Animal Models of Movement Disorders: Volume II (Neuromethods 62)

Neuromethods

Series Editor Wolfgang Walz University of Saskatchewan Saskatoon, SK, Canada



For further volumes: http://www.springer.com/series/7657

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Animal Models of Movement Disorders Volume II

Edited by

Emma L. Lane The Brain Repair Group, Welsh School of Pharmacy, Cardiff University, Cardiff, Wales, UK

Stephen B. Dunnett The Brain Repair Group, School of Biosciences, Cardiff University, Cardiff, Wales, UK

Editors Emma L. Lane The Brain Repair Group Welsh School of Pharmacy Cardiff University Cardiff, Wales, UK [email protected]

Stephen B. Dunnett The Brain Repair Group School of Biosciences Cardiff University Cardiff, Wales, UK [email protected]

ISSN 0893-2336 e-ISSN 1940-6045 ISBN 978-1-61779-300-4 e-ISBN 978-1-61779-301-1 DOI 10.1007/978-1-61779-301-1 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2011936395 © Springer Science+Business Media, LLC 2011 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or d ­ issimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper Humana Press is part of Springer Science+Business Media (www.springer.com)

Preface to the Series Under the guidance of its founders Alan Boulton and Glen Baker, the Neuromethods series by Humana Press has been very successful since the first volume appeared in 1985. In about 17 years, 37 volumes have been published. In 2006, Springer Science + Business Media made a renewed commitment to this series. The new program will focus on methods that are either unique to the nervous system and excitable cells or which need special consideration to be applied to the neurosciences. The program will strike a balance between recent and exciting developments like those concerning new animal models of disease, imaging, in vivo methods, and more established techniques. These include immunocytochemistry and electrophysiological technologies. New trainees in neurosciences still need a sound footing in these older methods in order to apply a critical approach to their results. The careful application of methods is probably the most important step in the process of scientific inquiry. In the past, new methodologies led the way in developing new disciplines in the biological and medical sciences. For example, Physiology emerged out of Anatomy in the nineteenth century by harnessing new methods based on the newly discovered phenomenon of electricity. Nowadays, the relationships between disciplines and methods are more complex. Methods are now widely shared between disciplines and research areas. New developments in electronic publishing also make it possible for scientists to download chapters or protocols selectively within a very short time of encountering them. This new approach has been taken into account in the design of individual volumes and chapters in this series. Neuherberg, Germany

Wolfgang Walz

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Preface Movement is controlled by the interaction of many component parts of the central nervous system, from myelinated motor neurons projecting from the spinal cord to the higher neural processes in cerebellum and basal ganglia. This produces a highly controllable, functional system. However, this finely integrated network can be disrupted by injury and a range of disease processes that lead to significant motor dysfunction. Damage to single elements of this circuitry, which result from both sporadic and genetic conditions, can cause profound alterations in motor function. In order to understand these disorders and thereby facilitate recovery and repair, it is necessary to translate in vitro findings and hypotheses into animal representations of both functional and dysfunctional systems. These animal models range in species from lower orders, such as Drosophila and Caenorhabditis elegans, through vertebrate species including fish, to mammals, such as rodents and nonhuman primates. Each model has its own profile of face, construct, and predictive validities, all of which must be considered when selecting the most appropriate for the experiment in hand. Similarly, the assessment methods used will depend on the species and the outcome variables that need to be assessed and must be similarly scrutinized for validity to answer the postulated hypothesis. In the first volume (Neuromethods, vol. 61), we introduced the variety of tools used in the assessment of motor function, highlighting their advantages and limitations and noting important technical considerations. We first take a look through the clinician’s perspective on animal models of disease, before exploring both simple (e.g., Drosophila) and more complex (rodent and nonhuman primate) model systems and reviewing the use of genetic manipulations, behavioral assessments, and the increasing use of imaging techniques. We then take a journey, descending through the central nervous system, describing animal models of disorders that target different levels of motor control. One interesting development found through the process of formulating this volume was the overlap in rodent behavioral techniques that are used across a range of motor disorders. Importantly, despite their wide use, each laboratory has its own approach to each behavioral technique. Many of the standard tests appear simple on first inspection, but a critical eye is required, and seemingly insignificant manipulations can produce critical differences in the outcomes and interpretation of the data produced. The first volume went on to consider dopaminergic influences over motor control ­typified in the motor disorder, Parkinson’s disease. The interest in this disorder, aided by significant developments such as the accidental discovery of a toxin, MPTP, that produces pathology similar to the disease process, and identification of specific genes involved in the familial form of the disease have led to an extraordinary array of animal models. Furthermore, we not only need to model the disease, but also the consequences of pharmacological treatment, the development of another dopaminergic motor phenotype, l-dopa-induced dyskinesia. In the present volume (Neuromethods, vol. 62), we expand these themes to cover ­animal models of other basal ganglia disorders such as Huntington’s disease and multiple systems atrophy, then through neo- and allo-cortical systems to describe models of ischemia and eye movement control. Descending through the cerebellum, there is a description of

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the role of this complex nucleus in the control of fine motor function, disorders that affect this “little brain” and how they are represented in vivo. The last section considers the role of spinal cord systems, including the modeling of spinal cord injury, demyelinating disorders, and amyotrophic lateral sclerosis. We would like to take this opportunity to thank the teams that contributed chapters and in particular to acknowledge the more junior members who are often those actually at the coalface of these experiments. We also regret the loss of some chapters (thankfully, very few), a consequence of the ever-increasing demands on the time of researchers. We hope that this text will be a valuable reference for those studying motor disorders by covering methodologies in detail and providing the information necessary to consider both the appropriate models and assessment tools that can most informatively answer the key experimental issues in our field. Cardiff, UK

Emma L. Lane Stephen B. Dunnett

Contents Volume I Series Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Part I  Generic Methods of Assessment   1 Why Cannot a Rodent Be More like a Man? A Clinical Perspective . . . . . . . . . . . . . . 3 Anne E. Rosser   2 Zebrafish as a Vertebrate Model Organism for Studying Movement Disorders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11 Maria Sundvik and Pertti Panula   3 Methodological Strategies to Evaluate Functional Effectors Related to Parkinson’s Disease Through Application of Caenorhabditis elegans Models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31 Kim A. Caldwell and Guy A. Caldwell   4 Effects of Alpha-Synuclein Expression on Behavioral Activity in Drosophila : A Simple Model of Parkinson’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55 Robert G. Pendleton, Xiaoyun C. Yang, Natalie Jerome, Ornela Dervisha, and Ralph Hillman   5 Neurological Evaluation of Movement Disorders in Mice . . . . . . . . . . . . . . . . . . . . . 65 Simon P. Brooks   6 Rodent Skilled Reaching for Modeling Pathological Conditions of the Human Motor System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87 Jenni M. Karl and Ian Q. Whishaw   7 High-Throughput Mouse Phenotyping . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109 Sabine M. Hölter and Lisa Glasl   8 MRI of Neurological Damage in Rats and Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 Mathias Hoehn   9 Quantification of Brain Function and Neurotransmission System In Vivo by Positron Emission Tomography: A Review of Technical Aspects and Practical Considerations in Preclinical Research . . . . . . . . . . . . . . . . . . . 151 Nadja Van Camp, Yann Bramoullé, and Philippe Hantraye

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10 Optical Approaches to Studying the Basal Ganglia . . . . . . . . . . . . . . . . . . . . . . . . . . 191 Joshua L. Plotkin, Jaime N. Guzman, Nicholas Schwarz, Geraldine Kress, David L. Wokosin, and D. James Surmeier 11 Electrophysiological Analysis of Movement Disorders in Mice . . . . . . . . . . . . . . . . . 221 Shilpa P. Rao, Véronique M. André, Carlos Cepeda, and Michael S. Levine

Part II Dopamine Systems 12 Genetic Models of Parkinson’s Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ralf Kühn, Daniela Vogt-Weisenhorn, and Wolfgang Wurst 13 6-OHDA Lesion Models of Parkinson’s Disease in the Rat . . . . . . . . . . . . . . . . . . . Eduardo M. Torres and Stephen B. Dunnett 14 6-OHDA Toxin Model in Mouse . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gaynor A. Smith and Andreas Heuer 15 Rotation in the 6-OHDA-Lesioned Rat . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Stephen B. Dunnett and Eduardo M. Torres 16 Of Rats and Patients: Some Thoughts About Why Rats Turn in Circles and Parkinson’s Disease Patients Cannot Move Normally . . . . . . . . . . . . . . . . . . . . . Gordon W. Arbuthnott 17 Comparing Behavioral Assessment of Sensorimotor Function in Rat and Mouse Models of Parkinson’s Disease and Stroke . . . . . . . . . . . . . . . . . . . . . . . Sheila M. Fleming and Timothy Schallert 18 Rodent Models of l-DOPA-Induced Dyskinesia . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hanna S. Lindgren and Emma L. Lane 19 Using the MPTP Mouse Model to Understand Neuroplasticity: A New Therapeutic Target for Parkinson’s Disease? . . . . . . . . . . . . . . . . . . . . . . . . . Giselle M. Petzinger, Beth E. Fisher, Garnik Akopian, Ruth Wood, John P. Walsh, and Michael W. Jakowec 20 The MPTP-Treated Primate, with Specific Reference to the Use of the Common Marmoset (Callithrix jacchus) . . . . . . . . . . . . . . . . . . . . . . . . . . . . Michael J. Jackson and Peter Jenner 21 Behavioral Assessment in the African Green Monkey After MPTP Administration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . D. Eugene Redmond Jr.

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Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 437

Volume II Preface to the Series . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . v Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . vii Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . xiii

Part I  Basal Ganglia   1 Behavioral Assessment of Genetic Mouse Models of Huntington’s Disease . . . . . . . Miriam A. Hickey and Marie-Françoise Chesselet   2 Excitotoxic Lesions of the Rodent Striatum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Máté D. Döbrössy, Fabian Büchele, and Guido Nikkhah   3 Combination Lesion Models of MSA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Daniela Kuzdas and Gregor K. Wenning   4 The Role of the Dorsal Striatum in Instrumental Conditioning . . . . . . . . . . . . . . . Mark A. Rossi and Henry H. Yin   5 3-Nitropropionic Acid and Other Metabolic Toxin Lesions of the Striatum . . . . . . Cesar V. Borlongan and Paul R. Sanberg   6 Functional Assessment of Subcortical Ischemia . . . . . . . . . . . . . . . . . . . . . . . . . . . Tracy D. Farr and Rebecca C. Trueman

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Part II Neo- and Allo-Cortical Systems   7 Functional Organization of Rat and Mouse Motor Cortex . . . . . . . . . . . . . . . . . . . G. Campbell Teskey and Bryan Kolb   8 Forebrain Circuits Controlling Whisker Movements . . . . . . . . . . . . . . . . . . . . . . . . Kevin D. Alloway and Jared B. Smith   9 An Approach to Understanding the Neural Circuitry of Saccade Control in the Cerebral Cortex Using Antidromic Identification in the Awake Behaving Macaque Monkey Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Kevin Johnston and Stefan Everling 10 Photothrombotic Infarction of Caudate Nucleus and Parietal Cortex . . . . . . . . . . . Toshihiko Kuroiwa and Richard F. Keep 11 Models of Rodent Cortical Traumatic Brain Injury . . . . . . . . . . . . . . . . . . . . . . . . Frances Corrigan, Jenna M. Ziebell, and Robert Vink 12 The Use of Commissurotomy in Studies of Interhemispheric Communication . . . . Ian Steele-Russell

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Part III Cerebellar and Brain Stem Systems 13 Genetic Models of Cerebellar Dysfunction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Robert Lalonde and Catherine Strazielle 14 Cerebellar Control of Fine Motor Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rachel M. Sherrard 15 Cerebellum and Classical Conditioning . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Richard F. Thompson 16 Assessments of Visual Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ma’ayan Semo, Carlos Gias, Anthony Vugler, and Peter John Coffey 17 The Role of the Pedunculopontine Tegmental Nucleus in Motor Disorders . . . . . . Nadine K. Gut and Philip Winn

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Part IV Spinal Cord Systems 18 Contusion Models of Spinal Cord Injury in Rats . . . . . . . . . . . . . . . . . . . . . . . . . . Kelly A. Dunham and Candace L. Floyd 19 Demyelination Models in the Spinal Cord . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Paul A. Felts, Damineh Morsali, Mona Sadeghian, Marija Sajic, and Kenneth J. Smith 20 Preparation of Spinal Cord Injured Tissue for Light and Electron Microscopy Including Preparation for Immunostaining . . . . . . . . . . . Margaret L. Bates, Raisa Puzis, and Mary Bartlett Bunge 21 Assessing Spinal Cord Injury . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gillian D. Muir and Erin J. Prosser-Loose 22 Precise Finger Movements in Monkeys . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Roger Lemon

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Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 435

Contributors Kevin D. Alloway  •  Center for Neural Engineering, Pennsylvania State University, State College, University Park, PA, USA Margaret L. Bates  •  The Miami Project to Cure Paralysis, University of Miami Miller School of Medicine, Miami, FL, USA Cesar V. Borlongan  •  Department of Neurosurgery and Brain Repair, Center of Excellence for Aging and Brain Repair, University of South Florida College of Medicine, Tampa, FL, USA Fabian Büchele  •  Laboratory of Molecular Neurosurgery, Department of Stereotactic Neurosurgery, University Freiburg - Medical Center, Freiburg, Germany Mary Bartlett Bunge  •  The Miami Project to Cure Paralysis, University of Miami Miller School of Medicine, Miami, FL, USA; Department of Cell Biology & Anatomy, University of Miami Miller School of Medicine, Miami, FL, USA; Department of Neurological Surgery, University of Miami Miller School of Medicine, Miami, FL, USA Marie-Françoise Chesselet  •  Department of Neurology, David Geffen School of Medicine, UCLA, Los Angeles, CA, USA Peter John Coffey  •  Department of Ocular Biology and Therapeutics, Institute of Ophthalmology, University College, London, UK Frances Corrigan  •  Centre for Neuroscience Research, School of Medical Sciences, University of Adelaide, Adelaide, SA, Australia Máté D. Döbrössy  •  Laboratory of Molecular Neurosurgery, Department of Stereotactic Neurosurgery, University Freiburg - Medical Center, Freiburg, Germany Kelly A. Dunham  •  Department of Physical Medicine & Rehabilitation, University of Alabama at Birmingham, Birmingham, AL, USA Stefan Everling  •  Department of Physiology & Pharmacology, Robarts Research Institute, University of Western Ontario, London, ON, Canada Tracy D. Farr  •  Department for Experimental Neurology, Center for Stroke Research Berlin (CSB), Department for Experimental Neurology, Charité University Medicine, Berlin, Germany Paul A. Felts  •  Centre for Anatomy and Human Identification, University of Dundee, Dundee, UK Candace L. Floyd  •  Department of Physical Medicine & Rehabilitation, University of Alabama at Birmingham, Birmingham, AL, USA Carlos Gias  •  Department of Ocular Biology and Therapeutics, Institute of Ophthalmology, University College, London, UK Nadine K. Gut  •  Strathclyde Institute of Pharmacy & Biomedical Sciences, University of Strathclyde, Glasgow, Scotland, UK Miriam A. Hickey  •  Department of Neurology, David Geffen School of Medicine, UCLA, Los Angeles, CA, USA

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Kevin Johnston  •  Centre for Neuroscience Studies, Queen’s University Kingston, Kingston, ON, Canada Richard F. Keep  •  Department of Neurosurgery, University of Michigan, Ann Arbor, MI, USA Bryan Kolb  •  Canadian Centre for Behavioural Neuroscience, University of Lethbridge, Lethbridge, AB, Canada Toshihiko Kuroiwa  •  Department of Neurosurgery, University of Michigan, Ann Arbor, MI, USA; Department of Clinical Laboratory, Namegata District General Hospital, Namegata, Ibaraki, Japan Daniela Kuzdas  •  Division of Clinical Neurobiology, Department of Neurology, Innsbruck Medical University, Innsbruck, Austria Robert Lalonde  •  Départment de Psychologie, Université de Rouen, Mont-Saint-Aignan, France Roger Lemon  •  Institute of Neurology, University College London, London, UK Damineh Morsali  •  Department of Neuroinflammation, Institute of Neurology, University College London, London, UK Gillian D. Muir  •  Department of Biomedical Sciences, Western College of Veterinary Medicine, University of Saskatchewan, Saskatoon, SK, Canada Guido Nikkhah  •  Laboratory of Molecular Neurosurgery, Department of Stereotactic Neurosurgery, University Freiburg - Medical Center, Freiburg, Germany Erin J. Prosser-Loose  •  Department of Biomedical Sciences, Western College of Veterinary Medicine, University of Saskatchewan, Saskatoon, SK, Canada Raisa Puzis  •  The Miami Project to Cure Paralysis, University of Miami Miller School of Medicine, Miami, FL, USA Mark A. Rossi  •  Centre for Cognitive Neuroscience, Duke University, Durham, NC, USA Ian Steele-Russell  •  Departments of Psychiatry and Behavioral Science, Neuroscience and Experimental Therapeutics, and Veterinary Integrative and Biomedical Science, Texas A&M University System Health Science Center and Texas A&M University, TX, USA Mona Sadeghian  •  Department of Neuroinflammation, Institute of Neurology, University College London, London, UK Marija Sajic  •  Department of Neuroinflammation, Institute of Neurology, University College London, London, UK Paul R. Sanberg  •  Department of Neurosurgery and Brain Repair, Center of Excellence for Aging and Brain Repair, University of South Florida College of Medicine, Tampa, FL, USA Ma’ayan Semo  •  Department of Ocular Biology and Therapeutics, Institute of Ophthalmology, University College, London, UK Rachel M. Sherrard  •  UPMC-Univ Paris 6 and CNRS, UMR7102 Neurobiologie des Processus Adaptatifs, Paris, France Jared B. Smith  •  Center for Neural Engineering, Pennsylvania State University, State College, PA, USA Kenneth J. Smith  •  Institute of Neurology, University College London, London, UK Catherine Strazielle  •  INSERM U954 and Faculté de Médicine, Université Henri Poincaré, Vandœuvre-lès-Nancy, France

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G. Campbell Teskey  •  Department of Cell Biology and Anatomy, Hotchkiss Brain Institute, University of Calgary, Calgary, AB, Canada Richard F. Thompson  •  Program in Neuroscience, University of Southern California, Los Angeles, CA, USA Rebecca C. Trueman  •  The Brain Repair Group, School of Biosciences, Cardiff University, Cardiff, Wales, UK Robert Vink  •  Centre for Neuroscience Research, School of Medical Sciences, University of Adelaide, Adelaide, SA, Australia Anthony Vugler  •  Department of Ocular Biology and Therapeutics, Institute of Ophthalmology, University College, London, UK Gregor K. Wenning  •  Division of Clinical Neurobiology, Department of Neurology, Innsbruck Medical University, Innsbruck, Austria Philip Winn  •  Strathclyde Institute of Pharmacy & Biomedical Sciences, University of Strathclyde, Glasgow, Scotland, UK Henry H. Yin  •  Centre for Cognitive Neuroscience, Duke University, Durham, NC, USA Jenna M. Ziebell  •  Centre for Neuroscience Research, School of Medical Sciences, University of Adelaide, Adelaide, SA, Australia

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Part I Basal Ganglia

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Chapter 1 Behavioral Assessment of Genetic Mouse Models of Huntington’s Disease Miriam A. Hickey and Marie-Françoise Chesselet Abstract Huntington’s disease (HD) is a monogenetic, neurodegenerative disease. It is fatal, and although treatments are available for minor symptomatic relief, it remains incurable. Careful study of models of HD remains critical for elucidation of disease mechanisms and for the development of therapeutics. Models that rapidly develop to end stage are useful to assess efficacy of therapeutics for neuroprotection in relatively short experiments. However, the striatum, an area of brain traditionally associated with degeneration in HD already shows extensive atrophy by the time the symptoms manifest in patients, indicating that substantial degeneration occurs in the years preceding clinical onset. Thus, it is vital to also study models that allow analysis of the early pre-manifest disease stage. This requires the development of sensitive output measures to reveal deficits before the onset of obvious anomalies. Here, we briefly review the characteristics of ­several mouse models of HD and outline methods for analysis of behavioral deficits in both severe fast-­ progressing models and for early stages of disease in slowly progressive models. Key words: Knock-in, Transgenic, Viral vector, Behavior, Aggregate, Neuropathology, Therapeutic, Motor, Cognitive, Affective

1. Introduction Huntington’s disease (HD) is an autosomal dominant, genetic disorder that causes progressive neurodegeneration in the central nervous system (CNS). The disease, which affects up to 1 in 10,000 people worldwide, is caused by expansion of a trinucleotide repeat encoding glutamine (CAG) within exon 1 of the HD gene (1), leading to the expression of mutant huntingtin protein. The striatum undergoes extensive degeneration in HD, from an early stage in disease progression (2) although many other areas also degenerate. Traditionally, the triad of symptoms that are associated with

Emma L. Lane and Stephen B. Dunnett (eds.), Animal Models of Movement Disorders: Volume II, Neuromethods, vol. 62, DOI 10.1007/978-1-61779-301-1_1, © Springer Science+Business Media, LLC 2011

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HD include progressive decline in motor control and cognitive function, and psychiatric disturbances (3). Motor signs include chorea, dystonia, dysarthria, and dysphagia. Over time, the chorea gives way to bradykinesia (slowed movement) and rigidity (4). Cognitive dysfunction includes poor planning and impaired visuospatial skills (5–7). Modified rating scales have been developed to chart psychiatric symptoms, with much success (8). Apathy and irritability appear to be sensitive to disease progression, while depression is highly prevalent (8–10) and suicide is frequent (11). Once diagnosed, the disease progresses relentlessly over a period of 15–20 years. Although some symptoms may be improved with pharmacological treatments there are currently no effective neuroprotective or disease-modifying agents. Recent data show that many deficits and pathological changes are present in gene carriers that do not yet display sufficient symptoms for diagnosis of disease onset. These signs include changes in white matter and deficits in visual working memory, in odor recognition and identification, variability in tongue protrusion force and in timing of finger taps, in addition to increased incidence of apathy and irritability (2, 10, 12–16). Indeed several of these endpoints may be used for clinical trials over as little as 1 year (16). Other neuropathological changes also precede the onset of typical motor features and are present in gene carriers many years prior to the predicted age of onset. These changes include intranuclear inclusions (aggregates of insoluble mutant huntingtin), reduced striatal and whole brain volume, cortical thinning, and white matter loss, as well as hypothalamic changes (2,  12, 17–20). Intriguingly, cortical thinning has been shown to correlate with Total Functional Capacity, a measurement used in clinical trials for monitoring disease progression (20). In addition, the areas of cortical thinning, but not striatal volume, were different in patients with differing clinical phenotypes (20). The classical “selectivity” of neuronal degeneration in HD is relative rather than absolute, as outlined above. This has implications for the behavioral and pathological analysis of models of this disorder. Importantly, HD patients share symptoms with other neurodegenerative diseases, including Parkinson’s disease (PD) and Alzheimer’s disease (AD). Dystonia (co-contraction of antagonistic muscles) can complicate both PD and HD (21, 22); akinesia, a cardinal sign of PD, also occurs in patients with HD (23); as mentioned, cognitive disruption is present and may even be a presenting symptom in HD patients, whereas it dominates the symptoms in AD and can be present in PD (10, 16, 24, 25); both PD and HD patients show high prevalence of affective disorders and depression (9, 26); finally sleep disturbances are present in HD, AD, and PD (27). Thus, despite the presence of different pathologies, one should not be surprised to find similar behavioral deficits in models of these disorders, and indeed behavioral

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tests used for models of PD and AD can be utilized in the investigation of models of HD. An important consideration is that in patients with HD, clinical symptoms do not manifest until approximately 1⁄3 to 1⁄2 of striatum has degenerated (28). Thus, profound degeneration is necessary to cause clinical signs traditionally associated with the disease. However, in most models the neuronal cell loss that “defines” the disease in patients occurs late or is even absent in the lifetime of the animal. Nevertheless, profound behavioral anomalies can be detected with sensitive behavioral tests, indicating that the mutation induces neuronal dysfunction before causing cell death, mirroring observations in patients (29). An important caveat is that tests most commonly used to assess motor skills such as open field and rotarod are not always sensitive enough to detect the earliest deficits in models of basal ganglia dysfunction. Indeed, these tests are primarily sensitive to cerebellar or motor neuron deficits. In this chapter, we will present a brief overview of the most often studied mouse models of HD and will review the tests we found most useful in our own experience to assess early motor deficits in genetic mouse models of HD, with an emphasis on considerations to keep in mind for the choice of test and its interpretation.

2. Mouse Models of HD Prior to the discovery of genetic mutations, the only models available for HD were based on the use of neurotoxins that kill more or less selectively the neurons that are lost in patients. Toxin-based rodent models for HD were primarily based on local injections of quinolinic acid into the striatum or peripheral 3-nitroproprionic acid (3-NP; reviewed by (30)). However, since HD is caused by a single genetic mutation with high penetrance, expression of the mutated gene has high construct validity and there is little justification for the use of toxin models at this time. While no animal model is perfect, several genetic models of HD exist that replicate many features of the human disease. Genetic models of HD in rodents fall into two categories: mice or rats expressing a fragment of huntingtin as a transgene,1 or by viral vector-mediated gene transfer; and those expressing full length huntingtin, as a transgene, or as a knock-in2 mutation. In particular, mouse models based on expression of the full-length mutant

 Transgene: exogenous fragments of DNA. Generally inserted into the host DNA. May be introduced via a viral vector. If present in the germline, can be used to generate a transgenic line. 2  Knock-in: targeted insertion of cDNA into specific site in host organism’s genome. 1

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HD gene provide a long timeframe in which to examine the earliest behavioral and pathological manifestations of the mutation. Transgenes or viral vectors most often express the mutated protein in the context of a normal load of wildtype endogenous huntingtin, although a few models have been generated on a null background. In addition, the expression levels and site of expression can vary depending on the promoter used for the transgene, or the site of injection of the viral vector. Virus vector-mediated gene transfer has also facilitated the development of genetic models of HD in non-human primates (31, 32). The rapid progression of the neurodegenerative phenotype in these models is probably due to a high level of expression of mutant htt, as a result of a combination of strong expression from a heterologous promoter and transduction of infected neurons with multiple copies of the virus vector. In another rAAV-based model of HD in rats, quantitative RT-PCR revealed mutant htt expression to be over 100-fold that of the endogenous rat htt mRNA (33). Thus, cumulative expression of expanded polyQ proteins throughout the lifetime of experimental animals is not required to induce neuronal cell death; rather, acute overexpression of polyQ is toxic to adult neurons in vivo. There are several points to consider when comparing these models. Shorter fragments of huntingtin are more toxic than longer fragments (see for review (34)). Wildtype huntingtin is protective and important for development (35), and transgenic mice result in higher levels of expression of total (wildtype and mutated) huntingtin. Many kinds of peripheral cells become dysfunctional in HD (36, 37). Therefore viral vector-based models may not reflect the entire disease accurately. In contrast to transgenic models, knock-in models carry a mutation within their own endogenous huntingtin homologue (Hdh) gene. Thus they express a full length protein, theoretically in the proper genomic and protein context. However, the extent of human sequences inserted with the mutation determines the protein context surrounding the CAG repeat expansion and whether or not part of the human promoter is inserted with the mutation may influence the regional and temporal level of expression of the mutated protein. Typically, full-length models result in a more slowly progressing phenotype, which may aid in delineating the important early steps in disease progression. Despite radical differences in the timelines of disease progression between models, significant convergence in phenotypes and pathology is observed, as detailed below. It is important to consider animal size as a factor for eventual analysis of neurosurgical interventions for HD, or for analysis of pharmacokinetics and pharmacodynamics, either within the brain or periphery. Transgenic rats are more amenable than mice in a variety of behavioral tests and imaging studies, and successful rat models of HD have been extensively characterized (38, 39). In addition, several types of large animal models for HD now exist,

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including sheep (40), pigs (41), and non-human primates (32). However, the cost and facilities required will likely be limiting factors in their use. In addition, the paucity of behavioral research on sheep and pigs may impede therapeutics testing; however, steps are being made to address this issue (42). The vast majority of HD research utilizes mice, and here, we focus on behavioral testing of mice. 2.1. R6/2 HD Transgenic Mice

The R6 lines were the first successful transgenic mouse models of HD to be generated (43) and the R6/2 line remains the best characterized and most widely used of the fragment models. Indeed, this model develops several features observed in full-length htt models, albeit within a much faster timeframe (44–46). R6/2 mice express exon 1 of the human HD gene with approximately 150 CAG repeats (43). It is important to test for repeat length in each colony and in the animals actually used for the experiment because genetic drift occurs in these mice. This is particularly important in light of recent studies showing amelioration of disease in mice with “hyper-expanded” repeats (>300) (47, 48). Gender is also important to consider as male R6/2 mice tend to develop disease over a shorter time course than females (49). The original R6/2 mice have a rapid disease course. By approximately 10 weeks, their body weight plateaus, and thereafter declines, and the mice die prematurely at 13–16 weeks of age (43, 50). However, life span varies with source colony and husbandry techniques (51). These mice and other models expressing a fragment of the mutant protein can show glucose intolerance and diabetes, and this is also an important symptom to monitor (52). Furthermore, at late stages of disease, temperature regulation becomes dysfunctional, resulting in low body temperatures (53). R6/2 mice develop progressive motor deficits. By 6–8 weeks of age (depending on the colony) R6/2 mice begin to show overt deficits in gait, tremor, and poor coat condition (44). Cognitive deficits in aspects of visuospatial and procedural memory in R6/2 mice reminiscent of human HD are apparent in tests using the Morris water maze, T-maze, or Pavlovian fear conditioning, although age and disease progression can impact strategies used to complete cognitive tasks (54–57). Emotional responses are also dysfunctional at early (44) and late stages (58). Thus, these mice represent a severe and fast progressing model of HD, which can be used for in vivo high throughput behavioral testing at ages prior to overt phenotype development and severe pathology (44), although the timeframe to overt phenotype development is extremely short. The aggressive phenotype in R6/2 mice facilitates behaviorand, survival-based assays of therapeutic strategies in a relatively short time frame (59). However, institution-specific veterinary rules may dictate that the animal must be euthanized when a specific amount of weight is lost, and often prohibits the use of survival as an endpoint measure for neuroprotective studies (60).

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A limitation of the model is that early disease-related deficits may not be experimentally accessible in this model. It is also important to consider that R6/2 mice often exhibit phenotypes atypical of adult-onset HD, including epilepsy, cardiac dysfunction, and neuromuscular junction abnormalities (43, 61, 62). Nevertheless, this model clearly demonstrates in  vivo toxicity of exon 1 of mutant huntingtin and offers a wealth of behavioral, molecular, cellular and pathological endpoints for analysis. R6/1 mice, a sister line generated initially with a smaller repeat length, provide a longer timeframe to disease onset (43, 63). There may be some variability in onset of R6/1 phenotype, in different colonies, however a congenic line on the C57Bl/6 background has accelerated onset of behavioral dysfunction (63–66). Indeed with increasing C57Bl/6 background, we and others have also observed acceleration of phenotype and pathology in HD knock-in models (see below) (45, 67). In particular, rotarod impairments (see below) and learning deficits in the Morris water maze appear by 2m of age in the B6 R6/1 line, indicating substantial early dysfunction. 2.2. Full-Length Models of HD

Full-length models of HD include transgenic and knock in models. Full length Htt transgenic models include expression via a yeast artificial chromosome (68, 69), and more recently, via a bacterial artificial chromosome (70). Several lines of YAC HD transgenic mice have been generated that express a full-length human genomic mutant HD gene. YAC HD72 mice, with 72 CAG repeats, exhibit slower disease progression resulting from the lower levels of expression and perhaps the shorter repeat length. YAC128 mice express full-length HD containing 128 CAG repeats under the control of the native promoter (69). YAC128 mice have a more pronounced behavioral phenotype than YAC HD72 mice and exhibit a biphasic motor profile, with open field hyperactivity at 3 months followed by hypoactivity at 12 months. YAC128 mice have a normal gait but are impaired on the rotarod task by 6 months of age (69). Cognitive impairment precedes and then parallels progressive motor impairment (71). Both YAC lines show nuclear localization of htt fragments, which coincides with motor dysfunction and occurs first and most abundantly in the striatum, the region most vulnerable in HD (72). In addition, YAC128 mice show ~18% neuronal cell loss specifically in the striatum by 12 months of age (69). Carrying a stable poly CAA-CAG (n = 97) repeat to decrease somatic instability, the BAC HD mice develop progressive open field and rotarod deficits (49, 70). They show cortical and striatal atrophy and “dark neurons” without neuronal loss. Huntingtin aggregates and even diffuse huntingtin nuclear staining occur only in old age. These mice show consistent levels of both full length and N-terminal cleaved huntingtin between the ages of 2 and 12 months, while behavioral deficits and neurodegenerative changes progress.

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Both YAC and BAC transgenic full-length models show a gain in body weight over time (49), in contrast to the hallmark loss in body weight observed in HD patients despite increased caloric intake (73). Interestingly, overexpression of both wildtype and mutant full length huntingtin is associated with weight gain (74), suggesting that it is the overall greater level of the protein in these models that causes the weight gain. Theoretically, knock-in (KI) models should provide an optimal model of human HD as these mice carry the mutation in its appropriate genomic and protein context; however, this depends on the specific construct used (75). A number of KI mouse models have been generated to date. Of the various KI models, CAG140 (45, 76) and Hdh150 (46, 77, 78) are the best characterized. The Hdh150 mice harbor an expanded CAG repeat inserted into mouse Hdh, while CAG140 mice express a chimeric HD/Hdh protein, with human mutant exon 1 and part of the human promoter. Thus in contrast to the Hdh150, in these mice the protein context of the mutation includes both human and mouse sequences and the level and pattern of expression of the mutant Htt may be modified by the presence of human sequences in the promoter region. All KI mice eventually develop some evidence of neuronal degeneration, in the form of dark neurons in HdhQ111 mice and axonal degeneration in HdhQ72/80 mice in striatal projection areas. However, only CAG140 (45) and Hdh150 (77) mice show frank striatal ­neuronal loss, at approximately 2 years of age. Optical density for DARPP-32 is reduced by 1 year of age in CAG140 mice. In contrast to the YAC and BAC mice, both Hdh150 and CAG140 lines lose weight (45, 46, 77) at late disease stages, again a feature of the human disease. Most KI mice show overt, spontaneous gait deficits by 2 years of age (45, 77, 79). This late stage may recapitulate the overt phenotype that develops in R6/2 mice at approximately 8 weeks of age. Old Hdh150 mice show transcriptome anomalies similar to those of late stage R6/2 mice (46). Thus, these latter models (CAG150, Q111) require 2 years to progress to a point that is reached in 12 weeks in the R6/2 model, demonstrating the long timeframe that can be utilized for therapeutic and disease mechanism testing. In contrast, the CAG 140 mice have a much faster disease progression than these other KI mice; however, it is still much more protracted than in the R6/2 transgenics. We have recently found transcriptional deficits in CAG140 homozygous KI mice by 4m of age, many months prior to onset of overt phenotype (80), in agreement with the presence of behavioral and pathological deficits in these mice as early as 1 month of age (45, 76). Even on an identical genetic background, CAG140 mice develop aggregates much sooner than Hdh150 mice (81). Diffuse staining for aggregated huntingtin in striatal neurons is present from approximately 2m of age in CAG140 mice, with inclusions present from 4m (76, 81). In contrast, a diffuse nuclear localization is only

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observed at 7m in Hdh150 mice, and inclusions are evident by approximately 10m (77). Diffuse staining is observed in 6-weekold homozygous HdhQ111 knock-in mice, microaggregates by 5m and inclusions by 10m (79).

3. Methodological Considerations When Using Mice as Models of Neurodegenerative Diseases

As shown above, and in many reports that examine the effect of strain of mouse on behavioral tests, strain is extremely important to control, especially when lines are maintained on a mixed background (82–84). Non-sibling, distantly related pairs should be used for breeding for these colonies and this requires careful documentation of family lineages. The advent of congenic3 strains does not alleviate the effect of strain, since regular “refreshing” is necessary to prevent genetic drift. When mice are maintained on a hybrid background, careful consideration of generation number is also required when comparing results. Gender is an obvious factor to take into account when conducting any analyses, and in particular in behavioral research (45, 84–88). Although clear gender differences have not been documented clinically in HD patients, it is important to consider that males have been demonstrated to show more accelerated disease progression in R6/2 mice (49). Environmental enrichment for laboratory animals is now often required by animal research oversight committees, and this has profound effects on phenotype expression and disease course and can complicate comparison of results across laboratories (49). Indeed, prolonged environmental enrichment is beneficial in HD mouse models (89, 90) and the type of enrichment provided to mice should be detailed in methods sections. In addition, several laboratories now provide additional husbandry to advanced stage models of HD, including lowered water spouts and moistened chow (51, 90). These accommodations extend survival and should also be outlined in the methods. Additionally, prior to any behavioral testing, consideration should be given to whether the test subjects should be handled. We frequently do not handle our mice prior to testing the open field. However, mice are handled subsequently, to reduce stress in more involved tests of motor function, for example, rotarod (see below). Another important consideration is that, as in humans, the disease in these animals is progressive. Many models show hyperactivity followed by hypoactivity, and cellular

 Congenic: To generate a congenic strain mice from two genetically different strains are bred together. Resulting progeny are then bred again to the desired strain, up to 10 times (usually). Speed congenics may reduce the time taken, and involves analysis of the progeny for genetic markers of the desired strain, thus allowing only progeny with the most genetic markers of the desired strain to move forward to breeding.

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deficits change with age in R6/2 mice (29, 69, 76). Therefore, one needs to test at different times, carefully match groups for ages and take this possibility into account during statistical analysis of longitudinal data. Consideration of the effect of age on performance of wildtype mice is critical, since the window of genotype deficit may be reduced, due to declining performance of wildtype animals (45, 91). Finally, mice are nocturnal animals and the diurnal phase during which testing is conducted can profoundly influence the results. Testing in the dark phase, when mice are more active may provide more sensitivity, due to the higher normal level of activity (44, 45, 76). For these tests, it is useful to have access to a reversed phase animal room. Otherwise, one may choose tests that can be performed during the light phase. It is also important to schedule these tests after the animals have habituated to the dark or light, i.e., outside of the first/last hour of darkness or light.

4. Tests for Spontaneous Phenotype in Genetic Mouse Models of HD

5. Tests for Analysis of Early Motor Deficits in Mouse Models of HD

Most mouse models of HD do not develop obvious behavioral anomalies in their home cage until late in the disease. This obvious behavior is sometimes referred to as “symptomatic” or perhaps more accurately as “manifest”. When present, spontaneous home cage behavioral anomalies can be scored on a non-parametric scale for the presence or absence of tremor, and presence or absence of an unsteady or uncoordinated gait, in addition to turning ability and the width of the hindlimb base, the extent of piloerection (indicative of poor coat maintenance) and clasping. Interestingly, the CAG140 KI mice show these signs when approximately 40% of neurons and volume are lost (45), akin to the 1⁄3 to 1⁄2 loss observed in patients when obvious signs manifest (28).

As discussed earlier, one distinct advantage of mouse models of HD is the ability to analyze early stages of the disease, that are equivalent to the long pre-manifest phase of the disease that is now recognized in patients (16). This requires the use of specialized tests that challenge the motor system. Remarkably, many of the same tests used in R6/2 mice are able to detect deficits in the more progressive CAG140 mouse model at a very early age (44, 45). We will describe these tests starting with the most commonly used and then those we have found to offer superior sensitivity and/or ease of use.

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5.1. Open Field Test

One of the most well-known and well-used behavioral tests, the open field test can be conducted in the dark or light phase; however, it is important to maintain consistency within groups, and to disclose the period of the light phase used for testing. Behavior in the open field can be analyzed with manual methods, using video recording of activity for later analysis of rearing and locomotion, in addition to grooming or other behaviors, offering a low-cost alternative. When the required equipment is available, however, open field behavior can be analyzed automatically usually by recording infra-red beam breakages to identify the position of the mouse in X, Y and Z axes. With careful calibration to ensure accurate quantification of rearing and locomotion, automation can contribute information on speed, area occupied, number and length of individual movements, etc., more easily than video analysis. As stated above, hyperactivity is noted in several models at early stages of disease and is followed by hypoactivity (69, 76, 92, 93), thus this test can be used over time, or at specific ages. An issue associated with the open field is that it relies upon spontaneous locomotion. Factor analysis by various authors have found that exploration (94) and anxiety (95) can account for a large amount of variability in rodent open field activity. Thus, it is advantageous to use other tests in parallel.

5.2. Rotarod

Rotarod analysis can be performed either in the light or dark phase. It is easy to use, relatively automated, and relatively reproducible as long as the conditions of use are clearly defined. This allows for a comparison of mouse models between different laboratories. Paradigms include fixed speeds, or accelerating speeds, incorporating smooth or grooved axles, all of which greatly affect sensitivity (96). The standard apparatus is purchased from Ugo Basile, Varese, Italy, and it is used to measure the latency of mice to fall from the rotating rod. Usually, a few mice “cling” to the axle, and to prevent bias these are removed after three consecutive rotations and the latency at that time is used as the score for that animal. Sometimes the axle of the rotarod is covered with smooth rubber (“smooth axle”) to reduce clinging and to make the task more difficult than on the grooved axle. Duration and frequency of trials as well as speed can differ and it is important to document all parameters used in the published reports. Unfortunately, this test is particularly sensitive to motor neuron or cerebellar deficits, which are typically absent in HD; accordingly, HD mice tend to only show anomalies on the rotarod at advanced stages of the disease. In our experiments, mice were trained to run on the smooth axle that is accelerated from 4 to 40 rpm over 10 min (3 trials per day for 4 days) (44, 45, see also 90). Even with this protocol, however, deficits in rotarod ­performance of R6/2 mice were not detected until after initial climbing, and open field-rearing deficits and deficits in CAG140 KI mice are very subtle (44, 45). Many other models have now shown deficits in rotarod performance, including KI mice (97) and

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the BAC transgenic lines (49, 70), although there is some ­variability in rotarod impairments in YAC128 transgenic mice (49, 71). 5.3. Climbing

Climbing behavior can be measured without expensive equipment, either during the light or the dark phases of the day, and shows very early alterations in mouse models of HD. We found deficits as early as 4.5 weeks in R6/2 mice (44) and 1.5 months in the CAG140 KI mice (45). Mice are placed at the bottom of wire cylinder cages (diameter, 10.5 cm, height, 15.5 cm) and spontaneous activity is videotaped for 5 min to quantify the number of climbs (defined as all four paws on the side of the cage). The same tapes can be used to quantify the number of rears made by the mouse to monitor general activity. This test has now been used in several other transgenic mice and reveals deficits in BAC and YAC 128 transgenic mice (98, 99). A caveat is that when tested during the dark phase, gender effects are noted, and group sizes must be designed accordingly (Fig. 1).

5.4. Wheel Running

Running wheels are widely used in research on circadian rhythms and are fully automated. They represent a significant initial investment but are valuable for preclinical drug testing because of the high power to detect differences between groups, the minimal investigator time involvement, and the wealth of information obtained. We have successfully developed the use of running wheels

Fig. 1. Climbing activity in C57Bl/6J mice at 6.5 weeks of age. Mice were tested either during dark phase, or light phase (not both). Mice climb more during their active phase (dark phase). However, gender differences are observed in climbing during the dark phase, which would necessitate large mixed gender group sizes, in order to observe genotype differences, or treatment effects. Larger mixed gender group sizes (or single gender groups) are not necessary if mice are tested during the day phase. Data are shown as mean ± SEM. Data were analyzed using a completely randomized ANOVA, followed by Fisher’s PSD post hoc tests. Effect of phase F(1,30) = 16.5, p