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Methods for neural ensemble recordings 2nd ed Edition Miguel A. L. Nicolelis

Methods for neural ensemble recordings 2nd ed Edition Miguel A. L. Nicolelis Methods for neural ensemble recordings 2nd ed Edition Miguel A. L. Nicolelis Methods for neural ensemble recordings 2nd ed Edition Miguel A. L. Nicolelis

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Methods for neural ensemble recordings 2nd ed Edition Miguel A. L. Nicolelis Digital Instant Download Author(s): Miguel A. L. Nicolelis ISBN(s): 9780849370465, 0849370469 Edition: 2nd ed File Details: PDF, 14.98 MB Year: 2008 Language: english
METHODS for NEURAL ENSEMBLE RECORDINGS SECOND EDITION © 2008 by Taylor & Francis Group, LLC
FRONTIERS IN NEUROSCIENCE Series Editors Sidney A. Simon, Ph.D. Miguel A.L. Nicolelis, M.D., Ph.D. Published Titles Apoptosis in Neurobiology Yusuf A. Hannun, M.D., Professor of Biomedical Research and Chairman/Department of Biochemistry and Molecular Biology, Medical University of South Carolina, Charleston, South Carolina Rose-Mary Boustany, M.D., tenured Associate Professor of Pediatrics and Neurobiology, Duke University Medical Center, Durham, North Carolina Methods for Neural Ensemble Recordings Miguel A.L. Nicolelis, M.D., Ph.D., Professor of Neurobiology and Biomedical Engineering, Duke University Medical Center, Durham, North Carolina Methods of Behavioral Analysis in Neuroscience Jerry J. Buccafusco, Ph.D., Alzheimer’s Research Center, Professor of Pharmacology and Toxicology, Professor of Psychiatry and Health Behavior, Medical College of Georgia, Augusta, Georgia Neural Prostheses for Restoration of Sensory and Motor Function John K. Chapin, Ph.D., Professor of Physiology and Pharmacology, State University of New York Health Science Center, Brooklyn, New York Karen A. Moxon, Ph.D., Assistant Professor/School of Biomedical Engineering, Science, and Health Systems, Drexel University, Philadelphia, Pennsylvania Computational Neuroscience: Realistic Modeling for Experimentalists Eric DeSchutter, M.D., Ph.D., Professor/Department of Medicine, University of Antwerp, Antwerp, Belgium Methods in Pain Research Lawrence Kruger, Ph.D., Professor of Neurobiology (Emeritus), UCLA School of Medicine and Brain Research Institute, Los Angeles, California Motor Neurobiology of the Spinal Cord Timothy C. Cope, Ph.D., Professor of Physiology, Wright State University, Dayton, Ohio Nicotinic Receptors in the Nervous System Edward D. Levin, Ph.D., Associate Professor/Department of Psychiatry and Pharmacology and Molecular Cancer Biology and Department of Psychiatry and Behavioral Sciences, Duke University School of Medicine, Durham, North Carolina Methods in Genomic Neuroscience Helmin R. Chin, Ph.D., Genetics Research Branch, NIMH, NIH, Bethesda, Maryland Steven O. Moldin, Ph.D, University of Southern California, Washington, D.C. © 2008 by Taylor & Francis Group, LLC
Methods in Chemosensory Research Sidney A. Simon, Ph.D., Professor of Neurobiology, Biomedical Engineering, and Anesthesiology, Duke University, Durham, North Carolina Miguel A.L. Nicolelis, M.D., Ph.D., Professor of Neurobiology and Biomedical Engineering, Duke University, Durham, North Carolina The Somatosensory System: Deciphering the Brain’s Own Body Image Randall J. Nelson, Ph.D., Professor of Anatomy and Neurobiology, University of Tennessee Health Sciences Center, Memphis, Tennessee The Superior Colliculus: New Approaches for Studying Sensorimotor Integration William C. Hall, Ph.D., Department of Neuroscience, Duke University, Durham, North Carolina Adonis Moschovakis, Ph.D., Department of Basic Sciences, University of Crete, Heraklion, Greece New Concepts in Cerebral Ischemia Rick C.S. Lin, Ph.D., Professor of Anatomy, University of Mississippi Medical Center, Jackson, Mississippi DNA Arrays: Technologies and Experimental Strategies Elena Grigorenko, Ph.D., Technology Development Group, Millennium Pharmaceuticals, Cambridge, Massachusetts Methods for Alcohol-Related Neuroscience Research Yuan Liu, Ph.D., National Institute of Neurological Disorders and Stroke, National Institutes of Health, Bethesda, Maryland David M. Lovinger, Ph.D., Laboratory of Integrative Neuroscience, NIAAA, Nashville, Tennessee In Vivo Optical Imaging of Brain Function Ron Frostig, Ph.D., Associate Professor/Department of Psychobiology, University of California, Irvine, California Primate Audition: Behavior and Neurobiology Asif A. Ghazanfar, Ph.D., Princeton University, Princeton, New Jersey Methods in Drug Abuse Research: Cellular and Circuit Level Analyses Dr. Barry D. Waterhouse, Ph.D., MCP-Hahnemann University, Philadelphia, Pennsylvania Functional and Neural Mechanisms of Interval Timing Warren H. Meck, Ph.D., Professor of Psychology, Duke University, Durham, North Carolina Biomedical Imaging in Experimental Neuroscience Nick Van Bruggen, Ph.D., Department of Neuroscience Genentech, Inc. Timothy P.L. Roberts, Ph.D., Associate Professor, University of Toronto, Canada The Primate Visual System John H. Kaas, Department of Psychology, Vanderbilt University Christine Collins, Department of Psychology, Vanderbilt University, Nashville, Tennessee Neurosteroid Effects in the Central Nervous System Sheryl S. Smith, Ph.D., Department of Physiology, SUNY Health Science Center, Brooklyn, New York © 2008 by Taylor & Francis Group, LLC
Modern Neurosurgery: Clinical Translation of Neuroscience Advances Dennis A. Turner, Department of Surgery, Division of Neurosurgery, Duke University Medical Center, Durham, North Carolina Sleep: Circuits and Functions Pierre-Hervé Luoou, Université Claude Bernard Lyon, France Methods in Insect Sensory Neuroscience Thomas A. Christensen, Arizona Research Laboratories, Division of Neurobiology, University of Arizona, Tuscon, Arizona Motor Cortex in Voluntary Movements Alexa Riehle, INCM-CNRS, Marseille, France Eilon Vaadia, The Hebrew University, Jerusalem, Israel Neural Plasticity in Adult Somatic Sensory-Motor Systems Ford F. Ebner, Vanderbilt University, Nashville, Tennessee Advances in Vagal Afferent Neurobiology Bradley J. Undem, Johns Hopkins Asthma Center, Baltimore, Maryland Daniel Weinreich, University of Maryland, Baltimore, Maryland The Dynamic Synapse: Molecular Methods in Ionotropic Receptor Biology Josef T. Kittler, University College, London, England Stephen J. Moss, University College, London, England Animal Models of Cognitive Impairment Edward D. Levin, Duke University Medical Center, Durham, North Carolina Jerry J. Buccafusco, Medical College of Georgia, Augusta, Georgia The Role of the Nucleus of the Solitary Tract in Gustatory Processing Robert M. Bradley, University of Michigan, Ann Arbor, Michigan Brain Aging: Models, Methods, and Mechanisms David R. Riddle, Wake Forest University, Winston Salem, North Carolina Neural Plasticity and Memory: From Genes to Brain Imaging Frederico Bermudez-Rattoni, National University of Mexico, Mexico City, Mexico Serotonin Receptors in Neurobiology Amitabha Chattopadhyay, Center for Cellular and Molecular Biology, Hyderabad, India Methods for Neural Ensemble Recordings, Second Edition Miguel A.L. Nicolelis, M.D., Ph.D., Professor of Neurobiology and Biomedical Engineering, Duke University Medical Center, Durham, North Carolina © 2008 by Taylor & Francis Group, LLC
CRC Press is an imprint of the Taylor & Francis Group, an informa business Boca Raton London New York Edited by Miguel A. L. Nicolelis Duke University Medical Center Durham, NC METHODS for NEURAL ENSEMBLE RECORDINGS SECOND EDITION © 2008 by Taylor & Francis Group, LLC
CRC Press Taylor & Francis Group 6000 Broken Sound Parkway NW, Suite 300 Boca Raton, FL 33487-2742 © 2008 by Taylor & Francis Group, LLC CRC Press is an imprint of Taylor & Francis Group, an Informa business No claim to original U.S. Government works Printed in the United States of America on acid-free paper 10 9 8 7 6 5 4 3 2 1 International Standard Book Number-13: 978-0-8493-7046-5 (Hardcover) This book contains information obtained from authentic and highly regarded sources. Reprinted material is quoted with permission, and sources are indicated. A wide variety of references are listed. Reasonable efforts have been made to publish reliable data and information, but the author and the publisher cannot assume responsibility for the validity of all materials or for the conse- quences of their use. No part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any information storage or retrieval system, without written permission from the publishers. For permission to photocopy or use material electronically from this work, please access www. copyright.com (http://www.copyright.com/) or contact the Copyright Clearance Center, Inc. (CCC) 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. CCC is a not-for-profit organization that provides licenses and registration for a variety of users. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation without intent to infringe. Library of Congress Cataloging-in-Publication Data Methods for neural ensemble recordings / [edited by] Miguel A.L. Nicolelis. -- 2nd ed. p. ; cm. -- (Frontiers in neuroscience) Includes bibliographical references and index. ISBN 978-0-8493-7046-5 (alk. paper) 1. Electroencephalography. 2. Microelectrodes. 3. Neurons. I. Nicolelis, Miguel A. L. II. Series: Frontiers in neuroscience (Boca Raton, Fla.) [DNLM: 1. Neurons--physiology. 2. Brain--physiology. 3. Electrophysiology--methods. 4. Microelectrodes. WL 102.5 M592 2008] QP376.5.M47 2008 616.8’047547--dc22 2007027468 Visit the Taylor & Francis Web site at http://www.taylorandfrancis.com and the CRC Press Web site at http://www.crcpress.com © 2008 by Taylor & Francis Group, LLC
To Pedro, Rafael, and Daniel, May your voyages be full of lore and fun © 2008 by Taylor & Francis Group, LLC
ix Contents Series Preface............................................................................................................xi Preface.................................................................................................................... xiii Editor .....................................................................................................................xvii Contributors ............................................................................................................xix Chapter 1 State-of-the-Art Microwire Array Design for Chronic Neural Recordings in Behaving Animals ........................................................1 Gary Lehew and Miguel A. L. Nicolelis Chapter 2 Surgical Techniques for Chronic Implantation of Microwire Arrays in Rodents and Primates......................................................... 21 Laura M. O. Oliveira and Dragan Dimitrov Chapter 3 Technology for Multielectrode MicroStimulation of Brain Tissue..................................................................................................47 Timothy Hanson, Nathan Fitzsimmons, and Joseph E. O’Doherty Chapter 4 Strategies for Neural Ensemble Data Analysis for Brain–Machine Interface (BMI) Applications.............................................................57 Miriam Zacksenhouse and Simona Nemets Chapter 5 Chronic Recordings in Transgenic Mice............................................83 Kafui Dzirasa Chapter 6 Multielectrode Recordings in the Somatosensory System.................97 Michael Wiest, Eric Thomson, and Jim Meloy Chapter 7 Chronic Recording During Learning...............................................125 Aaron J. Sandler Chapter 8 Defining Global Brain States Using Multielectrode Field Potential Recordings ........................................................................ 145 Shih-Chieh Lin and Damien Gervasoni © 2008 by Taylor & Francis Group, LLC
x Methods for Neural Ensemble Recordings, Second Edition Chapter 9 Multielectrode Recording in Behaving Monkeys ............................ 169 R. E. Crist and M. A. Lebedev Chapter 10 Neural Ensemble Recordings from Central Gustatory-Reward Pathways in Awake and Behaving Animals..................................... 189 Albino J. Oliveira-Maia, Sidney A. Simon, and Miguel A. L. Nicolelis Chapter 11 Building Brain–Machine Interfaces to Restore Neurological Functions .......................................................................................... 219 Mikhail A. Lebedev, Roy E. Crist, and Miguel A. L. Nicolelis Chapter 12 Conceptual and Technical Approaches to Human Neural Ensemble Recordings....................................................................... 241 Dennis A. Turner, Parag G. Patil, and Miguel A.L. Nicolelis © 2008 by Taylor & Francis Group, LLC
xi Series Preface Our goal in creating the Frontiers in Neuroscience Series is to present the insights of experts on emerging fields and theoretical concepts that are, or will be, in the van- guard of neuroscience. Books in the series cover genetics, ion channels, apoptosis, electrodes, neural ensemble recordings in behaving animals, and even robotics. The series also covers new and exciting multidisciplinary areas of brain research, such as computational neuroscience and neuroengineering, and describes breakthroughs in classical fields like behavioral neuroscience. We hope every neuroscientist will use these books in order to get acquainted with new ideas and frontiers in brain research. These books can be given to graduate students and postdoctoral fellows when they are looking for guidance to start a new line of research. Each book is edited by an expert and consists of chapters written by the leaders in a particular field. Books are richly illustrated and contain comprehensive bibliog- raphies. Chapters provide substantial background material relevant to the particu- lar subject. We hope that as the volumes become available, the effort put in by us, the publisher, the book editors, and individual authors will contribute to the further development of brain research. The extent to which we achieve this goal will be determined by the utility of these books. Sidney A. Simon, Ph.D. Miguel A.L. Nicolelis, M.D.,Ph.D. Series Editors © 2008 by Taylor & Francis Group, LLC
xiii Preface Almost 10 years ago, as the fresh-off-the-press volumes of the first edition of Meth- ods for Neural Ensemble Recordings reached the CRC booth just in time for the inaugural day of the Society for Neuroscience Meeting, there were already several laboratories around the world applying new approaches and technologies to chroni- cally record the simultaneous extracellular activity of small populations of single neurons in behaving animals. Yet, in those days a considerable number of neuro- physiologists were still not convinced that such a technique would bring significant benefits to the field. Those opinions were bluntly articulated in two peculiar encounters that I experi- enced during that time. In the first, one of the leading neurophysiologists of our time stopped by my SFN poster display to, according to him, kindly remind me that there was “no future and no career in neural ensemble recordings.” Instead, he suggested that I should simply give up this “foolish stuff” and come back to the “church of single unit recording.” Just a couple of years later, this grandfatherly and well-intended advice was pointedly reinforced by a NIH reviewer who, in response to a grant that John Chapin and I had submitted, wondered in despair “…why one needs to use space-age tech- nology to study the brain?…” What a difference 10 years make! Today, a whole generation of young graduate students and postdocs dive into the great adventure of systems neurophysiology by learning how to record from tens or even hundreds of single neurons simultaneously as soon as they enter a lab. Without a doubt, this young and innovative branch of neuroscience has been totally embraced by the next generation of neuroscientists. Those closely following the development of the field have also seen that, whereas a decade ago, most if not all studies employing chronic multi-electrode recording were performed only on rats, today the same basic technique is employed for studies in different species of non-human primates, such as owls, squirrels and Rhesus mon- keys, as well as mice (wild and transgenic strains). In the last few years, the method has found its way to neurosurgical suites where it is now used during intra-operative recordings in Parkinsonian patients whose symptoms can only be improved through a deep brain stimulator. If these dramatic changes were not enough, multiple technological breakthroughs in the fabrication of high-density microelectrode arrays, surgical implantation tech- niques, and multi-channel signal processing have significantly expanded the yield and longevity of chronic neural ensemble recordings in behaving animals. These advances have allowed the establishment of direct real-time brain–machine inter- faces, a new experimental paradigm that holds a lot of promise, not only in terms of basic neurophysiology research, but also as the potential core for the development of a new generation of neuroprosthetic devices aimed at restoring mobility and com- munication skills in severely disabled patients. © 2008 by Taylor & Francis Group, LLC
xiv Methods for Neural Ensemble Recordings, Second Edition Over the last 10 years, thanks to the generosity of some of the field’s pioneers who agreed to write a series of detailed methods-oriented chapters for the book, the first edition of Methods for Neural Ensemble Recordings has become a technical reference book for those who have taken the plunge into the field. As such, it has been a most rewarding experience for those of us who participated in the original volume to spot copies of the book, not nicely stored on book shelves, but worn and torn next to neurophysiological setups, spread all over the world. The second edition of this book comes to light during a very different time and environment in systems neuroscience. Partially liberated from having to maintain at any cost some of its classic dogmas, systems neurophysiology is once again thriving in both theory and practice. Furthermore, the ever growing consensus that distrib- uted populations of neurons, rather than single neurons, define the true functional unit of the central nervous system creates continuous demand for new technologies that can push multi-electrode recordings methods and neural ensemble physiology to new limits. Since 1994, when I joined the Department of Neurobiology at Duke, I have had the privilege to work and collaborate with a unique group of highly talented people who arrived at our laboratory with the goal of achieving just that: pushing the limits of our blossoming field and, during this process, shedding new light on the neuro- physiological principles that make ensembles of neurons perform the “business of the brain.” This book has been written by a group of these young collaborators, whose col- lective groundbreaking work covers most of the current areas of basic and clinical research that utilize multi-electrode recordings as the method of choice to probe brain circuits. The final product of this two year project, the second edition of Meth- ods in Neural Ensemble Physiology, is dedicated to all the high-school, undergradu- ate and graduate students, along with the postdoctoral fellows, technicians, research associates, collaborators, and visitors who, by joining our laboratory, made this jour- ney through distributed and dynamic brain circuits quite a thrill. I would like to thank Duke University, the Brain and Mind Institute at the Ecole Polytechnic Federale de Lausanne, the Edmond and Lily Safra International Insti- tute of Neuroscience of Natal, the National Institutes of Health, DARPA, and the Anne W. Deane Professorship Endowment for providing me with the time, space, support and resources for completing this and other projects that have considerably expanded the horizons of my life as a scientist in the last two years. I would like to sincerely thank our CRC editor, Barbara Norwitz, for waiting patiently for the completion of this project. I would also like to acknowledge the continuous support of my “older brother” and series co-editor, Sidney Simon, who invited me to edit the first edition of this book, 13 years ago, as my first assignment as a Duke assistant professor. We never stopped getting into trouble together after that. I also would like to thank my dear friend and collaborator, Susan Halkiotis, for dedicating 2 long years of her life to this project, and many more years to make sure that everything that crosses her desk, no matter how crazy or challenging the task, is taken care of with utmost care, perfection, and class. With fun on top! © 2008 by Taylor & Francis Group, LLC
Preface xv Finally, I would like to thank Laura Oliveira for thirty years of trusting and supporting the pursuit of my dreams, far from home, wherever they may take us, no matter the odds or personal costs. I can barely wait to see what the next ten years will bring! Miguel A. L. Nicolelis Natal, Brazil © 2008 by Taylor & Francis Group, LLC
xvii Editor Miguel A. L. Nicolelis, M.D., Ph.D., is the Anne W. Deane Professor of Neurosci- ence and professor in the departments of neurobiology, biomedical engineering, and psychology at Duke University, where he also serves as co-director of the Center for Neuroengineering. In addition, Dr. Nicolelis is scientific coordinator at the Edmond and Lily Safra International Institute of Neuroscience of Natal (ELS-IINN) in Natal, Brazil and taught neuroscience during 2006–2007 at École Polytechique Féderale de Lausanne in Lausanne, Switzerland. Dr. Nicolelis is a native of Sao Paulo, Brazil, where he received his M.D. and Ph.D. in neurophysiology from the University of Sao Paulo. He graduated from the University of Sao Paulo School of Medicine and was awarded the Oswaldo Cruz Prize for research, the highest honor awarded to a Brazilian medical student. After postdoctoral work at Hahnemann University, Dr. Nicolelis joined the faculty at Duke University in 1994. Dr. Nicolelis is interested in understanding the general computational principles underlying the dynamic interactions between populations of cortical and subcortical neurons involved in motor control and tactile perception. Although Dr. Nicolelis is best known for his study of brain–machine interfaces (BMI) for neuroprosthetics in human patients and nonhuman primates, he is also developing an integrative approach to studying neurological and psychiatric disorders by recording neuronal ensem- ble activity across different brain areas in genetically modified mice. Dr. Nicolelis believes that this approach will allow the integration of molecular, cellular, systems, and behavioral data in the same animal to produce a more complete understanding of the nature of the alterations associated with these disorders. Neuroscience laboratories in the U.S. and Europe have incorporated Dr. Nicole- lis’ experimental paradigm to study a variety of mammalian neuronal systems. His research has influenced basic and applied research in computer science, robotics, and biomedical engineering. This multidisciplinary approach to research has become widely recognized in the neuroscience community. Dr. Nicolelis was named as one of Scientific American’s Top 50 Technology Leaders in America in 2004 and has received a number of honors and awards includ- ing the Whitehead Scholar Award, DARPA Award for Sustained Excellence by a Performer; Ruth and A. Morris Williams, Jr., Faculty Research Prize; Whitehall Foundation Award; McDonnell-Pew Foundation Award; Duke University Thomas Langford Lectureship Award; the Ramon y Cajal Chair at the University of Mexico; and the Santiago Grisolia Chair at Catedra Santiago Grisolia. He has authored more than 120 manuscripts in scientific journals, and edited numerous books and special journal issues. He is frequently an invited speaker at scientific conferences and meet- ings throughout the world. © 2008 by Taylor & Francis Group, LLC
xix Contributors Dr. Roy Crist Department of Neurobiology Duke University Durham, North Carolina Dr. Dragan Dimitrov Monterey, California Dr. Kafui Dzirasa Department of Neurobiology Duke University Durham, North Carolina Mr. Nathan Fitzsimmons Department of Neurobiology Duke University Durham, North Carolina Dr. Damien Gervasoni Faculté de Médecine Laënnec University Claude Bernard Lyon, France Mr. Timothy Hanson Duke University Durham, North Carolina Dr. Mikhail Lebedev Department of Neurobiology Duke University Durham, North Carolina Mr. Gary Lehew Department of Neurobiology Duke University Durham, North Carolina Dr. Shih-Chieh Lin Department of Neurobiology Duke University Durham, North Carolina Mr. Jim Meloy Department of Neurobiology Duke University Durham, North Carolina Dr. Simona Nemets Faculty of Mechanical Engineering Technion–Israel Institute of Technology Haifa, Israel Dr. Miguel A. L. Nicolelis Department of Neurobiology Duke University Durham, North Carolina Mr. Joseph E. O’Doherty Department of Biomedical Engineering Duke University Durham, North Carolina Dr. Laura M. Oliveira Department of Neurobiology Duke University Durham, North Carolina Dr. Albino J. Oliveira-Maia Department of Neurobiology Duke University Durham, North Carolina Dr. Parag Patil Department of Neurosurgery University of Michigan Hospitals Ann Arbor, Michgan Dr. Aaron Sandler Department of Neurobiology Duke University Durham, North Carolina © 2008 by Taylor & Francis Group, LLC
xx Methods for Neural Ensemble Recordings, Second Edition Dr. Sidney A. Simon Department of Neurobiology Duke University Durham, North Carolina Dr. Eric Thomson Department of Neurobiology Duke University Durham, North Carolina Dr. Dennis Turner Division of Neurosurgery Duke Medical Center Durham, North Carolina Dr. Michael Wiest Department of Neurobiology Duke University Durham, North Carolina Dr. Miriam Zacksenhouse Faculty of Mechanical Engineering Technion–Israel Institute of Technology Haifa, Israel © 2008 by Taylor & Francis Group, LLC
1 1 State-of-the-Art Microwire Array Design for Chronic Neural Recordings in Behaving Animals Gary Lehew and Miguel A. L. Nicolelis INTRODUCTION Over the last two decades, many laboratories around the world have started to rely on microelectrode arrays formed by fine microwires, organized in different geometri- cal configurations, to chronically record the extracellular activity of populations of individual neurons in both anesthetized and behaving animals (Nicolelis et al. 1997, 2003; Lebedev et al. 2006; Verloop and Holsheimer 1984; Williams et al. 1999). As the field of chronic multielectrode recordings evolved, so did the designs of such microwire-based arrays. Indeed, during the last 13 years, our laboratory at the Duke University Cen- ter for Neuroengineering (DUCN) has specialized in producing a large variety of microwire array configurations that can now be utilized in a large variety of species (e.g., mice, rats, monkeys, and intraoperative human recordings). In particular, our efforts have been directed at producing arrays that can be utilized in experimental protocols demanding simultaneous recordings from large samples of single neurons CONTENTS Introduction................................................................................................................1 The DUCN Design and Fabrication Approach for Multisite, Chronic Neural Ensemble Recordings......................................................................................2 The Layered Approach ..............................................................................................4 Discretely Wired Approach .....................................................................................10 Building a Layered PCB Microwire Array..............................................................15 Building a Discretely Wired Array.......................................................................... 18 Conclusions..............................................................................................................20 References................................................................................................................20 © 2008 by Taylor & Francis Group, LLC
2 Methods for Neural Ensemble Recordings, Second Edition (e.g., 50 to 500), distributed across multiple cortical and subcortical brain sites in fully awake and behaving animals over long periods of time (months to years). The goal of this chapter, therefore, is to review the DUCN accumulated experience and describe its current state-of-the-art design and fabrication approach for producing high-quality microwire-based arrays for chronic, multisite, neural ensemble recordings. THE DUCN DESIGN AND FABRICATION APPROACH FOR MULTISITE, CHRONIC NEURAL ENSEMBLE RECORDINGS Over the years, the DUCN technical staff has abided by the principle that microelec- trode arrays for chronic single-neuron recordings have to be designed according to the main anatomical characteristics and contours of the specific brain area targeted in a particular experimental project. Thus, by collaborating with neurophysiologists specialized in working with different animal species, our technicians developed a large variety of microwire array (bundle) configurations for targeting different corti- cal and subcortical brain areas. This initiative proved to be essential for achieving optimal configurations that significantly increase the neuronal yield and longevity of our chronic recordings. In this design process, both 2-D and 3-D contour matching can be performed. In addition, different strategies have been implemented to fabri- cate adjustable microelectrodes. Such techniques allow microwire tips to be repeat- edly and selectively repositioned after surgery. Using this option, users can resample the neural area of interest once the physiological properties of a given populations of neurons have been accomplished. Among other advantages, such a data collection strategy increases considerably the overall neuronal sample obtained per recording site over the course of several weeks or months. In recent years, we have added stimu- lation electrodes, local ground reference electrodes, and cannula for drug injection to the basic configuration of such microwire arrays, increasing significantly the range of experimental manipulations that can be carried out once such multiple devices are chronically implanted in the animal’s brain. The basic configuration of the DUCN microelectrode arrays consist of insu- lated metallic conductors with an overall diameter in the range of 25 to 50 μm. The metallic conductor is selected based on corrosion resistance and degree of stiffness, because the wire is driven lengthwise. Tungsten or a stainless steel alloy is often used with an insulation coating of polyimide, Formvar, Isonel, or Teflon. The micro wires are processed to be as straight and stiff as possible. Using a variety of cutting devices, the tips of the wires can be cut blunt or at any desired angle (see Figure 1.1). The angle cut exposes more of the metallic conductor, which lowers the impedance of the electrode. This can reduce the ability to record from well-isolated single neurons. However, the sharply raked tip of the angle cut makes penetration through the brain tissues much easier and less traumatic, while producing less resistance as the length of the microwire travels through the tissue. At this level of design, multiple factors have to be balanced in order to produce an electrode that will yield viable long-term single unit recordings. For instance, not only must optimal impedance for single unit recordings be a consideration during electrode production, but care must be taken to produce an array that will penetrate © 2008 by Taylor & Francis Group, LLC
State-of-the-Art Microwire Array Design for Chronic Neural Recordings 3 tissue easily during the surgical insertion process. Conversely, relying solely on a fine tip microelectrode array (Schwartz 2004) to facilitate tissue penetration may increase the chance of encapsulation of the electrode tips by glia and extracellular protein deposits and thus diminish the ability to record neural signals beyond a few days or weeks. The lesson here is simple: Finding the right compromise among a large number of variables (electrode material, electrode tip shape, cut angle, etc.) is the single most relevant challenge facing developers of microelectrode arrays for chronic recordings. In the DUCN fabrication routine, electrode tips can be left bare, or they can be electroplated, either selectively or as a group, to reduce impedance. An electrode impedance-measuring device such as the A-M systems model 2900 is a useful tool to quantify the variables in electrode impedance, such as the effect of electroplating on impedance. The spacing or separation of the recording electrode tips are in the range of 200 to 1000 μm, center to center, depending on the brain area targeted, the animal species utilized, and the experimental protocol. As a design principle, our electrode arrays are fabricated as part of a complete investigative system. Moreover, a series of features are incorporated into the design of the array to improve the efficiency of the manufacturing, testing, and surgical implantation processes. For instance, reinforced breakaway tabs are used to provide a standard clamp point for electrode holders of the stereotaxic equipment used dur- ing surgery as well as assembly and testing operations. At the end of the implantation surgery, the tabs are easily removed. Another set of small breakaway tabs are used to hold alignment in assembly jigs as the microwires are positioned and bonded in place on the printed circuit board (PCB). These same tabs are also used later in the manufacturing process as a palette for conductive paint as it is applied at the microwire to printed circuit trace junction. Guide holes in the PCB are also used to accept alignment pins when multiple arrays are combined or stacked. FIGURE 1.1 Two 35-μm electrode tips, top is angle cut, bottom is blunt cut. © 2008 by Taylor & Francis Group, LLC
4 Methods for Neural Ensemble Recordings, Second Edition THE LAYERED APPROACH The various methods employed to manage and organize microwires into arrays can be classified as either layered or discretely wired. In some cases both approaches are used to combine microelectrode arrays on to a single-output connector to streamline the interface to the subject under investigation. The layered approach involves designing PCBs for the purpose of adapting the spacing of the electrode array to the spacing of the chosen connector and providing a platform on which the array components can be mounted. As shown in Figure 1.2, the plated through-hole pads facilitate the mounting of a surface mount connector, and the traces connected to each pad are positioned to mechanically and electrically bond to microwires in an array. The PCB designs are very specific and dedicated to a limited number of applications because the thick- ness of the board determines row spacing, and the printed circuit trace determines the spacing of electrodes within each row. Alignment jigs, specific to each array pattern and variation, are fabricated for use in the assembly process. The layered approach requires a significant initial invest- ment, and minor changes can be costly because the boards and jigs are specifically dedicated to a given application. The benefit of this approach is that higher production volumes are achievable, meeting the demand with repeatable high quality devices. The majority of layered designs are arrays with two row groups, combined with other assemblies to form multiple rows, or used singularly as a two-row array. Varia- tions of this design exist in many forms. Figure 1.2 illustrates a design in which two separate arrays are built onto a single board and combined onto a single connector. This design allows for implanting into two regions of the brain in a single step. Combining the signals in one output connector helps to streamline the interface to the subject. A further variation of this layered design incorporates remote arrays or satellite arrays. These satellite arrays are connected to the central array via flexible cable assemblies. With this design, the electrode arrays are positioned independently but utilize a single output connector. FIGURE 1.2 Two 4 × 4 electrode arrays share a common 32-channel connector. © 2008 by Taylor & Francis Group, LLC
State-of-the-Art Microwire Array Design for Chronic Neural Recordings 5 Figure 1.3 shows a layered array which contains the additional feature of being movable in depth after installation. Several millimeters of travel allow researchers to fine tune the electrode position and also extend the usefulness of the array. A further variation of this layered design incorporates remote arrays or satellite arrays. These satellite arrays are connected to the central array via flexible cable assemblies. With this design, the electrode arrays are positioned independently, but utilize a single- output connector. As shown in Figure 1.3, taken in midfabrication, a 2 × 8 electrode array is com- bined with a moveable 2 × 4 array and a remote or satellite 2 × 4 array (not pictured). All electrode wires in these arrays are 35 μm tungsten spaced at 250 μm. The spac- ing between the 2 × 8 and moveable 2 × 4 arrays is 300 μm. A 32-channel pream- plifier commonly referred to as a headstage, can be seen connected to the output connector of the electrode array. The headstage provides a buffer between the electrode and the recording equip- ment in terms of amplification, impedance matching, and filtering. It is sometimes necessary to fabricate electrodes with the headstage in place to allow for a work- ing clearance with electrode holders, stimulation connectors, moveable components, injection cannula, or other ancillary features. Figure 1.4 shows a custom-built electrode holder attached at the clamp points of a 2 × 16 array PCB. This board adapts the spacing of a 0.025 in. surface mount connector to the array pattern of 250 μm on centers. The board is FR4 composite material. The overall thickness dimension of the bare FR4, as seen at the right edge of the board, is a major determining factor in the spacing between rows of the array. The dark green area is a solder-masking agent for insulation. FIGURE 1.3 A 2 × 8 array and a movable 2 × 4 array. © 2008 by Taylor & Francis Group, LLC
6 Methods for Neural Ensemble Recordings, Second Edition The PCB shown in Figure 1.5 illustrates the manner in which microwires are mechanically bonded at the edge of the board and electrically bonded in a staggered formation along the gold-plated traces. The black area is a solder-mask agent used to provide electrical insulation. The tab to the right was used as a palette during the application of conductive paint and can be removed along with the tab on the left later in the fabrication process. The hole in each tab facilitates alignment if multiple assemblies are combined. The wires are 35 μm diameter tungsten with polyimide insulation. The insulation is removed at the silver paint locations to provide the best possible conductivity. FIGURE 1.5 A close-up view of microwire electrodes attached to a printed circuit board. FIGURE 1.4 Electrode holder clamping a 2 × 16 PCB prior to the installation of the surface- mount connector. © 2008 by Taylor & Francis Group, LLC
State-of-the-Art Microwire Array Design for Chronic Neural Recordings 7 With the layered approach, the assemblies, which form two rows of electrodes, are combined and connectors are installed on each PCB or bridged between boards. As shown in Figure 1.6, the assemblies can be stacked to combine high-density arrays, or used singularly as a two-row array. Fixtures or jigs are designed and constructed for the purpose of achieving align- ment of multiple PCBs or, in many cases, soft moldable clay can be used to tempo- rarily hold alignment while a permanent bond is made with adhesive. The example shown in Figure 1.6 is a 96-channel, 6 × 16 electrode array with 35 μm wire spaced at 250 μm on center. This array consists of three 2 × 16 assem- blies aligned and bonded with a coiled common ground wire in the foreground. A single set of electrode holder tabs are left in place, which will be removed in the final stages of surgery. The electrode holder tabs are reinforced by plated cop- per cladding, which provides a robust mechanical and electrical ground connection between the electrode and stereotaxic equipment during surgery. The 16-channel 4 × 4 array with 50 μm wires spaced at 250 μm shown in Figure 1.7, is formed by joining two 2 × 4 assemblies. A double row surface-mount nano connector spans the pads of both boards. The two boards are separated using a shim material to obtain the correct spacing. The staggered microwire termination pattern, used to promote ease of assembly, is visible through the clear epoxy coating. The coating is applied for insulation and additional mechanical integrity. A single set of holder tabs is left in place, and will be removed after installation. Multiple array designs can sometimes be included on the same PCB in the inter- est of promoting economy by design. Shown in Figure 1.8, two spacing variations FIGURE 1.6 96-channel, 6 × 16 array with 35 μm electrodes spaced at 250 μm in mid- fabrication. © 2008 by Taylor & Francis Group, LLC
8 Methods for Neural Ensemble Recordings, Second Edition of layered 4 × 4 arrays occupy a common electrode-printed circuit assembly, joined at the holder tabs. Guide plates, visible in the center, are also included in the design and will be removed for use in fabrication jigs or included in arrays with ultralong FIGURE 1.7 A 4 × 4 array completed and ready for testing. FIGURE 1.8 Multiuse layered array PCB design being fitted with microwires in an assem- bly jig. © 2008 by Taylor & Francis Group, LLC
State-of-the-Art Microwire Array Design for Chronic Neural Recordings 9 electrode wires that require floating guide plate supports. The board is shown mounted in an assembly jig and partially completed with one set of 35 μm electrode wires attached at corresponding traces of the board. Two sets of pads per trace are included to provide a universal mounting for the connector or for remote wiring of a satellite array. The jig is fabricated from a vast collection of the salvaged remains of routed FR4 panels in conjunction with the custom-drilled wire guides. The critical dimensional requirement of the bare FR4 thickness of a layered array is easily met by using the scrap materials from the production run. The supporting deep green-colored frame- work material in this case is FR4 material salvaged from other PCB production runs. The electrode featured in Figure 1.9 is a 32-channel, dual 4 × 4 layered array with electrodes spaced at 250 μm. The arrays are spaced at 2.5 mm. Guide tubes for drug injection cannula are fabricated from 23-gauge stainless steel tubing. The guide tubes are shown with inner pins fabricated from 0.013 in. stainless steel wire. The inner pins maintain a clear tract and are replaced with injection cannula when needed. The black material is a rubberized cyanoacrylate adhesive, which is used to pro- vide mechanical strength to the assembly, bonding the connector and guide tubes firmly to the PCB. The layered 128-channel array shown in Figure 1.10 illustrates how the separate assemblies are combined to provide a high density array. This array features 35 μm tungsten electrodes spaced at 250 μm and organized in eight rows of 16 electrodes. The electrode holder tabs have been removed on all assemblies except one, which will provide a clamp point for the entire assembly. The coiled ground wire is a com- mon ground connection to all four assemblies and is attached after the individual assemblies are combined. The connectors have been spaced to provide clearance to allow the four 32-channel headstages to simultaneously interface. This design features an offset between the array and connector to allow the array to be in close proximity to other arrays. This offset shifts the array footprint to coincide with the FIGURE 1.9 Dual 4 × 4 layered array with drug-injection cannula guides. © 2008 by Taylor & Francis Group, LLC
10 Methods for Neural Ensemble Recordings, Second Edition connector footprint along two planes in order to allow clearance for adjacent arrays or to shift the connector location to a more suitable location on the subject. DISCRETELY WIRED APPROACH The discretely wired approach is to form the array with straight microwires, mechan- ically bonding the microwires in the desired pattern and then routing the free ends of microwires to the connector. PCBs designed for this application are a tremendous benefit with regard to conformity in manufacturing, especially when small-scale, surface-mount nano-size connectors are used. This approach can be easily applied to achieve complex spacing and shapes, more so than with the layered approach. A single PCB design can be utilized for a multitude of array designs, because the printed circuit design is no longer directly related to the array pattern. In this case, the PCB provides two functions: to provide a structural component used to hold and position the assembly during manufacture and implantation, and to provide a mechanical and electrical bonding of the wire to the connector pin. With moveable designs, the PCB can also be used as the structural foundation for the moveable implements of the array. These boards also include features similar to boards used with layered arrays, such as tabs for clamping and holding the assem- bly, tabs for use as a palette, guide holes for alignment, and array patterns. Assembly jigs are designed and built specifically for each array pattern and can be partially or selectively loaded with microwire to yield variations within each FIGURE 1.10 128-channel 8 × 16 layered array. © 2008 by Taylor & Francis Group, LLC
State-of-the-Art Microwire Array Design for Chronic Neural Recordings 11 pattern. These arrays can be used in various combinations of satellite configurations or as a singular device. The 64-channel electrode array shown in Figure 1.11 was built using a discrete method and sacrificial wire guide. The wire guide in this case is used as a part of the assembly jig and is incorporated as part of the electrode assembly. The microwires are 35 μm diameter, spaced on a 1000 μm grid. Three separate PCBs are used in this example, one for each of the two connectors, and the third is to establish and maintain the spacing of the electrodes forming the wire guide. Epoxy is used as a conformal coating to provide insulation and mechanical integrity. A 32-channel independently positionable array is shown in Figure 1.12, during the assembly process attached to the jig. The pins on the left are 0.010 in. diameter tungsten pins; the wires on the right are 35 μm tungsten electrodes with a spacing of 1000 μm on center. In this moveable design, the electrode wire is attached at the tip of the pin, and a coil is formed around the pin to store the additional wire needed in travel. The pin is friction fit into a molded grid and transfers motion to the electrode wire. Silicon gel is molded at the electrode-to-pin interface to provide protection and allow for motion within. The electrode signal wires can be seen at the top of the picture routed to a 32-channel dual row connector. The electrodes can be positioned independently for peak efficiency. Several meth- ods can be used to move the electrodes. A simple manual pin-pushing device can be built with an outer guide tube and inner adjustable depth stop. This can be adjusted with the aid of a micrometer for exact depth settings. A variation on approach is to modify a micrometer, adding the outer guide tube and inner push rod directly to the micrometer as shown in Figure 1.13. A more complex variation of this method is illustrated in Figure 1.14 and Figure 1.15. FIGURE 1.11 (See color insert following page 140.) 8 × 8 electrode array fabricated using the discrete-wired method. © 2008 by Taylor & Francis Group, LLC
12 Methods for Neural Ensemble Recordings, Second Edition The device shown in Figure 1.14 is a miniature linear actuator, motorized and encoded to provide accurate movements of the electrode friction pin-drive system. A touch-down sensor in the drive mechanism allows the device to be used in a hand- held manner. The operator positions the guide tube over the electrode drive pin, bot- toming the guide tube in the electrode shell. FIGURE 1.13 A micrometer modified to provide movements to a friction pin moveable electrode. FIGURE 1.12 A discrete-wired array still attached to the assembly jig. © 2008 by Taylor & Francis Group, LLC
State-of-the-Art Microwire Array Design for Chronic Neural Recordings 13 Using the controller shown in Figure 1.15, the linear actuator is advanced until the inner drive rod contacts the electrode drive pin. Upon contact, the linear actuator is automatically stopped. The operator then enters the amount of travel and speed of travel desired at the electrode. A switch mounted on the actuator enables the control- ler to begin advancing the electrode at the discretion of the operator. The controller provides readout of the motion in microns, and has the capacity to store and retrieve position information, and communicate via RS-232. The electrode in Figure 1.16 is an example of a completed array that incorporates friction fit pins to independently position the electrodes. The main body of the array is fabricated by modifying a type F connector shell and forms the enclosure for the moveable components. The protective cap, threaded on the top of the body, has a FIGURE 1.14 A miniature linear actuator equipped to advance moveable electrodes. FIGURE 1.15 Hand-held electrode positioning system. © 2008 by Taylor & Francis Group, LLC
14 Methods for Neural Ensemble Recordings, Second Edition shaft that provides for stereotaxic attachment. This cap is replaced with a flat top cap after the array is firmly affixed in surgery. The remote-wired connector can be strategically positioned in surgery as needed to provide clearance. The electrode featured in Figure 1.17 is a 16-channel 2 × 8 moveable array using 15 μm tungsten wire. The electrode wires are drawn to a bundle and driven through a cast alignment block with a maximum deployment of 2.0 mm. Single-row connec- tors are used to connect to two 8-channel headstages. The electrode is shown prior FIGURE 1.17 A 16-channel moveable array used with dual 8-channel headstages. FIGURE 1.16 An independently positionable 32-channel array and stereotaxic-mounted cap. © 2008 by Taylor & Francis Group, LLC
State-of-the-Art Microwire Array Design for Chronic Neural Recordings 15 to completion without a conformal coating. Adjustments are made by rotating the slotted head screw at the top. The 16-channel electrode in Figure 1.18 has the moveable drive assembly built into the body of the connector. The drive mechanism is a 0–80 threaded screw, which advances the electrodes at the rate of 317.5 μm/r. The maximum travel for this electrode is 2.0 mm. BUILDING A LAYERED PCB MICROWIRE ARRAY The first step in building the DUCN microwire array is to design and fabricate the PCBs. With the design parameters such as the array pattern and connector pad layout established, the PCB design will be largely governed by these decisions. A PCB software tool is used to create the artwork files that are then used to manufacture the printed circuits. The files can be e-mailed to a PCB manufacturing facility for use in automated processing machines. Boards can be created, tested, and shipped within a matter of days. It is also possible to have the connectors installed by automated process, if desired. With the PCBs in hand, the next step is to fabricate a jig to assist in aligning the microwires to the board for bonding. The basic concept is to have two identical panels, drilled on centers to the array pattern with a center-mounted slotted beam in place to hold the PCB in the correct position relative to the panels. The microwires are loaded into the holes, a PCB is positioned into the slot, and the process of bonding the wires to the board can begin. Several assemblies can be produced before the jig will require reloading. The panels are drilled with slightly larger size holes than the outside diameter of the insulated microwire. A typical hole size is 90 μm for a 50 μm outside diameter (OD) microwire. The hole size can be critical; if the hole does not provide enough clearance for the OD variations of the wire, the insulation could be damaged and the jig will be more difficult to operate. The panels are usually FIGURE 1.18 A 16-channel moveable array used with a 16-channel headstage. © 2008 by Taylor & Francis Group, LLC
16 Methods for Neural Ensemble Recordings, Second Edition thin sheets of Delrin, FR4 composite material, or paper. The jig components can be included in the layout of the PCB or be built by hand. If the panels will be pro- duced by hand, a precision drill press with X, Y, and Z feed will be needed. A Servo model 7060 drill press and a Newport model 200 XY table (shown in Figure 1.6) are capable of producing good quality jig guide plates. It is helpful to specify that the scrap materials from the routed panels be shipped with the PCBs because the exact thickness of the boards is needed to be matched on the jig for optimum alignment. This scrap material can be used to fabricate the entire jig, or combined with other production runs as needed. With the microwires attached, the assemblies can be tested, and the connector and ground wire installed. A conformal coating is applied as an insulation barrier. In Figure 1.19, a 17 × 2 array is being assembled with a jig. The microwire array spac- ing is established by drilling the pattern in Delrin. A split beam holds the printed circuit directly over the array pattern as the microwires are attached to the board in a staggered length formation. The jig is then flipped and the process is repeated on the other side of the PCB. With both sides connected the board is removed from the jig slightly and the microwires cut to free the assembly. Several boards can be processed before the jig must be reloaded with wire. Micro drill bits in sizes ranging from 50 μm to well over 100 μm can be used in the fabrication of the jig. These drill bits are used with the drill press and translation stage shown in Figure 1.20. The drill press has a standard Albrecht keystone chuck, but for precision drilling a 1-mm-diameter collet will provide the best accuracy and ease of use with micro drill bits. The flute length versus diameter of commercially FIGURE 1.19 Microwires are held in alignment with a jig. © 2008 by Taylor & Francis Group, LLC
State-of-the-Art Microwire Array Design for Chronic Neural Recordings 17 available bits shown below indicates the maximum material thickness for a through- hole in the jig plate. The close-up view of a 90 μm drill bit in Figure 1.21 illustrates the flute length of 500 μm versus diameter. Due to their fragility, micro drill bits require very close tolerance drilling equipment to function. Drill Diameter (in microns) Flute Length (in microns) 50 400 60 400 70 400 80 500 90 500 100 700 110 700 FIGURE 1.20 Precision drilling equipment. © 2008 by Taylor & Francis Group, LLC
18 Methods for Neural Ensemble Recordings, Second Edition BUILDING A DISCRETELY WIRED ARRAY The first step in building this type of array is to fabricate a jig, which will be used to hold the microwires in alignment for a mechanical bonding process. The jig consists of two thin plates drilled to match the pattern of the array, mounted to a frame with approximately 25 mm space between. The microwires are loaded into the jig and mechanically bonded as an assembly. If the spacing is greater than 300 μm center to center, it can be helpful to use a third sacrificial plate to act as a carrier for the adhesive. This sacrificial plate helps to maintain the integrity of the pattern. The assembly can then be removed from the jig and mechanically bonded to a connector printed circuit assembly. The free ends of the microwires are then routed through plated through-holes, which are electrically connected to the conductor pins of the connector and mechanically bonded. The board is flipped to expose the microwire- plated hole junction. The microwires are trimmed to length, insulation is removed, and conductive paint or epoxy is applied at this junction. A conformal coating is applied to achieve an insulation barrier. The discrete wired jig shown in Figure 1.22 is designed to make a 4 × 8 array with electrode spacing of 500 μm and a row spacing of 800 μm. The jig is loaded with 35 μm wires, fixing to be bonded to form the array. Delrin blocks are used to help the adhesive span the distances between rows in order to maintain spacing integrity. The 16-channel adjustable electrode shown in Figure 1.23 features a 0.050 in. hex socket exposed at the top of the assembly, which is used to deploy the microwire electrodes from the cannula after being implanted over the target area. The adjustable mechanism is installed in an extruded brass square tube. The 0–80 threads of the mechanism provide incremental travel of 317.5 μm/r of the hex FIGURE 1.21 A 90 μm diameter drill bit typically used to drill microwire-guide arrays for jigs. © 2008 by Taylor & Francis Group, LLC
State-of-the-Art Microwire Array Design for Chronic Neural Recordings 19 socket. A single reinforced tab provides a clamp point for the stereotaxic electrode holder. The signal wires are terminated remotely to the connector PCB adapter of another array. FIGURE 1.22 Discrete-type jig loaded with 35 μm wires. FIGURE 1.23 A 16-channel moveable array shown with electrodes partially deployed. © 2008 by Taylor & Francis Group, LLC
20 Methods for Neural Ensemble Recordings, Second Edition CONCLUSIONS The design of a large variety of microwire array (bundle) configurations has enabled our laboratory to perform chronic, multielectrode recordings in a variety of animal species, including wild type and transgenic mice, rats, and owl and rhesus monkeys. More recently, the same approach has been translated into a new methodology for monitoring brain activity in patients subjected to neurosurgical procedures. This new generation of microelectrode arrays has pushed the limits of systems neurophysiology and allowed, for the first time, the simultaneous monitoring of the activity of hundreds of individual neurons, distributed across multiple, inter- connected cortical and subcortical structures that define particular neural circuits (e.g., somatosensory, motor, gustatory, etc.) for long periods of time (weeks to years, depending on the animal species and experimental protocol) in behaving animals. REFERENCES Lebedev, M.A. and Nicolelis, M.A.L. (2006). Brain machine interfaces: Past, present and future. Trends Neurosci 29: 536–546. Nicolelis, M.A.L., Ghazanfar, A.A., Faggin, B., Votaw, S., and Oliveira, L.M.O. (1997). Reconstructing the engram: simultaneous, multiple site, many single neuron record- ings. Neuron 18: 529–537. Schwartz, A.B. (2004). Cortical Neural Prosthetics. Annu Rev Neurosci 27: 487–507. Verloop, A.J. and Holsheimer, J. (1984). A simple method for the construction of electrode arrays. J Neurosci Meth 11: 173–178. Williams, J.C., Renmaker, R.L., and Kipke, D.R. (1999). Long-term neural recording char- acteristics of wire microelectrode arrays implanted in cerebral cortex. Brain Res Pro- tocols 4: 303–313. © 2008 by Taylor & Francis Group, LLC
21 2 Surgical Techniques for Chronic Implantation of Microwire Arrays in Rodents and Primates Laura M. O. Oliveira and Dragan Dimitrov And still we could never suppose that fortune were to be so friendly to us, such as to allow us to be perhaps the first in handling, as it were, the electricity concealed in nerves, in extracting it from nerves, and, in some way, in putting it under everyone’s eyes. Luigi Galvani, 1791 CONTENTS Introduction..............................................................................................................22 Differences Between Rodents and Primates Pertinent to Surgical Technique........23 Surgical Techniques for Rodents .............................................................................24 Preoperative Supplies and Room Preparation ..............................................24 Preoperative Animal Preparation.................................................................25 Anesthesia Techniques and Intraoperative Monitoring................................26 Electrode Specific for Rodents .....................................................................27 Implantation Techniques...............................................................................28 Brain Electrode Arrays......................................................................28 EMG Electrode Surgery .................................................................... 31 Postoperative Care........................................................................................32 Surgical Techniques for Primates............................................................................32 Preoperative Supplies and Room Preparation ..............................................32 Preoperative Animal Preparation.................................................................34 Anesthesia Techniques and Intraoperative Monitoring................................34 Electrode Specific for Primates....................................................................35 Implantation Techniques...............................................................................35 Exposure ............................................................................................35 Cortical Localization .........................................................................36 Drilling of Craniotomies....................................................................36 Opening of the Brain Coverings........................................................40 © 2008 by Taylor & Francis Group, LLC
22 Methods for Neural Ensemble Recordings, Second Edition INTRODUCTION The study of electrophysiology started with the work of Luigi Galvani (1737–1798), who was the first to provide evidence for the electrical nature of “the mysterious fluid” (at the time referred to as “animal spirits”). Galvani’s nephew, Giovanni Aldini (1762-1834), continued this line of inquiry in 1803, using Galvani’s and Alessandro Volta’s (bimetallic electricity) principles together, despite the fact that Volta did not believe in animal electricity. Carlo Mateucci (1818–1868) in Bologna and Emil Du Bois-Reymond (1818–1896) in Berlin described the phenomenon called “negative variation” when a galvanometer showed an unexpected decrease in current intensity during muscle contraction. The study of the electrophysiology of the nervous system began when Julius Berstein (1839–1917) proposed his theory of the nerve impulse as a wave of negativity (membrane theory of the nerve tissue). Later, using a galvanom- eter with one electrode in the gray matter and one on the skull surface (or electrodes in different points of the external surface of the brain), Richard Caton (1842–1926) recorded a feeble current in the brain. In 1870, for the first time, Gustav Fritsch (1838–1927) and Eduard Hitzig (1838–1907) inserted an electrode in the dura of a dog brain and stimulated the motor area, generating movement in the contralateral side of the animal’s body (Niedermeyer 1993; Piccolino 1998). The work of these and many other scientists marked the beginning of the study of the electrophysiology of the nervous system, opening doors to the possibility of stimulating different areas of the brain through electrical current and subsequently recording the brain electrical activity. Improvements in electrode manufacturing, the advent of modern acquisition equipment, and better surgical and asepsis techniques have provided us the ability to chronically implant multiple electrodes simultaneously in several areas of the brain in the same animal (Nicolelis, Baccala et al., 1995) and to study the interactions of populations of neurons (Nicolelis, Fanselow et al., 1997; Ghazanfar, Stambaugh et al., 2000). Upon the animal’s recovery from surgery, we have been able to record simultaneously from different brain areas of mice (Costa, Cohen et al., 2004), rats (Faggin, Nguyen et al., 1997; Ghazanfar and Nicolelis 1997; Nicolelis, Ghazanfar et al., 1997) and nonhuman primate brains (Nicolelis, Stam- baugh et al., 1999; Nicolelis, Dimitrov et al., 2003) for long periods of time, from a couple of months in rodents (Ghazanfar and Nicolelis 1997; Nicolelis, Ghazanfar et al., 1997) to up to years in non-human primates, such as owl monkeys (Nicolelis, Ghazanfar et al., 1998) and Rhesus monkeys (Nicolelis, Dimitrov et al., 2003). These recordings are carried out under several different experimental condi- tions and behavior tasks (Kralik, Dimitrov et al., 2001; Nicolelis and Ribeiro 2002). With chronically implanted multiple electrodes, it is also possible to record different Electrode Lowering............................................................................ 41 Skull and Wound Closure.................................................................. 41 Postoperative Care........................................................................................43 Conclusions and Future Directions..........................................................................44 Acknowledgments....................................................................................................45 References................................................................................................................45 © 2008 by Taylor & Francis Group, LLC
Surgical Techniques for Chronic Implantation of Microwire Arrays 23 layers of the same area of the brain (Chapin and Lin 1984) and study spatiotem- poral response of many neurons (Nicolelis and Chapin 1994). Microcannulae can be attached to the electrode arrays and are used to inject drugs in the areas of the implant during chronic experimental recordings (Shuler, Krupa et al., 2002). Chronically implanted electrodes offer unparalleled advantages for correlating neuronal activity and animal behavior. In our lab, these techniques were developed in rodents and later adapted to primates. Over the last several years, there have been significant strides in making rodent implantations more reliable, faster, and easier. We have identified and resolved many of the issues that now permit larger neuronal yields that last longer. As a consequence of continuous improvement in techniques, the length of time required for surgery has been reduced. At the same time, over the last 14 years, we have developed a surgical technique adapted to the unique features of primates. This has made primate implantations routine and reproducible. Here, we will describe detailed technical aspects of the current surgical implantation approach used in our laboratory at the Duke University Center for Neuroengineering (DUCN). Such a surgical protocol has evolved and benefited from almost two decades of accu- mulated experience on chronic multielectrode neural recordings (Nicolelis, Stam- baugh et al., 1999; Nicolelis, Dimitrov et al., 2003, Kralik, Dimitrov et al., 2001; Nicolelis and Ribeiro 2002). DIFFERENCES BETWEEN RODENTS AND PRIMATES PERTINENT TO SURGICAL TECHNIQUE The success in obtaining recordings from chronically implanted electrodes and how long they last depends fundamentally on the quality of the surgical implantation tech- nique. The ability to open small craniotomies, only large enough to fit the electrode array with minimal bleeding from the bone and from the meninges in very small animals such as mice and rats, requires practicing the techniques several times before attempting to gather good data using the implant of electrode arrays. This is especially true in cortex areas, which are close to the dura and are very sensitive to lesions. The surgical technique for implantation that we follow in our lab is similar for all species, but each species has its own peculiarities. For instance, the mouse head is very small and the skull is thin, requiring a delicate technique. Mouse surgeries require very small screws, drill bits, and custom-designed electrode arrays. Because of the small head size and bone thickness, inserting more than two small arrays of electrodes and more than two fixation screws per animal is not recommended. Compared to mice, the rat’s head is bigger and the bone is stronger, increasing the possibility of using bigger electrode arrays and reaching more cortical and subcorti- cal areas. Because rats are stronger than mice, they can tolerate larger amounts of the acrylate used to secure the electrodes in place, which increases the number of arrays that can be inserted in the brain (Faggin, Nguyen et al., 1997; Nicolelis and Ribeiro 2002). Cortical and subcortical localization in rodents is based on commercially available stereotactic atlases. Theoretically, implanting electrodes into rodents and primates should be very similar; however, in practice they are vastly different. For investigators accustomed © 2008 by Taylor & Francis Group, LLC
24 Methods for Neural Ensemble Recordings, Second Edition to the hardiness of rodents and trained in rodent implantation techniques, per- forming similar procedures on primates can seem overwhelming. For instance, the commitment of time, personnel, and lab resources is much greater for primate surgery. Furthermore, from the point of anesthesia, primates require much more atten- tion during surgery. Whereas rodents require only monitoring of a few physiologi- cal parameters and infrequent injections, maintaining a primate under anesthesia is much more labor intensive. Thus, for our primate surgeries, one member of the surgery team is dedicated to monitoring the animal throughout. This higher level of anesthesia technique is akin to pediatric anesthesia and requires specific equipment, planning, and personnel. Obvious differences in anatomical details include a thicker skull, thicker brain coverings, and a better-developed subdural space, all of which also influence the definition of the optimal surgical strategy for chronic multi-electrode implantation in nonhuman primates. The thicker skull requires forethought in terms of the appro- priate electrode length. The brain coverings including the dura and pia are better developed, more variable in their thickness and at least the dura requires wide open- ing with microsurgical instruments for electrode penetration. We have found that the pia and arachnoid layers of primates are more variable, often tougher, and more prone to dimpling than rodents, sometimes necessitating microsurgical opening, as well as paying careful attention to be sure penetration has occurred. The primate brain is more prone to problems with swelling and retraction due to fluctuations in CO2 levels in the blood that are primarily affected by ventilation of the animals’ lungs. Plans must be in place to deal with these issues intraoperatively. Overall, we have also observed that intraoperative neural recordings are more difficult in primates than they are in rodents. Much more attention must be paid to noise detection and reduction, especially because many more electrical devices nec- essary for anesthesia are involved. Primates are more prone to infection than rodents and require more attention to sterile technique throughout, with sterilization of all the instruments, the use of sterile gloves and gowns, and draping of the surgical field. For those unaccustomed to working in a sterile environment for long periods of time, this can present unfore- seen challenges and cost significant amounts of time. In summary, primate implantations require a team approach. It often takes sev- eral days to weeks to prepare and coordinate one’s lab prior to performing primate surgery, underscoring the critical importance of preoperative planning. SURGICAL TECHNIQUES FOR RODENTS PREOPERATIVE SUPPLIES AND ROOM PREPARATION Typically, our rodent surgeries are performed on a surgery table in a regular room in the laboratory or in a designated surgical room. The table is kept clean and unclut- tered and has the stereotaxic apparatus already installed (for rats we use the cat and small primates stereotaxic apparatus with a rat adaptor, and for mice, we use the stereotaxic apparatus for small rodents, both from David Kopf Instruments, Tujunga, © 2008 by Taylor & Francis Group, LLC
Surgical Techniques for Chronic Implantation of Microwire Arrays 25 California). Between the two stereotaxic bars, a rectangular platform made of plexi- glass is glued to a height adjustable stage where a warm pad is placed. This apparatus is used for the anesthetized animal for the duration of the surgery and immedi- ate postoperative recovery. Other essential pieces of equipment include a binocu- lar surgical microscope, a dental drill already installed in a source of compressed air, an amplifier connected to an audio monitor, an oscilloscope, and a microposi- tioner. The table, the stereotaxic apparatus and the plexiglass platform are cleaned before the surgery with 70% alcohol or Asseptiwipes (wipes moistened in a solu- tion of N-alkyl(68%C12, 32%C14) dimethyl ethyl benzyl ammonium chloride 0.125%, N-alkyl(60%C14, 30%C16, 5%C12, 5%C14) dimethyl ethyl benzyl ammonium chloride 0.125%, isopropyl alcohol 14.850%, and other ingredients. All our surgeries follow the National Institute of Health (NIH) and Associa- tion for Assessment and Accreditation of Laboratory Animal Care (AAALAC) guidelines for rodents that are expected to recover from anesthesia. These guide- lines include appropriate preoperative and postoperative care of the animals, asepsis (sterilization of the surgical tools, use of sterile gloves, mask, head covers, clean lab coat, and aseptic procedures), gentle tissue handling, minimal dissection of tissues, effective hemostasis, and correct use of suture materials and patterns when stitches are necessary (National Research Council 1996; Baumans, Remie et al., 2001). The surgical instruments (basically, four small hemostats, scalpel handle, Freer periosteal elevator, scissors, micro scissors, regular and micro tweezers, dental drill handpiece, special screw drivers, stainless steel wire cutters, and micro cup curette) and supplies (gauze, cotton-tip applicators, kimwipes, drill bits, small stainless steel screws, silicone cups for dental acrylic, and beakers) are sterilized by autoclave. All supplies that cannot be exposed to heat and humidity are sterilized by ethylene oxide gas (extrafine point markers, plastic rulers, electrode holders, electrodes, etc.). The microwire arrays (see chapter 1) are made of very delicate materials and cannot be sterilized by autoclave. Instead, these must be sterilized by ethylene oxide gas. The electrodes should be carefully packed in such a way that they are not dam- aged during handling, transportation to and from the sterilization facility, or by personnel helping in the surgery. Because of the length of these surgeries, all the equipment used during surgery should be routinely tested the day before the animal is anesthetized. To avoid unnecessary delays during surgery, it is imperative that all materials, equipment, and drugs for the surgery are readily available and in good working condition. A copy of the brain atlas appropriate to the animal undergoing implantation surgery is necessary to help in the location of the areas to be implanted. It is advisable to complete these steps the day before the surgery; however, they can be completed the day of the surgery prior to anesthetizing and preparing the animal. PREOPERATIVE ANIMAL PREPARATION Typically, all animals selected to undergo chronic implantation of multielectrode arrays have several days to acclimate in the new laboratory facility before surgery. Often, these animals are subjected to weeks of behavioral training before they are implanted with arrays of electrodes. Only animals in good health are subjected to the surgery. © 2008 by Taylor & Francis Group, LLC
26 Methods for Neural Ensemble Recordings, Second Edition In general, we work with Long–Evans adult rats, males (in general 300–350 g) or females (in general 250–300 g). However, depending on the studies we can use younger or older animals. Mice are chosen according to their genotype since our laboratory currently utilizes a variety of mutant animals and their wild-type litter- mates as control (Costa, Lin et al., 2006; Dzirasa, Ribeiro et al., 2006). Because rodents have a high metabolism and are not at risk for vomiting and aspirating, food is not withheld until up to 2 h before the surgery. Plans for anesthesia, surgery, and determining the coordinates of the areas to be implanted must be carefully studied ahead of time. This will help decrease the time of the surgery for electrode-array implantation. A record sheet should be started for the procedure prior to beginning anesthesia. This not only meets the requirements of AAALAC, NIH, and Institutional Animal Care and Use Committee (IACUCs), but also provides a record of the procedure, the implanted areas and their coordinates, the depth of the electrode for each array, and the position of the ground wires related to the position of the connector of the array. ANESTHESIA TECHNIQUES AND INTRAOPERATIVE MONITORING Following guidelines from Duke University’s IACUC and the advice of Duke’s veteri- nary staff, we use two anesthesia regimens for rat surgeries and one for mice surgeries. For rats: Pentobarbital IP Ketamine associated with Xylazine IM For mice: Ketamine IM associated with Xylazine IM Pentobarbital has a good hypnotic effect, but analgesia is obtained only when using high doses of the drug, which can cause cardiovascular and respiratory depression. The advantage of using Pentobarbital is that the effect of the anesthesia lasts longer than the Ketamine/Xylazine combination. Ketamine is a dissociative anesthetic that provides light surgical anesthesia with a short duration. When combined with sedatives or tranquilizers, the quality of the anesthesia is highly improved and the duration of the effect is increased. In general, in our surgeries supplemental doses of Pentobarbital are required every 2 h and Ket- amine every 1½ h. The supplemental doses for both drugs are from ⅓ to ½ of the original dose, but Xylazine is not supplemented unless the surgery lasts longer than 7–8 hours, which is very uncommon in rodent surgeries. For both anesthesia regimens (rats and mice), anesthetic induction is carried out in a chamber with a mixture of 5% Isoflurane and O2. In our experience, a good induction with Isoflurane helps the selected anesthesia regimen last longer, decreas- ing the need for frequent injections of supplemental doses of the anesthetic. Once the animal is deeply anesthetized with Isoflurane, an injection of the selected drug is administered (Duke University DLAR 1995; National Research Council 1996; Hellebrekers and Booij 2001). Following the injection of anesthesia, the animals © 2008 by Taylor & Francis Group, LLC
Surgical Techniques for Chronic Implantation of Microwire Arrays 27 may receive a dose of Atropine SQ to prevent or treat excessive secretion of the airways and to improve cardiac function. This is important especially with the use of Pentobarbital. For rats, if the surgery is expected to last for a longer period, small doses of sterile saline can be injected SQ, and IP injections of 10% dextrose solution will help to maintain hydration and blood glucose level during the surgery. In general, mice surgeries are much shorter than rat surgeries due to the small number of screws and arrays that are implanted, and extra volume or dextrose injections are not necessary. Once the animal is deeply anesthetized, it is placed in an area for preparation for surgery, apart from the operating theater. The animal’s head is shaved with small clippers, from the area just above the eyes to the back of the head and from ear to ear. If electromyography (EMG) electrode implants are planned, the skin on top of the target muscles is also shaved at this point. After the shaved hair is cleaned, the animal is transported to the surgery table and placed on the platform with a thermal pad to maintain the animal’s body temperature throughout the surgery. If an electrical pad is used, the pad should be covered to avoid direct contact with the animal in order to prevent burns to the animal’s skin in case the pad overheats, and a rectal temperature probe should be inserted. A good option is the Deltaphase® ther- mal pads (Braintree Scientific, Braintree, Massachusetts) that maintain temperature around 37ºC for about 5–6 h. This type of pad must be monitored and changed when the temperature decreases. After the animal is on a warm pad, it is mounted on the stereotaxic apparatus, using proper ear bars for rodents, placing the mouth and nose piece, and leaving the position of the head fixed until the top of the head is parallel to a flat surface. Once the rat head is secure, the sterile pack can be opened and the sterile materi- als should be kept in a sterile field or container. The skin of the head is cleaned three times using Iodophor (iodine soaps) or Chlorhexidine followed by alcohol 70%. The animal’s eyes are covered with an ophthalmic ointment to prevent corneal lesions since the animal loses the blinking reflex with the anesthesia. ELECTRODE SPECIFIC FOR RODENTS The arrays of electrodes to be implanted are made in-house and vary in number and distribution of electrodes. For the past nineteen years, our experiments have utilized arrays or bundles made of thin microwires (see chapter 1 for details). Some arrays can be made to reach more than one area of the brain. In general, for rat surgeries, the arrays are comprised of 32 electrodes (4 × 8), but they can also be made up of either 16 (2 × 8) or 64 (8 × 8) electrodes. The tips are cut either blunt or sharp. When cut sharp, the electrodes may be able to penetrate the brain without dissecting the dura. In this case, the length of each electrode may be slightly different (see Figure 2.1). If necessary, a little cut on the dura with a bent fine needle may help relieve the tension of the meninges and facilitate the implantation of the electrodes. Obviously, the length of the microwires varies for each surgery and depends on the brain area to be implanted. © 2008 by Taylor & Francis Group, LLC
28 Methods for Neural Ensemble Recordings, Second Edition IMPLANTATION TECHNIQUES Brain Electrode Arrays The length of the surgery for the implantation of microwire arrays varies from ani- mal to animal. It also depends on the number of arrays to be implanted, the location of such brain regions, and the experience of the surgeon. In general, the minimum time is about 3–4 h if only one or two arrays are to be implanted. Each additional array can add about 45–60 min to the total surgical time. It is essential not to rush the implantation procedure. In our accumulated experience, gentle and slow penetration of the brain tissue yields the best long-term results. During surgery, the status of anesthesia is checked periodically by pinching the inter-digit membrane of the hind paw or pinching the tails of the animal. Supplemen- tal doses of anesthesia are given as needed. The areas of the brain to be implanted also vary for each study and may vary even in the same study because it is not possible to reach all areas of interest for each animal, especially in mice. The surgical procedures described below have been very successful in the past. Following this approach has allowed our experimental animals to tolerate these implants very well, survive without any postsurgical com- plication, and provide high-quality recordings for months after the surgery. After the animal is anesthetized, mounted to the stereotax, and cleaned, the surgical team suits up for surgery in a lab coat or gown, mask, sterile gloves, and head cap. Before starting surgery, the status of the anesthesia is checked as described above. To alleviate pain during the incision, an injection of lidocaine 1% or Bipu- vocaine can be given under the skin at the site of incision. An incision is made on the midline of the scalp, from just above the eyes to the back of the head, and the skin is propped open. The borders of the skin are held open with small hemostats, and bleeding, which in general is very small, is cleaned with gauze or cotton-tip FIGURE 2.1 Sharp electrodes penetrating the dura in a rat surgery. Photo by Edgard Morya. © 2008 by Taylor & Francis Group, LLC
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Methods for neural ensemble recordings 2nd ed Edition Miguel A. L. Nicolelis

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    Methods for neuralensemble recordings 2nd ed Edition Miguel A. L. Nicolelis Digital Instant Download Author(s): Miguel A. L. Nicolelis ISBN(s): 9780849370465, 0849370469 Edition: 2nd ed File Details: PDF, 14.98 MB Year: 2008 Language: english
  • 7.
    METHODS for NEURAL ENSEMBLERECORDINGS SECOND EDITION © 2008 by Taylor & Francis Group, LLC
  • 8.
    FRONTIERS IN NEUROSCIENCE SeriesEditors Sidney A. Simon, Ph.D. Miguel A.L. Nicolelis, M.D., Ph.D. Published Titles Apoptosis in Neurobiology Yusuf A. Hannun, M.D., Professor of Biomedical Research and Chairman/Department of Biochemistry and Molecular Biology, Medical University of South Carolina, Charleston, South Carolina Rose-Mary Boustany, M.D., tenured Associate Professor of Pediatrics and Neurobiology, Duke University Medical Center, Durham, North Carolina Methods for Neural Ensemble Recordings Miguel A.L. Nicolelis, M.D., Ph.D., Professor of Neurobiology and Biomedical Engineering, Duke University Medical Center, Durham, North Carolina Methods of Behavioral Analysis in Neuroscience Jerry J. Buccafusco, Ph.D., Alzheimer’s Research Center, Professor of Pharmacology and Toxicology, Professor of Psychiatry and Health Behavior, Medical College of Georgia, Augusta, Georgia Neural Prostheses for Restoration of Sensory and Motor Function John K. Chapin, Ph.D., Professor of Physiology and Pharmacology, State University of New York Health Science Center, Brooklyn, New York Karen A. Moxon, Ph.D., Assistant Professor/School of Biomedical Engineering, Science, and Health Systems, Drexel University, Philadelphia, Pennsylvania Computational Neuroscience: Realistic Modeling for Experimentalists Eric DeSchutter, M.D., Ph.D., Professor/Department of Medicine, University of Antwerp, Antwerp, Belgium Methods in Pain Research Lawrence Kruger, Ph.D., Professor of Neurobiology (Emeritus), UCLA School of Medicine and Brain Research Institute, Los Angeles, California Motor Neurobiology of the Spinal Cord Timothy C. Cope, Ph.D., Professor of Physiology, Wright State University, Dayton, Ohio Nicotinic Receptors in the Nervous System Edward D. Levin, Ph.D., Associate Professor/Department of Psychiatry and Pharmacology and Molecular Cancer Biology and Department of Psychiatry and Behavioral Sciences, Duke University School of Medicine, Durham, North Carolina Methods in Genomic Neuroscience Helmin R. Chin, Ph.D., Genetics Research Branch, NIMH, NIH, Bethesda, Maryland Steven O. Moldin, Ph.D, University of Southern California, Washington, D.C. © 2008 by Taylor & Francis Group, LLC
  • 9.
    Methods in ChemosensoryResearch Sidney A. Simon, Ph.D., Professor of Neurobiology, Biomedical Engineering, and Anesthesiology, Duke University, Durham, North Carolina Miguel A.L. Nicolelis, M.D., Ph.D., Professor of Neurobiology and Biomedical Engineering, Duke University, Durham, North Carolina The Somatosensory System: Deciphering the Brain’s Own Body Image Randall J. Nelson, Ph.D., Professor of Anatomy and Neurobiology, University of Tennessee Health Sciences Center, Memphis, Tennessee The Superior Colliculus: New Approaches for Studying Sensorimotor Integration William C. Hall, Ph.D., Department of Neuroscience, Duke University, Durham, North Carolina Adonis Moschovakis, Ph.D., Department of Basic Sciences, University of Crete, Heraklion, Greece New Concepts in Cerebral Ischemia Rick C.S. Lin, Ph.D., Professor of Anatomy, University of Mississippi Medical Center, Jackson, Mississippi DNA Arrays: Technologies and Experimental Strategies Elena Grigorenko, Ph.D., Technology Development Group, Millennium Pharmaceuticals, Cambridge, Massachusetts Methods for Alcohol-Related Neuroscience Research Yuan Liu, Ph.D., National Institute of Neurological Disorders and Stroke, National Institutes of Health, Bethesda, Maryland David M. Lovinger, Ph.D., Laboratory of Integrative Neuroscience, NIAAA, Nashville, Tennessee In Vivo Optical Imaging of Brain Function Ron Frostig, Ph.D., Associate Professor/Department of Psychobiology, University of California, Irvine, California Primate Audition: Behavior and Neurobiology Asif A. Ghazanfar, Ph.D., Princeton University, Princeton, New Jersey Methods in Drug Abuse Research: Cellular and Circuit Level Analyses Dr. Barry D. Waterhouse, Ph.D., MCP-Hahnemann University, Philadelphia, Pennsylvania Functional and Neural Mechanisms of Interval Timing Warren H. Meck, Ph.D., Professor of Psychology, Duke University, Durham, North Carolina Biomedical Imaging in Experimental Neuroscience Nick Van Bruggen, Ph.D., Department of Neuroscience Genentech, Inc. Timothy P.L. Roberts, Ph.D., Associate Professor, University of Toronto, Canada The Primate Visual System John H. Kaas, Department of Psychology, Vanderbilt University Christine Collins, Department of Psychology, Vanderbilt University, Nashville, Tennessee Neurosteroid Effects in the Central Nervous System Sheryl S. Smith, Ph.D., Department of Physiology, SUNY Health Science Center, Brooklyn, New York © 2008 by Taylor & Francis Group, LLC
  • 10.
    Modern Neurosurgery: ClinicalTranslation of Neuroscience Advances Dennis A. Turner, Department of Surgery, Division of Neurosurgery, Duke University Medical Center, Durham, North Carolina Sleep: Circuits and Functions Pierre-Hervé Luoou, Université Claude Bernard Lyon, France Methods in Insect Sensory Neuroscience Thomas A. Christensen, Arizona Research Laboratories, Division of Neurobiology, University of Arizona, Tuscon, Arizona Motor Cortex in Voluntary Movements Alexa Riehle, INCM-CNRS, Marseille, France Eilon Vaadia, The Hebrew University, Jerusalem, Israel Neural Plasticity in Adult Somatic Sensory-Motor Systems Ford F. Ebner, Vanderbilt University, Nashville, Tennessee Advances in Vagal Afferent Neurobiology Bradley J. Undem, Johns Hopkins Asthma Center, Baltimore, Maryland Daniel Weinreich, University of Maryland, Baltimore, Maryland The Dynamic Synapse: Molecular Methods in Ionotropic Receptor Biology Josef T. Kittler, University College, London, England Stephen J. Moss, University College, London, England Animal Models of Cognitive Impairment Edward D. Levin, Duke University Medical Center, Durham, North Carolina Jerry J. Buccafusco, Medical College of Georgia, Augusta, Georgia The Role of the Nucleus of the Solitary Tract in Gustatory Processing Robert M. Bradley, University of Michigan, Ann Arbor, Michigan Brain Aging: Models, Methods, and Mechanisms David R. Riddle, Wake Forest University, Winston Salem, North Carolina Neural Plasticity and Memory: From Genes to Brain Imaging Frederico Bermudez-Rattoni, National University of Mexico, Mexico City, Mexico Serotonin Receptors in Neurobiology Amitabha Chattopadhyay, Center for Cellular and Molecular Biology, Hyderabad, India Methods for Neural Ensemble Recordings, Second Edition Miguel A.L. Nicolelis, M.D., Ph.D., Professor of Neurobiology and Biomedical Engineering, Duke University Medical Center, Durham, North Carolina © 2008 by Taylor & Francis Group, LLC
  • 11.
    CRC Press isan imprint of the Taylor & Francis Group, an informa business Boca Raton London New York Edited by Miguel A. L. Nicolelis Duke University Medical Center Durham, NC METHODS for NEURAL ENSEMBLE RECORDINGS SECOND EDITION © 2008 by Taylor & Francis Group, LLC
  • 12.
    CRC Press Taylor &Francis Group 6000 Broken Sound Parkway NW, Suite 300 Boca Raton, FL 33487-2742 © 2008 by Taylor & Francis Group, LLC CRC Press is an imprint of Taylor & Francis Group, an Informa business No claim to original U.S. Government works Printed in the United States of America on acid-free paper 10 9 8 7 6 5 4 3 2 1 International Standard Book Number-13: 978-0-8493-7046-5 (Hardcover) This book contains information obtained from authentic and highly regarded sources. Reprinted material is quoted with permission, and sources are indicated. A wide variety of references are listed. Reasonable efforts have been made to publish reliable data and information, but the author and the publisher cannot assume responsibility for the validity of all materials or for the conse- quences of their use. No part of this book may be reprinted, reproduced, transmitted, or utilized in any form by any electronic, mechanical, or other means, now known or hereafter invented, including photocopying, microfilming, and recording, or in any information storage or retrieval system, without written permission from the publishers. For permission to photocopy or use material electronically from this work, please access www. copyright.com (http://www.copyright.com/) or contact the Copyright Clearance Center, Inc. (CCC) 222 Rosewood Drive, Danvers, MA 01923, 978-750-8400. CCC is a not-for-profit organization that provides licenses and registration for a variety of users. For organizations that have been granted a photocopy license by the CCC, a separate system of payment has been arranged. Trademark Notice: Product or corporate names may be trademarks or registered trademarks, and are used only for identification and explanation without intent to infringe. Library of Congress Cataloging-in-Publication Data Methods for neural ensemble recordings / [edited by] Miguel A.L. Nicolelis. -- 2nd ed. p. ; cm. -- (Frontiers in neuroscience) Includes bibliographical references and index. ISBN 978-0-8493-7046-5 (alk. paper) 1. Electroencephalography. 2. Microelectrodes. 3. Neurons. I. Nicolelis, Miguel A. L. II. Series: Frontiers in neuroscience (Boca Raton, Fla.) [DNLM: 1. Neurons--physiology. 2. Brain--physiology. 3. Electrophysiology--methods. 4. Microelectrodes. WL 102.5 M592 2008] QP376.5.M47 2008 616.8’047547--dc22 2007027468 Visit the Taylor & Francis Web site at http://www.taylorandfrancis.com and the CRC Press Web site at http://www.crcpress.com © 2008 by Taylor & Francis Group, LLC
  • 13.
    To Pedro, Rafael,and Daniel, May your voyages be full of lore and fun © 2008 by Taylor & Francis Group, LLC
  • 14.
    ix Contents Series Preface............................................................................................................xi Preface.................................................................................................................... xiii Editor.....................................................................................................................xvii Contributors ............................................................................................................xix Chapter 1 State-of-the-Art Microwire Array Design for Chronic Neural Recordings in Behaving Animals ........................................................1 Gary Lehew and Miguel A. L. Nicolelis Chapter 2 Surgical Techniques for Chronic Implantation of Microwire Arrays in Rodents and Primates......................................................... 21 Laura M. O. Oliveira and Dragan Dimitrov Chapter 3 Technology for Multielectrode MicroStimulation of Brain Tissue..................................................................................................47 Timothy Hanson, Nathan Fitzsimmons, and Joseph E. O’Doherty Chapter 4 Strategies for Neural Ensemble Data Analysis for Brain–Machine Interface (BMI) Applications.............................................................57 Miriam Zacksenhouse and Simona Nemets Chapter 5 Chronic Recordings in Transgenic Mice............................................83 Kafui Dzirasa Chapter 6 Multielectrode Recordings in the Somatosensory System.................97 Michael Wiest, Eric Thomson, and Jim Meloy Chapter 7 Chronic Recording During Learning...............................................125 Aaron J. Sandler Chapter 8 Defining Global Brain States Using Multielectrode Field Potential Recordings ........................................................................ 145 Shih-Chieh Lin and Damien Gervasoni © 2008 by Taylor & Francis Group, LLC
  • 15.
    x Methods forNeural Ensemble Recordings, Second Edition Chapter 9 Multielectrode Recording in Behaving Monkeys ............................ 169 R. E. Crist and M. A. Lebedev Chapter 10 Neural Ensemble Recordings from Central Gustatory-Reward Pathways in Awake and Behaving Animals..................................... 189 Albino J. Oliveira-Maia, Sidney A. Simon, and Miguel A. L. Nicolelis Chapter 11 Building Brain–Machine Interfaces to Restore Neurological Functions .......................................................................................... 219 Mikhail A. Lebedev, Roy E. Crist, and Miguel A. L. Nicolelis Chapter 12 Conceptual and Technical Approaches to Human Neural Ensemble Recordings....................................................................... 241 Dennis A. Turner, Parag G. Patil, and Miguel A.L. Nicolelis © 2008 by Taylor & Francis Group, LLC
  • 16.
    xi Series Preface Our goalin creating the Frontiers in Neuroscience Series is to present the insights of experts on emerging fields and theoretical concepts that are, or will be, in the van- guard of neuroscience. Books in the series cover genetics, ion channels, apoptosis, electrodes, neural ensemble recordings in behaving animals, and even robotics. The series also covers new and exciting multidisciplinary areas of brain research, such as computational neuroscience and neuroengineering, and describes breakthroughs in classical fields like behavioral neuroscience. We hope every neuroscientist will use these books in order to get acquainted with new ideas and frontiers in brain research. These books can be given to graduate students and postdoctoral fellows when they are looking for guidance to start a new line of research. Each book is edited by an expert and consists of chapters written by the leaders in a particular field. Books are richly illustrated and contain comprehensive bibliog- raphies. Chapters provide substantial background material relevant to the particu- lar subject. We hope that as the volumes become available, the effort put in by us, the publisher, the book editors, and individual authors will contribute to the further development of brain research. The extent to which we achieve this goal will be determined by the utility of these books. Sidney A. Simon, Ph.D. Miguel A.L. Nicolelis, M.D.,Ph.D. Series Editors © 2008 by Taylor & Francis Group, LLC
  • 17.
    xiii Preface Almost 10 yearsago, as the fresh-off-the-press volumes of the first edition of Meth- ods for Neural Ensemble Recordings reached the CRC booth just in time for the inaugural day of the Society for Neuroscience Meeting, there were already several laboratories around the world applying new approaches and technologies to chroni- cally record the simultaneous extracellular activity of small populations of single neurons in behaving animals. Yet, in those days a considerable number of neuro- physiologists were still not convinced that such a technique would bring significant benefits to the field. Those opinions were bluntly articulated in two peculiar encounters that I experi- enced during that time. In the first, one of the leading neurophysiologists of our time stopped by my SFN poster display to, according to him, kindly remind me that there was “no future and no career in neural ensemble recordings.” Instead, he suggested that I should simply give up this “foolish stuff” and come back to the “church of single unit recording.” Just a couple of years later, this grandfatherly and well-intended advice was pointedly reinforced by a NIH reviewer who, in response to a grant that John Chapin and I had submitted, wondered in despair “…why one needs to use space-age tech- nology to study the brain?…” What a difference 10 years make! Today, a whole generation of young graduate students and postdocs dive into the great adventure of systems neurophysiology by learning how to record from tens or even hundreds of single neurons simultaneously as soon as they enter a lab. Without a doubt, this young and innovative branch of neuroscience has been totally embraced by the next generation of neuroscientists. Those closely following the development of the field have also seen that, whereas a decade ago, most if not all studies employing chronic multi-electrode recording were performed only on rats, today the same basic technique is employed for studies in different species of non-human primates, such as owls, squirrels and Rhesus mon- keys, as well as mice (wild and transgenic strains). In the last few years, the method has found its way to neurosurgical suites where it is now used during intra-operative recordings in Parkinsonian patients whose symptoms can only be improved through a deep brain stimulator. If these dramatic changes were not enough, multiple technological breakthroughs in the fabrication of high-density microelectrode arrays, surgical implantation tech- niques, and multi-channel signal processing have significantly expanded the yield and longevity of chronic neural ensemble recordings in behaving animals. These advances have allowed the establishment of direct real-time brain–machine inter- faces, a new experimental paradigm that holds a lot of promise, not only in terms of basic neurophysiology research, but also as the potential core for the development of a new generation of neuroprosthetic devices aimed at restoring mobility and com- munication skills in severely disabled patients. © 2008 by Taylor & Francis Group, LLC
  • 18.
    xiv Methods forNeural Ensemble Recordings, Second Edition Over the last 10 years, thanks to the generosity of some of the field’s pioneers who agreed to write a series of detailed methods-oriented chapters for the book, the first edition of Methods for Neural Ensemble Recordings has become a technical reference book for those who have taken the plunge into the field. As such, it has been a most rewarding experience for those of us who participated in the original volume to spot copies of the book, not nicely stored on book shelves, but worn and torn next to neurophysiological setups, spread all over the world. The second edition of this book comes to light during a very different time and environment in systems neuroscience. Partially liberated from having to maintain at any cost some of its classic dogmas, systems neurophysiology is once again thriving in both theory and practice. Furthermore, the ever growing consensus that distrib- uted populations of neurons, rather than single neurons, define the true functional unit of the central nervous system creates continuous demand for new technologies that can push multi-electrode recordings methods and neural ensemble physiology to new limits. Since 1994, when I joined the Department of Neurobiology at Duke, I have had the privilege to work and collaborate with a unique group of highly talented people who arrived at our laboratory with the goal of achieving just that: pushing the limits of our blossoming field and, during this process, shedding new light on the neuro- physiological principles that make ensembles of neurons perform the “business of the brain.” This book has been written by a group of these young collaborators, whose col- lective groundbreaking work covers most of the current areas of basic and clinical research that utilize multi-electrode recordings as the method of choice to probe brain circuits. The final product of this two year project, the second edition of Meth- ods in Neural Ensemble Physiology, is dedicated to all the high-school, undergradu- ate and graduate students, along with the postdoctoral fellows, technicians, research associates, collaborators, and visitors who, by joining our laboratory, made this jour- ney through distributed and dynamic brain circuits quite a thrill. I would like to thank Duke University, the Brain and Mind Institute at the Ecole Polytechnic Federale de Lausanne, the Edmond and Lily Safra International Insti- tute of Neuroscience of Natal, the National Institutes of Health, DARPA, and the Anne W. Deane Professorship Endowment for providing me with the time, space, support and resources for completing this and other projects that have considerably expanded the horizons of my life as a scientist in the last two years. I would like to sincerely thank our CRC editor, Barbara Norwitz, for waiting patiently for the completion of this project. I would also like to acknowledge the continuous support of my “older brother” and series co-editor, Sidney Simon, who invited me to edit the first edition of this book, 13 years ago, as my first assignment as a Duke assistant professor. We never stopped getting into trouble together after that. I also would like to thank my dear friend and collaborator, Susan Halkiotis, for dedicating 2 long years of her life to this project, and many more years to make sure that everything that crosses her desk, no matter how crazy or challenging the task, is taken care of with utmost care, perfection, and class. With fun on top! © 2008 by Taylor & Francis Group, LLC
  • 19.
    Preface xv Finally, Iwould like to thank Laura Oliveira for thirty years of trusting and supporting the pursuit of my dreams, far from home, wherever they may take us, no matter the odds or personal costs. I can barely wait to see what the next ten years will bring! Miguel A. L. Nicolelis Natal, Brazil © 2008 by Taylor & Francis Group, LLC
  • 20.
    xvii Editor Miguel A. L.Nicolelis, M.D., Ph.D., is the Anne W. Deane Professor of Neurosci- ence and professor in the departments of neurobiology, biomedical engineering, and psychology at Duke University, where he also serves as co-director of the Center for Neuroengineering. In addition, Dr. Nicolelis is scientific coordinator at the Edmond and Lily Safra International Institute of Neuroscience of Natal (ELS-IINN) in Natal, Brazil and taught neuroscience during 2006–2007 at École Polytechique Féderale de Lausanne in Lausanne, Switzerland. Dr. Nicolelis is a native of Sao Paulo, Brazil, where he received his M.D. and Ph.D. in neurophysiology from the University of Sao Paulo. He graduated from the University of Sao Paulo School of Medicine and was awarded the Oswaldo Cruz Prize for research, the highest honor awarded to a Brazilian medical student. After postdoctoral work at Hahnemann University, Dr. Nicolelis joined the faculty at Duke University in 1994. Dr. Nicolelis is interested in understanding the general computational principles underlying the dynamic interactions between populations of cortical and subcortical neurons involved in motor control and tactile perception. Although Dr. Nicolelis is best known for his study of brain–machine interfaces (BMI) for neuroprosthetics in human patients and nonhuman primates, he is also developing an integrative approach to studying neurological and psychiatric disorders by recording neuronal ensem- ble activity across different brain areas in genetically modified mice. Dr. Nicolelis believes that this approach will allow the integration of molecular, cellular, systems, and behavioral data in the same animal to produce a more complete understanding of the nature of the alterations associated with these disorders. Neuroscience laboratories in the U.S. and Europe have incorporated Dr. Nicole- lis’ experimental paradigm to study a variety of mammalian neuronal systems. His research has influenced basic and applied research in computer science, robotics, and biomedical engineering. This multidisciplinary approach to research has become widely recognized in the neuroscience community. Dr. Nicolelis was named as one of Scientific American’s Top 50 Technology Leaders in America in 2004 and has received a number of honors and awards includ- ing the Whitehead Scholar Award, DARPA Award for Sustained Excellence by a Performer; Ruth and A. Morris Williams, Jr., Faculty Research Prize; Whitehall Foundation Award; McDonnell-Pew Foundation Award; Duke University Thomas Langford Lectureship Award; the Ramon y Cajal Chair at the University of Mexico; and the Santiago Grisolia Chair at Catedra Santiago Grisolia. He has authored more than 120 manuscripts in scientific journals, and edited numerous books and special journal issues. He is frequently an invited speaker at scientific conferences and meet- ings throughout the world. © 2008 by Taylor & Francis Group, LLC
  • 21.
    xix Contributors Dr. Roy Crist Departmentof Neurobiology Duke University Durham, North Carolina Dr. Dragan Dimitrov Monterey, California Dr. Kafui Dzirasa Department of Neurobiology Duke University Durham, North Carolina Mr. Nathan Fitzsimmons Department of Neurobiology Duke University Durham, North Carolina Dr. Damien Gervasoni Faculté de Médecine Laënnec University Claude Bernard Lyon, France Mr. Timothy Hanson Duke University Durham, North Carolina Dr. Mikhail Lebedev Department of Neurobiology Duke University Durham, North Carolina Mr. Gary Lehew Department of Neurobiology Duke University Durham, North Carolina Dr. Shih-Chieh Lin Department of Neurobiology Duke University Durham, North Carolina Mr. Jim Meloy Department of Neurobiology Duke University Durham, North Carolina Dr. Simona Nemets Faculty of Mechanical Engineering Technion–Israel Institute of Technology Haifa, Israel Dr. Miguel A. L. Nicolelis Department of Neurobiology Duke University Durham, North Carolina Mr. Joseph E. O’Doherty Department of Biomedical Engineering Duke University Durham, North Carolina Dr. Laura M. Oliveira Department of Neurobiology Duke University Durham, North Carolina Dr. Albino J. Oliveira-Maia Department of Neurobiology Duke University Durham, North Carolina Dr. Parag Patil Department of Neurosurgery University of Michigan Hospitals Ann Arbor, Michgan Dr. Aaron Sandler Department of Neurobiology Duke University Durham, North Carolina © 2008 by Taylor & Francis Group, LLC
  • 22.
    xx Methods forNeural Ensemble Recordings, Second Edition Dr. Sidney A. Simon Department of Neurobiology Duke University Durham, North Carolina Dr. Eric Thomson Department of Neurobiology Duke University Durham, North Carolina Dr. Dennis Turner Division of Neurosurgery Duke Medical Center Durham, North Carolina Dr. Michael Wiest Department of Neurobiology Duke University Durham, North Carolina Dr. Miriam Zacksenhouse Faculty of Mechanical Engineering Technion–Israel Institute of Technology Haifa, Israel © 2008 by Taylor & Francis Group, LLC
  • 23.
    1 1 State-of-the-Art Microwire Array Designfor Chronic Neural Recordings in Behaving Animals Gary Lehew and Miguel A. L. Nicolelis INTRODUCTION Over the last two decades, many laboratories around the world have started to rely on microelectrode arrays formed by fine microwires, organized in different geometri- cal configurations, to chronically record the extracellular activity of populations of individual neurons in both anesthetized and behaving animals (Nicolelis et al. 1997, 2003; Lebedev et al. 2006; Verloop and Holsheimer 1984; Williams et al. 1999). As the field of chronic multielectrode recordings evolved, so did the designs of such microwire-based arrays. Indeed, during the last 13 years, our laboratory at the Duke University Cen- ter for Neuroengineering (DUCN) has specialized in producing a large variety of microwire array configurations that can now be utilized in a large variety of species (e.g., mice, rats, monkeys, and intraoperative human recordings). In particular, our efforts have been directed at producing arrays that can be utilized in experimental protocols demanding simultaneous recordings from large samples of single neurons CONTENTS Introduction................................................................................................................1 The DUCN Design and Fabrication Approach for Multisite, Chronic Neural Ensemble Recordings......................................................................................2 The Layered Approach ..............................................................................................4 Discretely Wired Approach .....................................................................................10 Building a Layered PCB Microwire Array..............................................................15 Building a Discretely Wired Array.......................................................................... 18 Conclusions..............................................................................................................20 References................................................................................................................20 © 2008 by Taylor & Francis Group, LLC
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    2 Methods forNeural Ensemble Recordings, Second Edition (e.g., 50 to 500), distributed across multiple cortical and subcortical brain sites in fully awake and behaving animals over long periods of time (months to years). The goal of this chapter, therefore, is to review the DUCN accumulated experience and describe its current state-of-the-art design and fabrication approach for producing high-quality microwire-based arrays for chronic, multisite, neural ensemble recordings. THE DUCN DESIGN AND FABRICATION APPROACH FOR MULTISITE, CHRONIC NEURAL ENSEMBLE RECORDINGS Over the years, the DUCN technical staff has abided by the principle that microelec- trode arrays for chronic single-neuron recordings have to be designed according to the main anatomical characteristics and contours of the specific brain area targeted in a particular experimental project. Thus, by collaborating with neurophysiologists specialized in working with different animal species, our technicians developed a large variety of microwire array (bundle) configurations for targeting different corti- cal and subcortical brain areas. This initiative proved to be essential for achieving optimal configurations that significantly increase the neuronal yield and longevity of our chronic recordings. In this design process, both 2-D and 3-D contour matching can be performed. In addition, different strategies have been implemented to fabri- cate adjustable microelectrodes. Such techniques allow microwire tips to be repeat- edly and selectively repositioned after surgery. Using this option, users can resample the neural area of interest once the physiological properties of a given populations of neurons have been accomplished. Among other advantages, such a data collection strategy increases considerably the overall neuronal sample obtained per recording site over the course of several weeks or months. In recent years, we have added stimu- lation electrodes, local ground reference electrodes, and cannula for drug injection to the basic configuration of such microwire arrays, increasing significantly the range of experimental manipulations that can be carried out once such multiple devices are chronically implanted in the animal’s brain. The basic configuration of the DUCN microelectrode arrays consist of insu- lated metallic conductors with an overall diameter in the range of 25 to 50 μm. The metallic conductor is selected based on corrosion resistance and degree of stiffness, because the wire is driven lengthwise. Tungsten or a stainless steel alloy is often used with an insulation coating of polyimide, Formvar, Isonel, or Teflon. The micro wires are processed to be as straight and stiff as possible. Using a variety of cutting devices, the tips of the wires can be cut blunt or at any desired angle (see Figure 1.1). The angle cut exposes more of the metallic conductor, which lowers the impedance of the electrode. This can reduce the ability to record from well-isolated single neurons. However, the sharply raked tip of the angle cut makes penetration through the brain tissues much easier and less traumatic, while producing less resistance as the length of the microwire travels through the tissue. At this level of design, multiple factors have to be balanced in order to produce an electrode that will yield viable long-term single unit recordings. For instance, not only must optimal impedance for single unit recordings be a consideration during electrode production, but care must be taken to produce an array that will penetrate © 2008 by Taylor & Francis Group, LLC
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    State-of-the-Art Microwire ArrayDesign for Chronic Neural Recordings 3 tissue easily during the surgical insertion process. Conversely, relying solely on a fine tip microelectrode array (Schwartz 2004) to facilitate tissue penetration may increase the chance of encapsulation of the electrode tips by glia and extracellular protein deposits and thus diminish the ability to record neural signals beyond a few days or weeks. The lesson here is simple: Finding the right compromise among a large number of variables (electrode material, electrode tip shape, cut angle, etc.) is the single most relevant challenge facing developers of microelectrode arrays for chronic recordings. In the DUCN fabrication routine, electrode tips can be left bare, or they can be electroplated, either selectively or as a group, to reduce impedance. An electrode impedance-measuring device such as the A-M systems model 2900 is a useful tool to quantify the variables in electrode impedance, such as the effect of electroplating on impedance. The spacing or separation of the recording electrode tips are in the range of 200 to 1000 μm, center to center, depending on the brain area targeted, the animal species utilized, and the experimental protocol. As a design principle, our electrode arrays are fabricated as part of a complete investigative system. Moreover, a series of features are incorporated into the design of the array to improve the efficiency of the manufacturing, testing, and surgical implantation processes. For instance, reinforced breakaway tabs are used to provide a standard clamp point for electrode holders of the stereotaxic equipment used dur- ing surgery as well as assembly and testing operations. At the end of the implantation surgery, the tabs are easily removed. Another set of small breakaway tabs are used to hold alignment in assembly jigs as the microwires are positioned and bonded in place on the printed circuit board (PCB). These same tabs are also used later in the manufacturing process as a palette for conductive paint as it is applied at the microwire to printed circuit trace junction. Guide holes in the PCB are also used to accept alignment pins when multiple arrays are combined or stacked. FIGURE 1.1 Two 35-μm electrode tips, top is angle cut, bottom is blunt cut. © 2008 by Taylor & Francis Group, LLC
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    4 Methods forNeural Ensemble Recordings, Second Edition THE LAYERED APPROACH The various methods employed to manage and organize microwires into arrays can be classified as either layered or discretely wired. In some cases both approaches are used to combine microelectrode arrays on to a single-output connector to streamline the interface to the subject under investigation. The layered approach involves designing PCBs for the purpose of adapting the spacing of the electrode array to the spacing of the chosen connector and providing a platform on which the array components can be mounted. As shown in Figure 1.2, the plated through-hole pads facilitate the mounting of a surface mount connector, and the traces connected to each pad are positioned to mechanically and electrically bond to microwires in an array. The PCB designs are very specific and dedicated to a limited number of applications because the thick- ness of the board determines row spacing, and the printed circuit trace determines the spacing of electrodes within each row. Alignment jigs, specific to each array pattern and variation, are fabricated for use in the assembly process. The layered approach requires a significant initial invest- ment, and minor changes can be costly because the boards and jigs are specifically dedicated to a given application. The benefit of this approach is that higher production volumes are achievable, meeting the demand with repeatable high quality devices. The majority of layered designs are arrays with two row groups, combined with other assemblies to form multiple rows, or used singularly as a two-row array. Varia- tions of this design exist in many forms. Figure 1.2 illustrates a design in which two separate arrays are built onto a single board and combined onto a single connector. This design allows for implanting into two regions of the brain in a single step. Combining the signals in one output connector helps to streamline the interface to the subject. A further variation of this layered design incorporates remote arrays or satellite arrays. These satellite arrays are connected to the central array via flexible cable assemblies. With this design, the electrode arrays are positioned independently but utilize a single output connector. FIGURE 1.2 Two 4 × 4 electrode arrays share a common 32-channel connector. © 2008 by Taylor & Francis Group, LLC
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    State-of-the-Art Microwire ArrayDesign for Chronic Neural Recordings 5 Figure 1.3 shows a layered array which contains the additional feature of being movable in depth after installation. Several millimeters of travel allow researchers to fine tune the electrode position and also extend the usefulness of the array. A further variation of this layered design incorporates remote arrays or satellite arrays. These satellite arrays are connected to the central array via flexible cable assemblies. With this design, the electrode arrays are positioned independently, but utilize a single- output connector. As shown in Figure 1.3, taken in midfabrication, a 2 × 8 electrode array is com- bined with a moveable 2 × 4 array and a remote or satellite 2 × 4 array (not pictured). All electrode wires in these arrays are 35 μm tungsten spaced at 250 μm. The spac- ing between the 2 × 8 and moveable 2 × 4 arrays is 300 μm. A 32-channel pream- plifier commonly referred to as a headstage, can be seen connected to the output connector of the electrode array. The headstage provides a buffer between the electrode and the recording equip- ment in terms of amplification, impedance matching, and filtering. It is sometimes necessary to fabricate electrodes with the headstage in place to allow for a work- ing clearance with electrode holders, stimulation connectors, moveable components, injection cannula, or other ancillary features. Figure 1.4 shows a custom-built electrode holder attached at the clamp points of a 2 × 16 array PCB. This board adapts the spacing of a 0.025 in. surface mount connector to the array pattern of 250 μm on centers. The board is FR4 composite material. The overall thickness dimension of the bare FR4, as seen at the right edge of the board, is a major determining factor in the spacing between rows of the array. The dark green area is a solder-masking agent for insulation. FIGURE 1.3 A 2 × 8 array and a movable 2 × 4 array. © 2008 by Taylor & Francis Group, LLC
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    6 Methods forNeural Ensemble Recordings, Second Edition The PCB shown in Figure 1.5 illustrates the manner in which microwires are mechanically bonded at the edge of the board and electrically bonded in a staggered formation along the gold-plated traces. The black area is a solder-mask agent used to provide electrical insulation. The tab to the right was used as a palette during the application of conductive paint and can be removed along with the tab on the left later in the fabrication process. The hole in each tab facilitates alignment if multiple assemblies are combined. The wires are 35 μm diameter tungsten with polyimide insulation. The insulation is removed at the silver paint locations to provide the best possible conductivity. FIGURE 1.5 A close-up view of microwire electrodes attached to a printed circuit board. FIGURE 1.4 Electrode holder clamping a 2 × 16 PCB prior to the installation of the surface- mount connector. © 2008 by Taylor & Francis Group, LLC
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    State-of-the-Art Microwire ArrayDesign for Chronic Neural Recordings 7 With the layered approach, the assemblies, which form two rows of electrodes, are combined and connectors are installed on each PCB or bridged between boards. As shown in Figure 1.6, the assemblies can be stacked to combine high-density arrays, or used singularly as a two-row array. Fixtures or jigs are designed and constructed for the purpose of achieving align- ment of multiple PCBs or, in many cases, soft moldable clay can be used to tempo- rarily hold alignment while a permanent bond is made with adhesive. The example shown in Figure 1.6 is a 96-channel, 6 × 16 electrode array with 35 μm wire spaced at 250 μm on center. This array consists of three 2 × 16 assem- blies aligned and bonded with a coiled common ground wire in the foreground. A single set of electrode holder tabs are left in place, which will be removed in the final stages of surgery. The electrode holder tabs are reinforced by plated cop- per cladding, which provides a robust mechanical and electrical ground connection between the electrode and stereotaxic equipment during surgery. The 16-channel 4 × 4 array with 50 μm wires spaced at 250 μm shown in Figure 1.7, is formed by joining two 2 × 4 assemblies. A double row surface-mount nano connector spans the pads of both boards. The two boards are separated using a shim material to obtain the correct spacing. The staggered microwire termination pattern, used to promote ease of assembly, is visible through the clear epoxy coating. The coating is applied for insulation and additional mechanical integrity. A single set of holder tabs is left in place, and will be removed after installation. Multiple array designs can sometimes be included on the same PCB in the inter- est of promoting economy by design. Shown in Figure 1.8, two spacing variations FIGURE 1.6 96-channel, 6 × 16 array with 35 μm electrodes spaced at 250 μm in mid- fabrication. © 2008 by Taylor & Francis Group, LLC
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    8 Methods forNeural Ensemble Recordings, Second Edition of layered 4 × 4 arrays occupy a common electrode-printed circuit assembly, joined at the holder tabs. Guide plates, visible in the center, are also included in the design and will be removed for use in fabrication jigs or included in arrays with ultralong FIGURE 1.7 A 4 × 4 array completed and ready for testing. FIGURE 1.8 Multiuse layered array PCB design being fitted with microwires in an assem- bly jig. © 2008 by Taylor & Francis Group, LLC
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    State-of-the-Art Microwire ArrayDesign for Chronic Neural Recordings 9 electrode wires that require floating guide plate supports. The board is shown mounted in an assembly jig and partially completed with one set of 35 μm electrode wires attached at corresponding traces of the board. Two sets of pads per trace are included to provide a universal mounting for the connector or for remote wiring of a satellite array. The jig is fabricated from a vast collection of the salvaged remains of routed FR4 panels in conjunction with the custom-drilled wire guides. The critical dimensional requirement of the bare FR4 thickness of a layered array is easily met by using the scrap materials from the production run. The supporting deep green-colored frame- work material in this case is FR4 material salvaged from other PCB production runs. The electrode featured in Figure 1.9 is a 32-channel, dual 4 × 4 layered array with electrodes spaced at 250 μm. The arrays are spaced at 2.5 mm. Guide tubes for drug injection cannula are fabricated from 23-gauge stainless steel tubing. The guide tubes are shown with inner pins fabricated from 0.013 in. stainless steel wire. The inner pins maintain a clear tract and are replaced with injection cannula when needed. The black material is a rubberized cyanoacrylate adhesive, which is used to pro- vide mechanical strength to the assembly, bonding the connector and guide tubes firmly to the PCB. The layered 128-channel array shown in Figure 1.10 illustrates how the separate assemblies are combined to provide a high density array. This array features 35 μm tungsten electrodes spaced at 250 μm and organized in eight rows of 16 electrodes. The electrode holder tabs have been removed on all assemblies except one, which will provide a clamp point for the entire assembly. The coiled ground wire is a com- mon ground connection to all four assemblies and is attached after the individual assemblies are combined. The connectors have been spaced to provide clearance to allow the four 32-channel headstages to simultaneously interface. This design features an offset between the array and connector to allow the array to be in close proximity to other arrays. This offset shifts the array footprint to coincide with the FIGURE 1.9 Dual 4 × 4 layered array with drug-injection cannula guides. © 2008 by Taylor & Francis Group, LLC
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    10 Methods forNeural Ensemble Recordings, Second Edition connector footprint along two planes in order to allow clearance for adjacent arrays or to shift the connector location to a more suitable location on the subject. DISCRETELY WIRED APPROACH The discretely wired approach is to form the array with straight microwires, mechan- ically bonding the microwires in the desired pattern and then routing the free ends of microwires to the connector. PCBs designed for this application are a tremendous benefit with regard to conformity in manufacturing, especially when small-scale, surface-mount nano-size connectors are used. This approach can be easily applied to achieve complex spacing and shapes, more so than with the layered approach. A single PCB design can be utilized for a multitude of array designs, because the printed circuit design is no longer directly related to the array pattern. In this case, the PCB provides two functions: to provide a structural component used to hold and position the assembly during manufacture and implantation, and to provide a mechanical and electrical bonding of the wire to the connector pin. With moveable designs, the PCB can also be used as the structural foundation for the moveable implements of the array. These boards also include features similar to boards used with layered arrays, such as tabs for clamping and holding the assem- bly, tabs for use as a palette, guide holes for alignment, and array patterns. Assembly jigs are designed and built specifically for each array pattern and can be partially or selectively loaded with microwire to yield variations within each FIGURE 1.10 128-channel 8 × 16 layered array. © 2008 by Taylor & Francis Group, LLC
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    State-of-the-Art Microwire ArrayDesign for Chronic Neural Recordings 11 pattern. These arrays can be used in various combinations of satellite configurations or as a singular device. The 64-channel electrode array shown in Figure 1.11 was built using a discrete method and sacrificial wire guide. The wire guide in this case is used as a part of the assembly jig and is incorporated as part of the electrode assembly. The microwires are 35 μm diameter, spaced on a 1000 μm grid. Three separate PCBs are used in this example, one for each of the two connectors, and the third is to establish and maintain the spacing of the electrodes forming the wire guide. Epoxy is used as a conformal coating to provide insulation and mechanical integrity. A 32-channel independently positionable array is shown in Figure 1.12, during the assembly process attached to the jig. The pins on the left are 0.010 in. diameter tungsten pins; the wires on the right are 35 μm tungsten electrodes with a spacing of 1000 μm on center. In this moveable design, the electrode wire is attached at the tip of the pin, and a coil is formed around the pin to store the additional wire needed in travel. The pin is friction fit into a molded grid and transfers motion to the electrode wire. Silicon gel is molded at the electrode-to-pin interface to provide protection and allow for motion within. The electrode signal wires can be seen at the top of the picture routed to a 32-channel dual row connector. The electrodes can be positioned independently for peak efficiency. Several meth- ods can be used to move the electrodes. A simple manual pin-pushing device can be built with an outer guide tube and inner adjustable depth stop. This can be adjusted with the aid of a micrometer for exact depth settings. A variation on approach is to modify a micrometer, adding the outer guide tube and inner push rod directly to the micrometer as shown in Figure 1.13. A more complex variation of this method is illustrated in Figure 1.14 and Figure 1.15. FIGURE 1.11 (See color insert following page 140.) 8 × 8 electrode array fabricated using the discrete-wired method. © 2008 by Taylor & Francis Group, LLC
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    12 Methods forNeural Ensemble Recordings, Second Edition The device shown in Figure 1.14 is a miniature linear actuator, motorized and encoded to provide accurate movements of the electrode friction pin-drive system. A touch-down sensor in the drive mechanism allows the device to be used in a hand- held manner. The operator positions the guide tube over the electrode drive pin, bot- toming the guide tube in the electrode shell. FIGURE 1.13 A micrometer modified to provide movements to a friction pin moveable electrode. FIGURE 1.12 A discrete-wired array still attached to the assembly jig. © 2008 by Taylor & Francis Group, LLC
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    State-of-the-Art Microwire ArrayDesign for Chronic Neural Recordings 13 Using the controller shown in Figure 1.15, the linear actuator is advanced until the inner drive rod contacts the electrode drive pin. Upon contact, the linear actuator is automatically stopped. The operator then enters the amount of travel and speed of travel desired at the electrode. A switch mounted on the actuator enables the control- ler to begin advancing the electrode at the discretion of the operator. The controller provides readout of the motion in microns, and has the capacity to store and retrieve position information, and communicate via RS-232. The electrode in Figure 1.16 is an example of a completed array that incorporates friction fit pins to independently position the electrodes. The main body of the array is fabricated by modifying a type F connector shell and forms the enclosure for the moveable components. The protective cap, threaded on the top of the body, has a FIGURE 1.14 A miniature linear actuator equipped to advance moveable electrodes. FIGURE 1.15 Hand-held electrode positioning system. © 2008 by Taylor & Francis Group, LLC
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    14 Methods forNeural Ensemble Recordings, Second Edition shaft that provides for stereotaxic attachment. This cap is replaced with a flat top cap after the array is firmly affixed in surgery. The remote-wired connector can be strategically positioned in surgery as needed to provide clearance. The electrode featured in Figure 1.17 is a 16-channel 2 × 8 moveable array using 15 μm tungsten wire. The electrode wires are drawn to a bundle and driven through a cast alignment block with a maximum deployment of 2.0 mm. Single-row connec- tors are used to connect to two 8-channel headstages. The electrode is shown prior FIGURE 1.17 A 16-channel moveable array used with dual 8-channel headstages. FIGURE 1.16 An independently positionable 32-channel array and stereotaxic-mounted cap. © 2008 by Taylor & Francis Group, LLC
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    State-of-the-Art Microwire ArrayDesign for Chronic Neural Recordings 15 to completion without a conformal coating. Adjustments are made by rotating the slotted head screw at the top. The 16-channel electrode in Figure 1.18 has the moveable drive assembly built into the body of the connector. The drive mechanism is a 0–80 threaded screw, which advances the electrodes at the rate of 317.5 μm/r. The maximum travel for this electrode is 2.0 mm. BUILDING A LAYERED PCB MICROWIRE ARRAY The first step in building the DUCN microwire array is to design and fabricate the PCBs. With the design parameters such as the array pattern and connector pad layout established, the PCB design will be largely governed by these decisions. A PCB software tool is used to create the artwork files that are then used to manufacture the printed circuits. The files can be e-mailed to a PCB manufacturing facility for use in automated processing machines. Boards can be created, tested, and shipped within a matter of days. It is also possible to have the connectors installed by automated process, if desired. With the PCBs in hand, the next step is to fabricate a jig to assist in aligning the microwires to the board for bonding. The basic concept is to have two identical panels, drilled on centers to the array pattern with a center-mounted slotted beam in place to hold the PCB in the correct position relative to the panels. The microwires are loaded into the holes, a PCB is positioned into the slot, and the process of bonding the wires to the board can begin. Several assemblies can be produced before the jig will require reloading. The panels are drilled with slightly larger size holes than the outside diameter of the insulated microwire. A typical hole size is 90 μm for a 50 μm outside diameter (OD) microwire. The hole size can be critical; if the hole does not provide enough clearance for the OD variations of the wire, the insulation could be damaged and the jig will be more difficult to operate. The panels are usually FIGURE 1.18 A 16-channel moveable array used with a 16-channel headstage. © 2008 by Taylor & Francis Group, LLC
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    16 Methods forNeural Ensemble Recordings, Second Edition thin sheets of Delrin, FR4 composite material, or paper. The jig components can be included in the layout of the PCB or be built by hand. If the panels will be pro- duced by hand, a precision drill press with X, Y, and Z feed will be needed. A Servo model 7060 drill press and a Newport model 200 XY table (shown in Figure 1.6) are capable of producing good quality jig guide plates. It is helpful to specify that the scrap materials from the routed panels be shipped with the PCBs because the exact thickness of the boards is needed to be matched on the jig for optimum alignment. This scrap material can be used to fabricate the entire jig, or combined with other production runs as needed. With the microwires attached, the assemblies can be tested, and the connector and ground wire installed. A conformal coating is applied as an insulation barrier. In Figure 1.19, a 17 × 2 array is being assembled with a jig. The microwire array spac- ing is established by drilling the pattern in Delrin. A split beam holds the printed circuit directly over the array pattern as the microwires are attached to the board in a staggered length formation. The jig is then flipped and the process is repeated on the other side of the PCB. With both sides connected the board is removed from the jig slightly and the microwires cut to free the assembly. Several boards can be processed before the jig must be reloaded with wire. Micro drill bits in sizes ranging from 50 μm to well over 100 μm can be used in the fabrication of the jig. These drill bits are used with the drill press and translation stage shown in Figure 1.20. The drill press has a standard Albrecht keystone chuck, but for precision drilling a 1-mm-diameter collet will provide the best accuracy and ease of use with micro drill bits. The flute length versus diameter of commercially FIGURE 1.19 Microwires are held in alignment with a jig. © 2008 by Taylor & Francis Group, LLC
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    State-of-the-Art Microwire ArrayDesign for Chronic Neural Recordings 17 available bits shown below indicates the maximum material thickness for a through- hole in the jig plate. The close-up view of a 90 μm drill bit in Figure 1.21 illustrates the flute length of 500 μm versus diameter. Due to their fragility, micro drill bits require very close tolerance drilling equipment to function. Drill Diameter (in microns) Flute Length (in microns) 50 400 60 400 70 400 80 500 90 500 100 700 110 700 FIGURE 1.20 Precision drilling equipment. © 2008 by Taylor & Francis Group, LLC
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    18 Methods forNeural Ensemble Recordings, Second Edition BUILDING A DISCRETELY WIRED ARRAY The first step in building this type of array is to fabricate a jig, which will be used to hold the microwires in alignment for a mechanical bonding process. The jig consists of two thin plates drilled to match the pattern of the array, mounted to a frame with approximately 25 mm space between. The microwires are loaded into the jig and mechanically bonded as an assembly. If the spacing is greater than 300 μm center to center, it can be helpful to use a third sacrificial plate to act as a carrier for the adhesive. This sacrificial plate helps to maintain the integrity of the pattern. The assembly can then be removed from the jig and mechanically bonded to a connector printed circuit assembly. The free ends of the microwires are then routed through plated through-holes, which are electrically connected to the conductor pins of the connector and mechanically bonded. The board is flipped to expose the microwire- plated hole junction. The microwires are trimmed to length, insulation is removed, and conductive paint or epoxy is applied at this junction. A conformal coating is applied to achieve an insulation barrier. The discrete wired jig shown in Figure 1.22 is designed to make a 4 × 8 array with electrode spacing of 500 μm and a row spacing of 800 μm. The jig is loaded with 35 μm wires, fixing to be bonded to form the array. Delrin blocks are used to help the adhesive span the distances between rows in order to maintain spacing integrity. The 16-channel adjustable electrode shown in Figure 1.23 features a 0.050 in. hex socket exposed at the top of the assembly, which is used to deploy the microwire electrodes from the cannula after being implanted over the target area. The adjustable mechanism is installed in an extruded brass square tube. The 0–80 threads of the mechanism provide incremental travel of 317.5 μm/r of the hex FIGURE 1.21 A 90 μm diameter drill bit typically used to drill microwire-guide arrays for jigs. © 2008 by Taylor & Francis Group, LLC
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    State-of-the-Art Microwire ArrayDesign for Chronic Neural Recordings 19 socket. A single reinforced tab provides a clamp point for the stereotaxic electrode holder. The signal wires are terminated remotely to the connector PCB adapter of another array. FIGURE 1.22 Discrete-type jig loaded with 35 μm wires. FIGURE 1.23 A 16-channel moveable array shown with electrodes partially deployed. © 2008 by Taylor & Francis Group, LLC
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    20 Methods forNeural Ensemble Recordings, Second Edition CONCLUSIONS The design of a large variety of microwire array (bundle) configurations has enabled our laboratory to perform chronic, multielectrode recordings in a variety of animal species, including wild type and transgenic mice, rats, and owl and rhesus monkeys. More recently, the same approach has been translated into a new methodology for monitoring brain activity in patients subjected to neurosurgical procedures. This new generation of microelectrode arrays has pushed the limits of systems neurophysiology and allowed, for the first time, the simultaneous monitoring of the activity of hundreds of individual neurons, distributed across multiple, inter- connected cortical and subcortical structures that define particular neural circuits (e.g., somatosensory, motor, gustatory, etc.) for long periods of time (weeks to years, depending on the animal species and experimental protocol) in behaving animals. REFERENCES Lebedev, M.A. and Nicolelis, M.A.L. (2006). Brain machine interfaces: Past, present and future. Trends Neurosci 29: 536–546. Nicolelis, M.A.L., Ghazanfar, A.A., Faggin, B., Votaw, S., and Oliveira, L.M.O. (1997). Reconstructing the engram: simultaneous, multiple site, many single neuron record- ings. Neuron 18: 529–537. Schwartz, A.B. (2004). Cortical Neural Prosthetics. Annu Rev Neurosci 27: 487–507. Verloop, A.J. and Holsheimer, J. (1984). A simple method for the construction of electrode arrays. J Neurosci Meth 11: 173–178. Williams, J.C., Renmaker, R.L., and Kipke, D.R. (1999). Long-term neural recording char- acteristics of wire microelectrode arrays implanted in cerebral cortex. Brain Res Pro- tocols 4: 303–313. © 2008 by Taylor & Francis Group, LLC
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    21 2 Surgical Techniquesfor Chronic Implantation of Microwire Arrays in Rodents and Primates Laura M. O. Oliveira and Dragan Dimitrov And still we could never suppose that fortune were to be so friendly to us, such as to allow us to be perhaps the first in handling, as it were, the electricity concealed in nerves, in extracting it from nerves, and, in some way, in putting it under everyone’s eyes. Luigi Galvani, 1791 CONTENTS Introduction..............................................................................................................22 Differences Between Rodents and Primates Pertinent to Surgical Technique........23 Surgical Techniques for Rodents .............................................................................24 Preoperative Supplies and Room Preparation ..............................................24 Preoperative Animal Preparation.................................................................25 Anesthesia Techniques and Intraoperative Monitoring................................26 Electrode Specific for Rodents .....................................................................27 Implantation Techniques...............................................................................28 Brain Electrode Arrays......................................................................28 EMG Electrode Surgery .................................................................... 31 Postoperative Care........................................................................................32 Surgical Techniques for Primates............................................................................32 Preoperative Supplies and Room Preparation ..............................................32 Preoperative Animal Preparation.................................................................34 Anesthesia Techniques and Intraoperative Monitoring................................34 Electrode Specific for Primates....................................................................35 Implantation Techniques...............................................................................35 Exposure ............................................................................................35 Cortical Localization .........................................................................36 Drilling of Craniotomies....................................................................36 Opening of the Brain Coverings........................................................40 © 2008 by Taylor & Francis Group, LLC
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    22 Methods forNeural Ensemble Recordings, Second Edition INTRODUCTION The study of electrophysiology started with the work of Luigi Galvani (1737–1798), who was the first to provide evidence for the electrical nature of “the mysterious fluid” (at the time referred to as “animal spirits”). Galvani’s nephew, Giovanni Aldini (1762-1834), continued this line of inquiry in 1803, using Galvani’s and Alessandro Volta’s (bimetallic electricity) principles together, despite the fact that Volta did not believe in animal electricity. Carlo Mateucci (1818–1868) in Bologna and Emil Du Bois-Reymond (1818–1896) in Berlin described the phenomenon called “negative variation” when a galvanometer showed an unexpected decrease in current intensity during muscle contraction. The study of the electrophysiology of the nervous system began when Julius Berstein (1839–1917) proposed his theory of the nerve impulse as a wave of negativity (membrane theory of the nerve tissue). Later, using a galvanom- eter with one electrode in the gray matter and one on the skull surface (or electrodes in different points of the external surface of the brain), Richard Caton (1842–1926) recorded a feeble current in the brain. In 1870, for the first time, Gustav Fritsch (1838–1927) and Eduard Hitzig (1838–1907) inserted an electrode in the dura of a dog brain and stimulated the motor area, generating movement in the contralateral side of the animal’s body (Niedermeyer 1993; Piccolino 1998). The work of these and many other scientists marked the beginning of the study of the electrophysiology of the nervous system, opening doors to the possibility of stimulating different areas of the brain through electrical current and subsequently recording the brain electrical activity. Improvements in electrode manufacturing, the advent of modern acquisition equipment, and better surgical and asepsis techniques have provided us the ability to chronically implant multiple electrodes simultaneously in several areas of the brain in the same animal (Nicolelis, Baccala et al., 1995) and to study the interactions of populations of neurons (Nicolelis, Fanselow et al., 1997; Ghazanfar, Stambaugh et al., 2000). Upon the animal’s recovery from surgery, we have been able to record simultaneously from different brain areas of mice (Costa, Cohen et al., 2004), rats (Faggin, Nguyen et al., 1997; Ghazanfar and Nicolelis 1997; Nicolelis, Ghazanfar et al., 1997) and nonhuman primate brains (Nicolelis, Stam- baugh et al., 1999; Nicolelis, Dimitrov et al., 2003) for long periods of time, from a couple of months in rodents (Ghazanfar and Nicolelis 1997; Nicolelis, Ghazanfar et al., 1997) to up to years in non-human primates, such as owl monkeys (Nicolelis, Ghazanfar et al., 1998) and Rhesus monkeys (Nicolelis, Dimitrov et al., 2003). These recordings are carried out under several different experimental condi- tions and behavior tasks (Kralik, Dimitrov et al., 2001; Nicolelis and Ribeiro 2002). With chronically implanted multiple electrodes, it is also possible to record different Electrode Lowering............................................................................ 41 Skull and Wound Closure.................................................................. 41 Postoperative Care........................................................................................43 Conclusions and Future Directions..........................................................................44 Acknowledgments....................................................................................................45 References................................................................................................................45 © 2008 by Taylor & Francis Group, LLC
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    Surgical Techniques forChronic Implantation of Microwire Arrays 23 layers of the same area of the brain (Chapin and Lin 1984) and study spatiotem- poral response of many neurons (Nicolelis and Chapin 1994). Microcannulae can be attached to the electrode arrays and are used to inject drugs in the areas of the implant during chronic experimental recordings (Shuler, Krupa et al., 2002). Chronically implanted electrodes offer unparalleled advantages for correlating neuronal activity and animal behavior. In our lab, these techniques were developed in rodents and later adapted to primates. Over the last several years, there have been significant strides in making rodent implantations more reliable, faster, and easier. We have identified and resolved many of the issues that now permit larger neuronal yields that last longer. As a consequence of continuous improvement in techniques, the length of time required for surgery has been reduced. At the same time, over the last 14 years, we have developed a surgical technique adapted to the unique features of primates. This has made primate implantations routine and reproducible. Here, we will describe detailed technical aspects of the current surgical implantation approach used in our laboratory at the Duke University Center for Neuroengineering (DUCN). Such a surgical protocol has evolved and benefited from almost two decades of accu- mulated experience on chronic multielectrode neural recordings (Nicolelis, Stam- baugh et al., 1999; Nicolelis, Dimitrov et al., 2003, Kralik, Dimitrov et al., 2001; Nicolelis and Ribeiro 2002). DIFFERENCES BETWEEN RODENTS AND PRIMATES PERTINENT TO SURGICAL TECHNIQUE The success in obtaining recordings from chronically implanted electrodes and how long they last depends fundamentally on the quality of the surgical implantation tech- nique. The ability to open small craniotomies, only large enough to fit the electrode array with minimal bleeding from the bone and from the meninges in very small animals such as mice and rats, requires practicing the techniques several times before attempting to gather good data using the implant of electrode arrays. This is especially true in cortex areas, which are close to the dura and are very sensitive to lesions. The surgical technique for implantation that we follow in our lab is similar for all species, but each species has its own peculiarities. For instance, the mouse head is very small and the skull is thin, requiring a delicate technique. Mouse surgeries require very small screws, drill bits, and custom-designed electrode arrays. Because of the small head size and bone thickness, inserting more than two small arrays of electrodes and more than two fixation screws per animal is not recommended. Compared to mice, the rat’s head is bigger and the bone is stronger, increasing the possibility of using bigger electrode arrays and reaching more cortical and subcorti- cal areas. Because rats are stronger than mice, they can tolerate larger amounts of the acrylate used to secure the electrodes in place, which increases the number of arrays that can be inserted in the brain (Faggin, Nguyen et al., 1997; Nicolelis and Ribeiro 2002). Cortical and subcortical localization in rodents is based on commercially available stereotactic atlases. Theoretically, implanting electrodes into rodents and primates should be very similar; however, in practice they are vastly different. For investigators accustomed © 2008 by Taylor & Francis Group, LLC
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    24 Methods forNeural Ensemble Recordings, Second Edition to the hardiness of rodents and trained in rodent implantation techniques, per- forming similar procedures on primates can seem overwhelming. For instance, the commitment of time, personnel, and lab resources is much greater for primate surgery. Furthermore, from the point of anesthesia, primates require much more atten- tion during surgery. Whereas rodents require only monitoring of a few physiologi- cal parameters and infrequent injections, maintaining a primate under anesthesia is much more labor intensive. Thus, for our primate surgeries, one member of the surgery team is dedicated to monitoring the animal throughout. This higher level of anesthesia technique is akin to pediatric anesthesia and requires specific equipment, planning, and personnel. Obvious differences in anatomical details include a thicker skull, thicker brain coverings, and a better-developed subdural space, all of which also influence the definition of the optimal surgical strategy for chronic multi-electrode implantation in nonhuman primates. The thicker skull requires forethought in terms of the appro- priate electrode length. The brain coverings including the dura and pia are better developed, more variable in their thickness and at least the dura requires wide open- ing with microsurgical instruments for electrode penetration. We have found that the pia and arachnoid layers of primates are more variable, often tougher, and more prone to dimpling than rodents, sometimes necessitating microsurgical opening, as well as paying careful attention to be sure penetration has occurred. The primate brain is more prone to problems with swelling and retraction due to fluctuations in CO2 levels in the blood that are primarily affected by ventilation of the animals’ lungs. Plans must be in place to deal with these issues intraoperatively. Overall, we have also observed that intraoperative neural recordings are more difficult in primates than they are in rodents. Much more attention must be paid to noise detection and reduction, especially because many more electrical devices nec- essary for anesthesia are involved. Primates are more prone to infection than rodents and require more attention to sterile technique throughout, with sterilization of all the instruments, the use of sterile gloves and gowns, and draping of the surgical field. For those unaccustomed to working in a sterile environment for long periods of time, this can present unfore- seen challenges and cost significant amounts of time. In summary, primate implantations require a team approach. It often takes sev- eral days to weeks to prepare and coordinate one’s lab prior to performing primate surgery, underscoring the critical importance of preoperative planning. SURGICAL TECHNIQUES FOR RODENTS PREOPERATIVE SUPPLIES AND ROOM PREPARATION Typically, our rodent surgeries are performed on a surgery table in a regular room in the laboratory or in a designated surgical room. The table is kept clean and unclut- tered and has the stereotaxic apparatus already installed (for rats we use the cat and small primates stereotaxic apparatus with a rat adaptor, and for mice, we use the stereotaxic apparatus for small rodents, both from David Kopf Instruments, Tujunga, © 2008 by Taylor & Francis Group, LLC
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    Surgical Techniques forChronic Implantation of Microwire Arrays 25 California). Between the two stereotaxic bars, a rectangular platform made of plexi- glass is glued to a height adjustable stage where a warm pad is placed. This apparatus is used for the anesthetized animal for the duration of the surgery and immedi- ate postoperative recovery. Other essential pieces of equipment include a binocu- lar surgical microscope, a dental drill already installed in a source of compressed air, an amplifier connected to an audio monitor, an oscilloscope, and a microposi- tioner. The table, the stereotaxic apparatus and the plexiglass platform are cleaned before the surgery with 70% alcohol or Asseptiwipes (wipes moistened in a solu- tion of N-alkyl(68%C12, 32%C14) dimethyl ethyl benzyl ammonium chloride 0.125%, N-alkyl(60%C14, 30%C16, 5%C12, 5%C14) dimethyl ethyl benzyl ammonium chloride 0.125%, isopropyl alcohol 14.850%, and other ingredients. All our surgeries follow the National Institute of Health (NIH) and Associa- tion for Assessment and Accreditation of Laboratory Animal Care (AAALAC) guidelines for rodents that are expected to recover from anesthesia. These guide- lines include appropriate preoperative and postoperative care of the animals, asepsis (sterilization of the surgical tools, use of sterile gloves, mask, head covers, clean lab coat, and aseptic procedures), gentle tissue handling, minimal dissection of tissues, effective hemostasis, and correct use of suture materials and patterns when stitches are necessary (National Research Council 1996; Baumans, Remie et al., 2001). The surgical instruments (basically, four small hemostats, scalpel handle, Freer periosteal elevator, scissors, micro scissors, regular and micro tweezers, dental drill handpiece, special screw drivers, stainless steel wire cutters, and micro cup curette) and supplies (gauze, cotton-tip applicators, kimwipes, drill bits, small stainless steel screws, silicone cups for dental acrylic, and beakers) are sterilized by autoclave. All supplies that cannot be exposed to heat and humidity are sterilized by ethylene oxide gas (extrafine point markers, plastic rulers, electrode holders, electrodes, etc.). The microwire arrays (see chapter 1) are made of very delicate materials and cannot be sterilized by autoclave. Instead, these must be sterilized by ethylene oxide gas. The electrodes should be carefully packed in such a way that they are not dam- aged during handling, transportation to and from the sterilization facility, or by personnel helping in the surgery. Because of the length of these surgeries, all the equipment used during surgery should be routinely tested the day before the animal is anesthetized. To avoid unnecessary delays during surgery, it is imperative that all materials, equipment, and drugs for the surgery are readily available and in good working condition. A copy of the brain atlas appropriate to the animal undergoing implantation surgery is necessary to help in the location of the areas to be implanted. It is advisable to complete these steps the day before the surgery; however, they can be completed the day of the surgery prior to anesthetizing and preparing the animal. PREOPERATIVE ANIMAL PREPARATION Typically, all animals selected to undergo chronic implantation of multielectrode arrays have several days to acclimate in the new laboratory facility before surgery. Often, these animals are subjected to weeks of behavioral training before they are implanted with arrays of electrodes. Only animals in good health are subjected to the surgery. © 2008 by Taylor & Francis Group, LLC
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    26 Methods forNeural Ensemble Recordings, Second Edition In general, we work with Long–Evans adult rats, males (in general 300–350 g) or females (in general 250–300 g). However, depending on the studies we can use younger or older animals. Mice are chosen according to their genotype since our laboratory currently utilizes a variety of mutant animals and their wild-type litter- mates as control (Costa, Lin et al., 2006; Dzirasa, Ribeiro et al., 2006). Because rodents have a high metabolism and are not at risk for vomiting and aspirating, food is not withheld until up to 2 h before the surgery. Plans for anesthesia, surgery, and determining the coordinates of the areas to be implanted must be carefully studied ahead of time. This will help decrease the time of the surgery for electrode-array implantation. A record sheet should be started for the procedure prior to beginning anesthesia. This not only meets the requirements of AAALAC, NIH, and Institutional Animal Care and Use Committee (IACUCs), but also provides a record of the procedure, the implanted areas and their coordinates, the depth of the electrode for each array, and the position of the ground wires related to the position of the connector of the array. ANESTHESIA TECHNIQUES AND INTRAOPERATIVE MONITORING Following guidelines from Duke University’s IACUC and the advice of Duke’s veteri- nary staff, we use two anesthesia regimens for rat surgeries and one for mice surgeries. For rats: Pentobarbital IP Ketamine associated with Xylazine IM For mice: Ketamine IM associated with Xylazine IM Pentobarbital has a good hypnotic effect, but analgesia is obtained only when using high doses of the drug, which can cause cardiovascular and respiratory depression. The advantage of using Pentobarbital is that the effect of the anesthesia lasts longer than the Ketamine/Xylazine combination. Ketamine is a dissociative anesthetic that provides light surgical anesthesia with a short duration. When combined with sedatives or tranquilizers, the quality of the anesthesia is highly improved and the duration of the effect is increased. In general, in our surgeries supplemental doses of Pentobarbital are required every 2 h and Ket- amine every 1½ h. The supplemental doses for both drugs are from ⅓ to ½ of the original dose, but Xylazine is not supplemented unless the surgery lasts longer than 7–8 hours, which is very uncommon in rodent surgeries. For both anesthesia regimens (rats and mice), anesthetic induction is carried out in a chamber with a mixture of 5% Isoflurane and O2. In our experience, a good induction with Isoflurane helps the selected anesthesia regimen last longer, decreas- ing the need for frequent injections of supplemental doses of the anesthetic. Once the animal is deeply anesthetized with Isoflurane, an injection of the selected drug is administered (Duke University DLAR 1995; National Research Council 1996; Hellebrekers and Booij 2001). Following the injection of anesthesia, the animals © 2008 by Taylor & Francis Group, LLC
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    Surgical Techniques forChronic Implantation of Microwire Arrays 27 may receive a dose of Atropine SQ to prevent or treat excessive secretion of the airways and to improve cardiac function. This is important especially with the use of Pentobarbital. For rats, if the surgery is expected to last for a longer period, small doses of sterile saline can be injected SQ, and IP injections of 10% dextrose solution will help to maintain hydration and blood glucose level during the surgery. In general, mice surgeries are much shorter than rat surgeries due to the small number of screws and arrays that are implanted, and extra volume or dextrose injections are not necessary. Once the animal is deeply anesthetized, it is placed in an area for preparation for surgery, apart from the operating theater. The animal’s head is shaved with small clippers, from the area just above the eyes to the back of the head and from ear to ear. If electromyography (EMG) electrode implants are planned, the skin on top of the target muscles is also shaved at this point. After the shaved hair is cleaned, the animal is transported to the surgery table and placed on the platform with a thermal pad to maintain the animal’s body temperature throughout the surgery. If an electrical pad is used, the pad should be covered to avoid direct contact with the animal in order to prevent burns to the animal’s skin in case the pad overheats, and a rectal temperature probe should be inserted. A good option is the Deltaphase® ther- mal pads (Braintree Scientific, Braintree, Massachusetts) that maintain temperature around 37ºC for about 5–6 h. This type of pad must be monitored and changed when the temperature decreases. After the animal is on a warm pad, it is mounted on the stereotaxic apparatus, using proper ear bars for rodents, placing the mouth and nose piece, and leaving the position of the head fixed until the top of the head is parallel to a flat surface. Once the rat head is secure, the sterile pack can be opened and the sterile materi- als should be kept in a sterile field or container. The skin of the head is cleaned three times using Iodophor (iodine soaps) or Chlorhexidine followed by alcohol 70%. The animal’s eyes are covered with an ophthalmic ointment to prevent corneal lesions since the animal loses the blinking reflex with the anesthesia. ELECTRODE SPECIFIC FOR RODENTS The arrays of electrodes to be implanted are made in-house and vary in number and distribution of electrodes. For the past nineteen years, our experiments have utilized arrays or bundles made of thin microwires (see chapter 1 for details). Some arrays can be made to reach more than one area of the brain. In general, for rat surgeries, the arrays are comprised of 32 electrodes (4 × 8), but they can also be made up of either 16 (2 × 8) or 64 (8 × 8) electrodes. The tips are cut either blunt or sharp. When cut sharp, the electrodes may be able to penetrate the brain without dissecting the dura. In this case, the length of each electrode may be slightly different (see Figure 2.1). If necessary, a little cut on the dura with a bent fine needle may help relieve the tension of the meninges and facilitate the implantation of the electrodes. Obviously, the length of the microwires varies for each surgery and depends on the brain area to be implanted. © 2008 by Taylor & Francis Group, LLC
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    28 Methods forNeural Ensemble Recordings, Second Edition IMPLANTATION TECHNIQUES Brain Electrode Arrays The length of the surgery for the implantation of microwire arrays varies from ani- mal to animal. It also depends on the number of arrays to be implanted, the location of such brain regions, and the experience of the surgeon. In general, the minimum time is about 3–4 h if only one or two arrays are to be implanted. Each additional array can add about 45–60 min to the total surgical time. It is essential not to rush the implantation procedure. In our accumulated experience, gentle and slow penetration of the brain tissue yields the best long-term results. During surgery, the status of anesthesia is checked periodically by pinching the inter-digit membrane of the hind paw or pinching the tails of the animal. Supplemen- tal doses of anesthesia are given as needed. The areas of the brain to be implanted also vary for each study and may vary even in the same study because it is not possible to reach all areas of interest for each animal, especially in mice. The surgical procedures described below have been very successful in the past. Following this approach has allowed our experimental animals to tolerate these implants very well, survive without any postsurgical com- plication, and provide high-quality recordings for months after the surgery. After the animal is anesthetized, mounted to the stereotax, and cleaned, the surgical team suits up for surgery in a lab coat or gown, mask, sterile gloves, and head cap. Before starting surgery, the status of the anesthesia is checked as described above. To alleviate pain during the incision, an injection of lidocaine 1% or Bipu- vocaine can be given under the skin at the site of incision. An incision is made on the midline of the scalp, from just above the eyes to the back of the head, and the skin is propped open. The borders of the skin are held open with small hemostats, and bleeding, which in general is very small, is cleaned with gauze or cotton-tip FIGURE 2.1 Sharp electrodes penetrating the dura in a rat surgery. Photo by Edgard Morya. © 2008 by Taylor & Francis Group, LLC
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