shaped termination area with rosette-type swellings in the granular layer. Terminals were distributed mainly in the apex of the target lobules. Rosette terminals of PN axons were clustered almost exclusively in aldolase C-positive compartments in a single or multiple lobules. This projection pattern of single pontocerebellar axons with axon collaterals to multiple longitudinal compartments is very similar to that of single LRN and DCN neurons and may represent a general feature of the cortical

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projection of single axons in the MF systems. Axons originating from the central, rostral, and lateral part of the PN projected mainly to specific lobules with multiple branches to the simple lobule, crus II and paramedian lobule, to crus I and dorsal paraflocculus, and to the ventral paraflocculus and lobule IXc, respectively. These findings indicated that the projection of single pontocerebellar axons was closely related both to lobular and longitudinal organization of the cerebellum.

General Features of Axonal Projection Patterns of Single CF and MF Neurons One of the common features of the branching patterns of single MF axons is that multiple longitudinal zones innervated by single MF axons are arranged mediolaterally in one to a few lobules of the Cx. In general, stem axons of MFs give rise to several primary collaterals successively to the Cx, while running medially toward the midline in the deep white matter rostral and dorsal to the CN (Fig. 20.10a). These primary collaterals arise perpendicularly from the stem axons and widely ramify mainly in the parasagittal plane of the folium, so that each MF collateral terminates in a relatively narrow longitudinal zone covering one to a few lobules. It is generally assumed that transverse, lobular subdivisions of the Cx are mainly based on the distribution of the MF input, whereas longitudinal divisions represent the output systems of the Cx in which both the PC zones and their target CNs are innervated by the common CF axons of adjacent neurons in a subnucleus of the IO (Fig. 20.10b). However, the main discontinuities of terminals of single MF axons are mediolateral rather than lobular, and concern the clustering of MF terminals in sagittal strips. This clustering of terminals of a single MF axon into sagittal strips may be a common feature of MF projections in general. These branching patterns of single MF axons may account for the “patchy-mosaic” or “fractured somatotopy” of granule cell cutaneous receptive fields in which skin surfaces are represented discontinuously in adjacent granule cells (multiple representations of the same receptive fields) (Welker 1987). Even though the distribution of cutaneous receptive fields looks patchy, the basic organization of the projection of single MF axons in the Cx should be regarded as multiple sagittal strips along longitudinal compartments of aldolase C expression and OC projection. The relationship between MF multiple zones and aldolase C-defined longitudinal zones was examined in several MF systems. MF projection zones of the LRN were aldolase C-negative in the paramedian lobule (Ruigrok and Cella 1995). Pontocerebellar MF terminals are located in aldolase C-positive bands, whereas most MF terminals originating from the GN and CuN are located in aldolase C-negative bands. Thus, the functional significance of cerebellar compartmentation defined from the viewpoint of aldolase C expression seems to be well correlated with the zones defined from the viewpoint of olivocerebellar, corticonuclear, and MF projections. However, there are some exceptions to this relationship. In lobules VIII and IX, spino- and trigeminocerebellar MFs terminate in patches, which are topographically related to the zebrin-negative zones, but the borders of some of the

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a

b

Aldolase C (+)

Aldolase C (-)

Midline CN Aldolase C (-)

Aldolase C (+) D midline

CN

M R

MFNucl IO

Aldolase C (-)

c

Aldolase C (-)

CN

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Aldolase C (+)

MFNucl IO

Fig. 20.10 Schematic summary diagrams showing characteristic features of the axonal projection patterns of a single MF neuron (a) and a single CF neuron (b) in the Cx and CN, and the relationship between MF and CF longitudinal zones innervated by single MF and CF neurons, respectively (c). (a) A single MF neuron terminates in multiple small longitudinal strip-like zones that are arranged successively in a mediolateral direction, often bilaterally. Nuclear collateral may or may not exist in different MF systems. Individual small longitudinal zones may correspond to “microcomplexes.” MF Nucl, nucleus from which a MF arises. (b) A single CF terminates in a longitudinal zone across one or more lobules, even the entire rostrocaudal length of the cerebellum. PCs within the zone project to the corresponding target nucleus that the same CF neuron innervates, and both the PCs and the target nucleus have the same aldolase C expression pattern (either aldolase C-positive or C-negative). (c) The relationship between longitudinal bandlike zones of MF terminals of a single neuron and longitudinal bands of the OC projection and PCs within the aldolase C-negative bands

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efferent modules or CF zones are located in the middle of a zebrin-positive or zebrin-negative zone, and some modules include both zebrin-positive and negative PCs (Gravel and Hawkws 1990). Some MF axons of the ECuN send axon collaterals to aldolase C-negative and aldolalse C-positive bands (Quy et al. 2011). Therefore, the relationship between terminal zones of single MF axons and bands defined by aldolase C expression will require further investigation for each MF system. One of the important questions for understanding cerebellar function based on morphology is how multiple longitudinal zones of terminals of an MF neuron are functionally related to the longitudinal zones identified with OC and corticonuclear projections and the aldolase C expression. Concerning the nuclear projection, it is obvious that the cortical and nuclear projections of a single MF axon does not exactly follow the so-called corticonuclear topographical relationship, since the cortical projection of a single MF neuron is generally more widely spread in the transverse direction than its nuclear projection, and the MF nuclear projection does not exist in all MF systems. Therefore, this situation does not exactly fit the classical microcomplex scheme of Ito (1984), in which collaterals of MF axons and PCs that receive input from the same MF axons converge onto common target nuclear output neurons. However, the concept of the microcomplex as a functional unit is still applicable by classifying microcomplexes into microcomplexes with and without nuclear MF collateral input. In this basic structure of the microcomplexes, a single MF neuron may innervate multiple microcomplexes in a single longitudinal compartment of the OC and olivonuclear projection, and multiple microcomplexes which are arranged in the transverse plane and belong to separate longitudinal compartments with the same aldolase C expression (Fig. 20.10c). Cerebellar granular cells have three to five dendrites, each of which receives one excitatory MF synapse (Palay and Chan-Palay 1974). Parallel fibers are 4–7 mm long end to end (Brand et al. 1976; Pichitpornchai et al. 1994). Therefore, the transverse arrangement of the MFs is further enhanced by the transverse orientation of the parallel fibers that link the MF input via granular cells with PCs, suggesting that the MF granule cell–parallel fiber system may distribute information to many PCs along a folial axis. However, granular cells within a microzone are mainly reached by MFs with the same receptive fields that have little convergence from MFs with different types of input, and their activation patterns primarily reflect the synchronized activity in presynaptic MFs driven by similar input (Jorntell and Ekerot 2006). MFs of the cuneocerebellar tract with specific receptive fields are distributed approximately along the longitudinal microzonal organization determined by CFs with specific cutaneous receptive fields, and the cutaneous parallel fiber receptive fields in PCs and interneurons are similar to those of MFs (Garwicz et al. 1998). Ekerot and Larson (1980) demonstrated a close correlation between the sagittal projection zones in the hemisphere of the anterior lobe activated by the MF of the cuneocerebellar path and the CF of the dorsal funiculus spino-olivocerebellar paths. This finding clearly provides electrophysiological evidence correlating the MF sagittal strips with the CF longitudinal band in a single longitudinal compartment defined by aldolase C expression. Since aldolase C-positive and C-negative

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bands alternate mediolaterally within lobules, the question remains as to how parallel fibers of granule cells innervated by single MF axons influence PCs in these two longitudinal bands. Whether a single MF axon innervates only adjacent or separate lobules depends on the MF axon and the MF system to which it belongs. Some cutaneous tactile MF projections are highly localized, while the LRN projection that conveys flexor reflex afferent inputs is rather widespread in a rostrocaudal direction. These physiological findings are consistent with the projection patterns of single MF axons in these MF systems. Single MF axons of the LRN tend to innervate adjacent or separate lobules, whereas those of the DCN tend to innervate only one lobule. This difference in the rostrocaudal extent of the sagittal bands of a single MF axon is probably related to the functional aspect of the single MF axon that reflects the somatotopy and the degree of independence or coordination of movements of limbs and axial muscles. The same may hold true for single CFs, since cortical segments that are separate in a rostrocaudal direction but within a single longitudinal band are often innervated by axon collaterals of a single IO neuron (Fig. 20.2) (Ekerot and Larson 1982). To understand the functional interaction between widely distributed PCs innervated by a group of adjacent single MF neurons and those innervated by single OC axons, more detailed information is needed as to how terminals of single PCs in separated cortical longitudinal zones are organized in the CN, and how the two afferent inputs are integrated as a final output at the PCs. Furthermore, for understanding the basic functional mechanism of MF systems in general, it will be necessary to specify the similarity and the difference of axonal projection patterns of single MFs in other MF systems such as the spinocerebellar and vestibular systems.

Functional Considerations From the studies on single axonal morphologies, it is clear that some MF systems emit axon collaterals to the CN, but others do not, or only a small portion of neurons in an MF system emit nuclear collaterals. In contrast, almost all of OC axons emit nuclear collaterals to the corresponding target CN. Eccles et al. (1974) showed that early excitation in the FN was not evoked from the dorsal spinocerebellar tract by hind limb inputs, but was evoked by spinocerebellar inputs via the LRN and the IO. The present anatomical findings are consistent with these electrophysiological findings. Therefore, it is important to identify the presence or absence of excitatory input to the CN by a nuclear collateral in each MF system, although it is still tacitly assumed that all afferent neurons in the MF systems have nuclear axon collaterals. During slow tracking movements of wrist flexion and extension, both PCs (Mano and Yamamoto 1980) and nuclear neurons in the DN and IP (Schieber and Thach 1985) were found to show a bidirectional increase of spike activity rather than a reciprocal pattern. In rapid alternating movements of a forelimb, CN cells showed reciprocal changes of activity above and below a resting mean frequency (Thach 1968; Wetts et al. 1985), whereas PCs had a bidirectional discharge pattern

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(predominantly bidirectional increase) (Mano and Yamamoto 1980). As exemplified in these cases, CN cells either increase or decrease their activity at the onset of and during movement. The neural mechanism of this increase or decrease of discharge frequency in cerebellar nuclear cells is an unsolved important question of cerebellar physiology (Shinoda et al. 1987, 1992, 1993, 1997). There are two possible pathways to produce this increase in spike firing in nuclear neurons (Fig. 20.11). If there is an excitatory input via axon collaterals of MF neurons to the CN (Fig. 20.11A), this excitatory input may cause an initial increase of nuclear cell discharge. In this case, PCs may either increase or decrease their firing, although most PCs are known to increase their activity at the onset of movement (Thach 1968; Mano and Yamamoto 1980). Only a few intracellular recording studies were performed in in vivo preparations to examine synaptic inputs to nuclear neurons from the precerebellar nuclei (Ito et al. 1970; Kitai et al. 1977; Shinoda et al. 1987). EPSPs were evoked by stimulation of the IO, the PN, and the NRTP, (Ito et al. 1970; Shinoda et al. 1987). Accordingly, excitatory inputs of extracerebellar origin may exist to increase nuclear cell discharge at the onset of or during movement. However, contrary to the common assumption that precerebellar neurons that project to the Cx emit axon collaterals to the corresponding CN, some MF neurons have no axon collateral to the CN (Fig. 20.11B). In this case, changes in nuclear cell firing rates will be caused solely by changes in PC firing rates; an increase in nuclear cell discharge will be caused by disinhibition through PCs that decrease their firing, since PCs are inhibitory (Fig. 20.11Ba). Physiologists usually searched for PCs that show an increase of activity in response to sensory stimuli or movements, but they have not paid much attention to the possible existence of PCs that show a decrease in activity. PCs have high spontaneous or background activity (60–80/s) (Thach 1968; Mano and Yamamoto 1980), and CN neurons have also spontaneous activity (30–40/s). Spontaneous simple spikes occur in isolated PCs in the absence of any parallel fiber input (Llina´s and Sugimori 1980; Raman and Bean 1999). The relatively high frequencies of simple spikes may establish optimal carrier frequencies necessary for effective modulation of PCs, and enable bidirectional responses in PCs and their target CN and VN, namely, an increase in firing due to either excitation or disinhibition and a decrease in firing due to either inhibition or disfacilitation. Interneurons in the molecular layer, basket and stellate cells, exert inhibitory control on PCs (Andersen et al. 1964; Eccles et al. 1967; Midtgaard 1992). Golgi cells in the granule cell layer indirectly influence simple spike discharge by inhibiting the discharge of granule cells (Eccles et al. 1967). Therefore, even excitatory inputs of MFs may suppress activity of PCs via these inhibitory interneurons. On the other hand, a decrease of activity in nuclear neurons was observed in alternating arm movements (Thach 1968; Wetts et al. 1985) and also in horizontal saccadic eye movements (Ohtsuka and Noda 1991) at the onset of and during movement. This decrease in activity in nuclear cells may be caused by an increase in activity of PCs, even though nuclear axon collaterals of MFs are present (Fig. 20.11A) or absent (Fig. 20.11Bb). It has been suggested that the background discharge of CN neurons is due to continuous excitatory input, or the membrane properties of CN neurons (Thach 1968). A study in a slice preparation

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Fig. 20.11 Two possible operational modes of cerebellar corticonuclear interaction in the CN in relation to MF input from the cerebral cortex. (A) A pathway with an MF collateral terminating on the CN. Nuclear output neurons increase or decrease their activity at the onset of and during movement, depending on the strength of PC inhibition. (B) a pathway without an axon collateral projecting to the CN. Nuclear output neurons increase their activity (Ba) or decrease their activity (Bb) at the onset of and during movement. Mx motor cortex, Ass Cx association cortex, IN inhibitory interneuron (basket and stellate cell), Gr cell granular cell, PT pyramidal tract (Modified from Shinoda et al. 1993)

supports the idea of the latter mechanism because the electroresponsive membrane properties of CN neurons contribute to the background discharge (Jahnsen 1986). Furthermore, Llina´s and M€ uhlethaler (1988) demonstrated the existence of a powerful low-threshold Ca2+-dependent spike which triggered a burst of fast

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Na+-dependent spikes in CN neurons. Therefore, changes of spike activity in CN neurons can be easily caused by slight modulation of the membrane potential by excitatory or inhibitory inputs superimposed on the underlying spontaneous activity. It is still unknown in the case of voluntary limb and eye movements whether nuclear output is generated only by PC input or by the interaction of MF or CF input and PC input. From the perspective of the function of target cells outside the cerebellum, information is not yet available as to which is more important – an increase or a decrease in firing rates of CN neurons. Further analysis is required of the axonal projection patterns of single MF neurons with different origins, such as the NRTP and spinocerebellar neurons, to understand the neural mechanisms of interaction of MF inputs in the Cx and CN for proper generation of cerebellar outputs to targets outside the cerebellum for control of movement and other functions. Acknowledgments This research was supported by a research grant from CREST (Core Research for Evolutional Science and Technology) of the Japan Science and Technology Agency to Y.S. and by Grants-in-Aid for Scientific Research from the Ministry of Education, Science and Culture of Japan to I.S.

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Visual Circuits from Cerebral Cortex to Cerebellum; The Link Through Pons

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Mitchell Glickstein

Abstract

This chapter reviews some anatomical, behavioral, and physiological studies that bear on the role of the sensory input to the cerebellum via the pons in general and the visual input in particular. The core of the argument is that in monkeys, the visual input comes from those extrastriate visual areas that are involved in motion detection and visual control of movement. Studies of other inputs and other species are cited when they are relevant to the more general issue of the nature of sensory input to the cerebellum. The pons occupies a prominent place at the base of the brain, extending from the caudal end of the midbrain to the rostral border of the medulla. Costanzo Varolio (aka Varolius) (1591) dissected the human brain from its ventral surface, thus giving him a clear view of the origin of the cranial nerves and the shape of the lower brainstem. It was Varolius who gave the pons its name, based on its resemblance to a bridge over a canal. Importantly subdivided into a dorsal and a ventral division, it is the ventral division that contains the pontine nuclei. One of the largest circuits through the human brain originates in the cerebral cortex and projects to the cerebellum by way of the pontine nuclei. The input to the pons arises from both sensory and motor areas of the cerebral cortex. Pontine inputs also originate from the superior colliculus and other brainstem structures, but it is the cortical input to the pons that is by far the largest. Anatomical, recording, and lesion-based studies help to understand the functions of these pathways. In this chapter, I will emphasize the visual input in relation to some of the more general principles about the nature of the cortico-ponto-cerebellar circuit. In addition to a purely visual input, visual information is also a constituent element of visuomotor

M. Glickstein Cell and Developmental Biology, University College London, Wolfson House, 4 Stephenson Way, London, NW1 2HE, UK e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 469 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_21, # Springer Science+Business Media Dordrecht 2013

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Fig. 21.1 Cross section through the frontal lobe of a tree shrew (Tupaia glis) showing the location of retrogradely labeled cells after HRP injection in the pons. The gray line in lamina V of cortex is a continuous band of labeled pyramidal cells

signals. My discussion is based mostly on vision and the work of my own lab. A much broader view is well summarized in a recent review by Thier and M€ ock (2006).

Which Cells in the Cerebral Cortex Have an Axon that Projects to the Pons? There are two related questions about the nature of the link between cerebral cortex and the pons. Which cells are the origin of the corticopontine fibers? Which areas of cortex provide an input to the pons? The first question is easily answered. The input to the pontine nuclei comes only from Lamina V pyramidal cells. Injection of horseradish peroxidase (HRP) into the pontine nuclei allows identification of the cells of origin of the axons that project to the pons. Often in such preparations, there is a single line within lamina V composed of corticopontine cells. In all cases that have been studied, it is only lamina V pyramidal cells that are labeled. Figure 21.1 shows a cross section through the frontal cortex of a tree shrew in which HRP had been injected into the pontine nuclei several days prior to the brain being prepared to identify labeled cortical cells. In some cases, cortical Lamina V may be divisible into a superficial layer Va and a deeper Vb. In the rat, all corticopontine fibers originate in Vb. Cells in lamina Va are labeled after an injection confined to the basal ganglia (Mercier et al. 1990).

Which Areas of Cortex Project to the Pons? Although all corticopontine cells are located in lamina V, the distribution and extent of such cells across the cortex varies greatly among mammals. In rats, all of the cerebral cortex projects to the pons (Legg et al. 1989). In monkeys, some cortical areas are without a pontine projection, and for some areas the projection is sparse (Glickstein et al. 1985). The great majority of corticopontine connections arise from

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a contiguous area of cortex that extends from the superior temporal sulcus caudally to the arcuate sulcus rostrally and from the corpus callosum medially to the superior temporal sulcus laterally (Fig. 21.2). Differences in the source of the projection from cortical visual areas help to interpret the functions of the cortico-ponto-cerebellar pathway. The extrastriate visual areas can be subdivided into two major groups: a dorsal-medial group that extends into in the parietal lobe, and a ventral-lateral group that extends into in the temporal lobe (Ungerleider and Mishkin 1982; Glickstein and May 1982). The dorsal-medial visual areas project heavily to the pons, the lateral-ventral areas do not. Differences between the two groups in the extent of their corticopontine projections reflect their differences in function. Lesions of the dorsal areas in monkeys produce a profound deficit in visual guidance of the wrist and fingers without impairing the animal’s ability to learn a visual discrimination task. Ventral lesions are without effect on visual guidance of movement, but severely impair visual discrimination learning (Glickstein et al. 1997).

What Is the Pathway of the Cortical Projection to the Pons? Corticopontine fibers descend in the internal capsule and collect ventrally at the base of the midbrain in the cerebral peduncles. In the human and monkey, corticopontine fibers constitute the great majority of the fibers in the cerebral peduncles. The cerebral peduncles also contain the axons of the pyramidal tract. One way to appreciate the relative importance of the cortico-cerebellar link is to compare the ratio of cross-sectional area of the peduncle to that of the pyramidal tract. Figure 21.3 illustrates the point. The section on the left is through the midbrain showing the fibers in the cerebral peduncle of a fiber-stained human brain. The section on the right is through the pyramidal tract in the same brain. Interruption of the pathway from cortex to the pons at the level of the caudal limb of the internal capsule in a stroke patient produced a profound impairment in skilled visuomotor performance by the contralateral hand, and a moderate impairment on the ipsilateral hand (Classen et al. 1995). The interpretation of the loss in this case was that the fiber lesion had interrupted an input to the pons from extrastriate cortical visual areas.

Where Do Corticopontine and Collicular Fibers Terminate in the Pons? In monkeys, the major targets of extrastriate visual areas are the dorsolateral and dorsal regions of the pontine nuclei (Glickstein et al. 1980). The location of functional borders within the pontine nuclei is relatively arbitrary. There are hints that the true organization is probably based on a central-peripheral axis within the pontine nuclei. In an elegant study of pontine termination from the cerebral cortex

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of rats, Leergaard et al. (2000) showed that the pattern of termination is based on a circular arrangement around the pyramidal tracts as they course through the pontine nuclei. The same principle may be present for terminations of fibers originating in cortical visual areas.

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Fig. 21.3 Fiber-stained human brain. Compare the volume of fibers in the cerebral peduncles with that of the pyramidal tract

There are other, noncortical sources of visual input to the pontine nuclei. In all animals studied, there is an input from both the deep “motor” laminae of the superior colliculus as well as from lamina III, the stratum opticum (Mower et al. 1979; Schwarz et al. 2005). In monkeys, the cortical and collicular projections to the pons terminate in adjacent, partially overlapping regions of the dorsolateral and dorsal regions of the pontine nuclei. In cats, the projections are to different areas. Fibers that project to the pons from the superior colliculus terminates in the dorsolateral pons, whereas fibers from the cerebral cortex terminate in ventromedial pons. The fact that the two inputs do not overlap allows for a comparison of their cerebellar targets and offers some insights into the difference in the functions of the two pathways. Cortically activated visual cells have as their major target the dorsal paraflocculus of the cerebellar hemispheres and adjacent uvula. The axons of cells that receive an input from the superior colliculus terminate principally in lobule VII, the oculomotor vermis.

Demagnification The receptive field properties of visual cells in the cat pons resemble those of the cortical cells that provide their input (Gibson et al. 1978). Both corticopontine and pontine receptive fields are powerfully influenced by the speed and direction of visual targets. They are relatively insensitive to differences in target shape. In both cases, the major difference between cortical and pontine receptive fields is in their size. Corticopontine cells in cat Area 18 have receptive fields that are up to a degree or two in cross section, while pontine receptive fields may encompass an entire visual hemifield. In the cerebral cortex of both monkeys and cats, the central portion of the visual field is represented in a much greater area than the periphery. The large size of some pontine receptive fields poses a challenge. Consider an object traveling across the

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cortical representation of the visual field. Because of the great difference in magnification from center to periphery, it would travel slowly across the cortical representation of the central visual field, and then more rapidly in the periphery. If cortex were represented uniformly in its projection to the pons, then optimal target speed would be different in the center and periphery of a large receptive field. The corticopontine pathway corrects by de-magnifying the visual input in cats (Cohen et al. 1981). In monkeys, there is a very small corticopontine projection from the peripheral field representation in Area 17. There is an orderly increase in the number of corticopontine cells as you go from the representation of the center of gaze to the periphery.

Fibers from the Cerebral Cortex and Colliculus Give off Collaterals In his Textura del Systema Nervioso (1899), Cajal illustrates a Golgi-stained section through the pontine nuclei. Each of the fibers gives off a collateral to the pons. Ugolini and Kuypers injected fast blue, a retrograde tracer into the pyramidal tract at the level of the medulla. In addition to the expected retrograde filling of pyramidal tract fibers, she described extensive collaterals of pyramidal tract fibers as they traversed through the pontine nuclei (Ugolini and Kuypers 1986). The same principle of collaterals may be present for corticopontine fibers. Corticopontine visual fibers in the cat give off collaterals to the superior colliculus (Baker et al. 1983). Many cortical visual cells that can be activated antidromically from the pontine nuclei are also activated antidromically by electrical stimulation in the superior colliculus. By analyzing the pattern of collision between responses activated from the two sites, it is possible to calculate the point at which the bifurcation occurs. One branch of these axons may play a direct role in motor control, the other branch in the corollary discharge.

What Are the Receptive Field Properties of Pontine Visual Cells? In cats, pontine visual cells appear to be activated solely by the magnocellular visual system. They are strongly influenced by the direction and velocity of moving targets, but relatively unaffected by their shape. These properties resemble closely those of the cortical cells which activate them (Gibson et al. 1978). Similarly, in monkeys, the extrastriate cortical visual areas that project to the pons are dominated by their magnocellular inputs.

Where and How Do the Axons of Pontine Cells Terminate on the Cerebellum? The distribution of pontocerebellar fibers can help to interpret the function of the corticopontine system. Anna Rosina and her colleagues (1980) studied

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the pattern of termination of pontine cells on the cerebellar cortex. While the majority of the pontocerebellar fibers terminate in the contralateral cerebellum, there is a definite ipsilateral projection. Following Bolk (Glickstein and Voogd 1995), we might speculate that the ipsilateral fibers are particularly involved in the coordination of movements that require cooperation between the two sides of the body, such as rotation of the neck or conjugate movement of the eyes.

Some Further Speculations on the Role of the Cortico-PontoCerebellar System In cats, there are inputs to the pontine nuclei from two distinct cortical visual areas: Area 18 (Visual II) and the lateral suprasylvian area. The pontine targets of the two cortical areas do not overlap extensively. Fibers originating in Area 18 terminate centrally within the pontine nuclei, close to the pyramidal tract fibers as they course through the pons. Fibers from lateral suprasylvian cortex terminate more peripherally within the pontine nuclei. This pattern of input is related to the distribution of axons from these two areas on the cerebellar cortex (Mower et al. 1979; Robinson et al. 1984). The axons of cells that receive an input from Area 18 terminate medially on the vermis-paravermal area of the cerebellar cortex. Pontine cells that relay the input from the lateral suprasylvian area terminate more laterally in the cerebellar hemispheres. Both classes of visual input are probably involved in the visual guidance of movement. The input from Area 18 may be involved in regulating whole body movements; the lateral suprasylvian area in independent use of the limbs.

Behavioral Evidence of the Function of the Corticopontine Link The corticopontine pathway is involved in the sensory control of movement. One example of that principle comes from a study of rats jumping. Rats can be readily trained to jump across a gap to reach a bit of food at the end of a runway. In the dark, rats use their whiskers to gauge the distance which must be crossed (Hutson and Masterton 1986). The somatosensory areas of the rat cerebral cortex, including the whisker barrel field sends a powerful input to the pontine nuclei. If this pathway is cut, rats will refuse to jump the gap in the dark, although jumping in the light is unaffected (Jenkinson and Glickstein 2000).

Collateral Fibers and the Corollary Discharge When a movement is executed, there is a corollary discharge which signals that movement. As Helmholtz pointed out, such a system would serve to maintain the perceptual stability of the world around us despite movement of the eyes or head.

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The pattern of branching by corticopontine fibers may well contribute to that function. Collateral branching of the fibers as they traversed the pons would provide an anatomical basis for the corollary discharge.

Receptive Fields and Visual Guidance; the Appearance of the Ground to a Walking Cat Occasionally a receptive field will seem uniquely appropriate for guiding a specific movement. Some pontine receptive fields are maximally activated by a large, textured target moving downward and outward in the visual field. A target of this sort resembles the appearance of the ground to a walking or running cat. Following David Lee (1980), the cell could be part of a circuit that calculates time to contact with an object in the visual field of a moving cat.

Cerebellum and Kinesie Paradoxale Kinesie paradoxale refers to the fact that patients suffering from Parkinson’s disease are often capable of skilled and coordinated movement. They may catch a ball, walk, or run if aided by an appropriate visual input. The corticopontine system could provide an appropriate visual input allowing for cerebellar control of those movements. The properties of pontine visual cells are similar to the visual inputs that promote such movements. The suggestion is that an intact cerebellar pathway may serve to control movement in a patient with nonfunctional basal ganglia (Glickstein and Stein 1991).

Cerebellar Localization and Current Problems One basic problem which is still poorly addressed is that of localization in the cerebellum. Except for lateralization, Luciani (1891) denied any localization in the cerebellar cortex. This chapter has presented some of the evidence for the role of one or another part of the cerebellum in sensory guided movement. There is good evidence, for example, for a the role of Vermis Lobule VII and caudal VI in saccadic adaptation (Barash et al. 1999) and Pavlovian conditioning (Thompson 1983; Yeo et al. 1984) showed that the critical area is hemispheric Lobule VI. The control of these functions is localized in definite and restricted regions of the cerebellum. The challenge is to understand the functions of the great mass of the cerebellar cortex, especially that of the hemispheres. One current view is that they are involved in some forms of cognition. But anatomical evidence leads me to question such a role. The visual input is from motion sensitive areas of the prestriate and adjacent cortex. There is virtually no input to the pons from the lateral-ventral extrastriate visual areas. There is a modest projection from prefrontal cortex, but the areas that project to the pons are mostly those related to eye movements. The

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situation of the cerebellar hemispheres is reminiscent of that of the corpus callosum in 1936 when Walter Dandy (1936) cut the callosum in a surgical patient without causing any obvious deficits. He wrote “This simple experiment at once disposes of the extravagant claims to function of the corpus callosum.” It didn’t.

References Baker J, Gibson A, Mower G et al (1983) Cat visual corticopontine cells project to the superior colliculus. Brain Res 265:222–232 Barash S, Melikyan A et al (1999) Saccadic dysmetria and adaptation after lesions of the cerebellar cortex. J Neurosci 19:1031–1039 Cajal SR (1899) Textura del Sistema Nervioso. Moya, Madrid Classen J, Kunesch F et al (1995) Subcortical origin of visuo-motor apraxia. Brain 118:1365–1374 Cohen J, Robinson F et al (1981) Cortico-pontine projections of the lateral suprasylvian cortex: De-emphasis of the central visual field. Brain Res 219:239–248 Dandy W (1936) Operative experience in cases of pineal tumor. Arch Surg 33:19–46 Gibson A, Baker J, Mower G, Glickstein M (1978) Corticopontine visual cells in area 18 of the cat. J Neurophysiol 41:484–495 Glickstein M, May J (1982) Visual control of movement: the visual input to the pons and cerebellum. In: Neff WD (ed) Contributions to sensory physiology. Academic, New York Glickstein M, Stein J (1991) Paradoxical movement in parkinson’s disease. Trends Neurosci 11:480–482 Glickstein M, Voogd J (1995) Lodewijk bolk and the comparative anatomy of the cerebellum. Trends Neurosci 18:206–210 Glickstein M, Cohen JL et al (1980) Corticopontine visual projections in macaque monkeys. J Comp Neurol 190:209–229 Glickstein M, May J, Mercier B (1985) Corticopontine projection in the macaque: the distribution of labelled cortical cells after large injections of horseradish peroxidase in the pontine nuclei. J Comp Neurol 235:343–359 Glickstein M, May J, Buchbinder S (1997) Visual control of the arm, the wrist, and the fingers; pathways through the brain. Neuropsychologia 36:981–1001 Hutson K, Masterton R (1986) The sensory contribution of a single vibrissa’s cortical barrel. Neurophysiol 56:1196–1223 Jenkinson E, Glickstein M (2000) Whiskers, barrels, and cortical efferent pathways in gapcrossing by rats. J Neurophysiol 84:1781–1789 Lee D (1980) The optic flow-field: the foundation of vision. Philos Trans R Soc 290:169–179 Leergaard TB et al (2000) Rat somatosensory cerebropontocerebellar pathways: spatial relationships of the somatotopic map of the primary somatosensory cortex are preserved in a threedimensional clustered pontine map. J Comp Neurol 422:246–266 Legg C, Mercier B, Glickstein M (1989) Corticopontine projection in the rat: the distribution of labelled cortical cells after large injections of horseradish peroxidase in the pontine nuclei. J Comp Neurol 286:427–441 Luciani L (1891) Il Cerveletto. Le Monnier, Florence Mercier B, Legg C, Glickstein M (1990) Basal ganglia and cerebellum receive different somatosensory information in the rat. Proc Natl Acad Sci USA 87:4388–4392 Mower G, Gibson A, Glickstein M (1979) Tectopontine pathway in the cat: laminar distribution of cells of origin and visual properties of target cells in dorsolateral pontine nucleus. J Neurophysiol 42:1–15 Robinson F, Cohen J et al (1984) Cerebellar targets of visual pontine cells in the cat. J Comp Neurol 223:471–482

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Rosina A, Provini L et al (1980) Ponto-neocerebellar axonal branching as revealed by double fluorescent retrograde labelling technique. Brain Res 195:461–466 Schwarz C, Horowski A, M€ ock M, Thier P (2005) Organization of tectopontine terminals within the pontine nuclei of the rat and their spatial relationship to terminals from the visual and somatosensory cortex. J Comp Neurol 484:283–298 Thier P, M€ock M (2006) The oculomotor role of the pontine nuclei and the nucleus reticularis tegmenti pontis. Prog Brain Res 151:293–320 Thompson R (1983) Neuronal substrates of simple associative learning; classical conditioning. Trends Neurosci 6:270–275 Ugolini G, Kuypers HG (1986) Collaterals of corticospinal and pyramidal fibres to the pontine grey demonstrated by a new application of the fluorescent fibre labelling technique. Brain Res 365:211–227 Ungerleider L, Mishkin M (1982) Two cortical visual systems. In: Ingle DJ et al (eds) Analysis of visual behavior. MIT Press, Cambridge Varolio C (1591) De nervis opticis. Wechel and Fischer, Frankfurt Yeo C, Hardiman M, Glickstein M (1984) Discrete lesions of the cerebellar cortex abolish the classically conditioned nictitating membrane response of the rabbit. Behav Brain Res. 60:99–113

Cerebellar Connections with Limbic Circuits: Anatomy and Functional Implications

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Gene J. Blatt, Adrian L. Oblak, and Jeremy D. Schmahmann

Abstract

There is an emerging body of evidence suggesting that the cerebellum participates in limbic-related functions including emotion and affect. The underlying connectivity of the cerebellar cortex and nuclei with limbic-related brain areas and associative and paralimbic cortices suggests widespread cerebellar influence on behaviors including the experience and expression of emotion, sadness and grief, integrative hypothalamic visceral/sensory functions, pain perception, modulation, and intensity due to noxious stimuli, as well as other nonmotor behaviors. The key anatomical relationships are the fastigial nucleus projections to the ventral tegmental area (VTA), cerebellar interconnections with the septum, hippocampus and amygdala, direct cerebellar connections with hypothalamic circuits that integrate somatic-, visceral-, and limbic-related activity, and indirect connections with the nucleus accumbens (NAcc), a mesolimbic dopaminergic structure that predicts activity in a reward paradigm in limbic-related structures. Additionally, the cerebellum is interconnected with cingulate cortices that play a role in motivation and emotional drive, and with associative and paralimbic regions of prefrontal, posterior parietal, superior temporal polymodal, and parahippocampal regions heavily implicated in high order

G.J. Blatt (*) • A.L. Oblak Anatomy and Neurobiology, Boston University School of Medicine, 700 Albany Street, Boston, MA, 02118, USA e-mail: [email protected], [email protected] J.D. Schmahmann Massachusetts General Hospital, Harvard Medical School, 175 Cambridge Street, Suite 340, Boston, MA, 02114, USA e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 479 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_22, # Springer Science+Business Media Dordrecht 2013

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processing important for the integration of cognition and emotion. These connections between cortical and subcortical areas of the limbic system with the cerebellum (vermis and fastigial nucleus in particular) are the likely anatomical underpinning of the demonstrated cerebellar influence on limbic-related behaviors in the clinical setting and in earlier behavioral and physiological studies. These cerebellar connections with cerebral limbic areas are also implicated in neurodevelopmental disorders such as autism which demonstrate neuropathology and aberrant neurochemistry in the cerebellar cortex and nuclei. Defining the vermis and fastigial nuclei as the probable location of the limbic cerebellum has relevance for future studies of cerebrocerebellar interconnections and functional coupling, and for therapeutic strategies that attempt to enhance cerebellar modulation of limbic-related structures in order to treat neuropsychiatric disorders.

Introduction Converging lines of evidence provide support for the notion that the cerebellum is essential to the distributed neural circuits involved in cognition and emotion as well as in motor control (Schmahmann 1997, 2010). Unlike the cerebral cortex which can be parcellated on the basis of its architectonic heterogeneity (Brodmann 1909), the main components of the cerebellar cortex are essentially uniform throughout (Eccles et al. 1967; Ito 1984), with the exception of selected neuronal elements such as unipolar brush cells in the vestibulocerebellum (Mugnaini and Floris 1994) and molecular identities of cerebellar cortical neurons (Schilling et al. 2008). In contrast to the cerebellar cortex which is repeating and paracrystalline in structure, the anatomical connections of the cerebellum with extracerebellar areas are remarkably heterogeneous. Defining these connections of the cerebellum with brain areas involved in higher function is therefore integral to the evolving understanding of cerebellar function. There is at present a fair understanding of the regional cerebellar anatomy devoted to motor behavior, and data now also exist suggesting that unique cerebellar modules contribute to the different processes that underlie cognition and emotion. Further, behaviors that fall within the domains of neuropsychiatry, i.e., personality, emotion, and affect, appear to be subserved by the limbic cerebellum (notably the cerebellar vermis and fastigial nucleus – Heath 1977; Schmahmann 1991, 2000, 2010; see review in Herna´ez-Gon˜i et al. 2010). Intellect and emotion are tightly interrelated however (LeDoux 1996; Barbas 2007), and so the areas of cerebellum involved in neuropsychiatric phenomena may extend beyond the cerebellar areas linked only with regions traditionally regarded as limbic. A central question therefore emerges when considering the newly appreciated wider role of the cerebellum, namely, which parts of cerebellum are relevant to these different functions, and specifically, where, if at all, is the limbic cerebellum that is devoted to the processing of emotion and affect?

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This chapter reviews anatomical and physiological studies demonstrating links between the cerebellum and cerebral areas that subserve cognition and emotion, and it draws on pertinent functional imaging investigations and patient studies to underscore the clinical relevance of these connections.

Defining the Limbic System Paul Broca (1824–1880) drew attention to the cingulate gyrus, hippocampus, and parahippocampal gyrus at the medial surface of the mammalian brain (Broca 1878), introducing the term limbic lobe (derived from the Latin word limbus or edge) because it formed the hem, or edge, of the medial part of the cerebral hemisphere. These structures, along with the olfactory bulb, were initially considered to be important for the sense of smell. James Papez (1883–1958) introduced the notion of a limbic system, a number of interconnected brain areas that subserved the experience and expression of emotions. He proposed “that the hypothalamus, the anterior thalamic nuclei, the gyrus cinguli, the hippocampus, and their interconnections constitute a harmonious mechanism which may elaborate the functions of central emotion, as well as participate in emotional expression” (Papez 1937). Regions now considered integral parts of the limbic system include the amygdala, cingulate cortex, fornix, hippocampus, hypothalamus, olfactory cortex, thalamus, brainstem, ventral tegmental area, and parts of the prefrontal cortex (MacLean 1949, 1954, 1969; Nauta 1958, 1986; Heimer et al. 2008). Subcortical areas such as the ventral striatum and the extended amygdala were not initially conceptualized as part of the limbic circuit but are now recognized as brain substrates essential for the experience and expression of emotion (Schreiner and Kling 1953; Bucy and Kluver 1955; Nauta and Domesick 1976). Converging evidence now suggests that, like the ventral striatum, parts of the cerebellum also contribute to the distributed neural circuits subserving emotion. And if the cerebellum has a role in affective experience and expression, it must be anatomically linked with limbic regions in order for the cerebellum to influence the constituent elements of affect. Competing theories of the neurobiology of emotion have existed for some time, including the James-Lange (Lange and James 1922) view that feelings arise from physiological responses to the environment and the Cannon-Bard theory (Cannon 1927) which holds that physiological responses and feeling states arise simultaneously. Whichever theory is correct, interactions between mood state and physical experience are dependent upon the anatomical interactions between regions concerned with feelings and those concerned with physiologically defined phenomena, and on the way these brain areas regulate, or react to, the internal and external environment. The experience of feeling is also fundamentally influenced by one’s conscious awareness and interpretation of one’s own mood (Damasio 1999). Emotion and cognition are thus interdependent, as reflected in the extensive interconnections between the limbic system and cortical association areas (see Pandya and Yeterian 1985; Barbas et al. 2003). It is therefore pertinent to the present

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discussion of the anatomic systems underlying cerebellar connections with the limbic system to briefly consider the cerebellar incorporation into the distributed neural circuits that support cognitive operations.

Cerebellar Connections with the Limbic System Anatomical tract tracing and physiological studies demonstrate that the cerebellum is interconnected with the limbic system. The fornix, mammillothalamic tract, median forebrain bundle, and cingulum bundle are closely related to classic limbic system structures including the anterior thalamic nucleus, hypothalamus, cingulate cortex, and hippocampus. The limbic regions are also connected with the ventral tegmental area (VTA), periaqueductal gray, and interpeduncular nucleus. In the cat, the fastigial nucleus projects to the VTA, interpeduncular nucleus, periaqueductal gray, and locus coeruleus; the interpositus and dentate nuclei project to the interpeduncular nucleus and VTA (Snider and Maiti 1975). The mammillary bodies are closely linked with the anterior thalamic nuclei through the mammillothalamic tract and are connected with the cerebellum through projections to the nuclei of the basilar pons (Haines and Dietrichs 1984; Aas and Brodal 1988). The cingulate cortex is important for the introspective feeling of emotions, and it also supports other behaviors including initiation, motivation, and goal-directed behaviors (Devinsky et al. 1995). The anterior cingulate cortex modulates autonomic activity and internal emotional responses as well as a cognitive domain that is involved in skeletomotor activity and responds to noxious stimuli. Vilensky and Van Hoesen (1981) demonstrated in the monkey that the cingulate projects to the cerebellum through the nuclei of the basis pontis. Furthermore, the rostral cingulate projects to the medial pontine nuclei and the caudal cingulate to more lateral regions. Brodal et al. (1991) studied the organization of the connections of the cingulate cortex and cerebellum using the retrograde tracer, wheat germ agglutininhorseradish peroxidase (WGA-HRP). Following injections of tracer into the nuclei of the basis pontis, retrogradely labeled cells were found in layer V throughout the cingulate cortex. Topographic organization within the pons of fibers originating in different parts of the cingulate gyrus was also demonstrated. Cingulopontine fibers were distributed in a patchy mosaic within a narrow band along the ventromedial aspect of the pontine nuclei. With the combined use of lesions in the cingulate gyrus and injections of WGA-HRP in the ventral paraflocculus, there was considerable overlap between terminal fibers originating in the cingulate gyrus and cells retrogradely labeled from the ventral paraflocculus. Physiological mapping studies have been used to investigate cerebellar connections with extracerebellar structures, showing that cerebellar stimulation produces a number of effects on limbic structures. In cats and rats, stimulation of the rostral vermis, fastigial nucleus and intervening midline folia of the cerebellum result in facilitation of units in the septal region, inhibition of units in the hippocampus, and a mixed pattern of physiological responses in the amygdala (Babb et al. 1974).

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However, stimulating the lateral hemispheres of the cerebellum or the dentate nucleus produces no change in activity in these areas, and stimulation of the posterior vermis leads to inconsistent facilitation of the septal nuclei with no change in the hippocampus. These findings support the notion of Heath et al. (1978) that the fastigial nucleus is an integral part of the network for emotion and epilepsy.

Cerebellar-Hypothalamic Circuits Bidirectional pathways link all four intracerebellar nuclei with the hypothalamus that is engaged in somatic-visceral integration (see Zhu et al. 2006 for review). These connections may play a role in feeding, cardiovascular, osmotic, respiratory, micturition, immune, emotion, and other nonsomatic regulations. It has been proposed that these cerebellar-hypothalamic pathways are essential modulators and coordinators that integrate visceral and behavioral responses (Zhu et al. 2006). This role may explain, in part, why many studies that investigate visceral functions also implicate the cerebellum. These direct cerebellar-hypothalamic circuits have been confirmed both anatomically (see Dietrichs et al. 1994; Haines et al. 1997 for reviews) and physiologically (Bratus and Ioltukhovskii 1986; Supple 1993; Wang et al. 1994). Horseradish peroxidase (HRP) labeling studies of cerebellar-hypothalamic circuits reveal that the posterior part of the dorsomedial hypothalamic nucleus receives direct projections from the cerebellum, with somewhat fewer projections to the posterior hypothalamus nucleus (Onat and Cavdar 2003). In addition to these direct projections, there are also indirect projections from the hypothalamus via the basilar pontine nuclei to the cerebellum. Liu and Mihailoff (1999) demonstrated hypothalamopontine projections in rat to the rostral medial and dorsomedial portions of the pons which in turn project to the paraflocculus and vermis of the cerebellum. These authors hypothesize that since cerebral cortical inputs including limbic cortical inputs target similar pontine nuclei, the hypothalamopontine system might integrate autonomic and/or limbic-related functions with movement or somatic-related activity or, more likely, that the cerebellum uses both the direct and indirect hypothalamic inputs to perform integrative somatic, limbic, and visceral functions.

Cerebellar Connections with Paralimbic and Neocortical Association Areas The cerebellum is connected to the cerebral cortex through a two-step process with the pontine nuclei serving as the conduit between the corticopontine pathway and the mossy fiber-mediated pontocerebellar pathway. The two-step pathway from the cerebellum back to the cerebral cortex has the thalamus as the intermediate step between the cerebellothalamic pathway and the thalamocortical pathway. Schmahmann and Pandya (1992) examined the associative and paralimbic pathways

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in rhesus monkey and described a particular ordering of corticopontine projections. Each cerebral cortical area commits fibers that course with a predictable trajectory in the cerebral white matter, and that end in a unique patch of terminations in the nuclei of the basis pontis. Prefrontal cortical projections arise in the dorsolateral prefrontal cortex (DLPFC) and dorsomedial prefrontal cortex and terminate in the rostral third of the medial pons. Posterior parietal projections arise from both gyral and sulcal cortices including the multimodal caudal regions and terminate throughout the rostrocaudal extent of the pons. Projections from the temporal lobe arise from multimodal regions of the cortex of the upper bank of the superior temporal sulcus and the superior temporal gyrus and terminate in the lateral and dorsal pontine regions. Paralimbic cortices in the cingulate gyrus concerned with motivation, emotion, and drive (Devinsky et al. 1995; Paus 2001) communicate with the cerebellum via their projections to the pontine nuclei (Brodal et al. 1991; Vilensky and van Hoesen 1981). The caudal inferior parietal lobule, multimodal regions of the superior temporal gyrus, and the posterior parahippocampal formation that are interconnected with paralimbic structures also contribute to the corticopontine projection (Schmahmann 1996; Schmahmann and Pandya 1997). The cortico-ponto-cerebellar projections in the nonhuman primate are extensive (Glickstein et al. 1985; Schmahmann and Pandya 1991, 1993, 1997; Schmahmann 1996). There is a notable topographic arrangement of corticopontine projections in the monkey. The DLPFC and medial prefrontal cortex project to the medial pons, a finding confirmed also by electrophysiological recordings in the rat (Watson et al. 2009); parietal cortex to the lateral pons; superior temporal lobe (including language and auditory areas) to the lateral and dorsolateral pons; and the parahippocampal gyrus (spatial memory) and parastriate areas project to the dorsolateral pons (Schmahmann and Pandya 1989, 1991, 1992, 1993, 1995, 1997; Schmahmann 1996). Cerebellar efferents from the intracerebellar nuclei, including the dentate, target the ventrolateral thalamic nucleus (VL), a structure which has been known for many years to perform sensorimotor transformations, and which in turn projects to primary motor cortex (M1). The VL thalamic nucleus is composed of sub-regions that project also to nonmotor cerebral areas including prefrontal and posterior parietal cortices (Percheron et al. 1996, for review). Other cerebellar-recipient thalamic nuclei include the intralaminar nuclei (central lateral, paracentral, centromedian, and parafascicular) that have projections to the posterior parietal cortex, multimodal regions of the upper bank of the superior temporal sulcus, prefrontal cortex, cingulate gyrus, and primary motor cortex (Schmahmann 1996, for review). Thus, the likely consequence of widespread cerebellar connectivity with the cerebral cortex is that different cerebellar regions modulate nonmotor functions including attentional processes, emotion, and cognitive-related domains (Ito 1993; Schmahmann 1996, 2001, 2004; Schmahmann and Sherman 1998; Strick et al. 2009, for review). Earlier physiological studies showed that, in contrast to arm and leg sensorimotor representations in cerebellar anterior lobe and lobule VIII, with face representation additionally in lobule VI (Snider and Eldred 1951), the anterior cingulate region projects to medial parts of crura I and II. Association cortices project to the posterior lateral cerebellar hemispheres (Allen and Tsukahara 1974), and in particular, the

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parietal lobe is linked with lobule VII, including crus I, crus II, and paramedian lobule VIIB (Sasaki et al. 1975). Transsynaptic viral tract-tracing studies in monkey confirmed and extended these physiological observations, demonstrating that whereas the motor cortex is linked with cerebellar lobules IV, V, and VI, the prefrontal and posterior parietal cortices are reciprocally interconnected with cerebellar lobules crus I and crus II (the hemispheric extensions of lobule VIIA), and they also have connections with cerebellar lobules VIIB and X (Hoover and Strick 1999; Middleton and Strick 2001; Kelly and Strick 2003; Strick et al. 2009 for review). The cerebral association areas are considerably expanded in human compared to monkey. This is reflected in the composition of the cerebral peduncle that conveys the corticopontine fibers. Using diffusion tensor magnetic resonance imaging, it appears that, whereas in the monkey, the majority of corticopontine fibers are derived from the motor system, the bulk of the corticopontine fibers in the human are derived from prefrontal regions (Ramnani 2006).

Observations from Neuroimaging Studies Functional Topography in the Cerebellum Activation of the cerebellum by tasks within multiple cognitive domains including executive, linguistic, mnemonic, attentional, and visuospatial have been demonstrated by positron emission tomography (PET) and functional magnetic resonance imaging (fMRI). Meta-analysis using an activation likelihood estimation approach reveals that there is a functional topography within the cerebellum (Stoodley and Schmahmann 2009). The sensorimotor cerebellum is located in the anterior lobe, parts of lobule VI, and lobule VIII, and the cognitive cerebellum is located in lobules VI and VII at the vermis and in the hemispheric extensions into lobule VI, as well as crura I and II of lobule VIIA, and lobule VIIB. Functional MRI of a range of cognitive and motor paradigms shed further light on the topographic arrangement of these different processes in the cerebellum (Stoodley et al. 2012). Five tasks investigating sensorimotor (finger tapping), language (verb generation), spatial (mental rotation), working memory (N-back), and emotional processing (viewing images from the International Affective Picture System [IAPS]) were conducted in nine healthy subjects. Finger tapping activated the ipsilateral anterior lobe (lobules IV-V) as well as lobules VI and VIII. Activation during verb generation was found in right lobules VII and VIIIA. Mental rotation activated left-lateralized clusters in lobules VII-VIIIA, VI, crus I, and midline VIIAt. The N-back task showed bilateral activation in right lobules VI, crus I, and left lobules VIIB-VIIIA. Cerebellar activation was evident bilaterally in lobule VI while viewing arousing vs. neutral images. This fMRI study provided support for topographic organization of motor execution vs. cognitive/emotional domains within the cerebellum, likely reflecting the anatomical specificity of cerebrocerebellar circuits underlying different task domains. There is also evidence for topographic organization of cerebellar nonmotor functions within a single individual (Stoodley et al. 2010), including activation by emotion-inducing images of the IAPS task in the cerebellar vermis and lateral hemispheres.

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Activation of the cerebellum in studies of the neural correlates of pain anticipation, thirst, hunger, and smell provide additional fMRI evidence for a limbic cerebellum consisting of the fastigial nucleus, vermis, and flocculonodular lobe. Cerebellar vermis activation is seen in neuroimaging studies investigating panic (e.g., Reiman et al. 1989), sadness, and grief (Beauregard et al. 1998; Gundel et al. 2003). Autonomic processing (Parsons et al. 2000), including the autonomic cardiovascular arousal that occurs during both exercise and mental arithmetic stressor tasks (Critchley et al. 2000), and air hunger (Evans et al. 2002) result in activation of posterior cerebellar regions in both the midline and lateral hemispheres.

Cerebellum and Pain Modulation Nociception is a complex behavior that recruits autonomic, sensorimotor, affective, and cognitive systems. Human pain imaging studies regularly demonstrate activation in the cerebellum, and investigations directed specifically at the role of the cerebellum in pain appreciation have only recently been performed. Anterior regions of the cerebellum are activated by the experience of pain (Becerra et al. 1999), whereas posterior regions are active during the anticipation of pain (Ploghaus et al. 1999). Different cerebellar regions are involved when processing one’s own painful experience (posterior vermis) as opposed to experiencing empathy for another’s pain (lobule VI; Singer et al. 2004). The activation of hemispheral lobule VI and vermal lobule VII is quite consistent across these studies of emotionally salient stimuli (Stoodley and Schmahmann 2009), and like the activation patterns seen within the cerebellum for cognitive tasks, the focus of the cerebellar activation varies according to the demands of the task. It is possible that hemispheric lobules VI and VII activation reflects more cognitive components of task performance (e.g., empathy), due to the connections of these regions with association cortices, and more limbic tasks (including autonomic processing) may particularly involve the posterior vermis, the putative limbic cerebellum. Helmchen et al. (2003) examined perceived pain intensity to a noxious and nonnoxious thermal stimulus. In contrast to innocuous stimuli, painful stimuli revealed activation in the intracerebellar nuclei, anterior vermis, and bilaterally in cerebellar hemispheric lobule VI. With the same noxious stimulus, there was differential cerebellar activation depending on the perceived pain intensity: high pain intensity ratings were associated with activation in ipsilateral hemispheric lobules III-VI, deep cerebellar nuclei, and in the anterior vermis (lobule III). This differential cerebellar activation pattern reflects a cerebellar role not only in somatosensory processing but also in perceived intensity of a painful stimulus, indicating a role for the cerebellum in the modulation of nociceptive circuits. Borsook and colleagues (2008) studied the effects of noxious thermal heat and brush applied to the face in a group of healthy subjects and patients with neuropathic pain and defined areas of activation in the cerebellum. The data indicate that different regions of the cerebellum are involved in acute and chronic pain processing. Heat produces greater contralateral activation compared with brush

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stimulation, whereas brush resulted in more ipsilateral/bilateral cerebellar activation. Further, innocuous brush stimuli in healthy subjects produced decreased cerebellar activation in lobules concerned with somatosensory processing. These data provided support for a dichotomy of innocuous stimuli/sensorimotor cerebellum activation versus noxious experience/cognitive/limbic cerebellum activation. These authors (Moulton et al. 2011) later demonstrated that the cerebellum may contain specific regions involved in encoding generalized aversive processing. In their study, aversive stimuli in the form of noxious heat and unpleasant pictures (as tested using the IAPS) were shown to activate overlapping areas in cerebellar lobules VI, crus I, and VIIB. Further, functional connectivity MRI (fcMRI) mapping indicated that cerebellar areas showing functional overlap with pain and viewing unpleasant pictures are interconnected with limbic system structures including the anterior hypothalamus, subgenual anterior cingulate cortex, and the parahippocampal gyrus.

Autonomic Influences The earlier physiological literature (Martner 1975) as well as clinical (Heath et al. 1979; Schmahmann and Sherman 1998; Levisohn et al. 2000; Schmahmann et al. 2007; Tavano et al. 2007) and behavioral studies in animals (Berman et al. 1978) suggest that the cerebellar vermis is involved in the regulation of a range of nonsomatic functions including cardiovascular control, thirst, and feeding behavior. Cerebello-hypothalamic circuits have been postulated to be a potential neuroanatomical substrate underlying this modulation. Demirtas-Tatlidede and colleagues (2011) tested the relationship between the cerebellar vermis and nonsomatic functions by stimulating the cerebellum noninvasively using transcranial magnetic stimulation. Theta burst stimulation (TBS) was applied to the vermis (identified with neuronavigation software), the right cerebellar hemisphere and the left cerebellar hemisphere in 12 healthy human subjects with the aim of modulating cerebellar activity. Following stimulation of the vermis, but not the cerebellar hemispheres, there was a mild but significant decrease in heart rate, and subjective ratings by subjects indicated a significant increase in thirst as well as a trend toward increased appetite. These observations are consistent with physiological and imaging data indicating a role for the cerebellum in the regulation of visceral responses and suggest a role for the vermis in somatovisceral integration, likely through cerebello-hypothalamic pathways.

Indirect Limbic Inputs Resting-state functional connectivity magnetic resonance imaging (fcMRI) studies have provided novel insights into the interactions of the cerebellum with the cerebral hemispheres in the human brain (Habas et al. 2009; Krienen and Buckner 2009; O’Reilly et al. 2010). These investigations demonstrate (physiological)

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connections of limbic and paralimbic areas of the cerebral hemispheres with the cerebellar vermis and with crura I and II of lobule VIIA. The cerebellum receives the majority of its inputs from the cerebral hemispheres by way of the pontine nuclei. Associative and limbic inputs to the cerebellum may be more widespread than originally conceptualized, however. Functional connectivity MRI reveals that spontaneous activity in the nucleus accumbens (NAcc), a major structure in the dopaminergic mesolimbic system, predicts activity from a reward paradigm in limbic structures including the orbital medial prefrontal cortex, amygdala, insula, and posterior parietal cortex as well as the cerebellum, despite an absence of direct connections between cerebellum and the NAcc (Cauda et al. 2011). This could help explain in part why limbic-cerebellar circuitry has been overlooked – its functional connectivity may include anatomical connections mediated by second or third stage areas.

The Cerebellum Implicated in Autism Spectrum Disorders (ASD) There is evidence that autism is associated with structural brain abnormalities that include the cerebellum. Postmortem studies of early infantile autism reveal decreased neurons in deep cerebellar nuclei and reduced numbers of Purkinje cells especially in posterolateral cerebellar cortex (e.g., Bauman and Kemper 1985). Using MRI to compare the brains of normal subjects with those of individuals with autism, Courchesne et al. (2001) reported that cerebellar vermal lobules VI and VII are significantly smaller in autistic individuals. These regions of the cerebellum are connected to brain areas that govern attention, arousal, and the assimilation of sensory stimulation. Certain characteristic features of autism, including sensitivity to sensory stimuli and repetitive behavior, may therefore be explained in part by these cerebellar structural abnormalities. In addition to neuropathology in the cerebellum in autism, GABAergic differences in the posterolateral cerebellar cortex in the crus II region have recently been reported (Yip et al. 2007, 2008, 2009). In adult postmortem subjects with autism, the remaining Purkinje cells and larger sized interneurons in the dentate nuclei contained decreased glutamic acid decarboxylase type 65 or 67 (GAD 65/67) whereas molecular layer interneurons such as basket cells contained increased GAD 67 compared to age-matched controls. Since the crus II region is a major recipient area for frontal lobe afferents via the pons, dysfunction of this portion of the lateral posterior hemisphere and dentate may influence autistic behaviors including social interactions, executive function, and affect. An fMRI study of young adults with autism found increased cerebellar activation by motor tasks, but decreased cerebellar activation in an attention paradigm (Allen and Courchesne 2003). These findings were thought to be related to deficient sensory tracking when stimuli appeared at a rapid rate, reflecting decreased cerebellar volumes in individuals with autism. Kates et al. (1998) studied a pair of monozygous twins, one of whom met criteria for strictly defined autism while the other showed restrictions in social interaction

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and play but did not meet these criteria. Both boys had smaller frontal lobes and superior temporal gyri compared to healthy controls, but the more affected twin had smaller cerebellar vermis lobules VI and VII as well as smaller caudate, amygdaloid, and hippocampal volumes, in line with data consistently suggesting a role for the cerebellum in autism. Children with attention-deficit/hyperactivity disorder have reduced volumes of the posterior inferior lobe of the cerebellum compared to age-matched control subjects, even adjusting for brain volume and IQ (Berquin et al. 1998). And adults with Down’s syndrome also have smaller cerebellar volumes than age-matched controls after controlling for total intracranial volume and total brain volume, a difference that did not appear to change over time in a small subset of patients followed serially (Aylward et al. 1997). Recently, neuropathology in the hippocampus and cerebellum was reported in three postmortem cases of fragile X syndrome, one of the autism spectrum disorders that results from a single gene defect, the fragile X mental retardation 1 (FMR1) gene with the FMR1 protein being absent or deficient in affected individuals (Greco et al. 2011). The size of the vermis was decreased preferentially in posterior lobules VI and VII, and there were decreased numbers of Purkinje cells throughout the cerebellum. Additionally, altered neuronal migration was evident both in the hippocampus and cerebellum, indicating the vulnerability of the two structures to a single gene defect.

Implications of a Cerebellar Role in Emotional Processing In the original description of the cerebellar cognitive affective syndrome (Schmahmann and Sherman 1997, 1998; Levisohn et al. 2000), patients whose pathology included the cerebellar vermis and midline regions containing the fastigial nucleus experienced personality changes characterized by either flattening of affect or disinhibition with inappropriate behavior. These behavioral phenomena following midline lesions are in accord with earlier physiological experiments demonstrating a relationship between the fastigial nucleus and complex behaviors such as sham rage, grooming repertoires, and predatory attack (Berntson et al. 1973; Reis et al. 1973; Ball et al. 1974). Changes in mood are reported in patients with what was previously known as dominantly inherited olivopontocerebellar atrophy (OPCA) (Kish et al. 1988), now recognized as one of a number of spinocerebellar ataxias. Patients experience higher depression scores compared to controls, and the depression scores correlate weakly with cognitive testing. In the sporadic form of OPCA now know as multiple systems atrophy cerebellar type (MSAc), 36% of patients experience pathological laughing and crying (PLC; Parvizi et al. 2007) at some point in the course, reflecting incongruity between the experience and expression of emotion. When the MSA is predominantly of the parkinsonian form (MSAp), the incidence of PLC is substantially lower, on the order of 3%. Both MSAc and MSAp are synucleinopathies, with the principal difference between them lying in the degree of cerebellar pathology.

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Imaging studies link the cerebellum to depression. Increased cerebral blood flow was noted in the cerebellar vermis in the study by Mayberg et al. (1999) who induced transient sadness in healthy volunteers and in patients with depression. An MRI voxel-based morphometric study (Peng et al. 2011) investigating gray matter density in patients with major depressive disorder (MDD) found volumetric reductions in the left cerebellar hemisphere along with lower volumes of gray matter of the frontal-limbic cortical areas including orbital frontal cortex (OFC) and dorsolateral prefrontal cortex (DLPFC). And resting-state fcMRI in patients with depression reveals that connectivity between the cerebellar vermis and the posterior cingulate cortex correlates with severity of the depression (Alalade et al. 2011). These results are supported by postmortem studies in MDD patients, which show reduced glial fibrillary acidic protein in the cerebellum (Fatemi et al. 2004). Deficits in behavior and emotions are apparent when the cerebellar vermis is involved, as occurs in patients with congenital malformations and/or tumors (Pollack et al. 1995; Schmahmann et al. 2007; Tavano et al. 2007; Hernaez-Goni et al. 2010). Indeed, the posterior vermis has been conceptualized as the limbic cerebellum (Schmahmann 1991, 2000), and vermal abnormalities appear to account in large part for the emotional disturbances, inappropriate behavior and changes in affect that characterize the cerebellar cognitive affective syndrome (Schmahmann and Sherman 1998; Schmahmann et al. 2007). In addition to its role in language, working memory, spatial processing, and regulation of mental activities, the cerebellum is implicated in the pathophysiology of developmental disorders including schizophrenia, stuttering, dyslexia, and autism spectrum disorders (Schmahmann 1991, 2000; Rapoport et al. 2000; Schutter and van Honk 2005; Stoodley 2011). The precise role of the cerebellum as well as the mechanisms by which the cerebellum modulates cognitive tasks remains a matter of ongoing study (Schmahmann 1997; Strick et al. 2009). In one recent report (Hopyan et al. 2010), the authors investigated whether children with cerebellar tumors could identify emotion in situations in which emotion in music and emotion in the accompanying lyrics were either congruent or incongruent. The study found that cerebellar patients had no problem with emotion identification, but they were impaired on the tasks that required cognitive control of emotion. Control of behavior, both motor and nonmotor, appears to be a central theme in the role of the cerebellum in nervous system function, as encapsulated in the dysmetria of thought theory (Schmahmann 1991, 2000) discussed elsewhere in this volume.

Therapeutic Implications There are potential therapeutic implications to the new appreciation that the cerebellar vermis may be the putative site of the limbic cerebellum contributing to the modulation of emotional processing. A recent pilot study of cerebellar vermal stimulation with noninvasive transcranial magnetic stimulation in patients with schizophrenia demonstrated that this approach is safe and well tolerated in a psychiatric population. It also provided preliminary evidence for clinical efficacy

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as measured by psychiatric rating scales (Demirtas-Tatlidede et al. 2010). More definitive studies of cerebellar neuromodulation in psychiatric and neuropsychiatric populations are awaited. Conclusions

The identification and delineation of cerebellar connections with cerebral cortical association areas was a critical step in the determination of the cerebellar role in nonmotor function. The clinical syndromes of cognitive impairment resulting from cerebellar damage reflect loss of cerebellar influence on the association areas with which cerebellum is interconnected. In a similar manner, neural circuits that subserve emotional processing are also linked with cerebellum, and likely to be relevant in the affective impairments that characterize some patients with cerebellar lesions. Evidence for a cerebellar influence on emotional processing is derived from anatomical studies as well as physiological and behavioral investigations in animal models, functional imaging studies in healthy subjects, and clinical observations in patients. The present view supported by these lines of inquiry is that the cerebellar vermis and fastigial nucleus represent the cerebellar limbic system and are engaged in the modulation of affective behavior. Further studies are needed to explore the connections of cerebellum with limbic and paralimbic regions within the cerebral cortex and subcortical areas and to further define the putative limbic-related regions in the cerebellum. It also remains to be shown how cerebellar limbic areas influence autonomic regions of the body directly or indirectly and the nature of cerebellar modulation of limbic behaviors. Further definition of these cerebellar limbic interactions will promote a deeper understanding of the clinical manifestations of cerebellar injury on affect and emotion, as reflected in the cerebellar cognitive affective syndrome and the neuropsychiatry of the cerebellum. Acknowledgments Supported in part by the Sidney R. Baer Jr., Birmingham, and MINDlink Foundations (JDS), Eunice Kennedy Shriver National Institute of Child Health and Human Development 5R01HD039459-06A1, and The Hussman Foundation (GJB). The assistance of Jason MacMore and Caitlin Clancy is gratefully acknowledged.

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Cerebellar Influences on Descending Spinal Motor Systems

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Tom J. H. Ruigrok

Abstract

The cerebellar nuclei, and to some extent the vestibular nuclei, mediate the ultimate result of cerebellar processing to the rest of the brain. Cerebellar output is directed to the diencephalon and to a score of brainstem regions. This chapter reviews the cerebellar nuclear projections to the brainstem areas that give rise to descending connections that can influence motor programming at spinal cord level, i.e., the reticulospinal, vestibulospinal, rubrospinal, tectospinal, and interstitiospinal pathways. In addition, cerebellar projections to other areas will be briefly considered. Although cerebellar output is structured by the modular internal organization of cerebellar circuitry and related olivocerebellar connections, it is concluded that the modular output, that is, output of individual cerebellar nuclei or parts thereof, still reaches many areas in the brainstem and diencephalon. In addition, multiple modules may converge their outputs indirectly to the same muscles. This suggests that multiple modules may each take part in different aspects of control of the same muscle or muscle group. Conversely, individual modules, due to the distributed nature of their outputs, may simultaneously affect several descending motor systems with the same ensuing goal. More detailed anatomical and physiological studies will be necessary to explore these possibilities.

Introduction The cerebellum, which is dominantly involved in the coordination, adaptation, and learning of motor behavior and, most likely, also participates in visceral, affective, and cognitive functions, exerts its influence on these functions by way of the

T.J.H. Ruigrok Department of Neuroscience, Erasmus Medical Center Rotterdam, 3000 CA, Rotterdam, The Netherlands e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 497 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_23, # Springer Science+Business Media Dordrecht 2013

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cerebellar nuclei. In addition, selected sets of Purkinje cells provide a direct input to specific regions of the vestibular nuclei. The specific connections of virtually all regions of the cerebellar nuclei with those parts of the thalamus that are connected to the primary and premotor cortices were already recognized and stressed in the early literature (Allen and Tsukahara 1974). However, the cerebellar nuclei also have a prominent direct impact on a number of premotor centers with connections to the spinal cord or lower brainstem. Here, a brief overview of the cerebellar nuclei will be provided followed by a review of their connections to these descending motor systems. Finally, these connections will be briefly assessed in relation to other regions that receive cerebellar output and their joint impact on cerebellar functioning and dysfunctioning.

The Cerebellar Nuclei The cerebellar nuclei in rodent and nonhuman primate are divided into four main nuclei: the medial, posterior interposed, anterior interposed, and lateral cerebellar nuclei (Fig. 23.1). These divisions are essentially based on cytoarchitectonic grounds. However, these basic nuclear divisions also agree with organizational features of the cerebellum. Purkinje cells projecting to each nuclear subdivision are organized into longitudinal strips. Jan Voogd, in his historic work (Voogd 1964), proposed that the medial cerebellar nucleus (MCN) receives its cortical afferents from the A zone, both interposed nuclei from the C-zone and the lateral cerebellar nucleus (LCN) from the D zone. The B zone, intercalated between A and C zones, targets the lateral vestibular nucleus. Presently, this basic scheme still holds true although the description of multiple subdivisions of the various zones has been refined considerably and has been shown to correspond with the description of several nuclear subdivisions (Apps and Hawkes 2009; Ruigrok 2011; Voogd and Glickstein 1998; Sugihara and Shinoda 2007). The remarkable reciprocal relation between the inferior olive and cerebellar nuclei has been noted by several authors and may form the basis of the olivocerebellar organization (Ruigrok 1997; Ruigrok and Voogd 1990, 2000). Here, a short general description of the cerebellar nuclei of the rat, cat, and macaque will be presented before detailing the projections from these nuclei to the rest of the brain (see Fig. 23.1). The MCN, in cat and macaque often referred to as the fastigial nucleus, is the most medially located nucleus. Its rostral half is found directly dorsomedial to the rostral half of the fourth ventricle. Caudally, it is found dorsal to the lateral part of the nodulus. In the rat, as in mice, the MCN is characterized by a prominent outgrowth that reaches up into the white matter, consists of generally large cells, and is known as the dorsolateral protuberance (DLP: Korneliussen 1968). The rostral tip of the MCN is continuous with the medial aspect of the superior vestibular nucleus. Caudally, the MCN is separated from both parts of the interposed nuclei by passing corticovestibular fibers. Located within these fibers are small groups of nuclear cells which are collectively known as the interstitial cell groups (ICG; Buisseret-Delmas et al. 1998). The ICG, based on their cortical and

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Fig. 23.1 Three-dimensional reconstructions of the left cerebellar nuclear complex of the rat (left), cat (middle), and macaque (right). Top panels depict the cerebellar nuclei in a rostral view (inverted image); middle panels show the nuclei in a dorsal (top) view; bottom panels in a caudal view. In the macaque the basal interstitial nucleus (BIN) is also shown. Note that the general organization of the nuclei is virtually similar. For abbreviations see List. Reconstructions were made with Neurolucida (MicroBrightfield, Inc, Williston, VT, USA) using serial microscopical sections (rat, cat) or sections from the macaque atlas by Paxinos et al. (2000). For abbreviations see list

efferent connections, seem to be mostly associated with the posterior interposed nucleus (PIN; Buisseret-Delmas et al. 1993; Pijpers et al. 2005). Although the ICG are not mentioned in the description of the cerebellar nuclei of other mammals, they can also be recognized in the cat. The PIN is found directly lateral to the caudal half of the MCN where it rests on the roof of the fourth ventricle. More rostrally and laterally, its borders with the anterior interposed and lateral cerebellar nuclei, respectively, are not always clear, especially in transverse sections. The rostromedial extension of the PIN continues as the interstitial cell groups (Buisseret-Delmas et al. 1993, 1998). In cat, the PIN includes a medial and lateral dorsal-ward protrusion. The anterior interposed nucleus (AIN) of all three mammalian species has a mediolaterally elongated shape. Within this nucleus a clear somatotopic pattern can be recognized with a representation of the hind limb located mediorostrally and the head region in its caudolateral aspect (e.g., see Lu et al. 2007; Pijpers et al. 2005). In the rat, a conspicuous bulge is noted at the border region between the AIN and the LCN. This dorsolateral hump (DLH) has been associated with either the

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Fig. 23.2 Serial plots (b–g) of microscopical sections depicting patterns of labeled varicosities (i.e., terminal arborizations) in the caudal brainstem after various injections (shown in a) with an anterograde tracer in the cerebellar nuclei of the rat. The boundaries of several nuclei and fiber tracts are indicated. Note the generally widespread distribution of projections in every case. For abbreviations, see list (Adapted from Teune et al. 2000)

AIN or the LCN (Voogd 2004; Korneliussen 1968). Recently, it has been shown to receive cortical afferents from a strip of Purkinje cells located between the D1 and D2 zones, which has been termed D0, and to receive climbing fiber collaterals that are derived from the dorsomedial group that is associated with the principal olive (Pijpers et al. 2005; Sugihara and Shinoda 2004, 2007). It is remarkable that the DLH provides a prominent ipsilateral descending projection to the lateral medullary reticular formation and adjacent spinal trigeminal nucleus (Fig. 23.2f; Bentivoglio and Kuypers 1982; Teune et al. 2000).

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The LCN, also referred to as dentate nucleus in cat and primates, is rostrally separated from the AIN by traversing fibers from the inferior cerebellar peduncle. In the rat, the LCN consists of the dorsolateral magnocellular part and a ventromedial parvicellular part. In the rhesus monkey, these areas actually form a gyrated, or toothed, area, with the medial sheath consisting of mostly small neurons, whereas the lateral sheath is made up of mostly large cells. Throughout the neuropil of all cerebellar nuclei, small cells are located that have been demonstrated to supply a GABAergic input to the inferior olive (De Zeeuw et al. 1989; Fredette and Mugnaini 1991; Teune et al. 1995). In the macaque, a loosely packed group of neurons, the basal interstitial nucleus (BIN, Fig. 23.1), is found ventral to the LCN that continues in a rostrolateral direction toward the floccular peduncle and which maintains reciprocal connections with the vestibulocerebellum (Langer 1985). In the cat and rat, neurons in a similar position are found but form an even less homogeneous nuclear contour (Ruigrok 2003).

Medial and Lateral Descending Motor Systems from the Brainstem Hans Kuypers divided the pathways descending to the spinal cord into three groups (Kuypers 1981, 1985). The first group consists of the corticospinal tract, which is subdivided into the crossed lateral and uncrossed medial descending tracts. A second group of descending tracts originates from the brainstem and is also divided into a medial and a lateral descending system based on the funicular course of the fibers and the termination pattern of the participating fibers within either the medial or lateral part of the intermediate region (i.e., layers V–VIII) of the spinal cord). Finally, a third group originates from the locus coeruleus and subcoeruleus as well as from the raphe nuclei and terminate diffusely throughout the gray matter of the spinal cord. This review will focus on the cerebellar influence of the second group of descending connections, that is, the reticulospinal, vestibulospinal, tectospinal, rubrospinal, and interstitiospinal tracts.

Reticulospinal Tracts The medial and medioventral pontomedullary reticular formation are at the origin of several long descending tract systems (Torvik and Brodal 1957). Delineation and identification of various subregions has proven to be difficult and may be different in different species. For example, in the rat, over 25 reticular regions have been shown to contain neurons with descending fibers (Newman 1985a, b). Here, we will only describe the cerebellar influence on the reticulospinal tracts in a general sense. It is well established that reticulospinal connections may follow several routes to the spinal cord (Nyberg-Hansen 1965; Peterson et al. 1975). Reticulospinal neurons located in the pontine and rostral medullary reticular formation mostly, but

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certainly not exclusively, follow a course by way of the ventral funiculus where they are found adjacent to the fibers of the medial longitudinal fascicle (mlf). They will terminate throughout the length of the spinal cord predominantly in the medial and central parts of the intermediate zone (i.e., in laminae VII–VIII). Additional reticulospinal tracts are found in the ipsilateral and contralateral ventrolateral funiculus. Both tracts convey axons of reticulospinal neurons from more caudal regions. Projections from these tracts are found throughout all laminae of the spinal cord. Individual reticulospinal axons can collateralize over considerable distances thereby affecting different regions of the body (Peterson et al. 1975). To further exemplify the heterogeneous nature of the reticulospinal tracts, it has been shown that the individual fibers may be excitatory (glutamatergic), inhibitory (containing GABA, glycine or both), or modulatory (e.g., serotonin) (Holstege and Kuypers 1987). At least some of the reticulospinal axons can have monosynaptic excitatory effects on motoneurons located throughout the length of the spinal cord (Shapovalov and Gurevitch 1970; Shapovalov 1972). From the above, it follows that the actions of the reticulospinal system are extremely diverse (for review, see Arshavsky et al. 1986). Indeed, reticulospinal systems have been suggested to be involved in maintaining and controlling ongoing motor activity, in the gating of somatosensory information to segmental as well as supraspinal levels, and in the control of autonomic activity including pain modulatory systems (Fields 2004; Saper 2004). The cerebellar nuclei are known to project prominently to the reticular formation (see Fig. 23.2), thereby involving the regions that give rise to the reticulospinal tracts. In fact the only nuclear region that does not seem to contribute significantly to pontomedullary reticular projections is the posterior interposed nucleus (PIN, Fig. 23.2, Table 23.1). Major inputs have been described to arise from specific areas of the MCN and LCN. In addition, in the rat, the nuclear region intercalated between the MCN and the interposed nuclei, that is, the ICG, and the transition area between the interposed nuclei and the LCN, that is, the DLH, also display projections to the pontomedullary reticular formation. The organization of cerebellar nuclear projections is highly specific as will be reviewed below.

Medial Cerebellar Nucleus Projections from especially the caudal aspect of the MCN have been shown to terminate in the dorsal aspect of the contralateral medial pontomedullary reticular formation by way of the uncinate tract in rat (Fig. 23.2d), cat, and monkey (Asanuma et al. 1983; Batton et al. 1977; Teune et al. 2000; Voogd 1964). In the monkey, the caudal region of the fastigial nucleus has been implicated in oculomotor functions subserving saccades (Noda 1991; Gonzalo-Ruiz et al. 1988). This area not only projects to the contralateral medial pontine reticular formation, known to be involved in saccade control (Enderle 2002), but also to the contralateral dorsomedial medullary reticular formation (Noda et al. 1990). This latter area would overlap the region from where neck muscles can be directly activated or

Medulla oblongata Inferior olive Lateral reticular nucl. Nucl. of the solitary tract Parasolitary nucl. Medullary reticular nucl. Paramedian nucl. Parvocell. retic. nucl. Gigantocell. retic. nucl. Lat. paragigantocell. nucl. Spinal trigem. nucl., oral part Spinal trigem. nucl., interpolar part Spinal trigem. nucl., caudal part Superior vestibular nucl. Lateral vestibular nucl. Medial vestibular nucl., rostral Medial vestibular nucl., caudal Spinal vestibular nucl. Nucl. prepositus hypoglossi Metencephalon Basal pontine nuclei -

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-

ICG (T82)

Table 23.1 Overview of brainstem and diencephalic regions of the rat where labeled varicosities were found after injection with an anterograde tracer in the part of the cerebellar nuclei indicated and, at least partly, depicted in Figs. 23.2 and 23.3. Large dots denotes dense labeling, intermediate dots indicate fair labeling, small dots indicate sparse labeling. Question mark indicates that the area was not available for analysis; i indicates that the labeling was found ipsilateral to the injection side (based on Teune et al. 2000) MCN PIN AIN LCN

23 503

Nucl. retic. tegm. pontis A5 noradrenergic group (?) Principal sens. trig. nucl. Caud. pont retic. nucl. Oral pont. retic. nucl. Central gray pons Pedunculopontine tegm. nucl. Parabrachial nucl. Mesencephalon Red nucl., parvicellular Red nucl., magnocellular Pararubral area Deep mesenceph. nucl. Superior colliculus, superficial Superior colliculus, intermediate Superior colliculus, deep Ventral tegmental area Dorsal raphe nucleus Ventral tegm. relay zone Medial access. oculomot. nucl. Interstitial nucl. of Cajal Nucl. of Darkschewitsch Nucl. parafascicularis prerubr. Periaquaductal gray

Table 23.1 (continued)

-

-

-

-

-

-

-

-

-

-

-

-

-

-

-

-

ICG (T82)

-

PIN Caudal (R100)

Rostral (T195) -

DLP (R127)

MCN

-

-

-

-

-

-

medial (T79)

-

-

-

-

lateral (T108) -

-

-

-

-

-

-

medial (R89)

AIN

-

-

-

-

-

lateral (R98)

-

-

-

-

-

DLH (R138)

-

-

-

-

-

-

rostral (T77)

LCN

-

-

-

-

ventral (R178)

-

-

-

-

-

caudal (T94)

504 T.J.H. Ruigrok

Nucl. of post. commissure Anterior pretectal nucl. Posterior pretectal nucl. Diencephalon Mammillary nuclei Lateral hypothal. area Dorsal hypothal. area Zona incerta Nucl. fields of Forel Ventral lat. geniculate nucl. Parafascicular thalam. nucl. Central medial thalam. nucl. Laterodorsal thalam. nucl. Posterior thalamic nucl. Ventroposterior thalam. group Ventromedial thalam. nucl. Ventrolateral thalam. nucl. Total number of areas With fair or dense labeling -

-

19 10

-

-

? 31 14

-

-

-

? 40 21

-

-

26 10

-

-

-

-

-

-

-

? 15 4

-

-

-

16 9

-

-

-

27 7

-

-

-

28 10

-

-

21 13

-

-

-

-

-

25 15

-

-

-

-

? 31 26

-

-

41 25

-

23 Cerebellar Influences on Descending Spinal Motor Systems 505

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T.J.H. Ruigrok

inhibited (Peterson et al. 1975). Ipsilateral reticular projections that originate from the MCN are conspicuously less dense. They enter the brainstem by way of the direct fastigiobulbar tract located medially adjacent to the superior cerebellar peduncle (Teune et al. 2000; Voogd 1964; Ruigrok et al. 1990). The dorsolateral protuberance (DLP) of the MCN, which is prominent in rodents, has been shown to more specifically reach intermediate and lateral (i.e., parvocellular) parts of the contralateral reticular formation (Fig. 23.2c; Rubertone et al. 1990; Teune et al. 2000). The rostral MCN seems to preferentially target the vestibular nuclei (see below) and sends only moderate projections to the intermediate (mediolateral) levels of the reticular formation and to the lateral paragigantocellular nucleus (Fig. 23.2b). Bagnal and coworkers (2009) recently provided evidence that the ipsilateral connections of the MCN are glycinergic whereas glutamatergic MCN neurons project contralaterally. By definition, the Purkinje cells that innervate the MCN belong to the A zone (Voogd 1964; Voogd and Glickstein 1998). More recently, the A zone has been subdivided into a number of subzones, each of which relate to a specific region of the MCN (Apps and Hawkes 2009; Voogd and Ruigrok 2004). Sugihara and Shinoda (2007) have suggested that three regions may be recognized in the MCN dealing with spinal (DLP and rostral MCN), eye movement (centrodorsocaudal MCN), and head orientation (centrocaudal MCN) aspects of motor control.

The Posterior Interposed Nucleus and the Interstitial Cell Groups The ICG of the rat, which are intercalated between the MCN and both interposed nuclei, are the target of the Purkinje cells of the X and CX zone (Buisseret-Delmas et al. 1993, 1998; Voogd and Ruigrok 2004). They have been shown to have projections to the contralateral pontomedullary reticular formation, where they mostly terminate in the gigantocellular reticular nucleus (Fig. 23.2e; Teune et al. 2000). In the rat, virtually no pontomedullary projections have been described that originate from the main body of the PIN.

The Anterior Interposed Nucleus and the Dorsolateral Hump The dorsolateral hump (DLH) is sometimes also incorporated as part of the LCN (Angaut and Cicirata 1990), because it maintains reciprocal connections with the dorsomedial group of the principal olive (Ruigrok 1997). Nevertheless, it was originally described as part of the AIN (Goodman et al. 1963; Korneliussen 1968) and the DLH and lateral part of the AIN give rise to the ipsilateral descending tract of the cerebellum (Bentivoglio and Kuypers 1982; Bentivoglio and Molinari 1985; Mehler 1967). Their efferents terminate mostly in the parvicellular regions of the ipsilateral pontomedullary reticular formation but also invade the deeper layers of the spinal trigeminal nucleus (Fig. 23.2f; Teune et al. 2000). Stimulation of the DLH

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induces movement of the lips, neck, and forelimbs (Cicirata et al. 1992; Angaut and Cicirata 1990), indicating perhaps that it is involved specifically in food manipulation. Indeed, the lateral and intermediate medullary reticular formation of the rat have been demonstrated to contain pre-oromotor neurons (Travers et al. 2000) and are involved with mastication and swallowing (Luo et al. 2001). More rostral regions of the lateral reticular region have been shown to be involved in vocalization in the squirrel monkey (Hannig and Jurgens 2006). The DLH receives its Purkinje fiber input from the D0 zone, which is intercalated between the D1 and D2 zones of lobules V–VII (Pijpers et al. 2005; Sugihara and Shinoda 2004).

The Lateral Cerebellar Nucleus The connections of the LCN with the pontomedullary reticular formation are well documented for rat, cat, and monkey (Chan-Palay 1977; Teune et al. 2000; Tolbert et al. 1980). Particularly dense projections are noted in the contralateral gigantocellular reticular nuclei (Fig. 23.2g). The projection originates mostly from the magnocellular dorsal aspects of the nucleus. This region seems to receive its Purkinje cell projections mostly from the D2 zone. Projections also reach the ipsilateral reticular formation and have been shown to activate reticulospinal neurons monosynaptically (Bantli and Bloedel 1975; Tolbert et al. 1980). The role of a disynaptic dentate-reticulo-spinal connection is not yet clear (Arshavsky et al. 1986).

Vestibulospinal Tracts Classically, the vestibular nuclear complex is divided into medial (MV), spinal (SpV), lateral (LV), and superior vestibular nuclei (SuV). In addition, several subgroups have been recognized in various animals (i.e., groups F, L, X, Y, and Z) (Brodal 1974; Highstein and Holstein 2006). Vestibulospinal tracts are divided into a medial vestibulospinal tract, which descends by way of the mlf, originates mainly from the MV, and reaches to cervical levels where its fibers terminate bilaterally; and a lateral vestibulospinal tract, which runs lateral to the mlf, originates from the LV and descends and terminates mostly ipsilaterally throughout the length of the spinal cord (Brodal 1974; Holstege and Kuypers 1982; Nyberg-Hansen 1964b; Highstein and Holstein 2006). Later studies in various mammals have indicated that the vestibulospinal organization may be more complex (Leong et al. 1984; Masson et al. 1991; Peterson and Coulter 1977) and that neurons from all major vestibular nuclei can contribute to vestibulospinal projections (for review, see Rubertone et al. 1995). In addition, apart from their termination in the ventromedial part of the intermediate zone of the spinal cord, projections to the dorsal horn have been described (Bankoul and Neuhuber 1992). Moreover, the medial vestibulospinal tract may carry both inhibitory and excitatory fibers whereas the LV neurons that contribute to the lateral vestibulospinal tract are excitatory.

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In addition to the vestibulospinal tracts, projections from especially the MV and SV are directed to the oculomotor centers where they can have excitatory or inhibitory influences. Some neurons, mostly located in the rostral, magnocellular part of the MV, collateralize to both the oculomotor centers as well as the cervical cord (Highstein and Holstein 2006; Ruigrok et al. 1995). Finally, vestibular connections to autonomic brainstem systems that may regulate cardiovascular responses are known (Balaban and Porter 1998; Highstein and Holstein 2006). The vestibular nuclei are intimately connected with the cerebellum. Not only is a large part of its output directed to the cerebellar cortex and nuclei, but it also receives a main input from the cerebellum. The cerebellar input to the vestibular complex is special because it is the only brainstem system that receives afferents from the cerebellar nuclei as well as directly from the cerebellar cortex.

Cerebellar Corticovestibular Projections Direct projections from the cerebellar cortex to the vestibular nuclei arise from the vermis (lateral A and B zones), from the caudal vermis (ventral uvula and nodulus), and from the flocculus and adjacent ventral paraflocculus (Voogd et al. 1996). The projections from the Purkinje cells of the B zone, which is mostly located in lobules I–VIa but also has a component in lobule VIII, to the neurons of the LV constitute the most direct influence of the cerebellar cortex on a descending pathway (Ito and Yoshida 1966). The B-zone-lateral vestibulospinal connection is mostly involved in the control of extensor or anti-gravity musculature (Arshavsky et al. 1986). Purkinje cell projections from the nodulus and ventral uvula can be found throughout most of the vestibular complex with the exception of the LV (Bernard 1987; Wylie et al. 1994). This region is implicated in the control of head movement. Floccular projections to the vestibular nuclei seem to terminate in a more orderly fashion. Purkinje cells that show modulation in their firing frequency upon visual stimulation around a vertical axis project predominantly to the magnocellular part of the MV, whereas Purkinje cells modulated by visual stimulation around a horizontal axis project to SuV and the Y-group (De Zeeuw et al. 1994; Schonewille et al. 2006; Tan et al. 1995; Balaban et al. 2000). Floccular function is predominantly attributed to the control of the vestibulo-ocular and optokinetic reflexes and its coordination with pursuit movements (Ilg and Thier 2008; Voogd and Barmack 2006). The Purkinje cells of the flocculus target a specific population of the MV, termed floccular target neurons, which have physiological characteristics resembling cerebellar projections neurons (Sekirnjak et al. 2003).

Cerebellar Nucleo-Vestibular Connections More indirect control of vestibular function is also induced by the Purkinje cells of the A zone by way of the MCN. Apart from the reticular connections mentioned above, prominent, usually bilateral, projections to virtually all components

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of the vestibular nuclei are found (Teune et al. 2000), which mostly, but not exclusively, originate from the rostral part of the MCN (Fig. 23.2, Table 23.1). In the mouse ipsilateral MCN projections, like the projections to the reticular formation, have been shown to be glycinergic, whereas contralateral vestibular projections are glutamergic (Bagnall et al. 2009). It is not known if cortical and MCN projections converge on the same vestibular regions or neurons. Sparse vestibular projections from the other cerebellar nuclei – the ICG, AIN, DLH, and LCN – are mostly ipsilateral (Delfini et al. 2000; Teune et al. 2000). These connections suggest that cerebellar processes are widely integrated with vestibular functions.

Rubrospinal Tract In most mammals the red nucleus (RN) can be recognized as a prominent rotund structure located centrally in the rostral part of the mesencephalic tegmentum. Usually, a distinction is made between a caudal magnocellular part (RNm) and a rostral parvicellular part (RNp). It should be recognized that this distinction relates to some extent to the differences in projections of both regions. The magnocellular part is the origin of the crossed rubrospinal tract. This part can be identified in the mesencephalon of most limb-carrying terrestrial vertebrates (ten Donkelaar 1988). In primates and carnivores the parvicellular part is associated mostly with the uncrossed projection to the inferior olive by way of the central tegmental tract. Recently, Miller and Gibson (2009) have proposed the additional term of parvicellular contralateral red nucleus (RNpc) to identify small- and medium-sized neurons that also project to the contralateral spinal cord or the contralateral brainstem (Pong et al. 2008). Thus, the axons of RNm and RNpc neurons cross at the level of the red nucleus, project mostly contralaterally, and basically differ from each other only with respect to the distance they have to travel to their respective main projection region, that is, caudal cord versus rostral cord or brainstem. On the other hand, RNp neurons, by definition, project to the ipsilateral olive by way of the central tegmental tract. The rodent red nucleus as defined by the atlas of Paxinos and Watson (1998) consists essentially only of RNm and RNpc neurons (Ruigrok 2004; Rutherford et al. 1984). In many other species, such as cat, macaque, and human, a “real” RNp has been identified (for review see Onodera and Hicks 2009). In addition, in these animals, as well as in the rodent, immediately rostral, medial, and dorsal to the RN several areas are located that basically surround the retroflex fascicle and carry names such as the nucleus of Bechterew, nucleus of Darkschewitsch, interstitial and rostral interstitial nucleus of the medial longitudinal fascicle, subparafascicular nucleus, and prerubral field. Homologies between various names that are presently in use are difficult to establish (e.g., see Onodera and Hicks 2009; Ruigrok 2004), but these nuclei mostly have in common the fact that they contain small neurons which project massively to the inferior olive. For this reason they have been collectively named as nucleus parafascicularis prerubralis in the rat (Carlton et al. 1982; Ruigrok 2004). Onodera and Hicks (2009)

510

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have provided evidence that the projection from this mesodiencephalic area to the inferior olive is topographically organized, and that some species differences may exist in whether these projections course by way of either the medial or central tegmental tracts. Rubrospinal fibers terminate mostly within the intermediate zone of the spinal cord; however, terminals are also found at cervical levels in lamina IX of monkey, cat, and rat (Miller and Gibson 2009) where they may sparsely terminate on motoneurons that innervate distal muscles (Al-Izki et al. 2008; Kuchler et al. 2002). Many authors have mentioned that the rubrospinal tract diminishes with the expansion of the corticospinal tract. In man, it is questioned if the rubrospinal tract, which may consist of only several hundreds of fibers, descends beyond the cervical level (for review see Onodera and Hicks 2009). The decrease in size of the rubrospinal tract is countered by an increase in the importance of the RNp and adjacent regions to the inferior olive. It has been speculated that the relative increase in the projection from the mesodiencephalic junction nuclei to the inferior olive or to specific parts thereof mimics the increased control of cerebellar and cortical structures over specific motor functions and, in man, may have enabled the advent of bipedalism and speech (Onodera and Hicks 1999, 2009). The function of the rubrospinal tract is still under debate. Selective lesion of this system produces only mild deficits in motor control. In cat and monkey, this affects grasping behavior and causes dragging of the dorsum of the paw during locomotion (Horn et al. 2002; Miller and Gibson 2009). In rat, mild changes in locomotion pattern have been described (Muir and Whishaw 2000; Whishaw et al. 1998). RNm stimulation seems to affect mostly distal extensor muscles in cat (Horn et al. 2002), whereas most RNm units display up-modulation during the swing phase in stepping decerebrate cats (Arshavsky et al. 1986, 1988).

Cerebellar Projections to the Red Nucleus The RNm is heavily targeted by the AIN (Fig. 23.3a, f–h, Table 23.1), fibers of which terminate in a somatotopic fashion (Daniel et al. 1987; Ruigrok 2004; Teune et al. 2000) on the somata and proximal dendrites of rubrospinal neurons (Ralston 1994). Although small changes may exist between different mammalian species it appears that the medial or mediorostral part of the AIN projects to the RN part that sends projections to the lumbar cord whereas the (caudo-)lateral-most part of the AIN targets RN regions that target the contralateral brain stem and upper cervical cord (Conde 1988; Daniel et al. 1987; Stanton 1980). The medial aspect of the PIN projects to the medial margin of the RNm in rat (Fig. 23.3a, e; Daniel et al. 1987) and cat (Robinson et al. 1987). This projection is less dense compared to the input derived from the AIN. RNm projections originating from the MCN and LCN are sparse or nonexistent; the latter mostly targeting regions immediately lateral and rostral to the RNm (Fig. 23.3).

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Tectospinal Tract The superior colliculus is associated mostly with visual input and is involved in directing gaze to objects of interest. This is performed by saccades, head movements, or a combination of both. The control of the superior colliculus over the paramedian pontine reticular formation from where saccades are initiated is well established in most mammals (Enderle 2002). However, in addition, arm or even whole body movements may be evoked by stimulation of the superior colliculus. These body movements are triggered by tectoreticular and tectospinal pathways (Harting 1977; Nyberg-Hansen 1964a; Rose and Abrahams 1978). The tectospinal component seems to be small in most mammals with the largest component to be found in carnivores (Meredith et al. 2001; Murray and Coulter 1982; Nudo and Masterton 1989). Most tectospinal neurons do not project beyond the cervical segments from where the neck muscles are innervated (Nudo and Masterton 1989). Most axons arise from the caudolateral quadrant and after decussating in the dorsal tegmental decussation descend contralaterally close to the mlf where they form the predorsal bundle. In cat, a small ipsilateral projecting group of tectospinal fibers has been described (Olivier et al. 1994). Although tectospinal cells may terminate directly on cervical motoneurons (Olivier et al. 1995), conclusive anatomical proof has not yet been obtained (Muto et al. 1996; Shinoda et al. 2006). Therefore, the most direct impact of superior colliculus on cervical neck motoneurons seems to involve at least a segmental or reticular relay (Kakei et al. 1994; Shinoda et al. 2006).

Cerebellar Nucleo-Tectal Connections The cerebellar nuclei have a profound and often neglected impact on the mesencephalic tectum. Cerebellar projections to this region mostly terminate in the intermediate and deep layers of the contralateral superior colliculus (Fig. 23.3, Table 23.1). In various animals they have consistently been shown to originate predominantly from two areas of the cerebellar nuclei. The lateral aspect of the PIN and adjacent area of the LCN supply the bulk of cerebellotectal projections (Hirai et al. 1982; Kurimoto et al. 1995; May and Hall 1986; May et al. 1990; Uchida et al. 1983). Both regions mediate information from the paraflocculus, which is known to receive mostly extrastriate visual information by way of the dorsolateral basal pons in rat, cat, and monkey (Gayer and Faull 1988; Glickstein et al. 1994; Robinson et al. 1984; Kralj-Hans et al. 2007). In the macaque, neurons in the lateral PIN and LCN have been shown to connect disynaptically to visually related regions in the posterior parietal cortex (Prevosto et al. 2010). Projections to the superior colliculus also arise from the caudal, oculomotor, half of the MCN (Fig. 23.3a, b). This part mostly receives cortical input from the oculomotor vermis (caudal lobule VI and lobule VII) and has been suggested to be specifically involved with the control and adaptation of saccades (Thier et al. 2002; Noda and Fujikado 1987; Fujikado and Noda 1987; Takagi et al. 1998). Interestingly, there are no projections from the MCN to the superior colliculus in the rabbit, and only sparse

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Fig. 23.3 Serial plots (b–k) of microscopical sections depicting patterns of labeled varicosities (i.e., terminal arborizations) in the rostral brainstem and mesodiencephalic junction after various injections (shown in a) with an anterograde tracer in the cerebellar nuclei of the rat (Adapted from Teune et al. 2000)

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projections in the gray squirrel (Uchida et al. 1983; May and Hall 1986), findings that seems to correlate well with the observation that the rabbit superior colliculus does not seem to be essential for generating either saccades or optokinetic nystagmus (Collewijn 1975). The lateral aspect of the AIN has also been reported to provide input to the superior colliculus in the rat and rabbit (Kurimoto et al. 1995; Uchida et al. 1983). In the cat and monkey, the MCN projections have been shown to terminate bilaterally in the superior colliculus, usually somewhat more superficially compared to the PIN projections which terminate strictly contralaterally and deeper in the intermediate layer (Kawamura et al. 1982; May et al. 1990). Both types of projections seem to make use of a different type of synapse as judged from their respective degeneration characteristics (Warton et al. 1983), and it has been suggested that they play different roles in the control of saccades. A clear topographic pattern, however, has not been described in a variety of mammals (Hirai et al. 1982; Kawamura et al. 1982; Kurimoto et al. 1995; May and Hall 1986; May et al. 1990; Noda et al. 1990; Uchida et al. 1983; Teune et al. 2000). The ventrolateral aspect of the caudal superior colliculus reportedly receives the densest cerebellar input (Fig. 23.3). However, cerebellar terminals are also found in tectal areas that contain large tectospinal neurons. Presently, no specific anatomical information seems to be available that positively identifies the collicular targets of the cerebellar output (Warton et al. 1983). Although GABAergic projections to the pretectum have been reported to arise from the lateral PIN and ventral LCN in the cat (Nakamura et al. 2006), no evidence for cerebellar inhibitory synapses in the superior colliculus has been found (Warton et al. 1983).

Interstitiospinal Tract The interstitiospinal tract originates from scattered large neurons located dorsomedial to the red nucleus and lateral to the oculomotor nuclei and periaqueductal gray within and surrounding the mlf. This poorly defined region is known as the interstitial nucleus of Cajal (INC). Fibers descend ipsilaterally by way of the mlf to the spinal cord where they terminate in laminae VII and VIII of the spinal gray. The interstitiospinal tract has excitatory monosynaptic contacts with neck musculature but also provides di- and polysynaptic mostly excitatory activation of back, fore- and hindlimb muscles (Fukushima et al. 1978; Holstege and Cowie 1989). The INC, apart from acting as origin of the interstitiospinal tract is also involved in the control of eye movements. Neurons participating in this function appear to form a different population from the cells that give rise to the interstitiospinal tract (Bianchi and Gioia 1995; Zuk et al. 1983). Projections from the INC also reach the oculomotor nuclei, the pontine reticular formation, and the vestibular nuclei. As such it plays an important role coordinating eye and head movements. The area directly rostral the INC, known as the rostral interstitial nucleus of the mlf and containing many parvalbumin-containing neurons, provides input to the INC and is specifically involved in the control of vertical eye movements (Buttner-Ennever 2006; Fukushima-Kudo et al. 1987; Fukushima 1991; Horn and Buttner-Ennever 1998; Tolbert et al. 1978b).

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Cerebellar Projections to the Interstitial Nucleus of Cajal The most prominent cerebellar projections to the INC arise from the oculomotor part of the MCN in rat (Fig. 23.3b), cat, and monkey (Noda et al. 1990; Teune et al. 2000; Sugimoto et al. 1982). However, it also receives an afferent contribution from other parts of the cerebellar nuclei (Table 23.1) (Chan-Palay 1977; Teune et al. 2000).

Cerebellar Projections to Other Areas Apart from the cerebellar projections to the five classic descending premotor tracts, all cerebellar nuclei also provide input to various regions of thalamus (Aumann et al. 1994; Chan-Palay 1977; Teune et al. 2000), which will not be reviewed here (but see Table 23.1). Also, many cerebellar nuclear neurons have been shown to provide input to the cerebellar cortex (Provini et al. 1998; Batini et al. 1992; Buisseret-Delmas and Angaut 1988; Tolbert et al. 1978b) some of which may be inhibitory (Uusisaari and Knopfel 2010; Batini et al. 1989). In addition, the cerebellar nuclei provide input to a number of precerebellar nuclei in the brainstem. Cerebellar nuclear inputs to the reticulotegmental nucleus of the pons and the basal pontine nuclei are well-known and have been shown to participate in potential functionally important reverberating circuitry involving cerebello-bulbo-cerebellar loops (Mock et al. 2006; Tsukahara et al. 1983). However, projections to nuclei that give rise to the descending motor tracts may also provide feedback to the cerebellum, that is, the vestibular nuclei and the reticular formation are an important source of mossy fiber input to the cerebellum. In addition, many rubrospinal fibers have been demonstrated to collateralize specifically to the AIN (Huisman et al. 1983). A more prominent connection to a precerebellar nucleus is formed by the cerebellar nuclear projections to the inferior olive. This connection has been shown to precisely match the collateral projection of the cerebellar climbing fibers to the cerebellar nuclei (Ruigrok and Voogd 1990, 2000), and to consist exclusively of GABAergic fibers (De Zeeuw et al. 1989; Fredette and Mugnaini 1991) that originate from a population of small neurons intermingled with the larger and generally excitatory projection neurons and to possess their own physiological characteristics (Fredette and Mugnaini 1991; Uusisaari et al. 2007). The population of small GABAergic neurons most likely exclusively targets the inferior olive (Teune et al. 1995; De Zeeuw and Ruigrok 1994). Another major target of cerebellar nuclear projections is formed by the nuclei that supply a major input to the inferior olive. This is especially true for the nuclei in the mesodiencephalic junction that participate in this cerebellar nucleomesodiencephalo-olivocerebellar loop (De Zeeuw and Ruigrok 1994; Ruigrok and Voogd 1995). In man, the main circuit is formed by the projections from the dentate nucleus to the parvicellular RN which, by way of the central tegmental tract, targets the principal olive. Although the function of this circuit is far from clear (Ruigrok 1997; Ruigrok and Voogd 1995; Hoebeek et al. 2010; Kennedy 1990), this so-called triangle of Guillain-Mollaret (Guillain et al. 1933) has received attention as lesions of this circuit induce myoclonus of the palatal muscles in particular (sometimes also

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involving pharyngeal, laryngeal, diaphragm, or extraocular muscles), and may result in conspicuous enlargement of particular parts of the inferior olive (Gautier and Blackwood 1961; Boesten and Voogd 1985; Ruigrok et al. 1990; Shaikh et al. 2010). Finally, cerebellar nuclear connections have been described that do not fall within any of the categories mentioned above (Table 23.1). Examples are formed by the cerebellar projections to hypothalamic areas, zona incerta, parvicellular reticular formation, parabrachial nuclei, and periaqueductal gray. (Teune et al. 2000; Zhu and Wang 2008; Haines et al. 1997). These connections suggest a wide impact of cerebellar processing involving various autonomic as well as pain functions.

Divergence of Cerebellar Projections The wide array of cerebellar targets described with anterograde techniques, even after small injections, suggests that individual nucleo-bulbar fibers may collateralize to multiple brainstem and diencephalic areas (Chan-Palay 1977; Teune et al. 2000). A survey of cerebellar nuclear targets based on a study with anterograde tracers in the rat is provided in Table 23.1 (based on Teune et al. 2000). A systematic survey of collateralization based on reconstructions of individual fibers has not been performed but based on available partial reconstructions (Shinoda et al. 1988), double retrograde tracing (Bentivoglio and Kuypers 1982; Lee et al. 1989; Gonzalo-Ruiz and Leichnetz 1987), and electrophysiological data (Tolbert et al. 1978a; Bharos et al. 1981), individual nucleo-bulbar neurons, as a rule, seem to be able to influence multiple areas simultaneously. Indeed, from Table 23.1 it can be seen that relatively small injections result in labeled varicosities within at least ten (and usually considerably more) different areas. The only exception is formed by the nucleo-olivary projection neurons, which, as mentioned above, constitute a separate population. From the available anterograde and retrograde tracing data it can be deduced that individual projection neurons in the cerebellar nuclei are likely to be involved in the control of several motor pathways. For example, a neuron in the AIN projecting to the magnocellular red nucleus may also influence, by way of ongoing projections to the ventrolateral and ventro-anterior region of the thalamus, motor output by way of the corticospinal tract. Simultaneously, activity may be fed back to the cerebellum by way of direct nuclear projections to the basal pontine nuclei, by way of rubrocerebellar collaterals of the rubrospinal tract, or by way of the cortico-ponto-cerebellar route. It will be obvious that a more elaborate knowledge on the circuitry involving individual neurons will be necessary in order to be able to evaluate the functional processing within these multiple circuits.

Convergence of Cerebellar Projections In contrast to the apparent divergence of cerebellar nuclear connections, a considerable convergence of cerebellar output to specific muscles has also been noted. Injections with the retrogradely and transneuronally transported rabies virus

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into several muscles have resulted in viral labeling of several locations within the cerebellar nuclei and cerebellar cortex (Graf et al. 2002; Morcuende et al. 2002; Ruigrok et al. 2008; Tang et al. 1999). Ruigrok et al. (2008) showed that viral injections in antagonistic muscles of hind and forelimb result in infection of several longitudinal strips of Purkinje cells which were partly overlapping (Fig. 23.4). Initial strips were observed in the vermis, but, with only slightly longer survival times, additional strips were found in paravermal and hemispheral parts of the cerebellum (Fig. 23.4). This suggests that multiple cerebellar modules, by way of their individual nuclear output, ultimately can converge to control the activity of the same muscle. Simultaneously, however, the overlap of labeled strips suggests that a single module, by way of its nuclear output, can influence the activity of several, and even antagonistic, muscles or muscle groups. It is clear that the latter characteristic, at least partly, is also based on the distributed nature of the termination of individual fibers such as those in the reticulospinal and rubrospinal tracts (Shinoda et al. 1977, 2006).

Functional Implications It has been suggested that the modular arrangement of cerebellar circuitry, characterized by strips of Purkinje cells converging upon a region within the cerebellar nuclei that is matched by the organization of the olivocerebellar climbing fiber system, lies at the basis of the functional working blocks of the cerebellum (Apps and Garwicz 2005; Ruigrok 2011). As is evident from the account sketched above, however, it is by no means clear how individual cerebellar modules interact with brainstem structures to result in functionally meaningful signals. Individual modules, for example, by way of their collateralizing output, are capable of simultaneously influencing numerous brainstem structures including several nuclei that have descending connections to the spinal cord. On the other hand, as shown with viral tracing techniques, multiple modules may participate in the control of the same muscle. This particular organization suggests that each cerebellar module may serve a specific function in the control of muscles. The interaction of multiple strips of Purkinje cells with several target areas has most clearly been demonstrated in the relatively simple system of floccular control on reflexive eye movements (van der Steen et al. 1994; Voogd and Wylie 2004; Ruigrok et al. 1992; Sugihara et al. 2004). However, more recent studies that attempted to study the contribution of individual modules to skeletomotor function suggest that a single module may affect a particular type of muscle control (Horn et al. 2010; Cerminara and Apps 2011; Pijpers et al. 2008); impairment of the C1 hind limb module influences the phaselocked modulation of reflexes during locomotion (Pijpers et al. 2008). Inactivation or lesion of various olivary regions, resulting in inactivation or severe functional impairment of related cerebellar nuclear areas, also results in highly specific deficits in motor control (Horn et al. 2010; Cerminara and Apps 2011). It therefore seems that different cerebellar modules have the potential to participate in the control of the same muscles that are used in different functional contexts.

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Fig. 23.4 Demonstration of involvement of multiple cerebellar modules in the control of single muscles using transneuronal retrograde transport of rabies virus in rat. a1: Pattern of Purkinje cell labeling in the lateral vermis 5 days after rabies injection in the ipsilateral gastrocnemius muscle. a2: 3-D reconstruction of the anterior lobe of this case indicating rabies labeled Purkinje cells (red) together with zebrin II labeled Purkinje cells (yellow). Note that the zebrin II labeled bands identified as p1–p6 can all be recognized. The position of the rabies labeled Purkinje cells between p2 and p3 (left hand white arrowhead) is identical to that of the B zone. In addition a contralateral strip of labeled Purkinje cells is noted just medial to the p2 zebrin II band (right hand white arrowhead), which corresponds to the location of the lateral A1 zone. b1,2: Similar to a1,2 after injection of rabies virus in the ipsilateral anterior tibial muscle. In this case, both the B and lateral A1 zone (arrowhead) are noted ipsilateral to the injection. c: Pattern of infection 6 days after injection of the gastrocnemius muscle. Note that the original zones can still be recognized but also have a mirror representation in the other cerebellar hemisphere. However, several additional zones are also recognized (arrowhead) in the paravermis and hemisphere (not shown). d: Similar to c for injection of the anterior tibial muscle (Modified from Ruigrok et al. 2008 and Ruigrok 2011)

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Clinical Implications It follows from this account that the impact of cerebellar malfunction may be diverse and, not infrequently, difficult to fully understand. Due to the distributed connectivity, focal damage to the vestibulocerebellar, vermal, paravermal, and hemispheral components of the cerebellum will result in various combinations of specific cerebellar disorders (Thach and Bastian 2004). Conversely, due to the nature of ongoing spinocerebellar degenerative disease, this may differentially influence a multitude of cerebellar modules. Likewise, cerebellar tumors or lesions almost invariably involve several adjacent modules in an incomplete way (i.e., the modules are organized in longitudinal cortical arrays which can be discontinuous and may be found in both the anterior and posterior lobes (Pijpers et al. 2006; Ruigrok 2011)). The resulting clinical syndrome, therefore, may be hard to predict or explain. Although it is now well accepted that the cerebellum may be involved in a large array of nonmotor brain functions, cerebellar disorders are usually recognized as a combination of specific motor deficits. Due to the double decussation of most cerebellofugal projections (i.e., the decussation of the superior cerebellar peduncle in the caudal mesencephalon, followed by the decussation of corticospinal and rubrospinal tracts), unilateral cerebellar deficits usually involve affected movements of the ipsilateral side of the body. Cerebellar-based handicaps in motor control commonly involve three symptoms: muscle weakness (hypotonia), ataxia, and tremor (Thach and Bastian 2004).

Hypotonia Sudden removal of cerebellar nuclear output in cases such as hemorrhage, ischemic infarction, or surgical excision, diminish the tonic excitatory drive to premotor regions. This is likely to result in a general reduction of activation of the related motoneuron pools, which will be reflected by a reduced muscle tone (hypotonia). Similarly, however, cerebellar lesions that do not involve the nuclei are likely to diminish the inhibitory drive of the cortical Purkinje cells on the cerebellar nuclei, thereby increasing nuclear output resulting in hypertonic responses. Hypotonia and hypertonia may (partly) disappear after a period of weeks or months, probably by adjustment of the tonic drive to premotor regions by other, unaffected, cerebellar or extracerebellar brain regions.

Ataxia Although ataxia is typical for cerebellar disorders, it is not easily characterized. It usually reflects the inability of a patient to execute voluntary movements in a “normal” way. Movements or parts thereof are not initiated at the correct moment, not terminated at the appropriate time, and not corrected adequately. This results in improperly conducted and dysmetric movements. To some extent hypo- and

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hypertonia of muscles or muscle groups form the foundation of these motor deficits. Because antagonistic and correcting activity is inadequately timed, oscillations easily occur around the intended target. In combination with hypotonia this may result in pendular reflexes. Due to dysmetria and failure to time movements, cerebellar patients also encounter problems with the execution of fast rhythmic movements (dysdiadochokinesis). When these disorders concern the mouth, pharynx, or larynx musculature it will result in cerebellar dysarthria that typically involves changes in rhythm and amplitude in combination with a careless or “slurred” pronunciation.

Intention Tremor This form of tremor is also very characteristic in cerebellar disorders and may be related to dysfunction of properly timed components of movements, resulting in a disturbance in the beat and rhythm of movements with tremor as a consequence. This form of tremor has been associated with the specific physiological properties of olivary neurons that may result in rhythmical and simultaneous firing of neuronal ensembles (Llina´s and Pare´ 1995). The tremorgenic action of harmaline, for example, has been reported to result in rhythmic activity of olivary neurons, which, by way of their effect on the Purkinje cells of the cerebellar cortex and subsequently on the cerebellar nuclear activity, will be transformed to rhythmical activity of reticulospinal, and potentially also other descending pathways (Llina´s and Volkind 1973).

Conclusions and Future Directions The cerebellum is usually appreciated as a structure with a uniform internal structure that will perform a particular type of information processing. Within this internal structure, based on the organization of corticonuclear and olivocerebellar connections, a number of parallel, longitudinally organized, modules can be recognized which form functional entities. The organization of the input to these modules and the organization of their output channels, therefore, will determine the type of information processed within a module and which structures will be informed of its result. The above account shows that the output of a particular module is directed to many regions in brainstem and diencephalon and may affect multiple centers with descending connections to the spinal cord. In addition, several modules may ultimately affect activity patterns of the same muscle or muscle group. In order to fully understand the cerebellar involvement in motor control and learning (and in other functions) it will be imperative to understand to what extent individual modules or micromodules control different aspects of controlling muscles. In addition, a precise description of the often di- and polysynaptic pathways involved will be necessary. The advent of new neuroanatomical techniques such as selective

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(modular) lesioning, transneuronal tracing, and single axon reconstructions are expected to greatly aid in fulfilling these requirements for further understanding cerebellar function and dysfunction.

List of Abbreviations in Text and Figures 7 12 AIN Amb AP Aq BIN BPN CG cp D DLH Ecu FF fr Gi IC ICG icp INC IO IV LCN LG LRN LV MCN mcp

Facial nucleus Hypoglossal nucleus Anterior interposed nucleus Ambiguus nucleus Anterior pretectal nucleus Aquaduct Basal interstitial nucleus Basilar pontine nuclei Central gray Cerebral peduncle Nucleus of Darkschewitsch Dorsolateral hump External cuneate nucleus Fields of Forel Retroflex fascicle Gigantocellular reticular nucleus Inferior colliculus Interstitial cell groups Inferior cerebellar peduncle Interstitial nucleus of Cajal Inferior olive Fourth ventricle Lateral cerebellar nucleus Lateral geniculate nucleus Lateral reticular nucleus Lateral vestibular nucleus Medial cerebellar nucleus Medial cerebellar peduncle

Md MG ml mlf Mo5 MV n7 NRTP PAG PCRt PF PIN PnC PnO py RN RNm RNp RNpc SC scp SN SO Sp5 SpV SuV ZI

Medullary reticular nucleus Medial geniculate nucleus Medial lemniscus Medial longitudinal fascicle Motor trigeminal nucleus Medial vestibular nucleus Facial nerve Nucleus reticularis tegmenti pontis Periaquaductal gray Parvicellular reticular formation Parafascicular thalamic nucleus Posterior interposed nucleus Caudal pontine reticular formation Oral pontine reticular formation Pyramidal tract Red nucleus Magnocellular red nucleus Parvicellular red nucleus RNp neurons with contralateral projections Superior colliculus Superior cerebellar peduncle Substantia nigra Superior olive Spinal trigeminal nucleus Spinal vestibular nucleus Superior vestibular nucleus Zona incerta

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Cerebellar Thalamic and Thalamocortical Projections

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Sharleen T. Sakai

Abstract

Although it is well known that the major output of the cerebellum is directed to the thalamus and ultimately to the cerebral cortex, the anatomical details and functional organization of this system remains unclear. Here, the current status of the cytoarchitecture of the motor thalamus, its afferents and efferent cortical projections are reviewed. The distribution of the cerebellothalamic and pallidothalamic projections to motor cortical areas is also discussed and the functional importance of these motor systems is highlighted.

Introduction Thalamus as the gateway to the cerebral cortex occupies a pivotal place in the processing of incoming and outgoing signals. Over the past 20 years, interest in the organization of the motor thalamus increased due to its role in the amelioration of tremor and rigidity following either thalamotomy (Ohye and Narabayashi 1979; Tasker et al. 1982; Ohye 1997) or deep brain stimulation (Benabid et al. 1996; Hubble et al. 1997; Starr et al. 1998). Despite the importance of the motor thalamus in motor control functions, details of its anatomical organization including its afferent and efferent connections still remain to be addressed. Thalamic studies are often stymied by the difficulty in defining and clearly delineating its constituent nuclei and their borders. The lack of agreement on thalamic terminology and nomenclature has also contributed to the confusion (for review, see Percheron et al. 1996, Ilinsky and Kultas-Ilinsky 2002; Jones 2007). Thalamic nuclei can be defined based on cytoarchitecture and chemoarchitecture but the term motor

S.T. Sakai Department of Psychology and Neuroscience Program, Michigan State University, 108 Giltner Hall, East Lansing, MI, 48824, USA e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 529 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_24, # Springer Science+Business Media Dordrecht 2013

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thalamus refers to the projection territory of the basal ganglia (efferent projections of the substantia nigra and globus pallidus) and the deep cerebellar nuclei. Since these projections may not strictly adhere to nuclear boundaries, comparisons made from different experiments and based on different species are problematic. This is due, in part, to the difficulty in reliably delineating the boundaries of the motor thalamus across mammalian species since both the number and cytoarchitectural details of the constituent subnuclei vary across species (Jones 1985, 2007). Many studies compared results across different experiments and animals in concluding that the cerebellum and basal ganglia projections distributed to separate thalamic nuclei and that the thalamocortical projections to different motor cortical areas arose from separate thalamic nuclei (Schell and Strick 1984; Alexander et al. 1986; Jones 1985, 2007; Percheron et al. 1996). The primary problem with such comparisons is the uncertainty of applying the same thalamic nuclear boundaries across experiments. One way to address this uncertainty is to eliminate the need to delineate thalamic nuclear boundaries by directly evaluating overlap of the cerebellar and basal ganglia projections with thalamocortical projection neurons using multiple neuroanatomical tracers in the same animal. Here, a review of the afferent and efferent connections based on anatomical studies utilizing axonal transport techniques in primates with a particular emphasis on the comparison of the cerebellothalamic and pallidothalamic projections is presented. Results from experiments using transneuronal labeling are presented elsewhere in this volume and will not be reviewed here.

Cyto- and Chemoarchitecture of the Motor Thalamus According to the terminology of Olszewski (1952), the motor thalamus typically consists of the ventral anterior nucleus (VA), ventral lateral nucleus pars oralis (VLo), ventral posterior lateral nucleus pars oralis (VPLo), ventral lateral nucleus pars caudalis (VLc), ventral lateral nucleus pars medialis (VLm), and area X (X) in the macaque monkey. The cytoarchitectonic distinctions between these nuclei are somewhat vague and not readily agreed to by others who have proposed either more or less thalamic subnuclei (Walker 1938; Hassler 1982; Percheron et al. 1996; Ilinsky and Kultas-Ilinsky 2002; Jones 2007). An alternative approach to defining and naming thalamic nuclei was based on the distribution of afferent connections. Jones (2007) employed this criterion in proposing a terminology that might be applicable to both primate and non-primate species in his simpler nomenclature of the motor thalamus: VA and two subdivisions of VL: anterior and posterior where VLa primarily refers to the pallidothalamic territory and VLp, the cerebellothalamic territory. Some researchers have argued that a single VL should not contain both pallidal and cerebellar territories and that the nigral and pallidal thalamic projections to its primary targets should be subdivisions of a single basal ganglia–related entity such as VA (Percheron et al. 1996; Ilinsky and Kultas-Ilinsky 2002). The nomenclatures of Olszewski, Jones, and Ilinsky are compared in Table 24.1. Based on these differences, it is clear that thalamic parcellation and

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Cerebellar Thalamic and Thalamocortical Projections

Table 24.1 Motor thalamic nomenclature Olszewski (1952) VApc Jones (2007) VApc Ilinsky and Kultas-Ilinsky (2002) VApc

VLo VLa VAdc

531

VPLo VLp VL

X(VLx) VLp VL

VLc VLp VL

nomenclatures remain far from standardized. In the following, the distinctive cytoand chemoarchitectonic features of the motor thalamus using Olszewski’s (1952) nomenclature and where directly applicable, the terminology of Jones (1985, 2007) is described. The ventral lateral nucleus pars oralis (VLo) of Olszewski (1952) in the macaque monkey primarily occupies the region caudal to the VA, rostral to the ventral posterior lateral thalamus (VPL), and lateral to the internal medullary lamina. Its cytoarchitecture is diverse consisting of multiple subnuclei depending on the author (Walker 1938; Hassler 1982; Percheron et al. 1996). The VLo corresponds to the ventral lateral nucleus anterior division (VLa) of Jones (1985) and consists of small-to-medium-sized, darkly stained cells packed irregularly as seen in Nissl preparations. Cells of VLo can be distinguished from the more posterior ventral posterior lateralis nucleus pars oralis (VPLo) because the latter contains larger cells with a sparser and more homogeneous distribution. Cells of VLo are also distinct from the ventral lateral nucleus pars caudalis (VLc) in that the VLc cells are darkly stained for Nissl and smaller. A medial division of VPL (area X) was also identified by Olszewski (1952) and is characterized by the presence of small, homogeneously distributed, lightly Nissl stained cells. These subnuclei,VPLo, VLx, VLc, described by Olszewski together form the single VLp nucleus of Jones. Since the differences between these subnuclei are variants on a common theme and together these nuclei form a cerebellar projection to the motor cortex, Jones (1985) suggested the single designation VLp. The subnuclei of the motor thalamus are difficult to distinguish based on Nissl cytoarchitecture alone but differential staining within the motor thalamus has been found using a variety of histochemical and immunocytochemical stains. Acetylcholinesterase (AChE) staining greatly facilitates the comparison between VLo and VPLo (Fig. 24.1). The VLo (VLa) stains dark for AChE contrasting sharply with the lighter AChE staining in VA rostrally and moderately stained VPLo (VLp) caudally. The differential AChE staining is most apparent in the owl monkey thalamus (Stepniewska et al. 1994; Sakai et al. 2000) in comparison to the macaque monkey thalamus (Sakai et al. 2003; Jones 2007). Differential immunocytochemical reactivity to the monoclonal antibody, CAT 301, is also found where high immunoreactivity is found in VPLo (VLp) and low in VLo (VLa) in the macaque monkey thalamus (Fig. 24.1) (Hendry et al. 1988; Stepniewska et al. 2003). Subnuclei of the motor thalamus are also immunoreactive for calcium-binding proteins. Low calbindin immunoreactivity is found in VPLo and VLx whereas VLo and VLc were moderately to strongly immunoreactive (Percheron et al. 1996; Stepniewska et al. 2003; Calzavara et al. 2005; Jones 2007). Jones reported high parvalbumin immunoreactivity in VLp and moderate to weak immunoreactivity in VLa whereas

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Fig. 24.1 Low power photomicrographs of a series of coronal sections through the macaque motor thalamus. (a) Brightfield photomicrograph of a section showing cerebellothalamic wheat germ agglutinin conjugated horseradish peroxidase (WGA-HRP) labeling in VPLo and the patches of pallidal biotinylated dextran amine (BDA) labeling in VLc. Asterisks denote the same blood vessel in A-E. (b) Major cytoarchitectonic features of the motor thalamus at this thalamic level in a cresyl violet stained section. (c) Darkfield photomicrograph of the same section shown in A. Patchy cerebellar and pallidal labeling in VLc interdigitate. (d) Acetylthiochoinesterase (AChE) chemoarchitecture stained section. (e) The adjacent section immunoreacted for CAT 301. Note that VPLo is immunopositive for CAT 301 (Modified from Sakai et al. (2003). Ascending inputs to the pre-supplementary motor area in the macaque monkey: cerebello- and pallido-thalamocortical projections. Thalamus Related Syst 2: 175–187 with permission)

Calzavara et al. (2005) found that parvalbumin immunoreactivity in VLo with dense and patchy immunoreactivity was found in VLx, VPLo, and VLc. The patchy immunoreactivity of dense and light staining, particularly in medial VLx, VLc and along the VLo and VPLo border lead Calzavara et al. (2005) to suggest that

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533

parvalbumin immunoreactivity is of limited value in delineating the motor thalamic subnuclei. In summary, cytoarchitectonic criteria combined with either AChE, CAT 301 or calbindin immunoreactivity enhance delineation of the motor thalamus, particularly the distinction between VLo and VPLo.

Afferents of the Motor Thalamus The motor thalamus can be defined based on its afferent projections from the cerebellum and globus pallidus. Many studies have examined the cerebellothalamic distribution in the macaque monkeys using silver degeneration methods following cerebellar lesions and anterograde axonal transport techniques (Kusama et al. 1971; Mehler 1971; Chan-Palay 1977; Stanton 1980; Kalil 1981; Asanuma et al. 1983a, b; Ilinsky and Kultas-Ilinsky 1987). There is general agreement that the cerebellothalamic projections distribute in a lamella-like arrangement composed of rod-like zones of axonal terminations from the dentate nucleus and more diffuse and focal terminations from interpositus and fastigial nuclei (Kalil 1981; Asanuma et al. 1983b; Mason et al. 2000). These terminal projections arise from the contralateral dentate and interpositus nuclei and the fastigial nuclei bilaterally (Stanton 1980; Kalil 1981; Asanuma et al. 1983a, b; Rouiller et al. 1994; Sakai et al. 1996). Both the dentate and interpositus nuclei densely project to contralateral thalamus while the projections from fastigial nucleus are sparser in comparison (Kalil 1981; Asanuma et al. 1983a, b). The cerebellar nuclei project to the motor thalamus in a topographic manner whereby anterior regions of the cerebellar nuclei primarily project to lateral motor thalamus and posterior parts of the cerebellar nuclei preferentially project to medial motor thalamus (Stanton 1980; Kalil 1981; Asanuma et al. 1983a). The dentate and interpositus nuclei give rise to fibers that project to overlapping thalamic domains but it is not currently known if these inputs converge onto the same thalamic neurons in the monkey. Although a dorsoventral topography from the dentate nucleus has been described (Middleton and Strick 1997), an analysis of the fibers from the ventral dentate has shown that they distribute throughout the VL region (VLp of Jones) (Mason et al. 2000). It has been suggested that each cerebellar nucleus contains a somatotopic body representation (Asanuma et al. 1983a; Middleton and Strick 1997; Jones 2007). Recent evidence suggests that a point to point somatotopy arising from each cerebellar nucleus to the motor thalamus may not completely characterize these projections. Based on small injections of retrograde tracers into the motor thalamus of the macaque monkey following electrophysiological identification of the face, forelimb, or hind limb representation, Evrard and Craig (2008) suggest that the cerebellar projections can be more aptly described as somatotopographic reflecting their finding that these projections both diverge and converge within the thalamus in a pattern that includes limited foci as well as broadly dispersed patches. This pattern of focal and widely distributed axonal fields was also reported using biotinylated dextran amine (BDA) labeling of the cerebellothalamic axons (Mason et al. 2000).

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These data suggest that the afferent information arising from the cerebellar nuclei include both detailed somatotopically organized information as well as more generalized topographical information. Taken together, this anatomical distribution may best reflect the information processing required in order to produce coordinated multi-joint movements (Evrard and Craig 2008). The majority of cerebellothalamic fibers cross midline at the brachium conjunctivum and travel anteriorly to the diencephalon. Dense bundles of cerebellar fibers turn dorsally coursing through the fields of Fo´rel in approaching caudal thalamus. At this level, a contingent of fibers continues dorsally through the zona incerta, traversing the ventral posterior inferior nucleus (VPI) and traveling on to caudal intralaminar nuclei and the mediodorsal (MD) nucleus. The main bundle of cerebellar fibers courses rostrally to the external medullary laminae to successively disperse at multiple caudorostral levels to the motor thalamus including VPLo, VLx, and VLc (Stanton 1980; Kalil 1981; Asanuma et al. 1983a; Mason et al. 2000). As noted earlier, analysis of cerebellothalamic terminals reveal two primary types of fibers: fibers with focal terminal fields and those with dispersed terminal fields (Mason et al. 2000). A single cerebellothalamic axon may emit several long branches with individual terminal fields consisting of clusters of elongated discs (Kalil 1981; Mason et al. 2000). An axon could give rise to as many as 29 terminal fields and close to 300 terminal boutons are associated with a single cerebellothalamic axon (Mason et al. 1996, 2000). Cerebellothalamic terminals are large, filled with round vesicles, and make asymmetrical contacts onto dendrites of thalamocortical projection neurons or interneurons (Harding and Powell 1977; Kultas-Ilinsky and Ilinsky 1991; Mason et al. 1996; Ilinsky and Kultas-Ilinsky 2002). The putative neurotransmitter is glutamate. Although no direct evidence is available in primates, a recent study reported VGlut2 immunoreactivity associated with the cerebellothalamic projections in the rat (Kuramoto et al. 2011). There is some disagreement as to the extent of cerebellar projections to VLo with some investigators reporting this projection (Kusama et al. 1971; Mehler 1971; Chan-Palay 1977; Stanton 1980; Kalil 1981) and others denying it (Percheron 1996; Asanuma et al. 1983a; Ilinsky and Kultas-Ilinsky 1987; Jones 2007). If cerebellothalamic projections distribute to rostral motor thalamus including VLo as suggested by single tracing studies, then the possibility remains that thalamus receives overlapping and possibly converging inputs from the globus pallidus and cerebellum. A direct assessment of this question was made by using two anterograde tracers, one injected into the globus pallidus and the other injected into the deep cerebellar nuclei. The distribution of the axonal labeling emanating from each source is then directly compared (Rouiller et al. 1994; Sakai et al. 1996) (Fig. 24.2). In general, the pallidothalamic projections distribute broadly throughout VLo with small patchy foci found rostrally in the ventral anterior nucleus pars principalis (VApc) and VLc. Cerebellothalamic territory extends anteriorly beyond the cellsparse zones of VPLo, VLx, and VLc. The double labeling method revealed some interdigitation of pallidothalamic and cerebellothalamic labeling in VLo, VLc, VLx, and VPLo (Rouiller et al. 1994; Sakai et al. 1996). These small interdigitating patches of pallidal and cerebellar projections are limited and occur preferentially

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535

VLc VLc

VLo

CI X VPLo VPLo CM

ZI R VLc

VLc

CI

CI

VLo

MD

X VPLo

X

WGA-HRP CM

VLm

63

MD

VPLo

Pc VLo ZI

87

VPI ZI

BDA

Fig. 24.2 Line drawings of coronal sections showing the distribution of cerebellothalamic (black) and pallidothalamic projections(red) in motor thalamus. Cerebellar labeling is a result of WGAHRP injections into the contralateral cerebellar nuclei and BDA injections were made into the internal segment of the globus pallidus (GPi). Major cytoarchitectonic delineations are shown for each thalamic level in the corner insets. In section 63, the cerebellar labeling is dense and patchy in VPLo and VLx and pallidal labeling is present in VLo and VLc. More posteriorly in thalamus as seen in section 87, the cerebellothalamic projections are prominent in VPLo, VLx and VLc while pallidal projections decline in VLc (From Sakai et al. (1996) Comparison of cerebellothalamic and pallidothalamic projections in the monkey (Macaca fuscata): a double anterograde labeling study. J Comp Neurol 368: 215–228 with permission)

along border zones between nuclei. Although zones of interdigitating inputs were observed in close apposition to the proximal dendrites and soma of the same neuron, this was very rare. Based on electrophysiological evidence, it is unlikely that single thalamic neurons receive converging inputs from the cerebellum and globus pallidus (Yamamoto et al. 1984; Nambu et al. 1988, 1991; Anderson and Turner 1991; Jinnai et al. 1993). While it seems clear that the projections arising from the globus pallidus and those arising from the cerebellar nuclei primarily distribute to separate thalamic territories, individual thalamic nuclei receive differentially weighted inputs from these sources (Rouiller et al. 1994; Sakai et al. 1996). One explanation of these findings is that finger-like cell groupings characteristic of VPLo extend rostrally and irregularly into VLo (Asanuma et al. 1983a; Calzavara et al. 2005; Jones 2007) and may account for some of the discrepancies in the reports of the cerebellothalamic distribution. At the same time, the small foci of cerebellar labeling observed in the most rostral motor thalamus seem unlikely to be rostral

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extensions of VPLo (Rouiller et al. 1994; Sakai et al. 1996). Nonetheless, areas of overlapping pallidal and cerebellar projections were rare (Sakai et al. 1996). Finally, it should be noted that VLc is a nucleus that receives a patchy and complementary pattern of labeling (Rouiller et al. 1994; Sakai et al. 1996). In this regard, it is of interest that a direct correspondence between the cerebellothalamic territory and negative calbindin immunoreactivity is found in VPLo and much of VLx (Calzavara et al. 2005). These authors found a complementary pattern of patchy cerebellar projections and areas of calbindin-poor immunoreactivity in VLc. These results suggest that calbindin immunohistochemistry may be helpful in delineating the cerebellar territory without regard to the constraints imposed by cytoarchitectonic analysis (Calzavara et al. 2005). Although the bulk of the cerebellothalamic projections target the motor thalamus, cerebellar axons also distribute to MD and the intralaminar nuclei. Dense bundles of cerebellar fibers ascend through the fields of Forel, pass through the centrum medianum (CM), and distribute to the central lateral (CL) and to the paralamellar portion of MD (Asanuma et al. 1983a, b; Rouiller et al. 1994; Percheron et al. 1996; Sakai et al. 1996; Mason et al. 2000). The distribution of cerebellothalamic fibers arising specifically from the ventral dentate nucleus to MD is quite limited, and very little labeling is noted in MD other than its most lateral paralamellar portion (Mason et al. 2000).

Motor Thalamic Projections to Cortex The primary target of the motor thalamus is cortex lying anterior to the central sulcus. The general topography of the thalamocortical projections has been known for some years (Kievit and Kuypers 1977), but the extent of divergence and convergence of the thalamocortical projections as well as whether cerebellum or globus pallidus is a source of afferents to those projections is not completely known. The following will review the source of thalamocortical projections to the motor and premotor areas in nonhuman primates.

Projections to MI It has long been known that the primary cortical projection of the cerebellothalamic projections is to the primary motor cortex (MI) (for review, see Jones 2007) (Fig. 24.3). Typically, studies use microstimulation to map the body representation within MI (cytoarchitectonic area 4) in order to identify the sites for retrograde axonal tracer injections. These studies report that thalamic projections from VPLo, VLx, and VLc project to the primary motor cortex (Schell and Strick 1984; Wiesendanger and Wiesendanger 1985; Matelli et al. 1989; Darian-Smith et al. 1990; Tokuno and Tanji 1993; Rouiller et al. 1994; Morel et al. 2005). Moreover, when these experiments are combined with anterograde tracer injections into the cerebellar nuclei, the cerebellothalamic pathway is directly demonstrated revealing that the cerebellothalamic projections coincide with MI thalamocortical projections primarily in VPLo, but with decreasing coincidence of labeling in VLx, VLc, and

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SMA Pre-SMA M1

PMdc

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Fig. 24.3 Schematic drawing of the motor cortical areas shown on a dorsal view of the macaque monkey brain. The motor cortical areas include: the primary motor cortex (MI), rostral and caudal subregions of the dorsal (PMd) and ventral premotor areas (PMv), and the pre-supplementary motor (pre-SMA) and supplementary motor areas (SMA) located on the mesial surface of the hemisphere

VLo (Rouiller et al. 1994; Sakai et al. 2002; Stepniewska et al. 2003) (Fig. 24.4). In addition, a generalized somatotopic organization is observed with the face represented medially and the leg laterally. The idea that cortical areas receive mixed inputs derived from multiple thalamic nuclei was first proposed by Kievit and Kuypers (1977) and later by Darian-Smith et al. (1990). However, it was also proposed that ascending information from the cerebellum and globus pallidus project via parallel and separate pathways to the thalamic nuclei which in turn project to separate motor cortical fields including MI and the supplementary motor area (SMA) (Jones 1985; Ilinsky and Kultas-Ilinsky 1987; Alexander and Crutcher 1990). These results and other similar studies used retrograde tracers to label the thalamocortical neurons and then compared the distribution of labeling with reports of pallido- and cerebellothalamic projections (Schell and Strick 1984; Darian-Smith et al. 1990; Shindo et al. 1995). However, overlapping projections arising from the cerebellar territory and pallidal territory to a single cortical field had been proposed based on single labeling (Nambu et al. 1988, 1991; Matelli et al. 1989; Darian-Smith et al. 1990; Holsapple et al. 1991; Inase and Tanji 1995; Shindo et al. 1995) and multiple labeling experiments (Rouiller et al. 1994; Sakai et al. 1999, 2002). These latter studies showed that MI thalamocortical and cerebellothalamic projections overlap extensively, but regions of overlapping MI thalamocortical cells with pallidothalamic projections are also noted. The regions of overlapping cerebellar and pallidal thalamocortical projections tend to occur within the border areas particularly between VPLo and VLo (Fig. 24.4). As noted earlier, this labeling may be due to the difficulty in distinguishing the cell sparse interdigitating fingers of VPLo (Jones 2007), but the extent to which pallidothalamic projections reach MI remains controversial.

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Pre-SMA VApc SMA VLo

PMdr

GPi

PMdc VLc

PMv VLx

MI

Cerebellum

VPLo

Fig. 24.4 Schematic summary diagram showing the distribution of cerebellothalamic and pallidothalamic projections to motor thalamus and the thalamocortical projections to the motor cortical areas in the macaque monkey. The major input from the dentate and interpositus nuclei is to the contralateral VPLo, VLx, VLc while the major input from the internal segment of the globus pallidus (GPi) is the VLo and VApc. The motor thalamus provides primary input to the motor cortical areas: MI receives dense projections from VPLo, and also VLx and VLc, SMA receives dense input from VLo, pre-SMA receives primary input from both VApc and VLx, PMdr receives primary input from VApc and VLc while PMdc receives primary input from VApc and VLo, and PMv receives primary input from VLo and VPL. Motor cortical areas largely receive mixed and weighted input derived from GPi and cerebellum. The gradient in the projection densities is roughly indicated by the thickness of the arrows (Data from Morel et al. 2005 Divergence and convergence of thalamocortical projections to premotor and supplementary motor cortex: a multiple tracing study in the macaque monkey. Eur J Neurosci 21: 1007–1029; Sakai et al. 2002 The relationship between MI and SMA afferents and cerebellar and pallidal efferents in the macaque monkey. Somatosens Mot Res 19: 139–148; Sakai et al. 2003 Ascending inputs to the pre-supplementary motor area in the macaque monkey: cerebello- and pallido-thalamocortical projections. Thalamus Related Syst 2: 175–187 with permission)

Earlier studies noted that rostral MI, including the proximal forelimb representation, primarily receives thalamic projections from VLo whereas caudal MI, lying within the rostral bank of the central sulcus and containing the distal forelimb representation, primarily receives thalamic projections from VPLo (Matelli et al. 1989; Darian-Smith et al. 1990; Tokuno and Tanji 1993). In contrast, Holsapple et al. (1991) proposed that the caudal MI within the rostral bank of the central sulcus receives input predominantly derived from the globus pallidus via VLo. A preponderance of pallidothalamocortical projections to MI sulcal cortex has not been reported elsewhere, perhaps because few studies have systematically injected the sulcal cortex. However, coincidence of pallidothalamic projections

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with the digit representation of MI sulcal cortex was noted in VLo using multiple labeling techniques (Stepniewska et al. 2003).

Projections to SMA and pre-SMA Cerebellothalamic projections extend beyond MI to project to other motor cortical fields including premotor cortex and supplementary motor areas. The supplementary motor area was originally thought to reside within the mesial cortex anterior to MI and posterior to the frontal granular cortex (Penfield and Welch 1949; Woolsey et al. 1952). However, the traditional SMA has been further subdivided into the rostral pre-supplementary motor area (pre-SMA) and caudal SMA based on distinctive functional and anatomical features (Matsuzaka et al. 1992; Tanji 1994; Matsuzaka and Tanji 1996; Picard and Strick 1996; Sakai et al. 2003) (Fig. 24.3). The SMA is microexcitable cortex but at higher current thresholds than those effective in MI. In contrast, pre-SMA is less responsive to even higher microstimulation currents. Although previous work suggested that the SMA was primarily influenced by the basal ganglia outflow (Schell and Strick 1984), other studies reported thalamic afferents arising from multiple nuclei, including those nuclei that receive cerebellar input (Yamamoto et al. 1984; Wiesendanger and Wiesendanger 1985; Nambu et al. 1988, 1991; Darian-Smith et al. 1990; Matelli et al. 1989, 1996; Tokuno et al. 1992). The afferent distribution of the SMA thalamocortical cells was determined using a triple labeling paradigm whereby the pallidal and cerebellar afferents were labeled using two different anterograde tracers, and the SMA thalamocortical cells were labeled using a retrograde tracer following physiological identification of the hand/arm representation in SMA (Sakai et al. 1999). The interrelationship between the afferent sources and the retrogradely labeled neurons could be directly assessed using this paradigm. The SMA receives primarily afferents arising from VLo coincident with pallidal projections but it also receives some afferents from VLc, VLx, and VPLo, coincident with cerebellar projections (Fig. 24.4). Similar results were reported by Rouiller et al. (1994) based on multiple labeling methods. Because the thalamic projections to MI and SMA arise from overlapping regions, the possibility that these projections might originate from the same neuron required further study. The direct comparison of the ascending projections from the pallidal and cerebellar sources to MI and the supplementary motor area (SMA) using multiple labeling methods revealed that MI and SMA receive predominant thalamic input originating from the cerebellum and globus pallidus, respectively (Rouiller et al. 1994; Sakai et al. 2002) (Fig. 24.4). MI and SMA also received secondary afferent input but evidence of collateralized projections from thalamus to MI and SMA was rare (Darian-Smith et al. 1990; Rouiller et al. 1994; Shindo et al. 1995; Sakai et al. 2002; Morel et al. 2005). The projections to the pre-SMA have also been evaluated using multiple labeling techniques. The pre-SMA occupies the mesial cortex rostral to the SMA and is functionally distinct from SMA in that its neurons are responsive during movement preparation (Tanji 1994; Matsuzaka and Tanji 1996; Picard and Strick 1996) and in updating the temporal order of movement events (Shima and Tanji 1998, 2000).

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Using multiple labeling techniques, the pre-SMA receives ascending inputs from both the cerebellum and globus pallidus by way of the motor thalamus in both the owl monkey (Sakai et al. 2000) and macaque monkey (Sakai et al. 2003) (Fig. 24.4). These results were similar to those reported by Matelli and Luppino (1996) who used fluorescent tracers to retrogradely label these thalamocortical neurons. The pre-SMA inputs primarily arose from caudal VA in the pallidothalamic territory and VLx in cerebellothalamic territory.

Projections to the Premotor Cortex The premotor cortex consists of the cortex lying rostral to MI and is coincident with cytoarchitectonic area 6. The spur of the arcuate sulcus roughly divides the premotor cortex into dorsal and ventral subdivisions (Fig. 24.3). The dorsal premotor cortex (PMd) lies medial to the spur of the arcuate sulcus and extends as far medially as SMA while the ventral premotor cortex (PMv) lies lateral to the spur. In addition, the PMd and PMv can be further subdivided into rostral and caudal divisions. These subdivisions differ anatomically based on cytoarchitectonic and histochemical differences (Barbas and Pandya 1987; Matelli et al. 1985, 1989; Matelli and Luppino 1996; Kurata 1994; Stepniewska et al. 2007) and functionally (Kurata and Tanji 1986; Rizzolatti et al. 1988; Preuss et al. 1996). Recently, the distribution of thalamic afferents to the premotor subdivisions has been reevaluated using multiple labeling techniques in conjunction with quantitative methods. These analyses reveal that each premotor subdivision receives a predominant thalamic input and secondary, less dense afferents derived from multiple thalamic nuclei. The PMd receives afferents from motor thalamus including VLo, VLx, and VLc (Morel et al. 2005; Stepniewska et al. 2007) as well as VApc (Kurata 1994; Matelli and Luppino 1996; Rouiller et al. 1999; Morel et al. 2005; Stepniewska et al. 2007). A topographic shift in the distribution of thalamocortical projections is noted in comparison of the rostral and caudal PMd afferents (Fig. 24.4). Rostral PMd preferentially receives projections from VApc, VLc, and MD whereas caudal PMd preferentially receives projections arising from VLo (VLa in Morel et al. 2005) in the macaque monkey and VLa and VLx in the owl monkey (Stepniewska et al. 2007). Rostral and caudal sectors of PMv also receive differentially distributed afferents: rostral PMv receives predominant input from MD with less dense input from VApc, and VLo and caudal PMv receive predominant afferents from VLo and VLc (Morel et al. 2005). Others have noted significant input arising from VLx to PMv (Matelli et al. 1989; Rouiller et al. 1999; Stepniewska et al. 2007). Taken together, these data demonstrate that sectors of PM receive differentially weighted thalamic inputs. Morel and others (2005) speculated on the extent of divergence and convergence in the cortex by analyzing the degree of overlap and segregation in the thalamocortical projections. They suggest that the degree of thalamic overlap varies in the PM subdivisions with gradients of increasing projections from MD to rostral PM and from VLo and VPLo to caudal PM. To some extent, the degree of overlap in thalamus is related to the proximity of the cortical area. For example, greater overlap in thalamus was noted from adjacent cortical

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areas such as between rostral and caudal PMv (Morel et al. 2005), caudal PMd and SMA (Rouiller et al. 1999), and pre-SMA and rostral PMd (Rouiller et al. 1999). The thalamic inputs to the PM subdivisions cross cytoarchitectonic boundaries and arise from nuclei receiving afferent inputs from the cerebellum and the globus pallidus. The information from these sources is likely to overlap in the cortex. Since the predominant input to both VLx and VPLo originates from the cerebellum and these nuclei, in turn, provide afferents to PM subdivisions, these cortical subdivisions receive cerebellar inputs, albeit of differing strengths (Morel et al. 2005; Stepniewska et al. 2007). Similarly, since the nuclei, VApc, VLo, and VLc, all receive pallidal input and in turn, project to PM, these cortical regions also receive inputs derived from the globus pallidus. In this manner, the PM subdivisions receive mixed inputs from these sources.

Projections to Other Cortical Areas The cerebellar thalamic territory also projects to other cortical areas. The VLx gives rise to a small percentage of cells projecting to the frontal eye field (area 8) and even fewer to area 45 (Contini et al. 2010). The VLc projects to the posterior parietal cortex, in particular, the superior parietal lobule (Miyata and Sasaki 1983; Schmahmann and Pandya 1990). A small number of labeled cells in VLx and VLc were noted projecting to prefrontal cortical areas 9 and 46 using conventional neuroanatomical tracers (Middleton and Strick 2001). As described earlier, cerebellar afferents also distribute to the central lateral nucleus (CL) and the lateral part of MD (Fig. 24.2). Cells of the central lateral nucleus of the intralaminar group project to all of the motor areas and cells in lateral or paralamellar MD project to more rostral cortical areas including motor, frontal eye fields, and prefrontal cortex (Matelli et al. 1989; Darian-Smith et al. 1990; Schmahmann and Pandya 1990; Shindo et al. 1995; Matelli and Luppino 1996; Morel et al. 2005; Stepniewska et al. 2007; Contini et al. 2010). General Topography of Projections Each motor cortical area receives differentially weighted inputs arising from the thalamic territories receiving cerebellar and pallidal afferents (Fig. 24.4). These inputs represent a unique mixture of afferents arising from multiple thalamic nuclei. However, there is considerable overlap in the afferent distribution to adjacent cortical areas, especially from border zones between cortical areas. A general topography corresponding to functional gradients within the motor areas has been previously noted (Matelli et al. 1989; Matelli and Luppino 1996; Rouiller et al. 1999; Morel et al. 2005). Traditional views of the motor cortical areas propose that these areas are hierarchically organized with MI involved in movement execution and the remaining motor cortical areas engaged in higher order aspects of motor control. The PMd and PMv have been associated with different roles in the selection and planning of movement (Wise et al. 1997; Hoshi and Tanji 2007), while the SMA is involved with internally generated movement sequences (Mushiake et al. 1991; Tanji 1994; Shima and Tanji 1998). The pre-SMA is hypothesized to participate in movement preparation and in updating the temporal

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order of movement events (Matsuzaka et al. 1992; Matsuzaka and Tanji 1996; Shima and Tanji 1998, 2000). Despite the distinctly hierarchical functions attributed to each of these areas, recent accounts suggest a modification of this view. Functional analyses of the results obtained from varying microstimulation parameters suggest that the motor cortex processes information required for activating multiple muscles and movements (Schieber 2001; Graziano 2006; Graziano and Afalo 2007). Rather than a focal somatotopic organization of hierarchically organized areas, the motor cortex contains multiple, overlapping, and fractured representations that are suggested to provide the substrate for the production of coordinated synergistic movements within a broad topography (Sanes and Donoghue 2000; Schieber 2001, 2002; Graziano 2006; Graziano and Afalo 2007). Anatomically, a distributed network of diverging and converging connections may well provide the necessary substrate (Schieber 2001; Graziano 2006; Graziano and Afalo 2007; Evrard and Craig 2008). The existence of widespread cortico-cortical projections, diverging corticospinal projections, and converging spinothalamic and cerebellothalamic afferents all have been proposed as links in a larger distributed network subserving flexibility in motor responses (Schieber 2001; Graziano 2006; Graziano and Afalo 2007; Evrard and Craig 2008). In addition, the differentially weighted ascending input originating from cerebellum and globus pallidus would be an important link in such a distributed network. The basal ganglia plays an important role in the acquisition of motor skills, the maintenance of motor routines, and procedural learning (Yin et al. 2009, for review, see Doyon et al. 2003). Both the cerebellum and the basal ganglia output nucleus, the globus pallidus, contribute to motor skill learning (Hikosaka 2002; Groenewegen 2003), but the mechanisms of how these structures interact is largely unknown. The differentially weighted thalamic output to the motor cortical areas may provide an important substrate for motor skill learning. Additional studies comparing the distributions of these projections to the motor cortical areas will help elucidate these mechanisms. For over a decade now, the cerebellum has been increasingly implicated in cognitive processing (Schmahmann 1996; Middleton and Strick 2001; Thach 2007; Ito 2008; Strick et al. 2009). While there is wealth of neuroimaging data in humans demonstrating a role for the cerebellum in higher order cognitive processing, the precise anatomical pathways subserving such functions remain elusive based on direct anatomical tracing methods. Cerebellar inputs largely project to MI, PM subdivisions, and pre-SMA via VPLo, VLx, and VLc. Cerebellothalamic projections distribute less densely to the VA, CL and MD, nuclei that in turn project to more rostral cortical regions including eye movement related areas such as supplementary eye field, frontal eye field, and rostral part of PMd (Shook et al. 1991; Rouiller et al. 1999; Morel et al. 2005). The cerebellar input to MD is largely confined to its lateral or paralamellar portion and projections to other parts of MD are quite sparse (Stanton 1980; Kalil 1981; Asanuma et al. 1983a; Sakai et al. 1996; Stepniewska et al. 2003). Paralamellar MD and CL project diffusely to sensorimotor cortex (for review, see Jones 2007).

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Since the preponderance of the cerebellar projections ultimately target premotor and motor cortical areas, it is likely that the main role of the cerebellothalamic projection system is to facilitate motor responses or action plans (see Glickstein 2007). At the same time, recent evidence suggests that prefrontal cortex may influence the motor thalamus by way of corticothalamic projections (Xiao et al. 2009), thus, providing a route for the cognitive mediation of motor plans.

Conclusions and Future Directions Tremendous progress has been made in detailing the cerebellothalamic and thalamocortical projections in nonhuman primates. The motor thalamus has been defined on the basis of newer chemoarchitectonic methods and correlated with the distribution and topography of these projections using a multitude of neuroanatomical tracing methods. Cerebellothalamic projections arise primarily from the contralateral dentate and interpositus nuclei and the fastigial nucleus bilaterally. These projections heavily distribute to VPLo with less dense projections to adjacent subnuclei including VLx, VLc, VLo, CL, and lateral MD. In turn, these nuclei give rise to projections to the motor cortical areas. Overall, the density of the thalamic projections to these areas varies, giving the impression that each cortical area receives differentially weighted afferents derived from the cerebellum and the globus pallidus, the second source of primary afferents of the motor thalamus. Converging input derived from the cerebellum and the globus pallidus to the motor cortical areas may provide crucial information for movement execution including motor skill learning. The thalamus occupies a pivotal position influencing cerebellar and pallidal access to the cerebral cortex but still little is known regarding the relative contributions of these structures to the overall motor network. Future studies combining multiple neuroanatomical tracers will help elucidate the details of the cortical processing of these inputs. These studies are crucial to our understanding of how motor output is influenced by ascending cerebellar and pallidal information.

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Nambu A, Yoshida S, Jinnai K (1991) Movement-related activity of thalamic neurons with input from the glohus pallidus and projection to the motor cortex in the monkey. Exp Brain Res 84:279–284 Ohye C (1997) Influence of clinical presentation on target selection. Adv Neurol 74:149–157 Ohye C, Narabayashi H (1979) Physiological study of presumed ventralis intermedius neurons in the human thalamus. J Neurosurg 50:290–297 Olszewski J (1952) The thalamus of the Macaca mulatta. Karger, Basel Penfield W, Welch K (1949) The supplementary motor area in the cerebral cortex of man. Trans Am Neurol Assoc 74:179–184 Percheron G, Francois C, Talbi B, Yelnik J, Fenelon G (1996) The primate motor thalamus. Brain Res Rev 22:93–181 Picard N, Strick PL (1996) Motor areas on the medial wall; a review of their location and functional activation. Cereb Cortex 6:342–353 Preuss TM, Stepniewska I, Kaas JH (1996) Movement representation in the dorsal and ventral premotor areas in the owl monkeys: a microstimulation study. J Comp Neurol 371:649–676 Rizzolatti G, Camarda R, Fogassi L, Gentilucci M, Luppino G, Matelli M (1988) Functional organization of inferior area 6 in macaque monkey. II area F5 and the control of distal movement. Exp Brain Res 71:491–501 Rouiller EM, Liang F, Babalian A, Moret V, Wiesendanger M (1994) Cerebellothalamocortical and pallidothalamocortical projections to the primary and supplementary motor cortical areas: a multiple tracing study in macaque monkeys. J Comp Neurol 345:185–213 Rouiller EM, Tanne J, Moret V, Boussaoud D (1999) Origin of thalamic inputs to the primary, premotor, and supplementary motor cortical areas and to ara 46 in macaque monkeys: a multiple retrograde tracing study. J Comp Neurol 409:131–152 Sakai ST, Inase M, Tanji J (1996) Comparison of cerebellothalamic and pallidothalamic projections in the monkey (Macaca fuscata): a double anterograde labeling study. J Comp Neurol 368:215–228 Sakai ST, Inase M, Tanji J (1999) Pallidal and cerebellar inputs to thalamocortical neurons projecting to the supplementary motor area in Macaca fuscata: a triple-labeling light microscopic study. Anat Embryol (Berl) 199:9–19 Sakai ST, Stepniewska I, Qi HX, Kaas JH (2000) Pallidal and cerebellar afferents to presupplementary motor area thalamocortical neurons in the owl monkey: a multiple labeling study. J Comp Neurol 417:164–180 Sakai ST, Inase M, Tanji J (2002) The relationship between MI and SMA afferents and cerebellar and pallidal efferents in the macaque monkey. Somatosens Mot Res 19:139–148 Sakai ST, Stepniewska I, Qi HX, Kaas JH (2003) Ascending inputs to the pre-supplementary motor area in the macaque monkey: cerebello- and pallido-thalamocortical projections. Thalamus Relat Syst 2:175–187 Sanes JN, Donoghue JP (2000) Plasticity and primary motor cortex. Annu Rev Neurosci 23:393–415 Schell GR, Strick PL (1984) The origin of thalamic inputs to the arcuate premotor and supplementary motor areas. J Neurosci 4:539–560 Schieber MH (2001) Constraints on somatotopic organization in the primary motor cortex. J Neurophysiol 86:2125–2143 Schieber MH (2002) Motor cortex and the distributed anatomy of finger movements. Adv Exp Med Biol 508:411–416 Schmahmann JD (1996) From movement to thought: anatomic substrates of the cerebellar contribution to cognitive processing. Hum Brain Map 4:174–198 Schmahmann JD, Pandya DN (1990) Anatomical investigation of projections from thalamus to posterior parietal cortex in the rhesus monkey: a WGA-HRP and fluorescent tracer study. J Comp Neurol 295:299–326 Shima K, Tanji J (1998) Both supplementary and presupplementary motor areas are crucial for the temporal organization of multiple movements. J Neurophysiol 80:3247–3260

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Shima K, Tanji J (2000) Neuronal activity in the supplementary and presupplementary motor areas for temporal organization of multiple movements. J Neurophysiol 84:2148–2160 Shindo K, Shima K, Tanji J (1995) Spatial distribution of thalamic projections to the supplementary motor area and the primary motor cortex: a retrograde multiple labeling study in the macaque monkey. J Comp Neurol 357:98–116 Shook BL, Schlag-Rey M, Schlag J (1991) Primate supplementary eye field. II. Comparative aspects of connections with the thalamus, corpus striatum and related forebrain nuclei. J Comp Neurol 307:562–583 Stanton GB (1980) Topographic organization of ascending cerebellar projections from the dentate and interposed nuclei in Macaca mulatta: an anterograde degeneration study. J Comp Neurol 190:699–731 Starr PA, Vitek JL, Bakay RA (1998) Ablative surgery and deep brain stimulation for Parkinson‘s disease. Neurosurgery 43:989–1015 (see comments) Stepniewska I, Preuss TD, Kaas JH (1994) Architectonic subdivisions of the motor thalamus in owl monkey (Aotus trivirgatus): comparison of Nissl, AChE and CO patterns. J Comp Neurol 349:536–557 Stepniewska I, Sakai ST, Qi HX, Kaas JH (2003) Somatosensory input to the ventrolateral thalamic region in the macaque monkey: potential substrate for parkinsonian tremor. J Comp Neurol 455:378–395 Stepniewska I, Preuss TD, Kaas JH (2007) Thalamic connections of the dorsal and ventral premotor areas in New World owl monkeys. Neurosci 147:727–745 Strick PL, Dum RP, Fiez JA (2009) Cerebellum and nonmotor function. Annu Rev Neurosci 32:413–434 Tanji J (1994) The supplementary motor area in the cerebral cortex. Neurosci Res 19:251–268 Tasker RR, Organ LW, Hawrylyshyn P (1982) Investigation of the surgical target for alleviation of involuntary movement disorders. Appl Neurophysiol 45:261–274 Thach WT (2007) On the mechanism of cerebellar contributions to cognition. Cerebellum 6:163–167 Tokuno H, Tanji J (1993) Input organization of distal and proximal forelimb areas in the monkey primary motor cortex: a retrograde double labeling study. J Comp Neurol 333:199–209 Tokuno H, Kimura M, Tanji J (1992) Pallidal inputs to thalamocortical neurons projecting to the supplementary motor area: an anterograde and retrograde double labeling study in the macaque monkey. Exp Brain Res 90:635–638 Walker AE (1938) The primate thalamus. Chicago University Press, Chicago Wiesendanger R, Wiesendanger M (1985) Cerebello-cortical linkage in the monkey as revealed by transcellular labeling with the lectin wheat germ agglutinin conjugated to the marker horseradish peroxidase. Exp Brain Res 59:105–117 Wise SP, Boussaoud D, Johnson PB, Caminiti R (1997) Premotor and parietal cortex: corticocortical connectivity and combinatorial computations. Annu Rev Neurosci 20:25–42 Woolsey CN, Settlage PH, Meyer DR, Spencer W, Pinto Hamuy T, Travis AM (1952) Patterns of localization in precentral and “supplementary” motor areas and their relation to the concept of a premotor area. Res Publ Assoc Nerv Ment Dis 30:238–264 Xiao D, Zikopoulos B, Barbas H (2009) Laminar and modular organization of prefrontal projections to multiple thalamic nuclei. Neurosci 161:1067–1081 Yamamoto T, Noda T, Miyata M, Nishimura Y (1984) Electrophysiological and morphological studies on thalamic neurons receivng entopedunculo- and cerebellothalamic projections in the cat. Brain Res 301:231–242 Yin HH, Mulcare SP, Hila´rio MRF, Clouse E, Holloway T, Davis MI, Hansson AC, Lovinger DM, Costa RM (2009) Dynamic reorganization of striatal circuits during the acquisition and consolidation of a skill. Nat Neurosci 12:333–341

Cerebellar Outputs in Non-human Primates: An Anatomical Perspective Using Transsynaptic Tracers

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Abstract

Important insights into cerebellar function can be gained from an anatomical analysis of cerebellar output. Recent studies using transsynaptic tracers in nonhuman primates demonstrate that the output of the cerebellum targets multiple nonmotor areas in the prefrontal and posterior parietal cortex, as well as the motor areas of the cerebral cortex. The projections to different neocortical areas originate from distinct output channels within the cerebellar nuclei. The neocortical area that is the main target of each output channel is a major source of input to the channel. Thus, a closed-loop circuit represents the fundamental macro-architectural unit of cerebro-cerebellar interactions. The outputs of these circuits provide the cerebellum with the anatomical substrate to influence the control of movement and cognition. Similarly, it has been shown that discrete multisynaptic loops connect the basal ganglia with motor and nonmotor areas of the cerebral cortex. Interactions between cerebro-cerebellar and cerebro-basal ganglia loops have been thought to occur mainly at the level of the neocortex. More recently, neuroanatomical studies demonstrate that the anatomical substrate exists for substantial interactions between the cerebellum

A.C. Bostan (*) Center for the Neural Basis of Cognition, Systems Neuroscience Institute and Department of Neurobiology, School of Medicine, University of Pittsburgh, Pittsburgh, PA, 15261, USA e-mail: [email protected] P.L. Strick Pittsburgh Veterans Affairs Medical Center, Pittsburgh, PA, 15261, USA and Center for the Neural Basis of Cognition, Systems Neuroscience Institute and Department of Neurobiology, School of Medicine, University of Pittsburgh, Pittsburgh, PA, 15261, USA e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 549 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_25, # Springer Science+Business Media Dordrecht 2013

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and the basal ganglia in both the motor and nonmotor domains. These data, along with the revelations about cerebro-cerebellar circuitry, provide a new framework for exploring the contribution of the cerebellum to diverse aspects of behavior.

Introduction The neocortical areas that provide inputs to the cerebellum have been well established (Fig. 25.1) (Glickstein et al. 1985; Schmahmann 1996). On the other hand, the targets of cerebellar output are still in the process of being fully identified (Strick et al. 2009). Recent results from neuroanatomical studies using transsynaptic tracers in nonhuman primates indicate that cerebellar output targets both motor and nonmotor areas of the cerebral cortex. This feature of cerebellar output provides part of the neural substrate for the involvement of cerebellum not only in the generation and control of movement but also in nonmotor aspects of behavior. This chapter reviews new evidence about the areas of the cerebral cortex that are the target of cerebellar output. It describes the functional map that has recently been discovered within one of the major output nuclei of the cerebellum, the dentate nucleus. Furthermore, the chapter presents evidence that the fundamental unit of cerebro-cerebellar operations is a closed-loop circuit. Finally, it discusses the new anatomical evidence that the cerebellum and basal ganglia are interconnected. The classical view of cerebro-cerebellar interconnections is that the cerebellum receives information from widespread neocortical areas, including portions of the frontal, parietal, temporal, and occipital lobes (Fig. 25.1) (Glickstein et al. 1985; Schmahmann 1996). This information was then thought to be funneled through cerebellar circuits where it ultimately converged on the ventrolateral nucleus of the thalamus (e.g., Allen and Tsukahara 1974; Brooks and Thach 1981). The ventrolateral nucleus was believed to project to a single neocortical area, the primary motor cortex (M1). Thus, cerebellar connections with the cerebral cortex were viewed as means of collecting information from widespread regions of the cerebral cortex. The cerebellum was thought to perform a sensorimotor transformation on its inputs and convey the results to M1 for the generation and control of movement. According to this view, cerebellar output was entirely within the domain of motor control, and abnormal activity in this circuit would lead to purely motor deficits. Recent analysis of cerebellar output and function has challenged this view (e.g., Schell and Strick 1984; Middleton and Strick 1994, 1996a, b, 2000, 2001; Hoover and Strick 1999; Clower et al. 2001, 2005; Dum and Strick 2003; Kelly and Strick 2003; Akkal et al. 2007; Strick et al. 2009). It is now clear that efferents from the cerebellar nuclei project to multiple subdivisions of the ventrolateral thalamus (for a review, see Percheron et al. 1996), which, in turn, project to a myriad of neocortical areas, including regions of frontal, prefrontal, and posterior parietal cortex (Jones 1985). Thus, the outputs from the cerebellum influence more widespread regions of the cerebral cortex than previously recognized. This change in perspective is important because it provides the anatomical substrate for the output

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Fig. 25.1 Origin of projections from the cerebral cortex to the cerebellum. Top: The relative density of corticopontine neurons is indicated by the dots on the lateral and medial views of the macaque brain. Bottom: Histogram of relative density of corticopontine cells in different cytoarchitectonic areas of the monkey. Ai, As inferior and superior limbs of arcuate sulcus, respectively, CA calcarine fissure, CgS cingulate sulcus, CS central sulcus, IP intraparietal sulcus, LS lateral sulcus, Lu luneate sulcus, IO inferior occipital sulcus, PO parietal-occipital sulcus, PS principal sulcus, STS superior temporal sulcus (Adapted from Strick et al. (2009))

of the cerebellum to influence nonmotor as well as motor areas of the cerebral cortex. As a consequence, abnormal activity in cerebro-cerebellar circuits could lead not only to motor deficits but also to cognitive, attentional, and affective impairments. Prior neuroanatomical approaches for examining cerebro-cerebellar circuits have been hindered by a number of technical limitations. Chief among these limitations is the multisynaptic nature of these pathways and the general inability of conventional tracers to label more than the direct inputs and outputs of an area. To overcome these and other problems, neurotropic viruses have been used as transneuronal tracers in the central nervous system of primates (for references and a review, see Strick and Card 1992; Kelly and Strick 2000, 2003). Selected strains of virus move transneuronally in either the retrograde or anterograde direction (Zemanick et al. 1991; Kelly and Strick 2003). Thus, one can examine either the inputs to or the outputs from a site. The viruses used as tracers move from neuron to

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Fig. 25.2 Targets of cerebellar output. Red labels indicate areas of the cerebral cortex that are the target of cerebellar output. Blue labels indicate areas that are not the target of cerebellar output. These areas are indicated on lateral and medial views of the cebus monkey brain. The numbers refer to cytoarchitectonic areas. AIP anterior intraparietal area, AS arcuate sulcus, CgS cingulate sulcus, FEF frontal eye field, IP intraparietal sulcus, LS lateral sulcus, Lu lunate sulcus, M1 face, arm, and leg areas of the primary motor cortex, PMd arm arm area of the dorsal premotor area, PMv arm arm area of the ventral premotor area, PrePMd predorsal premotor area, PreSMA presupplementary motor area, PS principal sulcus, SMA arm arm area of the supplementary motor area, ST superior temporal sulcus, TE area of inferotemporal cortex (Adapted from Strick et al. (2009))

neuron exclusively at synapses, and the transneuronal transport occurs in a timedependent fashion. Careful adjustment of the survival time after a virus injection allows for the study of neural circuits composed of two or even three synaptically connected neurons. Virus tracing has been used to examine cerebellothalamocortical pathways to a wide variety of neocortical areas (Middleton and Strick 1994, 1996a, b, 2001; Lynch et al. 1994; Hoover and Strick 1999; Clower et al. 2001, 2005; Kelly and Strick 2003; Akkal et al. 2007) (Fig. 25.2).

Cerebellar Output Channels In an initial series of studies, virus was injected into physiologically defined portions of M1 and the survival time was set to label second-order neurons in the

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Fig. 25.3 Output channels in the dentate. The dots on representative coronal sections show the location of dentate neurons that project to a specific area of the cerebral cortex in the cebus monkey. The neocortical target is indicated above each section. Abbreviations are according to Fig. 25.2 (M1 primary motor cortex, PMv ventral premotor area). D dorsal, M medial (Adapted from Middleton and Strick (1996b))

deep cerebellar nuclei (Hoover and Strick 1999). In general, cerebellar projections to M1 originate largely from neurons in the dentate nucleus (75%), although a smaller component also originates from the interpositus (25%). Several studies have focused on the organization of the dentate nucleus. The dentate nucleus is a complex three-dimensional structure (Fig. 25.3). Results from different experiments can be displayed in a common framework on an unfolded map of the nucleus (Fig. 25.4) (Dum and Strick 2003). Virus transport following injections into the arm representation of M1 labeled a compact cluster of neurons in the dorsal portion of the dentate at mid-rostrocaudal levels (Figs. 25.2 and 25.3, far right panel, Fig. 25.4, top center panel). Virus transport from the leg representation of M1 labeled neurons in the rostral pole of the dorsal dentate (Figs. 25.2 and 25.4, top left panel), whereas virus transport from the face representation labeled neurons at caudal levels of the dorsal dentate (Figs. 25.2 and 25.4, top right panel). Clearly, each neocortical area receives input from a spatially separate set of neurons in the dentate, which has been termed an output channel (Middleton and Strick 1997). The rostral to caudal sequence of output channels to the leg, arm, and face representations in M1 (Fig. 25.4, top panels, Fig. 25.5) corresponds well with the somatotopic organization of the dentate previously proposed on the basis of physiological studies (e.g., Allen et al. 1978; Stanton 1980; Rispal-Padel et al. 1982; Asanuma et al. 1983; Thach et al. 1993). The region of the dentate that contains neurons that project to M1 occupies only 30% of the nucleus (Hoover and Strick 1999; Dum and Strick 2003). This implies that a substantial portion of the dentate projects to neocortical targets other than M1. To test this proposal and define the neocortical targets of the unlabeled regions of the dentate, virus was injected into selected premotor, prefrontal, and posterior parietal areas of the cortex (Fig. 25.2). Virus transport from the arm representations of the ventral premotor area (PMv) and the supplementary motor area (SMA) provided evidence that both neocortical areas are the targets of cerebellar output (Fig. 25.2) (Middleton and Strick 1997;

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M1 face (Jo18)

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Fig. 25.4 Unfolded maps of the dentate: output channels to different areas of the cerebral cortex in the cebus monkey. Top panels: Somatotopic organization of output channels to leg, arm, and face M1 in the dorsal dentate. Bottom panels: Ventral location of output channels to prefrontal cortex. The key below each diagram indicates the density of neurons in bins through the nucleus. Rostral is to the left. Abbreviations are according to Fig. 25.1 (M1 primary motor cortex, PreSMA presupplementary motor area) (Adapted from Dum and Strick (2003) (which includes a detailed description of the unfolding method))

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Fig. 25.5 Summary map of dentate topography. The lettering on the unfolded map indicates the neocortical target of different output channels in the cebus monkey. The location of different output channels divides the dentate into motor and nonmotor domains. Staining for monoclonal antibody 8B3 is most intense in the nonmotor domain. The dashed line marks the limits of intense staining for this antibody. The designation of the region marked by “?” is unclear. Abbreviations as in Fig. 25.2 (FEF frontal eye field, M1 primary motor cortex, PMv ventral premotor area, PreSMA presupplementary motor area, SMA supplementary motor area) (Adapted from Dum and Strick (2003) and Akkal et al. (2007))

Akkal et al. 2007). The output channels to these premotor areas are located in the same region of the dentate that contains the output channel to arm M1 (Figs. 25.3 and 25.5). It has been hypothesized that the clustering of output channels to M1 and the premotor areas in the dorsal region of the dentate creates a motor domain within the nucleus (Fig. 25.5) (Dum and Strick 2003). It has been shown that the dorsal premotor cortex (PMd) also receives inputs from the motor territory of the dentate (Hashimoto et al. 2010). Interestingly, the output channels to the arm representations of M1, PMv, PMd, and SMA appear to be in register within the dentate. This raises the possibility that the nucleus contains a single integrated map of the body within the motor domain. Virus transport following injections into prefrontal cortex revealed that some subfields are the target of dentate output, whereas others are not (Middleton and Strick 1994, 2001). Dentate output channels project to areas 9m, 9l, and 46d, but not to areas 12 and 46v (Figs. 25.2–25.5). Importantly, the extent of the dentate that is occupied by an output channel to a specific area of prefrontal cortex is comparable to that occupied by an output channel to a neocortical motor area (Fig. 25.4). Thus, it is likely that the signal from the dentate to prefrontal cortex is as important as its signal to one of the neocortical motor areas. In addition, dentate output channels to

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areas of prefrontal cortex are located in a different region of the nucleus than the output channels to the neocortical motor areas. The output channels to prefrontal cortex are clustered together in a ventral region of the nucleus that is entirely outside the motor domain. The output channels to prefrontal cortex are also rostral to the output channel that targets the frontal eye field (Lynch et al. 1994). Although the presupplementary motor area (PreSMA) has traditionally been included with the motor areas of the frontal lobe, evidence indicates that it should be considered a region of prefrontal cortex (for reviews, see Picard and Strick 2001; Akkal et al. 2007). In support of this proposal, virus transport from the PreSMA labeled an output channel in the ventral dentate where the output channels to areas 9 and 46 are located (Figs. 25.2 and 25.4, bottom, Fig. 25.5). This result illustrates that the topographic arrangement of output channels in the dentate does not mirror the arrangement of their targets in the cerebral cortex. For example, the PreSMA is adjacent to the SMA on the medial surface of the hemisphere (Fig. 25.2), but the output channels to the two neocortical areas are spatially separated from one another in the dentate (Fig. 25.5). Thus, the topographic arrangement of output channels in the dentate appears to reflect functional relationships between neocortical areas rather than the spatial relationships among them. Virus transport from regions of posterior parietal cortex demonstrated that some of its subfields are also the target of output channels located in the dentate (Figs. 25.2 and 25.5) (Clower et al. 2001, 2005). For example, area 7b, which in the cebus monkey is located laterally in the intraparietal sulcus, is the target of an output channel located ventrally in the caudal pole of the dentate (Fig. 25.5). A second region of posterior parietal cortex, the anterior intraparietal area (AIP), receives a focal projection from a small cluster of neurons that is located dorsally in the dentate at mid-rostro-caudal levels. In addition, the AIP receives a broadly distributed projection from neurons that are scattered in dentate regions that contain output channels to M1, the PMv, and perhaps other premotor areas. This creates a unique situation in which AIP may receive a sample of the dentate output that is streaming to motor areas in the frontal lobe, as well as input from its own separate output channel. However, area 7a, which is located on the inferior parietal lobule (Fig. 25.2), does not receive substantial input from the dentate or other cerebellar nuclei (Clower et al. 2001). There also is evidence that the medial intraparietal area (MIP) and ventral lateral intraparietal area (LIPv) are the targets of cerebellar output from the deep cerebellar nuclei (Prevosto et al. 2010). Currently, the information about cerebellar projections to other areas in the posterior parietal cortex is complex and incomplete. It is clear, however, that multiple areas of the posterior parietal cortex are the targets of output channels from the ventral dentate. Maps from individual experiments have been coalesced into a single summary diagram where the average location of each output channel is indicated (Fig. 25.5). This summary diagram emphasizes several notable features about the topographic organization of the dentate. A sizeable portion of the nucleus projects to parts of the prefrontal and posterior parietal cortex. The output channels to prefrontal and posterior parietal areas are clustered in a ventral and caudal region of the nucleus. Consequently, these output channels are spatially segregated from those in the

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dorsal dentate that target motor areas of the cortex. Thus, the dentate appears to be spatially subdivided into separate motor and nonmotor domains that focus on functionally distinct neocortical systems. Another feature emphasized by the summary diagram is that the neocortical targets for large portions of the dentate remain to be determined. The division of the dentate into separate motor and nonmotor domains is reinforced by underlying molecular gradients within the nucleus (Fortin et al. 1998; Pimenta et al. 2001; Dum et al. 2002; Akkal et al. 2007). Fortin et al. (1998) reported that immuno-staining for two calcium-binding proteins, calretinin and parvalbumin, is greatest in ventral regions of the squirrel monkey dentate. A monoclonal antibody, 8B3, which recognizes a chondroitin sulfate proteoglycan on subpopulations of neurons, also differentially stains the dentate in cebus monkeys and macaques (Pimenta et al. 2001; Dum et al. 2002; Akkal et al. 2007). Immunoreactivity for 8B3 is most intense in ventral regions of the dentate that project to prefrontal and posterior parietal areas of cortex. In contrast, antibody staining is least intense in the dorsal regions of the nucleus that project to the neocortical motor areas. These observations suggest that 8B3 recognizes a significant portion of the nonmotor domain within the dentate. Measurements indicate that approximately 40% of the nucleus is intensely stained by 8B3. This analysis does not include the caudal portion of the dentate (marked by a “?” in Fig. 25.5) because this region does not stain intensely for 8B3 and its neocortical target remains to be determined. However, based on its location, it is likely that this caudal region projects to a nonmotor area of the cerebral cortex. If this is the case, then the nonmotor domain of the dentate may represent as much as 50% of the nucleus in the cebus monkey. In the human, it has long been recognized that the dentate is composed of a dorsal, microgyric portion and a ventral, macrogyric portion (for references and illustration, see Voogd 2003). Compared with the microgyric dentate, the macrogyric dentate is reported to (a) develop later, (b) have smaller cells, (c) display a selective vulnerability in cases of neocerebellar atrophy, and (d) have a higher iron content. This last observation suggests that molecular gradients may exist within the human dentate as they do in the monkey dentate; however, this possibility remains to be tested. Comparative studies suggest that the dentate has expanded in great apes and humans relative to the other cerebellar nuclei (Matano et al. 1985). Furthermore, most of this increase appears to be due to an expansion in the relative size of the ventral half of the dentate (Matano 2001). This observation implies that the nonmotor functions of the dentate grow in importance in great apes and humans.

Macro-architecture of Cerebro-cerebellar Loops The neocortical areas that are the target of cerebellar output also project via the pons to the cerebellar cortex (Glickstein et al. 1985; Schmahmann 1996). This observation suggests that cerebro-cerebellar connections may form a closed-loop

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Fig. 25.6 Regions of cerebellar cortex that project to areas of cerebral cortex. The black dots on the flattened surface maps of the cerebellar cortex indicate the location of Purkinje cells that project to the arm area of M1 (left panel) or to area 46 (right panel) in the cebus monkey. The Purkinje cells that project to M1 are located in lobules that are separate from those that project to area 46. Nomenclature and abbreviations are according to Larsell (1970) (Adapted from Kelly and Strick (2003))

circuit. This concept has been tested for a representative motor area (the arm area of M1) and a nonmotor area (area 46 in the prefrontal cortex) (Kelly and Strick 2003). The anatomical evidence indicates that a specific region of the cerebellar cortex both receives input from and projects to the same area of the cerebral cortex. Retrograde transneuronal transport of rabies virus was used to define the Purkinje cells in cerebellar cortex that project to M1 or to area 46. The arm area of M1 receives input from Purkinje cells located mainly in lobules IV–VI of the cerebellar cortex (Fig. 25.6, left panel). In contrast, area 46 receives input from Purkinje cells located mainly in Crus II of the ansiform lobule (Fig. 25.6, right panel). There is no evidence of overlap between the two systems. Thus, the two areas of the cerebral cortex are the targets of output from Purkinje cells that are located in separate regions of the cerebellar cortex. Clearly, the separation of motor

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Fig. 25.7 Closed-loop circuits link the cerebellum with the cerebral cortex. Two topographically separate closed-loop circuits are illustrated. One interconnects the cerebellum with M1 and the other interconnects the cerebellum with area 46. In each loop, the neocortical area projects to a specific site in the pontine nuclei (PN), which then innervates a distinct region of the cerebellar cortex (CBM). Similarly, a portion of the dentate nucleus (DN) projects to a distinct region of the thalamus, which then innervates a specific neocortical area. Note that the neocortical area, which is the major source of input to a circuit, is the major target of output from the circuit. CBM cerebellar cortex, DN dentate, PN pontine nuclei, TH subdivisions of the thalamus (Adapted from Strick et al. (2009))

and nonmotor functions seen in the dentate nucleus extends to the level of the cerebellar cortex. In separate experiments, anterograde transneuronal transport of herpes virus was used to define the granule cells in cerebellar cortex that receive input from M1 or from area 46. The arm area of M1 projects to granule cells located mainly in lobules IV–VI, whereas area 46 projects to granule cells mainly in Crus II. Again, each cerebral cortical area projects to granule cells that are located in a separate region of the cerebellar cortex. Moreover, these findings indicate that the regions of the cerebellar cortex that receive input from M1 are the same as those that project to M1. Similarly, the regions of the cerebellar cortex that receive input from area 46 are the same as those that project to area 46. Thus, M1 and area 46 form separate, closed-loop circuits with different regions of the cerebellar cortex (Fig. 25.7). Altogether, these observations suggest that multiple closed loop circuits represent a fundamental macro-architectural feature of cerebro-cerebellar interactions. There are a number of important functional implications to these results. They suggest that the cerebellar cortex is not functionally homogeneous. Instead, the results imply that cerebellar cortex contains localized regions that are interconnected with specific motor or nonmotor areas of the cerebral cortex. In fact, it has been hypothesized that the map of function in the cerebellar cortex is likely to be as rich and complex as that in the cerebral cortex (Kelly and Strick 2003).

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As a consequence, global dysfunction of the cerebellar cortex can cause wide-ranging effects on behavior (e.g., Schmahmann 2004). However, localized dysfunction of a portion of the cerebellar cortex can lead to more limited deficits, which may be motor or nonmotor depending on the specific site of the cerebellar abnormality (e.g., Fiez et al. 1992; Schmahmann and Sherman 1998; Allen and Courchesne 2003; Gottwald et al. 2004). Thus, precisely defining the location of a lesion, a site of activation, or a recording site is as important for studies of the cerebellum as it is for studies of the cerebral cortex. As noted above, the neocortical targets for substantial portions of the dentate remain unidentified. In addition, fastigial and interpositus nuclei send efferents to the thalamus (Batton et al. 1977; Stanton 1980; Kalil 1981; Asanuma et al. 1983), and the neocortical targets of these deep nuclei remain to be fully determined. The closed-loop architecture described above enables us to make some predictions about additional neocortical targets of cerebellar output (Middleton and Strick 1998; Dum and Strick 2003; Kelly and Strick 2003). If closed-loop circuits reflect a general rule, then all of the areas of cerebral cortex that project to the cerebellum are the targets of cerebellar output. In addition to the neocortical areas that have already been investigated, the cerebellum receives input from a wide variety of higher-order, nonmotor areas. This includes areas of extrastriate cortex, posterior parietal cortex, cingulate cortex, and the parahippocampal gyrus on the medial surface of the hemisphere (Fig. 25.1) (Brodal 1978; Wiesendanger et al. 1979; Vilensky and van Hoesen 1981; Leichnetz et al. 1984; Glickstein et al. 1985; Schmahmann and Pandya 1991, 1993, 1997). If some or all of these areas turn out to be cerebellar targets, then the full extent of cerebellar influence over nonmotor areas of the cerebral cortex is remarkable and much larger than previously suspected. In discussing the neural substrate for a cerebellar influence over nonmotor functions, it is important to note the longstanding notion that the cerebellum is interconnected with the limbic system. Cerebellar stimulation can alter limbic function and elicit behaviors like sham rage, predatory attack, grooming, and eating (e.g., Zanchetti and Zoccolini 1954; Berntson et al. 1973; Reis et al. 1973). Cerebellar lesions can tame aggressive monkeys without creating gross motor abnormalities (Peters and Monjan 1971; Berman 1997). Classic electrophysiological evidence suggests that cerebellar stimulation, especially in portions of the fastigial nucleus and associated regions of vermal cortex, can evoke responses at limbic sites, including the cingulate cortex and amygdala (e.g., Anand et al. 1959; Snider and Maiti 1976). The major weakness in the cerebello-limbic hypothesis is the absence of a clear anatomical substrate that links the output of the cerebellum, and especially the fastigial nucleus, with limbic sites such as the amygdala. Although neuroanatomical evidence indicates that the deep cerebellar nuclei are interconnected with the hypothalamus (Haines et al. 1990), these connections do not appear sufficient to mediate all of the behavioral effects evoked by cerebellar stimulation. Thus, the circuits that link the output of the cerebellar nuclei with regions of the limbic system need to be explored using modern neuroanatomical methods.

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The Cerebellum Is Interconnected with the Basal Ganglia The loops that link the cerebellum with the cerebral cortex have traditionally been considered to be anatomically and functionally distinct from those that link the basal ganglia with the cerebral cortex (Doya 2000; Graybiel 2005). As the projections from the cerebellum and basal ganglia to the cerebral cortex are relayed through distinct thalamic nuclei (Percheron et al. 1996; Sakai et al. 1996), any interactions between cortico-cerebellar and cortico-basal ganglia loops were thought to occur primarily at the neocortical level. Results from recent anatomical experiments challenge this perspective and provide evidence for disynaptic pathways that link the cerebellum with the basal ganglia more directly. To explore whether the cerebellum projects to the basal ganglia, rabies virus was injected into a region of the putamen. The injection sites were localized largely to the sensorimotor territory of the striatum (Parent and Hazrati 1995a). The virus went through two stages of transport: retrograde transport to first-order neurons in the thalamus that innervate the injection site and then, retrograde transneuronal transport to second-order neurons in the deep cerebellar nuclei that innervate the first-order neurons (Fig. 25.8). The neurons in the cerebellar nuclei that were labeled by virus transport were located largely in the dentate nucleus. Thus, a major output of cerebellar processing, the dentate, projects via the thalamus to an input stage of basal ganglia processing, the putamen. In another series of experiments, rabies virus was injected into the external segment of the globus pallidus (GPe). The virus went through three stages of transport: retrograde transport of the virus from the injection site to first-order neurons in the striatum, retrograde transneuronal transport from these first-order neurons to second-order neurons in the thalamus, and retrograde transneuronal transport from the second-order neurons in the thalamus to third-order neurons in the deep cerebellar nuclei. Most of the labeled neurons in the cerebellar nuclei were confined to the dentate (Fig. 25.8). Thus, not only does the output from the cerebellum influence the striatum, but the target of this influence includes striatal neurons in the so-called indirect pathway which projects to GPe (e.g., DeLong and Wichmann 2007). The injections of rabies virus into GPe involved two different regions of the nucleus. The injection in one animal labeled neurons primarily in ventral and caudal regions of dentate. The injection site in the other animal was placed approximately 1 mm caudally in GPe and labeled neurons in more dorsal regions of dentate. These observations suggest that the projection from the dentate to the basal ganglia is topographically organized. Virus transport from the basal ganglia labeled neurons in both the motor and nonmotor domains of the dentate (Hoshi et al. 2005). These observations suggest that the cerebellar projection to the input stage of basal ganglia processing influences motor and nonmotor aspects of basal ganglia function. To explore whether the basal ganglia project to the cerebellum, rabies virus was injected into selected sites within the cerebellar cortex. The virus went through two stages of transport: retrograde transport of the virus from the injection site to

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Fig. 25.8 (continued)

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first-order neurons in the pontine nuclei, and then, retrograde transneuronal transport from these first-order neurons to second-order neurons in the subthalamic nucleus (STN) of the basal ganglia (Figs. 25.8 and 25.9). Rabies virus injections were placed in two areas within the hemispheric expansion of cerebellar lobule VII: the posterior aspects of Crus II (Crus IIp) and the hemispheric lobule VIIB (HVIIB). In all of these experiments, virus transport labeled substantial number of second-order neurons in the STN (Fig. 25.9). The second-order neurons labeled from virus injections into Crus IIp and HVIIB differed in their rostro-caudal and dorso-ventral distributions within the STN. The Crus IIp injections labeled larger numbers of neurons in ventromedial portions of rostral STN, whereas the HVIIB injections labeled larger numbers of neurons in the dorsal aspects of caudal STN (Fig. 25.9). Thus, a disynaptic connection links the STN with cerebellar cortex and this connection is topographically organized. The STN can be subdivided into sensorimotor, associative, and limbic territories based on its interconnections with regions of the globus pallidus and the ventral pallidum (Fig. 25.9) (Parent and Hazrati 1995b; Joel and Weiner 1997; Hamani et al. 2004). The results from rabies virus injections into cerebellar cortex provide evidence that the projections from the STN to the cerebellar cortex originate from all three of its functional subdivisions. Specifically, most of the STN neurons that project to Crus IIp were found in the associative territory, in regions that receive substantial inputs from the frontal eye fields and regions of the prefrontal cortex (Fig. 25.9) (Monakow et al. 1978; Stanton et al. 1988; Inase et al. 1999; Kelly and Strick 2004). In contrast, most of the STN neurons that project to HVIIB were found in the sensorimotor territory, in regions that receive substantial inputs from the primary motor cortex and premotor areas of the frontal lobe (Fig. 25.9) (Monakow et al. 1978; Nambu et al. 1996, 1997; Inase et al. 1999; Kelly and Strick 2004).

ä Fig. 25.8 Experimental paradigms and circuits interconnecting the cerebellum and basal ganglia: The left panel depicts the experimental paradigm and results from Hoshi et al. (2005), describing cerebellar output to the basal ganglia (orange circuit). Rabies virus was injected into the striatum. The virus went through two stages of transport: retrograde transport to first-order neurons in the thalamus that innervate the injection site and then, retrograde transneuronal transport to secondorder neurons in the dentate nucleus (DN) that innervate the first-order neurons. Striatal neurons that receive cerebellar inputs include neurons in the “indirect” pathway that send projections to the external globus pallidum (GPe). The right panel of the figure depicts the experimental paradigm and results from Bostan et al. (2010), describing basal ganglia output to the cerebellum (purple circuit). Rabies virus was injected into the cerebellar cortex. The virus went through two stages of transport: retrograde transport to first-order neurons in the pontine nuclei (PN) that innervate the injection site and then, retrograde transneuronal transport to second-order neurons in the subthalamic nucleus (STN) that innervate the first-order neurons. These interconnections enable two-way communication between the basal ganglia and the cerebellum. Each of these subcortical structures has separate parallel interconnections with the cerebral cortex (up and down large black arrows). The small black arrows in both panels indicate the direction of virus transport. DN dentate nucleus, GPe external segment of the globus pallidus, GPi internal segment of the globus pallidus, PN pontine nuclei, STN subthalamic nucleus (Adapted from Bostan and Strick (2010))

564 Fig. 25.9 STN projection to the cerebellar hemisphere: (a) Histogram of the rostrocaudal distribution of secondorder neurons labeled in the STN by retrograde transport of virus from Crus IIp (red bars) and HVIIB (blue bars). Missing bars correspond to missing sections. (b) Charts of labeled neurons in STN after rabies virus injections into Crus IIp (red dots) and HVIIB (blue dots) are overlapped to illustrate the topographic differences in distribution of STN secondorder neurons in the two cases. (c) Schematic representation of STN organization, according to the tripartite functional subdivisions of the basal ganglia (Parent and Hazrati 1995b; Joel and Weiner 1997; Hamani et al. 2004). (d) Schematic summary of the known connections between STN and areas of the cerebral cortex. C caudal, D dorsal, M medial, STN subthalamic nucleus (Adapted from Bostan et al. (2010))

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Therefore, the anatomical substrate exists for both motor and nonmotor aspects of basal ganglia processing to influence cerebellar function. The results from the transsynaptic tracer studies reveal the anatomical substrate for two-way communication between the cerebellum and the basal ganglia in both the motor and nonmotor domains. One prediction from these findings is that activity in one of these major subcortical systems may directly affect the function of the other. Similarly, the interconnections between the two structures may enable abnormal activity at one site to propagate to the other. Such interactions between the cerebellum and the basal ganglia are likely to have important implications for motor and nonmotor functions. They supply a framework for understanding cerebellar contributions to disorders such as Parkinson’s disease and dystonia that have traditionally been considered “basal ganglia disorders” (for a review, see Bostan and Strick 2010). Furthermore, the anatomical connections between the cerebellum and the basal ganglia provide a potential explanation for the presence of cerebellar involvement in studies that were explicitly designed to study the normal functions of the basal ganglia. For example, several imaging studies have examined whether regions of the basal ganglia and related neocortical areas display functional activation consistent with their involvement in temporal difference models of reward-related learning (O’Doherty et al. 2003; Seymour et al. 2004). It is noteworthy that robust cerebellar activation was present in these experiments along with activation in the dorsal and ventral striatum. The disynaptic connection between the cerebellum and the basal ganglia provide an anatomical substrate for reward-related signals in the basal ganglia to influence cerebellar function during learning, and vice versa. Thus, the two subcortical structures may be linked together to form an integrated network. Future work is needed to elucidate the functional characteristics of this network.

Summary and Conclusions The dominant view of cerebellar function over the past century has been that it is concerned with the coordination and control of motor activity through its connections with M1 (Brooks and Thach 1981). It is now apparent that a significant portion of the output of the cerebellum projects to nonmotor areas of the cerebral cortex, including regions of prefrontal and posterior parietal cortex. Thus, the anatomical substrate exists for cerebellar output to influence the cognitive and visuospatial computations performed in prefrontal and posterior parietal cortex (Clower et al. 2001, 2005; Middleton and Strick 2001). Furthermore, it has been shown that there are significant interconnections between the cerebellum and the basal ganglia in both the motor and nonmotor domains. Thus, the anatomical substrate exists for cerebellar output to influence the basal ganglia, and vice versa. As a corollary, abnormalities in cerebellar structure and function have the potential to produce multiple motor and nonmotor deficits by affecting various neocortical areas and subregions of the basal ganglia.

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The output to nonmotor areas of the cerebral cortex and basal ganglia originates specifically from a ventral portion of the dentate. This nonmotor region of the dentate is recognized by several molecular markers. Several authors have argued that ventral dentate and related regions of the cerebellar hemispheres are selectively enlarged in great apes and humans (Leiner et al. 1991; Matano 2001). Indeed, the enlargement of the ventral dentate in humans is thought to parallel the enlargement of prefrontal cortex. These observations have led to the proposal that the dentate participation in nonmotor functions may be especially prominent in humans (e.g., Leiner et al. 1991; Schmahmann and Sherman 1998). In recent years, concepts about cerebellar structure and function have changed radically. Not only is the cerebellum informed by neocortical information from multiple domains, but cerebellar output is directed at a variety of neocortical regions. As a consequence the output from the cerebellum can impact not only the generation and control of movement, but also cognition and affect. The anatomical evidence that the cerebellum exerts an influence over nonmotor function is complemented by results from neuroimaging studies and by the analysis of the deficits that accompany cerebellar lesions. Thus, it has become clear that the adaptive plasticity that the cerebellum provides for the generation and control of movement is also available for cognition and affect.

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Delineation of Cerebrocerebellar Networks with MRI Measures of Functional and Structural Connectivity

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Christophe Habas, William R. Shirer, and Michael D. Greicius

Abstract

In humans, resting-state functional connectivity MRI (fcMRI) allows precise in vivo delineation of the neocerebellum’s participation in well-segregated, nonmotor intrinsic connectivity networks (ICNs). These data reveal that the neocerebellum participates in several ICNs, including the default mode network (lobule IX), the salience network (lobule VI), and the right and left executive networks (crus I and II). Additionally, fcMRI permits an anatomical parcellation of the neocerebellum based on its specific functional links with the associative cortex. Lobules V, VII, IX, and especially crus I and II constitute a supramodal cognitive zone specifically interconnected with prefrontal, parietal, and cingulate neocortices. Structural connectivity using DTI-based tractography complements fcMRI data and confirms anatomical connections between the dentate nucleus, thalamus, and associative cortices. Taken together, these results support the theory that specific neocerebellar subregions are key nodes in parallel, multisynaptic, closed-loop circuits involved in executive, mnemonic, and affective functions.

C. Habas (*) Service de NeuroImagerie, CHNO des XV-XX, Universite´ Pierre et Marie Curie Paris 6, 28 rue de Charenton, 75012 Paris, France e-mail: [email protected] W.R. Shirer • M.D. Greicius Department of Neurology and Neurological Sciences, Functional Imaging in Neuropsychiatric Disorders (FIND) Lab, Stanford University School of Medicine, 300 Pasteur Drive, 94305-5101 Stanford, CA, USA e-mail: [email protected], [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 571 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_26, # Springer Science+Business Media Dordrecht 2013

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Introduction In primates, the cerebrocerebellar system is organized into discrete, parallel, multisynaptic, closed-loop circuits (Strick et al. 2009). A common gross anatomical connectivity pattern is shared by all these circuits. The cerebral cortex selectively projects via the pontine nuclei (PN) to the contralateral deep cerebellar nuclei, mainly the dentate nuclei (DN), and to the associated cerebellar cortex. The DN, in turn, send projections to the cerebral cortex via the contralateral thalamus (Vincent et al. 2003; Nieuwenhuys et al. 2007). These cerebro-ponto-cerebello-thalamocerebral networks specifically and differentially influence motor, premotor, and association cortices (Middleton and Strick 1997; Schmahmann and Pandya 1997). For instance, tracing methods demonstrate that dorsal, lateral, and ventral parts of the DN are specifically connected with frontal motor, premotor, and prefrontal cognitive regions, respectively (Middleton and Strick 2001; Dum and Strick 2003; Akkal et al. 2007) (Fig. 26.1). Moreover, motor areas preferentially connect with the anterior cerebellar lobe (lobules I to V) and lobule VIII (second cerebellar homunculus), whereas executive and limbic areas are mostly connected with the posterior lobe (lobules VI and VII), which is densely interconnected with the DN. From apes to humans, the telencephalization process is accompanied by an increasing number of nonmotor cerebral afferents reaching the neocerebellum (lobules VI and VII) (MacLeod et al. 2003; Whiting and Barton 2003). It can therefore be inferred that, in humans, the neocerebellum contributes to parallel associative cerebrocerebellar subsystems involved in various aspects of cognition and emotion (Schmahmann 2004). The hypothesis concerning topographic arrangement of motor and nonmotor function in th cerebellum was proposed by Schmahmann (1991, 2004) who regarded the cerebellum as a general modulator of cerebral activity, with the anterior and posterior cerebellum supporting and refining motor and cognitive performance, respectively. In support of this view, neuroimaging studies have shown specific cerebellar activation during emotion, language, working memory, and executive function (reviewed in: Stoodley and Schmahmann 2008), while clinical data have substantiated several, but sometimes variable, cognitive and affective impairments in patients suffering from focal cerebellar lesions (Schmahmann and Shermann 1998; Levisohn et al. 2000; Tedesco et al. 2011). Despite evidence from functional imaging and lesion studies, cerebellar involvement in cognition remains a matter of debate. For instance, some studies failed to find significant attentional or semantic impairment in cerebellar patients (Helmuth et al. 1997; Thier et al. 1999; Haarmeier and Thier 2007). Furthermore, in chronic patients, standard neuropsychological tests often turn up only minor (if any) cognitive impairments even though motor deficits are readily detected. These inconsistencies in cognitive impairments could be attributed to several factors, such as compensation for cerebellar disorders by unaffected cerebellar and cerebral regions or stronger involvement of the cerebellum during “early cognitive development rather than during cognitive performance in adulthood” (Timmann and Daum 2010).

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Fig. 26.1 Input and output organization of the cerebellar cortex (a) and dentate nuclei, the output cerebellar channel, (b) in the primate cortex as revealed by transneuronal tracers. (a1) Projections from Purkinje cells of the cerebellar cortex to the motor and association prefrontal cortices. (a2) Projections from the motor and prefrontal cortices to the granule cells of the cerebellar cortex. (b1) Dentate regions connected with the motor and association cortices. (b2) Cortical areas targeted by dentato-thalamic projections. (b3) Thalamic nuclei relaying dentate output to motor and association cortices (From Dum and Strick 2003; with authors’ permission)

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However, until recently, the nonmotor, probably genetically prewired cerebrocerebellar circuits subserving these cognitive and emotional functions could not be directly and completely identified in humans because of two limitations. First, the gold-standard histological tracing methods used in animals cannot be applied to humans. Second, standard fMRI studies using the general linear model only discriminate a limited number of highly task-specific brain areas whose blood oxygenation level–dependent (BOLD) signal time series closely mimics the temporal model of the experimental task (van Dijk et al. 2010). Thus, in those cases where, for example, a cerebellar region’s BOLD signal displays a complex or subthreshold relation to the task waveform, fMRI may underestimate the number of nodes comprising a task-specific network. Recently, two complementary MRI methods have been developed which overcome these limitations and allow for functional and structural identification of large-scale brain networks: resting-state functional connectivity MRI (fcMRI) and diffusion tensor imaging (DTI) tractography. fcMRI relies on temporal correlations between spontaneous low-frequency (0.01–0.1 Hz) fluctuations of the BOLD signal between spatially distinct but functionally related cortical and subcortical regions (Beckmann et al. 2005; Fox and Raichle 2007). Regions with synchronous spontaneous activity constitute intrinsic connectivity networks (ICNs) and may be linked by mono- or polysynaptic pathways (Greicius et al. 2008; Vincent et al. 2007). The degree to which these spontaneous fMRI signal fluctuations reflect ongoing conscious processing rather than, for example, nonconscious rhythmic waves of cortical excitability is a matter of continuing debate. Generally, two methods are used to extract these ICNs from raw fcMRI data. First, independent component analysis (ICA) is an exploratory, model-free, data-driven statistical method, which transforms the whole resting-state dataset into maximally independent spatial components (Beckmann and Smith 2004). Each component consists of a spatial map and its associated time series. In a typical ICA of an 8-minute resting-state fMRI dataset, there may be 40 components computed, of which 10–15 may represent genuine ICNs, based on their resemblance to well-characterized task-activation networks. The remaining 25–30 represent various noise sources that can also result in correlated BOLD signal fluctuations. The second method uses a region of interest (ROI) as a seed for a whole-brain correlation analysis. That is, the mean (or major Eigen) time series of an ROI is extracted and used as a regressor to search the brain for other voxels whose time series is significantly correlated with the ROI. This results in a map of functional connectivity to the ROI. The ICNs derived from resting-state fMRI studies have been shown to overlap to a large degree with maps of structural connectivity derived from DTI tractography analyses (Skudlarski et al. 2008; Greicius et al. 2009; van den Heuvel et al. 2009). Therefore, fcMRI identifies functionally related areas belonging to a common specialized structural network, whose anatomical architecture can only be established by tractography (within the limits of its spatial resolution and of its ability to detect fiber crossings). However, it is noteworthy that despite a strong correlation between functional and structural connectivity, especially concerning the DMN, topography of ICNs can be influenced by previous or ongoing cognitive

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processing (Hasson et al. 2009; Shirer et al. 2011) and can exhibit variability between sessions and individuals (Honey et al. 2009). Thus, ICNs may represent polysynaptic circuits and may continually reconfigure around the underlying anatomical skeleton (Honey et al. 2009). Resting-state connectivity of the cerebellar system has successfully been studied with both ROI-based and ICA-based methods. The ROI-based method was applied to the cerebellar cortex in order to delineate cerebellar subregions preferentially associated with the dentate nucleus as well as with motor, sensory, and associative cerebral cortical areas. ICA was used to demonstrate which cerebellar regions were associated with which ICNs.

Functional Connectivity ROI-Based fcMRI of the Dentate Nucleus Functional connectivity (Allen et al. 2005) was found between the left dentate nucleus, and (1) the right DN, (2) the cerebellar cortex bilaterally (anterior and posterior vermis and hemispheres), (3) the thalamus bilaterally (ventral anterior, ventral lateral) but right dorsomedial nucleus, (4) the striatum (caudate nuclei bilaterally and right putamen), (5) the right limbic cortex (insula, hippocampus, and parahippocampus), (6) the right posterior cingulate cortex, (7) the right medial occipital lobe, (8) the inferior parietal lobe (BA 39/40), (9) the right (para-) cingulate cortex (BA 24/32), (10) the dorsolateral prefrontal cortex bilaterally (BA 8/9/46), and (11) the frontal pole (BA 10). Functional connectivity was found between the right dentate nucleus, and (1) the left DN, (2) the bilateral cerebellar cortex (anterior and posterior vermis and hemispheres) including the fastigial/globose nuclei, (3) the thalamus bilaterally (ventral anterior, ventral lateral, dorsomedial), (4) the left hypothalamus, (5) the striatum (caudate nucleus, putamen, and pallidum bilaterally), (6) the right insula (BA 13), (7) the anterior cingulate cortex bilaterally (BA 24) with a right predominance, (8) the left occipital cortex (BA 19), and (9) the dorsolateral prefrontal cortex (BA 9/46 with an extension to BA 10). Although fcMRI data provide no information about the directionality of the connectivity, some of these regions may correspond to mono- or disynaptic targets of the DN. In monkeys, the DN projects to the motor (BA 4), premotor (BA 6 medial, i.e., (pre-)SMA, and lateral), prefrontal (BA 9/46), and posterior parietal associative brain areas via the thalamus (ventral lateral and dorsomedial) (Strick et al. 2009). The DN also targets the striatum (Hoshi et al. 2005) and hypothalamus (Haines et al. 1997). Therefore, fcMRI of the DN may have functionally traced associative dentato-thalamo-cortical, dentato-striatal, and dentato-hypothalamic circuits. The remaining areas functionally connected with the DN in this study could be polysynaptic (two or more) relays or phylogenetically new relays of these circuits. It cannot be ruled out that these regions also send afferents to DN and the overlying cerebellar cortex. However, these cerebral afferents reach the cerebellum via a relay in the PN (and bulbar olivary nucleus

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and reticular nuclei), which were not detected in this study. This could argue in favor of a preferential detection of functional connectivity of the cerebellar output channel. Alternatively, this lack of detection of the PN may be explained by a threshold problem, low sensitivity technique (1.5 T), or low spatial resolution.

ROI-Based fcMRI of the Cerebellar Cortex The first cerebellar fcMRI study was performed by He et al. (2004) examining connectivity between the anterior inferior cerebellum and the rest of the brain. However, no exact location of the cerebellar seed region was provided and, when referring to the figure, this region seems to be located inside the cerebellar white matter. Therefore, this study was not included in the current chapter.

ROI-Based fcMRI of the Cerebral Cortex Krienen and Buckner (2009) and O’Reilly et al. (2009) defined an anatomical parcellation of the cerebellar cortex based on their specific coherence with distinct cortical ROIs and using the probabilistic cerebellar atlas of Diedrischsen et al. (2009). Krienen and Buckner found correlations between the dorsolateral prefrontal cortex and lobule VII (crus I and II and VIIB, especially crus II); the medial prefrontal cortex and lobule VII (crus I); and the anterior prefrontal cortex and lobules VI and VII (crus I/II/VIIB/VIIIA) (Fig. 26.2). O’Reilly and colleagues corroborated these observations by showing that motor, premotor, and somatosensory cortices were correlated with the cerebellar anterior lobe (lobules V/VI/VIII), whereas the prefrontal cortex was functionally connected with the posterior lobe (paravermal and lateral hemisphere of lobule VIIA including crus I and II). Strong correlations were also detected between visual area MT and lobules V/VI/VIII; superior temporal gyrus including auditory areas with lobules V/VI; and inferior posterior parietal cortex with lobules VIIA (paravermis) and crus II. Therefore, two main zones were distinguished in the cerebellum: (1) a primary sensorimotor zone (lobules V/VI/VIII) containing overlapping sensory and motor domains in relation to somatomotor, visual, and auditory cortices, and (2) a supramodal zone (lobule VII) containing overlapping cognitive domains in relation to prefrontal and parietal cortices. The latter associative areas were also mapped in this study and comprise the posterior frontal medial gyrus (BA 8), the middle medial gyrus (BA 9/46), the frontal pole (BA 10), the inferior parietal lobule (BA 39), the medial superior parietal lobule (BA 7b), and the posterior cingulate cortex (BA 25). Moreover, the cerebellar supramodal zone can be further segregated according to its functional links with dorsolateral, medial, and anterior prefrontal cortex. It is noteworthy that the functional connectivity between prefrontal and parietal cortices and neocerebellum is not restricted to oculomotor regions such as frontal and parietal eye fields or vermian and paravermian parts of lobule VI/VII but rather involves multiple regions in prefrontal cortex, parietal cortex, and neocerebellum. Therefore,

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Fig. 26.2 Functional anatomic parcellation of the human cerebellum based on resting-state functional connectivity using ROIs located in motor cortex (MOT), dorsolateral prefrontal cortex (DLPFC), medial prefrontal cortex (MPFC), and anterior prefrontal cortex (APFC). Caudal view (top). Rostral view (middle). Dorsal view (bottom) (From Krienen and Buckner 2009; with authors’ permission)

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it cannot be claimed that prefronto/parieto-cerebellar interconnections exclusively relate to the oculomotor system (Doron et al. 2010). More recently, Buckner et al. (2011) seeded small regions within the cerebellum in order to precisely determine the functionally correlated topography in the cerebral cortex during resting state. In particular, they established that the cerebellum contains at least two topographically organized, inverted representations of the complete cerebrum, with the exception of primary visual and auditory cortices. The cerebral cortex, including somatomotor, premotor, and association areas, is functionally linked to (1) a homotopic map extending from the somatomotor anterior lobe to crus I and II and (2) a mirrorimage secondary map extending from crus I and II to lobule VIII. If crus I and II, in association with part of lobule VI, and lobules VIIB and IX, are in functional coherence with the association cortex, the border of crus I and II displayed strong

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correlations with the default-mode network. It was also found that the somatomotor map in the anterior lobe (lobules IV/V/VI) represents the foot, hand, and tongue in the rostral-to-caudal axis and was located close to the vermis. Therefore, medial lobule VI belongs to the somatomotor zone, while the lateral part of this lobule takes part in the supramodal zone.

ICA fcMRI The above-mentioned studies provide a functional anatomic parcellation of the cerebellar cortex and show correlation of the neocerebellum (lobule VII) with the associative neocortex. However, most of these ROI-based studies did not examine correlations between neocerebellum and other parts of the brain and thus could not identify and segregate all the relays contributing to distinct specialized cerebroneocerebellar networks. ICA is a method that examines whole-brain connectivity without the a priori identification of an ROI. Habas et al. (2009) applied ICA analysis to resting-state fMRI data and used an unbiased template-matching procedure to identify previously studied ICNs: (1) the default-mode network (DMN) (Greicius et al. 2003, 2004) involved in stream of consciousness, mental imagery, episodic memory retrieval, and self-reflection (Raichle et al. 2001); (2) the executive control network (ECN; divided by ICA into left and right ECNs) involved in working memory, attention, response selection, and flexibility (Seeley et al. 2007); (3) the salience network (Seeley et al. 2007) required for the processing and integration of interoceptive, autonomic, and emotional information; and (4) the sensorimotor network (Biswal et al. 1995). Distinct cerebellar contributions were found in each of these ICNs. The neocerebellum was shown to participate in (1) the DMN (lobule IX), (2) the right and left ECNs (crus I and II with a narrow extension in lobules VIIB and rostral IX), (3) the salience network (lobule VI with narrow extension in lobules VIIA: crus I and II, and VIIB), and (4) the sensorimotor network (lobules V and adjacent VI) (Fig. 26.3). Three other structures of the cerebrocerebellar system were also identified: the PN (DMN, ECN, salience network), the DN (sensorimotor, salience networks), and the red nucleus (sensorimotor, DMN, ECN, salience network) (Fig. 26.4). These results are in agreement with O’Reilly et al. (2009) and Krienen and Buckner’s (2009) data, highlighting functional connectivity of lobule VII (especially crus I and II) with dorsolateral and dorsomedial (BA 9/46) prefrontal and cingulate cortices (ECNs), and frontoinsular cortex (salience network) (Fig. 26.4). The ICA results, however, provide four new pieces of data. First, the cerebellar supramodal zone can be extended to the caudal part of lobule VI, which is included in the salience network and can be clearly dissociated from the more anterior sensorimotor part of the same lobule and to lobule IX participating in the DMN. This is in line with O’Reilly et al. (2009) who also found a correlation between the tonsilla and the prefrontal cortex. Second, the major part of lobule VIIA is devoted to the ECNs. These results are in accordance with the meta-analysis of cerebellar neuroimaging studies (Stoodley and Schmahmann 2008) showing involvement of lobule VI/VII (crus I) in nonmotor linguistic, spatial, and

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Delineation of Cerebrocerebellar Networks

Fig. 26.3 Cortical (top) and cerebellar (bottom) regions functionally linked within five intrinsic connectivity networks (somatomotor, left and right executive control, default-mode, and limbic salience networks) as revealed by independent component analysis (Habas et al. 2009)

Abbreviations: DMPFC dorsomedian prefrontal cortex; DLPFC dorsolateral prefrontal cortex; H hemisphere; INS inferior parietal cortex; M1 motor cortex RSC/PCC retrosplenial cortex/posterior cingulate cortex; SMA supplementary motor cortex; SPC superior parietal cortex; THAL thalamus.

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Fig. 26.4 Human cerebellar topography based on restingstate functional connectivity to the frontal cortex (a) (from Krienen and Buckner 2009) and (b) to five intrinsically connected networks including somatomotor, left and right executive control, defaultmode, and limbic salience networks (from Habas et al. 2009). In a: (pre-)motor cortex (yellow), parietal cortex (green), dorsal prefrontal cortex (red), and medial prefrontal cortex (orange). In b: sensorimotor network (purple), defaultmode network (yellow), right and left executive control network (red and blue, respectively), and salience network (green). Sps, superoposterior sulcus. This figure demonstrates the substantial overlap in cerebellar functional connectivity maps obtained with ROI- and ICA-based parcellation of the cerebellum. Convergent findings include cerebellar regions connected to the sensorimotor network (yellow in a/purple in b), the executive control network (red in a/red and blue in b), the default-mode network (yellow in b, especially in lobules IX), and the salience network (orange in a/green in b)

executive processes. Third, an extracerebellar relay such as the red nucleus also contributes to nonmotor circuits. This is in keeping with Nioche et al. (2009) and supports the view that during phylogeny, not only the cerebellum but also its associated nuclei have evolved in parallel with the neocortex (Ramnani 2006). Fourth, ICNs encompass cerebellar input (PN) and output channels (DN) so that an ICN may indeed represent loops reciprocally linking cerebellum and cerebral cortex (Fig. 26.5).

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Tractography fcMRI enables identification of the neural relays contributing to a given specialized network but cannot distinguish which relays are directly connected by monosynaptic links. In other words, functional connectivity defined by fcMRI cannot distinguish direct connectivity between two regions from indirect connectivity possibly mediated by a third region. Two recent studies have attempted to overcome this drawback by using DTI-based tractography, a method that allows for reconstruction of white matter tracts that directly link two neural regions. Habas and Cabanis (2007), and Doron et al. (2010) applied tractography to track corticopontocerebellar fibers. They found that orbitofrontal, prefrontal, pericentral, and temporal/occipital cortices project onto specific PN, which, in turn, project via the middle cerebellar peduncle to the cerebellar cortex. More precisely, Doron et al. (2010) only described prefrontal corticopontine fibers arisen from caudal and medial superior frontal gyrus and

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Fig. 26.6 DTI-based deterministic tractography of the dentate nuclei showing, on a sagittal slice, dentatothalamo-cortical projections ending within the prefrontal (yellow fibers) and superior parietal (green fibers) cortices (From Jissendi et al. 2008; with authors’ permission)

a small region of the medial prefrontal gyrus. However, collaterals of mossy fibers from the PN to deep cerebellar nuclei could not be tracked. Ramnani et al. (2005) compared the organizational origins of corticocerebellar fibers in the cerebral peduncle (crus cerebri) between macaques and humans. This DTI study found a larger contribution to the crus cerebri from the prefrontal cortex in humans than in macaques. Within the cerebellum, the deep cerebellar nuclei, mainly the DN, are targeted by the PN (mossy fibers), bulbar olivary nucleus (climbing fibers), and the cerebellar cortex (Granziera et al. 2009). The DN are directly connected with the red nucleus and indirectly via the ventral part of the thalamus (Habas and Cabanis 2007; Granziera et al. 2009), with the cerebral cortex: sensorimotor (M1/S1), temporal (Habas and Cabanis 2007), prefrontal (BA 9), and parietal (BA 7) (Jissendi et al. 2008) cortices (Fig. 26.6). Altogether, these tractography studies confirm connections in humans between the neocerebellum and the associative cerebral cortex and, in particular, closed loops between the neocerebellum, including the DN, and prefrontal and parietal cortices. However, because of their low spatial resolution, their partial coverage of the brain, their low sensitivity to discriminate fiber crossings, and their inability to follow trajectories within low anisotropic regions, these studies may underestimate both the number of neocortical areas involved in the cerebrocerebellar system and also the number of loops. Therefore, fcMRI and tractography (i.e., functional and structural connectivity) are complementary in deciphering functional networks. Conclusion

fcMRI enables functional anatomic parcellation of the cerebellum into wellsegregated subregions which participate in specific, functionally distinct, largescale cerebrocerebellar networks. DTI tractography provides a complementary approach which can be used to help distinguish direct from indirect connections in the functional maps. Using these approaches, the cerebellar cortex can be subdivided into a polymodal sensorimotor zone (lobules IV/V/VI and VIII) and

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a supramodal cognitive zone (lobules VI/VII and, especially, crus I and II, and lobule IX). Subregions of the cognitive neocerebellum take part in the DMN (lobule IX), the salience network (lobule VI), and the right/left ECNs (crus I and II). Strong functional links exist between crus I and II and prefrontal, parietal, and cingulate cortices, supporting the role of the most phylogenetically recent part of the cerebellum in executive and affective functions. These intrinsically connected networks variably include the DN and PN. Lack of detection of the PN/DN in certain circuits could be ascribed to the stringent statistical postprocessing of the fcMRI data. It also cannot be ruled out that connectivity between the cerebellum and other parts of the brain may be mediated by the bulbar olivary, lateral vestibular, and reticular nuclei, as well as via the striatum (Hoshi et al. 2005; Bostan et al. 2010). It is worth noting that no functional connectivity was detected for the posterior vermis, especially lobule VII involved in limbic emotional processing, regardless of the fcMRI method performed. Thus, while functional and structural connectivity approaches have helped us begin to delineate the functional anatomy of the cerebellum, there is still much work to do.

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Ramnani N, Behrens TEJ, Johansen-Berg H, Richter MC, Pinsk MA, Andersson JLR, Rudebeck P, Ciccarelli O et al (2005) The evolution of prefrontal inputs to the corticopontine system: diffusion imaging evidence from macaque monkeys and humans. Cereb Cortex 16:811–818 Schmahmann JD (1991) An emerging concept: the cerebellar contribution to higher function. Arch Neurol 48:1178–1187 Schmahmann JD (2004) Disorders of the cerebellum: ataxia, dysmetria of thought, and the cerebellar cognitive affective syndrome. J Neuropsychiatry Clin Neurosci 16:367–378 Schmahmann JD, Pandya DN (1997) The cerebrocerebellar system. Int Rev Neurobiol 41:31–60 Schmahmann JD, Shermann JC (1998) The cerebellar cognitive and affective syndrome. Brain 121:561–579 Seeley WW, Menon V, Schatzberg AF, Keller J, Glover GH, Kenna H, Reiss AL, Greicius MD (2007) Dissociable intrinsic connectivity networks for salience processing and executive control. J Neurosci 27:2349–2356 Shirer WR, Ryali S, Rykhlevskaia E, Menon V, Greicius MD (2011) Decoding subject-driven cognitive states with whole-brain functional connectivity patterns. Cereb Cortex 22:158–165 Skudlarski P, Jagannathan K, Calhoun VD, Hampson M, Skudlarska BA, Pearlson G (2008) Measuring brain connectivity: diffusion tensor imaging validates resting state temporal correlations. Neuroimage 43:554–561 Stoodley CJ, Schmahmann JD (2008) Functional topography in the human cerebellum: a metaanalysis of neuroimaging studies. Neuroimage 44:489–501 Strick PL, Dum RP, Fiez JA (2009) Cerebellum and nonmotor function. Annu Rev Neurosci 32:413–434 Tedesco AM, Chiricozzi FR, Clausi S, Lupo M, Molinari M, Leggio MG (2011) The cerebellar cognitive profile. Brain 134:3669–3683 Thier P, Haarmeier T, Treue S, Barash S (1999) Absence of a common functional denominator of visual disturbances in cerebellar disease. Brain 122( Pt 11):2133–2146 Timmann D, Daum I (2010) How consistent are cognitive impairments in patients with cerebellar disorders. Behav Neurol 21:1–21 van den Heuvel MP, Mandl RCW, Kahn RS, Pol HEH (2009) Functionally linked resting-state networks reflect the underlying structural connectivity architecture of the human brain. Hum Brain Mapp 30:3127–3141 van Dijk KR, Hedden T, Venkataraman A, Evans KC, Lazar SW, Buckner RL, (2010) Intrinsic functional connectivity as a tool for human connectomics: theory, properties, and optimization. J Neurophysiol 103:297–321 Vincent JL, Pratel GH, Fox MD, Snyder AZ, Baker JT, Van Essen DC, Zempel JM, Voogt J (2003) The human cerebellum. J Chem Neuroanat 26:243–252 Whiting BA, Barton RA (2003) The evolution of the cortico-cerebellar complex in primates anatomical connections predict patterns of correlated evolution. J Hum Evol 44:3–10

Radiographic Features of Cerebellar Disease: Imaging Approach to Differential Diagnosis

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O. Rapalino, Robert Chen, and R. G. Gonzalez

Abstract

The cerebellum shares many neuroanatomical, histochemical and neurophysiological features with other CNS structures and many generic processes affecting the cerebellum will exhibit similar imaging features as seen in supratentorial and brainstem regions. The cerebellum also has many particular features that are reflected in its selective involvement by multiple pathological processes with characteristic imaging features. The purpose of this chapter is to provide the reader with a general overview and differential diagnosis of the pathological processes involving the cerebellum with similar imaging features as seen in other CNS structures as well as describing particular conditions more selectively involving the cerebellum with more specific radiological appearance.

Normal CT and MR Appearance of the Cerebellum The cerebellar gray and white matter have x-ray attenuation and magnetic relaxation properties similar to the brainstem and supratentorial gray and white matter. On CT, gray matter is slightly hyperdense in comparison with the white matter. On MRI, gray matter, in relation to white matter, is also hyperintense on T2-weighted sequences and hypointense on T1-weighted images. The dentate

O. Rapalino (*) • R.G. Gonzalez Neuroradiology Division, Department of Radiology, Massachusetts General Hospital, 55 Fruit Street, 02114 Boston, MA, USA e-mail: [email protected] R. Chen Emergency Imaging Division, Department of Radiology, Massachusetts General Hospital, 55 Fruit Street, 02114 Boston, MA, USA e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 587 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_27, # Springer Science+Business Media Dordrecht 2013

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Fig. 27.1 Normal MR appearance of the cerebellum. Sagittal T1 (a) and axial T2 (b) weighted images demonstrate normal cerebellar volume and signal intensity arising from the gray and white matter. Notice that the cerebellar gray and white matter signal is similar to the gray and white matter arising from the brainstem and supratentorial brain. The susceptibility weighted sequence (c) demonstrates normal hypointense signal arising from the dentate nuclei

nuclei show evidence of progressive mineralization with advancing age, best seen on high-field MR imaging (Fig. 27.1). Specific normative data of ADC and FA values of cerebellar structures have been previously published showing a significant variability of these values among different cerebellar anatomic structures and between the cerebellum, brainstem, and supratentorial brain (Lee et al. 2009; Yoon et al. 2006). There are also significant regional differences in metabolite ratios within the cerebellum and between the cerebellum and the rest of the brain (Minati et al. 2010; Mascalchi et al. 2002; Costa et al. 2002; Jacobs et al. 2001), with the cerebellum showing increased absolute creatine levels and decreased

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Fig. 27.2 Unilateral cerebellar hypoplasia. Axial (a) and coronal (b) T2-weighted images demonstrate left-sided cerebellar volume loss when compared to the normal right side. Given the asymmetric nature of disease, an acquired insult to the cerebellum in utero was suspected

related NAA/Cr ratio(s) in comparison with the supratentorial gray matter and thalamus (Minati et al. 2010; Mascalchi et al. 2002; Costa et al. 2002; Jacobs et al. 2001; Safriel et al. 2005).

Decreased Cerebellar Volume: Is It Hypoplasia or Atrophy? A central conundrum in the radiological evaluation of cerebellar disease is the differentiation between cerebellar hypoplasia (suggestive of an in utero developmental process) (Figs. 27.2–27.5) and atrophy (more often related to an acquired disorder). Both conditions will produce reduced cerebellar volumes, but the presence of prominent cerebellar fissures and progressive volume loss will indicate cerebellar atrophy (Table 27.1) (Figs. 27.6, 27.7, 27.8).

Developmental Disorders Cerebellar developmental disorders can be classified based on their extent of anatomic involvement. Unilateral cerebellar disorders are typically acquired and result from specific insults to the developing cerebellum in utero, whereas bilateral disorders are typically congenital. Bilateral disorders can be subdivided into abnormalities with central predominance (midline or vermian malformations) or conditions with more diffuse involvement (Ten Donkelaar and Lammens 2009). Unilateral cerebellar hypoplasia can be secondary to in utero ischemia or

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Fig. 27.3 Cerebellar agenesis. Multiplanar T2-weighted HASTE sequences (a, b, c) were performed in utero, illustrating CSF signal intensity filling the posterior fossa. No cerebellar tissue is seen

hemorrhage selectively involving one of the cerebellar hemispheres. Cerebellar agenesis is a rare condition and the end of the spectrum of bilateral disruption of cerebellar development (Boyd 2010; Huissoud et al. 2009) (Fig. 27.2). Bilateral cerebellar hypoplasia may be related to genetic disorders or toxins (e.g., phenytoin) (Ten Donkelaar and Lammens 2009) (Figs. 27.3, 27.4, 27.5). There are different types of congenital malformations affecting the cerebellum, some of them already characterized at the molecular level (Table 27.2). The spectrum of congenital cerebellar abnormalities also include the Chiari malformation subtypes (Table 27.3) and certain cystic abnormalities of the posterior fossa (Table 27.5).

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Fig. 27.4 Bilateral cerebellar hypoplasia. Sagittal T2 (a), axial T2 (b), and coronal T1-weighted SPGR (c) sequences demonstrate significant volume loss involving both cerebellar hemispheres compatible with hypoplasia

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Fig. 27.5 Cerebellar hypoplasia associated with lissencephaly and Dandy-Walker variant. Sagittal T1 (a) and axial T2 (b, c) weighted images demonstrate some residual vermian tissue. There is normal positioning of the torcular herophili and a normal-sized posterior fossa, which distinguishes it from the more “classic” and severe Dandy-Walker malformation

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Table 27.1 Differentiation between cerebellar hypoplasia and atrophy Imaging feature Hypoplasia Cerebellar volume Decreased Cerebellar fissure (size) Normal to decreased Temporal evolution Static

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Atrophy Decreased Enlarged (increased) Static or progressive

Fig. 27.6 Mild cerebellar atrophy with nonprogressive cerebellar ataxia. Sagittal T1 (a) and axial T2 (b, c) weighted images illustrate mild increase in the size of the cerebellar fissures. Compare this with normal cerebellar volumes seen in Fig. 27.1b

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Fig. 27.7 Infantile neuronal ceroid lipofuscinosis. Sagittal T1 (a), axial T2 (b), and coronal T2-weighted images. There is evidence of increased T2 signal and atrophy of the cerebellum

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Fig. 27.8 Hereditary spastic paraplegia. Axial T2-weighted (a) and coronal T1-weighted (b) images demonstrate mild to moderate cerebellar atrophy. MR spectroscopy (c) shows mild decrease of NAA/Cr ratios compatible with neuronal injury/dysfunction

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Congenital Malformations of the Cerebellum Table 27.2 Congenital malformations affecting the cerebellum Condition Pathogenesis Key features Molar tooth malformations • Postsegmentation “Molar tooth” (Joubert syndrome–related malformation appearance of disorders, JSRD) (Alkan associated with midbrain, midline et al. 2009; Leao et al. 2010; ciliary dysfunction vermian cleft and Parisi 2009) (Fig. 27.9) dysplasia, thick • Involved genes: horizontal superior INPP5E, AHI1, cerebellar NPHP1, CEP290, peduncles, lack of TMEM67/MKS3, decussation of RPGRIP1L, superior cerebellar ARL13B, and fibers, central CC2D2A pontine tracts, and corticospinal tracts; batwing-shaped fourth ventricle, brainstem nuclear abnormalities Dandy- Classic DandyAbnormal Partial or complete Walker Walker development of vermian agenesis, complex malformation anterior and cystic enlargement or (DWM) (Alkan posterior of fourth ventricle, spectrum et al. 2009; Blaser membranous and posterior fossa 2007; Niesen 2002) portions of enlargement with (Figs. 27.10, 27.11) rhombencephalic superior roof (area displacement of the membranea) torcula herophili. (Niesen 2002) or Rarely associated fourth ventricular with fourth outlet foramina ventriculocele (Ten Donkelaar and Lammens 2009) Multiple genes linked, including Zic1 and Zic4 (Blank et al. 2011) Hypoplastic vermis Abnormal Variable hypoplasia with rotation (HVR) development of of cerebellar (Dandy-Walker anterior and vermis. Absence of posterior fossa variant) (Wong posterior enlargement or et al. 2012) membranous fourth ventricular (Fig. 27.12) portions of cystic dilatation rhombencephalic roof

Associated features Polymicrogyria, ocular and renal findings, hepatic fibrosis, ventriculomegaly, corpus callosum dysgenesis

Hydrocephalus, cortical dysplasia, heterotopia, abnormal myelin development, corpus callosum dysgenesis, callosal thinning, occipital cephalocele, septum pellucidum agenesis

(continued)

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Table 27.2 (continued) Condition Cerebellar hypoplasia (Figs. 27.4, 27.5)

Rhombencephalosynapsis (Alkan et al. 2009) (Figs. 27.13, 27.14)

Pathogenesis Key features Proliferation defects

Midbrain-hindbrain Vermian agenesis dorsoventral and fusion of patterning defect cerebellar hemispheres Focal cerebellar dysplasia Defects of late maturation and cortical organization Congenital muscular dystrophy Postsegmentation Cerebellar cyst-like (FMCD), Walker-Warburg malformation: changes, syndrome, muscle-eye-brain alpha-dystroglycan pontomedullary disease (MEB disease) glycosylation malsegmentation, defects polymicrogyria, delayed myelination Chiari I–IV malformation Tonsillar herniation (Figs. 27.15, 27.16, 27.17, associated with 27.18, 27.19, 27.20) variable degrees of dysmorphic features of the posterior fossa structures Lhermitte-Duclos disease Rare mass-like (LDD) (dysplastic abnormality gangliocytoma of the compatible with cerebellum) (Fig. 27.21) a cerebellar hamartoma

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Associated features Often with pontine hypoplasia, pontomedullary segmentation anomaly Septum pellucidum agenesis, corpus callosum dysgenesis

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Fig. 27.9 Joubert syndrome. Sagittal T1 (a), axial T2 (b), and coronal T2 (c) weighted images demonstrate a dysplastic appearance of the cerebellar hemispheres. The classic molar tooth configuration used to describe the appearance of the midbrain and superior cerebellar peduncles is best demonstrated on axial images (Fig. 27.10b), where the superior cerebellar hemispheres are elongated in the anteroposterior dimension. Additionally, the fourth ventricle demonstrates a “batwing appearance”

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Fig. 27.10 Dandy-Walker malformation. Sagittal T1 (a), axial T2 (b), and coronal T1-weighted SPGR (c) sequences reveal enlargement of the posterior fossa, elevation of the torcular herophili, and absence of normal vermian tissue. The fourth ventricle communicates with a large posterior fossa cyst. The supratentorial brain demonstrates marked macrocephaly with cerebral dysplasia and hydrocephalus

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Fig. 27.11 Dandy-Walker malformation. Sagittal (a) and axial (b, c) T2-weighted images demonstrate marked enlargement of the posterior fossa. Only a small amount of vermian tissue is present, and the fourth ventricle communicates with the enlarged retrocerebellar “cyst.” Note the elevated torcular herophili

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Fig. 27.12 Hypoplastic vermis with rotation (HVR) (previously known as Dandy-Walker variant). Sagittal T1 (a), coronal T1 (b), and axial T2 (c) weighted sequences demonstrate some residual vermian tissue superiorly. There is normal positioning of the torcular herophili and a normal-sized posterior fossa, which distinguishes it from the more “classic” and severe Dandy-Walker malformation

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Fig. 27.13 Rhombencephalosynapsis. Sagittal T1 (a), axial T2 (b), and coronal postcontrast T1 (c) weighted images demonstrate apparent fusion of the cerebellar hemisphere and absence of a normal cerebellar vermis, with continuous white matter tracts crossing the midline

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Fig. 27.14 Rhombencephalosynapsis. Companion case. Sagittal T1 (a), axial T2 (b), and coronal postcontrast T1 (c) weighted images also demonstrating fusion of the cerebellar hemispheres, with continuous white matter tracts crossing the midline and apparent absence of the cerebellar vermis

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Fig. 27.15 Chiari I. Sagittal T1 (a), sagittal phase-contrast (b), and axial T2 (c) weighted sequences demonstrate a “peg-like” configuration of the cerebellar tonsils, which have descended below the level of the foramen magnum by greater than 5 mm. Phase-contrast images demonstrate lack of flow-related signal within the posterior portion of the foramen magnum (b)

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Fig. 27.16 Chiari I. Sagittal T1 (a), axial T2 (b), and coronal T1 postcontrast (c) weighted images demonstrate a more severe example of cerebellar tonsillar descent, with the tonsils extending caudally to the C3 level and posterior to the upper cervical cord (a)

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Fig. 27.17 Chiari II. Sagittal T1 (a), axial T2 (b), and coronal T1-weighted SPGR (c) sequences reveal an elongated appearance of the brainstem and a pointed configuration of the cerebellar tonsils which protrude through the foramen magnum by greater than 5 mm. While the beaked configuration of the tectum is poorly seen secondary to crowding of the posterior fossa contents, the supratentorial abnormalities are well appreciated, which in this case included polymicrogyria and callosal dysgenesis

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Fig. 27.18 Chiari II malformation. Sagittal T1-weighted (a), axial T2-weighted (b), and axial CISS sequence (c) demonstrate cerebellar tonsillar herniation, a dysplastic and elongated configuration of the midbrain and the rostral aspect of a cervical syrinx

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Fig. 27.19 Chiari III. Sagittal T2 (a), axial T2 (b), and coronal T2 (c) weighted images demonstrate severe dysgenesis of the brainstem and cerebellum with a small posterior encephalocele, compatible with a Chiari III malformation

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Fig. 27.20 Chiari III. Scout image (a) and two axial (b, c) noncontrast CT images. Another case with demonstration of a prominent occipital encephalocele, dysgenesis of the brainstem, and low-lying cerebellar tonsils

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Fig. 27.21 Lhermitte-Duclos abnormality. Axial noncontrast CT (a), coronal CT (b), sagittal precontrast T1 (c), axial T2 (d), axial FLAIR (e), and coronal postcontrast T1 (f) demonstrate an irregular mass-like abnormality within the right cerebellar hemisphere. There is a thickened and striated appearance to the cerebellar folia, with associated mass effect upon the brainstem and fourth ventricle

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Chiari Malformations Spectrum Dr. Hans Chiari, an Austrian pathologist, first described the first two types of this spectrum of craniocervical malformations (Vannemreddy et al. 2010; Pillay et al. 1991; Chiari 1987; Tubbs and Cohen-Gadol 2010). There are now at least four wellrecognized conditions in this spectrum, ranging from cerebellar tonsillar herniation as seen with Chiari I malformation to complete agenesis of the cerebellum in Chiari IV. Table 27.3 Chiari malformations Condition Frequency Chiari I (Figs. 27.15, Frequent 27.16)

Chiari “1.5” (Tubbs et al. 2004)

Rare

Chiari II (Naidich et al. 1983) (Figs. 27.17, 27.18)

Rare

Chiari III (Sirikci et al. Very rare 2001; Aribal et al. 1996; Castillo et al. 1992) (Figs. 27.19, 27.20) Chiari IV Very rare

Key features Herniation of cerebellar tonsils (5 mm below the foramen magnum) Peg-like configuration of cerebellar tonsils

Herniation of cerebellar tonsils and brainstem through the foramen magnum (Tubbs et al. 2004) Caudal descent and herniation of cerebellar tonsils, fourth ventricle, and brainstem. Myelomeningocele, elongated midbrain with tectal beaking (due to collicular fusion), elongated pons, and medullary kinking

Low occipital or high cervical encephalocele Chiari II malformation features and widened foramen magnum Marked cerebellar hypoplasia or aplasia, no hindbrain herniation

Associated features Syringomyelia (50–75%) (Vannemreddy et al. 2010), hydrocephalus (3–10%) (Vannemreddy et al. 2010), basilar invagination and other craniocervical osseous anomalies (Vannemreddy et al. 2010) Syringomyelia (50%) (Tubbs et al. 2004)

Low-lying torcular herophili, hydrocephalus (90%), asymmetric ventricles, interhemispheric cysts (Wong et al. 2009), corpus callosum agenesis, absent septum pellucidum, polymycrogyria and cortical dysgenesis, colpocephaly, “beaten copper” appearance of skull vault, scalloping changes of skull base, craniocervical osseous anomalies, split cord malformations

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Cerebellar tonsillar hernitation is not a phenomenon unique to the Chiari malformations and also be seen with other disorders, including pathologies associated with decreased volume of the posterior skull base, increased mass effect or hypotension syndrome (Table 27.4).

Differential Diagnosis of Cerebellar Tonsillar Herniation Table 27.4 Conditions associated with cerebellar tonsillar herniation (Milhorat et al. 2010) Mechanism Conditions Developmental osseous restriction (Milhorat Chiari I malformation et al. 2010) Craniosynostosis Achondroplasia Acromegaly Paget’s disease Disproportionate cerebellar overgrowth Costello syndrome (Gripp et al. 2010; Tubbs and (Gripp et al. 2010; Martinez-Glez et al. 2010; Oakes 2003) Conway et al. 2007) Macrocephaly-capillary malformation (M-CM) (Martinez-Glez et al. 2010; Conway et al. 2007) Basilar invagination (Milhorat et al. 2010) Collagen disorders with occipitoatlantoaxial (atlantoaxial impaction) joint instability/hypermobility (Milhorat et al. 2007) Posttraumatic occipitoatlantoaxial joint instability Osteogenesis imperfecta Spinal cord tethering (Milhorat et al. 2010) Tethered cord syndrome Lumbosacral myelomeningocele (as seen with Chiari II malformation) Intracranial hypertension (Milhorat et al. 2010) Hydrocephalus Intracranial mass or hemorrhage (particularly in the posterior fossa) Intracranial/intraspinal hypotension syndrome CSF leak Dural ectasia

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Cystic Pathologies in the Posterior Fossa Table 27.5 Cystic abnormalities in the posterior fossa (Yildiz et al. 2006) Condition Pathophysiology and key features Associated features Dandy-Walker Variable cerebellar vermian Hydrocephalus (90%) (Yildiz malformation agenesis, cystic dilatation of et al. 2006) (Figs. 27.10, 27.11) fourth ventricle, and enlarged posterior fossa with rostral displacement of tentorium (Yildiz et al. 2006) Dandy-Walker variant Variable cerebellar vermian (Fig. 27.12) hypoplasia and prominent retrocerebellar cystic space freely communicating with a normal-size or mildly enlarged IV ventricle. No posterior fossa enlargement (Yildiz et al. 2006) Blake’s pouch cyst Infravermian cyst communicating Failure of regression of Blake’s with the fourth ventricle. Inferior pouch and fenestration of local mass effect on adjacent foramen of Magendie (Yildiz vermis or medial cerebellar et al. 2006). Hydrocephalus hemispheres. Normal position of tentorium Megacisterna magna Fluid collection in direct continuity with cisterna magna and without local mass effect Arachnoid cyst CSF fluid collection surrounded by (communicating and arachnoid layers. Localized fluid noncommunicating collection producing local mass types) (Fig. 27.22) effect on the adjacent cerebellar parenchyma

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Fig. 27.22 Arachnoid cyst. Sagittal T1 precontrast (e), axial T2 (b), and axial FLAIR (c) weighted images illustrate a large extra-axial posterior fossa mass superior to the right cerebellar hemisphere, which causes scalloping of the adjacent brain parenchyma. It is isointense to CSF on all precontrast pulse sequences and does not demonstrate any appreciable enhancement after contrast administration (a, d, f). The lesion is classic for an arachnoid cyst

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Acquired Disorders The cerebellum can also be affected by as broad spectrum of ischemic/hemorrhagic, infectious, inflammatory, metabolic and neoplastic processes as well as pathophysiological charges seen with functional nuroimaging studies. The following tables will provide a general overview and differential diagnosis of these pathologies (Table 27.6–Table 27.14).

Vascular, Ischemic, and Hemorrhagic Pathologies Table 27.6 Vascular pathologies Condition Hemorrhagic risk Capillary telangiectasia Rare (Guibaud et al. 2003; (Fig. 27.23) Bland et al. 1994; McCormick et al. 1993) Developmental venous Rare (McCormick et al. 1993; anomaly (Lee et al. 1996; Garner et al. 1991; Kovacs Garner et al. 1991; Wilms et al. et al. 2002, 2007; Kondziolka 1990; Senegor et al. 1983) et al. 1991) (Figs. 27.24, 27.25)

Dural arteriovenous malformations/fistulas (Fig. 27.26)

Arteriovenous malformations (Fig. 27.27) Cavernous angiomas (Fig. 27.28)

Key features Enhancing focus with similar features to other anatomic locations Occasionally associated with areas of gliosis or venous congestion (Fenzi and Rizzuto 2008) Confluent veins in a caput medusae configuration (Wilms et al. 1991) Rare (Satoh et al. 2001) Clustered vessels often associated with parenchymal areas of venous congestion or infarct (Lee et al. 2003; van Dijk and Willinsky 2003; Fujita et al. 2002) 2–3% per year (Wilkins 1985) Similar features to other anatomic sites 2.39% per year (Cantu et al. Similar features to other 2005) anatomic sites Radiation-induced cavernous hemangiomas have a more benign evolution (Lew et al. 2006)

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Fig. 27.23 Capillary telangiectasia. Axial T2 (a) and axial postcontrast T1 (b) illustrate faint brush-like enhancement within the right cerebellar hemisphere without corresponding T2 signal abnormality

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Fig. 27.24 Small developmental venous anomaly. Axial T2 (a), axial T1 postcontrast (b), and coronal postcontrast T1 (c) weighted images show the classic “caput medusae” like enhancement within the left medial cerebellar hemisphere, converging upon a single draining vein. A small punctate focus of corresponding T2 hyperintensity is seen

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Fig. 27.25 Large developmental venous anomaly. Axial (a), coronal (b), and sagittal (c) postcontrast MIP images from a CTA, as well as an axial source CTA image, reveal a large collection of small veins converging upon a larger venous trunk in the left cerebellar hemisphere. DVAs are typically associated with cavernous malformations

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Fig. 27.26 Arteriovenous fistula. Axial noncontrast CT (a), sagittal MIP from CTA (b), and digital subtraction angiogram (c, d) demonstrate a Cognard type IIA tentorial AVF with arterial feeders from bilateral superior cerebellar arteries with the right side dominant in supply. Venous drainage is via two cerebellar veins which drain into the straight sinus and torcula herophili. Notice the retrograde flow is only into the transverse and sigmoid venous sinuses and not the cortical veins, differentiating it from a Cognard type IIb; also, note the simultaneous opacification of the arterial and venous system, which is necessary to diagnose an arteriovenous fistula

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Fig. 27.27 Cerebellar arteriovenous malformation. Noncontrast axial CT demonstrates lobular hemorrhage within the right aspect of the brainstem as well as slightly hyperdense lobulations within the right cerebellar hemisphere (a). Postcontrast axial CT images (b, c) illustrate a large tangle of arterial and venous vessels within the right cerebellar hemisphere, corresponding to enlarged flow voids seen on the axial T2-weighted MR sequence (d). Digital subtraction angiogram (e) indicates the dominant arterial supply to the cerebellar AVM was via enlarged superior cerebellar arteries

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Fig. 27.28 Cavernous angioma. Axial T1 (a) weighted image reveals faint regions of intrinsic T1 hyperintensity in the region of the vermis, with small areas of enhancement (b). Axial T2 (c) and FLAIR (d) weighted sequences demonstrate corresponding T2 hypointensity, with marked blooming on the axial (e) and coronal (f) gradient echo sequences secondary to blood products. This is a typical appearance for a cavernous angioma

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Cerebellar Ischemic Conditions Table 27.7 Hypoxic/ischemic pathologies affecting the cerebellum Conditions Main features Arterial infarcts (Lin et al. 2008; Terao et al. Typically artery-to-artery embolism (33–60%) or 2005; Caplan 2005) (Figs. 27.29, 27.30, 27.31) cardioembolism (20–23%) (Terao et al. 2005; Kumral et al. 2005; Chaves et al. 1994) Hemorrhagic infarcts in 13% of cases (Terao et al. 2005) Most commonly involving the PICA (60%) and SCA (40%) territories (Terao et al. 2005; Min et al. 1999; Marinkovic et al. 1995) Venous infarcts (Fig. 27.32) Secondary to venous thrombosis in the posterior fossa, predominantly the straight and sigmoid sinuses (Ruiz-Sandoval et al. 2010) Arterial dissection Variable clinical presentation with infarcts, pseudoaneurysm formation, or subarachnoid hemorrhage (Sedat et al. 2007; Han et al. 1998; Mascalchi et al. 1997) Most commonly involving the vertebral artery, PICA, or SCA Vasculitis (Murakami et al. 2010; McLean Small peripheral infarcts suggestive of medium et al. 1993) and small size arteries (Provenzale and Allen 1996) Posterior reversible encephalopathy syndrome Cerebellum is affected in 33–53% of cases, more (Fig. 27.33) often in patients with autoimmune disorders (Fugate et al. 2010; Donmez et al. 2010; McKinney et al. 2007) Hypoxic-ischemic encephalopathy Cerebellar vermian involvement (particularly, its (Figs. 27.34, 27.35) anterior portion) has been described with severe hypoxia (Connolly et al. 2007; Chao et al. 2006; Sargent et al. 2004) Neurosyphilis (Fig. 27.36) Meningovascular syphilis can rarely involve the cerebellar arteries (Umashankar et al. 2004)

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Fig. 27.29 Right posterior inferior cerebellar artery infarction. Axial diffusion-weighted sequences (a, b, c) show marked DWI hyperintensity and restricted diffusion within the right cerebellar hemisphere in the distribution of the right posterior inferior cerebellar territory, with marked mass effect causing partial compression of the fourth ventricle and shift of the vermis to the left. Notice the sparing of the anterior and superior cerebellar vascular territories

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Fig. 27.30 Superior cerebellar artery infarction. Axial diffusion-weighted sequences (a, b, c) show areas of DWI hyperintensity and restricted diffusion within the right cerebellar hemisphere in the distribution of the superior cerebellar artery vascular territory. The PICA and AICA vascular distributions were spared

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Fig. 27.31 Anterior inferior cerebellar artery infarction. Axial diffusion-weighted sequences (a, b) reveal DWI hyperintensity within the right cerebellar hemisphere in the distribution of the anterior inferior cerebellar artery vascular territory. The SCA and PICA vascular distributions were spared

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Fig. 27.32 Cerebellar venous infarction. Axial DWI (a) and FLAIR (b) weighted images illustrate abnormal hyperintense signal within the right posterior cerebellar hemisphere. While this could conceivably represent an infraction within the distribution of the PICA, a CT venogram (c) revealed a hypodense filling defect within the right transverse and sigmoid sinuses, clarifying the etiology of this infarction as venous in origin

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Fig. 27.33 Posterior reversible encephalopathy syndrome (PRES). Axial FLAIR (a, b) and T2 (c) weighted images demonstrate hyperintense foci of signal abnormality within the cortical and subcortical white matter of the parieto-occipital lobes as well as the cerebellar hemispheres. PRES is thought to be an abnormality of autoregulation and typically affects the posterior circulation

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Fig. 27.34 Moderate hypoxic-ischemic injury. Axial DWI (a, b), axial T2-weighted (c), and axial CT without contrast (d). Extensive areas of abnormal DWI and T2 hyperintensity in the cerebral hemispheres with relative preservation of the cerebellum. The cerebellum is relatively resistant to global hypoxia-ischemia

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Fig. 27.35 Severe hypoxic-ischemic injury with cerebellar involvement. Axial DWI (a, c), axial T2 (b), and axial FLAIR (d) sequences. There are extensive areas of abnormal DWI and T2 hyperintensity in the cerebral and posterior cerebellar hemispheres

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Fig. 27.36 Neurosyphilis. DWI (a, b, c), coronal T1-weighted images (d, e), and axial FLAIR images. Scattered infarcts are seen in the DWI sequences and foci of leptomeningeal enhancement compatible with meningovascular neurosyphilis

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Trauma and Hemorrhagic Conditions Table 27.8 Traumatic and hemorrhagic pathologies affecting the cerebellum Parenchymal contusions and Cerebellar contusions typically secondary to falls (Jagodzinski hematomas (Fig. 27.37) et al. 1990). Cerebellum typically spared with high-speed motor vehicle trauma (Kirkpatrick 1983; Sato et al. 1987). Hematomas 24 h), and the majority (60%) suffer combined audiovestibular dysfunction (Lee et al. 2009). The prognosis for isolated AICA infarction is excellent with no deaths or severe disability at 3 months among 12 patients reported by Toghi et al. (1993).

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Fig. 28.7 AICA infarct. Diffusion MRI at the mid-pontine level reveals restricted diffusion diffusely involving the left middle cerebellar peduncle (asterisk) and the adjacent cerebellar hemisphere. There is variability in the extent of AICA cerebellar supply. The left AICA/PICA borderzone (dotted line) is shifted toward the PICA territory in this patient with a small left PICA. Note the two tiny foci of restricted diffusion in the contralateral cerebellar hemisphere

Superior Cerebellar Artery Infarction In a study of 60 patients with MRI confirmed SCA infarcts, lesions involved the lateral SCA cerebellar territory in 23%, the medial territory in 15%, combined medial and lateral territories in 15% (Fig. 28.8), cortical or deep borderzone territories in 30%, and combined SCA and other vascular territories in 17%; the latter commonly associated with coma and the rostral basilar syndrome (Kumral et al. 2005a). As with PICA infarction, large vessel atherosclerotic disease is more common than cardioembolism, with vertebral dissection the least common (Kumral et al. 2005a). Classically medial SCA infarcts have prominent truncal ataxia and mild limb ataxia, which is reversed with lateral SCA infarcts (Sohn et al. 2006). Cerebellar dysarthria is thought to arise from involvement of the rostral paravermian area (Urban et al. 2003), but may be seen in both lateral and medial SCA infarction (Sohn et al. 2006). The rostral trunk of the SCA supplies the vermian and paravermian surface, while the caudal trunk supplies the lateral hemispheric territory. These branches, in addition to the main SCA trunk, provide direct perforating branches to the cerebral peduncles, midbrain, and colliculi (Rhoton 2000). The marginal branch of the SCA supplies the superior petrosal cerebellar surface, anastomosing frequently with the rostral trunk of the AICA and providing perforators to the middle cerebellar peduncle (Rhoton 2000). The hemispheric branches of the SCA also anastomose with the hemispheric branches of the PICA. Compared to PICA infarcts, significant swelling from large SCA infarcts is thought to be less common (7% in one study (Kase et al. 1993)). Despite this fact, SCA infarcts were found to have worse outcomes than PICA or AICA strokes

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Fig. 28.8 SCA infarct. Diffusion MRI at the level of the superior cerebellar peduncles (asterisk) demonstrates acute infarct in the left SCA territory. The contralateral normal SCA territorial supply is outlined (dotted line)

with 61% rate of independent outcome and 13% rate of severe disability or death at 3 months (Tohgi et al. 1993). This may relate to the greater likelihood of SCA infarcts to involve the midbrain. If patients with infarction in the brainstem, thalamus, and occipital lobes are excluded, Kumral et al. found that 96% of SCA infarcts produced no or minor disability (Kumral et al. 2005a).

Hemorrhagic Diseases of the Posterior Fossa Aneurysmal Disease Saccular Aneurysms Saccular aneurysms occur at vessel branch points. Vertebrobasilar (VB) aneurysms account for 15% of saccular intracranial aneurysms, approximately 60% of which reside at the basilar terminus (BT) (Rhoton 2002). Posterior communicating artery (PCOM) aneurysms are generally considered as posterior circulation aneurysms. In the International Study of Unruptured Intracranial Aneurysms (ISUIA) (Wiebers et al. 2003) which included 4,060 patients, 828 patients had posterior circulation aneurysms, of which 42% were located at the PCOM, 34% were at the BT, and the remaining 24% at vertebrobasilar locations other than the BT (Wiebers et al. 2003). Given the proximity to cranial nerves and the brainstem, unruptured posterior circulation aneurysms commonly present with cranial nerve palsy or brainstem compressive symptoms. An oculomotor palsy, occurring in 20% of PCOM aneurysms (Golshani et al. 2010), may also be caused by posteriorly directed BT aneurysms compressing the oculomotor nerve as it enters the interpeduncular

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Fig. 28.9 Basilar apex aneurysm. Antero-posterior (Towne’s view) (a) and lateral (b) digital subtraction angiography of a 6 mm basilar apex aneurysm (arrow). The aneurysm projects posteriorly into the interpeduncular fossa which is well demonstrated on the CT angiogram (c). The aneurysm likely impinges on the left oculomotor nerve at its exit from the midbrain. The patient presented with an acute onset of diplopia from partial left oculomotor palsy

fossa (Fig. 28.9), or by superior cerebellar artery or posterior cerebral artery aneurysms as the oculomotor nerve passes in between these vessels (Ciceri et al. 2001; Rhoton 2002; Al-Khayat et al. 2005a; Peluso et al. 2007b). An acute presentation of oculomotor palsy suggests a morphological change in the aneurysm, and can herald impending rupture (Golshani et al. 2010). Similarly facial or vestibulocochlear nerve palsy may occur with AICA aneurysms, and lower cranial nerve palsies with PICA aneurysms (Peluso et al. 2007a, 2008). Unruptured posterior circulation aneurysms have a higher risk of rupture than anterior circulation aneurysms of the same size. According to ISUIA, 5-year cumulative rupture rates are 2.5%, 14.5%, 18.4%, and 50% for posterior circulation aneurysms less than 7, 7–12, 13–24, and 25 mm or greater, respectively, compared to 0%, 2.6%, 14.5%, and 40% for similar anterior circulation aneurysms (Wiebers et al. 2003). Moreover, posterior circulation aneurysms have worse clinical outcomes following rupture. Ruptured VB aneurysms have up to 68% mortality within the first 48 h, compared to 23% for anterior circulation aneurysms (Schievink et al. 1995). Similarly, patients with ruptured VB aneurysms have poorer clinical grades if they survive to hospital admission (Schievink et al. 1995). Based on ISUIA data, posterior circulation location is a predictor of poorer clinical outcome after both surgical and endovascular treatment (Wiebers et al. 2003). In many centers, posterior circulation aneurysms are preferentially managed endovascularly (Sanai et al. 2008). The International Subarachnoid Aneurysm Trial (ISAT) (Molyneux et al. 2005) demonstrated better neurological outcomes after endovascular coiling compared to microsurgical clipping for ruptured cerebral aneurysms equally suitable for either treatment. Of note, posterior circulation aneurysms were underrepresented in the ISAT trial because most centers considered it unethical to randomize such patients due to perceived high surgical risk, particularly with BT aneurysms (Molyneux et al. 2002). In a meta-analysis of 2,568 unruptured cerebral aneurysms treated surgically, of which 395 were in the posterior circulation, morbidity and mortality for non-giant

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(1 mM) are required for DSE/DSI (Brenowitz and Regehr 2003), which may not readily occur under physiological conditions. However, much smaller calcium increases are necessary if the signal is prolonged (Brenowitz et al. 2006), suggesting that patterns of activity that promote sustained dendritic calcium elevations may be important for DSE/DSI in vivo.

Synaptically Evoked Suppression of Excitation (SSE): Local Endocannabinoid Signaling Brief high-frequency (50–100 Hz) bursts of PF activity can evoke eCB release from PCs that transiently suppresses synaptic strength (SSE, Fig. 39.4d–f) (Brown et al. 2003). Here, glutamate released by PFs activates AMPA receptors, which locally depolarize dendrites and open voltage-gated calcium channels, and type-1 metabotropic glutamate receptors (mGLUR1s) that are coupled to Gq (Fig. 39.4e). Elevations of calcium and Gq activation are particularly effective at stimulating PLCb4 to produce DAG (Ohno-Shosaku et al. 2002; Hashimotodani et al. 2005). From this point SSE and DSE share a common mechanism (Maejima et al. 2001, 2005). Because PLCb4 is involved in SSE but not DSE, much smaller postsynaptic calcium increases are required for SSE than for DSE (Brenowitz and Regehr 2005). In contrast to DSE/DSI (Fig. 39.4c), SSE is restricted to synapses active during the PF burst because of localized mGluR1 activation, spatially restricted dendritic calcium increases, and limited extracellular spread of eCBs (Fig. 39.4f) (Brown et al. 2003; Brenowitz and Regehr 2005). ä Fig. 39.4 Global and local endocannabinoid signaling in Purkinje cells. (a–c) Brief depolarization of PCs from a holding potential of 60 mV to 0 mV robustly reduces synaptic strength that recovers less than a minute after the depolarization (DSE/DSI, A). Postsynaptic VGCCs are opened during the depolarization, activating an unidentified calcium-sensitive molecule that produces DAG. DAGLa then produces 2-AG that retrogradely activates CB1Rs on PF boutons, which reduce the probability of vesicular release. (c) Because dendritic calcium influx is widespread, eCBs are released globally. (d–f) Brief, high-frequency PF stimulation also reduces synaptic strength for tens of seconds (SSE, D). (e) Glutamate activates postsynaptic AMPA receptors (AMPARs) and Gq-coupled group I metabotropic glutamate receptors (mGluRs). Calcium influx through VGCCs and Gq synergistically activates PLCb4 that produces DAG. The remaining steps are the same as in (b). (f) Unlike DSE/DSI eCB release in SSE is restricted to activated synapses during induction

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Importantly, SSE can be influenced by the precise timing of CF activity (Brenowitz and Regehr 2005). PF-SSE is greatly enhanced when the CF is activated just following a PF burst (a 100 ms delay), but strong enhancement is not observed when the order is reversed. Regulation of dendritic calcium signaling is important for associative SSE. Paired CF and PF activity causes a robust calcium elevation localized near the PF input that far exceeds the linear sum of individual PF and CF calcium increases. The precise mechanism underlying the supralinear calcium increases in associative SSE remains unclear. Calcium release from internal calcium stores is not required for associative SSE, suggesting that the mechanism involves local regulation of membrane conductances (Martina et al. 2007) or the localized saturation of endogenous calcium buffers (Maeda et al. 1999). Given the importance of the CF as an error signal, eCB signaling may play a crucial role in converting CF activity into short-term associative SSE in vivo.

Long-Term Depression (LTD) In addition to the importance of eCB signaling in short-term depression at PF to PC synapses, eCBs are also involved in the induction of LTD at PF to PC synapses (PC-LTD) (Wang et al. 2000; Safo and Regehr 2005). When PF/CF pairing protocols similar to those used to induce associative SSE are repeated for several minutes, a form of eCB-mediated PC-LTD is induced. Like SSE, eCB-mediated PC-LTD is synapse-specific and requires postsynaptic calcium elevations, mGluR1 activation, and DAGL activity, implicating the involvement of 2-AG. Moreover, selective elimination of CB1Rs from PFs prevented LTD, indicating that retrograde eCB signaling is an important step in LTD induction (Carey et al. 2010). However, in contrast to eCB-mediated LTD in other brain areas (Gerdeman et al. 2002; Robbe et al. 2002; Chevaleyre and Castillo 2003; Sjostrom et al. 2003; Kreitzer and Malenka 2005) PF-LTD is expressed postsynaptically (Daniel et al. 1998; Ito 2001; Safo and Regehr 2005). It remains unclear how presynaptic CB1R signaling promotes postsynaptic LTD. One possibility is that CB1R activation stimulates nitric oxide (NO) production in PFs (Hillard et al. 1999). NO then diffuses into the postsynaptic cell where it is known to reduce the glutamate-sensitivity of AMPARs when combined with large postsynaptic calcium increases that can arise from conjunctive PF and CF stimulation (Shibuki and Okada 1991; Lev-Ram et al. 1997; Ito 2001). Future studies are required to clarify the precise role of eCB signaling in the induction of PC-LTD.

Long-Term Potentiation (LTP) Moderate frequency (8 Hz) trains of PF stimulation can evoke a presynaptic form of LTP at PF to PC synapses (PF-LTP) (Salin et al. 1996). PF-LTP is mechanistically similar to hippocampal mossy fiber LTP (MF-LTP) (Zalutsky and Nicoll 1990; Weisskopf et al. 1994) and involves calcium-stimulated adenylyl cyclase (Wayman

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Fig. 39.5 Endocannabinoid involvement in PF-LTP. (a) PF stimulation promotes calcium influx through R-type calcium channels, which activates calmodulin (CaM) and stimulates adenylyl cyclase (AC1/8) thereby elevating cyclic AMP (cAMP) levels. cAMP increases the probability of vesicular release. (b) When PFs and CFs are stimulated coincidently, eCBs are released from the postsynaptic cells that activate presynaptic CB1Rs, which inhibits cAMP production. Gbgmediated inhibition of R-type calcium channels is sufficient to block the calcium signal that stimulates AC1/8 (Myoga and Regehr 2011). Gai/o can also inhibit AC1/8 directly

et al. 1994; Storm et al. 1998; Wong et al. 1999; Wang et al. 2003) and elevations of presynaptic cAMP levels (Fig. 39.5a) (Chen and Regehr 1997). When 8 Hz PF stimulation is paired with CF activation, LTP is blocked in a CB1R-dependent manner (van Beugen et al. 2006). Here, eCBs are released from PCs during the paired protocol and activate CB1Rs on PFs. CB1R signaling inhibits adenylyl cyclase, thereby reducing the cAMP elevations that are produced by PF stimulation alone (Fig. 39.5b). There are multiple mechanisms that could mediate CB1R-dependent suppression of cAMP production. Gai/o could inhibit adenylyl cyclase directly. Alternatively, because the induction of PF-LTP involves calcium-dependent forms of adenylyl cyclase, Gbgi/o could reduce adenylyl cyclase activity indirectly by inhibiting voltage-gated calcium channels. It is known that presynaptic calcium influx in PF boutons is mediated by P-type, N-type, and R-type calcium channels (Brown et al. 2004). Moreover, blocking R-type calcium channels, but not other types of calcium channels that are present in PF boutons, prevents the induction of

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LTP (Myoga and Regehr 2011). These findings suggest that local calcium signals near R-type calcium channels activate adenylyl cyclase to induce LTP. Because CB1Rs inhibit R-type calcium channels (Brown et al. 2004), a reduction in calcium influx is sufficient to explain CB1R-mediated inhibition of PF-LTP. Moreover, other neuromodulators that regulate R-type calcium channels (Dittman and Regehr 1996) may also be important for controlling PF-LTP.

Endocannabinoid Signaling in Cerebellar Circuitry The flow of information from GrCs to PCs through the cerebellar cortex is regulated by interneurons (Palay and Chan-Palay 1974). GrCs cells make excitatory synapses onto molecular layer interneurons, which are subdivided into SCs and BCs. SCs are located in the distal molecular layer and inhibit PC dendrites, whereas BCs are located in the proximal molecular layer and inhibit PC cell bodies and axon initial segments. Together SCs and BCs provide feedforward inhibition to PCs and are important for controlling the precise timing of excitation in PCs. Golgi cells (GoCs) are located in the GrC layer and receive excitatory input from both mossy fibers and PFs. They provide feedback inhibition to GrCs and are important for controlling information entering the cerebellar cortex. eCB signaling has been implicated in both inhibitory circuits, but in distinct ways.

Parallel Fiber to Stellate Cell and Basket Cell Synapses SCs and BCs can also release eCBs to regulate the strength of their PF synaptic inputs. Briefly depolarizing SCs or BCs results in robust DSE (Beierlein and Regehr 2006) that shares a similar time-course and calcium dependence to DSE in PCs (Brenowitz et al. 2006; Myoga et al. 2009). Moreover, brief bursts of PF activity results in SSE (Beierlein and Regehr 2006). Interestingly, SSE in SCs is distinct from PCs because it depends on NMDA receptors as well as mGluR1. PF activity can also evoke LTD in SCs (SC-LTD) that is mediated by retrograde eCB signaling (Soler-Llavina and Sabatini 2006). The mechanism of eCB release in SC-LTD is distinct from PC-LTD in several ways. First, SC-LTD is dependent on calcium influx through calcium-permeable AMPA receptors in addition to the activation of mGluR1. Second, SC-LTD is expressed as a presynaptic reduction in glutamate release rather than a reduction in the glutamate sensitivity of AMPA receptors. These differences highlight the diversity of eCB signaling.

Parallel Fiber to Golgi Cell Synapses eCB signaling at PF to GoC synapses is very different from PF to SC synapses. Although these synapses are suppressed by the exogenous application of CB1R agonists, GoCs do not exhibit DSE or SSE (Beierlein et al. 2007). This suggests that

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GoCs do not readily release eCBs, but because CB1Rs are present on PF synapses onto GoCs these synapses could be suppressed by eCBs released by other types of cells. Differences in molecular machinery between GoCs and other cell types in the cerebellar cortex that underlie this difference remain unclear. However, the lack of eCB release from GoCs indicates that GrC activity is processed very differently by feedback and feedforward inhibitory circuits in the cerebellar cortex.

The Spread of Endocannabinoids in the Cerebellar Cortex The extent to which eCBs spread in extracellular space has important implications for the range of eCB signaling. Initial studies performed at room temperature suggested that eCBs could effectively diffuse from a cell because eCBs released by one PC could suppress synapses onto another PC tens of microns away (Vincent and Marty 1993). This distance is largely determined by temperature. At near-physiological temperature (34 C) DSE does not spread from one PC to another PC. Moreover, the synapse-specificity of SSE (Brown et al. 2003; Brenowitz and Regehr 2005) indicates that under physiological conditions the spread of eCBs is limited.

Endocannabinoid Signaling and Information Flow Through the Cerebellar Cortex To date, eCBs have been implicated at many stages of information flow through the cerebellar cortex (Fig. 39.6). PCs release eCBs that inhibit PF, CF, and inhibitory synaptic inputs, as well as the somata of neighboring SCs and BCs. SCs and BCs can release eCBs to inhibit their PF synaptic inputs. Although GoCs are ineffective at releasing eCBs, the synapses they receive may be sensitive to eCBs released by neighboring cells.

The Endocannabinoid System and Cerebellar Function The high levels of CB1R expression within the cerebellar cortex and the many physiological effects of cannabinoids on signaling within the cerebellum suggest that the eCB system is important for cerebellar function. This is supported by the finding that exogenous cannabinoids have profound effects on motor functions. Studies are also beginning to provide insight into the role of eCBs in cerebellardependent behaviors.

Human Studies In humans, exogenous cannabinoids such as THC can produce a number of motor effects that may involve the cerebellum. At socially relevant doses, humans display

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Fig. 39.6 Sites of endocannabinoid signaling in the cerebellar cortex. eCBs are released by PCs that inhibit their PF (a), CF (b), and inhibitory (c) synaptic inputs. Molecular layer interneurons (SCs and BCs) can also release eCBs that inhibit their PF synaptic inputs (d). eCBs released by PCs can additionally activate CB1Rs on SC dendrites and somata (e) that suppress spontaneous firing. Golgi cells (GoCs) cannot effectively release eCBs, but their PF synaptic inputs may be sensitive to eCBs released by other cells (f)

impairments in operating automobiles (Liguori et al. 1998) and airplanes (Janowsky et al. 1976). It is difficult to attribute these impairments to the cerebellum because these complex motor tasks require brain regions beyond the cerebellum. Studies of THC intoxication on simple motor tasks have returned inconsistent results (Heishman et al. 1990). However, a recent study found that the delay paradigm of eyeblink conditioning, which is thought to be exclusively cerebellar-dependent, is impaired in chronic marijuana users (Skosnik et al. 2008). These findings support a role for eCB signaling in cerebellar function, even though the mechanisms responsible for this impairment are unknown. Extensive studies of the CB1R antagonist rimonabant, a potential appetite suppressant and weight loss drug (Colleran and Pai 2007), revealed numerous side effects (Moreira and Crippa 2009), but motor impairments were not obvious.

Animal Models Knockout mice are beginning to provide insight into eCB signaling in cerebellardependent behaviors. Mice lacking mGluR1 (Aiba et al. 1994), Gaq (Offermanns et al. 1997), and PLCb4 (Miyata et al. 2001; Nakamura et al. 2004) have deficits in eyeblink conditioning and are also ataxic. These findings are consistent with a role for eCB synthesis in cerebellar function, but the behavioral deficits could reflect roles for these proteins in processes other than eCB signaling. Mice lacking CB1Rs exhibit largely normal motor performance (Zimmer et al. 1999). CB1R knockout mice have impaired delay eyeblink conditioning, which is compatible with findings

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from pharmacological studies (Kishimoto and Kano 2006). It is possible that an impairment of eCB-dependent LTD and LTP contributes to this deficit (Ito 1989; Safo and Regehr 2005; van Beugen et al. 2006; Schonewille et al. 2010). The lack of severe cerebellar-dependent behavioral impairment suggests that either the roles of eCB signaling are subtle, or that compensatory mechanisms mask the normal roles of eCBs. Selective elimination of CB1Rs from select populations of cells with temporal control will be a powerful tool in future studies of eCB signaling and behaviors mediated by the cerebellum (Carey et al. 2010).

Endocannabinoid Signaling in Other Cerebellar Circuits This chapter has focused on eCB signaling in the cerebellar cortex, but immunohistochemical studies indicate that the proteins involved in eCB synthesis and both CB1Rs and CB2Rs are present in the vestibular nuclei, the deep cerebellar nuclei (DCN), and the IO (Suarez et al. 2008). The DCN integrates PC activity and ascending inputs from MFs and CFs, is the sole output of the cerebellum, and could be an additional important locus of eCB signaling in the cerebellum, although no functional studies have been published. Retrograde eCB signaling has been demonstrated in the principle neurons of the IO, which are the source of CFs (Best and Regehr 2008). Within the IO, serotonin activates Gq-coupled 5-HT2A receptors to promote eCB release and suppression of glutamatergic synaptic inputs. This form of heterosynaptic plasticity suggests a general role for interactions between the serotonergic and eCB systems elsewhere in the brain, and also provides evidence that eCB signaling may be important for gating error signals that enter the cerebellar cortex. Conclusion

Following the discovery that eCBs mediate retrograde inhibition in the brain, intense study has elucidated many important mechanisms of eCB signaling in the cerebellum. It is now known that 2-AG mediates most retrograde signaling in the cerebellum. eCB signaling can result in short-term and long-term changes in synaptic efficacy, which has important implications for cerebellar-dependent learning. Moreover, eCBs are released by distinct populations of cells, which is important for understanding the manner in which the eCB system controls information flow through the cerebellar cortex. There are, however, numerous important questions that remain. What determines the duration of eCB signaling? Understanding the mechanisms that control the duration and magnitude of eCB signaling is of great importance for the eCB system as a therapeutic target. How do presynaptic CB1Rs regulate postsynaptic PC-LTD? What are the functional consequences of eCB signaling in the DCN? Perhaps the most important question is, with such prominent expression of molecules involved in the production and detection of eCBs and such prominent regulation of synapses throughout the cerebellum, what is the contribution of eCB signaling in the cerebellum to behavior? Why is so little known about the behavioral function of eCB signaling in the cerebellum? Will

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more refined behavioral tests reveal effects of eCB signaling on motor function or motor learning, or do compensatory mechanisms account for the lack of behavioral phenotypes? New genetic and molecular tools and behavioral studies will help to provide answers to these questions, and lead to new insights into the functional consequences of the eCB system in vivo.

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Purinergic Signaling in the Cerebellum

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Mark J. Wall and Boris P. Klyuch

Abstract

Purinergic signaling is a highly complex and evolutionarily conserved mechanism of extracellular communication in the brain that is involved in many physiological and pathological functions. The complexity of the system stems from the multitude of purine receptor subtypes and the large number of potential endogenous purine receptor ligands (ATP, ADP, UTP, UDP, UDP-glucose, and adenosine) which can either be directly released or arise from extracellular metabolism (and thus are potentially controlled by a variety of metabolizing enzymes). The chapter summarizes data on purinergic signaling in the cerebellum. Although much work has defined purine receptor distribution and the cellular effects of purine receptor activation, relatively little is known about how and when purines are released in the cerebellum, the role of purinergic signaling in cerebellar circuits, and the importance of purines in cerebellar motor control.

Introduction Purines are important extracellular signaling molecules that mediate diverse physiological effects, via cell-surface receptors, throughout the brain and periphery (for review see Burnstock 2007). Purine receptors can be divided into two classes: P1 receptors, which bind adenosine and P2 receptors, recognizing primarily ATP, ADP, UTP, and UDP. ATP can be released, like a conventional neurotransmitter, by exocytosis from neurons (for review see Pankratov et al. 2006) and from glia (Montana et al. 2006; Pangrsic et al. 2007; but see Hamilton and Attwell 2010) (Fig. 40.1). ATP is often

M.J. Wall (*) • B.P. Klyuch School of Life Sciences, University of Warwick, Gibbet Hill, Coventry, CV4 7AL, UK e-mail: [email protected], [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 947 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_40, # Springer Science+Business Media Dordrecht 2013

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co-released from neurons with classical neurotransmitters such as glutamate (Pankratov et al. 2007), GABA (Jo and Role 2002), noradrenaline (Burnstock and Holman 1961), and acetylcholine (Nurgali et al. 2003) (for review see Pankratov et al. 2006; Burnstock 2009). ATP can also be released via hemichannels (Pearson et al. 2005; Huckstepp et al. 2010) and via P2X7 receptors (Suadicani et al. 2006) (Fig. 40.1). Both necrotic and apoptotic cells can release ATP and ATP can be

VGCC AMP AC

ADP Ado

ATP

HC

A1

ADO

ATP NPP NTPD

e5’-N

A AD

NT INO

ATP

ADO

AMP

ADP

P2Y

P2X

PLC

P1 AC GIRK

2+

Ca

VGCC

Fig. 40.1 ATP and adenosine production, release and mechanism of action. Diagram illustrating effects and production of ATP and adenosine (blue-presynaptic cell, brown-postsynaptic cell). ATP can be released from neurons or glia by several mechanisms including exocytosis and via the opening of hemichannels. The released ATP activates P2X receptors, which are ligand-gated ion channels and also activates P2Y receptors which are G protein-coupled receptors and signal via second messengers (leading to increases in intracellular Ca2+ concentration). The ATP is broken down to adenosine (by NTPDases) which can then activate P1 receptors. Adenosine can also be directly released by transport out of cells or by other mechanisms in response to neural activity. P1 receptors are G protein-coupled receptors which can signal via adenylyl cyclase and can have effects on Ca2+ channels and GIRK K+ channels in pre- and postsynaptic cells. Adenosine is removed by transport into neurons and glia (equilibrative and concentrative transporters) and by metabolism (to inosine by adenosine deaminase). AC adenylate cyclase, ADA adenosine deaminase, Ado adenosine, ADP adenosine diphosphate, AMP adenosine monophosphate, ATP adenosine triphosphate, e50 -N - ecto-50 -nucleotidase, GIRK G protein-coupled inwardly rectifying K+ channels, HC hemichannels, Ino inosine, E-NPP ectonucleotide pyrophosphatase/phosphodiesterase, NT nucleotide transporter, E-NTPD ectonucleoside triphosphate diphosphohydrolase, P1 first type of purinoceptors (adenosine receptors), P2X purinoceptors (ligand-gated ion channels), P2Y purinoceptors (G protein-coupled), PLC phospholipase C, VGCC voltage-gated Ca2+ channels

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released by mechanical stimulation (stress and shear) and following pathogen invasion. Following release (and P2 receptor activation), ATP is metabolized to ADP (which is still active at some P2Y receptors) then to AMP (which is inactive) and finally to adenosine which activates P1 receptors (Fig. 40.1). The adenosine is removed from the extracellular space either by specific transporter proteins (equilibrative and concentrative) or further broken down to inactive metabolites (inosine and hypoxanthine, Fig. 40.1). Although current dogma suggests that adenosine arises only from extracellular breakdown of ATP, there is gathering evidence that adenosine release can also occur directly via reverse transport and adenosine maybe directly released by neural activity (evidence reviewed below).

ATP (P2) Receptors P2 receptors are expressed throughout the brain and periphery and have been localized to both neurons and glia (including astrocytes, oligodendrocytes, and microglia). Fast responses to ATP are mediated by the activation of P2X receptors, which are ligand-gated ion channels. Seven P2X receptor subunits have been cloned (P2X1-7) and can produce homomeric and heteromeric P2X receptors. P2X receptors are permeable to monovalent cations such as Na+ and K+ and divalent cations such as Ca2+. Influx of Ca2+ via P2X receptors can occur at the resting potential (unlike the NMDA receptor, which is blocked by Mg2+) and thus can provide a source of Ca2+ entry which can influence both neurons and glia. For example, the activation of P2X receptors on nerve terminals increases glutamate release, an effect that may result from increased Ca2+ influx (Khakh and Henderson 1998). Members of the P2X receptor family show many pharmacological and functional differences depending on subunit composition including channel conductance, kinetics of activation, desensitization and deactivation and fractional Ca2+ permeability. In some P2X receptors (P2X2, 4 and 7), following prolonged activation with ATP, the channel pore “dilates” allowing permeability to large cations such as NMDG+ (N-methyl-D-glucamine). For full review of the molecular physiology of the P2X receptor family see Khakh 2001; Egan et al. 2006; and Jarvis and Khakh 2009. Metabotropic P2Y receptors are G protein-coupled receptors with seven transmembrane domains, an extracellular NH2 terminus, and an intracellular COOH terminus. Eight P2Y receptor subtypes have been cloned (P2Y1,2,4,6,11,12,13,14). The P2Y nomenclature lacks several members, reflecting either the existence of nonmammalian orthologs or receptors having P2Y receptor sequence similarity but no apparent response to nucleotides. Most P2Y receptors bind to a single heterotrimeric G protein, with for example, P2Y1,2,4,6 binding to Gq/11 but some can bind to multiple G proteins. P2Y receptors show significant differences in their pharmacological and functional profiles. For example, some P2Y receptors are activated principally by nucleoside diphosphates (P2Y1,6,12), others are activated mainly by nucleoside triphosphates (P2Y2,4), others by both purine and pyrimidine nucleotides (P2Y2,4,6), and some by purine nucleotides alone (P2Y1,11,12).

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Some P2Y receptors stimulate and others inhibit the enzyme adenyl cyclase, and activation of P2Y receptors can increase intracellular Ca2+ by an effect on intracellular stores (Fig. 40.1). Functionally P2Y receptors can inhibit or enhance synaptic transmission and can regulate ion channels (such as voltage-gated K+ and Ca2+ channels). Signaling via P2Y receptor activation is further complicated, as many cells express multiple P2Y receptor subtypes. (For a comprehensive review of P2Y receptors see Hussl and Boehm 2006; von Kugelgen 2006).

P2 Receptor Expression in the Cerebellum The expression pattern of P2 receptors in the cerebellum is highly complex (like most other brain regions) and its analysis is complicated by the variety of different techniques used to identify receptor subtypes: receptor protein (immunohistochemistry, ligand binding), receptor function (electrophysiology, Ca2+ imaging), and receptor mRNA (RT-PCR, in situ hybridization, etc.). Data interpretation is further complicated by studies that have investigated receptor expression in cerebellar cultures (glial, neuronal, and mixed), which may have different expression patterns compared to whole cerebellum. There may also be differential P2 receptor expression patterns depending on the age of animal (for example see Xiang and Burnstock 2005; Amadio et al. 2007). Table 40.1 outlines the putative distribution of P2 receptors in the major elements of the rodent cerebellum. Table 40.1 P2 receptor expression in rodent cerebellum. A table outlining the expression of P2 receptor subunits by the major cell types of the cerebellum. The numbers refer to the references below Stellate/ Receptor Granule Purkinje Golgi basket Bergmann subunit cells cells cells cells glia Astrocytes Others/comments P2X1 1PF, 2c 1 9 1 P2X2 2c, 3, 14 3, 4c, 3 3, 5 5 wk 3 3 DCN, 13(no RNA 5 wk, (peduncle) for P2X2 detected in 6c, 14 cerebellum) P2X3 2c, 8, 8, 9 7 synaptosomes, 23 15c (no RNA for P2X3 in cerebellum) P2X4 2c, 6, 2, 5str, 10 5 5 wk, 10 13 DCN 5str, 13 10, 13str wk wk, 14 P2X5 11 wk 11 12 11 DCN, 13 (no RNA for P2X5 in cerebellum) P2X6 13 wk 5str, 5 very wk 13 DCN 13str, 14 P2X7 2c 20c 7 synaptosomes, 21(no RNA for P2X7 in cerebellum) (continued)

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Table 40.1 (continued) Stellate/ Receptor Granule Purkinje Golgi basket Bergmann subunit cells cells cells cells glia Astrocytes Others/comments P2Y1 2c, 15c, 16str, 22 17 (P7) 17 (P7) 17 NA fibers, 22 16 wk, 22,17 Lugaro cells P2Y2 18 12 P2Y4 2c 12, 14, 9 P2Y6 2c, 15c P2Y11 P2Y12 2c P2Y13 19c, 20c str defined as strong labeling in the reference, wk defined as weak labeling in the reference, DCN deep cerebellar nuclei, PF parallel fibers, c in cell culture, P7 labeling in postnatal day 7 rats, NA fibers noradrenaline containing fibers 1. Loesch and Burnstock (1998) Cell tissue Res 294:253–260 (measured protein) 2. Herva´s et al. (2003) J Neurosci Res 73:384–399 (measured mRNA) 3. Kanjhan et al. (1999) J Comp Neurol 407:11–32 (measured protein and mRNA) 4. Mateo et al. (1998) J Neurosci 18:1704–1712 (Ca2+ rise and pharmacology) 5. Rubio and Soto (2001) J Neurosci 21:641–653 (measured protein) 6. Garcia-Lecea et al. (1999) Neuropharm 38:699–706 (Ca2+ rise and pharmacology) 7. Herva´s et al. (2005) Biochem Pharm 70:770–785 (measured protein and Ca2+ rises) 8. Se´gue´la et al. (1996) J Neurosci 16:448–455 (measured mRNA) 9. Donato et al. (2008) Cell Calcium 44:521–523 (effects on synaptic transmission, pharmacology and protein measured) 10. Leˆ et al. (1998) Neuroscience 83:177–190 (measured protein) 11. Guo et al. (2008) Neuroscience 156:673–692 (measured mRNA and protein) 12. Brockhaus et al. (2004) Eur J Neurosi 19:2221–2230 (effects on synaptic transmission and pharmacology) 13. Collo et al. (1996) J Neuroscience 16:2495–2507 (measured mRNA) 14. Halliday and Gibb (1997) J Physiol 504:51P (electrophysiology) 15. Amadio et al. (2002) Neuropharm 42: 489–501 (measured mRNA) 16. Mora´n-Jime´nez et al. (2000) Mol Brain Res 78:50–58 (measured protein) 17. Amadio et al. (2007) BMC Develop Biol 7:77 (measured protein) 18. Loesch and Glass (2006) Methods Mol Biol 326:151–162 (measured mRNA) 19. Carrasquero et al. (2005) Purinergic Signal 1:153–159 (Ca2+ rises and pharmacology) 20. Carrasquero et al. (2009) J Neurochem 110:879–889 (Ca2+ rises and pharmacology) 21. Yu et al. (2008) Brain Research 1194:45–55 (measured mRNA) 22. Saitow et al. (2005) J Neurosci 25,2108–2116 (Ca2+ rises and pharmacology) 23. Chen et al. (1995) Nature 377:428–431 (measured mRNA)

Functional Effects of P2 Receptor Activation in the Cerebellum Several studies have measured increases in intracellular Ca2+ following P2 receptor activation in Purkinje cells (Kirischuk et al. 1996; Mateo et al. 1998; Garcia-Lecea et al. 1999), granule cells (Herva´s et al. 2003), cerebellar astrocytes (Carrasquero et al. 2009), and Bergmann glia (Kirischuk et al. 1995; Piet and Jahr 2007). The rise in Ca2+ could stem from the release of Ca2+ from intracellular stores (P2Y receptor

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activation, e.g., Kirischuk et al. 1995) or a combination of intracellular and extracellular Ca2+ entry (P2X receptor activation Herva´s et al. 2003). There are only a few studies where whole cell or single channel P2X receptor currents have been recorded in cerebellar cells: Purkinje cells (Halliday and Gibb 1997), granule cells (Halliday and Gibb 1997), and granular layer astrocytes (Carrasquero et al. 2009). Whole cell ATP currents recorded in Purkinje cells are very small (seen as an increase in background current noise) suggesting low receptor expression (Sharonova IN, personal communication). This is supported by the small number of patches (pulled from Purkinje cells) which contain P2X receptor channels (Gibb AJ, personal communication). No P2X receptor–mediated synaptic current has been recorded in the cerebellum.

Synaptic Actions of ATP Application of ATP increases the frequency of spontaneous postsynaptic currents (sPSCs) recorded from Purkinje cells in rat cerebellar slices indicating a presynaptic action on neurons or synaptic terminals (Fig. 40.2) (Brockhaus et al. 2004). This effect starts around the second postnatal week and is coincident with P2 receptor expression (Casel et al. 2005). These effects are produced by ATP acting at P2 receptors and not the result of ATP being broken down to adenosine and activating P1 receptors which would be expected to reduce sPSC frequency. In the third postnatal week, application of PPADs (P2 receptor antagonist) decreases sPSCs frequency, suggesting that there is a tone of extracellular ATP within the tissue, which tonically increases sPSC frequency (Casel et al. 2005). This is supported by the actions of ARL67156 (an ecto-ATPase inhibitor active in the cerebellum, Wall et al. 2008), which increases sPSC frequency (presumably by increasing the concentration of extracellular ATP). ATP affects the frequency of both glutamatergic sPSCs and GABAergic sPSCs suggesting presynaptic effects on both parallel fibers and basket cells (Fig. 40.2). These effects appear to involve both P2X and P2Y receptors (suggested as P2X5 and P2Y2, Brockhaus et al. 2004). More recent experiments, using a combination of electrophysiology and immunohistochemistry, have further investigated which presynaptic receptors are present on basket cells (Donato et al. 2008). Activation of presynaptic P2X receptors (containing P2X3 subunits) results in Ca2+ entry leading to a presynaptic facilitation of GABA release. Basket cell terminals also appear to express P2Y4 receptors, which inhibit the release of GABA (Donato et al. 2008). P2Y receptors (probably P2Y1) have also been shown to produce a long-term enhancement of GABAA receptor sensitivity in Purkinje cells through a Gq-mediated increase in intracellular Ca2+ concentration (Saitow et al. 2005). P2Y receptors also increase the frequency and amplitude of spontaneous GABAA-mediated inhibitory postsynaptic currents (IPSCs) recorded in Purkinje cells by increasing the excitability of interneurons and Lugaro cells (Saitow et al. 2005). Overall the effects of P2 receptor activation on cerebellar circuit function are complex as there are inhibitory and excitatory effects on both excitatory and inhibitory neurons with pre- and postsynaptic mechanisms.

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Fig. 40.2 ATP modulates both glutamatergic and GABAergic spontaneous postsynaptic currents (sPSCs) recorded in Purkinje cells. Whole cell patch clamp recordings were made from Purkinje cells in cerebellar slices from 14- to 19-day-old rats. The sPSCs were pharmacologically isolated into glutamatergic and GABAergic and then the effects of ATP on sPSC frequency were measured. (a) Graph plotting the frequency of sPSCs against time. Blocking GABAergic postsynaptic currents with bicuculline (10 mM) inhibited > 90% of the sPSCs, showing most of the sPSCs are GABAergic. The frequency of the spontaneous glutamatergic PSCs, remaining in bicuculline, was enhanced by ATP (100 mM). (b) To illustrate the effects of ATP more clearly, the ATP-mediated increase in the sPSC frequency is shown in a trace normalized to the frequency in bicuculline before ATP application. (c) Blocking of glutamatergic sPSCs with NBQX (5 mM), to isolate GABAergic postsynaptic currents, had only minor effects on sPSC frequency. The frequency of the isolated spontaneous GABAergic PSCs, which remained in NBQX, was increased by ATP (100 mM). (d) The histogram summarizes data from a number of slices and shows that the frequency of both NBQX-resistant GABAergic sPSCs and bicuculline-resistant glutamatergic sPSCs was significantly increased by ATP; *P < 0.05, *P < 0.005 versus control (Taken from Brockhaus et al. 2004)

Role of ATP in Glial Signaling Bergmann glia form large gap junction-coupled networks within the molecular layer of the cerebellum and appear crucial for motor coordination (Shibuki et al. 1996).

M.J. Wall and B.P. Klyuch

Fig. 40.3 (continued)

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The activation of P2Y receptors in Bergmann glia leads to an elevation of intracellular Ca2+ (Kirischuk et al. 1995). Electrical stimulation in the molecular layer elicits two component Ca2+ transients in Bergmann glia; a very small fast component due to AMPA receptor activation and a slower component as a result of P2Y receptor activation (Piet and Jahr 2007). The source of the ATP is suggested to be molecular layer interneurons, which can be activated by either direct electrical stimulation or by the activation of mGluR1 receptors following distal parallel fiber stimulation (Piet and Jahr 2007). Other potential sources of ATP were excluded because they either do not express mGlu1 receptors (parallel fibers) or when activated with climbing fiber stimulation (Purkinje cells) no Bergmann glia Ca2+ transients were detected (Piet and Jahr 2007). Release of ATP from Bergmann glia was discounted as the Ca2+ transients were abolished with tetrodotoxin (blocking action potentials) showing a neuronal component is involved in the ATP release (Piet and Jahr 2007). However, there is currently no direct evidence that molecular layer interneurons release ATP. More recent in vivo experiments have shown spontaneous waves of Ca2+ spread between coupled Bergmann glia and also spread through groups of velate protoplasmic astrocytes in the granule cell layer (Hoogland et al. 2009; Nimmerjahn et al. 2009). These waves can be initiated by the local application of ATP (Fig. 40.3a, d although this effect diminishes with repeated application Fig. 40.3f, ä Fig. 40.3 ATP-triggered transglial calcium waves in vivo. (a) A transglial calcium wave evoked in vivo by ejection of ATP (pipette concentration: 1 mM, 10 ms) into the molecular layer. The dye Alexa 594 (red) was included in the pipette solution to allow visualization of the pipette and ATP ejection. Cells were bolus loaded with the calcium indicator, Fluo-5 F (calcium signal: green) and SR101 was used to fluorescently stain Bergmann glia (red). In initial frames, there was a spreading red signal (ATP perfusion), followed by an increase in the green signal illustrating a rise in intracellular [Ca2+] in Bergmann glia. (b) The ATP-evoked Ca2+ signal has an elliptical domain oriented along the PF (parallel fiber) axis. This is a similar orientation to spontaneously occurring calcium waves. (c) Ca2+ waves can be triggered at different imaging depths after ATP ejection at the same depth. (d) Activation of velate astrocytes in the granule cell layer after ATP ejection in lower third of the molecular layer, using G-CaMP2 to visualize intracellular [Ca2+]. ATP perfusion is visualized with Alexa 594 (red) and the increase in intracellular [Ca2+] is green. (To visualize transglial waves more clearly, a nonreplicating adenovirus AdEasy-1 was used to express the calcium-sensitive fluorescent protein G-CaMP2 in Bergmann Glia). (e) The reduction of ATPtriggered transglial signals by PPADS (500 mM) shows that the ATP-evoked increase in intracellular [Ca2+] is via P2 receptors. The graphs plot the peak Ca2+ response (normalized to control values before PPADs) against time for a single recording (top panel) and against condition (ATP or ATP/PPADs) for a number of recordings (bottom panel). (f) There is a marked decrease in successive calcium responses after repeated applications of ATP (interval 20 s). The decrease in ATP response probably results from the incomplete refilling of intracellular Ca2+ stores which is characteristic of P2 receptor-mediated [Ca2+] increases. (g) Top panel, dependence of Ca2+ response amplitude to pulses of ATP injected at different time intervals (20–120 s). The Ca2+ responses were normalized to the first response. Bottom panel, responses recovered to half the size of the first response in an estimated 27  9 s (rat, fluo-5 F), 30  6 s (mouse, fluo-5 F), and 37  4 s (mouse, G-CaMP2), consistent with refilling times for internal Ca2+ stores. These results are consistent with a mechanism in which point-like release of ATP acts on P2 receptors to trigger transcellular calcium release waves (Modified from Hoogland et al. 2009)

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g) and can be blocked by application of a P2 receptor antagonist (Fig. 40.3e, Hoogland and Kuhn 2010; Hoogland et al. 2009). The Ca2+ waves persist when action potentials are blocked (Tetrodotoxin, TTX) and when glutamate receptors are blocked (with g-D-Glutamylglycine, g–DGG). Thus parallel fiber activity may not be the physiological pathway for triggering ATP release from Bergmann glia in vivo (Nimmerjahn et al. 2009). Such Ca2+ waves, in large networks of coupled Bergmann glia, are activated during motor behavior in freely behaving mice and are suggested to initiate changes in brain dynamics and blood flow (Nimmerjahn et al. 2009). However, the consequences of these glial waves to cerebellum function are not known.

Extracellular Metabolism of ATP Once released into the extracellular space, ATP is metabolized to adenosine by several classes of enzymes called ectonucleotidases or ecto-ATPases (of which there are many subtypes, reviewed by Zimmermann 2000). Ecto-ATPases are glycoproteins that are located on the outer surface of plasma membranes of neural, glial, and endothelial cells. As with any other brain region, there is strong expression of these enzymes within the cerebellum, and application of ATP to cerebellar slices results in the production of adenosine (Fig. 40.1) (Wall and Dale 2007). EctoATPase activity (measured from liberation of Pi) has been localized within axodendritic, axoaxonic, and dendrodendritic granule cell appositions and is associated with pre- and postsynaptic membranes (Zinchuk et al. 1999). Antibody labeling of the ecto-ATPase CD39 (NTPDase 1 converts ATP directly to AMP without liberating ADP) labeled soma and dendrites of Purkinje cells, with very weak labeling of the granule cell layer (Wang and Guidotti 1998). Within the molecular layer, the enzyme ecto-50 -nucleotidase CD73 (which converts AMP to adenosine) has been localized within glial cells and within parallel and climbing fiber synapses, although there are changes in expression (from neuronal to glial) during postnatal development (Schoen et al. 1991). In mixed cerebellar cultures, ecto-50 -nucleotidase is localized on granule cell and glial membranes (Maienschein and Zimmermann 1996). The breakdown of ATP to adenosine in the cerebellum can be reduced by a number of enzyme inhibitors such as ARL 67156 (Wall and Dale 2007), Evans Blue (Wall and Dale 2007), POM-1 (Wall et al. 2008), and a,bmethylene-ADP (Wall and Dale 2007) which differ in their selectivity and potency. However, it has proven extremely difficult to block all the cerebellar metabolism of ATP to adenosine.

Adenosine Signaling in the Cerebellum Adenosine is an important neuromodulator, involved in a number of physiological and pathophysiological processes in the central nervous system and in peripheral organs such as the heart and muscles (Latini and Pedata 2001; Boison 2006). The presence of adenosine (P1) receptors, high-level expression of enzymes which

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either produce or metabolize adenosine (ecto-50 -nucleotidase and adenosine deaminase correspondingly) suggest an important role for adenosine in cerebellar function (Braas et al. 1986; Geiger and Nagy 1986; Zinchuk et al. 1999). In recent work, adenosine was shown to be released in the cerebellum during hypoxia or glutamatemediated excitotoxicity, suggesting a neuroprotective role (Fatokun et al. 2008; Atterbury and Wall 2009). Adenosine is also released during neural activity (see below Wall and Dale 2007).

Adenosine (P1) Receptors in the Cerebellum The specific actions of adenosine are mediated through cell-surface receptors divided into four subgroups: A1, A2A, A2B, and A3 (Fredholm et al. 2001). All adenosine receptors are G protein-coupled receptors which are either inhibitory (A1 and A3) or excitatory (A2A and A2B). Activation of A1 and A3 receptors inhibits the activity of adenylate cyclase resulting in a reduction of cAMP production. A1 receptors open K+ channels (G protein-coupled inwardly rectifying potassium channels) and close voltage-gated Ca2+ channels (leading to hyperpolarization and inhibition of transmitter release) (Fig. 40.1). Activation of A2A and A2B receptors increases adenylate cyclase activity which can facilitate transmitter release and play a role in blood vessel dilatation (Fredholm et al. 2001). More detailed information about the general role of adenosine in the brain can be found in excellent reviews (Fredholm et al. 2001; Fredholm 2010). The A1 receptor is the most abundantly expressed adenosine receptor in the cerebellum (Goodman et al. 1983; Rivkees et al. 1995). However, the precise expression of A1 receptors in the cerebellum is not completely clear. Based on in situ hybridization and immunohistochemistry, several groups have shown that A1 receptors are expressed by granule cells (both soma and axons (parallel fiber) terminals), basket cells, and deep cerebellar nuclei. In the molecular layer, the basket cells are the most heavily labeled cell type, while labeling of parallel fibers is moderate (Reppert et al. 1991; Rivkees et al. 1995). Purkinje cells show very light labeling and Golgi cells show no labeling at all. Studies using mutant mice (Weaver and Reeler which, either lack granule cells or have displaced granule cells respectively) suggested that A1 receptors are only expressed by granule cells and are preferentially located on parallel fiber terminals (Goodman and Synder 1982; Goodman et al. 1983). The functional effects of A1 receptor activation in the cerebellum have been extensively studied using in vitro brain slice preparations (Takahashi et al. 1995; Dittman and Regehr 1996; Wall and Dale 2007; Atterbury and Wall 2009). However, the first functional effects of adenosine in the cerebellum were measured in vivo by Kocsis and co-authors who showed that activation of A1 receptors reduces glutamate release at parallel fiber but not climbing fiber synapses (Fig. 40.4a, b) (Kocsis et al. 1984). In later work, using whole cell patch clamp recordings from Purkinje cells in slices, it was shown that climbing fiber synapses are also regulated by adenosine, but the level of inhibition is much less compared to

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Fig. 40.4 Synaptic action of adenosine at parallel and climbing fibre Purkinje synapses in vivo and invitro. (a) Parallel fibre-induced extracellular field potentials recorded in vivo before and after application of adenosine (100 mM). The large response is the fibre volley and the slower potential (arrow) is the potential produced by glutamatergic parallel fibre excitatory postsynaptic potentials (EPSPs). The application of adenosine reduced the slow potential (*) with little effect on the volley. (b) There is no appreciable change in the extracellular field potential induced by climbing fibre stimulation in vivo before (top) or after (bottom) adenosine application (1 mM). (c) Superimposed parallel fibre excitatory postsynaptic currents (EPSCs) recorded in vitro from Purkinje cells in cerebellar slices. The largest amplitude EPSC is in control and increasing concentrations of adenosine (1, 10, and 100 mM) produced greater reductions in EPSC amplitude. (d) Superimposed climbing fibre EPSCs recorded in vitro from Purkinje cells in cerebellar slices. The largest amplitude EPSC is in control and increasing concentrations of adenosine (3, 30, and 300 mM) produce greater reductions in EPSC amplitude. (e) The graph plots parallel fibre-Purkinje cell EPSC amplitude against time during application of different adenosine concentrations for a single experiment. (f) The graph plots climbing fibre-Purkinje cell EPSC amplitude against time during application of different adenosine concentrations for a single experiment. Field potentials (a and b) were recorded in vivo from 180–250 g rats. EPSCs (c and d) were recorded from Purkinje cells in cerebellar slices from 10–15 day old rats using whole cell patch clamp (a and b) modified from (Kocsis et al. 1984). (c, d, e and f) modified from (Takahashi et al. 1995)

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parallel fiber synapses (Fig. 40.4c, d) (inhibition with high concentrations of adenosine 31% vs 78%, Takahashi et al. 1995). The difference between in vivo and in vitro studies may stem from differences in adenosine metabolism and recording techniques (high-resolution patch clamp vs extracellular). Metabolism of adenosine is probably much greater in vivo and thus maybe no exogenous adenosine reached the receptors on climbing fiber terminals in the studies of Kocsis et al. 1984. The difference in inhibition between parallel fiber and climbing fiber synapses reported from in vitro cerebellar slices may stem from a difference in A1 receptor potency (concentration of adenosine producing half maximal suppression 1.1 mM at parallel fiber synapses vs 3.4 mM at climbing fibers synapses, Takahashi et al. 1995) or could stem from differences in adenosine metabolism. To distinguish between metabolism and receptor potency, experiments need to be repeated at the two synapses using a non-hydrolysable analogue of adenosine such as CPA (N6cyclopentyladenosine). The activation of A1 adenosine receptors on parallel fiber terminals produces a small reduction ( 10%) in the amplitude of the presynaptic volley without changing its shape. This suggests that Ca2+ influx is reduced by a direct inhibition of presynaptic Ca2+ channels rather than indirectly altering Ca2+ influx via a reduction in action potential duration (Dittman and Regehr 1996). At parallel fiber synapses (but not at climbing fiber synapses) block of A1 receptors with a selective antagonist (such as 8-CPT or DPCPX) increases the amplitude of excitatory postsynaptic events recorded with either extracellular or patch clamp recording demonstrating continual adenosine-mediated inhibition and the presence of an extracellular adenosine tone (Takahashi et al. 1995; Wall and Dale 2007). Adenosine biosensors (Llaudet et al. 2003) were unable to measure a tone of adenosine directly in cerebellar slices but were able to detect inosine and hypoxanthine which arise from adenosine metabolism (Wall et al. 2007). The tone of A1 receptor activation, as measured by the use of A1 receptor antagonists, is developmentally regulated: it is much reduced in the cerebellum from younger animals (P8-14) although the levels of A1 receptor expression do not appear to change during development. Besides the inhibition of glutamate release, adenosine has also been shown to inhibit GABA release at Golgi cell-granule cell synapses (Courjaret et al. 2009). The authors showed that activation of adenosine A1 receptors reduces both the phasic GABAergic inhibitory postsynaptic currents (IPSCs) and the tonic ambient GABA current in granule cells and slightly increases their excitability. This is probably a direct effect on the release of GABA from Golgi cells rather than a reduction in parallel fiber glutamate release onto Golgi cells (although the authors did not block excitatory transmission during their experiments). This conflicts with labeling studies (see above) where no expression of A1 receptors was found on Golgi cells. Unlike parallel fibers synapses, there is no tonic activation of A1 receptors at these synapses. As adenosine reduces both inhibition and excitation, the overall effects of adenosine on cerebellar circuit function is potentially complex (Courjaret et al. 2009). There is no immunohistochemical evidence for expression of A1 receptors by Bergmann glia cells. However, activation of A1 receptors does reduce extrasynaptic parallel fiber transmission to Bergmann glia (Bellamy 2007). Unlike the parallel

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fiber-Purkinje cell synapses, blocking A1 receptors fails to increase the amplitude of extrasynaptic currents in Bergmann glia cells suggesting no tonic activation of these A1 receptors under basal conditions, although recordings were made from slices from young rats (Bellamy 2007). Recent studies have suggested that adenosine signaling (via A1 receptor activation) can modulate other neuromodulatory pathways in the cerebellum. A1 receptors expressed in Purkinje cells inhibit mGluR1 currents (induced by agonist application). This inhibition appears to be a direct effect of the A1 receptor as it does not require G protein signaling (Tabata et al. 2007). There is also evidence that adenosine A1 receptors can form functionally interacting complexes with MGlu1a subunits in rat cerebellum (Ciruela et al. 2001). Radioligand binding studies (with the A3 receptor agonist [125I]AB-MECA) suggested low-level expression of the A3 receptor in the cerebellum (Jacobson et al. 1993). However, subsequent studies suggested that [125I]AB-MECA also labels A1 receptors (Shearman and Weaver 1997) and thus the actual expression of A3 receptors in the cerebellum is not clear. To date, there is no strong functional evidence for A3 receptor expression in the cerebellum. The cerebellum of rodents contains high levels of the A1 adenosine receptor, possibly the A3 receptor, but not the excitatory A2A or A2B receptors (Jarvis et al. 1989; Fredholm et al. 2001). Several papers report a protective role for A2A receptors in cultured granular neurons (Fatokun et al. 2007, 2008). However, the physiological relevance of A2A receptors in the intact cerebellum has not been shown. There may be different receptor expression patterns in cultured neurons or it is possible that earlier studies did not detect their expression.

Adenosine Release In the brain, adenosine release has been observed experimentally following various manipulations including hypoxia, ischemia, hypoglycemia, seizures, electrical stimulation, high-extracellular K+ concentrations, application of glutamate, prolonged wakefulness, or apoptosis (Latini and Pedata 2001; Boison 2006). This adenosine could arise from the extracellular breakdown of ATP (see above) or could be released directly (via transporters or by other mechanisms). The mechanism by which adenosine is released in the cerebellum is still not clear. However, there are several possible pathways for its extracellular production (Latini and Pedata 2001; Wall and Dale 2008).

Does the Extracellular Adenosine Tone Arise from Extracellular ATP Metabolism? The metabolism of extracellular ATP to adenosine is catalyzed by several classes of enzyme encoded by at least three different gene families collectively called ectonucleotidases (Zimmermann 2000). This is a major mechanism of extracellular adenosine production in many brain areas (Fredholm et al. 2001; Latini and

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Pedata 2001) and as outlined above, there are high levels of ectonucleotidase expression within the cerebellum. If the cerebellar adenosine tone results from extracellular ATP breakdown, then reducing ATP metabolism should lower the tone and increase parallel fiber EPSP amplitude. However, this was only observed in one third of slices, with no effect in the remaining slices (Wall and Dale 2007; Courjaret et al. 2009). This result could stem from the low potency of ectonucleotidase inhibitors (Wall et al. 2007) rather than the adenosine tone coming from sources other than ATP. Casel et al. (2003) suggest that a tone of extracellular ATP exists within synapses onto Purkinje cells as P2 receptor antagonists reduce sPSC frequency (see above). It has not been possible to measure this ATP tone with biosensors placed within cerebellar slices even with ecto-ATPase inhibitors such as ARL67156 (Wall and Dale 2007). This suggests that the ATP is either highly localized or present at a low concentration. Potential sources of ATP within the cerebellum include Bergmann glia (Hoogland and Kuhn 2010) and molecular layer interneurons (Piet and Jahr 2007). Inducible transgenic mice have been developed that express a dominant-negative SNARE domain selectively in astrocytes and thus cannot release ATP by exocytosis (Pascual et al. 2005). Such mice would be potentially useful for studying ATP release in the cerebellum, but penetration of the transgene is poor in Bergmann glia (Klyuch and Wall unpublished observations).

Release of Adenosine in Cerebellar Cultures In their pioneering work, Philibert and Dutton 1989 and Schousboe et al. 1989 showed, using cerebellar cultures (either mixed neuronal cultures or granule cell cultures) combined with HPLC analysis, that adenosine can be released by a high-extracellular K+ concentration (30–50 mM) or by 10–100 mM glutamate application. This adenosine does not appear to originate from extracellular ATP metabolism (Schousboe et al. 1989). The adenosine efflux is mostly Ca2+dependent and occurs from cerebellar neurons not astrocytes (Philibert and Dutton 1989; Schousboe et al. 1989). Schousboe and colleagues also reported that adenosine release had similar dynamics to glutamate release which might indicate that cerebellar granule cells co-release glutamate and adenosine together (Schousboe et al. 1989). In later work, high-K+-evoked adenosine release was reported to be only partly (about 50%) dependent on Ca2+ influx (Sweeney 1996). This difference in the mechanism of K+-evoked adenosine release might be explained by different techniques of culture preparation and culture age. A recent study has demonstrated that purine release from cerebellar cultures is critically dependent on culture age and composition (Wall et al. 2010). Granule cell cultures and cultures of cerebellar astrocytes do not release purines (in response to a depolarizing stimulus) whereas mixed neuron and glial cultures reliably release purines. These studies, on cerebellar cultures, suggest that there are at least two distinct mechanisms by which adenosine is released: firstly, Ca2+ dependent which could involve vesicular release (Philibert and Dutton 1989; Schousboe et al. 1989); secondly, Ca2+-independent adenosine release which might occur via equilibrative

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nucleoside transporters (Philibert and Dutton 1989; Sweeney 1996). The involvement of equilibrative nucleoside transporters (ENT) was suggested by using specific ENT antagonists such as dipyridamole (DPR) and S-(p-nitrobenzyl)-6thioinosine (NBTI). Both DPR and NBTI produced a concentration-dependent reduction in Ca2+-independent K+-evoked adenosine release in cerebellar neuron cultures (Sweeney 1996). The data from cultures suggest it is possible to release adenosine from cerebellar cells with a strong depolarizing stimulus. However, it is difficult to extrapolate data from cultures using such strong stimuli to what actually occurs in the intact cerebellum under physiological conditions.

Release of Adenosine in Cerebellar Slices Molecular Layer Stimulation In our laboratory, purine microelectrode biosensors (Dale et al. 2005) were used to show that adenosine can be directly released from cerebellar slices (Wall and Dale 2007, 2008). Focal electrical stimulation delivered to the molecular layer causes adenosine release in a frequency-dependent manner. Maximal adenosine release occurs at 20 Hz with little increase at higher frequencies. With a constant stimulation frequency, an increase in stimuli number increases adenosine release (Fig. 40.5a) (Wall and Dale 2007, 2008). To check if adenosine is directly released or results from extracellular ATP metabolism, simultaneous ATP and adenosine biosensor recordings were made (Fig. 40.5b). Even with a relatively high concentration of adenosine detected (up to 1 mM) and in the presence of ecto-ATPase inhibitors, no ATP was detected. Thus unless ATP is broken down so quickly it cannot be detected (which seems unlikely), then the source of adenosine is not extracellular ATP breakdown (Wall and Dale 2007). Similar results (release of adenosine by neural activity with no ATP detected) have been reported in the hippocampus and at the Calyx of Held (reviewed by Wall and Dale 2008). By using the sodium channel blocker, tetrodotoxin (TTX) and Ca2+-free physiological saline, adenosine release was shown to be both action potential and Ca2+dependent (Fig. 40.5c, d). Furthermore adenosine release was not reduced by blocking adenosine transport via ENT1 and ENT2 (with DPR/NBTI) (Wall and Dale 2007). Thus the release mechanism appears to involve exocytosis and occurs either by the direct release of adenosine by exocytosis or the release of an interposed transmitter which causes downstream adenosine release (Wall and Dale 2007). This interposed transmitter does not appear to be glutamate or a number of other transmitter candidates (Fig. 40.5c). In searching for the source of adenosine, it was suggested that parallel fibers which run along the molecular layer are the most likely source of adenosine. The position of the stimulating and recording electrodes, along a beam of parallel fibers, are ideal to stimulate and record release from parallel fiber (Wall and Dale 2007, 2008). Furthermore, activation of presynaptic receptors (GABAB mGluR4 and A1 receptors) caused a reversible reduction in adenosine release. High expression of A1 and GABAB receptors and unique location of mGluR4 receptors on parallel fiber

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terminals (Mateos et al. 1998) strongly suggest that adenosine release either occurs from parallel fibers or at least their activity is required (Wall and Dale 2007). Stimulation in the molecular layer releases enough adenosine to activate A1 receptors and inhibit transmitter release at the parallel fiber synapse. Thus, this potentially represents an important feedback mechanism for controlling neural activity.

Do Climbing Fibers Release ATP/Adenosine? Application of a high K+ solution to cerebellar slices results in a Ca2+-dependent increase in the extracellular concentration of adenosine. Climbing fibers maybe the

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source of this adenosine release, as their ablation greatly reduces the amount of adenosine released (Do et al. 1991). Inhibition of ecto-50 -nucleotidase by a,bmethylene-ADP and GMP also decreased this release of adenosine by 50–60%, suggesting that a significant proportion of adenosine was derived from the extracellular metabolism of ATP (Do et al. 1991). Thus it appears, from this report, that climbing fibers either directly release ATP or activate a cellular component which release ATP, although there have been no follow-up studies.

Breakdown and Uptake of Adenosine Once released, adenosine is either inactivated by metabolism to inosine and hypoxanthine (by the enzymes adenosine deaminase and purine nucleoside phosphorylase respectively) or translocated into neurons or glia by ENT1 and/or ENT2 (equilibrative transporters) or concentrative transporters (CNTs, for which there are currently no inhibitors). After internalization, adenosine is phosphorylated to AMP (by adenosine kinase), maintaining low levels of intracellular adenosine and thus preventing efflux out of the cell by ENTs and providing an inward directional gradient to regulate extracellular concentrations. Nucleoside transporters, adenosine deaminase, and purine nucleoside phosphorylase are expressed at relatively high levels in the cerebellum, suggesting an effective system for adenosine inactivation (Geiger and Nagy 1986; Anderson et al. 1999a, b). In a recent study, it was shown that adenosine kinase is the major determinant of basal adenosine levels in cerebellar slices (Wall et al. 2007). Its inhibition (with iodotubercidin) markedly increased extracellular adenosine concentration and the amount of synaptic inhibition (at parallel fiber-Purkinje cell synapses) mediated by A1 receptors. Blocking transport of adenosine, with inhibitors of ENT1 and ENT2 (NBTI and dipyridamole) had erratic effects on extracellular adenosine concentration and often did not prevent the effects of adenosine kinase inhibition. This suggests that there must be an adenosine transporter present that is not blocked. Extracellular breakdown of adenosine by adenosine deaminase appeared to play only a minor role in controlling the concentration of extracellular adenosine (Wall et al. 2007). Basal concentrations of adenosine are lower in slices from immature rats, although the mechanisms for adenosine removal are still present (Atterbury and Wall 2009).

Hypoxia In common with many brain regions, hypoxia induces adenosine release in the cerebellum resulting in the reversible depression of synaptic transmission (parallel fiber-Purkinje cell) via A1 receptor activation (Atterbury and Wall 2009) and neuroprotection (Logan and Sweeney 1997). The release of adenosine (measured by biosensors and depression of synaptic transmission) is much faster in mature

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cerebellar slices compared to immature slices (Atterbury and Wall 2009). The source and mechanism of hypoxia-induced adenosine release in the cerebellum is currently unknown.

The Role of Purinergic Signaling in the Cerebellum and in Motor Control The precise role that purinergic signaling plays in cerebellar function is currently unclear. Experiments in which Bergmann glia (which utilize ATP signaling to produce Ca2+ waves) were transgenically removed from mice, showed defects in LTD and eye blink conditioning although motor coordination was unaffected (Shibuki et al. 1996). It is tempting to suggest that glial ATP signaling is therefore important for correct cerebellar function, but it could be that other roles of Bergmann glia (such as glutamate uptake) underlie the deficits observed in the transgenic mice. Global knockout of the adenosine A1 receptor in mice results in hyperalgesia, anxiety, and decreased hypoxic protection but had no effect on total locomotor activity or sensorimotor reflexes including visual placing reflex, wire rod test, and prehensility (Johansson et al. 2001). Thus A1 receptors appear to play little or no role in motor control (although this is not an inducible knockout and there may have been compensatory adaptation). There are several P2Y receptor knockouts (including P2Y1, P2Y2, P2Y4, and P2Y6) but there is no obvious cerebellar phenotype in these mice although subtle motor effects may not have been noticed (Leon et al. 1999; Matos et al. 2005; Bar et al. 2008). There is evidence that adenosine signaling in the cerebellum is involved in the ataxia produced by alcohol and cannabinoid intoxication (Dar 1997, 2000). For example, knock down of A1 receptors by intracerebellar injection of antisense oligodeoxynucleotide reduces ataxia induced by alcohol and cannabinoids (Dar and Mustafa 2002).

Conclusions and Future Work A great deal of work has defined the distribution of purine receptors and their effects on cellular function within the cerebellum but still many questions remain unanswered. What effects does purine signaling play on the cerebellar neural network and on cerebellar output? Mixed excitatory effects (P2X, P2Y) and inhibitory effects (A1 and P2Y) on both excitatory and inhibitory neurons and glia make this question difficult to answer. What is the source of extracellular ATP and adenosine? The presence of extracellular ATP has not been directly measured but has been inferred from experiments using antagonists (such as PPADs). These experiments suggest that ATP could arise from Bergmann glia (Hoogland and Kuhn 2010; Hoogland et al. 2009) and from molecular layer interneurons (Piet and Jahr 2007). Adenosine has been measured following electrical stimulation in the molecular layer. There is currently no evidence that this adenosine arises from ATP (Wall and Dale 2007, 2008) and thus maybe directly released. Adenosine can also be released by hypoxia although the cellular source is unclear (see Dale and

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Frenguelli 2009 for discussion of possible release mechanisms). To be able to precisely define the role of purinergic signaling in the cerebellum requires the development of new pharmacological and genetic tools. For example, to dissect apart the effects of ATP and adenosine there is a requirement for much more potent and selective ecto-ATPase inhibitors. The role of concentrative adenosine transporters is unclear (in the cerebellum and in other brain areas) as there are no selective inhibitors. Targeted, inducible knockouts of specific purinergic enzymes and receptors would also improve the understanding of their roles in the cerebellum. A major site of A1 receptor expression is the parallel fiber terminal. Linking an inducible targeted knockout (using the a6 GABAA receptor promoter, e.g., see Yamamoto et al. 2003) would allow their role in motor behavior to be defined. Acknowledgments Work in the author’s laboratory is funded by the Medical Research Council (UK). We are grateful for comments on the chapter by Professor Nicholas Dale, Professor Bruno Frenguelli, and Dr. Yuri Pankratov.

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Modulatory Role of Neuropeptides in the Cerebellum

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Georgia A. Bishop and James S. King

Abstract

The existence of neuropeptides in the central nervous system has been known for almost 40 years. Within the cerebellum approximately 22 neuropeptides have been identified. However, the functional role(s) for many of the different peptides in the cerebellum is largely unknown as their distribution in the cerebellum is not uniform and varies between species. Further, to date sufficient physiological analysis has not been carried out to determine the potential importance of cerebellar peptides. It has been proposed that peptides slowly modulate cerebellar circuits by acting through G protein-coupled receptors in contrast to the fast action of amino acid neurotransmitters that bind to ionotropic receptors. This effect, in turn, would influence the state of an entire neuronal circuit rather than a single neuron. This chapter will be focused on two members of the corticotropin-releasing factor (CRF) family of peptides, namely, corticotropin-releasing factor and urocortin because their distribution has been described in multiple species and their function has been investigated to a greater extent than any of the other peptides found in the cerebellum. A review of the distribution of these peptides as well as a discussion of their cognate receptors is included. Data on the functional role of both peptides is presented. It is clear that complex interactions take place between these peptides and other transmitters in the cerebellum. The physiological data suggest that CRF is essential for normal cerebellar-mediated motor performance. There is less data on the physiological role of urocortin (UCN) in the cerebellum. To date, it has been shown that UCN increases the spontaneous firing rate

G.A. Bishop (*) • J.S. King Department of Neuroscience, The Ohio State University, 333 W. 10th Avenue, 43210-1239 Columbus, OH, USA e-mail: [email protected], [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 971 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_41, # Springer Science+Business Media Dordrecht 2013

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of Purkinje cells. The mechanisms by which this occurs are yet to be determined. Considering there are many other peptides in the cerebellum, it becomes clear that modulation by these peptides must be seriously considered when analyzing and interpreting physiological data related to cerebellar circuitry. The classic view that only glutamatergic or GABAergic inputs are relevant when describing the effects on firing rate, patterns of activation, output of the cortex or nuclei are no longer valid. Peptides are an integral component of cerebellar function and must be considered to completely understand cerebellar function.

Introduction The existence of neuropeptides in the central nervous system has been known for almost 40 years with the initial studies being conducted in the early 1970s (see (Hokfelt 1991) for review). Over the last 40 years, the number of identified neuropeptides has increased from the initial 15–20 described in the 1970s to nearly 100 today (Ito 2009). Ito (2009) has recently written an excellent review that focuses on 22 neuropeptides that have been identified in the cerebellum. Several of these peptides such as atrial neuritic peptide, galanin, cerebellin, and motilin are expressed in cerebellar neurons including Purkinje cells, Golgi cells, and Lugaro cells. Others, including corticotropin-releasing factor, insulin-like growth factor 1, calcitonin gene–related peptide, Leu and Met-enkephalin, substance P, a-melanocytestimulating hormone, melanin-concentrating hormone, neuronal neurotensin, somatostatin, neuropeptide Y, and cholecystokinin are localized in mossy fiber and/or climbing fiber afferents. Finally, there is a plexus of varicose axons, possibly originating from the hypothalamus, which contain angiotensin II, dynorphin, Leu and Met-enkephalin, and orexin. In his review, Ito (2009) noted that the functional role(s) for many of the different peptides in the cerebellum is largely unknown as their distribution in the cerebellum is not uniform and varies between species. Further, to date sufficient physiological analysis has not been carried out to determine the potential functional importance of cerebellar peptides. Ito concluded that peptides slowly modulate cerebellar circuits by acting through G protein-coupled receptors in contrast to the fast action of amino acid neurotransmitters that bind to ionotropic receptors. His hypothesis was that peptides, based on their diffuse distribution, could have a tonic effect on a large number of target neurons. This effect, in turn, would influence the state of an entire neuronal circuit rather than a single neuron. However, he acknowledged that much work needs to be carried out before the functional roles of many of these peptides in cerebellar circuits can be defined. This chapter will be focused on the corticotropin-releasing factor (CRF) family of peptides. Their distribution has been described in multiple species and their function has been investigated to a greater extent than any of the other peptides found in the cerebellum.

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Corticotropin-Releasing Factor Family of Peptides The CRF family of peptides includes, among others, CRF, urocortin (UCN), urocortin II (UCN II), and urocortin III (UCN III) (Vaughan et al. 1995; Takahashi et al. 1998; Iino et al. 1999; Bittencourt et al. 1999; Baigent and Lowry 2000; Skelton et al. 2000; Lewis et al. 2001; Reyes et al. 2001; Million et al. 2002; Li et al. 2002; Swinny et al. 2002). Only CRF and UCN have been shown to be present in the cerebellum and thus, they will be the only members of this peptide family discussed in this chapter. Corticotropin-releasing factor (CRF) is a 41 amino acid peptide that was originally described as playing a critical role in the regulation of the pituitary-adrenal axis as well as in complementary stress-related endocrine, autonomic, and behavioral responses (Brown et al. 1982; Vaughan et al. 1995; Ito and Miyata 1999). However, it is now clear that CRF is not confined to the hypothalamus and the stress axis. There are extensive morphological and physiological data that establish the presence of and a role for CRF in cerebellar circuits in many mammalian species including, mouse, rat, cat, rabbit, opossum, sheep, nonhuman primates, and humans (Bloom et al. 1982; Olschowka et al. 1982; Merchenthaler et al. 1983; Merchenthaler 1984; Powers et al. 1987; Kitahama et al. 1988; Cummings et al. 1988; Mugnaini and Nelson 1989; Watabe et al. 1991; Potter et al. 1994; Yamano and Tohyama 1994; Cummings et al. 1994; Bishop and King 1999). UCN has 45% sequence homology with CRF in the rat (Vaughan et al. 1995). This family member has not been extensively studied in the cerebellum, but has been described in the rodent, monkey, and human (Takahashi et al. 1998; Swinny et al. 2002; Vasconcelos et al. 2003). Functionally, UCN does not appear to be physiologically involved in either the basal or stress-induced activation of the pituitary (Skelton et al. 2000; Richard et al. 2002). Rather, UCN appears to be a more potent suppressor of ingestive behavior (food and water intake) and a less potent inducer of anxiogenic behavior as compared to CRF.

Distribution Immunohistochemical studies have localized CRF in climbing fiber (Fig. 41.1a–d, block arrows) and mossy fiber (Fig. 41.1a–d, small arrow) afferents in the cerebellar cortex of several species (Cummings et al. 1983; Powers et al. 1987; Cha and Foote 1988; van den Dungen et al. 1988; Cummings 1989; Errico and Barmack 1993; Yamano and Tohyama 1994; Bishop 1998; Tian and Bishop 2003). In the cerebellar nuclei CRF-positive varicosities surround the somata of nuclear neurons (Fig. 41.1f, white block arrows). In addition to classic climbing and mossy fiber afferents, CRF has been identified in varicosities (Figs. 41.1c–e, arrowheads) that surround Purkinje cell somata and primary dendrites (Cummings et al. 1988). Similar varicose fibers also traverse the neuropil of the cerebellar nuclei (Fig. 41.1f, small white arrows), presumably terminating on the dendrites of nuclear neurons. Clearly, the climbing fibers originate from the inferior olive as this is the

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Fig. 41.1 Distribution of CRF in the cerebellum. (a–e) illustrates CRF immunoreactivity that is present in climbing fibers (a–c – block arrows), mossy fibers (a–d – small arrows), and varicose axons (c–e – arrowheads) within the cerebellar cortex. Varicose axons (f – white arrowheads) also are evident in the neuropil of the cerebellar nuclei and labeling is also evident around the nuclear neuron cell bodies (f – white block arrows). The calibration bar in a ¼ 50 mm and also applies to b, c, and e. The calibration bars in d ¼ 50 mm and in f ¼ 40 mm

sole source of this afferent system. Further, CRF positive neurons have been identified in the olivary complex (Powers et al. 1987; Barmack and Young 1988; Kitahama et al. 1988; Cummings 1989; Cummings et al. 1989; Errico and Barmack 1993). The CRF-immunopositive mossy fibers arise from several brainstem nuclei including the vestibular complex, the nucleus prepositus hypoglossi, the

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paramedian reticular nucleus, nucleus reticularis gigantocellularis, and raphe nuclei in the rabbit (Errico and Barmack 1993). The source of the beaded plexus of axons has not been experimentally identified. Ito (2009) suggests that the source for these axons is likely the hypothalamus. This remains to be established. There is no agreement in the literature as to the distribution of UCN in the cerebellum. Data suggest that UCN is present in isolated neurons in the cerebellum (Weitemier et al. 2005) of mice; however, the extent of labeling is strain dependent with the C57BL6 strain showing more neurons than DBA/2 J mice. The observation of UCN positive cells in the hemisphere is in agreement with the mRNA data in the Allen Brain Atlas published online by the Allen Brain Institute, which shows positive in situ hybridization labeling for UCN in cells located in the Purkinje cell layer and granule cell layer in the cerebellar hemisphere (Fig. 41.2b–c). These scattered cells could be neurons (e.g., Purkinje cells and/or granule cells) or glia such as Bergmann glia or astrocytes. Likewise, mRNA for UCN was detected in the cerebellar nuclei (Fig. 41.2d, e). Immunohistochemical data partially supports these mRNA data. In the rat (Bittencourt et al. 1999) no neuronal labeling is observed in the cerebellar cortex. However, sparse UCN-immunoreactive (ir) fibers in all three layers of the cerebellar cortex were reported. This labeling was observed primarily in the flocculus, with lesser accumulations in the paraflocculus and lingula. Our immunohistochemical data (unpublished observations) likewise show that UCN immunoreactivity (UCN-ir) is sparse. In the paraflocculus, UCN-IR profiles that resemble mossy fiber terminals are evident (Fig. 41.3b, white arrowheads) as are punctate profiles that distribute randomly in the neuropil of the granule cell layer. The most prominent immunohistochemical labeling is present in fine varicose fibers that enter the Purkinje cell layer to form a diffuse plexus of varicosities in juxtaposition to Purkinje cell bodies (Fig. 41.3a black arrows). There also appears to be labeling around Purkinje cell somata (Fig. 41.3a, arrowhead). UCN-ir also is present in neurons in the cerebellar nuclei (Fig. 41.3c, d), confirming the mRNA data noted above, as well as in varicosities that traverse the neuropil in the lateral cerebellar nucleus (Fig. 41.3c, small arrows). In the monkey, UCN-ir fibers were limited almost exclusively to the granule cell and Purkinje cell layers of the flocculus (Vasconcelos et al. 2003); UCN-ir was not identified in neurons within the monkey cerebellum. In humans (Takahashi et al. 1998), radioimmunoassay results indicate UCN is present in both the hemisphere and vermis in relatively high concentrations. RT-PCR data also documented mRNA for UCN in the hemisphere and vermis of the human cerebellum further supporting an intrinsic source of UCN in the cerebellum. In contrast to the majority of studies that show only limited amounts of UCN within the cerebellum, a study by Swinney et al. (2002) indicated a wider distribution for this peptide in Purkinje cell bodies and their dendrites throughout the cerebellum. The study also described UNC as being present in basket cells, stellate cells, parallel fibers, and climbing fibers. However, the light and electron microscopy data were not consistent and the antibody was uncharacterized in these studies. In agreement with other studies, punctate labeling was observed in cerebellar nuclei.

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Fig. 41.2 In situ hybridization for urocortin from online Allen Brain Atlas (http://mouse.brainmap.org/brain/Ucn.html?ispopup¼true) (a) is a graph that documents the gene expression level and expression density for urocortin in different regions of the brain. Compared to areas such as the hippocampus (HIP) and cortex (CTX) there is relatively little urocortin in the cerebellum (CB, dashed lines). Other areas examined include hippocampal formation (HPF), hypothalamus (HY), lateral septal complex (LSC), midbrain (MB), medulla (MY), olfactory bulb (OLF), pons (P), pallidum (PAL), retrohippocampal region (RHP), amygdaloid complex (cAMY), striatum (S), striatum-dorsal region (STRd), striatum ventral region (STRv), and thalamus (TH). (b) and (c) illustrate the distribution of mRNA for urocortin in the cerebellar cortex. Neurons in the Purkinje cell layer, either Purkinje cells or Bergmann glia, are positively labeled. Some labeling also is present in the granule cell layer. (d) and (e) illustrate the distribution of mRNA for urocortin in the nucleus interpositus anterior and posterior. Several neurons express the mRNA for this peptide. The areas enclosed by the boxes in b and d are shown at higher magnification in c and e, respectively

In summary, the general consensus from the literature is that UCN is present in scattered cell bodies located primarily in the flocculus and hemisphere of the cortex. Sparse immunoreactive labeling of fibers is present in the cerebellar cortex whereas UCN immunoreactivity is clearly present in the cerebellar nuclei. The source of the UCN-ir axons in the cerebellum is yet to be determined. They could be collaterals of Purkinje cells; this would only correlate with beaded fibers

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Fig. 41.3 Distribution of urocortin in the cerebellum. (a) illustrates urocortin immunoreactivity in varicose axons (black arrows) that distribute along the Purkinje cell layer. Some varicosities appear to surround Purkinje cell bodies (black arrowhead). (b) illustrates urocortin-positive mossy fibers (white arrows). Note that only a few scattered terminals are labeled. Scattered varicose axons (c, small arrows) are evident in the neuropil of the cerebellar nuclei. In addition, neurons in the cerebellar nuclei (c, d) are intensely immunolabeled. (d) is an image from a double label experiment in which urocortin (red) and calbindin (green) were tagged with different fluorophores. The calbindin antibody immunolabels the terminals of Purkinje cells which surround urocortinpositive nuclear neurons. The calibration bar in a ¼ 25 mm, in b ¼ 20 mm, and in c ¼ 20 mm. The calibration bar in c also applies to d

located in the Purkinje cell and granule cell layer in areas where UCN-positive Purkinje cells are present such as the hemisphere. It is likely that UCN-positive axons, similar to those expressing CRF, are derived, at least in part, from neurons in the brainstem. Based on studies that described the distribution of UCN-positive neurons (Bittencourt et al. 1999), there are several brainstem nuclei that could be extrinsic sources of UCN to the cerebellum including the locus coeruleus, vestibular nuclei, as well as reticular and raphe nuclei. Finally, UCN-positive neurons in the cerebellar nuclei (McCrea et al. 1977; Tolbert et al. 1978; Batini et al. 1989, 1992) may be an additional source of UCN collateral axons within the cerebellum.

CRF Receptors in the Cerebellum Two major CRF receptor subtypes (CRF-R1 and CRF-R2) have been identified and characterized (Potter et al. 1994; Chalmers et al. 1996). Both types are members of

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the class B subfamily of seven-transmembrane receptors that signal by coupling to G proteins (Dautzenberg and Hauger 2002) and stimulate the production of cAMP (Lovenberg et al. 1995b). Each receptor subtype has unique binding characteristics and a differential distribution in the brain. Although it binds to both types of receptors, CRF has a higher affinity for CRF-R1 than for CRF-R2 (De Souza 1995; Chalmers et al. 1996; Behan et al. 1996). In contrast, compared to CRF, UCN binds six times more strongly to membranes of cells expressing CRF-R1 and 40 times more strongly to those expressing CRF-R2(Vaughan et al. 1995) and is considered the putative endogenous ligand for this later receptor subtype (Skelton et al. 2000). For CRF-R1, there is one functional splice variant and several nonfunctional variants (Dautzenberg and Hauger 2002). The majority of the nonfunctional splice variants are found only in humans and all are deficient in ligand binding and/or signaling properties (Dautzenberg et al. 2001). In situ hybridization and immunohistochemical studies have localized the mRNA for CRF-R1 or the peptide itself in the rodent’s cerebellum (Chen et al. 1986, 2000; Chai et al. 1990; De Souza and Insel 1990; Potter et al. 1994; King et al. 1997; Primus et al. 1997; Bishop et al. 2000). In the mouse cerebellum (Bishop et al. 2000), CRF-R1 immunoreactivity is present on the somas and primary dendrites of most, if not all, Purkinje cells, as well as on radial glial cells, and scattered granule cells. Data from our lab in the rat confirms these observations (Fig. 41.4a–b). Further, neurons in the cerebellar nuclei of the rat also express CRF-R1 receptors (Fig. 41.4c). There are three functional splice variants of the type 2 CRF receptor referred to as CRF-R2a, CRF-R2b, and CRF-R2g. CRF-R2g is only found in humans and CRF-R2b is found primarily in the periphery in heart, lung, skeletal muscle, and the gastrointestinal tract (Lovenberg et al. 1995a). In contrast, CRF-R2a, is absent in the periphery, but has a major distribution in the CNS. Recently, a truncated isoform of CRF-2a has been described and is designated CRF-2a-tr (Miyata et al. 1999a, 2001; Dautzenberg et al. 2001); it also is located in the CNS. CRF-2a-tr contains the first three and a part of the fourth transmembrane domains of the fulllength type 2 CRF receptor. It is lacking the remaining membrane loops as well as the c-terminus. In tissue sections from both the rat and the mouse, the pattern of immunostaining for the full-length and truncated CRF-R2a antibodies is distinct. CRF-R2a is localized primarily to Purkinje cells (Fig. 41.4e, f), Bergmann glial cells throughout the cerebellum, as well as cells in the granule cell layer (Fig. 41.4d, small arrows) which could be astrocytes or granule cells (Bishop et al. 2006). In addition, it also is present in select populations of basket/stellate cells in the molecular layer as well as in scattered Golgi cells in the granule cell layer (Bishop et al. 2006). Large neurons within the cerebellar nuclei strongly label for the full-length receptor (Fig. 41.4g). However, as for cells in the cortex, not all cells in the nuclei are immunopositive suggesting selectivity in receptor expression. In contrast to the postsynaptic distribution of the full-length CRF-R2, the truncated isoform primarily shows an axonal or presynaptic distribution. The truncated receptor is localized to the initial segments of Purkinje cells

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Fig. 41.4 Distribution of the CRF-R1 and CRF-R2a receptor in the cerebellum. The type 1 CRF receptor (CRF-R1) is located primarily on the somata of Purkinje cells (a, b) and cerebellar nuclear neurons (c). The area indicated by the box in a is shown at higher magnification in b. CRF-R2a is evident over Purkinje cell dendrites (d, e) and granule cells (d small arrows). It also is present on the somata and dendrites of neurons in the cerebella nuclei (f). The calibration bar in a, d, e and f ¼ 50 mm; in b ¼ 25 mm; and in c ¼ 20 mm

(Fig. 41.5a, c, e), Golgi cells, basket cells, and nuclear neurons (Fig. 41.5b, yellow arrows) (Bishop et al. 2000). The identity of the initial segment arising from a Purkinje cell was confirmed using calbindin as a double label to identify Purkinje cells (Fig. 41.5c–e). In the molecular layer, the truncated isoform is located in parallel fibers and their axon terminals, based on electron microscopic data. In addition, CRF-R2a-tr also is localized in Purkinje cell terminals in the cerebellar nuclei (Bishop et al. 2000; Lee et al. 2004; Tian et al. 2006).

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Fig. 41.5 Distribution of CRF-R2a-tr in the cerebellum. The truncated isoform of the type 2 CRF receptor (CRFR2a-tr) is evident in the base and initial axonal segments of Purkinje cells (a, c – yellow arrows). Purkinje cells are clearly identified by calbindin labeling (d, e) and CRFR2a-tr in double label paradigms. The truncated isoform of the receptor also is present in the initial segments of neurons in the cerebellar nuclei (b – yellow arrows). The calibration bar in a ¼ 50 mm; in b ¼ 40 mm; and in c ¼ 50 mm. The calibration bar in c also applies to d and e

Functional Roles of CRF in the Cerebellum Several studies have shown that CRF has an effect on general locomotor activity (Swerdlow et al. 1986a, b; Monnikes et al. 1992; Lowry et al. 1996; Lai and Siegel 1997; Bittencourt and Sawchenko 2000; Contarino et al. 2000; Ohgushi et al. 2001; Dirks et al. 2002; Timofeeva et al. 2003; Bale and Vale 2004). However, based on the techniques used, it was not possible to show that these effects involved

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cerebellar circuitry. The initial report that demonstrated a functional role for CRF in the cerebellum showed that the exogenous application of CRF enhanced the firing rate of Purkinje cells in the intact animal via several distinct interactions (Bishop 1990). First, CRF potentiated the excitatory effects of amino acids such as aspartate or glutamate. In addition, CRF blocked the inhibitory effect of GABA providing a second mechanism for increasing the firing rate of Purkinje cells. Finally, CRF shortened or eliminated the period of suppression produced by activation of climbing fibers in the cerebellar cortex. It was of interest that, although CRF interacted with several known amino acid neurotransmitters, by itself it had little to no effect on the firing rate of Purkinje cells. These were the first data to suggest that CRF played a role as a neuromodulator in cerebellar circuits. Insights into the mechanism of action of CRF in the cerebellum were provided by Fox and Gruol (1993). Their study provided the first data on the mechanism by which CRF enhanced the firing rate of Purkinje cells. They demonstrated that CRF produced a dose-dependent reduction in the amplitude of the afterhyperpolarization (AHP) which followed a current-induced spike train. Similar findings had been described earlier in the hippocampus (Aldenhoff et al. 1983). This earlier study had demonstrated that CRF decreased the AHP in hippocampal neurons and that the likely mechanism was by a direct effect in closing calcium-dependent potassium channels. It was proposed that similar effects on potassium channels occurred in the cerebellum as well. Taken together, these early studies documented that CRF did play a role in modulating Purkinje cell activity and thus could influence cerebellarmediated locomotor behavior. Further data on the functional role of CRF in the cerebellum was provided by Miyata et al. (Miyata et al. 1999b). They carried out studies to define a role for CRF released from climbing fibers derived from the inferior olivary complex. This report concluded that CRF was critical in mediating cerebellar long-term depression (LTD) at the parallel fiber-Purkinje cell synapse. They concluded that CRF primarily played a role in the induction of LTD and that it was not involved in the long-term maintenance of the depressed activity. Further, their study determined that the effects on parallel fiber firing rate was dependent on activation of the protein kinase C second messenger pathway. This may seem to be a contradiction to earlier studies that demonstrated enhanced activation of Purkinje cells following CRF application in the intact animal (Bishop 1990) and in cultured neurons (Fox and Gruol 1993). However, this apparent discrepancy can be resolved based on the following observations. First, in the earlier studies the effects were measured on spontaneous simple spike activity and did not involve activation of climbing fibers prior to application of the peptide. Several studies on LTD (reviewed in (Linden and Connor 1993; Ito 2001) have documented the need for conjunctive activation of both the climbing fiber and parallel fiber pathway to elicit the response. In the absence of climbing fiber input, or the use of a massive depolarization of the Purkinje cell through the microelectrode, it is unlikely that LTD was induced in the paradigm in which CRF was applied exogenously. Miyata et al. (1999b) concluded that CRF had a very specific role in inducing only the early component of LTD; it did not play a role in sustaining the suppression of

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activity. The temporal resolution of the extracellular technique may not have been sufficient to detect a slight initial decrease in activity induced by application of the peptide. Fox and Gruol (1993) also analyzed the effect of CRF on the firing rate of Purkinje cells. Although not significant, they did describe a decrease in the simple spike firing rate after application of CRF. As it was not statistically significant, they did not pursue this further. Taken together, the data from each of these studies suggests that CRF plays a complex role in modulating Purkinje cell activity. In the absence of climbing fiber activation and a presumed lack of LTD induction, CRF clearly enhances the simple spike firing rate of Purkinje cells. However, if there is conjunctive activation of the climbing fiber pathway, and CRF is released, there is an opposite effect in that the presence of the peptide induces a decrease in the activity of parallel fibers that were simultaneously active. Thus, the level of activity in climbing fibers plays a major role in determining the modulatory role of CRF in cerebellar circuits. In addition to being essential for induction of LTD at the parallel fiber synapse, CRF also regulates excitatory transmission at the climbing fiber-Purkinje cell synapse referred to as climbing fiber LTD (CF-LTD) (Schmolesky et al. 2007). This study found that exogenous application of CRF transiently induced reductions in both the climbing fiber-evoked excitatory postsynaptic current, as well as in the second component of the classic complex spike and the afterhyperpolarization induced by activation of the climbing fiber afferent system in an in vitro slice preparation. The findings of this study indicate an additional level of complexity to the physiological role of CRF in the cerebellum. The mechanism is complex and involves activation of both the PKC and PKA second messenger pathways. Data indicate inhibitors of the PKC pathway reduce the effect of CRF upon the CF-EPSC as well as on CF-induced LTD. The PKA pathway is essential for the CRFmediated reduction of climbing fiber–evoked excitatory postsynaptic currents (CF-EPSC) as well as the decreased amplitude of the associated afterhyperpolarization and the induction of CF-LTD. The effects on the second component of the complex spike and the afterhyperpolarization were all related to a CRF-mediated decrease in the amplitude of the CF-EPSC. Reduction in the EPSC impairs CF-evoked calcium influx as levels of depolarization are decreased. Schmolesky et al. (2007) concluded that there is an important link between CRF-induced reduction of climbing fiber EPSCs and the induction of parallel fiber–mediated LTD. They indicated that plasticity at the CF-Purkinje cell synapse would have a major impact on ongoing cerebellar processing. By decreasing the influence of climbing fibers there is a concomitant decrease in parallel fiber– mediated LTD as it has been shown that the massive depolarization induced by climbing fiber activation is required to block the excitatory effect of parallel fibers that were active at the same time. A decrease in climbing fiber input would result in enhanced activity in the subset of Purkinje cells contacted by these afferents. They also found that low basal levels of endogenous CRF was not sufficient to induce CF-LTD. This led them to suggest that activity-driven release of higher concentrations of CRF is likely necessary to have a significant impact upon

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glutamatergic transmission at the CF-PC synapse. This observation correlates well with the studies that document changes in CRF expression and release with increased activation of the climbing fiber pathway (Tian and Bishop 2003; Beitz and Saxon 2004). Several studies have shown that enhancing activity in the cerebellar afferent pathways alters the level of CRF expression in the cerebellum (Tian and Bishop 2003; Beitz and Saxon 2004). One study (Tian and Bishop 2003) concluded that stimulation of cerebellar afferents induces a significant change in both the distribution and number of CRF immunolabeled climbing fibers and mossy fibers. In the other study (Beitz and Saxon 2004), use of microdialysis probes verified that application of harmaline, a drug that potently activates olivocerebellar axons, results in significant increases in CRF release into the extracellular fluid compared to pre-harmaline levels. Taken together, these data suggest that the modulatory effects ascribed to CRF could have a greater influence when the firing rate of climbing fibers was increased. This would be consistent with previous studies that demonstrated that peptides are released primarily when afferents are stimulated at high rates or axon terminals are powerfully depolarized (Verhage et al. 1991). In a companion study (Tian et al. 2008), the effects of enhanced firing rate in climbing fibers also was shown to alter the distribution of the type 1 CRF receptor. As noted above, and by others, CRF-R1 is localized primarily on Purkinje cell somata and the adjacent portion of their primary dendrites. In addition, Bergmann glial cells, granule cells, Golgi cells, basket cells, and stellate cells also express CRF-R1, although to a lesser extent. Increased activation of climbing fibers within their physiological range of activity had little effect on CRF-R1 expression in Purkinje cells. The primary effect was an increase in receptor expression in the processes of Bergmann glial cells with a concomitant decrease in expression in basket and stellate cells. These observations suggest that there is a complex interaction between CRF and its receptors in modulating the activity of neurons in the cerebellar cortex. It is of interest that there is an apparent ligand-receptor mismatch with respect to the localization of CRF in climbing fibers as there is a paucity of CRF-R1 on the intermediate and distal dendrites of Purkinje cells as they extend into the molecular layer; the sites where climbing fibers synapse. This suggests that CRF released from climbing fibers may not have a direct effect on modulating synaptic input to these processes. This is further confirmed by the fact that there was little change in the distribution of the receptors on the somata of Purkinje cells, suggesting that they were not affected by the increased levels of CRF documented in the previously discussed studies (Tian and Bishop 2003; Beitz and Saxon 2004). However, increased levels of the peptide mediated by increasing the firing rate of these afferents did alter receptor expression on other neurons in the molecular layer, including stellate and basket cells and of even more interest on Bergmann glial cells. This suggests that the effects of CRF on Purkinje cell activity may be mediated by interactions with these other cellular elements in the cerebellum. Additional studies are needed to better define these potential direct effects of CRF released from climbing fibers on GABAergic interneurons and Bergmann glia.

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There are data to suggest that CRF could have a physiological effect on glial elements in the cerebellum. Patch clamp data (unpublished observations) indicate that application of CRF hyperpolarizes the membrane potential of Bergmann glia with a concomitant 7% increase in resistance indicating decreased conductance across the membrane (Fig. 41.6a–c). Additional studies are needed to more clearly define the mechanisms by which CRF alters Bergmann glial cell polarization and how this impacts the physiological response of Purkinje cells. That does leave a question as to the role of the CRF receptors localized to the Purkinje cell soma. It is possible, as discussed below, that these receptors are bound primarily by CRF released from the varicose axonal plexus that distributes along the Purkinje cell layer. Alternatively, they could be bound by UCN in areas where this peptide is present. Taken together, these studies demonstrate that CRF plays a role in modulating cerebellar output. Further, it is an essential component of long-term depression which is associated with motor learning. Finally, based on unpublished observations from our laboratory, we suggest a direct effect of this family of peptides on motor performance. If the CRF antagonist, astressin, is injected into the cerebellar nuclei, there is a significant decrease in motor performance on the rotorod, a paradigm that is highly correlated with cerebellar deficits (Fig. 41.6d, e). Thus, from cellular to behavioral studies, it appears that CRF is essential for normal cerebellar-mediated motor performance.

Functional Role of UCN in the Cerebellum Several studies (reviewed in (Gysling et al. 2004)) suggest that UCN has functions that are distinct from those ascribed to CRF. Whereas CRF is the major regulator of the hypothalamo-pituitary-adrenal axis in response to stress, UCN appears to primarily play a role in feeding behavior and may elicit anxiety-like behaviors in response to stress (Skelton et al. 2000). This effect on motor activity could occur at ä Fig. 41.6 Effects of CRF family of peptides on Bergmann glial cells and on motor performance: In patch clamp recordings, Bergmann glial cells were localized between Purkinje cells and injected with Lucifer yellow (a, b). The fluorescent dye spread into the radial processes as well as the appendages in the molecular layer. Resting membrane potential was recorded while passing 300 pA current pulses through the electrode. The resulting voltage pulse across the membrane is depicted as the downward deflection in the RMP record and provides information on changes in resistance across the membrane. Application of 0.1 mM CRF hyperpolarized the membrane with an associated 7% increase in resistance as determined by an increase in the amplitude of the voltage pulses. UCN II also hyperpolarized the membrane but was not as effective. UCN had little effect on the resting membrane potential. d and e are histograms that illustrate the effects induced by injection of astressin, a potent antagonist of the type 1 and type 2 CRF receptors, into the cerebellar nuclei. Significant performance deficits were observed at all speeds tested in a constant speed rotorod paradigm as well as in an accelerating rotarod paradigm (p DNA. . . recursion (after discarding the obsolete notions of Central Dogma and Junk DNA). Note, that for the purposes of simplicity Icon 4 shows the recursion as a single circular line – but it symbolizes multicomponential (vectorial) entities. The cardinally important Generalization of Recursion (from neuroscience to genetics) is the concept introduced here that the coding DNA vectors (many “exons” acting together), when transcribed, create RNA vectors that are of contravariant valence, since their translation into protein vectors creates physical objects. However, when protein vectors are signaled (measured) by noncoding DNA via bonding not only to homeodomains but also to ncDNA vectors, they are transcribed into another RNA vector, this time of covariant (sensory) valence. Thus, a recursion, similar to one shown in Icon 3 converges into the Eigenstates of the recursion in the genome, and the cRNA and ncRNA Eigenvectors produce the metric, comprising the functional geometry of the genome function. If the recursion converges to follow the Weyl’s Law on Fractal Quantum Eigenstates, the genomic recursion switches the growth of fractal protein structures (such as a Purkinje neuron, shown in Icon 5) into the next step of recursive hierarchy. The physiological process requires canceling (methylating) ncDNA segments perused in the recursion (see Fig. 61.9), such that the ncDNA fractal segments, governing growth according to FractoGene are not overused. It follows, that hypo-methylation and incorrect chromatin modulation could permit an uncontrolled (cancerous) growth as shown in Fig. 61.9 (yellow “cookie”). For further details, consult the original papers containing Icons 1–5 and the text of this review, relating the seminal concept of generalization of recursion described in Fig. 61.2 with the fractal recursive iteration shown in Fig. 61.9

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happen, science needs to specify the mathematics that underlie “a biological system theory” (Bertalanffy 1934). Identification is essential for both the neural network and for the underlying genome, including the suggestion here that they are conceptually identical. Further, as it is suggested here, science needs to move away from a “one-to-one” and “arrow-type” mapping toward the “many-to-many more” and “recursive” and dual representations. This is important not just for theory, but for entire industries. The “Big Pharma” model of “one gene, one disease, and one billion dollar pill” is obsolete for over a decade because of a simplified and incorrect “one-to-one” assumption. Now the future lies in the generalization of covariant and contravariant neural network representations for the genome-epigenome (hologenome) system. The cerebellar biological neural networks, as shown, provide a precedent for this mathematical insight that is also applicable to genomics.

Generalization of Recursion from Cerebellar Neuroscience to Genomics; Covariant and Contravariant RNA Functors and Their Eigenstates As it was shown over three decades ago in Pellionisz and Llina´s (1980) if using nonCartesian (generalized, non-orthogonal) coordinate systems (see Icon 1 in Fig. 61.2), invariants (such as displacement) are represented in with a dual valence. The orthogonal projection-components, named covariant tensor-components in mathematics by Sylvester (1853), can be independently established; however, covariant components do not physically assemble the invariant. In turn “motor expressions,” expressed as interdependent parallelogram-type coordinates, that he called contravariant tensors, do assemble the object in a physical manner. It is cardinal in mathematics of generalized coordinates (tensor geometry) that a matrix can convert the “covariant sensory intention vectors” into “contravariant motor execution vectors” (see Panels C-F of Icon 1). The matrix that does this is the manyto-many interconnection-system of a massively parallel neural network of the cerebellum. Thus, the cerebellar sensory-motor coordination is accomplished by the conversion via the metric tensor. The metric comprises the geometry of the nonCartesian multidimensional space-time, embedding both sensory and motor events. This perhaps difficult but cardinal concept of sensory- and motor components as coand contravariant vectors was most lucidly encapsulated by Anderson (1990) pp. 351–355, in Anderson et al. (1990). If dual, covariant, and contravariant functors, shown in Fig. 61.2 Panel 3, are freely let to recur (when proprioception vectors are directly used by recursion as if they were execution vectors, without the cerebellar cortex, see Icons 1–2 in Fig. 61.2), they converge into the Eigenstates (where the normalized covariantand contravariant representations are identical – while in general they are different). Finding the Eigenvectors characterizing the Eigenstates by free recursion (that in sensorimotor systems is manifested in uncoordinated, oscillatory movements) is essential, since the metric tensor (and its inverse, or Moore-Penrose Pseudo-inverse

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for overcomplete space (Pellionisz 1984)), capable of converting covariants to contravariants, is obtained as a matrix-product of the Eigenvectors. In category theory, covariant and contravariant as well as mixed valence of functors (vectors and generalized vectors, tensors, are just one specific type of functors; they relate invariants to coordinate axes) are both well established and reaffirmed (Francis 2008). Herein, with the conceptual guidance of Icons 3–4 in Fig. 61.2, the dual representation is generalized to the interpretation of RNA system in Genomics. There is no debate that the so-called amino-acid-coding RNA-s (cRNA-s, as a multicomponent, vectorial entity) physically aggregate physical objects (proteins). Thus, the valence of cRNA-s is contravariant, similar to motor vector components that also have to assemble the physical object. The contravariant cRNA vectors, however, via RNA self-replication (Glasner et al. 2000) are available not only to construct proteins, but to interact (interfere) with the rather different covariant multicomponent ncRNA vectors. These measurements, what protein systems are already built, arise when proteins bind to noncoding DNA (both in intergenic and intronic sequences) involving transcription factors (Kornberg and Baker 1992). Through the arising ncRNA functors (multicomponent vectors), the already built proteins are thus “measured” not just by a single sequence, referred to as “homeoproteins” generated by a “homeodomain” (Foucher et al. 2003), but a single protein-component is signaled by the many components of even a single but multicomponent ncRNA covariant vector. Compare the concept to covariant sensory vectors providing independent measures of motor events in Icon 1 of Fig. 61.3. Putting the RNA system here into a new conceptual framework, also re-defining the role of intronic and intergenic “noncoding” (formerly, “Junk”) DNA, recalls earlier metaphors. Interpreting the RNA system as a “hidden layer,” an implication referring to interconnections known in neural nets for decades Mattick (2005) phased out his earlier metaphor that conceptually compared the RNA system to the man-made “operating system of computers” (Mattick 2001). Recently, even “genomic matrix” relating to fractals and chaos (Petoukhov and He 2010) and even “RNA matrix” approaches emerged (Izzo et al. 2011). However, the co- and contravariant valences of RNA functors have not been recognized to date. This generalization of valence of functors from sensory- and motor vectors to covariant as well as contravariant RNA multicomponent entities provides an opportunity to approach the role of RNA systems in coordinated genome function in a novel manner; that is, both conceptually and mathematically already identified in living systems (cerebellar neuronal networks) that the genome-epigenome system is known to generate. Appreciation of the valences of RNA functors opens new vistas beyond approaching the RNA-metric from a mindset that moves the perspective of science beyond man-made technologies like operating systems of computers (Mattick 2001). Looking at the RNA system in a new light as “the metric tensor of protein building genic sequences regulated by protein sensing noncoding sequences,” the RNA system is conceptually likened to a “genomic cerebellum.” First of all, this permits deploying already proven advanced geometric (thus software-enabling) analysis of experimental results of genomics. Second, the perspective on evolution

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is affected by recalling the shifting metaphor by Mattick from “operating system” to “hidden layer” (2001 vs 2005) and his reference that the RNA system serves a “coordinated genome expression.” One cannot help noticing that the “invention” through evolution of the physically separate, additional cerebellar neural network (with the shark) provided for a new class of more highly coordinated vertebrates. The conceptual equivalence is noted, therefore, that much of single-cell organisms

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contain a minimal amount of “noncoding DNA” – thus appear to operate with minimal covariant ncRNA, similar to organisms before the cerebellum appeared, permitting only an imprecise, un-coordinated execution of genomic commands. As the amount of noncoding (regulatory) sequences hyperescalated, the emerging RNA-metric permitted the coordinated growth and governance of complex (also multicellular) organisms. This new interpretation of the RNA system is to be compared to Mattick’s referral to “the Cambrian explosion” (Mattick 2004). At the least, identification of a common advanced geometry intrinsic to living systems makes “System Theory” approaches to genomic systems mathematically explicit. A more remote but an inevitable goal for the use of a common advanced geometry is to accelerate the unification of genomics and neuroscience. It is fully realized that building this seminal idea into a robust school of thought will require significant time and resources.

Recursive Algorithms Rule Both Vector–Matrix and Fractal Representations Algorithms based on recursion (see Icon 4 of Fig. 61.2 from Pellionisz (2008a, b)) share the fundamental property that each state of the system is deduced from its previous states. Recursion, in itself, does not discriminate analog (e.g., traditional feedback) mechanisms from digital deduction as, for example, in the sequence of Fibonacci numbers, where each subsequent integer is the sum of the previous two. The metric tensor characterizes the non-Euclidean geometry with integer dimensions, established by recursion of covariants to contravariants to compose the metric from Eigendyads (Pellionisz and Llina´s 1985). The embedding Minkowski-spacetime manifold, however is “smooth,” mathematically speaking it is derivable. However, Purkinje neurons show a non-Euclidean, moreover, a discrete geometry with fractal (non-integer) dimension (Pellionisz 1989). Realization that the same cerebellum utilizes recursion of dual vectors, as well as its main type of neurons, the Purkinje cells are built by an also recursive, but by a rather different fractal iterative recursion (see Icon 5 of Fig. 61.2; Pellionisz 2002, 2003) a cardinal question arose ever since the fractal model of Purkinje neurons (Pellionisz 1989). The question became even more vexing with the FractoGene concept, stating that fractal DNA governs growth of fractal organelles such as the Purkinje neuron; fractal organs such as the lung, circulatory systems; and organisms such as the Cauliflower Romanesca pictured in Pellionisz (2008a). The question was if the vector–matrix and fractal representations are in a mathematical conflict with one-another, or rather, if they reveal another profound dualism, similar to one already encountered in physics. The question was also conceptual regarding not only the mathematics, but also possibly referring to a “language.” The “early wave” of looking at fractality of DNA suspected it as a “language” (Flam 1994). The concept of a “language,” however, does not appear to be consistent with the concept of “sensorimotor coordination.” Resolution of the question became easier once the “hint” that fractality reflects

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a “language” was dismissed (Chatzidimitriou-Dreismann et al. 1996). Section on “The Genome is Fractal! Proof of Concept and the Basis of Generalization: Whole Genome Analysis Reveals Repetitive Motifs Conforming to the Zipf-Mandelbrot Parabolic Fractal Distribution Law of the Frequency/Ranking Diagram” shows below that the established fractality of the genome conceptually supports FractoGene, fractal growth of Purkinje cells governed by fractal DNA. Both in the DNA and in networks of neurons, the fractality characterizes the geometry in a consistent manner. The question was settled by Bieberich (1999) (see his Figure reproduced as Fig. 61.3 in this chapter) to show a conceptual consistency of fractal and vector– matrix representations. Thus, a geometric characterization of sensorimotor function and the geometry of the Purkinje neurons that implement smooth (derivable) function by non-derivable fractals are not only compatible, but mutually convertible. The revelation by Bieberich (1999) was not entirely surprising, given the known fact in physics that light can be seen as a wave-phenomenon, or particle-phenomenon, depending on the theory of Schr€ odinger or Heisenberg. Thus, the Bieberich-diagram is intellectually rather pleasing. Even more intriguing is its extension toward fractal internal representation (consciousness) in Bieberich (2011). Based on insights from fractal modeling of Purkinje neuron (Pellionisz 1989), utilities could be developed based on the of fractality of both DNA and the organelles, organs and organisms grown by the genome, the concept of FractoGene by Pellionisz (2002, 2003, 2006). The FractoGene algorithmic approach to the whole genome provided quantitative predictions that could be verified or refuted by experimentation; moreover the “Fugu Prediction of FractoGene” (that the 1/8 of the noncoding DNA of fugu compared to that of the human should result in a “fractal primitive” dendritic tree in the fugu) was supported by experimental results (Simons and Pellionisz 2006a, b).

Tensor Network Theory: Vector–Matrix Recursion as Basis of the Cerebellum Acting as a Sensorimotor Metric Tensor Recursion of sensory to motor vectors (and the generalization of valence of RNA functors) was characterized by Icon 2 of Fig. 61.2 as an essential procedure to converge into Eigenvectors, with their matrix-product comprising the geometry in the metric tensor. With the example of encyclopedic Fig. 61.4 of this chapter from Pellionisz (1987), it is shown how such metric is the basis of an entire system of gaze control, stabilizing the head by the vestibulocollic sensorimotor neural network. Icons 2–3 of Fig. 61.2 showed that sensory functors could recur directly, used in an unchanged manner, as motor functors. However, the recursion would result in an oscillation converging into Eigenvectors. In the cerebellar sensorimotor system, the Eigenvectors are imprinted in the inferior olive (Pellionisz and Llina´s 1985). In turn, as shown in Fig. 61.4 here, Eigenvectors from the inferior olive give rise to their matrix-product implemented by the neuronal network of cerebellar cortex. The scheme shown in Fig. 61.4 stabilizes gaze (head position) by a two-step operation: First, there is a covariant embedding from a symbolically

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Fig. 61.4 Tensor network model of the vestibulocollic reflex, embodying a covariant intention to contravariant motor execution transformation via the cerebellar neuronal network (From Pellionisz (1987)). For details, see the original publication and the text below. This figure also serves as the inspiration of the seminal concept of generalization of TNT to Genomics. The generalization is based on the fact that a physical object of the head movement is both measured by the covariant sensory vector that converted both in dimensionality and covariant to contravariant valence. Likewise, the genome expresses physical objects (proteins) both by protein-coding codons (in a contravariant manner), that can be measured by similar (but noncoding triplets, wherein the detection is covariant), but in order to attain quantum fractal eigenstates of stable protein systems a many-to-many RNA converter is needed. The RNA system is, therefore, conceptually equivalent to the sensory-motor transformer of the cerebellum

two-dimensional sensory vector into an also covariant, but higher (figuratively, 3) dimensional motor intention vector (i) – that would go directly to (mis)serve as an imprecise execution vector (since motor vectors must be contravariant; (i) should be (e)). Through the ascending mossy fibers, the (i) covariant intention vector is both converted into the (e) contravariant vector (negative, since Purkinje cells are inhibitory), that with the mossy fiber collateral (i) vector in the cerebellar nuclei constitutes an output vector (ie). Thus, the brain stem would send out instead of the covariant intention vector (i) the proper e ¼ i(ie) precise contravariant execution vector. This architecture explains why the entire sensorimotor would work (as for a dysmetric patient; even Purkinje cell affected only by alcohol) with intentions directly executed, but the additional neural network that was a nifty improvement as an addition to the brain of the shark makes a dysmetric direct execution of intentions into one that matches the physical geometry of the executor system (in this case, muscles) with its internal metrical representation.

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Coordination of an entire sensorimotor architecture is presented here to illuminate how non-trivial the generalization of contra- and covariant cRNA functors directly recursing into ncRNA covariant functors is. Such direct recursion is excellent for finding the Eigenstates of a DNA > RNA > PROTEIN recursive system, but the multicomponent RNA Eigenvectors must interact in an all-to-all manner, by means of RNA interference, not just of one component, Fire et al. (1998), but in a many-to-many multicomponent manner. Also, the sensorimotor coordination scheme is to illuminate why RNA interference is “silencing” – conceptually similar to the inhibitory effect of cerebellar Purkinje cells. Development of the school of functional geometry of a comprehensive system of coordinated genome function, comparably to that of a sensorimotor apparatus, requires a long-term program. One of the most difficult questions is if the genomic recursion obeys the Fractal Weyl’s Law on Fractal Quantum Eigenstates (see Shepelyansky (2008), originally Weyl (1912)). This question will be discussed in the section on “Future Directions.”

Fractals Are Pervasive in Nature; Both the Cerebellar Brain Cells and the DNA are Fractal Objects Mandelbrot (1983) coined the term “fractal” in his epoch-making book only about a quarter of a century ago, but the impact of identifying fractal geometry intrinsic to Nature is already profound.

The Zipf-School Suspected that the DNA Contained a Fractal Language The first “hints” that the A, C, T, and G nucleotide sequences of DNA (especially of noncoding DNA) possibly harbored a (mathematical) “language” was published before the epoch of “massive whole genome sequencing,” in 1994 in Science (see Fig. 61.1 in Flam 1994). Its original caption: “Line of evidence. Plotting frequency against rank of arbitrary ‘words’ in noncoding yeast DNA yields the linear plot found in human language” reveals the key word “arbitrary.” Note that “words” of the noncoding DNA were three to eight bases, sampled in an unjustified manner. Neither graph appeared to conform to the straight “Line of evidence” of Zipf’s law. The study reported by Flam was based on a comparison with the empirical “Zipf’s law,” which applies to natural languages Zipf (1949). The distribution of frequencies (actual occurrences) of words in a large corpus of data versus their rank is generally a power-law distribution, with exponent close to one. Zipf’s law is thus an experimental law, not a theoretical one. Zipf-like distributions are commonly observed, in many kinds of phenomena. However, the causes of Zipf-like distributions in real life are a matter of some controversy, with DNA being no exception. While the early observations applied to DNA in 1994 were found worthy of reporting in Science and were widely heralded that “something interesting was lurking in the junk (DNA),” the “Zipf-test” was inconclusive. Review by Simons and Pellionisz (2006a) pointed out that investigators failed to detect “well-defined

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scaling or fractal exponents” (Chatzidimitriou et al. 1996) or “any signs of hidden language in noncoding DNA” (Bonhoeffer et al. 1997). Empirical law aside, the biggest problem was the definition of “words” in the DNA. First, Harvard linguistics professor Zipf (1902–1950) established his “law,” based on observations on the English language, in which “words” are taken for granted. He found that in text samples the frequency of any word was roughly inversely proportional when plotted against the rank of how common each word was; the frequency of the k-th most common word in a text was roughly proportional to 1/k. Plotting both frequency and rank on a logarithmic scale, “Zipf’s law” was expected to yield a declining linear graph also for “words” of the DNA. When applying this natural language linguistics to DNA the results were not entirely convincing (Fig. 61.1 of Flam 1994). The problem was not only that the graphs did not quite conform to the linear Zipf’s law. IIt tsaasdIt is unacceptable that the definition in the noncoding DNA was completely and explicitly arbitrary. Of course, there was no definition at that time of what A, C, T, and G strings might constitute “words.” In the analysis conducted by Mantegna et al. (1994): “when the group arbitrarily divided up their samples of junk (DNA) into “words” between 3 and 8 bases long and applied the Zipf test, the telltale linear plot emerged.” Looking at the reproduced Fig. 61.1 of Flam (1994), the plot (for noncoding DNA “words” open squares on a log-log scale) starts fairly close to linear, but drops off remarkably at the tail end. The original Flam diagram of the Zipf’s law for DNA was even more controversial when it was applied to the “coding regions” of the DNA (see graph of open circles in Fig. 61.1 from Flam 1994). Here, Flam claimed that the Zipf’s law “failed” – and the reason cited was that “The coding part (of the DNA) has no grammar – each triplet of bases corresponds to an amino acid in a protein. There’s no higher structure to it.” Today, both the “definition” of arbitrarily picked three to eight letter strings for “words” and the “axiom” that there is no higher structure to coding DNA appear demonstrably dogmatic. Zipf’s law is most easily observed by scatterplotting the data, with the axes being log(rank order) and log(frequency). The simplest case of Zipf’s law is a “1/f function.” Given a set of Zipf-like distributed frequencies, sorted from most common to least common, the second most common frequency will occur 1/2 as often as the first. The nth most common frequency will occur 1/n as often as the first. However, this cannot hold precisely true, because items must occur an integer number of times: there cannot be 2.5 occurrences of a word. Nevertheless, over fairly wide ranges, and to a fairly good approximation, many natural phenomena obey Zipf’s Law.

The Genome is Fractal: Grosberg-School Suspected that the DNA Showed Fractal Folding The classic book of the mathematician who coined the word “fractal” (as a measure of dimension of roughness of results of recursive procedures), by Mandelbrot (1983) generated a huge impetus into the direction of pulling away from looking at the genome as a language, and looking at fractals more as the “geometry of nature.” The twin schools of thought, toward approaching the structure of the

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genome – and the protein structures whose development it governs, manifested in the seminal work by Grosberg et al. (1988, 1993) to claim that the folding of DNA strands were fractal. Decades later, as an eminent example how established methods of biochemistry can be used to support paradigm-shifts, the Science cover article appeared (Erez-Lieberman et al. 2009), in effect the Science Adviser to the US President, Eric Lander appealing “Mr. President, the Genome is Fractal!” Inspired by the Hilbert-curve, a recursive folding that provides the much needed propensities. First, it is knot-free to permit uninterrupted transcription. Second, it is ultra-dense to enable squeezing the 2-m-long DNA strand into the nucleus of a cell with 6 mm diameter. Remarkably, the Hilbert-curve is capable of filling the entire space available, and in its 3D form its fractal dimension is 3.0. Third, it also provides the advantage that is paramount for The Principle of Recursive Genome Function, Pellionisz (2008a, b) that the DNA can be read not only serially, from one end to the thread to the other, but because all segments of the DNA are in maximal proximity to one-other, they can also be read in parallel.

The Perez-School Shows that the DNA is Fractal at DNA, Codon- and Full Chromosome Set and Whole Genome Levels The Perez-school of study of recursive systems was interdisciplinary (Perez 2011b) and showed first results in 1988 (Perez 1988a, 1991).The fractal nature of A, T, C, and G coding or noncoding nucleotide sequences, chromosomes and genomes was evidenced over two decades (see review Perez 2011a). Details, for example, Perez (1991) and Marcer (1992) are comprised in two books (Perez 1997, 2009a). The results spanning from recursive studies through DNA and full genome analysis, including full set of chromosome levels, Perez (2008) is likely to be a serious candidate to the measure of “Abstract DNA Roughness” as proposed in section “Public Domain Agenda in Industrialization of Genomics: Local and Global Fractal Dimension as a Standard Definition for Optimally Distinguishing Cancerous and Control Genomes Based on Their Abstract Measure of “Roughness”.” Fractals to DNA Numerical Decoding: Toward the Golden Ratio “Small is beautiful.” Inspired by the recursive “Game of Life” (Gardner 1970) using the largest computers in the time a cellular automata a large random 0/1 cell populations was run in 1988 (Perez 1988a and 2009b). After 110 parallel network iterations, with a recursive single-line code, a “clown” pattern (see Panel 1 of Fig. 61.5) emerged from the small seven cells “U” (see upper left corner of Panel 1 of Fig. 61.5 from Perez (1988a)). A strong illustration of « small is beautiful » is the discovery of a predictive formula of the Mendeleev’s Elements periodic table architecture (Perez 2009a, b). The “Fractal Chaos” Artificial Neural Network In the 1980s, various parallel artificial neural networks were explored (Perez 1988a 1988b), with a particular interest in discrete waves and by fractals. The fractal chaos is summarized by right-bottom Panel 5 of Fig. 61.5. In the dynamics of the fractal, a curious focal point seems to emerge: the “Golden ratio.” The fractal network also provides “de´ja vu” recall memory and holographic-like memory (Perez 1990a;

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Fig. 61.5 Examples from the Perez-School of Recursive Results. Panel 1: “Clown” emerging from U (upper left corner), citing original recursions in 1988 by Perez (Reproduced from Perez (2008a)). Panel 2: DNA supracode and recursive Fibonacci series: 1 1 2 3 5 8 13 21 34 55 89 . . . Example of resonances in HUMC1A1 gene (Reproduced from Perez (2011a)). Panel 3: Chromosome 1–8. The Evidence of Binary Proteomics Code (red) and Modulated Proteomics Code (blue) at the Whole Human Genome Scale. Green: Genomic, Red: Proteonomic (Reproduced from Perez (2011a)). Panel 3: Chromosome 9-Y (Reproduced from Perez (2011a)) Panel 5: Perez (2010) n. Fractals to DNA numerical decoding: the Golden ratio. Evidence of Golden ratio hypersensitivity in a specific region of the “Fractal Chaos” recursive neural network model (From original figure from (Perez 1997), reproduced on Web (Perez 2009b))

Perez and Bertille 1990). At that time chaos in the DNA was also searched, but it is discrete; A, T, C, and G bases could be coded by integers, while chaos theory is based on real numbers. Note that the ratio between 2 Fibonacci integers is near to the Golden ratio. This raised the question of an integer-based chaos theory. Indeed, a hypersensitivity of the fractal for inputs based on recursive Fibonacci numbers was demonstrated (Perez 1990b).

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“DNA SUPRACODE” Overview A connection between DNA coding regions sequences as gene sequences A, T, C, and G patterned proportions and Golden ratio–based Fibonacci/Lucas integer numbers were proposed (Perez 1991; Marcer 1992, see also Fig. 61.5. Panel 2). Correlation samples were established in genes or gene-rich small genomes with evolution or pathogenicity (example of HIV genome particularly; see the book Perez (1997)). “Resonances” were analyzed, where a resonance is a Fibonacci number of contiguous A, T, C, and G nucleotides (i.e., 144). If this sub-sequence contains exactly 55 bases T and 89 bases C, A, or G, this set was called a “resonance.” Thousands of resonances were discovered (see upper right corner of Panel 2 of Fig. 61.5 from Perez (1991)): in HIV – the whole genome is long of about 9,000 bases – there are resonances overlapping about two third of the genome. In Single-Stranded DNA Human Genome, Codons Population are Fine-Tuned in Golden Ratio Proportions A new Bioinformatics bridge between Genomics and Mathematics emerged (Perez 2010). This “Universal Fractal Genome Code Law” states that the frequency of each of the 64 codons across the entire human genome is controlled by the codon’s position in the Universal Genetic Code table. The frequency of distribution of the 64 codons (codon usage) within single-stranded DNA sequences was analyzed. Concatenating 24 Human chromosomes, it was demonstrated that the entire human genome employs the well-known universal genetic code table as a macro-structural model. The position of each codon within this table precisely dictates its population. So, the Universal Genetic Code Table not only maps codons to amino acids, but also serves as a global checksum matrix. Frequencies of the 64 codons in the whole human genome scale are a self-similar fractal expansion of the universal genetic code. The original genetic code kernel governs not only the micro-scale but the macro-scale as well. Particularly, the six folding steps of codon populations modeled by the binary divisions of the “Dragon fractal paper folding curve” show evidence of two attractors. The numerical relationship between the attractors is derived from the Golden ratio. It was demonstrated that: 1. The whole Human Genome Structure uses the Universal Genetic Code Table as a tuning model. It predetermines global codons proportions and populations. The Universal Genetic Code Table governs both micro- and macro-behavior of the genome. 2. The Chargaff’s second rule from the domain of single A, T, C, and G nucleotides was extended to the larger domain of codon triplets. 3. Codon frequencies in the human genome were found to be clustered around two fractal-like attractors, strongly linked to the Golden ratio 1.618 (Perez 2010). A Strange Meta-Architecture Organizes Our 24 Human Chromosomes A curious interaction network was found among our 24 human chromosomes (Perez 2011a) (see Fig. 61.5, Panels 3–4 for human Chromosomes 1–8 and 9-Y,

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respectively). It was proven that the entire human genome codon population is fine-tuned around the “Golden ratio” (Perez 2010). Across the entire human genome, there appears to be an overall balance in the whole single-stranded DNA. This digital balance fits neatly around two attractors with predominant values of 1 and (3-Phi)/2, where Phi is the Golden ratio. Yet, the same analysis applied individually to each of the 24 chromosomes of humans and to each of the 25 chromosomes of the chimpanzee which reveals a 99.99% correlation between both genomes but diversity and heterogeneity particularly in the case of our chromosomes 16 17 19 20 and 22 (see the book “Codex Biogenesis,” Perez (2009a)). Thus, a paradox emerges. The same analysis shows a global unity across the genome, whereas, applied to each of the constituent chromosomes of this same genome a great heterogeneity between these chromosomes is revealed. With the objective to analyze this paradox in greater depth, a meta-structure was discovered that overlaps all 24 human chromosomes. It is based on a set of strong numerical constraints based particularly on Pi, Phi and integer numbers such as 2, 3, etc. A functionality of this fine-tuned structure appears: the structure is 90% correlated with the density of genes per chromosome from the Human Genome project. It is 89% correlated with the chromosome’s permeability to intrusion by retroviruses like HIV, 94% with CpG density, and 62% with SNP inserts/deletes. Finally, a classification network of the 24 human chromosomes was discovered, including one measuring scale, ranging from 1/Phi (chromosome 4) to 1/Phi + 1/Pi (chromosome 19), which is both correlated with the increasing density of genes and permeability to the insertion of external viruses or vaccines. Unifying All Biological Components of Life: DNA, RNA, Proteins A powerful basic Pi, Phi based numerical projection law of the C O N H S P bioatoms average atomic weights were established (Perez 2009a), and methods will be published in a forthcoming paper (Perez 2012). An integer-based code unifies the three worlds of genetic information: DNA, RNA, and Protein-aggregating amino acids. Correlating, synchronizing, and matching Genomics/Proteomics global patterned images in all coding/noncoding DNA sequences, all biologic data is unified from bio-atoms to genes, proteins, and genomes. This code applies to the whole sequence of human genome, and produces generalized discrete waveforms. In the case of the whole double-stranded human genome DNA, the mappings of these waves fully correlate with the well-known Karyotype alternate dark/gray/light bands. This “unification of all biological components” is illustrated in Panels 3–4 of Fig. 61.5 (Perez 1988a). A complete proof of self-similarity within the whole human genome is provided by Perez (2008). In this “binary code” which emerges from whole human DNA, the ratio between both bistable states is exactly equal to “2” (the space between two successive octaves in music). As shown in Perez (2008) the Top State is exactly matching with a Golden ratio, the Bottom State is also related to the Golden ratio. If PHI ¼ 1.618, it is the Golden ratio, and is phi ¼ 0.618 ¼ 1/PHI, then the “Top” level ¼ phi ¼ 1/PHI and the “Bottom” level ¼ phi/2 ¼ 1/2 PHI. Top/Bottom ¼ 2.

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Neural Net Elements are Fractal: Purkinje Neuron Fractal Model About the same time as the Grosberg-school of thought devoted itself to the analysis of fractal folding of DNA, the School of Recursive Function developed a fractal structural model of a dendritic arborization (Pellionisz 1989). The seminal concept of “recursion” to the DNA to build a fractal neuron is explicitly argued in point 3.1.3 of that paper: “Neural Growth: Structural Manifestation of Repeated Access to Genetic Code”: “One of the most basic, but in all likelihood rather remote, implication of the emerging fractal neural modeling is that it corroborates a spatial ‘code-repetition’ of the growth process with the repetitive access to genetic code. This conceptual link between the two meta-geometries of double helix and ‘fractal seed’ may ultimately lead to precisely pinpointing those exact differences in the ‘genetic’ code that lead to a differentiation to Purkinje-, pyramidal cell, Golgi-cell or other type of specific neurons. It must be emphasized, however, that establishing a rigorous relation of these ‘code sequences’ to the genetic code that underlies the morphogenesis of differentiated neurons may be far in the future.” The Genome is Fractal! Proof of Concept and the Basis of Generalization: Whole Genome Analysis Reveals Repetitive Motifs Conforming to the Zipf-Mandelbrot Parabolic Fractal Distribution Law of the Frequency/Ranking Diagram This chapter decidedly expands on this point to provide support to the generalization, to further detailing a study heralded earlier on the fractality of a whole DNA (Pellionisz 2006; Simons and Pellionisz 2006b; Pellionisz 2009a). With a rapidly increasing number of species in which the whole genome is sequenced and DNA is fully available moreover “motif discovery methods” are increasingly available. See the TEIRESIAS algorithm by Rigoutsos and Floratos (1998), the MEME and MAST algorithms by Bailey and Gribskov (1998), and GEMODA algorithm by Jensen et al. (2006), and Kyle et al. (2006), repetitive “motifs” lend themselves as natural units serving as “words.” This raises not only the necessity, but a possibility to revisit the original Zipf’s law analysis (Flam 1994; Mantegna et al. 1994). In the study reported here, the recently found short, repetitive sequences (“Pyknon”-s) described by Rigoutsos et al. (2006) are used as more natural “words” than completely arbitrarily picked three to eight nucleotide sequences. In the human DNA, they found about 128,000 short, repetitive sequence elements, apparently indiscriminately distributed over coding as well as noncoding regions of the DNA. Therefore, there is no need, indeed no basis to separate “words” occurring in the DNA either in the regions of “genes” or what used to be called “junk” DNA. In addition, the short, repetitive sequence motifs mined by the TEIRESIAS algorithm Rigoutsos and Floratos (1998) showed no apparent difference in occurrence either in the “coding” or “noncoding” regions. Using the web-interface by the Group of Rigoutsos at IBM Watson Research Center http://cbcsrv.watson.ibm.com/Tspd.html a “pyknon-type” motif discovery

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Fig. 61.6 Zipf-Mandelbrot Parabolic Fractal Distribution Curve of short repetitive DNA sequences in the whole genome of Mycoplasma genitalium. Frequency as a function of rank is parabolic on a log-log scale, after Pellionisz (2009a). See detailed explanation the reference and in the text below

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was made for the whole genome of the Mycoplasma genitalium, the smallest DNA known Fraser et al. (1995). The web-interface returned the list of short, repetitive DNA sequences in the order of their ranking (as integers) with the frequency of occurrence (also as integers). Results immediately lend themselves to a log-log plotting of the frequency (y) against ranking (x), as seen in the graph below. Figure 61.6 shows the frequency (y) plotted against ranking (x) of “PyknonLike-Elements” short repetitive sequences (PLE-s) of the whole DNA of Mycoplasma genitalium. Results reveal a “Zipf-Mandelbrot Parabolic Fractal Distribution.” Both frequencies and occurrences are shown on a log-log scale. Note that the actual distribution is distinctly different from the linear Zipf’s law. More detailed analysis of the above results reveals by standard curve-fitting that the data can be modeled by the generalization of Zipf’s Law, defined as the Zipf-Mandelbrot Parabolic Fractal Distribution. The Zipf-Mandelbrot function is given by f ðk; N; q; sÞ ¼

1=ðk þ qÞs HN;q;s

where HN,q,s is given by HN;q;s ¼

N X i¼1

1 : ði þ qÞs

This may be thought of as a generalization of a harmonic number. In the limit as N approaches infinity, this becomes the Hurwitz zeta function z(q,s). For finite N and q ¼ 0 the Zipf-Mandelbrot law becomes Zipf’s law. For infinite N and q ¼ 0 it becomes a Zeta distribution.

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1

2

3

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Fig. 61.7 Curve-fitting (in purple) of the frequency (y) against ranking (x) of “Pyknon-type” short repetitive sequences of the whole DNA of Mycoplasma genitalium (in blue). The curve reveals a Zipf-Mandelbrot “Parabolic Fractal Distribution” that can be approximated by the quadratic polynomial of y ¼ 0.052x2  0.0015x + 1.71, after Pellionisz (2009b); see detailed explanation in the reference and in the text below

In the Parabolic Fractal Distribution, the logarithm of the frequency or size of entities in a population is a quadratic polynomial of the logarithm of the rank; standard curve-fitting approximates the data with the quadratic polynomial y ¼ 0.052x2  0.0015x + 1.71. As in typical cases, there is a so-called King effect where the highest-ranked item(s) tend(s) to exhibit a significantly greater frequency or size than the model predicts on the basis of the other items. Data by the Rigoutsos et al. (2006) motif discovery reveal the Zipf-Mandelbrot Parabolic Fractal Distribution curve of frequency against ranking of short repetitive sequences in the entire genome (full DNA) of a free-living organism. It is noteworthy that for the analysis no distinction between the “protein-coding” and “nonprotein-coding” DNA segments need to be made. Nonetheless, one might argue that since the DNA of the Mycoplasma genitalium contains only 6 months) tg/tg mice have swellings, irregular microtubules, and an increase in cytoplasmic organelles (Meier and MacPike 1971; Ryo et al. 1993). Many of the morphological alterations are progressive with an onset between 3 and 5 weeks, the same time frame as the appearance of Purkinje cell dendritic abnormalities and the onset of ataxia (Green and Sidman 1962; Seyfried and Glaser 1985; Ryo et al. 1993).

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In the tg/tg mouse, the terminal noradrenergic axons of the locus coeruleus increase in all areas, including the cerebellum (Levitt and Noebels 1981). There is a concomitant increase in norepinephrine levels (Levitt and Noebels 1981; Levitt et al. 1987). The number and size of the neurons in the locus coeruleus are unaltered. An increase in basal cAMP levels in the neocortex of the tg/tg mouse has been suggested to reflect the enhanced norepinephrine innervation (Tehrani and Barnes 1995). Tyrosine hydroxylase (TH) and its mRNA are abnormally high in the Purkinje cells of both young and adult tg/tg mice (Hess and Wilson 1991; Austin et al. 1992; Fletcher et al. 1996). Increased TH expression is limited to the soma and dendrites of Purkinje cells but is widespread throughout the cerebellum. In normal mice, the expression in Purkinje cells is transient, with only small numbers of Purkinje neurons retaining TH into adulthood. The increased TH expression in the tg/tg mouse occurs in a zebrin II parasagittal banding pattern but zebrin II staining is normal (Hawkes and Herrup 1995; Fletcher et al. 1996; Abbott et al. 1996). The expression of n-nitric oxide synthase (n-NOS), both mRNA and protein, is elevated in the adult tg/tg mouse (Rhyu et al. 2003). The calretinin mRNA in the granule cell layer and ryanodine receptor type 1 in Purkinje cells decrease in the adult animal (Cicale et al. 2002). How these changes in TH, n-NOS, or the genes involved with calcium regulation affect cerebellar function is unclear. In the adult tg/tg mouse there are marked changes in GABAA receptors. In the cerebral cortex, GABAA receptor function decreases and there are changes in the subunit composition and binding properties that may contribute to the absence seizures (Tehrani and Barnes 1990, 1995). In the cerebellum there is a 40% reduction in the number of GABAA receptors in the granular layer with a large component of the loss in a6 and g2 subunits (Kaja et al. 2007a). Whether these changes in GABAA receptors are a primary deficit or compensatory has not been studied. However, the reduction in GABAA receptors has the potential to contribute to abnormal signal processing in the cerebellar cortex.

Tottering Mouse and P/Q-Type Ca2+ Channel Dysfunction At the biophysical level, the mutation in the tg/tg mouse results in a loss of P/Q-type channel function. There is a 30–40% reduction in the Ca2+ current in Purkinje cells with little alteration in the channel kinetics (Wakamori et al. 1998). The P/Q-type channels in the tg/tg mouse do not show a shift of activation to more depolarized voltages as found in EA2 patients and other EA2 mouse models (Wakamori et al. 1998; Mori et al. 2000; Guida et al. 2001; Jen et al. 2001; Wappl et al. 2002). It has been suggested that a decrease in the expression of functional channels in the tg/tg mouse may contribute to the reduction in Ca2+ current (Pietrobon 2010). The P/Q-type Ca2+ channels have a wide distribution in the nervous system and are found in the presynaptic terminals, soma, and dendrites of neurons (Mintz et al. 1992; Westenbroek et al. 1995; Fletcher et al. 1996). High to moderate levels of expression are found in the cerebellum, cerebral cortex, hippocampus, and olfactory bulb. Based on alternative splicing at multiple loci in the gene, as well as

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functional differentiation of CaV2.1 channels due to combination with various auxiliary subunits, there is extensive variation in the properties of P/Q-type Ca2+ channels throughout the brain and spinal cord (Mermelstein et al. 1999; Luvisetto et al. 2004). Given their location at presynaptic terminals, the P/Q-type Ca2+ channels are major contributors to the release of neurotransmitters at CNS synapses (for review, see Catterall 1998; Pietrobon 2002; Pietrobon 2010). Although N (Cav2.2)- and R (Cav2.3)-type Ca2+ channels work in concert with P/Q-type channels, the latter appear to be more efficiently coupled to the mechanisms governing vesicle release (Mintz et al. 1995; Wu et al. 1999; Li et al. 2007). In part, the increased efficiency reflects the preferential location of P/Q-type channels near release sites at some synapses (Wu et al. 1999). There is abundant expression of P/Q-type Ca2+ channels in cerebellar granule and Purkinje cells (Mintz et al. 1992; Westenbroek et al. 1995). Located presynaptically at the parallel fiber–Purkinje cell synapse, P/Q-type channels generate approximately 50% of the calcium currents in granule cells (Randall and Tsien 1995). P/Q-type Ca2+ channels dominate synaptic transmission at the parallel fiber– Purkinje cell and climbing fiber–Purkinje cell synapses (Mintz et al. 1995). Inhibitory synaptic transmission between Purkinje cells and their targets in the cerebellar nuclei is almost exclusively dependent on P/Q-type channels in the mouse by the third week (Iwasaki et al. 2000). Similarly, P/Q-type channels are exclusively operative at inhibitory synapses between molecular layer interneurons and Purkinje cells, between Purkinje cells via their axon collaterals, and Golgi cell inhibitory synapses on the granule cells (Forti et al. 2000; Stephens et al. 2001; Lonchamp et al. 2009). Therefore, a reduction in P/Q-type channel function has the capacity to alter fast synaptic transmission throughout the cerebellum. In many CNS synapses the dominance of P/Q-type channels mediating synaptic transmission is developmental and well-functioning by the third week (Iwasaki et al. 2000), just prior to, or at the time of onset of symptoms in the tg/tg mouse (Green and Sidman 1962; Meier and MacPike 1971). Parallel fiber–Purkinje cell synaptic transmission is decreased in adult tg/tg mice but only mildly decreased in younger mice (Matsushita et al. 2002). This agerelated decrease in parallel fiber–Purkinje cell synaptic transmission is consistent with the development of the behavioral phenotype. A decrease in parallel fiber– Purkinje cell synaptic transmission in adult mice was also found in vivo (Chen et al. 2009). However, one slice study failed to find a change in the excitatory postsynaptic current (EPSC) evoked by parallel fibers (Zhou et al. 2003). Paired-pulse facilitation is normal at the parallel fiber–Purkinje cell synapse, implying that evoked glutamate release is intact (Matsushita et al. 2002). Although P/Q-type Ca2+ channels are expressed in the presynaptic climbing fiber terminals (Mintz et al. 1995), the EPSC amplitude of the climbing fiber–Purkinje cell synapse is not altered in the tg/tg mouse (Matsushita et al. 2002). This suggests that the deficits in presynaptic transmitter release are not as uniform or pervasive as predicted. The release of glutamate from parallel fibers in the tg/tg mouse is controlled by N-type channels as assessed by the fraction of the excitatory postsynaptic potentials (EPSPs) blocked by o-conotoxins in contrast to P/Q-type channels in wild-type

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animals (Zhou et al. 2003). The increase in N-type Ca2+ channels on the presynaptic parallel fibers is thought to account for the increase in baclofen inhibition and the prolongation of heterosynaptic depression (Zhou et al. 2003). Adrenergic modulation is greatly increased at the parallel fiber–Purkinje cell synapse. Noradrenaline and other a-2 noradrenergic agonists inhibit parallel fiber–Purkinje cell synaptic transmission and this inhibition is potentiated in tg/tg mice. Although the location of the responsible receptors is unclear, the increased sensitivity is interesting given the increased noradrenergic innervation (Levitt and Noebels 1981; Noebels 1984), and that stress is a trigger for the motor attacks in the tg/tg mouse (Campbell and Hess 1998; Fureman et al. 2002). Alterations in synaptic transmission occur throughout the nervous system in the tg/tg mouse. Excitatory but not inhibitory synaptic transmission is reduced in the ventrobasal thalamic nucleus (Caddick et al. 1999). There is a profound reduction in presynaptic P/Q-type Ca2+ currents in CA3-CA1 hippocampal synaptic transmission; however, this loss is compensated for by an increase in N-type currents (Qian and Noebels 2000). At the neuromuscular junction the changes include increased run-down of high-rate evoked release and an increase in spontaneous acetylcholine (ACh) release (Plomp et al. 2000). However, these alterations are subclinical as the dynamics of saccadic eye movements are normal in young tg/tg mice, suggesting that neuromuscular transmission is functionally intact (Stahl et al. 2006). The lack of clinically evident neuromuscular deficits reflects the upregulation of R-type and N-type Ca2+ channels that compensates for the reduction in P/Q-type channel function (Pardo et al. 2006; Kaja et al. 2007b). In general, compensation for loss of P/Q-type currents by other voltage-gated Ca2+ channels mitigates the deficits in synaptic transmission in the tg/tg mouse. Potentially as important as altered synaptic transmission for cerebellar dysfunction in the tg/tg mouse, P/Q-type channels are the dominant type of voltage-gated Ca2+ transmitter on Purkinje cells and account for 85–95% of their total Ca2+ current (Mintz et al. 1992; Usowicz et al. 1992; Jun et al. 1999). The loss of P/Q-type Ca2+ currents in Purkinje cells precedes the development of the tg/tg phenotype and is due to a failure in the normal switch from L-type to P/Q-type currents that occurs in the first 2 postnatal weeks (Wakamori et al. 1998; Erickson et al. 2007). Both parallel fiber and climbing fiber inputs result in postsynaptic activation of these channels on Purkinje cells (Hartmann and Konnerth 2005). The P/Q-type Ca2+ channels on Purkinje cells coordinate AMPA receptor activation with voltage-dependent calcium flux (Denk et al. 1995). These Ca2+ currents are integral to the function of the Purkinje cell. For example, the increased intracellular Ca2+ in Purkinje cells is essential to parallel fiber–Purkinje cell synaptic plasticity, both long-term potentiation and long-term depression (for reviews see Ito (2001), Jorntell and Hansel (2006)). The level of Ca2+ is thought to act as the switch that regulates whether parallel fiber input evokes long-term potentiation or long-term depression (Jorntell and Hansel 2006). The P/Q-type Ca2+ channels play an important role in the spontaneous, intrinsic firing of Purkinje cells via their coupling to KCa channels (Womack and Khodakhah 2002). Reduction in P/Q-type Ca2+ current increases the firing and irregularity of

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Purkinje cell simple spikes, and blocking results in bursting followed by silence. Loss of intrinsically driven simple spike firing has been shown in cerebellar slices from the tg/tg mouse as well in the leaner and ducky mutants. In both of these latter models, P/Q-type currents are reduced, although the mutation in the Cacna1a gene effects the a1A subunit in the leaner and the a2d2 subunit in the ducky (Lorenzon et al. 1998; Barclay et al. 2001), suggesting that irregularity in Purkinje cell firing is a common aspect of P/Q-type Ca2+ channelopathies. Therefore, altered P/Q-type Ca2+ channels on Purkinje cells likely play a major role in the cerebellar deficits in the tg/tg mouse (Stahl et al. 2006). The misregulation of Ca2+ signaling in the tg/tg mouse extends beyond the loss of P/Q-type channel function. L-type Ca2+ channels are upregulated in the tg/tg mouse in Purkinje cells but not in cerebellar granule cells or nuclear neurons (Campbell and Hess 1999; Erickson et al. 2007). In the Cav2.1 null mouse, which does not have a paroxysmal motor phenotype, L-type current increases in granule cells (Fletcher et al. 2001). With the exception of the nucleus of the brachium of the inferior colliculus, other CNS regions do not exhibit a significant increase in L-type Ca2+ channels or binding in the tg/tg mouse. There is no evidence for changes in a12.1 mRNA levels (Doyle et al. 1997). The importance of this upregulation is that L-type Ca2+ channel blockers prevent the episodic dystonia and low frequency oscillations in the cerebellar cortex and agonists induce the attacks and the oscillations (Campbell and Hess 1999; Chen et al. 2009). Clearly, the upregulation of L-type Ca2+ channels is involved in the episodic motor phenotype of the tg/tg mouse.

Cerebellar Circuit Function and Episodic Abnormalities There have only been a few studies examining the properties of the cerebellar circuitry in the tg/tg mouse in vivo. Simple spike firing of Purkinje cells exhibits increased variability in the tg/tg mice, and the increased variability is induced by blocking P/Q-type channels in wild-type mice (Hoebeek et al. 2005; Walter et al. 2006). The tg/tg mice also exhibit shorter pauses in simple spike firing after the complex spike, the so-called inactivation period. Simple spike firing irregularity is present during the compensatory eye movements in response to sinusoidal optokinetic stimulation, although the depth of modulation is normal. Spontaneous climbing fiber discharge and their spatial tuning during optokinetic stimulation do not differ in the tg/tg and wild-type mice. As discussed above, the Purkinje cell terminals in the cerebellar nuclei show progressive structural damage (Hoebeek et al. 2008). Electrophysiological studies reveal that stimulation in the cerebellar cortex evokes inhibition in nuclear neurons with a normal threshold and latency. However, the spontaneous firing of nuclear neurons is greater and more irregular than in wild-type animals, pointing to altered synaptic transmission between Purkinje cells and their targets. One of the most prominent and intriguing aspects of the channelopathies is the episodic CNS dysfunction (Kullmann 2002; Ryan and Ptacek 2010). It is unknown

T.J. Ebner and G. Chen

Fig. 67.1 (continued)

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how a chronic reduction in P/Q-type Ca2+ channel function produces intermittent cerebellar ataxia in EA2 patients or the episodic dystonia in the tg/tg mice. Changes in parallel fiber synapses, alterations in parallel fiber–Purkinje cell synaptic transmission, and increased variability in simple spike firing are inherently “static” abnormalities. While likely contributing to the baseline ataxia or changes in oculomotor reflexes, it is not obvious that these abnormalities underlie the transient attacks of dystonia. During the motor attacks, one would predict large, transient disruptions in motor circuits, including the cerebellum, as found with immediate early gene expression (Campbell and Hess 1998). Flavoprotein optical imaging revealed a novel form of activity in the cerebellar cortex of the tg/tg mice in vivo: episodic, low frequency oscillations (Fig. 67.1a–b, Chen et al. 2009) were observed in the cerebellar cortex of tg/tg mice (Chen et al. 2009). The oscillations consist of large amplitude increases and decreases of fluorescence at 0.04–0.08 Hz that suggest large fluctuations in cell activity (Reinert et al. 2004, 2007). A region of a folium oscillates in phase within a narrow range of frequencies suggesting that a population of neurons is in synchrony (Fig. 67.1c, Chen et al. 2009). However, neighboring regions oscillate at varying mean frequencies and are not in phase. Oscillations at these frequencies have not been observed in wild-type animals or other murine models of cerebellar disease. The oscillations are spontaneous, transient, and propagate (Chen et al. 2009). Spontaneously, the oscillations increase in amplitude in a region, spread to neighboring regions, and then subside over 30–120 min. When strong oscillations are present, the beam-like response evoked by parallel fiber stimulation is markedly reduced, suggesting that cerebellar cortical physiology is highly abnormal during the oscillations. The spontaneous firing of cerebellar Purkinje cells and unidentified cerebellar neurons also exhibit low frequency oscillations in the tg/tg mouse (Fig. 67.1d, Chen et al. 2009). These oscillations likely contribute to the

ä Fig. 67.1 (a) Sequential images of the cerebellar cortex show spontaneous oscillations in an anesthetized tg/tg mouse. Large amplitude oscillations are present in the paramedian lobule (PML) and lower amplitude oscillations in Crus I and II. Each image shows pseudocolored pixels that are above the background fluorescence level (DF/F). Time from image acquisition onset is indicated in the upper right corner of each image. (b) Timecourse of DF/F obtained from three regions of interest (ROIs) indicated in the first image of (a) (colored boxes of 20  20 pixels). (c) Pixel based spectral analysis shows the frequency maps for the same experiment. Each map is superimposed on a background image of the cerebellar cortex. (d) Example of the spontaneous firing of a Purkinje cell in a tg/tg mouse. (e) Power maps of the awake tg/tg mice during baseline and dystonia induced by caffeine. (f) Frequency distributions from a single mouse for the optical (bars) and hamstring EMG recordings (lines) during dystonia (red) and baseline (blue) periods. Inset shows regular, low frequency bursts in the hamstring EMG during dystonia. (g) Diagram of hypothesized mechanism of how the oscillations in Purkinje cells (PC) may effect the deep Cerebellar nuclei (DCN) and contribute to the episodic dystonia (with permission from Chen et al. (2009))

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variability of Purkinje cell firing in the tg/tg mouse (Hoebeek et al. 2005; Walter et al. 2006). The oscillations are likely generated within the cerebellar cortex. Blocking AMPA receptors, the dominant ionotropic glutamate receptors at parallel fiber–Purkinje cell and climbing fiber–Purkinje cell synapses (Konnerth et al. 1990; Perkel et al. 1990), did not significantly alter the oscillations (Chen et al. 2009). Furthermore, stimulation of the parallel fibers over a wide range of parameters did not initiate or modify the oscillations. Removal of Ca2+ from the bathing solution significantly reduces the power and area of the oscillations. Blocking L-type Ca2+ channels reduces the oscillations and L-type Ca2+ channel agonists increase the oscillations, demonstrating that L-type Ca2+ channels play a major role in the generation of the oscillations. The low frequency oscillations are also present in the cerebellar cortex of awake tg/tg mice (Chen et al. 2009). Imaging in the awake tg/tg mouse shows that caffeine not only triggers the motor attacks but also accentuates the oscillations (Fig. 67.1e, Chen et al. 2009). During the periods of dystonia, there are bursts of EMG activity at the same low frequencies observed in the cerebellar cortex (Fig. 67.1f, Chen et al. 2009). In the baseline period, there is little significant coherence between the optical and EMG signals. During caffeine-induced episodes of dystonia, the coherence level increases. Therefore, the cerebellar cortical oscillations increase and are coupled with the abnormal EMG activity during the motor attacks. The low frequency oscillations in the cerebellar cortex provide a mechanism for generating the episodic neurological dysfunction in the tg/tg mouse. Both the oscillations and the motor attacks are transient and triggered by caffeine (Fureman et al. 2002). Although the cellular mechanisms of the oscillations are not known, a plausible mechanism is the interplay between Ca2+ entering through the L-type Ca2+ channels and Ca2+ released from inositol triphosphate (IP3) and ryanodinesensitive intracellular stores in Purkinje cells (Kano et al. 1995; Finch and Augustine 1998; Carter et al. 2002). The synchronization and propagation of the oscillations occur over 30–120 min, consistent with the time course of the motor attacks (Green and Sidman 1962; Noebels and Sidman 1979). The low-frequency bursts in the EMG activity during the attacks of dystonia and the increase in the coherence between the cerebellar and EMG activity suggest that the abnormal cerebellar oscillations are integral to the episodic motor attacks. It was hypothesized that the reduction in functioning P/Q-type channels and the upregulation of the L-type channels in the cerebellum leads to low-frequency, baseline oscillations in Purkinje cells (Fig. 67.1d, Chen et al. 2009). Triggering factors such as stress, ethanol, or caffeine increase the amplitude of the baseline oscillations which, in turn, could synchronize groups of cells and the oscillations propagate to neighboring regions. The mechanisms of synchronization and propagation are unknown. The oscillations in Purkinje cell output will be transmitted downstream to the cerebellar nuclei (Campbell and Hess 1998). Due to the extensive convergence of Purkinje cells on nuclear neurons, the synchronization of Purkinje cell activity over a region of the cerebellar cortex is likely to be amplified

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at the level of the cerebellar nuclei. The targets of this highly abnormal cerebellar output, including the red nucleus, motor thalamus, and motor cortex will, in turn, be extensively modulated (Campbell and Hess 1998) and provide a powerful drive to the muscles through descending projections to the spinal cord to generate the episodic dystonia.

Triggers of Episodic Dystonia Triggers are a conspicuous feature of the episodic dystonia in tg/tg mice and the ataxia in EA2 patients. Given that the episodic motor attacks in the tg/tg mouse require functioning Purkinje cells (Campbell et al. 1999; Erickson et al. 2007), and the dependence on upregulation of L-type Ca2+ channels on these cells (Campbell and Hess 1999), intracellular Ca2+ homeostasis in Purkinje cells is likely to play a role. A hypothesized common pathway for the triggers is activation of ryanodine receptors (RyR) and mobilization of internal Ca2+ stores from the endoplasmic reticulum (Fureman et al. 2002; Fureman and Hess 2005; Chen et al. 2009). The xanthine compounds, such as caffeine and theophylline, evoke motor attacks in several rodent models of dystonia and in tg/tg mice (Fureman et al. 2002; Raike et al. 2005). Caffeine evokes large increases in intracellular Ca2+ from RyR sensitive stores in Purkinje cells but not from parallel fibers (Kano et al. 1995; Carter et al. 2002). Stress, both physiological and psychological, is a powerful trigger. Physical restraint and changes in the environment reliably induce the episodic dystonia in tg/tg mice (Kaplan et al. 1979; Syapin 1983; Campbell and Hess 1998). The locus coeruleus and noradrenergic neurotransmission have been widely implicated in mediating the effects of stress (for review see Sved et al. (2002); Berridge (2008)). Stress increases the firing of neurons in the locus coeruleus (Abercrombie and Jacobs 1987). The locus coeruleus has widespread norepinephrine projections throughout the CNS including the cerebellar cortex (Olson and Fuxe 1971; Pickel et al. 1974). Stimulation of the locus coeruleus elicits a prolonged inhibition of Purkinje cell firing (Hoffer et al. 1973; Woodward et al. 1979). However, norepinephrine has complex excitatory and inhibitory effects on cerebellar neurons depending on the specific adrenergic receptor and cell type involved (Kondo and Marty 1997; Saitow et al. 2000; Hirono and Obata 2006). Blocking a-adrenergic receptors decreases the frequency of the motor attacks but not the severity or duration (Fureman and Hess 2005). Interestingly, b-adrenergic agents have no clear effect on the motor attacks, although these receptors are located on Purkinje cells and molecular layer inhibitory interneurons (Kondo and Marty 1997; Hirono and Obata 2006). In contrast, a-adrenergic receptors are located on cerebellar interneurons and Bergmann glia (Kulik et al. 1999; Hirono and Obata 2006). As described above, there is hyperinnervation by the locus coeruleus in the tg/tg mouse and upregulation of TH in the cerebellum (Levitt and Noebels 1981; Noebels 1984; Hess and Wilson 1991; Austin et al. 1992). However, lesions of the locus coeruleus do not affect the episodic dystonia (Noebels 1984; Campbell et al. 1999),

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suggesting that the hyperinnervation does not underlie how stress acts as a trigger for the episodic dystonia. The mechanisms by which stress and norepinephrine initiate motor attacks in the tg/tg mouse remain to be determined.

Assessment of EA2 Therapies in tg/tg Mouse The tg/tg mouse also serves as a model system to develop and test new therapies for EA2. Until recently the most effective therapy has been acetazolamide, a blocker of carbonic anhydrase (Griggs et al. 1978; Strupp et al. 2007). Although the mechanism of action of acetazolamide is unknown, it has been hypothesized to alter Purkinje cell excitability and increased Purkinje cell–nuclear neuron synaptic transmissions (Strupp et al. 2004). Recently, nonselective blockers of voltagegated potassium channels (e.g., 4-aminopyridine) have been shown to significantly reduce the frequency and severity of the episodic cerebellar symptoms in EA2 (Strupp et al. 2004). Potassium channel blockers also reduce the frequency but not the duration of the episodic motor attacks in the tg/tg mouse (Weisz et al. 2005). Initially, it was suggested that 4-aminopyridine restores Purkinje cell inhibitory drive on the cerebellar nuclei (Kalla et al. 2007; Strupp et al. 2008). However, the frequency of Purkinje cell firing and Purkinje cell–nuclear neuron synaptic transmission is normal (Hoebeek et al. 2005, 2008). The irregularity in Purkinje cell simple spike firing of the tg/tg mouse has been linked to the loss of KCa channel function, particularly the small conductance type (Walter et al. 2006). In tg/tg mice, activation of the SK channel can regularize the simple firing of Purkinje cells in the slice and improve rotorod performance (Walter et al. 2006; Alvina and Khodakhah 2010). Importantly, activation of SK channels does not increase Purkinje cell firing rate but does increase the postsynaptic response to parallel fiber input (Alvina and Khodakhah 2010). The KCa channel activator, chlorzoxazone, given orally also reduces the severity, frequency, and duration of stress-induced episodic dystonia, although chlorzoxazone does not prevent the attacks (Alvina and Khodakhah 2010). Therefore, the tg/tg mouse has the potential to serve as a model for the screening of new EA2 therapies.

Summary The tg/tg mouse serves as a powerful model of a human channelopathy, EA2. The tg/tg mouse has and will continue to provide insights into the mechanisms and treatment of episodic neurological dysfunction across the molecular, cellular, and systems levels. However, there remains a great deal to learn. After 50 years, researchers are still trying to decipher the neuronal basis of episodic motor attacks and how the attacks are initiated. Answering these questions in this mouse model of a Cav2.1 channelopathy would not only benefit EA2 patients but also provide a template to understand episodic neurological dysfunction more generally.

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Jaap J. Plomp and Arn M. J. M. van den Maagdenberg

Abstract

The natural mutant mouse rolling Nagoya has an uncoordinated gait and frequently displays sideways body rolls. The mutation underlying this autosomal recessively inherited severely ataxic motor phenotype is present in Cacna1a, the gene encoding the pore-forming a1 subunit of CaV2.1 type voltage-gated Ca2+ channels. This type of channel is crucially involved in neuronal Ca2+ signaling and in neurotransmitter release from nerve terminals at many central synapses and, in the periphery, at the neuromuscular junction. This chapter reviews the phenotypic, motor behavioral, histological, biochemical, neurophysiological, and electrophysiological findings in this mouse mutant. Human neurological diseases exist which are associated with CaV2.1 dysfunction (“Ca2+-channelopathy”), either due to CACNA1A mutation or autoimmune attack. The relevance of the rolling Nagoya mouse mutant as a model for these diseases is discussed.

Introduction Rolling Nagoya (RN) is a natural mutant mouse with a recessive neurological phenotype characterized by severe ataxia and frequent lurching (reviewed in Plomp et al. 2009). The mutant was discovered and first described in 1973 (Oda 1973). In the following years, the phenotype and cerebellum morphology have been studied in detail. Investigations on this mouse mutant intensified since the year

J.J. Plomp (*) Departments of Neurology and Molecular Cell Biology – Group Neurophysiology, Leiden University Medical Centre, Research Building, S5P, NL-2300 RC Leiden, The Netherlands e-mail: [email protected] A.M.J.M. van den Maagdenberg Departments of Neurology and Human Genetics, Leiden University Medical Centre, NL-2300 RC Leiden, The Netherlands M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 1541 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_68, # Springer Science+Business Media Dordrecht 2013

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2000 when the causative mutation was found to reside in the Cacna1a gene (Mori et al. 2000). This gene codes for the pore-forming a1 subunit of voltage-gated Ca2+ channels of the CaV2.1 type, which have been previously named P/Q-type channels. Neuronal voltage-gated Ca2+ channels are multi-subunit membrane proteins that translate a depolarizing voltage change of the plasma membrane into cellular influx of Ca2+ along its electrochemical gradient. The ensuing (local) cytosolic increase in Ca2+ concentration can trigger a multitude of secondary signaling cascades. Although being present to a certain degree in neuronal cell somata membrane (Mintz et al. 1992), CaV2.1 type channels preferentially localize to the presynaptic nerve terminals of many central synapses where they play an important role in mediating the release of neurotransmitter (Westenbroek et al. 1995, 1998). In most central nerve terminals, including those in the cerebellum, CaV2.1 channels act in concert with CaV2.2 (N-type) and CaV2.3 (R-type) channels (Mintz et al. 1995). CaV2.1 channels also are present at synapses in the peripheral nervous system. One important site is the neuromuscular junction (NMJ), which serves to transmit impulses from motor neuron onto skeletal muscle fibers. At these synapses, CaV2.1 is the sole channel type responsible for the Ca2+ influx mediating neurotransmitter release (Uchitel et al. 1992). Interestingly, mutations in the orthologous human gene CACNA1A have been shown to underlie disease in patients suffering from rare inherited forms of migraine and ataxia (Ophoff et al. 1996; Zhuchenko et al. 1997). Furthermore, CaV2.1 channels were shown to be the autoimmune targets at the NMJ in the neuroimmune disorder Lambert-Eaton myasthenic syndrome (LEMS) (Lennon et al. 1995), which is characterized by muscle weakness and is sometimes accompanied by cerebellar ataxia. The discovery of human disease-associated CACNA1A mutations makes the finding of the RN mutation in Cacna1a of particular interest because it implies that RN may be a relevant model for such diseases and could be used in drug studies. Besides RN, an array of other CaV2.1 mouse mutants has been discovered (for reviews, see Felix 2002; Pietrobon 2005) that also may have such modeling roles for human CaV2.1-channelopathies. This chapter will provide a detailed overview of the neurochemical, physiological-, and morphological studies performed in the hetero- and homozygous RN mouse and will discuss the relevance of the RN mouse for human ataxia, migraine and neuromuscular synapse dysfunction.

Phenotypic Description of the Rolling Nagoya Mouse The RN mutant mouse was identified in Nagoya, Japan, more than 35 years ago by Oda when experimenting with matings between two different inbred strains, called SIII and C57Bl/6Nga (Oda 1973, 1981). The autosomal recessive RN mutation was subsequently back-crossed onto a C3Hf/Nga background to remove a reduced fertility characteristic present within the SIII strain (Tamaki et al. 1986). It appeared that the RN mutation was a new allele of the tottering locus, which had been mapped to chromosome 8 (Oda 1981). Tottering is another natural mutant mouse, now known to harbor a recessive Cacna1a mutation (Doyle et al. 1997; Fletcher et al. 1996).

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Fig. 68.1 Some phenotypic characterizations of rolling Nagoya mice. (a) Photograph of a homozygous RN mouse rolling on its back. Normal wild-type littermate at the top of the panel. (b) Ataxic gait with sideways roll of a homozygous RN mouse. (c) Reduced body weight of homozygous RN mice at 3 months of age (Kaja et al. 2007c). (d) Body weight gain during the first three postnatal weeks of homozygous RN (rol/rol), heterozygous RN (rol/+) and wild-type mice (+/+) mice (n ¼ 10), showing a growth lag of homozygous RN mice starting from day 8. Wild-type and heterozygous mice have identical body weight gain. (e) Spectrum of neonatal hind limb reflex scores, from fully normal (score 4) to fully abnormal (score 1) in a tail-suspension test. (f) Mean scores of groups of mice (n ¼ 10), showing impaired hind limb positioning starting around postnatal day 12. * p < 0.05, *** p < 0.001 (Panels d, e, and f reprinted from Takahashi et al. (2010), Copyright 2010, with permission from Elsevier)

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The most obvious phenotype in homozygous RN mice is the clear ataxic gait, predominantly visible as abnormal cyclic movements of the hind limbs, and frequent sideways roll-overs (Fig. 68.1a, b). RN mice do not display body or limb tremor, either during movement or at rest. Neither do they have overt symptoms of epilepsy, in contrast to many of the other natural Cacna1a mouse mutants (Felix 2002). Body

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weight of adult RN mice is reduced by 25–30% (Nakamura et al. 2005; Kaja et al. 2007c) (Fig. 68.1c). Weight gain lag commences at about 8 days postnatally, as can be detected when comparing large groups of homozygous RN neonatal mice with heterozygous and wild-type animals (Takahashi et al. 2010) (Fig. 68.1d). The motor deficits of the hind limbs and balancing difficulties become clearly noticeable upon visual inspection between postnatal days 10 and 14, and can be detected as early as 8 days after birth if specific neuromuscular tests are performed (Takahashi et al. 2010). A simple test is to observe and score the posture of the hind limbs when the (neonatal) mouse is suspended by the tail. RN mice spread their hind limbs much less far than the unaffected littermates (Fig. 68.1e, f). It is unknown which kind of (brain) dysfunction is represented by this abnormal behavioral response. The motor symptoms in adult RN males make coitus more difficult, resulting in reduced reproducibility of homozygous RN mice. RN females are fertile but produce less surviving offspring due to reduced nursing abilities (Oda 1981). More successful breeding of the RN strain is achieved by heterozygous matings, roughly resulting in Mendelian ratios (25%/50%/25%) of wild-type, heterozygous and homozygous RN offspring. During a few weeks after the onset of symptoms there is an increased chance of death of homozygous RN mice. However, once a homozygous RN mouse has successfully gone through this stage, it has a normal life span (Oda 1981), an observation which is confirmed in breedings in other laboratories. Motor performance of adult homozygous RN mice has been studied in great detail (Tamaki et al. 1986). Compared to unaffected littermates (heterozygous RN and wild-types), they are less well able to sustain hanging onto a horizontal wire. Furthermore, they fall off more easily when placed nose-downwards on a thick vertical rope or descend the rope in an immature way, normally only to be observed in very young unaffected littermates. Similarly, RN mice are less well able to walk on a narrow (12 mm) path 50 cm above the ground, with frequent falls as a consequence. In gait analysis from footprint patterns, RN mice appear to have a more waddling stride with now and then performing double steps with the same limb. In swimming tests, RN mice swim slowly and clumsy with disturbed rhythm and inconsistent hind limb paddling. Muscle weakness due to NMJ dysfunction following from disturbed neurotransmitter release as a consequence of the RN mutation in presynaptic CaV2.1 channels (see below), is another important feature of RN mice (Kaja et al. 2007c). RN mice deliver about 60% less pulling force in grip-strength measuring experiments, as compared to wild-type controls. Muscle fatigability of RN mice can be demonstrated when RN mice are placed on a mesh that can be inverted so that the mice will hang upside-down. Hanging times of RN mice are in the range of 10 s, whereas almost all wild-type mice complete the maximum recording period of 300 s. Assessment of the ability of neonatal mice to sustain hanging with their hind limbs on the edge of a test-tube shows that muscle weakness develops from about postnatal day 8 onwards (Takahashi et al. 2010). Thus, a combination of ataxia and (fatigable) muscle weakness underlies the disturbed gait of RN mice. The heart rate of RN mice is decreased by 20% and less sensitive to injection of the CaV2.1 blocker o-Agatoxin-IVA (Ohba et al. 2009). Also, the data of the authors

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(unpublished) revealed a 35% reduced breathing rate. These observations suggest a dysregulation of the autonomic nerve system due to the RN mutation. Furthermore, RN mice seem to have a higher pain threshold, as homozygous and heterozygous RN mice (3 months old) show reduced sensitivity to painful stimuli (Fukumoto et al. 2009). Heterozygous RN mice display no overt neurological symptoms and thus far it was thought that the RN phenotype was purely recessive and that the presence of one wild-type allele was sufficient to prevent the motor symptoms. However, a recent detailed study of young (2 months old) and aged (22 months old) groups of heterozygous mice reveal that heterozygous mice can exhibit clear neuromuscular deficits at old age (Takahashi et al. 2009b). Interestingly, in the hippocampus and forebrain of young heterozygous mice, wild-type CaV2.1-a1A mRNA expression is higher than that of RN CaV2.1-a1A, while in aged mice this becomes the opposite, although this is not the case in the perirhinal cortex (Takahashi et al. 2009a; Takahashi and Niimi 2009). The shifting balance of the expression of wild-type and RN alleles in some brain areas in heterozygotes may underlie the appearance of motor dysfunction at older age. It may also underlie the deficits in some aspects of memory formation that occur in aged heterozygous mice (Takahashi et al. 2009a). In general, the phenotype in RN mice can be rated as being of intermediate severity, compared to that of other natural CaV2.1 mouse mutants, with the ataxia being more severe than in tottering, but less severe than in leaner mice and no signs of epilepsy, which is clearly present in the latter mutants (Noebels 1984). Furthermore RN mice do not exhibit the paroxysmal dyskinesia seen in tottering mice (Green and Sidman 1962). Interestingly, compound mutant mice with RN and tottering alleles show the abnormal gait of the RN mousse, but not the typical epileptiform seizures of the tottering mouse.

CaV2.1 Channels and the Rolling Nagoya Mutation in Cacna1a Voltage-gated Ca2+ channels are key protein structures in neuronal cell membranes. They transduce electrical signals into cellular influx of Ca2+, which acts as a second messenger in many processes such as regulation of excitability, transmitter release, gene regulation and axonal growth. CaV2.1 channels belong to the group of highvoltage-activated Ca2+ channels that also includes CaV1 (L-type), CaV2.2 (N-type) and CaV2.3 (R-type) channels (Ertel et al. 2000). With their localization in the membranes of both neuronal cell bodies and in particular presynaptic terminals (Westenbroek et al. 1995, 1998) (Fig. 68.2a), CaV2.1 channels are involved in many neuronal Ca2+ signaling pathways, including those leading to gene expression (Sutton et al. 1999) and, above all, they are crucial factors in neurotransmitter release mechanisms in both the central and the peripheral nervous system. CaV2.1 protein and mRNA are abundantly distributed throughout the brain, with a particularly high expression in the cerebellum (Mori et al. 1991; Starr et al. 1991; Westenbroek et al. 1995; Ludwig et al. 1997) (Fig. 68.2b). In the periphery, CaV2.1 channels are present at presynaptic motor nerve terminals at the NMJ (Day et al. 1997; Westenbroek et al. 1998). As are other CaV channels, CaV2.1 are

1546 Fig. 68.2 Histological characterization of wild-type and rolling Nagoya mouse brains. (a) Punctuated immunohistochemical staining on the dendritic area of a cerebellar Purkinje cell of a normal mouse brain, indicating predominant presence of a1A protein at synapses. Scale bar 50 mm. (b) Autoradiographic analysis of the presence of CaV2.1-a1A mRNA in the normal mouse brain. Note the pronounced presence of signal in the cerebellum (arrow). (c) Low level of tyrosine hydroxylase staining in adult wild-type mouse cerebellum. (d) Persistent high level tyrosine hydroxylase expression in adult RN mouse cerebellum (Panel a used with permission of the Society for Neuroscience, from (Westenbroek et al. 1995). Panel b used with permission of the Society for Neuroscience, from Ludwig et al. (1997); permission conveyed through Copyright Clearance Center, Inc. Panels c and d used with permission of the Society for Neuroscience, from Mori et al. (2000); permission conveyed through Copyright Clearance Center, Inc)

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Fig. 68.3 A schematic representation of the mouse CaV2.1 Ca2+ channel with indication of the protein localization of amino acid changes due to Cacna1a mutations. (a) The CaV2.1 channel is a heteromultimeric complex composed of an a1 pore-forming subunit which conducts Ca2+ ions upon opening and accessory a2d, b and g subunits that have a modulatory role. (b) Transmembrane topology of the CaV2.1-a1 protein, with the location of the rolling Nagoya (RN) arginine-to-glycine mutation at position 1262 (R1262G) in the voltage-sensing S4 segment of the third repeating domain. Also indicated are the localizations of the mutations of other natural Cacna1a mouse mutants such as tottering and leaner (Doyle et al. 1997; Fletcher et al. 1996), rocker (Zwingman et al. 2001), tg-4J and tg-5J (Miki et al. 2008) and wobbly (Xie et al. 2007), all characterized by ataxia and epilepsy at various severity levels, and two knock-in mouse models harboring human familial hemiplegic migraine type 1 (FHM1) mutation R192Q or S218L in Cacna1a (Van Den Maagdenberg et al. 2004; Van Den Maagdenberg et al. 2010)

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heteromultimeric complexes composed of a pore-forming a1 subunit and a number of associated auxiliary subunits (Fig. 68.3a), each encoded by a distinct gene (Benarroch 2010). The CaV2.1-a1 subunit is associated with proteins of the a2d, b and g families (although for the latter family this is under debate (Dolphin 2009)). These auxiliary subunits are important for trafficking of the CaV2.1 channel into the neuronal membrane and for modulation of the biophysical and electrophysiological properties of the CaV2.1 channel (Dolphin 2009). The CaV2.1-a1A pore forming subunit is encoded by the gene Cacna1a, which is located on mouse chromosome 8. The RN mutation in this gene is a C-to-G change at nucleotide position 3784, resulting in a charge-neutralizing amino acid change from a highly conserved arginine to glycine at position 1262 (Mori et al. 2000) (Fig. 68.3b). This R1262G mutation localizes in one of the channel’s voltage sensors (in the fourth transmembrane segment of the third repeating domain) and this disturbs the characteristic pattern of positively charged amino acids which is important for the voltage-sensing function of the channel. A large number of mutations in the human CACNA1A has been identified in recent years and shown to underlie several human neurological disorders, including inherited forms of migraine, episodic ataxia and epilepsy (Van Den Maagdenberg et al. 2007; Benarroch 2010). Similarly, as with RN, many other neurological mouse mutants were shown to carry Cacna1a mutations (Felix 2002; Pietrobon 2005) (Fig. 68.3b).

Histological Analyses of Rolling Nagoya Brain Areas RN brain anatomy and morphology as well as the expression and distribution of neurotransmitter receptors have been studied in detail, with special focus on the cerebellum. From early on, there has been a controversy on the existence of cerebellar atrophy and apoptosis. While some of the older studies showed a reduced cerebellar volume and weight and a reduction in the total number of granule, basket and superficial stellate cells, others found no differences between RN cerebella and those from control mice (for summary overview, see Introduction of Tomoda et al. 1992). In later years, the matter has been re-addressed but this has not definitely resolved the issue. In 3–4 weeks old RN mice, normal cerebellar anatomy without signs of apoptosis was observed (Mori et al. 2000). Deep cerebellar nuclei of 4–8 months old RN mice had a normal cell density (Sawada et al. 2001). Again, others have reported contrasting results in that they observed (cerebellar granule cell) apoptosis in 4 months old RN mice (Rhyu et al. 1999) and, especially in the anterior lobe, in 3 weeks old RN mice (Suh et al. 2002). The reasons for these different observations still remain unclear. In deep cerebellar nuclei, increased numbers of CaV2.1-a1 positive neurons have been demonstrated. Such a phenomenon may possibly occur as a compensatory response to reduced CaV2.1 activity due to the RN mutation (see below) (Sawada et al. 2001). Tyrosine hydroxylase (TH) is normally expressed only in catecholaminergic neurons in the adult phase and during a developmental phase in some

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non-catecholaminergic neurons. In cerebellar Purkinje cells of normal mice, TH is expressed transiently from the third to fifth postnatal week after which it decreases to a low level, only to increase again at old age (Fujii et al. 1994; Hess and Wilson 1991). RN mice show persistent ectopic TH expression in their cerebellum (Mori et al. 2000; Sawada et al. 1999, 2001) (Fig. 68.2c, d). This phenomenon appears in a typical parasagittal striping pattern (Sawada and Fukui 2010; Sawada et al. 2010), partly overlapping the staining pattern for zebrin II, a Purkinje cell marker. Interestingly, no enzymatically active form of TH, i.e., phosphorylated at serine residue 40, was identified in the RN cerebellum (Sawada et al. 2004), suggesting that there is no aberrant catecholamine synthesis and release. Ectopic and sustained TH expression is also found in cerebella of the other CaV2.1 mouse mutants tottering (Hess and Wilson 1991) and leaner (Austin et al. 1992), but is also present in the dilute lethal mutant which is also ataxic but has no mutation in Cacna1a (Sawada et al. 1999). This indicates that abnormal TH expression is not uniquely associated with the presence of a CaV2.1 mutation. Because the Ca2+ concentration in Purkinje cells is an important determinant of TH expression (Fureman et al. 1999; Brosenitsch and Katz 2001), it is more likely that disturbed intracellular Ca2+ level is the common factor in the ectopic TH expression in Cacna1a- and dilutelethal mutants. In case of the Cacna1a-mutated mice this may stem either directly from CaV2.1 dysfunction, or may be indirectly mediated by aberrant Ca2+ influx through upregulated CaV1 (L-type) Ca2+ channels (Sawada and Fukui 2001). Besides ectopic TH expression, levels of other neuronal proteins or peptides are changed in the RN cerebellum. The neuropeptide corticotropin-releasing factor (CRF) is present in increased amounts in some climbing fibers as well as in mossy fibers and inferior olive neurons (Sawada et al. 2001, 2003; Ando et al. 2005). The increases in CRF in climbing fibers correlate with TH-positive Purkinje cells (Sawada et al. 2001). CRF is widely expressed throughout the CNS where it has neuromodulator roles, including that of increasing the glutamate sensitivity and reducing the g-aminobutyric acid (GABA) sensitivity in Purkinje cells (Bishop et al. 2000). Furthermore, it can potentiate CaV1 currents (Kanno et al. 1999), which may be in turn of influence on the ectopic TH expression in the RN cerebellum. Expression of ryanodine receptors type 1 and 3 is altered in the RN cerebellum (Sawada et al. 2008). These are Ca2+-induced Ca2+ release channels present on the membrane of the endoplasmic reticulum which allow for Ca2+ release from the endoplasmatic reticulum into the cytosol. The ryanodine receptor alterations may thus affect several Ca2+-dependent processes, including ectopic TH expression. Neurotransmitter receptor autoradiography studies have shown reduced levels of GABAA and adenosine A1 receptors in the cerebellum and of A1 receptors in the cerebral cortex and caudate-putamen of RN mice. Furthermore, benzodiazepine binding sites were found reduced in the cerebral cortex and increased in the CA1 subfield of the hippocampus (Onodera et al. 1988). Morphological studies showed abnormally shaped Purkinje cell dendritic spines and single parallel fibers varicosities making multiple synaptic contacts, not observed in the wild-type (Rhyu et al. 1999). Morphology of cerebellar Purkinje cell dendritic spines of RN mice has been investigated recently in more detail with electron microscopy (Oda et al. 2010).

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Spines at tertiary branches appear to be decreased in density by about 15% and in length by about 25%. Conversely, spines at proximal dendrites are more dense in RN than wild-type. This differential change may parallel the differential effects found in synaptic transmission between synapses of climbing and parallel fibers (see below) (Matsushita et al. 2002). Based on behavioral, histological, and physiological analyses (see below), the motor symptoms in RN mice are generally typified as cerebellar ataxia. However, some components of the movement difficulties are also compatible with extrapyramidal dysfunction. This has led to biochemical and morphological investigations of relevant brain areas in RN. Increased local cerebral glucose utilization (indicating enhanced neuronal activity) in the basal ganglia (including the globus pallidus, entopeduncular nucleus, substantia nigra reticulate and subthalamic nucleus) as well as electrophysiological abnormalities recorded in the globus pallidus have led to the hypothesis that motor disturbances of RN mice may perhaps be also due to striatal dysfunction (Kato et al. 1982; Tomoda et al. 1992). In addition, radiochemical studies have shown increased preproenkephalin and preprotachykinin mRNA in the striatum (Taniwaki et al. 1996). On the basis of these results it may be speculated that the RN mouse is an experimental models for basal ganglia dysfunction, besides being a model for cerebellar ataxia. More research is clearly needed to shed light on how (combined) striatal and cerebellar dysfunction causes motor dysfunction in RN mice. Taken together, there is much histological and biochemical evidence of altered expression levels of a multitude of intracellular and membrane proteins in many structures of the RN brain. These changes may in principle all contribute to motor dysfunction but must be secondary (developmental or compensatory) phenomena resulting from the primary defect in RN, namely a disturbed Ca2+ signaling due to the dysfunction of CaV2.1 channels resulting from the RN missense mutation. The most prominent presence of CaV2.1 channels in the normal cerebellum (Ludwig et al. 1997; Mori et al. 1991; Starr et al. 1991; Westenbroek et al. 1995) (Fig. 68.2b) and the histological changes observed in RN suggest a major role for this area in the motor dysfunction of RN mice.

Effect of the Rolling Nagoya Mutation on CaV2.1 Channel Electrophysiology Expression levels and kinetic parameters of Ca2+ channels can be determined in single cells using voltage-clamp techniques, allowing for measurement of the electrical currents that are caused by Ca2+ flux through the channels (for textbook on these techniques, see (Hille 2001)). The consequences of the RN mutation on CaV2.1 channel function have been investigated with cellular electrophysiological methods in primary Purkinje cell cultures obtained from RN mice as well as in a heterologous expression system, i.e., baby hamster kidney cells that also stably express the auxiliary subunits a2d and b1a (Mori et al. 2000). In these type of experiments, Ba2+ instead of Ca2+ is often used as charge carrier in order to increase the amplitude of the currents and to circumvent Ca2+-induced deleterious processes in

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the cell. A drawback of transfection experiments in non-neuronal cells is the unnatural expression level and the lack of the native neuronal environment of the channel, which may contain important modulatory factors. Furthermore, channel inactivation kinetics in the presence of Ba2+ are known to differ considerably from those in the presence of the in vivo charge carrier Ca2+. The first drawback is not present with the voltage-clamp study of mutant mouse channels in the soma of cerebellar cells, either cultured or in a brain slice. The cerebellum is suitable to study the functional consequences of Cacna1a mutations, since large part of the total Ca2+ current in cerebellar Purkinje and granule cells is mediated via CaV2.1 channels (about 90% and 45%, respectively (Mintz et al. 1992; Randall and Tsien 1995)). The whole cell peak current density in baby hamster kidney cells expressing RN-mutated CaV2.1 channels was reduced by nearly 75%, compared with the wild-type control (Mori et al. 2000). Furthermore, the mutation affected the voltage-dependence of activation of CaV2.1 channels, shifting the midpoint of activation (i.e., the voltage at which half of the channel population is opened) by 10 mV in the positive direction. The slope factor of the activation voltage curve (a measure for the steepness of the relationship between voltage and channel opening) was increased, demonstrating a shallower voltage dependence. The voltage-dependence of inactivation (i.e., the closure of channels in spite of the persistent present of the voltage stimulus) of RN CaV2.1 channels was unaffected in these experiments. The results of these expression studies were largely confirmed in native cerebellar Purkinje cells from RN mice, showing a positive shift (8 mV) of the midpoint of the voltage of activation and an increase in slope factor (Fig. 68.4a) and reduced current density (25%, Fig. 68.4b). However, the inactivation voltage midpoint was shifted by 9 mV in the positive direction. Together, these changes in electrophysiological behavior of RN-mutated CaV2.1 channels shows that the R1262G mutation, which is localized in the voltage sensor, makes the channel less sensitive to voltage changes, resulting in diminished CaV2.1 activity in Purkinje and other cells expressing this channel. This likely forms the initial factor in the cascade that ultimately results in the ataxia of RN mice.

Effects of the Rolling Nagoya Mutation on Cellular Neurophysiological Behavior Aberrant Action Potential Firing Pattern in Cerebellar Purkinje Cells Intracellular voltage measurements at Purkinje cell bodies in RN brain slices show a disturbed firing pattern of action potentials upon stimulation with large depolarizing currents (Mori et al. 2000). The repetitive firing of Na+ action potentials was aborted due to interspike depolarization (Fig. 68.4c, d), reminiscent of the effect of blocking Ca2+-activated K+-channels by Cd2+ in wild-type neurons. Ca2+activated K+-channels are important for post-spike repolarization and are presumably activated by the Ca2+ influx through CaV2.1 channels on the soma and dendritic tree of the Purkinje cell. Apparently, reduced CaV2.1 function in RN Purkinje cell dendritic tree and/or soma leads to less activity of Ca2+-activated

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Fig. 68.4 Electrophysiological characterizations in rolling Nagoya mice. (a) The activation voltage curve of relative Ca2+ current in dissociated RN mouse cerebellar Purkinje cells is shifted by about 10 mV in the positive direction and the steepness of the curve is somewhat less, indicating reduced voltage sensitivity of the RN-mutated CaV2.1 channel. (b) Reduced Ca2+ current density in dissociated cerebellar Purkinje cells from RN mice, measured with voltage-clamp methods. (c) Sustained firing of repetitive action potentials upon long lasting depolarization in wild-type cerebellar Purkinje cells. (d) Aborted repetitive firing in RN cerebellar Purkinje cells. (e) Changes in CaV2.1-mediated ACh release at the RN mouse diaphragm neuromuscular synapse. Upper traces: increased spontaneous ACh release, measured as miniature endplate potential frequency, at RN NMJs. Ten sweeps of 1-s recording are superimposed. Lower traces: smaller endplate

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K+-channels and thus to impaired repolarization of neuronal action potentials. This makes it more difficult for subsequent action potentials to be triggered, impairing high-frequency spiking. Obviously this will influence the signal encodings involved in motor coordination regulation by the cerebellar neuronal network. Reduced current through RN-mutated CaV2.1 channels was further indicated by the observation that Ca2+ spikes were hard to evoke in RN cells and were not followed by burst of Na+ influx-mediated action potentials, normally seen in wild-type neurons (Mori et al. 2000). Together these findings with experimental stimulation strongly suggest that in vivo the RN mutation impairs the neuronal firing behavior of Purkinje cells in response to (integrated) synaptic excitation. This will likely affect cerebellar neuronal network function with resulting ataxia. Similar experimental observations have been made in other CaV2.1-mutants such as the tottering and leaner mouse (Hoebeek et al. 2005; Walter et al. 2006). These findings do not exclude additional contribution of brain areas other than the cerebellum in causing the movement abnormalities of RN mice. In that respect it is interesting that spontaneous firing rate of globus pallidus neurons in the basal ganglia is increased, likely resulting from a diminished inhibitory input (Tomoda et al. 1992).

Dysfunction of Cerebellar Synapses and the Neuromuscular Junction Because of the mutation in CaV2.1-a1 and the preferential and abundant localization of this channel at cerebellar nerve terminals (Westenbroek et al. 1995) (Fig. 68.2a), it is most likely that cerebellar synapses in RN mice have aberrant transmission which contributes to the motor dysfunctions. Cortical cerebellar Purkinje cell dendrites receive extensive (excitatory) synaptic input from nerve terminals of climbing and parallel fibers and there are many other synaptic connections within the cerebellum neuronal network (Watanabe 2008). Neurochemistry studies in cerebellar homogenates showed increased free concentration of neurotransmitters glycine, serotonin and dopamine, decreased glutamate concentration and increased or unchanged noradrenalin concentration (Muramoto et al. 1981, 1982; Nakamura et al. 2005). Furthermore, expression and activity of the catecholamine synthesis pathway enzyme TH was found increased (see above), although this likely concerns an enzymatically non-active form and thus does not result in aberrant catecholamine neurotransmitter synthesis or release (Sawada et al. 2004). While these biochemical studies, together with the histological cerebellar synaptic studies described above, roughly indicate neurotransmitter deficits in the RN ä Fig. 68.4 (continued) potentials in RN NMJs, indicating reduction of evoked ACh release (f) Increased decrement of the compound muscle action potential recorded in RN mouse foot muscle upon 10 Hz supramaximal stimulation of the sciatic nerve. Note the lower initial amplitude in RN muscle. (Panels a–d used with permission of the Society for Neuroscience, from Mori et al. (2000); permission conveyed through Copyright Clearance Center, Inc. Panels e and f modified from (Kaja et al. 2007c), with permission from John Wiley & Sons, Inc)

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cerebellum, they did not provide detailed insight in possible dysfunctions in neurotransmission at specific synapses. Cerebellar synaptic function in RN mice has been characterized with direct and detailed electrophysiological measurements in only one study (Matsushita et al. 2002). Voltage-clamp recordings were made in brain slices of the glutamatergic synaptic currents originating from neurotransmission in parallel fiber as well as climbing fiber synapses on Purkinje cells. There was reduction of excitatory postsynaptic currents at parallel fiber synapses, with increased paired-pulse facilitation, suggestive of a low neurotransmitter release probability. Experiments with Ca2+ channel type-selective toxins showed that presynaptic Ca2+ influx at wildtype parallel fiber synapses is jointly mediated by CaV2.1, -2.2 and presumably 2.3 type channels, with CaV2.1 being the predominant type. CaV2.1 contribution in RN parallel fiber synaptic transmission is somewhat reduced while that of CaV2.2 and 2.3 is somewhat increased. The situation in climbing fiber synapses is completely different: excitatory postsynaptic currents in these synapses are enhanced rather than decreased and they decay at a slower pace, compared to wild-type. Specific CaV2.1 contribution to neurotransmitter release is clearly reduced, while that of CaV2.2 is increased. Further pharmacological and electrophysiological analyses indicated that the increased and broadened excitatory postsynaptic currents are rather due to increased postsynaptic sensitivity of the glutamate receptors than to increased presynaptic release of glutamate. Apparently, disturbed Ca2+ homeostasis in RN cerebellar Purkinje cells leads to a differential regulation of glutamate receptors at parallel and climbing fiber synapses. NMJ dysfunction is to be expected in Cacna1a-mutant mice because of the prominent role of CaV2.1 channels at intramuscular motor nerve terminals where they mediate the release of acetylcholine (ACh) (Westenbroek et al. 1998; Uchitel et al. 1992). In a series of studies, this hypothesis was tested with detailed electrophysiological methods in several mice strains mutant in Cacna1a or genes coding for accessory subunits (Plomp et al. 2000; Kaja et al. 2005, 2007a, b, 2010), including RN (Kaja et al. 2007c). At RN NMJs, a large reduction (50–75%, depending on the muscle type) of the nerve stimulation-evoked ACh release was present (Fig. 68.4e). This reduction in evoked ACh release was accompanied by a threefold increase of spontaneous ACh release, measured as miniature endplate potential frequency (Fig. 68.4e). These effects on synaptic electrophysiology were of intermediate magnitude at heterozygous RN NMJs, demonstrating gene-dosage dependency of these parameters. Most likely, the opposing effects on evoked and spontaneous release result from a complex effect of the mutation on different functional channel parameters, allowing for increased Ca2+ influx at resting potential while limiting Ca2+ influx upon depolarization by a nerve impulse. RN is the only Cacna1a mouse mutant so far in which such opposing effects have been found by us. There seems no compensatory expression of non-CaV2.1 channels at RN motor nerve terminals, in view of the ability of the selective CaV2.1 channel blocker o-agatoxinIVA to reduce evoked ACh release by 95%, equally in wild-type and RN synapses. Severely reduced evoked ACh release at NMJs is the most likely cause of the muscle weakness and fatigue observed in grip strength and inverted grid hanging tests of RN mice. Muscle weakness due to NMJ malfunction was further substantiated by the

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finding of a reduced and decrementing compound muscle action potential with in vivo electromyography (Fig. 68.4f). Furthermore, enhanced d-tubocurarine (a reversible ACh receptor antagonist) sensitivity in ex vivo muscle contraction experiments confirmed a reduced safety factor of neuromuscular synaptic transmission in RN nerve-muscle preparations. Altogether, these NMJ studies strongly indicate that the gait abnormality of RN mice is likely due to a combination of ataxia and muscle weakness. Therefore, the RN mouse may model, besides ataxia, aspects of the NMJ dysfunction in the human neuroimmunological disease LEMS (see below), in which presynaptic CaV2.1 channels at the NMJ are targeted by autoantibodies, causing muscle weakness (Lennon et al. 1995).

The Rolling Nagoya Mouse as a Model for Human CaV2.1 Channelopathies? Ataxia The severe ataxic motor behavior of RN mice is well characterized (Tamaki et al. 1986; Takahashi et al. 2010) and is not paralleled by epilepsy, as present in many other Cacna1a-mutant strains such as tottering, leaner, rocker and null-mutants (Felix 2002; Pietrobon 2002). In view of this, the RN mouse may be a valuable ataxia model, suitable for testing the anti-ataxic properties of (experimental) drugs, especially in the context of human CACNA1A mutation-related cerebellar ataxia (Ophoff et al. 1996; Soong and Paulson 2007; Jen et al. 2007). Such studies are needed in view of the inadequacy of the current drug treatment of ataxia (Ogawa 2004). Only a few compounds (such as acetazolamide in episodic ataxia type 2) have been shown beneficial in human ataxia but none of them has been studied in a controlled or comparative way (Jen et al. 2007), leaving efficacy and mechanism of action relatively unknown. Surprisingly few anti-ataxic drug studies have been performed using the RN mouse mutant as ataxia model. Two studies have shown anti-ataxic effects of thyrotropin-releasing hormone and synthetic analogues (with only minor hormonal activity) in RN mice, possibly due to yet undefined neuroprotective or metabolic effects on RN brain areas (Kinoshita et al. 1995; Nakamura et al. 2005). It would be of interest to test the effect of compounds acting on Ca2+-activated K+-channels, in view of the likely involvement of these channels in the less well sustained action potential firing of cerebellar Purkinje cells of RN (Mori et al. 2000) and firing abnormalities in other ataxic Cacna1a-mutant mice (Hoebeek et al. 2005; Walter et al. 2006). A potential drawback of the use of RN mice in ataxia drug studies is that the ataxic motor symptoms are “contaminated” by (fatigable) muscle weakness due to NMJ malfunction (Kaja et al. 2007c), limiting the intensity at which neuromuscular performance can be tested. Migraine Migraine is a common, neurovascular brain disorder of disabling attacks of headache and associated neurological symptoms (International Headache Society 2004).

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Human CACNA1A gene mutation causes familial hemiplegic migraine type 1 (FHM1) (Ophoff et al. 1996), a rare subtype of migraine with aura with transient hemiparesis during the aura phase (International Headache Society 2004). FHM is autosomal dominantly inherited and considered as a model for the common forms of migraine in view of the similarity of disease symptoms. Moreover, the majority of FHM patients also have normal attacks of migraine without hemiplegia. Interestingly, some 20% of FHM1 patients concomitantly suffer from permanent cerebellar ataxia, which is also the prominent neurological phenotype in RN mice. In view of these similarities, the RN mouse might be considered a model for (familial hemiplegic) migraine. However, electrophysiological studies of FHM1-mutated CaV2.1-transfected cells (Pietrobon 2005; Van Den Maagdenberg et al. 2007) and dissociated cerebellar neurons from Cacna1a knockin mice carrying the FHM1 CACNA1A mutations R192Q or S218L (Van Den Maagdenberg et al. 2004, 2010) (Fig. 68.3b) indicate that the consequences of FHM1 mutations on CaV2.1 channel function are in several important respects opposite to those of the RN mutation. Whereas FHM1 mutations increase neuronal Ca2+ influx through CaV2.1 channels by causing a shift of channel activation voltage in the negative direction and an increase in CaV2.1 current density, the RN mutation causes reduced CaV2.1 current density and a shift of activation voltage in the positive direction (Mori et al. 2000). In view of these differences, the RN mouse seems not a faithful model for (familial hemiplegic) migraine.

Lambert-Eaton Myasthenic Syndrome As described above, the neuromuscular electrophysiological analyses and muscle strength tests of RN mice (Kaja et al. 2007c) showed that there is a certain similarity with the paralytic autoimmune disease LEMS, where auto-antibodies target presynaptic CaV2.1 channels at the NMJ. The electrophysiological hallmark of NMJs of LEMS patients is severely reduced ACh release leading to small endplate potentials, as demonstrated with intracellular electrophysiological measurements in biopsied muscle (Lambert and Elmqvist 1971). A similar electrophysiological phenotype is present in RN mouse NMJs. Likewise, reduced ACh release has been shown at biopsy NMJs of three congenital myasthenic syndrome patients without anti-CaV2.1 antibodies or identified CACNA1A mutation, but with symptoms of ataxia (Maselli et al. 2001) and, furthermore, at biopsy NMJs of two episodic ataxia type 2 patients with CACNA1A truncation mutations (Maselli et al. 2003). Conversely, a proportion of the LEMS patients have accompanying symptoms of cerebellar ataxia (Mason et al. 1997; Lorenzoni et al. 2008; Titulaer et al. 2008). Results of clinical electromyographical studies performed in LEMS patients resemble in some respects the experimental electromyography findings in RN mice, in that there is a low initial compound muscle action potential which decrements during low frequency nerve stimulation (1–10 Hz). Thus, although different causes (i.e., genetic mutation of CaV2.1 vs autoimmunity against this channel), underlie the paralytic symptoms in RN mice and LEMS patients, these similarities indicate that RN mice can serve as a non-immunological model for aspects of LEMS, in particular the function of the NMJ. RN mice could be useful for drug studies aiming to improve treatment of NMJ dysfunction.

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Conclusions and Future Directions In conclusion, the RN R1262G mutation in the Cacna1a gene, encoding CaV2.1-a1 pore-forming subunit, causes changes in the electrophysiological behavior of CaV2.1 channels. One of the most distinct features of the RN-mutated channel, compared to the wild-type, is a reduced voltage sensitivity of the channel. This presumably leads to reduced Ca2+ influx in cell bodies of cerebellar and other neurons that express the channel, causing disturbed Ca2+ signaling leading to aberrant expression of many neuronal proteins and possibly also to the apoptosis of some neurons. As CaV2.1 channels are crucial components in the neurotransmitter release machinery at many synapses, synaptic transmission is likely to be disturbed, especially in brain areas that express high levels of CaV2.1, such as the cerebellum. Together these complex phenomena underlie the ataxia of RN mice. It is yet unclear if and to which extent non-cerebellar regions such as the basal ganglia contribute to the motor symptoms and whether cerebellar atrophy and apoptosis is an important factor. Analysis of in vivo neuromuscular function together with electrophysiological synaptic studies at the NMJ indicate that RN mice suffer from (fatigable) muscle weakness, besides the ataxia. The RN mouse mutant may serve as a model for human ataxia in studies to characterize the efficacy of experimental anti-ataxia drugs. Although the RN mouse may not be a very good model for common forms of migraine and not even for FHM1 (defined by CACNA1A mutations), it may be useful in the experimental study of new antiataxic drugs and drugs that restore disturbed NMJ function. Acknowledgments The studies of A.M.J.M. v.d. M are supported by the Centre for Medical Systems Biology (CMSB) in the framework of the Netherlands Genomics Initiative (NGI).

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Ataxic Syrian Hamster

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Kenji Akita

Abstract

A spontaneous genetic model of cerebellar ataxia in the Syrian hamster (Mesocricetus auratus) is described. Breeding data indicate that the ataxic condition is inherited as an autosomal recessive trait. The homozygous mutant hamsters are smaller in size than the unaffected littermates but have a normal appearance. Both homozygous males and females are fertile, and females are able to nurture litters. They develop a progressive but moderate ataxia beginning at 7 weeks of age; however, they live a normal life span. The major pathologic change in the ataxic mutants is significant cerebellar atrophy, including a rapid and substantial loss of Purkinje cells. Despite the obvious corticocerebellar atrophy, the general structure of the cerebellum is well retained. In addition, most other regions in the brain appear normal by light microscopy. The degeneration of cerebellar Purkinje cells starts after the third postnatal week in mutants and peaks around the fifth week; they then lose almost all Purkinje cells by around 18 months old. They also exhibit a slow and moderate reduction in granule-cell density, probably as a consequence of the primary loss of Purkinje cells. In the homozygous hamster brain, expression of Nna1, the gene responsible for the Purkinje cell degeneration (pcd) phenotype in mice, is suppressed. A phenotypic comparison of the ataxic hamsters with the pcd mutant mice suggests that influence of the causal allele in ataxic hamsters is considerably milder than those of most of the alleles found in the pcd mice. Thus, this ataxic Syrian hamster is a unique animal model of cerebellar ataxia with a severity distinct from any of the pcd phenotypes in mice.

K. Akita (*) Biomedical Institute, Research Center, Hayashibara Biochemical Laboratories, Inc., 675-1 Fujisaki, Naka-ku, Okayama, 702-8006, Japan e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 1563 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_69, # Springer Science+Business Media Dordrecht 2013

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Introduction To date, numerous laboratory animals with neurological mutations have been described. These animal models are characterized by selective neuronal loss and/ or dysfunctions resulting from discrete, genetically induced lesions, and they have served as valuable experimental models for the developmental, differentiational, and degenerative phases of neuronal systems (Heintz and Zoghbi 2000). Hereditary ataxias are characterized by loss of balance and motor coordination because of dysfunction of the cerebellum and its afferent and efferent connections. No effective treatment is available yet for most ataxic syndromes or related neurodegenerative disorders (Evidente et al. 2000). Several mutant alleles for ataxic mice have been identified so far, and some of them are genetically and pathologically well characterized (Lalonde and Strazielle 2007). These strains are considered models for neuronal degeneration or hereditary spinocerebellar ataxia and have been used in neurological investigations and attempts to develop therapeutics. At Hayashibara Biochemical Laboratories (HBL; Okayama, Japan), Syrian hamsters have been bred for more than 30 years for the mass production of human-origin bioactive glycoproteins such as interferon (IFN)-a, IFN-g, and tumor necrosis factor-a. In the production process, newborn hamsters are transplanted with human lymphoblastoid or myelomonocytic cells subcutaneously, and the cells are allowed to grow for weeks in the immunosuppressed hamsters to form cell lumps. The cellular masses are excised and dispersed; then the human cells are induced to produce bioactive glycoproteins. To meet this purpose, selective breeding of fertile and large Syrian hamsters had been conducted for more than two decades, and closed colonies were established. In this chapter, a spontaneous ataxic mutation in the Syrian hamster found and established at HBL is introduced. To date, only a few neurological mutations have been reported in hamsters, including models of paroxysmal dystonia or epilepsy, peripheral neuropathy, and Parkinsonism (Homburger and Peterson 1987). The ataxic mutation in the Syrian hamster is characterized by substantial corticocerebellar atrophy with rapid loss of the cerebellar Purkinje-cell population, occurring after the third postnatal week. Importantly, expression of Nna1 (nervous system nuclear protein induced by axotomy 1), the causal gene of the Purkinje cell degeneration (pcd) mutation reported in mice, is almost completely suppressed in the brain of mutant hamsters (Akita et al. 2007; Akita and Arai 2009). A history of the discovery, general properties, and possible applications of this ataxic mutant are described. The similarities and discrepancies between the ataxic hamster and wellcharacterized pcd mutant mice are discussed as well.

Origin of Discovery and Hereditary Mode In 2001, two male ataxic hamsters were found out of nine littermates in one line within a closed colony for breeding at HBL. They showed typical clinical signs of ataxic gait, including slight trembling of the head. The hindlimb locomotor defects

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1565 #08102

#0812

n=2 n=2

n=6 n=5 Box a

Box b

n=5 n=3

n=3

n=7 n=5

n=8 n=6 n=6 n=9

Fig. 69.1 A family tree of the ataxic Syrian hamsters (extracted). Arrows indicate the first two ataxic males. Male, square; female, circle. Because of space limitations, the number of offspring is shown below the characters in some parts. Affected and unaffected animals are indicated in closed and open characters, respectively. Within unaffected animals, expected carriers are marked by an X in the characters. Certified unaffected females (wild-types) are shown in double circles. Homozygous crossbreeding yielded the ataxic offspring without exception (Box a). In addition, heterozygous crossbreeding of the mutant hamsters yielded the expected Mendelian ratio of 1:3 for affected (recessive homozygotes) versus unaffected (heterozygotes plus wild-types) (Box b)

were marked compared with those of the forelimbs. Although distinguishable ataxia (unsteady walking and stumbling) developed around 2 months of age, no other specific abnormalities were found in their appearance and behavior. The family’s breeding records were retraced in detail, and then hereditary components of the ataxia were investigated by mating sibling pairs for several generations. Figure 69.1 shows a family tree of the mutant pedigree. This clearly explained that the genetic mode of the ataxic phenotype was autosomal recessive. The ataxic phenotype was fixed by the repeated mating of homozygous and/or heterozygous mutants, and the mutant line was established 2 years later.

Breeding and General Properties The ataxic mutant line of the Syrian hamster has not been commercially supplied or deposited to other research institutes so far. This strain has been breeding only at HBL, Inc.

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As a result of selective breeding, HBL’s Syrian hamsters are larger than those supplied from commercial breeders. The average body weight of normal hamsters in the closed colonies is approximately 40 g at the time of weaning (third postnatal week) and 160 g at 10 weeks old. The body size of the ataxic mutants is comparable to that of wild-types at the time of weaning, but they are about 20% smaller at 10 weeks of age (approximately 130 g). They have an otherwise normal appearance, exhibit no other peculiar disease symptoms, and live a normal lifespan (
2 years) under conventional breeding conditions. Both homozygous males and females are fertile. The ataxic hamsters apparently breed well when young but, thereafter, the breeding rate declines. Three-month-old homozygous pairs recorded breeding success rates of 70–80%, comparable to age-matched wild-type controls. In contrast, the rate decreased to 20–30% when the mutant hamsters reached 6 months of age. Thus, a line of mutant hamsters was maintained by crossing pairs of young homozygotes. It is unclear why the breeding performance declines with advancing age compared with the wild-type; however, the progression of discordant movement in the mutant hamsters would be one explanation for unsuccessful matings because no specific failures have been found in the reproductive organs of adult mutants of either sex. For good breeding success rates, the mating period should be extended from 4 days to up to 2 weeks for the mutant pairs. In the case of young pairs of ataxic mutants, the average number of litters is approximately 10, and the male-to-female ratio of the litters is almost 1:1. Although the average number of litters is slightly smaller than that in normal pairs, these breeding data are almost comparable to those of the parental wild-type colonies. Homozygous mothers are able to nurture litters normally, and pups can be weaned in the third postnatal week, comparable to normal hamsters. To reduce environmental stresses, group housing is better during their juvenile period. The major clinical sign of the mutant hamster is a moderate ataxia of gait, including unsteady walking and stumbling as well as a slight trembling of the head. When suspended by its tail, the affected mutant twists its trunk to only one side, whereas a normal hamster extends its trunk and limbs. But postural reflex movements are normal. Symptoms become obvious in the mutants around 7 weeks of age and fixed after another 1 or 2 weeks, and then they progress slowly. With progression, the gait becomes broad-based with frequent loss of balance (Fig. 69.2b), and at 20 months of age, the mutants stumble frequently (Fig. 69.2c). Because of their discordant movements, careful attention should be paid to care of old mutants, particularly in their food and water intake. Other than these ataxia-related symptoms, no other behavioral abnormalities have been confirmed, including circadian coordination or body-temperature regulation. Hibernation is an intriguing physiological phenomenon in the Syrian hamster. In this animal, the domination of endogenous circannual rhythms is not strong, allowing hibernation at any time of the year by exposing them to winterlike conditions (Arai et al. 2005). Thus, it was examined whether the ataxic mutants still retain the integrated physiological functions required for successful hibernation. By keeping the mutant hamsters under a condition of short

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a

b Wild-type

Mutant

c Frequency (/min)

25 20 15 10 5 0

5 7 12 80 Age of the hamsters (weeks old)

Fig. 69.2 Ataxic phenotype of the homozygous mutant hamsters. (a) A photograph of an ataxic homozygous mutant (left) and a wild-type hamster (right) at 6 months of age. There are no distinguishable differences between them except for body size. (b) Paw-print gait analysis of female wild-type (left) and ataxic (right) hamsters at the age of 9 months. Footprints (forepaws, black; hind paws, red) were left on a sheet of paper, while hamsters walked through a narrow tunnel. The wild-type hamster exhibited a regular and even pattern of footprints, whereas the mutant hamster showed irregular, uneven strides. An irregularly loaded and broad-based pattern of hind paws demonstrated abnormalities upon ambulation. (c) Stumbling frequency of ataxic hamsters. The number of stumblings or topplings was counted in a clear breeding cage using the homozygous female mutants at the indicated ages. Unstable movements became obvious at the seventh postnatal week with progressive deterioration in mutant hamsters. In contrast, wild-type hamsters did not exhibit such symptoms. Data are presented as mean  SD (n ¼ 5) (Adapted from Akita and Arai 2009)

photoperiod (light:dark ¼ 2:22 h) and low ambient temperature (4 C), they started to hibernate after 2–3 months of an acclimatization period as wild-type hamsters did. The ataxic mutants repeated torpor (¼ hypothermic state) and arousal cycles just as normal hamsters did for about 2 months. The post hoc analysis of their body temperature by subcutaneously transplanted thermologgers showed that the body-temperature oscillations were not significantly different between normal hamsters and ataxic mutants. This suggests that

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there are no critical deficits in the function of the heat center located in the hypothalamic region as well as the thermoregulatory system, including the sensory nerve system, adipose tissues, muscular system, and endocrine organs. Biochemical and neurochemical characterizations of ataxic mutant hamsters have not yet been carried out so far.

Histology From a macroscopic perspective, the histological abnormality of the ataxic mutants is limited to the cerebellum. There are no morphological malformations; in addition, no grossly distinguished abnormalities are seen in the skeletal system, muscular system, or splanchnic organs. The adult ataxic hamsters exhibited significant atrophy in the cerebellum compared with wild-type hamsters (Fig. 69.3A, a and b). The average wet weight of cerebellum in the adult mutant female hamster was 93 mg (n ¼ 3, 19 weeks old), which is an approximately 50% reduction compared with that in age- and sexmatched wild-type hamsters (180 mg on average, n ¼ 10). The cerebellar volume of the mutants was also significantly smaller. In spite of this, the general structure of the cerebellum in the mutant hamsters was well retained. There was a remarkable reduction in the cerebellar cortical region, that is, a reduction in the thickness of both molecular and granule-cell layers; however, the area of the deep cerebellar nuclei was not changed significantly. With the exception of the cerebellum, the central nervous system of both young and older mutants appeared normal. The average wet weight of the cerebrum was 750 mg (n ¼ 3, 19 week-old females), which is almost comparable to that in the age- and sex-matched wild-type animals (780 mg, n ¼ 10). By the time of weaning (third postnatal week), Purkinje cells have settled and formed the Purkinje-cell layer between the molecular and granular layers in the cerebellar cortical region. At this stage, cerebellar Purkinje cells of the mutant hamsters appeared normal in their location as well as their density; however, the Purkinje-cell number decreased over the next several weeks. A remarkable degeneration of Purkinje cells was observed at 5 weeks of age in the ataxic mutants judged by a degenerative neuron-specific fluorescent dye, FluoroJadeB (Chemicon, USA) (Fig. 69.3B, a), suggesting that the most critical period of Purkinje-cell degeneration in the ataxic hamsters is around 5 weeks of age. By the time ataxic hamsters reached 18 months of age, almost all Purkinje cells disappeared in all lobules of the cerebellum (Fig. 69.3A, d). In contrast to the rapid degeneration of Purkinje cells, a slow and moderate reduction of cerebellar granule cells was also observed in older mutants (Fig. 69.3B, b). At 5 and 7 weeks of age, granule-cell densities in the wild-type controls and mutants were comparable, implying normal formation of the granule-cell stratum in the young mutants. But the granule-cell density of the mutants was reduced to approximately 70% at 18 months of age. These observations suggest that a progressive, although mild, degeneration of cerebellar granule cells occurs in the mutant hamsters probably because of lack of support from

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Fig. 69.3 Degeneration of cerebellar Purkinje cells in ataxic hamsters. (A) Histological findings of the midsagittal cerebella of wild-type (a, c) and ataxic mutant Syrian hamsters (b, d) at the age of 18 months. Upper panels indicate a substantial atrophy in the cerebellum of the ataxic hamster (b) compared with the wild-type control (a). Arrows indicate Purkinje cells in the normal cerebellum (c). Almost all Purkinje cells had disappeared in the cerebellum of the mutants (d). Midsagittal sections (5 mm) were stained with hematoxylin and eosin. G granule-cell layer, M molecular layer. Bars indicate 600 mm for the upper panels and 30 mm for the lower panels, respectively. (B) Time courses of the number of cerebellar Purkinje cells and granule cell density in the cerebellar midsagittal section of wild-type (open circle) and mutant hamsters (closed circle). Two sections from each animal, three animals per group were examined. (a) The number of Purkinje cells was counted in all lobules of the cerebellar midsagittal sections. Degenerating Purkinje cells were also counted with degenerating-neuron-specific FluoroJade B staining. Open and closed bars indicate the number of degenerating Purkinje cells in the wild-type and mutant hamsters, respectively. Data are presented as mean  SD. (b) In contrast to a rapid loss of Purkinje cells, the density of granule cells decreased very slowly in the mutant hamsters. The density of granule cells was calculated by counting the number of granule cells within the middle part of the granule cell layer and is expressed in cells/2,500 mm2. Data are presented as mean  SD (Adapted with permission from Taylor & Francis)

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Purkinje cells. Because only light-microscopic examinations have been performed in the mutant hamsters to date, detailed and minute microscopic observations would shed light on other pathological perspectives in the ataxic mutants.

Genetics Most of the murine mutations affecting the cerebellum are viewed as recessive, including pcd, staggerer, reeler, nervous, scrambler, yotari, rolling mouse Nagoya, tottering, and leaner (Lalonde and Strazielle 2007), with lurcher and weaver as exceptions. Because there is no allelism or linkage test for the hamster, a simple comparative testing of gene expression was done. The expression profile of five previously identified ataxia-related genes (pcd/Nna1, weaver/Girk2, staggerer/ Rora, reeler/Rln, and lurcher/Grid2) was examined in wild-type and mutant hamsters by semiquantitative RT-PCR and northern hybridization analysis. All these genes were expressed abundantly in the cerebellum of wild-type Syrian hamsters; however, only Nna1 was significantly silenced in the cerebellum of mutant hamsters. The suppression of Nna1 gene expression in the homozygous mutant was seen not only in the cerebellum, but also in the whole brain (Fig. 69.4a). On the other hand, the complete suppression of Nna1 was not likely because nested PCR produced a barely detectable level of Nna1 gene transcript from the whole brain cDNA template of the homozygous mutants. These results strongly suggest that silenced Nna1 gene expression is implicated in the primary loss of cerebellar Purkinje cells in mutant hamsters, similar to the previously reported pcd mutant mice. The determination of the allele responsible for the ataxic phenotype of mutant hamsters requires more detailed examination; however, it is probable that the pcd-type mutation is at least involved in the pathogenesis of ataxia in the mutant hamsters. Figure 69.4b shows the Nna1 expression profile in 12 major organs from an adult wild-type male hamster. A 4-kb Nna1 transcript was expressed abundantly in the testis and whole brain, with low levels detected in skeletal muscle, and negligible expression detected in other organs (the heart, liver, stomach, kidney, lung, small intestine, spleen, thymus, and adrenal). In addition to the 4-kb transcript, a testisspecific smaller transcript (
1-kb) was detected; this appeared to be an alternatively spliced form of the Nna1 transcript. The size of the larger Nna1 transcript in the hamster brain is comparable to that in mice (Fernandez-Gonzalez et al. 2002). Because the hamster-specific nucleotide probe corresponding to exons 1–6 of the Nna1 gene sequence in mice was used in the northern analysis, the smaller transcript is expected to contain these regions. The possibility of cross reaction of the probe to other Nna1-like genes (named cytosolic carboxypeptidases; CPPs) seems unlikely because the corresponding region is absent in CPP genes of mice according to a sequence alignment analysis. Therefore, it is conceivable that the alternative splicing occurred downstream of exon 6. The functional significance of the Nna1 splicing variant in the testis remains unclear. Further, there still have been no decisive data concerning the location and manner of the mutation in Nna1 in the ataxic hamsters; neither is it understood how the mutated allele affects the phenotype.

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Fig. 69.4 Nna1 expression in wild-type and mutant hamsters (3.5 months old). (a) The Nna1 transcript(s) in the cerebellum, whole brain, and testis was examined using a hamster-specific DNA probe (580 bp) corresponding to exons 1–6 in the mouse. A 4-kb transcript was absent in the brain of the mutant hamsters, whereas a 4-kb and a 1-kb transcript were detected in the testis of both wild-type (WT) and mutant (MT) hamsters. Lower panels show the loading control (glyceraldehydes-3-phosphate dehydrogenase, gapdh). Note that the exon numbering is based on the current mouse Nna1 structure (Wang and Morgan 2007). (b) Expression of the Nna1 gene in the major tissues of a wild-type male hamster. Ht heart, Br whole brain, Lv liver, Mu skeletal muscle, Sm stomach, Kd kidney, Lg lung, Si small intestine, Sp spleen, Th thymus, Ts testis, and Ad adrenal (Adapted with permission from Taylor & Francis)

Possible Applications A detailed understanding of disease progression and neuronal degeneration in the ataxic Syrian hamster will enable its use as an animal model of cerebellar ataxia similar to ataxic mutant mouse strains. Based on the histological profile, the degeneration of Purkinje cells in the ataxic mutant hamsters starts at the third or fourth postnatal week and peaks around the fifth week (Fig. 69.3B). It precedes the emergence of ataxic symptoms by a few weeks. The ataxic hamsters gradually deteriorate over several weeks, suggesting that it would be possible to monitor the progress of disease using weekly behavioral tests. Although hamsters are often used in circadian, reproductive, and endocrine regulation research, they are infrequently used in neurological behavioral assays. Several behaviors were examined including the rota-rod, inclined plane task, grid walking, open field, and others. It was found that a modified rota-rod test was able to evaluate movement defects even in a very early phase of ataxia with good sensitivity and reproducibility. Figure 69.5 shows a typical result of the modified rota-rod test with the homozygous mutants and ageand sex-matched wild-type control hamsters. Even before the emergence of gait ataxia at 7 weeks of age, the mutant hamsters obviously exhibited inferior motor coordination compared with wild-types in this test. The rota-rod performance significantly and rapidly worsened over time. The time spent on the rotating rod became much shorter, and finally, no mutants could walk on the rotating rod. In this test, a rapid decline of motor performance seems to occur before the cerebellar neuronal loss (Fig. 69.3B), suggesting that the disturbance or dysfunction of

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Fig. 69.5 A modified rota-rod test using Syrian hamsters. Each hamster was placed on a rotating rod (Ф 60 mm) at a constant speed of 6 rpm, and the time spent on the rod was recorded with an upper limit of 180 s. At 3 weeks of age, hamsters received a training session to familiarize with the apparatus. Starting from the next week (4 weeks of age), the session was carried out once a week up to 10 weeks of age. One session consisted of six consecutive trials, and the sixth trial was used for evaluation. Data of homozygous mutants (open circles; dotted line) and age- and sex-matched wild-type control hamsters (closed circles; solid line) are presented as mean  SEM (n ¼ 6 in each group)

synaptic transmission precedes the histologically distinguishable degeneration of Purkinje cells. Therefore, compounds that can improve cerebellar synaptic transmission or support neuronal survival, particularly Purkinje cells, are expected to delay the progression of ataxia in the mutant hamsters. Other than rota-rod, inclined plane task and grid walking could be used to detect the locomotive deficits in the mutant hamsters. Thus, considering therapeutic interpretation, it would be possible to evaluate the efficacy of neuroprotective compounds using these assay systems.

pcd Mutation in Mice Purkinje cell degeneration (pcd) is a mouse mutant that is characterized by postnatal degeneration of selective cell types. More than 30 years have passed since the first description of the original pcd mutant mouse from The Jackson Laboratory (Mullen et al. 1976). During the intervening period, the pcd neurological mutant has been studied extensively as one animal model for hereditary ataxia in humans (Wang and Morgan 2007). In this section, a brief overview of the pcd mutation in mice is described to gain a better understanding of the ataxic hamster.

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According to the latest version (September 27, 2010) of a web database provided by The Jackson Laboratory concerning phenotypic alleles, 12 independent phenotypic alleles have been identified to date: 6 spontaneous alleles (pcd1J, pcd2J, pcd3J, pcd5J, pcd7J, and pcd8J), 5 chemically induced alleles (pcd4J, pcd6J, pcdBtlr, pcdm2Btlr, and pcdbabe), and 1 allele created by random gene disruption (pcdJWG). Although the severity of each allele varies, the cardinal features of the mutants are similar. Homozygotes of the original pcd mice (pcd1J) have a normal appearance but are smaller in size than their unaffected littermates (Mullen et al. 1976). The lifespan of the mutants appears to be normal. These animals exhibit an abrupt and almost complete loss of cerebellar Purkinje cells and a progressive degeneration of granule cells, probably as a consequence of primary degeneration of Purkinje cells (Triarhou et al. 1985). The cerebellum of the mutant mice is essentially normal prior to the second postnatal week, but most Purkinje cells degenerate rapidly over the subsequent 3 weeks. Concomitant with the abrupt loss of Purkinje cells, locomotive discordance becomes obvious during the third to fourth postnatal week. In addition to the distinct degeneration of the cerebellar Purkinje and granule cells, the pcd1J mice also display progressive, partial degeneration profiles in other brain regions. In the thalamus, for example, selected populations of thalamic neurons begin to degenerate at 7–9 weeks of age (O’Gorman and Sidman 1985). Neurons in both the inferior olivary complex and the deep cerebellar nuclei of the pcd1J mice appear to be affected by the Purkinje-cell loss because these neurons normally establish synaptic contacts with the Purkinje cells (Ghetti et al. 1987). Indeed, secondary, transneuronal degeneration profiles, i.e., a reduction in cell number and substantial cell atrophy, are observed in these regions of the old mutants. Of the sensory organs, photoreceptor cells in retina (Mullen et al. 1976; La vail et al. 1982) and mitral neurons in olfactory bulbs (Mullen et al. 1976; Greer and Shepherd 1982) are affected in the pcd1J mice. Degenerative profiles become obvious in the retina after 5 months and in olfactory bulbs after 4 months. In nonneural systems, defective spermatogenesis is one of the major features of pcd1J mice. Male pcd1J mice are sterile due to the reduced number, reduced motility, and structural abnormalities of the sperm (Mullen et al. 1976; Handel and Dawson 1981). Recent studies of pcd have demonstrated that Nna1 is the causal gene responsible for the pcd phenotypes and that Nna1 protein loss-of-function is directly linked to the pcd phenotypes. Nna1 was originally identified as a gene involved in axon regeneration in spinal motor neurons (Harris et al. 2000). It encodes a putative zinccarboxypeptidase that contains nuclear localization signals and an ATP/GTP binding motif. Nna1 and its related genes (Nna1-like genes or CPPs) are reported to constitute the M14D subfamily, a new subfamily of the M14 family of metallocarboxypeptidases; thus, Nna1 protein is thought to act as a peptidase (Rodriguez de la Vega et al. 2007). Nna1 has been mapped to mouse chromosome 13 and is composed of 25 exons (Fernandez-Gonzalez et al. 2002). In adult wild-type mice, a 4-kb Nna1 transcript is expressed mainly in the brain, testis, and heart. Five additional Nna1-like genes (CCP2
6) were also abundantly expressed in testis and brain (Kalinina et al. 2007). The co-expression of Nna1 and Nna1-like genes in

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several regions of the brain in mice implies functional redundancy among these related genes. The basis of Purkinje cell death in pcd has not been resolved; however, recent studies suggest that Nna1 loss-of-function results in elevated autophagy responses, including a mitochondrial autophagy known as “mitophagy” in neurons (Chakrabarti et al. 2009; Berezniuk et al. 2010). In the pcd mutant mice, Nna1 expression in the brain was significantly suppressed. It was undetectable in pcd1J and pcd2J and significantly lower in pcd3J than in the wild-type control (Fernandez-Gonzalez et al. 2002). Although the Nna1 transcript was not reduced in the brain of pcd5J mice, the level of Nna1 protein was dramatically decreased (Chakrabarti et al. 2006). Furthermore, Nna1 protein was undetectable in the brain of pcd1J mice as well as pcd3J mice (Wang and Morgan 2007). On the other hand, Purkinje cell–specific expression of Nna1 transgene successfully rescued the neuronal cell loss as well as the ataxia in the pcd3J mice (Wang et al. 2006). It was also found that the presence of the zincbinding domain of the Nna1 protein was necessary to prevent the degeneration of retinal photoreceptor cells and cerebellar ataxia in pcd5J mice (Chakrabarti et al. 2008). To date, spontaneous genetic mutations within Nna1 have been analyzed in four independent pcd alleles (pcd1J, pcd2J, pcd3J, and pcd5J; Fernandez-Gonzalez et al. 2002; Chakrabarti et al. 2006). Each allele exhibits quite different mutational features. In pcd2J, a
7.8-kb insertion within intron 13 of Nna1 was found to lead to a reduced level of Nna1 mRNA. In contrast, a
12.2-kb deletion between intron 6 and exon 9 was found in pcd3J. This mutation resulted in the formation of aberrant mRNA and was, therefore, expected to produce a truncated protein. The mutation observed in pcd5J was a 3-nucleotide (GAC) insertion in exon 18 that caused an aspartic acid insertion near the zinc-carboxypeptidase domain. Although the mutation site was not within the peptidase domain, this change was considered to destabilize the protein. In pcd1J, no genetic mutation was assigned within the coding region. Nevertheless, the Nna1 transcript was undetectable in the brain and was significantly decreased in the testis of this mutant compared with the wild-type mice. On the basis of these observations, it is now considered that the mutation in pcd1J mice exists in the regulatory region of Nna1.

Comparison to pcd Mutant Mice Both the ataxic hamsters and the pcd mice exhibit ataxia as an autosomal recessive trait. The general appearance and characteristics are also similar, namely, a normal appearance and smaller body weight than littermate controls. A strict longevity study has not yet been conducted in the ataxic hamsters; however, the mutant strain has been recorded to live for more than 2 years. It would appear that mutant hamsters have a normal lifespan under conventional breeding conditions, similar to the pcd mutant mice. The pcd mouse is characterized by a moderate ataxia beginning in the third to fourth postnatal week. An exception is the pcd2J mutant, which is reported to

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develop ataxia later than pcd1J and pcd3J mice; however, the timing of the onset was unspecified (Fernandez-Gonzalez et al. 2002). The mutant hamsters develop ataxia at the seventh postnatal week, which is 3–4 weeks later than pcd mice. Hamsters are born after the shortest gestation of any eutherian mammal (16 days); however, both the timing of weaning and the onset of puberty in inbred strains of mice and the Syrian hamster are very close (Silver 1995; Balk and Slater 1987). Considering this, it seems likely that the brain develops at a similar pace in these two animal species during early adulthood. Therefore, it is assumed that the timing of the onset of ataxia is substantially different between pcd mice and the ataxic hamsters. Time courses of Purkinje-cell loss in these two animal models provide reasonable support for this notion. In pcd mice, the death of Purkinje cells occurs within a relatively brief period of time. Purkinje-cell death begins in pcd1J mice at 18 days of age. In 22- and 24-day-old mutants, 25–50% of the Purkinje cells in the cerebellum had been lost. By 29 days of age, they had lost >90% of the Purkinje cells (Mullen et al. 1976). In the ataxic hamsters, however, there was no significant sign of Purkinje-cell death at 18–20 days old (third postnatal week); approximately 80% of Purkinje cells were alive at 32–34 days old (fifth postnatal week); and more than 30% were still alive at 46–48 days old (seventh postnatal week) in the mutant hamsters (Fig. 69.3B, a). Thus, degeneration of Purkinje cells progresses considerably slower in the ataxic hamsters compared with the pcd1J mice, likely delaying ataxia development in the mutant hamsters. The reason for this is unknown; however, a partial functional rescue from the loss of Nna1 by other genes, for example, Nna1-like genes, is speculated. Both the pcd mutant mice and ataxic hamsters exhibit a significant loss of Purkinje cells in their early adulthood; however, observations indicate a dissimilar profile of secondary or late-onset degeneration in these two animal models. As mentioned before, the reduction of granule-cell density in the ataxic hamster seems moderate compared with that in the pcd1J mice, in which approximately 95% of granule cells degenerated by 20 months of age (Triarhou 1998). Other than reduction of granule-cell density, the pcd mutant mice also exhibit lateonset degeneration of deep cerebellar nuclei, inferior olivary complex, thalamic neurons, retinal photoreceptor cells, and mitral neurons in olfactory bulb. The degenerative profile in these regions of the old ataxic hamsters (>1 year old) was still subtle. For example, the thickness of the outer nuclear layer in retina did not significantly differ between the wild-type and homozygous mutant hamsters (10–12 rows in each group). Because these secondary or late-onset degenerations progress slowly in the pcd mice, a more intensive study using older mutant hamsters may be required. There are still no data concerning the localization of Nna1 mRNA expression within the brain of Syrian hamsters. In normal mice, Nna1 mRNA is prominent not only in Purkinje cells, but also in the mitral cells of the olfactory bulb, thalamic neurons, and retinal photoreceptor cells. In most strains of the pcd mutant mice, specific neurons in these areas are found to degenerate progressively. Although the Nna1 transcripts are detectable in the brain of pcd3J and pcd5J mice, it is considered that the functional Nna1 protein is not produced in pcd3J or not retained in pcd5J.

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This indicates that Nna1 protein loss-of-function is responsible for the pcd phenotypes as described above. On the other hand, the ataxic hamsters do not exhibit distinctive neurodegenerative features, except for those related to the cerebellar Purkinje cells and granule cells, even though Nna1 mRNA was undetectable in the brain of ataxic mutants. This discrepancy is possibly because of the difference in the localization of Nna1 expression and/or the functional redundancy between Nna1 and Nna1-like genes. These possibilities can be explored by examining the localization of mRNA expression of Nna1 and Nna1-like genes in hamsters. Male infertility in the pcd mutants may be an important characteristic that is useful for assessing the severity of the mutated allele because the testicular expression of Nna1 in sterile strains was significantly reduced (Fernandez-Gonzalez et al. 2002). The testicular Nna1 transcript was reduced approximately 20-fold in pcd1J mice and was significantly reduced in abundance and size in pcd3J. In contrast, reduced but stable expression of Nna1 was confirmed in the testis of fertile pcd2J mice. From these results, it was assumed that the testicular expression of Nna1 is related to successful spermatogenesis in the pcd mice. More recently, western blotting has confirmed that no Nna1 protein is detectable in the testis of either pcd1J or pcd3J mice (Wang and Morgan 2007). As shown in Fig. 69.4a, the ataxic hamsters abundantly express Nna1 mRNA in testis. In this respect, it appears plausible that ataxic hamsters with strong Nna1 expression in the testis are fertile. Reportedly, only pcd2J mice exhibit a milder phenotype such as delayed development of ataxia and no degeneration of thalamic neurons similar to the ataxic hamsters. These observations suggest that male infertility is correlated with the severity of the mutated allele, although further comparative studies using other pcd strains are necessary to confirm this.

Conclusions and Future Directions The ataxic Syrian hamster bears many similarities to the pcd mutant mice, and it appears certain that the pcd-type mutation is involved in ataxia pathogenesis in the mutant hamsters because Nna1 expression is almost completely suppressed in the brain. Although more detailed investigations will be required to obtain conclusive proof, the ataxic hamster can be assumed to be an animal model homologous to the pcd mutant mice. The ataxic phenotype of the mutant hamsters develops more slowly and mildly than that reported for most alleles of the pcd mice; therefore, the ataxic hamster might be a preferable alternative experimental model for cerebellar ataxia. Preliminary observations of some behavioral tests using the ataxic mutants also suggested that it was applicable to the evaluation of therapeutics for ataxic symptoms. Given that pcd is one of the well-investigated mutated alleles for hereditary cerebellar ataxia and that pcd mutant mice have yielded significant insights into the treatment of chronic neurodegenerative disorders, the ataxic Syrian hamster could also be applied in related research in the near future.

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References Akita K, Arai S (2009) The ataxic Syrian hamster: an animal model homologous to the pcd mutant mouse? Cerebellum 8:202–210 Akita K, Arai S, Ohta T et al (2007) Suppressed Nna1 gene expression in the brain of ataxic Syrian hamsters. J Neurogenet 21:19–29 Arai S, Hanaya T, Sakurai T et al (2005) A novel phenomenon predicting the entry into a state of hibernation in Syrian hamsters (Mesocricetus auratus). J Vet Med Sci 67:215–217 Balk MW, Slater GM (1987) Care and management. In: van Hoosier GL, McPherson CW (eds) Laboratory hamsters. Academic, Orland Berezniuk I, Sironi J, Callaway MB et al (2010) CCP1/Nna1 functions in protein turnover in mouse brain: implication for cell death in Purkinje cell degeneration mice. FASEB J 24:1813–1823 Chakrabarti L, Neal JT, Miles M et al (2006) The Purkinje cell degeneration 5 J mutation is a single amino acid insertion that destabilizes Nna1 protein. Mamm Genome 17:103–110 Chakrabarti L, Eng J, Martinez RA (2008) The zinc-binding domain of Nna1 is required to prevent retinal photoreceptor loss and cerebellar ataxia in Purkinje cell degeneration (pcd) mice. Vision Res 48:1999–2005 Chakrabarti L, Eng J, Ivanov N et al (2009) Autophagy activation and enhanced mitophagy characterize the Purkinje cell of pcd mice prior to neuronal death. Mol Brain 2:24 Evidente VGH, Gwinn-Hardy KA, Caviness JN et al (2000) Hereditary ataxias. Mayo Clin Proc 75:475–490 Fernandez-Gonzalez A, La Spada AR, Treadaway J et al (2002) Purkinje cell degeneration (pcd) phenotypes caused by mutations in the axotomy-induced gene, Nna1. Science 295: 1904–1906 Ghetti B, Norton J, Triarhou LC (1987) Nerve cell atrophy and loss in the inferior olivary complex of “Purkinje cell degeneration” mutant mice. J Comp Neurol 260:409–422 Greer CA, Shepherd GM (1982) Mitral cell degeneration and sensory function in the neurological mutant mouse Purkinje cell degeneration (PCD). Brain Res 235:156–161 Handel MA, Dawson M (1981) Effects on spermiogenesis in the mouse of a male sterile neurological mutation, Purkinje cell degeneration. Gamete Res 4:185–192 Harris A, Morgan JI, Pecot M et al (2000) Regenerating motor neurons express Nna1, a novel ATP/GTP-binding protein related to zinc carboxypeptidase. Mol Cell Neurosci 16:578–596 Heintz N, Zoghbi HY (2000) Insights from mouse models into the molecular basis of neurodegeneration. Annu Rev Physiol 62:779–802 Homburger F, Peterson J (1987) Experimental biology: genetic models in biomedical research. In: van Hoosier GL, McPherson CW (eds) Laboratory hamsters. Academic, Orland Kalinina E, Biswas R, Berezniuk I (2007) A novel subfamily of mouse cytosolic carboxypeptidases. FASEB J 21:836–850 La Vail MM, Blanks JC, Mullen RJ (1982) Retinal degeneration in the pcd cerebellar mutant mouse. I. Light microscopic and autoradiographic analysis. J Comp Neurol 212:217–230 Lalonde R, Strazielle C (2007) Spontaneous and induced mouse mutations with cerebellar dysfunctions: behavior and neurochemistry. Brain Res 1140:51–74 Mullen RJ, Eicher EM, Sidman RL (1976) Purkinje cell degeneration, a new neurological mutation in the mouse. Proc Nat Acad Sci USA 73:208–212 O’Gorman S, Sidman RL (1985) Degeneration of thalamic neurons in “Purkinje cell degeneration” mutant mice. I. Distribution of neuron loss. J Comp Neurol 234:277–297 Rodriguez de la Vega M, Sevilla RG, Hermoso A (2007) Nna1-like proteins are active metallocarboxypeptidases of a new and diverse M14 subfamily. FASEB J 20:851–865 Silver LM (1995) Reproduction and breeding. In: Silver LM (ed) Mouse genetics: concept and applications. Oxford University Press, New York Triarhou LC (1998) Rate of neuronal fallout in a transsynaptic cerebellar model. Brain Res Bull 47:219–222

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Triarhou LC, Norton J, Alyea CJ et al (1985) A quantitative study of the granule cells in Purkinje cell degeneration (pcd) mutant. Ann Neurol 18:146 Wang T, Morgan JI (2007) The Purkinje cell degeneration (pcd) mouse: an unexpected molecular link between neuronal degeneration and regeneration. Brain Res 1140:26–40 Wang T, Parris J, Li L et al (2006) The carboxypeptidase-like substrate-binding site in Nna1 is essential for the rescue of the Purkinje cell degeneration (pcd) phenotype. Mol Cell Neurosci 33:200–213

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Marco Molinari, Maria Teresa Viscomi, and Maria G. Leggio

Abstract

Hemicerebellectomy (HCB) is characterized by ablation of half of the vermis with one cerebellar hemisphere, including the deep cerebellar nuclei, while sparing the vestibular nuclei and all surrounding structures. This approach has been adopted widely by many groups mainly in rats in various contexts of research. The purpose of this chapter is to review old and recent data focusing on morphological as well as functional data obtained in this model in addressing cerebellar function and brain plasticity mechanisms.

Introduction The Hemicerebellectomy (HCb) Model Since the admirable description by Ramon and Cajal, the well-organized structure of the cerebellar system has fascinated researchers and provided an appropriate

M. Molinari (*) Laboratory of Experimental Neurorehabilitation Unit A – Ataxia Laboratory, I.R.C.C.S. Santa Lucia Foundation, Via Ardeatina 306, 00179 Rome, Italy e-mail: [email protected] M.T. Viscomi Laboratory of Experimental Neurorehabilitation, I.R.C.C.S. Santa Lucia Foundation, Via Ardeatina 306, 00179 Rome, Italy e-mail: [email protected] M.G. Leggio Neurorehabilitation Unit A – Ataxia Laboratory, I.R.C.C.S. Santa Lucia Foundation, Via Ardeatina 306, 00179 Rome, Italy and Department of Psychology, University of Rome La Sapienza, Via dei Marsi 78, 00185 Rome, Italy e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 1579 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_70, # Springer Science+Business Media Dordrecht 2013

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Fig. 70.1 Hemicerebllectomy. Upper part: macrophotograph of a corona1 section through the cerebellum and brain stem of an adult rat that has received a right HCb. Note the absence of the right cerebellar hemisphere, with complete sparing of the left cerebellar nuclei and brain stem structures. Lower part: microphotograph of a Nisslstained section, showing retrograde degeneration in the inferior olive 21 days after right HCb. Note that the left side of the IO shows decreased neuronal density

locus with which the cellular or systemic functional hypothesis of neural function can be tested. To this end, the classical lesion method – damaging a structure to test its function – has been applied many times using small selective lesions, as well large lesions that remove the entire cerebellum. The ablation of half of the cerebellum by hemicerebellectomy (HCb) has been adopted widely by many groups in various contexts of research. The purpose of HCb is to remove half of the vermis with one cerebellar hemisphere, including the deep cerebellar nuclei, while sparing the vestibular nuclei and all surrounding structures (Fig. 70.1). This approach is simple, effects low mortality, and has a high degree of reproducibility. Because of the crossed input–output cerebellar organization, HCb damages the axons of all neurons of the contralateral inferior olive (IO) and pontine nuclei (Pn) and nearly deprives the contralateral cerebral cortex of cerebellar input. HCb has been performed experimentally, primarily in rat and mouse and occasionally in cat (Kolodziejak et al. 2000; Tarnecki 2003) and monkey (Barrionuevo et al. 1978; Bialowas et al. 1984). HCb has been applied primarily in behavioral, neurophysiological, and morphological studies. HCb clearly affects motor behavior, but to a lesser extent than complete cerebellar lesions (Molinari et al. 1990; Petrosini et al. 1990), allowing behavior to be tested on land and in water, which is not possible after complete cerebellar ablation. Further, because HCb is unilateral and based on the complete, crossed organization

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of cerebellar connections, the same specimen, comprising an experimental and control side, can be examined physiologically and morphologically. In human pathology, cerebellar lesions are often unilateral, and experimental HCb findings in animals can be compared with “natural” HCb in humans due to stroke, bleeding, trauma, and surgery (Molinari et al. 2008).

HCb and Experimentally Induced Neuroplasticity Before the plasticity of the central nervous system was demonstrated, HCb-based studies were instrumental in demonstrating functionally relevant, lesion-induced structural changes. The HCb approach highlighted the importance of the rewiring of connections for functional recovery (Tsukahara et al. 1983) and the impact of the developmental time frame during which a lesion develops – the “age at lesion” effect (Castro 1978). Several groups have used HCb to determine the mechanisms of postlesional brain plasticity. With regard to anatomical brain organization, disparate plastic changes occur between unlesioned animals and after adult or neonatal HCb. Further, the substantial reorganization of cerebellar circuits that occurs after lesions form early in development compared with in adulthood allows correlations between changes in connectivity and recovery. The importance of brain plasticity in functional recovery has been instrumental in modifying the general view of a brain that is unable to recover after damage. Castro and colleagues observed abnormal cerebellar projections to the ipsilateral red nucleus and ventral thalamus after neonatal HCb in rats (Castro 1978). Several groups have documented aberrant projections to the red nucleus and thalamus after HCb in the early postnatal period in describing the axonal collateralization of aberrant cerebellothalamic projections to the ipsilateral thalamus (Molinari et al. 1986) and the synaptic organization of the cerebellorubral synapses that sprout (Gramsbergen and Ijkema-Paassen 1982). These aberrant ispilateral projections maintain the topographic specificity of the normal contralateral route, at the least for the cerebellorubral projection (Naus et al. 1984). HCb that is performed early in development induces rewiring in not only spared cerebellar efferents but also in systems that project to the cerebellar stations. In adulthood or perinatally, HCb induces unilateral retrograde degeneration in the major precerebellar stations: the inferior olive, pontine nuclei, vestibular nuclei, and various brain stem nuclei (see below). These retrograde phenomena deprive many pathways of their natural targets. When such a loss occurs in adulthood, the pathway that is deprived degenerates, spreading trans-synaptically. Conversely, when the target is lost during development, the connections are rewired substantially. Neonatal HCb is associated with anomalous increases in crossed sensorimotor cortico-pontine (O’Donoghue et al. 1987) and rubro-olivary projections (Swenson and Castro 1982). This pattern is also observed in ascending pathways. Spinal projections to the Deiters’ nuclei are crossed. After early HCb, the surviving

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Deiters’ nucleus receives increased amounts of ipsilateral spinal fibers (Castro and Smith 1979). Notably, these plastic changes do not involve the systems that originate from the surviving precerebellar nuclei. Specifically, the rubro-spinal, vetibulo-spinal, and reticulo-spinal pathways do not undergo significant changes after neonatal HCb (Petrosini et al. 1988).

HCb as a Model for Studying Axotomy-Induced Neurodegeneration Functional impairments that develop after CNS lesions are associated with axonal damage in common pathologies, such as multiple sclerosis (Trapp and Nave 2008), stroke (Dihne et al. 2002), and spinal and brain trauma (Bramlett and Dietrich 2007). In addition to interrupting the flow of information, axonal damage affects neuronal parent cell bodies in regions that are remote but functionally connected to the primary lesion. These degenerative phenomena are termed “remote damage” and have been studied in the cerebellar system by several groups (Strata et al. 2001; Buffo et al. 2003; Viscomi et al. 2009b). The well-described anatomical efferent-afferent organization of the cerebellar peduncles and the density of the major afferent sources in two well-defined brain stem structures – the IO and Pn – constitute an optimal model that can be used to examine mechanisms of remote damage. Damage to the cerebellar cortex and nuclei or sections of the peduncles affects olivary and pontine axons due to direct axonal lesions and deprives these nuclei of cerebellar inputs. When the lesion affects all Pn and IO axons, extensive neuronal death occurs (Fig. 70.1). Neuronal death persists for approximately 2 months, during which the olivary and pontine neuronal cell populations decline to between 5% and 15% of prelesional values (Buffo et al. 1998; Viscomi et al. 2004). Notably, degeneration phenomena do not develop all the same time. At any time during the 2 months time frame, neurons in various degenerative states are present. Because axonal damage is induced at only one time, disparities in the degeneration state of neurons indicate that various groups of precerebellar neurons respond differently to axonal loss. After axonal injury, the precerebellar neuronal somata undergo alterations, such as chromatolysis, downregulation of basophilic cytoplasmic substances, nuclear eccentricity, nuclear and nucleolar enlargement, cell swelling, and dendrite retraction. Within hours, coincident with early chromatolytic changes, biochemical reactions develop, shifting the cells from a “transmitting” to a “degenerative/ regenerative” state (Florenzano et al. 2002; Buffo et al. 2003). Differences in the type of axonal damage have marginal effects on degeneration. Cell death after peduncolotomy and HCb is similar, peaking in the first week and degenerating at a slower but stable rate afterwards (Viscomi et al. 2009b). This biphasic time course suggests that at least two death mechanisms are active. Based on observations after cortico-spinal tract section (Hains et al. 2003), necrosis-like death mechanisms appear to occur in the first week, after which slower apoptotic mechanisms develop (Viscomi et al. 2004).

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Consistent with the two-mechanism hypothesis, different death-associated proteins are induced at various times. In the inferior olive, c-Fos is expressed 6 and 24 h after development of a lesion, after which c-Jun and Jun-D upregulate for up to 2 months (Buffo et al. 1998). The expression of ionotropic purinergic receptors (P2XR) and neuronal nitric oxide synthase (nNOS) changes between 7 days and 2 months after HCb (Florenzano et al. 2002). Notably, the coexpression of nNOS and P2XR in the precerebellar neurons that survive longer support a model of prosurvival, functional interactions between these molecules (Florenzano et al. 2008).

HCb and Endocannabinoids Evidence from the last decade has implicated the endocannabinoid system (ECS) in CNS physiology and pathology. The ECS is a ubiquitous lipid signaling system with homeostatic functions, comprising at least two receptors – cannabinoid receptor type-1 (CB1R) and type-2 (CB2R) – their endogenous ligands, the endocannabinoids (eCBs); and the proteins that mediate their transport, synthesis, and degradation (Bari et al. 2010). The ECS has been demonstrated to govern neuronal cell fate (Galve-Roperh et al. 2008) and neuroinflammation in various neurodegenerative diseases (Rossi et al. 2010). In particular, CB2 receptors mediate neuroprotective activity specifically through a series of glia-dependent anti-inflammatory actions (Ashton and Glass 2007; Fernandez-Ruiz et al. 2008). CB2R has rarely been observed in neurons and is expressed primarily in microglial cells (Carrier et al. 2004; Gong et al. 2006) and astrocytes (Sheng et al. 2005; Onaivi et al. 2006). Although CB2R expression has been observed in central and peripheral neurons (see (Viscomi et al. 2010) for references), its function in them is unknown (Chin et al. 2008). Recent data using the HCb model have implicated neuronal CB2R in neuroprotection. In the cerebellar system, CB2R is expressed in cortical cerebellar neurons; in the two chief precerebellar stations – the IO and Pn – CB1R is highly expressed while little CB2R is present. HCb changes this pattern dramatically, on which approximately 50% of the precerebellar neurons express CB2R. Under these conditions, in contrast to findings of nearly exclusive glial expression of CB2R (Cabral et al. 2008), HCb-induced CB2R exists only in neurons, not in astrocytic or microglial cells (Fig. 70.2). Experimental modulation of CB2R has demonstrated the function of CB2R newly expressed receptors and the importance of their signals in influencing the death/survival choices of axotomized neurons. Although the modulation of CB1R does not affect survival rate, selective CB2R agonists clearly protect neurons; conversely, CB2R antagonists increase their degeneration. These effects also have a bearing on recovery; CB2R stimulation, but not of CB1R, improves neurological outcomes (Viscomi et al. 2009a). HCb is the only in vivo experimental approach in which CB2R expression can be induced in neurons, allowing one to determine the mechanisms of CB2R

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Fig. 70.2 Confocal images of the pontine nuclei after HCb. Upper row: Double P2X1/nNOS immunofluorescence. Middle row: Triple CB2R/DAPI/ GFAP immunofluorescence. Lower row: Triple CB2R/ DAPI/OX-42 immunofluorescence

intracellular signaling in neuroprotection. Exploiting this technique has helped define CB2R-related intracellular signaling – the effects of CB2R activation on mitochondria and Cyt-c release are mediated by p-Akt phosphorylation (Fig. 70.3) (Viscomi et al. 2009a).

HCb and Neuron-Glia Cross Talk in Neurodegeneration The importance of neuron-glia cross talk in influencing cell death and survival is largely recognized (Owens et al. 2005; Zipp and Aktas 2006), and relationships between glial activation, inflammation, and neuronal death have been established (Block and Hong 2005). In this context, the communication channels of neuron-glia cross talk must be identified. The HCb model has been instrumental in clarifying glial function in remote degeneration. After HCb of precerebellar nuclei, neuronal degeneration occurs concomitantly with astrocytic and microglial activation. Both, neuronal degeneration and glial activation are evident by 7 days, peaking at 3 weeks and decreasing thereafter (Viscomi et al. 2008a). Neuronal damage due to gliosis is often mediated by cytokines, particularly IL-1b, which is secreted by microglia and increases neuronal damage (Loddick and Rothwell 1996) and glial proliferation (Giulian et al. 1988). Notably, in the HCb model, IL-1b is secreted by activated astrocytes but not by microglia (Viscomi et al. 2008a). Glucocorticoids, such as methylprednisolone sodium succinate (MPSS), are used widely to control CNS inflammation, and their effects in controlling flogosis and neurodegeneration have been examined after HCb (Viscomi et al. 2008a). Although MPSS is effective in reducing HCb-induced flogosis, it improves

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CELL DEATH

release Cyt-c

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Fig. 70.3 Cannabinoid-mediated neuroprotection. Schematic of the hypothesized neuroprotective mechanisms that are activated by CB2R induction in neurons that have been axotomized by HCb. CB2-dependent stimulation of the PI3K/Akt pathway inhibits Cyt-c release and subsequent cell death, improving cell survival (Data from Viscomi et al. 2009a)

neuronal survival only at high doses (50 mg/kg). High-dose MPSS markedly reduces degeneration, but its effect is not permanent. When the treatment is suspended, degeneration and flogosis resume. In the HCb model, the secretion of IL-1b by astrocytes constitutes the critical link between glial response and cell death. Conversely, microglial cells, which are sensitive to CB2R modulation, appear to have a secondary function in this model. The inhibition of microglial but not astrocytic activation by minocycline does not reduce cell death rates (Viscomi et al. 2008b). This lack of efficacy conflicts with reports of neuroprotection that is induced by microglial inhibition in several experimental models (Guimaraes et al. 2010; Copeland and Brooks 2010). Particularly, in stroke, minocycline has been shown to be effective in animal models (Cho et al. 2007) as well as in patients (Hess and Fagan 2010). The lack of neuroprotection by minocycline in the HCb model is not unique, microglial inhibition has been shown to be not protective in various models (Diguet et al. 2004; Stefanova et al. 2004; Lee et al. 2010). Whether activated microglia induce beneficial or harmful effects remains to be determined (Block and Hong 2005).

HCb at Various Developmental Stages and Motor Recovery Cerebellar development is not complete at birth, and important steps of cerebellar development occur postnatally (Altman and Winfree 1977). The effects of HCb in neonatal rats, compared with the same lesion in adult animals, demonstrate the importance of the “age at lesion” effect with regard to the motor syndrome after development of a cerebellar lesion (Petrosini et al. 1990; Molinari et al. 1990; Petrosini et al. 1992).

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HCb in the early postnatal period (postnatal Day 1) affects motor development, although not all aspects of motor behavior are affected similarly (Petrosini et al. 1990). The emergence of the quadruped stance, placing reactions, and the ability to swim develop normally despite the cerebellar lesion. The development of other motor competence skills is delayed, but they recover nearly completely. This group is represented by the development of righting reflexes, cliff avoidance, geotaxic reaction, pivoting, and crawling. Conversely, complex functions, such as crossing a narrow path or remaining suspended on a wire, are permanently impaired after HCb (Petrosini et al. 1990). Thus, after early HCb, different motor competences are affected differently. The most peculiar effect of early HCb is characterized by a normal development followed by the development of an impairment at a later stage. This phenomenon is called “growing into deficits” (Leonard and Goldberger 1987) and is evidenced by the progressive reduction in grasping ability and development of a directional bias in the vestibular drop response. Overall, the most significant event in the “growing into deficit” phenomenon is the shift in postural asymmetry after HCb from the side of the lesion to the contralateral side in the third postnatal week (Petrosini et al. 1990). These differences in the occurrence of motor symptoms after early postnatal HCb are consistent with the progressive involvement of archi- and neocerebellar structures in motor control as the postnatal central nervous system develops. In adulthood, HCbed rats develop disparate motor patterns according to the age at which the cerebellar lesion was induced, as evidenced between rats that were HCbed early after birth, at weaning, or in adulthood (Molinari et al. 1990). When HCb is performed at birth, adult rats present with slight extensor hypotonia, contralateral to the side of the lesion, and have efficient locomotion. If the same lesion is created in adulthood, the clinical manifestations are more extensive; the rats show severe extensor hypotonia, ipsilateral to the side of the lesion, that is associated with a wide base and ataxia, impairing locomotion (Fig. 70.4). HCb at weaning has less severe effects on motor features in adulthood than after HCb in adult rats. The differences in clinical presentation that develop after the same lesion, HCb, is induced at various developmental stages are related to disparities in the intensity and characteristics of neuroplasticity throughout development (Petrosini et al. 1992; Gramsbergen 1993). Thus, the HCb model can be used to determine the biological substrate of the “age at lesion” phenomenon that is observed in many clinical conditions. Motor abilities differ depending on whether HCb has been performed at birth or in adulthood. Adult rats with neonatal lesions perform better on several motor tests. Early lesions are associated with more efficient locomotion, prompting a debate over whether these differences are due to the severity of the damage or whether they represent the outcomes of different motor strategies. Neonatal cerebellar lesions are associated with extensive remodeling of cerebellar connections (Castro 1978; Gramsbergen and Ijkema-Paassen 1982; Castro and Mihailoff 1983; Molinari et al. 1986; O’Donoghue et al. 1987; Gramsbergen

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Fig. 70.4 The “time at lesion” effect of HCb. Effects of right HCb at different developmental stages on posture at adulthood (Data from Molinari et al. 1990)

and Ijkema-Paassen 1991). Thus, the organization of the motor system in adult rats that have received HCb at birth differs from the physiological one; such differences might be reflected by the motor strategies that are used in a given task. To test this hypothesis, Molinari and Petrosini (Molinari and Petrosini 1993) analyzed the kinematics of recovered spontaneous locomotion after neonatal or adult HCb. Rats that received HCb in adulthood had impaired locomotion; wide base, irregular asymmetric stepping, tilt to the side of the lesion, and limb hyperflexion were the hallmarks of their gait. Adult rats that received HCb at birth had efficient locomotion, characterized by spinal flexion with the head held low, high stepping during swing, and symmetric regular and hypometric stepping. As shown in Table 70.1, the functional differences in gait after HCb at various developmental stages are attributed to the use of disparate compensatory motor strategies and are not due to differences in the extent of similar symptoms.

HCb and Maze Learning The cerebellum has been demonstrated to mediate high-order cognitive functions (see Molinari and Leggio, this volume), as shown in studies on lesions and function in humans. The most widely adopted animal model has entailed the analysis of spatial functions after cerebellar lesions. Spatial ability requires the close integration of sensory and motor information and is tested in rodents using several maze

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Table 70.1 Comparison of gait in adult rats hemicerebellectomized at birth or in adulthood Gait feature Neonatal HCb Adult HCb Posture Dorsally arcaute spinal cord Tilted to the lesion side Swing phase Bilateral high-stepping Bilateral hyperflexion Stance phase Symmetrical Ipsilesional extensor hypotonia Interlimb coordination Preserved Altered Step symmetry Preserved Lost Step sequence Preserved Altered Step length Hypometric Irregular

tasks, such as the Morris water maze (MWM), a commonly used test in cerebellar research (Morris 1984). In this test, rats are trained to search and find an escape platform into a tank that is filled with opaque water. The test can be solved using different strategies and behavioral analyses, revealing the cognitive functions that are adopted and their function (D’Hooge and De Deyn 2001). HCbed rats have been subjected to the MWM by many groups (Petrosini et al. 1998; Colombel et al. 2004). In the MWM HCbed rats are severely impairment in coping with the task. When placed in the tank, lesioned rats swim aimlessly at the periphery of the pool and make no attempt to explore it. Eventually and abruptly, they leave the periphery, crossing the center of the pool. In such a traversing, a rat might find the escape. Repetition of the test, although it might increase the rate of escape, does not shorten the latency of escape. This behavior differs from that of control rats. Once placed in the pool unlesioned animals reach the platform using rather tortuous routes, but as the sessions progress, they change strategies quickly, using progressively more efficient search routes to avoid sectors that have been explored; soon, they are able to swim directly to the correct location, with dramatic reductions in escape latencies. In contrast to unlesioned rats, HCbed animals fail to develop progressively more efficient strategies that effect a faster solution in the MWM and remain fixed to the initial strategy. Notably, this pattern is true only if the animals are naı¨ve to the task. If rats are trained in the MWM before the cerebellar lesion is induced, their performance is unaffected. All MWM scores are similar before and after development of the cerebellar lesion, even if the platform location has changed. These findings suggest that cerebellar damage does not affect the processing of spatially relevant data directly. Instead, it blocks the acquisition of the procedural competence that is needed to master the task (Petrosini et al. 1998). The function of the cerebellum in maze learning has been demonstrated in observational learning. Observational learning in animals has been the focus of controversy among scholars and it has been object of a recent review (Burke et al. 2010). Observational learning refers to the capacity to acquire a competence solely by viewing it, without direct involvement in its performance. As discussed, HCbed rats are unable to solve the MWM, but if healthy rats are allowed to watch conspecifics perform the MWM – for instance, by observing them from a grid that is suspended over the pool – subsequent lesioning of their

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cerebellum does not impair their ability to solve the MWM (Petrosini et al. 2003). Conversely, if the cerebellum is lesioned before rats observe the MWM, observational training does not improve their performance on the MWM. Thus, the link between HCb and observational learning paradigms implicates the cerebellum in solving the MWM; specifically, cerebellar processing is not critical for mastering spatial information but is essential for acquiring procedures by performing or observing them. Further, the HCb/MWM paradigm allows one to test the model of cerebellar sequence processing within the domain of spatial procedural learning experimentally (see Molinari and Leggio, this volume).

HCb and Changes in Motor Cortical Physiology It has been well known since Dow and Moruzzi’s work, The Physiology and Pathology of the Cerebellum, that cerebellar output affects motor cortex excitability (Dow and Moruzzi 1958), as confirmed using noninvasive neurophysiological techniques in humans (Di Lazzaro et al. 1994). Physiologically, stimulation of the intact cerebellum reduces the excitability of the contralateral motor cortex. The deep cerebellar nuclei have facilitatory effects, whereas the cerebellar cortex, by inhibiting the activity of the nuclei, blocks cerebellar facilitation of the cerebral cortex. Subjects with cerebellar pathologies have impaired excitability and enhanced inhibition in the motor cortex (Di Lazzaro et al. 1995; Liepert et al. 1998; Tamburin et al. 2004). Notably, clinical studies have implicated the cerebellum in processing nonmotor cortices, such as the somatosensory cortex (Restuccia et al. 2001, 2007). In a series of elegant studies based on HCb in rats, Manto and colleagues examined cerebellar processing in somatosensory-driven modulation of motor cortex excitability (Luft et al. 2005; Oulad Ben Taib et al. 2005; Oulad Ben Taib and Manto 2008; Helmich et al. 2010). Sustained somatosensory stimulation increases motor cortex excitability, mediated by interconnections between motor and somatosensory cortices (Luft et al. 2002). Manto and colleagues (Oulad Ben Taib et al. 2005) assessed the function of the cerebellum in somatosensory modulation of the motor cortex by selective unilateral lesioning of the deep cerebellar nuclei in rodents. Ipsilateral cerebellar intranuclear administration of alcohol, TTX, and deep cerebellar stimulation impaired the enhancements in contralateral motor responses, which is normally observed during repetitive somatosensory stimulation. These findings implicated cerebellar processing of incoming sensory information in the control of cerebral cortical excitability. Cerebellar control might act through the cerebello-thalamo-cortical loop (Molinari et al. 2005) or through influences on subcortical strucutures. The cerebello-rubro-spinal loop is a significant subcortical route that sustains communication between the sensory and motor systems (Tarnecki 2003). Specifically, the magnocellular red nucleus responds to sensory and motor stimuli and constitutes an important route of motor control. The activity of the magnocellular red nucleus is governed by the cerebellum. Whereas lesions of the cerebellar cortex increase the

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spontaneous discharge rate of rubral neurons, HCb that involves the DCN reduces spontaneous activity but does not affect the rubral response latency to limb somatosensory stimulation significantly (Tarnecki et al. 2001). Thus, by lesioning the DCN and the cerebellar cortex, HCb decreases the excitability of the two motor targets of the cerebellar output system – the controlateral motor cortex and magnocellular red nucleus. Notably, the HCb-induced reduction in cortical excitability in the motor cortex can be antagonized by transcranial direct current stimulation (Oulad Ben Taib and Manto 2009). Further, low-frequency repetitive stimulation of the motor cortex with periods of peripheral repetitive stimulation can recover the capacity of the motor cortex to be modulated by somatosensory stimuli that is lost after HCb (Oulad Ben Taib et al 2005). Thus, findings from the rat HCb model allow cerebellar impairments in humans to be examined using modern neurophysiological approaches (Torriero et al 2011). Conclusions

HCb is an animal model that has provided important insights into cerebellar function and mechanisms of brain plasticity. Despite its long history, dating back to Luciani’s work in 1891, (Manni and Petrosini 1997), HCb remains a widely used model and continues to generate novel findings in neurobiology (Viscomi et al. 2010) on cerebellar function (Mandolesi et al. 2010) and corticocerebellar interplay (Oulad Ben Taib and Manto 2009).

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Gramsbergen A, Ijkema-Paassen J (1982) CNS plasticity after hemicerebellectomy in the young rat. Quantitative relations between aberrant and normal cerebello-rubral projections. Neurosci Lett 33:129–134 Gramsbergen A, Ijkema-Paassen J (1991) Increased cell number in remaining cerebellar nuclei after cerebellar hemispherectomy in neonatal rats. Neurosci Lett 124:97–100 Guimaraes JS, Freire MAM, Lima RR, Picanco-Diniz CW, Pereira A, Gomes-Leal W (2010) Minocycline treatment reduces white matter damage after excitotoxic striatal injury. Brain Res 1329:182–193 Hains BC, Black JA, Waxman SG (2003) Primary cortical motor neurons undergo apoptosis after axotomizing spinal cord injury. J Comp Neurol 462:328–341 Helmich RC, Siebner HR, Giffin N, Bestmann S, Rothwell JC, Bloem BR (2010) The dynamic regulation of cortical excitability is altered in episodic ataxia type 2. Brain 133:3519–3529 Hess DC, Fagan SC (2010) Repurposing an old drug to improve the use and safety of tissue plasminogen activator for acute ischemic stroke: minocycline. Pharmacotherapy 30:55S–61S Kolodziejak A, Dziduszko J, Niechaj A, Tarnecki R (2000) Influence of acute cerebellar lesions on somatosensory evoked potentials (SEPs) in cats. J Physiol Pharmacol 51:41–55 Lee JH, Tigchelaar S, Liu J, Stammers AM, Streijger F, Tetzlaff W, Kwon BK (2010) Lack of neuroprotective effects of simvastatin and minocycline in a model of cervical spinal cord injury. Exp Neurol 225:219–230 Leonard CT, Goldberger ME (1987) Consequences of damage to the sensorimotor cortex in neonatal and adult cats. I. Sparing and recovery of function. Brain Res 429:1–14 Liepert J, Wessel K, Schwenkreis P, Trillenberg P, Otto V, Vorgerd M, Malin JP, Tegenthoff M (1998) Reduced intracortical facilitation in patients with cerebellar degeneration. Acta Neurol Scand 98:318–323 Loddick SA, Rothwell NJ (1996) Neuroprotective effects of human recombinant interleukin-1 receptor antagonist in focal cerebral ischaemia in the rat. J Cereb Blood Flow Metab 16: 932–940 Luft AR, Kaelin-Lang A, Hauser TK, Buitrago MM, Thakor NV, Hanley DF, Cohen LG (2002) Modulation of rodent cortical motor excitability by somatosensory input. Exp Brain Res 142:562–569 Luft AR, Manto MU, Ben Taib NO (2005) Modulation of motor cortex excitability by sustained peripheral stimulation: the interaction between the motor cortex and the cerebellum. Cerebellum 4:90–96 Mandolesi L, Foti F, Cutuli D, Laricchiuta D, Gelfo F, De Bartolo P, Petrosini L (2010) Features of sequential learning in hemicerebellectomized rats. J Neurosci Res 88:478–486 Manni E, Petrosini L (1997) Luciani’s work on the cerebellum a century later. Trends Neurosci 20:112–116 Molinari M, Petrosini L (1993) Hemicerebellectomy and motor behaviour in rats. III. Kinematics of recovered spontaneous locomotion after lesions at different developmental stages. Behav Brain Res 54:43–55 Molinari M, Bentivoglio M, Granato A, Minciacchi D (1986) Increased collateralization of the cerebellothalamic pathways following neonatal hemicerebellectomy. Brain Res 372:1–10 Molinari M, Petrosini L, Gremoli T (1990) Hemicerebellectomy and motor behaviour in rats. II. Effects of cerebellar lesion performed at different developmental stages. Exp Brain Res 82:483–492 Molinari M, Restuccia D, Leggio MG (2005) Cerebellar information flow in the thalamus: implications for cortical functions. Thalamus Relate Syst 3:141–146 Molinari M, Chiricozzi F, Clausi S, Tedesco A, De Lisa M, Leggio M (2008) Cerebellum and detection of sequences, from perception to cognition. Cerebellum 7:611–615 Morris R (1984) Developments of a water-maze procedure for studying spatial learning in the rat. J Neurosci Meth 11:47–60 Naus CG, Flumerfelt BA, Hrycyshyn AW (1984) Topographic specificity of aberrant cerebellorubral projections following neonatal hemicerebellectomy in the rat. Brain Res 309:1–15

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O’Donoghue DL, Kartje-Tillotson G, Castro AJ (1987) Forelimb motor cortical projections in normal rats and after neonatal hemicerebellectomy: an anatomical study based upon the axonal transport of WGA/HRP. J Comp Neurol 256:274–283 Onaivi ES, Ishiguro H, Gong JP, Patel S, Perchuk A, Meozzi PA, Myers L, Mora Z, Tagliaferro P, Gardner E, Brusco A, Akinshola BE, Liu QR, Hope B, Iwasaki S, Arinami T, Teasenfitz L, Uhl GR (2006) Discovery of the presence and functional expression of cannabinoid CB2 receptors in brain. Ann NY Acad Sci 1074:514–536 Oulad Ben Taib N, Manto M (2008) Reinstating the ability of the motor cortex to modulate cutaneomuscular reflexes in hemicerebellectomized rats. Brain Res 1204:59–68 Oulad Ben Taib N, Manto M (2009) Trains of transcranial direct current stimulation antagonize motor cortex hypoexcitability induced by acute hemicerebellectomy. J Neurosurg 111: 796–806 Oulad Ben Taib N, Manto M, Laute MA, Brotchi J (2005) The cerebellum modulates rodent cortical motor output after repetitive somatosensory stimulation. Neurosurgery 56:811–820 Owens T, Babcock AA, Millward JM, Toft-Hansen H (2005) Cytokine and chemokine interregulation in the inflamed or injured CNS. Brain Res Brain Res Rev 48:178–184 Petrosini L, Molinari M, Gremoli T, Granato A (1988) Neonatal versus adult hemicerebellectomy: a behavioral and anatomical analysis. In: Flohr E (ed) Postlesional neural plasticity. Springer, Berlin, pp 213–220 Petrosini L, Molinari M, Gremoli T (1990) Hemicerebellectomy and motor behaviour in rats. I. Development of motor function after neonatal lesion. Exp Brain Res 82:472–482 Petrosini L, Molinari M, Gremoli T, Granato A (1992) Neonatal versus adult hemicerebellectomy: a behavioral and anatomical analysis. Post Neural Plast 213–220 Petrosini L, Leggio MG, Molinari M (1998) The cerebellum in the spatial problem solving: a co-star or a guest star? Prog Neurobiol 56:191–210 Petrosini L, Graziano A, Mandolesi L, Neri P, Molinari M, Leggio MG (2003) Watch how to do it! New advances in learning by observation. Brain Res Brain Res Rev 42:252–264 Restuccia D, Valeriani M, Barba C, Le Pera D, Capecci M, Filippini V, Molinari M (2001) Functional changes of the primary somatosensory cortex in patients with unilateral cerebellar lesions. Brain 124:757–768 Restuccia D, Della MG, Valeriani M, Leggio MG, Molinari M (2007) Cerebellar damage impairs detection of somatosensory input changes. A somatosensory mismatch-negativity study. Brain 130:276–287 Rossi S, Bernardi G, Centonze D (2010) The endocannabinoid system in the inflammatory and neurodegenerative processes of multiple sclerosis and of amyotrophic lateral sclerosis. Exp Neurol 224:92–102 Sheng WS, Hu S, Min X, Cabral GA, Lokensgard JR, Peterson PK (2005) Synthetic cannabinoid WIN55, 212-2 inhibits generation of inflammatory mediators by IL-1beta-stimulated human astrocytes. Glia 49:211–219 Stefanova N, Mitschnigg M, Ghorayeb I, Diguet E, Geser F, Tison F, Poewe W, Wenning GK (2004) Failure of neuronal protection by inhibition of glial activation in a rat model of striatonigral degeneration. J Neurosci Res 78:87–91 Strata P, Buffo A, Rossi F (2001) Regenerative events in the olivocerebellar pathway. Restor Neurol Neurosci 19:95–106 Swenson RS, Castro AJ (1982) Plasticity of meso-diencephalic projections to the inferior olive following neonatal hemicerebellectomy in rats. Brain Res 244:169–172 Tamburin S, Fiaschi A, Marani S, Andreoli A, Manganotti P, Zanette G (2004) Enhanced intracortical inhibition in cerebellar patients. J Neurol Sci 217:205–210 Tarnecki R (2003) Responses of the red nucleus neurons to limb stimulation after cerebellar lesions. Cerebellum 2:96–100 Tarnecki R, Lupa K, Niechaj A (2001) Responses of the red nucleus neurons to stimulation of the paw pads of forelimbs before and after cerebellar lesions. J Physiol Pharmacol 52:423–436

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Torriero S, Oliveri M, Koch G, Lo GE, Salerno S, Ferlazzo F, Caltagirone C, Petrosini L (2011) Changes in cerebello-motor connectivity during procedural learning by actual execution and observation. J Cogn Neurosci 23:338–348 Trapp BD, Nave KA (2008) Multiple sclerosis: an immune or neurodegenerative disorder? Annu Rev Neurosci 31:247–269 Tsukahara N, Fujito Y, Kubota M (1983) Specificity of the newly-formed corticorubral synapses in the kitten red nucleus. Exp Brain Res 51:45–56 Viscomi MT, Florenzano F, Conversi D, Bernardi G, Molinari M (2004) Axotomy dependent purinergic and nitrergic co-expression. Neuroscience 123:393–404 Viscomi MT, Florenzano F, Latini L, Amantea D, Bernardi G, Molinari M (2008a) Methylprednisolone treatment delays remote cell death after focal brain lesion. Neuroscience 154: 1267–1282 Viscomi MT, Latini L, Florenzano F, Bernardi G, Molinari M (2008b) Minocycline attenuates microglial activation but fails to mitigate degeneration in inferior olive and pontine nuclei after focal cerebellar lesion. Cerebellum 7:401–405 Viscomi MT, Oddi S, Latini L, Pasquariello N, Florenzano F, Bernardi G, Molinari M, Maccarrone M (2009a) Selective CB2 receptor agonism protects central neurons from remote axotomy-induced apoptosis through the PI3K/Akt pathway. J Neurosci 29:4564–4570 Viscomi MT, Florenzano F, Latini L, Molinari M (2009b) Remote cell death in the cerebellar system. Cerebellum 8(3):184–191 Viscomi MT, Oddi S, Latini L, Bisicchia E, Maccarrone M, Molinari M (2010) The endocannabinoid system: a new entry in remote cell death mechanisms. Exp Neurol 224:56–65 Zipp F, Aktas O (2006) The brain as a target of inflammation: common pathways link inflammatory and neurodegenerative diseases. Trends Neurosci 29:518–527

Section 6 Symptoms of Cerebellar Disorders in Human

Cerebellar Motor Disorders

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Giuliana Grimaldi

Abstract

Many of the clinical manifestations of cerebellar damage were described at the turn of the nineteenth and twentieth centuries by the pioneers of the cerebellar physiopathology, including Luciani, Babinski, and Holmes. Cerebellar diseases result in lack of coordination and disturbances of accuracy of movements, causing a constellation of symptoms and motor signs which can be grouped into four categories: oculomotor disturbances, speech deficits, deficits of limb movements, and abnormalities of gait and posture. Instability of gaze and nystagmus, hypermetria/hypometria of saccades, saccadic pursuit, skew deviation (ocular misalignment), and disorders of vestibulo-ocular reflex/optokinetic responses are the main oculomotor alterations observed in cerebellar diseases. Gaze-evoked nystagmus is the most common form of nystagmus encountered in disorders of the cerebellum. Ataxic dysarthria has a typical scanning quality; it is often explosive, with a staccato rhythm and a nasal character. Ataxic speech tends to become slow with slurring, and words may be unintelligible. Cerebellar damage typically results in impairment of performance of limb movements. Ataxia of limbs includes: dysmetria, decomposition of movement, dysdiadochokinesia, cerebellar tremor, isometrataxia, disorders of muscle tone (both hypotonia and cerebellar fits), loss of check and rebound, abnormal handwriting, and megalographia. Tremor in cerebellar diseases is mainly composed of low frequency oscillations, usually with a kinetic component. Kinetic tremor is often associated with a concomitant postural tremor. Cerebellar patients have increased body sway and a broad-based stance due to the inability to maintain the body in a stationary position (ataxia of stance). Similarly, gait in cerebellar patients is irregular and broad based.

G. Grimaldi Unite´ d0 Etude du Mouvement (UEM), Neurologie - ULB Erasme, 808 Route de Lennik, Bruxelles, 1070, Belgium e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 1597 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_71, # Springer Science+Business Media Dordrecht 2013

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Successive steps are spaced in a staggering way and followed by corrections or falls. Rhythm is distorted and speed is often reduced. Walking trajectory veers erratically with difficulties in initiation, stops, or turns.

Introduction This chapter provides a description of the motor disorders caused by cerebellar dysfunction. Lack of coordination and disturbances of accuracy of movements – the cardinal features of cerebellar diseases – involve ocular movements, speech articulation, and limb and trunk movements. Clinical motor deficits can be grouped into the following four categories: oculomotor disturbances, dysarthria, ataxia of limbs, and ataxia of stance and gait abnormalities. These signs may occur in isolation or in various combinations. Knowledge of the anatomical and functional role of cerebellar structures is fundamental to the understanding of the clinical deficits. The relationship between the topographical localization of a given cerebellar lesion and the resulting symptoms has been the topic of many studies (Table 71.1 and Fig. 71.1). A cerebellar functional topography has been proposed. The statement that motor tasks are mainly localized to the anterior cerebellar lobe has been sustained by the meta-analysis of functional neuroimaging studies (Stoodley and Schmahmann 2009). A recent study in patients with isolated cerebellar infarctions has demonstrated that lateropulsion, vertigo, and nystagmus are more common in patients with a lesion in the caudal vermis, while lesions of the anterior paravermis are associated with dysarthria and limb ataxia (Ye et al. 2010) (see also the section on “Topography of Clinical Deficits” for details). Cerebellar dysfunction results from a large number of etiologies including congenital malformations and dysmorphologies (Ten Donkelaar and Lammens 2009; Dennis et al. 2010); genetically transmitted ataxias (Manto and Marmolino 2009); infections (Amlie-Lefond and Jubelt 2009); immune-mediated (Hadjivassiliou et al. 2008; Ishikawa and Kobayashi 2010) and paraneoplastic (Grant and Graus 2009) disorders; toxic agents (Manto and Jacquy 2002a, b); and vascular (Konczak et al. 2010), neoplastic (Sarrazin 2006), and traumatic conditions (Krauss et al. 1995). Nevertheless, these different causes of cerebellar disease manifest with similar clinical presentations. The cerebellum has also a role beyond motor control. Large unilateral or bilateral cerebellar lesions involving the posterior lobe manifest with cognitive, affective, and behavioral abnormalities (Fig. 71.2). Briefly, the dysmetria of thought hypothesis illustrates the parallelism between motor and cognitive cerebellar control: “in the same way that cerebellum regulates the rate, force, rhythm, and accuracy of movements, so does it regulate the speed, capacity, consistency, and appropriateness of cognitive and emotional processes” (Schmahmann 1991; Schmahmann et al. 2007). Current models suggest that motor and nonmotor networks connecting the cerebellum with cortical areas operate independently in closed and segregated loops. Nevertheless, the existence of a functional cross talk between motor and cognitive cerebellar networks is suggested by clinical evidence showing an influence of cognitive activation on the cerebellar motor control (Salih et al. 2010).

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Table 71.1 Motor symptoms of cerebellar lesions Symptom Description Nystagmus Rhythmic oscillatory movements of one or both eyes. A fast/slow component in opposite directions is observed Dysmetria of saccades Inaccurate saccades with over/ and saccadic pursuit undershooting of the target. Both undershoots and overshoots may be observed. Fixation deficits (instability of fixation, oscillations) Abnormal gain of the VOR (vestibulo-ocular reflex) Limb dysmetria

Decomposition of movement

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Main localization of lesions Flocculus/paraflocculus Uvula and nodulus Dorsal vermis/Fastigial nucleus Flocculus/paraflocculus Uvula and pyramid

Inability to maintain the eyes motionless during a fixation task Compensative rotation during eye/head movements

Flocculus/paraflocculus Uvula and pyramid

An error in trajectory due to a disturbed range, rate, and/or force of the movement. Most often composed of hypermetria during fast movements Decomposition of multi-joint tasks or socalled complex movements into elemental movements

Dentate nucleus Interpositus nucleus Lateral cerebellar cortex

Ataxic stance

Broad-based stance with increased body sway

Ataxic gait

Irregular, broad based, and unsteady gait. Patients tend to fall.

Dysarthria

Explosive nasal speech with a typical scanning aspect

Dentate nucleus and interposed nuclei Intermediate zone Medial and intermediate cerebellum Fastigial and interposed nuclei Flocculo-nodular lobe Posterior inferior cerebellar vermis (abnormal tandem gait) Superior vermis Superior paravermal region Intermediate cerebellar cortex Dentate nucleus

Adapted from Grimaldi and Manto (2011b)

Contributions of Luciani, Babinski, and Holmes Important contributions have been provided since the nineteenth century. In 1891, Luigi Luciani (1840–1919) published the results of his experimental studies on cerebellum. He observed, in primates and dogs, three elemental deficits ipsilateral to cerebellar lesion, formulating his triad of cerebellar symptoms: atonia, asthenia, astasia to which he later added a fourth sign: dysmetria (Manni and Petrosini 1997).

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Fig. 71.1 Medio-lateral subdivision of the cerebellum into a spinocerebellar zone and a cerebrocerebellar zone. The main connection sites of the fastigius nucleus (dark grey), the interpositus nucleus (light grey) and the dentate nucleus (black) are shown on an unfolded cerebellum. Functional roles are summarized into grey boxes (From Grimaldi and Manto (2011b), with permission)

Joseph Babinski (1857–1932) introduced the concept of asynergia to coin the decomposition of movement and gave the first description of the adiadochokinesia (Babinski 1902). In the early twentieth century, Gordon Holmes (1876–1965) described in detail the clinical deficits due to cerebellar lesions in soldiers of the First World War (Holmes 1917) and grouped these signs and symptoms into five categories: hypotonia, static tremor, asthenia, fatigability, astasia which included dysmetria and intention tremor. The clinical description of motor cerebellar disorders cannot be truly provided without citing the works of these pioneers. In particular, Holmes’ description of cerebellar function/dysfunction still exerts major influences over contemporary neurological texts (Haines and Manto 2007).

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Fig. 71.2 Illustration of the anatomical dichotomy of cerebellar structures (unfolded cerebellum) controlling motor/sensory tasks versus cognitive tasks. Foci of activation are illustrated. Abbreviations: H hemispheric, Tons tonsil, Flocc flocculus, D declive, F folium, T tuber, Pyr pyramis, Uv uvula, Nod nodulus (From Grimaldi and Manto (2011b), with permission)

Symptoms The most common complaints of patients presenting lesions restricted to the cerebellum are gait difficulties (lack of balance), headache, dizziness, limb clumsiness (incoordination of the extremities), speech difficulties (inarticulate speech), blurred vision, feebleness, and fatigability. Cerebellar symptoms may also include impaired handwriting and cognitive impairments. Patients affected with acute cerebellar diseases, as stroke and tumor with edema, complaint, in particular, headache with nausea and vomiting (Manto 2010). Headache may be the sole symptom in case of tumor, abscess or cerebellar stroke, and is often reported very early in patients presenting with acute cerebellar diseases. The pain may be restricted to the parietal or occipital regions, or to a region around or behind the eye (mono- or bilaterally). Neck stiffness may be also present in association with occipital pain. Headaches can be influenced by postural changes. Any child complaining of headache in the morning, whether or not it is associated with nausea, vomiting (including the projectile vomiting), clumsiness or gait difficulties, should be investigated rapidly for a possible cerebellar tumor. Intracerebellar hemorrhage occurring in hypertensive patients may present as a severe, abrupt, dull, headache associated with gait ataxia, and vomiting (Manto 2010). Vertigo (which patients may describe with the less specific term, Dizziness) is an illusion of rotatory or linear movement due to a functional or lesional disturbance which may occur at any level of the vestibular system, from peripheral to central

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Table 71.2 Central vs peripheral vestibular syndrome Central Features of vertigo Rotatory Feeling of dizziness, disequilibrium, lightheadedness Sways and oscillations Lateropulsion usually toward the side of the lesion Features of nystagmus Multidirectional Conjugate More severe with gaze toward the side of the lesion Effect of fixation on No effect nystagmus Effect of Dix-Hallpike Immediate and persistent maneuver increase of vertigo

Effect of head and trunk movements Concomitant acoustic symptoms and signs Concomitant neurological symptoms and signs

Peripheral Dizziness

May be present

Lateropulsion usually away from the side of the lesion Rotatory/horizontal Dissociate More severe with gaze away from the side of the lesion More intense when fixation is removed by the Frenzel lenses Increase of nystagmus in the horizontal plane Temporary increase of vertigo after a latency of 1–40 s Increase of the severity of nystagmus Often reported

Often present

Usually absent

May induce a slight change

Adapted from Bonavita and De Simone (2004), Karatas (2008)

connections. Central etiologies may account for 20–40% of causes (Tilikete and Vighetto 2009). Such an erroneous perception of self-motion or object-motion can also be described as an unpleasant distortion of static gravitational orientation (Dieterich 2007). Dizziness and nausea/vomiting were the most common symptoms (after gait difficulties and headache), in a prospective series of 115 cases with cerebellar disorders (Manto 2002). Sudden onset of rotatory vertigo might be the only presenting symptom of a cerebellar infarct. In this case, the clinical features may closely mimic an acute peripheral labyrinthine disorder. However, the absence of nystagmus, nystagmus that changes direction with different eye positions, and normal caloric responses may be suggestive of a cerebellar infarct (Masson et al. 1992). The differential diagnosis of central and peripheral vestibular syndrome can thus be narrowed by the physical examination, including evaluation for nystagmus and the Dix–Hallpike maneuver (Table 71.2). Peripheral vestibular nystagmus is usually rotatory/horizontal, is more intense when fixation is removed by the Frenzel lenses, and is most pronounced on gaze away from the side of the vestibular involvement. In contrast, central nystagmus is more evident with gaze directed toward the site of the cerebellar lesion. The Dix–Hallpike maneuver may elicit nystagmus and vertigo in cases of benign paroxysmal positional vertigo (BPPV) (Manto 2002).

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The complexity of vertigo is also related to the fact that vertigo is associated with numerous pathologies, both at the central and peripheral level. This includes: (a) cerebellar disease, central positional vertigo, vertebrobasilar ischemia and orthostatic hypotension, basilar migraine, paroxysmal ataxia/dysarthria (Dieterich 2007); (b) BPPV, Menie`re’s disease, vestibular neuritis, labyrinthitis; (c) and many medications and psychiatric disorders which can cause dizziness (Post and Dickerson 2010). Clumsiness is another typical symptom in cerebellar patients. Patients often report that they have difficulties performing activities of daily living with their limbs, especially the hands and the fingers. Manipulation of small objects is difficult to perform

Oculomotor Disturbances The cerebellum participates in visual fixation, binocular alignment, saccade accuracy, generation and maintenance of smooth ocular pursuit, and vestibulo-ocular reflex (VOR) modulation (Leigh and Zee 2006). Therefore, instability of gaze and nystagmus, hypermetria/hypometria of saccades, saccadic pursuit, skew deviation (ocular misalignment), and disorders of VOR and optokinetic responses are the main oculomotor alterations observed in cerebellar diseases. The instability of gaze is estimated clinically by holding the index finger in front of the patient at a distance of about 30 cm and in a lateral position (no more than 30 ), in upward and downward position. Nystagmus consists of rhythmic oscillatory movements of one or both eyes, with a fast and a slow component in opposite directions. Gaze-evoked nystagmus (GEN) is the most common form of nystagmus in patients with disorders of the cerebellum (Gilman et al. 1981). GEN is a rhythmic oscillation of the eyes while attempting to maintain eccentric gaze (Cannon and Robinson 1987). It should be noted that initial difficulty maintaining eccentric gaze with a few beats of nystagmus may be physiological, and occur also at small angles of gaze (Whyte et al. 2010). Downbeat nystagmus (DBN) may produce disabling oscillopsia (illusion of visial motion). DBN results from focal lesions affecting primarily the cerebellar flocculus and/or paraflocculus which are engaged in (tonic) inhibition of the superior vestibular nucleus and its excitatory efferent tract, but not the downward vestibular system (Pierrot-Deseilligny and Milea 2005). DBN is often present in Chiari malformation. Periodic alternating nystagmus (PAN) can also be associated with this condition (Korres et al. 2001). Subacute cerebellar ataxia associated with anti-GAD antibodies (anti- glutamic acid decarboxylase) may be characterized by abnormal eye movements such as PAN, DBN, and slow vertical saccades (Tilikete et al. 2005). Spontaneous nystagmus (in primary gaze), GEN, PAN, and saccadic intrusions into pursuit eye movements (as micro-saccadic oscillations and square wave saccadic intrusions) are reported in patients affected with ataxia-telangiectasia. Cerebellar involvement in this autosomal recessive disorder is characterized by degeneration of Purkinje neurons, resulting in disinhibition of the caudal fastigial oculomotor

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a

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Contact with target

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Fig. 71.3 (continued)

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region thus affecting the saccade generating mechanisms, and the vestibular nuclei causing nystagmus (Shaikh et al. 2009). Lesions of the midline zone of the cerebellum (together with lesions of the midbrain and the lower brainstem, and drug overdose) may produce upbeat nystagmus (UBN). UBN is a primary position nystagmus with the fast phase in the upward direction (Manto 2002). Latency, precision, and velocity of saccades are estimated by asking the patient to look laterally at first one and then the other finger located in each temporal visual field (Manto 2010). Indeed, the saccades are rapid eye movements used to voluntarily move gaze from one target of interest to another (conjugate eye movements). Brainstem and cerebellar populations of neurons contribute to the generation of saccades. In particular, the cerebellum plays a role in steering and stopping the saccades, thus determining their accuracy (Ramat et al. 2007). Posterior vermis and the caudal fastigial nucleus, to which it projects, are the structures controlling speed, accuracy, and consistency of saccades. Moreover, the caudal fastigial nucleus is necessary for the recovery of saccadic accuracy (Robinson and Fuchs 2001). Patients affected by Friedreich ataxia commonly demonstrate ocular motor abnormalities including fixation instability and saccadic dysmetria. There is a significant correlation between saccadic latency and disease severity in this autosomal recessive disorder (Hocking et al. 2010). Patients affected by vascular cerebellar lesions show reduced acceleration during the initiation phase of conjugate smooth pursuit, diminished smooth pursuit gain, and saccadic hypometria. They also have impaired slow vergence (simultaneous movements of the eyes in opposite directions), which causes ineffective dysconjugate eye movements and consequently difficulties in maintaining a stable binocular vision. The cerebellar vermis controls slow vergence (Sander et al. 2009). A study conducted on a group of patients suffering from primary cerebellar degenerative diseases undertaking a walkway task, demanding precise foot placement at each step, and a visual fixation task, requiring only eye movements, revealed that cerebellar patients have dysmetric saccades when attempting to fixate a target during a walking task (Fig. 71.3). Such similarities between the oculomotor deficits displayed by patients during the visual fixation task and during walking indicate that the latter are not merely a consequence of ataxic gait (Crowdy et al. 2000). Ocular hypermetria is defined as an inaccurate saccade with overshooting of the target. This is very suggestive of a cerebellar lesion (Selhorst et al. 1976). Saccadic pursuit is a stepwise decomposition of pursuit movements. It is a common but nonspecific finding in cerebellar patients. ä Fig. 71.3 Raw data from two test walks. (a) Complete walk for a typical control subject. (b) Complete walk for a typical cerebellar patient. High levels on the foot contact traces represent times of foot contact with a particular stepping stone (L2, R5 etc.) and the numbers refer to the stone’s place in the sequence along the walkway. Positive values on the horizontal eye movement trace represent eye movements left of center in degrees, negative values right of center. In (b), dashed boxes highlight example dysmetric saccades. The inset in (b) shows left and right saccade onsets identified by the automatic saccade ID program (From Crowdy et al. (2000), with permission)

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Skew deviation (ocular misalignment) produces vertical diplopia – a static ocular alignment in which one eye is higher than the other – due to lesions in the peripheral vestibular apparatus, medulla, cerebellum, thalamus, and vestibular cortex (Manto 2002). A clinical test to differentiate skew deviation from trochlear nerve palsy has been proposed: a decrease 50% of the vertical deviation from the upright to supine position during the so-called upright-supine test is suggestive of skew deviation and warrants investigation for a possible lesion in the posterior fossa as the cause of vertical diplopia (Wong 2010). Unilateral lesions in caudal parts of the cerebellum, such as an infarct in the territory of the posterior inferior cerebellar artery, may result in partial ocular tilt reaction (OTR). OTR is a brainstem otholitic – ocular reflex, characterized by a combination of head tilt, conjugate eye cyclotorsion, skew deviation, and impairment of vertical perception (Manto 2002). The vestibulo-ocular reflex (VOR) maintains steady vision (stable retinal image) during rapid movements of the head and the body, by inducing eyes movement simultaneous and as fast as the head movement but in the opposite direction. Healthy subjects present a cancelation of the VOR during the rotating chair test: the subject is seated on a chair – which rotates at constant velocity – and fixates an object that moves synchronously with head movement (i.e., maintains the arms outstretched and fixates his thumbs). Normal cancelation of the VOR suppresses the motion of the environment and the healthy subjects’ eyes stay fixed on the object. On the opposite, cerebellar diseases lead to an impaired suppression of the VOR (Dichgans et al. 1978) resulting in an inability to suppress the motion of the environment, and consequently the occurrence of saccadic intrusions. Indeed, during the rotating chair test (vestibular suppression test), cerebellar patients reveal intermittent deviations of the eyes with corrective saccades. Moreover, patients with cerebellar disorders (in particular with lesions to the flocculus), present an increased ratio between head and eye velocities (increased gain of the VOR) (Robinson 1976).

Dysarthria and Other Speech Deficits Speech motor control is under the supervision of two separate networks constituting a preparative and an executive loop, respectively. The cerebellum contributes to both these pathways (Riecker et al. 2005). Dysarthria is a speech motor disorder that may occur in several diseases involving not only the cerebellum, but also the pyramidal (Donkervoort and Siddique 2009) and the extrapyramidal systems. Dysarthria may occur in several neurological diseases and, on the basis of its nosological frame, it may present with different features. For instance, speech deficits related to Parkinson’s Disease (extrapyramidal dysarthria) are often called hypokinetic dysarthria and can be characterized by abnormal prosody (monopitch, monoloudness), hypophonia, reduced stress, imprecise consonants, and inappropriate silences (Martnez-Sa´nchez 2010). Cerebellar lesions disrupt the coordination of prosody, articulation, phonation, temporal regulation, and fluency of speech production (Kent et al. 1979). As a consequence,

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clarity, rhythm, and fluency of speech are impaired in cerebellar disorders. This form of dysarthria is often called ataxic dysarthria. In most instances, ataxic speech is explosive, with a staccato rhythm and a nasal character. Words may become unintelligible because of the severity of the temporal dysregulation of muscles activities. Speech tends to become slow with slurring (Kent et al. 1997). The scanning aspect – the most readily recognizable deficit in cerebellar dysarthria – is characterized by hesitations, accentuation of some syllables, omission of appropriate pauses, and addition of inappropriate pauses (Zentay 1937; Gilman et al. 1981). The motor speech deficits can be grouped into three clusters: (1) articulatory inaccuracy – associated with inaccuracy of repetitive movements – that determines imprecise consonants, irregular articulatory breakdowns, and distorted vowels; (2) prosodic excess – related to motor slowing – which is characterized by excess and equal stress, prolonged phonemes, slow speech rate, and prolonged intervals between words and syllables; (3) phonatory/prosodic insufficiency – associated with hypotonia of speech musculature – including harshness, monopitch, and monoloudness (Brown et al. 1970; Darley et al. 1969). Functional MRI (fMRI) studies in healthy volunteers show that the cerebellar representation of the tongue and orofacial muscles corresponds to that of the area involved in patients with cerebellar dysarthria due to focal vascular lesions (Urban et al. 2003). Impairment of the upper cerebellar hemisphere in the territory of the superior cerebellar artery results in dysarthria (Ogawa et al. 2010). Combined parametric data (measurements at the acoustic speech signal, tracking of articulatory movements and fMRI studies) have contributed to redefine the concept of ataxic dysarthria and indicated the cerebellum to support several aspects of speech processing. Among these: acceleration of orofacial gestures, timing (coordination) of complex articulatory sequences, and control of brainstem reflexes monitoring respiratory and laryngeal muscle activity. Besides the superior aspects of the cerebellum to mediate the speech motor functions a role in the correct speech perception has been also identified (Ackermann et al. 1992; Ackermann and Hertrich 2000). Lesions of either cerebellar hemisphere can cause dysarthria. Positron emission tomography (PET) studies on regional cerebral blood flow changes during speech production in a group of subjects with hereditary ataxia (SCA1, SCA5) have revealed that the greatest decline in blood flow occurs in the cerebellar regions, whereas blood flow increases in the classic cortical speech area (Broca’s area). Such increase in cortical flow has been considered a compensatory mechanism for cerebellar degeneration. Nevertheless, compensation is not complete. Syllable timing shifts in the direction of equal syllable duration (Sidtis et al. 2010). Dysarthria is a typical feature of Friedreich’s ataxia (FRDA), and EMG activities recorded during the production of nonsense utterances (papapa/bababa. . .) in these patients show a prolongation of anticipatory and overall muscular activities. This abnormality might be an elemental feature of dysmetria and might reflect also some degrees of dysdiadochokinesia (see below). Given that FRDA is a spinocerebellar neurodegenerative disorder, it may be expected that the dysarthria is not purely ataxic (i.e., associated with impairment of the cerebellum and its

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connections), but rather involves a combination of ataxic/spastic/flaccid components. A recent study has shown that FRDA speech deficits are consistent with ataxic dysarthria and cerebellar lesions. Indeed, the deficits consist of mild impairments including consonant articulatory imprecision, reduced pitch variation, loudness maintenance, reduced phrase length, reduced breath support for speech, and hypernasality, thus resulting in a mild reduction in sentence intelligibility and communicative efficiency (Folker et al. 2010). Dysprosody is a disturbance in the melodic aspect of speech that may also occur in cerebellar patients. Comprehension is spared and paraphrasias are generally absent in cerebellar diseases (Manto 2002).

Dysphagia Dysphagia is a disruption in the swallowing process which may occur in many brain diseases including stroke, tumors, bulbar, and pseudobulbar paralysis, amyotrophic lateral sclerosis, multiple sclerosis, multisystem degenerations, Parkinson’s disease, Huntington’s disease, myasthenia gravis and myasthenic syndromes, myopathies, and peripherial neuropathies (Olszewski 2006). fMRI and the studies of regional cerebral blood flow using PET have demonstrated that volitional swallowing involves the basal ganglia and the cerebellum, as well as cortical structures of the brain. The cerebellum is activated bilaterally, especially on the left side (Suzuki et al. 2003; Zald and Pardo 1999). Investigation of swallowing function in patients with multiple system atrophy (MSA) with a clinical predominance of cerebellar symptoms (MSA-C) has revealed that, in addition to the role of the parkinsonism, the cerebellar dysfunction affects coordination of the tongue and is related to the swallowing impairment (Higo et al. 2005).

Ataxia of Limbs Ataxia of limbs may be defined as unsteadiness or incoordination of limbs, encompassing an impairment of the control of force and timing of movements. These abnormalities generate errors in speed, range, rhythm, starting, and stopping motor activity (Walker 1990). Cerebellar damage typically results in impairment in performance of limb movements. Unlike patients with impaired position sense, cerebellar patients are still able to describe the direction of the movement without looking at the moving limb (Lechtenberg 1993). The main deficits involving limbs are dysmetria, dysdiadochokinesia, postural and kinetic tremor, and decomposition of movement. As a general rule, motor deficits are lateralized to the side of the cerebellar lesion. MRI provides meaningful information in terms of correlating the cerebellar lesion site and the clinical data in subjects with degenerative as well as focal cerebellar disorders. For instance, limb ataxia is correlated with atrophy of the intermediate and lateral cerebellum in patients with cerebellar cortical degeneration (Timmann et al. 2008). A study on 39 patients following cerebellar stroke and grouped by

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stroke location has demonstrated that cerebellar stroke does not always result in motor impairment. In particular, the cerebellar motor syndrome resulted from stroke in the anterior lobe, while strokes involving posterior lobe produce minimal (lobule VI) or any (VII–X) motor impairment (Schmahmann et al. 2009). Lesionsymptom mapping studies on patients with cerebellar stroke are consistent in showing that the motor signs are mostly associated with lesions in paravermal regions of lobules IV/V and the deep cerebellar nuclei (see below “Topography of Clinical Deficits”) (Schoch et al. 2006). Moreover, it has been highlighted that in the acute stage of cerebellar stroke, arm movements mainly present abnormal slowness (similar to bradykinesia) and not incoordination, with fast recovery (Konczak et al. 2010). When these patients are asked to perform the movements as fast as possible, they often overshoot the target and may also show an ability to increase the speed of motion. Therefore, the slowness of movement might be due to a compensatory phenomenon aiming at reducing the overshoot. Dysmetria is an error in trajectory due to a disturbed range, rate, and force of movement (Holmes 1917, 1922; Gilman et al. 1981). In most cases, dysmetria occurs both for proximal and distal joints and is often followed by corrective movements (Hore et al. 1991). Both hypermetria and hypometria occur in cerebellar patients, even though hypometria is less common. Hypermetria refers to the overshoot of the target (Fig. 71.4) and is largest when the movement is made as fast as possible and when the inertia of the moving limb is increased (Manto and Jacquy 1994; Manto et al. 1995a, b). Hyperventilation increases hypermetria in patients with spinocerebellar ataxia type 6 (SCA 6) (Manto and Bosse 2003). Hypermetria is often associated with a delayed onset latency of the antagonist EMG activity (Fig. 71.5). This neurophysiological finding as well as the severity of hypermetria correlates well with the AS20 score, a clinical rating score of ataxia (Manto 2010). Lesions of the dentate nuclei are typically associated with an overshoot of the target and a decomposition of multi-joint movements (Thach et al. 1992; Bastian et al. 1996). Decomposition of movement into elementary components is due to a lack of synergy between joints, resulting in a lack of fluidity in motion and in ataxic movements (Topka et al. 1998). Decomposition of movement may be assessed by the index-to-wrist maneuver: the patient performs a pointing movement toward the wrist of the examiner which is maintained horizontally at the height of the patient’s shoulder and at an approximate distance of 85% of the patient’s upper limb’s height (Manto 2002). In this maneuver, cerebellar patients demonstrate asynchronous movements of the shoulder and the elbow. For slow multi-joint movements, decomposition is manifested by errors in the direction and rate of the movement. Decomposition of movement is often accompanied by an inability to generate independent finger movements. The attempt to move the thumb and the index alone induces successive flexion of the other fingers (“sign of the piano”); this sign is also detectable by asking the patient to perform repeated tapping of the index against the thumb (Manto 2002). Deficits in adaptation of the interaction torques generated during a multi-degree of freedom human arm have been demonstrated in cerebellar patients and could account for the decomposition (Topka and Massaquoi 2002).

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Fig. 71.4 Top panels: illustration of fast single-joint voluntary movements in a control subject (a) and in a cerebellar patient (b). Motion is typically hypermetric in the patient, overshooting the target located at 0.4 rad from the starting position. Bottom: illustration of the structures of the cerebellum associated with hypermetric ballistic movements. The lobules associated with an inability to adapt to inertia are indicated also. Abbreviations: C central, Culm culmen, H hemispheric, Tons tonsil, Flocc flocculus (the paraflocculus is illustrated above the flocculus), D declive, F folium, T tuber, Pyr pyramis, Uv uvula, Nod nodulus. (From Grimaldi and Manto (2011b), with permission)

Dysdiadochokinesia (also called adiadochokinesia) denotes the inability to perform rapid successive movements (Babinski 1902), resulting in irregular and slow alternating sequential movements. The successive pronation/supination task is typically used to evaluate this deficit. In advanced cases, dysdiadochokinesia often results in an abnormal sway of the elbow, and the alternate character of the task may therefore not be detectable (Manto 2002). Dysrythmokinesia is one of the characteristics of adiadochokinesia. It consists of a disturbed rhythm of movements contrasted with relatively preserved accuracy of movements. Dysrythmokinesia can be detected in a tapping test. Cerebellar tremor. Tremor in cerebellar disease is mainly composed of low frequency oscillations. There is usually a kinetic component, which is often

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AGO1 AGO2 FCR

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ECR Latency

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Fig. 71.5 Triphasic pattern of electromyographic (EMG) activities in a control subject (left) and in a cerebellar patient exhibiting hypermetria (right). In the control subject, the first agonist burst (AGO1) is followed by a burst in the antagonist muscle (ANTA), followed by a second burst in the agonist muscle (AGO2). In the cerebellar patient, three EMG deficits are observed: the rate of rise of EMG activities is depressed, the onset latency of the antagonist EMG activity is delayed and the two agonist bursts are not demarcated. FCR: flexor carpi radialis; ECR: extensor carpi radialis. EMG traces are full-wave rectified and averaged (n ¼ 10 movements) (From Manto (2009), with permission)

associated with a concomitant postural tremor (Rondot and Bathien 1995). The term “action tremor” refers to any tremor produced by voluntary contraction of muscles. It includes postural, isometric, and kinetic tremor (Grimaldi and Manto 2008). Tremor may be bilateral, but in most cases oscillations are observed ipsilateral to the cerebellar lesion. Isometric tremor occurs when a voluntary muscle contraction is opposed by a rigid stationary object (Findley and Koller 1995). Kinetic tremor appears during the execution of a movement and is usually maximal as the limb approaches the target (Holmes 1939). Kinetic tremor tends to involve predominantly the proximal musculature (Gilman et al. 1981; Lechtenberg 1993) and decreases with inertia, unlike cerebellar dysmetria (Chase et al. 1965; Hewer et al. 1972). Kinetic tremor has a frequency between 2 and 7 Hz in the large majority of cases. In most cases also, oscillations are perpendicular to the main direction of the intended movement. Kinetic tremor is evaluated by the following maneuvers: (1) finger-to-nose test: patient performs movements of one upper limb with the hand initially on the ipsilateral thigh and then touching the nose with the index; (2) finger-to-finger test: patient touches the examiner’s finger, which is moved and stopped in different locations in space; and (3) knee-tibia test: this test is executed in the supine position. The patient raises one leg and places the heel on the contralateral knee, which is kept motionless. The patient slides the heel down the tibial surface in a regular way toward the ankle. The heel is then raised again up to the resting knee (Manto 2002). Postural tremor appears during postural tasks. Its frequency is usually between 4 and 12 Hz (Fig. 71.6 and 71.7). Tremor appears immediately, but increases in amplitude after a few seconds in the line of gravity. It is tested clinically by the following maneuvers: (1) the arm outstretched task: holding the upper limbs outstretched with the hands in supination, parallel to the

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Fig. 71.6 Monoaxial accelerometry in essential tremor. The patient is asked to maintain the upper limbs horizontally parallel to the floor. A peak frequency of 7.5 Hz is clearly identified on the power spectrum (From Grimaldi and Manto (2008), with permission)

floor at the height of the shoulder; (2) index-to-index test: the patient is asked to maintain the two index fingers medially, pointing at each other at a distance of about 1 cm; forearms are maintained horizontally at the height of the shoulders; and (3) the heel-to-knee test: the patient is asked to maintain the heel on the knee for several seconds (Stewart and Holmes 1904; Holmes 1922). The oscillations appearing during the heel-to-knee test rapidly evolve into lateral sways in severe cases. A typical 3 Hz leg tremor may be observed during the sustained leg elevation with 90 flexion in the knee and hip joint, and is often suggestive of atrophy of the anterior cerebellar lobe in alcoholic patients (Fig. 71.8). Generally, action tremor is suggestive of anterior lobe cerebellar pathology; however, it may be observed also in diffuse cerebellar diseases such as idiopathic late-onset cerebellar atrophy (ILOCA) or hereditary spinocerebellar ataxias (SCAs) (Silfverski€old 1977; Mauritz et al. 1979; Harayama et al. 1983; Manto et al. 1996). Eye closure and body displacements tend to enhance the oscillations. Postural tremor in cerebellar disease (Table 71.3) can be further described as (see Table 71.3): (a) precision tremor – usually due to lesions of cerebellar nuclei – has a frequency of 2–5 Hz, occurring during the execution of precision tasks and involving the distal musculature; (b) asthenic tremor – in case of hemispherical lesion – precipitated by fatigue; (c) axial postural tremor; and (d) midbrain tremor associated with midbrain lesions (Brown et al. 1997). A postural tremor of the shoulder may be precipitated by fatiguing tasks in patients with a large cerebellar malformation involving a cerebellar hemisphere. Isometrataxia is the inability to maintain constant forces during skilled tasks requiring hand or finger use (Mai et al. 1989). It is tested by asking the patient to

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3.40 Hz

POSTURAL TREMOR–CEREBELLAR STROKE 0.30 V2/sec Accelerometer - Upper limb outstreched

10 sec Flexor Carpi Radialis - Rectified EMG 3.37 Hz 10 sec 0.02 V2/sec

Fig. 71.7 Postural tremor in a patient presenting a cerebellar stroke in the territory of the superior cerebellar artery, involving the outflow tract of the cerebellar nuclei. Data from a monoaxial accelerometer and from a surface electromyographic (EMG) sensor are shown, as well as the corresponding power spectra (From Grimaldi and Manto (2008), with permission)

exert a slight and constant pinch force (using his index finger and thumb) on the lateral part of the examiner’s thumb, which will feel an irregular pressure in absence of tremor of the hand. Isometrataxia is an underestimated cerebellar deficit. Isometric tremor often masks isometrataxia. Nevertheless, isometrataxia can be differentiated from the isometric tremor thanks to two major features: it is

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Fig. 71.8 3-Hz body sway in chronic alcoholic cerebellar degeneration. (a) Representation of the center of pressure during a Romberg test using a pressure platform. (b) Displacements of the center of pressure in orthogonal axis X-Y. (c) Superimposition of power spectral density curves in the anterior-posterior axis. The arrows shows the 3-Hz tremor. (d) Time-frequency representation of the oscillations (From Manto (2010), with permission)

Table 71.3 Various presentations of cerebellar postural tremors Tremor type Precipitants Frequency (Hz) Midbrain tremor Any posture 2–5 Asthenic cerebellar tremor Fatigue/weakness Irregular Precision cerebellar tremor Accurate 2–5 placements Cerebellar axial postural tremor Any posture 2–10 Cerebellar proximal exertional tremor Prolonged exercise 3–4 Adapted from Brown et al. (1997)

Distribution Distal > proximal Proximal + distal Distal Proximal > distal Proximal > distal

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not a rhythmic phenomenon and it occurs during slight contractions. Isometrataxia may be observed in the first hours after a cerebellar stroke in the territory of the posterior inferior cerebellar artery (Manto 2002). Disorders of muscle tone. Muscle tone is assessed by passively moving the wrists, elbows, shoulders, ankles, knees, and hips of the patient and by grasping the patient’s forearm and shaking the relaxed arm. Hypotonia is usually associated with severe cerebellar damage. The decline in resistance to the passive manipulation of limbs tends to be more pronounced in proximal joints. Pendular tendon reflexes may be observed, notably at the knee, characterized by the distal leg continuing to swing (like a pendulum) five or more times following elicitation of the patella tendon reflex (Holmes 1922, 1939). Differential diagnosis of cerebellar hypotonia includes extensive brainstem lesions, spinal shock, anterior horn cell disease,poyradiculitis/polyneuropathy, and floppy syndrome in children (Manto 2010). Disorder of muscle tone in cerebellar injury may also present as spasms associated with intermittent opisthotonos (Stewart and Holmes 1904). Such cerebellar fits occur in patients with posterior fossa tumors, Chiari malformations, and stroke involving the cerebellar cortex but sparing the nuclei (Manto 2010). The mechanism is thought to be extensor tone disinhibition (Sprague and Chambers 1953). Paroxysmal facial contractions may be observed in ganglioglioma of the cerebellum (Chae et al. 2001) and in children with hamartoma of the floor of the fourth ventricle (Manto 2010). Loss of check and rebound. Impaired check causes a large movement called excessive rebound. It is assessed by asking the patient to maintain the upper limbs extended with the hands pronated. The examiner exerts a tap on the wrist thus causing a large displacement of the limb, immediately followed by an overshoot at the initial position followed by oscillations (Manto 2002). The lack of check can be evaluated during the Stewart-Holmes maneuver (Stewart and Holmes 1904): the patient is asked to perform a forceful flexion of the elbow while the examiner attempts to extend the joint. When the examiner abruptly releases the forearm, the arm flexion continues unopposed and the patient is at risk of hitting him/herself in the shoulder, chest, or face. This of course should be prevented by the examiner placing his/her other arm between the patient’s limb and face prior to abruptly releasing the patient’s limb that is being tested. Handwriting abnormalities. Cerebellar patients show difficulties in handwriting. This is assessed by asking the patient to write standard sentences, and it includes drawing the Archimedes’ spiral. The subject is comfortably seated in front of a table, and a sheet of paper is fixed on the table with tape to avoid motion artifacts. The subject executes the task with his dominant hand without timing requirements (Grimaldi and Manto 2008). Writing is irregular. Megalographia (also called macrographia) is another clinical sign of cerebellar dysfunction. Handwriting is abnormally large (Frings et al. 2010). The observation of letters “unequal in size and irregularly spaced” has been reported by Holmes (1917) in patients with cerebellar lesions following gunshot injuries.

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Ataxia of Stance and Gait Ataxia of stance is characterized by an inability to maintain the body in a stationary position. Body sway is increased and the trunk tends to lurch from side to side or to drift laterally (Gilman et al. 1981). Cerebellar patients have a charactersitically broad-based stance with increased body sway. They may also show distorted anticipatory adjustments and defective postural responses to external forces (Horak and Diener 1994). Oscillations occur in the anterior-posterior plane (3 Hz sway due to lesions of the anterior cerebellar lobe), in the lateral plane, or are rotatory. A low frequency sway (40 years), slowly progressive ataxia with associated tremor and sensory neuropathy (Verbeek et al. 2004). Clinical features included gait and limb ataxia, with variable dysarthria, slow saccades, ocular dysmetria, and decreased vibratory sense below the knees. Four affected individuals showed hyperreflexia, two of whom also had extensor plantar responses. Only one patient, with a disease duration of 23 years, was wheelchair bound. MRI of that patient showed severe cerebellar atrophy. Postmortem examination of one patient showed frontotemporal atrophy, atrophy of the cerebellar vermis, pons, and spinal cord. There was neuronal loss in the cerebellar vermis, dentate nuclei, and inferior olives, but not in the pons. There was also thinning of the cerebellopontine tracts and demyelination of the posterior and lateral columns of the spinal cord. Following disease localization on chromosome region 20p13-p12.3 (Verbeek et al. 2004), missense mutations in the PDYN gene encoding prodynorphin were identified in the original family as well as in other three Dutch families with progressive ataxia (Bakalkin et al. 2010). SCA23 appears to be a rare (0.5%) cause of ADCAs in the European population. Prodynorphin is a precursor protein for opioid neuropeptides, alpha-neoendorphin, and dynorphins A (Dyn A) and B (Dyn B). Although the pathogenic mechanisms remain to be elucidated, alterations in Dyn A activities and/or impairment of secretory pathways by mutant prodynorphin may lead to glutamate neurotoxicity, which underlies Purkinje cell degeneration and ataxia (Bakalkin et al. 2010). SCA25 SCA25 has been described in a single French family with spinocerebellar ataxia and sensory involvement (Stevanin et al. 2004). This disease demonstrates significant intrafamilial phenotypic variability, with the clinical picture ranging from pure

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sensory neuropathy with little cerebellar involvement to a Friedreich’s ataxia-like phenotype. Some individuals presented with cerebellar ataxia and a sensory neuropathy in infancy; others manifested prominent sensory neuropathy with mild ataxia at an older age. In three of seven individuals, vomiting and gastrointestinal features were the presenting symptoms. Based on this family, SCA25 shows reduced penetrance. MRI showed significant global cerebellar atrophy without brainstem involvement. Sural nerve biopsy of one patient showed loss of myelinated fibers and EMG showed sensory involvement. Linkage studies mapped the mutated gene to the locus 2p21-p13 (Stevanin et al. 2004) but the responsible gene has not been identified yet.

SCA26 SCA26 was described only in a single Norwegian family so far (Yu et al. 2005). Clinically, the affected members presented with pure cerebellar ataxia and dysarthria. Age at onset ranged from 26 to 60 years. MRI showed atrophy of the cerebellum sparing the pons and medulla. Anticipation was not observed. The disease was mapped to chromosome 19p13.3 adjacent to the Cayman ataxia and SCA6 loci. The region consists of 3.3 Mb with approximately 100 known and predicted genes (Yu et al. 2005). SCA27 SCA27 is characterized by impaired cognitive abilities and slowly progressive ataxia. It was described in a Dutch family with 14 patients, who presented with childhood-onset postural tremor and slowly progressing ataxia evolving from young adulthood (Brusse et al. 2006; van Swieten et al. 2003). In several of these patients, orofacial dyskinesia, suggesting basal ganglia affection, was present. Neuropsychological testing additionally revealed intellectual decline, behavioral problems (aggressive outbursts), and cognitive impairment (memory deficits and executive dysfunction). MRI revealed only moderate cerebellar atrophy, but could be notably normal in the patients with a shorter disease duration (Brusse et al. 2006; van Swieten et al. 2003). SCA27 is caused by mutations in the fibroblast growth factor 14 gene (FGF14) (Brusse et al. 2006; Dalski et al. 2005; van Swieten et al. 2003). The occurrence of a frameshift mutation (Dalski et al. 2005) and a chromosomal translocation disrupting FGF14 (Misceo et al. 2009) suggests a mechanism of haploinsufficiency. FGF14 is required for the correct expression and regulation of the voltage-gated Na+ channel; loss of FGF14 function has been shown to reduce excitability of hippocampal neurons and impair spontaneous and repetitive firing in Purkinje neurons (Laezza et al. 2007; Shakkottai et al. 2009). SCA28 SCA28 was first described in a four-generation Italian family presenting with juvenile-onset, and slowly progressive gait and limb ataxia, dysarthria, hyperreflexia at lower limbs, nystagmus, and ophthalmoparesis (Cagnoli et al. 2006; Mariotti et al.

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2008). Mean age at onset was around 20 years, starting with imbalanced standing, gaze-evoked nystagmus, and mild gait uncoordination. Later on, slow saccades, CPEO, ptosis, and exaggerated deep tendon reflexes develop (Mariotti et al. 2008). SCA28 is caused by mutations in the AFG3L2 gene located on chromosome 18p11.22 (Di Bella et al. 2010). Missense mutations have been identified in more than ten European families (Cagnoli et al. 2010; Di Bella et al. 2010; Edener et al. 2010). Age at onset is variable (range: 6–60 years) but disease most often begins in midlife with slowly progressive gait ataxia. The clinical phenotype is characterized by an ataxic syndrome, frequently associated with progressive ophthalmoplegia, ptosis, and pyramidal hyperreflexia with Babinski signs and altered vibration sense; less frequently, parkinsonian or dystonic features are also reported. EMG did not show evidence of polyneuropathy. In some rare cases, SCA28 mutations are associated with reduced penetrance (Di Bella et al. 2010). Cerebral MRI shows isolated cerebellar, mostly vermian, atrophy (Cagnoli et al. 2010; Di Bella et al. 2010; Edener et al. 2010). Recently, a homozygous mutation has been identified in two siblings with a distinct phenotype characterized clinically by lower extremity spasticity, peripheral neuropathy, ptosis, oculomotor apraxia, dystonia, cerebellar atrophy, and progressive myoclonic epilepsy (Pierson et al. 2011). AFG3L2 is a component of the m-AAA metalloprotease complex, located in the internal mitochondrial membrane. Interestingly, the homologous protein paraplegin is also a component of the same mitochondrial complex and causes recessive spastic paraplegia type 7 (SPG7), a neurodegenerative disease closely related to spinocerebellar ataxia (Casari et al. 1998). The m-AAA metalloprotease complex is part of the mitochondrial protein quality control system, participating in both the degradation of incorrectly folded protein and the assembly of components of the inner membrane, including complexes of the respiratory chain (Tatsuta and Langer 2008). SCA28 is the first ADCA caused by the alteration of a mitochondrial protein. So far only missense mutations have been described, the vast majority of which occur in the highly conserved proteolytic domain (Cagnoli et al. 2010; Di Bella et al. 2010; Edener et al. 2010). In a cellular model, the mutations impair AFG3L2 proteolytic activity, cytochrome c oxidase activity, and eventually cell respiration (Di Bella et al. 2010). A dominant-negative effect has been proposed for the heterozygous mutations causing SCA28 (Di Bella et al. 2010). Although AFG3L2 and paraplegin were found to be selectively highly expressed in human Purkinje cells and primary motor neurons, respectively (Di Bella et al. 2010), the pathogenic mechanism of the specific and distinct neurodegenerative phenotype linked to paraplegin/SPG7 or AFG3L2 mutations remains unknown.

SCA29 In a four-generation Australian family with congenital nonprogressive ataxia associated with cognitive impairment, Dudding and colleagues (Dudding et al. 2004) identified a locus on chromosome 3p26 with overlap with the SCA15 locus. Examination of the family showed that all affected members had gait ataxia and cognitive disability with variable features of dysarthria, dysmetria, dysdiadochokinesia, nystagmus, dystonic movements, and cerebellar hypoplasia on

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imaging. Clinical signs of pyramidal tract dysfunction and sensory changes were absent. No further genetic study has been reported for this family. It remains to be elucidated whether SCA29 and SCA15/16 represent the same disorder.

SCA30 SCA30, thus far, appears to be a primarily cerebellar type ataxia (ADCA-III). Storey and colleagues described an Anglo-Celtic family from Australia with a slowly progressive, late-onset pure cerebellar ataxia disorder (Storey et al. 2009). The mean reported age at onset was 52 years (range 45–76). The phenotype was characterized by relatively pure, slowly evolving gait and appendicular ataxia with mild-to-moderate dysarthria. Four individuals had mild lower limb hyperreflexia; none had evidence of neuropathy. All patients had hypermetric saccades with normal vestibuloocular reflex gain. MRI of two patients showed cerebellar atrophy with preservation of the brainstem. Linkage analysis identified a novel locus on chromosome 4q34.3-q35.1 (Storey et al. 2009) but the responsible gene has not been identified yet. SCA31 This ADCA, previously called “chromosome 16q22.1-linked spinocerebellar ataxia” is now termed “SCA31.” This disorder was originally reported from Japan in a study of six families (Nagaoka et al. 2000). The genetic locus lies on 16q22 within the SCA4 critical region, but the phenotype is different in that there is no sensory motor polyneuropathy in the Japanese patients. The core clinical features have been slowly progressive pure cerebellar ataxia (ADCA-III) with onset in late adult life (mean age at onset ¼ 60 years) variably associated with sensorineural hearing loss (Ishikawa et al. 2005; Nagaoka et al. 2000; Ouyang et al. 2006; Owada et al. 2005). Many patients have brisk tendon reflexes (Ouyang et al. 2006). There was no evidence for a polyneuropathy by nerve conduction studies (Nagaoka et al. 2000). Previously, a C/T substitution in the 50-untranslated region of the puratrophin-1 gene (16C > T in PLEKHG4) (Ishikawa et al. 2005) or a disease-specific haplotype just upstream of PLEKHG4 was used for the diagnosis of SCA31. The disease was found to segregate with a heterozygous 16C > T transition in the 50 untranslated region of the PLEKHG4 gene on chromosome 16q22, which codes for pleckstrin homology domain-containing protein, also known as puratrophin-1 (Ishikawa et al. 2005). Subsequent studies identified affected individuals without the 16C > T transition, indicating that the causative mutation is likely to be centromeric to the C > T transition (Amino et al. 2007). Very recently, SCA31 was found to be associated with a peculiar insertional mutation, a 2.8- to 3.5-kb-long (TGGAA)n pentanucleotide repeat inserted in 900 kb complex region containing other (TAAAA)n, (TAGAA)n pentanucleotide repeats. Since (TAAAA)n and (TAGAA)n were also found in controls, only (TGGAA)n is considered pathogenic (Sakai et al. 2010; Sato et al. 2009). The length of the SCA31 repeat insertion inversely correlated with patient age of onset, and an expansion was documented in a single family showing anticipation (Sato et al. 2009). The SCA31

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repeat insertion was excluded in the SCA4 sensory ataxia families (Edener et al. 2011), clearly demonstrating that SCA31 and SCA4 are distinct disorders caused by different genes at the same locus (see above). The SCA31 complex repeats appeared to be inserted in an intron shared by two different genes, BEAN (Brain Expressed, Associated with Nedd4) and TK2 (thymidine kinase 2), transcribed in opposite directions. Aggregations of RNA containing transcribed inserted repeats were found in the nuclei of patients’ Purkinje cells, suggesting that RNA-mediated gain-of-function toxicity is a likely pathogenic mechanism (Sato et al. 2009). So far, SCA31 has been described only in Japan, where it is one of the most common ADCAs (in Nagano, it accounts for 42% of ADCA families), and is probably caused by a single founder mutation (Sakai et al. 2010; Sato et al. 2009).

SCA32 Online Mendelian Inheritance in Man (OMIM) reports a Chinese family with an autosomal dominant form of spinocerebellar ataxia associated with cognitive impairment and azoospermia (Jiang et al. 2010). There was a broad range of age at onset, particularly among females. Those with onset of ataxia before age 40 years showed cognitive impairment. Brain MRI showed cerebellar atrophy. All affected males were infertile and had azoospermia with testicular atrophy. Linkage to chromosome 7q32-q33 was reported (MIM 613909). SCA35 SCA35 is characterized by cerebellar ataxia, upper limb involvement, and spasmodic torticollis. The disorder has been described in a four-generation Chinese family in which nine individuals developed spinocerebellar ataxia with a mean age at onset of 43.9 years (range, 40–48 years) (Wang et al. 2010). Early features included walking difficulty, ataxia, and cerebellar dysarthria, while upper limb ataxia occurred later. There was slow progression, and most patients needed a walking aid or became wheelchair-dependent after about 10 years. Additional features included tremor, hyperreflexia, extensor plantar responses, spasmodic torticollis, ocular dysmetria, and position sense defects. None of the patients had nystagmus, ophthalmoplegia, peripheral neuropathy, or cognitive impairment. MRI showed mild-to-severe cerebellar atrophy. SCA35 was mapped on chromosome 20p13-p12.2. Whole exome sequencing of four affected members allowed to identify a heterozygous missense mutation in the TGM6 gene encoding transglutaminase 6. A different heterozygous missense mutation in the same gene was found in a second unrelated Chinese family with a similar phenotype (Wang et al. 2010). Interestingly, the disease-associated region identified in SCA35 (20p13-12.2) is in a similar position to the region mapped in SCA23 (Verbeek et al. 2004) and SCA36 (Kobayashi et al. 2011), which are caused by mutations in the PDYN and NOP56 genes, respectively. The pathogenic mechanisms underlying SCA35 are still unknown. TGM6 is a member of the transglutaminase family, four of which (TGM1-3 and TGM6) have been shown to be expressed in the human brain (Jeitner et al. 2009). Cerebral

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transglutaminases catalyze the covalent attachment of glutaminyl residues and various other amine-bearing compounds, all of which have an impact on the formation of stable soluble and insoluble polymers (Jeitner et al. 2009). Several evidences link transglutaminases to neurodegenerative ataxias. Much higher activity of transglutaminases and transglutaminase-catalyzed products has been observed in several neurodegenerative disorders (Jeitner et al. 2009). TGM6 is predominantly expressed by a subset of neurons in the central nervous system, including Purkinje cells and TGM6 deposits are present in the cerebellum of patients with gluten ataxia (Hadjivassiliou et al. 2008). Finally, mouse studies have further shown that intraventricular injection of anti-transglutaminase antibodies provoked ataxia in mice (Boscolo et al. 2010).

SCA36 SCA36 has been described in Japanese families presenting with cerebellar ataxia associated with motor neuron involvement (Kobayashi et al. 2011). All affected individuals started their ataxic symptoms (gait ataxia, truncal instability, dysarthria, and limb incoordination) in their fifth to sixth decade of life (mean: 52.8 years). An unusual feature for SCA was the motor neuron involvement in patients with longer disease duration. Affected individuals developed tongue atrophy with fasciculation, although swallowing function was relatively preserved. Skeletal muscle atrophy and fasciculation in the limbs and trunk were observed in advanced cases. Most patients had hyperreflexia, but none had severe lower limb spasticity or extensor plantar responses. Brain MRI showed mild cerebellar atrophy. Nerve conduction studies were normal, whereas EMG showed neurogenic changes only in cases with skeletal muscle atrophy, indicating a lower motor neuropathy. The pattern of muscle involvement and progression differed from that seen in amyotrophic lateral sclerosis (ALS). SCA36 was mapped to chromosome 20p13 in a similar position to the region mapped in SCA23 (Verbeek et al. 2004) and SCA35 (Wang et al. 2010). Candidate gene sequencing and repeat analysis did not find any mutation in the PDYN and TGM6 genes but identified a pathogenic heterozygous hexanucleotide repeat expansion (GGCCTG) in intron 1 of the NOP56 gene. Normal GGCCTG repeat sizes ranged from 3 to 8. Expanded repeats in patients ranged from 1,500 to 2,500 units. There was no inverse correlation between age at onset and number of repeats and no obvious anticipation in the pedigrees. Overall, 9 (3.6%) unrelated cases were found among 251 cohort patients. Intranuclear RNA inclusions (RNA foci) were found in SCA36 patients’ cells. Several evidences indicate that RNA foci play a role in the etiology of SCA and other neurodegenerative diseases through sequestration of specific RNA-binding proteins (Daughters et al. 2009; Wojciechowska and Krzyzosiak 2011). Gel-shift assays showed that the SCA35 expanded GGCCUG repeat selectively bound and sequestered the RNA-binding protein SFRS2. In addition, transcription of miRNA MIR1292, located within intron 1 of NOP56, was significantly decreased in lymphoblastoid cells of SCA36 patients. Altogether, the findings indicate that pathogenesis of SCA36 is likely to involve a toxic RNA gain of function (Kobayashi et al. 2011).

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DRPLA Dentatorubral-pallidoluysian atrophy (DRPLA) is named for the pathologic findings associated with the disease (Oyanagi 2000). First described in 1982 in Japan, it occurs with highest frequency in the Japanese population (third cause of SCA in Japan with a prevalence of 0.2–0.7/100,000) (Takano et al. 1998). A limited number of cases were reported from European countries, North America, and Turkey (Whaley et al. 2011). Onset of the disease varies from early childhood to late adulthood (>70 years) with a mean age around 45 years (Tsuji 2012). DRPLA can cause a wide range of clinical manifestations, including ataxia, choreoathetosis, dystonia, ballismus, myoclonus, epilepsy, and dementia (Takahashi et al. 1988). The clinical phenotype depends on age at onset and repeat length. The most constant clinical findings in DRPLA are cerebellar ataxia, dysarthria, and progressive dementia. These features are present in almost all patients irrespective of age of onset and repeat length. Patients with disease onset before the age of 20 years and large expansions show the clinical syndrome of progressive myoclonus epilepsy. Some of them have opsoclonus. In these patients, the differential diagnosis includes Lafora disease, Unverricht–Lundborg disease, neuronal ceroid lipofuscinosis, and MERRF (Tsuji 2012). In patients with later disease onset and shorter expansions, myoclonus and seizures are less prominent. Instead, many of these patients have involuntary choreic or dystonic movements and psychiatric abnormalities, including personality changes, hallucinations, and delusional ideas. Thus, the adult form of DRPLA needs to be differentiated from Huntington disease and SCA17. In the late-onset form, ataxia can occur years before dementia or chorea develops (Yabe et al. 2002). Most patients also have oculomotor abnormalities, including gazeevoked nystagmus, broken up smooth pursuit, square wave jerks, and vertical gaze palsy (Ikeuchi et al. 1995). Neuropathologically, there is degeneration of the dentate nucleus, with its projection to the red nucleus, and the external pallidum, with its projection to the subthalamic nucleus. Usually, the dentatorubral system is more severely affected. Atrophy may be also present in other basal ganglia nuclei, the thalamus, and the inferior olives. Involvement of the pontine tegmentum has been also found, and this appears to correlate with oculomotor abnormalities (Oyanagi 2000). MRI shows atrophy of cerebellum, brainstem, and cerebrum, with high-intensity signals in the pallidum and cerebral white matter on T2weighted images (Tsuji 2012). DRPLA is caused by an expanded CAG repeat (polyglutamine) in the coding region of the gene (ATN1) encoding atrophin-1, a protein of unknown function (Koide et al. 1994). The normal allele has 6–35 repeats, but individuals affected with DRPLA have 48–93 repeats (Tsuji 2010) (Fig. 101.2, see figure legend for details). If inherited from the paternal lineage, DRPLA alleles may undergo further expansion causing anticipation. The gene product, atrophin-1, has an unknown function. Immunohistochemical studies on patients’ tissues have demonstrated that diffuse accumulation of mutant atrophin-1 in the neuronal nuclei, rather than the formation of neuronal intranuclear inclusions (NIIs), was the predominant pathologic condition and involved a wide range of central nervous system regions (Yamada et al. 2001). Evidence suggests that atrophin-1 is a component of

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a transcription corepressor complex and that polyglutamine-induced toxicity in DRPLA may be mediated by repression of CREB-dependent transcriptional activation (Shimohata et al. 2005).

Episodic Ataxias Episodic ataxias (EAs) are a clinically and genetically heterogeneous group of rare autosomal dominant diseases characterized by recurrent, paroxysmal episodes of incoordination and imbalance, giddiness, and vertigo, often with associated progressive ataxia (Table 101.4). The symptoms are mainly cerebellar in origin but some of them present with additional abnormalities during the attacks. The incidence of EA is likely to be less than 1/100,000 (Jen et al. 2007). EAs are currently classified by their genetic etiologies. Like SCAs, they are labeled as EA followed by a number to denote the distinct chromosomal locus. Seven distinct subtypes (EA1-7) have been described, with six loci and four genes identified thus far. It is quite likely, however, that the number of phenotypes and mutated genes will grow further. Despite the significant genetic heterogeneity, the majority of clinical cases result from two recognized entities, EA1 and EA2 (Jen et al. 2007). So far, the genes identified in episodic ataxias code for membrane proteins including voltage-gated ion (K+ and Ca2+) channels and a neurotransmitter (glutamate) transporter. The genetic identification of these genes broadened the clinical spectrum of episodic ataxia, now known to be variably associated with epilepsy, dystonia, hemiplegic migraine, myasthenia, and even intermittent coma (Jen et al. 2007).

EA1 EA1 is the second most common episodic ataxia and clinically characterized by brief attacks of ataxia and dysarthria, and interictal myokymia (i.e., twitching of small muscles around the eyes or in the hands). Disease onset occurs in early childhood. The episodes last seconds to minutes and are typically triggered by exercise, emotional stress, or startle (Jen et al. 2007). The interictal myokymia (also termed “neuromyotonia”) may be detected clinically or may only be apparent at electromyography (Brunt and van Weerden 1990). Brain MRI is normal. EA1 is clinically heterogeneous, even within kindreds. Graves and colleagues observed variable phenotypes in monozygotic twins, thereby implicating a role for nongenetic factors on the severity of the disease (Graves et al. 2010). Clinically, EA1 can be differentiated from EA2 (see below) by the shorter duration of the attacks, the presence of myokymia, and the absence of interictal ataxia and nystagmus. EA1 is caused by mutations in the KCNA1 gene on chromosome 12q13, encoding the voltage-gated potassium channel Kv1.1 (Browne et al. 1994). To date, 19 missense mutations and 1 nonsense mutations have been reported (Jen et al. 2007; Pessia and Hanna 2010).

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Table 101.4. Autosomal dominant cerebellar ataxias: genetic and clinical features of episodic ataxias Age at Main clinical onset MIMb Disease Genea Chromosomal locus Mutation symptoms (range) Number EA1 KCNA1 12p13 Point Muscle spasms, 2–15 160120 mutations interictal myokymia and jerking movements, chorea at onset Attack duration: seconds to minutes Response to acetazolamide: occasional EA2 CACNA1A 19p13 (allelic to Point Downbeat 2–20 108500 spinocerebellar mutations nystagmus, ataxia SCA6 and Deletions dysarthria, vertigo, familial hemiplegic muscle weakness, migraine FHM1) migraine. Interictal ataxia and nystagmus Attack duration: hours to days Response to acetazolamide: yes EA3 Unknown 1q42 – Myokymia, 1–42 606554 migraine, tinnitus, vertigo, dysarthria Attack duration: 1 min to 6 h Response to acetazolamide: yes EA4 / Unknown Unknown – Vertigo, diplopia. 23–60 606552 PATX Interictal nystagmus and saccadic smooth pursuit Attack duration: hours Response to acetazolamide: no EA5 CACNB4 2q22-23 Point Vertigo. Interictal 3–19 601949 mutations nystagmus, ataxia, epilepsy Attack duration: hours Response to acetazolamide: transient (continued)

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Table 101.4. (continued)

Disease Genea EA6 SLC1A3

EA7

Unknown

Main clinical Chromosomal locus Mutation symptoms 5p13 Point Cognitive mutations impairment. Interictal epilepsy, migraine, ataxia, motor delayed milestones Attack duration: hours/day Response to acetazolamide: no 19q13 – Vertigo, dysarthria, muscle weakness Attack duration: hours/days Response to acetazolamide: not reported

Age at onset MIMb (range) Number 90% of probands positive for polyglutamine ADCA do have a dominant family history (Marelli et al. 2011a). The small percentage of the patients with a genetically confirmed polyglutamine disease, who did not have a clear dominant familial history, had at least another affected subject in the family or the family history was ambiguous due to adoption, unknown paternity, or premature death of the parents (Marelli et al. 2011a). Thus, a search for polyglutamine expansion is not warranted in true sporadic cases, in whom the frequency of positive genetic tests is expected to be quite low (2–22%) (Brusse et al. 2007; Klockgether 2008).

Treatment At the moment, there are no proven therapies for SCAs. Repeatedly, attempts have been made to relieve ataxia but no confirmed pharmacological treatment is yet available (Ogawa 2004). Adaptive rehabilitation directed at maximizing functional capacity and symptom management continue to play a prominent role in managing these patients. There are aspects of these disorders that can be relieved, often dramatically, by medications. Examples include dopaminergic drugs for parkinsonian phenotypes, drugs to relieve spasticity and to treat sleep-related symptoms. There have been trials targeting neurotransmitter function of various types (e.g., physostigmin and serotoninergic and cholinergic drugs), with modest results (Gazulla and Modrego 2008; Ogawa 2004). Some efficacy has been reported for thyreotropin-releasing hormone, D-cycloserine, and azetazolamide in SCA6, but well-designed studies are lacking (Ogawa 2004). Severity and frequency of the unsteadiness of attacks in EAs may be reduced by acetazolamide (Jen 2008). Particularly, patients with EA2 can be dramatically responsive to acetazolamide, resulting in a decrease of frequency, duration, and severity of attacks. Acetazolamide is usually started at 125–250 mg/day and then increased up to 1,000 mg/day if needed and tolerated (Jen 2008). Side effects, such as tingling, numbness, decreased appetite, altered taste, impaired concentration and memory, or kidney stones have to be encountered. Nonpharmacologic interventions are extremely important. Physical and occupational therapists can recommend changes needed for safety at home, establish routines to minimize debilitation, advice on the use of adaptive equipment, assess driving safety, and provide “hands-on” psychologic support. Speech and language pathologists can provide testing and interventions to decrease the risk from swallowing abnormalities.

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Future Perspectives Data from animal and cell models show that future therapeutic strategies may rely on silencing gene expression, increasing the protein clearance, reducing the toxicity of the mutated protein, or influencing downstream pathways activated by the mutant protein or transplantation (Bauer and Nukina 2009; Underwood and Rubinsztein 2008). As pathogenic information is clarified, treatments directed at specific aspects of these cellular cascades may be developed. A promising strategy in the laboratory has been to reduce the expression of the mutated allele in some CAG/polyglutamine expansion disorders such as SCA3/MJD and SCAl, based on the toxic gain-offunction hypothesis (Xia et al. 2004). This has been achieved silencing the disease gene by RNA interference (RNAi). The therapeutic value of this approach in SCAs has already been shown in SCA1 transgenic disease models (Xia et al. 2004). However, numerous problems related to safety, delivery, and dosage remain to be solved before RNAi will become a viable alternative to conventional pharmacological approaches. A rational approach and a promising strategy for the development of therapies is the selection of an appropriate drug target based on an understanding of the molecular pathogenic mechanisms (e.g., effects on transcription, protein or RNA aggregation, oxidative stress, and intracellular signaling) which lead to neurodegeneration. This approach is greatly facilitated by the availability of transgenic animal models. Examples of drug candidates that have emerged from this approach are the rapamycin ester temsirolimus (SCA3) (Menzies et al. 2010) and dantrolene (SCA3) (Chen et al. 2008). Given the central role of protein aggregation in pathogenesis, the search for antiaggregation compounds may yield a compound that acts on all polyglutamine diseases. Lithium therapy has shown promise in a transgenic model of SCAl and this may be related to its effects on transcriptional regulation (Watase et al. 2007). Another avenue for modifying transcriptional dysregulation has been the use of HDAC inhibitors to improve the chromatin changes induced by the polyglutamine proteins (Steffan et al. 2001; Ying et al. 2006).

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Xia H, Mao Q, Eliason SL, Harper SQ, Martins IH, Orr HT et al (2004) RNAi suppresses polyglutamine-induced neurodegeneration in a model of spinocerebellar ataxia. Nat Med 10:816–820 Yabe I, Sasaki H, Kikuchi S, Nonaka M, Moriwaka F, Tashiro K (2002) Late onset ataxia phenotype in dentatorubro-pallidoluysian atrophy (DRPLA). J Neurol 249:432–436 Yabe I, Sasaki H, Takeichi N, Takei A, Hamada T, Fukushima K et al (2003) Positional vertigo and macroscopic downbeat positioning nystagmus in spinocerebellar ataxia type 6 (SCA6). J Neurol 250:440–443 Yamada M, Wood JD, Shimohata T, Hayashi S, Tsuji S, Ross CA et al (2001) Widespread occurrence of intranuclear atrophin-1 accumulation in the central nervous system neurons of patients with dentatorubral-pallidoluysian atrophy. Ann Neurol 49:14–23 Yamada M, Sato T, Tsuji S, Takahashi H (2008) CAG repeat disorder models and human neuropathology: similarities and differences. Acta Neuropathol 115:71–86 Yamashita I, Sasaki H, Yabe I, Fukazawa T, Nogoshi S, Komeichi K et al (2000) A novel locus for dominant cerebellar ataxia (SCA14) maps to a 10.2-cM interval flanked by D19S206 and D19S605 on chromosome 19q13.4-qter. Ann Neurol 48:156–163 Ying M, Xu R, Wu X, Zhu H, Zhuang Y, Han M et al (2006) Sodium butyrate ameliorates histone hypoacetylation and neurodegenerative phenotypes in a mouse model for DRPLA. J Biol Chem 281:12580–12586 Yu GY, Howell MJ, Roller MJ, Xie TD, Gomez CM (2005) Spinocerebellar ataxia type 26 maps to chromosome 19p13.3 adjacent to SCA6. Ann Neurol 57:349–354 Zhang Y, Snider A, Willard L, Takemoto DJ, Lin D (2009) Loss of Purkinje cells in the PKCgamma H101Y transgenic mouse. Biochem Biophys Res Commun 378:524–528 Zhuchenko O, Bailey J, Bonnen P, Ashizawa T, Stockton DW, Amos C et al (1997) Autosomal dominant cerebellar ataxia (SCA6) associated with small polyglutamine expansions in the alpha 1A-voltage-dependent calcium channel. Nat Genet 15:62–69 Zoghbi HY, Orr HT (2000) Glutamine repeats and neurodegeneration. Annu Rev Neurosci 23:217–247 Zoghbi HY, Orr HT (2009) Pathogenic mechanisms of a polyglutamine-mediated neurodegenerative disease, spinocerebellar ataxia type 1. J Biol Chem 284:7425–7429 Zortea M, Armani M, Pastorello E, Nunez GF, Lombardi S, Tonello S et al (2004) Prevalence of inherited ataxias in the province of Padua, Italy. Neuroepidemiology 23:275–280 Zuhlke C, Dalski A, Schwinger E, Finckh U (2005) Spinocerebellar ataxia type 17: report of a family with reduced penetrance of an unstable Gln49 TBP allele, haplotype analysis supporting a founder effect for unstable alleles and comparative analysis of SCA17 genotypes. BMC Med Genet 6:27

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Stefano Di Donato, Daniele Marmolino, and Franco Taroni

Abstract

Mitochondria, double-membrane organelles, are the major site of energy production and vital components of all eukaryotic cells because of the presence in their inner membrane of the respiratory chain (RC) which accomplishes oxidative phosphorylation (OXPHOS). In addition to respiration, mitochondria carry out diverse tasks which include import of proteins, ions, and metabolites, anaplerotic and degradative metabolic reactions, organelle dynamics, and signaling for apoptosis. Despite this multiplicity of functions, the term “mitochondrial disorders” is currently ascribed to OXPHOS diseases. Given the complexity of its biochemistry and of its peculiar dual genetic control, proper OXPHOS is a process that requires the assembly of numerous different proteins coded either by the mitochondrial genome or by the nuclear genome, and the orchestrated function of the five respiratory chain enzyme complexes packaged in the RC, the special hetero-multimeric structure of the inner mitochondrial membrane. Hence, mutations in mitochondrial DNA genes or in nuclear DNA genes encoding integral proteins of the RC subcomplexes, their regulatory and assembly factors, and the set of proteins that complete and regulate cellular

S. Di Donato (*) Fondazione IRCCS Istituto Neurologico C., Besta Via Celoria, 11–20133 Milano, Italy e-mail: [email protected] D. Marmolino Laboratoire de Neurologie expe´rimentale, Universite´ Libre de Bruxeles (ULB), Route de Lennik 808 Campus Erasme - 1070, Bruxelles, Belgium e-mail: [email protected] F. Taroni Fondazione IRCCS Istituto Neurologico C., Besta Via Celoria, 11–20133 Milano, Italy and Department of Diagnostics and Applied Technology, Unit of Genetics of Neurodegenerative and Metabolic Diseases Istituto Neurologico “Carlo Besta”, Via Celoria 11, I-20133 Milan, Italy e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 2269 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_102, # Springer Science+Business Media Dordrecht 2013

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bioenergetics, cause OXPHOS dysfunction and a corresponding variety of genetic diseases with heterogeneous clinical manifestations. This chapter focuses on mitochondrial disorders that express significant pathology in the cerebellum and in its long-tract connections with the peripheral organs, particularly the neuromuscular and osteoarticular systems, and manifest with spinocerebellar ataxia as the prominent symptom or sign.

Introduction to Mitochondrial Functions Mitochondria are ubiquitous in eukaryotic cells and are the site of oxidative phosphorylation (OXPHOS). These cytoplasmic organelles, which display an amazing plasticity in shape appearing as bean-like organelles or extended reticular networks, are thought to have arisen about 1.5–2 billion years ago from a symbiotic association between a glycolytic motile proto-eukaryotic cell and an oxidative alpha-protobacterium (Wallace 2010). Relics of this symbiotic event in the mitochondria of the contemporary eukaryotes are the double-membrane structure, the circular mitochondrial DNA (mtDNA) with its unique genetic code and its specific transcription and translation systems, the presence of transmembrane carriers for ions, metabolites, and proteins, the numerous and diverse degradative and biosynthetic pathways, the existence of proteins enabling organelle dynamics including fusion and fission of mitochondria, the presence of cell-destructive pathways (apoptosis), and the respiratory function (Di Donato 2000). It is hypothesized that, as mitochondrial metabolism became progressively integrated between the glycolytic primordial cell and the engulfed organelles, along billion years of life, the symbiosis maturated through specific interactions to establish the consolidation of the definitive and effective paths of cell metabolism, and their genetic control (Wallace 2010). Among these interactions are the nucleo-mitochondrial communications concerning the assembly of a properly functioning RC, and the critical role of mitochondria in the control of cell death (Poyton and McEwen 1996). This complexity is reflected by the belief that mitochondria contain between 900 and 1,300 different proteins (Chan 2006; Di Mauro and Schon 2008). In the medical literature, the term “mitochondrial disorders” is to a large extent applied to the clinical syndromes associated with abnormalities of the common final pathway of the mitochondrial energy metabolism, i.e., OXPHOS. Faulty OXPHOS may be due to overall dysfunction of the respiratory chain (RC), the hetero-multimeric structure embedded in the inner mitochondrial membrane, or can be associated with single or multiple defects of the five complexes forming the RC. From the genetic standpoint, the RC is a unique structure of the inner membrane formed by means of complementation of two separate genetic systems: the nuclear genome (nDNA) and the mitochondrial genome (mtDNA) (Zeviani and Di Donato 2004). To exemplify the aforementioned complex functions of mitochondria and to give a comprehensive insight into the OXPHOS pathologies described below, this chapter will: (a) briefly summarize the current information on some non-OXPHOS-linked organelle

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autonomous functions relevant to cerebellar pathology; and (b) describe more extensively the structure, molecular genetics, and function of the RC in the completion of OXPHOS.

Transport Systems Mitochondria are virtually impermeable to any substance and they accordingly hold specific import and export systems allowing the traffic of essential molecules such as proteins, ions, and metabolites across the inner and outer membranes (Huizing 1998). 1. Protein Transport. Most proteins imported to mitochondria are synthesized with a cleavable NH2-terminal targeting sequence and are sorted to their correct mitochondrial location by the dynamic interaction of distinct receptor and transport systems in the outer and inner membranes, assisted by folding and unfolding proteins and protein motors (Schatz and Dobberstein 1996). Practically, newly synthesized proteins to be targeted to mitochondria are unfolded by chaperon proteins and directed to integral outer membrane proteins that function as protein import receptors, the outer membrane translocators (TOMs). These receptors recognize chaperon-bound but not chaperon-free proteins. Thereafter, the translocating peptide is moved across the phospholipids’ bilayer of the inner membrane through a hydrophilic hetero-oligomeric ad hoc channel composed of integral proteins of the inner membrane, the translocators of the inner membrane (TIMs). Concomitantly, heat-shock proteins (HSPs), such as HSP70, act as ATP-driven translocation motors that bind and drive the imported polypeptide (Schatz and Dobberstein 1996; Koehler et al. 1998). Other proteins, as the inner membrane mitochondrial AAA metallo-proteases, contribute to generate fully functional imported proteins (Langer 2000). These proteases not only accomplish protein quality control and degrade non-assembled or damaged inner membrane protein, but also mediate maturation of the newly imported proteins; some among these mitochondrial proteases play an important role in maintaining cerebellar integrity (Martinelli et al. 2009; Di Bella et al. 2010). 2. Substrate Carriers. To maintain the electrochemical gradient generated by the electron transfer chain (mETC) the inner membrane is almost impermeable in physiologic conditions. The matrix space contains in fact a highly selected set of small molecules, whereas the intermembrane space is chemically equivalent to the cytosol (Huizing 1998; Palmieri et al. 2000). To function properly, the inner membrane holds a variety of proteins that transport various substrates and products into and out the matrix. This large family of proteins includes the adenine nucleotide translocator(s) (ANT), the uncoupling proteins (UCPs), and carriers for phosphate oxoglutarate, citrate, succinate-fumarate, oxaloacetatesulfate, carnitine, and ornitine (Palmieri et al. 2000; Stepien et al. 1992; Ricquier and Bouillard 2000). ANT is the most abundant integral protein of the inner membrane and catalyzes the transmembrane exchange of extramitochondrial ADP with ATP generated at the matrix side of the inner membrane in the process of OXPHOS. Notably, efficient ANT function contributes to the control of

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respiration (Kaukonen et al. 2000). Mitochondrial metabolism is also controlled by Ca++ uptake, a process that is mediated essentially, but not exclusively, by the calcium uniporter, a channel able to transport Ca++ and Sr++ but not Mg++ (Rizzuto et al. 2000). This uniporter regulates the influx of Ca++ into the matrix according to electrochemical gradient of Ca++. Besides, inter-mitochondrial solute exchange and calcium influx into mitochondria from the endoplasmic reticulum (de Brito and Scorrano 2008) are also dependent on the recently identified profusion proteins which include mitofusin 1 and mitofusin 2, and OPA1 (Chen et al. 2007; de Brito and Scorrano 2008), inner membrane-linked GTPases which are important players in apoptosis and in cerebellar homeostasis (see below) (Frezza et al. 2006).

Degradative Metabolic Pathways The inner mitochondrial membrane and the matrix space hold fundamental functions linked to energy production. These complex functions consist in substratespecific pathways which degrade glucose-derived pyruvate, fatty acid, and amino acid to acetyl CoA. This in turn is the principal substrate of the Krebs’ cycle, a unique final metabolic path which degrades acetyl CoA to CO2 and reducing equivalents (Stryer 1988). Genetic alterations of these paths are associated with human diseases generally presenting in infancy or childhood. Some of these disorders, such as the various forms of PDH deficiency, show cerebellar pathology and ataxia (Di Donato 2000). 1. Pyruvate and alpha-keto amino acid oxidation. The multimeric enzyme pyruvate dehydrogenase complex (PDHC) is responsible for the oxidation of glucosederived pyruvate to acetyl CoA. Other multimeric systems such as the alphaketoglutarate dehydrogenase (KGDH) and the branched-chain ketoacid dehydrogenase (BCKADH) are present in the inner membrane and devoted to the oxidation of amino acids. 2. Fatty acid oxidation. Mitochondrial oxidation of lipids is a complex process that requires a series of enzymatic reactions. In mammals, the mitochondrial carnitinepalmitoyl transferase enzyme system, in conjunction with the carnitine/ acylcarnitine translocase, provides the active carnitine-dependent mechanism whereby long-chain acyl-CoAs are transported from the cytosol into the mitochondrion where beta-oxidation occurs. The final step of each cycle in the beta-oxidation spiral is the release of two molecules of acetyl CoA and a fatty acyl-CoA which is two-carbon atoms shorter. Complete catabolism of longchain acyl-CoAs in mitochondria is accomplished by the action of two distinct, albeit coordinated, beta-oxidation systems (Di Donato and Taroni 2008). 3. The citric acid cycle. The tricarboxylic acid cycle (TCA) or citric acid cycle is the fundamental degradative pathway of the mitochondrial matrix that brings to completion the oxidation of acetyl CoA, the final molecule of the metabolic fate of carbohydrate, fatty acid, and amino acid. The mechanistics of the TCA cycle are based on a complex series of reactions that start from acetyl CoA,

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NAD+, FAD, GDP, Pi, and water to give rise to CO2, NADH, FADH2, GTP, H+, and CoA (Stryer 1988). The citric acid cycle is strictly aerobic, and ATP is generated by the oxidation of NADH and FADH2 in the respiratory chain, respectively two ATP per each FADH2, and three ATP per NADH, as electrons are transferred to O2 (Stryer 1988).

Mitochondrial Dynamics Mitochondria, often depicted in the past as static kidney- or bean-shaped organelles, are dynamic in nature (Chan 2006). As a matter of fact any given mitochondrion is not a discrete organelle because it will fuse or merge, soon or after in its life, with other mitochondria. Mitochondrial fusion not only results in the mixing of outer and inner membrane but also of the matrix content, including the mitochondrial DNA organized in the matrix in discrete units called nucleoids (Wallace 2010; Chan 2006). Fusion and the opposite event of mitochondrial splitting – dubbed fission (Youle and Karbowski 2005), do not simply occur at cell division but are constant features of the cell life, so that the identity of any mitochondrion is transient. The relevance of this concept in mitochondrial genetics is patent as one of its major consequences is that a given mitochondrion within a heteroplasmic cell, i.e., a cell containing more than unique species of mitochondrial DNA, will likely contain both mutant and wild-type DNA (Wallace 2010; Chan 2006). Mitochondrial fusion, coupled with the opposing process of fission, controls the morphology of mitochondria and plays important roles in cell biology. In mammals, three distinct inner membrane GTPases are required for mitochondrial fusion: Mfn1, Mfn2, and OPA1 (Chan 2006, 2007). Mitochondrial fission is principally driven by the cytosolic dynamin-related protein DRP1 that, under activation, binds to the outer membrane protein FIS1 and then drives mitochondrial splitting. Proteins driving fission are crucial actors in mitochondrial-mediated apoptosis (Youle and Karbowski 2005). Neurons have a particular dependence on the precise control of mitochondrial dynamics, and neurodegenerative diseases are associated with mutations in Mfn2 and OPA1 genes (Chan 2007). Noteworthy, a model of cerebellar neurodegeneration in mice was generated by removing Mfn2 from the cerebellum (Chen et al. 2007). During development and after maturity, Purkinje cells of these mice required Mfn2 for dendritic outgrowth, spine formation, and cell survival, and Mtf2-null Purkinje cells showed aberrant mitochondrial distribution, ultrastructure, and electron transport chain activity. This model intriguingly provides a molecular mechanism for the dependence of respiratory activity on mitochondrial fusion (Chen et al. 2007).

Oxidative Phosphorylation and the Respiratory Chain From the genetic standpoint, the mitochondrial RC is unique as it is formed by means of complementation of two separate genetic systems: the nuclear genome

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and the mitochondrial genome. The nuclear genome encodes most of the approximately 88 protein subunits of the respiratory complexes and most of the mtDNA replication and expression systems, whereas the mitochondrial genome encodes 13 RC subunits, 22 mitochondrial-specific tRNAs, and two RNA components of the mitochondrial translational apparatus (Saraste 1999; Di Donato 2009). Remarkably, the mtDNA of multicellular animals all retain roughly the same 13 OXPHOS polypeptide genes (Wallace 2007, 2010) (see below). Hence, mitochondrial disorders due to OXPHOS defects include both Mendelian-inherited and cytoplasmicinherited diseases. These diseases are mostly expressed in high-energy demanding non-mitotic tissues such as heart, skeletal muscle, and the central and peripheral nervous system. The cerebellum, because of its high content of specialized postmitotic neurons with intricate fibers’ connection exemplified by the Purkinje cell, the most complex and integrated neural system in the human species, is especially sensitive to impaired OXPHOS (Chen et al. 2007).

RC Function Most of the cell’s energy is generated by OXPHOS, a process requiring the orchestrated action of five respiratory enzyme complexes packaged in a special structure of the inner mitochondrial membrane, the mitochondrial RC (Saraste 1999). In terms of function, the RC carries out two main reactions which operate in an integrated fashion: the exoergonic transfer of electron equivalents from the reduced electron carriers NADH and FADH2 to molecular oxygen, a process coupled to proton translocation across the inner membrane, and the endoergonic ATP synthesis, driven by the energy primarily stored as an electrochemical proton gradient (Saraste 1999; Di Donato 2000). The first two linked events of respiration, i.e., electron transfer and proton pumping, are carried out by the mETC, a functional supramolecular structure located in the lipid bilayer of the membrane composed by four complexes (complex I–IV). In humans, complex I or NADHubiquinone oxidoreductase, one of the four proton translocating complexes of the RC (complex I, III, IV, and V), accomplishes the oxidation of NADH derived by the oxidation of fatty acids, pyruvate and amino acids, and contains seven subunits encoded by the mtDNA and approximately 38 nuclear-encoded subunits (Smeitink et al. 2006). Complex II or succinate-ubiquinone oxidoreductase, which accomplishes the oxidation of FADH2 derived from the Krebs’ cycle and beta-oxidation of fatty acids and branched-chain amino acids, is composed by four subunits, all encoded by the nuclear genome. Complex III or ubiquinol-ferricytochrome c oxidoreductase holds one subunit, cytochrome b, encoded by the mitochondrial genome and ten subunits encoded by the nuclear genome. Complex IV or cytochrome c oxidase is composed by 13 subunits, three of which are encoded by mtDNA (COX I–III) and ten by nuclear DNA. In addition, mETC contains two highly hydrophobic, mobile, small electron carriers, coenzyme Q 10 and cytochrome c, both synthesized by nuclear genes (Zeviani and Di Donato 2004; Smeitink et al. 2006). Synthesis of ATP from ADP, the second fundamental reaction of OXPHOS, is a process performed by complex V or ATP synthase. ATP synthase is composed by two mtDNA-encoded subunits (ATPase six and

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eight), and 13 nDNA-encoded subunits (Fig. 102.1). As mentioned, the proton electrochemical gradient generated at the mETC level drives the condensation of ADP and inorganic phosphate into ATP. Hence, the fundamental reaction of life, i.e., oxygen activation and the conservation of energy in cell respiration is essentially a function of the integrity of the inner membrane RC (Saraste 1999). The energy that is released as electrons pass through complexes I, III, and IV is used to transport protons out across the mitochondrial inner membrane to generate a membrane electrochemical potential (D-P). Electron transfer and ATP synthesis are coupled, or linked (Wallace 2010; Belevich et al. 2006). In fact, the RC works as a proton pump which generates a proton gradient and a membrane potential of about 180 mV across the inner membrane with a negative polarity at its matrix side. The proton gradient is utilized by the ATP synthase to phosphorylate matrix ADP. During this process, the proton gradient is decreased and this activates respiration, i.e., electron transfer. The competence of the RC, i.e., the effectiveness of ATP generation compared to caloric intake, is called coupling efficiency, and reflects both the efficiency by which complexes I, III, and IV convert the oxidation of reducing equivalents into D-P and the efficiency by which complex V converts D-P into ATP (Wallace 2010). Tightly coupled OXPHOS indicates a coupling efficiency that maximizes ATP generation per calorie intake (Wallace 2010; Belevich et al. 2006). Remarkably, all the proton-translocating complexes of OXPHOS (complexes I, III, IV, and V, see Fig. 102.1) must be balanced to ensure that one complex is not disproportionally permeable to protons and thus shorts D-P. This is achieved by having the core electron and proton transport genes retained on a single piece of non-recombining DNA, the exclusively maternally inherited mtDNA (Wallace 2007, 2010). Since the RC is intrinsically a complex structure, sophisticated molecular and biochemical interactions are required to accomplish maintenance of its integrity and full activity. The following considerations synthesize the complexity of our respiratory machinery: (a) The RC is the only structure in the animal world under the dual genetic control of the chromosomal-endowed nuclear genome and of the organelle-endowed mitochondrial genome. A practical genetic consequence is that RC disorders in humans are associated with both Mendelian inheritance and cytoplasmic maternal inheritance (Zeviani and Di Donato 2004). (b) Gene expression from the two genomes must be strictly regulated and correlated to the variable energy demand of the cell, a phenomenon implying a stringent cross talk between the two genomes to generate the adequate ATP synthesis (Poyton and McEwen 1996). The vehicles for such intergenomic communication are both nuclear-coded proteins that variably signal to and influence the replication, transcription, translation and fission–fusion of the mitochondrial apparatus, and metabolic signals that are released from the mitochondrion and get in touch with the cell nucleus to inform about derangements from physiological rates of respiration (Di Donato 2009). Oxygen tension is an important factor in regulating nuclear responses which include the induction of hypoxia-inducible-factor, a master regulator of oxygen-sensitive

H+

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Fig. 102.1 (continued)

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Homeostatic effects Growth factor signaling Activation of uncoupling proteins mtDNA replication

Toxic effects Lipid peroxidation Protein oxidation mtDNA damage

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gene expression (Piruat and Lopez-Barneo 2005). Also, reactive oxygen species (ROS) might play a regulatory action in some conditions related to respiratory chain dysfunction (Baughman and Mootha 2006). (c) Genetics of mtDNA differ from that of nuclear DNA firstly because mtDNA, endowed in mitochondria in matrix structures dubbed nucleoids, is present in each cell in hundreds or thousands of copies (polyploidy) at difference from nDNA which is present in non-gametic cells with two copies inherited respectively from each parent. Also, mtDNA is maternally inherited and only the mother transmits her oocyte mtDNA to all of her offspring, and her daughters transmit their mtDNA to the next generation (Zeviani and Di Donato 2004; Taylor and Turnbull 2005). Normally, the mitochondrial genotype of an individual is composed of a single mtDNA species, a condition known as homoplasmy. However, the propensity of mtDNA to mutate at high rates can determine a condition known as heteroplasmy, where the wild-type and the mutant genomes coexist intracellularly. The pathogenic outcome of heteroplasmic mutations depends upon a threshold effect that dictates which mutation load in each tissue is compatible with a safe respiratory capacity of the cell and thus the phenotypic expression of a mtDNA-associated character (Zeviani and Di Donato 2004) (see above the concept of heteroplasmy in section “Mitochondrial Dynamics”). The actual transmission of potential deleterious characters associated with mutant heteroplasmic mtDNA is conditioned by the fact that during embryogenesis and tissue differentiation, including the production of primary oocytes, the distribution of mutant and wild-type mtDNA is stochastic. Furthermore in oogenesis only a restricted number of mtDNA molecules are randomly transferred into each oocyte and at oocyte maturation a rapid replication of this mtDNA pool occurs (Taylor and Turnbull 2005; Ankel-Simons and Cummins 1996). Hence, in principle, it cannot be foreseen whether maternal transmission of mutant mtDNA will cause disease, or be relatively safe. In this context of genetic uncertainty, and of putative risk for survival of animal species because of deleterious mtDNA ä Fig. 102.1 Creative drawing of the respiratory chain, major mitochondrial pathways including savage paths for ROS (OH.), and the human mitochondrial DNA. Respiratory chain complexes. Mitochondrially encoded subunits, embedded in the midst of nuclear-encoded subunits, are shown in different colors: complex I subunits ¼ blue; complex III subunit: green; complex IV subunits ¼ red; complex V subunits: yellow. Mitochondrial DNA is drawn as circles inside mitochondria: Its schematic gene content is on the right. myt genes : complex I genes are in blue; complex III cytb gene is in green; complex IV genes are in red; complex V genes are in yellow. syn genes: tRNA genes are in gray. rRNA genes are in purple. ISC: Fe-Sulfur centers present in complex I, II, III of the respiratory chain; OH. Ion : a reactive oxygen species (ROS) toxic to protein, lipid DNA, and the FeS centers (upper side of figure). Abbreviations: ADP adenosine 5-diphosphate, ATP adenosine 5-triphosphate, FAD/FADH2 oxidated/reduced flavin adenine dinucleotide, FeS iron-sulfur centers, GPx glutathione peroxidase, NAD/NADH oxidated/ reduced nicotinamide adenine dinucleotide, OH– hydroxyl ion, Q/QH2 oxidated/reduced ubiquinone, SOD2 mitochondrial superoxide dehydrogenase, TCA tricarboxylic acid cycle (Courtesy of Dr Loredana Lamantea, Division of Molecular Neurogenetics)

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mutations, nature somehow developed protective strategies in the mammalian ovary that, throughout poorly understood mechanisms, systematically eliminate the proto-oocytes that harbor the most severe and harmful mtDNA mutations (Fan 2008; Wallace 2010). (d) OXPHOS is not a perfect device since an estimated 0.2% of the oxygen consumed during respiration is not fully reduced to water but is only partially reduced to reactive oxygen intermediates (superoxide anion, O2., and hydrogen peroxide, H2O2), which can be converted to the highly reactive hydroxyl ion (OH.) (Smeitink et al. 2006). These intermediates, referred to collectively as ROS, are generated at two main sites of the RC, the NADH dehydrogenase (complex I) and the ubiquinol-coenzyme Q oxidoreductatase (complex III), and are toxic to the cell. In normal conditions, ROS can be rescued by the mitochondrial enzymes manganese superoxide dismutase and glutathione peroxidase (Fig. 102.1). There is some evidence that ROS concentrations in sensitive cells might shape the phenotype of mitochondrial diseases due to mutations of mt DNA, and thus contribute to uncover and partially explain the poor correlation between mutation and phenotype in these disorders (MorenoLoshuertos et al. 2006).

Cerebellar Disorders due to Defects of Nuclearly Encoded Mitochondrial Proteins Friedreich Ataxia (FRDA) Friedreich ataxia (FRDA) is the most common hereditary ataxia, accounting for about half of genetic ataxias in white populations (Harding 1983). It is an autosomal-recessive neurodegenerative disorder estimated to affect 4.7:100,000– 1.8:100,000 people of Western European origin with an estimated heterozygous carrier rate of 1:60–1:110 (Delatycki et al. 2000; Schulz et al. 2009) (Table 102.1). This rate seems to decrease in Northern European populations to as low as 1:100,000–0.13:100,000 in Denmark, Sweden, Norway, and Finland (Schulz et al. 2009). The disorder is also found in individuals of North African, Middle Eastern, or Indian origin but is virtually nonexistent in sub-Saharan Africans, Asians, and Amerindians (Labuda et al. 2000; Schulz et al. 2009). The disease is named after Nikolaus Friedreich, Professor of Pathology and Therapy in Heidelberg (Germany) who first reported the condition in nine patients from three families between 1863 and 1877 (Delatycki et al. 2000). He described the essential findings of the disease: degenerative atrophy of the posterior columns of the spinal cord leading to progressive ataxia, sensory loss, and muscle weakness, often associated with scoliosis, foot deformity, and heart disease. Despite Friedreich’s accurate account of the disease, the complete clinical spectrum of FRDA and the features that distinguished this disease from other ataxia syndromes have been controversial for more than a century, until the Que´bec Collaborative Group in 1976 (Geoffroy et al. 1976) and Harding in 1981 (Harding 1981) defined the essential clinical criteria for diagnosis of FRDA.

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Table 102.1 Cerebellar disorders due to defects of nuclearly encoded mitochondrial proteins Disorder Gene Inheritance Clinical presentation Friedreich ataxia FXN Autosomal Onset  25 years (FRDA) recessive (typical, 80% of cases) Adult-onset variants: 25–50 years (20% of cases) Progressive cerebellar ataxia Sensory neuropathy Amyotrophy Pyramidal signs Cardiomyopathy (60% of cases) Diabetes (10–30% of cases) MRI: Atrophy of cervical spinal cord Cerebellar atrophy only in advanced cases MEMSA POLG Autosomal Onset: infancy to adulthood (Myoclonic recessive Progressive cerebellar ataxia epilepsy Sensory neuropathy myopathy sensory Myoclonus ataxia) Progressive external ophthalmoplegia (after 30 years) Epilepsy Episodes of epilepsy/encephalopathy mtDNA deletion/depletion in muscle MRI: Cerebellar atrophy T2 hypersignal in thalamus, cerebellar white matter, inferior olivary nuclei SANDO POLG Autosomal Juvenile-adult onset (Sensory ataxic C10orf2 recessive Sensory ataxia neuropathy Sensory neuropathy dysarthria Dysarthria ophthalmoparesis) Ophthalmoplegia MRI: Cerebellar atrophy Thalamic lesions MIRAS POLG Autosomal Onset: 5–40 years (mostly 30 years) (Mitochondrial recessive Cerebellar ataxia Recessive Ataxia Sensory neuropathy Syndrome) Nystagmus Dysarthria Ophthalmoplegia Tremor Cognitive decline Myoclonus Epilepsy mtDNA deletions in muscle MRI: Cerebellar white-matter changes Cerebellar atrophy (hemispheric or vermian) T2 hypersignal in thalamus, and dentate and inferior olivary nuclei (continued)

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Table 102.1 (continued) Disorder Gene Inheritance Clinical presentation IOSCA C10orf2 Autosomal Onset: 7 years) Sensorineural deafness (>7 years) Epilepsy/encephalopathy mtDNA depletion in muscle MRI: Cerebellar white-matter changes Cerebellar atrophy Olivopontocerebellar atrophy ADCK3 Autosomal Onset in childhood Ataxia with recessive Progressive cerebellar ataxia coenzyme Q10 deficiency Seizures () Developmental delay () Mental retardation () Reduced CoQ10 in muscle MRI: Cerebellar atrophy Responds to coenzyme Q10 (ubiquinone) supplementation XLSA/A ABCB7 X-linked Onset in early childhood (X-linked Nonprogressive cerebellar ataxia sideroblastic Mild sideroblastic anemia and microcytosis anemia with Slow progression in adulthood ataxia) MRI: Cerebellar atrophy SCA28 AFG3L2 Autosomal Juvenile onset (Spinocerebellar dominant Cerebellar ataxia ataxia Ophthalmoparesis type 28) Slow saccades Increased tendon reflexes Pyramidal signs MRI: Cerebellar atrophy

As described by Friedreich, progressive, unremitting ataxia is the principal feature of the disease. Gait instability and generalized clumsiness are typical presenting symptoms. The onset is usually around puberty, but it may vary from 2 to 3 years of age to later than 25 years of age (Pandolfo 2008a). As defined by Harding (Harding 1981), the cardinal clinical features are (1) autosomal-recessive inheritance, (2) onset before 25 years of age, (3) progressive limb and gait ataxia, (4) absence of tendon reflexes in the lower extremities, (5) electrophysiologic evidence of axonal sensory neuropathy followed by (within 5 years of onset), (6) dysarthria, (7) areflexia at all four limbs, (8) distal loss of position and vibration sense, (9) extensor plantar response, and (10) pyramidal weakness of the legs. Patients lose the ability to walk 10–15 years after disease onset and need

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a wheelchair for their daily activities (Harding 1981; D€urr et al. 1996; Delatycki et al. 2000). Although heart failure is the commonest cause of death, cardiomyopathy, along with scoliosis and foot deformity, is not an essential feature (Harding 1981). Patients who exhibit all these clinical features listed by Harding are considered to have the “typical” or “classic” form of the disease. As with age of onset, there is a great variability in other clinical features, including rate of progression, severity, and duration of the disease (Harding 1993; D€urr et al. 1996; Montermini et al. 1997). Patients may be confined to a wheelchair in their early teens or may still be ambulant in their late thirties. Cardiac complications may be severe enough to cause early death or may be minimal or absent. Other variable neurologic findings include oculomotor abnormalities, nystagmus, optic atrophy (approximately 30%) with decreased visual acuity in 13–18% of patients (Delatycki et al. 2000; Fortuna et al. 2009), and sensorineural hearing loss (8–39%) (Delatycki et al. 2000; Alper and Narayanan 2003). The most common abnormality of ocular movement is fixation instability with square-wave jerks (SWJs) (Fahey et al. 2008). Ophthalmoparesis is not observed. More than half of the patients manifest progressive scoliosis, and about half manifest pes cavus, pes equinovarus, and clawing of the toes (Harding 1981). Amyotrophy of the small muscles of the hand and distal muscles of the leg and foot is common (Harding 1981; Alper and Narayanan 2003). Diabetes mellitus is found in 10–30% of patients, and glucose intolerance in 25–40% (Harding 1981; Finocchiaro et al. 1988; Delatycki et al. 2000; Alper and Narayanan 2003; Santos et al. 2010). A combination of insulin resistance and inadequate insulin response contributes to diabetes in FRDA; both forms are likely to be a direct consequence of the mitochondrial dysfunction that occurs in this disease (Pandolfo 2008b). Cardiomyopathy is evident in around two-thirds of patients with FRDA and is primarily symmetric concentric hypertrophic cardiomyopathy, although some patients exhibit asymmetric septal hypertrophy (Harding 1993; Isnard et al. 1997). Electrocardiogram is almost always abnormal with widespread T-wave inversions with or without signs of ventricular hypertrophy, indicating that subclinical heart disease is almost universal in FRDA. After discovery of the FRDA gene in 1996 (Campuzano et al. 1996), direct testing for the FRDA mutation of patients with autosomal-recessive or sporadic ataxia led to the characterization of several “atypical” FRDA variants (Montermini et al. 1997). While the vast majority (93–96%) of patients with typical features of FRDA (Harding 1981) are homozygous for a GAA-repeat expansion in the first intron of the FRDA gene (D€ urr et al. 1996; Montermini et al. 1997), some patients not manifesting “classic” or “typical” FRDA according to Harding criteria also are homozygous for the expanded trinucleotide repeat (D€urr et al. 1996; Montermini et al. 1997). Atypical variants represent approximately 25% of genetically proven FRDA cases (Bidichandani et al. 2000) and include the Acadian type, late-onset FRDA (LOFA), and FRDA with retained reflexes (FARR). The Acadian type is observed in a specific population of French origin (Cajuns) living in North America (Louisiana) and is distinguished from typical FRDA by its milder course and lower incidence of cardiomyopathy (Montermini et al. 1997).

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Age of onset older than 20 years can be found in 15–20% of patients with a FRDA phenotype (De Michele et al. 1994). LOFA cases have all the features of typical FRDA, but disease onset is after 25 years of age (De Michele et al. 1994; D€urr et al. 1996; Montermini et al. 1997; Gellera et al. 1997). Patients with delayed onset, arbitrarily subdivided into late-onset FRDA (LOFA; 25–39 years) and verylate-onset FRDA (VLOFA; 40 years up to 60 years) (Bidichandani et al. 2000), usually exhibit mild clinical impairment, slower progression of disease, and fewer secondary complications or associated manifestations. Disease progression, as indicated by years from onset to becoming confined to wheelchair, is slower in patients with late-onset FRDA. As compared with patients with typical FRDA, LOFA and VLOFA patients show an increased occurrence of lower limb spasticity and retained reflexes, decreased skeletal abnormalities, and no evidence of cardiomyopathy on echocardiograms (Bhidayasiri et al. 2005). FARR is a variant in which patients fulfill the Harding’s diagnostic criteria (Harding 1981), except that tendon reflexes are preserved in lower limbs (Palau et al. 1995; Klockgether et al. 1996). Sensory axonal neuropathy may or may not be evident on neurophysiologic tests. The FRDA clinical features are generally present but less pronounced. FARR patients may also exhibit electrocardiogram abnormalities and echocardiographic evidence of hypertrophic cardiomyopathy. There may be coexistence of typical FRDA phenotype in some families (Palau et al. 1995; Klockgether et al. 1996). Neurophysiological investigations are useful, albeit nonessential in the genetic era, for diagnosis. Nerve conduction studies characteristically show an early and severe peripheral sensory axonal neuronopathy with small or absent sensory action potentials, slightly decreased sensory conduction velocities, and absent spinal somatosensory evoked potentials (Harding 1993; Zouari et al. 1998). Motor conduction velocities are normal or mildly reduced (Delatycki et al. 2000). Magnetic resonance imaging (MRI) reveals atrophy of the cervical spinal cord and signal abnormalities in the posterior and lateral columns, with essentially normal brainstem, cerebellum, and cerebrum (Wullner et al. 1993; Mascalchi et al. 1994). In contrast to other hereditary ataxias, progressive cerebellar degeneration is not a feature of FRDA and usually occurs in more advanced cases. Albeit rare, mild cerebellar vermian atrophy may be observed in late-onset presentations after age 25 years (Bhidayasiri et al. 2005). In general, however, brain MRI can be useful in the diagnostic procedure, because the presence of cerebellar atrophy might point to forms of hereditary recessive ataxia other than Friedreich ataxia. FRDA patients should be referred to a cardiologist on an annual basis. Cardiac involvement with left ventricular hypertrophy but normal ejection fraction can be documented by electrocardiography (ECG) and/or echocardiography in approximately two-thirds of patients (Casazza and Morpurgo 1996; Kipps et al. 2009). ECG shows widespread T-wave inversion in virtually all patients, ventricular hypertrophy in most patients, conduction disturbances in about 10% of patients, and supraventricular ectopic beats and atrial fibrillation in occasional cases (Schulz et al. 2009).

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Neuropathological changes are also specific for FRDA and differentiate this disorder from other hereditary ataxias (Lamarche et al. 1984). Loss of large primary neurons in the dorsal root ganglia is a prominent and early finding and represents the hallmark of this disease. As the disease progresses the posterior columns of the spinal cord degenerate. This is associated with atrophy of the spinocerebellar tracts, the corticospinal motor tracts of the spinal cord, and the large sensory fibers in peripheral nerves (Lamarche et al. 1984; Koeppen 1998). Clarke’s column is atrophied, but motor neurons of the anterior horn are unaffected. In the cerebellum, the cortex is spared until late in the course of the disease, when Purkinje cell loss can be observed, but the dentate nuclei are severely atrophied (Lamarche et al. 1984; Koeppen 1998). Altogether, neuropathological abnormalities indicate that FRDA is a disease of the sensory systems that provide afferent information from the periphery to the brain and cerebellum for appropriate (efferent) movement control. Because FRDA is common, the diagnosis is often based on the “classic” triad of progressive cerebellar dysfunction, hypoactive knee and ankle jerks, and adolescent or preadolescent onset. However, since genetic testing has become widespread available, Friedreich ataxia should be considered in all patients with sporadic or recessive ataxia, with the exception of those with severe olivopontocerebellar atrophy on neuroimaging (Schols et al. 1997). FRDA is caused by a large GAA-trinucleotide repeat expansion in the first intron of the FXN gene on chromosome 9q13–21 (Campuzano et al. 1996; Pandolfo 2006). This distinctive and unique mutation is found in 98% of patient chromosomes. The gene encodes a 210-amino acid protein termed “frataxin”. Normal individuals have a repeat size of 6–36 triplets, whereas FRDA patients carry expanded alleles with 70–1,700 repeats, most commonly 600–900 GAA (Schmucker and Puccio 2010). More than 95% of FRDA patients carry the expansion on both alleles. Expansion size is inversely correlated with age at onset and confinement to a wheelchair, and directly correlated with the incidence of cardiomyopathy (D€ urr et al. 1996). A small proportion of patients (4%) are compound heterozygotes harboring point mutations or microdeletions on one allele and a GAA expansion on the other allele (Gellera et al. 2007). More than 40 different FXN micromutations (missense, nonsense, frameshift, splice-site, and one 2.8-kb deletion) have been reported so far (Santos et al. 2010). In the large majority of cases, they are private mutations but some (those affecting the ATG codon, G130V, I154F, R165C, and W173G) have been identified in more than one family. Micromutations are usually associated with typical FRDA phenotype but atypical features are reported for some of them (Santos et al. 2010). For example, the relatively frequent G130V is associated with a milder clinical course (Bidichandani et al. 1997). So far, no FRDA patient homozygous for a point mutation has been identified, suggesting that some functional frataxin protein is necessary for normal development and survival (Cosse´e et al. 2000; Puccio 2009) and that the GAA expansion might be a relatively mild mutation as it allows the synthesis of some frataxin that is structurally and functionally normal because the coding sequence is not involved in the mutation. It cannot be excluded that the absence of patients

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homozygous for point mutations is due to their rarity or to an atypical phenotype not recognized as FRDA. The GAA-repeat expansion inhibits transcription of the gene (Ohshima et al. 1998) whereas point mutations adversely affect protein expression, function, and/or stability (Cavadini et al. 2000; Gellera et al. 2007). DNA containing expanded GAA repeats can adopt a triple helical structure (“sticky DNA”) that directly interferes with transcriptional elongation (Wells 2008). Alternatively, GAA-repeat expansion may mediate gene silencing through epigenetic changes leading to heterochromatin formation, a mechanism that may have relevant implications for therapy (Gottesfeld 2007). Whichever the pathomechanism, cell cultures and tissues derived from patients have decreased amounts of both frataxin mRNA and protein. The degree of reduction correlated with the size of the GAA repeat (Campuzano et al. 1997; Gellera et al. 2007). In patients homozygous for the expansion, the size of the smaller allele is more closely inversely correlated with both age at onset and the severity of the clinical phenotype (D€urr et al. 1996). This is consistent with greater residual transcriptional competence of the allele with fewer GAA repeats (Bidichandani et al. 1998). So this allele is primarily responsible for the residual amount of normal frataxin and has a major role in modulating disease severity (Gellera et al. 2007). Frataxin is a mitochondrial protein conserved through evolution (Cho et al. 2000; Pandolfo and Pastore 2009), the precise function of which is still not fully understood. Many studies have consistently indicated that it participates in multiple iron-dependent mitochondrial pathways (Pandolfo and Pastore 2009; Schmucker and Puccio 2010). The most commonly accepted pathophysiological consequences of frataxin deficiency are a severe disruption of iron-sulfur (Fe-S) cluster (ISC) biosynthesis (Martelli et al. 2007), mitochondrial iron overload coupled to cellular iron dysregulation (Babcock et al. 1997; Puccio et al. 2001), and an increased sensitivity to oxidative stress (Babcock et al. 1997; Wong et al. 1999). The tight link between these cellular pathways makes it difficult to identify the primary cause of the pathogenic process. The first evidence of a link between frataxin and iron metabolism was the identification of a mutant yeast (S. cerevisiae) strain with increased intracellular iron and a deletion of the frataxin homologue Yfh1p (Babcock et al. 1997). This condition was reminiscent of previous observations of increased iron content in FRDA patient hearts (Lamarche et al. 1980). In the mutant yeast, the tenfold increase in intramitochondrial iron content was accompanied by OXPHOS deficiency and increased sensitivity to oxidants. Early biochemical investigations in heart biopsies of FRDA patients demonstrated deficiencies of the ISC-containing subunits of the mitochondrial respiratory chain complexes I (CI), II (CII), and III (CIII) and of ISC-containing proteins aconitases (R€otig et al. 1997). These findings were confirmed in the Yfh1-deleted yeast (Duby et al. 2000). An “iron toxicity” model was initially proposed and these ISC enzyme defects were thought to be a consequence of oxidative stress induced via the Fenton reaction by mitochondrial iron accumulation. However, data obtained in animal models indicate that mitochondrial iron accumulation is a distal consequence of an earlier, proximal effect of frataxin deficiency. Complete deletion of the frataxin

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gene in a knockout mouse model leads to early embryonic lethality with no evidence of abnormal iron storage in embryo cells, indicating an important role for frataxin during development that seems to be independent of the accumulation of iron in mitochondria (Cosse´e et al. 2000). Further studies using conditional mouse models with tissue-specific frataxin deletions demonstrated that the primary deficit in the disease is the ISC protein deficiency followed by secondary mitochondrial iron accumulation, with no overt sign of oxidative stress damage (Puccio et al. 2001; Seznec et al. 2005). A large body of evidence now indicates a major role for frataxin in the biogenesis of cellular ISC (Schmucker and Puccio 2010). Frataxin has been shown to directly interact with the main Fe-S biosynthetic complex, ISCU/NFS1/ISD11 (Schmucker et al. 2011; Tsai and Barondeau 2010; Wang and Craig 2008) and frataxin deficiency is now widely thought to cause primary deficits of mitochondrial and extramitochondrial ISC-containing proteins in human, mouse, and yeast (Martelli et al. 2007, M€uhlenhoff et al. 2002, Schmucker and Puccio 2010; Stehling et al. 2004). Although a role for oxidative stress in the pathogenesis of FRDA was proposed very early (Wong et al. 1999) and numerous studies have demonstrated an impaired response of antioxidant enzymes in cell lines and model organisms (Chantrel-Groussard et al. 2001; Llorens et al. 2007), to date, there is no conclusive evidence that increased oxidative stress has a major pathogenic role in frataxin-deficient cells or tissues (Schmucker and Puccio 2010; Seznec et al. 2005). Several therapeutic strategies have been or are being developed to counteract the effects of frataxin deficiency. Some pharmacological compounds such as antioxidants or iron chelators have shown potential efficacy in improving some of the symptoms of the disease and are currently in clinical testing. However, despite the efforts, the best care for patients with Friedreich ataxia has not yet been defined according to evidence-based criteria (Schulz et al. 2009). Mitochondrial dysfunction has been addressed in several open-label, non-placebo-controlled trials, and randomized placebo-controlled trials that employed idebenone (Pandolfo 2008b; Schulz et al. 2009), a synthetic analogue of coenzyme Q10 (ubiquinone) with potent antioxidant activity (Gillis et al. 1994). Since idebenone also stimulates OXPHOS and ATP production by facilitating the flux of electrons along the mitochondrial respiratory chain, it may contribute to ameliorate the electron transport impairment due to ISC protein deficiency occurring in patients with FRDA. Idebenone was found to decrease cardiac hypertrophy and improve cardiac function in mouse models (Seznec et al. 2004), with beneficial effects observed in patients at dose levels of 5 mg/kg per day (Mariotti et al. 2003). A 6-month, double-blind, placebocontrolled phase-II clinical trial (NICOSIA) using higher doses (5 mg, 15 mg, and 45 mg/kg per day) of idebenone showed evidence of dose-dependent improvement in secondary neurological end points compared with placebo (Di Prospero et al. 2007). By contrast, a more recent phase-III double-blind, randomized, placebocontrolled intervention trial (IONIA) did not provide evidence for efficacy of idebenone (10–20 mg/kg per day) in the treatment of ataxia in ambulatory FRDA pediatric patients (Lynch et al. 2010). To date, there is no evidence of a significant benefit of idebenone on neurological symptoms associated with FRDA.

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Furthermore, also the clinical relevance of the positive effect on left ventricular mass still has to be assessed (Kearney et al. 2009). However, since idebenone and other drugs that improve mitochondrial function (e.g., EPI-A0001) or increase antioxidant defense levels (e.g., pioglitazone, MitoQ) still offer promise for FRDA treatment, larger studies of longer duration may be needed to ultimately assess their neurological efficacy. Other current experimental approaches address iron-mediated toxicity. Deferiprone, an orally active, blood-brain barrier permeable iron chelator can potentially redistribute iron from the mitochondria to other cellular compartments and to blood transferrin (Sohn et al. 2008). In a preliminary open-label study in a small group of FRDA patients, deferiprone (20–30 mg/kg per day) showed some improvement of neurological symptoms (Boddaert et al. 2007). A large multicenter phase-II double-blind, randomized, placebo-controlled trial (sponsored by ApoPharma, Canada) is currently under way. Finally, drugs increasing the amount of frataxin are excellent candidates for a rational approach to FRDA therapy (Marmolino and Acquaviva 2009). Several compounds have been assessed for their ability to increase frataxin cellular levels, including erythropoietin (Boesch et al. 2008), histone deacetylase (HDAC) inhibitors (Gottesfeld 2007), and PPAR-g agonists (Marmolino et al. 2009). HDAC inhibitors are compounds that may reverse gene silencing induced by GAA expansion by facilitating chromatin opening. Indeed, they have shown promise in increasing frataxin levels and reducing neuronal pathology in dorsal root ganglia in FRDA mouse models (Sandi et al. 2011) and are therefore expected to enter clinical tests soon.

POLG-Related Ataxias DNA-polymerase g (pol g) is the enzyme that replicates and repairs mtDNA (Kaguni 2004). Pol g is a heterotrimer composed of one catalytic subunit (pol gA, encoded by POLG), containing the polymerase and exonuclease activities, and two accessory subunits (pol gB, encoded by POLG2) thought to be important for processivity (Yakubovskaya et al. 2006). Since the first report in 2001 (Van Goethem et al. 2001), over 120 pathogenic mutations have been described in the gene encoding the catalytic pol gA subunit (POLG) [Human DNA Polymerase Gamma Mutation Database (NIEHS Mitochondrial DNA Replication Group): http://tools.niehs.nih.gov/polg/] and these are associated with a wide spectrum of mtDNA depletion or deletion neurological syndromes ranging from adult-onset myopathies to severe infantile encephalopathies. Six major phenotypes are currently associated with POLG mutations (Wong et al. 2008; Cohen et al. 2010): (1) autosomal dominant progressive external ophthalmoplegia (adPEO: generalized myopathy and often variable degrees of sensorineural hearing loss, axonal neuropathy, ataxia, depression, parkinsonism, hypogonadism, and cataracts) (Van Goethem et al. 2001; Lamantea et al. 2002), (2) autosomal recessive progressive external ophthalmoplegia (arPEO: progressive

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weakness of the extraocular eye muscles resulting in ptosis and ophthalmoparesis without associated systemic involvement) (Lamantea et al. 2002), (3) Alpers– Huttenlocher syndrome (AHS: childhood-onset progressive severe encephalopathy with intractable epilepsy and hepatic failure) (Naviaux et al. 2004), (4) childhood myocerebrohepatopathy spectrum (MCHS: childhood onset, developmental delay or dementia, lactic acidosis, and myopathy with failure to thrive) (Wong et al. 2008), and two autosomal-recessive ataxic syndromes: (5) myoclonic epilepsy myopathy sensory ataxia (MEMSA), and (6) ataxia neuropathy spectrum (ANS) disorders.

Myoclonic Epilepsy Myopathy Sensory Ataxia (MEMSA) Previously referred to as spinocerebellar ataxia with epilepsy (SCAE) (Tzoulis and Bindoff 2009; Tzoulis et al. 2006), MEMSA now describes the spectrum of severe disorders with ataxia, epilepsy, and myopathy, with or without ophthalmoplegia (Cohen et al. 2010; Van Goethem et al. 2003b) (Table 102.1). Onset may vary between infancy and adulthood. Ataxia is universally present and results from a combined cerebellar and peripheral sensory dysfunction, producing a clinical picture with nystagmus, scanning dysarthria, midline and appendicular ataxia. The vast majority of patients (98%) also develop features of a peripheral neuropathy with diminished tendon reflexes and distal sensory impairment. Epilepsy develops in later years in >60% of patients, often beginning focally in the right arm and then spreading to become generalized. The myopathy may be distal or proximal, and, as in the other POLG spectrum disorders, it may also present as exercise intolerance. It may occur with or without ragged-red fibers (Van Goethem et al. 2003b). The disease should be suspected in patients with juvenile-onset, progressive spinocerebellar ataxia and sensory neuropathy, myoclonus, late (30 years) development of PEO, epilepsy (epilepsia partialis continua in one side of the body, visual symptoms, generalized tonic–clonic seizures, status epilepticus), episodes with epilepsy and progressive encephalopathy. MRI shows cerebellar cortical atrophy, dentate atrophy, high T2 signal focal lesions in thalamus, cerebellar white matter, and inferior olivary nuclei. Patients may develop acute, focal, T2 hyperintense cortical lesions, mostly occipital, but also frontal or parietal. Lesions evolve mirroring epilepsy/encephalopathy episode severity and are associated with bad prognosis (Tzoulis and Bindoff 2009; Tzoulis et al. 2006). MEMSA is inherited as an autosomal-recessive trait and is most commonly associated with A467T and W748S mutations in the linker region of pol gA. These mutations, which are relatively rare in non-Scandinavian populations (Craig et al. 2007), ultimately lead to secondary damage of the mtDNA in the form of point mutations, multiple deletions, and quantitative depletion (Zsurka et al. 2008). Survival is worse in patients carrying the A467T and W748S mutations (compound heterozygous) and best in A467T homozygotes (Tzoulis et al. 2006). The cornerstone of therapy is antiepileptic treatment (Engelsen et al. 2008). However, the seizures are often refractory to conventional therapy, including

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anesthesia. Sodium valproate is strongly contraindicated in POLG disorders or suspected POLG disorders due to a significant risk for development of severe liver failure (Stewart et al. 2010; Tzoulis et al. 2006).

Ataxia Neuropathy Spectrum (ANS) Ataxia neuropathy spectrum includes an overlapping clinical spectrum of disorders organized around ataxia and neuropathy in the absence of significant muscle weakness or myopathy (Hakonen et al. 2005; Milone et al. 2008; Winterthun et al. 2005; Wong et al. 2008). ANS embraces the phenotypes also referred to as mitochondrial recessive ataxia syndrome (MIRAS) (Hakonen et al. 2005) and sensory ataxia neuropathy dysarthria and ophthalmoplegia (SANDO) (Milone et al. 2008), and may partly overlap with the SCAE/MEMSA phenotype (Wong et al. 2008). Since POLG patients have multiple overlapping manifestations (ataxia is usually a combination of central and peripheral disease and epilepsy is often present), it may ultimately be difficult to classify patients into specific syndromes (MIRAS, SANDO, or SCAE). SANDO is a clinical entity characterized by the triad of sensory ataxic neuropathy, dysarthria, and ophthalmoparesis (Fadic et al. 1997) (Table 102.1). It is a juvenile-adult onset disease most prominent and presenting feature is sensory ataxia. In spite of the peripheral nature of ataxia, MRI may show mild atrophy of the cerebellum and medulla or thalamic lesions (Milone et al. 2008; Van Goethem et al. 2003a). Ragged-red fibers and mtDNA deletion or depletion are not mandatory features (Milone et al. 2008; Van Goethem et al. 2003a). POLG mutations have been found to be scattered throughout the entire coding region. However, the disease is genetically heterogeneous with at least another gene, C10orf2 (previously PEO1/TWINKLE), involved (Hudson et al. 2005). MIRAS is the most frequent recessive ataxia in Finland, with a carrier frequency of 1:125, and has a high prevalence in Norway (Hakonen et al. 2005). Clinical manifestations, which may start between 5 and 41 years of age, are characterized by cerebellar ataxia, nystagmus, dysarthria, ophthalmoplegia, tremor, cognitive decline, and myoclonus (Hakonen et al. 2005; Van Goethem et al. 2004; Winterthun et al. 2005) (Table 102.1). Loss of vibratory and position perception is commonly seen. The onset of the disease is most often at 30 years of age, with sensory axonal polyneuropathy or balance disturbances, followed by progressive cerebellar ataxia. Unlike SANDO, epilepsy is a frequent manifestation in MIRAS, being the presenting symptom in many cases, with both partial and generalized seizures, sometimes becoming refractory to antiepileptic drugs and evolving to status epilepticus. As previously mentioned for MEMSA, sodium valproate is strongly contraindicated for its hepatotoxicity (Stewart et al. 2010). Brain MRI shows bilateral white-matter changes in the cerebellar hemispheres, mild hemispheric or vermian cerebellar atrophy, and T2-weighted hypersignal on thalamus, and dentate and inferior olivary nuclei (Hakonen et al. 2005; Van Goethem et al. 2004;

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Winterthun et al. 2005). Nerve conduction studies also demonstrate axonal, less often demyelinating, sensory neuropathy. Elevated protein might be detected in CSF. Muscle biopsy is not diagnostic, but multiple mtDNA deletions may be detected in muscle (Winterthun et al. 2005) and brain (Hakonen et al. 2008). The p.A467T and p.W748S POLG mutations are responsible for most cases of this disorder (Hakonen et al. 2005; Tzoulis et al. 2006), with all Finnish patients being homozygous for the p.W748S mutation associated in cis-position with the p.E1143G polymorphism (Hakonen et al. 2005). There is no genotype–phenotype correlation. The two mutations are very rare in non-Scandinavian patients with adult-onset ataxia (Craig et al. 2007).

Infantile-Onset Spinocerebellar Ataxia (IOSCA) Infantile-onset spinocerebellar ataxia (IOSCA) is an autosomal-recessive severe disorder currently identified only in Finland (Table 102.1). It is characterized by normal development until age 9–18 months, followed by onset of ataxia, muscle hypotonia, loss of deep-tendon reflexes, and athetosis (Lonnqvist et al. 1998; Nikali et al. 2005). Ophthalmoplegia and sensorineural deafness develop by age 7 years. By adolescence affected individuals are profoundly deaf and no longer ambulatory; sensory axonal neuropathy, optic atrophy, autonomic nervous system dysfunction, and hypergonadotrophic hypogonadism in females become evident. Epilepsy can develop into a serious and often fatal encephalopathy with myoclonic jerks or focal clonic seizures that progress to epilepsia partialis continua followed by status epilepticus with loss of consciousness (Lonnqvist et al. 2009). Conventional antiepileptic drugs (phenytoin and phenobarbital) are ineffective in most patients. As for POLG mutations, sodium valproate may cause severe hepatotoxicity (Lonnqvist et al. 2009). Nerve conduction studies and nerve biopsy demonstrate a severe, mostly sensory, axonal neuropathy. Sensory ganglia are more severely affected than motor neurons (Lonnqvist et al. 1998). Neuroimaging studies at early stages of disease demonstrate reduced size of cerebellar hemispheres which progresses to more widespread olivopontocerebellar atrophy (Nikali et al. 2005) with corresponding atrophic changes seen on pathology (Lonnqvist et al. 1998). Muscle biopsy is not diagnostic but mtDNA depletion may be seen in this tissue as well as in brain and liver. Unlike MIRAS, multiple mtDNA deletions are not observed (Hakonen et al. 2008). IOSCA is caused by mutation in the C10orf2 gene (previously PEO1/TWINKLE), which encodes twinkle, a specific mitochondrial helicase involved in DNA replication, and one of its smaller isoform, twinky, whose function is currently unknown (Nikali et al. 2005). The same founder mutation (p.Y508C) is detected in almost all Finnish patients with IOSCA. Similar to POLG, C10orf2 mutations are also associated with different phenotypes, such as autosomal-recessive Alpers-like early-onset encephalopathy with mtDNA depletion (Hakonen et al. 2007) and autosomal dominant progressive external ophthalmoplegia (Spelbrink et al. 2001).

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Ataxia with Coenzyme Q10 Deficiency Coenzyme Q10 (CoQ10, also known as ubiquinone) is an essential lipid-soluble electron carrier in the mitochondrial respiratory chain – it carries electrons from complex I and complex II to complex III – and an important antioxidant (Quinzii et al. 2008). Deficiency of CoQ10 is a clinically and genetically heterogeneous spectrum of autosomal-recessive disorders, generally responsive to CoQ10 supplementation, which include multisystemic manifestations as well as pure CNS involvement (Montero et al. 2007; Quinzii and Hirano 2010) (Table 102.1). Four major clinical phenotypes have been recognized: (1) encephalomyopathy characterized by a triad of recurrent myoglobinuria, brain involvement, and ragged-red fibers; (2) early-infantile multisystemic disease typically with prominent nephropathy and encephalopathy; (3) pure myopathy; (4) cerebellar ataxia with marked cerebellar atrophy. The ataxic phenotype is the most common presentation of CoQ10 deficiency (Montero et al. 2007). It is characterized by childhood-onset progressive cerebellar ataxia, cerebellar atrophy, and markedly decreased levels of CoQ10 in muscle and fibroblasts (Lamperti et al. 2003; Lagier-Tourenne et al. 2008). Associated symptoms include seizures, developmental delay, mental retardation, and pyramidal signs (Lamperti et al. 2003; Montero et al. 2007; Mollet et al. 2008). Some patients may present exercise intolerance and elevated serum lactate (Lagier-Tourenne et al. 2008; Mollet et al. 2008). The adult-onset form of ataxia and CoQ10 deficiency is usually associated with hypergonadotrophic hypogonadism (Montero et al. 2007). Diagnosis is based on reduced amount of CoQ10 in muscle, as plasma CoQ10 levels are usually normal (Montero et al. 2007). Muscle histopathology is essentially normal and brain MRI discloses global cerebellar atrophy. Some patients present an excellent response to CoQ10 while others show only a partial improvement of some symptoms and signs or no improvement at all. In a recent study, patients with ataxia and CoQ10 deficiency responded to ubiquinone supplementation (30 mg/kg per day) with a statistically significant reduction of the ataxia rating scale (ICARS) score after 2 years of treatment (Pineda et al. 2010). Overall, CoQ10 deficiency is the mitochondrial encephalomyopathy with the best clinical response to CoQ10 supplementation, highlighting the importance of an early identification of this disorder (Montero et al. 2007). Five different nuclear genes involved in CoQ10 biosynthesis have been associated with primary CoQ10 deficiency (Quinzii and Hirano 2010). Mutations in COQ2, PDSS1, PDSS2, and COQ9 have been identified in patients with the infantile multisystemic phenotypes. To date, ADCK3 (previously CABC1) is the only gene known to cause the cerebellar ataxic phenotype (Lagier-Tourenne et al. 2008; Mollet et al. 2008). Patients with ADCK3 mutations may be distinguished from other recessive ataxias by the presence of cerebellar atrophy with history of exercise intolerance in childhood and elevated serum lactate at rest or after moderate exercise. Some patients presented seizures (epilepsia partialis continua) and did not respond to ubiquinone supplementation (Mollet et al. 2008).

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X-Linked Sideroblastic Anemia with Ataxia (XLSA/A) Sideroblastic anemia with ataxia is a rare X-linked recessive disorder characterized by nonprogressive ataxia that is apparent during early childhood, mild sideroblastic anemia with hypochromia and microcytosis not requiring transfusion, elevated free erythrocyte protoporphyrin levels, and lack of excessive parenchymal iron deposition (Allikmets et al. 1999; Hellier et al. 2001) (Table 102.1). Some heterozygous females may express mild hematologic manifestations, but none exhibits ataxia. Neurological features, that include motor delay and dysarthria, are nonprogressive until adulthood when slow progression becomes evident (Hellier et al. 2001). MRI shows severe cerebellar atrophy. XLSA/A is due to mutations in the mitochondrial ATP-binding cassette transporter ABCB7 gene on chromosome Xq13 (Allikmets et al. 1999). Unlike the more common X-linked sideroblastic anemia (XLSA) caused by mutations in the ALAS2 gene, XLSA/A does not respond to pyridoxine supplementation (Bekri et al. 2000).

Autosomal Dominant Spinocerebellar Ataxia Type 28 (SCA28) Spinocerebellar ataxia type 28 (SCA28) is one of the most recently characterized forms of autosomal dominant spinocerebellar ataxias (SCAs) and the only autosomal dominant ataxia caused by mutations in a nuclear gene encoding a mitochondrial protein (Di Bella et al. 2010; Mariotti et al. 2008) (Table 102.1). The SCA28 neurological phenotype is characterized by juvenile-adult onset (range: 1–79 years of age), slow disease progression, eye movement abnormalities, and, in some cases, pyramidal signs (Cagnoli et al. 2010; Mariotti et al. 2008). The disease course is slowly progressive with most of the patients remaining ambulant in their late sixties. The first symptoms are unbalance in standing, mild gait ataxia, and limb incoordination. Eye movement abnormalities are frequently observed. In patients with short disease duration, persistent gaze-evoked nystagmus is the prevalent finding, while in patients with more than 20 years of disease duration, slow saccades, ophthalmoparesis, and ptosis are predominantly observed (Mariotti et al. 2008). Other frequent neurological signs are the presence of increased tendon reflexes and muscle tone, without overt spasticity, in lower limbs. Some patients exhibit focal dystonia, parkinsonism, or cognitive impairment (Cagnoli et al. 2010). Nerve conduction studies, visual, auditory, somatosensory, and motor-evoked potentials are normal. Brain MRI shows cerebellar atrophy particularly evident in the superior vermis. Muscle histopathology is normal with no ragged-red fibers, no mtDNA deletions, and normal respiratory chain activity (Mariotti et al. 2008). The disease is caused by heterozygous missense mutations in the gene encoding the inner mitochondrial membrane metalloprotease AFG3L2 (ATPase family gene three-like two) (Di Bella et al. 2010). AFG3L2 is highly and selectively expressed in human cerebellar Purkinje cells (Di Bella et al. 2010). Along with paraplegin, a cognate mitochondrial protease the loss of which causes the recessively inherited

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form of hereditary spastic paraplegia SPG7 (Casari et al. 1998), AFG3L2 forms an evolutionarily conserved protein complex (m-AAA) in the inner mitochondrial membrane with ATPase and metalloprotease activities that function in the maintenance of the mitochondrial proteome and the biogenesis of respiratory chain complexes (Tatsuta and Langer 2008). The vast majority of mutations are located in the highly conserved protease domain of the protein (Cagnoli et al. 2010; Di Bella et al. 2010; Edener et al. 2010).

Cerebellar Disorders Due to Defects of MtDNA Heteroplasmic Point Mutations Myoclonic Epilepsy with Ragged-Red Fibers (MERRF) Myoclonic epilepsy with ragged-red fibers (MERRF) is typically maternally inherited. Intra-familial variability has been described in terms of age of onset and severity of the deficit (Chinnery et al. 1997). Clinically the disorder is characterized by myoclonic seizures, epilepsy, cerebellar ataxia, and myopathy with ragged-red fibers, behavioral and cognitive deficits, hearing loss, short stature, and optic atrophy (Shoffner et al. 1990). Multiple symmetric lipomatosis (Morbus Madelung) generally anticipate onset of neurological symptoms. Patients may also exhibit cardiomyopathy, oculomotor deficit with pigmentary retinopathy, and diabetes mellitus. EEG analysis in patients reveals abnormal background activity which progresses slowly as the disease evolves. Generalized spike-and-wave discharges at 3–5 Hz are common. Focal epileptiform discharges may occur (So et al. 1989). Ragged-red fibers are present in the muscle of almost all MEERF patients. Cerebellar ataxia is a common feature of MERRF syndrome and may be even the presenting manifestation in some patients (Table 102.2–102.3). In a study, patients with MERRF carrying the A8344G mutation presented slow progressive cerebellar Table 102.2 Mitochondrial diseases associated with ataxia Syndrome mtDNA genes mutations associated with frequent ataxia MERRF MILS KSS NARP mtDNA genes mutations associated with infrequent ataxia MELAS LS HAM

Type of ataxia Cerebellar ataxia Cerebellar ataxia Cerebellar ataxia Sensory ataxia Cerebellar ataxia Cerebellar ataxia Sensory mixed ataxia

MERRF myoclonic epilepsy and ragged-red fibers, MILS maternally inherited Leigh syndrome, KSS Kearns–Sayre syndrome, NARP neurogenic muscle weakness, ataxia, and retinitis pigmentosa, MELAS mitochondrial encephalomyopathy, lactacidosis, stroke-like episodes, LS Leigh syndrome

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Table 102.3 Main ataxias associated with a mitochondrial DNA mutation (Adapted from Manto 2010) Disorder Phenotype Neuropathology Kearns–Sayre syndrome Onset < 20 Neuronal loss and gliosis (KSS) Progressive external in basal ganglia ophtalmoplegia Spongy degeneration of Pigmentary retinophathy white matter Progressive cerebellar syndrome Decreased expression of Poor growth mtDNA-encoded proteins Hearth block in neurons of the dentate Increased protein levels in CSF nuclei Mitochondrial Stroke-like episodes (particular Infarct-like lesions encephalomyopathy, lactic lesions in the parieto-occipital Gliosis acidosis, and stroke-like lobes) Demyelination episodes (MELAS) Lactic acidosis Spheroids Ragged-red fibers Degeneration o the Dementia posterior columns and Headache spinocerebellar tract Seizures Accumulation of abnormal Deafness mitochondria in smooth Ataxia muscles and endothelium Vomiting of blood vessels Myoclonic epilepsy with Myoclonus Neuronal loss and gliosis ragged-red fibers (MERRF) Epilepsy in dentate nuclei, inferior Muscle weakness olives, posterior columns, Cerebellar ataxia and spinocerebellar tract Deafness Dementia Neuropathy, ataxia, and Neuropathy Cerebral and cerebellar retinitis pigmentosa (NARP) Cerebellar ataxia atrophy Retinitis pigmentosa Lesions in basal ganglia

symptoms (Ito et al. 2008) with cerebellar, brain stem, and cerebral atrophy of mild degree. Atrophy of the superior cerebellar peduncles was more visible than atrophy of the middle cerebellar peduncles. Furthermore, in previous studies, cerebral atrophy, cerebral white matter T2 hyperintensities, striatal T2 hyperintensities, pallidal atrophy with calcification, and cerebellar atrophy were already reported in patients with MERRF (Huang et al. 2002). Moreover, all these studies showed degeneration of the brain stem and dentate nuclei as well as the basal ganglia, cerebellar cortices, and spinal cord (Fukuhara 1991). Interestingly, the N-acetylaspartate/creatine ratio was found decreased in the cerebellum in some cases, whereas it was preserved in other studies (Watanabe et al. 1998). A patient carrying the A8344G point mutation showed an unusual presentation of MERRF, including cerebellar ataxia, myoclonus, and cervical lipomas, compatible with Ekbom’s syndrome. He had a history of slowly progressive dysarthria, dizziness, and gait disturbances. Later, he developed shortterm memory deficits, progressing to disorientation for time and place. Gait ataxia and dysarthria as well as cognitive deficits were present. General examination revealed symmetrical, posterior, cervical lipomas. Neurologically the patient presented cognitive dysfunction, moderate dysarthria with slow speech, and nystagmus. Moderate,

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intermittent segmental action myoclonus was present in the upper extremities. Tone was slightly reduced in the upper and lower limbs and rebound was present in the upper limbs. Synergy, trajectory, and placement of the limbs were abnormal with dysmetria and dysdiadocokinesia that were moderate in the upper and mild in the lower extremities. Stretch reflexes were hypoactive but symmetric and plantar responses were flexor, while vibration and position senses were normal. Gait and stance were ataxic and only two to three steps were possible with tandem gait. CT scan showed cerebellar atrophy and extensive symmetrical lipomas in the posterior cranio-cervical area, while, MRI demonstrated cerebellar atrophy. Moreover, CSF analysis (including lactate level), EEG (with photostimulation), electromyography, and nerve conduction studies were normal. Serum creatine kinase and lactic acid were increased. Muscle biopsy showed both ragged-red fibers and cytochrome oxidasenegative fibers, compatible with mitochondrial myopathy. In summary, in MERRF, neuronal loss occurs in the cerebellar dentate nuclei, globus pallidus, and red nuclei, substantia nigra, inferior olivary nuclei, and cerebellar cortex (Oldfors et al. 1995). Posterior columns, spinocerebellar tracts, and Clarke columns are also degenerating in the spinal cord (Fukuhara 2008). Brain imaging may demonstrate cerebral atrophy, calcium deposits in basal ganglia, and signal changes in the white matter on brain MRI (Di Mauro et al. 2002). The most common mutation of mtDNA in MERRF is the A8344G transition in the tRNALys gene (Wallace et al. 1988). Other mutations in the same gene are associated with MERRF (Silvestri et al. 1992), MERRF/MELAS syndrome (see below), or other syndromes. In MERRF muscle, A8344G mutation is associated to Complex IV deficiency, although complex I can also be affected. Ragged-red fibers are invariably detected in the muscle biopsy. Positive correlation between the severity of the disease, age at onset, mtDNA heteroplasmy, and reduced activity of respiratory chain complexes in skeletal muscle has been shown. Genotype– phenotype correlation between MERRF syndrome and the A8344G mutation is low. Moreover, A8344G mutation can be present in Leigh’s syndrome, isolated myoclonus, familial lipomatosis, isolated myopathy, and a variant neurological syndrome characterized by ataxia, myopathy, hearing loss, and neuropathy. Conventional antiepileptic drugs are used for seizures. Physical therapy to improve any impaired motor function and standard pharmacologic therapy for cardiac symptoms are also of current use. Levetiracetam, clonazepam, zonisamide, and valproic acid (VPA) have been used to treat myoclonic epilepsy; however, VPA may cause secondary carnitine deficiency and should be avoided or used with L-carnitine supplementation. Coenzyme Q10 (100 mg 3 per day) and L-carnitine (1,000 mg 3 per day) are often used in hopes of improving mitochondrial function.

Mitochondrial Encephalomyopathy with Lactic Acidosis and Stroke-like Episodes (MELAS) Mitochondrial myopathy, encephalopathy, lactic acidosis, and stroke (MELAS) syndrome is a progressive neurodegenerative disorder (Tay et al. 2006). It is characterized by stroke-like episodes due to focal brain lesions generally localized

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a

b

c

d

Fig. 102.2 Ragged-red fibers in MELAS. (a) Modified Gomori trichrome stain showing several ragged-red fibers (arrowhead). (b) Cytochrome c oxidase stain showing Type-1 lightly stained and Type II fibers, darker fibers, and a few fibers with abnormal collections of mitochondria (arrowhead). Note cytochrome c oxidase-negative fibers as usually seen in mitochondrial encephalopathy, lactic acidosis, and stroke-like episodes (MELAS). (c) Succinate dehydrogenase staining showing a few ragged blue fibers and intense staining in the mitochondria of the blood vessels (arrow). (d) Electron microscopy showing abnormal collection of mitochondria with paracrystalline inclusions (arrowhead), osmiophilic inclusions (large arrowhead), and mitochondrial vacuoles (small arrowhead). (Adapted from Abu-Amero et al. 2009)

in the parieto-occipital lobes, lactic acidosis, and ragged-red fibers (Fig. 102.2) (Di Mauro et al. 2002). In infancy, MELAS is associated with developmental delay, learning disability, or attention deficit prior to the development of the first stroke that can occur between 4 and 15 years (Fayssoil 2009). Stroke-like episodes are the hallmark feature of this disorder (Testai and Gorelick 2010). Progressive encephalopathy leading to dementia or failure to thrive may be also present (Hirano and Pavlakis 1994). Generally, patients start with seizures, recurrent headaches, anorexia, and vomiting. Exercise intolerance or proximal limb weakness can be the initial manifestation, followed by generalized tonic–clonic seizures. Seizures are often associated with stroke-like episodes of transient hemiparesis or cortical blindness that may produce altered consciousness. The cumulative residual effects of the stroke-like episodes gradually impair motor abilities, vision, and mentation.

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Sensorineural hearing loss adds to the progressive decline of these individuals. Furthermore, migrainous headaches occur in the majority of affected individuals. In MELAS multiple organs are affected, as demonstrated by large-scale symptoms: myoclonus and ataxia (Table 102.2–102.3) (Petruzzella et al. 2004), episodic coma, optic atrophy, cardiomyopathy, pigmentary retinopathy, ophthalmoplegia, diabetes mellitus, hirsutism, gastrointestinal dysmotility, and nephropathy. Interestingly, during stroke-like episodes, brain MRI shows areas of increased T2 signal, typically involving the posterior cerebrum and not conforming to the distribution of major arteries. Slow spreading of the stroke-like lesions in the weeks following the first symptoms can be documented by T2-weighted MRI (Iizuka et al. 2003). In addition, diffusion-weighted MRI shows increased apparent diffusion coefficient (ADC) in the stroke-like lesions of MELAS, in contrast to the decreased ADC seen in ischemic strokes (Kolb et al. 2003). Neuropathological studies in patients with a not “classic” MELAS phenotype show diffuse atrophy of the cerebellar cortex, gliosis of cerebellar white matter, and cactus formation in Purkinje cells with mitochondria accumulation (Tanahashi et al. 2000). Severe sporadic cerebellar ataxia can be also observed in some rare cases. In these patients, neurologic examination reveals ataxic gait, dysartric speech, and bilateral dysmetry on finger-to-nose, mild proximal muscle weakness, and hypotrophy in the upper limb girdle muscles (Petruzzella et al. 2004). Proprioception and all sensory modalities result preserved with normal lactate and creatine kinase content. Electromyographically, myopathic features are observed in proximal upper and lower limb muscles. In the brain MRI shows cortical and cerebellar atrophy. Histochemical analysis reveals the presence of ragged-red fibers in the skeletal muscle. Examination of MELAS’ CNS showed diffuse gliosis in the white matter, diffuse atrophy of the cerebral and cerebellar cortices, and cactus formation of Purkinje cells as described (Tanahashi et al. 2000). These lesions cannot be explained by the sole angiopathy, and strongly suggest other mechanisms involved, such as cytopathy. Ohama et al. (1987) used the term “mitochondrial angiopathy” to explain that abnormal accumulation of mitochondria in vascular endothelial cells and smooth muscle cells which could be responsible for the infarct-like lesions. Electron microscopic study confirmed that mitochondrial angiopathy is a unique and common change in MELAS brain. However, not all CNS lesions of MELAS can be attributed to abnormal vascular functions. Sano et al. (1995) have reported uncoupling between cerebral oxygen metabolism and cerebral blood flow with reduced fractional oxygen extraction, indicating hyperemia related to the malfunction of mitochondria in aerobic energy production. The fact that cultured cells derived from a MELAS patient exhibited a respiratory deficiency suggests that an intrinsic metabolic abnormality contributes to the neurological disorder of MELAS (Kobayashi et al. 1991). In fact, pathological changes in patients suggest neuronal metabolic dysfunction: cactus formation in the cerebellum of some cases, diffuse cortical atrophy and diffuse gliosis, and grumose degeneration in rare case. Subsequently, it turned out that Purkinje cell-expanded dendrites filled with mitochondria made up this structure (Morgello et al. 1988; Robain et al. 1988). Cactus formations found in metabolic and neurodegenerative disorders strongly suggest that degeneration is

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caused by a direct neuropathic or neurotoxic effect but not by ischemia. These findings, together with multiple areas of necrosis and diffuse gliosis in the cerebellum, suggest that the cerebellar function of MELAS patients should be evaluated carefully. In conclusion, all pathological changes reported in the current literature, including mitochondrial angiopathy, diffuse cortical atrophy, diffuse gliosis in the white matter, and cactus formations of Purkinje cells suggest that an intrinsic mitochondrial malfunction causes neuronal damage. MELAS syndrome has been associated with maternally inherited, or sporadic, heteroplasmic point mutations located in the tRNALeu(UUR) gene (Tanahashi et al. 2000). The most common mutation affecting 80% of patients is the A3243G (Mehrazin et al. 2009). Additionally, a lower number of patients have a heteroplasmic T3271C point mutation in the terminal nucleotide pair of the anticodon stem of the tRNALeu(UUR) gene. MELAS is also associated to the G13513A mutation in the ND5 gene and POLG deficiency. The MELAS disorder–associated human mitochondrial tRNALeu(UUR) mutation causes aminoacylation deficiency and a concomitant defect in translation initiation. Respiratory enzyme activities in MELAS revealed that more than one-half of the patients have complex I or complex I + IV deficiency. Thus, complex I deficiency probably correlates with MELAS. Moreover, this downregulation may ultimately lead to reduced respiratory chain activity caused by impaired translation of UUG-rich genes such as ND6 (component of complex I) (Borner et al. 2000; Shanske et al. 2008; Sasarman et al. 2008). Further, studies revealed that the A3243G mutation produces a severe combined respiratory chain defect in myoblasts, with almost complete lack of assembly of complex I, IV, and V, and a slight decrease of assembled complex III. This assembly defect occurs despite a modest reduction in the overall rate of mitochondrial protein synthesis (Deschauer et al. 2007). Translation of some polypeptides is decreased, and evidence of amino acid misincorporation is noted in others (Jacobs et al. 2000). No specific treatment for MELAS exists. Sensorineural hearing loss has been treated with cochlear implantation and seizures respond to traditional anticonvulsant therapy. Diabetes mellitus is managed by dietary modification, oral hypoglycemic agents, or insulin therapy. L-arginine showed promise in treating stroke-like episodes. Migraine headaches and cardiac manifestations are treated in the usual manner. Coenzyme Q10 and its analogue idebenone have been beneficial in some individuals.

MELAS/MEERF Overlapping Syndrome Several members of three-generation kindred from Italy were affected by a maternally inherited syndrome characterized by features of both MERRF and MELAS (Nakamura et al. 1995). Clinically, symptoms such as myoclonus epilepsy, neural deafness, and ataxia were variably associated with stroke-like episodes and/ or migrainous attacks. Morphologically, numerous MELAS-associated SDHstained vessels were observed in muscle biopsies, either alone or in combination with ragged-red fibers, the morphological hallmark of MERRF. Sequence analysis revealed the presence of a single heteroplasmic T8356C mutation, in the tRNA(Lys)

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gene. In another study, Nakamura and colleagues (Nakamura et al. 1995) found a family with a MELAS/MERRF overlap syndrome carrying the mutation in the tRNAser(UCN) gene. Patient had myoclonus epilepsy, hearing impairment, ataxia, mental deterioration, myopathy, lactic acidosis, and basal ganglia calcifications.

Neuropathy, Ataxia, and Retinitis Pigmentosa (NARP) Neuropathy, ataxia, and retinitis pigmentosa (NARP) is a maternally inherited mitochondrial disorder. The main clinical manifestations include proximal neurogenic muscle weakness with sensory neuropathy, ataxia (Table 102.2–102.3), pigmentary retinopathy, seizures, learning difficulties, and dementia (Holt et al. 1990). Onset of symptoms, particularly ataxia and learning difficulties, is often in early childhood. Diffuse cerebral and cerebellar atrophy, with symmetric lesions of the basal ganglia in more severe cases underlie NARP (Berkovic et al. 1993; Uziel et al. 1997). Other clinical features include short stature, sensorineural hearing loss, progressive external ophthalmoplegia, cardiac conduction defects (heart block), and a mild anxiety disorder (Santorelli et al. 1997). Visual symptoms may be the only clinical feature in similar cases. Individuals with NARP can be relatively stable for many years, but may suffer episodic deterioration, often in association with viral illnesses. Electromyography and nerve conduction studies may demonstrate peripheral neuropathy (which may be a sensory or sensorimotor axonal polyneuropathy). Cerebellar atrophy is the most frequent finding in NARP syndrome, followed by periventricular white matter involvement, in some cases associated with increased bilateral lenticular nuclei intensity on T2-weighted images (Uziel et al. 1997). Bilateral symmetric hypodensities in the basal ganglia are typical of Leigh’s syndrome or bilateral putaminal necrosis (Desguerre et al. 2003). Particularly, neuropathological examination in a patient with NARP (mtDNA T8993G mutation) revealed moderate cerebral atrophy and marked atrophy of the cerebellum and brain stem, as well as small confluent cysts in the putamen (Rojo et al. 2006). Additionally, the involvement of the visual pathways, in addition to the atrophy of the optic nerves and tracts, was characterized by loss of neurons and astrocytic gliosis in the lateral geniculate nucleus and superior colliculus. The cerebellum showed focal areas of necrosis with loss of Purkinje cells and massive dropout of granule cells. Many remaining Purkinje cells showed axonal torpedoes filled with phosphorylated neurofilaments, accompanied by marked loss of neurons in the inferior olivary and dentate nuclei. The involvement of the auditory pathways was manifested as a moderate loss of neurons in the dorsal and ventral cochlear nuclei, while the olfactory bulbs and tracts were atrophic. This was accompanied by marked neuronal depletion of the olfactory bulb. Multiple and confluent cystic lesions were seen in the putamen together with massive neuron loss and gliosis, while a moderate neuron loss and gliosis were also observed in the caudate. The disease is usually caused by a heteroplasmic mutation in the ATPase six subunit gene (T8993G) (Holt et al. 1990). The same mutation has also been found in a subset of patients with Leigh’s syndrome. A transition in the same position (T8993C) has also been described in patients (Sciacco et al. 2003), while a further mutation in the same position (T8993C) is associated to a milder

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phenotype (De Vries et al. 1993). Muscle biopsy results negative for ragged-red fibers staining. Heteroplasmy levels directly correlate with the severity of phenotype. In fact, high mutant mtDNA levels result in a full Leigh’s syndrome phenotype. NARP and Leigh’s syndrome may coexist in the same family. Individuals with NARP can also have a detectable mutation at mt-ATP6 nucleotide 8993. In vitro cells and tissues biopsies carrying the T8993G mutation are associated with a reduction of ATP synthesis (Carelli et al. 2003). The T8993G and T8993C mutations show a substantial genotype–phenotype correlation as individuals with T8993G mutant loads below 60% are usually asymptomatic, or have only mild pigmentary retinopathy or migraine, though a few asymptomatic adults with mutant loads of up to 75% have been reported (Tatuch et al. 1992). By contrast, individuals with moderate levels (70–90%) of the T8993G mutation present with the NARP phenotype, whereas those with mutant loads above 90% have maternally inherited Leigh syndrome. T8993C is a less severe mutation than T8993G, and virtually all symptomatic individuals have T8993C mutant loads of more than 90%. Treatment for NARP is supportive and includes use of sodium bicarbonate or sodium citrate for acidosis and antiepileptic drugs for seizures. Dystonia is treated with benzhexol, baclofen, tetrabenazine, and gabapentin alone or in combination or by injections of botulinum toxin. Anti-congestive therapy may be required for cardiomyopathy. Nutritional assessment of daily caloric intake and adequacy of diet is done regularly. Psychological support for affected individuals and family is essential.

Maternally Inherited Leigh Syndrome (MILS) Ataxia is present in nearly all patients with MILS. However, the phenotype is clinically heterogeneous. For example, the mitochondrial ATP6 mutation in the same family can be clinically associated to a late-onset MILS or NARP (Childs et al. 2007). The T8993C is associated to MILS phenotype at early infancy (Debray et al. 2007). Predominant ataxia and neuropathy were found in a family carrying the T9185C mutation with a heteroplasmy rate >90% (Castagna et al. 2007). Other ATP6 mutations may also cause the disease and ataxia may be the only manifestation (Campos et al. 1997). Other mitochondrial genes frequently mutated in MILS are the ND1-6, ATP6, COXIII, and tRNALys genes (Finsterer 2008), with a mutation load that correlates positively with the severity of the phenotype (Santorelli et al. 1993). Hearing Loss-Ataxia-Myoclonus (HAM) The heteroplasmic insertion 7472insC in the tRNASer(UCN) gene is responsible of the clinical phenotype in patients. HAM was firstly described as a maternally inherited syndrome characterized by a combination of sensorineural hearing loss, ataxia, and myoclonus in a large kindred from Sicily (Tiranti et al. 1995). Hearing loss was the most widespread and sometimes the only symptom. Moreover, the degree of heteroplasmy in blood and muscle correlate with the clinical phenotype, while homoplasmic mutant hybrids showed decreased complex I activity, low oxygen consumption, and high lactic acid output, indicating faulty oxidative

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phosphorylation. This phenotype was later confirmed in other pedigrees with a different number of clinical signs: isolated hearing loss, epilepsia partialis continua and ataxia, or MERRF phenotype (Jaksch et al. 1998). Given the increasing frequency at which the 7472insC has been found, the search for this mutation should become part of the routine screening of mitochondrial encephalomyopathies and/or maternally inherited hearing loss (Hutchin and Cortopassi 2000). Recently a homoplasmic mutation T7512C in the tRNASer(UCN) gene, has been reported in a family with a combination of sensorineural hearing loss, myoclonus epilepsy, ataxia, severe psychomotor retardation, short stature, and diabetes mellitus (Ramelli et al. 2006). Therapeutically, ubiquinone and antioxidants can be beneficial.

Other Mitochondrial Disorders with Ataxia The heteroplasmic tRNAIle gene mutation G4284A was found in a patient in association with truncal ataxia, dysarthria, severe hearing loss, mental retardation, ptosis, ophthalmoparesis, distal myoclonus, and diabetes mellitus. Complex I and IV activities were low in the muscle (Corona et al. 2002). Ataxia was also a phenotypic manifestation of the C14680A mutation in a patient with exercise intolerance, weakness and lactic acidosis, who showed a mosaic pattern of succinate dehydrogenase staining on muscle biopsy (Casali et al. 1999).

Large-Scale Rearrangements of Mitochondrial DNA Kearns–Sayre Syndrome (KSS) Kearns–Sayre Syndrome is a rare sporadic neuromuscular-ocular disorder (Kearns et al. 1958). Major clinical features include: chronic progressive external ophthalmoplegia (CPEO), pigmentary retinopathy (salt–pepper like appearance), pigmentary degeneration of the retina, cardiac conduction defect such as heart block, elevated cerobrospinal fluid protein, and cerebellar dysfunction with ataxia (Ishikawa et al. 2000). Serum and cerebrospinal fluid lactate and pyruvate levels are usually increased and muscle biopsy analysis shows ragged-red fibers. KSS is often combined with endocrinological symptoms (see below). Signs and symptoms normally appear by the age of 20. Children with KSS usually appear normal at birth, with male and female equally affected. Generally, the first physical characteristic is growth retardation. In addition, ptosis due to weakness of the elevator palpebrae superioris is seen during infancy. Other eye muscles may be affected later on and eventually result in CPEO. This syndrome also affects muscle of face, throat, neck, and/or shoulders, yet never affects the pupils. Usually, both the horizontal and upward gazes are involved while the downward gaze is often spared (Simaan et al. 1999). Sometimes it can mimic myasthenia gravis, but the serum creatine kinase levels may be slightly elevated (Ishikawa et al. 2000). Most individuals with KSS have visual difficulties due to the atypical retinitis pigmentosa. The ERG is usually normal or it shows mildly attenuated A and B

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wave amplitudes. Skeletal muscle biopsy shows typical ragged-red fibers in almost all patients. KSS patients die of abnormal cardiac conduction within the His bundle and because of syncopal episodes. Complete heart block can occur and His bundle record show trifascicular block (Kearns et al. 1958; Berenberg et al. 1977). Neurologic abnormalities occurring in KSS include nystagmus, ataxia, hearing loss, and dementia. The neuropathology of KSS is chiefly characterized by status spongiosus. Spongiform degeneration may be observed in the white matter of the cerebrum and cerebellum, and the gray matter is preferentially affected in the brainstem (Sparaco et al. 1993). Lesions may affect cranial nuclei, including the oculomotor nuclei. Brain stem lesions in the medulla may account for the respiratory distress in patients (Tanji et al. 1999). Furthermore, intracranial calcifications are common and CSF protein content is usually elevated (Tanji et al. 1999). A moderate loss of Purkinje cells and spongiform degeneration of the cerebellar white matter has also been observed. Moreover, MRI shows abnormal T2 high signal intensity in the deep gray matter nuclei, the cerebellar and the subcortical white matter. Together these evidences suggest that mitochondrial abnormalities in the dentate nucleus in conjunction with loss of Purkinje cells and spongiform degeneration of the cerebellar white matter is important in the genesis of the cerebellar dysfunction in KSS (Tanji et al. 1999). In some patients multiple endocrine dysfunction is observed, and include: hypoparathyroidism, diabetes mellitus, and/or primary failure of the ovaries or testes (Kearns et al. 1958; Ishikawa et al. 2000), which may result in short stature, a delay in reaching puberty, excessive fatigue, and/or muscle cramps. A deficiency of coenzyme Q10 in the serum and muscles can be also observed. The exact cause of Kearns–Sayre Syndrome is not yet known. Most cases are due to mitochondrial mutations that spontaneously occur within oocyte or zygote. Most of the literature suggests that KSS is caused by large-scale heteroplasmic deletions from 1.3 to 8.0 kb of mtDNA, which include subunits of the oxidative phosphorylation enzymes and several tRNA genes (Ota et al. 1994; Blok et al. 1995). Duplications have also been observed (Poulton et al. 1989) representing a possible distinctive feature from individuals with CPEO. As most patients present large-scale deletions of mitochondrial DNA gene sequence, analysis of regions surrounding the mtDNA deletion breakpoint in KSS individuals revealed putative vertebrate topoisomerase II sites suggesting that direct repeat sequences, together with putative topoisomerase II sites, may predispose certain regions of the mitochondrial genome to deletions (Blok et al. 1995). Single large-scale rearrangements of mtDNA found in many sporadic cases of KSS, can be detected in human oocytes, but, it is not yet clear why the mutations are not transmitted through female gametes to progeny. The severity of the clinical phenotype in patients with mitochondrial deletions depends on the distribution of normal and mutant mitochondria (degree of heteroplasmy). Additionally, the nature of the mitochondrial mutation appears to directly affect the clinical phenotype and the reduction in the absolute amount of wild-type mtDNA may play a significant pathogenic role (Moraes et al. 1995).

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Treatment of heart problems in KSS may require a pacemaker or antiarrhythymic drug therapy. Surgery may be used to correct visual problems. Separate treatment options for associated disorders (diabetes mellitus or hypoparathyroidism) may be necessary. In some cases, treatment may include hormone replacement therapies. Clinical trials with CoQ10 for the treatment of KSS and other mitochondrial cytopathies have shown potential benefits. L-Carnitine and vitamin therapy including coenzyme Q10, riboflavin, and vitamins C and K have also been tested.

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X-Linked Ataxias

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Josef Finsterer

Abstract

Hereditary ataxias are genetically and phenotypically heterogenous and follow various types of transmission. Hereditary ataxias which are X-linked are also a heterogeneous group of disorders but share the X-linked transmission and the development of cerebellar or sensory ataxia. In X-linked ataxias, ataxia may be either a dominant feature of the phenotype or only a minor feature among several others. X-linked ataxias with ataxia as the dominant feature include X-linked sideroblastic anemia with ataxia (XLSA) and fragile X tremor/ataxia syndrome (FXTAS). The number of X-linked disorders with non-dominant ataxia is increasing, and it is quite likely that further disorders of this type will be identified on the molecular level. Though therapy of X-linked ataxias is only symptomatic, there is ongoing research to develop more causal approaches. To further stimulate such research, it is important to clearly delineate the various phenotypes and to analyze details about the pathogenetic background so far achieved. Early recognition of X-linked ataxias is important since it has an impact on the treatment and the genetic counseling of affected patients.

Introduction X-linked ataxias are genetic disorders presenting with ataxia as the only or dominant clinical feature of the disorder, or in which ataxia is a minor or inconsistent clinical feature of the disease. Ataxia may be of the cerebellar or sensory type. Among the hereditary ataxia syndromes none is characterized by pure ataxia. The most well-known disorders, in which ataxia dominates the presentation,

J. Finsterer Danube University Krems, Postfach 20, 1180 Vienna, Austria e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 2313 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_103, # Springer Science+Business Media Dordrecht 2013

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Table 103.1 X-linked ataxias Disorder Gene Ataxia is the dominant feature XLSA ABCB7 FXTAS FMR1 Ataxia is not the dominant feature Arts syndrome PRPS1 PLP1 disorders PLP1 CMT1X GJB1 MECP2 duplication MECP2 Rett syndrome MECP2 Joubert syndrome TMEM216, CEP290 TMEM67, RPGRIP1L CC2D2A Adrenoleucodystrophy ABCD1 MTS TIMM8A XLMR

CUL4B

PM, deletion

PDH deficiency XLMR

PDHA1 SLC9A6

PM PM, deletion, duplication

Mutation

Features in addition to ataxia

PM CGG 59–200

Anemia, UMNS Dementia, PS, neuropathy

PM PM PM Duplication PM PM

Retardation, hypoacusis PMD, spastic paraplegia 2 Neuropathy, dysarthria Dystonia, spasticity, dementia Autism, epilepsy, paraparesis Hypotonia, retardation, OP

PM, deletion PM

UMNS, hypogonadism Hypoacusis, optic atrophy, dystonia Dysmorphism, retardation, hypogonadism Lactacidosis; Leigh syndrome Retardation, epilepsy, microcephaly

PM point mutation, UMNS upper motor neuron signs, PS Parkinson syndrome, PMS Pelizaeus Merzbacher disease, OP ophthalmoparesis

include the fragile X tremor/ataxia syndrome (FXTAS) and X-linked sideroblastic anemia with ataxia (XLSA). In most of the other cases, ataxia is one among other clinical manifestations of the disease (Table 103.1).

Disorders with Ataxia as the Dominant Feature X-Linked Sideroblastic Anemia with Ataxia (XLSA) XLSA is a rare syndromic mitochondrial disorder (MID), characterized by mild, early-onset sideroblastic anemia with hypochromia and microcytosis, associated with cerebellar ataxia and incoordination (Allikmets et al. 1999; Hellier et al. 2001). Transmission of the disease follows an X-linked trait with the consequence that only males are affected and that only females transmit the disease (female carriers). Only few families have been reported so far.

Etiology and Pathogenesis XLSA is caused by mutations in the mitochondrial ATP-binding cassette transporter ABCB7 located on chromosome Xq13.1-q13.3 (Maguire et al. 2001). The normal allelic variant comprises 16 exons (Bekri et al. 2000). In yeast,

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the corresponding gene plays a central role in the maturation of cytosolic iron-sulfur cluster-containing proteins (Bekri et al. 2000). There are indications that the protein transports components required for the maturation of cytosolic iron-sulfur clusters from the mitochondrion to the cytosol (Napier et al. 2005). Like frataxin, ABCB7 is involved in the biosynthesis of iron-sulfur clusters. Point mutations have been detected in exons 5–16 of the gene and in the intron/exon boundaries. Mutations are detected by direct sequencing and are of the partial loss of function type. Mutations so far indentified include the c. 1,200 T > G (p.Ile400Met), c. 1,231 G > T (p.Val411Leu), and c. 1,297 G > A (p.Lys433Glu) substitutions.

Clinical Presentation Since XLSA is a X-chromosomal disorder, the full phenotype is found only in male patients. The dominant clinical feature of male XLSA patients is ataxia of the cerebellar type. Ataxia is either non-progressive (Pagon et al. 1985) or slowly progressive (Hellier et al. 2001). In addition to trunk/gait ataxia, clinical neurologic examination may show dysmetria, dysdiadochokinesia, or mild dysarthria and intention tremor. In single patients, ataxia may improve with time (Pagon et al. 1985) but usually ataxia progresses on from the fifth decade of life (Hellier et al. 2001). Single patients develop upper motor neuron signs in the lower limbs, manifesting as exaggerated tendon reflexes, non-sustained ankle clonus, and extensor plantar responses. Rarely, patients develop nystagmus, hypometric saccades, or strabismus. Cognitive functions are usually not affected but occasionally learning disability may be seen. Some patients develop depression and a single patient with psychosis was reported (Hellier et al. 2001). Anemia does not become symptomatic. Diagnosis The diagnosis is based on the individual and family history, clinical examination, eventually neuropsychological testing, blood chemical examinations, imaging studies, and genetic studies. Blood cell counts may reveal hypochrome and microcytic anemia with marked poikilocytosis, reticulocytosis, and heavy stippling. Siderocytes may be found in affected males and occasionally in female carriers. The hematocrit and the mean corpuscular erythrocyte volume may be significantly reduced (Pagon et al. 1985). Serum concentrations of free erythrocyte protoporphyrin are elevated. Bone marrow examination may show iron stores in the form of ring sideroblasts. Cerebral imaging may be normal or may show moderate to severe cerebellar atrophy or hypoplasia (Raskind et al. 1991). Serum iron parameters are normal (Hellier et al. 2001; Pagon et al. 1985). Differentials Differential diagnoses that have to be considered include: (a) X-linked spastic paraplegia, which is not associated with anemia; (b) refractory anemia with ring sideroblasts, which is characterized by excessive iron accumulation within mitochondria of erythrocytes (Boultwood et al. 2008); (c) sideroblastic anemia with glutaredoxin-5-deficiency due to mutations in the GLRX5 gene

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(Camaschella 2009); and (d) X-linked sideroblastic anemia without ataxia but increased serum iron and transferrin saturation and parenchymal iron overload by the second or third decade, which is due to mutations in the ALAS2 gene (Fleming 2002).

Carriers Clinical neurologic examination is normal in female carriers. Female carriers have a normal hematocrit but may show a dimorphic blood smear with both hypochrome, microcytic and normal red blood cells. Bone marrow examination may show ring sideroblasts as in affected males. Free erythrocyte protoporphyrin serum levels are normal. If these structural abnormalities are absent, identification of female carriers requires identification of the disease-causing mutation in at least one family member. If an affected male is not available, sequencing of the regions of interest is recommended. Management Affected males profit from physical therapy for ataxia and spasticity. In the advanced stages of the disease, patients may benefit from ankle fixation orthoses and walkers. Speech therapy may improve intelligibility from dysarthria. Computed devices may help to manage difficulties with handwriting. Targeted genetic testing or screening for suspected point mutations in the ABC7 gene is available on a custom basis. Genetic Counseling Males who carry the mutation are clinically affected. Affected males pass the mutation to all their daughters but not to their sons. The father of an affected male will neither be affected nor a carrier. Female carriers are clinically unaffected and have a 50% chance to transmit the mutation in each pregnancy. If the mother of an affected male does not carry the mutation, the mutation has to be classified as de novo. If a mother has two affected sons but does not carry the mutation, germline mosaicism may be the cause. The risk of the sibs of a proband depends on the carrier status of the mother. Male sibs carrying the mutation will be affected; female sibs carrying the mutation will be carriers. The probands maternal aunts may be at risk of being a carrier or of being affected. Testing of female relatives at risk, prenatal testing, and preimplantation genetic diagnosis is possible if the mutation is known.

Fragile X Tremor/Ataxia Syndrome (FXTAS) FXTAS is clinically characterized by progressive cerebellar ataxia and intention tremor. Additionally, cognitive decline and neuropathy may develop. FXTAS is a trinucleotide disorder which follows an X-linked trait of inheritance. The prevalence of FXTAS is unknown.

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Etiology and Pathogenesis FXTAS is a trinucleotide disorder caused by an intronic CGG-repeat expansion between 59 and 200 repeats (premutation) in the FMR1 gene on chromosome Xq27.3 accompanied by aberrant methylation (Saul and Tarleton 1998; Strom et al. 2007). Males with >200 repeats develop a different phenotype, fragile X mental retardation syndrome (no transcription of FMR1). Only patients carrying the premutation exhibit FXTAS. Alleles up to 58 repeats are transmitted in a stable fashion without increase or decrease in the repeat number (Saul and Tarleton 1998). In normal alleles (5–40 repeats), the CGG-repeats are interrupted by an AGG triplet after every nine or ten CGG-repeats. The AGG triplets are assumed to maintain repeat integrity by preventing DNA strand slipping during replication (Saul and Tarleton 1998). In intermediate alleles (41–58 repeats), the number of uninterrupted CGG-repeats is an important predictor of repeat instability (Saul and Tarleton 1998). Increasingly longer, uninterrupted, pure repeats are more likely to become unstable (Eichler et al. 1994). The penetrance increases with age. The CGGexpansion is most readily tested with PCR and the methylation status by Southern blotting. Only in single cases retraction of the CGG-repeat between generations has been reported (Vits et al. 1994). Only rarely, the syndrome may be due to point mutations. Clinical Presentation FXTAS is characterized by late-onset, progressive cerebellar ataxia and intention tremor. Additional features include loss of short-term memory, deficits in executive functions, working memory deficits, cognitive decline, and dementia (Bacalman et al. 2006; Grigsby et al. 2006; Jacquemont et al. 2007; Louis et al. 2006). During the disease course parkinsonism, proximal weakness of the lower limbs, peripheral neuropathy, or autonomic dysfunction may additionally develop. The three cardinal clinical features include progressive intention tremor, ataxia, and cognitive decline, starting in the sixth decade of life (Adams et al. 2008; Berry-Kravis et al. 2007). Only 4% of the patients exhibit all three features, 20% two features, and half of the patients all three features (Adams et al. 2008). Onset is around 60 years of age, usually with tremor as the initial manifestation. Ataxia follows after 2 years and life expectancy ranges between 5 and 25 years after onset (Leehey et al. 2007). Diagnosis The diagnosis is based on the clinical presentation, as outlined above, imaging findings, and genetic testing. MRI typically shows symmetric regions of T2-hyperintensities in the middle cerebellar peduncles and adjacent cerebellar white matter (peridentate white matter) or non-specific symmetric signal changes in the cerebral white matter but normal pons and basal ganglia (Fig. 103.1) (Mascalchi 2008). The cerebellar volume may be decreased and the ventricular volume increased. The neuroradiological findings appear to correlate with the CGG-repeat length (Cohen et al. 2006). Genetic testing, which is readily available in many laboratories, is recommended if there is a clinical suspicion. The diagnosis is established as “definite” if an appropriate CGG-expansion, typical MRI

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Fig. 103.1 Axial T2weighted images showing hyperintense signals in the middle cerebellar peduncle bilaterally. Additionally, atrophy of the cerebellar hemispheres can be seen. (Reproduced from Ishii et al. 2010)

demyelination of the cerebellar peduncles or the brainstem, and either tremor or ataxia are present. The diagnosis is “probable” if one major neuroradiological finding and one minor clinical sign (parkinsonism, working memory deficits, executive cognitive function deficit) or two major clinical manifestations are present (Grigsby et al. 2006). The diagnosis is “possible” if a minor neuroradiological manifestation or one major clinical sign is present (Grigsby et al. 2006). The expansion is usually detected by PCR and Southern blot. PCR may fail to detect alleles in the upper permutation range and full mutation alleles. The methylation status of the gene can be assessed by “methylation PCR.” Deletion/duplication analysis can be performed by quantitative PCR, real-time PCR, multiplex ligation-dependent probe amplification (MLPA), or by array GH.

Differentials All neurological disorders presenting with the categories parkinsonism, tremor, ataxia, dementia, autonomic dysfunction, or stroke have to be considered as differentials (Biancalana et al. 2005; Hall et al. 2005). Management No specific treatment is available. Treatment is restricted to supportive care for gait disturbance or cognitive deficits. Genetic Counseling A carrier of the premutation may have a mother or father who is also mutation carrier. The mother of a male with a premutation has a premutation allele or an intermediate allele. Women and men carrying premutations are at risk for FXTAS.

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The risk of the sibs depends on their gender, the gender of the carrier parent, and the size of the expanded allele in the carrier parent. Males who carry the premutation inherit it to all their daughters but not their sons. Male transmission of the premutation may result in a small increase in the CGG-repeat number but not in a full mutation. All daughters of transmitting males are unaffected premutation carriers. Female premutation carriers have a 50% risk of transmitting a premutation allele to each offspring. The risk of inheriting a premutation allele to offspring of an individual with intermediate alleles ranging from 41 to 58 repeats is higher than that of the general population. In case a child is diagnosed with fragile X-syndrome, and in case his mother carries a premutation allele, his maternal grandfather is at risk to develop FXTAS. Prenatal testing of fetuses at risk for a FMR1 mutation can be performed on DNA from amnion cells. Preimplantation diagnosis may be available in families with known FMR1 mutation.

Genetic Disorders with Ataxia as a Non-dominant Feature Ataxia is a non-dominant phenotypic feature in a number of X-linked neurodegenerative disorders. These disorders include phospho-ribosyl-pyrophosphate synthetase 1 (PRPS1)-superactivity, proteolipid protein-1 (PLP1) disorders, X-linked Charcot-Marie-Tooth (CMT1X) disease, methyl-CgP binding protein-2 (MECP2) duplication syndrome, Rett syndrome, Joubert syndrome, X-linked adrenoleucodystrophy, Mohr–Tranebjaerg syndrome, X-linked mental retardation (XLMR) syndrome, and X-linked pyruvate-dehydrogenase (PDH) deficiency.

Phospho-Ribosyl-Pyrophosphate Synthetase 1 (PRPS1)-Superactivity (Arts Syndrome) Arts syndrome is part of the PRPS1-related spectrum of disorders due to missense mutations in the PRPS1 gene. PRPS1 superactivity manifests clinically as mental retardation, delayed motor development, muscle hypotonia, optic atrophy, profound hypoacusis, hyperuricemia, increased risk of infection, and ataxia (de Brouwer et al. 2010). Onset of most abnormalities of Arts syndrome is before the age of 2 years. PRPS1 catalyzes the first step of the synthesis of nucleotides, which not only serve as building blocks of nuclear acids but also as cofactors in cellular signaling and metabolism (de Brouwer et al. 2008, 2010).

Proteolipid Protein 1 (PLP1) Disorders PLP1 disorders are a heterogeneous group of X-linked diseases due to mutations in the PLP1 gene, which encodes a structural component of the myelin sheath (Fattal-Valevski et al. 2009). They manifest clinically in infancy or early childhood with nystagmus, hypotonia, and cognitive impairment, which consecutively

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progress to spasticity and ataxia with shortened lifespan (Garbern and Hobson 1999). The clinical presentation may emerge into distinct phenotypes such as Pelizaeus–Merzbacher disease or spastic paraplegia 2. Pelizaeus–Merzbacher disease is characterized by progressive psychomotor delay, nystagmus, spastic quadriplegia, and cerebellar ataxia. Cerebral MRI typically shows marked demyelination. Variable clinical expression within a family may be seen in patients carrying the novel PLP15 missense mutation c. 619 T > C.

X-Linked Charcot-Marie-Tooth (CMT1X) Disease CMT1X disease is a demyelinating hereditary neuropathy due to missense mutations in the GJB1 gene, which encodes the gap junction protein connexin32 (Siskind et al. 2009). Usually, these patients present with distal motor weakness, wasting of the distal lower limb muscles, and foot deformities. Some patients additionally develop central nervous system involvement, manifesting as loss of language, dysarthria, temporary facial weakness, truncal instability, or appendicular ataxia (Siskind et al. 2009).

Methyl-CgP Binding Protein 2 (MECP2) Duplication Syndrome The MECP2 duplication syndrome is clinically characterized by early childhoodonset learning disability, which is followed by progressive dystonia, spasticity, and ataxia (McWilliam et al. 2010). Life expectancy is reduced. Three affected children from one family died at age 9, 14, and 19 years respectively. Three carriers presented with learning difficulties, epilepsy, or psychosis (McWilliam et al. 2010). The phenotype in these patients was due to duplications in the MECP2 gene (McWilliam et al. 2010; Lugtenberg et al. 2009).

Rett Syndrome Rett syndrome, predominantly affecting girls, is an X-linked neurodevelopmental disorder characterized by mental retardation with impaired cognition and adaptability with autistic behavior, stereotypic hand movements, seizures, and ataxia (Kondo and Yamagata 2002). Single patients develop dysmorphism, slowly progressive spastic paraparesis, distal wasting, postural tremor of the upper limbs, or dysarthria (Dotti et al. 2002). Rett syndrome is due to missense or nonsense mutations in the methyl-CgP binding protein 2 (MECP2). MECP2 mutations are the cause of MECP2-related disorders, which in addition to Rett syndrome include variant Rett syndrome and maniac-depressive psychosis, pyramidal signs, parkinsonism, and macro-orchidism (PPM-X) syndrome (Christodoulou and Ho 2001).

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Fig. 103.2 Axial T1weighted image showing a deepened interpeduncular fossa (open arrowhead) and abnormal superior cerebellar peduncles (thick arrowheads), constituing the “molar tooth sign.” (Reproduced from Brancati et al. 2010)

Joubert Syndrome Joubert syndrome is a congenital syndrome characterized by multiple anomalies, of which the molar tooth sign, a complex midbrain-hindbrain malformation, is the dominant feature (Fig. 103.2) (Brancati et al. 2010). Other neurological abnormalities include muscle hypotonia, developmental delay, intellectual disability, disturbed eye movements, neonatal hypoventilation, and ataxia (Brancati et al. 2010). Involvement of non-neurological systems includes retinal dystrophy, nephrolithiasis, hepatic fibrosis, and polydactyly. Joubert syndrome is one of the ciliopathies, due to mutations in proteins responsible for the primary cilium of the centromere. So far, mutations in ten different genes have been detected (Table 103.1). Though transmission usually follows an autosomal recessive mode, some families with X-linked transmission have been described.

X-Linked Adrenoleukodystrophy X-linked adrenoleukodystrophy is a rare neurological disorder with a wide range of phenotypic expression (Li et al. 2010). X-linked adrenoleukodystrophy is clinically characterized by adult-onset spino-cerebellar ataxia, manifesting as dysarthria, cerebellar ataxia, and mild spastic paraparesis. Additionally, adrenal and gonadal impairment are prominent features of the phenotype (Vianello et al. 2005). MRI of the cerebrum and spinal cord may show atrophy of the cerebellum and upper cervical spinal cord (Li et al. 2010) and cerebral demyelination (Vianello et al. 2005). The disorder is due to point mutations or deletions in the ABCD1 gene (Li et al. 2010).

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Mohr–Tranebjaerg Syndrome Mohr–Tranebjaerg syndrome is an X-linked MID with pre- or postlingual sensorineural hearing loss since early childhood, and slowly progressive dystonia and ataxia since the teens (Tranebjærg 2003). On from the early 20s optic atrophy occurs, and on from the early 40s dementia develops (Tranebjærg 2003). Carriers may present with mild hypoacusis or late-onset focal dystonia. Mohr–Tranebjaerg syndrome results from point mutations in the TIMM8A gene or Xq22 deletion (Tranebjærg 2003).

X-Linked Mental Retardation (XLMR) Syndrome XLMR syndrome is due to point mutations or deletions in the CUL4B gene and manifests clinically as mental retardation, minor facial dysmorphism, short stature, delayed puberty, hypogonadism, macrocephaly, pes cavus, and gait ataxia (Isidor et al. 2010). In a patient with XLMR due to a deletion in the CUL4B gene, abnormalities of the aortic valve were additionally reported.

X-Linked Pyruvate-Dehydrogenase (PDH) Deficiency PDH deficiency is most commonly due to missense mutations, deletions, or duplications in the PDHA1 gene encoding the E1 subunit of the PDH complex (Bachmann-Gagescu et al. 2009; Ostergaard et al. 2009). The PDH complex is responsible for the decarboxylation of pyruvate to acetyl-CoA. The clinical presentation ranges from neonatal lactacidosis to severe leucencephalopathy (Leigh syndrome) (Bachmann-Gagescu et al. 2009). Less severe cases may present with intermittent ataxia exclusively (Bachmann-Gagescu et al. 2009; Ostergaard et al. 2009). The genotype-phenotype correlation is poor. Carbohydrate-free diet together with thiamine, carnitine, and vitamin E may have a beneficial effect with an excellent outcome in single patients.

X-Linked Mental Retardation, Microcephaly, Epilepsy, and Ataxia Syndrome This syndrome is a recently described condition clinically characterized by microcephaly, epilepsy, absent speech, and ataxia. The phenotype resembles that of Angelman syndrome, clinically characterized by developmental delay with minimal or absent use of words, happy demeanor with disposition for frequent smiling and spontaneous laughter, and ataxia (Gilfillan et al. 2008). Patients develop an open mouth with frequent drooling, abnormal food-related behavior, microcephaly, and flexed arms. In more than 80% of the cases, seizures develop before the age

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of 3 years. Clinical onset is at birth. The syndrome is due to point mutations in the SLC9A6 gene, one of more than 100 genes responsible for X-linked mental retardation syndromes (Gilfillan et al. 2008). Conclusion

X-linked ataxias are a heterogeneous group of disorders, which have in common the X-linked transmission and the development of cerebellar or sensory ataxia. Ataxia may be either a dominant feature in these disorders or only one collateral abnormality among several others. Those in which ataxia dominates are the X-linked sideroblastic anemia with ataxia (XLSA) and the fragile X tremor/ ataxia syndrome (FXTAS). The number of X-linked disorders with minor manifestations of ataxia is increasing, and it is quite likely that further disorders of this type will be detected on the molecular level. Though therapy in these conditions is only symptomatic, there is ongoing research to develop more causal approaches. For this purpose, it is important not only to describe the phenotypes and its variability in detail but also to fully clarify the pathogenetic background of all these disorders. Early recognition is not only useful for the treatment but also for genetic counseling.

References Adams SA, Steenblock KJ, Thibodeau SN, Lindor NM (2008) Premutations in the FMR1 gene are uncommon in men undergoing genetic testing for spinocerebellar ataxia. J Neurogenet 22:77–92 Allikmets R, Raskind WH, Hutchinson A et al (1999) Mutation of a putative mitochondrial iron transporter gene (ABC7) in X-linked sideroblastic anemia and ataxia (XLSA/A). Hum Mol Genet 8:743–749 Bacalman S, Farzin F, Bourgeois JA et al (2006) Psychiatric phenotype of the fragile X-associated tremor/ataxia syndrome (FXTAS) in males: newly described fronto-subcortical dementia. J Clin Psychiatry 67:87–94 Bachmann-Gagescu R, Merritt Ii JL, Hahn SH (2009) A cognitively normal PDH-deficient 18-year-old man carrying the R263G mutation in the PDHA1 gene. J Inherit Metab Dis (in press) Bekri S, Kispal G, Lange H, Fitzsimons E, Tolmie J, Lill R, Bishop DF (2000) Human ABC7 transporter: gene structure and mutation causing X-linked sideroblastic anemia with ataxia with disruption of cytosolic iron-sulfur protein maturation. Blood 96:3256–3264 Berry-Kravis E, Abrams L, Coffey SM et al (2007) Fragile X-associated tremor/ataxia syndrome: clinical features, genetics, and testing guidelines. Mov Disord 22:2018–2030 Biancalana V, Toft M, Le Ber I et al (2005) FMR1 premutations associated with fragile X-associated tremor/ataxia syndrome in multiple system atrophy. Arch Neurol 62:962–966 Boultwood J, Pellagatti A, Nikpour M et al (2008) The role of the iron transporter ABCB7 in refractory anemia with ring sideroblasts. PLoS One 3:e1970 Brancati F, Dallapiccola B, Valente EM (2010) Joubert syndrome and related disorders. Orphanet J Rare Dis 5:20 Camaschella C (2009) Hereditary sideroblastic anemias: pathophysiology, diagnosis, and treatment. Semin Hematol 46:371–377 Christodoulou J, Ho G (2001) MECP2-related disorders [updated 2 Apr 2009]. In: Pagon RA, Bird TC, Dolan CR, Stephens K (eds) GeneReviews [Internet]. University of Washington, Seattle; 1993–. http://www.ncbi.nlm.nih.gov/bookshelf/br.fcgi?book¼gene&part¼rett

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Cohen S, Masyn K, Adams J et al (2006) Molecular and imaging correlates of the fragile X-associated tremor/ataxia syndrome. Neurology 67:1426–1431 de Brouwer APM, Duley JA, Christodoulou J (2008) Arts syndrome. In: Pagon RA, Bird TC, Dolan CR, Stephens K (eds) GeneReviews [Internet]. University of Washington, Seattle; 1993–. http://www.ncbi.nlm.nih.gov/bookshelf/br.fcgi?book¼gene&part¼arts de Brouwer AP, van Bokhoven H, Nabuurs SB, Arts WF, Christodoulou J, Duley J (2010) PRPS1 mutations: four distinct syndromes and potential treatment. Am J Hum Genet 86:506–518 Dotti MT, Orrico A, De Stefano N et al (2002) A Rett syndrome MECP2 mutation that causes mental retardation in men. Neurology 58:226–230 Eichler EE, Holden JJ, Popovich BW, Reiss AL, Snow K, Thibodeau SN, Richards CS, Ward PA, Nelson DL (1994) Length of uninterrupted CGG repeats determines instability in the FMR1 gene. Nat Genet 8:88–94 Fattal-Valevski A, DiMaio MS, Hisama FM et al (2009) Variable expression of a novel PLP1 mutation in members of a family with Pelizaeus-Merzbacher disease. J Child Neurol 24:618–624 Fleming MD (2002) The genetics of inherited sideroblastic anemias. Semin Hematol 39:270–281 Garbern JY, Hobson GM (1999) PLP1-related disorders [updated 16 Mar 2010]. In: Pagon RA, Bird TC, Dolan CR, Stephens K (eds) GeneReviews [Internet]. University of Washington, Seattle; 1993–. http://www.ncbi.nlm.nih.gov/bookshelf/br.fcgi?book¼gene&part¼pmd Gilfillan GD, Selmer KK, Roxrud I et al (2008) SLC9A6 mutations cause X-linked mental retardation, microcephaly, epilepsy, and ataxia, a phenotype mimicking Angelman syndrome. Am J Hum Genet 82:1003–1010 Grigsby J, Leehey MA, Jacquemont S et al (2006) Cognitive impairment in a 65-year-old male with the fragile X-associated tremor-ataxia syndrome (FXTAS). Cogn Behav Neurol 19:165–171 Hall DA, Berry-Kravis E, Jacquemont S et al (2005) Initial diagnoses given to persons with the fragile X associated tremor/ataxia syndrome (FXTAS). Neurology 65:299–301 Hellier KD, Hatchwell E, Duncombe AS, Kew J, Hammans SR (2001) X-linked sideroblastic anaemia with ataxia: another mitochondrial disease? J Neurol Neurosurg Psychiatry 70:65–69 Ishii K et al (2010) A Japanese case of fragile-X-associated tremor/ataxia syndrome (FXTAS). Inter Med 49:1205–1208 Isidor B, Pichon O, Baron S, David A, Le Caignec C (2010) Deletion of the CUL4B gene in a boy with mental retardation, minor facial anomalies, short stature, hypogonadism, and ataxia. Am J Med Genet A 152A:175–180 Jacquemont S, Hagerman RJ, Hagerman PJ, Leehey MA (2007) Fragile-X syndrome and fragile X-associated tremor/ataxia syndrome: two faces of FMR1. Lancet Neurol 6:45–55 Kondo I, Yamagata H (2002) Mutation spectrum and genotype-phenotype correlation of MECP2 in patients with Rett syndrome. No To Hattatsu 34:219–223 Leehey MA, Berry-Kravis E, Min SJ et al (2007) Progression of tremor and ataxia in male carriers of the FMR1 premutation. Mov Disord 22:203–206 Li JY, Hsu CC, Tsai CR (2010) Spinocerebellar variant of adrenoleukodystrophy with a novel ABCD1 gene mutation. J Neurol Sci 290:163–165 Louis E, Moskowitz C, Friez M, Amaya M, Vonsattel JP (2006) Parkinsonism, dysautonomia, and intranuclear inclusions in a fragile X carrier: a clinical-pathological study. Mov Disord 21:420–425 Lugtenberg D, Kleefstra T, Oudakker AR et al (2009) Structural variation in Xq28: MECP2 duplications in 1% of patients with unexplained XLMR and in 2% of male patients with severe encephalopathy. Eur J Hum Genet 17:444–453 Maguire A, Hellier K, Hammans S, May A (2001) X-linked cerebellar ataxia and sideroblastic anaemia associated with a missense mutation in the ABC7 gene predicting V411L. Br J Haematol 115:910–917 Mascalchi M (2008) Spinocerebellar ataxias. Neurol Sci 29(suppl 3):311–313 McWilliam C, Cooke A, Lobo D, Warner J, Taylor M, Tolmie JL (2010) Semi-dominant X-chromosome linked learning disability with progressive ataxia, spasticity and dystonia

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associated with the novel MECP2 variant p.V122A: akin to the new MECP2 duplication syndrome? Eur J Paediatr Neurol 14:267–269 Napier I, Ponka P, Richardson DR (2005) Iron trafficking in the mitochondrion: novel pathways revealed by disease. Blood 105:1867–1874 Ostergaard E, Moller LB, Kalkanoglu-Sivri HS et al (2009) Four novel PDHA1 mutations in pyruvate dehydrogenase deficiency. J Inherit Metab Dis (in press) Pagon RA, Bird TD, Detter JC, Pierce I (1985) Hereditary sideroblastic anaemia and ataxia: an X linked recessive disorder. J Med Genet 22:267–273 Raskind WH, Wijsman E, Pagon RA, Cox TC, Bawden MJ, May BK, Bird TD (1991) X-linked sideroblastic anemia and ataxia: linkage to phosphoglycerate kinase at Xq13. Am J Hum Genet 48:335–341 Saul RA, Tarleton JC (1998) FMR1-related disorders [updated 18 May 2010]. In: Pagon RA, Bird TC, Dolan CR, Stephens K (eds) GeneReviews [Internet]. University of Washington, Seattle; 1993–. http://www.ncbi.nlm.nih.gov/bookshelf/br.fcgi?book¼gene&part¼fragilex Siskind C, Feely SM, Bernes S, Shy ME, Garbern JY (2009) Persistent CNS dysfunction in a boy with CMT1X. J Neurol Sci 279:109–113 Strom CM, Huang D, Li Y, Hantash FM, Rooke J, Potts SJ, Sun W (2007) Development of a novel, accurate, automated, rapid, high-throughput technique suitable for population-based carrier screening for fragile X syndrome. Genet Med 9:199–207 Tranebjærg L (2003) Deafness-dystonia-optic neuronopathy syndrome [updated 24 Mar 2009]. In: Pagon RA, Bird TC, Dolan CR, Stephens K (eds) GeneReviews [Internet]. University of Washington, Seattle; 1993–. http://www.ncbi.nlm.nih.gov/bookshelf/br.fcgi? book¼gene&part¼ddon Vianello M, Manara R, Betterle C, Tavolato B, Mariniello B, Giometto B (2005) X-linked adrenoleukodystrophy with olivopontocerebellar atrophy. Eur J Neurol 12:912–914 Vits L, De Boulle K, Reyniers E, Handig I, Darby JK, Oostra B, Willems PJ (1994) Apparent regression of the CGG repeat in FMR1 to an allele of normal size. Hum Genet 94:523–526

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Mitsunori Yamada

Abstract

The neuropathology of major types of cerebellar system degeneration is described in this chapter. Multiple system atrophy (MSA) is a major, nonhereditary spinocerebellar degeneration, characterized pathologically by the olivopontocerebellar atrophy, striatonigral degeneration, and autonomic nervous system degeneration in any combination. The occurrence of glial cytoplasmic inclusions in the oligodendrocytes is a pathologic hallmark of MSA, and involves multiple brain regions including the pontocerebellar tracts and internal capsules, as well as the other affected systems. Spinocerebellar ataxias (SCAs) contain various types of neurodegenerative diseases, and are characterized pathologically by the combined degeneration of the cerebellar cortex and other brain regions. Polyglutamine diseases are a major group of SCA, and there is a significant correlation between CAG-repeat lengths in the causative genes and disease severities. The formation of neuronal intranuclear inclusions is a characteristic feature of polyglutamine diseases, and is found in brain regions specific to each disease. The chromosome 16q-linked autosomal dominant cerebellar ataxia/spinocerebellar ataxia type 31 is a newly discovered cerebellar degeneration, and characterized pathologically by a peculiar eosinophilic structure which is surrounding the remaining Purkinje cell bodies.

Introduction The cerebellum can be affected in various pathologic conditions, such as neurodegenerative disorders, ischemic conditions, nutritional deficiencies, metabolic

M. Yamada Department of Clinical Research, National Hospital Organization, Saigata National Hospital, 468-1 Saigata, Ohgata-ku Johetsu-city, Niigata, 949-3193, Japan e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 2327 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_104, # Springer Science+Business Media Dordrecht 2013

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disorders, inflammatory diseases, and paraneoplastic syndromes. The major cell type affected in each disease condition is the Purkinje cell. Granule cells are less frequently affected, but may be the main target in several conditions, such as Creutzfeldt-Jakob disease or methyl mercury poisoning. In this chapter, the neuropathology of major types of cerebellar system degeneration is described. This includes conventional and molecular neuropathology, which has been established by the discovery of causative gene mutations or proteins.

Spinocerebellar Degeneration Sporadic Disorders Multiple System Atrophy Multiple system atrophy (MSA) is a nonhereditary, adult-onset disorder, characterized clinically by parkinsonism, ataxia, and autonomic failure, in any combination (Burn and Jaros 2001). The term “multiple system atrophy” was first proposed by Graham and Oppenheimer (1969), based on the finding that sporadic olivopontocerebellar atrophy (OPCA), striatonigral degeneration (SND), and Shy-Drager syndrome can coexist both clinically and pathologically. Recent studies categorize MSA into two phenotypes: (1) MSA-P is the category of MSA where parkinsonism predominates, and (2) MSA-C is the category of MSA where cerebellar ataxia predominates (Gilman et al. 1999; Gilman et al. 2008). Grossly, there is obvious atrophy of the ventral part of the pons, with marked degeneration of the middle cerebellar peduncles (Fig. 104.1). The cerebellar cortex and inferior olive are also atrophic. Shrinkage with discoloration is noticed in the putamen (Fig. 104.2), and this change is commonly accentuated in its posterolateral part. The substantia nigra is depigmented (Fig. 104.3). In patients with clinical features of OPCA/MSA-C type, atrophy is more prominent in the pontocerebellar system than the striatonigral system. In patients with clinical features of SND/MSA-P type, it is typical that the striatonigral system is more severely affected. Histologically, neuron loss is commonly observed in the pontine nuclei, inferior olive, putamen, substantia nigra, intermediolateral nucleus of the spinal cord, and autonomic ganglia. In the cerebellar cortex, Purkinje cells are variably depleted (Fig. 104.4). The flocculus is typically less affected. In contrast to the Purkinje cell pathology, granule cells are preserved. Neuron loss may be observed in the locus ceruleus, vestibular nuclei, Onuf’s nucleus, and sensory ganglia. A pathological hallmark of MSA is the presence of glial cytoplasmic inclusions (GCIs) in the oligodendrocytes (Fig. 104.5) (Papp et al. 1989; Nakazato et al. 1990). GCIs are immunohistochemically positive for a-synuclein (Fig. 104.6) (Arima et al. 1998; Tu et al. 1998; Wakabayashi et al. 1998), and appear in multiple brain regions including the motor cortex, cerebral white matter subjacent to the motor cortical areas, globus pallidus, internal capsule, olfactory bulbs, and reticular formation of the brainstem, as well as in the striatonigral and olivopontocerebellar systems.

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Fig. 104.1 Multiple system atrophy. The brain shows severe atrophy of the brainstem and cerebellum

Fig. 104.2 Multiple system atrophy. The coronal section of the cerebrum through the level of the mammillary bodies shows shrinkage with discoloration of the putamen

The optic nerve usually lacks the inclusions. Filamentous inclusions positive for a-synuclein are also found in the neuronal cytoplasm, axons, and nucleus in the affected regions (Fig. 104.7). It is interesting that Purkinje cells are excluded from the molecular pathology.

2330 Fig. 104.3 Multiple system atrophy. The midbrain shows marked depigmentation of the substantia nigra (upper panel), and the ventral pons is severely atrophic (lower panel)

Fig. 104.4 Multiple system atrophy. Severe Purkinje cell loss is evident. Hematoxylin and eosin stain

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Fig. 104.5 Multiple system atrophy. Glial cytoplasmic inclusions are evident in oligodendrocytes. GallyasBraak silver stain

Fig. 104.6 Multiple system atrophy. Glial cytoplasmic inclusions are positive for a-synuclein. Immunostain

Late Cortical Cerebellar Atrophy Late cortical cerebellar atrophy (LCCA) is a nonhereditary progressive disorder, characterized clinically by late-onset purely cerebellar ataxia (Marie et al. 1922). Grossly, the cerebellum shows atrophy of the cerebellar cortex that is usually accentuated in the superior half of the vermis. Histologically, Purkinje cell loss is predominant in this region, and the cerebellar hemispheres are less affected. In some cases, Purkinje cell loss may involve extensive cortical regions in the vermis and hemispheres, or may be predominant in the cerebellar hemispheres (Tsuchiya et al. 1994). In LCCA, it is common that neuronal loss is also noticed in the inferior olive, particularly in the dorsomedian part.

Hereditary Disorders Spinocerebellar Ataxia Type 1 Spinocerebellar ataxia type 1 (SCA1) is a dominantly inherited form of spinocerebellar degeneration caused by expansion of a CAG repeat in the SCA1

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Fig. 104.7 Multiple system atrophy. Accumulation of asynuclein is evident in the cytoplasm and nuclei of neurons. Immunostain

gene localized to chromosome 6p23 (Zoghbi and Orr 1995). The numbers of CAG repeat units in patients with SCA1 range from 40 to 81, and correlate inversely with the age at onset and disease severity. The clinical features in the early stages of SCA1 are characterized by progressive ataxia, pyramidal impairment, and oculomotor palsy, followed in the later stages by amyotrophy and sensory disturbances. Cognitive function typically remains intact. Neuropathologic studies have revealed atrophy of the brainstem and spinal cord, which are more marked in patients with juvenile onset. The cerebellum may be atrophic, but the cerebrum typically appears normal. The brain weight mostly ranges from 1,100 g to 1,200 g (Iwabuchi et al. 1999). In the cerebellum, Purkinje cells are mildly to moderately depleted, but in some patients the cerebellar cortex appears almost normal. Torpedoes are occasionally observed in the granular layer. The cerebellar dentate nucleus also shows mild to moderate neuronal loss, associated with grumose degeneration, which is characterized by accumulation of numerous eosinophilic and argyrophilic granular materials around the somata and dendrites of dentate neurons. In the brainstem, neuronal loss is observed in the pontine nuclei and inferior olivary nucleus, although this is relatively mild in comparison with that found in SCA2. The substantia nigra, red nucleus, and cranial nerve nuclei including the vestibular and oculomotor nuclei are often affected. The spinal cord shows apparent neuronal loss in the anterior horn and Clarke’s column with degeneration of the spinocerebellar tracts. The posterior column shows various degrees of degeneration. In the cerebrum, no apparent change is evident in the cortex, white matter, striatum or thalamus, although mild to moderate degeneration is often observed in the outer segment of the globus pallidus. The inner segment of the globus pallidus may be involved in some patients (Uchihara et al. 2006). Neuronal intranuclear inclusions (NIIs) are observed in broad areas of the brain (Duyckaerts et al. 1999), such as the cerebral cortex, striatum, globus pallidus, substantia nigra, pontine nuclei, reticular formation, inferior olive, dentate nucleus, and spinal anterior horn, with the highest incidence in the pontine nuclei. They are

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eosinophilic and spherical, and present singly or occasionally in pairs in a nucleus. Their sizes may vary even in the same patient, ranging from ~1 to 3 mm in diameter. NIIs are inconspicuous using hematoxylin-eosin staining, but easily detected by immunohistochemistry for ubiquitin, ataxin-1 (a protein construct of the SCA1 gene), and expanded polyglutamine stretches using a monoclonal antibody 1C2 (Trottier et al. 1995). It should be noted that no inclusion has been found in Purkinje cells, which are an essential cell type depleted in SCA1 brains. The lack of prominent NIIs in Purkinje cells is also observed in the other CAG repeat diseases including SCA2, Machado-Joseph disease, and DRPLA (Hayashi et al. 1998; Koyano et al. 2002), even though mutant proteins accumulate diffusely in their nuclei. No inclusions have been reported in glial cells.

Spinocerebellar Ataxia Type 2 Spinocerebellar ataxia type 2 (SCA2) is a dominantly inherited neurodegenerative disease caused by expansion of a CAG repeat in the SCA2 gene localized to chromosome 12q24.1 (Imbert et al. 1996; Orozco et al. 1989; Pulst et al. 1997; Sanpei et al. 1996). The numbers of CAG repeat units in patients with SCA2 range from 35 to 64. The clinical features in the early stages of SCA2 are characterized by progressive ataxia, diminished tendon reflexes, and slow eye movement, followed in the later stages by amyotrophy, sensory disturbance, involuntary movements, and mental deterioration. Neuropathologic studies have revealed atrophy of the cerebellum and pontine base. The substantia nigra is depigmented. The brain weight mostly ranges from 690 g to 1,265 g (Durr et al. 1995; Iwabuchi et al. 1999; Orozco et al. 1989). In the cerebellum, Purkinje cells and granule cells are moderately to severely depleted, but the dentate nucleus is typically spared. In the brainstem, the pontine nuclei, inferior olive, and substantia nigra are severely affected. Moderate degeneration is also detected in the red nucleus. The involvement of the spinal anterior horn and dorsal column is variable. Mild degeneration may be encountered in the basal ganglia, thalamus, and cerebral cortex in some patients. Ataxin-2 has a cytoplasmic localization in normal brain, and the SCA2 gene is expressed in Purkinje cells and some specific groups of brainstem and cortical neurons (Huynh et al. 1999). Purkinje cells in SAC2 patients also possess many cytoplasmic granules immunopositive for ataxin-2 and expanded polyglutamine stretches (Huynh et al. 2000). These intracytoplasmic granules are negative for ubiquitin. In contrast to the other CAG repeat diseases, NII formation is not prominent in SCA2. Ubiquitinated NIIs have been found only in 1–2% of pontine neurons. They are also detectable in the other affected regions such as the substantia nigra, inferior olive, globus pallidus, and cerebral cortex, but not in Purkinje cells (Koyano et al. 1999). Machado-Joseph Disease/Spinocerebellar Ataxia Type 3 Machado-Joseph disease/spinocerebellar ataxia type 3 (MJD/SCA3) is a dominantly inherited multisystem neurodegenerative disorder characterized by variable combinations of cerebellar ataxia, pyramidal signs, dystonic extrapyramidal symptoms, peripheral neuropathy with amyotrophy, nystagmus, eyelid

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Fig. 104.8 Machado-Joseph disease/spinocerebellar ataxia type 3. Atrophy is evident in the brainstem. The cerebellum is relatively preserved

retraction, external ophthalmoplegia, and facial fasciculations (Rosenberg 1992; Sudarsky and Coutinho 1995). Dystonia is often prominent in younger patients. The disorder is due to an unstable CAG repeat on chromosome 14q32.1. The numbers of CAG repeat units in patients with MJD range from 56 to 84. The brain weight of affected patients mostly ranges from 1,000 to 1,300 g. In most cases, there is obvious atrophy of the brainstem (Fig. 104.8) and spinal cord, and the substantia nigra is depigmented (Fig. 104.9). The cerebellum may also be atrophic due to loss of white matter volume (Fig. 104.10). Atrophy with brownish discoloration is occasionally evident in the globus pallidus and subthalamic nucleus. The main lesions in MJD are located in the spinocerebellar system and cerebellar dentate nucleus (Iwabuchi et al. 1999; Yamada et al. 2004). In the spinal cord, Clarke’s column generally shows severe neuronal loss with marked degeneration of the spinocerebellar tracts. Severe degeneration is also detected in the anterior horn, with consequent degeneration of the anterior spinal roots and skeletal muscles of the extremities. The involvement of the spinal posterior horn and dorsal column is variable and usually mild. In most cases, no apparent abnormality is detected in the corticospinal tract. In the brainstem, mild to moderate neuronal loss is detectable in the pontine nuclei, with accentuation in the caudal region. Neuronal depletion is also evident in the substantia nigra, reticular formation, accessory cuneate nucleus, and cranial nerve nuclei including the nuclei of the external ocular muscles, and hypoglossal and vestibular nuclei. Although variable degrees of degeneration may be present in the red and dorsal column nuclei, the inferior olive is typically spared. The cerebellar cortical neurons are preserved in most cases, but minimal loss of Purkinje cells and occasional torpedoes are encountered in some patients. The cerebellar white matter is atrophic and shows myelin pallor due to degeneration of the pontocerebellar and spinocerebellar fibers. The dentate nucleus shows

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Fig. 104.9 Machado-Joseph disease/spinocerebellar ataxia type 3. The midbrain shows marked depigmentation of the substantia nigra (upper panel). The pons shows moderate atrophy (lower panel)

Fig. 104.10 MachadoJoseph disease/ spinocerebellar ataxia type 3. The cerebellar hemisphere shows diffuse myelin pallor in the white matter. Kl€uverBarrera stain

moderate to severe loss of neurons with grumose degeneration. In the globus pallidus, the internal segment is more severely affected. Severe neuronal loss is also detectable in the subthalamic nucleus. The thalamus may display mild degeneration, especially in the centromedian nucleus; however, no significant degeneration is detected in the striatum or cerebral cortex.

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Fig. 104.11 MachadoJoseph disease/ spinocerebellar ataxia type 3. Intranuclear inclusions are evident in neurons of the pontine nuclei. Immunostain for expanded polyglutamine stretches with 1C2 monoclonal antibody

NII formation is found in the affected brain regions (Fig. 104.11), and shows a relatively high incidence among the CAG repeat diseases (Paulson et al. 1997; Schmidt et al. 1998). In addition, as in other polyglutamine disorders, NIIs are detectable in unaffected regions including the cerebral cortex, thalamus (especially the intralaminar nucleus), striatum, lateral geniculate body, inferior olive, and dorsal root and sympathetic ganglia (Yamada et al. 2001a; Yamada et al. 2004). The distribution is generally wider in patients with a longer expansion of the CAG repeat. No inclusion has been observed in Purkinje cells (Koyano et al. 2002; Yamada et al. 2004). NIIs are spherical and eosinophilic, and vary in size from ~1 to 4 mm. They are present in the nucleus as a single structure or frequently as doublets. Immunohistochemistry reveals that NIIs are positive for ubiquitin, ataxin3, expanded polyglutamine stretches, and several transcription factors. Ultrastructurally, NIIs are non-membrane bound, and contain a mixture of granular and filamentous structures, the latter being approximately
12–15 nm in diameter and organized in random but sometimes parallel arrays. There is a relationship between NIIs and nuclear structures such as promyelocytic leukemia protein nuclear bodies and coiled bodies (Yamada et al. 2001a). The widespread occurrence of NIIs suggests that neurons are affected in the polyglutamine pathogenesis of MJD to a much greater extent than has been recognized by conventional neuropathologic studies. A neuropathologic study of a MJD patient, who was suspected to have died at an early stage of the disease, indicated that extensive formation of NIIs may be an early pathologic change, and that NII formation is related to phenotypic expression in the disease (Yamada et al. 2004). In contrast to the frequent formation of NIIs, diffuse nuclear immunolabeling for expanded polyglutamine stretches is a rare finding in MJD brains. In addition to NIIs, affected neurons possess many cytoplasmic granules immunolabeled with 1C2. Electron microscopy has shown that the granules are a subset of lysosomes (Yamada et al. 2002a). The appearance of this cytoplasmic pathology involves many brain regions with a distribution pattern generally similar to that of NII, suggesting that mutant proteins in the MJD brain are involved in both the ubiquitin/proteasome and endosomal/lysosomal pathways for protein degradation in

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different intraneuronal compartments. The cytoplasmic pathology is also observed in Purkinje cells of SCA6, but absent in SCA17. No pathologic changes related to polyglutamine pathology have been reported in glial cells or visceral organs.

Spinocerebellar Ataxia Type 6 Spinocerebellar ataxia type 6 (SCA6) is an autosomal dominant neurodegenerative disorder caused by expansion of a CAG repeat in the a1A voltage-dependent calcium channel gene (Ikeuchi et al. 1997; Ishikawa et al. 1997; Zhuchenko et al. 1997). The number of CAG repeats in SCA6 patients ranges from 20 to 33. It should be noted that CAG-repeat expansion in SCA6 is smaller than usual lengths of CAG repeats in other polyglutamine diseases. The main clinical feature is characterized by late-onset rather pure cerebellar ataxia. Grossly, there is marked atrophy in the cerebellum. The other brain regions are usually spared. Histologically, severe Purkinje cell loss with Bergmann’s gliosis is a consistent feature in the cerebellum. The loss is typically conspicuous in the cerebellar vermis, especially in the superior vermis. The cerebellar hemispheres show milder loss of the cell type. Granule cells are also affected, although the degree is milder than that of Purkinje cells. The dentate nucleus is usually spared. Mild degeneration of the inferior olive may be seen in some patients. No obvious changes have been reported in the other brain regions (Ishikawa et al. 1999). Small aggregates immunopositive for expanded polyglutamine stretches are present mainly in the cytoplasm but also in the nuclei of Purkinje cells (Ishikawa et al. 2001). Spinocerebellar Ataxia Type 7 Spinocerebellar ataxia type 7 (SCA7) is a dominantly inherited spinocerebellar degeneration characterized by retinal-cerebellar atrophy, and caused by expansion of a CAG repeat in the SCA7 gene localized to chromosome 3p12-13 (Enevoldson et al. 1994, Gouw et al. 1994, 1995; Holmberg et al. 1995; Martin et al. 1994; Neetens et al. 1990). The SCA7 repeat is one of the most unstable CAG repeats known, and the number of repeats in patients ranges from 38 to 460 (van de Warrenburg et al. 2001). There is a marked variability in age at onset and severity of the symptoms. The main clinical features include a decrease of visual acuity, progressive cerebellar ataxia, dysarthria, and dysphagia. Typically, no dementia or epilepsy is noted. Patients with extremely long CAG repeat stretches show juvenile or infantile onset, more rapid disease progression, and a broader spectrum of phenotypes than those with the adult onset form. Although the SCA7 gene products are expressed throughout the brain and retina, neurodegeneration is restricted in some regions. Grossly, the brains of SCA7 patients show atrophy of the optic pathways and cerebellum. Histologically, the retinas disclose severe degeneration of the pigmented epithelium and loss of photoreceptors, bipolar cells, and ganglion cells, with consecutive degeneration from the optic nerves to optic radiations including the lateral geniculate bodies. In the cerebellum, degeneration is observed in the cortex (Purkinje cell dominant), spinocerebellar and olivocerebellar tracts, and dentate nucleus. Although the inferior olive is generally involved, the

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degeneration of the ponto-cerebellopetal system is variable. The pyramidal pathways and motor neurons in the brainstem and spinal cord are also affected. Degeneration may be evident in the subthalamic nucleus, globus pallidus, and substantia nigra in some patients. The cerebral cortex and thalamus are typically free from degeneration. NIIs are detected in the affected brain regions with a relatively high incidence in the inferior olive (Holmberg et al. 1998). Interestingly, NIIs are also observed in areas of the cerebral cortex such as the supramarginal gyrus and insula. In addition to NIIs, 1C2 immunostaining reveals cytoplasmic granular staining in neurons in some brain regions including the supramarginal gyrus, hippocampus, thalamus, geniculate body, and pontine nuclei, and is not always dependent on NII formation. No pathologic changes are detectable in glial cells. In spite of the severe phenotype, infantile-onset SCA7 patients show relatively limited neuronal degeneration in the cerebellum and retina (Carpenter and Schumacher 1966; de Jong et al. 1980; Ryan et al. 1975; Traboulsi et al. 1988).

Spinocerebellar Ataxia Type 8 Spinocerebellar ataxia type 8 (SCA8) is a hereditary neurodegenerative disorder caused by expansion of a CTG repeat in the 30 untranslated region of a gene localized to chromosome 13q21 (Koob et al. 1999). Affected individuals show progressive gait and limb ataxia, dysarthria and nystagmus, with variable ages at onset (Day et al. 2000; Ikeda et al. 2000). Neuropathologic studies have revealed relatively pure atrophy of the cerebellum (Ito et al. 2006). Histologically, severe loss of Purkinje cells is the most prominent finding. The remaining Purkinje cells are atrophic and occasionally show somatic sprouts. Neuronal loss is also detectable in the inferior olive and substantia nigra. Although SCA8 is not a CAG repeat disease, it is interesting that 1C2-positive intranuclear inclusions and pan-nuclear staining are found in Purkinje, medullary, and dentate neurons from human SCA8 brains (Moseley et al. 2006). These inclusions are also positive for ubiquitin. 1C2-positive granular structures are detected in the cytoplasm of Purkinje cells (Ito et al. 2006). Glial cell involvement is not seen. The Chromosome 16q-Linked Autosomal Dominant Cerebellar Ataxia/Spinocerebellar Ataxia Type 31 The chromosome 16q-linked autosomal dominant cerebellar ataxia/spinocerebellar ataxia type 31 (16q-ADCA/SCA31) is a form of spinocerebellar ataxia common in Japan, and clinically characterized by late-onset purely cerebellar ataxia (Ishikawa and Mizusawa 2010). Grossly, brains of 16q-ADCA patients show obvious atrophy of the cerebellum. The cerebrum and brainstem appear normal. Histologically, the cerebellum shows moderate to severe loss of Purkinje cells. In contrast to the other spinocerebellar ataxias, it is remarkable that the remaining Purkinje cells in 16qADCA are associated with a peculiar eosinophilic structure which is surrounding Purkinje cell bodies (Fig. 104.12). The centered Purkinje cells are often shrunken. Immunohistochemically, it has been demonstrated that the eosinophilic structure is

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Fig. 104.12 The chromosome 16q-linked autosomal dominant cerebellar ataxia/ spinocerebellar ataxia type 31. A peculiar eosinophilic structure (arrow) is evident around the Purkinje cell body. Hematoxylin and eosin stain

mainly composed of somatic sprouts of Purkinje cells and presynaptic terminals of unknown origin. Astrocytic processes are present at the outside margin of the structure.

Dentatorubral-Pallidoluysian Atrophy Dentatorubral-pallidoluysian atrophy (DRPLA) is an autosomal dominant neurodegenerative disorder caused by expansion of a CAG repeat in the DRPLA gene located on chromosome 12p13.31. The number of CAG repeat units in DRPLA patients ranges from 49 to 84. Intergenerational instability is more pronounced in paternal transmission. DRPLA patients show various symptoms, such as myoclonus, epilepsy, ataxia, choreoathetosis, and dementia, and the combinations of these symptoms depend on the age at onset (Naito 1990). Patients with earlier onset (generally below the age of 20 years) show progressive myoclonus, epilepsy, and mental retardation (juvenile type, as classified by Naito). Patients showing late disease onset (over the age of 40 years) predominantly show cerebellar ataxia and dementia (late-adult type). Patients in whom the disease appears between the third and fifth decades belong to an intermediate type, and usually show ataxia and choreoathetosis (early-adult type). There is a reverse correlation between the age at onset and CAG repeat length. In contrast to the considerable heterogeneity in clinical presentation, the neuropathology of the DRPLA brain shows a relatively uniform pattern of lesion distribution, with combined degeneration of the dentatorubral and pallidoluysian systems (Fig. 104.13). The globus pallidus and subthalamic nucleus (Luys body) show consistent loss of neurons with astrocytic gliosis. In the globus pallidus, neuronal depletion is more severe in the lateral segment than in the medial segment. The dentate nucleus also shows loss of neurons, and the remaining atrophic neurons frequently exhibit grumose degeneration. Degeneration of the red nucleus is typically mild. In general, pallidoluysian degeneration is more marked than that of the dentatorubral systems in the juvenile type, and the reverse situation is observed in the late-adult type. Mild degeneration may be seen in the cerebral cortex, especially in patients showing juvenile onset.

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Fig. 104.13 Dentatorubralpallidoluysian atrophy. The brainstem and cerebellum are atrophic

In the case of infantile onset with 80 CAG repeats, neuronal depletion occurs in multiple brain regions including the cerebral cortex, striatum, inferior olive, and cerebellar cortex (Ohama et al. 1995). Diffuse myelin pallor of the cerebral and cerebellar white matter is often reported in aged patients. Morphometric analysis has revealed a decreased number of glial cells in the affected white matter (Yamada et al. 2002b). Despite the restricted nature of the brain lesions, it is characteristic that the amount of central nervous system (CNS) tissue is significantly reduced throughout the brain and spinal cord. Brain weights of DRPLA patients often become less than 1,000 g (Naito and Oyanagi 1982). Most of the brain regions lacking obvious neuronal loss show an increase of neuronal density due to atrophy of the neuropil. Thickening of the cranium is often observed in patients with juvenile onset. NII formation in the DRPLA brain is not restricted to the dentatorubral and pallidoluysian systems, but involves multiple regions including the cerebral cortex, substantia nigra, and pontine nuclei. Although the distribution is widespread, the incidence of neurons with inclusions is relatively low, and ranges from ~1 to 3% even in the dentate nucleus. NIIs in DRPLA are immunohistochemically positive for atrophin-1, expanded polyglutamine stretches, ubiquitin, and transcription factors (Yamada et al. 2001b, c). Intranuclear inclusions are also detectable in glial cells (Hayashi et al. 1998; Yamada et al. 2002b), as well as in non-neuronal tissues such as the kidney and pancreas (Yamada et al. 2001b). Immunohistochemistry with 1C2 antibody shows that diffuse accumulation of mutant atrophin-1 in neuronal nuclei is the predominant pathologic condition (Fig. 104.14), rather than NII

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Fig. 104.14 Dentatorubralpallidoluysian atrophy. Mutant proteins with expanded polyglutamine stretches are diffusely accumulated in the neuronal nuclei of the cerebellar dentate nucleus. Arrows indicate intranuclear inclusions. Immunostain for expanded polyglutamine stretches with 1C2 monoclonal antibody

Fig. 104.15 Filamentous inclusion positive for expanded polyglutamine stretches (arrow) is evident in the neuronal cytoplasm of the cerebellar dentate nucleus. Immunostain for expanded polyglutamine stretches with 1C2 monoclonal antibody

formation, and involves a wide range of CNS regions including the dentatorubral and pallidoluysian systems (Yamada et al. 2001b). The extent and frequency of neurons showing the diffuse nuclear pathology changes markedly and strikingly depending on the CAG repeat length, suggesting that neuronal dysfunction caused by mutant protein accumulation, rather than neuronal depletion, is responsible for the development of various clinical features in DRPLA. Neuronal nuclei with accumulation of mutant atrophin-1 show deformity with marked nuclear membrane indentations (Takahashi et al. 2001). Immunohistochemistry with 1C2 antibody also reveals the presence of granular staining in the neuronal cytoplasm, with a distribution pattern resembling that of diffuse nuclear staining (Yamada et al. 2004). In addition to NII formation, filamentous inclusions are also observed exclusively in the cytoplasm of dentate nucleus neurons (Fig. 104.15) (Yamada et al. 2000). The morphology of these structures is indistinguishable from the skeinlike inclusions observed in motor neurons in amyotrophic lateral sclerosis;

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Fig. 104.16 Early-onset ataxia with ocular motor apraxia and hypoalbuminemia/ataxiaoculomotor apraxia type 1. The cerebellum is severely atrophic

however, they are immunohistochemically positive for atrophin-1, expanded polyglutamine stretches and ubiquitin, but negative for TDP-43, the TAR DNAbinding protein 43. Light and electron microscopic features of NIIs in DRPLA are essentially similar to those of MJD.

Friedreich’s Ataxia Friedreich’s ataxia (FRDA) is an autosomal recessive neurodegenerative disorder caused by expansion of GAA repeats in the frataxin gene located on chromosome 9q13-21.1. The number of GAA repeat units in FRDA patients ranges from 120 to 1,700. Symptoms of FRDA usually appear around the beginning of the second decade of life, and include ataxia, sensory loss, and muscle weakness, as well as the common signs of scoliosis, foot deformity, and hypertrophic cardiomyopathy (Pandolfo 2009). Diabetes may be developed in some patients. Grossly, the spinal cords of FRDA patients are usually small, with atrophy of the posterior nerve roots. Histologically, the spinal cord shows marked degeneration of the posterior columns and corticospinal tracts. There is severe loss of neurons in the Clarke’s column with degeneration of the dorsal spinocerebellar tracts. The accessory cuneate nucleus in the medulla also shows severe loss of neurons, with degeneration of the ventral spinocerebellar tracts. The neuronopathy in the dorsal root ganglia causes the loss of the peripheral sensory nerve fibers. Cranial nerves, such as the trigeminal, glossopharyngeal, and vagus nerves, are also affected. Purkinje cell loss is usually minimal, but neuron loss is detectable in the cerebellar dentate nucleus. Most

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Fig. 104.17 Early-onset ataxia with ocular motor apraxia and hypoalbuminemia/ataxiaoculomotor apraxia type 1. The cerebellar cortex shows severe Purkinje cell loss. Hematoxylin and eosin stain

Fig. 104.18 Early-onset ataxia with ocular motor apraxia and hypoalbuminemia/ataxiaoculomotor apraxia type 1. Degeneration is evident in the posterior columns and spinocerebellar tracts. The corticospinal tracts are also affected. Kl€uver-Barrera stain

patients exhibit loss of the Betz and other pyramidal neurons in the motor cortex. Degeneration may be evident in the subthalamic nucleus, globus pallidus (the external segment), substantia nigra, auditory nuclei, and retinal ganglion cells (Pandolfo 2008).

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Early-Onset Ataxia with Ocular Motor Apraxia and Hypoalbuminemia/Ataxia-Oculomotor Apraxia Type 1 Early-onset ataxia with ocular motor apraxia and hypoalbuminemia/ataxiaoculomotor apraxia type 1 (EAOH/AOA1) is an autosomal recessive hereditary ataxia caused by mutations in the APTX gene (Date et al. 2001; Moreira et al. 2001). EAOH/AOA1 is characterized by early-onset slowly progressive ataxia, ocular motor apraxia, peripheral neuropathy, and hypoalbuminemia. MRI imaging exhibits atrophy of the cerebellum, particularly of the cerebellar vermis. Neuropathologically, the cerebellum shows marked atrophy (Fig. 104.16), and severe Purkinje cell loss is the most characteristic feature in the brains of EAOH/AOA1 (Fig. 104.17). The loss is more accentuated in the cerebellar vermis. Degeneration is also apparent in the posterior columns and spinocerebellar tracts (Fig. 104.18). Neuron loss is detectable in the spinal anterior horns and dorsal root ganglia. There are no obvious pathologic changes in the cerebral cortex or basal ganglia. No inclusions have been reported in neurons or glial cells (Tada et al. 2010).

Conclusions and Future Directions The discovery of causative proteins or gene mutations has triggered the development of novel neuropathology in spinocerebellar degenerations. It is now likely that the dynamics of the molecular-dependent lesion distribution may be responsible for a variety of clinical phenotypes in ataxic diseases. To understand the pathogenesis of each disease, it will be necessary that a pivotal role of the molecular pathology should be clarified in the development of selective neuronal degeneration in the central nervous system.

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General Management of Cerebellar Disorders: An Overview

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Winfried Ilg and Dagmar Timmann

Abstract

Cerebellar disorders primarily effect motor functions and can lead to significant and serious restrictions in activities of daily living. Possibilities for medical interventions are rare and limited to specific diseases and symptoms. Furthermore, motor rehabilitation for patients suffering from cerebellar damage is challenging, since the cerebellum is known to play an important role for the execution as well as for the (re)learning of precise movements. This chapter reviews the state of the art in medical intervention and rehabilitation, focusing on presenting new results on motor rehabilitation in cerebellar disease. Recent studies indicate that even in the case of degenerative cerebellar diseases intensive and continuous motor training can reduce ataxia symptoms and increase motor performance relevant to daily living. In addition, current studies in the area of motor learning – in combination with modern imaging techniques – in cerebellar disease are described. These results offer promising perspectives for a deeper understanding of remaining motor learning capacities in cerebellar disease, and thus might help in the future to optimize motor rehabilitation for individual patients.

W. Ilg (*) Section Computational Sensomotorics, Department of Cognitive Neurology, Hertie Institute for Clinical Brain Research, Centre for Integrative Neuroscience, University of T€ ubingen, Frondsbergstrasse 23, 72070 T€ ubingen, Germany e-mail: [email protected] D. Timmann Department of Neurology, University of Duisburg-Essen, Hufelandstrasse 55, 45147 Essen, Germany e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 2349 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_105, # Springer Science+Business Media Dordrecht 2013

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Introduction Cerebellar dysfunction can induce a variety of motor impairments including upper and lower limb movement, oculomotor control, balance, and walking (Bastian 1997, 2011; Diener and Dichgans 1996; Holmes 1939). Causes for cerebellar impairments can be various, including stroke, cerebellar tumors, multiple sclerosis, and degenerative disease. Functional recovery heavily depends on the cause and site of the lesion. Within this spectrum, degenerative cerebellar diseases are especially hard to treat, since – despite greatly improved understanding of the genetic underpinnings (Klockgether 2011; Sch€ ols et al. 2004) – no cure or drug treatments to ameliorate ataxia or decelerate disease progression are yet available. Furthermore, motor rehabilitation is also challenging for this patient population, since the cerebellum is known to play a functional role in motor learning. Therefore, poor recovery or low benefit of physiotherapeutic training may be a consequence of damaging structures critically involved in the relearning of motor skills (Bastian 2006; Thach and Bastian 2004). However, recent results deliver pieces of evidence that patients with degenerative cerebellar diseases can benefit from intensive and continuous motor training. This chapter will review the state of the art of medical intervention and rehabilitation, focusing on presenting new results and developments in the field of motor rehabilitation and motor adaptation in cerebellar disease. Other relevant reviews with different focuses can be found in several articles (see Marsden and Harris 2011; Mills et al. 2007, who focus on rehabilitation in multiple sclerosis, Cassidy et al. 2009 for an excellent review of clinical management; Urbscheit and Oremland 1995; Martin et al. 2009; and Trujillo-Martin et al. 2009).

General Predictions of Functional Recovery The cause, site, and extent of brain lesions are generally thought to be important predictors of the degree of functional recovery. For example, functional deficits seem more marked following a hemorrhage compared to an ischemic infarct, but have better chances of recovery if survived. Outcome has only been systematically studied in cerebral hemorrhage, but is likely the same if the hemorrhage occurs within the cerebellum (Weimar et al. 2003). In focal cerebellar lesions, either due to tumor surgery or stroke, lesion site appears to be more important than extent. Lesion extent does not predict long-term outcome. Rather, functional recovery is worse in lesions affecting the deep cerebellar nuclei and its output pathway, the superior cerebellar peduncles (Kelly et al. 2001; Konczak et al. 2005; Schoch et al. 2006). Because the latter are almost exclusively supplied by the superior cerebellar artery (Amarenco 1991), stroke in the territory of the superior cerebellar artery seems to have a worse prognosis than stroke affecting the posterior and anterior inferior cerebellar arteries. Furthermore, additional extra-cerebellar stroke lesions involving the cerebral hemispheres or brain stem also negatively affect recovery of walking and other functional capacities after stroke (Gialanella et al. 2005).

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In progressive degenerative cerebellar ataxias neuronal loss is caused by genetic factors with autosomal dominant inheritance as in the spinocerebellar ataxias (SCA) (Sch€ ols et al. 2004) or a recessive trait like in Friedreich’s ataxia (FA) (Durr et al. 1996). On the other hand, many sporadic cases of cerebellar degeneration escape the detection of causative factors and are currently classified as sporadic adult onset ataxia of unknown etiology (SAOA) (Klockgether 2010). Although clinical presentation is variable in many subtypes, SCA and SAOA mostly present with predominantly cerebellar ataxia, whereas in FA, afferent deficits are generally the major cause of ataxia. In general, degenerative ataxias seem to be the most difficult group of ataxias to treat, due to the progressive nature. In addition, virtually all parts of the cerebellum are affected although degeneration is frequently most prominent in the midline (Schulz et al. 2010). In contrast, ataxia following stroke, neurosurgery, or trauma affects only circumscribed regions of the cerebellum, but leaves other regions intact. These regions may compensate for the defective parts. In addition, in case of focal lesions, effects of neural plasticity are likely more effective because there is no competition with ongoing progressive neurodegeneration. Whereas patients with focal lesions clearly improve in motor functions over time, patients with degeneration slowly deteriorate (Schoch et al. 2007). Thus, in patients with progressive degenerative diseases, it would be a major achievement if they stayed on the current status of motor function as long as possible or if progression of functional impairment was slowed down. In patients with focal lesions, on the other hand, it is currently unknown to what extent functional improvement is due to spontaneous recovery or interventions like physiotherapeutical training.

Medical Intervention Medication is of restricted use in cerebellar ataxia. Focal cerebellar disorders, such as cerebellar stroke, tumors or lesions in multiple sclerosis, are treated according to the respective evidence-based clinical practice guidelines. Causal treatment of cerebellar degenerations is not available with the rare exceptions of a few, autosomal recessive ataxias with known metabolic defects (Fogel and Perlman 2007). As yet, no medication is known which ameliorates the clinical symptoms of cerebellar ataxia except treatment of attacks in episodic ataxias and treatment of downbeat nystagmus (see Aminopyridines below). More effective medications are available to treat accompanying extra-cerebellar symptoms in degenerative cerebellar disease such as extrapyramidal disorders, bladder dysfunction, and migraine. Over the years, a number of different drugs have been tested for symptomatic treatment of cerebellar ataxia (for recent reviews see Revuelta and Wilmot 2010; Trujillo-Martin et al. 2009). Most studies describing positive effects were openlabel trials and performed in a small number of patients. As yet, none of these positive effects could be replicated in subsequent studies using larger patient populations and a randomized placebo-controlled double-blind clinical trial design. Noneffective treatments include serotonergic drugs (5-hydroxytryptophane and

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buspirone), nutritional supplements (creatine and L-carnitine), antioxidants (vitamin E, coenzyme Q10, idebenone), and others (amantadine, gabapentin). Different antiepileptic drugs have been tried to treat cerebellar tremor, e.g., clonazepam, levetiracetam, primidone, and topiramate. None showed clear effects. In more recent years, idebenone, riluzole, and aminopyridines have been in the focus of interest as possible treatment options in cerebellar ataxia. Their possible therapeutical indications and results of drug trials will be discussed in more detail. Idebenone in Friedreich’s ataxia. Friedreich’s ataxia is characterized by a loss of frataxin. Loss of frataxin leads to respiratory chain defects and accumulation of iron in the mitochondria and, as a consequence, increased oxidative stress (Schulz et al. 2009). Idebenone, a short-chain derivative of coenzyme Q10, is both an antioxidant and electron carrier and is therefore thought to be of particular help in Friedreich’s ataxia (Mancuso et al. 2010; Schulz et al. 2009). A number of studies showed that echocardiographic parameters of cardiac hypertrophy improved using conventional doses of idebenone (Mariotti et al. 2003). A more recent study suggested that high dosages may also ameliorate neurological symptoms (Di Prospero et al. 2007). As a consequence idebenone was approved for treatment of Friedreich’s ataxia in Canada under the trade name of Catena ®. However, the results of two subsequent Phase III randomized placebo-controlled double-blind drug trials in comparatively large study populations failed to support the effectiveness of high-dosage idebenone both for neurological as well as cardiac study endpoints (MICONOS (Mitochondrial Protection with Idebenone In Cardiac Or Neurological Outcome Study); IONIA (Idebenone effects On Neurological ICARS Assessments); Lagedrost et al. 2011; Lynch et al. 2010). Thus, different to initial belief, idebenone (and other antioxidant therapies) does not appear a treatment option in Friedreich ataxia. Based on the known pathomechanisms of the disease, other treatment options are currently evaluated or developed, such as iron-chelating agents to increase the frataxin level (erythropoietin, histone deacetylase inhibitors) and other gene-based strategies (Kearney et al. 2009). As yet, an effective medical treatment of Friedreich’s ataxia is lacking. Riluzole. Riluzole is thought to be a neuroprotective agent that has been administered in different neurodegenerative disorders with varying success. Using a randomized, placebo-controlled, and double-blinded study design, Ristori and coworkers (2010) found positive effects in 38 patients with various forms of chronic ataxias. Fifty milligrams of Riluzole given twice daily was followed by significant effects on a clinical ataxia rating score (ICARS) after 8 weeks of treatment. A high percentage of individual patients (13/19) showed a significant reduction of the ICARS score after 8 weeks (>5 points; maximum ICARS score ¼ 100). Patients suffered from autosomal dominant (spinocerebellar ataxia, SCA, type 1, 2, and 28), recessive (Friedreich’s ataxia), and X-chromosomal ataxias (Fragile X tremor ataxia syndrome); immunological disease (Anti-GAD, Anti-Yo); sporadic disease (sporadic adult onset ataxia, SAOA; multiple system atrophy type C, MSA-C); and multiple sclerosis. Thus, patients differed in primary location of the disease (cerebellar cortex, nuclei, or spinocerebellar pathways) and etiology. Although a drug which is of benefit for any kind of cerebellar ataxia is

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highly desirable, it is hard to imagine that it exists. Furthermore, a previous study in a large group of patients with MSA (almost 400; most of them suffering from MSA-P) did not show any beneficial effects, neither on ameliorating symptoms nor in slowing disease progression (Bensimon et al. 2009). It remains to be tested whether the positive results of Ristori and coworkers can be replicated in a larger and more homogenous study population using a longer treatment period than 8 weeks. Aminopyridines. In recent studies, aminopyridines proved to be effective in treating both episodic ataxia type 2 (EA2) and nystagmus (downbeat- and upbeat nystagmus) (see Strupp and Brandt 2009 for a review). Aminopyridines are potassium channel blockers which were commonly believed to increase the inhibitory drive of cerebellar Purkinje cells. According to recent animal data, they restore the diminished precision of pacemaking in Purkinje cells (Alvina and Khodakhah 2010). Both 4-aminopyridine and 3,4-diaminopyridin have been used and, except for differences in dosages, effects appear to be the same. In EA2, acetazolamide is long been known to reduce attacks (Griggs et al. 1978). 4-Aminopyridine is a useful alternative in cases where acetazolamide lost its effect or severe side-effects occur (e.g., recurrent kidney stones). 4-Aminopyridine has been shown both to reduce attacks (Strupp et al. 2011) and to ameliorate interictal cerebellar signs (Schniepp et al. 2011). Downbeat nystagmus (DBN) leads to unsteadiness of gait and vertigo. Blurred vision and oscillopsia are additional symptoms (Strupp and Brandt 2009). Idiopathic DBN refers to cases with unknown etiology and secondary DBN to cases with known etiology. Secondary DBN is most frequently due to cerebellar degeneration. 3,4-Diaminopyridine reduces DBN in approximately 50% of the patients (Strupp et al. 2003). Treatment worked best in patients without structural lesions of the cerebellum and brainstem (i.e., idiopathic DBN). Positive effects have also been described in SCA6 and other hereditary ataxias (Tsunemi et al. 2010). Results about possible improvement of accompanying ataxic symptoms are contradictory in the literature (Sprenger et al. 2005; Tsunemi et al. 2010). Upbeat nystagmus is a less common type of nystagmus in cerebellar degeneration. 4-aminopyridine appears to be effective (Glasauer et al. 2005). Placebo-controlled group studies, however, are missing. Use of aminopyridines is off-label. A retarded form of 4-aminopyridine has been approved as Ampyra ® (Fampridin) for treatment of gait disorders in multiple sclerosis in the United States in 2009. 3,4-Diaminopyridine has been approved as Firdapse ® (Amifampridin) for treatment of Lambert-Eaton syndrome in Europe in 2010. Treatment is generally well tolerated. Similar to acetazolamide transient perioral or digital paraesthesias, headache and nausea are possible side effects. In higher dosages, aminopyridines can cause cardiac arrhythmias and seizures.

Cognitive Rehabilitation Neuroanatomical, functional imaging, and human lesion studies suggest involvement of the cerebellum in cognitive function (for recent reviews see Stoodley and

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Schmahmann 2009; Strick et al. 2009). In addition, many degenerative cerebellar disorders with extra-cerebellar involvement present with neuropsychological abnormalities (Burk 2007). The cerebellum appears to contribute particularly to executive functions and language. Deficits are more marked in acute than in chronic disease. Whereas dysfunction is generally subtle in adults with chronic cerebellar disease and appears more marked in childhood disorders, significant intellectual disability is reported in pre- and perinatal cerebellar disorders (Timmann and Daum 2010 for review). Thus, neuropsychological assessment should be part of the clinical workup in patients with cerebellar disorders, in particular in children, and acute disease. Cognitive dysfunction may require rehabilitation. As yet, however, studies examining the effect of neuropsychological training in cerebellar disease are sparse. Two case reports in patients suffering from acute cerebellar hemorrhage report no or modest improvement of executive functions following cognitive training (Maeshima and Osawa 2007; Schweizer et al. 2008). In contrast, remarkable improvement of executive dysfunction was observed in patients with acute ischemic stroke regardless of whether cognitive training was administered (Pierscianek et al. 2007).

Motor Rehabilitation Impairments in Motor Performance and the Adaptation of Movements The cerebellum is involved in the control of various kinds of motor behavior in speech, oculomotor control, limb movements, and balance. The functional role of the cerebellum is not the generation but more the shaping and fine-tuning of movements. Therefore, cerebellar damage does not cause loss of movement, but instead leads to abnormalities in movement characterized by increased variability and poor accuracy (Bastian 2006). In the case of limb movements, and in particular goal-directed movements of the upper extremities, typical ataxia symptoms are dysmetria (hypermetria and hypometria), cerebellar tremor, and dyssynergia, the lost ability to move joints simultaneously (Bastian et al. 1996; Manto 2009; Topka et al. 1998; Urbscheit and Oremland 1995; Vilis and Hore 1980). Similarly, cerebellar ataxic gait is typically characterized by an increased step width, variable foot placement, irregular foot trajectories, and a resulting instable stumbling walking path (Diener and Dichgans 1996; Holmes 1939; Ilg et al. 2007; Morton and Bastian 2004) with a high risk of falling (van de Warrenburg et al. 2005). The underlying control mechanisms for the execution of accurate movements are suggested to involve the implementation of internal forward models within the cerebellum, which predict sensory consequences of actions (Bastian 2006; Kawato 1999; Wolpert et al. 1998). Thus, it has been proposed that the cerebellum calculates a current “state estimate” – by combining sensory information about the last known position of the limb with predictions of its responses to recent movement

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commands – which is used to accurately plan and control goal-directed movements (Miall et al. 2007). Such mechanisms play a particularly important role in movements in which predictive control strategies are necessary. Examples for these are fast or ballistic movements when reactive control mechanism are not sufficient because of time delays in sensory signals. Also in multi-joint movements, predictive control is generally involved for the predictive compensation of joint interaction forces (Bastian et al. 1996). Therefore, it is well known that impairments in limb control become more pronounced in multi-joint limb movements and for increased movement velocity (Topka et al. 1998). Further impairments of cerebellar patients related to predictive control mechanism are dysfunctions in the adjustment of grasping forces (Brandauer et al. 2008; Nowak et al. 2002) and even cerebellar tremor, which is suggested to be caused by missing predictive signals from the cerebellum to the motor cortex (Vilis and Hore 1980). Within its functional role on shaping and fine-tuning of movements, the cerebellum is well known to be important for the adaptation of motor patterns to changing conditions. Therefore, the internal forward models are suggested to be adapted to the new conditions by practice-dependent motor learning based on sensory prediction errors (see Bastian 2006; Tseng et al. 2007). Importantly, motor adaptation to changing conditions is constantly required in everyday life, e.g., by adapting arm control to its new dynamics when holding an object (e.g., a cup), adjustment of grip force to an empty or full bottle, or to adjust leg control when wearing heavy shoes in the winter. Even in the adaptation of motor patterns to the effects of muscle fatigue, the cerebellum is suggested to be involved (Barash et al. 1999; Golla et al. 2008). Impairments of cerebellar patients in (short-term) practice-dependent motor learning have been shown for various motor tasks including the adaptation of limb movement control to additional loads (Ilg et al. 2008; Manto et al. 1995), visuo-motor adaptation (Deuschl et al. 1996; Martin et al. 1996; Werner et al. 2009), force-field adaptation (Maschke et al. 2004; Rabe et al. 2009; Smith and Shadmehr 2005), and adaptation of gait pattern to a split-belt treadmill (Morton and Bastian 2006). In addition, it has been shown that cerebellar patients are impaired in the automaticity of recently practiced movements (Lang and Bastian 2002). However, all these experiments show the impairments in short-term motor adaptation. The capabilities of such patients to adapt or to relearn movements over a longer duration of time have not yet been examined. Thus, whether such patients have lost the ability of practice-dependent motor learning or whether they require longerduration or higher-intensity training to learn still remains an open question (Morton and Bastian 2007). Recent studies argue in favor of the latter hypothesis. They found that cerebellar patients showed motor adaptation in longer-term trials and identified different interacting adaptation processes on different timescales (Chen-Harris et al. 2008; Smith et al. 2006; Xu-Wilson et al. 2009). It was suggested in these studies that the cerebellum might be more responsible for fast adaptation. Instead, slower adaptation processes could be less dependent on the cerebellum (see section “Further Studies on Mechanisms of Motor Adaptation and Rehabilitation”).

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Animal Cerebellar Lesion Models Indicate Long-Term Adaptation and Effects of Motor Training Results from animal studies support the hypothesis that in spite of cerebellar impairments especially long-term improvements of motor functions are possible. Barash et al. (1999) studied the effects of small lesions of the oculomotor vermis which impaired the ability of monkeys to rapidly adapt saccadic eye movements. However, the capability of slow recovery from dysmetria was still preserved. Since the examined lesions were small, it is not clear, whether the long-term adaptation process was accomplished by compensation of adjacent areas in the cerebellum or from other brain areas. In addition, animal studies showed, that degeneration processes in the cerebellum could be decelerated by intensive motor training. Studies on rats and mice showed that motor training prevents or reduces cerebellar degeneration caused by alcoholic-induced degeneration disease, age and also SCA 1 (Carro et al. 2001; Larsen et al. 2000; Fryer et al. 2011). Furthermore, studies give evidence for the hypothesis that exercising demanding and new complex multi-joint movements outperforms “pure exercise” in preventing cerebellar degeneration as much as possible (Black et al. 1990; Kleim et al. 2007; Klintsova et al. 2004).

Motor Rehabilitation in Human Cerebellar Disease Since the cerebellum has been described as a key player in learning and adaptation of motor patterns, the benefit from physiotherapeutic training for patients with cerebellar lesions is still under debate. In particular for patients suffering from degenerative cerebellar disease, it has been suggested that these patients may not benefit from physiotherapy in terms of relearning movements and increasing their coordination capability (Bastian 1997; Morton and Bastian 2007). Instead, the training of compensation strategies has been proposed such as decomposing multi-joint movements into simpler movements limited to single joints. However, other authors argue that this approach leads to a complete degeneration of remaining multi-joint coordination capabilities (Panturin 1997). Relatively few clinical studies have evaluated physiotherapeutic interventions for patients with cerebellar ataxia. Furthermore, many are single case studies with different types of cerebellar disease and severity of ataxia. This heterogeneity in patient populations makes it very difficult to compare and evaluate intervention methods in existing studies. Physiotherapeutical treatment (including balance and gait training, general strengthening and the utilization of various senses) has been performed on different patient groups suffering from vestibular and vestibulocerebellar dysfunction (Brown et al. 2006). As result, the cerebellar group (consisting of patients with cerebellar degeneration as well as stroke) profited the least from the intervention (compared to patients with peripheral or central vestibulopathy or extra-cerebellar stroke).

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Although, these results further illustrate the more difficult prospects for the rehabilitation of cerebellar patients compared to other neurological diseases, some studies have shown benefits for retraining posture and balance control. Using increasingly demanding balance and gait tasks, improvements were reached for increased postural stability in clinical measures and less dependency on walking aids in everyday life (Balliet et al. 1987; Gill-Body et al. 1997). Locomotion training on treadmills with (Cernak et al. 2008; Freund and Stetts 2010) or without (Vaz et al. 2008) body-weight support has been proposed in particular for patients with more severe ataxia, who are not able to walk freely. In these studies, increased functional mobility with mobility aids has been reported. Recently, the benefits of intensive coordinative training in degenerative cerebellar disease were examined systematically in an intraindividual case–control design (Ilg et al. 2009, 2010). Sixteen patients suffering from progressive ataxia due to cerebellar degeneration (n ¼ 10) or degeneration of afferent pathways (n ¼ 6) have been tested. All patients were able to walk a distance of 10 m with or without walking aid. The strategy of physiotherapeutical intervention was to activate and demand control mechanisms for balance control and multi-joint coordination. Furthermore, the intervention trained the patients’ ability to select and use visual, somatosensory, and vestibular inputs to preserve and retrain patients’ capability for reacting to unforeseen situations and for avoiding falls as much as possible. Important principles in the motor training are (1) to train a small amount of exercises with a high amount of repetitions, (2) to include movements with relevance in all day living, and (3) to train increasingly demanding movements (from static to dynamic balance, from slow to fast movements, from single joint movements to complex multijoint coordination). These are common principles in neurorehabilitation (e.g., Platz et al. 2001; Taub et al. 1999). The physiotherapeutic program consisted of a 4-week course of intensive training with three sessions of 1 h per week. Exercises included the following categories: (1) static balance, e.g., standing on one leg; (2) dynamic balance, e.g., sidesteps, climbing stairs; (3) whole-body movements to train trunk-limb coordination (see Fig. 105.1a); (4) steps to prevent falling and falling strategies; and (5) movements to treat or prevent contracture (see for more details Ilg et al. 2009). Patients were examined four times: 8 weeks before intervention, immediately before and after the intervention period, and after 1 year for follow-up assessment. Significant improvements in motor performance and reduction of ataxia symptoms were observed in clinical scores (SARA, Scale for the assessment and rating of ataxia (Schmitz-H€ubsch et al. 2006); ICARS (Trouillas et al. 1997)) after training and were significantly sustained at follow-up assessment (see Fig. 105.1b). The natural disease progression of degenerative cerebellar ataxias is 0.6–2.5 points per year on the SARA scale (ranging from 0 to 40) depending on genotypes (Jacobi et al. 2011). This implies that the average improvement achieved by training (4.9 SARA points after intervention and 3.1 SARA points after 1 year for the cerebellar group) is equivalent to gaining back functional performance of 2 or more years of disease progression.

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Fig. 105.1 (a) Snapshot of a demanding exercise: training of dynamic balance and multi-joint coordination. (b) Group data of the clinical ataxia score SARA before training intervention (BT), after the 4 weeks training intervention (AT), and for follow-up assessment after 1 year (F1J). (c) Illustration of an experiment testing dynamic balance capacities. Patients have to compensate the perturbation of the accelerating treadmill (accelerating phase 1s) by anterior directed steps. The red and the blue characters show the same patient before (red) and after (blue) the intervention period. After intervention patients were able to compensate the perturbation in a more efficient and secure way. (d) Quantitative measurement for the body sway in the perturbation experiment for the three assessment points (Ilg et al. 2009, 2010)

In addition, increased goal attainment scores (Kiresuk et al. 1994) showed a substantial retention of training effects for individual important activities of daily life. According to their disease stage, patients selected (in interaction with the physiotherapist) subjectively relevant goals for their individual goal before the intervention. Examples for such goals were (1) walking over a distance of about 300 m without a walking aid or a helping person and (2) walking a distance of 30 m with a full cup without spilling. Five levels of goal attainment were defined. After intervention, the patients scored his or her subjective outcome. Subjective improvements after intervention and even after 1 year were on average greater than expected before intervention. These results indicated that despite a gradual decline of motor performance and gradual increase of ataxia symptoms due to progression of underlying neurodegeneration, patient benefits can be meaningful for their everyday life and persisted after 1 year. Further results of this study were as follows: (1) specific

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improvements in balance and coordination shown by quantitative movement analysis on static and dynamic balance tasks (see Fig. 105.1c, d and Ilg et al. 2009 for details) make it unlikely that the observed effects have been mediated predominantly by nonspecific mechanisms such as improved cardiovascular endurance. (2) Continuous training is crucial in particular for patients with degenerative diseases. Importantly, the degree of long-term retention depended on training: training correlated significantly with differences in SARA scores at 1 year follow-up. (3) Patients with predominant cerebellar ataxia revealed more distinct improvement than patients with afferent ataxia in most outcome measures. This discrepancy is likely caused by a loss of afferent information in these patients, which inhibits necessary inputs for adequate cerebellar processing.

Discussion Current Practice in Motor Rehabilitation Recent results of clinical studies give evidence that intensive motor training including exercises for multi-joint coordination and balance can reduce ataxia symptoms in patients suffering from degenerative cerebellar disease equivalent to gaining back functional performance of two or more years of disease progression (Ilg et al. 2010). Thereby, continuous training seems in particular crucial for patients with degenerative diseases for stabilizing improvements and should become a standard of care. However, motor rehabilitation in degenerative cerebellar disease remains a challenge and requires a careful analysis of the patient’s current motor capacities and condition in everyday life. In general, a combination of restorative and compensatory techniques may be utilized; the relative emphasis depends on the severity of cerebellar ataxia and its pattern of progression (Bastian 1997; Cassidy et al. 2009; Marsden and Harris 2011). Such compensatory techniques can include (1) replacing rapid multi-joint movements with slower movements with sequential single joints movements (Bastian 1997), (2) the rehearsal of eye movements for goal-directed stepping (Crowdy et al. 2002), or (3) the training of more cognitive strategies to adjust to different movement conditions (Taylor et al. 2010). Best evidence for beneficial interventions exist for coordination and balance training (Balliet et al. 1987; Gill-Body et al. 1997; Ilg et al. 2009), which is (1) adapted on the severity of the ataxia and (2) consists in increasingly demanding exercises for multi-joint coordination and balance. These approaches, to activate the remaining capabilities for balance and coordination, should be used as long as possible, in order to potentially decelerate the process of degeneration of functional motor capabilities (which has not yet been shown in human but only in animal studies). However, these types of exercises might be limited to stages of disease, in which the patients are able to stand without help and have also some gait capabilities. In more severe cases, in which free standing and walking is not possible anymore, treadmill training (Cernak et al. 2008; Freund and Stetts 2010;

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Vaz et al. 2008) with potential weight support may be helpful to increase walking capabilities (with the use of mobility aids) and to preserve general fitness as far as possible. The use of mobility aids is dependent on disease stage and individual preferences. Clinical observation suggests that some individuals with ataxia find light touch contact – e.g., with the use of Nordic poles – more useful as a strategy than conventional walking aids like traditional walking sticks (Cassidy et al. 2009). This strategy would also help to use and train remaining balance capacities. In more severe cases, the use of walkers for ensuring the safeness of mobility will be the first choice. Furthermore, in severe cases of upper limb ataxia, when basic activities of daily living like eating are impaired, the use of orthotics – in order to stiffen mechanically several degrees of freedom – can help to improve functional performance (Cassidy et al. 2009; Gillen 2002).

Open Questions in Cerebellar Motor Rehabilitation The above-described first positive results gained in motor rehabilitation of patients suffering from degenerative cerebellar disease raise a lot of new questions. These relate on the one hand to further clinical studies, in order to evaluate and to compare the efficiency of training approaches in larger patient populations and to find reliable predictors of training benefits. On the other hand, further research in motor adaptation is needed, in order to understand the mechanisms of improving functional performance.

Further Clinical Studies The majority of studies, which have shown improvements in motor performance, focused on ambulatory patients, which are able to walk with or without walking aid. Thus, further studies are needed to examine whether patients with more severe impairments would also benefit from physiotherapeutical training (adjusted to their impairments, e.g., for arm movements) or whether the capacity to improve motor performance relies on a specific level of residual cerebellar integrity. Motor Rehabilitation for Upper Extremities There is a clear lack of rehabilitation studies for upper limb movements in cerebellar disease. Although upper limb rehabilitation is very common in stroke patients (e.g., Platz et al. 2001; Taub et al. 2003), there are only few attempts to transfer similar approaches to cerebellar ataxia. Richards et al. (2008) applied modified constraint-induced movement therapy protocols for goal-directed arm movements to three chronic stroke patients suffering from ataxia due to extra-cerebellar lesions (basal ganglia, thalamus). Participants improved on either the Fugl-Meyer Test (Fugl-Meyer et al. 1975) or the Wolf Motor Function Test (Wolf et al. 1989) and increased their daily use of the impaired upper extremity. In principle, the concept of motor training with increasingly demanding coordination exercises could also be transferred to goal-directed upper limb movements.

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It would be highly interesting, whether an intensive and continuous training of manipulating different objects (inducing different arm dynamics) could lead to improvements in adjusting upper limb movements to changing conditions relevant for all day living. Another approach to train these adaptation capabilities could be the use of a robotic manipulandum to exercise upper limb movements with different perturbations like velocity-dependent force fields (Vergaro et al. 2011). The benefits and relevance of these approaches for patients’ all day living and the transfer to different movements has to be evaluated in clinical studies.

Long-Term Studies and Quality of Life In general, there is a fundamental need of long-term studies to evaluate the benefits of physiotherapeutical treatments in patients’ everyday life. Although there exist currently no specific instruments capable of detecting differences in the quality of life for cerebellar patients, the use of questionnaires commonly applied in chronic disease has been proposed (see Trujillo-Martin et al. 2009 for review). Two recent studies examined the application of health-related quality of life scales in degenerative cerebellar ataxia (Schmitz-H€ ubsch et al. 2010) and Friedreich Ataxia (Paulsen et al. 2010), which quantified the dimensions of mobility, usual activities, pain, depression, and self-care. Results show that in particular ataxia severity, extent of noncerebellar involvement, and the presence of depressive syndrome are the most important predictors of subjective health status for these patients (Schmitz-H€ubsch et al. 2010). Further studies have to examine in how far such scales can help to evaluate future therapeutical intervention studies in terms of the relevance for daily routine. When performing long-term studies with patients suffering from degenerative disease, one has to keep in mind that progression in degeneration is different for specific types of diseases (e.g., for SCA 1,2,3,6; see Jacobi et al. 2011). Thus, long-term benefits have to be regarded in the light of natural history studies for specific diseases. Prediction of Intervention Benefits A very important question for effective rehabilitation is to find reliable predictors for the outcome and potential benefit of physiotherapeutical motor training for individual patients. Further studies have to show whether functional ataxia scores describing the current status of ataxia (i.e., ICARS or SARA) or whether more sophisticated quantifications of neural degeneration and lesion sites using modern brain imaging methods can be exploited for a reliable outcome prediction. It was shown for cerebellar patients with focal lesions after tumor resection or stroke that lesion site is critical for motor recovery and lesions affecting the deep cerebellar nuclei are not fully compensated (Konczak et al. 2005; Schoch et al. 2006). In a similar way, involvement of deep nuclei may also be a crucial factor in motor rehabilitation for degenerative cerebellar disease. Thus, patients with a degeneration predominantly affecting the cerebellar cortex (i.e., SCA6) may benefit more (and longer in progress of degeneration) from motor rehabilitation compared to patients suffering from a degeneration predominantly of the deep nuclei (i.e., SCA3, Friedreich ataxia). Another indicator in degenerative disease based on brain imaging could be cerebellar volume of all or specific parts of the cerebellum, which has been

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shown to correlate negatively with severity of ataxia symptoms (Richter et al. 2005). Recent MRI studies deliver first evidence that the various subtypes of degenerative cerebellar disease lead to region-specific degeneration patterns in the cerebellar cortex as well in the deep nuclei (Du et al. 2010; Ying et al. 2009). Deeper knowledge in disease-specific degeneration patterns could also help to develop more efficient motor training approaches for individual patients.

Future Studies on Mechanisms of Motor Adaptation and Motor Rehabilitation Although it has been shown by quantitative movement analysis that patients can improve in specific motor behaviors like multi-joint coordination and dynamic balance (Ilg et al. 2009), there is no direct evidence that these functional improvements are related specifically to motor learning (e.g., a more efficient use of sensory signals could also cause the functional improvements on dynamic balance). Therefore, further studies are needed in order to clarify (1) whether associated changes in neural substrates could be identified either within the cerebellum and its connections or in other brain structures compensating the cerebellar deficit and (2) in which way the functional improvements are related to capabilities in motor learning.

Modern Brain Imaging Techniques Modern structural and functional brain imaging techniques are of possible help. Voxel-based morphometry (VBM) has been shown to reveal training-related changes (Draganski et al. 2004). Thus, VBM might be one method to show to what extent neural changes take place in cerebellar and extra-cerebellar motor circuits, or whether areas related to non-motor, e.g., cognitive strategies (frontal lobe), may step in. Resting state functional MRI and diffusion tensor imaging (DTI) are further options. These techniques are able to show functional connectivity and trainingrelated network alterations. For example, alterations in networks including the cerebellum have been identified, which correlate to the degrees of motor adaptation in visuomotor adaptation tasks (Albert et al. 2009; Della-Maggiore et al. 2009). The Relationship Between Short-Term Motor Adaptation and Motor Rehabilitation There are several remaining questions concerning the relationship between common motor adaptation paradigms and motor training in rehabilitation: (1) Could cerebellar patients show motor adaptation in long-term studies over weeks with high intensities? (2) Could the short-term adaptation capability be a valuable predictor of the outcome of long-term motor rehabilitation? (3) Can results be transferred from motor adaptation paradigms to optimize motor rehabilitation for these patients? Recent studies examined in more detail under which constraints cerebellar patients could show motor adaptation. Thus, it has been shown in the paradigm of force-field adaptation in goal-directed arm movements (Review in Shadmehr et al.

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2010) that patients with severe cerebellar degeneration can adapt significantly better to perturbations when induced gradually over a period of time instead of suddenly at once (Criscimagna-Hemminger et al. 2010). Based on these results, it has been hypothesized (Izawa et al. 2010) that cerebellar patients might be able to adapt motor pattern on the basis of trial-and-error learning (also known as reinforcement learning), which is especially associated with the basal ganglia (Schultz et al. 1997). Imaging studies suggest that the involvement of the cerebellum in reinforcement learning (see Swain et al. 2011) is dependent on the type of error feedback (Tanaka et al. 2004). In tasks with direct and immediate feedback, the cerebellum seems to be less involved (compared to tasks, where future rewards have to be predicted for a sequence of motor actions). Consistently, it has been shown that cerebellar patients benefit from direct cueing when performing sequence learning tasks (Spencer and Ivry 2009). Further studies following up on those methods could deliver valuable hints for optimizing motor learning exercises also in the framework of physiotherapeutical motor training.

Stimulation Techniques Very recently, studies showed first promising results in a new direction, namely, in noninvasive stimulation of the cerebellum by using transcranial direct current stimulation (tDCS) (Galea et al. 2009; Jayaram et al. 2011). The authors have shown that cerebellar tDSC can facilitate short-term visuomotor adaptation. Whether this method could have significant potential implications for patients with cerebellar disease or could be more useful to patients with damage outside the cerebellum – in order to facilitate the adaptation process initiated by the cerebellum – is not clear yet and currently under investigation. Conclusion

In conclusion, the rehabilitation in particular for degenerative cerebellar disease will remain a challenge for patients, physicians, and therapists. Recent advances in both clinical rehabilitation and research on motor adaptation in cerebellar disease will provoke further studies and hopefully lead to broader knowledge in this challenging field of motor rehabilitation and finally to an improvement of the patients’ quality of life.

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Antoni Matilla-Duen˜as, Carme Serrano, Yerko Iva´novic, Ramiro Alvarez, Pilar Latorre, and David Genı´s

Abstract

In the last decade, substantial scientific progress has enabled a better understanding of the pathogenesis of cerebellar diseases and the improvement of their diagnoses. Extensive preclinical work is expanding the possibilities of using experimental models to analyze disease-specific mechanisms and to approach candidate therapeutic strategies to create a rationale for clinical trials that might finally lead to successful treatment. At present, drug treatment of cerebellar disorders has shown limited effectiveness and current treatment is primarily

A. Matilla-Duen˜as (*) Department of Neurosciences, Basic, Translational and Molecular Neurogenetics Research Unit, Health Sciences Research Institute Germans Trias I Pujol (IGTP), Universitat Auto`noma de Barcelona, Ctra. de Can Ruti, Camı´ de les escoles s/n, 08916 Badalona (Barcelona), Spain e-mail: [email protected], [email protected] C. Serrano Neurology Service, Hospital de Martorell, 08760 Barcelona, Spain e-mail: [email protected] Y. Iva´novic Monte Alto Rehabilitation Medical Center, (Madrid), Private Practice, Calle Monte Alto 25, 28223, Madrid, Spain and National Reference Care Centre for People with Rare Diseases and Their Families–CREER–(Burgos), IMSERSO, Calle Bernardino Obrego´n, 09001 Burgos, Spain e-mail: [email protected], [email protected] R. Alvarez • P. Latorre Neurodegeneration Unit, Neurology Service, University Hospital Germans Trias i Pujol (HUGTP), Badalona (Barcelona), Spain e-mail: [email protected], [email protected] D. Genı´s Neurodegenerative Diseases Unit, University Hospital of Girona Dr. Josep Trueta, Girona, Spain e-mail: [email protected] M. Manto, D.L. Gruol, J.D. Schmahmann, N. Koibuchi, F. Rossi (eds.), 2371 Handbook of the Cerebellum and Cerebellar Disorders, DOI 10.1007/978-94-007-1333-8_106, # Springer Science+Business Media Dordrecht 2013

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supportive. Until effective and selective pharmacological treatment leading to better quality of life as well as increased survival of patients with cerebellar diseases is found, physical and sensory rehabilitation techniques are revealing effective approaches for improving the patient’s quality of life. The objective of this chapter is to provide an updated summary of the treatments currently available for cerebellar disorders, in particular for spinocerebellar ataxias, and to discuss the new emerging therapeutic strategies that are resulting from the intensive ongoing basic and translational research devoted to cerebellar diseases.

Introduction Damage to the cerebellum has been associated with a wide range of movement disorders including incoordination, reduced manual dexterity, postural instability, and gait disturbances (Manto 2008). Because ataxia is the most common neurological deficit resulting from dysfunction of the cerebellum, this chapter will mostly focus on treatment of ataxic disorders. Cerebellar ataxic movement patterns result from the impairment of the timing and duration of muscle activation or the magnitude and scaling of force production during voluntary movement, as the cerebellum is thought to be instrumental in these crucial elements of motor control. Drug treatment of cerebellar diseases has shown limited effectiveness, and therefore treatment is primarily supportive. With very few exceptions, such as in those ataxias associated with vitamin E and CoQ deficiencies, there are no disease-modifying therapies for cerebellar diseases. However, medications such as 5-hydroxytryptophan, clonazepam and others that will be discussed herein have been reported to have limited benefits in a few cerebellar conditions (Ogawa 2004; Matilla-Duen˜as et al. 2006; Ferrara et al. 2009; Manto and Marmolino 2009; Trujillo-Martin et al. 2009; Table 106.1). Very recently there have been encouraging advances in clinical ataxia research. Collaborative study groups throughout the world have developed and validated ataxia rating scales and instrumented outcome measures and have begun to rigorously define the natural history of these diseases, thus laying the foundation for well-designed clinical trials (Schmitz-Hubsch et al. 2010). Hereditary ataxias may have certain clinical features that respond very well to symptomatic medical therapy. Parkinsonism, dystonia, spasticity, urinary urgency, sleep pathology, fatigue and depression are all common in many of the different ataxia subtypes and very often respond to pharmacologic intervention as in other diseases. Much of the clinical interaction between the neurologist and the ataxia patient focus on identifying and treating these symptoms. As in the cases of infarctions, hemorrhages and neoplasms, surgical and medical treatment, radiotherapy or chemotherapy to treat the original cause of the cerebellar diseases is commonly followed by physical therapy. Treatment of the core clinical feature of these diseases – ataxia – is thus predominantly rehabilitative (reviewed in Watson 2009). The value of good physical therapy far exceeds any potential benefit from medications that a physician might prescribe to improve balance and coordination. Furthermore, speech and

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Table 106.1 Summary of the existing treatments in the spinocerebellar ataxias. Only those treatments in clinical trials or about to be tested in patients have been included Ataxia Treatment Status Conclusions/observations Autosomal recessive ataxias Friedreich’s ataxia Idebenone 5–20 mg/kg/ Completed Stabilization and possible day reduction of the left ventricular mass. Reduction of the progression of cerebellar manifestations during the early stages of the disease, but this evidence appears controversial Idebenone 500 mg/kg/ Completed No effects on cardiomiopathy or day neurological symptoms Pioglitazone Ongoing clinicaltrials.gov Deferiprone Completed Mild clinical improvement Erythropoeitin Completed Ataxia rating scales and frataxin levels improved CEPO Ongoing clinicaltrials.gov HDACis PreA study to evaluate the clinical pharmacokinetic and safety profiles will be initiated shortly Replacement strategies PreTAT-fused frataxin clinical Polyamides PreIncrease frataxin levels in vitro clinical Gene therapy PreUnder experimentation clinical Ataxias with vitamin E Vitamin E Treatment Ataxia and mental retardation are deficiency of choice reversed if treated early. In older individuals, disease progression can be halted RR-a-tocopherol Completed Stabilization of clinical symptoms with 800 mg/kg/day Ataxias with Coenzyme Q10 Treatment Slows progression of ataxia Coenzyme Q10 of choice deficiency Abetalipoproteinemia Vitamin E together with Treatment Initial treatment is crucial to a-tocopherol of choice avoid progression of the disease. Massive oral doses of atocopherol (100–150 mg/kg) are required Cerebrotendinous CDCA 15 mg/kg/day Treatment Decreases cholestanol levels xanthomatosis of choice leading to improvement of neurological symptoms Refsum’s disease Westminster-Refsum Treatment Lowers phytanic acid levels and diet of choice improves symptoms (continued)

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2374 Table 106.1 (continued) Ataxia Treatment Autosomal dominant ataxias SCA1 Lithium SCA2 NMDA antagonists, Deep brain stimulation Amantadine, dopaminergic, anticholinergic drugs SCA3 Clonazepam, Buspirone, Hydroxytryptophan, Lamotrigine, Tandospirone Amantadine, dopaminergic, anticholinergic drugs Varenicline SCA6 Acetazolamide, Gabapentin FXTAS Varenicline Memantine Dystonia Botulinum toxin Intention tremor Benzodiazepines, b-blockers, chronic thalamic stimulation Muscle cramps Magnesium, quinine, mexiletine Myoclonus Piracetam

Restless legs syndrome Saccadic intrusions Spasticity

Episodic ataxias EA1 EA2

Attack rates

Status

Conclusions/observations

Completed Clinical benefits. Adverse effects Completed Some benefits regarding ataxic symptoms Completed Alleviate tremor, bradkykinesia or dystonia Completed Some benefits regarding ataxic symptoms

Completed Alleviate tremor, bradkykinesia or dystonia Ongoing Completed Some benefits regarding ataxic symptoms Ongoing Ongoing Completed Clear benefits. Small dosage Completed Symptoms ameliorate

Completed Symptoms ameliorate

Completed Symptoms ameliorate. Also used to treat dementia/cognitive decline Dopaminergic treatment, Completed Clear benefits rotigotine, tilidine Memantine Completed Clear benefits Baclofen, memantine, Completed Clear benefits tizanadine with dopamine treatment Botulinum toxin Completed Clear benefits. Small dosage Carbamazepine, valproic Completed Clear benefits acid, ACTZ ACTZ Treatment Clear benefits. It should not be of choice prescribed to patients with liver, renal or adrenal insufficiency 4-aminopyridine, CHZ Completed Clear benefits 3,4-diaminopyridine Completed Improves down-beat nystagmus Carbazepine, sulthiame Completed Reduce the frequency of attacks, but the response is heterogeneous

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swallowing are often affected. In more severe cases, aspiration risk can be very significant and life threatening. Routine monitoring of swallowing by speech therapists, often including modified barium swallowing tests, is indicated in most patients.

Treatments for Autosomal Recessive Spinocerebellar Ataxias The heterogeneity of this group of diseases, which includes an extraordinary variety of gene mutations originating from different pathogenic mechanisms, places the task of reviewing all the potential or hypothetical future treatments beyond the scope of a single chapter. Therefore this chapter limits the review to the most common autosomal recessive ataxias together with those in which successful therapeutic options have been explored.

Friedreich’s Ataxia (FA) Biochemical investigations have revealed the role of frataxin, the protein associated with FA, in the assembly of iron–sulfur clusters (ISCs) in the mitochondrion (reviewed in Pandolfo and Pastore 2009). A GAA-triple repeat expansion in intron 1 of the frataxin (FXN) gene inhibiting frataxin expression is the most common type of mutation causing FA. As a consequence of frataxin deficiency, iron is accumulated in mitochondria leading to a loss of mitochondrial function. Cells from patients with FA become highly sensitive to oxidants causing further mitochondrial damage and respiratory chain dysfunction. Therefore, antioxidants such as idebenone, coenzyme Q10 and iron chelators such as deferroxiamine have been used in attempting to reduce oxidative stress (Pandolfo 2008; Schulz et al. 2009). Erythropoietin (EPO) is now being employed as a treatment for frataxin deficiency whereas histone-deacetylase inhibitors and gene therapy are experimental treatments currently being investigated. Idebenone is the antioxidant that has been the most widely used drug in FA treatment since the initial report of its successful use in the reduction of the left ventricular mass of three FA patients with cardiac hypertrophy (Rustin et al. 1999). In one of the first open-label trials in which nine patients were treated with 5 mg/kg/ day, cerebellar improvement was considered to be notable in mildly symptomatic patients after the first 3 months of therapy. Treatment during the early stages of the disease was found to reduce the progression of cerebellar manifestations (Artuch et al. 2002). A prospective open trial designed to study cardiac hypertrophy in FA patients found a reduction in the left ventricular mass of more than 20% in 50% of patients (Hausse et al. 2002). Another prospective study showed that idebenone did not halt the progression of ataxia, but significantly reduced the cardiac hypertrophy in six of eight patients as revealed by cardiac ultrasound studies (Buyse et al. 2003). In a 1-year, randomized, placebo-controlled trial of idebenone in 29 FA patients, significant reductions of interventricular septal thickness and left ventricular mass in the idebenone group were found (Mariotti et al. 2003). A randomized, double-blind,

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placebo-controlled trial comparing the effects of three different doses of idebenone (5, 15 and 45 mg/kg) over a 6-month period did not find significant differences between any of the employed clinometric scale total scores (Di Prospero et al. 2007). However, when wheelchair-bound patients were excluded, a significant improvement in the ICARS score was obtained suggesting a dose-related response in scales scores. Another large open-labeled prospective survey studying 104 FA patients during a median period of 5 years (88 treated with idebenone at 5 mg/kg/day) found that the total ICARS score worsened over the follow-up period in both the treated and non-treated groups (Ribai et al. 2007). The left ventricular mass index decreased in the treated group as well as the ejection fraction suggesting that idebenone may not have significantly altered the neurological progression. Even the beneficial effects of idebenone on cardiomyopathy in this study were questioned and are still currently an object of debate. In an open-labeled prospective study with 24 FA patients treated at different doses of idebenone ranging between 5 and 20 mg/kg/day and a long-term follow-up of between 3 and 5 years, differences were found between pediatric and adult patients (Pineda et al. 2008). While no changes were found in the clinical scale results and cardiac measurements of children after 5 years of follow-up, ICARS scores in adult patients increased after 3 years and cardiac echographic measurements remained stable. The authors concluded that idebenone leads to the stabilization of cardiomyopathy in both the groups albeit only pre-pubertal children obtain neurological benefit from the drug. Based on the results of this study, the age of therapy initiation would therefore appear to be an important factor. Less optimistic results were obtained in another study of a cohort of 35 patients over a 5-year period (Rinaldi et al. 2009). The authors reported that at the end of the study period the group without left ventricular hypertrophy (LVH) before treatment therapy had increased interventricular septum and posterior wall thickness while there was no change in the group with LVH before treatment. A Cochrane review concluded that no RCTs using idebenone or other drug therapy have shown significant benefits in the neurological symptoms associated with Friedreich’s ataxia (Kearney et al. 2009). Idebenone, on the other hand, showed a positive effect on left ventricular mass of the heart. In a 6-month randomized, double-blind, placebo-controlled intervention trial of 70 patients three arms of treatment, low dose idebenone, high dose and placebo, were compared (Lynch et al. 2010). Although there were differences between both placebo and treated groups favoring idebenone treatment, they were not statistically significant. The most recent reported results are obtained from the MICONOS study, a large, randomized, double-blind, placebo-controlled trial testing the efficacy and safety of the three doses of idebenone (Catena ®/Sovrima ®) and placebo over a 12-month treatment period. The primary endpoint of the study, mean change in the ICARS score from baseline, has not revealed significant differences between the active dose arms and placebo. Secondary endpoints also failed to reveal statistically significant differences between the placebo and active dose groups, and even cardiac benefit could not be proved. Although there are a few completed and ongoing studies with idebenone in FA (http://clinicaltrials.gov/) it seems clear that a significant response

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of neurological parameters to high doses of the drug will not be obtained, although an effect on cardiomyopathy might be expected in some cases. Simultaneous treatment with both coenzyme Q(10) and vitamin E in low and high doses were compared in a randomized, double-blind clinical trial in 50 patients over a 2-year period. Serum CoQ(10) and vitamin E levels, which had previously been determined to be low in these patients, reached normal values as a result of the treatment. The primary and secondary end points were not significantly different between the therapy groups. Comparison of the ICARS scores with cross-sectional data showed an overall 49% improvement. There were no differences between both the groups in the end points. The best predictor of a positive clinical response to this double therapy was low serum CoQ(10) and vitamin E levels (Cooper et al. 2008). A 2-year prospective, randomized double-blind trial of pioglitazone versus placebo is currently ongoing (http://clinicaltrials.gov/). Pioglitazone is a peroxisome proliferator-activated receptor g (PPARg) ligand that induces the expression of enzymes involved in the mitochondrial metabolism, including the superoxide dismutase. This drug appears to counteract the disabled recruitment of antioxidant enzymes in FA patients. Iron chelators such as deferiprone have been used in a short study of nine patients in which this drug successfully reduced labile iron levels in the dentate nuclei (Boddaert et al. 2007). The noted clinical improvement was mild. A larger scale study has been undertaken (Velasco-Sanchez et al. 2010). Iron chelators such as 2-pyridylcarboxaldehyde 2-thiophenecarboxyl hydrazone (PCTH) appear more effective than conventional radical scavengers in in vitro assays (Lim et al. 2008), albeit their efficacy needs to be proven in patients. After finding that recombinant human erythropoietin (rhuEPO) significantly increases frataxin expression levels in in vitro studies (Sturm et al. 2005), an open-label clinical pilot study to evaluate the safety and efficacy of rhuEPO was designed. Eight adult FA patients received 2,000 IU rhuEPO three times a week subcutaneously for 6 months. The scores in different ataxia rating scales and frataxin levels improved significantly after treatment, and the values measuring oxidative stress decreased (Boesch et al. 2007; Boesch et al. 2008). Because of the side effects of EPO on hematopoiesis and tumor growth, efforts to develop EPO derivative molecules avoiding binding the erythropoietin receptor resulted in the synthesis of carbamylated erythropoietin (CEPO). This drug has proven neuroprotective and increases the production of frataxin to the same levels as rhuEPO. A safety study on this drug is in process (http://clinicaltrials.gov/). Varenicline is a partial nicotinic receptor agonist which has been recently used in a clinical trial to investigate its effects on neurological clinical features in FA (Zesiewicz et al. 2009). However, the study with this drug was halted before completion due to concerns about its safety and insufficient evidence of efficacy in patients. Histone deacetylase inhibitors (HDACi) revert silent heterochromatin to an active chromatin conformation and therefore have been evaluated as molecules reverting gene expression. HDAC inhibition is a persistent reversible phenomenon which in theory should permit the intermittent administration of the drug. Such

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a regimen would minimize toxic side effects by reducing drug exposure while at the same time allowing sustained up-regulation of frataxin protein levels. This is an important consideration since uncommon but serious side effects had been reported with the use of other HDACi in the past. HDACi have been proven to increase frataxin protein levels in FA cells with a silent mutant frataxin gene (Herman et al. 2006; Rai et al. 2008, 2010). Among these HDACi, pimelic diphenylamide HDACi have stood out as efficient up-regulators of frataxin expression (Rai et al. 2010). A study to evaluate the pharmacokinetic and safety profiles of this new generation of HDACi will be initiated shortly in FA patients. The HIV-1 transactivator of transcription (TAT), an arginine-rich cell penetrant peptide, is being exploited in replacement therapy strategies for transducing fulllength proteins not only across the cell membrane, but also into intracellular organelles including the mitochondrion (Del Gaizo and Payne 2003; Vyas and Payne 2008; Rapoport and Lorberboum-Galski 2009). Very recently, a pre-clinical trial has proven successful in delivering synthetic TAT-fused frataxin (TATfrataxin) to the mitochondrial matrix in FA mice. After the treatment, TAT-frataxin did not induce inflammatory response in FA mice when injected chronically over 2 months, and FA mice increased their life span significantly (R.M.Payne, unpublished results). Based on the empiric observations that some polyamide compounds such as beta-alanine-linked pyrrole-imidazole polyamides bind GAA/TTC tracts with high affinity and disrupt the intramolecular DNA-associated regions, they were tested to examine their ability to increase frataxin gene transcription in cell cultures. This has proven to increase frataxin protein levels (Burnett et al. 2006). Similar effects have been obtained with pentamidine and related small molecules (Grant et al. 2006; Gottesfeld 2007). Different approaches have focused on the high capacity of the herpes simplex virus type 1 (HSV-1) amplicon vectors expressing the entire 80 kb FRDA genomic locus to successfully transduce onto FA patient frataxin-deficient fibroblasts in a FA mouse model (Gomez-Sebastian et al. 2007; Lim et al. 2007). Other innovative strategies designed to place exogenous frataxin protein into the mitochondria of patients include the intravenous administration of frataxin previously encapsulated with peptides in nanoparticles.

Ataxias with Vitamin E and Coenzyme Q10 Deficiencies The treatment of choice for the ataxia with vitamin E deficiency is lifelong highdose oral vitamin E supplementation. Some symptoms including ataxia and mental deterioration can be reversed if treatment is initiated early in the disease process. In older individuals, disease progression can be stopped, but deficits in proprioception and gait unsteadiness generally remain (Gabsi et al. 2001; Mariotti et al. 2004). With treatment, plasma vitamin E concentrations can become normal. No therapeutic studies have been performed on a large cohort to determine optimal dosage and evaluate outcomes. Reported doses of vitamin E range from 800 mg to 1,500 mg or 40 mg/kg body weight in children.

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The used vitamin E preparations are the chemically manufactured racemic form, all-rac-a-tocopherol acetate or the naturally occurring form, RRR-a-tocopherol. It is not currently known whether affected individuals should be treated with all-raca-tocopherol acetate or with RRR-a-tocopherol. It is known that alpha-tocopherol transfer protein (alpha-TTP) stereoselectively binds and transports 2R-a-tocopherols. For some ATTP mutations, this stereoselective binding capacity is lost and affected individuals cannot discriminate between RRR- and SRR-a-tocopherol (Traber et al. 1993; Cavalier et al. 1998). In this instance, affected individuals would also be able to incorporate non-2R-a-tocopherol stereoisomers into their bodies if they were supplemented with all-rac-a-tocopherol. Since the potential adverse effects of the synthetic stereoisomers have not been studied in detail, it seems appropriate to treat with RRR-a-tocopherol, despite the higher cost. Several studies have reported the stabilization of the clinical symptoms and even some improvement with twice daily doses of 800 mg of RRR-a-tocopherol in patients presenting ataxia with vitamin E deficiency (Amiel et al. 1995; Yokota et al. 1997; Martinello et al. 1998). Furthermore, fat-enriched meals are recommended in these patients. Oral antioxidant therapy with CoQ10 has slowed the progression of ataxia in patients who are specifically deficient in those components (Hirano et al. 2006; Pineda et al. 2010).

Abetalipoproteinemia The disease in patients with abetalipoproteinemia is directly related to vitamin E deficiency (reviewed in Kayden 2001). Substitution therapy with vitamin E presents complex problems due to severe intestinal misabsorption and requires a close dietetic control to take into account all the aspects involved in the management. In this disease, the lipoproteins transporting a-tocopherol are missing. An early onset treatment with vitamin E is crucial to avoid or halt the progression of the neurological symptoms. Initial parenteral administration of vitamin E is recommended followed by massive oral doses of a-tocopherol at 100–150 mg/kg. Plasma levels of vitamin E often fail to reflect the whole body content of vitamin E and the adequacy of vitamin replacement may be difficult to gauge from serum concentrations. However, this dose has been shown to improve the overall neurological state of the patient if treatment is started early (Zamel et al. 2008). Vitamin A, D and K and other dietary supplements must be given. The disease requires lifelong controls to avoid or minimize secondary nervous system damage.

Cerebrotendinous Xanthomatosis (CTX) Cerebrotendinous xanthomatosis (CTX) is a lipid storage disease in which several bile alcohols, particularly cholestanol, are accumulated. Treatment with chenodeoxycholic acid (CDCA) decreases the cholestanol levels and this leads to

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the improvement of cognition, psychiatric, motor clinical and neurophysiological parameters (Berginer et al. 1984). As observed with other metabolic ataxias, earlier treatment usually results in better results. The recommended doses are 15 mg/kg/ day in three daily doses. Other treatments include the use of pravastin, a 3-hydroxy3-methylglutaryl (HMG)-CoA reductase inhibitor or a combination of both chenodeoxycholic acid and pravastin (Salen et al. 1994; Verrips et al. 1999). More aggressive treatments of CTX with LDL-apheresis have shown contradictory results (Ito et al. 2003; Dotti et al. 2004).

Refsum’s Disease Refsum’s disease features are directly related to the progressive deposition of phytanic acid in different tissues. Thus, the main objective for treatment has been to lower the phytanic acid levels mainly by providing the Westminster-Refsum diet (Baldwin et al. 2010). As patients with Refsum’s disease may suffer with severe clinical exacerbations, therapeutic lipapheresis has been considered in these conditions (Gutsche et al. 1996; Weinstein 1999). Liver cell transplantation is under consideration as a treatment option in Refsum’s disease patients (Sokal et al. 2003; Najimi and Sokal 2005).

Treatments for Autosomal Dominant Spinocerebellar Ataxias There are currently no known effective pharmacologic treatments to reverse or even substantially reduce motor disability caused by cerebellar degeneration in most of the autosomal dominant spinocerebellar ataxias (SCAs) or related cerebellar disorders, although some benefits on ataxic and non-ataxic symptoms have been reported in a few therapeutic clinical trials (extensively reviewed in Ogawa 2004; MatillaDuen˜as et al. 2006; Manto and Marmolino 2009; Trujillo-Martin et al. 2009). Some benefits regarding ataxic symptoms have been reported with acetazolamide and gabapentin in SCA6 (Nakamura et al. 2009), 5-hydroxytryptophan, clonazepam, buspirone or tansodpirone, sulfamethoxazole/trimethoprim or lamotrigine in SCA3, NMDA modulators or antagonists, and deep brain stimulation in SCA2 with tremor or FXTAS (Ferrara et al. 2009). Amantadine, dopaminergic and anticholinergic drugs have been used to alleviate tremor, bradykinesia or dystonia in SCA2 and SCA3 (Botez et al. 1991; Tuite et al. 1995; Buhmann et al. 2003). Varenicline is currently being tested in SCA3 (Zesiewicz and Sullivan 2008) and FXTAS. Restless legs and periodic leg movements in sleep usually respond to dopaminergic treatment, tilidine or rotigotine (Schols et al. 1998; Hening et al. 2010; Zintzaras et al. 2010). Spasticity in SCAs is effectively treated with the GABA analogue baclofen, tizanadine or memantine when combined with dopaminergic treatment. In selected cases where other treatments have failed, botulinum toxin has been successfully used to treat dystonia and spasticity in SCA3 (Freeman and Wszolek 2005), although caution and small dosage are recommended since unusually severe

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and long-lasting muscular atrophy occurs in some SCA3 patients with this treatment because of subclinical involvement of motor neurons in the anterior horn in the degenerative process. Intention tremor has been ameliorated with benzodiazepines, b-blockers or chronic thalamic stimulation. Muscle cramps, which are often present at the onset of the condition in SCAs 2, 3, 7, and DRPLA, are alleviated with magnesium, quinine or mexiletine (Kanai et al. 2003). Piracetam have been used to treat myoclonus and/or dementia/cognitive decline (De Rosa et al. 2006; Kanai et al. 2007; Ince Gunal et al. 2008). In spite of the lack of effectiveness in the treatment of ataxia symptoms in most SCAs, treatment in some spinocerebellar ataxias has proven successful. Furthermore, most autoimmune cerebellar ataxias, such as anti-glutamic acid decarboxylase (GAD)-antibody-positive cerebellar ataxia and gluten ataxia, have proven to be treatable with intravenous immunoglobulin administration (Lock et al. 2006; Nanri et al. 2009). In the remaining ataxias, physiotherapy is currently being used as an effective treatment alternative. Ataxia improves with daily autonomous training of gait and stance in combination with physiotherapy. Other neurological symptoms such as dysarthria and dysphagia warrant logopedic treatment to maintain the ability to communicate and to prevent pneumonia from aspiration. A clinical trial with the aim of assessing the safety, tolerability and the effects of lithium in SCA1 has recently been completed, and patients are being recruited to assess lithium carbonate therapy in SCA2 and SCA3. Albeit there are clinical benefits with lithium treatment, common side effects include muscle tremors, twitching, ataxia and hypothyroidism. Long-term use of lithium has been linked to hyperparathyroidism, hypercalcemia (bone loss), hypertension, kidney damage, nephrogenic diabetes insipidus (polyuria and polydipsia), seizures, and weight gain (Tredget et al. 2010). Although lithium or a bioactive analogue may have promising potential to benefit ataxia patients, clinical and biological responses to a range of doses throughout an extended time period need to be carefully evaluated and monitored in any forthcoming clinical trial. Other ongoing clinical trials in SCAs include the NMDA receptor antagonist memantine in fragile X tremor and ataxia syndrome (FXTAS). Memantine has been successfully used to treat saccadic intrusions in spinocerebellar ataxias (Serra et al. 2008).

Treatments for Episodic Ataxias Several different drugs are reported to improve symptoms in EA1 and EA2, but so far there have been no controlled studies documenting or comparing efficacy of these different drugs. Carbamazepine, valproic acid and acetazolamide (ACTZ) have proven effective for EA1 (Eunson et al. 2000; Klein et al. 2004); and ACTZ (Griggs et al. 1978), 4-aminopyridine (Strupp et al. 2004; Strupp et al. 2008) and chlorzoxazone (CHZ) (Alvina and Khodakhah 2010a) have been effective in EA2 cases. The response to acetazolamide is often dramatic in EA2 (Griggs et al. 1978; Jen et al. 2004), and is considered the treatment of choice. ACTZ should not be

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prescribed to individuals with liver, renal or adrenal insufficiency. Acetazolamide, a carbonic-anhydrase (CA) inhibitor, may reduce the frequency and severity of the attacks in some but not all affected individuals with episodic ataxias. Chronic treatment with ACTZ may result in side effects including paresthesias, rash and formation of renal calculi. Antiepileptic drugs (AEDs) such as carbamazepine may significantly reduce the frequency of the attacks in responsive individuals; however, the response is heterogeneous as some individuals are particularly resistant to drugs (Eunson et al. 2000). Anticonvulsant drugs such as sulthiame may reduce the attack rates. During this treatment, abortive attacks were still noticed lasting a few seconds and troublesome side effects were paresthesias and intermittent carpal spasm (Holtmann et al. 2002). The potassium channel blocker 4-aminopyridine has been found to be effective in stopping attacks in patients with EA2 (Strupp et al. 2004; Alvina and Khodakhah 2010b). Furthermore, 3,4-diaminopyridine was demonstrated in a placebocontrolled study to improve down-beat nystagmus, which is often observed in patients with EA2 (Strupp et al. 2003).

Emerging Therapeutic Strategies In the last decade, intensive scientific research has been devoted to identify molecular pathways underlying cerebellar neurodegeneration (Matilla-Duen˜as et al. 2010) with the aims of discovering and establishing effective and selective therapeutic strategies to treat cerebellar diseases. Among them, a few innovative approaches yielding promising results are being investigated at the preclinical and in some cases at the clinical level including the use of RNA interference (RNAi) aiming to inhibit the expression of mutated polyglutamine-proteins in those SCAs caused by expanded polyglutamine mutations, prevention of protein misfolding and aggregation by over-expression of chaperones and by pharmacological treatments, and the regulation of gene expression by treatment with histone deacetylase inhibitors (HDACi). Intracerebellar injection of vectors expressing short hairpin RNAs was shown to selectively decrease the expression of mutant proteins and profoundly improve motor coordination, restore cerebellar morphology and prevent the characteristic intranuclear aggregated inclusions in Purkinje cells in SCA1 transgenic mice (Xia et al. 2004). While these results show that RNAi therapy improves cellular and behavioral characteristics in pre-clinical trials, its application in patients to protect or even reverse disease phenotypes shall be delayed until proper toxicity tests are assessed. Another target, molecular chaperones provide a first line of defense against misfolded, aggregation-prone proteins. Many studies have analyzed the effects of chaperone over-expression on inclusion body formation and toxicity of pathogenic polyQ fragments in cell culture, and it is clear that overexpression of molecular chaperones might prove beneficial for the treatment of cerebellar diseases (Muchowski and Wacker 2005). They prevent inappropriate

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interactions within and between non-native polypeptides, enhance the efficiency of de novo protein folding, and promote the refolding of proteins that have become misfolded as a result of the mutations and cellular stress (Chan et al. 2000). Chemical and molecular chaperones might also prevent toxicity by blocking inappropriate protein interactions, by facilitating disease protein degradation or sequestration or by blocking downstream signaling events leading to neuronal dysfunction and apoptosis. The first proof of concept studies supporting such idea was performed using Congo Red, thioflavine S, chrysamine G and Direct Fast yellow which proved to be effective in suppressing molecular aggregation in vitro and in vivo and ameliorate symptoms (Heiser et al. 2000; Sanchez et al. 2003), albeit their efficacy in vivo is limited by their variable abilities to cross the blood–brain barrier and bioviability such that proper pharmacologic analogues may need to be developed for further clinical considerations. Other low molecular mass chemical chaperones, such as the organic solvent dimethylsulfoxide (DMSO) and the cellular osmolytes glycerol, trimethylamine n-oxide and trehalose, appear to ameliorate cell death triggered by mutant ataxin-3 by increasing its stability in their native conformation (Yoshida et al. 2002). Trehalose was identified in an in vitro screen for inhibitors of polyglutamine aggregation, and its administration reduces brain and cerebellar atrophy, improves motor dysfunction and extends the lifespan of mice resembling the polyglutamine disorder Huntington’s disease (Tanaka et al. 2004). In vitro experiments suggest that the beneficial effects of trehalose result from its ability to bind and stabilize polyglutamine-containing proteins. More recently, a new generation of small chemical compounds that directly target polyQ aggregation without significant cytotoxicity have been identified in high-throughput screens using cell-free assays or by targeting cellular pathways (Heiser et al. 2002; Zhang et al. 2005). These compounds decrease molecular aggregation in cultured cells and brain slices and can rescue neurodegeneration in a drosophila model, although no effect was detected in mouse models possibly due to bioviability issues of the compounds. By a different mechanism, a small molecule that acts as a co-inducer of the heat shock response by prolonging the activity of heat-shock transcription factor HSF1, arimoclomol, significantly improves behavioral phenotypes, prevents neuronal loss, extends survival rates and delays disease progression in a mouse model of neurodegeneration (Kieran et al. 2004). Similarly, activation of heat-shock responses with geldanamycin inhibits aggregation and prevents cell death (Rimoldi et al. 2001). This suggests that pharmacological activation of the heat shock response may therefore be an effective therapeutic approach for treating neurodegenerative diseases. However, excessive upregulation of chaperones might lead to undesirable side effects, such as alterations in cell cycle regulation and cancer (Mosser and Morimoto 2004). Therefore, a delicate balance of chaperones will likely be required for a beneficial neuroprotective effect. For instance, chemical or molecular chaperones, used in combination with a pharmacological agent that up-regulates the synthesis of molecular chaperones, might be a valid therapeutic approach for treating spinocerebellar ataxias caused by polyglutamine expansions. Aggregate formation has also been successfully targeted with inhibitors of transglutaminase, such as

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cystamine, which reduces apoptotic cell death and alleviates disease symptoms (Dedeoglu et al. 2002; Karpuj et al. 2002). Compounds directly targeting mitochondrial function such as coenzyme Q10 (Shults 2003); creatine (Ryu et al. 2005) and tauroursodeoxycholic acid (TUDCA) (Keene et al. 2002); or autophagy, such as the mTor inhibitor rapamycin and various analogous (Ravikumar et al. 2004), have proven effective at reducing cellular toxicity in animal models, and are currently being tested in clinical trials in a few ataxia subtypes (Menzies and Rubinsztein 2010). Caspase activation, which usually precedes neuronal cell death, has been targeted by inhibiting their expression, recruitment and consequent activation onto “apoptosome-like signaling structures” or by enzymatic inhibitors all of which include minocycline, zVADfmk, CrmA, FADD DN and cystamine (Ona et al. 1999; Sanchez et al. 1999; Lesort et al. 2003). In general, the inhibitors of the different caspases have been shown to decrease microglia activation, prevent disease progression, delay onset of symptoms, enhance inclusion clearance and extend survival rates in several mouse and cell models of neurodegeneration (Ona et al. 1999; Chen et al. 2000; Lesort et al. 2003). Other agents promoting the clearance of mutant proteins in the CNS or which are Ca2+ signaling blockers and stabilizers, such as specific inhibitors of the NR2B-subunit of N-methyl-D-aspartate glutamate receptors, blockers/antagonists of metabotropic glutamate receptor mGluR5 and inositol 1,4,5-trisphosphate receptor InsP3R1 such as remacemide; intracellular Ca2+ stabilizers such as dantrolene; dopamine stabilizers such as mermaid-ACR-16; dopamine depleters and agents inducing anti-excitotoxic effects such as riluzole; or agents which alleviate cognitive components such as horizon-dimebon; appear to be at least partially beneficial for the treatment of some neurological symptoms in spinocerebellar ataxias (Gauthier 2009; Liu et al. 2009; Mestre et al. 2009). A recent clinical trial with riluzole showed a reduction of the ICARS score in patients with a wide range of cerebellar disorders (Ristori et al. 2010). Neuroprotective drugs such as olesoxime have proven to increase microtubule dynamics, reestablish neuritic outgrowth, improve myelination and prevent apoptotic factor release and oxidative stress in amyotrophic lateral sclerosis (ALS) and spinal muscular atrophy (SMA) (Bordet et al. 2007), and are potential drugs to be tested in ataxias. Inhibition of potassium channels with 3,4-diaminopyridine has proven efficient in normalizing motor behaviors in young SCA1 mice and in restoring normal Purkinje cell volume and dendrite spine density and the molecular layer thickness in older SCA1 mice. Aminopyridines, such as fampridine and diaminopyridine, increase PC excitability and are also efficient for treating down-beat nystagmus (Strupp et al. 2008; Alvina and Khodakhah 2010b; Tsunemi et al. 2010). Peroxisome biogenesis dysregulation has been identified as the underlying causative deficits in some cerebellar diseases in which bile acid supplements and dietary restriction of phytanic acid are indicative (Regal et al. 2010), and therefore treatment targeting this pathway has the potential of being explored. The roles that some proteins implicated in cerebellar diseases play in transcription and, more importantly, the effects mediated by some of their co-transcriptional regulators in the suppression of cytotoxicity are being used as targets to modulate

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the pathological effects, thus opening the path for new therapeutic strategies for treating some spinocerebellar conditions. Recent progress in histone deacetylase (HDAC) research has made possible the development of inhibitors for specific HDAC family proteins and these compounds could prove effective candidates for the treatment of spinocerebellar ataxias (Dokmanovic and Marks 2005; Thomas et al. 2008). Neuroprotective and neurorestoration strategies addressing specific bioenergetic defects might hold particular promise in the treatment of spinocerebellar conditions. Drugs, such as rasagiline, a selective irreversible monoamine oxidase B inhibitor, have been shown efficient in protecting neuronal cells against apoptosis through induction of the pro-survival Bcl-2 protein and neurotrophic factors providing an experimental rationale for rasagiline as a disease-modifying molecule (Naoi et al. 2009). Rasagiline is expected to enter a few phase 3 clinical trials shortly. Recent alterations of the insulin growth factor (IGF-1) pathway have been reported to be implicated in SCA1, SCA3 and SCA7 (Gatchel et al. 2008; Saute et al. 2010), suggesting that in vivo neuroprotection exerted by IGF-1 potentially through the PP2A-regulated PI3K/Akt signaling pathway, could potentially be used to halt cerebellar neurodegeneration (Fernandez et al. 2005; Leinninger and Feldman 2005). Clinical trials with IGF-1 on AT and SCA3 patients are underway. Gene therapy and stem cell and grafting approaches are being experimentally considered for treating spinocerebellar neurodegenerations (Chintawar et al. 2009; Erceg et al. 2010; Louboutin et al. 2010). Delivery of proteins or compounds by viral vectors onto the cerebellum represents one such gene therapeutic approach (Louboutin et al. 2010). Vectors used are capable of transducing neurons and microglia very effectively and thus can be used for gene delivery targeting the cerebellum in vivo. Neural cell replacement therapies are based on the idea that neurological functions lost during neurodegeneration could be improved by introducing new cells that can form appropriate connections and replace the function of lost neurons. This cell replacement therapeutic strategy, although potentially effective, is still in early experimental stages (Erceg et al. 2010), since the use and the process for reprogramming human somatic cells from accessible tissues, such as skin or blood, to generate functional “disease- and patient-specific” neurons from embryonic-like induced pluripotent stem cells (iPSCs) present several technical challenges (Saha and Jaenisch 2009; Tenzen et al. 2010). Since neurogenesis does occur in the adult nervous system, another approach is based on the stimulation of endogenous stem cells in the brain, cerebellum or spinal cord to generate new neurons. Studies to understand the molecular determinants and cues to stimulate endogenous stem cells are underway (Gage 2002). A recent study by Lee and colleagues suggested slowed progression of patients presenting the cerebellar subtype (MSA-C) of multiple systemic atrophy who had been treated with mesenchymal stem cell grafts (Lee et al. 2008). Although promising and further preclinical work is necessary to define the molecular mechanisms underlying these effects, one is only starting to learn the potential and challenges of these emerging therapies, especially their efficacy in treating human cerebellar neurodegeneration.

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Physical Therapy in Cerebellar Diseases The cerebellum integrates sensory input, mainly proprioceptive and vestibular, with voluntary motor action to assure coordinated and automated timing, duration and amplitude of muscle activity in normal movement. It guarantees equilibrium and vestibular-oculomotor control (midline cerebellar structures), accurate limb movement (cerebellar hemispheres and outflow tracts) and modulated speech (paramedian structures). It also plays a role in motor learning. Cerebellar damage typically results in varying degrees of instability of stance and gait, clumsy target maneuvers, slowed alternating movements, postural or action tremor of trunk or limbs, decreased muscle tone, slurred speech, dizziness, and nystagmus or saccade inaccuracies among other oculomotor signs. In addition, bladder/sphincter dysynergia causes frequent or urgent incontinence. SCA patients may suffer all symptoms or just some of them but unfortunately they will be progressive in most patients. Since few pharmacological options are available, most treatments rely heavily on rehabilitation therapy including exercise/physical therapy programs and speech and swallow evaluation and training. However, the cerebellum is known to play a crucial role in both motor control and motor learning; therefore, the benefit of physiotherapeutic training has been long time under dispute for patients with degenerative ataxia. In this regard, impairment of cerebellar patients in practice-dependent motor learning has been shown for various motor tasks (Maschke et al. 2004). Additionally, most of the research has been done with case studies or case series with heterogeneous populations, interventions and outcomes (Martin et al. 2009) and no information regarding the long-term effectiveness of physiotherapy is available. Nevertheless, rehabilitation programs clearly improve quality of life and motor performance and reduces ataxia symptoms in SCA patients (Class III evidence) (Ilg et al. 2009), however, and because of the low prevalence of these diseases, no double-blind, randomized, controlled trials have been performed to demonstrate the actual value of such interventions and there is insufficient evidence to support the efficacy of any specific therapy. Based on the principle that symptomatic treatments can be useful independent of the etiology of the problem, much of the work done with Friedreich’s ataxia patients and much of the progress achieved in this disease is currently applied to treat SCA patients.

Physical Therapy Examination A comprehensive natural history is a critical feature when examining SCA patients since these diseases can involve multiple systems, thus data about previous interventions and surgeries should be collected and cardiovascular and musculoskeletal systems carefully reviewed (Maring and Croarkin 2007). Additionally, gaining insights into the psychosocial factors is essential in the interview process since the progressive nature of the symptoms could subjectively influence an individual’s perception of his or her quality of life to various degrees (D’Ambrosio et al. 1987).

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The examination should consist of a complete history, a throughout review of the systems, and the implementation of the best available tests and measures to describe patients’ impairment and functional limitations. In routine clinical settings, traditional measurements of symmetry, range of motion and muscle strength are good indicators of specific impairments of the musculoskeletal system. Cranial nerves should also be tested for signs of impairment of ocular movements, acuity and visual field deficits, hearing loss, dysarthria and dysphagia. A complete test of sensory system is recommended since sensory neuropathy may be present and may contribute to the ataxia symptoms (Perlman 2004; Maring and Croarkin 2007). Finally, testing velocity and the independence of the gait are easy to achieve and represent important functional measurements in SCA patients. Such a complete physical therapy exam would eventually set the risk of falling and would prevent accompanying injuries and erosion of self-confidence (Perlman 2004). Composite rating scales have been proposed in order to improve reliability and validity of performance measures including the International Cooperative Ataxia Rating Scale (ICARS), the Ataxia Clinical Rating Scale, the Ataxia Functional Composite Scale, the Brief Ataxia Rating Scale, the Functional Ataxia Scoring Scale, the Inherited Clinical Rating Scale, the Northwestern University Disability Scale and the Scale for the Assessment and Rating of Ataxia (SARA). All of them show good interrater and test–retest reliabilities. Among them, only the International Cooperative Ataxia Rating Scale, the Ataxia Clinical Rating Scale, and the Inherited Ataxia Clinical Rating Scale demonstrated a good relationship between score and disease duration (Maring and Croarkin 2007). ICARS is the more widely used clinical scale up to date and it has been correlated with cerebellar volume measures in patients with pure cerebellar degeneration (Richter et al. 2005). However, internal validity is unclear and newer scales such as SARA can be completed faster and may have better construct validity (Schmitz-Hubsch et al. 2006). Other tools reported in single cases follow-up or cohort studies include the Berg Balance Test, the timed unsupported stance test, the Functional Ambulatory Category (FAC) test, the 10-m walk test, the Outpatient Physical Therapy Improvement in Movement Assessment Log (OPTIMAL), the transverse abdominal thickness, and the kinematic analysis and isometric endurance (Maring and Croarkin 2007; Freund and Stetts 2010). Recently, quantitative movement analysis of the gait, and static and dynamic balance tasks have revealed a specific behavior in patients with degenerative ataxia and intensive coordinative training. Essentially, patients with cerebellar ataxia showed significant improvement in intralimb coordination, balance control in gait and balance tasks as opposed to patients with afferent ataxia.

Physical Therapy Intervention The aim of physical therapy is to maintain the individual’s independence in all environmental contexts for as long as possible. The physical therapist will contribute to educate patients and family members about the effects of the disease on

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function of life style, potential interventions and realistic expectations about them. However, there is little evidence regarding specific physical therapy interventions in these patients, therefore programs shown to be beneficial in other patient populations with ataxia could be reasonably recommended (Sliwa et al. 1994; Perlmutter and Gregory 2003; Harris-Love et al. 2004). Those programs include aerobic fitness, maintenance of biomechanical alignment and counseling for assistive or adaptive devices which would preserve independence of mobility. Most authors agree that the main working goals in physical therapy are to develop strategies to optimize sensorial information, to improve balance in stance by postural reaction and postural stabilization, to develop strategies for an independent gait, to improve the quality and control of movement in different body postures, to exercise against resistance to improve hypotonia as well as to adjust motor control, to calibrate the motor control of speech and to improve coordination. Motor coordination can be trained using the Frenkel’s method (Vaz et al. 2008; Martin et al. 2009). Essentially, Heinrich Sebastian Frenkel designed a method to improve motor control through repetitive exercises. In general, treatment is recommend early in the course of the disease, the patient should start with easy and wide exercises and once they are perfectly performed, the next level of complexity would be recommended. All exercises should be performed with open and closed eyes, fast movements should precede the slow ones and proximal joints and trunk should be approached from the beginning. A report of progression should be registered. Frenkel’s method should be complemented with a Bobath concept approach to physiotherapy. The Bobath concept incorporates a bio-psycho-social approach and is based upon recovery as opposed to compensation (Graham et al. 2009). The main principles are: (1) human motor behavior is based upon continuous interaction between the individual, the environment and the task; (2) the individual focuses on the goal rather than the specific movement in the acquisition of motor skills; and (3) learning and adaptation of motor skill involve a process associated with practice and experience. Contemporary practice in the Bobath concept utilizes a problemsolving approach to the individual’s clinical presentation and personal goals. Treatment guides the individual toward efficient movement strategies for task performance. The method in Bobath concept focuses particularly in two interdependent aspects: the integration of postural control and task performance and the control of selective movement for the production of coordinated sequences of movement. Intervention is directed at analyzing and optimizing all factors contributing to efficient motor control. The Bobath concept also seeks to utilize appropriate sensory input to influence postural control and the internal representation of a postural body schema. One of the main strategies for improving postural control in relation to gravity and the environment is the alignment of body segments in relation to each other and the base of support. Selective movement and movement patterns will be accessed by facilitating task-specific patterns of muscle activation and the therapist aims to utilize afferent input to re-educate the internal reference systems to enable the patient to have more movement choices and greater efficiency of movement.

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In addition to both the Frenkel method and Bobath concept, single case reports have shown the benefit of trunk stabilization training and loco motor training using body-weight support on a treadmill (Cernak et al. 2008; Freund and Stetts 2010). In conclusion, individualized physical therapy, including traditional and biopsycho-social methodology, is proposed as one of the main symptomatic treatments for SCA patients. Therapy will have reasonable goals since small steps may represent a great motivation and may increase adherence to treatment leading to a real and sustainable change in patients’ lives and their families.

Concluding Remarks and Future Directions As with other cerebellar diseases, the spinocerebellar ataxias are devastating neurological diseases for which currently there are no effective and selective pharmacological treatments available that reverse or even substantially reduce motor disability caused by the cerebellar neurodegeneration. The recent progress on the understanding of cerebellar diseases has been possible through the combined efforts of worldwide international academic networks. However, further experimental and clinical research is needed to further understand their pathogenesis, validate and define the role of the updated clinical diagnostic criteria, and to enhance the assessment of the disease. This research will also help to develop novel supportive neuroimaging methods and other clinical investigations that might improve the diagnostic precision and facilitate early diagnosis and treatment. Importantly, more effort is necessary to define disease-modifying therapeutic strategies. Currently, physical therapy is the sole form of intervention that can improve walking ataxia in affected individuals and effectiveness of physiotherapy for adults with cerebellar dysfunction is currently under assessment (reviewed in Watson 2009). A more recent study where patients with variable forms of cerebellar degenerative disease were subjected to “intensive coordinative training” showed improvement in the ataxia and balance clinical scales, indicating that rehabilitation may be of real benefit to ataxic individuals (Ilg et al. 2009). Similarly, in a specific rehabilitation program including foot sensory stimulation, and balance and gait training, 24 ataxic patients with clinically defined sensory ataxia improved their balance with better results in dynamic conditions (Missaoui and Thoumie 2009). These studies are of particular interest because they showed how individuals with cerebellar damage can learn to improve their movements, recover the control of their balance and proprioceptive contributions enabling them to achieve personally meaningful goals in everyday life after proper training. Until effective and selective pharmacological treatment which ultimately should lead to better quality of life and increased survival of patients with cerebellar diseases, physical and sensory rehabilitation are meanwhile revealing effective approaches for improving the patient’s quality of life. Taken together, all the data resulting from the most recent intensive research highlight that providing effective treatments to ataxia patients is no longer an utopia, but it is possible in the foreseeable future.

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Acknowledgments Dr. Ivelisse Sanchez’s helpful comments and suggestions are kindly acknowledged. Dr. Antoni Matilla’s scientific research on ataxias is funded by the Spanish Ministry of Science and Innovation (BFU2008-00527/BMC), the Carlos III Health Institute (CP08/00027), the Latin American Science and Technology Development Programme (CYTED) (210RT0390), the European Commission (EUROSCA project, LHSM-CT-2004503304), and the Fundacio´ de la Marato´ de TV3 (Televisio´ de Catalunya). We are indebted to the Spanish Ataxia Association (FEDAES), the Spanish Federation for Rare Diseases (FEDER), and the ataxia patients for their continuous support and motivation. Antoni Matilla is a Miguel Servet Investigator in Neurosciences of the Spanish National Health System.

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Index

A ABP. See AMPAR-binding protein (ABP) Abscess, 2029, 2031 Acadm, 132 Accessory motor features, 2155–2157 Accn1, 139 Acetazolamide, 1234, 1555 3-acetylpyridine (3-AP), 267, 370–371 Achaete/scute like 1, 24 Actinopterygian fish, 1465, 1476 Action potential, 1027–1030, 1032–1034 Action sequences, 1704 Action tremor, 1611, 1612 Activation voltage, 1551, 1552, 1556 Activities of daily living (ADL), 1790–1792, 1795 Acyl-CoA dehydrogenase, 132 AD. See Alzheimer’s disease (AD) Adaptive filter, 1258, 1265, 1270, 1272, 1316 ADCA. See Autosomal dominant cerebellar ataxia (ADCA) Adenosine, 947–949, 952, 956–966 ADHD. See Attention-deficit hyperactivity disorder(ADHD) Adiadochokinesia, 1600, 1610, 1671 ADL. See Activities of daily living (ADL) Adolase C, 440, 444–448, 454–459 ADP. See After depolarizing potential (ADP) Adrenal steroid hormone, 330–331 Adrenergic receptors, 903–905, 906, 908–910, 1533 Adult neurogenesis, 1455–1456 Affective, 735–738, 746–748, 755 Affective processing, 1701 After depolarization, 1062, 1063 After depolarizing potential (ADP), 1030 After hyperpolarization, 1062–1063 After hyperpolarizing potential (AHP), 1032 2-AG. See 2-Arachidonoylglycerol (2-AG) Age at lesion effect, 1581, 1585

Agenesis, 1858 Age-related decline, 909 Aggregation, 2211–2213, 2232, 2240, 2251 Aging, 908–910 Agrammatic speech, 1723, 1724 Agrammatism, 1711 AHP. See After hyperpolarizing potential (AHP) AICAs. See Anterior inferior cerebellar arteries (AICAs) Aifm1, 137 AIS. See Axon initial segment (AIS) Akt2, 132, 134, 137 Alcohol, 108, 2081, 2086, 2090, 2093, 2107 Alcoholism, 713, 726 Aldolase C, 400 Allen brain atlas, 133, 135, 139 Allopregnanolone, 997–999, 1001, 1003, 1005–1007 Alzheimer’s disease (AD), 2157 Amino acid neurotransmitters, 1508 Aminopyridines, 2351–2353 Amos, 26 AMPA, 835, 836, 854, 856, 857, 870, 1069, 1070 AMPAR-binding protein (ABP), 864–865 AMPA receptors, 797–798, 801, 814–819, 822 Amphibian, 1430, 1431 Amygdala, 1181, 1182, 1774, 1775 Analysis-synthesis filter, 1319–1322, 1330 Anandamide, 929–932 Anatomy, 1774 Androgen receptor (AR), 332 Androgens, 320, 331, 332 Aneurysms, 679, 680, 691–698, 703, 704 Angiography, conventional, 1964, 1965, 1972, 1975, 1976 Animal experimentation, 1428, 1436 Animal model, 1564, 1571, 1572, 1575, 1576 Ankyrin, 245 2397

2398 Anterior and posterior interpositus, 1719 Anterior inferior cerebellar arteries (AICAs), 345, 350, 352, 681, 1629, 1960, 1964–1970 Anterior interposed nucleus (AIN), 379, 392, 397, 398, 401, 402, 407, 413, 417, 421, 422, 499, 506–507 Anterior intraparietal area (AIP), 552, 556 Anterior lobe, 1177, 1180, 1184, 1185 Anti-ataxic drug, 1555 Anticipation (related to dynamic mutations), 1810 Anticonvulsivants, 2086, 2088 Anti-Hebbian, 1258, 1267, 1268, 1272 Antineoplastics, 2080, 2088, 2089 Antipsychotics, 1741 Antithyroid drug, 326 Anxiety, 1512, 1731, 1733, 1740 3-AP. See 3-acetylpyridine (3-AP) Aplasia, 1859, 1860, 1861, 1864 Apoptosis, 1504, 1505, 1509, 1548, 1557 Aprosodia, 1729 APTX gene, 2344 AR. See Androgen receptor (AR) 2-Arachidonoylglycerol (2-AG), 929, 930, 931, 934, 935, 940 ARCA. See Autosomal recessive cerebellar ataxia (ARCA) Archimede’s spiral, 1615 Area 46, 554, 558, 559 Arm movements, 1282, 1286–1288, 1293 Arm tremor, 2154 Aromatase, 331 Arousal, 1182, 1185 ARSACS. See Autosomal recessive spastic ataxia of Charlevoix-Saguenay (ARSACS) Arteriovenous malformation, 697–699 Artery basilar, 345, 680–684, 686–687, 694, 695, 696 posterior inferior cerebellar, 680, 688, 693, 1629, 1630, 1728, 1960–1968, 1970–1972 superior cerebellar, 345, 353–355, 680, 692, 1629, 1630, 1722, 1728, 1960, 1962–1971 vertebral, 680–683, 686, 696, 699 Artery to artery embolism, 1964, 1965, 1967 Articulation, 1191–1194 Articulatory, 744, 751, 752, 753 As-c complex, 26 Ascl1, 24, 25, 28, 32–34, 36

Index Ascorbate, 715, 718, 723 Association areas, 483–485 Associative eyelid conditioning, 794, 802, 803 Asthenic tremor, 1612 Astrocytes, 1583, 1584, 1585 Astrocytoma, 725, 1729 Astronauts, 1242, 1245, 1246, 1250, 1251 AT. See Ataxia telangiectasia (AT) Ataxia, 518–519, 830, 836, 1482, 1483, 1487, 1490, 1491, 1493, 1494, 1509, 1512, 1514, 1522–1525, 1531, 1533, 1541, 1542, 1543, 1547, 1548, 1550, 1551, 1553, 1555–1557, 1670–1674, 1676, 1677, 1680, 1682, 1683, 1721, 1722, 1735–1737, 1901, 2144, 2155, 2156, 2166, 2168, 2194, 2272, 2278–2294, 2296–2301 Ataxia and non-ataxia symptoms, 2380 Ataxia of gait, 1601, 1616–1617, 1621, 2354 Ataxia of limbs, 1598, 1608–1615 Ataxia-oculomotor apraxia type 1, 2342–2344 type 2, 723 Ataxia of stance, 1598, 1616–1617 Ataxias, 1242–1243, 1250 Ataxia scales, 2387 Ataxia telangiectasia (AT), 1802–1805 Ataxic disorders, 1735–1736 Ataxic dysarthria, 1607, 1608 Ataxic hemiparesis, 1669–1683 Ataxic syrian hamster, 1435 Ataxin-1, 2333 Ataxin-2, 2333 Atention, 1725–1737, 1741 Atherosclerosis, 1681–1683 Atlas of the cerebellar nuclei, 1641, 1646, 1650, 1651 Ato, 26, 28 Atoh1, 4, 24, 27–31, 33, 36, 76, 92, 132–133 Atoh5, 27 Atoh1/Math1, 62 Atonal, 24, 27–29 Atonal homolog 1, 24, 27 ATP, 947–949, 952, 953, 955, 956, 960–966 ATP/GTP binding motif, 1573 Atrophin-1, 2340 Atrophy, 426 Attention, 1777, 1778, 1862–1864 Attentional control, 1733, 1735 Attention-deficit hyperactivity disorder (ADHD), 909, 910, 922, 1183, 1184, 1734, 1778

Index Atypical neuroleptics, 1911 Atypical psychosis, 1733 Auditory thalamus, 1178, 1181, 1182 Autism, 922, 923, 1014, 1043, 1334, 1778, 1896 Autism spectrum disorders (ASD), 488–489, 1733–1735 Autistic-like, 1737 Auto antibody, 882, 890, 891 Autonomic, 483, 1723, 1738 Autonomic failure, 2328 Autophagy, 1505, 1506 Autosomal dominant, 2194 Autosomal dominant cerebellar ataxia (ADCA), 721–723, 1617, 1800, 1805–1806 Autosomal recessive, 1565, 1574 Autosomal recessive cerebellar ataxia (ARCA), 1800, 1802–1805, 1809–1811 Autosomal recessive spastic ataxia of Charlevoix-Saguenay (ARSACS), 723 Aversive processing, 1720 Avian embryo, 1431 Axial postural tremor, 1612, 1614 Axon, 2160, 2161 Axonal competition, 1023 Axonal swelling, 2161 Axon initial segment (AIS), 243 Axonogenesis, 154–156 Axotomy, 1582 A zone, 398, 401, 404, 406–408 A2 zone, 399, 409, 411, 412, 418, 421

B BA. See Basilar artery (BA) Babinski’s sign, 1670, 1671 Bacterial, 2028, 2031 Bad, 137 Balance disorder, 2350, 2354, 2356–2358, 2360, 2362 Ballistic movement, 845, 846 Baltic myoclonus, 1924–1929 BARS. See Brief ataxia rating scale (BARS) Basal ganglia, 549, 550, 561–566, 1550, 1557 Basal interstitial nucleus, 379, 380 Basic-helix-loop-helix type (bHLH), 25, 27, 29, 31, 32, 76 Basilar apex (or terminus) aneurysms, 693 Basilar artery (BA), 345, 680–684, 686–687, 694, 695, 696

2399 Basilar artery occlusion, 686 Basilar dolichoectasia, 694, 695 Basis pontis, 1718, 1726 Basket cells, 78, 208, 222, 243–245, 809–812, 883, 884, 890, 897, 899, 900, 918, 1500, 1501, 1504, 2160, 2162, 2163, 2169 Bax, 1505 Bayesian inference, 1302, 1311 BDNF. See Brain-derived neurotrophic factor (BDNF) Bechterew’s nucleus, 394–398, 408, 423 Behavioral neurology, 1717 Benedikt’s syndrome, 685 Benign paroxysmal positional vertigo (BPPV), 1602, 1603 Bergmann glia, 107, 975, 976, 983, 984 Bergmann gliosis, 2163, 2166 Beta, 383, 384, 386, 391, 392, 395, 402, 411, 412, 414–416 bHLH, 93. see Basic-helix-loop-helix type (bHLH) Bim, 136, 137 Binominal data, 1644 Bioinformatics, 312–314 Biparous, 26 Bipolar, 1732, 1735 Bipolar disorder, 1778, 1780 Bistability, 1331–1333 Biventer, 347 BMPs. See Bone morphogenetic proteins (BMPs) Bone morphogenetic proteins (BMPs), 24, 29, 64, 77 Borden-Shucart classification, 700 BPPV. See Benign paroxysmal positional vertigo (BPPV) Brachium conjunctivum, 1471 Brain-derived neurotrophic factor (BDNF), 123, 1002 Brain imaging, 1193, 1195 Brain stem, 1222, 1223, 1225, 1874, 1875, 1881, 1882, 2031, 2032, 2034, 2163–2167 Brain stem compression, 1960, 1966, 1970–1972, 1977 Brain tumor, 1670, 1675 Breeding success rate, 1566 Brief ataxia rating scale (BARS), 1787, 1789–1790 Broca’s area, 1192 Burst firing, 1043 B zone, 398, 399, 404, 406, 407

2400 C 1C2, 2333, 2336, 2338, 2340, 2341 Ca2þ and calmodulin dependent kinase II (CaMKII), 887, 889 Ca2þ channels, 117, 118, 1422, 1423, 1526–1529, 1531–1533 Cacna1a, 135, 137, 1542, 1543, 1545, 1547–1549, 1551, 1554–1557 Cacna1a gene, 1529 Cacna2d2, 136 Cacng2, 135 Cadherins, 867 Ca2þ dyes, 116, 118 Caffeine, 121, 122, 1522, 1524, 1531, 1532, 1533 CAG repeat, 2331, 2333, 2334, 2336–2341 CAG repeat disease, 370 Ca2þ homeostasis, 1554 Calb1, 134, 139 Calbindin, 883 Calcineurin phophatase (CaN), 123 Calcium, 775, 794, 797, 1117, 1118, 1121, 1123 Calcium-activated potassium channels, 1062, 1066 a-Calcium/calmodulin-dependent kinase type II (aCamKII), 799 Calcium influx, 931, 932, 934, 936, 937 Calmodulin-dependent protein kinase (CaMK), 123 Calr3. See Calreticulin 3 (Calr3) Calreticulin 3 (Calr3), 136 Calretinin, 172, 885, 1471 CaMK. See Calmodulin-dependent protein kinase (CaMK) CaMKII. See Ca2þ and calmodulin dependent kinase II (CaMKII) cAMP. See Cyclic AMP (cAMP) CaN. See Calcineurin phophatase (CaN) Cancer, 1382, 1408–1414 Cannabinoid, 887, 888 Cannabinoid receptor type 1 (CB1-R), 800 Carbonic anhydrase, 1523, 1534 Carboxypeptidase, 1570, 1573, 1574 Cardioembolic stroke, 1681, 1683 Cardioembolism, 1960, 1967–1969 Carp, 1430, 1431 Carrier frequency, 1805 Carrier testing, 1807 Ca2þ signaling, 1529 Ca2þ signaling pathway, 123 Caspase-3, 1505 Ca2þ spikes, 116

Index Cat, 1430, 1436 Category fluency, 1710 Cato, 26 Cav2.1, 1066 CaV2.1, 1542, 1544, 1545, 1547–1557 CaV3.1, 1066 CaV2.1-a1, 1545–1548, 1553, 1557 CaV2.1-channelopathies, 1542 CB1-R. See Cannabinoid receptor type 1 (CB1-R) CCA. See Cerebellar cortical atrophy (CCA) CCAS. See Cerebellar cognitive affective syndrome (CCAS) Cck, 131 Ccna2 (CyclinA2), 134 Ccnd2 (CyclinD2), 132 CCP, 1573 CCTC. See Cortico-cerebellar-thalamiccortical (CCTC) brain circuit Cdk5, 137 cDNA, 1389 Cebus, 552–558 Cell fate specification, 23, 183 Central, 355 Central dogma, 1385, 1389, 1405 Central lobe, 1470, 1472 Central tegmental tract, 394, 423 Cerebellar, 1959 Cerebellar agenesis, 1737, 1741 Cerebellar artery aneurysms, 693, 696 Cerebellar ataxia, 1564, 1571, 1574, 1576, 1674, 1680, 2040, 2120, 2123, 2125, 2131, 2132, 2351, 2352, 2356, 2357, 2359–2361 Cerebellar atrophy, 1728, 1734 Cerebellar chip, 1330 Cerebellar cognitive affective syndrome (CCAS), 489, 736, 1722–1723, 1724, 1726, 1728, 1730, 1731, 1732, 1735, 1737, 1739, 1741, 1754, 1757, 1760, 1764, 1776, 1897 Cerebellar cortex (Cx), 239, 242, 249, 438, 557–561, 563, 895, 896, 897, 899, 900, 904, 906, 907, 916–919, 922, 1582, 1589, 2017 Cerebellar cortical atrophy (CCA), 721–723 Cerebellar cortical degeneration, 1632 Cerebellar degeneration, 1499, 1506, 1515 Cerebellar degenerative disorders, 1705 Cerebellar development, 181, 1448 Cerebellar fits, 1615 Cerebellar GABAergic interneurons, 186 Cerebellar GABAergic neurons, 63

Index Cerebellar GABAergic projection neurons, 186 Cerebellar glomeruli, 239 Cerebellar glutamatergic neurons, 66 Cerebellar hemorrhage, 1728, 1734, 1735 Cerebellar hypoplasia, 1737, 1741 Cerebellar infarction, 687 Cerebellar lesions, 1223–1225, 1226, 1229, 1230, 1235, 1703, 1705–1709, 1711 Cerebellar lobules, 485 Cerebellar malformations, 190, 193, 196 Cerebellar metastases, 2040, 2043 Cerebellar modulation, 1738, 1741 Cerebellar mutants, 238 Cerebellar mutism, 1730, 1732, 1753–1765 Cerebellar neurogenesis, 149 Cerebellar nuclei (CN), 75, 180, 377–426, 498–501, 550, 553, 556, 561, 882, 883, 884, 885, 887, 888, 891, 973–978, 980, 986, 1111, 1502, 1503, 1508, 1524, 1525, 1527, 1529, 1531, 1532, 1533, 1629, 1632, 1635–1637, 1639, 1641, 1642, 1645, 1646, 1649–1652 Cerebellar-plus syndrome, 2144 Cerebellar rhombic lip, 62 Cerebellar slice, 115, 117, 118, 121 Cerebellar stroke, 1629–1630, 1644, 1721, 1722, 1727, 1730, 1731 Cerebellar tremor, 1610, 1614, 2352, 2354, 2355 Cerebellar tumors, 1630–1631, 1729–1731 Cerebellar ventricular zone, 62, 68–69, 181 Cerebellar white matter, 355 Cerebellar zones, 390, 398–400 Cerebelless, 32 Cerebellin, 248 Cerebellitis, 726, 2031, 2035 Cerebello-bulbo-cerebellar loop, 514 Cerebello-mesencephalic cistern, 355 Cerebellopontine angle, 351 Cerebellothalamic, 530, 532–540, 542, 543 Cerebellum-like structures, 1289 Cerebral cortex, 1772, 1775, 1780 Cerebro-cerebellar, 735–739, 745, 746, 748, 751, 753–756, 1131, 1718, 1720, 1722, 1741 Cerebrocerebellum, 1755 Cerebro-pontine, 1133, 1135, 1137 Cerebrum, 1812, 1914–1916 CF. See Climbing fibers (CF) CF-LTD. See Climbing fiber LTD (CF-LTD) cGMP. See Cyclic GMP (cGMP) Channelopathy, 1522–1523, 1534

2401 Chaos, 1391, 1398–1400 Chiari malformations, 1737, 1874 Chick embryo, 1432 Chicken, 1430, 1432 Children, 1888–1891 Chlorzoxazone, 1534 Cholesterol, 994–996, 998, 1003, 1006, 1007 Cholesterol metabolism, 134 Choline, 715, 717, 718, 721, 724 Choroid plexus, 347, 348 Chromosome 13, 1573 Chromosome 16q-linked autosomal dominant cerebellar ataxia/spinocerebellar ataxia type 31 (16q-ADCA/SCA31), 2338–2339 Cingulate, 1719, 1720 Cingulate cortex, 481, 560 Citrate synthase (CS), 132 CK. See Creatine kinase (CK) Classical conditioning, 1511 Claude’s syndrome, 685 Climbing fiber LTD (CF-LTD), 982 Climbing fiber (CF) neuron, 81 Climbing fiber response, 897, 910 Climbing fibers (CF), 238, 239, 242, 258, 282, 361–367, 400, 438, 442, 793, 837, 882, 883, 885, 886, 887, 896, 897, 900, 901, 905, 907, 910, 972–975, 981–984, 1014–1023, 1031, 1035–1041, 1060–1062, 1067, 1068, 1071–1073, 1081–1086, 1089, 1090, 1092–1094, 1112–1116, 1122, 1132, 1135, 1137–1139, 1141–1143, 1145–1148, 1177–1179, 1181, 1258, 1262, 1263, 1265, 1268–1273, 1290, 1319–1322, 1326, 1328–1330, 1332, 1361, 1363, 1364, 1370, 1371, 1467, 1472, 1527–1529, 1532, 1549 branching, 406, 408 Clinical features, 2152–2158 Clinical heterogeneity, 2155 Clinical scales, 1786 Clinical studies, 1787 Clock, 1060, 1067, 1073 Closed-loop circuits, 550, 559, 560 Clumsiness, 1601, 1603 CN. See Cerebellar nuclei (CN) CN-GABA-interneuron, 76, 80 CN-GABA-ION neuron, 75 CN-Glu neuron, 76, 77, 79 Coactivator, 320, 322, 326 Cochlear nucleus, 83 Coculture, 272, 273

2402 Coenzyme Q10, 1804, 1811 Cognard, 700–702 Cognition, 480, 566, 755, 1701, 1703–1704, 1705 Cognitive, 735–738, 743–745, 748, 754, 755, 756, 1861, 1862, 1864, 1866–1868, 2155, 2157, 2158 Cognitive cerebellum, 1720 Cognitive double-hit, 1737 Cognitive flexibility, 1736 Cognitive operations, 1191, 1192 Cognitive overshoot, 1731, 1733 Cognitive rehabilitation, 2353–2354 Coiled bodies, 2336 Coma, 1961, 1971, 1973 Compartmental model, 1033 Compensatory strategies, 2359 Complex behavior, 1348–1354 Complex spike (CS), 363, 1060, 1063–1064, 1066, 1067, 1071, 1072, 1287, 1320–1322 Computational, 1360 Computed tomography, 1675 Computer model, 1042 Conditioned response (CR), 1176–1185 Conditioned stimulus (CS), 1119, 1176–1184 Conductance, 854, 858, 860, 868, 869 Connections, 735–738, 744, 746, 748, 755 Connectivity, 768–770 Connexin 36, 1023 Consciousness, 1354 Continuous data, 1644 Continuous training, 2350, 2359, 2361 Contravariant, 1382, 1383, 1385, 1389–1393, 1395, 1411 Conventional volumetric analysis, 1646–1649 Convergence, 770 Convergence of cerebellar projections, 515–516 Coordinated Genome Function, 1383, 1391, 1410, 1414 Coordination, 2156 Corepressor, 320 Cornichon, 858, 860, 863–864 Corollary discharge, 1258, 1261, 1263, 1265, 1267, 1271 Corona radiata, 1673–1674, 1676, 1677, 1683 Corpus cerebelli, 1464–1476 Cortex, 1580, 1582, 1589, 1673, 1674, 1676–1677, 1680, 1683 Cortical modules, 166

Index Corticocerebellar atrophy, 1564 Cortico-cerebellar-thalamic-cortical (CCTC) brain circuit, 1184 Corticonuclear microcomplex, 1722 Corticonuclear projection, 398–400 Corticopontine, 1719 Cortico-rubral projection, 397 Corticospinal tract, 501, 510, 515, 1673, 1674, 1676, 1678 Corticovestibular projection, 508 Coupling, 1025, 1026, 1028, 1031–1037, 1040, 1042 Covariance learning rule, 1318, 1321 Covariant, 1382–1383, 1386, 1387, 1389–1391, 1395, 1396, 1411 CR. See Conditioned response (CR) Cranial tremor, 2154 Craniotomy, 1971, 1973, 1977 Creatine, 715, 717, 718, 720, 721, 724, 725 Creatine kinase (CK), 720 Creutfeldt-Jakob disease (CJD), 2035, 2036 CRF-2a-tr, 978–980 CRF-R1, 977, 978, 979, 983 CRF-R2a, 978, 979, 984, 986 CRF receptor, 977–980, 983, 984, 986 Crista cerebelli, 1466 Critical period, 266, 273–275 cRNA, 1382, 1389, 1391, 1396 Cronbach’s a, 1786, 1789, 1790, 1793, 1795 Crossed cerebellar diaschisis, 1680–1681, 2067, 2071 Cross-modal transfer, 1119 Crus II, 558, 559, 563, 564 CS. See Complex spike (CS); Conditioned stimulus (CS); Citrate synthase (CS) Computed tomography (CT), 587, 609, 610, 619, 620, 626, 628, 632, 633, 655, 663, 665, 1970, 1973–1976 CT angiography, 683, 695, 703 CTG repeat, 2338 CT-scan, 2028, 2029, 2032 Culmen, 355 Curly cells, 1018–1019 Cx. See Cerebellar cortex (Cx) Cyclic AMP (cAMP), 930 Cyclic GMP (cGMP), 121 Cyclic guanosine monophosphate, 797 Cyclic nucleotide, 121, 122 Cytochrome oxidase, 1506, 1507 Cytochrome P450 17a-hydroxylase/c17,20lyase (P45017a,lyase), 997, 998, 1002 Cytochrome P450 aromatase (P450arom), 997, 1002

Index Cytochrome P450 side-chain cleavage enzyme (P450scc), 995, 997, 1000, 1002 Cytokines, 1574 C zone, 398, 404–408

D Dandy–Walker malformation, 1737, 1887 DAO. See Dorsal accessory olive (DAO) Darkschewitsch nucleus, 394 DBN. See Downbeat nygstagmus (DBN) DC. See Dorsal cap (DC) DCK. See Dorsal cap of kooy (DCK) DCN. See Deep cerebellar nuclei (DCN); Dorsal column nucleus (DCN) Decerebrated cat, 1436 Declarative memory, 1710, 1711 Declive, 347, 355 Decomposition of movement, 1599, 1600, 1608, 1609 Decompression, 1881, 1882, 2035 Decorrelation control, 1322, 1331 Deep cerebellar nuclei (DCN), 16, 62, 239, 246–247, 896, 905, 917, 922, 1012, 1040, 1060, 1061, 1070, 1073, 1080–1084, 1086, 1093, 1568, 1573, 1575, 1580, 1589, 1718, 1722, 1773, 1776, 2350. See also Cerebellar nuclei Degeneration, 2162, 2168, 2169 Degenerative ataxias, 1631 Degenerative disease, 2350, 2351, 2356, 2357, 2359–2363 Dehydroepiandrosterone (DHEA), 998, 1001 Dejerine’s syndrome, 682 Dementia, 1726, 2157 Demyelinating disease, 1675 Dendrites, 281, 767, 768, 769, 770, 772, 773, 777, 780 Dendritic growth, 999–1006 Dendritic lamellar bodies, 1025, 1026 Dendritic spines, 241, 1549 Dendritogenesis, 160–161, 163, 165 De novo mutation, 1802, 1806 Dentate nucleus (DN), 180, 196, 348, 378, 438, 550, 553, 554, 555, 556, 557, 559–561, 565, 1288, 1289, 1719, 1722 Dentatorubropallidoluysian atrophy (DRPLA), 1805, 1806, 1810, 1924–1926, 1935–1938, 2333, 2334, 2340–2342, 2381 Dentato-thalamo-cortical pathway, 1192 Depolarization-induced potentiation of inhibition (DPI), 887

2403 Depolarization-induced suppression of excitation, 932, 934–935, 937 Depolarization-induced suppression of inhibition (DSI), 887, 931–934 Depression, 489, 1724, 1732, 1735, 1740, 2158 Descending vestibular nucleus (DVN), 359 Desensitization, 117, 854, 861, 869 Detrusor-sphincter dyssynergia, 2124 Development, 994–997, 999, 1001, 1003–1006, 1014–1023, 1857–1864, 1867 Developmental disorder, 923 d2 glutamate receptor (GluD2), 241 DHEA. See Dehydroepiandrosterone (DHEA) Diaschisis, 1754, 1761 Dichotomy of cerebellar linkage, 1720 Dichotomy of cerebellar motor versus cognitive/limbic, 1722 Differential diagnosis, 587–674 Diffusion, 1914–1915 Diffusion tensor imaging (DTI), 1649 Diffusion-weighted imaging (DWI), 1671, 1674, 1676–1677, 1681–1683, 1975 d2 ionotropic glutamate receptor, 984 Direct current stimulation (tDCS), 2363 Disability, 2152, 2154, 2158 Disinhibition, 1723, 1726, 1728, 1729, 1731, 1732, 1733, 1740 Disorders of muscle tone, 1615 Disorders of vestibulo-ocular reflex, 1597 Disruption, 1858, 1859, 1860, 1865, 1867 Dissecting aneurysms, 694, 695 Dissection, 1960, 1961, 1976 Distal intracranial posterior circulation territory, 343 Distributed neural circuits, 1720 Divergence, 770 Divergence of cerebellar projections, 515 Dix-Hallpike maneuver, 1602 Dizziness, 1601, 1602, 1603 DLH. See Dorsolateral hump (DLH) DMCC. See Dorsomedial cell column (DMCC) DN. See Dentate nuclei (DN) Dominant ataxia, 1432–1434 DON. See Dorsal octavolateral nucleus (DON) Doppler, 1976 Dorsal accessory olive (DAO), 381, 383, 1016, 1018, 1026, 1027 Dorsal cap of kooy (DCK), 383, 1018, 1024, 1026, 1027, 1030, 1060, 1064, 1073 Dorsal cochlear nucleus (DCN), 1259, 1260 Dorsal column nucleus (DCN), 454–455, 1026

2404 Dorsal funiculus spino-olivary climbing fiber path, 388 Dorsal light response, 1475 Dorsal motor nucleus of the vagus, 350 Dorsal octavolateral nucleus (DON), 1259, 1260 Dorsal premotor area (PMd), 552, 553 Dorsal premotor cortex, 1285, 1286 Dorsal Y-group, 361 Dorsolateral hump (DLH), 499, 506–507 Dorsomedial cell column (DMCC), 361, 363, 381, 383, 386, 392, 395, 402, 404, 405, 1018, 1026, 1027 Dorsomedial group, 381, 383 Downbeat nygstagmus (DBN), 1603, 2351, 2353 DPI. See Depolarization-induced potentiation of inhibition (DPI) Dreher (dr), 66, 129 Drosophila, 25–27, 29 D. melanogaster, 24 DRPLA. See Dentatorubralpallidoluysian atrophy (DRPLA) DSI. See Depolarization-induced suppression of inhibition (DSI) DTI. See Diffusion tensor imaging (DTI) DTI tractography, 574, 582 Ducky, 136 Dural arteriovenous fistula, 699–702 Duret hemorrhage, 2058 DVN. See Descending vestibular nucleus (DVN) DWI. See Diffusion-weighted imaging (DWI) 25-Dx, progesterone receptor membrane component 1 (PGRMC1), 10003 Dynamic balance, 2357–2359 Dynamic clamp, 1116, 1117 Dynamic mutation, 1800, 1802, 1805, 1810 Dysarthria, 1598, 1599, 1603, 1606–1608, 1617, 1620, 1721–1723, 1730, 1758, 1787, 1862, 1961, 1963, 1967, 1969, 1971, 2194, 2216, 2221, 2225–2229, 2232, 2235, 2245, 2279, 2280, 2287, 2288, 2291, 2293, 2300 ataxic, 1607, 1608 hypokinetic, 1606 Dysdiadochokinesia, 1607, 1608, 1610 Dysgraphia, 1711 Dyslexia, 1334 Dysmetria, 490, 1305–1307, 1598, 1599, 1600, 1605, 1607, 1608, 1609, 1611, 1670, 1679, 1721, 1722, 1738, 1739, 1965, 1967, 1969, 2354, 2356 Dysmetria of thought hypothesis, 1598

Index Dysmetria of thought theory, 1738, 1739 Dysphagia, 1608, 2222, 2226, 2228, 2232 Dysprosody, 1608 Dysrhythmia, 2167, 2168 Dysrythmokinesia, 1610 Dyssynergia, 2354 Dystonia, 565, 1244, 1524, 1529, 1531–1534 Dystroglycan, 249 D zone, 398, 405, 407, 417

E EA. See Episodic ataxia (EA) EA2. See Episodic ataxia type 2 (EA2) Early infantile autism, 1734, 1735 Early-onset ataxia with ocular motor apraxia and hypoalbuminemia/ataxiaoculomotor apraxia type 1 (EAOH/AOA1), 2342–2344 ECBs. See Endocannabinoids (ECBs) ECN. See External cuneate nucleus (ECN) Edema, 1960, 1971, 1972, 1974 EDH. See Epidural hematoma (EDH) EEG, 1928, 1931, 1933–1035, 1937, 1938, 1941–1942, 1945–1946, 1948, 1951 Efference copy, 1324, 1325, 1326, 1328 Efferent neuron, 1466–1472 EGL. See External granular layer (EGL) Electrical coupling, 884, 1062, 1068 Electrical stimulation, 1777, 1779, 1780 Electrical synapse, 1031, 1032 Electromyographic recordings (EMG), 1524, 1531, 1532, 1556 Electrophysiology, 1550–1551, 1554, 2152, 2156, 2158–2159, 2161 Electrosensory, 1258–1261, 1265, 1267, 1272, 1273 Electrosensory lobe (ELL), 1259, 1260 ELL. See Electrosensory lobe (ELL) Emboliform nucleus, 79, 184, 189, 191, 197, 355, 1718, 1719, 1722 Embryonic development, 65, 69 EMG. See Electromyographic recordings (EMG) Eminentia granulalis, 1464–1466, 1468, 1473 Emotion, 480, 756, 1182, 1185, 1701, 1772 Emotional control, 1731, 1733, 1735, 1739 Emotional dysregulation, 1731 Emotional lability, 1730 Emotional pacemaker, 1732 Emotion attribution, 1734 Emotion regulation, 1777, 1779

Index Encephalitis, 2031, 2032, 2034 Encephalomyopathy, 2290, 2292–2297 Endocannabinoids (ECBs), 800, 802, 927, 929–941, 1068, 1583 Endocrine, 2010 Endothelin-1 receptor, 1509 Endplate, 1552, 1554, 1556 En2 gene in autism, 131 Engrailed genes (En1 and En2), 129 Enterovirus, 2033, 2035 Environment, 2080, 2085, 2104 Environmental influences, 320 Ependymoma, 725, 1729 EphA4, 131 Ephrin, 1015 Epidemiology, 2147–2148, 2152–2153 Epidural hematoma (EDH), 2056, 2058, 2060, 2063, 2065, 2070 Epilepsy, 1543, 1545, 1547, 1548, 1555, 1675, 1683, 2279, 2280, 2287–2289, 2292–2294, 2297, 2298, 2300 Episodic ataxia (EA), 722, 890, 1800, 1806, 1807, 1812 Episodic ataxia type 2 (EA2), 1522, 1523, 1526, 1531, 1533, 1534, 1555, 1556 Episodic neurological disorder, 1522, 1532, 1534 Equilibrium, 1242, 1243 ERa. See Estrogen receptor-a (ERa) ERb. See Estrogen receptor-b (ERb) Error-based learning, 1711 Error signal, 1041, 1317, 1318, 1321, 1322, 1325 Essential tremor, 2152 Estradiol, 998, 1001, 1002, 1004–1007 Estradiol (E2), 331 Estrogen receptor-a (ERa), 333 Estrogen receptor-b (ERb), 1002 Ethanol, 121, 122 Ethnicity, 1812 Eurydendroid cells, 1445, 1447, 1454, 1455, 1466, 1468 EVD. See Extraventricular drainage (EVD) Event timing, 1212 Everyday living, 2357, 2360, 2361 Evolution, 1464, 1476, 1772, 1775 Excitatory postsynaptic currents, 922, 1554 Excitotoxic, 1504 Executive, 1862–1865, 1867 Executive functions, 745–748, 754, 1709, 1723–1729, 1735, 1736, 1739 Expanded polyglutamine stretches, 2333, 2336, 2337, 2340–2342

2405 Explant, 273, 274 Exploration behavior, 1512 Exploration strategies, 1706 Expressive language delay, 1737 External cuneate nucleus (ECN), 80 External granular layer (EGL), 108–110, 115–116, 118 Extinction, 1119 Extracellular recordings, 830, 839 Extrapolation, 1435 Extrapyramidal, 1550, 2196, 2197, 2210, 2216, 2217 Extraventricular drainage (EVD), 1971, 1976, 1977 Eyeblink conditioning, 1112, 1118–1123, 1125, 1175, 1208, 1210, 1211 Eye-blink reflex, 1434 Eyelid conditioning, 842 Eyelid response, 1511 Eye movements, 1285, 1287

F Facial nucleus, 1179 Familial hemiplegic migraine type 1 (FHM1), 1547, 1556 FARS. See Friedreich ataxia rating scale (FARS) FAS. See Fetal alcohol syndrome (FAS) FASD. See Fetal alcohol spectrum disorder (FASD) Fastigial nucleus, 184, 190, 348, 378, 442, 480, 498, 502, 560, 1288, 1720, 1722, 1731, 1732, 1740 Fastigial oculomotor region (FOR), 1162–1164 Fast synaptic transmission, 773–776 Fate map, 4 fcMRI. See Functional connectivity magnetic resonance imaging (fcMRI) Fear conditioning, 1475 Feedback, 1282, 1284–1286, 1718, 1719 Feedback inhibitory circuit, 831, 834, 837, 846 Feed forward, 1282, 1284, 1718, 1719 Feed forward control, 1284 Feed forward inhibitory circuit, 834 Feed forward processes, 1303, 1305 FEF. See Frontal eye field (FEF) Fetal alcohol spectrum disorder (FASD), 121, 1183 Fetal alcohol syndrome (FAS), 1183 FGF8, 24, 34 Fgf8, 8–10, 1016

2406 FHM1. See Familial hemiplegic migraine type 1 (FHM1) Finger-to-finger test, 1611 Finger-to-nose test, 1611 Fish, 1442 Fissure horizontal, 355 Floccular complex, 1156–1161 Flocculus, 345, 350–352, 380, 508, 1328, 1332, 1333, 1774 FL purkinje cells, 1160, 1168 fMRI. See Functional magnetic resonance imaging (fMRI) Focal lesions, 1705 Folium, 355, 1531 FOR. See Fastigial oculomotor region (FOR) Foramen magnum, 347, 1874–1876, 1878–1881 Force fields, 1283–1286, 1288, 1291 Forward models, 1168, 1169, 1258, 1259, 1270–1272, 1282, 1291, 1297, 1299, 1300, 1302, 1304, 1308, 1311, 1324 Founder mutation, 1803, 1811 Fractal, 1384, 1389, 1391–1400, 1402–1407, 1409–1415 Fractal defect, 1382, 1407, 1408, 1412 Fractogene, 1389, 1393, 1394 Fragile X mental retardation 1 gene (FMR1), 1900 Fragile X mental retardation protein (FMRP), 1900 Fragile X syndrome (FXS), 1896, 1900 Fragile X tremor/ataxia syndrome (FXTAS), 1807, 1810, 2374, 2380, 2381 Frataxin, 2342 FRDA. See Friedreich’s ataxia (FRDA) Friedreich ataxia rating scale (FARS), 1787, 1790–1792, 1795, 1796 Friedreich’s ataxia (FRDA), 723, 1382, 1406, 1407, 1605, 1607, 1608, 1736, 1800, 1802–1805, 1810, 2342, 2351, 2352, 2361, 2373, 2375–2378, 2386 Frontal cortex, 1775 Frontal eye field (FEF), 395–397, 422–425, 552, 555, 556, 563 Frontal systems, 1726 Functional connectivity, 571, 574–581, 739, 740, 746, 749 Functional connectivity magnetic resonance imaging (fcMRI), 1720 Functional magnetic resonance imaging (fMRI), 741, 1720, 1727, 1775, 1776 Functional recovery, 1581

Index Functional redundancy, 1573, 1576 Functional tests, 1791, 1794 Fusiform aneurysms, 694–696 FXTAS. See Fragile X tremor/ataxia syndrome (FXTAS)

G GAA repeats, 2342 GABAA receptors, 113, 122, 123, 238, 885–891, 1526 GABAB-R. See Gamma-aminobutyric acid receptor type B (GABAB-R) GABAB receptor, 885–887, 889, 890 GABAergic, 1102–1109 GABAergic interneurons, 207 GABAergic neurogenesis, 32 GABAergic neurons, 76–78, 378, 390, 397, 416 GABA release, 812, 814, 817–822 Gabra6, 139 GAD. See Glutamate decarboxylase (GAD); Glutamic acid decarboxylase (GAD) Gain control, 844, 847 Gait, 1242–1244, 1510, 1514 Gait abnormality, 2155, 2156 Gait analysis, 567 Gait disturbance, 1966 Galanin, 910 Gamma-aminobutyric acid (GABA), 238, 244, 246, 715, 716, 721, 771, 774–776, 780, 783, 833–835, 838, 1070, 1071, 1112, 1466, 1468, 1470, 1471, 1472, 1476, 1508, 1899, 2163, 2168 Gamma-aminobutyric acid receptor type B (GABAB-R), 795 Gamma-band activity, 845 Ganglionic layer, 1466–1468, 1470, 1471, 1472, 1476 Gap junctions, 846, 1023–1026, 1031, 1033, 1035, 1038, 1039, 1061, 1062, 1066–1067, 1070–1073 Gas1, 132 Gating, 900, 902 Gaucher disease (GD), 1924, 1926, 1939–1942, 1951 Gaze-evoked nystagmus (GEN), 1603 Gbx2, 8, 10–12, 129, 1014, 1015 GCIs. See Glial cytoplasmic inclusions (GCIs) GCL. See Granule cell layer (GCL) GD. See Gaucher disease (GD) GEN. See Gaze-evoked nystagmus (GEN)

Index Gene, 302, 2194, 2198, 2200, 2202, 2205, 2207, 2208, 2212, 2215, 2216, 2218, 2221, 2223–2247, 2249, 2251 Gene array, 128, 131 Gene expression, 46 Gene networks, 127 Gene paint, 139 Generalized coordinates, 1390, 1406 Genetic, 1858, 1859, 1861 Genetic approach, 1227 Genetic counseling, 1800, 1808–1810, 2316, 2318–2319, 2323 Genetic disease, 2205–2208 Genetic epidemiology, 1806 Genetic fate mapping, 33, 65, 183 Genetic model, 1442, 1448, 1456, 1457 Genetic mutation, 1522 Genetic testing, 1800, 1804, 1806, 1812–1814 Genomic action, 1003 Geographic group, 1805 Gephyrin, 244 Gerstmann-Str€aussler-Scheinker disease, 722, 2035 GFAP. See Glial fibrillary acidic protein (GFAP) Giant fusiform aneurysms, 695, 696 Giant potentials, 1928, 1931 GIRK channels, 771, 782 Girk2 gene, 128, 136, 137, 1570 Gli2, 30, 132 Glia, 1583, 1584 Glial cytoplasmic inclusions (GCIs), 2328 Glial fibrillary acidic protein (GFAP), 996 Globose nucleus, 184, 189, 191, 197, 355, 378, 379, 1718, 1719, 1722 Globus pallidus, 530, 533–543 external segment, 563 Glomeruli, 769, 770, 831–837 Glucocorticoid receptor (GR), 330, 331 Glucocorticoids, 330–331, 1584 Glucose, 714, 715, 717, 719–724 GluD2. See d2 glutamate receptor (GluD2) GluRd2, 1500, 1503, 1505, 1506 Glutamate, 113–115, 243, 714–718, 720–724, 771, 775, 776, 794, 795, 797–799, 802, 803, 1069, 1071, 1112, 1113, 1117, 1124, 1503, 1507, 1508 Glutamate decarboxylase (GAD), 890 Glutamate receptor interacting protein (GRIP), 856, 864–866, 869 Glutamate receptor interacting protein ½, 797, 798

2407 Glutamate receptors, 241, 1500, 1503, 1507, 1508, 1532 Glutamatergic, 1102–1105, 1107–1109 Glutamatergic neuron, 76 Glutamatergic projection neurons, 187 Glutamatergic synapses, 793(Found only in abstract) Glutamic acid decarboxylase (GAD), 1899, 1902 Glutamine, 714–717, 720–724 Glutathione, 715, 718, 721 Gluten ataxia, 1986, 1987, 1991–1998 Glycine, 238, 246, 833, 834, 838 Glycinergic, 1102–1107 Glycinergic neurons, 378, 420 Golden ratio, 1398–1401 Goldfish, 1430, 1431, 1465, 1466, 1468–1470, 1472–1476 Golgi cells (GO), 78, 208, 238, 239, 245, 246, 767–770, 774–778, 780, 781, 783, 830, 831, 833–839, 841–847, 882–886, 889, 890, 918, 1158, 1159, 1467 Gonadal hormone, 331–335 GR. See Glucocorticoid receptor (GR) Gracile lobule, 347, 348 Graft, 272 Grafting study, 80, 83 Granular layer, 1525, 1526 Granule cell clonal expansion, 29 Granule cell density, 1563, 1569, 1575 Granule cell development, 28–31 Granule cell layer (GCL), 238, 239, 245, 1466, 1467, 1469, 1568, 1569 Granule cells (GC), 16, 17, 66, 67, 76, 90, 97–98, 107, 238, 282, 286, 290, 291, 829–837, 841, 846–847, 882–884, 886, 889, 891, 917, 918, 1003, 1006, 1320–1322, 1331, 1365–1369, 1372, 1466, 1467, 1469, 1501, 1504, 1505, 1513, 1524, 1526, 1527, 1529, 1568, 1569, 1575, 1576 Grasping, 1657–1665 Gravity, 1242 Gray matter, 1912–1915, 1917 Grb2, 132, 134 Grid2, 135, 137, 263–266, 1570 GRIP. See Glutamate receptor interacting protein (GRIP) Grip force, 1658, 1659, 1663, 1664 Grm1, 134, 140 Grooming, 1512 Group Y, 379 G substrate, 798

2408 H Handwriting abnormalities, 1615 Harlequin, 137 Harmaline, 1025, 1027, 1030, 1043, 1063, 1064, 1066, 1072, 2159 Headache, 1601, 1602 Head tremor, 2154, 2156, 2159 Hearing loss, 1961, 1967 Hebbian, 1121 Heel-to-knee test, 1612 Hemangioblastoma, 2043, 2044 Hemicerebellectomy, 1705 Hemicerebellum, 268, 271 Hemiparesis, 1671–1672, 1678 Hemispheres, 1857, 1860, 1864, 1867, 1868 Hemorrhage, 1960, 1971–1974, 1977 Hemorrhagic stroke, 1670, 1674–1675 Hemorrhagic transformation, 1971, 1972 Hepatitis A, 2033 Hereditary ataxia, 1800, 1802, 1809, 1813, 1814 Heterochronic, 273, 275 Heterologous synapses, 247, 248 Heteroplasmy, 1808 Heterotopic, 2160, 2162 Hibernation, 1566 Hindbrain, 4, 8–9, 61, 64–66, 69–70, 80 Hindbrain choroid plexus, 64–66, 69–70 Hippocampus, 1545, 1549 Hodgkin-Huxley model, 1033 Homeodomain, 389, 1391 Homeoprotein, 1391 Horizontal fissure, 355 Hormone receptors, 320 Hormones, 2010, 2012, 2014, 2015, 2020 Horner’s syndrome, 682 Hoxa2, 91 Hox genes, 1014, 1016, 1017 Human evolution, 1338 Huntington’s disease, 1736 HVIIB. See Lobule, hemispheric VIIB (HVIIB) Hydrocephalus, 1888–1890, 1960, 1971–1974, 1977 6-Hydroxydopamine (6-OHDA), 900, 907, 908 3b-Hydroxysteroid dehydrogenase/D5-D4isomerase (3b-HSD), 996, 997, 1000, 1002 Hypergravity, 1242, 1247–1250 Hypermetria, 1599, 1603, 1605, 1609, 1611, 1620 Hypermetria of saccades, 1603

Index Hyperpolarization, 1117, 1118, 1121, 1122 Hyperpolarization-activated cation channel, 1062 Hypertonia, 518 Hypesthesia, 1672 Hypokinetic dysarthria, 1606 Hypometria, 1603, 1605, 1609 Hypometria of saccades, 1603 Hypoplasia, 1858–1860, 1868 Hypothalamo-pituitary-adrenal (HPA) axis, 330 Hypothalamus, 81, 515, 1720, 1774 Hypothyroidism, 323 Hypotonia, 518, 1600, 1607, 1615, 1671, 1863

I ICARS. See International cooperative ataxia rating scale (ICARS) ICC. See Intraclass correlation coefficients (ICC) ICNs. See Intrinsicic connectivity networks (ICNs) Idebenone, 2352 Idiopathic late onset cerebellar ataxia (ILOCA), 1800, 1808, 1809, 2144 IGL. See Internal granular layer (IGL) ILOCA. See Idiopathic late onset cerebellar ataxia (ILOCA) Imaging, 587–674 Imitation, 1338, 1349, 1353, 1354 Immediate-early genes, 801 Implicit learning, 1707 Inborn error of metabolism, 725 Incidence, 2153 Index-to-index test, 1612 Index-to-wrist maneuver, 1609 Indirect pathway, 1331 Infarct, 1670, 1672, 1673, 1675–1678, 1681 Infarction, 1959–1977 Infection, 2028–2035 Infectious diseases, 1675 Infectious mononucleosis (EBV), 2033 Inferior medullary velum, 347 Inferior olivary complex, 1573, 1575 medial accessory olive (MAO), 380, 1016, 1018, 1019, 1026, 1027 subnucleus, 380, 394, 396, 397, 402, 407, 411–416, 421, 422 Inferior olivary nuclei, 1060, 1718 Inferior olive (IO), 268, 270, 271, 378, 438, 442, 498, 842, 882, 883, 885, 1013–1043, 1059–1074, 1080, 1081,

Index 1083, 1084, 1086, 1088, 1089, 1092–1094, 1112, 1114, 1115, 1124, 1133, 1141, 1143, 1145–1147, 1177, 1178, 1181, 1290, 1320, 1328, 1329, 1332, 1472–1474, 1501, 1503–1505, 1507, 1513, 1514, 1580, 1581 Inferior olive nucleus (ION), 76, 80 Information coding, 1338, 1350–1351, 1353, 1354 Inhibition, 829, 831, 834, 836, 837, 838, 841, 842, 843, 846–847, 1113, 1115–1117, 1120, 1121, 1123, 1368–1371 Inhibitory, 2163, 2167 Inhibitory interneuron, 208–210, 223–227 Inhibitory postsynaptic currents (IPSCs), 952 Inositol triphosphate (IP3) receptor, 996 In situ branch atheromatous disease, 1964, 1967 Instability of gaze, 1603 Intelligence quotient (IQ), 1726 Intention tremor, 1671, 2154–2156, 2167 Internal auditory arteries, 350–352 Internal capsule, 1670, 1673–1678, 1680, 1683 Internal clock, 1205 Internal granular layer (IGL), 108, 111 Internal models, 1193, 1194, 1196, 1282, 1298–1303, 1333, 2354, 2355 Internal segment of the globus palidus (GPi), 563 Internal stores, 795 International cooperative ataxia rating scale (ICARS), 1787–1790, 1792, 1793, 1795, 1796, 2352, 2357, 2361 International Study of Unruptured Intracranial Aneurysms (ISUIA), 691, 692 International Subarachnoid Aneurysm Trial (ISAT), 692 Interneurons, 378, 809–814, 816–822 Interposed nucleus, 1289 Interpositus, 1176, 1177, 1179, 1180 Interpositus nucleus, 180, 184, 190, 442, 448, 449, 560, 1119 Interrater reliability, 1786, 1788, 1795 Interstitial cell groups, 378, 379, 399, 402, 413–415, 422–424, 498, 499, 506 Interstitial nucleus of Cajal (INC), 513, 514 Interstitiospinal tract, 513 Intra-arterial therapy, 687, 703 Intracellular staining, 439 Intraclass correlation coefficients (ICC), 1786, 1788, 1793–1795 Intracranial vertebral arteries, 343 Intranuclear inclusions, 2166, 2167

2409 Intraparenchymal hemorrhage, 693, 704 Intraparietal cortex, 397 Intravenous thombolysis, 687 Intrinsic electroresponsiveness, 771–773 Intrinsic excitability, 842 Intrinsicic connectivity networks (ICNs), 571, 574, 575, 578, 580, 581, 583 Intrinsic program, 111–113 In utero electroporation, 76 Inverse dynamics model, 1283, 1287, 1288 Inverse models, 1299 In vitro explant experiments, 70 In vivo imaging, 1453, 1456, 1457 IO. See Inferior olive (IO) Iodothyronine deoidinase, 324 ION. See Inferior olive nucleus (ION) Ion channel, 1522, 2243 IQ. See Intelligence quotient (IQ) IS. See Ischemic stroke (IS) ISAT. See International Subarachnoid Aneurysm Trial (ISAT) Ischemia, 1693–1694 Ischemic stroke (IS), 1670–1672, 1674–1676, 1681–1683 Isochronic, 274 Isometrataxia, 1612, 1613 Isometric contraction, 845 Isometric tremor, 1611, 1613 Isthmic organizer, 1014–1016, 1448–1450 Isthmus, 15 ISUIA. See International Study of Unruptured Intracranial Aneurysms (ISUIA) Itpr1, 134

J Joubert syndrome (JS), 196–197, 1737 Junk DNA, 1385, 1391, 1398, 1402 Juxtacellular recording, 839

K Kainate receptors, 787, 799, 836 Kainite, 854–858, 864, 868–869 Kalman filter, 1300, 1301 Kcnj6 gene, 136 Kinematics, 1282, 1285, 1287–1289 Kinetic tremor, 1608, 1611, 2152–2155 Knee-tibia test, 1611 KRIP6, 858, 869, 870 Kuru, 2035

2410 L Lactate, 715, 717, 719–724 Lactic acidosis, 2287, 2293–2298, 2300 Lafora disease (LD), 1924, 1926, 1946, 1951 Lambert-Eaton, 1542, 1556 Language, 736, 738, 743–745, 747–750, 755, 1192, 1193, 1195, 1196, 1705, 1709–1711, 1720, 1724, 1725, 1727–1729, 1732, 1737, 1739, 1862, 1863, 1865–1868 Large-artery atherosclerosis, 1681–1683 Large-scale mouse mutagenesis, 1482 Late cortical cerebellar atrophy (LCCA), 2331 Lateral cerebellar nucleus (LNC), 378–380, 392, 402, 404, 413, 498, 507 Lateral descending motor system, 501 Lateral reticular nucleus (LRN), 80, 358, 364, 447, 449–453 Lateral vestibular nucleus (LVN), 368, 392, 398, 402, 405, 415, 498 Lateral vestibulospinal tract, 507 LBW. See Low birth weight (LBW) LC. See Nucleus locus coeruleus (LC) LCCA. See Late cortical cerebellar atrophy (LCCA) LD. See Lafora disease (LD) Leading process, 109 Leaner, 135, 1545, 1547, 1549, 1553, 1555 Learning, 1111, 1192, 1348–1350, 1352–1354, 1362–1364, 1370, 1371, 1861–1864, 1866 Learning rule, 1317, 1318, 1320, 1321, 1323, 1331 Learning strategies, 2356 Legionellosis, 2033 Leg tremor, 2154, 2155 L7-En2 transgenic mouse, 131 Lesion delineation, 1633, 1634 Lesions, 2165–2168 Letter fluency, 1709 Lewy bodies, 2163, 2164, 2165, 2167 Lewy neuritis, 2165, 2166 L7 gene, 134, 139 Lhx1, 35, 138, 139 Lhx5, 35 Limb ataxia, 1787 Limbic cerebellum, 1720, 1731, 1741, 1775 Limbic system, 481–482, 1720, 1772, 1777, 1779 Lineage trace, 82 Linearity, 1786–1788, 1793 Linguistic deficits, 1717 Lmx1a, 29, 66, 129

Index Lmx1b, 68, 1016 LNC. See Lateral cerebellar nucleus (LNC) Lngitudinal band, 441, 443–447, 452, 458, 459 Lobe anterior, 1177, 1180, 1184, 1185 central, 1470, 1472 electrosensory, 1259, 1260 vestibulolateral, 1466, 1467, 1468, 1470 Lobule cerebellar, 485 gracile, 347, 348 hemispheric VI, 1176, 1179, 1180, 1182, 1185 hemispheric VIIB, 563, 564 semilunar, 347, 355 Lobus caudalis, 1463, 1464, 1466, 1468, 1470 Locked-in syndrome, 683, 686 Locus coeruleus, 1524, 1526, 1533 Long lasting depression, 841, 842, 843 Long-term adaptation, 2356 Long-term depression (LTD), 794–803, 855, 856, 863, 864, 865, 866, 868, 886–888, 921, 922, 932, 935, 937, 940, 984, 1116, 1120, 1122–1124, 1263, 1268 Long-term potentiation (LTP), 794, 799, 801, 803, 860, 861, 886–888, 919, 935–937, 940, 1121, 1122 Loss of check, 1615 Loss-of-function, 1573, 1574, 1576 Low birth weight (LBW), 1840, 1843, 1844, 1846 Lox. See Lysyl oxidase (Lox) LRN. see Lateral reticular nucleus (LRN) LTD. See Long term depression (LTD) L-tetraiodothyronine, thyroxine (T4), 321 LTP. See Long term potentiation (LTP) L-triiodothyronine (T3), 321 L-type channel, 1532 Lugaro cells, 245, 830–831, 835, 837–838, 842, 883–885, 917, 918 Lurcher, 135, 136, 263, 1499, 1570 Luria test, 1735 LVN. See Lateral vestibular nucleus (LVN) Lysyl oxidase (Lox), 136

M Macaque monkey, 531–533, 537, 538, 1430, 1435 Machado–Joseph disease/spinocerebellar ataxia type 3 (MJD/SCA3), 1735, 2333–2337 Macrographia, 1615

Index Macrogyric dentate, 426 Mad3, 136 Magnetic resonance imaging (MRI), 578, 713, 1225, 1675, 1677, 1681–1682, 1819, 1822, 1829, 1831, 1833, 1841–1843, 1845, 1847, 1875, 1912, 1964, 1968, 1972, 1974, 2028, 2030, 2032, 2036, 2159, 2168, 2198–2202, 2218, 2219, 2221–2224, 2226–2243, 2246–2249 Magnetic resonance spectroscopy (MRS), 713, 714, 718, 723, 2029, 2032 Male-to-female ratio, 1566 Malformation(s), 1819, 1858–1860, 1863, 1866 Malformation syndromes, 1737 Mandelbrot, 1396, 1404, 1412 MAO. See Medial accessory olive (MAO) MAPK. See Mitogen-activated protein kinase (MAPK) Marmoset, 1430, 1435 Marr, 1360, 1361, 1364, 1366, 1368, 1371, 1374 Marr-Albus theory, 1114 Mash1, 24, 77, 79, 81 Maternal deprivation (MD), 331 Maternal inheritance, 2275, 2277, 2292, 2297–2300 Math1, 24, 132–133 Math1 (Atoh1), 181 Maze learning, 1587 Measles, 2032, 2033 Mechanism, 1964, 1965, 1967–1971, 1973, 1978 Mechanistic, 2161 Medaka, 1430, 1431 Medial cerebellar nucleus, 413 Medial descending motor system, 501 Medial extension of the ventral paraflocculus, 415 Medial intraparietal area (MIP), 556 Medial tegmental tract, 394, 422, 423, 425 Medial vestibular nucleus (MVN), 368, 379, 392, 398, 401, 414, 416 Medial vestibulospinal tract, 507 Medical intervention, 2350–2353 Medulla oblongata, 343, 682, 683, 688 Medullary stroke, 683 Medulloblastoma (MB), 29, 30, 725, 1729, 1732, 2040, 2043 Megalographia, 1615 Meis1, 138–140 Membrane diffusion, 862

2411 Membrane excitability, 811, 814, 818, 821–822 Memory, 1702, 1703, 1710, 1711, 1862, 1864–1866 Mental flexibility, 1724, 1726, 1727 Merlin, 131 MERRF. See Myoclonic epilepsy and ragged red fibers (MERRF) Mesodiencephalic junction, 1026, 1027, 1032, 1041 Metabotropic glutamate receptor type-1 (mGluR1), 134, 135, 140, 264, 266–267, 797 Metaiodobenzylguanidine, 2126 Metric tensor, 1381, 1383, 1386, 1389, 1390, 1391, 1393, 1394, 1406 MF. See Mossy fiber (MF) Microcircuit (cerebellar), 1319–1322, 1324, 1327, 1330, 1331, 1332 Microexplant cultures, 111, 116, 118, 120, 121 Microglial cells, 1583, 1584 Microgravity, 1242, 1245–1248 Microgyric dentate, 426 Microiontophoresis, 896, 897, 899 Microzone, 1113, 1329, 1330, 1332–1334 Midbrain, 343, 344, 352, 355, 682, 684–685, 690–692 Midbrain-hindbrain, 1858, 1860 Midbrain-hindbrain organizer, 12–14 Midbrain tremor, 1612, 1614 Middle intracranial posterior circulation territory, 343 Mid-hindbrain-boundary, 1448, 1450, 1451 Midline, 1018, 1020, 1022, 1025 Mifepristone (RU486), 999 Migraine, 1542, 1547, 1548, 1555–1557 Migration, 96, 1014–1018 Miller Fisher syndrome, 1986, 2000–2002 Mineralocorticoid receptor (MR), 330 Mineralocorticoids, 330 Minocycline, 1584, 1585 MIP. See Medial intraparietal area (MIP) Mitochondria, 1525, 2196, 2208, 2216, 2231, 2238 Mitochondrial inheritance, 1813 Mitogen-activated protein kinase (MAPK), 797, 798 Mitral cell, 1575 Mixed inhibition, 834 ML. See Molecular layer (ML) MLIs. See Molecular layer inhibitory interneuron (MLIs)

2412 Models, 768, 770, 779–784, 1362, 1364, 1366–1368, 1370, 1374 animal, 1564, 1571, 1572, 1575, 1576 compartmental, 1033 computer, 1042 forward, 1168, 1169, 1258, 1259, 1270–1272, 1282, 1291, 1297, 1299, 1300, 1302, 1304, 1308, 1311, 1324 genetic, 1442, 1448, 1456, 1457 Hodgkin-Huxley, 1033 internal, 1193, 1194, 1196, 1282, 1298–1303, 1333, 2354, 2355 inverse dynamics, 1283, 1287, 1288 inverse, 1299 non-mammal animal, 1430 rodent animal, 1432 shared circuits, 1338, 1349, 1354 transgenic mouse, 715, 723 Modified Rankin scale, 1683 Modified version of the International Cooperative Ataxia Rating Scale (MICARS), 1721 Modulation, 916–922 Module, 516 Molecular deficits, 2384 Molecular layer (ML), 64, 108, 110, 239, 1466, 1468–1470, 1472, 1476, 1569, 2162 Molecular layer inhibitory interneuron (MLIs), 799 Monitoring, 1773 Monocarboxylate transporter (MCT), 324 Monoinnervation, 259, 263, 269, 272, 273 Mood, 1777, 1778 Moore-Penrose, 1390 Mormyrid, 1430, 1431 Mormyrid fish, 1465–1466, 1468, 1470–1472 Morphogenesis, 1442 Morphology, 1103 Morris water maze (MWM), 1705 Mossy fiber (MF), 238–240, 358–367, 438, 767–771, 774–780, 782, 831, 833–836, 839, 882–885, 897, 899, 916, 918, 921, 972–975, 977, 983, 1113, 1115, 1121–1123, 1133, 1135, 1137, 1138, 1147, 1148, 1177–1179, 1181, 1287, 1289, 1319–1321, 1328, 1330, 1467, 1472 Mossy fiber (MF) neuron, 81 Motivation, 1772 Motor, 550–566, 735–738, 740, 742–745, 747–749, 751, 755, 1360–1363, 1366, 1373, 1374, 1773, 1775, 1777, 1860–1864, 1866

Index Motor adaptation, 1309, 1310, 2350, 2355, 2360, 2362–2363 Motor behavior, 830, 843, 844 Motor command, 1282–1285, 1287, 1288 Motor control, 1300, 1303, 1304, 1311, 1473, 1475 Motor cortex, 396, 397, 421, 422, 424, 425, 426, 462, 531, 536–540, 542, 1222, 1223, 1226, 1227, 1229–1231, 1234, 1235 Motor features, 2152, 2155–2157, 2160 Motor functions, 1243, 1250 Motor incoordination, 2372 Motor learning, 770, 778, 783, 794, 802, 803, 1037, 1038, 1040, 1041, 1114, 1120, 1122, 1123, 1228, 1229, 1233, 1282, 1285, 1286, 1291, 1308, 1510, 1514, 2156–2157, 2350, 2355, 2362, 2363 Motor neuron, 2201, 2216, 2217, 2234, 2238, 2241 Motor performance, 1037, 1038 Motor rehabilitation, 2350, 2354–2359 Motor skills, 1510 Motor speech, 1758, 1759 Motor thalamus, 529–542 Mouse, 62, 108 Mouse mutants, 70, 1542, 1543, 1545, 1547, 1549, 1554, 1555, 1557 Movement, 550, 566 Movement disorders, 2350, 2354–2357, 2359–2362, 2372 MR. See Mineralocorticoid receptor (MR) MR angiography, 682, 703 MRI. See Magnetic resonance imaging (MRI) MRS. See Magnetic resonance spectroscopy (MRS) MSA. See Multiple system atrophy (MSA) MSA-C, 2328 MSA-P, 2328 Multi-innervation, 258 Multi-joint coordination, 2356–2359, 2362 Multiple system atrophy (MSA), 723, 1608, 1731, 1802, 1809, 2145, 2146, 2148, 2328 Mumps, 2032, 2033 Murine oligodendroglia, 2129 Muscle activity, 1287, 1288 Mutant, 1449, 1457 Mutant TR, 329 Mutism, 1723, 1724, 1730–1732, 1754–1755, 1757–1759, 1764 MWM. See Morris water maze (MWM) Myasthenic syndrome, 1542, 1556

Index Mycn, 132, 134 Myoclonic epilepsy and ragged red fibers (MERRF), 1924–1926, 1929–1931, 1950 Myoclonus, 1923–1931, 1933, 1934, 1937, 1938, 1941, 1944, 1946–1951, 2279, 2287, 2288, 2293, 2294, 2296–2300 Myo-inositol, 715–718, 720–725

N N-acetylaspartate, 715, 721, 724 n-BCA. See n-butyl cyanoacrylate (n-BCA) n-butyl cyanoacrylate (n-BCA), 699 ncDNA, 1381, 1389 ncRNA, 1382, 1389, 1391, 1393, 1396 NE. See Norepinephrine (NE) Necrosis, 1506 Negative image, 1259, 1265, 1267, 1269, 1270, 1272 Neocerebellum, 571, 572, 576, 578, 582, 583, 1338–1344, 1348–1351, 1353, 1354 Neonatal life, 994, 997–1001, 1003–1007 Nephrin, 1017 Nerve terminals, 1542, 1545, 1553, 1554 N-ethylmaleimide sensitive fusion protein (NSF), 865, 866 NETO2, 858, 869 Network, 304 Neural tube, 1014–1017 Neurexin, 249 Neurobehavioral, 1718, 1723, 1726, 1728, 1729, 1732, 1734, 1741 Neurobiology, 770–771 Neurochemical profile, 715, 719–721, 725, 726 NeuroD, 28, 29, 31–32 NeuroD2, 132 Neurodegeneration, 717, 718, 723, 1583–1585, 2205, 2211, 2212, 2215, 2227, 2229–2231, 2238, 2241, 2251, 2382–2385, 2389 Neurodegenerative, 2144, 2158, 2162, 2167 Neuroepithelium, 76 Neurofascin, 245 Neurofilament, 2161 Neurog1, 28, 33–36 Neurog2, 24, 28, 33–36 Neurogenin1, 24 Neurogenin2, 24 Neurogranin, 884 Neuroimaging, 735, 1203, 1205–1207, 1209–1210, 1212, 1214, 1858, 1868,

2413 2152, 2159, 2160, 2162, 2168, 2216, 2218–2221, 2248 Neuroinflammation, 1583 Neuromodulator, 903, 981 Neuromodulatory, 896, 897, 899, 903 Neuromuscular junction, 142, 1553–1555 Neuronal a-synuclein immunoreactivity, 2127 Neuronal cell migration, 108 Neuronal ceroid lipofuscinosis (NCL), 1924–1926, 1931–1935, 1951 Neuronal circuit, 999, 1003, 1005–1007 Neuronal death, 1582, 1584 Neuronal growth, 1001 Neuronal intranuclear inclusions (NIIs), 2332, 2333, 2336, 2338, 2340–2342 Neuronal loss, 2120, 2127 Neuronal nitric oxide-synthase, 797, 798 Neuronal protection, 334 Neuron-glia crosstalk, 1584–1585 Neurons, 2159, 2161, 2164, 2166, 2167 Neuron types, 1102 Neuropathy, 2279, 2280, 2282, 2286–2289, 2293, 2294, 2298–2299 Neuropeptides, 971 Neuroplasticity, 1581–1582, 1587 Neuroprotection, 1001–1003, 1583, 1585 Neuropsychiatric, 1733, 1734, 1739, 1740, 1741 Neuropsychiatry, 1731 Neuropsychological testing, 1723, 1728 Neurosteroids, 995–1001, 1005–1007 Neurotoxic, 2089, 2090, 2096, 2097, 2098, 2101, 2104 Neurotransmitter, 897 Neurotransmitter release, 1542, 1544, 1545, 1554, 1557 Neurotrophic factors, 1002, 1004 Neurotrophin, 1507 Neurotrophin-3 (NT-3), 1004 NF2, 131 NFkB, 132, 134, 137 Ngn1, 24, 79, 81 Ngn2, 24 Nictitating membrane, 1178, 1180 Niemann-Pick type C (NP-C) mouse, 1003 NIIs. See Neuronal intranuclear inclusions (NIIs) Nitric oxide, 777, 797, 798 NMDA, 771, 775–777, 780, 781, 783, 784, 834–836, 1069–1070 NMDA receptor, 113–115, 121, 266, 795, 797, 887, 888, 889, 1004 N-Myc, 132

2414 Nna1, 136, 137, 1564, 1570, 1571, 1573–1576 Nodulus, 347, 351, 363, 380, 414–416, 498, 508, 1774 Nodulus-uvula, 1156, 1158 Noise cancellation, 1322, 1323, 1325, 1327, 1331 Nongenomic action, 1004 Non-genomic mechanism, 331 Non-genomic thyroid hormone action, 324 Non-mammal animal model, 1430 Nonmotor, 550, 551, 555, 557, 559, 560, 561, 565, 566, 1862 Non-progressive cerebellar ataxia, 1731 Non-progressive episodic ataxias, 2380 Noradrenaline, 1508, 1528 Noradrenergic axons, 1526 Norepinephrine (NE), 895 Normalization, 1634, 1635, 1637, 1649, 1651, 1652 Notch, 27, 29 Notch signaling, 27 Novel strategy, 1710 NSF. See N-ethylmaleimide sensitive fusion protein(NSF) N-type channel, 1527 NTZ, 30, 31, 33, 35 Nuclear collateral, 438, 453, 455, 456, 458, 460 Nuclear localization signal, 1573 Nuclear receptor (NR), 320, 321, 330 Nuclear transitory zone, 30 Nucleo-olivary, 1080, 1082, 1085, 1087 Nucleo-olivary pathway, 378, 398, 416, 419, 421, 423, 424 b-nucleus, 1018, 1026, 1027 Nucleus anterior interposed, 379, 392, 397, 398, 401, 402, 407, 413, 417, 421, 422, 499, 506–507 basal interstitial, 379, 380 Bechterew’s, 394–398, 408, 423 cochlear, 83 Darkschewitsch, 394 dentate, 180, 196, 348, 378, 438, 550, 553, 554, 555, 556, 557, 559–561, 565, 1288, 1289, 1719, 1722 descending vestibular, 359 dorsal cochlear, 1259, 1260 dorsal column, 454–455, 1026 dorsal octavolateral, 1259, 1260 emboliform, 79, 184, 189, 191, 197, 355, 1718, 1719, 1722 external cuneate, 80 facial, 1179

Index fastigial, 184, 190, 348, 378, 442, 480, 498, 502, 560, 1288, 1720, 1722, 1731, 1732, 1740 globose, 184, 189, 191, 197, 355, 378, 379, 1718, 1719, 1722 inferior olive, 76, 80 interposed, 1289 interpositus, 180, 184, 190, 442, 448, 449, 560, 1119 interstitial cell groups, 378, 379, 399, 402, 413–415, 422–424, 498, 499, 506 interstitial nucleus of cajal (INC), 513, 514 lateral cerebellar, 378–380, 392, 402, 404, 413, 498, 507 lateralis valvulae, 1473–1475 lateral reticular, 80, 358, 364, 447, 449–453 lateral vestibular, 368, 392, 398, 402, 405, 415, 498 locus coeruleus, 896, 900, 901, 903, 904, 906–908, 910 medial accessory olive (MAO), 380, 1016, 1018, 1019, 1026, 1027 medial cerebellar, 413 medial vestibular, 368, 379, 392, 398, 401, 414, 416 microgyric, 426 parasolitary, 359, 363, 391, 392 pontine gray, 80 posterior interposed, 378, 380, 395, 402, 414, 420, 422–424, 499, 502, 506 prepositus hypoglossi, 390–392 principal olivary, 1720 red nucleus, 509, 510, 515, 1177, 1179, 1181, 1581, 1589 reticulotegmental, 80 subthalamic, 563, 564 superior vestibular, 358, 359, 363, 368, 379, 380, 414, 416 trigeminal, 1026, 1027 ventral anterior, 530, 534 ventral lateral, 530, 531 vestibular, 1111 Nucleus reticularis tegmenti pontis (NRTP), 439, 455 Null mutant, 76 Nystagmus, 513, 1598, 1599, 1602, 1603, 1605, 1617, 1963, 1965–1967

O Oatp. See Organic anion transporter (Oatp) Observational learning, 1588 Observation-learning, 1707

Index Obsessive compulsive disorder, 1723, 1732, 1735 Ocular drift, 1243 Ocular misalignment, 1603, 1606 Ocular motor control, 843 Ocular tilt reaction, 1606 Oculomotor abnormalities, 2156, 2168 Oculomotor disorders, 1787 Oculomotor disturbances, 1598, 1603–1606 Oculomotor function, 1245, 1250 Oculomotor plant, 1325, 1326 Oculomotor system, 1510, 1511 Oculomotor vermis, 1156, 1162–1164 6-OHDA. See 6-hydroxydopamine (6-OHDA) OKR. See Optokinetic reflex (OKR) Olfactory bulb, 1573, 1575 Olig2, 33, 34, 1471, 1476 Olig3, 81, 82, 1017 Olivary axon, 259, 261, 267, 268, 271 Olivary glomeruli, 1026–1027 Olivary pseudohypertrophy, 2071, 2072 Olivocerebellar, 1132, 1133, 1139, 1146, 1147 Olivocerebellar degeneration, 1499 Olivocerebellar loop, 1073 Olivocerebellar pathway, 268 Olivocerebellar projection, 398–416 Olivocerebellar system, 1060, 1064, 1070, 1072–1074 Olivopontocerebellar atrophy (OPCA), 721, 723, 2120, 2122, 2127, 2328 Olivopontocerebellar system, 2127 Onco-neuronal antibodies, 2045, 2048, 2050 Ontogenesis, 768–770 Onyx, 699, 701, 702 Opsoclonus myoclonus, 1986, 1987, 1991, 2002, 2003 Opsoclonus myoclonus syndrome, 1731 Optical imaging, 1531 Optic tectum, 1465, 1473, 1476 Optimal motor control, 1310 Optogenetics, 1124 Optokinetic, 363, 370 Optokinetic reflex (OKR), 906, 907, 1156, 1157, 1160 Orbicularis oculi, 1178 Organic anion transporter (Oatp), 324 Orofacial muscles, 1192 Oscillations, 773, 778, 784, 844–847, 1145, 1531, 1532 Otoliths, 358, 369 Otx1, 129 Otx2, 8, 10–12, 91, 1014, 1015

2415 Oxidative phosphorylation (OXPHOS), 2270, 2271, 2274, 2275, 2278, 2284, 2285 Oxidative stress-induced cell death, 331 OXPHOS. See Oxidative phosphorylation (OXPHOS)

P p53, 1505 PACA. See Primary autoimmune cerebellar ataxia (PACA) PACAP (pituitary adenylate cyclase-activating polypeptide), 116–117 Pain, 486–487 Palatomyoclonus, 1014, 1043 Pallidothalamic, 530, 534, 535, 537, 538, 540 PAN. See Periodic alternating nystagmus (PAN) Pancreatic transcription factor 1a (Ptf1a), 4, 18, 32–35, 63, 68, 70, 76, 92, 133, 139, 181, 1017 Panic disorder, 1728, 1733 Paraflocculus, 414–418, 422 Parahippocampal gyrus, 1720 Paralimbic, 483–485 Parallel fibers (PF), 239–242, 263, 282, 291–293, 360, 363, 768–770, 772, 784, 794, 882–884, 886, 887, 1178, 1258–1263, 1267, 1268, 1269, 1272, 1289, 1320, 1321, 1331, 1361, 1363–1365, 1367–1373, 1466, 1468, 1472, 1476, 1525, 1527, 1528, 1531–1534, 1549, 1550, 1554 Paraneoplastic cerebellar degeneration (PCD), 2040, 2045, 2050 Parasagittal band, 270 Parasolitary nucleus (Psol), 359, 363, 391, 392 Paravermis, 1192 Parietal cortex, 1285 Parkinsonism, 2155 Parkinson’s disease, 565 Parvalbumin, 883, 884 Parvocellular red nucleus, 390, 394–398, 408, 415, 420–425 Parvovirus B19 infection, 2033 Patch clamp, 767, 768, 779 Patched (Ptc), 30, 132 Pathogenesis, 2211–2243 Pathogenetic, 1858 Pathologic laughing and crying, 1730 Pathologies, 782–783 Pathophysiology, 2152, 2160–2168 Pax2, 24, 34, 35, 210, 1016

2416 Pax6, 81, 132, 139, 188, 195, 327 PC. See Purkinje cell (PC) PCA. See Paraneoplastic cerebellar degeneration (PCA) PCL. See Purkinje cell layer (PCL) Pcna, 134 Pcp2(L7) gene, 134, 139 Peduncles, 1774, 1778, 1779 Pedunculo-cerebellar, 355 Pedunculotomy, 267 Pendular tendon reflexes, 1615 Perception, 1775 Periaquaductal gray, 1774, 1776 Pericellular nest, 259, 261, 269 Perinatal critical period, 332 Perinatal hypothyroidism, 322, 323, 326 Periodic alternating nystagmus (PAN), 1603 Peripheral stimulation, 841–842 Perseveration, 1723, 1724, 1733, 1737, 1739 Persistent sodium current, 1117 Personality, 2168 Pervasive developmental disorder, 1734 Petrosal surface, 351 PFS. See Posterior fossa syndrome (PFS) PGN. See Pontine gray nucleus (PGN) Phase reset, 842 Phenotype, 307, 310 Phonation, 1191 Phonemic cluster, 1710 Phosphocreatine, 715 Phospholipase A2, 797, 798 Phosphorylation, 856, 857, 860, 862–865 Photoreceptor cell, 1573, 1574, 1575 Physiotherapy, 2356, 2381, 2386, 2388, 2389 PICA. See Posterior inferior cerebellar artery (PICA) PICK1, 856, 864–865, 869, 870 Picture arrangement subtest, 1704 Pilocytic astrocytomas, 2040, 2043 Pinceau, 243 PKA. See Protein kinase A (PKA) Plasticity, 855–856, 859, 861, 862, 864, 865, 867, 868, 870, 1060, 1113, 1114, 1118–1124, 1227–1229, 1232, 1233, 1235, 1371 PO. See Principal olive (PO) Polyamine, 869 Polyglutamines, 2382, 2383 Polyglutamine tract, 1800 Pons, 343, 344, 348, 350, 352, 354, 469, 682–684, 688, 689, 704, 1674 Pontine gray nucleus (PGN), 80

Index Pontine nuclei (PN), 439, 455–457, 469, 482, 559, 563, 1177, 1178, 1181, 1580, 1581, 1584 Pontocerebellar circuit, 1731 Pore-forming a1 subunit, 1547, 1557 Positron emission tomography, 1682 Posterior cerebral arteries (PCAs), 343, 681, 685, 692, 693 Posterior circulation, 343 Posterior communicating arteries, 353 Posterior communicating artery aneurysms, 693 Posterior fossa, 1857, 1858, 1874–1877, 1881, 1887–1891, 1964 Posterior fossa syndrome (PFS), 1730–1732, 1754–1763 Posterior fossa trauma, 2056 Posterior inferior cerebellar artery (PICA), 680, 688, 693, 1629, 1630, 1728, 1960–1968, 1970–1972 Posterior inferior cerebellar artery aneurysms, 693 Posterior interposed nucleus (PIN), 378, 380, 395, 402, 414, 420, 422–424, 499, 502, 506 Posterior parietal cortex, 556, 565 Post infectious cerebellitis, 1723, 1731, 1986, 1989, 1998–2000, 2002 Postinhibitory rebound, 1123 Post-mitotic, 79 Postmitotic neurons, 107 Postmortem studies, 2155, 2157, 2165 Postnatal development, 239, 246 Postsynaptic density, 797 Postural tremor, 1611–1614, 2153–2155 Posture, 1222 Posture and gait, 1787–1789 Potassium channels, 1534 PP1, 887, 889 PP2B (calcineurin), 887, 889 P/Q-type Ca2þ channel, 135, 1066 P/Q-type channel, 1523, 1525–1529, 1532 PR. See Progesterone receptor (PR) Precerebellar nuclei, 514 Precerebellar systems, 80 Precision tremor, 1612 Prediction, 1702, 1703 Predictive errors, 1311 Prefrontal cortex, 554–556, 558, 563, 566 Premature birth, 1737 Premotor cortex, 422, 424, 539, 540 Presupplementary motor area (PreSMA), 532, 538, 539, 552, 554, 556

Index Pretectal complexes, 1026 Pretecto-olivary projection, 393 Pretectum, 309, 392–394, 412, 422 Preterm infants, 1734 Primary afferent, 358–360 Primary autoimmune cerebellar ataxia (PACA), 1986–1991, 1997, 1998 Primary dendrite, 1466–1470 Primary motor cortex (M1), 552, 553, 563, 1286 Primate, 1430, 1435–1436 Principal olivary nucleus, 1720 Principal olive (PO), 404, 1016, 1018, 1026, 1027 Prion diseases, 2035–2036 Prism adaptation, 1308, 1309, 1311 Procedural competence, 1588 Procedural learning, 1707, 1709 Procedural memory, 1711 Procedures, 1705–1707, 1711, 1712 Progesterone, 996–1007 Progesterone receptor (PR), 1000, 1003 Progressive myoclonic epilepsy, 1923 Projection maps, 149, 167 Projection neurons, 1102–1109 Promyelocytic leukemia protein nuclear bodies, 2336 Proneural genes, 24 Protein interacting with C-kinase 1, 797 Protein kinase A (PKA), 887, 889 Protein kinase C, 795, 797 Protein kinase G type 1, 797 Protein phosphatise, 797, 798 Protein toxicity, 2211, 2214, 2251 Proximal exertional tremor, 1614 Proximal intracranial posterior circulation territory, 343 Pseudobulbar palsy, 1730 Pseudochoreoathetosis, 1672 Psychiatric, 2155, 2158 Psychiatric disorders, 331 Psychopathology, 1772, 1778, 1779, 1780 Psychosis spectrum disorders, 1733, 1735 Ptc. See Patched (Ptc) PTEN, 137 Ptf1a. See Pancreatic transcription factor 1a (Ptf1a) Purkinje cell (PC), 4, 16–18, 44, 63, 64, 68, 69, 77, 90, 97–98, 115, 168, 238, 245–246, 258, 282–283, 360, 438, 558, 769, 770, 773, 783, 784, 793, 830, 831, 833, 837, 839, 841, 843, 882–891, 896–908, 917, 918, 972, 975–986, 995–1007, 1020,

2417 1022, 1023, 1029, 1032–1033, 1035–1038, 1040, 1041, 1060–1064, 1067, 1071–1073, 1081, 1083, 1084, 1086–1088, 1090, 1093, 1112–1117, 1119, 1123, 1124, 1157–1160, 1162–1165, 1167, 1244, 1245, 1248–1250, 1287–1289, 1292, 1319–1321, 1328–1333, 1361, 1363–1374, 1384, 1389, 1393–1396, 1402, 1409, 1412, 1466–1467, 1470–1472, 1476, 1499–1508, 1513–1515, 1524–1529, 1531–1534, 1546, 1549–1555, 1564, 1568–1576, 1897–1899, 2017, 2083, 2085, 2086, 2091, 2095, 2096, 2100, 2102, 2105, 2106, 2166–2169, 2382 Purkinje cell degeneration (pcd), 136, 1483, 1564, 1570, 1572–1576 Purkinje cell layer (PCL), 110–111, 239 Pursuit, 1156–1160, 1162, 1164–1168, 1288, 1289 Putamen, 561 Putaminal structures, 2125 P2X receptors, 948, 949, 952 Pyramidal sign, 1670, 1671, 1679 Pyramidal tract, 1670, 1677 Pyramis, 347 P2Y receptors, 948–950, 952, 955

Q Q fever, 2033 Quadrupedal locomotion, 1436 Quail-chick chimeras, 1432 Quality of life, 2361, 2363

R Rabbit, 1430, 1432, 1434 Rabies virus, 515, 517 Radial migration, 108–110, 115, 116 Ragged-red fibers, 2287, 2288, 2291–2294, 2296, 2297, 2300, 2301 Rasagiline, 2120, 2130 Rat, 994, 996, 999, 1003, 1004, 1006, 1007 Rate of migration, 108, 113, 114 RCM. See Rostral Cerebellar Malformation (RCM) Reading, 743–745 Rebound, 1615 Rebound burst, 1118 Rebound depolarization, 919, 921, 1062–1063 Rebound potentiation (RP), 887

2418 Receptive field, 831, 843, 844 a1 Receptor, 905 a2 Receptor, 906 b Receptor, 904–909 Receptors, 768–771, 777, 783, 784, 853 Recessive ataxia, 1432, 1434, 2373, 2375 Recovery, 2350–2351, 2356, 2361 Recursion, 1384–1387, 1389, 1390, 1393, 1394, 1396, 1404, 1409, 1410 Recursive genome function, 1383 Red nucleus (RN), 509, 510, 515, 1177, 1179, 1181, 1581, 1589 Reeler (rl), 128, 1570. See also Rln Reelin, 150, 334, 1899, 1900, 1902 Reference signal, 1325, 1326 Reflex, 508, 516, 519 Reinnervation, 258 Remote damage, 1582 Repeat, 2194, 2198–2202, 2205–2209, 2211, 2215, 2218, 2223–2231, 2234, 2239–2242, 2246, 2250 Reptile, 1430, 1431 Resistance to thyroid hormone, 328, 329 Resonator, 1033 Respiratory chain, 2270, 2273–2278, 2284, 2285, 2290–2292, 2294, 2297 Resting membrane potential, 123 Resting state, 571, 574–578, 580 Resting state analysis, 2362 Rest tremor, 2152, 2155 Reticular formation, 500 Reticulospinal tract, 501–502 Reticulotegmental nucleus (RTN), 80 Retina, 1573–1575 Retinal slip, 1325, 1326, 1328 Retinoic acid, 66, 71, 1014–1017 Retinoic acid-related orphan receptor (ROR), 326 Retinoid X receptor (RXR), 321 Retrograde messengers, 931, 932 Reward, 1774 Rey-Osterreith Figure, 1727 Rhombencephalon, 1464, 1472, 1473 Rhombencephalosynapsis (RES), 197–198, 1737 Rhombic lip, 16, 18, 92, 181, 1015–1017, 1022 lower, 16 upper, 16, 24, 28, 31–33, 35 Rhombomere, 63, 64, 76, 80, 83, 1014–1017 Rhythm, 1206, 1207 Rhythmic, 2156 Rhythmic activity, 1063–1064

Index Rifampicin, 2120, 2130 Riluzole, 2352 Rituals, 1737 rl. See Reeler (rl) Rln, 1570 Robo3, 1020, 1022 Rodent animal models, 1432 Rolling Nagoya, 1542 Roof plate, 8, 9, 76 Rora (RORa), 133–134, 161–162, 1570 Rostral cerebellar malformation (RCM), 132 Rotarod, 1571, 1572 Rotarod test, 1244 Rotavirus gastroenteritis, 2033 RP. See Rebound potentiation (RP) RTN. See Reticulotegmental nucleus (RTN) R-type channel, 1528 Rubella, 2032, 2033 Rubrospinal tract, 509–510 Ryanodine receptors, 1526, 1533, 1549

S SA. See Spinal atrophy (SA) Saccade, 843, 844, 1156, 1158, 1162–1164, 1166–1168, 1285 Saccade adaptation, 1167 Saccadic pursuit, 1599, 1603 SAH. See Subarachnoid hemorrhage (SAH) SAOA. See Sporadic adult-onset ataxia (SAOA) SAP-97, 866, 867, 869 SARA. See Scale for the assessment and rating of ataxia (SARA) sc, 26 SCA. See Spinocerebellar ataxia (SCA); Superior cerebellar artery (SCA) Scale for the assessment and rating of ataxia (SARA), 1787, 1790, 1792–1795, 2357–2359, 2361 Schizophrenia, 490, 726, 1183, 1184, 1334, 1734, 1735, 1741, 1778, 1780, 1907 Screening, 1456 Scripts, 1704–1705 Secreted signals, 61, 64–66 Seizures, 1896, 1898, 1900, 1923, 1927–1930, 1933–1935, 1937, 1938, 1941, 1944, 1945, 1947, 1949–1951 Sema6a, 132 Semantic cue retrieval, 1704, 1709, 1710 Semicircular canals, 363, 365, 366, 369 Semilunar lobule, 347, 355

Index Sensorimotor, 735–741, 745, 746, 748–750, 755, 756 Sensorimotor cerebellum, 1720 Sensorimotor coordination, 1383, 1384, 1387, 1393, 1396, 1406 Sensorimotor learning, 1210–1211 Sensory, 1362, 1363, 1372, 1373 Sensory ataxia, 1801, 2279, 2287–2288 SEP. See Somatosensory evoked potentials (SEP) Septum, 1774 Sequence processing, 1588 Sequencing, 1702 Serial reaction time task (SRTT), 1707, 1708 Serotonin, 915, 1068 Serotonin 5-HT1A receptor, 370 Serotonin receptor, 916 Set shifting, 1723, 1724, 1728 Sex-dependent, 1243, 1250 Sexual differentiation, 330 Sexual dimorphism, 1250 Sexually dimorphic neurogenesis, 332 Shared circuits model, 1338, 1349, 1354 SHH. See Sonic hedgehog (SHH) Short latency response(SLR), 119 Short-term adaptation, 2362 Short-term depression (STD), 1122 Short-term synaptic plasticity, 802 Shy-drager syndrome (SDS), 2120, 2328 Sialidosis, 1924, 1926, 1946–1948, 1951 Signaling centers, 62, 70 Signal processing, 1317, 1322, 1325, 1330, 1331, 1333 Signal to noise ratio, 897, 901, 903, 905 Sign of the piano, 1609 Silent synapses, 1331 Simple spike (SS), 363, 981, 982, 1287, 1288, 1289, 1320, 1321, 1328, 1332 Simulation, 1035, 1038, 1040, 1042, 1366 Simultanagnosia, 1723, 1724 Single-photon emission computed tomography, 1680–1683. See also SPECT Skew deviation, 1603, 1606 Slc1a6, 134 Slc6a5, 139 Slices, 768 Slit, 1018, 1020 SLR. See Short latency response (SLR) Smith predictor, 130, 1311 Smooth pursuit, 1156–1158, 1160, 1164 Snca, 139, 140 Social cognition, 1733

2419 Social cues, 1733, 1737 Social emotion, 1734, 1736 Social skill set, 1714, 1733–1735 Soluble-Guanylyl-Cyclase, 797, 798 Soma, 109, 112 Somatosensory evoked potentials (SEP), 1679–1681, 1702 Somatosensory mismatch negativity, 1703 Somatosensory modulation, 1589 Somatostatin, 115–116 Somatotopical localization, 386 Somatotopy, 739–741, 1719 Song, 1191 Sonic hedgehog (SHH), 30, 65, 69–70, 89, 132, 139 Sox9, 34 Space, 1242 Spasticity, 1671 Spatial, 736, 738, 742, 745, 747–750, 755 Spatial cognition, 1724 Spatial dysgraphia, 1711 Spatial functions, 1705 Spatially unbiased atlas template of the human cerebellum (SUIT), 1635, 1637, 1649 Spatial procedural learning, 1588 SPECT, 1760–1762, 1764 Speech, 1191, 1712 Spermatogenesis, 1573, 1576 Spinal atrophy (SA), 721–723 Spinal cord, 2385 Spinocerebellar ataxia (SCA) 1, 2, 3, 6, 7, 8, 370, 721–724, 890, 1429, 1432, 1631, 1633, 1649, 1743, 1800, 1803–1807, 1809–1812, 1901, 2270, 2280, 2283, 2287, 2289, 2291–2292, 2294, 2331–2333, 2337, 2338, 2351, 2352, 2373–2375, 2380–2381, 2383–2385, 2389 Spinocerebellar syndromes, 1735 Spinocerebellar tracts, 1718, 1719 Spinogenesis, 241 Spino-olivocerebellar, 1132, 1147 Sporadic adult-onset ataxia (SAOA), 1632, 1633, 1809 Sporadic ataxia, 1432, 1434 Staggerer (sg), 133, 134, 137, 241, 326, 1570 StAR. See Steroidogenic acute regulatory protein (StAR) Stargazer, 135, 136 Stargazing, 854, 856, 857, 861, 869 State estimation, 1300 State estimator, 1333 State prediction, 1298–1305, 1307, 1308, 1311

2420 STD. See Short term depression (STD) Stellate cell, 77, 208, 222, 223, 241, 243, 245, 247, 799, 809, 882–884, 886, 887, 897, 899, 900, 904, 918, 1466, 1468 Steroidogenic acute regulatory protein (StAR), 997, 1001, 1002 Steroid receptor coactivator (SRC)-1, 325 Steroids, 320 Stewart-Holmes maneuver, 1615 STO. See Subthreshold oscillations (STO) Straight cells, 1018–1019 Stress, 1522, 1524, 1528, 1532–1534 Stress responses, 330 Striatonigral degeneration (SND), 2120, 2127, 2328 Stroke, 1581, 1582, 1585, 1693, 1694, 1721–1731, 1737, 1740, 1960, 2292–2297 Structural magnetic resonance imaging, 1778 Structural plasticity, 1123 Subarachnoid hemorrhage (SAH), 695, 703, 2056, 2059, 2065, 2069, 2070 Subdural hematoma, 2057, 2061, 2064, 2066, 2073 Submarginal strand, 1018 Suboccipital cerebellar surface, 344 Sub occipital decompressive craniectomy, 687 Subthalamic nucleus, 563, 564 Subthreshold oscillations (STO), 1028–1032, 1036, 1038, 1042, 1062–1068, 1070–1074 Subtraction analysis, 1638, 1640–1644 Subtype, 77, 83 SUIT. See Spatially unbiased atlas template of the human cerebellum (SUIT) Sulcus, 353, 355 Superficial siderosis of the CNS, 2072, 2073 Superior cerebellar artery (SCA), 345, 353–355, 680, 692, 1629, 1630, 1722, 1728, 1960, 1962–1971 Superior colliculus, 511 Superior surface, 344 Superior vestibular nucleus (SVN), 358, 359, 363, 368, 379, 380, 414, 416 Supplementary motor area (SMA), 532, 537–539, 552–556 Supramodal, 1717 Swaying (sw), 129 Swimming, 1464, 1475, 1476 Symmetrical LTD and LTP, 1331 Synapse, 1542, 1546, 1550, 1552–1557 Synapse development, 238 Synapse elimination, 258–261

Index Synapse formation, 258, 263–264 Synapse maturation, 271–272 Synapses, 238 Synapse specific, 249, 1119, 1121 Synapse stabilization, 264 Synaptic competition, 242, 262, 270, 273, 275, 291–293 Synaptic plasticity, 768, 775–778, 794, 799, 802–803, 812–817, 822, 916, 922 Synaptic strength, 793, 794, 802, 803 Synaptic transmission, 915, 1527–1529, 1531, 1534 Synaptogenesis, 241, 244, 247, 249, 258, 994, 999–1007 Synchronization, 1067, 1204, 1205 Synchrony, 1031–1033, 1035–1038 a-Synuclein, 2164–2166, 2328, 2329, 2331, 2332 Syrian hamster, 1564 Syringomyelia, 1874–1876, 1878, 1880, 1883

T T3. See L-triiodothyronine (T3) T4. See L-tetraiodothyronine, thyroxine (T4) Tactile, 1372, 1373 Tactile defensiveness, 1733, 1737 Tamoxifen, 1001, 1004, 1005 Tangential migration, 109, 111, 115 Targetome, 128–129 TARPs. See Transmembrane AMPAR regulatory proteins (TARPs) Taurine, 715, 717, 720–725 Tbr1, 187 Tbr2, 188, 193, 195 tDCS. See Direct current stimulation (tDCS) Teaching signal, 1317, 1318, 1320–1327, 1330, 1333 Tecto-olivary projection, 393, 394, 411 Tectospinal tract, 511 Tectum, 392–394, 422 Telencephalon-cerebellar pathway, 1473 Teleost, 1430, 1442, 1445, 1447, 1456, 1464 Temporal processing, 1206, 1209, 1214, 1215 Tensor network theory (TNT), 1382, 1386–1390, 1394 Tentorial, 343, 344 Testis, 1570, 1571, 1573, 1574, 1576 Testosterone, 331 Test-retest reliability, 1786, 1788, 1793, 1794 TH. See Tyrosine hydroxylase (TH) Thalamic neuron, 1576 Thalamocortical, 530

Index Thalamus, 343, 550, 559, 560, 561, 563, 1719, 1735, 1774, 1775 Thanatophoric dysplasia (TD), 197, 198 Theory of mind, 1733, 1734 Therapeutic strategies, 2372, 2382–2385, 2389 Therapies, 1786, 1787, 1790, 1792 Theta, 1120 4th ventricle roof plate, 62, 64–66, 68–69 Thyroid, 2010, 2014, 2017, 2021 Thyroid hormone, 321–335 Thyroid hormone receptor (TR), 133, 137, 321 Thyroid hormone response element (TRE), 322 Thyrotropin (TSH), 327, 329 Thyroxine, 321 Time-lapse recording, 119 Time perception, 1210 Timing, 776, 780, 781, 783, 784, 1112, 1115, 1118–1121, 1125, 1361–1363, 1370, 1371, 1703, 1704 Titubation, 1616 TMS. See Transcranial magnetic stimulation (TMS) TNT. See Tensor network theory (TNT) Tonsil, 345, 347 Tool, 1291 Topographic organization, 1717 Topography, 735, 1598, 1609, 1617–1621 Torpedoes, 2160–2162, 2166, 2168 Tottering (tg), 135, 1542, 1545, 1547, 1549, 1553, 1557 Tottering mouse, 1522 Toxicity, 2081, 2090, 2092, 2095, 2097, 2099, 2101, 2107 tPA, 136 TR. See Thyroid hormone receptor (TR) Trace conditioning, 1119, 1120, 1176 Trafficking, 855–859, 861–866, 869 Trailing process, 109, 110, 113 Transcranial magnetic stimulation (TMS), 1203, 1206, 1286–1287, 1306, 1307, 1741, 1777 Transcription factors, 76, 181, 192, 193, 1383, 1391, 2336, 2340 Transduction, 770–771 Transgenic mouse model, 715, 723 Transmembrane AMPAR regulatory proteins (TARPs), 857–860, 862, 863, 866, 869 Transmitter release, 919 Transneuronal tracers, 551 Transneuronal tracing, 520 Transneuronal transport, 424 Transsynaptic tracers, 550 Trauma, 1670, 1675, 1682

2421 TRE. See Thyroid hormone response element (TRE) Treadmill training, 2359 Tremor, action, 1611, 1612 arm, 2154 asthenic, 1612 axial postural, 1612, 1614 cerebellar, 1610, 1614, 2352, 2354, 2355 cranial, 2154 essential, 2152 fragile X tremor/ataxia syndrome (FXTAS), 1807, 1810, 2374, 2380, 2381 head tremor, 2154, 2156, 2159 intention, 1671, 2154–2156, 2167 isometric, 1611, 1613 kinetic, 1608, 1611, 2152–2155 leg, 2154, 2155 midbrain, 1612, 1614 postural, 1611–1614, 2153–2155 precision, 1612 proximal exertional, 1614 rest, 2152, 2155 TR gene knockout, 322, 327, 328 Triangle of Guillain-Mollaret, 514 Trigeminal nerve, 355 Trigeminal nuclear complex, 1177, 1178 Trigeminal nucleus, 1026, 1027 Trigemino-oliovary projection, 386, 394 Triggers, 1522, 1528, 1532, 1533, 1534 Triiodothyronine, 321 Trinucleotide repeat, 1800, 1810 Triplet expansion, 1936 Trophic influence, 1738 Trpc3, 139, 140 T-type calcium channel, 1062–1063, 1066, 1117 Tuber, 347 Turtle, 1430, 1431 Tyrosine hydroxylase (TH), 1546, 1548

U UBC. See Unipolar brush cells (UBC) Ubiquitin, 2333, 2336, 2338, 2341, 2342 Ubiquitinated, 2166, 2333 UBN. See Upbeat nystagmus (UBN) Unc5c, 132 Uncinate tract, 502 Unconditioned response (UR), 1176–1180 Unconditioned stimulus (US), 1116, 1118, 1176–1184

2422 Unipolar brush cells (UBC), 62, 63, 66, 67, 238, 239, 834, 1468 Unipolar disorder, 1780 Universal cerebellar impairment, 1738 Universal cerebellar transform, 1738 Unverricht-lundborg disease (ULD), 1924–1929, 1948, 1950 Upbeat nystagmus (UBN), 1605 UR. See Unconditioned response (UR) Urocortin, 973, 976, 977 Urocortin II, 973 US. See Unconditioned stimulus (US) Uvula, 347, 358, 411, 414, 415

V Vaccination, 2031, 2033 Validation, 1786, 1793 Valvula cerebelli, 1465, 1467, 1468, 1470, 1472–1476 Varicella, 2033, 2034 VBM. See Voxel-wise morphometry (VBM) VDCC. See Voltage dependent Ca2þ channels (VDCC) Velocity storage, 1156, 1161 Venous thrombosis, 1974 Ventral anterior nucleus, 530, 534 Ventral lateral interparietal area (LIPv), 556 Ventral lateral nucleus, 530, 531 Ventral paraflocculus, 1157, 1158, 1160 Ventral premotor area (PMv), 552, 553, 555 Ventral striatum, 1774, 1776 Ventral tegmental area, 481 Ventricular zone (VZ), 24, 25, 29, 33–36, 76 Ventrolateral outgrowth (VLO), 381, 383, 1018, 1026, 1027, 1030 Verbal fluency, 1709, 1723, 1724, 1727, 1728, 1735, 1736, 1737, 1739 Verbal production, 1192 Vergence, 1605 Vermis, 345, 347, 480, 718–724, 726, 1244, 1248, 1719–1724, 1727–1732, 1734, 1735, 1740, 1774–1778, 1857, 1860, 1864, 1866–1868, 1897, 1900 Vertebral artery, 680–683, 686, 696, 699 Vertebrate neurogenesis, 27 Vertebrobasilar artery junction, 350 Vertebrobasilar dilatative arteriopathy (VBDA), 694 Vertigo, 1598, 1601, 1602, 1603, 1961, 1962, 1965–1967, 1969, 1977 Vestibular, 1580, 1581, 1582, 1586 Vestibular ataxia, 1801

Index Vestibular cerebellum, 1243, 1248 Vestibular nuclei, 350, 351, 498, 882, 884, 888 Vestibular nucleus, 1111 Vestibular reflex, 830 Vestibular stimulation, 843, 844 Vestibular system, 358, 360, 1222, 1224–1226, 1235 Vestibulocerebellar system, 1242, 1246 Vestibulocerebellum, 501, 834, 844, 1157, 1158, 1161, 1774, 1775 Vestibulolateral lobe, 1466, 1467, 1468, 1470 Vestibuloocular reflex (VOR), 794, 802, 906, 907, 1114, 1120, 1123, 1156–1161, 1164–1166, 1168, 1324, 1326, 1599, 1603, 1606 Vestibulospinal tracts, 507–508 Virus, 2032, 2033 Virus tracing, 562 Visceral functions, 483 Visoumotor adaptation, 2355, 2362, 2363 Visual acuity 1C2, 2333, 2336, 2338, 23340, 23341 NII, 2332, 2333, 2336, 2338, 23340, 23342 Visual spatial processing, 1717 Visuomotor integration, 1308, 1309 Visuospatial, 1862–1864, 1866, 1867 Visuospatial learning, 1707, 1712 Vitamin C, 718 VLO. See Ventrolateral outgrowth (VLO) Voltage-clamp, 1550, 1551, 1554 Voltage dependent Ca2þ channels (VDCC), 135 a1AVoltage-dependent calcium channel, 2337 Voltage-dependent calcium channels, 264–266 Voltage-gated Ca2þ channels, 1542, 1545 Voltage-Gated-Calcium-Channel, 797 VOR. See Vestibuloocular reflex (VOR) VOR adaptation, 1157, 1164, 1165, 1328 Voxel-based lesion-symptom mapping, 1721 Voxel-wise morphometry (VBM), 1645, 1646, 1649 Voxel-wise statistical mapping, 1543–1545

W o-agatoxin-IVA, 1544, 1554 Waggler, 135 Walking, 1284 Walking aids, 2357, 2358, 2360 Wallenberg’s syndrome, 350, 682 Wavelets, 1028, 1029 WCST. See Wisconsin Card Sort Test (WCST) Weaver (wv), 128, 241, 247, 1570

Index Weber’s syndrome, 685 Wechsler adult intelligence scale-revised, 1704 Weightlessness, 1242, 1245, 1246 Weyl Law, 1382, 1389, 1396, 1411 White matter, 1912–1917 Whooping cough, 2033 Wisconsin Card Sort Test (WCST), 1725, 1735 WNT, 24 Wnt-1, 7, 12, 82, 129, 1016, 1017 Word retrieval, 1710 Working memory, 735, 738, 742, 745–754, 1195, 1196, 1702, 1710, 1711, 1720, 1723, 1724, 1726, 1727, 1732, 1736, 1737, 1739 Writing, 1711–1712

X Xath1, 28 Xath5, 28 X-chromosomal, 2315 Xenopus laevis, 28, 31

2423 X-linked ataxia, 1807, 1809, 1810, 1811 XneuroD, 28 X zone, 398, 399, 407–409

Y Y zone, 399, 407, 409, 421, 422

Z Zebra finch, 1430, 1432 Zebra fish, 1430, 1435, 1442, 1466, 1471, 1476 Zebrin, 268, 271, 399, 445, 457, 459 Zebrin II, 44, 1470, 1471 Zic1, 29 Zinc-binding domain, 1574 Zipf-Mandelbrot Parabolic Fractal Distribution, 1382, 1394, 1402–1405, 1406, 1412 Zone, 1132, 1135, 1137, 1139, 1143, 1145, 1146, 1328, 1329, 1332