Functional Neural Transplantation IV Translation to Clinical Application, Part B [1st Edition] 9780128138793

Table of contents :
Content:
Series PagePage ii
CopyrightPage iv
ContributorsPages v-ix
PrefacePages xxv-xxviiS. Dunnett, A. Björklund
Chapter 1 - Stem cell transplantation for spinal cord injury repairPages 1-32Paul Lu
Chapter 2 - Plasticity and regeneration in the injured spinal cord after cell transplantation therapyPages 33-56Satoshi Nori, Masaya Nakamura, Hideyuki Okano
Chapter 3 - Transplantation of GABAergic interneurons for cell-based therapyPages 57-85Julien Spatazza, Walter R. Mancia Leon, Arturo Alvarez-Buylla
Chapter 4 - Rebuilding CNS inhibitory circuits to control chronic neuropathic pain and itchPages 87-105Joao M. Braz, Alex Etlin, Dina Juarez-Salinas, Ida J. Llewellyn-Smith, Allan I. Basbaum
Chapter 5 - From transplanting Schwann cells in experimental rat spinal cord injury to their transplantation into human injured spinal cord in clinical trialsPages 107-133Mary B. Bunge, Paula V. Monje, Aisha Khan, Patrick M. Wood
Chapter 6 - Recruitment of endogenous CNS stem cells for regeneration in demyelinating diseasePages 135-163Natalia A. Murphy, Robin J.M. Franklin
Chapter 7 - Progenitor cell-based treatment of glial diseasePages 165-189Steven A. Goldman
Chapter 8 - Pluripotent stem cells and their utility in treating photoreceptor degenerationsPages 191-223Nozie D. Aghaizu, Kamil Kruczek, Anai Gonzalez-Cordero, Robin R. Ali, Rachael A. Pearson
Chapter 9 - Stem cell-derived retinal pigment epithelium transplantation for treatment of retinal diseasePages 225-244Britta Nommiste, Kate Fynes, Victoria E. Tovell, Conor Ramsden, Lyndon da Cruz, Peter Coffey
Chapter 10 - Transplantation of reprogrammed neurons for improved recovery after strokePages 245-263Zaal Kokaia, Daniel Tornero, Olle Lindvall
Combined IndexPages 265-287

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Serial Editor

Vincent Walsh Institute of Cognitive Neuroscience University College London 17 Queen Square London WC1N 3AR UK

Editorial Board Mark Bear, Cambridge, USA. Medicine & Translational Neuroscience Hamed Ekhtiari, Tehran, Iran. Addiction Hajime Hirase, Wako, Japan. Neuronal Microcircuitry Freda Miller, Toronto, Canada. Developmental Neurobiology Shane O’Mara, Dublin, Ireland. Systems Neuroscience Susan Rossell, Swinburne, Australia. Clinical Psychology & Neuropsychiatry Nathalie Rouach, Paris, France. Neuroglia Barbara Sahakian, Cambridge, UK. Cognition & Neuroethics Bettina Studer, Dusseldorf, Germany. Neurorehabilitation Xiao-Jing Wang, New York, USA. Computational Neuroscience

Academic Press is an imprint of Elsevier 50 Hampshire Street, 5th Floor, Cambridge, MA 02139, United States 525 B Street, Suite 1800, San Diego, CA 92101–4495, United States The Boulevard, Langford Lane, Kidlington, Oxford OX5 1GB, United Kingdom 125 London Wall, London, EC2Y 5AS, United Kingdom First edition 2017 Copyright # 2017 Elsevier B.V. All rights reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopying, recording, or any information storage and retrieval system, without permission in writing from the publisher. Details on how to seek permission, further information about the Publisher’s permissions policies and our arrangements with organizations such as the Copyright Clearance Center and the Copyright Licensing Agency, can be found at our website: www.elsevier.com/permissions. This book and the individual contributions contained in it are protected under copyright by the Publisher (other than as may be noted herein). Notices Knowledge and best practice in this field are constantly changing. As new research and experience broaden our understanding, changes in research methods, professional practices, or medical treatment may become necessary. Practitioners and researchers must always rely on their own experience and knowledge in evaluating and using any information, methods, compounds, or experiments described herein. In using such information or methods they should be mindful of their own safety and the safety of others, including parties for whom they have a professional responsibility. To the fullest extent of the law, neither the Publisher nor the authors, contributors, or editors, assume any liability for any injury and/or damage to persons or property as a matter of products liability, negligence or otherwise, or from any use or operation of any methods, products, instructions, or ideas contained in the material herein. ISBN: 978-0-12-813879-3 ISSN: 0079-6123 For information on all Academic Press publications visit our website at https://www.elsevier.com/books-and-journals

Publisher: Zoe Kruze Acquisition Editor: Kirsten Shankland Editorial Project Manager: Ana Claudia Garcia Production Project Manager: Magesh Kumar Mahalingam Cover Designer: Mark Rogers Typeset by SPi Global, India

Contributors Nozie D. Aghaizu UCL Institute of Ophthalmology, London, United Kingdom Robin R. Ali UCL Institute of Ophthalmology, London, United Kingdom Arturo Alvarez-Buylla The Eli and Edythe Broad Center of Regeneration Medicine and Stem Cell Research, University of California, San Francisco, San Francisco, CA, United States Anne-Catherine Bachoud-L
evi Assistance Publique-H^ opitaux de Paris, Centre de R
ef
erence Maladie de Huntington, Service de Neurologie, H^ opital Henri Mondor-Albert Chenevier; INSERM U955, Equipe 01 Neuropsychologie Interventionnelle; Universit
e Paris Est, Facult
e de M
edecine, Cr
eteil; D
epartement d’Etudes Cognitives, Ecole Normale Sup
erieure, PSL* Research University, Paris, France Roger A. Barker Wallenberg Neuroscience Center, Lund University, Lund, Sweden; Wellcome Trust-MRC Cambridge Stem Cell Institute and John van Geest Centre for Brain Repair, University of Cambridge, Cambridge, United Kingdom Allan I. Basbaum University of California—San Francisco, San Francisco, CA, United States Anders Bj€orklund Wallenberg Neuroscience Center, Lund University, Lund, Sweden Joao M. Braz University of California—San Francisco, San Francisco, CA, United States Vania Broccoli San Raffaele Scientific Institute; CNR-Institute of Neuroscience, Milan, Italy Mary B. Bunge The Miami Project to Cure Paralysis; Department of Cell Biology; Department of Neurological Surgery, University of Miami Leonard M. Miller School of Medicine, Miami, FL, United States Monica Busse Centre for Trials Research, College of Biomedical & Life Sciences, Cardiff University, Cardiff, United Kingdom Melissa K. Carpenter Carpenter Group Consulting, Seattle, WA, United States Susanne Clinch Centre for Trials Research, College of Biomedical & Life Sciences; Brain Repair Group, School of Biosciences, Cardiff University, Cardiff, United Kingdom

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Contributors

Peter Coffey Institute of Ophthalmology; NIHR Biomedical Research Centre, Moorfields Eye Hospital NHS Foundation Trust, London, United Kingdom; Center for Stem Cell Biology and Engineering, Neuroscience Research Institute, University of California, Santa Barbara, Santa Barbara, CA, United States Lyndon da Cruz Institute of Ophthalmology; NIHR Biomedical Research Centre; Moorfields Eye Hospital NHS Foundation Trust, London, United Kingdom Mate D. D€ obr€ ossy University Freiburg—Medical Centre, Freiburg, Germany Elsa Diguet CEA, DSV, Molecular Imaging Research Center (MIRCen), Fontenay-aux-Roses; Institut de Recherches Servier, Neuropsychiatry Unit, Croissy sur Seine, France Stephen B. Dunnett Brain Repair Group, School of Biosciences, Cardiff University, Cardiff, United Kingdom Alex Etlin University of California—San Francisco, San Francisco, CA, United States Robin J.M. Franklin Wellcome Trust-Medical Research Council Cambridge Stem Cell Institute; University of Cambridge, Cambridge, United Kingdom Kate Fynes Institute of Ophthalmology, London, United Kingdom Steven A. Goldman Center for Neuroscience, University of Copenhagen Faculty of Health and Medical Sciences, Copenhagen, Denmark; Center for Translational Neuromedicine, University of Rochester Medical Center, Rochester, NY, United States Anai Gonzalez-Cordero UCL Institute of Ophthalmology, London, United Kingdom Genevieve Gowing Board of Governors Regenerative Medicine Institute, Cedars-Sinai Medical Center; Cedars-Sinai Medical Center, Los Angeles, CA, United States Magdalena G€ otz Physiological Genomics, Biomedical Center; SYNERGY, Excellence Cluster of Systems Neurology, Biomedical Center, Ludwig-Maximilians University Munich, Planegg; Institute of Stem Cell Research, Helmholtz Center Munich, Munich, Germany

Contributors

Philippe Hantraye CEA, DSV, Molecular Imaging Research Center (MIRCen); CNRS, CEA, Paris-Sud University, University of Paris-Saclay, Neurodegenerative Diseases Laboratory (UMR9199), Fontenay-aux-Roses, France Dina Juarez-Salinas University of California—San Francisco, San Francisco, CA, United States Aisha Khan The Interdisciplinary Stem Cell Institute, University of Miami Leonard M. Miller School of Medicine, Miami, FL, United States Agnete Kirkeby Wallenberg Neuroscience Center; Lund Stem Cell Center, Lund University, Lund, Sweden Zaal Kokaia Laboratory of Stem Cells and Restorative Neurology, Lund Stem Cell Center, Lund, Sweden Kamil Kruczek UCL Institute of Ophthalmology, London, United Kingdom Tilo Kunath MRC Centre for Regenerative Medicine, Institute for Stem Cell Research, School of Biological Sciences, The University of Edinburgh, Edinburgh, United Kingdom Meng Li Cardiff University Neuroscience and Mental Health Research Institute, School of Medicine; Cardiff University School of Biosciences, Cardiff, United Kingdom Olle Lindvall Laboratory of Stem Cells and Restorative Neurology, Lund Stem Cell Center, Lund, Sweden Ida J. Llewellyn-Smith Cardiovascular Medicine, Human Physiology and Centre for Neuroscience, Flinders University, Bedford Park, SA, Australia Paul Lu Veterans Administration San Diego Healthcare System; University of California, San Diego, CA, United States Walter R. Mancia Leon The Eli and Edythe Broad Center of Regeneration Medicine and Stem Cell Research, University of California, San Francisco, San Francisco, CA, United States Paula V. Monje The Miami Project to Cure Paralysis; Department of Neurological Surgery, University of Miami Leonard M. Miller School of Medicine, Miami, FL, United States

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Contributors

Natalia A. Murphy Wellcome Trust-Medical Research Council Cambridge Stem Cell Institute; University of Cambridge, Cambridge, United Kingdom Masaya Nakamura Keio University School of Medicine, Tokyo, Japan Ammar Natalwala MRC Centre for Regenerative Medicine, Institute for Stem Cell Research, School of Biological Sciences, The University of Edinburgh; Translational Neurosurgery Group, Western General Hospital, Crewe Road South, Edinburgh, United Kingdom Britta Nommiste Institute of Ophthalmology, London, United Kingdom Satoshi Nori Keio University School of Medicine, Tokyo, Japan Hideyuki Okano Keio University School of Medicine, Tokyo, Japan Malin Parmar Wallenberg Neuroscience Center; Lund Stem Cell Center, Lund University, Lund, Sweden Rachael A. Pearson UCL Institute of Ophthalmology, London, United Kingdom Conor Ramsden Institute of Ophthalmology; NIHR Biomedical Research Centre, Moorfields Eye Hospital NHS Foundation Trust, London, United Kingdom Anne E. Rosser Cardiff University Neuroscience and Mental Health Research Institute, School of Medicine; Cardiff University School of Biosciences, Cardiff, United Kingdom Julien Spatazza The Eli and Edythe Broad Center of Regeneration Medicine and Stem Cell Research, University of California, San Francisco, San Francisco, CA, United States Lorenz Studer The Center for Stem Cell Biology, Developmental Biology Program, Memorial Sloan Kettering Cancer Center, New York, NY, United States Clive N. Svendsen Board of Governors Regenerative Medicine Institute, Cedars-Sinai Medical Center; Cedars-Sinai Medical Center, Los Angeles, CA, United States Soshana Svendsen Board of Governors Regenerative Medicine Institute, Cedars-Sinai Medical Center; Cedars-Sinai Medical Center, Los Angeles, CA, United States

Contributors

Jun Takahashi Center for iPS Cell Research and Application, Kyoto University, Kyoto, Japan Daniel Tornero Laboratory of Stem Cells and Restorative Neurology, Lund Stem Cell Center, Lund, Sweden Olof Torper Physiological Genomics, Biomedical Center; SYNERGY, Excellence Cluster of Systems Neurology, Biomedical Center, Ludwig-Maximilians University Munich, Planegg; Institute of Stem Cell Research, Helmholtz Center Munich, Munich, Germany Victoria E. Tovell Institute of Ophthalmology, London, United Kingdom Nadja Van Camp CEA, DSV, Molecular Imaging Research Center (MIRCen); CNRS, CEA, Paris-Sud University, University of Paris-Saclay, Neurodegenerative Diseases Laboratory (UMR9199), Fontenay-aux-Roses, France Patrick M. Wood The Miami Project to Cure Paralysis; Department of Neurological Surgery, University of Miami Leonard M. Miller School of Medicine, Miami, FL, United States

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Preface The purpose of this book is to provide a series of critical reviews on recent progress in the field of neural transplantation with a focus on functional aspects of cell transplantation for brain repair. After 35 years of basic research and initial early attempts at clinical application the last 5–7 years has witnessed a major step change in both the range of clinical applications being considered and the refinement of alternative strategies and cell sources under consideration. In particular, recent advances in the sophistication of stem cell biology and differentiation are providing realistic prospects of new cell therapies for neurodegenerative diseases and injury reaching clinical reality. This volume represents the fourth in a series of review volumes that we have edited at 6- to 10-year intervals following the emergence of the field of neurotransplantation in the brain over four decades. All but the first volume (Functional Neural Transplantation, Raven Press, 1994) have been published as volumes in Elsevier’s Progress in Brain Research series (Functional Neural Transplantation II, PBR vol. 127, 2000, and Functional Neural Transplantation III parts 1 and 2, PBR vols. 200 and 201, 2011). The field of functional neural transplantation has developed dramatically over that period, from the early experimental forays establishing viable techniques for cell survival in the adult rodent brain in the 1970s, the first clear examples of functional transplantation in rodent models of neurodegeneration in the 1980s, early attempts at applying fetal cell transplantation in patients in the 1990s, to the present mature phase of sophisticated translational and clinical trial research. This includes an explosion of new opportunities arising from different, more flexible and powerful cell sources, notably pluripotent stem cells. The authors are among the current world leaders in this rapidly developing field. Consequently, the resulting volumes, like the previous three volumes in the series, provides authoritative reviews of the current state of play in this fast emerging and high profile field of cell replacement therapies. In particular, harnessing the power and potential of stem cells holds promise to transform the range of clinical conditions that are now within realistic prospect of “brain repair.” As we are passing the 20th anniversary of Jamie Thomson’s first human embryonic stem cell line, and the 10th anniversary of Yamanaka and Takahashi’s determination of specific factors for reprogramming of adult human cells to a pluripotent stem cell state, there has been remarkable progress in the understanding of the principles for differentiation of precursor and postmitotic cells of diverse neuronal and glial phenotypes types, which are viable for transplantation, replacing our previous exclusive dependence on fetal cells for cell replacement therapy. Advances in developmental biology are now, for the first time, allowing effective and efficient control of cell fate specification from pluripotent cell sources, accompanied by dramatic technical advances in standardization, reliability, and quality control necessary to underpin clinically acceptable 21st century medicinal products. Indeed, there are now major efforts worldwide to address the critical safety issues that need to be resolved prior to any clinical application.

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The chapters have been commissioned to provide authoritative scholarly representation of the current frontline of translational research across a broad range of applications of emerging cell transplantation technologies in regenerative medicine, applied to the brain and spinal cord, with a focus on neurodegenerative diseases and injury. Although the book was planned as a single volume, the detailed accounts provided by the authors have required publication in two parts, Part A and Part B, as consecutive volumes in the Progress in Brain Research Series. Nevertheless, we provide a unified contents list and index in order to retain the integrity of the whole. In the first chapter of part A, as editors, we discuss the principles of functional repair and recovery and outline our present understanding of the various mechanisms by which a cellular transplant may impact on the functional state of the host, be it an experimental animal or a patient. Understanding alternative mechanisms of recovery is an essential first stage in defining the task requirements for any new cell therapy and sets the demands of what is required to achieve functional efficacy in each specific disease indication. This is followed by a series of chapters that focus on the current and emerging states of the new technologies for ex vivo and in vivo reprogramming of neurons to different cell fates, characterization of alternative delivery vectors and stem cell lines, and the demands of regulatory compliance for taking any new therapy to the clinic. In the subsequent chapters the major clinical indications for which cell therapies are currently under investigation are reviewed. This starts in part A with two wellestablished basal ganglia disorders, Parkinson’s disease and Huntington’s disease, that have been the first to undergo clinical translation. Here, the experimental strategy is explicitly reparative, i.e., to replace lost neurons with new cells of the same neuronal phenotype with the capacity to integrate functionally into host brain, repair damage circuits, and restore lost function. In each case, we review the status of ongoing clinical trials using existing cell sources as well as new strategies for differentiating clinical grade cell therapy products from human stem cell sources. Cell therapy trials in these diseases have set the benchmarks for clinical trial design, including patient assessment strategies, motor and cognitive testing, and sophisticated imaging. They have also paved the way for the development and implementation of rehabilitation strategies, not just to promote strategies for compensation but with the capacity to impact graft survival, differentiation, and circuit repair. Next, in Part B, follows a series of chapters addressing the recent and significant advances in spinal cord injury and repair. The long-distance regeneration of precisely targeted axon pathways represents a field where sustained careful experimental analysis over many years is now getting close to implementation in clinical trials, building, in particular, on the capacity of central and peripheral glial cells to support damaged axons, promote regeneration, and enhance circuit repair, as well as neuroprotection and trophic support. New sources of neuroglia derived from pluripotent stem cells provide interesting alternatives to conventional oligodendrocyte and Schwann cells, but face the same biological, regulatory, and safety challenges as in applications in the brain.

Preface

A major area of advance for stem cell-based therapies has emerged over the last decade in the eye. Remarkable progress has been achieved in functional cell replacement in experimental animals using retinal pigment epithelium (RPE) and photoreceptor cells, and authentic cells of both types have been effectively, generated from pluripotent stem cells. There is a realistic prospect that the first demonstrably effective stem cell therapy in neurological medicine will emerge from ongoing trials using RPE cell replacement to treat blindness due to age-related macular degeneration. Finally, to conclude Part B, we consider another major cause of neurological illness worldwide, i.e., the prospects for cell-based treatment of stroke and related neurovascular diseases. This is an area more than any where there have been spurious claims worldwide of novel stem cell therapies. This has raised false hopes for vulnerable patients of magic cures in the absence of any credible scientific foundation and promoted a form of “stem cell tourism” that threatens to overshadow the important progress that is taking place in academic research centers worldwide. The final chapter provides a critical review of what is now possible to achieve and assesses the realistic hopes of true progress in this widespread disease. Functional Neural Transplantation IV, Part A and Part B, is targeted to inform active basic and translational research scientists working in the field, although we anticipate that the topic will attract a much broader readership among neuroscientists and clinicians interested in the current state of stem cell biology, cell therapy, and regenerative medicine. Readers can expect a balanced and realistic assessment of what is now possible, and the likely advances in the near future years, along with a critical appraisal of the limits of current therapies, and a debunking of the unsubstantiated claims. S. Dunnett, Cardiff, UK A. Bj€orklund, Lund, Sweden 13th February 2017

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CHAPTER

Stem cell transplantation for spinal cord injury repair

1 Paul Lu1

Veterans Administration San Diego Healthcare System, San Diego, CA, United States University of California, San Diego, CA, United States 1 Corresponding author: Tel.: +1-858-534-8857; Fax: +1-858-534-5220, e-mail address: [email protected]

Abstract Stem cells, especially neural stem cells (NSCs), are a very attractive cell source for potential reconstruction of injured spinal cord though either neuroprotection, neural regeneration, remyelination, replacement of lost neural cells, or reconnection of disrupted axons. The later have great potential since recent studies demonstrate long-distance growth and connectivity of axons derived from transplanted NSCs after spinal cord injury (SCI). In addition, transplanted NSCs constitute a permissive environment for host axonal regeneration and serve as new targets for host axonal connection. This reciprocal connection between grafted neurons and host neurons constitutes a neuronal relay formation that could restore functional connectivity after SCI.

Keywords Stem cells, Neural stem cells, Spinal cord injury, Axonal growth, Synaptic connection, Neuronal relay

1 INTRODUCTION Spinal cord injury (SCI) not only damages gray matter neurons that control local motor and sensory function but also the white matter axons that carry signal to and from brain with the rest of the body. Once injured, it is irreversible, since the injured adult central nervous system (CNS) is unable to spontaneously regenerate, often resulting in permanent functional deficits below the level of injury (Ramon, 1928). Therefore, reconstruction of the injured spinal cord and improvement of motor, sensory, and autonomic function is the ultimate goal of SCI research. One strategy for reconstruction of injured spinal cord is stem cell transplantation. Just like the typical clinical stem cell transplant that often refers to bone marrow Progress in Brain Research, Volume 231, ISSN 0079-6123, http://dx.doi.org/10.1016/bs.pbr.2016.11.012 © 2017 Elsevier B.V. All rights reserved.

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CHAPTER 1 Stem cell transplantation for spinal cord injury repair

transplant for replacement of diseased bone marrow, the main purpose of stem cell transplantation for SCI is to replace lost neurons and glia. In addition, reconnection of disrupted axons is vitally important since the major deficiency of SCI results from these injured axons rather than from local neuronal and glial loss. According to these criteria, only neural stem cells (NSCs) can fulfill these purposes since they can differentiate into neurons and glia for replacement of lost neurons and glia, and differentiated neurons can serve as interneurons or relay neurons that could reconnect the disrupted axons (Bonner and Steward, 2015; Fig. 1). In addition, NSCs can be differentiated specifically into oligodendrocyte progenitor cells (OPCs) for the purpose of remyelination after SCI (Watson and Yeung, 2011). Just like any other stem cells, NSCs may exhibit neuroprotection effects if transplanted at an acute stage after injury. Besides NSCs, bone marrow stromal cells or mesenchymal stem cells (MSCs) are a very popular stem cell source for neuroprotection after SCI due to their easy isolation from adult bone marrow and current usage in clinics (Dasari et al., 2014). Traditionally, NSCs and/or neural progenitor cells (NPCs) are mostly isolated from embryonic or fetal CNS (Reynolds and Weiss, 1992; Temple, 1989). NSCs can also be generated from embryonic stem (ES) cells (McDonald et al., 1999). The recent rapid development of induced pluripotent stem cells (iPSCs) technology greatly expands the stem cell source for generation of NSCs, (Takahashi et al., 2007; Tsuji et al., 2010). Similarly, forced expression of certain transcription factors can directly convert fetal or adult somatic cells into neurons (induced neurons, iNs) or NSCs (iNSCs; Hong et al., 2014; Kozhich et al., 2013; Ring et al., 2012; Vierbuchen et al., 2010). The advantages to use NSCs derived from iPSCs or iNs

FIG. 1 Diagram of a neuronal relay formation by transplanted NSCs after SCI. Descending supraspinal axons (red) regenerate into and make synaptic connections with grafted neurons (green) in the lesion site. Grafted neurons extend their axons (green) into caudal host spinal cord and make synaptic connection with host neurons (red). Illustrated by Audelia Arasheben.

2 MSCs for SCI repair

and iNSCs are that it not only eliminates the ethical concern of embryo or fetal tissue usage but also greatly reduces the risk of immune rejection since these NSCs can be generated in a patient-matched manner. In this chapter, I will briefly review transplantation of MSCs for SCI repair first, then, systematically review NSC transplantation for SCI repair, including the sources and origin of NSCs, and their differentiation potential in the model of SCI. Then I will focus on integration of grafted neurons in the injured spinal cord, with an emphasis on axonal growth and connectivity from transplanted neurons, and host axonal regeneration and connectivity with transplanted neurons. Finally, I will discuss the functional outcomes and the future challenges of NSC transplant for SCI repair.

2 MSCs FOR SCI REPAIR 2.1 SOURCES OF MSCs MSCs can be isolated from many different adult tissues, such as bone marrow, adipose tissue, inner organs and blood vessels, and from those fetal life-support systems, such as amniotic fluid and membrane, umbilical cord, or placenta (Foudah et al., 2014; Lopez-Verrilli et al., 2016). The diverse resources definitely increase supply of MSCs for transplantation. However, MSCs from different sources may have different features that greatly influence neural repair (Lopez-Verrilli et al., 2016).

2.2 MECHANISM OF MSC TRANSPLANT FOR SCI REPAIR One key feature of SCI is inflammation created by invasion of immune system cells, such as white blood cells. Studies show that MSCs possess immunomodulation and neuroprotection features, by attenuating reactive astrocytes and activated macrophages/microglia and increasing sparing of white matter (Abrams et al., 2009; Osaka et al., 2010; Ribeiro et al., 2015; Seo et al., 2011). This immunomodulation effect can be achieved even by intravenous (Osaka et al., 2010; Seo et al., 2011) or intrathecal (Cizkova et al., 2011) delivery of MSCs. In addition, MSCs can form a permissive cellular substrate for promotion of host axonal growth. A previous study shows that transplanted MSCs form guiding strands that attract host supraspinal motor axonal regeneration (Hofstetter et al., 2002). The neural regenerative effect could result from expression of neurotrophic factors by MSCs (Ribeiro et al., 2015). Indeed, our lab genetically modified MSCs to overexpress brain-derived neurotrophic factor (BDNF) that promotes the extent of host axonal regeneration into the MSC graft after SCI (Lu et al., 2005). The combination of BDNF-expressing MSC grafts with additional BDNF viral delivery below injury and cAMP administration into brainstem reticular motor neurons enables reticulospinal motor axons to regenerate not only into the MSC graft but also to cross the glial scar and regenerate into distal spinal cord beyond the injury site (Lu et al., 2012a).

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CHAPTER 1 Stem cell transplantation for spinal cord injury repair

2.3 NEURONAL DIFFERENTIATION POTENTIAL OF MSCs MSCs are typical adult multipotent stem cells that can differentiate into mesodermal lineages, such as osteoblasts, chondrocytes, myocytes, and adipocytes (Raff, 2003). Some early studies, however, report transdifferentiation of bone marrow-derived cells or MSCs into neuron-like cells, both in vitro (Woodbury et al., 2000) and in vivo (Brazelton et al., 2000; Mezey et al., 2000), even before reprogramming adult somatic cells into iPSCs (Takahashi et al., 2007). The theory underlying this transdifferentiation phenomenon is adult stem cell plasticity (Raff, 2003) or the existence of the so-called multipotent adult progenitor cells (MAPCs) in MSCs (Jiang et al., 2002). However, later studies demonstrate that transdifferentiation is an artifact of cell fusion in vivo (Terada et al., 2002; Ying et al., 2002). Similarly, our study along with others demonstrates artifact of in vitro differentiation of MSCs into neurons (Bertani et al., 2005; Lu et al., 2004; Neuhuber et al., 2004). It is interesting that the original in vitro neuronal induction of MSCs was achieved by simple chemicals or small molecules that could reprogram MSCs into neural cells. However, we knew very little about reprogramming genes, especially transcription factors and their related signal transductions, at that early time. The introduction of the iPSC technology allowed one to reprogram or convert any somatic cells, including MSCs, into pluripotent stem cells (Barzilay et al., 2009). iPSCs can be routinely differentiated into NSCs (Yuan et al., 2011). Recently, Cheng et al. (2014b) were the first to report the generation of chemically induced neural stem cells (ciNSCs) from several somatic cells by a cocktail of three small molecules. In addition, two studies have reported the successful generation of mouse and human neurons from dermal fibroblasts by an all-chemical approach (Hu et al., 2015; Li et al., 2015). It is quite possible that these small molecules could reprogram MSCs into NSCs or neurons for repair of SCI.

2.4 CLINICAL TRIALS OF MSCs FOR SCI There are many clinical trials of bone marrow-derived stem cell transplant for SCI probably due to the clinical usage of them to treat other diseases and their easy accessibility (Bhanot et al., 2011; Cheng et al., 2014a,b; Kakabadze et al., 2016; Oh et al., 2016). Bone marrow-derived stem cells were either directly injected into SCI site or adjacent to the injury site (Bhanot et al., 2011; Cheng et al., 2014a,b) or infused intrathecally (Kakabadze et al., 2016; Oh et al., 2016). Most studies claim some functional benefits although there are no controls in most of these trials. Therefore, the beneficial effect of MSC transplant on SCI patients is questionable. Some studies do report limited efficacy (Oh et al., 2016). It is encouraged that new clinical trials should follow the clinical guidelines for SCI for measurement of the safety or efficacy (Steeves et al., 2007).

3 NSCs for SCI repair

3 NSCs FOR SCI REPAIR 3.1 SOURCES OF NSCs AND THEIR DIFFERENTIATION POTENTIAL 3.1.1 NSCs from embryonic/fetal CNS tissue Fetal CNS tissue, especially rat embryonic spinal cord containing NSCs and NPCs, has been transplanted into SCI sites for therapeutic application long before NSCs had been identified and isolated in culture (Reier et al., 1986). The advantage of fetal CNS tissue for grafting is that its integrity and differentiation closely resemble the in vivo neural cell development. The disadvantage is that it requires fresh tissue for each graft and solid tissue grafts must be placed into the lesion site through a large opening of an otherwise closed contusion injury. In addition, the orientation of fetal tissue pieces is random and their integration with host may be variable. Alternatively, fetal/embryonic CNS tissue can be freshly dissociated as cell suspensions and directly injected into the site of the SCI (Giovanini et al., 1997; Lu et al., 2012b; Wictorin and Bj€ orklund, 1992). Both fresh fetal CNS tissue and freshly dissociated fetal CNS cells have the potential for full neural cell differentiation and can differentiate into both neurons and glia after transplantation into the injured spinal cord (Giovanini et al., 1997; Kadoya et al., 2016; Lepore and Fischer, 2005; Lu et al., 2012b; Wictorin and Bj€ orklund, 1992). We quantified transplanted neural cells differentiated from freshly isolated embryonic day 14 spinal cord and found approximately 28% grafted cells became NeuN-positive neurons, 27% APC + oligodendrocytes, and 16% GFAP + astrocytes (Lu et al., 2012b). NSCs were first isolated and cultured from rat embryonic forebrain in 1989 (Temple, 1989). These cells are multipotent and responsive to epidermal growth factor and basic fibroblast growth factor and therefore defined as CNS stem cells (Kitchens et al., 1994; Reynolds and Weiss, 1992, 1996). Late, multipotent NSCs (also termed as neuroepithelial cells) can be isolated as early as embryonic day 10.5 in the caudal neural tube in rats (Kalyani et al., 1997) and they can generate restricted precursor cells, named neuronal-restricted precursors (NRPs) and glialrestricted precursors (GRPs; Mayer-Proschel et al., 1997), which can fully differentiate into mature neuron and glial cells in vitro, respectively. The ability to culture NSCs in vitro has opened a new door for stem cell transplantation study since they can be expanded to produce a large quantity of cells and can be manipulated to differentiate into certain phenotypes of neurons or glia, or to express a transferred gene (Lian Jin et al., 2011; Pallini et al., 2005; Rietze and Reynolds, 2006). However, whether cultured NSCs can maintain their stem cell characters to differentiate into both neurons and glia both in vitro and in vivo, especially in vivo in injured spinal cord, is a critical question. An early study by Cao et al. (2001) showed that transplantation of cultured NSCs derived from rat fetal CNS tissue into either intact or injured spinal cord differentiates only into glial lineages, although cultured NSCs can differentiate into both neurons and glia in vitro. Similarly, engrafted NSCs derived from mouse embryo differentiate exclusively toward the astrocytic phenotype without neuronal or oligodendrocytic phenotypes or remain

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undifferentiated in the adult injured mouse spinal cord (Pallini et al., 2005). The loss of neuronal differentiation from cultured rodent NSCs transplanted into the injured spinal cord in vivo was confirmed by Setoguchi et al. (2004) and Abematsu et al. (2010), unless blocked by genetic modification of NSCs expressing noggin or by the administration of valproic acid for 7 days postgrafting. A potential mechanism is the presence of inflammatory cytokines that restrict NSC differentiation after grafting into injured spinal cord (Ricci-Vitiani et al., 2006). Downregulation of necessary genes for neuronal differentiation in the adult injured spinal cord environment might be another cause, since enhancement of transcriptional activity by valproic acid (Abematsu et al., 2010) or by inhibition of bone morphogenetic protein that plays important roles in neural development (Setoguchi et al., 2004) promotes neuronal differentiation in vivo. The differentiation of multipotent NSCs is influenced by the injured spinal cord environment, as described earlier. Does this injured environment also affect differentiation of more restricted precursor cells, such as NRPs? Cao et al. (2002) demonstrated that neuronal differentiation is inhibited in the injured spinal cord, although NRPs can differentiate into different types of neurons in the normal adult rat spinal cord. The authors suggest that manipulation of the injured spinal cord environment may be necessary to facilitate neuronal differentiation. In a different study, Han et al. (2002) culture NRPs for a short period of time over 1–2 weeks and transplanted them into intact spinal cord. Grafted NRPs differentiate exclusively into mature neurons and integrate well with the host neural cells. In addition, grafting of a mixture of NRPs and GRPs into a SCI enables neuronal differentiation and maturation. Emerging glial cells derived from GRPs appear to support survival and neuronal differentiation of NRPs within the nonneurogenic injured spinal cord environment (Lepore and Fischer, 2005). NSCs can also be isolated and cultured from human fetal CNS tissue. Uchida et al. (2000) first isolated human NSCs from fetal brain tissue and purified them using fluorescence-activated cell sorting of the human cell surface marker CD133. Human NSCs were cultured as neurospheres and transplanted into rostral and caudal parenchymal tissue surrounding a thoracic contusion injury site in immunodeficient mice (Cummings et al., 2005). In contrast to cultured rodent NSC transplants, which differentiate exclusively or mostly into glia, a quarter of human cells become Tuj1-positive neurons that form synaptic connections with host neurons 4 months postgrafting. The remaining grafted human cells differentiate into CC1/ APC-positive oligodendrocytes. Similar results were reported by the same group when human fetal CNS-derived NSCs were transplanted into chronic SCI sites in mice (Salazar et al., 2010). Besides the NSCs derived from human fetal brain, a different human NSC line (566RSC) was generated from fetal spinal cord and cultured as a monolayer for multiple passages (Yan et al., 2007). The majority (75%) of human NSCs differentiate into Tuj1-positive neurons when grafted into intact immunodeficient rat spinal cord. When the same human NSC line was grafted into a T3 transection site in our previous study (Lu et al., 2012a,b), the grafted cells completely fill lesion cavity after 2 months and the majority (57%) of cells differentiate into

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NeuN-positive neurons. In addition, Iwanami et al. (2005) transplanted human NSCs in a primate model of SCI and showed differentiation of neurons and glia and functional improvement. Such studies of human NSC transplants in animal model of SCI bring them close to clinical application. Indeed, the first two studies described earlier lead to clinical trials of human NSC transplantation for SCI (Curtis et al., 2016; Tsukamoto et al., 2013). In addition, a Korean group recently conducted a clinical trial by transplantation of human fetal brain-derived NSCs into the injured spinal cord after traumatic cervical SCI (Shin et al., 2015).

3.1.2 NSCs from adult CNS tissue Adult CNS tissue contains NSCs in neurogenic regions such as the hippocampus and the subventricular zone (Palmer et al., 1995). Adult NSCs can be isolated from spinal cord and ventricular neuroaxis (Weiss et al., 1996) and then cultured and expanded to obtain a large quantity of cells for transplantation studies. Although adult NSCs can differentiate into both neurons and glia in vitro, grafted adult NSCs in the injured spinal cord differentiate exclusively (Cao et al., 2001; Karimi-Abdolrezaee et al., 2006; Mothe et al., 2008; Vroemen et al., 2003) or mostly (Hofstetter et al., 2005; Wilcox et al., 2014) into glia. However, adult NSCs can differentiate into neurons when grafted into neurogenic regions, such as the hippocampus, indicating that local environments dictate the fate of stem cells (Song et al., 2002). The predominantly glial differentiation of adult NSCs limits their therapeutical potential as a neuronal replacement therapy. However, adult NSCs can function for neuroprotection, remyelination, or the promotion of host axonal regeneration after SCI (Karimi-Abdolrezaee et al., 2006; Mothe et al., 2008; Wilcox et al., 2014).

3.1.3 NSCs from ES cells Embryonic stem cells (ESCs) are derived from the inner cell mass of the early blastocyst and are able both to proliferate for a long period of time and to differentiate into varieties of cell types, including neural cells (Lukovic et al., 2012; Nistor et al., 2011). There are many lines of ESC lines now available from mouse and human, but very few from rats (Li et al., 2008), although rats serve as a very popular animal model for SCI. McDonald et al. (1999) first transplanted neural-differentiated mouse ESCs into rat SCI site and showed differentiation of a small number of neurons and glia. The neurogenic potential of mouse ESC-derived NSCs in injured spinal cord is controversial. While many studies show differentiation of neurons after transplantation of mouse ESC-derived NSCs or neural progenitors into mouse SCI sites (Kumagai et al., 2009; McCreedy et al., 2014), some studies show mouse ESC-derived NSCs primarily differentiate into glia when transplanted into the injured or demylinated spinal cord (Rowland et al., 2011; Salewski et al., 2015a), which is similar to that seen using cultured rodent primary NSCs from fetal CNS tissue, as discussed earlier. The McDonald group specifically generated OPCs from mouse ESCs and used them for remyelination studies (Liu et al., 2000).

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Besides oligodendrocytes, ESCs can be induced to differentiate into regionalspecific neural progenitors, including spinal cord motor neuron progenitors (MNPs; Erceg et al., 2008; Hu and Zhang, 2009; Nistor et al., 2011). When human MNPs are transplanted into cervical SCI sites in adult rats, host neuronal survival and axon growth are enhanced, indicating a neuroprotective effect (Rossi et al., 2010). When human MNPs are transplanted into thoracic transection site in combination with OPCs, grafted human cells become oligodendrocytes and motor neurons that exhibit electrophysiological activity and promote behavioral improvement (Erceg et al., 2010). However, there is no evidence showing that grafted human motor neurons extend their axons into peripheral nerve and innervate muscles (Erceg et al., 2010; Rossi et al., 2010). Thus, it is still a challenge to generate functional neurons from transplanted NSCs or progenitors in the injured adult spinal cord. We have transplanted human ESCs-derived NSCs into injured spinal cord and a large proportion of them differentiate into neurons, including some serotonergic (5-HT) neurons, which integrate into host CNS circuitry (Lu et al., 2012b; Zhao et al., 2013). It is interesting to characterize the detailed phenotypes of these human neurons and their specific connection with host neurons.

3.1.4 NSCs from iPSCs NSCs can be generated from iPSCs using a similar procedure as for ESCs (Chambers et al., 2009). Differentiated NSCs can be enriched or purified using fluorescenceactivated cell sorting (FACS) of NSC surface markers (Kozhich et al., 2013; Yuan et al., 2011). Before the enrichment procedure, Tsuji et al. (2010) identified “safe” clones of NSCs derived from murine iPSCs by examining their teratomaforming activity after transplantation into the NOD/SCID mouse brain. When safe iPSCs-derived NSCs are transplanted into the injured spinal cord in mice, they differentiated into both glial and neuronal lineages without teratoma formation. Similarly, human iPSC-derived NSCs grafted as neurospheres into SCI sites in mice differentiate into both glia and neurons and promote behavioral improvement (Nori et al., 2011). Human iPSCs can be differentiated into NSCs in a regionalspecific pattern and transplanted as caudalized human iPSC-derived NSCs into an early chronic SCI site in rats, where they generate both glia and neurons but fail to promote functional improvement (Nutt et al., 2013). NSCs in the latter study do not survive well in the lesion cavity and only a small proportion (11%) of grafted cells differentiate into neurons 2 months postgrafting (Nutt et al., 2013). We recently grafted FACS-purified human iPSC-derived NSCs into an immunodeficient rat C5 lateral hemisection site, where the grafted cells survived well, and the majority (71%) differentiated into NeuN + neurons that made connections with host neurons (Lu et al., 2014a,b). The generation of both neurons and glia from iPSCs in the injured spinal cord is of great significance since iPSCs are generated from adult somatic cells without the ethical concerns associated with embryo or fetal tissue usage, and they can also potentially be transplanted autologously to avoid immune rejection.

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Recently, however, several studies reported overgrowth or tumor formation after transplantation of human iPSC-derived NSCs in both SCI and other neurological disease models (Katsukawa et al., 2016; Koyanagi-Aoi et al., 2013; Lo´pez-Serrano et al., 2016; Miura et al., 2009; Nori et al., 2015; Okubo et al., 2016). This could be due to reactivation of the initially silenced transgenes (Choi et al., 2014), adult genomic instability (Ruiz et al., 2015), or generation of early-stage NSCs that aggressively divide for a long time (Katsukawa et al., 2016; Nori et al., 2015; Okubo et al., 2016). We observed a similar phenomenon of graft overgrowth or expansion in several human iPSC-derived NSC lines, when they were transplanted in injured spinal cord for a relative long term over 3 months (P. Lu et al., unpublished data). The risk of graft expansion and tumor formation prevents clinical translation of such a promising cell type for treatment of SCI and other neurological diseases and disorders.

3.1.5 Direct reprogramming of somatic cells into neurons or NSCs Besides differentiation of NSCs from iPSCs or ESCs, NSCs or neurons can be directly converted/reprogrammed from embryonic and adult somatic cells. The pioneering work by Vierbuchen et al. (2010) demonstrated that a combination of three factors, Ascl1, Brn2, and Myt1l, suffice to rapidly and efficiently convert mouse embryonic and postnatal fibroblasts into functional neurons in vitro. One year later, the same group used three factors in combination with NeuroD1, to convert human fetal and postnatal fibroblasts into iNs that exhibit typical neuronal morphologies and express multiple neuronal markers (Pang et al., 2011). Expression of additional factors can promote conversion of human fibroblasts into specific phenotypes of neurons, such as spinal motor neurons (Son et al., 2011). As we discussed early, mouse and human dermal fibroblasts can be converted into neurons in vitro by an all-chemical approach (Hu et al., 2015; Li et al., 2015). The advantage of the iN approach is that it directly converts somatic cells into neurons without a neuronal progenitor cell stage, therefore eliminating potential problems of overproliferation of induced NSCs or contaminated iPSCs in vivo. Besides iNs, induced neural stem cells (iNSCs) can also be generated from mouse and human fibroblasts by a single factor, Sox2 (Ring et al., 2012) or Oct4 (Mitchell et al., 2014). The same reprogramming factor Sox2 is used to convert astrocytes into neurons in vivo in both adult brain and adult injured spinal cord (Niu et al., 2013; Su et al., 2014). Similarly, a single neural transcription factor, NeuroD1, can also convert reactive glia after brain injury into glutamatergic neurons, and NG2-positive oligodendrocytes into GABAergic neurons (Guo et al., 2014). The later studies are particularly interesting since this reprogramming technique can convert the inhibitory nature of endogenous astrocytes from glial scar into functional neurons that could bridge the injured site for reconnectivity of otherwise disconnected spinal cord neurons. iNSCs can also be generated from mouse (Han et al., 2012; Kim et al., 2014; Thier et al., 2012) or monkey (Lu et al., 2013) fibroblasts using a combination of several reprogramming factors and can be stably expanded for many passages in vitro (Han et al., 2012; Kim et al., 2014; Thier et al., 2012). Moreover, iNSCs

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do not go through a pluripotent cell state, and thus reduce the risk for tumor formation after transplantation (Kim et al., 2014; Lu et al., 2013). Although most of the reprogramming studies of somatic cells into iNs or iNSCs are in vitro, the same technique can be applied ex vivo. Indeed, Torper et al. (2013) first demonstrated that engineered human fibroblasts and human astrocytes expressing inducible forms of neural reprogramming genes could be converted into neurons when the reprogramming genes are activated after transplantation into adult rat brain. Hong et al. (2014) generated iNSCs from mouse embryonic fibroblasts and transplanted them into a rat model of contusive SCI site. They showed differentiation of iNSCs into both neurons and glia and synaptic connectivity of grafted neurons with host neurons, which could enhance functional recovery (Hong et al., 2014). Such an ex vivo neural reprogramming and in vivo neural conversion has several advantages over traditional NSC transplantation. First, the somatic cells such as fibroblasts survive better than NSCs, especially within the injured CNS. Second, transplantation into patients takes much less time than iPSC-derived NSCs, since the neural conversion occurs in vivo after transplantation. Third, it is safe, since generation of iNs does not involve the intermediate stage of NSCs, which could overproliferate. However, the bottleneck for application of iNs or iNSCs is the low efficiency of generation, and mature iNs generated in vitro cannot be transplanted into SCI sites. Overcoming such a bottleneck could bring iNs or iNSCs closer to clinical translation (Su et al., 2014; Torper et al., 2013).

3.2 NSCs OR NPCs FOR NEURAL PROTECTION AND REMYELINATION After the initial damage to spinal cord, there is a secondary injury that further damages neuronal and glia cells surrounding the initial injury site. This secondary injury can last weeks or even months (Silva et al., 2014). NSC transplant at acute or subacute phases could potentially provide neuroprotection to reduce this injury. The potential mechanism behind this neuroprotective effect is that NSCs may ameliorate T-cell receptor-mediated T-cell activation and inhibit signaling of inflammatory cytokines in immune cells (Cusimano et al., 2012; Fainstein et al., 2008). Indeed, Emga˚rd et al. (2014) grafted human spinal cord-derived NSCs into a rat lumbar contusion injury site and demonstrated that the survival of host neurons in NSC grafted groups is significantly higher than in the control, indicating a neuroprotective effect. Indeed, this neuroprotective effect only occurs in NSC acutely or subacutely grafted groups, but not in the chronic group. Similarly, overexpression of L1 molecule in mouse ES cell-derived NSC aggregates rescues degenerated host motor neurons and parvalbumin-positive interneurons (Cui et al., 2011). Besides neuroprotection of host neurons surrounding the injury site, NSC transplants derived from mouse ESCs can rescue spared neural tissue at the site of injury itself (Salewski et al., 2015a). In addition, NSCs can secrete neurotrophic factors that protect host neuronal degeneration and promote host axonal regeneration after injury. Our early work demonstrated that NSCs of the C17.2 line constitutively secrete significant quantities of

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several neurotrophic factors in vitro, such as nerve growth factor (NGF), BDNF, and glial cell line-derived neurotrophic factor (GDNF) (Lu et al., 2003). The production of these neurotrophic factors continues in vivo when these cells are grafted into SCI site and supports extensive growth of host axons of known sensitivity to these growth factors (Lu et al., 2003). Recently, Nori et al. (2011) reveal the expression of neurotrophic factors (NGF, BDNF, and hepatocyte growth factor, HGF) by grafted human iPSC-derived NSCs in a mouse SCI model, which may be associated with host neuronal survival and host axonal regeneration. Similarly, Romanyuk et al. (2015) report that human iPSC-derived NSCs express NGF, BDNF, FGF8, and GDNF, which may associate with the sparing of spinal cord tissue and significant early functional improvement. Demyelination is a consequence of prolonged and dispersed oligodendrocyte cell death after SCI (Plemel et al., 2014). Transplantation of NSCs or glial progenitors, especially OPCs, can remyelinate these demyelinated axons to restore their functional conductivity. OPCs can be obtained and enriched from mouse ESCs using a specific differentiation protocol (Liu et al., 2000). When transplanted into demyelinated adult rat spinal cord or myelin-deficient shiverer (shi/shi) mutant mice, mouse ESC-derived OPCs differentiate into mature oligodendrocytes that are able to myelinate host axons (Liu et al., 2000). Later, a similar protocol was applied to generate human OPCs derived from human ESCs (Keirstead et al., 2005). Transplantation of human OPCs into subacute contusive SCI site promotes remyelination and enhances motor function. This study leads the first clinical trials of human ESCs for treatment of SCI (Eaton et al., 2015). Unfortunately, the clinical trial was stopped by the sponsor, Geron Corporation, due to financial constraints. A new study was reported recently to confirm feasibility of transplantation of human ESC-derived oligodendrocyte progenitors in rodent SCI model (Priest et al., 2015). Similar remyelination of host axons by transplanted NSCs derived either from ESCs (Salewski et al., 2015a) or iPSCs (Salewski et al., 2015b) has recently been reported. Myers et al. (2016) have recently reviewed remyelination studies by various cell engraftments, including NSCs and progenitors, and found an inconsistency among replications and so questioned the readiness for clinical translation.

3.3 AXONAL GROWTH AND CONNECTIVITY FROM NSC GRAFT For reconstitution of injured spinal cord, NSCs must differentiate into both neuronal and glial cells after transplantation, especially into neurons, which are necessary for neuronal replacement therapy. One of the hallmark features of a functional neuron is its connectivity to target cells by an axon and axonal terminals. Therefore, a successful neuronal replacement therapy for SCI requires integration of grafted neurons into host CNS circuity, including axonal growth and connectivity from donor neurons to host neurons. Reier et al. (1986) first demonstrated axonal projection from embryonic day 14 rat fetal spinal cord tissue graft using wheat germ agglutinin-conjugated horseradish peroxidase (WGA-HRP) as a tracer. In a later comprehensive study, both

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anterograde (WGA-HRP and Phaseolus vulgaris leucoagglutinin [PHA-L]) and retrograde tracer (Fluoro-Gold) were used to identify axonal projection from the graft (Jakeman and Reier, 1991). Graft-derived axons do project as far as 4–5 mm into host CNS tissue. However, the majority of these efferent fibers are confined to the immediate vicinity of the host–graft boundary. Although anterograde labeling can directly reveal axonal projections from grafted neurons, it only partially labels grafted neurons in certain locations. In addition, both anterograde and retrograde labeling may be subject to certain degree of artifact if leakage to surrounding tissue occurs or the injection is mistargeted (Steward et al., 2007). Later, the same group utilized pseudorabies virus for transneuronal labeling to test for host–graft neuronal connectivity. When the virus was injected into the phrenic nerve ipsilateral to the lesion/graft site, neurons similar to phrenic motor neurons are labeled in the graft, indicating synaptic connections from the grafted neurons onto host neurons within the phrenic circuit (White et al., 2010). The newly developed intrinsic cellular markers and reporters, such as green fluorescent protein (GFP) and alkaline phosphatase (AP), are more reliable methods for labeling axons emerging from neural implants than tracer injections since they are stable and easily identified. Fischer’s group was among the first to use the cell reporter, AP, to trace axons arising from rat fetal spinal cord tissue graft in the injured adult spinal cord, and observed axonal outgrowth for as long as 5 mm from the grafts, although there was considerable variation among animals (Lepore and Fischer, 2005). They also found that grafts containing a combination of NRPs and GRPs in the injured spinal cord failed to extend appreciable numbers of axons into the host spinal cord unless exogenous neurotrophins were also administered adjacent to the implant to stimulate outgrowth (Bonner et al., 2011; Lepore and Fischer, 2005). Besides using reporter genes from transgenic animals, NSCs or neural progenitors can be transduced to express these reporter genes using retroviral or lentiviral vectors (Blits et al., 2005; Vroemen et al., 2003). Unfortunately, most GFPtransduced rat NSCs were seen to differentiate into glia in the injured spinal cord, which limits their use to study axonal growth and connectivity from grafted NSCs with host neurons (Blits et al., 2005; Vroemen et al., 2003). Besides the above reporter genes, species-specific axonal markers may also be used to identify axonal extension and connectivity from xenografted NSCs. For example, Wictorin and Bj€ orklund (1992) grafted human fetal spinal cord cell suspension into adult rat spinal cord one segment above or below a partial lesion site and used a human-specific neurofilament marker to demonstrate human axonal extension into host white matter for distances up to 10 mm by 3–4 months postgraft. Along with their previous study of human axonal growth in adult lesioned rat brain (Wictorin et al., 1990), this finding, along with other studies (Foster et al., 1985; Nornes et al., 1983), is the first to report long-distance axonal growth in the inhibitory environment of the adult CNS. Although this study shows long-distance growth of human NSCs in the injured adult spinal cord, the human cells are implanted into normal cord parenchyma one segment (3 mm) above or below the lesion (Wictorin and Bj€ orklund, 1992). Whether human NSCs implanted into centers of SCI can survive,

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differentiate into neurons, extend their axons into host spinal cord, and establish synaptic connectivity is unknown from these early studies. Another example is an early study by Li and Raisman (1993) who grafted embryonic mouse hippocampal neurons into the intact rat cervical dorsal column where the main corticospinal tract (CST) is located. Using a mouse-specific axonal marker, M6, they found that grafted mouse neurons extend long, straight, and uniform axons into rat host spinal cord white matter for distances as long as 8 mm. The extension of the donor axons was confirmed by retrograde labeling with horseradish peroxidase. The graft-derived axons were intermingled with the host myelinated axons. Thus, this study is an early indicator that adult myelin is not necessarily inhibitory to the extension of developing axons. The drawback of the M6 marker is that it labels only early-stage axons and becomes ineffective when the grafts survive longer than 6–7 weeks (Li and Raisman, 1993). Recently, Grealish et al. (2014) use another human-specific NCAM marker to label human ESC-derived dopamine neurons grafted in a rat Parkinson disease model, which clearly showed long-distance axonal projection from midbrain to forebrain. This study indicates that grafted neurons have an intrinsic ability for growth and can innervate correct targets even in the injured adult environment. Recently, our lab has focused on the characterization of axonal growth and connectivity from grafted NSCs implanted in the injured spinal cord (Hou et al., 2013; Kadoya et al., 2016; Lu et al., 2012b, 2014a). Our hypothesis is that early-stage neurons from NSC implants have an intrinsic capability to extend their axons and connect with the host neurons even in the inhibitory environment of the injured adult spinal cord (Schwab et al., 2005). We first used a stable transgenic Fischer 344 rat line expressing a GFP reporter gene for NSC isolation to unequivocally label grafted cells, in order to follow their axonal growth and connectivity. Fischer 344 rats are inbred so that transplantation between animals of the same strain is isogenic, and does not require immunosuppression. We then transplanted freshly dissociated NSCs and NPCs from embryonic day 14 spinal cord or brainstem without in vitro culture to maximize neuronal differentiation potential and to reduce glia differentiation in vivo (Cao et al., 2001). In addition, we embedded freshly dissociated NSCs into a fibrin matrix containing growth factor cocktails to retain NSCs at the transected spinal cord lesion site and to support their survival, differentiation, and integration, since nonembedded NSCs survive poorly in severely injured spinal cord (Medalha et al., 2014). Finally, we transplanted NSCs 1–2 weeks postinjury to avoid the peak period of inflammation that could diminish the survival of NSCs in the lesion center. This subacute transplantation time is clinically relevant. Grafted rat NSCs consistently filled up transection sites and differentiated into abundant neurons and glia. More importantly, graft-derived neurons extended large numbers of axons into host rat spinal cord in both rostral and caudal direction (Fig. 2; Lu et al., 2012b). The axons of grafted neurons extended more than 20 mm (seven spinal segments) into the host spinal cord in rostral direction and 27 mm (nine segments) in caudal direction. We quantified 29,000 GFP-labeled axons emerging from the graft in the caudal direction (0.5 mm caudal to graft at T3 transection site) in a typical recipient host spinal cord. However, the density gradually decreases over

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FIG. 2 Extensive long-distance axonal outgrowth from neural stem cell grafts. (A) GFP and NeuN immunolabeling reveals that GFP-expressing neural stem cell grafts robustly extend axons into the host spinal cord rostral and caudal to the T3 complete transection site (caudal shown) over the 12-mm length of the horizontal section. (B and C) Higher magnification views from boxed area in (A). Extensive regions of the host spinal cord contain graft-derived projections in white matter (WM) and gray matter (GM). Inset shows that GFP-labeled projections arising from grafts express neurofilament (NF), confirming their identity as axons. Scale bar: (A) 550 mm; (B and C) 60 mm.

distance. In addition, we quantified 22% donor axons that were myelinated by host oligodendrocytes at 3 mm caudal to the graft (unpublished data). A time-course study demonstrated that the graft-derived axons initially traveled along host white matter and then made collateral branches for innervation of host gray matter neurons. We also transplanted human NSCs derived from fetal spinal cord, ESCs, or iPSCs into an adult rat SCI site (Lu et al., 2012b, 2014a). These cultured human NSCs were confirmed to be true NSCs by immunolabeling with the NSC markers, nestin and SOX2, in vitro before transplant (Lu et al., 2014a). Cultured human NSCs were transduced with lentiviral vectors expressing GFP reporter gene for their in vivo identification. We used immunodeficient rats as recipients for human cell xenograft to avoid problems associated with immunosuppression. This is especially relevant for iPSC-derived NSC grafts since iPSCs are potentially destined for autologous transplantation without immunosuppression. Again, we embedded human NSCs in fibrin matrices containing growth factor cocktails in order to retain them at the lesion site and to support their survival, growth, and differentiation in the injured spinal cord. When examined 7 weeks to 3 months postgraft, the majority of human NSCs were seen to have differentiated into neurons in vivo. More importantly,

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grafted human neurons extended very large numbers of human axons into host CNS for very long distances (Fig. 3; Lu et al., 2012b, 2014a). We quantified 20,500 axons from human iPSC-derived NSC graft per hemicord and this number is 41% higher than the number of axons emerging from grafts of rat-derived NSC cells. Indeed, the density of graft-derived human axons appears to be qualitatively equal to that of rodent host axons in some regions of white matter even at three spinal segments below the implantation site (Fig. 3D and E; Lu et al., 2014a). The total distance traversed by human axons corresponded to more than 26 spinal segments, exceeding 9 cm in length, much longer than has been seen for rat NSC-derived axons. Indeed, for example, human iPSC-derived NSCs from the C5 lesion site extended their axons through adult host white matter as far caudally as the distal lumbar spinal cord, and as far rostrally as the frontal cortex and olfactory bulb; human axons essentially

FIG. 3 Axonal extension of human iPSC-derived neural stem cells in sites of spinal cord injury. (A–C) Remarkably large numbers of GFP-labeled axons (colocalized with Tuj1, see insets) extend caudally into the host spinal cord (B) white matter and (C) gray matter (region of NeuN labeling). (D) GFP, MBP, and NeuN triple labeling of a coronal section three segments (C8) caudal to the graft shows dense distribution of human axons predominantly on right, lesioned side of the spinal cord. (E) Higher magnification of (D) from lateral white matter demonstrates remarkably high number of human axons interspersed in white matter. Scale bar: (A) 600 mm; (B and C) 32 mm; (D) 250 mm; (E) 20 mm.

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extended the entire length of the adult rat neuraxis. Both the density and distance of human axons traveled in adult injured spinal cord are unprecedented, to the best of our knowledge. These findings support our hypothesis that early-stage neurons have an intrinsic capability to extend their axons for connectivity even in the inhibitory environment of injured adult spinal cord. Graft-derived axons not only traveled for long distances but also make synaptic connection with host neurons. Both immunofluorescent labeling and electron microscopy revealed synaptic connections between graft-derived axons and host neurons, including host motor neurons (Fig. 4; Lu et al., 2012b). Graft-derived axonal terminals were observed around both host neuronal soma and dendrites, and seen to made abundant synaptic contacts with the host neurons. The growth and connectivity of human axons in adult injured spinal cord is confirmed by other studies. Zhao et al. (2013) grafted primitive neural stem cells (pNSCs) from human ESCs and demonstrated that grafted pNSCs survived, and differentiated into both glia and neurons that extended axons for long distances through the scar tissue at the graft–host interface and into the host spinal cord. Interestingly, some grafted human pNSCs were seen to differentiate into serotonergic neurons that extend 5-HT-positive processes into host spinal cord (Zhao et al., 2013), resembling those endogenous raphe-spinal axons derived from the brainstem. Similarly, grafts of human ESC-derived NSCs into a spinal compression injury site in Sprague–Dawley rats extended large numbers of axons for long distances into the host spinal cord, even though subject to immunosuppression (van Gorp et al., 2013). Although grafted human neurons can extend their axons for long distances and make synaptic connections with rodent host neurons, human neurons may still be immature and not fully functional after a few months postgraft in rodent SCI site,

FIG. 4 Synapse formation of graft-derived axons with host neurons. (A) A z-stack image triple labeled for GFP for graft-derived axons, synaptophysin (Syn, inset), and ChAT, indicating coassociation of rat NSC graft-derived axons with a synaptic marker in direct association with host motor neurons (arrowhead indicates one of the several examples). (B) Electron microscopy confirms that DAB-labeled GFP-expressing axon terminals form synapses (arrows) with host dendrites. Arrowhead indicates a separate, host–host synapse. Scale bar: (A) 8 mm; (B) 200 nm.

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since human CNS development and maturation is extended over a much longer period of time than in rodents (Semple et al., 2013). For example, the expression of neurofilament in the developing human CNS starts as late as gestational week 25 (>6 months) (Pundir et al., 2012) comparing to the embryonic day 15 in rodent brain (Chan et al., 1997). Similarly, synaptogenesis in human brain starts around gestational 20 weeks and takes years to complete comparing to rodent brain where synaptogenesis starts after birth (total 3 weeks for rodent gestation) and lasts only a few weeks (Semple et al., 2013). In addition, the maturation of oligodendrocytes and initiation of myelination in human brain starts at 23–36 gestational weeks comparing to postnatal days 1–3 in rodent (Semple et al., 2013). The prolonged developmental period of human neurons could limit their functional testing in rodent models of SCI, since the life span of rodent is relatively short. Alternatively, human NSCs can be tested in primate or other large animal models of SCI for long-term studies in order to fully appreciate their potential for differentiation, maturation, and functional connectivity.

3.4 HOST AXONAL REGENERATION AND CONNECTIVITY WITH NSC GRAFTS A neuronal relay requires not only outgrowth and connectivity of the grafted neurons with host neurons but also relies on ingrowth/regeneration and connectivity of host axons with grafted neurons (Fig. 1). Our hypothesis is that NSCs may provide a permissive environment for adult host axonal regeneration. On the other hand, host injured adult neurons may not have sufficient intrinsic growth capability to grow/ regenerate compared to young grafted neurons (Sun and He, 2010). Indeed, Iwashita et al. (1994) demonstrate extensive host neuronal regrowth cross a fetal spinal cord tissue graft for reconstruction of lesioned neonatal spinal cord. Such a robust host axonal growth/regeneration can only be achieved in neonatal animals, but not at the adult stage (Bregman et al., 1997). Although several studies have reported regeneration of adult host axons, including rubrospinal, reticulospinal, raphe-spinal, propriospinal, and sensory axons, into grafts of embryonic spinal cord tissue or ESCderived NSCs placed in sites of SCI (Bonner et al., 2011; Bregman et al., 1997; Jakeman and Reier, 1991; Lu et al., 2003), the distance and density of regenerating host adult axons into the grafts is typically modest. Our studies show similar results of modest ingrowth/regeneration of host supraspinal axons, including anterogradelabeled reticulospinal axons and 5-HT-immunolabeled serotonergic axons (Lu et al., 2012b, 2014a). In contrast to other host axonal tract systems that can only modestly regenerate into NSC graft, CST axons can robustly regenerate into NSC grafts. We recently reported that NSC or NPC grafts enable the extensive and consistent regeneration of CST axons into sites of severe SCI (Kadoya et al., 2016; Fig. 5). Indeed, CST axons can regenerate into complete spinal cord transection sites filled with an NSC graft. This is in contrast to early reports that CST axons can only grow through spared tissue bridges (Liu et al., 2010; Zukor et al., 2013). Successful

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FIG. 5 Host corticospinal tract axonal regeneration. (A and B) Host corticospinal tract (CST) axons labeled with BDA robustly regenerate into GFP-expressing neural stem cell grafts in site of C3 dorsal column lesion. (C and D) Higher magnification views from (B) showing (C) penetration of CST axons from main tract into NSC graft, and (D) cluster of regenerated CST axons within NSC graft. Dashed lines in (C) indicate host (H) and graft (G) interface. Scale bar: (A and B) 300 mm; (C and D) 60 mm.

regeneration of CST requires grafts to be driven toward caudalized (spinal cord), rather than rostralized, fates. Human ESC-derived NSCs also support regeneration of lesioned CST axons in rats and in immunosuppressed nonhuman primates (unpublished observations). The ability to robustly regenerate CST axons achieves a major milestone on the path to developing a translationally relevant SCI treatment for humans since CST is the most important projection for voluntary movement in humans. NSC grafts not only serve as a permissive cellular substrate to support host axonal regeneration but also function as new targets for regenerated axons to innervate or connect, since NSCs can differentiate into neurons. Indeed, we found that host axons formed bouton-like structures that are closely associated with grafted neurons and expressed the presynaptic marker synaptophysin, suggesting synaptic connections with grafted neurons (Kadoya et al., 2016; Lu et al., 2012b, 2014a). On the other hand, the connectivity of host axons with graft-derived neurons may limit continuous growth of regenerated host axons. This could explain why the regenerated descending host motor axons extend only into the rostral portion of the NSC grafts. Besides the spontaneous regeneration of host axons into NSC grafts, several studies have delivered neurotrophic factors to enhance the regeneration of injured adult axons into neural tissue grafts (Bregman et al., 1997; Lian Jin et al., 2011; Lu et al., 2003). Bregman et al. (1997) infused several neurotrophins (BDNF, NT-3, NT-4, and

3 NSCs for SCI repair

CNTF) individually at the site of injury and transplantation, and demonstrated increases in the extent of serotonergic, noradrenergic, and corticospinal axonal ingrowth within the transplants. NT-3 overexpression, delivered by ex vivo gene therapy into grafted NSCs, significantly enhanced host axonal regeneration into the NSC grafts, involving by both local sensory and motor axons (Lu et al., 2003). These results demonstrate that delivery of neurotrophins appears to be an effective tool to enhance host axonal regeneration. Since neurotrophins also affect NSC growth and differentiation, the time, route, duration, and dose of neurotrophin delivery all need to be carefully considered. Besides neurotrophins, other strategies that can enhance axonal regeneration are combined with NSC graft to increase therapeutic benefits (Hwang et al., 2011; Wang et al., 2011). For example, Hwang et al. (2011) combined NSC graft with the delivery of NT3 and chondroitinase, which cleaves chondroitin sulfate proteoglycans, at the interface between spinal cord and implanted NSCs and showed better host axonal growth into NSC transplant than was seen in the individual treatment controls. In addition, NSC survival, migration, and differentiation are enhanced in combination groups. A similar multiple strategy also promotes axonal integrity and plasticity in chronic SCI (Karimi-Abdolrezaee et al., 2010). Wang et al. (2011) cografted NSCs overexpressing TrkC with Schwann cells overexpressing NT-3 into the transected rat spinal cord. They found an enhancement of host serotonergic axonal growth in the cografting group compared to the individual treatment controls. Lowry et al. (2012) combined NSC grafts with long-term delivery of sonic hedgehog (Shh) from biodegradable microspheres and demonstrated enhancement of sprouting and regeneration of host supraspinal axons. To enhance the intrinsic growth capacity of adult neurons, He’s group recently identified several molecular pathways, including PTEN/mTOR, Jak/STAT, and DLK/JNK, which are associated with the regenerative capacity of adult neurons (Lu et al., 2014b). It will be interesting to combine these new strategies with NSC grafting to enhance host axonal regeneration and connectivity.

3.5 FUNCTIONAL OUTCOMES NSC transplantation not only reconstructs the injured spinal cord anatomically but may also bring functional benefits. The functional benefits could result from their intrinsic properties of neuroprotection, remyelination, and/or the formation of functional neuronal relays. The latter is unique to NSC transplants, which involves neuronal components of the transplant that reciprocally grow and connect with host neurons (Fig. 1). Therefore, we will focus on discussion of functional benefits from neuronal relay formation. Although many studies show various degrees of functional recovery after NSC transplant in SCI model, few explore the potential mechanism of functional neuronal relay formation. Abematsu et al. (2010) transplanted mouse fetal brain-derived NSCs along with valproic acid to promote neuronal differentiation in the mouse contusive injury model and showed that transplant-derived neurons reconstructed broken neuronal circuits by both receiving and sending synaptic connections to host neurons.

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Such a neuronal reconstruction dramatically enhances hind limb function. Ablation of the transplanted cells completely abolishes function recovery, confirming that the NSC transplant directly contributes to the functional recovery (Abematsu et al., 2010). Besides motor function, Bonner et al. (2011) have demonstrated the formation of sensory neuronal relays between injured dorsal column sensory axons and the denervated dorsal column nuclei by grafted mixtures of NRPs/GRPs. To promote graftderived axonal growth and connection with target neurons, they deliver BDNF by lentiviral vectors injected in the dorsal column nuclei. BDNF dramatically promotes growth and connection of graft-derived axons with target neurons. In addition, host sensory axons also regenerate into the graft, although the number is modest. The reciprocal connections between grafted and host neurons result in stimulus-evoked c-Fos expression in the grafted neurons and evoked responses in the target neurons after sciatic nerve stimulation with increased latency, indicating formation of a neuronal relay (Bonner et al., 2011). To demonstrate functional neuron relay formation for descending motor systems, we used the most severe injury model, complete upthoracic transection. Our anatomical analysis clearly shows long-distance growth and connectivity of graft-derived axons, modest ingrowth and connectivity of host reticulospinal axons (Lu et al., 2012b), and robust regeneration and connectivity of corticospinal tract axons (Kadoya et al., 2016). This reciprocal growth and connectivity of graft-derived neurons with host neurons could establish a novel neuronal relay that crosses a complete transection site. Indeed, we observed modest improvement in hindlimb locomotion in rats that received NSC grafts comparing to controls (Lu et al., 2012b). Retransection of spinal cord above the NSC graft abolished functional improvement, indicating that functional benefit requires host inputs into the graft from spinal cord and brain above the graft. Electrophysiological measures confirmed the formation of neural transmission across the lesion site, with responses detected that exhibited prolonged latency and altered waveforms, compared to intact animals. Again, electrophysiological transmission is abolished by retransection of the spinal cord above the NSC graft. These findings demonstrate that neuronal relays can be a new mechanism to support functional recovery after NSC transplantation in SCI.

4 FUTURE PERSPECTIVE The therapeutic application of NSCs for the reconstruction of injured spinal cord have great potential, especially at the present time when there are varieties of sources for NSC generation, especially from iPSCs and the direct conversion of adult somatic cells to NSCs. Transplanted NSCs not only exhibit therapeutic benefits for neuroprotection and remyelination but also can potentially serve as functional neuronal relays that can reconstruct the severely damaged spinal cord. Despite the great progress has been made in recent years, there is still a lack of consistent and meaningful improvement in histological and behavioral outcomes

4 Future perspective

after stem cell transplantation in the injured spinal cord. There are several challenges for NSC transplantation for SCI repair. First, transplanted NSCs must survive in injured spinal cord and differentiate into neural cells to exhibit their therapeutic effects. For neuroprotection and remyelination, NSCs can be transplanted either in the injury site or surrounding the injury site (Piltti et al., 2013). For neuronal relay formation, however, NSCs must be transplanted into the injury center to completely fill up and bridge damaged spinal cord for reconnectivity of separated spinal cord segments. Survival of NSCs in the injured spinal cord, especially at the center of a severe injury, is a challenge due to the inhospitable injury environment (Medalha et al., 2014). Immunorejection is another factor to be considered, when donor NSCs are allergenic or xenografted. Therefore, an efficient method to improve NSC survival, differentiation, and integration is needed. In our studies, transplantation of NSCs with growth factor cocktails embedded in fibrin matrix consistently supports grafted NSC survival, differentiation, and integration (Kadoya et al., 2016; Lu et al., 2012b, 2014a). Second, transplanted NSCs may have safety risks, such as graft expansion and tumor formation, since NSCs are stem cells that have a capacity to proliferate even after transplantation. This is especially true for iPSC-derived NSCs, since they are reprogrammed adult cells that bear more risk of tumor formation than fetal CNSderived or ESC-derived NSCs (Koyanagi-Aoi et al., 2013). An efficient method to prevent graft expansion and tumor formation while preserving the grafted neural cells is needed for safe use of iPSC-derived NSCs for the treatment of SCI and other neurological diseases and disorders. Another risk factor associated with NSC transplantation is that of allodynia or pain (Hofstetter et al., 2005; Macias et al., 2006). Suppression of glial differentiation or increase of neuronal differentiation may overcome this drawback (Davies et al., 2008; Hofstetter et al., 2005). Third, although several studies, including ours, demonstrate potential neuronal relay formation, the efficiency of such a relay to transmit neurotransmitters crossing injured spinal cord may be low. This is due to the lack of developmental cues in the adult CNS to organize grafted neurons toward an appropriate cytoarchitecture, and to guide graft-derived axons to their appropriate targets. In addition, the phenotypes of appropriate relay neurons to transmit neuronal signals crossing injured spinal cord are unknown, and identification and transplantation of such relay neurons may enhance functional outcomes. Guidance of graft-derived axons using neurotrophins or guidance molecules to the appropriate host target neurons could also enhance functional outcomes. Furthermore, enhancing the intrinsic adult neuronal growth state with neurotrophic factors or other means could enhance host axonal regeneration and strengthen the functional neuronal relay formation. Finally, intensive rehabilitation may reshape newly generated circuits and therefore promote functional recovery, since training and activity influence new circuit formation during development, particularly by activity-dependent stabilization of new connections and pruning of weak connections.

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ACKNOWLEDGMENTS This work was funded by grants from the Veterans Administration, NIH (NS09881), the Craig H. Neilsen Foundation, the Dr. Miriam and Sheldon G. Adelson Medical Research Foundation, and the California Institute for Regenerative Medicine.

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CHAPTER

Plasticity and regeneration in the injured spinal cord after cell transplantation therapy

2

Satoshi Nori, Masaya Nakamura, Hideyuki Okano1 Keio University School of Medicine, Tokyo, Japan Corresponding author: Tel.: +81-3-5363-3747; Fax: +81-3-3357-5445, e-mail address: [email protected]

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Abstract Spinal cord injury (SCI) typically damages the long axonal tracts of the spinal cord which results in permanent disability. However, regeneration of the injured spinal cord is approaching reality according to the advances in stem cell biology. Cell transplantation therapy holds potential to lead to recovery following SCI through some positive mechanisms. Grafted cells induce plasticity and regeneration in the injured spinal cord by promoting remyelination of damaged axons, reconstruction of neural circuits by synapse formation between host neurons and graft-derived neurons, and secreting neurotrophic factors to promote axonal elongation as well as reduce retrograde axonal degeneration. In this review, we will delineate (1) the microenvironment of the injured spinal cord that influence the plasticity and regeneration capacity after SCI, (2) a number of different kinds of cell transplantation therapies for SCI that has been extensively studied by researchers, and (3) potential mechanisms of grafted cell-induced regeneration and plasticity in the injured spinal cord.

Keywords Spinal cord injury, Cell transplantation therapy, Plasticity, Regeneration, Remyelination, Neuronal relay, Neurotrophic support

1 INTRODUCTION The adult mammalian central nervous system (CNS), which includes the brain and the spinal cord, has been recognized as a representative example of an organ in which regeneration is difficult and does not have enough plasticity (Cajal, 1928). A combination of factors, such as the inhibitory character of CNS myelin and injury-induced glial scars, the insufficient trophic support, and the small regenerative ability of endogenous adult neural stem/progenitor cells (NSPCs)

Progress in Brain Research, Volume 231, ISSN 0079-6123, http://dx.doi.org/10.1016/bs.pbr.2016.12.007 © 2017 Elsevier B.V. All rights reserved.

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CHAPTER 2 Plasticity and regeneration in the injured spinal cord after cell graft could be attributed to the lack of regenerative properties of the spinal cord. However, recent progress in the field of spinal cord injury (SCI) research especially on cell transplantation therapy demonstrated that the regeneration and plasticity of the injured spinal cord might really be possible (Barnabe-Heider and Frisen, 2008; Nakamura and Okano, 2013; Okano and Yamanaka, 2014; Okano et al., 2003). The microenvironment of the injured spinal cord can influence the plasticity and regeneration capacity after SCI. There are three phases of SCI, acute phase, subacute phase, and chronic phase. The acute phase of SCI is not suitable for cell transplantation, because it is considered as the inflammatory phase due to the upregulation of inflammatory cytokines, excitatory neurotransmitters, and free radicals. The differentiation capacity of grafted cells can be affected by the inflammatory environment and motor functional recovery was not observed in animal models of SCI (Cao et al., 2001). Moreover, the chronic phase is also not a good option for cell transplantation. Because the glial scar formation occurred in the chronic phase of SCI, and it prevents axonal regeneration. Therefore, the subacute phase is considered as the ideal time window for cell transplantation (Nishimura et al., 2013; Ogawa et al., 2002; Okano, 2002; Okano et al., 2003). There are several kind of cells used for cell transplantation therapy. NSPCs have been one of the most widely examined stem cells in the therapeutic study of SCI. The efficacy and safety of the transplantation of NSPCs into injured spinal cord have been widely demonstrated in several animal models (Cummings et al., 2005; Iwanami et al., 2005; Ogawa et al., 2002; Okada et al., 2005). Although NSPCs for neural regeneration after SCI may be promising, clinical application has not yet been realized in Japan due to the ethical issues with the use of NSPCs, which had to be collected from the brains of aborted fetuses. To avoid this ethical problem, NSPCs derived from induced pluripotent stem cells (iPSCs) have been tested for SCI transplantation and increasing experimental evidence supports their therapeutic potential (Kawabata et al., 2016; Kobayashi et al., 2012; Nori et al., 2011; Tsuji et al., 2010). However, there is reported to be a risk of tumorigenesis after transplantation of iPSC-derived NSPCs (Nori et al., 2015). A further study is now underway to prevent tumorigenesis after transplantation. Cell transplantation therapy can exert plastic changes by (A) remyelination, i.e., remyelinating the damaged axon to facilitate the nerve conduction after SCI; (B) reconstructing of neural circuits, i.e., graft-derived neurons act as novel interneurons between the injured axons and the denervated neurons distal to the injury; and (C) neurotrophic support, i.e., grafted cells produce neurotrophic factors that can enhance axonal sparing, neuronal survival, plasticity, and regeneration. These three mechanisms work in a synergic manner to facilitate plasticity and regeneration of injured spinal cord after cell transplantation therapy. This review outlines the present state of several different types of cell transplantation therapy for SCI and furthermore we discuss the mechanisms of regeneration and plasticity after cell transplantation therapy in the injured spinal cord.

2 Optimal timing of cell transplantation and plasticity after SCI

2 OPTIMAL TIMING OF CELL TRANSPLANTATION AND PLASTICITY AFTER SCI Plasticity after cell transplantation is affected by the microenvironment of the injured spinal cord. Various cytokines affect the cell fates of NSPCs after transplantation. A previous study showed that the levels of inflammatory cytokines (tumor necrosis factor (TNF)a, Interleukin (IL)-1a, IL-1b, and IL-6) peak 6–12 h after injury and remain elevated till 4 days after injury (Nakamura et al., 2003). Although these inflammatory cytokines are known to have neurotoxic as well as neurotrophic actions in the injured spinal cord, extremely high expression of these cytokines within 7 days after injury is neurotoxic. These inflammatory acute phase of SCI are not suitable for cell transplantation because of not only the inflammatory cytokines but also excitatory neurotransmitters and free radicals. IL-1b and IL-6 are rapidly increased in the injured spinal cord at this time point, they would induce Jak/Stat-signaling (Bonni et al., 1997) and grafted NSPCs mostly differentiate into astrocytic fates and do not contribute to functional recovery (Cao et al., 2001). Kumamaru et al. directly assess the in vivo biology of acutely engrafted NSPCs by using RNA sequencing. The grafted NSPCs showed beneficial effects on SCI, such as neuroprotection and neurohumoral secretion. Secreted substances such as platelet-derived growth factor, transforming growth factor beta 1 (TGF-b1), and IL-10 have critical roles in preventing neural apoptosis through the PI3K-Akt/NF-kB pathway (Kaltschmidt et al., 2005; Romashkova and Makarov, 1999; Zhou et al., 2009). Nerve growth factor (NGF) and brain-derived neurotrophic factor (BDNF) prevent activation of the mitochondrial apoptotic pathway through the P75NTR-mediated NF-kB pathway (Mattson and Meffert, 2006; Sofroniew et al., 2001). However, in vivo secretory activity of NSPCs was significantly different from their in vitro activity. Notably, overall transcriptional activity, external signal transduction, and neural differentiation of grafted NSPCs were significantly downregulated in the acute SCI environment compared with the grafted NSPCs in the uninjured spinal cord. They concluded that grafted NSPCs are vulnerable to the environmental force (Kumamaru et al., 2012). On the other hand, our study showed that transplantation of NSPCs into rat cervical SCI 9 days after injury resulted in behavioral improvement in upper extremities as compared to that in the control group. The grafted cells differentiated into neurons, astrocytes, and oligodendrocytes. Furthermore, we observed synapse formation between graft-derived neurons and host neurons (Ogawa et al., 2002). Tarasenko et al. (2007) transplanted the human NSPCs which had been treated with fibroblast growth factor-2 (FGF-2), heparin, and laminin into the rats SCI at the same day or 3 or 9 days after SCI. The best results with optimized survival rate, neuron and oligodendroglia differentiation, and motor functional recovery were observed in the 9-day postinjury transplantation group. They also claimed that the functional improvement is related with the transplantation time point after SCI. This apparent difference indicated that the allowance of a therapeutic time window after SCI is important for NSPC transplantation into the injured spinal cord. The acute

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CHAPTER 2 Plasticity and regeneration in the injured spinal cord after cell graft inflammatory phase lasts up to 1 week after injury in rodent SCI, and this period should be avoided for NSPC transplantation. On the other hand, about 2 weeks or more after SCI, a glial scar formation prevents axonal regeneration. Our previous study showed the extent of glial scar formation and the characteristics of inflammation are the most notable difference between the subacute injured microenvironment and the chronic injured microenvironment. The transplantation of NSPCs at the subacute phase of SCI, but not at the chronic phase, can promote functional recovery (Nishimura et al., 2013). Therefore, the optimal time window for NSPC transplantation is considered to be 1–2 weeks after SCI in a rodent model (the subacute phase; Nishimura et al., 2013; Ogawa et al., 2002; Okano, 2002; Okano et al., 2003).

3 CELL TRANSPLANTATION FOR NEURAL REGENERATION AND PLASTICITY 3.1 NEURAL STEM/PROGENITOR CELLS NSPCs were first demonstrated in the subventricular zone of the mouse in 1989 (Temple, 1989) and methods for culturing were established by Reynolds and Weiss (1992). NSPCs can be maintained in vitro culture in spherical bodies to proliferate, called a “neurosphere” (Reynolds and Weiss, 1992). This culture system has shed light on the molecular biological characteristics of NSPCs in the developmental process as well as in the CNS of adult mammals (Alvarez-Buylla et al., 2001; Doetsch et al., 1999; Eriksson et al., 1998; van Praag et al., 2002). NSPCs were capable of self-renewal and differentiating into neurons, astrocytes, and oligodendrocytes in vitro and in vivo (Reubinoff et al., 2001). Transplantation of NSPCs has been widely examined in the therapeutic study of SCI. Previous studies showed many different strategies including reconstruction of neural circuits by synapse formation, remyelination, and trophic factor secretion provided by the transplanted cells (Abematsu et al., 2010; Cummings et al., 2005; Hawryluk et al., 2012; Yasuda et al., 2011). Ogawa et al. (2002) transplanted rat fetal NSPCs into rat cervical SCI and showed behavioral improvement in skilled forelimb movement (pellet retrieval test) as compared to that in the control group. The grafted NSPCs differentiated into neurons, astrocytes, and oligodendrocytes and synapse formation between graft-derived neurons and host neurons was observed. Cummings et al. (2005) transplanted human NSPCs into nonobese diabetic-severe combined immunodeficient (NOD-SCID) mice thoracic SCI model. Grafted cells survived, migrated, and differentiated into neurons and oligodendrocytes, and were associated with motor functional recovery. The electron microscopic analysis demonstrated that remyelination by the grafted cells and synapse formation between grafted cells and mouse host neurons. Since the selective ablation of grafted cells by diphtheria toxin abolished the recovery, the survived grafted cells played a critical role in motor functional recovery. To test proof of concept on the effectiveness of NSPC transplantation for SCI in nonhuman primates, Iwanami et al. (2005)

3 Cell transplantation for neural regeneration and plasticity

transplanted human aborted fetal brain-derived NSPCs into nonhuman primate (common marmoset) SCI model. The transplanted cells survived and differentiated into neurons, astrocytes, and oligodendrocytes. Significantly better functional recovery was observed in NSPC-grafted mice compared with that in the control group. These results led to strong expectations that fetal brain-derived NSPCs are very useful as cell sources for transplantation in humans. However, there are still no prospects for clinical application of NSPCs in Japan due to ethical issues with the use of NSPCs derived from the brains of aborted fetal tissues.

3.2 EMBRYONIC STEM CELL-DERIVED NEURAL STEM CELLS Embryonic stem cells (ESCs; Evans and Kaufman, 1981; Martin, 1981; Thomson and Marshall, 1998) are pluripotent stem cells that can be derived from the inner cell mass of the early blastocyst. ESCs have the ability to differentiate into lineages derived from all three germ layers (Evans and Kaufman, 1981; Martin, 1981; Thomson et al., 1998). As the ESCs have the capability to differentiate into all cell types, the most common strategy to restore the neural function after SCI is the transplantation of various neural cells derived from ESCs. Previously, several reports were published to generate NSPCs, motor neurons, and oligodendrocyte progenitor cell (OPC)-like cells from ESCs. To verify the neural regenerative capability of ESC-derived cells, transplantation of these cells into various animal SCI models were also performed. Okada et al. (2008) used mouse ESCs derived from the inner cell mass to induce NSPCs with high plasticity that are present at a relatively early stage of development, and succeeded in constructing a neural development model culture system in vitro that mimics neural development and reflects its temporal and spatial specificity. This culture system eliminates the requirement for leukemia inhibitory factor to maintain the ESCs in their undifferentiated state, and by using floating cultures it allows the formation of embryoid bodies (EBs), which contain cells derived from the three germ layers. These EBs contain relatively early stage NSPCs, by growing EBs in floating cultures containing serum-free neural stem cell culture medium supplemented with FGF-2. Furthermore, the addition of a low concentration of Noggin plays an important role in forebrain formation, and the addition of a low concentration of retinoic acid increases the efficiency of neurosphere formation. Primary neurospheres formed in this manner can be subcultured as secondary and tertiary neurospheres. Although primary neurospheres produce neurons almost exclusively, secondary and tertiary neurospheres produce astrocytes and oligodendrocytes as well as neurons. Furthermore, by adding Noggin or changing the concentration of retinoic acid during EB formation, NSPCs with anterior–posterior axis region-specific properties can be induced. Similarly, dorsal–ventral region-specific control has been achieved by adding the ventralization factor sonic hedgehog or the dorsalization factor bone morphogenic protein-4 or Wnt3a to the culture during the formation of primary neurospheres. These results suggest that the regional specificity of NSPCs could be regulated by adding the right factor at the right time during the culture.

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CHAPTER 2 Plasticity and regeneration in the injured spinal cord after cell graft The efficacy of ESC transplantation into the injured spinal cord was demonstrated by McDonald et al. (1999). EBs formed from mouse ESCs and transplanted into the rat SCI model resulted in good functional recovery (McDonald et al., 1999). However, the risk of tumor formation associated with transplantation of less-differentiated EBs cannot be ignored. Keirstead et al. (2005) established a method of human ESCs to form OPCs-like cells by using a culture medium that contained a factor that promotes oligodendrocyte differentiation. After transplantation of these cells into the injured spinal cords of rats, they observed the remyelination of demyelinated axons and motor functional recovery. Kumagai et al. (2009) transplanted primary and secondary neurospheres induced from mouse ES cells via EB formation using the culture system of Okada et al. Interestingly, primary neurospheres showed neuron-dominant differentiation, whereas the secondary neurospheres differentiated into neurons, astrocytes, and oligodendrocytes is the outcomes were consistent with their in vitro characteristics: transplantation of secondary neurospheres significantly prevented the atrophy and demyelination of the injured spinal cord, and the motor functional recovery was only observed in the secondary neurosphere transplantation group. These results suggest that it is preferable to transplant NSPCs generating both neurons and glial cells. However, because the ESCs were established from excess embryos from infertility treatment, the ethical issues still remain in using these cells.

3.3 iPSC-DERIVED NSPCs iPSCs are somatic cells that have been reprogrammed by transducing such genes as Oct4, Sox2, Klf4, and c-Myc into mouse or human fibroblasts (Okita et al., 2007; Takahashi and Yamanaka, 2006; Takahashi et al., 2007). They are pluripotent stem cells that have the same proliferative ability and differentiation potential as ESCs. It appears possible to generate iPSCs from somatic cells, a technology that is expected to resolve the ethical problems that hinder the use of ESCs. NSPCs can be induced by iPSCs by using the same culture system used in ESCs. Previously, Miura et al. (2009) assessed the safety of each mouse iPSC (miPSC) clone by transplanting miPSC-derived NSPCs (miPSC-NSPCs) into the brains of NOD-SCID mice. Next, Tsuji et al. (2010) used a miPSC clone whose safety had been confirmed (clone 38C2) to examine the effectiveness of treating SCI by transplanting 38C2-derived NSPCs into the lesion site in a mouse model. The 38C2iPSC-derived SNSs (38C2-SNSs) transplantation was performed during the subacute stage at 9 days after SCI. Approximately 20% of the transplanted 38C2-SNSs survived within the injured spinal cord and differentiated into definable CNS cell types. Severe atrophic change and demyelination were significantly diminished in the group treated with 38C2-SNSs. Quantification of LFB-positive areas at the lesion epicenter revealed that 38C2-SNS-grafted mice demonstrated a significantly larger myelinated area than PBS-treated control mice. Detailed observation indicated that graft (38C2-SNS)-derived oligodendrocytes contributed to remyelination. Furthermore, evaluation of hindlimb motor function on the Basso mouse scale (BMS)

3 Cell transplantation for neural regeneration and plasticity

revealed significantly better recovery of function in the group treated with miPSC-NSPCs than in the control groups (PBS-treated and fibroblast-transplanted groups). Human iPSCs (hiPSCs), which do not entail ethical concerns, may become a preferred cell source for regenerative medicine in human patients. Nori et al. (2011) used a hiPSC clone (201B7) established by transducing four reprogramming factors (Oct4, Sox2, Klf4, and c-Myc) into adult human fibroblasts (Takahashi et al., 2007). The 201B7 clone was differentiated into NSPCs (201B7-NSPCs) and was transplanted into the injured spinal cord of NOD-SCID mice to evaluate their therapeutic effect. The grafted cells survived and differentiated into neurons, astrocytes, and oligodendrocytes. Approximately 50% of the grafted 201B7-NSPCs differentiated into neurons, of which 70% differentiated into g-aminobutyric acid (GABA)-ergic neurons. Furthermore, immunoelectron microscopic examination revealed graft-derived presynaptic and postsynaptic structures as well as synapse formation between host neurons at the injured site. To electrophysiologically evaluate functional recovery, motor evoked potential (MEP) was measured. MEP waves were detected in 201B7NSPC-grafted group, but not in the vehicle control group. Moreover, axonal regeneration, angiogenesis, and tissue preservation were also observed. As a result, BMS, rotarod test, and treadmill gait analysis showed good improvement in the lower extremity motor function in the 201B7-NSPC-grafted group compared with control group. The greatest potential drawback of hiPSC-based therapies is their potential for tumorigenicity. Therefore, follow-up was continued for 112 days after SCI, which revealed that the functional recovery was maintained with no tumor formation. To examine the effectiveness and safety of transplantation of hiPSC-NSPCs for SCI in primates, Kobayashi et al. (2012) moved on to 201B7-NSPC transplantation for treatment of SCI in common marmosets. Grafted 201B7-NSPCs survived and differentiated into neurons, astrocytes, and oligodendrocyte progenitor cells. Grafted 201B7-NSPCs enhanced axonal growth, myelination, and angiogenesis, thereby promoting motor functional recovery after SCI. Furthermore, there was no tumor formation for 12 weeks after transplantation. Taken together, once preevaluated for safety, hiPSC-NSPCs are a potential cell source for the treatment of SCI in the clinic. It is important to address safety issues especially with regards to tumorigenicity when considering the clinical use of hiPSC-NSPCs. Although we reported the safety and therapeutic potential of 201B7-NSPCs (Nori et al., 2011), the NSPCs derived from a different hiPSC clone, 253G1—which was established by transducing three reprogramming factors (Oct4, Sox2, and Klf4) into adult human fibroblasts (Nakagawa et al., 2008)—showed tumor formation after long-term observation (Nori et al., 2015). Since the 253G1 clone was generated without c-Myc, it was initially speculated that 253G1-NSPCs would be safer than 201B7-NSPCs. Nori et al. (2015) showed grafted 253G1-NSPCs survived, differentiated into three neural lineages, and the grafted mice exhibited temporary motor functional recovery for up to 47 days after transplantation. However, it was

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CHAPTER 2 Plasticity and regeneration in the injured spinal cord after cell graft followed by a gradual deterioration in motor function, accompanied by tumor formation. The tumors consisted of Nestin + undifferentiated neural cells and they exhibited activation of OCT4- and KLF4 transgenes. Transcriptome analysis also revealed that epithelial–mesenchymal transition may have contributed to the progression of tumors. Based on these findings, integration-free iPSCs (Okita et al., 2008, 2011) should be chosen to avoid transgene-induced tumorigenesis. As a step toward clinical applications, integration-free iPSC-NSPC transplantation into immune-deficient animals has been initiated. Moreover, transplantation into immune-deficient animals, along with subsequent long-term observation, should be used to determine the safety of the grafted cells.

3.4 MESENCHYMAL STROMAL CELLS Mesenchymal stromal cells (MSCs) are a kind of multipotent stem cell initially identified from the bone marrow (Friedenstein et al., 1974; Prockop, 1997). The population of MSCs is 0.001–0.01% of the total population of nucleated cells in the bone marrow (Pittenger et al., 1999). Human MSCs are easily obtained from bone marrow by iliac crest puncture. They are biologically safe and have been widely applied for transplantation in patients suffering from hematological cancer. The definition of multipotent MSCs is that (A) they are plastic-adherent when cultured in standard conditions; (B) they express CD105, CD73, and CD90, and lack the expression of CD45, CD34, CD14 or CD11b, CD79a or CD19, and HLA-DR surface molecules; and (C) they differentiate into osteoblasts, adipocytes, and chondrocytes in vitro (Jiang et al., 2002; Pittenger et al., 1999). MSCs promise a minimally invasive, autologous source of cells for transplantation and have been used in cell transplantation therapies for many diseases. The therapeutic potential of MSCs for SCI is still controversial. In general, the benefits of transplantation of MSCs are thought to be indirect environmental modification rather than direct translineage conversion of MSCs to neural lineage cells. Although neural differentiation is possible in vitro (Rismanchi et al., 2003), it is still controversial whether it has any meaningful effect in adult tissue in vivo (Castro et al., 2002; Mezey et al., 2003). Previous studies showed that transplantation of MSCs provides tissue sparing via neurotrophic paracrine effects and posttrauma inflammation regulation (Akiyama et al., 2002; Ankeny et al., 2004). Nakajima et al. (2012) reported that grafted MSCs significantly upregulated the level of IL-4 and IL-13 expression and downregulated the level of TNF-a and IL-6. The changes of inflammation factors resulted in the shifting of macrophage phenotype from M1 to M2. Previous reports also showed that MSCs can mobilize endogenous NSPCs (Mahmood et al., 2004a) as well as provide scaffolding for elongating axons (Ankeny et al., 2004; Hofstetter et al., 2002). Taken together, transplantation of MSCs promotes motor functional recovery in several studies. However, Tetzlaff et al. (2011) showed the inconsistency of these positive effects in a systematic review, among six human MSC transplantation studies: three reported benefits and three found no difference after transplantation.

3 Cell transplantation for neural regeneration and plasticity

3.5 OLFACTORY ENSHEATHING CELLS Olfactory ensheathing cells (OECs) possess special properties which resemble peripheral nervous system (PNS) Schwann cells and CNS astrocytes (Ramon-Cueto and Avila, 1998). OECs are present in the olfactory epithelium, they exhibit lifelong proliferative capacity, and they normally act as intermediary glial cell. They mediate the transition between axons in the PNS olfactory mucosa and their synapses in the CNS olfactory bulb (Doucette, 1991). Like Schwann cells, OECs can produce similar axon growth molecules and promote axonal regeneration, although the remyelination ability is poorer than Schwann cells (Lakatos et al., 2003; Techangamsuwan et al., 2008). Previous study showed the bridging effect of transplanted OECs on regenerated axons of from dissected dorsal root into spinal cord (Li et al., 2004). Another study showed that grafted OECs preserved the electrophysiological function of circuitry with the evoked cord dorsum potentials and sensorimotor cortex potentials in the region of dorsal column lesion (Toft et al., 2007). Although transplantation of OECs has been reported effective for SCI, the mechanism of the regeneration is still not clear. The previous studies have shed doubt on the transplantation of OECs promoted the motor functional recovery after SCI. Lu et al. (2006) reported that no significant axon growth promoting effect as well as no bridging effect of corticospinal axons in the OEC-transplanted group. Collazos-Castro et al. (2005) also showed neither regeneration of corticospinal tract nor motor functional recovery after transplantation of OECs into rats cervical SCI. Consistent with these animal studies that showed mixed result, clinical trials of OEC transplantation for human SCI have shown inconsistent result (Chhabra et al., 2009; Lima et al., 2006, 2010; Mackay-Sim et al., 2008).

3.6 SCHWANN CELLS Schwann cells (SCs) populate the growth-permissive PNS and are the myelinating cells of the PNS. They play important roles in postinjury nerve regeneration by supporting axonal outgrowth and remyelination and act as a bridge. Some previous studies reported that following SCI, the CNS generates Schwann or Schwann-like cells which extrude PNS myelin but not CNS myelin (Blakemore and Franklin, 2008; Gillespie et al., 1994). Zawadzka et al. (2010) revealed that the majority of Schwann cell-like cells originate from endogenous CNS precursors after induced demyelination by fate mapping analysis. At the same time, some PNS-derived mature Schwann cells dedifferentiated into immature Schwann cells via c-Jun mediated pathways and they migrate from the periphery into the lesion site (Nagoshi et al., 2011; Zawadzka et al., 2010). Another study showed that after transplantation into a demyelinated spinal cord slice ex vivo, neurotrophic factors which include NGF, BDNF, and ciliary neurotrophic factor expressed by SCs promote the survival and regeneration of damaged axons (Park et al., 2010). Furthermore, transplanted SCs also expressed many kinds of cell adhesion molecules and extracellular matrix proteins such as N-cadherin,

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CHAPTER 2 Plasticity and regeneration in the injured spinal cord after cell graft integrins, laminin, neural cell adhesion molecule (N-CAM), L1, contactin, and collagens (Ghosh et al., 2012; Pierucci et al., 2009). Recently, genetically modified SCs, combinational strategies involving SCs with matrices and scaffolds, and cotransplantation of SCs with MSCs have also been attempted. Deng et al. (2011) showed that grafted genetically modified glial cell line-derived neurotrophic factor (GDNF)-overexpressing SCs suppressed the expression of glial fibrillary acidic protein (GFAP) and chondroitin sulfate proteoglycans (CSPGs) of reactive astrocytes. This study indicated a therapeutic strategy to control the reactive astrocyte-induced inhibitory environment and to promote axonal regeneration, remyelination as well as motor functional recovery after SCI. Patel et al. (2010) reported the injectable matrices which composed of laminin and collagen showed the improvement of SCs survival, graft angiogenesis, and axonal in growth compared with control group. Ban et al. (2011) reported the effectiveness of cotransplantation of SCs with MSCs. They showed significantly more regenerated axons as well as reduced number of astrocytes in the lesion epicenter in cograft group. Moreover, electron microscopic examination revealed completely reconstructed myelin sheaths in the cotransplantation group.

4 PLASTICITY AND REGENERATION AFTER CELL TRANSPLANTATION THERAPY Above-mentioned cell transplantation therapy is the promising therapy for SCI patients. There are three main mechanisms by which transplanted cells may facilitate plasticity and regeneration. Grafted cells can exert plastic changes by (A) remyelinating the damaged axons, (B) reconstructing of neural circuits, and (C) tissue sparing by trophic factors of neuroprotection (Fig. 1). Many previous studies indicated cell transplantation therapies most often focus on replacing the glial environment. Schwann cells, OECs, pluripotent cell-derived OPCs, and NSPCs have inherent glial and myelinating properties (Blakemore and Franklin, 2008; Keirstead et al., 2005; Li et al., 2004; Yasuda et al., 2011). On the other hand, some recent studies focus on reconstructing of neural circuits (Abematsu et al., 2010; Fujimoto et al., 2012; Nori et al., 2011). In other reports, motor functional recovery has been reported without any obvious replacement of neural cells. For example, no glial cells or neurons were generated by MSCs. The mechanism here appears to be indirect, such as providing trophic support, modulating the inflammatory response, or providing a substrate for axonal growth (Parr et al., 2007).

4.1 REMYELINATION Remyelination was achieved by grafted CNS oligodendrocytes or grafted PNS SCs. OPCs can be efficiently derived from murine and human ESCs (Brustle et al., 1999; Nistor et al., 2005) and give rise to mature myelinating oligodendrocytes when transplanted to the CNS (Brustle et al., 1999; Keirstead et al., 2005).

4 Plasticity and regeneration after cell transplantation therapy

A

B

C

FIG. 1 Three main mechanisms of plasticity and regeneration in the injured spinal cord by cell transplantation therapy. (A) Remyelination of the demyelinated axons by graft-derived oligodendrocytes or Schwann cells. (B) Reconstructing of neural circuits (relay formation) by synapse formation between graft-derived neurons and host neurons, including the insertion of a graft-derived neuron between the injured axon and the target neuron. (C) Neurotrophic factors secreted by grafted cells to spare tissues, reduce the damage of the spinal cord, and promote axonal growth.

Previously, Yasuda et al. (2011) showed the contribution of remyelination to functional recovery following contusive SCI in mice. To isolate the effect of remyelination from other possible regenerative effects of the grafted cells, we transplanted shi-NSPCs which were obtained from myelin-deficient shiverer mutant mice alongside wild-type (wt-)NSPCs. Except for their myelinating potential, shi-NSPCs showed the same behavior as wt-NSPCs in vitro and in vivo. Myelin basic protein (MBP) was not expressed by shi-NSPCs-derived oligodendrocytes in vitro and formed much thinner myelin sheaths in vivo compared with wt-NSPCs-derived oligodendrocytes. Although shi-NSPC-grafted mice showed some locomotor and electrophysiological recovery, significantly better recovery was observed in wt-NSPC-grafted mice. These findings showed the biological importance of remyelination by the graft-derived cells for motor functional and electrophysiological recovery after the transplantation of NSPCs. In a similar study was performed by another laboratory, Salewski et al. (2015) transplanted the wt-mouse iPSC-derived NSPCs (wt-iPSC-NSPCs) and shi-mouse iPSC-derived NSPCs (shi-iPSC-NSPCs) into mouse thoracic contusion injury. They showed significantly better motor functional and electrophysiological recovery in the wt-iPSC-NSPCs group than in

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CHAPTER 2 Plasticity and regeneration in the injured spinal cord after cell graft the shi-iPSC-NSPCs group. The compact and multilayer remyelination conveyed by wt-iPSC-NSPCs group contrasted with the inefficient myelination of shi-iPSC-NSPCs group. They concluded the importance of remyelination for SCI repair and regeneration. Unlike the mouse cells, human NSPCs and human-iPSC-NSPCs predominantly differentiate into neurons and less mature oligodendrocytes in vitro as well as in vivo (Kobayashi et al., 2012; Nori et al., 2011; Romanyuk et al., 2015). Therefore, Numasawa-Kuroiwa et al. (2014) developed the induction method for oligodendroglial differentiation of hiPSC-NSPCs. This hiPSC-OPC-enriched NSPCs differentiated into oligodendrocytes, as well as into neurons and astrocytes. Kawabata et al. (2016) transplanted hiPSC-OPC-enriched NSPCs into the injured spinal cord of NOD-SCID mice to evaluate the therapeutic potential. Grafted hiPSC-OPCenriched NSPCs differentiated into oligodendrocytes (34.5%), astrocytes (36.1%), and neurons (26.1%) in mouse spinal cord. Graft-derived oligodendrocytes migrated broadly into host white matter and contributed to remyelination of spared axons. Moreover, graft-derived neurons formed synapses with host mouse neurons in the injured spinal cord, the reconstruction of neuronal circuits, were also observed. The hiPSC-OPC-enriched NSPC transplanted group showed significantly better motor and electrophysiological functional recovery compared with the control group. By simply comparing the behavioral data of hiPSC-OPC-enriched NSPCs and conventional hiPSC-NSPCs, their recovery is equivalent. Therefore, further studies using larger animal SCI models may be needed for assessing the differences in functional recovery between these two cells. Other reports showed that SCs and OECs also had the ability to wrap injured axons in peripheral myelin (Pinzon et al., 2001; Takami et al., 2002). However, they required trophic adjuvants to functionally remyelinate host axons.

4.2 RECONSTRUCTION OF NEURAL CIRCUITS This strategy involves the formation of neuronal relays between injured long tract axons and denervated neurons. Graft-derived neurons in the injured spinal cord function as novel interneurons between the injured axons and the denervated neurons distal to the injury (Abematsu et al., 2010; Bonner et al., 2011; Fujimoto et al., 2012; Nori et al., 2011). This approach has some advantages that injured axons do not need to regenerate long distances and axon growth is placed on the grafted neurons, which can be selected for the ability to extend long axons in vivo (Bonner and Steward, 2015). Previously, Nori et al. (2011, 2015) transplanted hiPSC-NSPCs into the injured spinal cord of mice and showed the motor functional recovery after transplantation. Approximately 50% of the grafted cells differentiated into neurons and grafted bIII tubulin-positive/human nuclear protein-positive cells in parenchymal location were contacted by synaptic boutons of the host origin neurons, suggesting that the host axons innervated hiPSC-derived neurons. Immunoelectron microscopic examination revealed graft-derived presynaptic and postsynaptic structures as well as synapse

4 Plasticity and regeneration after cell transplantation therapy

formation between host neurons at the injured site. MEP waves could be detected in the hiPSC-NSPC-grafted group, but not in the control group, suggesting that the neurons derived from the grafted cells functioned as interneurons in the mouse spinal cord and contribute to the reconstruction of neural circuits. Abematsu et al. (2010) transplanted the mouse fetal NSPCs to show neuronal relay formation in mice severe thoracic contusion injuries. After transplantation, NSPCs mainly differentiated into neurons with the systemic valproic acid treatment. Electron microscopy revealed that the graft-derived neurons rostral to the graft received synapses from the host and that graft-derived axons extended caudally from the transplant to form synapses with host neurons below the injury site. They also injected the WGA-expressing adeno-associated virus into the motor cortex and showed evidence of relay formation based on transsynaptic transport of WGA. They showed WGA granules in neurons in the ventral horn throughout the spinal cord in uninjured control mice. In mice with thoracic contusion injuries, WGA granules were observed only in the neurons above the injury, whereas not in the neurons below the injury. On the other hand, in NSPC-grafted mice, they showed WGA granules in neurons above and below the injury. They also showed significantly better functional recovery in NSPC-grafted mice, which was eliminated when grafted cells were ablated with diphtheria toxin. Hence, graft-derived neurons contributed to the relay formation in the injured spinal cord as well as improved motor function. Consistently, the same group showed similar results with hiPSC-derived neuroepithelial-like stem cells (Fujimoto et al., 2012). Yokota et al. (2015) focused on the intraspinal neural networks connecting each spinal cord segment; the propriospinal system. They transplanted mouse fetal NSPCs into mouse thoracic SCI model and applied laser microdissection to investigate the transcriptional activity of grafted NSPCs. The grafted NSPCs promoted the reorganization of propriospinal circuit through synapse formation between the grafted NSPCs and the host neurons. Next, they injected the excitotoxin N-methyl-D-aspartic acid (NMDA) into mice spinal cord to ablate the host spinal cord neurons, and then 7 days after ablation they produced SCI and transplanted NSPCs. Interestingly, the synaptogenic potential of the grafted NSPCs was abolished and the efficacy of the reorganization of propriospinal circuits was negated by the selective ablation of host neurons. According to these results, they concluded that the accelerated reorganization of propriospinal circuits by NSPC transplantation is important for motor functional recovery after SCI.

4.3 NEUROTROPHIC SUPPORT Neurotrophic support is also an important role of stem and associated cell transplantation after SCI. Stem cells secrete neurotrophic factors that can enhance axonal sparing, neuronal survival, plasticity, and regeneration (Teng et al., 2002, 2006). The previous studies on axonal regeneration traditionally focused on production of neurotrophins; NGF, BDNF, neurotrophin-3 (NT-3), and neurotrophin-4/5 (NT-4/5; Brock et al., 2010; Jakeman et al., 1998; Jones et al., 2001; Tuszynski,

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CHAPTER 2 Plasticity and regeneration in the injured spinal cord after cell graft 1997; Tuszynski et al., 1996). The neurotrophins are a family of four structurally related proteins that are important in developmental events such as cell survival, cell differentiation, and axonal growth in both the PNS and CNS (Harvey et al., 2015). BDNF has been reported to be important in many reparative capacities in the CNS, including axonal sprouting, neurogenesis, neuroprotection, myelination, and synaptic plasticity (Bamber et al., 2001; Blesch and Tuszynski, 2007; Brock et al., 2010). NT-3 has been reported to enhance the axonal regeneration and sprouting in the CNS (Alto et al., 2009). NGF and NT-4/5 have been the least tested in SCI studies. Although NGF was initially reported to enhance growth of sensory (including nociceptive) axons (Tuszynski et al., 1994), other studies showed axonal growth with improved motor functional recovery (Feng et al., 2009; Huang et al., 2006). Previous studies demonstrated that the effect of NT-4/5 was on neuroprotection and axonal growth after SCI (Blesch et al., 2004; Bregman et al., 1997). Moreover, cytokine growth factors, TGF-b family members, GDNF, as well as hepatocyte growth factor (HGF) all promote outgrowth of axons after SCI (Glazova et al., 2009; Jones et al., 2001; Kitamura et al., 2007, 2011; Moalem et al., 2000; Peng et al., 2003). It is well recognized that the effectiveness of MSC therapy in SCI is a result of indirect environmental modification. MSCs secrete BDNF, NGF, vascular endothelial growth factor (VEGF), TGF-b, and HGF. They are known to have positive effects such as axonal outgrowth, neuronal and glial survival, and angiogenesis (Crigler et al., 2006; Hu et al., 2005; Mahmood et al., 2004b; Parr et al., 2007). It is also reported that graft-derived astrocytes produce several neurotrophic factors and angiogenic growth factors. These factors promoted the regrowth of axons as well as endogenous repair (Hofstetter et al., 2002; Kumagai et al., 2009; Mense et al., 2006; Nori et al., 2011; Tsuji et al., 2010; Yoshida et al., 2002). Previously, Nori et al. (2011) showed that hiPSC-NSPC-derived astrocytes are closely associated with 5-hydroxytrypamine (5-HT)-positive fibers and neurofilament (NF-H)-positive fibers, suggesting that graft-derived astrocytes promoted noncellautonomous effects for facilitating the regeneration or regrowth of host axons. From the result of an RT-PCR expression analysis of graft- and host-derived neurotrophic factors (NGF, BDNF, and HGF), expression of these factors was observed in both the grafted human cells as well as in the host mouse spinal cord. It is interesting to know that not only grafted hiPSC-NSPCs secrete these factors by themselves but they also promote secretion from the host mouse tissue. Furthermore, transplantation of hiPSC-NSPC-enhanced angiogenesis and tissue sparing after SCI. Both the results of RT-PCR and histological analysis showed that grafted hiPSC-NSPC-derived astrocytes as well as host astrocytes expressed VEGF, suggesting that the transplantation of hiPSC-NSPCs promoted VEGF expression in both host- and graft-derived astrocytes (Nori et al., 2011). Hence, motor functional recovery after cell transplantation therapy was achieved not only by cell-autonomous mechanisms but also by noncell-autonomous (neurotrophic) mechanisms.

5 Conclusions

5 CONCLUSIONS The main goal of cell transplantation therapy is the regeneration of neurons and glial cells that undergo cell death after SCI. Cell transplantation therapies are the most successful therapeutic approach for SCI, enabling improved motor functions in several animal models. Previous researchers have been trying to reconstruct the histologically impaired spinal cord structure. There are three possible mechanisms by which grafted cells may facilitate regeneration and plasticity in the injured spinal cord. (A) Grafted cells promote remyelination through oligodendroglial cell replacement, leading to neuronal and spinal cord tissue sparing. Remyelination of damaged axons influences motor and sensory systems and they can preserve sublesion functions after SCI. (B) Reconstruction of neural circuits by grafted neuronal cells is also a promising strategy for the repair of SCI. A neuronal relay aims at adding new neurons at the injured spinal cord to restore communication along the long tracts of the spinal cord. However, continued technical improvement will be needed to improve specificity of connectivity, and successful functional outcome and reproducibility. (C) Neurotrophic factors secreted by grafted cells can enhance axonal sparing, neuronal survival, plasticity, and regeneration in the injured spinal cord. Moreover, grafted cells can also promote the secretion of the neurotrophic factors from the host cells in the injured spinal cord. Therefore, grafted cells in the injured spinal cord have not only autocrine but also paracrine ability. Previously, many researchers tried to achieve motor functional recovery at the chronic phase of SCI by NSPC transplantation. However, in most of the studies, no significant recovery of motor function has been obtained (Cusimano et al., 2012; Karimi-Abdolrezaee et al., 2006; Kusano et al., 2010; Nishimura et al., 2013; Parr et al., 2007). Nishimura et al. (2013) showed that the distribution of chronically grafted NSPCs was restricted compared with NSPCs grafted at the subacute phase, and the motor functional recovery was not observed in chronic group. The microenvironment of chronic injured spinal cord appears to be unfavorable for the therapeutic mechanisms of NSPC transplantation due to extensive glial scarring and the phenotype of infiltrating macrophages. Kumamaru et al. (2012) demonstrated that chronically grafted NSPCs had a capacity for differentiating into neurons and oligodendrocytes, and for secreting neurotrophic factors. However, motor functional recovery of chronic SCI animals was not observed. The failure of the chronic transplantation was not due to the lack of therapeutic potential of grafted cells but the refractory state of the chronic SCI environment. CSPGs are involved in glial scar formation and they are considered to be the most prevalent axonal growth inhibitors. Previous reports showed that digesting CSPGs by chondroitinase ABC (ChABC) promoted axonal growth and the migration of grafted NSPCs (KarimiAbdolrezaee et al., 2010). It is also notable that combination of rehabilitation (treadmill training) enhanced the effect of NSPCs-transplantation for chronic SCI mice model (Tashiro et al., 2016). Therefore, cell transplantation therapies for chronic SCI should be combined with the environmental modification therapies such as

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CHAPTER 2 Plasticity and regeneration in the injured spinal cord after cell graft ChABC administration and rehabilitation, and that would improve tissue plasticity and regeneration in the injured spinal cord.

ACKNOWLEDGMENTS We thank Dr. Akimasa Yasuda for illustrations. Work in the author’s laboratory was supported by grants from the JST-CIRM collaborative program; Grants-in-Aid for Scientific Research from JSPS and the Ministry of Education, Culture, Sports, Science, and Technology of Japan (MEXT); the Project for Realization of Regenerative Medicine and Support for Core Institutes for iPS Cell Research from the MEXT; a grant for the Research Center Network for Realization of Regenerative Medicine from the MEXT and A-MED; the Kanrinmaru project from Keio University; Research Fellowships for Young Scientists from the Japan Society for the Promotion of Science; Keio Gijuku Academic Development Funds; and a Grant-in-Aid for Scientific Research on Innovative Areas (Comprehensive Brain Science Network) from the MEXT.

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CHAPTER

Transplantation of GABAergic interneurons for cell-based therapy

3

Julien Spatazza, Walter R. Mancia Leon, Arturo Alvarez-Buylla1 The Eli and Edythe Broad Center of Regeneration Medicine and Stem Cell Research, University of California, San Francisco, San Francisco, CA, United States 1 Corresponding author: Tel.: 415-514-2348; Fax: 415-514-2346, e-mail address: [email protected]

Abstract Many neurological disorders stem from defects in or the loss of specific neurons. Neuron transplantation has tremendous clinical potential for central nervous system therapy as it may allow for the targeted replacement of those cells that are lost in diseases. Normally, most neurons are added during restricted periods of embryonic and fetal development. The permissive milieu of the developing brain promotes neuronal migration, neuronal differentiation, and synaptogenesis. Once this active period of neurogenesis ends, the chemical and physical environment of the brain changes dramatically. The brain parenchyma becomes highly packed with neuronal and glial processes, extracellular matrix, myelin, and synapses. The migration of grafted cells to allow them to home into target regions and become functionally integrated is a key challenge to neuronal transplantation. Interestingly, transplanted young telencephalic inhibitory interneurons are able to migrate, differentiate, and integrate widely throughout the postnatal brain. These grafted interneurons can also functionally modify local circuit activity. These features have facilitated the use of interneuron transplantation to study fundamental neurodevelopmental processes including cell migration, cell specification, and programmed neuronal cell death. Additionally, these cells provide a unique opportunity to develop interneuron-based strategies for the treatment of diseases linked to interneuron dysfunction and neurological disorders associated to circuit hyperexcitability.

Keywords Interneuron, Ganglionic eminence, Transplantation, Plasticity, Epilepsy

1 INTRODUCTION Normal brain function requires balanced levels of excitation and inhibition. In the mammalian cerebral cortex, these functions are respectively attributed to excitatory glutamatergic pyramidal cells and inhibitory interneurons expressing GABA Progress in Brain Research, Volume 231, ISSN 0079-6123, http://dx.doi.org/10.1016/bs.pbr.2016.11.005 © 2017 Elsevier B.V. All rights reserved.

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CHAPTER 3 Transplantation of GABAergic interneurons for cell-based therapy (g-aminobutyric acid) that together represent about 20% of all cortical cells (Lodato and Arlotta, 2015). While pyramidal neurons make long-range connections within and outside the cortex, interneurons synchronize the activity of local projection neuron ensembles and gate excitatory and inhibitory inputs that they receive (Klausberger and Somogyi, 2008; Klausberger et al., 2003; Lewis et al., 2012; Vogels and Abbott, 2009). Interneurons are thus considered to be the main cellular components for the control of brain excitability. Accordingly, a wide range of neurological and psychiatric disorders stem from cortical interneuron dysfunction (Marı´n, 2012). Notably, these conditions include epilepsy, schizophrenia, and autism, and have been referred to as interneuropathies (Kato and Dobyns, 2005). Neuronal transplantation has been extensively studied as a potential therapeutic strategy for the treatment of various neurological conditions. For such an approach to be successful, the candidate cell should be able to disperse following transplantation and functionally integrate within the diseased host circuitry in a manner that recapitulates the properties of the endogenous cells targeted for replacement. However, most cell types display very little dispersal upon transplantation in the postnatal central nervous system (CNS) (Dunnett and Bj€orklund, 2012; Gage, 2012; Lindvall et al., 1990), which is a prerequisite for the functional integration of the transplants. The discovery of the origin of telencephalic interneurons in mice (Anderson et al., 1997; Tamamaki et al., 1997), as well as the capacity for their precursors to functionally integrate as inhibitory interneurons upon transplantation in the postnatal mouse brain (Alvarez-Dolado et al., 2006; Wichterle et al., 1999), provided an opportunity to test the potential of interneuron transplantation as a therapy for interneuropathies and other conditions associated to circuit hyperexcitability. Here, we summarize advances in the emerging field of interneuron biology and transplantation and also review some work on the potential clinical relevance of interneuron transplantation. First, we briefly summarize telencephalic interneuron development and discuss their behavior upon transplantation in the postnatal mouse CNS. We then touch upon how transplantation has been used for the study of CNS development and eventually examine the disease-modifying properties of interneuron transplants from studies based on mouse models of epilepsy, Parkinson’s disease (PD), Alzheimer’s disease (AD), and psychiatric disorders.

2 DEVELOPMENT OF TELENCEPHALIC GABAergic INTERNEURONS 2.1 TANGENTIAL MIGRATION The ability of interneurons to migrate after heterochronic transplantation into the postnatal mouse brain likely stems from the extensive migration they undergo during development. Excitatory and inhibitory cortical neurons emerge from two distinct compartments in the developing brain. Excitatory neurons are produced locally in the ventricular zone of the pallium and invade the cortex radially using the radial-glial scaffold as a migratory substrate (G€otz and Huttner, 2005; Molyneaux et al., 2007).

2 Development of telencephalic GABAergic interneurons

In contrast, cortical inhibitory neurons are generated outside of the cortex in the ventral telencephalon and must migrate tangentially over long distances to reach their final position in the cortex (Anderson et al., 1997; de Carlos et al., 1996; DeDiego et al., 1994; Tamamaki et al., 1997; Wichterle et al., 2001). It is precisely this capacity to migrate across the radial-glial scaffold that may allow young interneurons to disperse through the postnatal brain parenchyma, making them a strong candidate for transplantation and cell-based therapy in the CNS. While the subcortical origin of telencephalic interneurons and their migratory route to reach the cortex was originally described in mouse, nonradial migration from the ventral forebrain also applies to interneurons in the developing primate cortex (Hansen et al., 2013; Ma et al., 2013), which challenges earlier reports suggesting a cortical origin for these cells (Jones, 2009; Letinic et al., 2002; Yu and Zecevic, 2011).

2.2 ORIGINS AND DIVERSITY In the mammalian cortex, inhibitory interneurons are less numerous than pyramidal cells by a ratio of 1:5. However, this small population of local circuit nerve cells displays high diversity in shape and function. Deciphering the meaning and origin of this interneuron diversity is key for our understanding of how information is processed in the brain and how defects in specific interneuron circuits may give rise to diseases. Consequently, many groups have worked toward characterizing this cellular diversity as well as its functional relevance for cortical circuit physiology. Accordingly, interneurons can be classified in more than 20 subtypes based upon various criteria (partially overlapping for some of them) including morphology, physiology, patterns of local connectivity, and molecular identity (DeFelipe et al., 2013; Gonchar and Burkhalter, 1997; Markram et al., 2004; Petilla Interneuron Nomenclature Group et al., 2008). While such an approach is necessary to fully appreciate the complexity of this heterogeneous cell population (DeFelipe et al., 2013; Petilla Interneuron Nomenclature Group et al., 2008), it is noteworthy that the expression of the calcium-binding protein parvalbumin (PV), the neuropeptide somatostatin (SST), and the ionotropic serotonin receptor 5HT3a (5HT3aR) defines three nonoverlapping groups of cells that account for nearly 100% of interneurons in the mouse primary somatosensory cortex (Rudy et al., 2011). Subtype identity is dictated by the spatiotemporal origin of cortical interneurons during development (Butt et al., 2005; Flames et al., 2007; Fogarty et al., 2007; Gelman et al., 2009; Ghanem et al., 2007; Miyoshi et al., 2010; Xu, 2004). The majority of mouse cortical interneurons are generated between E10.5 and E16.5 by progenitors located in the ventricular and subventricular zones of the subpallium, within the ganglionic eminences. This highly proliferative compartment of the embryonic brain can be anatomically and molecularly divided into three regions, namely, the lateral-, the medial-, and the caudal ganglionic eminences (LGE, MGE, and CGE, respectively). While it is commonly accepted that the LGE does not contribute to the mouse cortical interneuron population (Wichterle et al., 2001; Wonders and Anderson, 2006), the MGE and CGE are the two major sources of cortical

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CHAPTER 3 Transplantation of GABAergic interneurons for cell-based therapy interneurons and give rise to anatomically and functionally distinct subsets of cells (Anderson et al., 2001; Butt et al., 2005; Fogarty et al., 2007; Lavdas et al., 1999; Miyoshi et al., 2010; Nery et al., 2002, 2003; Rubin et al., 2010; Wichterle et al., 2001). MGE and CGE also produce interneurons that migrate to other brain regions, including striatum, septum, hippocampus, and amygdala, thus illustrating the heterogeneity and importance of these germinal zones. MGE-derived neurons represent  60–70% of all cortical interneurons in rodents. These cells express either PV or SST and are born for the most part during early neurogenesis and preferentially locate to deep layers of the neocortex (Anderson et al., 2001; Marı´n, 2013; Wichterle et al., 2001; Xu et al., 2008). From a molecular standpoint, MGE-derived interneurons are specified by transcription factors including the Dlx genes, Lhx6, Sox6, and Nkx2.1 (Chedotal and Rijli, 2009; Flandin et al., 2011; Kessaris et al., 2014; McKinsey et al., 2013; Sussel et al., 1999; Vogt et al., 2014). In contrast to the early production of MGE-derived interneurons, interneuron generation in the mouse CGE has been shown to peak at around E16.5 (Miyoshi et al., 2010). Progenitors in the CGE express the orphan nuclear receptors COUP-TF I/II (Kanatani et al., 2008) and generate  30% of mouse cortical interneurons (Miyoshi et al., 2010; Nery et al., 2002; Rudy et al., 2011). CGE-derived neurons represent a very heterogeneous pool of cells expressing vasoactive intestinal polypeptide (VIP) and calretinin (CR) as well as a group of cells that do not express VIP and include neurogliaform reelin (RLN)-expressing cells (Rudy et al., 2011). Virtually all CGE-derived interneurons express 5HT3aR in the neocortex (Lee et al., 2010; Vucurovic et al., 2010). CGE-derived neurons mostly target the superficial layers of the neocortex independently of their time of birth (Lee et al., 2010; Miyoshi et al., 2010). Interestingly, more than half of human cortical interneurons are thought to originate from CGE progenitors (Hansen et al., 2013), which could reflect the evolutionary expansion of the upper layers of the cortex that are highly enriched in late-born CGE-derived neurons (Hansen et al., 2013; Miyoshi et al., 2010). In addition to the major contributions from both MGE and CGE, the preoptic area (POA) accounts for  10% of all cortical interneurons (Gelman et al., 2009). This group includes some neuropeptide Y (NPY)-expressing multipolar cells, as well as some PV- and SST-positive cells. Two distinct progenitor domains have been identified so far in the POA, one expressing Nkx5.1 and another Dbx1 (Gelman et al., 2009, 2011).

3 TRANSPLANTATION AND THE STUDY OF BRAIN DEVELOPMENT The initial studies that unraveled the subpallial origin of cortical interneurons were mostly based on dye labeling of discrete groups of cells in cultured mouse brain slices (Anderson et al., 1997; Tamamaki et al., 1997). Before the advent of genetic fate mapping techniques, transplantation allowed for the in vivo confirmation of migratory routes and also provided valuable information on the fate and functions of cortical interneurons. Additionally, transplantation studies demonstrated the

3 Transplantation and the study of brain development

Neonate transplant

I I

II LGE MGE Interneuron progenitors

II

E13.5 mouse CGE Adult transplant

FIG. 1 Heterochronic transplantation of interneuron progenitors. The MGE or CGE is dissected from the embryonic mouse brain. The MGE is anatomically separated from the LGE by a large sulcus; the CGE is a caudal extension of both LGE and MGE. Dissociated cells from these ganglionic eminences can be transplanted using beveled glass needles into both neonatal and adult nervous system (see text). MGE and CGE interneuron progenitors have the ability to migrate and differentiate into multiple interneuron subtypes that become integrated into functional circuits; dispersal is more robust in the permissive neonatal brain.

remarkable ability for embryonic MGE and CGE cells to functionally integrate into both neonatal and adult host circuits (Fig. 1), and also provided key information on many aspects of interneuron development.

3.1 INTERNEURON INTRINSIC DEVELOPMENTAL PROGRAM The extraordinary migratory potential of MGE cells was first demonstrated in vitro (Wichterle et al., 1999). Using embryonic mouse brain explants grown in matrigel, MGE-derived neuroblasts were found to migrate extensively, as opposed to cells derived from neocortical explants. Upon homotopic and isochronic transplantation in utero using ultrasound guided injection, MGE cells were shown to migrate dorsally perpendicular to the radial-glial scaffold via both the neocortical subventricular and marginal zones. These homotopic and isochronic MGE transplant-derived cells primarily populated the neocortex but also contributed significantly to the globus pallidus, the striatum, the amygdala, and the CA1 region of the hippocampus (Wichterle et al., 2001). Transplanted MGE cells persisted into adulthood and mostly differentiated into aspiny local interneurons immunoreactive for GABA, PV, and SST, illustrating that the fate of interneurons was determined prior to their exit of the ganglionic eminence (Flames et al., 2007; Fogarty et al., 2007; Wonders et al., 2008). In contrast, LGE transplant-derived cells were found to migrate ventrally and anteriorly to give rise to medium spiny neurons in the striatum, nucleus accumbens, and olfactory tubercle, as well as granule and periglomerular cells in

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CHAPTER 3 Transplantation of GABAergic interneurons for cell-based therapy the olfactory bulb (Wichterle et al., 2001). Interestingly, upon isochronic transplantation in the MGE, LGE cells did not modify their migratory behavior and remained in the ventral forebrain, with very few cells populating the neocortex, thus suggesting that at least some aspects of the development of ventral forebrain neuronal progenitors are intrinsically determined. Heterochronic transplantation studies further confirmed that transplanted immature interneurons from the ganglionic eminences preserve their internal developmental program (Fig. 1). First, when injected in the postnatal brain, MGE cells display an initial highly migratory phase reminiscent of their distant origins (Fig. 2). Accordingly, transplant-derived cells have been shown to migrate distances up to 2.5 mm in the adult rodent brain (Davis et al., 2015; De la Cruz et al., 2011; Hunt et al., 2013; Martı´nez-Cerden˜o et al., 2010) and 5 mm in the neonate (Alvarez-Dolado et al., 2006; Southwell et al., 2010). Second, in line with the postnatal maturation of interneurons in vivo (Okaty et al., 2009), following their migratory phase, transplanted

CGE MGE

CC

Transplant: E13.5 MGE or CGE MGE 6DAT Host: P2

PV

RLN

SST

VIP

CC

P8

MGE 20DAT

MGE 35DAT

CGE 35DAT

P22

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FIG. 2 Transplant-derived interneuron development in the heterochronic environment. MGE and CGE progenitors were transplanted into the cortex of a P2 host. At 6 days posttransplantation (6DAT), many MGE and CGE transplant-derived cells are found within the superficial layers of the cortex and are tangentially oriented, a behavior reminiscent of endogenous interneuron migration within the marginal zone of the developing neocortex (top). A large number of cells have also started to invade the cortex at 6DAT and display a radial orientation (top). At this stage, virtually all transplant-derived cells display a typical migratory morphology, with a long leading process and a short trailing process (bottom). At 20DAT, many transplanted interneurons have undergone programmed cell death. The transplanted MGE cells that survive usually stay clear of cortical layer I (as opposed to transplanted CGE cells), distribute across all cortical layers (top), and display a more mature morphology (bottom). At 35DAT, the vast majority of MGE transplant-derived cells differentiate into GABAergic interneurons expressing either PV or SST. CGE transplants give rise to many neurogliaform neurons that express RLN and to VIP-expressing interneurons. Neurogliaform interneurons mostly localize to layer I. CC, corpus callosum. Scale bar: 50 mm.

3 Transplantation and the study of brain development

MGE interneurons develop over the course of several weeks in the heterochronic environment before acquiring mature morphology, marker expression, and electrophysiological properties (Alvarez-Dolado et al., 2006; Howard and Baraban, 2016; Southwell et al., 2010) (Fig. 2). Finally, while transplanted MGE cells can develop in regions they normally migrate to, they similarly differentiate into mature interneurons in regions of the CNS they are not fated to populate. For instance, upon heterochronic transplantation in the spinal cord, MGE precursor cells migrate away from the injection site, survive, and eventually display molecular marker expression, morphology, and electrophysiological properties similar to those of cortical interneurons (Bra´z et al., 2012, 2014, 2015). Recent results have shown that the development of CGE cells is also intrinsically determined. Indeed, despite the known ontogeny of cortical interneurons since the late 1990s, it is only recently that studies addressed whether CGE cells, like their MGE counterparts, might be amenable to transplantation in the postnatal brain (Fig. 1). Not surprisingly, CGE transplant-derived cells were found to migrate extensively following heterochronic transplantation in the neonatal brain, with a dispersal similar to that of MGE cells (Hunt and Baraban, 2015; Larimer et al., 2016), thus suggesting that tangential migration of ventral forebrain interneuron precursors is a determinant factor for their dispersal upon transplantation. However, conflicting results were found following transplantation in the adult brain, with one report describing the failure of CGE cells to disperse in the mature cortex (Davis et al., 2015) and another study showing accentuated dispersal of CGE cells compared to that of MGE cells (Isstas et al., 2016). Such inconsistencies may be due to the lack of a clear anatomical distinction between MGE/LGE and CGE (Fig. 1). As the CGE is a caudal extension of both LGE and MGE, using the most rostral aspect of the CGE for transplantation may give rise to grafts enriched in LGE-derived cells that exhibit poor dispersal (Wichterle et al., 1999). However, in line with genetic fate mapping studies (Miyoshi et al., 2010; Rudy et al., 2011), CGE transplant-derived interneurons were more likely to localize to cortical layer I (Larimer et al., 2016) and express VIP, CR, and RLN (Hunt and Baraban, 2015; Isstas et al., 2016; Larimer et al., 2016) (Fig. 2). Altogether, this work shows that the fates of both MGE and CGE cells are instructed by developmental programs established in the embryo and that these grafts can be used to complement existing circuits with specific subsets of interneurons. Starting at 35 days after transplantation, transplanted MGE progenitors display electrophysiological properties and intrinsic firing patterns similar to those of endogenous MGE-derived interneurons (Howard and Baraban, 2016; Larimer et al., 2016). Their synaptic integration was first illustrated using electron microscopy (Baraban et al., 2009; Southwell et al., 2010; Wichterle et al., 1999) and was confirmed by intracellular recordings on acute brain slices showing spontaneous and evoked postsynaptic currents (Alvarez-Dolado et al., 2006; Howard and Baraban, 2016; Martı´nez-Cerden˜o et al., 2010; Southwell et al., 2010), thus demonstrating that grafted cells receive functional inputs from host neurons. Paired recordings further showed inhibitory synapses made by transplanted MGE or CGE cells onto host

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CHAPTER 3 Transplantation of GABAergic interneurons for cell-based therapy pyramidal neurons (Howard and Baraban, 2016; Larimer et al., 2016; Southwell et al., 2012), as well as reciprocal inhibitory connections between CGE transplant-derived cells and host interneurons (Larimer et al., 2016). In agreement with their endogenous inhibitory function in the CNS and their functional integration upon transplantation, grafted MGE-derived interneurons can modify synaptic inhibition in the host brain (Alvarez-Dolado et al., 2006; Baraban et al., 2009; Bra´z et al., 2012; Howard et al., 2014; Southwell et al., 2010) and are thus of a great clinical interest for the manipulation of inhibition in disorders that display circuit hyperexcitability (Chohan and Moore, 2016; Southwell et al., 2014; Tyson and Anderson, 2014). The therapeutic potential of CGE transplants has not been as extensively evaluated as that of MGE transplants so far. However, the expanded repertoire of transplantable interneurons allowed by these grafts will undoubtedly offer further insight into interneuron development.

3.2 INTERNEURON FATE AND SURVIVAL Cortical interneuron precursor transplantation has been used to study the origin of interneuron subtypes. While MGE and CGE transplantation showed the variety of cortical interneurons generated by ventral forebrain progenitors, grafts of microdomains of the MGE provided insight into precursor cell heterogeneity. First, marker expression analysis revealed that each subregion of the subpallium can be further divided into distinct progenitor domains (Flames et al., 2007). In order to address whether these domains indeed corresponded to functionally distinct progenitor pools, homotopic and isochronic in utero transplants of the most dorsal and the most ventral domains of the MGE were generated. The neurochemical composition of the transplants was analyzed at postnatal day 14 (P14) and showed a strong bias for the generation of SST interneurons from the most dorsal region, as opposed to a bias for the production of PV interneurons by the ventral region (Flames et al., 2007). These results were corroborated using neonatal transplants (Inan et al., 2012; Wonders et al., 2008). Neonatal transplantation of microdomains has also been very useful to study the ontogeny of specific interneuron subtypes. Chandelier cells have attracted much attention due to their ability to both depolarize and hyperpolarize pyramidal neurons (Glickfeld et al., 2009; Szabadics et al., 2006; Woodruff et al., 2009, 2011) and their potential implication in schizophrenia (Howard et al., 2005). While genetic fate mapping techniques show that virtually all chandelier cells are generated in the MGE (Xu et al., 2008), transplantation experiments demonstrate that there is a strong bias for the production of these cells by ventral MGE progenitors at late stages of neurogenesis (Inan et al., 2012). These findings were further confirmed using a tamoxifen-dependent lineage tracing strategy showing that chandelier cell production by Nkx2.1-positive progenitors peaks at E16.5 (Taniguchi et al., 2013). Taken together, these findings suggest that cortical interneuron diversity may stem from the heterogeneity in progenitor populations found in the embryo. The mechanisms of interneuron lamination in the neocortex have also been studied using transplantation techniques (Pla et al., 2006). While the secretion of RLN by

3 Transplantation and the study of brain development

Cajal Retzius cells (D’Arcangelo et al., 1995; Ogawa et al., 1995; Soriano et al., 2005) is required for the proper lamination of the neocortex (Caviness, 1982), homotopic and isochronic transplants of MGE cells lacking the intracellular adaptor Dab1 required for RLN signaling revealed normal lamination of the mutant MGE cells, suggesting that cortical interneuron lamination does not depend on cell-autonomous RLN signaling. By contrast, WT MGE cells that are transplanted in the MGE of Dab1-deficient embryos fail to adopt a normal lamination and display a distribution that highly correlates that of misplaced pyramidal cells. Considering that interneurons invade the cortical plate after pyramidal cells have reached their final position and that synchronically born interneurons and pyramidal cells tend to locate in the same cortical layers, this work provides evidence that interneurons laminate in the cortex using cues presented by synchronically born pyramidal cells (Pla et al., 2006). This notion has been reinforced by (i) the aberrant lamination of both PV and SST interneurons in Fezf2-null mice that are lacking layer V corticofugal projection neurons (Lodato et al., 2011) and (ii) the recruitment of additional inhibitory synapses from PV interneurons by layer II–III callosal neurons converted into corticofugal projection neurons following Fezf2 overexpression (Ye et al., 2015). Nevertheless, the molecular nature of factors that govern interneuron positioning remains unknown. Finally, interneuron transplantation has been employed to address the rules governing waves of programmed cell death that occur in the developing CNS and help sculpt neural circuits (Southwell et al., 2012). Throughout the development of the nervous system, great numbers of neurons are eliminated at a time that coincides with synaptogenesis (Dekkers et al., 2013). In the case of mouse cortical interneurons, this occurs at around P7. While the role played by supernumerary cells during CNS development is still unknown, it was widely accepted that such a selection is driven by limited trophic support for which developing neurons would have to compete: the so-called neurotrophin hypothesis (Hamburger and Levi-Montalcini, 1949; LeviMontalcini, 1949). If we were to extend this notion to MGE transplants, the host brain should thus be able to only accommodate a finite number of transplanted interneurons. In contrast, transplantation studies strongly suggest that interneuron survival is independent from signals arising from the host (Southwell et al., 2012). Upon transplantation in the neonatal brain, MGE cells display developmental apoptosis coinciding with their own age, and not that of the host, and therefore asynchronously from endogenous interneurons (Fig. 2). Interestingly, the proportion of interneurons undergoing cell death remained constant across grafts of various sizes and was similar to the extent of cell death found among endogenous interneurons during normal development. Additionally, transplanted interneuron cell death was found to be independent of the neurotrophin receptor TrkB and was temporally recapitulated by cultured MGE cells in vitro. Altogether this work indicates that interneuron developmental programmed cell death is intrinsically determined and that interneuron survival does not depend on extrinsic cues from the host cells. However, it is possible that transplanted interneuron survival could depend on competition for survival signals emanating from interneurons themselves. Self-regulated cell death, at the

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CHAPTER 3 Transplantation of GABAergic interneurons for cell-based therapy individual cell level or at the population level, offers unique advantages for interneuron transplantation as the number of surviving transplanted cells is not adjusted with respect to the population of host interneurons, but mostly by the number of transplanted cells themselves. This work suggests that grafted cells execute their own endogenous program to determine the timing of cell death and the final number of surviving interneurons.

4 TRANSPLANTATION AND CORTICAL PLASTICITY In recent years, interneuron transplantation has been used to induce cortical plasticity in mature animals that normally exhibit minimal plasticity (Fig. 3) (Davis et al., 2015; Isstas et al., 2016; Larimer et al., 2016; Southwell et al., 2010; Tang et al., 2014). This line of work has reinforced the importance of the intrinsic programs that govern interneuron development and their integration upon transplantation. The model chosen for this work is the mouse visual system that displays a developmental critical period of plasticity for ocular dominance during which thalamic afferents compete for space and synaptic strength in the binocular zone of the primary visual cortex (Espinosa and Stryker, 2012; Wiesel and Hubel, 1963). During the critical period, but neither before nor after, imbalancing visual inputs by suturing one eye induces a shift in cortical responsiveness in favor of the open eye (Fagiolini et al., 1994; Prusky and Douglas, 2003). The induced rewiring of intracortical connectivity eventually leads to a loss of visual acuity for the closed eye, which mimics amblyopia, a condition found in humans and that affects 4% of the population (Hensch, 2005). The opening of the critical period is governed by the maturation of cortical GABAergic circuits (Di Cristo et al., 2007; Fagiolini and Hensch, 2000; Hanover et al., 1999; Hensch et al., 1998; Iwai et al., 2003; Kanold et al., 2009; Katagiri et al., 2007; Sugiyama et al., 2008). Accordingly, plasticity can be triggered ahead of time by promoting interneuron development (Di Cristo et al., 2007; Hanover et al., 1999; Sugiyama et al., 2008) or by pharmacologically enhancing inhibitory transmission in the visual cortex (Fagiolini and Hensch, 2000; Fagiolini et al., 2004). Interestingly, heterochronic transplantation of immature interneurons in the neonatal (Southwell et al., 2010) or adult (Davis et al., 2015) brain induces a second period of plasticity in the recipient visual cortex (Fig. 3). In both studies monocular deprivation was found to induce ocular dominance plasticity in MGE transplant recipients only if performed 5 weeks after the transplantation of E13.5 MGE cells, well after the endogenous critical period has ended (Fig. 3). Interestingly, the age of the transplanted cells at the time of transplant-induced plasticity corresponds to that of host interneurons when endogenous ocular dominance plasticity reaches its maximum, suggesting that interneuron intrinsic developmental programs regulate critical period timing. As MGE grafts primarily generate PV and SST interneurons, Tang et al. (2014) sought to determine the respective contribution of these two interneuron populations in MGE transplant-induced plasticity. Using the R26-GDTA allele that allows ablation of specific cell types through Cre-dependent diphtheria toxin alpha subunit

4 Transplantation and cortical plasticity

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FIG. 3 Immature interneuron transplantation and therapeutic applications. (Top) Immature interneurons can be obtained directly from the embryonic MGE or in vitro from embryonic stem (ES) or induced pluripotent stem (IPS) cells directed to differentiate into MGE-like progenitors. Interneurons have been transplanted into multiple regions of the CNS, including the striatum, neocortex, hippocampus, and spinal cord. Transplanted interneurons display disease-modifying activity in animal models of Parkinson’s disease, Alzheimer’s disease, epilepsy, schizophrenia, anxiety, spasticity, chronic pain, and neuropathic itch. (Bottom) Interneuron transplantation has also been used to study and manipulate cortical plasticity. The timing of native critical period of plasticity in the mouse visual cortex is dictated by the maturation of endogenous interneurons. Ocular dominance plasticity peaks at around P30 when inhibitory neurons are approximately 35 days of age (35D). Upon transplantation into both neonatal and adult visual cortex, interneurons induce ocular dominance plasticity when they reach a similar cellular age at approximately 35 days after transplantation (35DAT). Transplant-induced plasticity allows functional recovery of visual acuity in mouse models of developmentally acquired amblyopia. These findings suggest that interneuron development is governed by molecular programs established in the embryo and that these programs are retained and executed by embryonic interneurons upon heterochronic transplantation.

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CHAPTER 3 Transplantation of GABAergic interneurons for cell-based therapy expression, PV and/or SST cells were specifically eliminated from MGE transplants. Surprisingly, PV- and SST-depleted transplants each induced robust plasticity, thus indicating that both PV and SST interneurons can induce ODP. However, elimination of both PV and SST cells prevented transplant-induced plasticity, suggesting that a sufficient number of either cell type has to be present for MGE transplants to induce plasticity. It has been suggested that PV interneurons are a central hub for cortical plasticity gating (Bernard et al., 2016; Beurdeley et al., 2012; Chattopadhyaya et al., 2004; Fagiolini et al., 2004; Huang et al., 1999; Kuhlman et al., 2013; Maffei et al., 2006; Miyata et al., 2012; Pizzorusso et al., 2002; Spatazza et al., 2013; Sugiyama et al., 2008; Takesian and Hensch, 2013; YazakiSugiyama et al., 2009). However, the role of SST interneurons in this system had been overlooked so far. The work by Tang et al. (2014) does not preclude the possibility that the induction of plasticity by transplanted SST interneurons may be mediated in part by their action on host PV cells. The demonstration that SST interneurons can also induce plasticity highlights the power of transplantation experiments to decipher the cellular mechanisms of plasticity. The ability of SST neurons to induce cortical plasticity raised the question as to whether interneurons are in general capable of reopening sensory critical periods. To test this hypothesis, recent studies tested whether CGE-derived interneurons might be competent (Davis et al., 2015; Isstas et al., 2016; Larimer et al., 2016). The CGE gives rise to a pool of interneurons different from those originating in the MGE (Rudy et al., 2011). Just like MGE transplants, grafted CGE cells disperse and differentiate in the heterochronic environment. However, in contrast to the MGE, CGE-derived interneurons do not reactivate plasticity (Davis et al., 2015; Isstas et al., 2016; Larimer et al., 2016) despite being functionally integrated in the host brain (Larimer et al., 2016). One study found that PV and SST interneurons account for 20% of CGE transplant-derived cells (Larimer et al., 2016), suggesting that some young interneurons generated in the MGE migrate through the CGE (Butt et al., 2005). Such mixed transplants were found to reopen plasticity, although plasticity induction is solely attributable to PV and SST interneurons as it is fully abolished following genetic ablation of these cells from CGE transplants (Larimer et al., 2016). Taken together, these results suggest that transplant-induced plasticity is restricted to MGE-derived PV and SST interneurons. It is unlikely that transplantinduced plasticity results from increased inhibition as pharmacological enhancement of inhibition does not trigger plasticity after the critical period (Fagiolini et al., 2004). The inability for CGE-derived neurons to induce plasticity also suggests that the mechanisms required to home the transplanted cells into the host cortical network are not enough to elicit functional reorganization. Endogenous critical period closure has been associated with the expression of molecular brakes that stabilize mature cortical networks (Bavelier et al., 2010; Morishita et al., 2010; Sajo et al., 2016). Transplanted PV and SST cells may alter the expression of such brakes and thus allow host cells to rewire upon sensory deprivation. Undoubtedly, the study of both MGE and CGE transplants provides a powerful new tool to investigate the molecular

5 Disease-modifying properties of MGE transplants

and synaptic mechanisms enabling transplant-induced plasticity. The induction of plasticity by heterochronically transplanted interneurons also offers an opportunity to modify neural circuits for clinical gains.

5 DISEASE-MODIFYING PROPERTIES OF MGE TRANSPLANTS Removing plasticity brakes in adulthood holds great promises for lifelong learning and the treatment of neurodevelopmental disorders (Bavelier et al., 2010; Takesian and Hensch, 2013). Studies of transplant-induced cortical plasticity suggest that young interneuron transplantation could be employed to promote the functional reorganization of cortical networks following brain injury or trauma. While this hypothesis remains to be tested, it is supported by recent results showing that MGE transplantation in the adult visual cortex rescues amblyopia acquired upon visual deprivation in juvenile mice (Davis et al., 2015). Whether the plasticity-inducing effect of MGE transplants can be similarly applied to brain regions other than the visual cortex remains unknown. For example, the basolateral amygdala displays a critical period of plasticity during which fear memories can be erased by extinction training (Gogolla et al., 2009). Importantly, plasticity in the amygdala shares many features with that in the visual cortex, suggesting that common molecular and cellular determinants may be gating plasticity in both systems. Most notably, amygdala plasticity is constrained by the expression of perineuronal nets (Gogolla et al., 2009) and can be reactivated by antidepressant drugs (Karpova et al., 2011). It will be interesting to investigate whether interneuron transplantation can modify fear memory resiliency, which bears strong therapeutic implications for patients suffering from posttraumatic stress disorders. Interestingly, MGE cell transplants have been shown to reduce anxiety levels in WT animals (Valente et al., 2013). Interneuropathies constitute a wide range of neurological disorders that directly result from interneuron dysfunctions (Kato and Dobyns, 2005). Whether they are caused by a reduction in interneuron number or more specific deficits in the firing properties of individual neurons, these syndromes share impaired GABAergic transmission (Kato and Dobyns, 2005; Marı´n, 2012). Other conditions such as PD, Huntington’s disease, or neuropathic pain originate from imbalance between excitation and inhibition levels secondary to defects in other neuronal populations (Kato and Dobyns, 2005; Marı´n, 2012; Southwell et al., 2014). These diseases are all associated with network hyperexcitability, which led to the proposal that interneuron transplantation could be used to restore inhibition and thus alleviate the symptoms observed in animal models of these conditions (Fig. 3).

5.1 SCHIZOPHRENIA Schizophrenia has been associated with impaired GABA signaling (Inan et al., 2013; Marı´n, 2012). Injection of the NMDA receptor antagonist phencyclidine (PCP) triggers schizophreniform cognitive deficits in both healthy humans

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CHAPTER 3 Transplantation of GABAergic interneurons for cell-based therapy (Javitt and Zukin, 1991) and rodents (Mouri et al., 2007). PCP is thought to primarily act on NMDA receptors localized to cortical interneurons (Korotkova et al., 2010), which would result in altered activity of projection neurons in the prefrontal cortex (PFC). In order to address whether cortical interneurons could potentially help in the treatment of schizophrenia-related symptoms, MGE transplants were performed in the neonatal mouse brain 6 weeks before PCP acute administration (Tanaka et al., 2011). Interestingly, PFC transplants can prevent PCP-induced cognitive deficits, as opposed to visual cortex transplants that are ineffective in this system. Immediate early gene expression in PFC projection neurons was increased in MGE recipients, suggesting that MGE transplant benefits on PCP-induced deficits are not simply linked to modulation of PFC neuron activity. Disinhibition in the hippocampus is also thought to be determining schizophrenia-related positive symptoms (e.g., delusions and hallucinations) and psychosis (Heckers and Konradi, 2010; Schobel et al., 2013). To study this aspect of the disease, Gilani et al. (2014) used the cyclin D2 (CCND2) genetic mouse model (Glickstein et al., 2007) that displays a reduction in hippocampal interneurons and increased hippocampal output in vivo. Interestingly, MGE transplants can restore normal hippocampal activity and rescue the hippocampus-related cognitive deficits exhibited by this model. These findings point to the importance of interneuron development and survival in the pathogenesis of psychotic disorders and demonstrate the procognitive effects of interneuron-based strategies, which could benefit treatment-resistant patients that demonstrate hippocampal hyperactivation at rest.

5.2 EPILEPSY Epilepsy is a heterogeneous neurological disorder affecting more than 50 million people and characterized by repeated episodes of seizure activity (de Boer et al., 2008). While known genetic mutations are responsible for a small proportion of cases (Pandolfo, 2013), epilepsy can also develop as a result of traumatic brain injury, stroke, tumor, or surgery (Chang and Lowenstein, 2003). Hyperexcitability is a key feature of epilepsies and often reflects impaired inhibition, in both animal models (Cossart et al., 2001; Sloviter, 1987) and human patients (de Lanerolle et al., 1989; Mathern et al., 1995). Seizure reduction following interneuron transplantation was first demonstrated in a genetic mouse model of epilepsy displaying severe spontaneous seizures by the second to third postnatal week of age (Baraban et al., 2009). MGE progenitors were transplanted in the neonatal cortex and seizure events were recorded by video EEG starting 30 days after transplantation. A 90% reduction in seizure events was observed in grafted mutant animals over the course of a month-long monitoring period. In another model of acquired epilepsy (Hammad et al., 2015), neonatal transplantation of MGE cells also yielded a significant decrease in the frequency and duration of epilepsy episodes, as early as 3 weeks posttransplantation. Concomitantly, transplantation seemed to promote survival of the mutant animals, as 80% of the MGE graft recipients survived up to 4 months compared to an average survival of 29 days for control animals. Taken together, these findings demonstrate that neonatal MGE grafts can have a prophylactic effect in two distinct congenital seizure disorders.

5 Disease-modifying properties of MGE transplants

MGE precursor cells also demonstrated efficacy in mouse models of induced epilepsy. Calcagnotto et al. (2010) injected MGE cells in the mouse neonatal cortex and employed maximum electroconvulsive shock (MES) in adult mice to address whether MGE grafts could protect against the induction of tonic seizures. MES acutely induces a single seizure and is often used as a platform to screen antiepileptic drugs. Upon MES induction at 2 months posttransplant, the incidence of tonic seizure was significantly lower in the MGE grafts group compared to controls. Accordingly, animal survival rate was also increased among transplant recipients compared to controls. Interneuron transplantation in the adult cortex also reduces seizure propagation as indicated by local field potential measurement upon focal administration of 4-aminopyridine (4-AP), a potent convulsant and potassium channel blocker (De la Cruz et al., 2011). Here, the beneficial impact of MGE grafts on epileptiform activity was detected as early as 2.5 weeks posttransplant and was also found to be independent of the extent of transplanted cell survival, which suggests that a critical amount of grafted cells is required for optimal tuning of neuronal inhibition (Southwell et al., 2010; Tang et al., 2014). Upon saporin-induced elimination of hippocampal interneurons, MGE transplantation restored inhibitory postsynaptic currents onto CA1 pyramidal cells and reduced pharmacologically induced seizure susceptibility of the grafted animals (Zipancic et al., 2010). The impact of interneuron transplantation on spontaneous recurrent seizures was also tested in the pilocarpine mouse model of temporal lobe epilepsy (Henderson et al., 2014; Hunt et al., 2013). In both studies, hippocampal MGE transplants were performed approximately 2 weeks after the animals reached status epilepticus. In the hippocampus of epileptic mice, grafted MGE cells differentiate into GABAergic interneurons, acquire mature electrophysiological properties, and form functional synapses onto endogenous granule cells. MGE transplant-induced seizure suppression was found at 60 days after transplantation. Interestingly, prolonged video EEG monitoring of grafted animals showed that this effect did not persist in time despite the presence of functional transplanted cells (Henderson et al., 2014), which raises the question of the long-lasting effect of MGE grafts on the control of seizure phenotype. Taken together, these findings indicate that local interneuron precursor transplants have a potent impact on seizures. However, further work is required to determine the molecular and cellular mechanisms of transplant-induced seizure suppression in order to guide the development of transplant-based strategies for the treatment of refractory seizures. Inconsistencies in the timing of MGE graft efficacy suggest that those mechanisms may not be conserved across seizure models. For instance, while the kinetics of seizure suppression observed in the pilocarpine model strongly suggest a role for synaptic mechanisms (Henderson et al., 2014; Hunt et al., 2013), the early effects observed in the 4-AP model (De la Cruz et al., 2011) would indicate a possible role for nonsynaptic mechanisms. MGE grafts enhance both synaptic and extrasynaptic inhibition (Baraban et al., 2009). Interestingly, the requirement of extrasynaptic GABA-A receptors for the transplantmediated dampening of seizure propagation in the 4-AP model was recently shown

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CHAPTER 3 Transplantation of GABAergic interneurons for cell-based therapy (Jaiswal et al., 2015). Given the heterogeneity of MGE transplant-derived interneurons, it will be important to identify whether specific subtypes may prove therapeutic for specific forms of seizures.

5.3 PARKINSON’S DISEASE Interneuron transplantation as a cell-based therapeutic strategy has also been tested in a mouse model of PD (Martı´nez-Cerden˜o et al., 2010). PD affects a large population worldwide and is characterized by motor impairments as well as cognitive and autonomic dysfunctions. The motor symptoms result from the degeneration of substantia nigra pars compacta dopaminergic neurons that normally extend axonal projections to the striatum. Reduced dopamine release in the striatum induces a cascade of neurotransmitter release imbalance that inhibits the output of the basal ganglia and leads to motor dysfunctions (DeLong and Wichmann, 2007). Striatal GABAergic interneurons have been shown to gate basal ganglia output (Tepper and Bolam, 2004) and, consequently, worsen striatal imbalance in the dopaminedepleted striatum (Mallet et al., 2006), thus contributing to the pathophysiology of PD. These findings indicate that striatal inhibition can be used as a nondopaminebased lever to alleviate some of the symptoms characteristic of PD. The 6-hydroxydopamine (6-OHDA) rat model recapitulates the deterioration of the nigrostriatal pathway observed in PD patients. MGE progenitors transplanted into the striatum of 6-OHDA-treated adult rats are able to disperse, differentiate into GABAergic interneurons, synaptically integrate locally, and survive for up to a year (Martı´nez-Cerden˜o et al., 2010). Importantly, MGE grafts were able to dampen the motor deficits displayed in this animal model of PD. Of note, overall locomotor activity of naı¨ve control animals was increased following striatal MGE transplantation, thus suggesting that added interneurons can also exert a strong influence on striataldependent behaviors in an intact environment. The mechanisms of MGE cell effects in this system remain unknown and further work will be required for their identification. Previous studies in rodents have reported amelioration of PD-associated symptoms upon modulation of basal ganglia activity by enhancement of GABA stimulation (Lee et al., 2005; Luo, 2002; Winkler et al., 1999). Accordingly, motor deficits were likely improved as a consequence of transplant-induced increase in striatal inhibitory transmission. Alternatively, it is plausible that interneuron transplantation induces secondary changes mediating behavioral improvements or provides exogenous trophic support to remaining dopaminergic processes in the striatum. Finally, 25% of MGE transplant-derived cells differentiate into oligodendrocytes in the striatum, which raises the possibility of additional nonneuronal trophic support.

5.4 ALZHEIMER’S DISEASE Approximately 40 million people are affected by AD, a number that is predicted to triple by 2050 (Wimo et al., 2013). Memory deficits in patients suffering from AD are associated to an excitation–inhibition imbalance in the dentate gyrus that leads to

5 Disease-modifying properties of MGE transplants

hippocampal hyperactivity (Huang and Mucke, 2012). While amyloid-b (Ab) overproduction or accumulation leads to interneuron dysfunction (Palop et al., 2007; Verret et al., 2012), the expression of apolipoprotein (apo) E4, a strong genetic risk factor for AD, causes hippocampal hyperactivity in humans (Filippini et al., 2009) and a progressive decrease in hilar interneuron number in mice (Andrews-Zwilling et al., 2010; Leung et al., 2012; Li et al., 2009). As a result of aberrant neural network activity, AD patients have also been found to display increased incidence of epileptic events (Amatniek et al., 2006). Given the GABAergic dysfunctions observed both in patients and in mouse models of AD, it was tested whether interneuron replacement therapy leads to improvement of both cognitive and behavioral deficits in two widely used AD mouse models (Tong et al., 2014). Bilateral MGE transplantation was performed in the hilus of aged apoE4 knock-in mice. Transplanted cells were found to disperse throughout the hilus, extend dendrites into the molecular layer of the dentate gyrus, predominantly differentiate in GABAergic interneurons expressing SST, and survive for at least 90 days. Grafted cells functionally integrated and increased inhibitory transmission onto excitatory granule cells, thus compensating for the reduction of hilar interneurons in these mice. MGE cell transplantation rescued learning and memory deficits in apoE4 knock-in mice, both with and without Ab plaques, to levels similar to those of wild-type mice. These findings suggest that interneuron replacement could ameliorate AD-related symptoms. The lack of significant modification of Ab levels and plaques suggests that cognitive improvements are mediated by restored synaptic inhibition, although trophic support emanating from the transplanted cells onto remaining hilar interneurons cannot be ruled out. In this context it would be interesting to test whether early wild-type interneuron transplantation could have a prophylactic effect on interneuron decrease in apoE4 knock-in mice. Cell-based strategies for the treatment of AD appear as a viable approach given that wild-type MGE transplant-derived cells can survive in the apoE4-Ab toxic environment.

5.5 NEUROPATHIC PAIN Neuropathic pain is provoked by nerve injury and is characterized by both allodynia (where nonnoxious stimuli are painful) and hyperalgesia (where pain behaviors caused by normally painful stimuli are increased). While the complex molecular, biochemical, and cellular changes that occur upon nerve injury are central for both mechanical and thermal hypersensitivity, how they contribute to the pain described by patients is not fully resolved. It is accepted that peripheral nerve injury results in decreased GABAergic neurotransmission within the spinal cord dorsal horn. A loss of inhibitory interneurons has been reported in the lesioned spinal cord (Moore et al., 2002; Scholz et al., 2005), as well as the reduced expression of postsynaptic GABAA receptors (Fukuoka et al., 1998), decreased GABA release (Lever et al., 2003), and diminished glutamic acid decarboxylase (GAD) expression (Eaton et al., 1998; Lever et al., 2003). In agreement with the long-standing idea that disinhibition in the spinal dorsal horn could underlie neuropathic pain (Loeser and Ward, 1967), GABA agonists have been found to improve allodynia and hyperalgesia (Munro et al., 2009).

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CHAPTER 3 Transplantation of GABAergic interneurons for cell-based therapy A recent set of studies explored whether transplantation of MGE cells in the spinal cord could mitigate the behavioral features of two mouse models of neuropathic pain (see also chapter “Rebuilding CNS inhibitory circuits to control chronic neuropathic pain and itch” by Braz et al.). Transplanted MGE cells can survive in the adult spinal cord for at least 6 months, differentiate into GABAergic interneurons that display a “cortical” signature, and functionally integrate within the host spinal circuitry. Importantly, mechanical responsiveness is returned to baseline levels following MGE transplantation in mouse models of both sciatic nerve lesion where animals develop a severe hypersensitivity (Bra´z et al., 2012) and chemotherapy-induced neuropathic pain (Bra´z et al., 2015). In the latter, MGE cells deficient for the vesicular GABA transporter (VGAT) failed to rescue the pain behavior, which suggests that synaptic GABA release is important for MGE transplant-induced behavioral improvements in the paclitaxel model. As observed in other studies (De la Cruz et al., 2011; Southwell et al., 2010; Tang et al., 2014), there was no correlation between transplanted interneuron number and functional effect (Bra´z et al., 2012). Taken together, this work highlights the efficacy of MGE cell transplants for the management of neuropathic pain. Considering the known side effects of the traditional pharmacological approaches used in patients, interneuron-based therapy may thus represent a promising avenue for future therapeutic strategies.

6 CONCLUSION The ontogeny of ventral telencephalon interneurons undoubtedly endows this cell population with the remarkable ability to disperse upon transplantation in the neonatal, juvenile, or even adult brain. Following migration, grafted interneurons have the ability to differentiate and synaptically integrate into functional circuits. Grafted interneurons can modify the activity of host target cells, which has strengthened the potential of interneuron-based therapies for the treatment of various conditions characterized by hyperexcitability. While the numerous preclinical studies discussed here offer promises for such approaches to be taken out of the laboratory and into the clinic, specific limitations still remain to be addressed. Notably, the mechanisms by which MGE transplants control target cell activity need further investigation. Moreover, a better understanding of each disease’s etiology will be required so that the composition of the transplants may be adapted to achieve optimal efficacy. The generation of safe human interneurons will also be crucial for clinical transition. Recent reports demonstrating the in vitro production of transplantable MGE-like interneurons from human pluripotent stem cells (Maroof et al., 2013; Nicholas et al., 2013) have generated great excitement, which was further reinforced by studies showing therapeutic efficacy for such cells in mouse models of disease (Cunningham et al., 2014; Fandel et al., 2016; Liu et al., 2013). Fundamental knowledge has also been gained from the remarkable ability of young interneurons to disperse and functionally integrate upon heterochronic transplantation. Here too, we anticipate that transplantation of interneurons will continue

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revealing basic insights about how these neurons find their way through complex and heterogeneous environments, how they choose to make connections with other neurons, and how they determine whether they survive or die. Additionally, the ability of transplanted interneurons to induce juvenile-like plasticity offers a powerful tool to study basic mechanisms of critical periods of plasticity. Future work may also reveal what cell-intrinsic information within young interneurons endows them with their unique ability to migrate through the parenchyma of the postnatal brain.

ACKNOWLEDGMENTS We would like to thank Marianna Di Lullo and Shawn F. Sorrells for their careful reading of this manuscript and useful comments. We apologize to the authors of many additional relevant papers we could not cite here due to space constraints. Work in the Alvarez-Buylla’s laboratory is supported by the NIH (EY025174, NS028478, HD032116) and the John G. Bowes Research Fund. A.A.B. is the Heather and Melanie Muss Endowed Chair of Neurological Surgery at UCSF. A.A.B. is on the scientific advisory board and is co-founder of Neurona Therapeutics.

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Rebuilding CNS inhibitory circuits to control chronic neuropathic pain and itch

4

Joao M. Braz*, Alex Etlin*, Dina Juarez-Salinas*, Ida J. Llewellyn-Smith†, Allan I. Basbaum*,1 *University of California—San Francisco, San Francisco, CA, United States Cardiovascular Medicine, Human Physiology and Centre for Neuroscience, Flinders University, Bedford Park, SA, Australia 1 Corresponding author: Tel: +1-415-476-5270; Fax: +1-415-476-5270, e-mail address: [email protected]

†

Abstract Cell transplantation offers an attractive alternative to pharmacotherapy for the management of a host of clinical conditions. Most importantly, the transplanted cells provide a continuous, local delivery of therapeutic compounds, which avoids many of the adverse side effects associated with systemically administered drugs. Here, we describe the broad therapeutic utility of transplanting precursors of cortical inhibitory interneurons derived from the embryonic medial ganglionic eminence (MGE), in a variety of chronic pain and itch models in the mouse. Despite the cortical environment in which the MGE cells normally develop, these cells survive transplantation and will even integrate into the circuitry of an adult host spinal cord. When transplanted into the spinal cord, the cells significantly reduce the hyperexcitability that characterizes both chronic neuropathic pain and itch conditions. This MGE cell-based strategy differs considerably from traditional pharmacological treatments as the approach is potentially disease modifying (i.e., the therapy targets the underlying etiology of the pain and itch pathophysiology).

Keywords Cell transplant, Spinal cord, Pain, Itch, Dorsal horn, GABA, Medial ganglionic eminence

1 INTRODUCTION The management of chronic pain and itch relies predominantly on pharmacotherapy. Depending on the etiology of the chronic pain condition, e.g., whether it is associated with tissue injury and inflammation or develops after frank damage to peripheral or CNS neurons, relief can be achieved by blocking transmission of the pain message (e.g., local anesthetics), increasing inhibitory controls, Progress in Brain Research, Volume 231, ISSN 0079-6123, http://dx.doi.org/10.1016/bs.pbr.2016.10.001 © 2017 Elsevier B.V. All rights reserved.

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particularly at the spinal cord (e.g., opioids, serotonin–norepinephrine reuptake inhibitors, gabapentinoids) or decreasing peripheral sensitization (e.g., antiinflammatory drugs). An alternative is to focus on the biochemical and molecular basis of the CNS sensitization process that exacerbates pain, produces hypersensitivity and pain in response to normally innocuous stimuli (allodynia) and likely contributes to persistence of the pain (chronification). Recent studies, in fact, have identified over a hundred molecules (from receptors to second messenger systems, integrins, etc.) that contribute to this central sensitization process (Basbaum et al., 2009; Bra´z et al., 2014b; Julius and Basbaum, 2001). Of particular interest are preclinical studies suggesting that selective targeting of some of these molecules can be effective in reducing ongoing pain or preventing chronic pain from developing in the first place. Unfortunately, the great majority of these molecules participate in biochemical processes throughout the nervous system. As a result, attempts to translate these preclinical findings to patients are often plagued by adverse side effects that limit the doses that can be used clinically. In many respects, the same limitations exist in the management of chronic itch. Most importantly, although antihistamines are most commonly prescribed, many chronic itch conditions are completely unresponsive to these drugs (Silverberg et al., 2016). Recent studies have identified many different pruritogens, including several that are histamine independent (Cevikbas et al., 2011; LaMotte et al., 2014); but once again therapy is often associated with significant adverse side effects, particularly when the pharmacotherapeutics are administered systemically. Clearly, new approaches to the management of persistent pain and itch are needed. Here, we describe a very different strategy, one that attempts to repair a dysfunctional nervous system (rather than mitigating its symptoms) using transplantation of precursors of inhibitory interneurons. Cell transplantation is a very attractive alternative to pharmacotherapy for central nervous system disorders. Specifically, transplants provide a continuous, local delivery of therapeutic compounds, which avoids many of the adverse side effects associated with systemically administered drugs. Most importantly, if the transplants integrate into the central circuitry of the host, then the therapeutic molecules can be targeted to specific circuits, rather than flooding the system, as occurs with an implanted pump. Given that one of the major features of chronic pain (Sandkuhler, 2007; Woolf and Salter, 2000) and/or itch (Cevikbas et al., 2011; Ross, 2011) is increased excitability of spinal cord dorsal horn neurons, cell transplants that can locally deliver a relevant pharmacotherapy (e.g., an inhibitory neurotransmitter) provides an optimal strategy to decrease the chronic pain- or itch-induced hypersensitivity that characterizes both conditions. Among the earliest approaches using transplants were studies by Sagen and colleagues (Hentall and Sagen, 2000; Sagen et al., 1990), who reported long-term pain relief in preclinical studies after intrathecal delivery of adrenal chromaffin cells (Eaton et al., 1999; Michalewicz et al., 1999; Yu et al., 1998). Clinical trials in patients with cancerrelated pain using human adrenal medullary tissue, although very preliminary, were encouraging (Winnie et al., 1993). That approach, however, involved transplants that

3 MGE cell transplants to treat neuropathic pain

delivered compounds (norepinephrine and enkephalin) diffusely, into cerebrospinal fluid (CSF). For this reason, the approach is more comparable to an intrathecal pump. In this chapter, we describe experiments in which compromised spinal cord circuits can be restored by transplantation of precursors of GABAergic interneurons. The cells integrate into host circuits and can significantly ameliorate several chronic pain and itch conditions (Basbaum and Bra´z, 2016).

2 MEDIAL GANGLIONIC EMINENCE-DERIVED INHIBITORY INTERNEURONS Most, if not all inhibitory interneurons in the cerebral cortex derive from two regions of the ventral telencephalon (Marin and Rubenstein, 2001): the caudal (CGE) and medial (MGE) ganglionic eminence. During early development, MGE cells migrate dorsally into the cerebral cortex, where they mature into different morphological subtypes of interneuron. Seventy percent of these interneurons express somatostatin and/or parvalbumin, two markers of cortical GABAergic neurons (Wichterle et al., 1999). When transplanted into neonatal mouse cortex, embryonic MGE cells migrate extensively and differentiate into GABAergic interneurons (Alvarez-Dolado et al., 2006), integrate into host neuronal circuitry, and inhibit host neurons. In fact, Calcagnotto et al. (2010) demonstrated that MGE-derived interneurons integrate into host hippocampal circuitry, even when a pathological environment is created by chemical ablation of host hippocampal inhibitory interneurons. Most importantly, by replacing the ablated GABAergic interneurons, the MGE transplants restored normal hippocampal synaptic inhibition. In another study, seizure reduction in a mouse epilepsy model was attributed to a selective increase of GABAA-mediated inhibition of neurons in the transplanted cortical tissue (Baraban et al., 2009). As there are important similarities between seizures in epilepsy, a consequence of a hyperexcitable cerebral cortex after loss of inhibition, and the neuropathic pain and itch phenotypes, which also arise from hyperexcitability, it is not surprising that both are managed by anticonvulsants. The commonality in the treatment modality provided a strong rationale for our series of studies that assessed the utility of MGE cells, which are progenitors of GABAergic inhibitory interneurons, to “treat” chronic pain and itch. Our basic premise is that neuropathic pain and itch are not symptoms of some other neurological disease. Rather these conditions constitute neurological diseases in and of themselves.

3 MGE CELL TRANSPLANTS TO TREAT NEUROPATHIC PAIN Peripheral nerve injury-induced neuropathic pain is a major clinical problem involving altered spinal cord processing of pain messages (Gilron et al., 2015). Among these changes is a loss of inhibition, which leads to an increased excitability (hypersensitivity) of spinal cord dorsal horn neurons. These changes contribute not only to

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the development of spontaneous pain, but also to the mechanical allodynia, i.e., pain provoked by normally innocuous mechanical stimulation, which characterize these conditions. In the clinical setting, mechanical allodynia is particularly problematic, significantly reducing quality of life in these patients. There is now considerable evidence that neuropathic pain results from a loss of GABAergic inhibitory control, particularly at the level of the spinal cord. For example, release of GABA in the spinal cord dorsal horn is reduced after chronic constriction or traumatic periphery nerve injury (Moore et al., 2002). The expression of the GABA synthesizing enzyme, glutamic acid decarboxylase (GAD), is also decreased after partial peripheral nerve injury (Bra´z et al., 2012), resulting in a loss of inhibitory tone. Others have reported decreased GABA-mediated inhibition due to nerve injuryinduced degeneration of GABAergic spinal interneurons (Scholz et al., 2005; Sugimoto et al., 1990), reduced primary afferent input to GABAergic dorsal horn neurons (Kohno et al., 2003; Polga´r and Todd, 2008), downregulation of GABAergic receptors (Castro-Lopes et al., 1993; Eaton et al., 1998; Fukuoka et al., 1998; Ibuki et al., 1997), or a microglial-derived and BDNF-mediated shift in the chloride gradient in dorsal horn “pain” transmission neurons (De Koninck, 2007). The consequence of each of these events is increased spontaneous firing of dorsal horn neurons and responsiveness of “pain” transmission neurons to normally innocuous stimuli. Regardless of the origin of the decreased inhibition, pharmacological restoration or increase of GABAergic transmission unquestionably reduces both nerve injuryinduced hyperalgesia and allodynia (Malan et al., 2002). For example, profound analgesia can be achieved by activating spinal GABAA or GABAB receptors, in various models of inflammatory and neuropathic pain (Knabl et al., 2008; Yaksh et al., 2016; Zeilhofer et al., 2015). Pharmacological regulation of endogenous GABA control mechanisms, however, is not straightforward. For example, baclofen (a GABAB agonist), although previously used in the management of trigeminal neuralgia (Fromm et al., 1984) and although effective in many animal models of neuropathic pain (Hwang and Yaksh, 1997; Malan et al., 2002), has not proven useful for the treatment of neuropathic pain in patients and is often discontinued because of the side effect of sedation. Based on the reported utility of MGE cell transplants for the management of seizures, we asked whether MGE cell transplantation can restore spinal cord GABAergic inhibitory function, to a level that would ameliorate the neuropathic pain phenotype. We recognized that translating MGE cell transplants to the spinal cord faced several hurdles. First, the spinal cord environment for growth of the transplanted cells is completely different from the natural MGE cell environment, namely cerebral cortex. Second, although early transplant studies were performed in neonatal animals; it was not clear that the adult host would be as accepting of the transplants. Finally, because transplanted cells often migrate great distances, it was conceivable that even a successful spinal cord transplant might, over the long-term, result in migration of the cells to nontargeted areas, e.g., the contralateral spinal cord, distant spinal cord segments, and even supraspinal loci. In fact, none of the initial concerns materialized in our experiments.

3 MGE cell transplants to treat neuropathic pain

3.1 MGE CELLS AMELIORATE NEUROPATHIC PAIN We first examined the utility of the transplantation approach in the spared nerve injury (SNI) model of neuropathic pain in the mouse. In this model, two of the three branches of the sciatic nerve are transected, sparing the tibial nerve. After SNI surgery, mice rapidly develop a profound and long-lasting mechanical allodynia (Shields et al., 2003). Reflex withdrawal of the hindpaw to an innocuous mechanical stimulus, transmitted via the isolated tibial nerve, is readily induced. Normally, only noxious (painful) stimuli evoke hindpaw withdrawal. One day after the peripheral nerve injury, at which point the mechanical allodynia is manifest, we transplanted MGE cells isolated from mice in which GFP production was driven by the GAD67 promoter (Tamamaki et al., 2003) into the dorsal horn of the lumbar enlargement of the spinal cord (Bra´z et al., 2012, 2016). Our intent was to “treat” the area that receives inputs from the injured sciatic nerve. Control mice received only the medium that was used to transplant the cells. We then tested the mechanical threshold of transplanted and control mice weekly. In control mice, we found that the mechanical allodynia persisted for a month. In contrast, there was a gradual reduction of the SNI induced mechanical hypersensitivity in the MGE-transplanted mice, with an almost complete reversal by 4 weeks. Importantly, the recovery was not associated with reduced motor function, indicating that the transplants did not adversely influence ventral horn motoneurons. Next, we examined the therapeutic value of MGE cell transplants in a different neuropathic pain model, in which there is both mechanical and thermal (heat) hypersensitivity (Bra´z et al., 2015). In these experiments, we transplanted the MGE cells in a mouse model of chemotherapy (paclitaxel)-induced neuropathic pain, which is characterized by profound thermal as well as mechanical hypersensitivity (Flatters and Bennett, 2006; Smith et al., 2004). Paclitaxel is commonly used in cancer therapy but among its major adverse side effects is a severe neuropathic pain that is particularly prominent in the hands and feet (Lipton et al., 1989; Wasserheit et al., 1996). In our studies, systemic injection of paclitaxel induces bilateral, profound, and long-lasting (at least 4 weeks) mechanical allodynia and thermal hyperalgesia. As we observed following traumatic peripheral nerve injury, we recorded a significant reduction of the paclitaxel-induced hypersensitivity soon after transplantation (Fig. 1). Heat-evoked hindpaw withdrawal thresholds returned to baseline (preinjury) levels between the 1st and 2nd week posttransplantation. Mechanical thresholds also recovered, but the recovery was more gradual. Significantly, the transplant did not overcome the neuropathic pain phenotype in the contralateral hindlimb, which clearly indicates that the transplants exert a topographically localized recovery of inhibitory control. As expected, in control animals that received only medium (i.e., no MGE cells), the mechanical allodynia and thermal hyperalgesia persisted throughout the 4 weeks of observation. Importantly, in these studies, we also identified GABA as the critical neurotransmitter involved in the MGE-mediated therapeutic effects. To document the critical contribution of GABA, we transplanted MGE cells deficient for VGAT, the vesicular GABA transporter. These cells, which

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FIG. 1 Spinal cord transplantation of MGE cells reverses the thermal and mechanical hypersensitivity that develops in a model of chemotherapy (paclitaxel)-induced neuropathic pain. (A–B) Systemic administration of paclitaxel induces long-lasting and bilateral heat hyperalgesia (A) and mechanical allodynia (B). Both mechanical and heat hypersensitivity persisted for the 4-week observation period in animals that received an injection of transplant medium, without cells (control, black). In contrast, both mechanical and heat thresholds returned to baseline in animals that received an intraspinal transplant of the MGE cells (MGE, white). MGE cells deficient for the vesicular GABA transporter (VGAT), which can synthesize but not release GABA, were completely ineffective against both heat and mechanical hypersensitivity (MGE/VGAT, gray). This finding underscores the critical contribution of GABA to the restoration of inhibitory control. *P < 0.05; ***P < 0.001.

synthesize GABA but can no longer release it, survived after transplantation but were completely ineffective against both the mechanical and thermal hypersensitivity. Surprisingly, MGE transplants were not effective in a model of inflammatory pain (Bra´z et al., 2012). In these studies, we transplanted MGE cells into the lumbar spinal cord of adult mice; 4 weeks later the mice received an intraplantar injection of formalin, a chemical algogen that produces pain behaviors that mimic postoperative pain. We found that formalin-induced nocifensive behaviors (licking, biting) were not significantly different in medium- or MGE-transplanted mice. Because the pain behavior

3 MGE cell transplants to treat neuropathic pain

provoked by the formalin is short-lived, the studies actually assessed the prevention of inflammatory pain, rather than management of ongoing pain. Clearly, the transplants were not prophylactic in this tissue injury model, a finding that differs greatly from the surprising prophylactic effect that we observed in a neuropathic pain model. Although we need to assess different tissue injury-induced chronic pain models, the results in the formalin test suggest that MGE transplants may only be effective in models of nerve injury-induced neuropathic pain. Finally, it was of great interest to observe that when transplanted into naı¨ve, uninjured mice, MGE cells did not alter baseline mechanical thresholds. These results are in sharp contrast with the analgesic effects of administration of GABA agonists into the spinal CSF (Kaneko and Hammond, 1997), which readily increases baseline pain responses under normal conditions and reduces nocifensive behaviors in the formalin test. This difference also distinguishes transplantmediated restoration of inhibitory control from that produced by a pump that continuously releases GABA or a GABA mimetic in a circuit-independent manner.

3.2 MGE CELLS INTEGRATE EXTENSIVELY INTO HOST SPINAL CORD CIRCUITRY In parallel studies that combined anatomical, molecular, and electrophysiological approaches, we demonstrated that underlying the MGE-mediated behavioral improvement is a remarkable integration of the MGE transplants into the circuitry of the host spinal cord (Bra´z et al., 2012; Etlin et al., 2016). Indeed 4 weeks after transplantation, MGE cells not only survived but also differentiated into mature and functional GABAergic interneurons, extending long, ramified processes throughout the dorsal horn of the spinal cord. Note that the cells were transplanted prior to their assuming a neuronal phenotype (i.e., they were at a neuronal progenitor stage). Within 2 weeks of transplantation, however, we found that most GFP + MGE cells expressed the neuronal marker NeuN and were immunoreactive for GABA. MGE cells also expressed two markers of subpopulations of GABAergic interneurons: parvalbumin and neuropeptide Y (Laing et al., 1994; Rowan et al., 1993). Importantly, however, they did not express Pax2, a marker of spinal cord GABAergic interneurons (Maricich and Herrup, 1999), which suggests that the cells retain neurochemical features of their embryonic origin, namely the cortex. Consistent with this conclusion, we found that a significant percentage of the transplanted cells expressed somatostatin, a marker of cortical (Urban-Ciecko and Barth, 2016), but not spinal cord, GABAergic interneurons. In fact, somatostatin marks a subpopulation of excitatory interneurons in the spinal cord (Proudlock et al., 1993). We concluded that the spinal cord environment did not alter the neurochemical features of the transplants during their maturation. Furthermore, we noted that the morphology of the transplanted cells more closely resembles cortical rather than spinal cord GABAergic interneurons. Specifically, the MGE cells have round cell bodies with extensively ramified dendrites that extend in all directions, a morphology very different from the majority of spinal cord GABAergic interneurons that have fusiform cell bodies with dendrites that extend rostro-caudally (Bra´z et al., 2014a,b,c; Spike and Todd, 1992).

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Using transneuronal tracing approaches, we also demonstrated that the MGE transplants fully integrate into host spinal cord circuitry (Bra´z et al., 2012). Using a transgenic mouse developed in our laboratory (Bra´z et al., 2002), it is possible to trigger expression of the transneuronal tracer, wheat germ agglutinin (WGA), in a Cre-recombinase-dependent manner. Using this tracing method, we demonstrated that transplanted MGE cells receive inputs from most subtypes of primary afferents, including the subpopulation of small, nociceptive C-fibers that express the capsaicin-sensitive channel, TRPV1. Importantly, these presynaptic connections appear to be functional. Thus, MGE cells could readily be activated to express Fos in response to a peripheral noxious stimulus (hindpaw injection of formalin). Finally, using both WGA and the transneuronal, retrograde viral vector pseudorabies (Jasmin et al., 1997), we found that transplanted MGE cells established connections with a variety of spinal cord neurons, including projection neurons of the superficial dorsal horn that terminate in the parabrachial nucleus of the brainstem. Taken together, these studies suggested that MGE cells functionally integrate into the host spinal cord circuitry, including dorsal horn “nociceptive” circuits. We recognize, however, that there is not complete agreement as to the mechanism of transneuronal transfer of WGA from one neuron to another. Whether this involves synaptic communication could not be answered in our earliest studies. Therefore, to determine unequivocally that synaptic connections are formed between the host and the transplanted cells (and vice-versa), we turned to electrophysiology and electron microscopy.

3.3 FUNCTIONAL AND ANATOMICAL EVIDENCE FOR SYNAPTIC CONNECTIVITY OF TRANSPLANTED MGE CELLS Using a spinal cord slice preparation, we first showed that transplanted MGE cells have neuronal properties characteristic of inhibitory interneurons (Etlin et al., 2016). The cells readily discharged in response to intracellular injection of current and we recorded EPSCs from most MGE cells in response to electrical stimulation of an attached dorsal root. Based on conduction velocity and ability of the MGE cells to follow electrical stimulation of the dorsal root, we concluded that MGE cells receive both mono- and polysynaptic inputs from high threshold Ad- and C-fibers. Using an optogenetic approach involving light-evoked activation of TRPV1 primary afferent terminals that express channelrhodopsin (ChR2), we confirmed the presence of a primary afferent input. In a particularly revealing experiment, we transplanted MGE cells that express ChR2 into the dorsal horn (Fig. 2) and recorded evoked IPSCs in a small number of host neurons following light-evoked stimulation of the slice, which activated the MGE cells. We recorded responsive postsynaptic neurons throughout the superficial dorsal horn and in deeper laminae. Thus, the transplants receive primary afferent inputs and also influence dorsal horn host neurons. We also recorded postsynaptic responses to MGE-ChR2 cell activation in the presence of bicuculline, a selective GABA-A receptor antagonist. In all cases, the antagonist abolished or significantly reduced the evoked IPSCs, indicating that the GABA-A receptor mediates a major component of the therapeutic effect of transplants. Taken together, these

3 MGE cell transplants to treat neuropathic pain

FIG. 2 MGE cells inhibit host dorsal horn neurons. (A) The schematic illustrates a spinal cord slice from a mouse that underwent a partial sciatic nerve injury 7 days before transplantation of MGE cells that express channelrhodopsin (ChR2) and green fluorescent protein (GFP; Etlin et al., 2016). MGE/ChR2+/GFP+ and host neurons, from which patch clamp recordings were made, are denoted as green and black, respectively. In this configuration, a flash of blue light (460 nm, 20 ms) delivered through the microscope objective activates a large population of the MGE/ChR2+/GFP+ cells. (B) Optogenetically evoked IPSCs recorded in a host neuron after stimulation of MGE/ChR2+/GFP+ cells before (Control), after 5 min incubation in the GABA-A antagonist, bicuculline (Bic, 20 mM), and after a subsequent 20 min wash (Wash). The IPSC is abolished by bicuculline and partially recovers, indicating that the inhibitory effect of the transplant is mediated, at least in part, by a GABAergic action at the GABA-A receptor.

results provide strong evidence that MGE cells indeed establish functional GABAergic connections with the host circuitry. To establish conclusively that MGE cells make synaptic contacts with neurons of the host, we also processed spinal cord tissue with MGE transplants for electron microscopy (Fig. 3). Five to six weeks after transplantation, we identified extensive synaptic interactions between GFP-immunoreactive dendrites derived from the MGE cells and host spinal cord elements (Bra´z et al., 2014c) that are presynaptic to the MGE cells. Host inputs to MGE-derived neurons included putative primary afferent terminals, which had a characteristic scalloped appearance, and made multiple asymmetric synaptic contacts with both host dendrites and with dendrites that derive from the transplanted cells. We also found many examples in which axon terminals that derive from the transplanted cells formed synapses with host dendrites and cell bodies. Of particular interest is that the synaptic connectivity of MGE cells transplanted into the dorsal horn is dynamic and evolves over time. For example, at 6 months posttransplantation, not only were the transplanted neurons still readily detected in the spinal cord, but also there was a doubling of the number of host synapses onto GFP-positive MGE-derived dendrites. These findings demonstrate that there is remarkable and ongoing remodeling of host synapses in response to the transplant. With a view both to assess the GABAergic neurochemistry of the MGE cells and to examine possible interactions between host GABAergic neurons and the transplanted cells, we also combined immunoperoxidase labeling of the transplanted cells with immunogold localization of GABA-immunoreactivity. These studies

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FIG. 3 Synaptic integration of transplanted MGE cells into host circuitry. (A) Light microscopic image illustrates transplant-derived GFP-immunoreactive neurons throughout the substantia gelatinosa (SG) and in deep dorsal horn. (B) Electron microscopic image illustrates a host axon terminal (AH) that forms a symmetric synapse (arrowheads) on a GFP-immunoreactive, transplant-derived dendrite (DT). Scale bar 50 mm in A and 500 nm in B.

documented synaptic contacts between immunogold-labeled, host GABAergic interneurons, and GFP-immunoreactive MGE cells, whose terminals were also shown to be GABA-immunoreactive. This synaptic arrangement demonstrates that the transplants are under inhibitory control from the host. Somewhat unexpectedly, we also found that MGE cells are synaptically connected, i.e., communicate with one another. These latter findings support our contention that the transplanted cells, rather than recapitulating the normal organization of host GABAergic interneurons, make random synaptic connections. We assume that only a subset of the new connections are functional, depending on the target cell as well as the nature of the receptors expressed upon neurons that receive inputs from the transplanted cells. We speculate that the input from host neurons to MGE cells is also random, but that remains to be established. Taken together, these results suggest that the mechanisms underlying the therapeutic effects of the transplants cannot be attributed solely to direct inhibition of dorsal horn excitatory interneurons and projection neurons or by presynaptic inhibition of nociceptive primary afferents. Rather disinhibitory circuits may be part of the complex networks created by and with the transplanted cells.

3.4 IS THERE AN ENDOGENOUS GABAstat THAT REGULATES MGE-DERIVED INHIBITORY CONTROL? Consistent with earlier studies (Moore et al., 2002), we found that GAD65, but not GAD67 mRNA levels were significantly reduced in the spinal cords of SNI mice, ipsilateral to the injury (Bra´z et al., 2012). One week after the transplant, GAD levels did not differ from baseline (i.e., before surgery). In other words, MGE cells normalized GAD expression, returning it to endogenous preinjury levels, thereby reversing an injury-induced change that is a critical contributor to the persistent pain. Taken

4 Cell transplants for the management of chronic itch

together with the fact that the MGE cells had no effect on baseline mechanical thresholds when transplanted into uninjured mice, our results suggest that the MGE cells do not act as a chemical pump continuously releasing GABA into the milieu. Rather, we hypothesize that there is a feedback control from the host, in effect, a GABAstat that regulates the inhibitory control that derives from the transplant and maintains it at normal, endogenous levels. Note that our electrophysiological studies determined that the great majority of transplanted cells were spontaneously active, even in the absence of peripheral nerve injury. In other words, even though the cells are clearly functional, the fact that they do not alter baseline withdrawal thresholds likely reflects a regulatory adaptation of the host to the transplant.

3.5 MGE CELLS PREVENT THE DEVELOPMENT OF MECHANICAL ALLODYNIA Most unexpectedly, we found that when transplanted prior to a peripheral nerve injury, the MGE cells completely prevented the mechanical hypersensitivity that reliably develops following injury (Etlin et al., 2016). In these experiments, naı¨ve adult mice underwent spared nerve injury 4 weeks after they received an intraspinal MGE transplant. As expected, 1 day after injury, in mice in which we only injected medium, we recorded a marked reduction of the mechanical threshold, ipsilateral to the injury. In contrast, mice that received MGE transplants never developed mechanical allodynia. Thus, it appears that the MGE transplants maintain a level of inhibition that would normally be reduced following nerve injury. Presumably, this reflects the action of the proposed GABAstat. Hence, whatever mechanism underlies the pathophysiological impact of peripheral nerve injury on host GABAergic interneurons, the injury does not appear to have a negative effect on the transplanted cells. These observations, taken together with the fact that the MGE cells retain their cortical phenotype and fully integrate in a “foreign” tissue, suggest that the transplants are less susceptible to environmental influences induced by injury.

4 CELL TRANSPLANTS FOR THE MANAGEMENT OF CHRONIC ITCH Although the experience of itch and pain is clearly distinct, recent studies demonstrate fascinating areas of interaction (Akiyama and Carstens, 2014; Liu and Ji, 2013; Schmelz, 2015). For example, scratching can relieve itch, but it may be painful. Also, morphine, the most effective pharmacological approach for the management of severe pain, can provoke itch in both rodents (Moser and Giesler, 2014) and humans (Szarvas et al., 2003). Although recent studies found evidence for itchand pain-specific circuitry in the spinal cord dorsal horn (Akiyama and Carstens, 2013; Han et al., 2013; Imamachi et al., 2009; Kim et al., 2011; Liu et al., 2009, 2011; Mishra and Hoon, 2013; Sun and Chen, 2007; Wang et al., 2013), there is clearly significant evidence for interactions, which undoubtedly contribute to the

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opposing opioid effects. Of particular interest for our studies of MGE cell transplants for the control of chronic neuropathic pain is that the pathophysiology of some chronic itch conditions bears striking similarities to what underlies neuropathic pain. For example, similar to neuropathic pain, loss of inhibitory controls in the spinal cord is a major contributor to neuropathic itch. Specifically, Ross et al. (2010) reported that mice with a deletion of the gene that codes for the transcription factor, Bhlhb5, develop self-inflicted lesions due to excessive scratching, a condition presumed to result from chronic itch. These authors demonstrated that underlying this exacerbated scratching was a loss of a large population of dorsal horn inhibitory interneurons that coexpress GABA and dynorphin (Chiang et al., 2016; Kardon et al., 2014). In another study, using a mouse model of dry skin-induced chronic itch, Akiyama et al. (2011) reported that both GABA and glycine antagonists can block the scratching-induced inhibition of spontaneous firing of dorsal horn neurons. Finally, patients implanted with a baclofen pump developed severe pruritus when the baclofen was withdrawn, presumably due to an acute loss of GABAB-mediated inhibitory controls (Ben Smail et al., 2005). Given the commonalities in the pathophysiology of neuropathic pain and itch (i.e., loss of GABAergic inhibition), we hypothesized that increasing spinal cord GABA signaling via MGE cell transplantation would also have therapeutic effects against chronic itch.

4.1 MGE CELLS REDUCE SPONTANEOUS SCRATCHING AND RESOLVE SKIN LESIONS IN Bhlhb5 MUTANT MICE To test this hypothesis, we examined the consequences of transplanting MGE cells into the spinal cord of the Bhlhb5 mutant mice. In these experiments, we transplanted the cells in spinal cord segments that receive inputs from skin regions with lesions and recorded spontaneous scratching weekly. Two weeks after transplantation, not only did we observe a significant reduction of the excessive scratching, but there was also a dramatic resolution of the skin lesions (Fig. 4; Bra´z et al., 2014a,b). In contrast, mice that only received medium never recovered. Surprisingly, however, and in contrast with our results in the traumatic nerve injury (SNI) model of neuropathic pain, the MGE cells did not overcome the very large decrease of GAD levels that resulted from the loss of inhibitory interneurons in the Bhlhb5 mutant mice. These findings suggest that it is the exquisite axonal arborization and integration of the transplants into the host circuitry, rather than the number of surviving transplanted cells, that are critical for the therapeutic effect of the transplants. These findings reinforce the idea that intraspinal transplantation of GABAergic precursors is a viable and effective strategy to treat both neuropathic itch and pain.

4.2 MGE TRANSPLANTS ARE ALSO EFFECTIVE AGAINST CHRONIC, INFLAMMATORY ITCH As noted earlier, the MGE cell transplants did not influence nocifensive behaviors in a model of inflammatory pain (namely hindpaw injection of formalin). As there are both neuropathic itch and inflammatory chronic itch conditions, we also investigated

4 Cell transplants for the management of chronic itch

FIG. 4 Spinal cord transplantation of MGE cells in Bhlhb5 mutant mice restores GABAergic inhibitory control, decreases spontaneous scratching and resolves the associated skin lesions. (A) Mice deficient for the transcription factor Bhlhb5 develop self-inflicted lesions due to excessive scratching. If untreated, the skin lesions never resolve. (B–C) Within the 1st week of MGE cell transplantation into spinal cord segments that receive inputs from skin dermatomes with lesions, there is significant improvement ipsilateral to the transplant. In some cases, the improvement includes regrowth of hair in previously denuded regions.

whether enhancing GABAergic inhibitory controls is therapeutic against nonneuropathic chronic itch conditions, i.e., conditions not associated with a loss of GABAergic inhibitory control. In these studies, we transplanted the MGE cells into the spinal cord of transgenic mice in which there is a peripheral overexpression of the cytokine IL31 (IL31Tg mice). The phenotype observed in these mice results from an interaction of IL31 with primary afferent neurons that express the IL31 receptor

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(Cevikbas et al., 2014) and is presumed to model atopic dermatitis, a histamineindependent chronic itch condition (Budenkotte et al., 2010; Kim et al., 2016). Similar to the Bhlhb5 mutant mice, the IL31Tg mice develop severe skin lesions because of excessive scratching. Our preliminary results indicate that IL31Tg mice that received MGE cells exhibit less scratching and as a consequence have significantly reduced skin lesions (Bra´z et al., 2016). These results are yet another important demonstration of the broad therapeutic effects of the MGE cell transplantation approach.

5 TRANSLATING PRECLINICAL TRANSPLANTATION STUDIES TO THE CLINIC Although promising, the MGE transplantation approach, in its present form, cannot be translated directly to the clinic. The next step is to demonstrate that human MGE cells or an equivalent source of fetal, human inhibitory interneurons can replicate the mouse MGE antihyperalgesic and/or antipruritic effects. Recent reports used intrathecal (Vaysse et al., 2011) or intraspinal (Mukhida et al., 2007) GABA-expressing transplants of human neural precursor cells to reduce the mechanical hypersensitivity associated with partial peripheral nerve injury. Those studies were promising, but significant hurdles remain. For example, embryonic human progenitor cells, whether immortalized or not, require expansion in vitro. After multiple passages, some properties of the transplanted cells may change, which reduces the likelihood of their differentiating into neurons (Jain et al., 2003). Also of great concern is the potential development of tumors after transplantation. Finally, and of particular importance to addressing long-term pain and itch management, is the relatively poor survival rate of the transplants. The explanation for the limited survival is not clear, but perhaps occurs because there are significant differences in the properties of the transplanted tissue, which will vary depending on the source of the cells, the species, and age of the donor cells as well as culture conditions. Encouraging results, however, have recently been reported using embryonic or induced pluripotent cells, which differentiated into functional subtypes of GABAergic interneurons and successfully integrated into the host circuitry (Chambers et al., 2009; Nicholas et al., 2013).

6 CONCLUSIONS Our studies show that MGE cells are effective against a variety of chronic pain (chemotherapy- and peripheral nerve injury-induced neuropathic pain) and itch (neuropathic and inflammatory) models in the mouse. This cell-based approach has broad therapeutic utility and is particularly effective when loss of inhibitory controls is a major contributor to the condition. Cell transplantation differs considerably from traditional pharmacological treatments. Because transplantation is directed at the underlying etiology of the condition, it is potentially disease modifying. Among the therapeutically relevant properties of transplanted MGE cells are high survival rate,

References

stability and safety, differentiation into functionally integrated mature interneurons, and rescue of GABAergic inhibitory control. Clearly, the next step is to determine the extent to which cell transplantation can be translated to the clinical management of chronic pain and itch.

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From transplanting Schwann cells in experimental rat spinal cord injury to their transplantation into human injured spinal cord in clinical trials

5

Mary B. Bunge*,†,{,1, Paula V. Monje*,{, Aisha Khan§, Patrick M. Wood*,{ *The Miami Project to Cure Paralysis, University of Miami Leonard M. Miller School of Medicine, Miami, FL, United States † Department of Cell Biology, University of Miami Leonard M. Miller School of Medicine, Miami, FL, United States { Department of Neurological Surgery, University of Miami Leonard M. Miller School of Medicine, Miami, FL, United States § The Interdisciplinary Stem Cell Institute, University of Miami Leonard M. Miller School of Medicine, Miami, FL, United States 1 Corresponding author: Tel.: +1-305-243-4596; Fax: +1-305-243-3923, e-mail address: [email protected]

Abstract Among the potential therapies designed to repair the injured spinal cord is cell transplantation, notably the use of autologous adult human Schwann cells (SCs). Here, we detail some of the critical research accomplished over the last four decades to establish a foundation that enables these cells to be tested in clinical trials. New culture systems allowed novel information to be gained about SCs, including discovering ways to stimulate their proliferation to acquire adequately large numbers for transplantation into the injured human spinal cord. Transplantation of rat SCs into rat models of spinal cord injury has demonstrated that SCs promote repair of injured spinal cord. Additional work required to gain approval from the Food and Drug Administration for the first SC trial in the Miami Project is disclosed. This trial and a second one now underway are described.

Keywords Spinal cord injury, Spinal cord repair, Human Schwann cells, Schwann cell transplantation, Schwann cell clinical trials Progress in Brain Research, Volume 231, ISSN 0079-6123, http://dx.doi.org/10.1016/bs.pbr.2016.12.012 © 2017 Elsevier B.V. All rights reserved.

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1 INTRODUCTION In 1975, Richard Bunge proposed that glial cells taken from the nervous system and isolated and purified in tissue culture could then be transplanted for central nervous system (CNS) repair (Bunge, 1975). The foundation for this suggestion was the then recent development of culture methods to obtain purified populations of Schwann cells (SCs), the glial cells that provide ensheathment and myelin for axons in the peripheral nervous system (PNS). This vision drove our efforts for many years to learn more about SC biology to (1) develop improved protocols to obtain large numbers of rat and human SCs in culture, (2) test the efficacy of SC transplantation into injured rat spinal cord for repair, and, finally, (3) to translate our findings to autologous human SC clinical trials. It is the goal of this chapter to summarize some of these milestones: the development of procedures to prepare appropriate populations of human SCs for transplantation into humans with spinal cord injury (SCI), the results of implanting SCs into injured rat spinal cord, and the path required to gain approval from the Food and Drug Administration (FDA) to conduct autologous human SC clinical trials. It is not our focus here to review other SC trials in Iran (Oraee-Yazdani et al., 2016; Saberi et al., 2008, 2011; Yazdani et al., 2013) and China (Chen et al., 2014; Zhou et al., 2012). The Saberi and Zhou team trials have been compared by Guest et al., 2013. This is the first time that the history of the development of protocols to obtain human SCs for clinical use and the preclinical work required to obtain FDA approval for our first trial have been detailed. Issues that have been most important to the initiation of our first clinical study have been raised already (Guest et al., 2013). The populations of neurons and fibroblasts as well as SCs from rat dorsal root ganglia, and the possibility to maintain these in isolation and in coculture, enabled numerous studies that revealed new information about SC biology (Bunge, 2016). One example was the discovery of a molecular signal on the surface of axons that caused SCs to proliferate (Wood and Bunge, 1975). Another example was the discovery that the extracellular matrix that surrounds the SC-axon units plays a key role in SC development and function (Eldridge et al., 1989). These earlier studies have been reviewed previously (Bunge and Bunge, 1983; Bunge et al., 1983, 1990). Later studies that found the importance of cyclic AMP (cAMP) levels and heregulin in SC proliferation, key in acquiring adequate numbers of SCs for transplantation investigation, are summarized in this chapter. Obtaining SCs from sources other than peripheral nerve are reviewed briefly here as well, in view of the new trends that enable the use of stem cells as an alternative source of transplantable SCs.

2 ADVANTAGES OF PRIMARY SCs FOR CELL THERAPY IN SCI SCs were likely candidates for transplantation to promote CNS repair for many reasons. It is the SC in peripheral nerve that leads to normal repair following injury. In addition, a particularly relevant report by Richardson et al. (1980) demonstrated that

2 Advantages of primary SCs for cell therapy in SCI

a piece of peripheral nerve implanted into a complete gap in adult rat spinal cord elicited regeneration of axons from the central neuronal somata into the nerve implant they bordered. This was a key discovery because it showed that if the environment is appropriate, CNS neurons will regenerate axons in the damaged spinal cord, and an appropriate milieu was provided by the SC-rich peripheral nerve. Regeneration of CNS axons into peripheral nerve implants has also been demonstrated in nonhuman primates (Levi et al., 2002). SCs are readily accessible in biopsies of adult peripheral nerve. They are known to produce growth factors, including diverse neurotrophic factors, and extracellular matrix molecules that foster axon growth (Bunge, 1994) and provide new myelin, including for CNS axons. Upon transplantation, they can serve as a scaffold for the regeneration of axons across a cystic injury site. They can be generated in large numbers in culture and also in culture be genetically engineered to insert transgenes to express critical molecules for repair when transplanted (Golden et al., 2007). A strong rationale for choosing SC transplantation was that they could be transplanted autologously after removal of a peripheral nerve biopsy from a spinal cord injured person and then purified and expanded in number in culture prior to transplantation (Bunge and Wood, 2012). Mature SCs are among the few known cell types able to respond to injury by undergoing dedifferentiation and self-renewal during adulthood. This property, maintained throughout life, plays an instrumental role during nerve regeneration and restoration of myelin after nerve damage (Jessen and Mirsky, 2016; Jessen et al., 2015). In fact, it has been shown that SCs can retain their capacity to self-renew, foster axon growth, and remyelinate axons even after repeated nerve injury events (Thomas, 1970). Both myelinating and nonmyelinating SCs can dedifferentiate and convert into a repair phenotype that fills the bands of Bungner (reviewed in Jessen and Mirsky, 2016; Jessen et al., 2015). This intrinsic plasticity of adult nerve SCs is also manifested when SCs are cultured in vitro. SCs have the capacity to digest their own myelin and proliferate extensively within a dissected tissue explant of peripheral nerve (reviewed in Scherer and Salzer, 2001). In vitro, they are able to do so without the aid of other types of cells (Fernandez-Valle et al., 1995). Given their potential for self-renewal, adult nerve-derived SCs are endowed with an exceptional capacity for survival and expansion in vitro. From a practical perspective, this feature of the adult SC provides a unique advantage for the culturing of primary cells. Early studies showed that SCs require stimulation with very specific factors to proliferate (Raff et al., 1978a,b) and that these factors are bound to the axonal membrane (Salzer et al., 1980a,b; Wood and Bunge, 1975). SCs stop dividing once these factors are removed from the culture medium (Raff et al., 1978a) or deprived of the trophic support provided by their contact with axonal factors (Salzer and Bunge, 1980; Wood and Bunge, 1975). Cultures of isolated primary SCs are phenotypically and genetically stable over several passages. They also maintain their ability to foster axon growth and form a myelin sheath in vitro (Morrissey et al., 1995b) and after transplantation into the CNS or PNS (Fortun et al., 2009; Levi and Bunge, 1994; Levi et al., 1994). As opposed to stem cells, SCs are lineage-committed cells and cannot differentiate into a phenotype different from that of the SC. Though

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significant plasticity has been observed in the response of adult SCs to nerve damage, phenotypic transformations are restricted to developmental stages within the SC lineage (reviewed in Jessen et al., 2015). This remarkable feature of the SC provides an additional level of safety for their use in vivo.

3 SC PROLIFERATION: CUES FOR ACHIEVING EXPANSION BY USING HEREGULIN AND cAMP-STIMULATING AGENTS Inducing SCs to actively proliferate in vitro has been key to the successful expansion of cultured SCs to the high numbers required for transplantation. When SCs are isolated from nerve, the cells can survive for prolonged periods of time in vitro. As mentioned earlier, isolated SCs remain quiescent and do not proliferate unless stimulation with axons is reestablished (Wood and Bunge, 1975) or specific growth factors that mimic the effect of axons/axonal signals are provided directly into the culture medium (Morrissey et al., 1995c; Raff et al., 1978a,b). As opposed to other types of cells, cultured SCs do not typically proliferate in response to serum factors. Maximal levels of SC proliferation can be achieved when soluble heregulin, or heregulin-containing extracts, are provided in combination with agents that increase the intracellular levels of cAMP rather than alone (Dong et al., 1997; Levi et al., 1995; Porter et al., 1986; Raff et al., 1978b). In fact, it has been suggested that in combination heregulin and cAMP are sufficient to mimic the strong promitogenic effect provided by axon-derived signals (Morrissey et al., 1995c). The neuregulins comprise an extensive family of growth factors that serve as specific ligands for membrane receptors of the ErbB/HER family. These receptors belong to the superfamily of tyrosine kinase receptors and are closely related to the epidermal growth factor receptor. ErbB receptors signal through the activation of various intracellular kinase cascades (Alroy and Yarden, 1997). In SCs, heregulin elicits mitogenesis through the activation of heterodimers of the ErbB receptor consisting of two subunits: ErbB2 and ErbB3 (reviewed in Garratt et al., 2000). Together, ErbB2 and ErbB3 constitute a powerful signaling complex. Whereas the ligand binding domain of ErbB3 recognizes and binds with high affinity to heregulin, the tyrosine kinase activity of ErbB2 performs the function of transactivation and activates intracellular signal transduction cascades (reviewed in Citri et al., 2003; Stern, 2008). cAMP is a ubiquitous intracellular second messenger that results from the activation of membrane-bound or soluble forms of the adenylyl cyclase, the enzyme responsible for the synthesis of cAMP from ATP (Ladilov and Appukuttan, 2014). In SCs, cAMP acts via activation of its downstream effector protein kinase A (PKA) to modulate the rate of S-phase entry (Bacallao and Monje, 2013; Kim et al., 1997; Monje et al., 2006, 2008). Addition of cAMP-stimulating agents or agonists of PKA to the culture medium synergistically increases SC proliferation (S-phase entry) in the presence of heregulin but not when provided alone (Monje et al., 2008). By directly phosphorylating ErbB2, PKA synergistically enhances and prolongs the

4 Other sources of SCs

duration of the state of activation (tyrosine phosphorylation) of heregulin-stimulated ErbB2/ErbB3 heterocomplexes. In turn, this action of PKA contributes to synergistically increase the activation of kinase cascades downstream of heregulin-activated ErbB connected to the progression into the S-phase (Monje et al., 2008). Mechanistically, this mode of signaling crosstalk between cAMP-PKA and ErbB signaling is a highly efficient way to control cell proliferation. SCs are among the very few cell types that respond to cAMP elevation with an enhancement in proliferation rates. The action of heregulin in SC proliferation is also very specific. Albeit other growth factors such as platelet-derived growth factor and fibroblast growth factor can exert a mitogenic effect in cultured isolated SCs, and their action on proliferation is synergistically increased by costimulation with agents that increase cAMP (Kim et al., 2001; Monje et al., 2009; Stewart et al., 1991), their effect on proliferation is relatively modest compared to that of heregulin. No other factor has been identified so far that parallels the ability of heregulin to promote SC survival, lineage specification, migration, and proliferation (Garratt et al., 2000). Current protocols to culture human SCs have included heregulin and cAMPstimulating agents in combination to maintain the cells in the cell cycle and achieve a fast and sustainable rate of proliferation (Casella et al., 1996; Levi et al., 1995; Monje et al., 2006). The use of this mitogenic combination maximizes the expansion potential of the relatively low number of primary SCs that are typically recovered from enzymatic dissociation of adult nerve. The series of discoveries underlying the requirement and biological activity of these two key mitogens have been critical in deciphering the best possible conditions to isolate and expand human SCs for clinical trials. A narrative of the sequence of events that led to these discoveries is provided in subsequent pages.

4 OTHER SOURCES OF SCs In recent years, SC derivation from sources other than peripheral nerve has become increasingly popular. These technologies exploit the unlimited self-renewal capacity of different populations of stem cells, along with the potential to convert these into SCs, to overcome limitations in the production of primary cells for transplantation and other uses. Several studies have addressed the use of directed in vitro differentiation to generate SCs from skin-derived neural precursors (Biernaskie et al., 2006) and various types of mesenchymal stem cells, primarily those derived from the bone marrow (Brohlin et al., 2009; Keilhoff et al., 2006a,b,c) and adipose tissue (Kingham et al., 2007). Evidence indicates that induced pluripotent stem cells can also be driven to acquire a SC phenotype (recently reviewed in Ma et al., 2015). These accessible stem cell sources provide the advantage to derive human SCs in an autologous way. Of note, human SCs from skin-derived neural precursors and bone marrow stromal cells engineered via directed in vitro differentiation have been transplanted into the injured spinal cord and peripheral nerve in experimental animal models with promising results (Biernaskie et al., 2007; Shimizu et al., 2007).

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Regardless of the stem cell of origin, most available methods use one or more steps of chemical induction to drive the final conversion of multi- or pluripotent stem cell-derived cells into lineage-committed SCs (reviewed in Lehmann and Hoke, 2016; Ma et al., 2015). Some protocols include an intermediate step of neural crest derivation prior to SC conversion. Still, most reported methods rely on treatment with heregulin, provided alone or together with cAMP-stimulating agents such as forskolin, to drive final specification into the SC lineage and subsequent propagation in vitro. Multiple examples in the literature have shown that these converted SCs display characteristics of peripheral nerve SCs (e.g., Sakaue and Sieber-Blum, 2015), based on various features including an elongated (bipolar) morphology, the expression of SC-specific markers such as the calcium-binding protein S100 and glial fibrillary acidic protein, and responses to mitogenic factors. How much they resemble those obtained directly from nerves, however, is for the most part an open question. Another important question is whether these SCs have the capacity to differentiate fully and to efficiently form myelin either in vitro or after transplantation. Due to these limitations and gaps in our current understanding about how in vitro-converted SCs promote nerve tissue repair, stem cell-derived SCs are not yet ready for testing in clinical cell therapy (Lehmann and Hoke, 2016). Presently, the isolation of adult nerve-derived primary SCs through traditional methods is the preferred and safest way to procure SCs for use in humans. First, there is a long history of work on transplantation of primary SCs that have pointed to the feasibility and success of this approach. Second, as pointed out earlier, adult nerve-derived SCs are lineage-committed cells and their potential for transformation or derivation into an undesirable phenotype is extremely low. Nevertheless, the emerging field of SC conversion is advancing swiftly, and different options for a faster and safer production of SCs may soon become available. Stem cells may not be necessary, as evidenced by the possibility to directly convert human fibroblasts into SCs by transdifferentiation in vitro using small molecule multikinase inhibitors (Thoma et al., 2014). It is expected that at the current pace, SC conversion technologies would allow mass production of autologous human SCs without the need to surgically remove the patient’s nerve.

5 SC TRANSPLANTATION STUDIES IN RAT SCI MODELS With the availability of culture techniques to generate substantial numbers of SCs, studies began in earnest in the 1990s to assess the efficacy of SCs to repair the injured rat spinal cord, utilizing two SCI models, complete transection with a SC bridge and contusion (Bunge and Wood, 2012; Fortun et al., 2009; Tetzlaff et al., 2011). The development of the complete transection model by Dr. Xiao Ming Xu was critical to prove that SC transplantation led to regeneration of axons as opposed to sprouting or survival (Xu et al., 1995, 1997). When the stumps of the severed spinal cord were inserted into each end of a polymer channel that contained a cord of SCs and that

6 Development of the clinically relevant protocol

spanned a complete gap in the spinal cord, axons regenerated from both rostral and caudal stumps into the SC bridge (Xu et al., 1997). Study of contusion injury (Takami et al., 2002) was important due to its clinical relevance. SCs to be transplanted were always extracted from adult nerve, also for clinical relevance. In both models, the implant resembled peripheral nerve in that the SCs myelinated or ensheathed axons that were present in the implant. The implanted SCs were observed to reduce cyst size and protect tissue around the implant from secondary injury (in contused spinal cords), support axon growth into the implant, and form myelin around the implant axons. Modest improvement in hindlimb movements was noted in implanted contused rats. Whereas about 5000 SC-myelinated axons were present in SC grafts in contused rats, about half that number could be seen in the contusion lesion when SCs had not been transplanted, indicating endogenous SC ingress. Consideration of the multiple deleterious effects in the spinal cord tissue following injury led to interest in testing numerous combinatorial strategies in both injury models to improve SC efficacy. Combination of SCs with administration of a steroid, methylprednisolone; a variety of growth factors (primarily neurotrophins, in some cases generated after genetic manipulation of the SCs to be transplanted); olfactory ensheathing glia; an enzyme, chondroitinase; or elevation of cAMP all led to improvement in outcome measures compared to SC transplantation alone (Bunge, 2016; Fortun et al., 2009; Tetzlaff et al., 2011). With these combinatorial treatments, more SC-myelinated axons populated the graft, more axons from neurons above the cord were in the implant, and locomotion of the paralyzed rats was further improved. In the case of activating both trk B and C receptors to simulate BDNF and NT-3 delivery, there was up to a fivefold increase in graft volume and SC and myelinated axon number compared to SCs alone (Golden et al., 2007). Total axons (myelinated and ensheathed) were estimated to reach 75,000 in the combination group. Moreover, triple combinations were more effective than double ones. For example, in a recent study in which SC implants were combined with neurotrophins and/or chondroitinase, myelinated axon number was substantially increased, more supraspinal neurons responded, walking scores were improved, and hindlimb pain was reduced to a greater extent in the full combination compared to SCs with either growth factor or enzyme (Kanno et al., 2014, 2015). All these studies in rat models of SCI (reviewed previously by Bunge, 2016; Fortun et al., 2009; Tetzlaff et al., 2011) contributed to gaining approval from the FDA for a clinical trial to test the safety of transplanting autologous human SCs into SCI sites.

6 DEVELOPMENT OF THE CLINICALLY RELEVANT PROTOCOL FOR MANUFACTURING AUTOLOGOUS HUMAN SCs SCs were first isolated in cultures of embryonic rodent sensory ganglia that had been treated with antimitotic agents to remove nonneuronal cells. Following withdrawal of the antimitotic agents, the few SCs that had survived the treatment began to

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proliferate, repopulate the axons covering the culture surface, and form myelin, indicative of their normal potential for differentiation. Removal of the centrally located explant(s) containing the neuronal somata left cultures with nearly pure SCs (Wood, 1976; Wood and Bunge, 1975). The initial sensory ganglia cultures were grown and manipulated in small (22 mm diameter) fluorocarbon dishes coated with rat tail collagen to provide an adhesive substratum. Each of these cultures would have contained no more than 100,000–200,000 SCs that did not undergo further proliferation after removal of the neuron-containing explants. In fact, in the absence of neurons, the number of SCs declined over time. The property of strict regulation of proliferation by neuronal contact is a defining feature of normal SCs. It is an everlasting testament to the optimism and perspicacity of Richard Bunge that he recognized in these isolated SCs the potential for vast expansion in number and that they might someday be useful as implants for the repair of nervous system injury or disease (Bunge, 1975). The following describes the many new discoveries that were made and the technical obstacles that required resolution before his vision could be realized. The key contributions of the numerous researchers who overcame these obstacles are recognized.. The narrative is a history of the more than 30 years of work leading to the production of clinical grade autologous SCs from humans.

6.1 MAKING SCs PROLIFERATE IN CULTURE The first question was how to make the SCs proliferate, a problem soon resolved by two discoveries in the late 1970s (Brockes et al., 1979; Raff et al., 1978b). The first discovery was the observation that SCs isolated from neonatal rat sciatic nerve could be stimulated to low levels of proliferation by adding to the medium agents that elevated the intracellular levels of cAMP. These agents were dibuturyl-cAMP, a membrane permeable analog of cAMP, or cholera toxin, an irreversible activator of adenylyl cyclase. The second discovery was that SC proliferation could be increased further by adding brain or pituitary extract (Brockes et al., 1979). The factor in these extracts that stimulated SC division was purified to homogeneity and named glial growth factor (GGF; Brockes et al., 1981). These basic findings that SCs could be expanded in number following isolation in culture, and that this could be accomplished by stimulation with cAMP and GGF, were the critical first steps toward the development of the protocol we use today for the manufacture of human SC implants.

6.2 THE BROCKES PROTOCOL: FIBROBLAST DEPLETION TO PURIFY SC CULTURES The complete method developed by Brockes et al. (1979), proven to be an effective method for generating large numbers of highly purified rat SCs, has been widely used in diverse studies of SC biology. Because our present human SC protocol has been developed by a number of modifications of this original rat SC protocol, the Brockes protocol is described here in more detail. The SC cultures prepared according to the

6 Development of the clinically relevant protocol

Brockes method were derived by collagenase-mediated dissociation of early postnatal rat sciatic nerves. Because SCs are not fully mature at this stage, with less extensive processes and relatively small size compared to their adult counterparts, they readily survived tissue dissociation. The myelin-free cell preparations obtained from nerve dissociation were plated onto uncoated tissue culture flasks for further processing and expansion of cell numbers in medium supplemented with serum and cholera toxin. The starting cultures contained fibroblasts that would gradually outnumber the SCs. Methods were thus developed for purifying the SCs at the outset of culture. In serum-containing medium, the fibroblasts divide rapidly, in contrast to the SCs which divide much more slowly. Because rapidly dividing cells are susceptible to killing by antimitotic agents, it was possible to deplete the cultures of fibroblasts by treatment with cytosine arabinoside, a pyrimidine compound that inhibits DNA synthesis. Even more effective was an immunological approach. Fibroblasts express the antigenic marker Thy1-1 on their surfaces, making them highly susceptible to treatment with Thy1-1 antibodies followed by complement-mediated cell lysis. These methods effectively reduced the impact of fibroblast contamination in SC cultures, resulting in purities greater than 95% SCs.

6.3 THE PORTER PROTOCOL: ELIMINATION OF CHOLERA TOXIN AND MODIFICATION OF THE CULTURE SUBSTRATUM An important advance toward a more clinically relevant protocol was made with the elimination of cholera toxin from the Brockes SC protocol (Porter et al., 1986). Although cholera toxin was effective for raising cAMP levels in SCs, its effects on adenylyl cyclase were irreversible (Cassel and Pfeuffer, 1978; Gill and Meren, 1978), thereby causing concern that long-term effects, such as a permanent activation of some level of cell division, might diminish the value of SCs in repair. A possible alternative to cholera toxin, forskolin, that reversibly activates adenylyl cyclase, had been reported (Seamon and Daly, 1981). When cultures of SCs were prepared essentially as described in the Brockes protocol, forskolin, in conjunction with pituitary extract or partially purified GGF, was found to promote vigorous SC division (Porter et al., 1986). A further advance made by these investigators (Porter et al., 1986) was to coat the culture dishes with poly-L-lysine (PLL) to improve adhesion and retention of the SCs to the dishes, compared to the uncoated dishes used in the original Brockes protocol (Brockes et al., 1979). On the PLL-coated dishes, the SCs could proliferate to confluency and be maintained with consecutive passaging of cells for up to 4 months with excellent retention of functional capacities (Porter et al., 1986). After expansion under these conditions, the SCs ceased division upon removal of mitogenic factors from the medium, indicating that continual exposure to mitogens did not induce transformation or tumorigenesis. In addition, the expanded SCs exhibited a normal pattern of protein secretion and an undiminished ability to myelinate axons in vitro (Porter et al., 1986).

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6.4 TRANSFORMATION OF SCs WITH EXTENDED PASSAGES From the beginning of this research, there had been concern that prolonged exposure to mitogens might induce permanent functional changes in the SCs, including transformation. It is, therefore, of considerable interest to know whether SCs that had been cultured for significantly longer periods in the presence of mitogens would exhibit altered functional capabilities. The elimination of cholera toxin from the medium and its replacement by forskolin allowed rapid propagation of postnatal SCs in vitro (Porter et al., 1986). These SCs maintained their normal phenotype over several passages in forskolin-containing medium. Only when SCs were cultured for longer than 4 months (12 passages or 24 population doublings) did they begin to exhibit increased levels of proliferation in the absence of added mitogens, possibly due to the onset of secretion of autocrine mitogenic factor(s) by the SCs (Porter et al., 1987). This aberrantly high proliferation increased with continued culture and passaging in the presence of mitogens. The SCs cultured for 8 months (24 passages) still showed some ability to form myelin, but the myelinating capacity was completely lost by 16 months of culture with mitogens when the population was considered to be immortalized (Porter et al., 1987). These immortalized cells were subsequently shown to generate malignant tumors when transplanted into immune-deficient animals (Langford et al., 1988). For cells destined to be used clinically, the demonstration that neonatal rat SCs could be transformed by prolonged exposure to mitogens indicated a need to monitor the cells’ ability to maintain growth control. In the current clinical trials underway at the Miami Project to Cure Paralysis testing human SC transplantation safety in spinal cord injured subjects, the SCs are exposed to mitogens only through four passages (12–16 population doublings). This number of doublings is well below the number at which the loss of growth control was observed in cultures of rat SCs.

6.5 THE CHALLENGE OF ISOLATING SCs FROM ADULT RAT NERVE The Porter protocol using forskolin with GGF or pituitary extract, which contains GGF activity (Brockes et al., 1981), thus appeared to be effective for the generation of large enough populations of highly purified rat SCs for both biochemical and in vivo studies. Ideally, to be relevant for clinical application, the SC implant would be autologous, prepared from cells isolated from nervous tissue harvested from the person receiving the implant. Because usually these would be adults, there was a need to establish that the protocol would be effective for generating large numbers of cells from an adult nerve. The conditions for enzymatic digestion described in the Porter protocol had been optimized for use with immature postnatal sciatic nerves. In adult rat (sciatic) nerve, many of the SCs myelinate axons, forming extensive membranous sheaths around the axon. In addition, adult nerve contains robust epineurial and perineurial connective tissue and tightly packed collagen fibrils; as a result, direct dissociation of such nerves after incubation with digestive enzymes leads to extensive loss of SCs. In early experiments applying the Porter protocol to adult rat

6 Development of the clinically relevant protocol

nerves, cell yields were extremely low, even if serum was included in the incubation mixture to slow the process of digestion and the release of single cells (Scarpini et al., 1988). Earlier research focusing on the isolation of SCs from adult human nerves had shown that SC yields could be improved by the use of a protocol in which the nerves were first carefully stripped of connective tissue sheaths and blood vessels, and then cut into short segments that were explanted in culture on uncoated plastic dishes (Askansas et al., 1980). These investigators found that after the explants became attached to the dish, cells (including some SCs but large numbers of fibroblasts) migrated out of the explants onto the plastic. After 2–3 weeks, the explants were picked up and transferred to new dishes; these new cultures were maintained until a new migration of cells out of the explants had occurred. This replating of explants was repeated several times and led to a progressive depletion of the fibroblast population. When most of the cells in the outgrowth exhibited morphology typical of SCs (after about 14 weeks), the tissue explants were discarded and the adherent SC outgrowth was harvested. Whereas the SC purity was high (>90%) and the yield was sufficient for small scale biochemical experiments, this procedure (Askansas et al., 1980) did not appear to be adequate to generate large enough populations for large scale or in vivo transplantation experiments.

6.6 THE MORRISSEY–KLEITMAN PROTOCOL: INCREASING ADULT SC EXPANSION BY USING MULTIPLE REPLATING OF NERVE EXPLANTS AND DELAYED EXPLANT DISSOCIATION The findings reported by Askansas et al. (1980) demonstrated that the culturing of nerve explants with multiple explant replatings to deplete fibroblasts would result in better yields of viable SCs than could be obtained by immediate dissociation of the nerve tissue. In addition, these studies showed that, after the multiple replatings, the nerve explants still retained abundant viable SCs that might be readily recovered by enzymatic digestion of the explants themselves. Under supervision by Dr. Naomi Kleitman, a more effective protocol for the isolation and growth of adult rat SCs was devised by combining serial replating and enzymatic (collagenase–dispase) treatment of the explants. After the delay of multiple replatings, the explants were dissociated using a combination of enzymatic digestion and gentle trituration to render highly viable cells (Morrissey et al., 1991). In the Morrissey–Kleitman protocol, adult rat nerves were cut into lengths of 3–4 cm. Fascicles containing the axons, SCs, and endoneurial fibroblasts were extracted from within the nerve’s connective tissue by pulling the individual fascicles with fine forceps under a dissecting microscope. These fascicles were recut into short segments that were explanted onto plastic dishes for culture, including multiple explant replatings, as described by Askansas et al. (1980). The initial culture period of explant replating was shortened to 5–6 weeks compared to the Askansas et al. method. When the cellular outgrowth around the explants appeared to be primarily SCs (as determined by their slender bipolar shape), the explants were collected and then dissociated using a mixture of collagenase and

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dispase. This procedure yielded 20,000 cells/mg of nerve, a dramatic increase over previous results (800–1000 cells/mg). The culture over several passages of the purified adult SCs on PLL-coated dishes in medium containing GGF and forskolin (Morrissey et al., 1991) produced high yields, adequate for experiments requiring millions of cells for the first time. Because the percentage of SCs in the starting suspension was 95% or higher, additional treatments to remove fibroblasts were not necessary. This protocol, shown to work well using nerves from adult rats, appeared to be potentially valuable for generating large numbers of adult-derived human SCs in culture.

6.7 THE MORRISSEY–KLEITMAN–LEVI PROTOCOL: REPLACEMENT OF GGF WITH RECOMBINANT HEREGULIN AND ADDITION OF CHOLERA TOXIN BACK INTO THE MITOGENIC COCKTAIL When the Morrissey protocol (Morrissey et al., 1991) was used for human nerve tissue, the growth of human SCs observed using forskolin and GGF as mitogens was slower than expected, and fibroblast overgrowth became a serious problem (Morrissey et al., 1991, 1995a,b). Fortunately, by that time, another laboratory working with adult human SCs had devised a method that stimulated rapid and selective growth of human SCs while inhibiting fibroblast growth (Rutkowski et al., 1992, 1995). This method consisted of pretreating the nerves for 7 days with cholera toxin before enzymatic dissociation. This combination of delayed dissociation with exposure of the intact human nerves to cholera toxin increased the yield from 20,000 to 60,000 cells/mg of nerve at purities of about 90% SCs. The SCs were then plated onto PLL-coated dishes and cultured in serum-containing medium with a mitogen cocktail containing cholera toxin, forskolin, and human recombinant GGF. The combination of cholera toxin and forskolin markedly suppressed fibroblast proliferation while allowing relatively rapid SC proliferation. Using this protocol, the Rutkowski team reported larger numbers of human SCs than had been previously obtained, but proliferation slowed after 12 doublings and the cells exhibited morphological changes indicative of senescence, including extensive spreading and nuclear enlargement or fragmentation (Rutkowski et al., 1992, 1995). Based on the Rutkowski reports, cholera toxin was added back to the mitogenic cocktail used in the Morrissey–Kleitman protocol for human SCs. Another modification was the replacement of GGF (of bovine origin) with a recombinant human heregulin peptide (Levi et al., 1995). With the triple combination of cholera toxin, forskolin, and heregulin, large enough numbers of highly pure populations of human SCs were generated to provide for studies of the functional capability of the cultured cells to myelinate when transplanted in vivo. The human SCs were transplanted into gaps in the sciatic nerves of immune-deficient rats and mice and were shown to produce thinner than normal myelin sheaths typical of regenerated nerves (Levi and Bunge, 1994; Levi et al., 1994). In addition, transplantation of human SCs into the nude rat completely transected spinal cord led to robust regeneration of axons into the implant and some improvement in locomotor function (Guest et al., 1997). These

6 Development of the clinically relevant protocol

results provided evidence that human SCs could be exposed to mitogens for four passages and retain functional properties (such as myelination) that could improve nervous system repair. In summary, the complete Morrissey–Kleitman–Levi protocol for adult human nerves was as follows: (1) on the day of harvest, the intact nerves (containing connective tissue, fat, blood vessels as well as the actual fascicles containing axons ensheathed by SCs, perineurial cells, and endoneurial fibroblasts) were cut into short lengths (3–4 cm); (2) the fascicles were cleanly extracted by inserting forceps into the middle of each fascicle and pulling it free of the nerve; (3) the fascicles thus obtained (also 3–4-cm in length) were floated in uncoated culture dishes in serum-containing medium without growth factors (i.e., heregulin or cAMPstimulating agents); (4) after a period of time ranging from 1 to 2 weeks, the fascicles were minced to produce explants a few mm in length; (5) the explants were placed in new uncoated dishes containing a minimal amount of medium, allowing the explants to attach to the dish; (6) the cultures were subsequently monitored by phase contrast microscopy as described by Askansas et al. (1980) to evaluate the cellular composition of the outgrowth emanating from the explants; (7) when the outgrowth became confluent, the explants were picked up and moved to new dishes; (8) steps 6 and 7 were repeated until the outgrowth emanating from the explants was free of fibroblasts; (9) the explants were then collected and incubated overnight with a mixture of collagenase and dispase in serum-containing medium; (10) the explants were completely dissociated by gentle trituration through a pipette, yielding a suspension that was typically 90% SCs; and (11) the suspension was plated on PLL-coated dishes in medium containing heregulin, forskolin, and cholera toxin for further expansion up to four passages.

6.8 THE CASELLA PROTOCOL: DELAYED DISSOCIATION, CULTURE ON LAMININ, AND ELIMINATION OF CHOLERA TOXIN Though very promising, the protocol developed by Morrissey et al. (1991) and modified by Levi et al. (1995) was still not ideal for the manufacture of a SC product that could be safely used in the clinic. The initial period for maintaining and transferring the nerve explants was long and labor intensive, increasing the risk of loss through contamination. Division of cells following dissociation was relatively slow, and the amount of expansion obtained appeared limited; beyond the fourth passage the cells showed an increased tendency to aggregate and detach from the substratum. With prolonged expansion, SC division slowed relative to fibroblast division, and the cultures became contaminated with an unacceptable number of fibroblasts. Finally, adhesion and maximal growth on the PLL substratum required the presence of cholera toxin. As mentioned earlier, this mitogen exerts irreversible or slowly reversible effects on adenylate cyclase activity (Cassel and Pfeuffer, 1978). Altogether, these disadvantages diminished the potential of the protocol for clinical applications and for gaining FDA approval for clinical use.

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For these reasons, new studies were undertaken with three specific objectives: (1) to determine whether the period of maintenance of the fascicles and explants could be shortened and simplified without sacrificing starting cell yield and purity; (2) to define effective culture conditions not requiring cholera toxin for maximal growth; and (3) to increase the limits of expansion to which human SC propagation in culture could be driven (Casella et al., 1996). To achieve these goals, individual steps in the protocol were examined to ascertain their necessity and effectiveness. There was already good evidence that delaying dissociation of the explants dramatically improved the yield of viable SCs in the cell suspension obtained by dissociation. In addition, the proliferation of SCs within the explants maintained in the presence or absence of mitogens was evaluated. The relationship between the conditions of maintenance of nerve explants prior to dissociation and parameters such as cell yield, purity, and the initial rate of SC proliferation in culture was also evaluated (Casella et al., 1996). These experiments revealed that SCs within explants divided at a low rate during the first 3 weeks of culture; this proliferation falls to near zero during the fourth week. Electron microscopic analysis showed that medium containing heregulin/ forskolin exerted trophic effects on SC morphology within explants during the first 2 weeks of culture. The SCs in explants treated with heregulin and forskolin were larger, contained fewer lysosomes, and possessed less heterochromatin than those in nontreated explants. In addition, less myelin debris was present in explants treated with mitogens. Upon dissociation of the explants, the cell yield, SC purity, and proliferation rate following dissociation were all increased. The data supported the strategy of using an initial period of culture of fascicles in the presence of heregulin and forskolin for 2 weeks prior to enzymatic digestion. This modification considerably shortened the period required to generate a SC product. Next, the proliferation of the human SCs following dissociation of fascicles was studied in detail (Casella et al., 1996). In particular, studies were conducted to compare the growth of human SCs on a variety of substrata, including PLL but also collagen and laminin, two extracellular proteins occurring naturally in nerve. SC growth in medium with heregulin and forskolin was more rapid on either laminin or collagen than on PLL and, when grown on either collagen or laminin, the SCs did not show a tendency to cluster or detach from the culture dish. Passaging the cells at confluence, however, was considerably more difficult when SCs were grown on a collagen substratum because an extra step of collagenase treatment was required to release the cells from the dish, and a final step of mild trypsinization was needed to achieve a single cell suspension. In contrast, when the cells were grown on a laminin substratum, only a brief treatment with trypsin was needed to release the cells and achieve a single cell suspension. Laminin, therefore, became the substratum of choice for human SCs. Because human SCs grown on collagen or laminin substrata showed no tendency to form clusters or detach from the dish at later passages, tests were carried out to determine if cholera toxin could be eliminated from the medium without decreasing growth rates. For these experiments, SCs were isolated from explants that had been

6 Development of the clinically relevant protocol

maintained for 2 weeks either in medium containing heregulin and forskolin, or in medium containing heregulin, forskolin, and cholera toxin. In these experiments, the growth rates (and subsequent total yields of SCs) were significantly higher in the absence of cholera toxin than in its presence (Casella et al., 1996). Based on this outcome, cholera toxin was again eliminated from the mitogen cocktail used to grow human SCs. In accordance with these results, in the Casella protocol, the following modifications were made to the Morrissey–Levi protocol for the harvest and expansion of human SCs: (1) the cleaned nerve fascicles were kept in suspension in serum-containing medium with forskolin and heregulin for 1–2 weeks; (2) the fascicles were treated with enzymes and dissociated without explantation and multiple replating of explants, thus eliminating these steps, (3) the cell suspension was plated on laminin-coated tissue culture dishes; and (4) the SCs were expanded using serum-containing medium with 2 micromolar forskolin and 10 nanomolar heregulin. The SCs were replated to new dishes each time they achieved confluence. The Casella protocol reliably provided up to a 1000-fold expansion of the initial population at purities of >90% SCs.

6.9 THE ATHAUDA PROTOCOL: MANUFACTURE OF A CLINICAL GRADE HUMAN SC PRODUCT The protocols described earlier were not yet suitable for use in trial subjects. Adaptation of methods and reagents was needed to meet FDA standards for the manufacture of clinical grade cellular products. Multiple studies were performed to convert our laboratory protocols for isolation and culture of human SCs into those that could be used in clinical trials. The extensive work described here was performed by, or under the supervision of, Dr. Gagani Athauda in a designated facility at The Miami Project to Cure Paralysis at the University of Miami Miller School of Medicine following standards of Good Laboratory Practice or in a certified current Good Manufacturing Practice (cGMP) facility at the Diabetes Research Institute, also at the Miller School of Medicine. This work has not been previously published due to intellectual property considerations. Standard Operating Procedures (SOPs) were written describing each step of the manufacturing process, from the acquisition of nerve to the formulation of the final product to be used in human subjects. In all, more than 30 SOPs were required to describe the complete manufacturing process. Because the reagents used up to and including those of Casella et al. (1996) were research grade reagents, alternative sources of reagents likely to meet FDA approval had to be identified and their activity in supporting SC growth demonstrated. Experiments were done to optimize the use of nerve dissociation enzymes, heregulin, laminin, and fetal bovine serum from different sources. Quantitative determinations were made of cell yields, SC purity, sterility, viability, and the rate of cell growth over several passages. During the course of these studies, a new procedure was developed for the automated quantification of cell numbers and purities. In addition, SC purities were improved by the development of a selective adhesion protocol,

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not involving the use of antimitotic agents or antibodies. Acceptable reagent substitutes were identified that met the criteria of being pathogen-free and/or being of GMP origin and that supported expected SC growth rates, yields, and purities. Additional studies included tests to determine if the final product was free of reagents used in the manufacturing process; the final SC product was reliably free of residual heregulin and laminin. The final product was composed of >90% SCs based on the expression of known SC marker proteins such as S-100 and the human nerve growth factor receptor p75. Remaining cells were identified as S100 negative, p75 negative fibroblasts. Chromosomal abnormalities were not detected in any of the products tested. For all these studies, cadaveric human sural nerves were obtained from the University of Miami Bone Bank or from the Miami-Dade County Medical Examiner’s Office. The complete human SC manufacture process was validated by the processing of sural nerves from three organ donors with the cooperation of surgeons at the University of Miami Life Alliance Organization. The sural nerves were received and processed in an FDA-approved cGMP facility. Based on the results of these studies, the manufacturing process was approved by the FDA for use in a Phase 1 SC clinical trial in 2012. Richard Bunge’s vision was being realized nearly 40 years later.

7 CLINICAL RESEARCH FOR SPINAL CORD INJURY It is imperative that trials are conducted on human subjects (Steeves et al., 2007). They are research studies that explore whether a medical strategy, treatment, or device is safe and effective for humans. The primary purpose of clinical trials is research, which requires that all research studies follow strict scientific standards (Zarin and Tse, 2008). These standards protect patients and help produce reliable and reproducible results. Clinical trials are one of the final stages of a series of carefully conducted animal research experiments (Easterbrook et al., 1991). The trials show how the approach affects a living body and whether it is a viable solution to pursue for human beings. Clinical trials that are not carefully planned and conducted should be avoided to minimize the risks and give a false sense of hope. SCOPE, the Spinal Cord Outcomes Partnership Endeavor, is an academic-industry partnership founded in 2006. Its mission is to enhance the development of clinical trial and clinical practice protocols that will validate therapeutic intervention for SCI leading to the adoption of improved best practices (Scope-sci.org). Its Steering Committee consists of representatives from Federal agencies, corporate partners, funding foundations, and clinical experts. SCOPE presents instructional courses at the ASIA and the International Spinal Cord Society annual meetings. About 282,000 people suffer from SCI in the United States. Among these, about half are tetraplegic. These individuals need extensive support and lifelong care, which may translate into as much as $3 million in health care costs over the lifetime of the individual (National Spinal Cord Injury Statistical Center, 2016). A SCI may occur following an accident, where a sudden traumatic blow to the spine fractures or

7 Clinical research for spinal cord injury

dislocates vertebrae. The damage to spinal cord tissue begins at the moment of impact due to displaced bone fragments, disc material, or ligaments (Middleton et al., 2008). This includes damage to the axons that carry signals up and down the spinal cord, between the brain and the rest of the body. With a complete SCI, axons cannot send signals beyond the injury with the consequence of no sensory and/or motor function remaining below the injury. With an incomplete injury, there will be some level of sensory and/or motor function below the injury. Better response time by first responders after an accident, more effective emergency care, and new modes of rehabilitation have improved the chances of a better outcome. New repair strategies such as cell transplantation to modify the deleterious injured spinal cord milieu are urgently needed to further improve outcome. SCI trials are complicated for two reasons: (a) spontaneous recovery and (b) the placebo effect. Soon after SCI, patients may be completely paralyzed. Spontaneous recovery is observed in patients who heal without any treatment; in rare cases, patients demonstrate remarkable recovery and regain their near normal health. But for others, the rate of recovery is greatest during the first 3 months of injury but often may continue for a year or more. It is difficult to determine whether the recovery has resulted from the treatment (especially if the treatment was dispensed immediately after the injury) or due to spontaneity. From this perspective, trials in more stabilized chronic subjects allow an easier assessment of improvement due to the treatment. The placebo effect plays a major role because patients often falsely believe in the positive impact of the treatment they receive and report improvement in their condition.

7.1 PRECLINICAL STUDIES TO GAIN FDA APPROVAL FOR A SC TRIAL In late 2007, it seemed feasible to explore the possibility of gaining FDA approval for a SC transplantation trial. Since 1990, we at the Miami Project to Cure Paralysis had generated extensive data demonstrating that SC transplantation supports spinal cord repair in rodents with SCI. We also had developed an efficient method for procuring large, essentially pure populations of human SCs from adult peripheral nerve for autologous transplantation (see above). Using mitogen-expanded SCs in severe combined immunodeficient mice and athymic nude rats, we demonstrated that human SCs survived and were capable of enhancing axonal regeneration and forming myelin after transplantation into animals with sciatic nerve transection (Levi and Bunge, 1994; Levi et al., 1994, 2002) and spinal cord transection (Guest et al., 1997). Nearly 50 people became involved in the preparation and execution of the first autologous adult human SC clinical trial: SC transplantation experts, neurosurgeons, SC preparers (in a cGMP facility), pain researchers, animal care and behavioral testing workers, nurses, psychologists, SCI educators, and regulatory consultants. A subgroup started meeting in December, 2007, to plan the approach. With time it was learned that the FDA would need to know if, after transplantation of human and rat SCs into rats, tumors formed, how far the transplanted SCs distributed, whether the cells caused toxicity, if the cells survived for 6 months and what the

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appropriate cell dose would be (including the minimum toxic dose). The exact mode of SC injection into the epicenter of the subject’s spinal cord lesion needed to be determined. The SC “product” needed to be prepared and characterized in a cGMP facility. Every substance used and every step performed to prepare the cells needed to be approved by the FDA. In order to answer these questions adequately for the FDA, 1000 additional rats were subjected to contusion injury, transplanted with SCs, and analyzed. Our rat SCI model experiments were supplemented with new information from spinal cord injured Yucatan minipigs and nonhuman primates that strengthened the Investigational New Drug (IND) application to the FDA. The larger minipig spinal cord, compared to that of the rat, enabled key studies to determine cell dose and SC injection parameters appropriate for human use (Benavides et al., 2011; Guest et al., 2011, 2013; Solano et al., 2011). Moderate thoracic SCI was reproducibly generated in 15–20 kg minipigs; the resulting injury approximated a severe incomplete human SCI. A safe delivery technique (involving a stereotaxic injection device) for implanting cells into the injured cord was developed. Highly purified, autologous, adult primate sural nerve SCs were transplanted into the uninjured cervical spinal cord and also into brainstem lesions in primates (Benavides et al., 2010; Maldonado et al., 2008; Tovar et al., 2009). Cell survival for 35–42 days and 6 months was observed as the SCs had been modified genetically to express green fluorescent protein. SC myelin was found in the implants. Migration of labeled cells did not exceed 2 mm from the site of transplantation. Much discussion was conducted to decide on the assessment methods to adequately evaluate the trial subject for a year following transplantation. Preparation of the application to the FDA was challenging. As an academic institution, millions of dollars to prepare for and conduct the trial needed to come from generous donors. The larger number of rats required for the preclinical studies necessitated hiring new personnel and training them. There was an enormous amount of paper work to generate the SOPs required for every step in every protocol. The characteristics of SCs manufactured in a cGMP facility, using only materials approved by the FDA, were delineated. The outcome of every rat and its cause of death had to be documented in writing. The entire brain and spinal cord of every rat required sectioning in order to scrutinize the potential migration of the transplanted SCs from the site of the implantation. The eyes of a certified Pathologist were required to search all the sections for tumors. Reports detailing all these preclinical preparations are in the process of being submitted for publication at this writing. The IND application was filed with the FDA in September 2011.

7.2 REGULATORY REQUIREMENTS TO MANUFACTURE SCs FOR TRIALS The FDA regulations require IND holders to follow the Code of Federal Regulations and utilize the University of Miami cGMP compliant cell manufacturing facility. This facility is registered with the FDA and accredited by the Foundation of

7 Clinical research for spinal cord injury

Accreditation for Cellular Therapy. The cGMP compliant cell manufacturing activities took place in an International Organization of Standards Class 7 production suite. To qualify the facility, three lots of human SC product were manufactured using the methods and controls described in the IND application and approved by the FDA. The facilities, four production suites totaling 2500 ft.2, have restricted access and all essential equipment is monitored through an alarm system. The Quality Assurance team maintains a comprehensive and controlled documentation system which includes document change control and approval records, as well as document histories. Documentation in this system includes SOPs, Certificates of Analysis, specifications for critical materials, supplies and reagents, and master batch production records.

7.3 THE FIRST SC CLINICAL TRIAL AT THE MIAMI PROJECT The trial was released from clinical hold by the FDA in July 2012 and by approval from the University of Miami Institutional Review Board in October, 2012. The first transplantation of the FDA-approved Phase 1 clinical trial to evaluate the safety of autologous adult human SCs injected into the injury epicenter of six subjects with subacute SCI was performed in December, 2012. The trial was an open-label, unblinded, nonrandomized, nonplacebo controlled study with a dose escalation design. The primary end point was to evaluate safety over 5 years following SC administration at one of three doses (5, 10, or 15 million cells, 92.2–98.7% SC purity, in 50, 100, or 150 mL culture medium) within 72 days of injury to participants with complete thoracic SCI. The primary trial center was the University of Miami/Jackson Memorial Hospital, where all baseline testing, transplantation surgeries, and follow-up were performed. Participant screening and suitability (for autologous donors) were determined as per Good Clinical Practices and documented. This involved the review of relevant medical records for risk factors and, pertinent to tissue processing, clinical evidence of relevant communicable disease agents and diseases. A two-stage informed consent design was utilized. Consent was obtained according to the Declaration of Helsinki for individuals expected to be eligible for screening to determine full eligibility based on the inclusion and exclusion criteria. This screening included an interview with a clinical psychologist specialized in SCI to ensure understanding of the nature and intent of the trial. The sural nerve harvest was then performed. Participants continued their standard inpatient rehabilitation course during the 3–5 weeks required for cell processing. During that time, participants also received education from multiple study team members about SCI in general, about SCs, and about the clinical trial in detail. Within 5 days prior to the scheduled transplantation, electrophysiology was performed to seek evidence for incomplete injury by assessing somatosensory and motor-evoked potentials. A second informed consent was then obtained for the remaining baseline testing (including a second interview with the clinical psychologist), SC transplantation, and outcome assessments during the first week posttransplantation and thereafter.

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7.4 SC PROCESSING AND TRANSPORT TO THE TRANSPLANTATION SITE A 15-cm segment of sural nerve was removed from the subject for SC preparation. The nerve was dissected and the fascicles dissociated, essentially as described above. The culture medium contained Dulbecco’s modified Eagle’s medium, fetal bovine serum, forskolin, human recombinant heregulin, L-glutamine, and gentamicin. The cells from the fascicles were washed and plated onto laminin-coated plates. The cells were fed with culture medium every 3 days until reaching 80–90% confluency. At that time, cells were enzymatically dissociated and subcultured until reaching passage 2. The viable cell counts of final harvests before transplantation were determined. The final cell product was washed with serum-free culture medium, collected by centrifugation, and maintained in the cold until transplantation. Several controls were employed throughout the manufacturing process to ensure that the product was essentially free of process-related contaminants. These controls included the wash steps and release testing of the final product. An aliquot of the SC product was preserved to identify the SCs and contaminating fibroblasts to assess the purity of the final product. The participant-specific SC product was transported to the hospital operating room for administration after checking the preset lot release testing criteria approved by the FDA. The final product was delivered in vials that were stored in temperature controlled, validated containers at a concentration of 100,000 cells/mL. The final product was assessed for cell counts and viability and found to be usable for 8 h.

7.5 TRIAL OUTCOMES The trial outcomes showed that it is feasible to perform a peripheral nerve harvest within 5–30 days of injury and perform an intraspinal transplant within 4–7 weeks of injury, even in a neurologically complete spinal cord injured paraplegic subject. These six subjects were monitored for 1 year during which time they showed no adverse effects specifically linked to the nerve harvest or cell transplant. Clinically significant neuropathic pain and muscle spasticity were no greater than expected for a natural course cohort. There was no evidence of additional spinal cord damage. Whereas no motor functional improvement was noted, there was evidence of conduction through the lesion based on electrophysiological evaluation. One subject became sensory incomplete by 6 months posttransplantation. It was concluded in this Phase 1 subacute trial that the transplantation of autologous SCs into the lesion epicenter is safe. Outcome assessments are continuing for a total of 5 years.

7.6 THE NEXT TRIAL A second autologous SC trial is underway in the Miami Project. This Phase 1 trial is being conducted to test safety of SC transplantation in subjects who have been spinal cord injured for at least a year. As mentioned earlier, any advantages of the cell

Acknowledgments

therapy may be more clearly seen than in subacutely injured persons as the subjects have likely stabilized. Other differences from the first trial are that subjects will be limited to those manifesting lesions no more than 3-cm long and no greater than 2 mL in volume. This should enable, with up to 200 million transplanted SCs, the lesion to be filled, providing a bridge across the lesion to promote new axon growth. The 10 participants will have been diagnosed as complete thoracic (2), incomplete thoracic (nonambulatory) (2), complete cervical (2) or incomplete cervical (4) and will be recruited into the trial in this order. Extensive work in the Miami Project has demonstrated the value of combining additional interventions with the transplantation of SCs to better improve outcome after injury (see above). This has led to the inclusion in this trial of a plan of exercise conditioning and locomotor training for 3 months before and 6 months after autologous SC transplantation. Future trials will involve other combination strategies with SC transplantation.

7.7 NEXT STEPS IN SC MANUFACTURE AND QUALITY ASSURANCE A next step to be able to qualify for a Phase III clinical trial is to develop in vitro potency assays to evaluate the biological activity of the human SCs prior to transplantation. Another important development to serve a number of clinical trials is to arrange for large-scale GMP manufacture of human SCs. Our long-term goal and our dedication to this vision involve a central processing facility, where production and further manufacturing processes will take place on a large scale. We will develop a scale-up bioreactor production method for clinical grade SCs that meets or surpasses our current clinical production needs. We will test bioreactor technology to identify the one that is ideal to make this process scalable while meeting or exceeding all quality assurance requirements. We will validate our bioreactor results through: (a) cell viability growth and morphology evaluation; (b) expression of cell surface markers and growth curves; and (c) karyotyping and cytokine profile analyses.

8 CONCLUSION It is feasible and safe to obtain a sural nerve biopsy, expand the number of SCs from the biopsy in a cGMP cell culture facility, transplant the cultured SCs into the spinal cord of a spinal cord injured subject, and monitor these subjects for a year or more with a variety of outcome measures.

ACKNOWLEDGMENTS The authors are grateful to all the staff, students, and postdoctoral fellows who contributed to the studies mentioned here. The team of Adrianna Brooks, Risset Rodriguez, and Maxwell Donaldson working with Drs. Gagani Athauda, Aisha Khan, and Patrick Wood performed

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outstanding work to develop the protocols for and the preparation of human SCs for clinical trial. The generation of rat SCs over many years was accomplished by Yelena Pressman. The Animal, Imaging, Electron Microscopy, and Viral Vector Cores in the Miami Project were crucial to accomplishing the fundamental and preclinical SC investigations. Anil Lalwani, a Regulatory Consultant, and Dr. Kim Anderson were helpful in many stages of gaining FDA approval for our first clinical trial. Dr. Anderson also offered comments to improve this chapter. Other key participants for the trials are W. Dalton Dietrich, PhD; James Guest, MD, PhD; Allan Levi, MD, PhD; Damien Pearse, PhD; Patrick Wood, PhD; Aisha Khan, MBA, PhD; Marina Dididze, MD, PhD; and Diana Cardenas, MD. Funding from the NINDS, the Miami Project, the Buoniconti Fund, the Christopher and Dana Reeve Foundation International Research Consortium, the Hollfelder Foundation, the State of Florida, and the Craig Neilsen Foundation (P.V.M.) is most gratefully acknowledged. M.B.B. is the Christine E. Lynn Distinguished Professor of Neuroscience. We thank Erika Suazo for excellent word processing.

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Takami, T., Oudega, M., Bates, M.L., Wood, P.M., Kleitman, N., Bunge, M.B., 2002. Schwann cell but not olfactory ensheathing glia transplants improve hindlimb locomotor performance in the moderately contused adult rat thoracic spinal cord. J. Neurosci. 22, 6670–6681. Tetzlaff, W., Okon, E.B., Karimi-Abdolrezaee, S., Hill, C.E., Sparling, J.S., Plemel, J.R., Plunet, W.T., Tsai, E.C., Baptiste, D., Smithson, L.J., Kawaja, M.D., Fehlings, M.G., Kwon, B.K., 2011. A systematic review of cellular transplantation therapies for spinal cord injury. J. Neurotrauma 28, 1611–1682. Thoma, E.C., Merkl, C., Heckel, T., Haab, R., Knoflach, F., Nowaczyk, C., Flint, N., Jagasia, R., Jensen Zoffmann, S., Truong, H.H., et al., 2014. Chemical conversion of human fibroblasts into functional Schwann cells. Stem Cell Rep. 3, 539–547. Thomas, P.K., 1970. The cellular response to nerve injury. 3. The effect of repeated crush injuries. J. Anat. 106, 463–470. Tovar, D.F., Maldonado, A.L., Benavides, F., Rodriguez, M.L., Guest, J., 2009. Analyses of lesion volume, demyelination and neuronal injury following stereotaxic radiofrequency lesioning in spinal cord and brain. Soc. Neurosci. Abs. 55, 3. Wood, P.M., 1976. Separation of functional Schwann cells and neurons from normal peripheral nerve tissue. Brain Res. 115, 361–375. Wood, P.M., Bunge, R.P., 1975. Evidence that sensory axons are mitogenic for Schwann cells. Nature 256, 662–664. Xu, X.M., Guenard, V., Kleitman, N., Bunge, M.B., 1995. Axonal regeneration into Schwann cell-seeded guidance channels grafted into transected adult rat spinal cord. J. Comp. Neurol. 351, 145–160. Xu, X.M., Chen, A., Guenard, V., Kleitman, N., Bunge, M.B., 1997. Bridging Schwann cell transplants promote axonal regeneration from both the rostral and caudal stumps of transected adult rat spinal cord. J. Neurocytol. 26, 1–16. Yazdani, S.O., Hafizi, M., Zali, A.R., Atashi, A., Ashrafi, F., Seddighi, A.S., Soleimani, M., 2013. Safety and possible outcome assessment of autologous Schwann cell and bone marrow mesenchymal stromal cell co-transplantation for treatment of patients with chronic spinal cord injury. Cytotherapy 15, 782–791. Zarin, D.A., Tse, T., 2008. Medicine. Moving toward transparency of clinical trials. Science 319 (5868), 1340–1342. http://dx.doi.org/10.1126/science.1153632. Zhou, X.H., Ning, G.Z., Feng, S.Q., Kong, X.H., Chen, J.T., Zheng, Y.F., Ban, D.X., Liu, T., Li, H., Wang, P., 2012. Transplantation of autologous activated Schwann cells in the treatment of spinal cord injury: six cases, more than five years of follow-up. Cell Transplant. 21 (Suppl. 1), 39–47.

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Recruitment of endogenous CNS stem cells for regeneration in demyelinating disease

6

Natalia A. Murphy*,†, Robin J.M. Franklin*,†,1 *Wellcome Trust-Medical Research Council Cambridge Stem Cell Institute, Cambridge, United Kingdom † University of Cambridge, Cambridge, United Kingdom 1 Corresponding author: Tel.: +44-1223-337642, e-mail address: [email protected]

Abstract Demyelinating diseases, such as multiple sclerosis (MS), are responsible for a significant portion of the neurological disability burden worldwide, especially in young adults. Demyelination can be followed by a spontaneous regenerative process called remyelination, in which new myelin sheaths are restored to denuded axons. However, in chronic demyelinating disease such as MS, this process becomes progressively less efficient. This chapter reviews the biology of remyelination and the rationale and strategies by which it can be enhanced therapeutically in acquired demyelinating disease.

Keywords Remyelination, Adult stem cells, OPC, Demyelination, Myelin, Oligodendrocyte, Multiple sclerosis

1 INTRODUCTION Despite the existence of adult central nervous system (CNS) stem and progenitor cells, the fully developed mammalian CNS is commonly viewed to have little to no regenerative capacity. However, the response to demyelination provides an excellent example of complete regeneration within the adult CNS. This chapter aims to discuss the CNS endogenous regeneration potential and ways of enhancing it, focusing specifically on demyelinating disease. First, we review the biology of myelination, and remyelination in response to a demyelinating insult. Next, we present the most important examples of Progress in Brain Research, Volume 231, ISSN 0079-6123, http://dx.doi.org/10.1016/bs.pbr.2016.12.013 © 2017 Elsevier B.V. All rights reserved.

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demyelinating disease and discuss why and how in these cases remyelination fails. Finally, we discuss the rationale and strategies of augmenting the endogenous regenerative potential of the CNS, and review recent advances in this field. We describe how targeting endogenous stem cells for regeneration is a viable and attractive alternative to cell transplant therapies in demyelinating disease.

2 OVERVIEW: MYELINATION AND REMYELINATION 2.1 THE MYELINATED CNS: AN EVOLUTIONARY MILESTONE Myelination, the ensheathment of axons in a compacted, lipid rich, multilayered insulating membrane, was an evolutionary milestone for craniate vertebrates—it allowed them to develop complex predatory and escape behaviors, while increasing their body size (Zalc et al., 2008). Myelin serves a multitude of functions in the CNS. Crucially, the myelin sheath creates a region of high resistance and low capacitance around the myelinated part of the axon, enabling saltatory conduction (Felts et al., 1997) and increasing axonal transmission speed 20–100 times (Nave and Werner, 2014), while optimizing neurotransmitter efficiency. As well as accelerating conduction, myelination offers trophic (Griffiths et al., 1998; Lappe-Siefke et al., 2003) and metabolic (F€ unfschilling et al., 2012; Lee et al., 2012) support to the axon, making it an essential neuroprotective element of the CNS. Finally, changes in myelination throughout adult life have recently been proposed as a mechanism of functional plasticity within the adult CNS, implicated in memory and learning (reviewed in Bergles and Richardson, 2016; Tomassy et al., 2016). These functions, together with the fact that white matter makes up over 40% of the human brain (Snaidero and Simons, 2014), point toward an integral role of myelin in the adult mammalian brain, as evidenced by the devastating effects of demyelinating diseases (see later).

2.2 DEVELOPMENTAL MYELINATION Myelin in the CNS is produced by oligodendrocytes. These cells arise from oligodendrocyte progenitor cells (OPCs) which proliferate, migrate, and differentiate, extending cytoplasmic sheaths toward axons which they are to myelinate (reviewed in Emery and Lu, 2015). Once axonal contact is established, oligodendrocyte sheaths enwrap the axon and compact around it forming mature myelin (reviewed in Simons and Nave, 2016). Each oligodendrocyte can extend up to 80 myelinating processes (Chong et al., 2012), and almost never myelinates neighboring paranodes within the same axon (Young et al., 2013). In rodents, myelination peaks around postnatal day 14 and is largely completed by day 60 (Rivers et al., 2008). In contrast, in humans developmental myelination is prolonged and peaks around the 3rd–5th year of life (Yeung et al., 2014), continuing into the second decade (Giedd et al., 1999).

2.3 THE OPC OPCs arise in multiple independent populations from distinct domains of the neuroepithelium during development (reviewed in Fancy et al., 2011a) and migrate extensively to populate the CNS and differentiate into oligodendrocytes. A proportion

2 Overview: Myelination and remyelination

of OPCs remains in the adult CNS in the undifferentiated state. These cells, characterized by the expression of NG2, A2B5, and Pdgfra (Horner, 2000; Pringle, 1992; Raff et al., 1983) are widespread in the gray and white matter, and amount to 5–8% of all the cells in the mature CNS (Levine et al., 2001). They are the main proliferating cell population in the CNS, and an essential reservoir of progenitors for remyelination (see later). As such, they have been viewed as adult stem cells of the CNS (Crawford et al., 2014). Indeed, they continue to proliferate throughout life, demonstrating robust capacity for self-renewal. Lineage tracing studies have confirmed their multipotency, showing that OPCs can differentiate into oligodendrocytes and astrocytes in development, and into oligodendrocytes, astrocytes, and Schwann cells (the neural crest-derived myelinating cells of the PNS) in adult life (Kang et al., 2010; Rivers et al., 2008; Zawadzka et al., 2010; Zhu et al., 2008). These features provide a strong basis to consider the OPC as a type of adult stem cell, a helpful framework within which to study their behavior in the context of cancer, aging, and regeneration. OPCs can also be generated in adult life from neural stem cells of the subventricular zone (SVZ) and have been shown to contribute to remyelination in the vicinity of the SVZ (Jablonska et al., 2010; Nait-Oumesmar et al., 1999). However, as the structure of the human SVZ is very different to the rodent one (Fietz et al., 2012; Lim and Alvarez-Buylla, 2016), the relevance of this finding to human remyelination is unclear. Finally, as well as a central role in myelination and remyelination, OPCs have been shown to play physiological roles in the resting adult CNS, notably in modulating synaptic transmission (discussed in Dimou and Gallo, 2015).

2.4 REMYELINATION: THE DEFAULT RESPONSE TO A DEMYELINATING INSULT In response to demyelination, the adult CNS regenerates myelin sheaths restoring saltatory conduction (Smith et al., 1979) and reversing functional deficits (Duncan et al., 2009; Jeffery and Blakemore, 1997; Liebetanz and Merkler, 2006) in a process called remyelination (Fig. 1). This is a unique example of regeneration in an otherwise poorly regenerating CNS and is the default outcome in both experimental models of demyelination and in naturally occurring CNS diseases, including those of humans (Lasiene et al., 2008; Patrikios et al., 2006; Smith and Jeffery, 2006). Remyelination is achieved by OPCs, and broadly follows the principles of developmental myelination, with some exceptions (Fancy et al., 2011a). In response to myelin and oligodendrocyte damage, resident astrocytes and microglia produce factors which initiate the inflammatory response and activate OPCs within and around the damaged area. Activated OPCs undergo a change in morphology (Levine and Reynolds, 1999), upregulate specific activation genes, reexpress developmental markers (Fancy et al., 2004; Moyon et al., 2015; Watanabe et al., 2004), and become more responsive to mitogens produced by surrounding cells (Hinks and Franklin, 1999). Using environmental cues, activated OPCs proliferate and migrate into and within the lesion. This constitutes the recruitment phase of remyelination. In the next phase of remyelination, OPCs exit the cell cycle and differentiate into

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Myelinated CNS white matter tract A Axon

OPC

Oligodendrocyte

Demyelinating insult

Demyelination

B

Remyelination Recruitment C

Differentiation D

Remyelinated CNS white matter tract E

FIG. 1 Remyelination in the central nervous system. The diagram shows a simplified scheme of white matter remyelination in the mammalian CNS, along with cross-sectional images of a (Continued)

3 Demyelination

oligodendrocytes. These cells establish contact with denuded axons, and extend cytoplasmic processes, which wrap around them and form a compact myelin sheath. Apart from the smallest diameter myelinated axons, remyelinated axons are characterized by a thinner myelin sheath relative to their diameter (i.e., higher g ratio). The functional significance of this finding is currently unknown. The g ratio appears to be unchanged if the remyelination is carried out by SVZ-derived OPCs, although this may be due to the small diameter of the axons of the corpus callosum (Stidworthy et al., 2003).

3 DEMYELINATION 3.1 MYELIN DISORDERS Many myelin disorders have been described in human and nonhuman species (reviewed in Duncan and Radcliff, 2016). These can be broadly divided into primary genetic disorders of myelination, acquired inflammatory (including autoimmune) disorders, toxic disorders, and traumatic loss of myelin. The myelin loss that follows primary axon degeneration is sometimes called secondary demyelination, but this is a misleading term for what should be referred to as Wallerian degeneration. Genetic disorders of myelination can be due to inherent errors in lipid metabolism (e.g., X-linked adrenoleukodystrophy), mutations of essential myelin genes (e.g., Pelizaeus–Merzbacher disease), lysosomal storage disorders (e.g., Krabbe’s disease), or mitochondrial mutations (e.g., Kearns–Sayre syndrome). As well as frank demyelination, these disorders often involve abnormalities in developmental myelination and lead to hypomyelination or dysmyelination. As the causes of these diseases are inherent to the oligodendroglial lineage, they are not amenable to endogenous remyelination enhancement and will not be discussed further.

FIG. 1—Cont’d myelinated, demyelinated, and remyelinated white matter tract, stained with toluidine blue to visualize myelin. Myelinated white matter tracts contain directionally aligned axonal fibers myelinated by oligodendrocytes (purple), and a widespread population of OPCs (blue) (A). Other cells present (microglia and astrocytes) have been omitted for clarity. Insult to oligodendrocytes or the myelin itself leads to demyelination—leaving axons denuded (B). In response to this remyelination ensues. Resident OPCs migrate to the lesion site and proliferate in the first phase of remyelination called recruitment (C). Subsequently, the OPCs differentiate into oligodendrocytes (D), which establish contact with axons in order to ensheath them with myelin. These newly formed oligodendrocytes recreate the myelin sheaths (E), which are significantly thinner than the original sheaths, but nonetheless restore saltatory conduction and reverse any functional deficits which arise due to demyelination.

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3.2 CONSEQUENCES OF DEMYELINATION True demyelinating disease is characterized by denuded but otherwise intact axons. This can happen either because the myelin sheaths themselves have been damaged, or due to death of oligodendrocytes that produce and maintain them. A common pathological alteration of the myelin sheath is vacuolation—fluid seen in extracellular space between layers of myelin. Vacuolation may in turn lead to myelin sheaths becoming dissociated from the axon, resulting in axon denudation. Denuded axons temporarily stall action potential transmission (Felts et al., 1997) and the axonal membrane must be modified to allow for slower and more energy demanding, continuous action potential conduction. In addition, naked axons are more vulnerable, as they no longer have the trophic and metabolic support and barrier of the surrounding myelin sheath. If the axon is not quickly remyelinated it remains vulnerable to degeneration, which in turn leads to irreversible functional deficits (Franklin et al., 2012).

3.3 ACQUIRED DEMYELINATING DISORDERS The most common demyelinating disease in humans, affecting 2–2.5 million people worldwide (Milo and Kahana, 2010), is multiple sclerosis (MS). MS is an autoimmune disease of the brain, optic nerve, and spinal cord, where focal lymphocytic infiltration leads to damage of myelin and axons (Compston and Coles, 2008). After an initial relapsing–remitting stage of the disease, it almost invariably progresses to a degenerative stage where chronic demyelination leads to axonal loss and permanent functional decline (Nave and Trapp, 2008), resulting in major disability and a 5–10 year reduction of life expectancy. The cause of MS is not currently known, but likely represents a complex interaction of environmental circumstance and genetic predisposition, as both environmental and hereditary risk factors have been identified. Although the pathology of MS has been extensively studied, the disease mechanisms also remain elusive. This may in part be due to the intrinsic heterogeneity of the disease between individuals and between lesions within a single individual. However, regardless of these diverse pathological mechanisms, a common feature is failure of remyelination resulting in neurodegeneration. Primary demyelination is also a feature of neurological disorders including traumatic spinal cord injury and ischemic disease. It is likely that in both cases primary demyelination is followed by remyelination (albeit occurring at a rate that is age dependent). Whether there is persistent demyelination leading to axonal degeneration in either situation is unclear and in our view unlikely.

3.4 EXPERIMENTAL MODELS OF DEMYELINATION In order to study the mechanisms of remyelination, the progression of inflammatory demyelinating disease and the potential for remyelination enhancement via therapeutic interventions, various experimental models have been developed.

3 Demyelination

Most of these models are based either on toxin-induced demyelination, or on immune-induced inflammation with variable degrees of demyelination. Toxin-induced demyelination can be achieved systemically or focally (reviewed in Blakemore and Franklin, 2008). Focally induced toxic demyelination has proven an invaluable tool for studying the biology of the remyelination process. It is usually achieved by microinjecting ethidium bromide or lysolecithin into large white matter tracts of rodents. This creates local reproducible lesions, which undergo synchronous demyelination and remyelination. These are sometimes described as noninflammatory models of demyelination; however, this is incorrect since, as with all injury models, they trigger an innate immune response, and the lysolecithin model also features a small involvement of adaptive immune cells. Such lesions are invaluable for dissecting different aspects of the remyelination process and how it responds to perturbations. However, these lesions remyelinate very efficiently, and although remyelination can be slowed by using aged animals (see later), it always proceeds to completion. Systemically delivered toxin-induced demyelination by oral administration of cuprizone is an alternative toxin model. The basic mode of action of cuprizone, a copper chelator, is presumed to be inhibition of oxidative phosphorylation. The reasons for its selectivity to particular regions of white matter are unknown, and it is presumed to influence other cells and processes in the body. This model is dependent on the species and strain of the animal used, the dosage of cuprizone, and the administration regimen. Repeat exposure can result in a model of chronic demyelination in which the acute inflammatory response is much subsided (Mason et al., 2004), a feature which together with the ease of administration makes it a popular although not necessarily optimum model. The second group of demyelination models aims to mimic the immunopathogenesis of MS, rather than isolate the neurobiology of the regenerative response to demyelination. The main model in this group is experimental allergic/autoimmune encephalomyelitis (EAE), although valuable viral models also exist (e.g., Theiler’s Murine Encephalomyelitis Virus). The basis of EAE is to induce an adaptive immune response to myelin, and this is done by immunization of the animal with myelin, or myelin proteins with the addition of Complete Freund’s Adjuvant (CFA). This creates lesions that partially resemble those of MS. Many versions of EAE exist, some developed to specifically mimic certain aspects of MS (e.g., relapsing–remitting EAE, secondary progressive EAE). Although helpful in testing therapeutic interventions that modulate the maladaptive immune driver of disease, EAE models are not helpful in studying remyelination per se. All these models have led to important discoveries and therapeutic developments (see later). However, it is often highlighted that species differences may lead to setbacks (although this is perhaps more true of immune cells than those of the CNS). Recently, an interesting new experimental model has been developed based on use of a humanized mouse. Goldman and colleagues have generated mice completely myelinated by human oligodendrocytes and populated by human OPCs by transplanting human neural progenitor cells into neonatal mice (reviewed

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in Goldman et al., 2015). It has been proposed, that inducing demyelination in these mice would enable direct observation and experimental perturbation of human OPC-mediated remyelination (Dietz et al., 2016).

4 FAILURE OF REMYELINATION 4.1 WHY DOES REMYELINATION FAIL? As well as disease-specific reasons for remyelination failure (see later), the efficiency of the process is dependent on a host of generic (nondisease specific) characteristics (Franklin and Ffrench-Constant, 2008) like genetic background (Bieber, 2008), age (Shields et al., 1999), and gender (Li et al., 2006). Of these factors, age seems to be the most influential, and the most important for the progression of MS (Confavreux and Vukusic, 2006). All regenerative processes decrease with age, largely due to decreased function of adult somatic stem and progenitor cells (Sousounis et al., 2014), and remyelination is no exception. Both recruitment and differentiation phases of remyelination are slower in aged animals (Sim et al., 2002).

4.2 AT WHAT STAGE DOES REMYELINATION FAIL? In order for efficient remyelination to occur, activated OPCs must be recruited into the lesion and differentiate into myelinating oligodendrocytes. Failure at any point of this progression will result in a delay or arrest in remyelination. Numerous studies have attempted to elucidate the crucial point at which the process fails. It is conceivable that remyelination would fail due to an age-associated decline in the numbers of adult OPCs. However, there is no good evidence to support this and the density of OPCs appears to remain stable throughout adult life (e.g., Sim et al., 2002). A second possible reason for remyelination failure is that there is a failure of recruitment. This possibility is illustrated by the existence of MS lesions completely devoid of OPCs (Boyd et al., 2013) which is perhaps related to the presence of antibodies against the OPC antigen NG2 (Niehaus et al., 2000). Aged OPCs are recruited more slowly into lesions, possibly because they take more time to respond to growth factors and are less responsive to migration cues (Sim et al., 2002). The lesion size will also influence the time it requires for OPC recruitment, especially if they are all recruited from outside the lesion boundary (Chari et al., 2003). However, though undoubtedly affected by aging, at least in rodents failure of recruitment does not seem to be the rate-limiting step of remyelination, as augmentation of recruitment into lesions in aged animals does not cause an improvement of the rate of remyelination (Woodruff et al., 2004). This study points to the possibility that OPC differentiation is the rate-limiting step of remyelination with aging—a hypothesis further strengthened by the existence of chronic MS lesions with OPCs and preoligodendrocytes seemingly unable to differentiate or myelinate (Chang et al., 2002;

4 Failure of remyelination

Kuhlmann et al., 2008). Due to this, many studies aiming to find a therapeutic target enhancing remyelination have concentrated on promoting OPC differentiation. A recent hypothesis based on observations on human OPCs (discussed in Dietz et al., 2016) proposes that OPC differentiation efficiency depends on OPC density in the lesioned area. It has been noticed, that transplanted human OPCs tend to myelinate the axons of shiverer mice efficiently only once they reach a density of >30,000 cells/mm2. As density-dependent differentiation has been described in vitro (Rosenberg et al., 2008) and a mechanism for OPCs sensing their local density has been proposed (Hughes et al., 2013), it is a possible mechanism of differentiation regulation. The authors propose a lower limit of OPC density of 10,000 cells/mm2 needed for differentiation to continue, based on OPC densities in normal adult white matter, periplaque regions, and in regions of chronic demyelination. If this is the case, then experimental rodent models of remyelination almost never fall short of this limit; therefore, density is never the rate-limiting step in those models. Though not directly proven, this model is interesting as it functionally links the recruitment and differentiation phases of remyelination.

4.3 REMYELINATION FAILURE: INTRINSIC PROPERTIES OF REMYELINATING CELLS VS EXTRINSIC PROPERTIES OF THE ENVIRONMENT Regardless of the point at which remyelination fails, the failure of this process can always be traced to intrinsic changes within the OPC or extrinsic disturbances within the environment. Though a clear distinction between the two is difficult, it is a helpful framework with which to discuss remyelination failure. As adult stem cells show a deregulated epigenetic and transcriptional profile with aging (Chambers et al., 2007; Liu et al., 2013), we can expect a similar process to be going on within the adult OPC. Indeed, changes within the OPC itself have been documented—aged OPCs transplanted into a lesion environment show a slower recruitment compared to younger controls (Chari et al., 2003) and in older mice OPCs are more likely to differentiate into astrocytes in lesions (Doucette et al., 2010). Moreover, epigenetic dysregulation in aged OPCs has also been observed (Shen et al., 2008). In aged OPCs, insufficient recruitment of HDACs (histone deacetylases) to promoter regions of differentiation inhibitors like Hes5 leaves them active and stalls differentiation. As well as intrinsic disturbances, studies have identified a myriad of extrinsic sources which might contribute to remyelination failure (van Wijngaarden and Franklin, 2013). This is often a consequence of the presence of inhibiting factors within the lesion, or the absence of essential promoting factors that are normally present within the remyelinating environment.

4.3.1 Dysregulation of the innate immune response The innate immune response is crucial for efficient remyelination (reviewed in McMurran et al., 2016; Miron and Franklin, 2014). Following a demyelinating injury, microglia and peripheral macrophages migrate into the lesion attracted by

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factors released by damaged cells. They phagocytose myelin debris and secrete factors stimulating OPC recruitment and differentiation. With aging, both of these functions are reduced (Miron et al., 2013; Natrajan et al., 2015; Ruckh et al., 2012). This stalls remyelination in a twofold manner: first, by reducing the amount of factors directly regulating OPC function and, second, by reducing the efficiency of phagocytic removal of myelin debris, which inhibits differentiation (Kotter et al., 2005). These effects can be experimentally mimicked by reducing macrophage number or activation, or by injecting myelin debris into a lesion (Kotter, 2006; Kotter et al., 2001).

4.3.2 Dysregulation of the migratory cues Both in development and in remyelination, OPCs use migratory cues to find the area they are to (re)populate. For example, class 3 semaphorins have been described to regulate developmental OPC migration in an antagonistic fashion—SEMA3A serving as a repellent and SEMA3F as an attractant (Spassky et al., 2002). Both were found to be expressed in MS brains and in response to experimental demyelination (Williams et al., 2007). The imbalance of these factors could be the reason for remyelination failure, especially in MS lesions devoid of OPCs (Boyd et al., 2013). In addition to these guidance functions, SEMA3A inhibits OPC differentiation in vitro and in CCP lesions (Syed et al., 2011), further implicating overexpression of this molecule as a potential cause of remyelination failure.

4.3.3 Inhibitory extracellular matrix molecules within the lesion The adult CNS is devoid of a conventional extracellular matrix (ECM); however, ECM molecules are expressed in development and reexpressed following CNS insults (Colognato and Tzvetanova, 2011; Zhao et al., 2009). Glycosaminoglycans (GAGc) are among the reexpressed ECM molecules within lesions. Two types of GAGs are implicated in inhibiting remyelination: hyaluronans and chondroitin sulfate proteoglycans (CSPGs). Hyalouronan is found accumulating in spinal cord injury and MS lesions (Back et al., 2005). In response to demyelination, high-molecular weight hyaluronan is synthesized by astrocytes in MS and EAE. Oligodendrocyte-expressed hyaluronidases attempt to degrade it, and the products of this degradation interact with the TLR2 receptors on oligodendroglia leading to OPC maturation arrest (Preston et al., 2013; Sloane et al., 2010). CSPGs, also found in MS lesions (Sobel and Ahmed, 2001; van Horssen et al., 2006), have similarly been shown to arrest OPC process outgrowth and remyelination through interacting with the PTPs receptor (Pendleton et al., 2013). Importantly, this inhibition can be reduced by administration of chondroitinase ABC (Lau et al., 2012; Pendleton et al., 2013) or by inhibiting CSPG synthesis (Keough et al., 2016), proving that the CSPG-mediated differentiation block was reversible.

4.3.4 Axon–oligodendrocyte interactions It seems likely that the longer an axon remains demyelinated, the more vulnerable to degeneration it becomes (Franklin et al., 2012). Demyelinated axons express PSA-NCAM on their surface, a neural cell adhesion molecule that must become

4 Failure of remyelination

downregulated if efficient myelination is to occur (Charles et al., 2000; Fewou et al., 2007). PSA-NCAM is expressed on the surface of 11–19% of demyelinated axons in MS lesions (Charles et al., 2002), potentially inhibiting their remyelination. Recently, it has also been shown that in order to remyelinate efficiently, axons must be electrically active and release glutamate. Blocking neuronal activity, axonal vesicular release or AMPA receptors in demyelinated lesions reduces remyelination and increases the number of undifferentiated OPCs within lesions (Gautier et al., 2015).

4.4 EFFICIENT REMYELINATION: THE ROLE OF CELL SIGNALING PATHWAYS Remyelination failure often involves the dysregulation of molecular pathways essential for oligodendrocyte differentiation and myelination. These pathways integrate extrinsic information from the extracellular signaling molecules with the intrinsic state of the cell, to drive, permit, or inhibit efficient remyelination. For this reason, many have been researched and targeted for therapeutic enhancement of remyelination. When cell signaling pathways involved in remyelination control, such as the Wnt pathway (reviewed in Guo et al., 2015), the Notch pathway (Brosnan and John, 2008), or the BMP pathway (Grinspan, 2015) began to be discovered, it was hoped that simple inhibition or enhancement of a given pathway would enhance remyelination in disease states (Fancy et al., 2010). Increasingly however, the complexity, crosstalk, and nonbinary nature of these signals is beginning to be recognized, and the reality is much more challenging. A summary of the role of Wnt in OPC differentiation illustrates this. The canonical Wnt signaling pathway is highly conserved and crucial for the specification, differentiation, and growth of many cells (van Amerongen and Nusse, 2009). In mammals, extracellular Wnt ligands bind to Frizzled receptors and their coreceptors, and through the LRP5 and LRP6 proteins act to stabilize intracellular b-catenin (which, in the absence of Wnt signal is complexed and targeted for proteasomal degradation). b-catenin can either remain attached to the cell membrane, or translocate into the nucleus and interact with TCF/LEF transcription factors to stimulate gene expression. Strong evidence emerged to suggest that canonical Wnt signaling inhibits OPC differentiation in remyelination, and therefore inhibiting the pathway would likely enhance remyelination. An in situ-based transcription factor screen of the lysolecithin spinal cord lesion revealed expression of Tcf4 in Olig2-positive cells during remyelination, implicating Wnt pathway involvement. Indeed, constitutive activation of b-catenin in Olig2cre/DA-Cat mice delayed remyelination of lysolecithin lesions (Fancy et al., 2009), while Axin2 (one of b-catenin degradation complex factors) double knockout mice also showed delayed remyelination in comparison to wild type controls, despite exhibiting normal OPC recruitment (Fancy et al., 2011b). Moreover, treatment of wild type mice with XAV939—an Axin2 stabilizer (and thereby a promoter of b-catenin degradation), significantly increased the rate of OPC differentiation within lysolecithin lesions (Fancy et al., 2011b). These studies,

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together with similar findings in developmental myelination and reports of increased Wnt pathway gene and protein expression in the context of MS lesions (Han et al., 2008; Lock et al., 2002) indicate inhibitory effects of high Wnt tone on OPC differentiation. However, addition of Wnt stimulates oligodendrocyte maturation in culture, while overexpression of nucleus-located b-catenin increases proteolipid protein promoter activity (Tawk et al., 2011). Furthermore, remyelination is stalled in Tcf4 null mice (Ye et al., 2009). In an attempt to reconcile these seemingly contradictory results, a model where Tcf4 triggered opposing actions depending on the whether it was coupled with b-catenin or HDAC1/2 has been proposed—as histone deacetylases were found to compete with b-catenin for binding to Tcf4 (Ye et al., 2009). Alternatively, it has been hypothesized that Tcf4 activation is not necessarily always coupled to Wnt/ b-catenin signaling and increased Tcf4 activation in OPCs failing to differentiate could be a secondary effect rather than a cause of differentiation failure in the first place (Hammond et al., 2015). A generalized model of Wnt signaling in remyelination has been proposed by Guo and colleagues, postulating that the outcome of Wnt signaling depends on the differentiation stage of the cell. The transition of OPC to immature oligodendrocyte is hypothesized to require a low level Wnt tone, and so a pathologically high Wnt tone will cause differentiation inhibition (Fancy et al., 2014). Meanwhile further maturation to myelinating oligodendrocyte is proposed to require b-catenin-independent Tcf4 activation, uncoupling Tcf4 action and expression from Wnt tone (Guo et al., 2015). To further complicate matters, noncanonical Wnt pathways could also play a role in myelination, and Wnt signaling has been proven to crosstalk with the Akt/mTOR pathway in the oligodendrocyte lineage (Tyler et al., 2009) and doubtless with many more. As with the Wnt pathway, many other critical signaling pathways have been discovered to modulate remyelination in a diverse, context-dependent manner (Gaesser and Fyffe-Maricich, 2016), and although a predominant effect is often present, such pleiotropism presents challenges for developing therapeutic interventions.

5 ENHANCING ENDOGENOUS STEM CELLS: CURRENT AND FUTURE THERAPIES In establishing effective therapies for myelin disease, two strategies have been pursued: (1) enhancing endogenous remyelination and (2) providing exogenous myelinating cells by cell transplantation. In the context of intrinsic disorders of myelin assembly or metabolism, providing an exogenous source of myelination is the logical approach. However, in the case of acquired demyelinating disorders, remyelination failure is likely more due to pathological changes of the lesion environment, than a failure of the remyelinating cells themselves. Crucially, intrinsic changes in failing

5 Enhancing endogenous stem cells: Current and future therapies

OPCs have been proven to be, in principle, reversible and could be overcome by environment modification (Ruckh et al., 2012). Therefore, it makes sense to attempt enhancement of endogenous remyelination when considering therapies for acquired demyelinating disorders. Although the endogenous enhancement approach circumvents many problems related with cell transplantation (such as optimizing cell delivery, considering the ethics, immunological implications and sustainability of the cell source, etc.), it comes with its own challenges. For any remyelination enhancing therapy to be effective, it must be delivered efficiently to the site of action (i.e., past the blood–brain barrier), act specifically on the cells of interest, and reliably trigger the appropriate response. As with all remyelinating therapies, an objective, standardized, and relatively noninvasive way of assessing outcomes must be available in order to compare and evaluate outcomes of proposed treatments against each other and the current clinical gold standard.

5.1 REJUVENATION AS AN APPROACH TO ENHANCE REMYELINATION Aging is one of the crucial contributory factors of remyelination failure. Experiments using heterochronic parabiosis of old and young animals have proven that lesions in old animals can be efficiently remyelinated by old OPCs, provided they are exposed to a young environment (Ruckh et al., 2012). These experiments further demonstrated that the rejuvenation effects were in part due to increased myelin debris phagocytizing efficiency of young macrophages and that serum-derived factors from the young circulation also contribute to the rejuvenation effect. Much recent research has centered on the hallmarks of aging and interventions designed to reverse or slow it (Lo´pez-Otı´n et al., 2013). It seems likely that aging will affect the inherent behavior of adult OPCs and indeed, age-related epigenetic dysregulation of OPCs has been demonstrated (Shen et al., 2008). Physiological interventions resulting in decreased anabolic signaling (like dietary restriction and endurance training) or drugs that mimic such interventions, extend lifespan in animals (Fontana et al., 2010; Harrison et al., 2009), and slow the progression of numerous age-related disease processes including neurodegeneration (Duan and Ross, 2010). Given that these strategies have been demonstrated to enhance the function of other adult stem cells (Cerletti et al., 2012; Chen et al., 2003; Yilmaz et al., 2012) it seems very likely that these interventions will be beneficial in enhancing remyelination.

5.2 THE TRANSLATIONAL PATHWAY: FROM BENCH TO BEDSIDE Many remyelination enhancing drugs currently in preclinical testing or clinical trials are the result of a systematic search for therapeutic targets. The approach is based on the identification of nonredundant pathways and critical downstream effectors, augmentation or inhibition of which leads to enhancement of remyelination. These

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pathways, or their modulators, are often identified through a screen-based study. If the validation data are promising, the finding is further tested in various experimental disease models, often where the translatability of the modulating compounds, the routes of delivery, and potential side effects are beginning to be explored. Finally, if animal studies show a robust effect, the experimental compound is taken to clinical trials to assess the safety, tolerability, and efficacy in humans.

5.2.1 Lingo1 inhibition In search for pathways inhibiting axon regeneration, a receptor complex-inhibiting neurite outgrowth in response to myelin debris was described. This complex, composed of the Nogo66 and p75NTR receptors, was found to inhibit neurite outgrowth through RhoA inhibition (Fournier et al., 2001). In an attempt to identify the NgR1 complex activators by screening uncharacterized CNS-specific proteins, the LRR and Ig domain-containing Nogo Receptor interacting protein—LINGO-1, was found (Mi et al., 2004). Later, this protein was shown to inhibit oligodendrocyte differentiation via homophilic extracellular interactions and by blocking activation of ErbB2 (Lee et al., 2014; Mi et al., 2005). Lingo1 is expressed in the CNS and its inhibition using multiple approaches in culture enhances OPC differentiation (Mi et al., 2005). Lingo1 inhibition was also found to enhance dorsal root ganglion (DRG) coculture myelination (Lee et al., 2007). Furthermore, OPCs from Lingo1 knockout animals exhibited enhanced differentiation in vitro (Mi et al., 2005). In animal studies, Lingo inhibition was found to enhance remyelination in lysolecithin and cuprizone models of demyelination. Lingo1 knockout mice were found to exhibit resistance to EAE development, while inhibiting Lingo1 in wild type EAE mice, mitigated disease severity leading to improved regeneration and function (Mi et al., 2007, 2009). However, increased remyelination efficiency in Lingo1 knockout mice has not been reported neither has there been a systematic analysis of Lingo1 expression in oligodendrocyte lineage cells in the various models of CNS demyelination and remyelination. Based on the preclinical data, BIIB033, a fully humanized IgG1 monoclonal anti Lingo1 antibody was developed. The antibody was found to be safe and well tolerated in Phase I clinical trials, and two phase II trials were set up. The first, a trial of anti Lingo1 therapy in optic neuritis, failed to reach its primary endpoint (retinal fiber thickness preservation), although the therapy did show improvement in latency of Visually Evoked Potentials, which was one of the trials secondary endpoints (NCT01721161). The second study, a Phase II BIIB033 trial in relapsing–remitting MS (NCT018641487) was recently reported to have missed its primary endpoint.

5.2.2 RXR agonists Retinoid X nuclear receptors (RXRs) and especially RXRg, were found to have increased levels of expression in transcriptional profiling of remyelinating focal toxic lesions, clustering together with other genes related to oligodendrocyte differentiation. Upon further analysis, it was revealed that pharmacologically antagonizing

5 Enhancing endogenous stem cells: Current and future therapies

RXRg inhibited OPC differentiation and myelination in vitro. Furthermore, stimulation of RXRg through administration of the RXR agonist, 9cis retinoic acid (9cRA) accelerated remyelination in aged animals (Huang et al., 2011). Subsequently, it was shown that activation of the VDR–RXR heterodimer enhances OPC differentiation in vitro, and pharmacologically blocking VDR impaired remyelination, and partially abrogated the 9cisRA effect on OPC differentiation in vitro (de la Fuente et al., 2015). These findings underscore the importance of RXR signaling in the regenerative phase of MS, as well as the well-documented role of vitamin D in disease susceptibility (Burton and Costello, 2015). However, RXR signaling in the CNS is not restricted only to oligodendrocyte lineage cells. A recent study by Natrajan et al. (2015) has shown that RXR also influences macrophages in the CNS. Previously, it had been established that aged macrophages and monocytes within the lesion are not as efficient in myelin debris phagocytosis as young monocytes (Ruckh et al., 2012; see earlier). When the transcriptional profile of these cells was compared to young ones, the expression levels of genes related to the RXR pathway were found to be significantly decreased. Furthermore, when RXR signaling was stimulated in these cells, through treatment with bexarotene (an RXR agonist), these cells displayed a rejuvenated phenotype, which much improved myelin clearance. As predicted by these experiments, when animals with monocyte-specific RXRa knockout were subject to a focal demyelinative lesion, remyelination was delayed, most probably due to a delayed clearance of myelin debris (Natrajan et al., 2015). This study provided a second, indirect, oligodendroglia independent way in which RXR signaling stimulation could augment remyelination. On top of this, stimulating RXR in EAE was shown to improve EAE symptoms, via T cell modulation (Chandraratna et al., 2016). As RXR agonists are extensively studied for therapeutic use in cancer and metabolic disease, with one, bexarotene, already in clinical use, RXR signaling seems to be a very attractive therapeutic target for enhancing remyelination, especially due to its multitargeted beneficial actions. No clinical trials have been publicized to date, but experimental evidence makes a clinical trial very likely in the foreseeable future.

5.3 DRUG REPURPOSING FOR REMYELINATION An alternative way of finding interventions enhancing remyelination is to screen libraries of already existent FDA-approved compounds or track comorbid patient data for potential remyelinating effects of drugs already in clinical use. This approach has the advantage of circumventing the need for safety evaluation of the treatment, and greatly facilitates the introduction into clinical trials. However, this often means the mechanisms of action for the intervention are unclear at the time of the discovery, and sometimes these compounds turn out to act in ways other than enhancing remyelination. One interesting example of this is fingolimod (FT7720). Fingolimod was the first approved oral disease-modifying drug for relapsing–remitting MS (Brinkmann et al.,

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2010). It was assumed to act though attenuating inflammation in MS, as it was shown to inhibit lymphocyte dissemination from secondary lymphoid organs in EAE, through chronic binding and irreversible internalization of sphingosine-1-phosphate receptors (Kataoka et al., 2005; Papadopoulos et al., 2010). However, further experiments also revealed its unexpected cytoprotective effects on oligodendroglia. Fingolimod was found to protect oligodendrocytes in culture and in toxic lesions (Kim et al., 2011; Miron et al., 2008) and to enhance remyelination in cerebellar slice cultures (Miron et al., 2010). In light of these findings, and the fact that fingolimod slows brain atrophy in RRMS (Barkhof et al., 2014), a phase III clinical trial of fingolimod on primary progressive MS was set up. Unfortunately, it failed to find a therapeutic effect (Lublin et al., 2016). However, the established therapeutic benefits of fingolimod may well be due to its action on oligodendroglia as well due to its immunomodulatory properties. A systematic approach was recently employed in a large-scale phenotypic screen of drugs with a proven clinical safety profile. These compounds were tested on mouse epiblast stem cell-derived OPCs, and screened for extent of differentiation enhancement in vitro, by image quantification of myelin protein expression. This screen resulted in the identification of many compounds including clobetasole and miconasole—both in current clinical use for other indications. Follow-up studies confirmed differentiation enhancement by these drugs in organotypic cerebellar slice cultures, developmental myelination, and in a toxin demyelination model, and proposed a mechanism of action for both drugs, promoting them as likely targets for clinical studies (Najm et al., 2015). Another high-throughput screen, conducted on primary rat optic nerve-derived OPC cultures similarly aimed to find OPC differentiation enhancing drugs by measuring MBP expression in cultured cells treated with around 100,000 different active compounds (Deshmukh et al., 2013). Benztropine, a muscarinic antagonist used clinically in Parkinson’s disease treatment, was found among the most effective differentiation accelerating substances, and its effects were confirmed in EAE and cuprizone animal models. Antimuscarinic compounds were also implicated as myelination enhancers in a novel screening approach using micropillar arrays, engineered specifically to optimize high-throughput OPC differentiation screens (Mei et al., 2014). Here, the authors validated the efficacy of Clemastine, an antihistamine with antimuscarinic properties, and confirmed that it promoted remyelination in the rodent lysolecithin model. Clesmastine is currently undergoing phase 2 clinical trials for relapsing-remitting MS and neuromyelitis optica (NCT02040298, NCT02521311).

5.4 AUTOANTIBODIES: THE SOLUTION FROM WITHIN An interesting approach to enhancing remyelination is through the use of autoantibodies. These naturally occurring antibodies, present in the serum in the absence of stimulation by foreign antigens, have been found in almost all vertebrate species. First reported in 1966 (Boyden, 1966), antibodies are regarded as systemic

6 Concluding remarks

surveillance molecules that have been found to prevent autoimmunity by enforcing B-cell central tolerance induction (Nguyen et al., 2015). In contrast to conventional antibodies, autoantibodies bind to diverse antigens with low affinity and exhibit a short half-life. Remyelination enhancing autoantibodies were discovered by accident, when antibodies against myelin components were found to enhance remyelination in TMV-infected animals (Lang et al., 1984; Rodriguez et al., 1987) rather than worsen the disease. Following from this, autoantibodies to oligodendrocytes and myelin were found in human serum samples. After the confirmation of remyelination enhancement in vivo by two of the discovered autoantibodies, the var sequence of one of them was cloned to generate a humanized monoclonal antibody rHIgM22 (Mitsunaga et al., 2002; Warrington et al., 2007). The antibody was found to act by inhibiting apoptosis and oligodendrocyte differentiation as part of an extracellular complex containing PDGFra, aVb3 integrin, and Lyn kinase (Watzlawik et al., 2010). rHIgM22 promoted survival and proliferation of OPCs in vitro, but only in mixed brain cultures and not in pure OPC cultures, underscoring the importance of factors released by other glial cells in the antibody complex activation (Watzlawik et al., 2013). Phase I trials confirmed the predictions of these antibodies being very safe and well tolerated. The presence of Ab at the lesion site (and hence their efficient crossing of the blood–brain barrier) was confirmed by finding the Ab in CSF samples of every patient in the trial (NCT01803867). Following the success of safety trials, phase II trials are highly anticipated, especially as rHIg22 is one of the few clinically trialed remyelination enhancers that seem to act by promoting OPC survival and enhancing proliferation, rather than supporting differentiation.

6 CONCLUDING REMARKS Remyelination is a remarkable example of CNS regeneration. Studying the biology of this process, along with the conditions in which it fails, has led to important discoveries that are increasingly likely to become translatable into disease-modifying therapies (Table 1). Crucially, in acquired demyelinating disorders, it seems that remyelination failure is not insurmountable, and enhancing endogenous remyelination by therapeutic intervention may soon be achievable. A continued effort to identify the essential pathways and downstream effectors of the OPC differentiation process is needed to increase the range of potential therapeutic targets. Although we now know more than ever before about remyelination, questions still remain. Is remyelination as robust as developmental myelination in terms of supporting long-term axonal integrity? Can remyelination capacity be irreversibly exhausted? Is Schwann cell remyelination in the CNS a desired phenomenon? These and other questions will need to be answered, before we can fully comprehend the nature and harness the function of endogenous remyelination.

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Table 1 Compounds Currently Undergoing Clinical Trials With Possible Effects on Endogenous Remyelination Intervention Lithium carbonate

Clemastine fumarate

General Description and Mode of Action

Evidence of Influence on Remyelination

Used to treat manic phase of bipolar disorder. Mode of action uncertain, though presumed to act by inhibiting the activity of PKC in the brain Antihistamine and anticholinergic used for treating allergy

LiCl shown to stimulate myelin gene expression in oligodendrocytes and promote remyelination in tissue slices (Meffre et al., 2015) Identified to enhance oligodendrocyte differentiation in screen, and validated to enhance remyelination in vivo in focal toxic demyelination (Mei et al., 2014) Hypothesized to enhance myelin formation (Sedel et al., 2016)

Biotin (MD1003)

A.k.a vitamin B7, a coenzyme for carboxylase enzymes essential for fatty acid metabolism and gluconeogenesis

Quetiapine fumarate

Atypical antipsychotic and mood stabilizer, used to treat schizophrenia, bipolar disorder, and depression. Mode of action remains uncertain Humanized autoantibody shown to increase oligodendorcyte survival and hypothesized to enhance remyelination Synthetic form of thyroid hormone. TH acts in a diverse manner on many cells to regulate and integrate the metabolism of the entire organism

rHIgM22

Liothyronine

Shown to enhance remyelination in cuprizone-induced demyelination (Zhang et al., 2012) and EAE (Mei et al., 2012) Promotes OPC survival in vitro, in the presence of other glia cells (Watzlawik et al., 2013) Thyroid hormone has been shown to promote OPC differentiation in vitro and to enhance remyelination in EAE (reviewed in Calzà et al., 2005)

Clinical Status Completed pilot study in progressive MS (NCT01259388). Awaiting results

Ongoing phase 2 trials in RRMS and optic neuritis (NCT02040298, NCT02521311) Two studies have shown a beneficial effect in progressive MS (Sedel et al., 2015; Tourbah et al., 2016). Phase 3 trials ongoing (NCT02936037) Recently started phase 1/2 trial (NCT02087631)

NCT01803867 phase 1 study awaiting results. NCT02398461 study currently recruiting participants Two phase 1 studies recruiting participants: NCT02506751, NCT02760056

Domperidone

GSK239512

Bexarotene

vx15-2503

BIIB061 Olesoxime GNbAC1

Peripherally selective dopamine D2 receptor agonist, used to relieve nausea, increase prolactin release, or increase GI peristalsis Synthetic histamine H(3) receptor antagonist Retinoid X receptor gamma agonist. RXR family of nuclear receptors are transcriptional regulators in virtually every cell in the body Humanized antisemaphorin 4D antibody. SEMA4D (CD100) is an axonal guidance molecule Biogen proprietary compound, undisclosed mechanism of action Cholesterol-like synthetic mitochondrial pore modulator Humanized antibody against envelope protein of MSRV (multiple sclerosis-associated retrovirus)

Prolactin enhances OPC differentiation and remyelination in mice (Gregg et al., 2007)

Two phase 2 studies ongoing: NCT02308137, NCT02493049

In vitro screen on primary rat OPCs identified this drug as enhancing OPC differentiation (GSK) Upregulating RXRg signaling was shown to accelerate remyelination in focal toxic demyelinating lesion (Huang et al., 2011)

Phase 2 trial (NCT01772199), small positive effects observed (Schwartzbach et al., 2016) Ongoing phase 1 trial (EU2014003145-99)

SEMA4D has been shown to inhibit oligodendrocyte differentiation (Yamaguchi et al., 2012) and induce process collapse (Giraudon et al., 2004) Reported to be an “oral remyelinating compound” (Biogen) Shown to accelerate remyelination in animal models (Magalon et al., 2012) The viral envelope protein has been shown to inhibit OPC differentiation (Kremer et al., 2013)

Phase 1 trial completed, awaiting results (NCT01764737)

Phase 1 trial completed, awaiting results (NCT02521545) Phase 1 trial awaiting results (NCT01808885) Phase 2 study currently recruiting participants (NCT02782858)

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ACKNOWLEDGMENTS We would like to thank Ludiovica Di Canio, Bjoern Neumann, and Alerie Guzman de la Fuente for help in reviewing the content and form of this chapter.

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Yeung, M.S.Y., Zdunek, S., Bergmann, O., Bernard, S., Salehpour, M., Alkass, K., et al., 2014. Dynamics of oligodendrocyte generation and myelination in the human brain. Cell 159 (4), 766–774. € Yilmaz, O.H., Katajisto, P., Lamming, D.W., G€ ultekin, Y., Bauer-Rowe, K.E., Sengupta, S., et al., 2012. mTORC1 in the Paneth cell niche couples intestinal stem-cell function to calorie intake. Nature 486 (7404), 490–495. Young, K.M., Psachoulia, K., Tripathi, R.B., Dunn, S.-J., Cossell, L., Attwell, D., et al., 2013. Oligodendrocyte dynamics in the healthy adult CNS: evidence for myelin remodeling. Neuron 77 (5), 873–885. Zalc, B., Goujet, D., Colman, D., 2008. The origin of the myelination program in vertebrates. Curr. Biol. 18 (12), R511–R512. Zawadzka, M., Rivers, L.E., Fancy, S.P.J., Zhao, C., Tripathi, R., Jamen, F., et al., 2010. CNS-resident glial progenitor/stem cells produce Schwann cells as well as oligodendrocytes during repair of CNS demyelination. Cell Stem Cell 6 (6), 578–590. Zhang, Y., Zhang, H., Wang, L., Jiang, W., Xu, H., Xiao, L., et al., 2012. Quetiapine enhances oligodendrocyte regeneration and myelin repair after cuprizone-induced demyelination. Schizophr. Res. 138 (1), 8–17. Zhao, C., Fancy, S.P.J., Franklin, R.J.M., ffrench-Constant, C., 2009. Up-regulation of oligodendrocyte precursor cell alphaV integrin and its extracellular ligands during central nervous system remyelination. J. Neurosci. Res. 87 (15), 3447–3455. Zhu, X., Hill, R.A., Nishiyama, A., 2008. NG2 cells generate oligodendrocytes and gray matter astrocytes in the spinal cord. Neuron Glia Biol. 4 (1), 19–26.

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Progenitor cell-based treatment of glial disease

Steven A. Goldman1 Center for Neuroscience, University of Copenhagen Faculty of Health and Medical Sciences, Copenhagen, Denmark Center for Translational Neuromedicine, University of Rochester Medical Center, Rochester, NY, United States 1 Corresponding author: Tel.:+1-585-275-9550; Fax: +1-585-276-2298, e-mail addresses: [email protected]; [email protected]

Abstract Diseases of glia, including astrocytes and oligodendrocytes, are among the most prevalent and disabling, yet least appreciated, conditions in neurology. In recent years, it has become clear that besides the overtly glial disorders of oligodendrocyte loss and myelin failure, such as the leukodystrophies and inflammatory demyelinations, a number of neurodegenerative and psychiatric disorders may also be causally linked to glial dysfunction and derive from astrocytic as well as oligodendrocytic pathology. The relative contribution of glial dysfunction to many of these disorders may be so great as to allow their treatment by the delivery of allogeneic glial progenitor cells, the precursors to both astroglia and myelin-producing oligodendrocytes. Given the development of new methods for producing and isolating these cells from pluripotent stem cells, both the myelin disorders and appropriate glial-based neurodegenerative conditions may now be compelling targets for cell-based therapy. As such, glial cell-based therapies may offer potential benefit to a broader range of diseases than ever before contemplated, including disorders such as Huntington’s disease and the motor neuron degeneration of amyotrophic lateral sclerosis, which have traditionally been considered neuronal in nature.

Keywords Glial progenitor, Oligodendrocytic progenitor, Demyelinating disease, Leukodystrophy, Multiple sclerosis, Huntington’s disease, Cell transplant

1 INTRODUCTION Oligodendrocytes are the sole source of myelin in the adult CNS, and their loss or dysfunction is at the heart of a wide variety of diseases of both children and adults. In children, the hereditary leukodystrophies accompany cerebral palsy as major sources Progress in Brain Research, Volume 231, ISSN 0079-6123, http://dx.doi.org/10.1016/bs.pbr.2017.02.010 © 2017 Elsevier B.V. All rights reserved.

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of neurological morbidity. In adults, oligodendrocytic loss and demyelination contribute to diseases as diverse as multiple sclerosis, white matter stroke, and spinal cord injury (Roy et al., 2004). In addition, demyelination is also noted in degenerative disorders as varied as normal aging, Huntington’s and Alzheimer’s diseases, while oligodendrocytic pathology has been associated with disorders as diverse as amyotrophic lateral sclerosis (Lee et al., 2012) and schizophrenia (Tkachev et al., 2003). As a result, the myelin disorders are especially attractive targets for cell-based therapeutic strategies, as are the related disorders of astrocytes and glial progenitor cells (GPCs) that include dysmyelination as part of their presentation. Several recent studies have supported the readiness with which axons can remyelinate after either congenital or acquired demyelination, if provided myelinogenic cells (Duncan et al., 2009; Windrem et al., 2008). Human GPCs—also referred to as either oligodendrocyte progenitor cells (OPCs) or NG2 cells (Nishiyama et al., 2009)—can generate both oligodendrocytes and astrocytes, and are thus promising reagents by which to concurrently restore myelin to demyelinated regions of the diseased or injured CNS, while addressing the disorders of astrocytic function that so often attend white matter disease.

2 GPCs IN VIVO GPCs arise from neural stem cells (NSCs) of the ventricular subependyma and disperse widely throughout the central nervous system, pervading both gray and white matter (Roy et al., 1999). While the presence of GPCs in the adult human brain was inferred in several early studies that identified immature oligodendroglia in adult brain tissue (Armstrong et al., 1992; Gogate et al., 1994), mitotic bipotential GPCs were first isolated from the adult human brain using CNP2 promoter-based fluorescence-activated cell sorting (Roy et al., 1999). This study and others revealed that human glial progenitors comprise roughly 3% of all cells in the adult forebrain and are the major mitotic neural phenotype of the adult brain (Roy et al., 1999; Scolding et al., 1998). Subsequent work demonstrated that human GPCs generate both major macroglial phenotypes, astrocytes, and oligodendrocytes, and that they may do so in vivo as well, in a context-dependent fashion (Windrem et al., 2004). As the principal source of new oligodendrocytes, GPCs are thus responsible for remyelination in the demyelinated adult CNS (Tripathi et al., 2010; Zawadzka et al., 2010). As such, human GPCs are functionally synonymous with OPCs, to which they are also referred. Yet human GPCs appear to remain bipotential until their last division, and can give rise to both astrocytes and oligodendrocytes until that point. In vitro, they can manifest even broader lineage competence, as at least some fraction of adult human GPCs remain potentially neurogenic as well, in a density- and context-dependent fashion (Goldman, 2003; Nunes et al., 2003). Indeed, GPCs do not necessarily comprise a homogeneous pool; on the contrary, recent work by

3 Identifying optimal donor cell phenotypes for treating myelin disorders

Castelo-Branco and colleagues has revealed substantial molecular heterogeneity within the oligodendroglial lineage in rodents (Marques et al., 2016), as has been suggested in humans as well (Leong et al., 2014). Nonetheless, since there are no compelling data arguing for oligodendrocyte-restricted progenitors in humans, for consistency’s sake in this review, we will designate all of these glial progenitor phenotypes as GPCs, best identified empirically by the surface epitopes by which they have been isolated.

3 IDENTIFYING OPTIMAL DONOR CELL PHENOTYPES FOR TREATING MYELIN DISORDERS Disorders of myelin require extensive tissue repair, and in the case of the pediatric leukodystrophies, even whole neuraxis myelination. While endogenous glial progenitors can remyelinate demyelinated lesions to some degree, the mitotic exhaustion and functional depletion of endogenous glial progenitors that may occur in acquired demyelination ultimately limit the extent of spontaneous remyelination (Franklin and Ffrench-Constant, 2008), thus necessitating the introduction of exogenous glial progenitors as therapeutic vectors. Yet to be safe and effective as therapeutic vectors, transplantable GPCs must be reliably deliverable in both purity and quantity (Roy et al., 2004). The surface antigen-based purification of human GPCs, based on their selective expression of gangliosides recognized by monoclonal antibody A2B5 (Nunes et al., 2003; Roy et al., 1999), first allowed the isolation of both adult and fetal GPCs, which allowed their evaluation in animal models of both adult demyelination and congenital hypomyelination (Windrem et al., 2002, 2004). These studies revealed that while both fetal and adult human-derived GPCs were able to myelinate dysmyelinated brain tissue, adult GPCs did so more rapidly and efficiently, but manifested less expansion and migratory potential in vivo. In contrast, fetal GPCs emigrated more widely and engrafted more efficiently than did adult cells, and exhibited context-dependent differentiation as astrocytes or oligodendrocytes, and their tissue sources were more readily available, all of which suggested their potential therapeutic utility in a broad array of myelin disorders (Fig. 1). Moreover, the isolation of human tissue-derived GPCs allowed the assessment of their gene expression patterns, dominant signaling pathways and homeostatic self-renewal mechanisms (McClain et al., 2012; Sim et al., 2006, 2011), work that has enabled the ex vivo pretransplant modulation of the fate of these cells, further increasing their utility as transplantable vectors. Yet despite the attractiveness of fetal GPCs as therapeutic vectors, they remain finite in both initial number and expansion competence, necessitating their periodic reacquisition from new donor tissues. Fetal tissue suffers further limitations as a practicable cell source, given the unpredictability of tissue acquisition, limited gestational window of appropriate samples, and ongoing debate as to the use of fetal human-derived cells as clinical reagents. In light of these considerations, pluripotent

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Cell sources Human-induced pluripotent stem cells Patientderived skin cells or fibroblasts

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Hereditary leukodystrophies Congenital dysmyelinations Pelizaeus-Merzbacher disease Lysosomal storage diseases Tay-Sachs and Sandhoff’s gangliosidoses Krabbe’s disease Metachromatic leukodystrophy Mucopolysaccharidoses Niemann-Pick A Nonlysosomal diseases Adrenoleukodystrophy Canavan’s disease Vanishing white matter disease Cerebral palsy Periventricular leukomalacia Spastic diplegias of prematurity

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Autoimmune demyelination Multiple sclerosis Transverse myelitis Optic neuritis Vascular leukoencephalopathies Subcortical stroke Diabetic leukoencephalopathy Hypertensive leukoencephalopathy Spinal cord injury Inflammatory demyelination Radiation injury Neurodegenerative disorders Huntington disease Amyotrophic lateral sclerosis

FIG. 1 Glial progenitor cell sources, phenotypes, and clinical targets. Glial progenitor cells (GPCs) may be directly sorted from tissue, or produced from either human embryonic stem cells (hESCs) or human-induced pluripotential cells (hiPSCs), and then immunoselected based on their expression of either gangliosides recognized by monoclonal antibody (mAb) A2B5, or of

3 Identifying optimal donor cell phenotypes for treating myelin disorders

stem cells, including both human embryonic stem cells (hESCs) and induced pluripotent cells (iPSCs), have emerged in recent years as the more feasible sources of transplantable myelinogenic GPCs (Douvaras et al., 2014; Hu et al., 2009; Wang et al., 2013) (Fig. 2). Following the first report of myelination in the injured spinal cord by mouse ESCs (Brustle et al., 1999), oligodendrocytes derived from hESCs were similarly directed to generate myelin in vivo (Izrael et al., 2007; Nistor et al., 2005). Yet, while ground-breaking, these studies did not isolate GPCs or oligodendrocytes prior to transplant, nor did they follow animals over the time frames required to ensure the stability of the engrafted cells, of concern since any incidentally transplanted hESCs or undifferentiated derivatives may retain the potential for undesired expansion after implantation (Roy et al., 2006). Given this concern for tumorigenesis, stringent purification of lineage-restricted GPCs may be needed to ensure their safe use. Yet this point remains controversial; in 2009, the FDA approved a phase 1 safety trial evaluating the use of hESC-derived GPCs in spinal cord injury without such purification; although the trial was halted in 2011, its sponsor reported that its cessation was not related to safety, and described no serious adverse events attributable to the grafts (Priest et al., 2015). As a result, while the need for pretransplant isolation of terminally differentiated phenotypes remains unsettled, the overall safety profile of hESC-derived GPCs seems potentially acceptable. Human ESC-based therapy suffers also from the possibility of allograft rejection, and hence the need for immunosuppression in graft recipients. Enthusiasm has thus developed for the use of autologous grafts of myelinogenic GPCs derived from human-induced pluripotent cells (hiPSCs), potentially—though not assuredly (Zhao et al., 2011)—obviating the need for immune suppression. These cells are generated by the reprogramming of somatic cells to a pluripotent ground state, by the forced expression of a set of transcription factors that instruct stem cell phenotype (Belmonte et al., 2009). iPSCs were first generated from mouse (Takahashi and Yamanaka, 2006) and human (Yu et al., 2007) fibroblasts, and have since been differentiated into a variety of phenotypes, including neurons (Wernig et al., 2008),

CD140a/PDGFaR. The CD140a phenotype includes all potential oligodendrocytes, while the tetraspanin CD9 and sulfatide-directed mAb O4 identify progressively more oligodendrocytebiased fractions (Douvaras et al., 2014; Sim et al., 2011). In contrast, CD44 recognizes a more astrocyte-biased fraction (Liu et al., 2004). The choice of tissue-, hESC-, or iPSC-derived GPCs depends upon whether allogeneic or autologous grafts are desired. Whereas autologous grafts of iPSC-derived GPCs might obviate the need for immunosuppression, their generation may take months, and their use in the hereditary leukodystrophies would first require correction of the underlying genetic disorder in the donor cell pool; at present, such genetic disorders of myelin would be better approached with allografted tissue- or hESC-derived GPCs. Figure adapted from Osorio, M.J., Goldman, S.A., 2016. Glial progenitor cell-based treatment of the childhood leukodystrophies. Exp. Neurol. 283, 476–488.

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astrocytes (Hu et al., 2009), and oligodendrocytes (Czepiel et al., 2011). Methods established for generating GPCs from hESCs have proven effective with human iPSCs as well, and yield GPCs that are highly myelinogenic in vivo (Wang et al., 2013). This capability allows us to reasonably anticipate the use of iPSC-derived oligodendrocytes for autologous treatment, especially for nongenetic vascular, traumatic, and inflammatory demyelinations. Importantly though, iPSC-derived GPCs share many of the risks of those derived from hESCs, including aberrant differentiation and tumorigenesis (Mattis and Svendsen, 2011). In addition, iPSCs retain epigenetic marks of the cells from which they derive (Stadtfeld et al., 2010), so that their cell type of origin may influence their differentiation competence (Polo et al., 2010). Indeed, iPSCs may differ from one another in their lineage competence even when sourced from the same individual and tissue, making their standardization difficult. Of note, recent studies have reported the direct induction of glial progenitors and oligodendrocytes from fibroblasts as well (Najm et al., 2013; Yang et al., 2013). By thereby avoiding the need for pluripotent intermediates, such direct induction of GPCs may accelerate the production of transplantable cells, while mitigating their risk of tumorigenesis. While the lack of expandability of the largely postmitotic oligodendrocytes generated through this approach may still limit its practical utility, future advances may permit the direct induction of mitotic GPCs, which might yet prove a clinically feasible source of autologous myelinogenic cells (Goldman, 2013).

4 PEDIATRIC MYELIN DISORDERS AS TARGETS OF PROGENITOR CELL-BASED THERAPY Tens of thousands of children in the United States suffer from diseases of myelin loss. These include the metabolic demyelinations such as adrenoleukodystrophy; the lysosomal storage disorders, such as metachromatic leukodystrophy (MLD), the neuronal ceroid lipofuscinoses, gangliosidoses, and Niemann-Pick and Krabbe’s diseases; the hypomyelinating diseases, such as Pelizaeus-Merzbacher disease; the myelinoclastic disorders, including vanishing white matter disease, Alexander’s disease, and Canavan’s disease (Powers, 2004); and most common of all, periventricular leukomalacia and cerebral palsy (Silbereis et al., 2010). Their mechanistic heterogeneity notwithstanding, all of these conditions include the prominent loss of oligodendrocytes and myelin, highlighting their attractiveness as potential targets for cell replacement (see Fig. 1).

4.1 METABOLIC AND STORAGE DISORDERS OF MYELIN In some hereditary disorders of the white matter, such as Canavan’s disease and many of the lysosomal storage diseases, oligodendrocytes are principal targets of misaccumulated toxic substrates and their abnormal metabolic products, with

4 Pediatric myelin disorders as targets of progenitor cell-based therapy

demyelination an early and inauspicious hallmark of disease. In others, such as Krabbe’s disease, oligodendrocytes are essentially bystanders, killed by toxic metabolites generated by cells deficient in one or more critical enzymes (Powers, 2004). In yet others, such as Alexander’s disease and vanishing white matter disease, myelin loss may be caused by astroglial pathology (Bugiani et al., 2011; Dietrich et al., 2005). Given their heterogeneous etiologies, a common treatment platform for these disorders has proven elusive. Yet since GPC engraftment is both widespread and associated with astrocytic as well as oligodendrocytic production, GPCs would seem an especially promising vehicle for dispersing astrocytes and oligodendrocytes throughout otherwise diseased and/or enzyme-deficient brain parenchyma. The lysosomal storage disorders present especially attractive targets in this regard since wild-type lysosomal enzymes may be released by donor cells, and taken up by deficient host cells through the mannose-6-phosphate receptor pathway (Urayama et al., 2004), by which lysosomal enzymes released from wild-type donor cells may be transported to enzyme-deficient neighbors, permitting local correction of disease-specific metabolic disturbances. By this means, a relatively small number of donor glia may provide sufficient enzymatic activity to correct the underlying enzymatic deficit and storage disorder of a much larger number of host cells (Jeyakumar et al., 2005). The cell-based rescue of enzymatically deficient host cells was first described using wild-type NSCs transplanted into a mouse model of mucopolysaccharidosis type VII, in which neonatally implanted cells restored beta-glucuronidase enzymatic function in the recipient forebrain (Buchet et al., 2002; Meng et al., 2003; Snyder et al., 1995). Human NSCs proved similarly effective in achieving enzyme replacement in the b-hexosaminidase-deficient Sandhoff mouse, with corresponding functional benefits (Lee et al., 2007). In the same vein, NSCs engineered to overexpress sphingomyelinase, engrafted into sphingomyelinasedeficient Niemann-Pick type A mice, and yielded substantial reductions in misaccumulated sphingomyelin (Shihabuddin et al., 2004). Similarly, when NSCs were engrafted into a mouse model of neuronal ceroid lipofuscinosis (NCL), the cells dispersed broadly and ameliorated the lipofuscin misaccumulation of these animals (Tamaki et al., 2009). On that basis, a clinical trial to assess the use of human NSC allografts in treating infantile and late infantile NCL was undertaken (Selden et al., 2013). This phase 1 safety trial did not address therapeutic endpoints, but its initiation speaks to the efforts that may be anticipated in developing neural stem and GPCs as vehicles for intracerebral enzyme replacement in the metabolic leukodystrophies. The intracerebral delivery of GPCs would thus seem an especially promising approach for treating those enzyme deficiencies associated with early demyelination, which may require both enzyme replacement and structural remyelination. MLD, for example, is characterized by deficient expression of arylsulfatase A, which results in sulfatide misaccumulation and oligodendrocyte loss. Experimental models of MLD have responded well to GPC grafts, with broad dispersal and integration as well as enzymatic rescue and sulfatide clearance (Givogri et al., 2006).

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Krabbe’s disease, characterized by galactocerebrosidase deficiency and early demyelination, may prove similarly amenable to GPC-based enzymatic repletion and myelin restoration (Kondo and Duncan, 2016). When children with presymptomatic or early stage Krabbe’s disease were transplanted with umbilical cord stem cells, they manifested slower disease (Escolar et al., 2005). Yet the intracerebral infiltration of umbilical cord stromal derivatives is modest, suggesting that treatment of these children with GPCs, which are able to achieve parenchymal dispersal and infiltration as well as structural remyelination, might comprise an especially promising treatment strategy.

4.2 DISORDERS OF MYELIN FORMATION AND MAINTENANCE The experimental assessment of GPCs as vectors for remyelination has proceeded most aggressively in animal models of congenital hypomyelination. In an early study of cell-based myelin repair, mouse NSCs were transplanted into newborn shiverer mice, a hypomyelinated mutant deficient in myelin basic protein, and yielded context-dependent myelination (Yandava et al., 1999). Even earlier studies had shown that tissue grafts of fetal white matter could elaborate oligodendrocytes and myelinate the adult shiverer spinal cord, though on a more geographically limited basis (Gout et al., 1988; Gumpel et al., 1989). On the basis of these foundational studies, we transplanted immunosorted human GPCs into neonatal shiverers, so as to assess the relative myelinogenic potential of purified populations of human glial progenitors (Windrem et al., 2004). When delivered as highly enriched isolates, fetal human GPCs spread widely throughout the brain, developing as astrocytes and oligodendrocytes in a context-dependent fashion. The donorderived oligodendrocytes generated ultrastructurally mature myelin that effectively ensheathed host shiverer axons and formed nodes of Ranvier, which allowed the restoration of normal transcallosal conduction velocities in the transplanted mice (Windrem et al., 2004). By using a five-site injection protocol to achieve broader dispersal of GPCs, we next established cell engraftment throughout the entire neuraxis, with myelination of the spinal cord and roots as well as the entire brain, brainstem, cerebellum, and cranial nerve roots (Windrem et al., 2008). This was associated with substantially prolonged survivals in transplanted mice, with phenotypic recovery and frank rescue of a large minority. These data strongly suggested the feasibility of neonatal GPC implantation in treating childhood disorders of myelin formation and maintenance. Later studies refined the criteria for selecting myelinogenic progenitors, by identifying the PDGFa receptor epitope CD140a as recognizing the entire population of oligodendrocyte-competent progenitors (Sim et al., 2011). CD140a-sorted GPCs proved superior to those selected on the basis of A2B5 in both their efficiency and extent of myelination, and were highly migratory; they have thus supplanted A2B5-defined cells as a preferred cellular vector for therapeutic remyelination.

4 Pediatric myelin disorders as targets of progenitor cell-based therapy

5 days ESC medium w/o FGF2

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NF MBP FIG. 2 Stem cell-derived glial progenitor cell-mediated myelination of a dysmyelinated host. (A) This schematic outlines the multistage protocol by which GPCs, oligodendrocytes, and astrocytes may be generated from pluripotent cells. (B–F) Representative images taken at serial stages of glial differentiation, with the serial expression of selected marker proteins noted at each stage. (G) Three months after neonatal transplant into hypomyelinated shiverer mice, human-induced pluripotent cell (hiPSC)-derived GPCs have matured as myelinating, myelin basic protein (MBP)-expressing oligodendrocytes (MBP, green; human nuclear antigen, red). (H) The hiPSC-derived oligodendrocytes ensheath mouse axons (neurofilament, red; MBP, green). (I) Human iPSC-derived oligodendrocytes can myelinate (Continued)

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4.3 THE DILEMMA OF DISEASE-SPECIFIC DOSING The intracerebral delivery of GPCs may thus prove a viable approach to the treatment of a wide variety of enzymatic and storage diseases of myelin, as well as for the disorders of myelin misproduction or failure. In practice though, individual treatment regimens will need to be tailored to specific disease phenotypes and stages. Few data are available as to the numbers or proportion of wild-type cells required to achieve correction of enzymatic activity and substrate clearance in any storage disorder, and these values may need to be empirically derived for every disease target. Similarly, the extent of myelination required for effective treatment remains wholly speculative, as is the extent and duration of immunosuppression required for allograft acceptance; these parameters too may vary with disease phenotype. These caveats notwithstanding, NSC implantation has already been assessed as a means of myelin replacement in Pelizaeus-Merzbacher disease, in a phase 1 study that reported safety albeit unclear efficacy with the dose and delivery method chosen (Gupta et al., 2012); one may anticipate that future efforts will proceed to assess the efficacy of GPC grafts in this and related disorders. As these efforts proceed, with more dedicated myelinogenic donor cell phenotypes, more refined delivery methods and increasingly informed dose estimations, one may similarly expect that both the relative efficacy and limitations of GPC delivery in alleviating the childhood myelin disorders will be definitively established.

5 ADULT DISEASE TARGETS OF GPC-BASED TREATMENT In adults, oligodendrocytic loss contributes to diseases as diverse as hypertensive and diabetic white matter loss, traumatic spinal cord and brain injury, and multiple sclerosis (MS) and its variants. In addition, oligodendrocytic loss is prominent in the degenerative dementia associated with age-related white matter loss. All of these are potential targets of GPC replacement therapy, although the adult disease environment may limit this approach in ways not encountered in pediatric disease targets. For instance, the chronically ischemic brain tissue of diabetics with small vessel disease may require aggressive treatment of the underlying vascular insufficiency before any cell FIG. 2—Cont’d the entire brain of shiverer mice, which do not otherwise express MBP (green). (J) The myelin generated by hiPSC oligodendrocytes is ultrastructurally normal, exhibiting major dense lines and thick myelin sheaths. The use of such serial and distinct stages of growth factor exposure, paired with more extended periods of differentiation, have led to the production of highly enriched populations of human GPCs, that are highly efficient at myelinogenesis in vivo while manifesting no evident tumorigenesis. Scale: (B–E), 100 mm; (F), 25 mm; (G), 100 mm; (H), 10 mm; (J), 100 nm. Images taken from Wang, S., Bates, J., Li, X., Schanz, S., Chandler-Militello, D., Levine, C., Maherali, N., Studer, L., Hochedlinger, K., Windrem, M., et al., 2013. Human iPSC-derived oligodendrocyte progenitor cells can myelinate and rescue a mouse model of congenital hypomyelination. Cell Stem Cell 12, 252–264; figure adapted from Goldman, S.A., Kuypers, N.J., 2015. How to make an oligodendrocyte. Development 142, 3983–3995.

5 Adult disease targets of GPC-based treatment

replacement strategy may be considered. Similarly, the inflammatory disease environments of MS as well as many of the leukodystrophies present their own challenges, which need to be overcome before cell-based remyelination can succeed (Franklin and Ffrench-Constant, 2008; Ip et al., 2006). Nonetheless, current disease-modifying strategies for treating both vascular and autoimmune diseases have advanced to the point where transplant-based remyelination of adult targets may now be feasible.

5.1 PROGENITOR CELL THERAPY FOR MULTIPLE SCLEROSIS Interest in cell-based remyelination has focused on MS, a debilitating disease characterized by both inflammatory myelinolysis and degenerative axonal loss. The attraction of MS as a therapeutic target derives from its high incidence and prevalence, with more than 300,000 cases in the United States alone. MS has been a difficult target for cell therapy, given its relapsing course and the limitations of introducing new cells into an inflammatory environment. Nonetheless, contemporary immune modulating treatments have substantially diminished disease recurrence, making cell replacement a tenable repair strategy. Natalizumab (anti-a4 integrin), alemtuzumab (anti-CD52), rituximab (anti-CD20), fingolimod (a sphingosine-1-phosphate receptor modulator), dimethylfumarate, and terflunamide (a pyrimidine synthesis inhibitor), have all been associated with significant reductions in relapse rate (Weinstock-Guttman and Ramanathan, 2012), reflecting diminished episodes of radiographically and clinically significant central inflammation. These advances in the immunomodulatory control of MS suggest that attention may now shift from disease attenuation to the repair of demyelinated lesions, and to the prevention of the later progressive neurodegenerative phase that occurs in MS, designated secondary progressive MS. The latter condition, characterized by sustained axonal loss and an attendant, inexorable loss of function, long after acute inflammatory events have attenuated, may reflect a loss of axonal support by local oligodendroglia (Lee et al., 2012; Saab et al., 2016). As such, progressive MS might comprise an especially attractive target for glial replacement, since oligodendrocyte engagement with otherwise denuded axons might serve to preserve neuronal viability even in the absence of remyelination. The intracerebral delivery of GPCs into demyelinated brain may thus offer tangible benefits by oligodendroglial replacement and axonal engagement per se, as well as by myelin repair.

5.2 PROGENITOR CELL THERAPY FOR ADULT STRUCTURAL DEMYELINATIONS Besides the autoimmune demyelinations, the adult CNS is subject to demyelination from a broad variety of traumatic, vascular, and metabolic insults. Traumatic brain injury, white matter stroke, age-related white matter disease, and the leukomalacias associated with hypertension and diabetes are all potential targets of glial replacement for the purpose of white matter repair, and yet each of these potential disease targets needs to be separately modeled, given the vastly different disease

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environments and hence host constraints of each. Yet, to the extent that the pathology of these disorders is limited to oligodendrocytes and astrocytes, with preserved vascular function and neuronal architecture, they too may be amenable to GPC-based remyelination, as might also be iatrogenic causes of demyelination such as radiation therapy (Fox et al., 2014; Piao et al., 2015). To be sure, the complexity of the disease environment may make adult targets less approachable than their pediatric counterparts, especially so in aged subjects (Franklin and Ffrench-Constant, 2008; Ruckh et al., 2012). Moreover, previously stably myelinated but now-demyelinated axons may have very different thresholds and permissiveness toward remyelination than their never-before myelinated, hypomyelinated, or briefly myelinated pediatric counterparts. Indeed, recent studies have emphasized that improved understanding of the molecular environment of demyelinating foci, and the transcriptional response of OPCs to that environment, may offer significant insight into improving the efficiency and timing of GPC transplantation (Moyon et al., 2015). Such studies of the disease-specific tissue environment in demyelinating disorders are especially important given the relatively smaller data base regarding glial progenitor grafts into the adult CNS; whereas, the dispersal and ultimate myelination of glial progenitors introduced into neonatal and young animals is now well established, and forms the basis of the use of this strategy in the childhood myelin disorders as noted, the experimental basis for remyelination of adult-demyelinated brain is less robust. To be sure, when human GPCs were transplanted directly into lysolecithin-demyelinated lesions in the adult rat brain, the cells matured as oligodendrocytes and myelinated residual host axons, if with lower efficiency than in congenitally hypomyelinated brain (Windrem et al., 2002), and more recent studies have supported the broad migration of human GPCs introduced into the normal adult rat brain (Osorio and Goldman, 2016). Similarly, in a novel model of axon-sparing demyelination in adult cats, remyelination occurred efficiently from endogenous progenitors (Duncan et al., 2009). More recently, remyelination of demyelinated adult spinal cord lesions by human iPSCderived GPCs has been noted by van Evercoorn and colleagues (Mozafari et al., 2015), work that establishes the permissiveness of adult-demyelinated axons to remyelination by pluripotent cell-derived GPCs. Together, these and other studies argue that remyelination of adult-demyelinated axons is clearly feasible. Nonetheless, it is equally clear that any cell-based strategies for treating adult demyelination will require not only disease modification, but rigorous stratification to define those patients with a tissue environment permissive for donor cell integration, and sufficient axonal preservation to benefit from this approach.

5.3 REMYELINATION OF SPINAL LESIONS Besides the myelinated tracts of the brain, the ascending sensory and descending motor tracts of the spinal cord are frequent victims of demyelination, whether from MS, neuromyelitis optica, or segmental injuries. In efforts to remyelinate

6 Human glial chimeric mice reveal human-selective aspects

the contused rat spinal cord, implanted GPCs have been found to disperse and generate both astrocytes and myelinogenic oligodendrocytes (Han et al., 2004). Similarly, hESC-derived oligodendrocytes can remyelinate demyelinated cord lesions (Nistor et al., 2005), with functional benefit in experimental models (Sharp et al., 2010). On the basis of those observations, a first-in-man safety trial was performed using hESC-derived GPCs transplanted into patients with high-grade thoracic cord lesions (Priest et al., 2015). No serious adverse events or donor-derived tumorigenesis was noted in this trial, which was thus reassuring as to the potential of hESC GPC-based neurological therapeutics. That said, the therapeutic potential of a solely remyelinative strategy in patients with such high-grade spinal lesions is unclear, given the concurrent loss of segmental neurons and long tract axons in such lesions, and the compromised vascular and inflammatory environment of the injured spinal cord. Nonetheless, such GPC grafts may hold great promise in carefully selected patients with isolated segmental demyelination. As such, the previously noted recent demonstration of focal remyelination of the injured spinal cord by iPSC-derived GPCs (Mozafari et al., 2015), not only provides generic support for the use of these cells as therapeutic vectors but also offers insight into the parameters that define the migration and myelination competence of human GPCs in the specific environment of the spinal cord, and which may thereby be modulated to ensure graft success.

6 HUMAN GLIAL CHIMERIC MICE REVEAL HUMAN-SELECTIVE ASPECTS OF BOTH GLIAL FUNCTION AND DYSFUNCTION When hypomyelinated mutant mice are engrafted neonatally with human GPCs, the donor cells mature as both myelinating oligodendrocytes and fibrous astrocytes, ultimately yielding mice with a substantially humanized white matter (Windrem et al., 2008). Large numbers of human donor cells also remain as progenitors, which over time predominate, displacing and ultimately replacing the endogenous mouse glial progenitor pool. This competitive advantage of human over murine glial progenitors is evident in wild-type as well as in hypomyelinated mice, such that in the setting of normal glial turnover the human GPCs also give rise to gray matter astrocytes, eventually resulting in substantial astrocytic as well as oligodendrocytic humanization of the recipient rodent brains. The result of these events is that the xenografted mouse brains can become substantially humanized in their glial constituents (Windrem et al., 2008, 2014). These human glial chimeras lend themselves to the investigation of questions never-before approachable, for lack of an appropriate in vivo model of human glial function. In particular, these mice permit assessment of the species-specific contributions of human glia to neural network function. Astrocytes clearly play

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a central role in synaptic efficiency and plasticity in mammals (Kang et al., 1998). Hominid evolution in particular has been attended by increasing astrocytic complexity, which may have contributed greatly to the evolution of higher cognitive functions in primates. Human astrocytes are larger and far more fibrous than those of infraprimate mammals, and include vastly more encompassed synapses within their individual geographic domains (Oberheim et al., 2006, 2009). Accordingly, when neonatal mice were engrafted with astrocyte-biased human GPCs, resulting in the colonization of the host brains with human glia, the resultant glial chimeras exhibited substantially enhanced activity-dependent plasticity and learning (Han et al., 2013). This finding strongly suggested that human astrocytes might play a much larger role in neural processing than do those of rodents. Yet by doing so, this observation also highlighted the potential for glial pathology to wreak especial havoc within human neural circuits, with attendant implications for the human neurodegenerative disorders. As such, this study suggested that human-specific astrocytic specializations might contribute not only to human cognitive evolution but also to the appearance of human-selective neurodegenerative and psychiatric disorders.

7 GLIAL TRANSPLANT-MEDIATED AMELIORATION OF NEURODEGENERATIVE DISORDERS Among the human-selective disorders that might reflect glial human evolution are a number of prototypic neurodegenerative and neuropsychiatric disorders. Glial pathology has already been noted to contribute to a broad set of neurodegenerative and neuropsychiatric diseases traditionally considered disorders of solely neuronal dysfunction (Di Giorgio et al., 2008; Verkhratsky and Parpura, 2015). Amyotrophic lateral sclerosis in particular was first identified as having a substantial glial component to its etiology (Di Giorgio et al., 2007, 2008; Meyer et al., 2014; Yamanaka et al., 2008), with a failure in astrocytic glutamate transport and disrupted potassium homeostasis, as well as defective oligodendroglial metabolite transfer (Lee et al., 2012), each potentially contributing to the loss of motor neurons and other large pyramidal neurons in this disorder. Indeed, while a number of genetic bases for ALS have been identified, glial dysfunction may be a shared trait that confers selective neuronal vulnerability across the spectrum of ALS-associated motor neuronopathies. For instance, TDP43 proteinopathy-associated ALS has been shown to exhibit primary astrocytic pathology with secondary motor neuron dysfunction (Serio et al., 2013), highlighting the noncell autonomous nature of neuronal loss in even this quintessentially neuronal degeneration. Huntington’s disease (HD) is another such nominally neurodegenerative disorder, to which glial pathology also appears to make a significant causal

7 Glial transplant-mediated amelioration of neurodegenerative disorders

contribution. HD is an autosomal dominant disorder characterized by abnormally long CAG repeat expansions in the first exon of the Huntingtin gene. The encoded polyglutamine expansions of mutant huntingtin protein disrupt its normal functionsand protein–protein interactions, ultimately yielding widespread neuropathology, most rapidly evident in the neostriatum (Waldvogel et al., 2015). Yet despite the pronounced loss of striatal medium spiny neurons in HD, and evidence of glial dysfunction (Shin et al., 2005; Tong et al., 2014), few studies have investigated the specific contribution of glial pathology to the HD phenotype. This lack of understanding of the role of glia in HD—reflecting our broader ignorance of the role of glial dysfunction in the neurodegenerative disorders (Verkhratsky et al., 2014)—has highlighted the lack of in vivo models that permit the separate interrogation of glial and neuronal functions in HD, particularly so in humans. To address this fundamental gap in our knowledge, we used human glial chimeric mice to assess the role of human striatal glia in the pathogenesis of HD, by comparing the behavior and MSN physiology of mice xenografted at birth with mutant HD-expressing human hGPCs to their normal HTT hGPC-engrafted controls (Benraiss et al., 2016). In this study, the motor behavior of immunodeficient mice neonatally xenografted with hGPCs produced from mutant HD hESCs (48 CAG) was compared to that of controls engrafted with hGPCs derived from a sibling line of unaffected hESCs (18 CAG). The HD GPC-engrafted mice manifested impaired motor learning relative to control hGPC-engrafted mice, and exhibited increased neuronal input resistance and excitability, relative to those of mice engrafted with normal HTT (23 CAG)-transduced striatal glia (Fig. 3A–C). Since striatal chimerization with HD-derived glia proved sufficient to recapitulate aspects of disease phenotype in normal mice, one might anticipate neonatal chimerization of HD mice with normal hESC-derived glia might delay or rescue aspects of disease phenotype. Using R6/2 transgenic HD mice as hosts (Mangiarini et al., 1996), Benraiss and colleagues found that the substantial replacement of diseased striatal glia with wild-type human glia indeed resulted in a slowing of disease progression and corresponding increment in survival in transplanted R6/2 mice (Fig. 3D–I; Benraiss et al., 2016). Of note, the glial donor cells were isolated on the basis of CD44, the hyaluronan receptor, so as to capture a more astrocyte-biased donor cell population. This neonatal engraftment by CD44-defined glia yielded a transplantassociated fall in neuronal input resistance, and a corresponding drop in interstitial K+ in the R6/2 striatum, with an attendant rescue of the otherwise hyperexcitable phenotype of R6/2 striatal neurons (Fig. 3D and E; Benraiss et al., 2016). Together, these studies indicated a critical role for glial pathology in the progression of HD. As such, they suggest the potential for glial cell replacement as a therapeutic strategy in HD, and more broadly, to other neurodegenerative diseases in which glial pathology might be causally contributory.

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FIG. 3 Glial progenitor grafts slow disease progression, stabilize neuronal excitability, and extend survival in mouse models of Huntington’s disease. (A) Schematic outlines the preparation and delivery of normal CD44-defined astroglial progenitor cells into the striata of neonatal HTT mutant R6/2 (120Q)  rag1 / mice. (B) CD44-sorted hGPCs colonized the R6/2  rag1 / striatum. Striatal engraftment of the R6/2 mice by CD44-sorted hGPCs was robust and dense. Donor-derived cells increased as a function of time, such that transplanted hGPCs expanded to largely replace host glia in the striata and ventral forebrains of recipient R6/2 mice by 20 weeks. Scale: 1 mm. (C) By 20 weeks after neonatal graft, the donor

8 Human glial involvement in the neuropsychiatric disorders

8 HUMAN GLIAL INVOLVEMENT IN—AND POTENTIAL RESCUE OF—THE NEUROPSYCHIATRIC DISORDERS The therapeutic benefit in animal models of glial replacement in HD raises the possibility that glial cell replacement may prove of therapeutic value not only for diseases of oligodendrocytes and myelin but also for primarily astrocytic disorders as well. HD is not alone in manifesting with relative hypomyelination and concurrent defects in cognition typically ascribed as cortical in nature. A broad variety of neurodegenerative and neuropsychiatric disorders also share concurrent deficits in central white matter structure and cortical function that might suggest glial pathology. Indeed, astroglial pathology in particular might be especially germane to those neural

hGPCs (human nuclear antigen, red) integrated as astrocytes (GFAP, green) or persisted as GPCs (PDGFaR and olig2, green), but did not give rise to neurons; no overlap was seen of hN and neuronal NeuN. Scale: 25 mm. (D) Chimerization with normal glia partially normalized MSN physiological function. Whole-cell I-clamp recordings from rag1 / wild-type, CD44 hGPC-engrafted rag1 / wild types, R6/2  rag1 / mice, and CD44-engrafted rag1 / mice revealed that the input resistance Rinput, was significantly higher in R6/2  rag1 / striatal neurons than in wild-type  rag1 / controls, but was partially restored to normal in R6/2 mice chimerized with normal CD44-sorted hGPCs. (E) Normal glial engraftment reduces interstitial K+ levels in the R6/2 striatum. Potassium electrodes were used to measure the interstitial levels of striatal K+ in both wild-type mice and R6/2 littermates at 16 weeks of age (4 days), with and without neonatal intrastriatal transplants of CD44-sorted hGPCs. Untreated R6/2 mice manifested significantly higher levels of interstitial K, which were restored to normal in R6/2 mice neonatally engrafted with hGPCs. In contrast, hGPC engraftment did not influence the interstitial K+ levels of wild-type mice. (F) Striatal involution of R6/2 mice was slowed by normal glial engraftment. The neostriata of R6/2 HD mice typically shrink in volume with age. Human GPC engraftment attenuated this process, in that R6/2 mice manifested larger striatal volumes than unengrafted R6/2 mice by 16 weeks of age, with preservation of R6/2 striatal volumes at levels no different than wild-type controls. (E, F) Means  SEM; ** and ***p < 0.01 and 0.0001, one-way ANOVA. (G) Chimerization with normal glia slows motor loss and extends survival of R6/2 mice. Linear regression revealed that the rate of rotarod-assessed motor deterioration of R6/2 mice was significantly slower in mice engrafted with human GPCs than in untreated mice (p < 0.001 by ANOVA). (H) Treatment with hGPCs slowed cognitive decline in R6/2 mice. SmartCube testing (Psychogenics), a multimodal assessment of cognitive function (Benraiss et al., 2016), revealed a significant difference between sham-treated wild-type (WT) and R6/2 mice by 8 weeks of age, with significant functional preservation in the R6/2 mice when treated neonatally with normal hGPCs. In (H), each dot represents a mouse. The center, small, and large ellipses represent the mean, standard error, and standard deviation, respectively, of the composite features for each group. See Benraiss et al. (2016) for detail. (I) R6/2 mice whose striata were engrafted with human GPCs survived significantly longer than unengrafted mice (p < 0.01, Mantel-Cox log-rank test).

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diseases that are fundamentally unique to humans, such as schizophrenia (Tkachev et al., 2003), whose phylogenetic appearance may parallel that of human astrocytic evolution. A number of authors have pointed to the causal contribution of both astroglial and oligodendroglial pathology, and white matter abnormalities more broadly, in schizophrenia (Davis et al., 2003; Haroutunian et al., 2014; Hof et al., 2002; Nave and Ehrenreich, 2014; Takahashi et al., 2011; Verkhratsky and Parpura, 2015; Voineskos et al., 2013; Wang et al., 2015). The previously noted observation that astroglial replacement can attenuate the behavioral phenotype of HD transgenic mice suggests the potential for such glial replacement in cognitive disorders with substantial glial pathology, especially so in those disorders such as schizophrenia that may suffer both astrocytic and oligodendrocytic dysfunction. Whether healthy donor glial progenitors can replace diseased host glia in adolescents and adults with such conditions remains unclear, and will likely need to be investigated on a disease-bydisease basis. Moreover, should this strategy be clinically feasible, the ethical issues involved in potentially ameliorating a behavioral condition, and hence potentially influencing an adult personality, by means of an intracerebral graft of allogeneic stem cell-derived glia—to wit, those of another person—may prove significant (Hermeren, 2015). Nonetheless, the very possibility of improving cognition and ameliorating both neurodevelopmental and psychiatric pathology by means of glial transplantation offers both breath-taking new opportunities for cell-based therapeutics of the CNS.

9 CONCLUSIONS Overall then, a stunning variety of recent studies have converged to highlight the disorders of glia as especially promising initial targets for cell-based therapy of neurological disease. Using a common strategy of GPC implantation, a broadly inclusive set of both pediatric and adult disorders of the central nervous system may now prove amenable to cell-based repair. These disorders include not only the myelin disorders for which glial cell replacement was first intended, but also the entire spectrum of disorders involving glial pathology, and especially so those involving white matter disease including both astrocytic and oligodendrocytic elements. This broader range of disorders includes not only the structural and metabolic dysmyelinations, but also the entire spectrum of neurodevelopmental and neuropsychiatric diseases to which astrocytic and oligodendrocytic pathology might causally contribute.

ACKNOWLEDGMENTS Work discussed in the Goldman lab was supported by NIMH, NINDS, and grants from CHDI Foundation, Adelson Medical Research Foundation, Mathers Charitable Foundation, Novo Nordisk Foundation, Lundbeck Foundation, National Multiple Sclerosis Society, and the New York State Stem Cell Research Program (NYSTEM).

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Pluripotent stem cells and their utility in treating photoreceptor degenerations

8

Nozie D. Aghaizu1, Kamil Kruczek, Anai Gonzalez-Cordero, Robin R. Ali, Rachael A. Pearson1 UCL Institute of Ophthalmology, London, United Kingdom Corresponding authors: Tel.: +44-207-608-6981; Fax: +44-207-608-6991 (N.D.A.); Tel.: +44-207-608-4022; Fax: +44-207-608-6991 (R.A.P.), e-mail address: [email protected]; [email protected]

1

Abstract Age-related macular degeneration and inherited retinal degenerations represent the leading causes of blindness in industrialized countries. Despite different initiating causes, they share a common final pathophysiology, the loss of the light sensitive photoreceptors. Replacement by transplantation may offer a potential treatment strategy for both patient populations. The last decade has seen remarkable progress in our ability to generate retinal cell types, including photoreceptors, from a variety of murine and human pluripotent stem cell sources. Driven in large part by the requirement for renewable cell sources, stem cells have emerged not only as a promising source of replacement photoreceptors but also to provide in vitro systems with which to study retinal development and disease processes and to test therapeutic agents.

Keywords Photoreceptor cells, Retina, Blindness, Retinal dystrophies, Cones, Stem cells, Transplantation, Embryonic stem cells, Induced pluripotential stem cells

1 INTRODUCTION The processing and computation of sensory visual information are an essential element of how many animals interact with their environment. The eye is a highly specialized sensory organ equipped to detect visual stimuli coming from the environment and to convert them into electrical signals. These are then relayed to higher visual centers in the brain part of the central nervous system (CNS). Clich
ed connections between the eye and the brain (soul) have been in use long before modern research discovered an anatomical and developmental link between

Progress in Brain Research, Volume 231, ISSN 0079-6123, http://dx.doi.org/10.1016/bs.pbr.2017.01.001 © 2017 Elsevier B.V. All rights reserved.

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the retina and the brain (reviewed in London et al., 2012). The neural retina is part of the CNS and in vertebrates it can be regarded as an extension of the brain. In fact, both originate from the same germ layer, the neuroectoderm. During early development, mitotic cells of the rostral neural plate are specified to form the eye field (see review by Chow and Lang, 2001), which eventually gives rise to the neural retina and its resident photoreceptors—the principal light sensing cells contributing to image forming vision. The eye’s accessibility and its relatively simple neuronal circuit architecture have made the retina a popular model tissue to study both CNS development and translational approaches. Photons are detected by the outer segments of the photoreceptors located within the outer nuclear layer (ONL). This initiates the phototransduction signaling cascade, which converts light energy into an electrical stimulus. This stimulus is passed on by synaptic transmission, first to the secondary interneurons of the inner nuclear layer (INL) and then to the retinal ganglion cells (RGCs) in the ganglion cell layer (GCL), the axons of which fasciculate to form the optic nerve, along which the visual information is relayed to higher visual centers. Approximately 30% of all sensory input in the human brain is visual in nature (Dowling, 1987; Rodieck, 1998) and the loss of vision due to injury and/or disease in the retina, although rarely life-threatening, represents a severely debilitating condition. Robust regeneration and repair mechanisms only exist within lower vertebrate species. By contrast, the mammalian retina, like other parts of the mammalian CNS, is generally ill-equipped to perform regeneration and self-repair. Thus, once retinal neurons, especially the photoreceptors and their support cells, are profoundly damaged or lost due to injury/disease, permanent vision impairment typically follows. This has motivated researchers to develop novel therapeutic strategies with varying applicability, depending on the nature of the underlying retinal condition. These include, among others, cell transplantation, gene therapy, immunotherapy, and retinal prosthetics. Pluripotent stem cell (PSC) research represents a promising new approach in the field of retinal therapy, providing a renewable source of cells for transplantation, as well as a source of cells for in vitro disease modeling and drug screening/testing strategies. In recent years, efforts by several research groups have demonstrated that PSCs can be differentiated into retinal cells, including photoreceptors, by recapitulating in vivo developmental retinogenesis programs. Retinal cells generated in this way have become increasingly more like their in vivo counterparts in terms of differentiation/maturation state, morphology, and function. In this review, we aim to provide an overview of the potential utility of PSCs for treating visual impairment due to retinal degeneration, focusing on PSC-derived photoreceptors.

2 RETINAL DEGENERATION Retinal degeneration describes a cohort of debilitating conditions characterized by the progressive loss of photoreceptors and neuronal remodeling. Broadly, these conditions can be classified into three groups: rod-degenerative forms, mixed

2 Retinal degeneration

rod/cone-degenerative forms, or debris-associated forms, e.g., mertk defects and light damage (Jones and Marc, 2005). Various environmental factors, traumatic injury, as well as inherited genetic disorders, can lead to retinal degeneration. In industrialized countries, age-related macular degeneration (AMD) is now the leading cause of untreatable blindness in people of 50 + years (National Eye Institute, 2015). In addition to an age-related disease etiology, macular degeneration also occurs as a juvenile onset inherited form, Stargardt disease. The macula at the center of the human retina includes the cone-rich fovea that is essential for high acuity vision; photoreceptor degeneration in the macula, therefore, results in impaired central vision and loss of visual acuity. AMD is a multifactorial disease with both environmental and genetic etiologies. The major risk factors are advanced age, smoking, and genetic predispositions related to the alternative complement pathway (Age-related Eye Disease Study Research Group, 2000; Haines et al., 2005; Smith et al., 2001). AMD may manifest as either dry (geographic) or wet (neovascular). Dry AMD starts with chronic, low-grade inflammation in the macular neural retina, which eventually causes the RPE underlying the macula and the adjacent membrane, the Bruch’s membrane, to degenerate. This subsequently results in photoreceptor degeneration within the macula (Curcio et al., 1996; Jager et al., 2015). Patients with wet AMD additionally present with choroidal neovascularization, which penetrates through Bruch’s membrane and RPE into the subretinal space and the neural retina. The newly formed blood vessels are relatively fragile and prone to become permeabilised, which may lead to the accumulation of subretinal fluid/hemorrhages, retinal detachment, and photoreceptor death (Jager et al., 2015). Many retinal diseases are directly due to a single underlying inherited genetic defect. There are presently over 200 genes that have been linked to inherited retinal degeneration diseases, many of which are specifically expressed in photoreceptors or in the closely associated RPE cells (Daiger et al., 2013; Hartong et al., 2006). The encompassing term for those diseases characterized by initial loss of rod photoreceptor function is retinitis pigmentosa (RP); it is a group of inherited retinal dystrophies that triggers the loss of photoreceptors either directly or because of RPE damage. RP clinically manifests along a broad spectrum, depending on the affected gene and the nature of the mutation. Some patients become symptomatic during childhood, while others only develop symptoms in mid adulthood. The rate of disease progression is just as variable, with some patients becoming fully blind within a few years, while other patients never reach that state in their life time (Hamel, 2006; Hartong et al., 2006). Typically, RP patients first suffer from impaired night vision as rod photoreceptors are frequently lost first. Subsequently, patients may lose their peripheral vision, followed by central visual field deficits and eventually blindness. Retinal degenerations thus have a heterogeneous clinical disease presentation culminating in the loss of photoreceptors cells occupying the ONL. The INL and GCL remain largely intact for extended periods of time, although significant synaptic remodeling (Jones and Marc, 2005; Marc et al., 2003; Strettoi et al., 2003) and gliosis (Barber et al., 2013; Hippert et al., 2015; Rodrigues et al., 1987) may manifest during RP. Nonetheless, the fact that the inner neural retina can persist during retinal disease

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provides opportunity for clinical intervention to restore vision. Cone dystrophy is a term for those disorders characterized by loss of cone photoreceptor function. In many of these conditions, the rods can remain largely unaffected (Michaelides et al., 2004; Simunovic and Moore, 1998). In addition to these two groups of disorders, there are some conditions that are characterized cone–rod dystrophy, in which cones degenerate first followed by rod degeneration (Hamel, 2007; Michaelides et al., 2006).

3 THERAPEUTIC AVENUES FOR RETINAL DISEASES 3.1 IMMUNOTHERAPY Until 2006, the only treatment available for AMD was surgical removal of neovascular vessels in patients with wet AMD (Hawkins et al., 2004) or macular repositioning within the retina to place the macula over a less affected area of RPE (MacLaren et al., 2005; Mruthyunjaya et al., 2004). Major progress in combating neovascularization in wet AMD came with the administration of antibodies against the secreted proangiogenic molecule, vascular endothelial growth factor A (VEGF-A). These antibodies were shown to slow disease progression while preserving visual acuity in patients and, in some cases, to reverse visual loss (Rosenfeld et al., 2006; Spaide et al., 2006; Tah et al., 2015). However, a substantial number of wet AMD patients continue to present with declining vision coupled with macular atrophy despite anti-VEGF treatment (Rofagha et al., 2013). Furthermore, the majority of AMD patients have dry AMD and are thus unable to benefit from antineovascular immunotherapies. Other potential targets for immunotherapy for wet AMD include PDGF (Kaiser, 2013) and cytokine signaling (Doyle et al., 2015), whereas potential targets for dry AMD are components of the complement system (Gemenetzi and Lotery, 2015).

3.2 GENE THERAPY Inherited retinal degeneration can be caused by a plethora of genetic defects and have varying prospects for treatment by gene therapy (reviewed by Boye et al., 2013; MacLaren et al., 2016; Smith et al., 2009). Monogenic, recessive gene defects are particularly attractive candidates for gene therapy, since gene supplementation with a normal copy of the gene should, in theory, restore function and ameliorate the disease (reviewed by Smith et al., 2009). Monogenic, autosomal dominant disorders, where the defective gene results in the generation of a toxic product, may necessitate both the suppression of the affected gene and gene supplementation (Ku and Pennesi, 2015). Gene supplementation therapy for recessive defects may be challenging where the size of the therapeutic gene, e.g., ABCA4, exceeds the packaging capacity of adeno-associated viral (AAV) vectors, currently the most effective of viral vectors for retinal gene therapy. The development of dual AAV vectors to deliver the gene in two parts may overcome this problem (Colella et al., 2014; Trapani et al., 2014).

3 Therapeutic avenues for retinal diseases

Alternatively, expression of neuroprotective factors might limit the severity of retinal neurodegeneration, although to date, this has not been a very effective approach. To date, gene therapy has been shown to be partially effective for the treatment of the RPE65 form of Leber’s congenital amaurosis. The RPE65 gene is specifically expressed in RPE cells and encodes retinoid isomerohydrolase. Mutations in this gene cause RPE dysfunction leading to absence of rod function and night blindness and secondary loss of rod and cone photoreceptors. In phase I/II clinical trials, targeting affected RPE by subretinal injection of an AAV vector carrying nonmutant RPE65 cDNA, resulted in improvements in some objective measurements of rod sensitivity and visually guided mobility in dim light (Bainbridge et al., 2008; Maguire et al., 2009). Gene therapy targeting many other retinal dystrophies that affect the photoreceptors themselves is also at the preclinical or clinical stage (Edwards et al., 2016). With rapid progress being achieved in the field of gene editing, especially the emerging potential of CRISPR/Cas9 technology (Jinek et al., 2012), the in vivo correction of genetic disorders by gene therapy may eventually be a feasible treatment option (Maeder and Gersbach, 2016). Ultimately, however, gene therapy is limited by the requirement for the presence of target cells and is most effective when intervention occurs before there is extensive degeneration.

3.3 ELECTRONIC RETINAL PROSTHESIS Even in advanced RP, when most or all the photoreceptors have died, the inner neural retinal layers harboring the amacrine, bipolar retinal ganglion, horizontal, and M€uller cell somata are still present (Jones and Marc, 2005; Marc et al., 2003; Strettoi et al., 2003). This has been exploited in therapeutic approaches that employ an electronic retinal prosthesis. To date, several different groups have tested in clinical trials the implantation of various light-detecting electrical devices (e.g., Argus®II, Alpha-IMS, IMI, IRIS, and EPI-RET 3). These devices generate and pass on electrical stimuli to the remaining retinal circuitry (Luo and Da Cruz, 2014). Some patients who received implants were able to produce implant-mediated visual perception in daily life (Stingl et al., 2013), identify letters and short words (da Cruz et al., 2013), and detect motion in visual tasks (Dorn et al., 2013) with long-lasting improvements observed sometimes years after the implantation (Ho et al., 2015). These are promising results but are tempered by the current limitations in the achievable spatial resolution of the electrical devices.

3.4 REPAIR BY CELL TRANSPLANTATION 3.4.1 Transplantation of donor-derived single cell photoreceptor suspensions As an alternative to the implantation of electronic photosensitive devices, the transplantation of biological material has been extensively explored. The inner neural retina (INL–GCL) that remains following photoreceptor degeneration could also be supplied with visual information by the transplantation of healthy photoreceptors

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to replace those that have degenerated (Fig. 1). Ideally, transplanted cells will integrate into the recipient retina, form synapses with bipolar/horizontal cells, and attain mature photoreceptor morphology. All these aspects are required to detect photons, perform the visual cycle/phototransduction cascade, and to pass on electrically encoded signals to the downstream interneurons. This strategy, however, has the

FIG. 1 Transplantation of isolated photoreceptor precursors in suspension as a strategy for cell replacement therapy in the retina. Photoreceptor precursors at an appropriate developmental stage can be harvested from eyes of donor mice (A) or retinal organoids derived from pluripotent stem cells (B). Most commonly photoreceptor precursors are marked with a fluorescent reporter such as Nrl.GFP in transgenic donor mice (C) or Crx.GFP in a mESC line (D). Donor tissue is enzymatically dissociated into suspension, from which required cells are purified by means of fluorescence-activated cell sorting (FACS) (E) or magnetic-activated cell sorting (MACS). A single cell suspension of purified donor cells is then injected into the subretinal space of recipient mice (F). Engraftment is usually assessed by histology of treated eyes posttransplantation (G). Following successful injection, a donor cell mass (cm) is found surviving in the subretinal space of the recipient animal. Panel (F) was reproduced with permission from Jayakody, S.A., Gonzalez-Cordero, A., Ali, R.R., Pearson, R.A., 2015. Cellular strategies for retinal repair by photoreceptor replacement. Prog. Retin. Eye Res. 46, 31–66. http://dx.doi.org/10.1016/j.preteyeres.2015.01.003.

3 Therapeutic avenues for retinal diseases

distinct advantage of potentially recreating (at least some of ) the preexisting retinal circuitry for optimal processing of visual information. Numerous studies over the past few decades have attempted to address the challenge of replacing lost photoreceptors in the degenerating retina. Several studies focused on the transplantation of neural stem cells (e.g., Banin et al., 2006; McGill et al., 2012) or retinal progenitor cells (RPCs) (e.g., Klassen et al., 2004; Luo et al., 2014). While these cells survive quite well, their ability to generate differentiated retinal cell types, particularly photoreceptors, following transplantation appears limited. In the 1990s, Gouras and colleagues, and Lund and colleagues, reported on transplantation of isolated postnatal photoreceptors derived from donor animals into the RCS rat or the C3Hrd1/rd1 mouse. In these animal models of retinal degeneration, there is an almost complete loss of endogenous photoreceptors. Combined, these studies indicated that transplanted photoreceptors were able to extend nascent outer segment-like structures toward the RPE and putative presynaptic structures toward cells of the INL (Gouras et al., 1991a,b,c; Kwan et al., 1999). However, despite these findings it was uncertain whether transplanted photoreceptors could restore visual function. A number of other studies over the past decade have attempted to address this challenge (MacLaren et al., 2006; Pearson et al., 2012). We reported the recovery of some visual function in stationary night-blind mice (Gnat1
/
) following subretinal transplantation of donor-derived rod photoreceptors (Pearson et al., 2012). Gnat1
/
mice possess a largely intact ONL, populated by nonfunctional rod photoreceptors. To isolate purified populations of donor-derived rod photoreceptors, most of the studies discussed in this section used the transgenic Nrl.GFP+/+ mouse as a source, in which GFP expression is under transcriptional control of the neural retinal leucine zipper gene (Nrl) promoter, which becomes transcriptionally activated specifically in rod photoreceptors shortly after terminal mitosis (Akimoto et al., 2006). Following the transplantation of Nrl.GFP+ postmitotic rod precursor cells (MacLaren et al., 2006), GFP + cells were found correctly located within the host Gnat1
/
retina. In terms of morphology, the labeled rod photoreceptors had nuclei that were correctly located within the ONL, single outer processes that terminated in inner and outer segments pointing toward the RPE as well as single inner processes that terminated in the OPL as axon terminals. These harbored presynaptic release machinery, correctly opposed by the postsynaptic dendrites of bipolar and horizontal cells (Bartsch et al., 2008; Lakowski et al., 2010; MacLaren et al., 2006; Pearson et al., 2012). Most importantly, nearly all of the GFP+ cells within the host retina expressed rod a-transducin, the protein normally missing from the endogenous Gnat1
/
photoreceptors. The improvement in visual function could be detected on several levels in these mice, which normally lack rod-mediated (scotopic) vision: following transplantation and upon stimulation with scotopic light, single cell recordings showed that GFP+ cells within the recipient retina were responsive to dim light (in contrast to neighboring GFP negative cells); RGCs, as well as the visual cortex, showed light-dependent responses; and lastly, the mice also demonstrated behavior mediated by scotopic

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vision in a visual cue-based water maze and optokinetic head tracking tests. Crucially, the degree of functional restoration was dependent upon high numbers of GFP + photoreceptors within the host ONL (MacLaren et al., 2006; Pearson et al., 2012; Warre-Cornish et al., 2014). Similar observations have been reported following the transplantation of rod precursors into the Crx
/
model of retinal degeneration (Neves et al., 2016). As was collectively demonstrated by Gust and Reh (2011), MacLaren et al. (2006), and Pearson et al. (2012), committed, but not fully differentiated, rod photoreceptor precursor cells isolated from P4–8 Nrl.GFP+/+ donor mice by fluorescence-activated cell sorting (FACS) yielded the best transplantation outcomes. On the other hand, using rod photoreceptors derived from mice younger than P0 or older than P10, or RPCs resulted in poor transplantation outcomes. Mature Nrl.GFP+/+ rod photoreceptors show a markedly reduced survival potential compared to immature ones in vitro (R.A. Pearson and A. Kalargyrou, unpublished observations), which might be one factor for their poor transplantation potential. Lamba and colleagues described a conserved immune modulatory mechanism important for tissue repair; platelet-derived growth factor (PDGF)-like signaling induced mesencephalic astrocyte-derived neurotrophic factor (MANF) in innate immune cells. MANF was reported to promote neuroprotection and tissue repair and improved the success of photoreceptor replacement therapies (Neves et al., 2016). This study also showed that Nrl.GFP+/+ donor-derived photoreceptors, similar to those described by Pearson and colleagues (2012), were able to drive visual responses in the Crxtvrm65 model of rapid retinal degeneration, as assessed by ERG (Neves et al., 2016). Successful transplantation thus seems to rely on the fact that the donor cell population is specifically enriched for postmitotic rod photoreceptors and that donor cell survival is key to transplantation outcome. Much of previous research has focused on replacing rod photoreceptors. This has been driven largely by pragmatic reasons, the most prominent being that the model of choice, the mouse, is nocturnal and thus has a rod-dominant retina. However, human vision is primarily dependent on cone, rather than rod photoreceptors. Cones are vital for visual acuity, color, and daylight vision. Diseases that lead to their loss, such as AMD, are particularly devastating. However, since even the human retina consists of only 5% cones (Sz
el et al., 1996), it is reasonable to expect to restore significant cone-mediated function with the replacement of relatively few cones. We provided the first report of cone transplantation (Lakowski et al., 2010), using a Crx.GFP transgenic line that labels developing Crx-expressing rod and cone photoreceptors (Samson et al., 2009). Transplantation of embryonic-stage Crx-positive (Crx +) donors, the majority committed to a cone fate, led to the presence of a few thousand reporter-labeled cells within the host retina. A few of these cells resembled cones, but the majority had rod-like morphology. The mixed nature of the Crx.GFP donor population (rods and cones) left open the question of whether the preponderance of rod-like cells was due to plasticity in the fate of the donor photoreceptors or the result of more successful integration of the rod precursors (vs the cones) present within the

3 Therapeutic avenues for retinal diseases

mixed population. Recently, the groups of Ader and Wallace have independently reported the transplantation of purified populations of cone or cone-like cells. Again, many of these resembled rods in their morphology (Santos-Ferreira et al., 2014; Smiley et al., 2016), an observation we have also made in our own investigations (R.A. Pearson, unpublished findings). Ader and colleagues also reported expression of cone markers within these rod-like cells and assessed the functionality of these cells, using multielectrode array recordings on retinas from Pde6ccpfl1/cpfl1 recipients, reporting a significant increase in light-evoked activity in retinas that had received transplanted cones, compared with those that had received rods or a sham injection (Santos-Ferreira et al., 2014).

3.4.2 Cytoplasmic material transfer Until very recently, the commonly held understanding was that transplanted donor photoreceptors migrated into and integrated within the host retina. Two recent studies, one by ourselves (Pearson et al., 2016) and the other by Ader and colleagues (Santos-Ferreira et al., 2016) have challenged this notion. These independent studies have shown that while a small number of donor cells integrate, as previously observed (MacLaren et al., 2006), the majority remain in the subretinal space, where they engage in a process of cytoplasmic material transfer of RNA and/or protein between donor and host photoreceptors. The underlying cellular mechanisms remain to be elucidated but they do not involve classic processes of cell–cell fusion, as seen between transplanted bone marrow-derived progenitor cells and diverse cell types in several tissues (Terada et al., 2002; Wang et al., 2003), and no translocation of the nucleus occurs. The method of transfer is sufficiently robust to allow host photoreceptors to express a wide variety of proteins that they are otherwise missing (e.g., rod a-transducin, rhodopsin, peripherin-2), in addition to the fluorescent reporters (GFP, DsRed) used to tag the donor cells. These findings have far reaching implications and will require a significant reinterpretation of the existing literature. Not least, we must determine if material transfer is specific to the intact retina and/or whether cell integration can occur at greater rates in degenerated retinae, where barriers such as the outer limiting membrane may be compromised and may aid integration (and/or material transfer) (Barber et al., 2013; Pearson et al., 2010; West et al., 2008). Moreover, identifying the cellular mechanisms behind cytoplasmic material transfer may present an unexpected novel approach to targeting and treating diseased host cells.

3.4.3 Transplantation of donor-derived photoreceptors into severely degenerated retinae The proof-of-concept studies described earlier have typically involved transplantation into slowly degenerating or stationary models of retinal degeneration, the rationale being to give the donor cells the best chance of rescuing visual function without the considerable complexities presented by the severely degenerated retina. Nonetheless, end-stage disease is likely to represent the most suitable target for the first

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clinical trials. Moreover, in the absence of host photoreceptors, cytoplasmic material transfer cannot occur, so it is important to determine if restoration of vision can be achieved through establishment of functional connectivity between transplanted photoreceptor cells and host INL. Some of the earliest studies to address this were those by Lund and colleagues, who transplanted donor-derived postnatal retinal cells into the PDEbrd1/rd1 mouse model of rapid degeneration and showed donor cells in close apposition with the host inner retinal neurons and some indication of supporting visual function, as measured by increased light avoidance behavior at low light levels, compared to control animals (Kwan et al., 1999). More recent studies have revisited this, transplanting donor-derived retinal cells into PDEbrd1/rd1 (Barber et al., 2013; Singh et al., 2013) or P347S mouse models (Eberle et al., 2012). All three studies provided evidence for survival of donor cells with no migration into the remaining retinal tissue. However, the donor cells formed (largely disorganized) clusters within the host subretinal space. Some evidence for expression by the donor cells of the proteins missing in the host retina was seen in each of the studies. Singh and colleagues further examined the functionality of these donor cells using a variety of tests including electroretinography (ERG), light/dark box tests, pupillary reflex, and cortical imaging, that suggested functional improvement following transplantation (Singh et al., 2013). It should be noted, however, that the model used in this study was a strain of PDEbrd1/rd1 retinal degeneration mice that we have recently shown to harbor a second mutation, in the Gpr179 gene, which affects ON-bipolar cells, the prime target of rod signaling (Nishiguchi et al., 2015). Further experiments are, therefore, required to confirm the circuitry used by donor rods to drive visual function in this model.

3.4.4 Transplantation of retinal sheets Besides transplanting photoreceptors by injection of a cell suspension, the transplantation of full thickness retinal sheets has also been investigated (see Seiler and Aramant, 2012, for a comprehensive review). Conceivably, the establishment of synaptic connectivity between graft retinal sheet and recipient retina might be more difficult to establish than that following transplantation of single cell suspension photoreceptors alone. Whether one results in better visual outcome and transplant survival has yet to be determined in a systematic comparative study (Zarbin, 2016). However, donor retinal sheets derived from embryonic rats that were transplanted subretinally into adult recipient rats have been reported to form outer segments toward the recipient RPE (Seiler and Aramant, 1998) and to help sustain light-evoked activity in the superior colliculus of the S334ter-3 retinal degeneration rat at advanced stages of degeneration, compared to sham injected controls (Seiler et al., 2010). Radtke, Aramant and Seiler and colleagues have also provided several reports of improved visual function and visual acuity following transplantation of retinal sheets derived from perinatal human retina in several patients with RP and AMD, although further studies with more extensive tests of visual function will be required (Radtke et al., 1999, 2008; see Seiler and Aramant, 2012).

4 Pluripotent stem cells

4 PLURIPOTENT STEM CELLS PSCs have the ability to self-renew indefinitely and to differentiate into cells of all three germ layers (endoderm, mesoderm, and ectoderm), including retinal cells and, more specifically, photoreceptors (De Los Angeles et al., 2015). They lend themselves to in vitro studies of development as well as disease modeling and high-throughput drug screening, while also providing a renewable source of transplantable material that could eliminate the dependence on donor-derived material. PSCs can be derived from the inner cell mass of preimplantation stage blastocysts, as was demonstrated initially with mouse embryonic stem cells (mESCs) (Evans and Kaufman, 1981; Martin, 1981) and, subsequently, with human ESCs (hESCs) (Thomson et al., 1998) (Fig. 2). Pluripotency can also be reintroduced into differentiated somatic cells via the forced expression of four transcription factors usually associated with the stem cell state: Oct4, Sox2, cMyc, and Klf4 (Takahashi and Yamanaka, 2006). Such iPSCs, although derived from postembryonic stages, were shown to be similar, if not identical, in various aspects such as transcriptional profile, epigenetic landscape, differentiation potential, and mutational load, to ESCs, which are considered to be the gold standard in pluripotency characteristics (Bilic and Izpisua Belmonte, 2012; Chin et al., 2009; Jaenisch et al., 2010; Mayshar et al., 2010). iPSCs have been generated from diverse cells and tissues such as fibroblasts (Takahashi and Yamanaka, 2006), keratinocytes (Aasen et al., 2008), blood (Loh et al., 2009), hematopoietic stem cells (Eminli et al., 2009), and fetal RPE (Hu et al., 2010). It has emerged, however, that the efficiency of iPSC generation decreases with increasing source cell/tissue differentiation status (Eminli et al., 2009). Furthermore, iPSCs generated from a given somatic cell/tissue type retain the epigenetic memory of the previous differentiated state, thus, favoring the redifferentiation along a related lineage (Jaenisch et al., 2010). Thus, reliable and robust differentiation protocols that favor the production of the desired differentiated cells are required and, where iPSC technology is involved, choosing an appropriate somatic source cell/tissue may also have to be taken into consideration (Jayakody et al., 2015). The following sections describe some of the recent advances in the field of ESC/iPSC-derived retinal differentiation.

4.1 PIONEERING WORK IN NEURAL SPECIFICATION AND RETINAL DIFFERENTIATION ESCs, as well as iPSCs, can be maintained in a mitotic stem cell state or be driven to differentiate into all three germ layers. To achieve the latter, they are typically stimulated to enter certain lineage pathways by exposing them to defined modifications in the in vitro culture environment, which may include the following: changes in the concentration of growth factors, inhibitors and pharmacological agents; alterations in serum, nutrient, and O2 concentration; changes in the extracellular matrix (ECM) milieu as well as the forced expression/repression of relevant genes (Kurosawa, 2007). In addition, the cells may be grown in the classic 2D monolayer approach

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FIG. 2 Stem cell sources for in vitro retinal research and cell therapy. Most research into cell replacement in the retina has focused on using photoreceptor cells (D) isolated from donor mouse retina (A and C). Search for expandable and renewable cell sources triggered interest in defining the capacity of various stem cells for retinal repair. Pluripotent or multipotent stem cells can be derived from various sources (A–C, G, and H), including inner cell mass of the blastocyst (H) (ESCs), adult eye (A) (ocular stem cells), or other adult tissues (G) (iPSCs). These ocular stem cells consist of cells of the iris and ciliary body (B), stem €ller cells (F). All these stem cell cell features can be also triggered in RPE (E), and Mu populations can be cultured in vitro and directed into specific retinal lineages as required through appropriate culture conditions generating cell populations relevant for cell therapy development as well as in vitro disease modeling. Importantly, retinal morphogenesis can be recapitulated in the 3D culture of pluripotent stem cells with the formation of optic cups (I) that contain laminated retinal tissue (J). Typically, ocular stem cells show a much restricted differentiation capacity compared to ESCs or iPSCs.

4 Pluripotent stem cells

or in self-assembling 3D stem cell aggregates, called embryoid bodies (EBs) and grown in suspension, or a combination of these two methods. The in vitro differentiation of PSCs into a desired cell type likely mirrors the situation in vivo; stem cells gradually lose their pluripotency while gaining properties associated with the mature differentiated state, which is orchestrated by various signals (summarized in Fig. 3B). During early development, mitotic cells of the rostral neural plate are specified to form the eye field (Chow and Lang, 2001), which eventually give rise to the neural retina and its resident photoreceptors. This step-wise instruction appears to have to be recapitulated to produce neural retinal cells (including photoreceptors) in vitro (Fig. 3A). Pioneering in vitro work by the Sasai group showed that mESCs can be specified toward a rostral neural plate fate (Watanabe et al., 2005). They used a 3D, serum-free, floating culture of EB-like aggregates system (SFEB), which, in the absence of caudalising signals, gave rise to rostral telencephalic neuronal precursor cells. More specifically, the SFEB cultures were exposed to antagonists of Wnt and Nodal signaling (Dkk-1 and LeftyA ! SFEB/ DL), both of which are required for the establishment of the anterior–posterior axis during development (Glinka et al., 1998; Schier and Shen, 2000). It is worth noting the lack of serum in the culture media (knockout serum replacement (KSR) was used instead). While this removed uncertainty from their differentiation protocols (undefined serum components and batch-to-batch differences), it also represents a step toward good manufacturing practices (GMP) standards. The same group subsequently reported the induction of an RPC fate (Rx+/Pax6+/ Ki67+) by adding fetal calf serum (FCS) and activin at later stages of SFEB/DL culture (Fig. 3C), now referred to as SFEB/DLFA (Ikeda et al., 2005). However, RPC induction was relatively inefficient (16% of cells in mESC-derived cultures). With the goal of increasing the in vitro production of photoreceptors, the same group in a later study isolated and cultured mESC- and hESC-derived Rx+ RPCs followed by exposure to the Notch inhibitor DAPT (Fig. 3C) (Osakada et al., 2008, 2009). This promotes RPC cell cycle exit, giving rise to neural retinal cells usually born at the developmental stage at which Notch inhibition is introduced (Jadhav et al., 2006); photoreceptors are born in great numbers during a broad developmental time window (Young, 1985) and their production should thus be favored. The cultured Rx+ RPCs were also treated with soluble factors previously identified to promote photoreceptor genesis: aFGF, bFGF, SHH, retinoic acid, and taurine (Levine et al., 2000; Osakada et al., 2008, 2009). Cells expressing more mature photoreceptor markers such as the opsins were found in both mESC- and hESC-derived cultures, but differentiation usually required significantly longer for human compared with mouse stem cells, in keeping with the longer developmental periods seen in vivo. The photoreceptor differentiation protocol developed by the Sasai group was perhaps the first to demonstrate the step-wise differentiation of PSCs into photoreceptors using (almost exclusively) strictly defined culture conditions. However, other slightly different, sometimes less defined, protocols by other groups have also led to the in vitro production of photoreceptors; e.g., the rostralization/ neuralization of mESCs with insulin–transferrin–selenium fibronectin (ITSFn)

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FIG. 3 Directed differentiation of pluripotent stem cells toward retinal tissues. Physiologically relevant retinal tissue can be obtained through recapitulation of major retinal morphogenesis steps by directed differentiation of pluripotent stem cells (A). In most protocols, stem cells are aggregated into an embryoid body cluster, which subsequently undergoes neural induction in appropriate culture conditions. Neural progenitors need to acquire a forebrain identity to be able to form an eye field-like neuroepithelium. Addition of Matrigel is often used to facilitate the formation of thick and rigid retinal neuroepithelium. Once an eye field-like domain is specified, 3D culture allows evagination of an optic vesicle structure, which folds into a bilayered optic cup as occurs in vivo. Both cell intrinsic genetic programs, as well as input from soluble factors and basement membrane components, drive this sequence in vivo (B). Many soluble and extracellular matrix components were utilized in differentiation protocols to date in an attempt to faithfully recapitulate this sequence of morphogenetic events and their use is schematically represented in (C). Figure 3 was adapted with permission from Jayakody, S.A., Gonzalez-Cordero, A., Ali, R.R., Pearson, R.A., 2015. Cellular strategies for retinal repair by photoreceptor replacement. Prog. Retin. Eye Res. 46, 31–66. http://dx. doi.org/10.1016/j.preteyeres.2015.01.003.

4 Pluripotent stem cells

and bFGF followed by the coculture with postnatal retinal cells (Zhao et al., 2002) or the rostralization/neuralization of hESCs and human iPSCs (hiPSCs) with Dkk-1 and noggin (BMP antagonist) as well as eye induction with IGF-1 (Lamba et al., 2006, 2010). The latter protocol relies on Matrigel for large parts of the culturing period, thus causing issues for GMP compliance. The Gamm group have developed a differentiation protocol for hESCs and hiPSCs that remarkably produces photoreceptors using standard off-the-shelf neuronal media and without the addition of exogenous factors at initial culture stages, reliant largely on the increased endogenous biosynthesis of Dkk-1, noggin, and FGFs (Fig. 3C) (Meyer et al., 2009). This appears to be stimulated by the culture conditions, i.e., the choice of media and switching from suspension to adherent and back to suspension culture. Collectively, these studies revealed the potential for generating retinal cells from PSCs of mouse and of human origin (embryonic and induced). While these were significant advances, the photoreceptor cells generated still lacked the ability to reach maturity, particularly the robust generation of outer segments, in vitro.

4.2 GROWING RETINAL ORGANOIDS DERIVED FROM PSCs In the reports discussed thus far, photoreceptors had been generated in vitro either in 2D adherent cultures or in EB aggregate suspension cultures. Both lack the capacity to produce photoreceptors within the context of an intact retinal neuroepithelium, which may impact on their differentiation. This changed with the ground-breaking study by the Sasai group, when they grew their mESCs as quickly aggregating SFEB cultures with KSR, nonessential amino acids, pyruvate, 2-mercaptoethanol and, crucially, with added Matrigel growth factor reduced (GFR) (Fig. 3C) Eiraku et al. (2011). Under these conditions, the SFEB cultures spontaneously formed self-organized retinal epithelia, which, in addition to expressing RPC markers, remarkably showed in vivo-like morphogenetic movements (evagination and invagination) in the absence of neighboring tissues usually found within the developing eye, such as the surface ectoderm and lens (Fig. 3A). RPCs even displayed interkinetic nuclear migration—a migratory behavior of progenitor cells usually only observed within intact epithelia (Norden et al., 2009; Pearson et al., 2005). In vitro retinogenesis progressed even further following the manual separation of the retinal neuroepithelial optic cups from the remaining EB: in the presence of all-trans retinoic acid and taurine, the optic cups circularized into optic vesicles, which became stratified and contained all retinal neurons and M€uller glia within the correct strata. Of clinical relevance, self-organized neural retina could also be generated from hESCs, although this required prolonged culturing times compared to mESCs and further rostralising signals resulting from addition of the Wnt inhibitor, IWR1e, the Hh/smoothened agonist SAG and FCS (Fig. 3C) (Nakano et al., 2012). Other groups have subsequently reported the production of iPSC-derived optic vesicles (Jin et al., 2012; Mellough et al., 2012), some achieving optic vesicle formation using the Meyer protocol (Meyer et al., 2009), i.e., without Matrigel

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and without adding exogenous factors at initial culturing stages, but relying on the endogenous biosynthesis of rostralising and eye field-inducing signals (Phillips et al., 2012; Reichman et al., 2014). Recently, Zhong et al. (2014) addressed one of the main hurdles of in vitro photoreceptor production: complete photoreceptor maturation including outer segment formation. They modified the Meyer protocol (Meyer et al., 2009; Phillips et al., 2012), which now also relied on the presence of Matrigel (GFR), FCS, taurine, and all-trans retinoic acid to stimulate photoreceptor maturation in hiPSC-derived, long-term optic vesicle cultures (Fig. 3C). Although the protocol is not fully xeno-free, it supports the differentiation of apical connecting cilia on the photoreceptors and, occasionally, primitive outer segment-like formations containing stacked disk. Remarkably, these rudimentary structures formed even in the absence of proximal RPE, but whether an intact photoreceptor–RPE interface is required for full outer segment maturation remains to be seen. Functionally, a small proportion of photoreceptors (2/13) were even responsive to light, as determined by perforatedpatch recordings. In subsequent years, an increasing number of groups, including ours, have reported the successful in vitro generation of (predominantly murine) stem cellderived photoreceptors from optic vesicles, many displaying ciliogenesis and, in some cases, nascent outer segment formation (Chen et al., 2016; Decembrini et al., 2014; Gonzalez-Cordero et al., 2013; Lowe et al., 2016; Mellough et al., 2015; Parfitt et al., 2016; Zhou et al., 2015). These more recent studies mostly utilized permutations of the Sasai group protocols (Eiraku et al., 2011; Nakano et al., 2012). To date, however, no protocols exist for complete in vitro photoreceptor maturation, supporting the full elaboration of functional outer segments to the same extent as in vivo retinogenesis. This necessitates the continued optimisation of current differentiation protocols (Fig. 3C). Among previous such optimisations, Decembrini et al. (2014) demonstrated that hyperoxia applied from the onset of photoreceptor differentiation onward protects newly born photoreceptors from apoptosis. Mellough et al. (2015) included IGF-1 into their protocols to stimulate optic cup genesis similar to previous studies on EBs (Lamba et al., 2006, 2010). Additionally, IGF-1 appears to be important for the maintenance of viable photoreceptors and further accelerates hESC-derived optic vesicle maturation; as previously mentioned, hPSC-derived photoreceptor differentiation is considerably more time-consuming compared with mouse stem cell differentiation (Osakada et al., 2008, 2009). Lowe et al. (2016) managed to bypass the manual separation of optic cups from the remaining EBs in a two-step process: hESC-derived, Matrigel (GFR)-treated, and differentiating aggregates were allowed to spontaneously attach as monolayers with formed tight junctions. Subsequently, dispasemediated detachment of the monolayers and floating culture led to the self-formation of retinal organoids with morphologically and ultrastructurally advanced ciliogenesis and outer segment formation compared with previously published protocols.

5 Challenges for PSC research

Future studies will require further optimisations to the differentiation protocols to enhance in vitro photoreceptor maturation, particularly robust and widespread outer segment formation, to reduce the amount of manual manipulation involved in EB dissections, and to improve banking and storage once the tissue has reached the desired stage. Many areas of the protocol require optimisation: retinal organoid genesis may benefit from enhanced nutrient absorption and improved general culture conditions in spinning bioreactors in a similar way as cerebral organoids (Lancaster et al., 2013; reviewed in Ovando-Roche et al., in press). Another issue to address could be a more complete recapitulation of ocular development, by stimulating the in vitro generation of neural retina in conjunction with surrounding tissues such as surface ectoderm, lens, and RPE (see Hayashi et al., 2016; Kuwahara et al., 2015; Songstad et al., 2015). This may enable photoreceptors to mature beyond currently observed levels.

5 CHALLENGES FOR PSC RESEARCH Many challenges still need to be addressed to unleash the full potential of PSCs in clinical applications such as retinal cell therapy, as well as for the in vitro modeling of development and disease. Some of these challenges were recently discussed by Chader and Young (2016). One challenge is to develop protocols that are capable of further biasing the differentiation of RPCs within PSC-derived retinal organoids toward specific photoreceptor fates (cone subtypes and rods)—the retinal cell type affected the most in many retinopathies. More than just their enrichment, it will be important to maximize the level of cellular maturity and function that can be achieved from PSC differentiation protocols. Murine and human stem cell-derived photoreceptors have been shown to express a number of photoreceptor markers, although not all protocols yield expression profiles equivalent to the corresponding in vivo stages (Lamba et al., 2010; West et al., 2012; see Gonzalez-Cordero et al., 2013). Future studies with hPSC-derived photoreceptors will need to demonstrate their similarity to photoreceptors in vivo using techniques such as microarray analysis. A better understanding of the gene regulatory networks governing the cellular and functional maturation of photoreceptors could lead the way to new growth factor/inhibitor or tissue culture treatment regimens during the differentiation of PSCs. Although retinal organoids are intrinsically capable of displaying some degree of photoreceptor maturation (Chen et al., 2016; Decembrini et al., 2014; Gonzalez-Cordero et al., 2013; Lowe et al., 2016; Mellough et al., 2015; Parfitt et al., 2016; Zhou et al., 2015), complete photoreceptor differentiation encompassing the establishment of connecting cilia combined with robust elaboration of outer segments has so far not been achieved in vitro. It remains to be seen whether ocular organoids (rather than retinal organoids), which in addition to neural retinal tissue, also contain surface ectoderm, lens, and RPE, could better support full photoreceptor maturation.

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The pioneering work on stem cell-derived photoreceptor generation was based on mESC and hESC studies (Osakada et al., 2008, 2009) and these still represent the most advanced donor cell source, in terms of our ability to generate apparently developmentally normal retinal structures. However, hiPSCs have some logistical and ethical advantages (Chader and Young, 2016). For hiPSCs to realize their full potential, further studies will need to focus on optimizing hiPSC manufacturing protocols that result in efficient and reproducible differentiation into retinal organoids and photoreceptors. This will require systematic studies to identify the best somatic cell source to convert into hiPSC lines, given factors such as epigenetic memory (Jaenisch et al., 2010), as well as optimisations of reprogramming protocols to efficiently obtain footprint-free iPSCs (i.e., without viral integration of the stem cell transcription factor expression constructs) (Malik and Rao, 2013). Following reprogramming, the differentiation of hiPSCs, or indeed hESCs, into retinal organoids/photoreceptors is very time-consuming, certainly when compared with the differentiation of mouse PSCs (Nakano et al., 2012; Osakada et al., 2008). Although accelerated protocols would be desirable, it remains to be determined whether full functionality can be guaranteed under such conditions. Important for future clinical application are the continued efforts to achieve GMP-compliant protocols for all the processes involved, including generating the cell source, differentiation, storage, and distribution (Solomon et al., 2015). Due to the long time required to differentiate human PSCs into photoreceptors, appropriate storage methods for photoreceptors and/or retinal organoids have to be developed, so that they can be obtained on demand (Jayakody et al., 2015). Furthermore, completely xeno-free differentiation protocols should be developed to avoid any potential risks of zoonoses, activation of animal retroviruses, or immune rejection following cell therapy. Immune rejection may continue to be a risk even when cell therapies from allogenic PSCs are to be produced under GMP conditions due to mismatched human leukocyte antigens (HLA) donor and recipient haplotypes. Recently, efforts have been made to generate hiPSC banks to match HLA haplotypes of large proportions of the general population (Taylor et al., 2012). This is an enormous challenge with significant financial cost and commercial risk, which will likely require concerted and multinational efforts to be overcome (Bravery, 2015; Solomon et al., 2015) but could have enormous benefits. Patient-derived hiPSCs destined for autologous therapy circumvent the need for HLA-matching. RP-causing mutations in such patient-derived hiPSCs can be rectified in vitro using CRISPR/Cas9 gene editing technology, providing the prospect of genetically repaired and differentiated hiPSCs being used in future transplantation therapies (Bassuk et al., 2016). Additionally, the differentiation of patient-derived hiPSC into retinal organoids lends itself to easily accessible and manipulatable in vitro disease modeling as well as targeted high-throughput drug screens (Parfitt et al., 2016; Tucker et al., 2013; Wahlin et al., 2014). Promising as they may be, such individually tailored hiPSC transplantation therapies would currently still be challenged by high cost and time demands.

6 Clinical prospects

6 CLINICAL PROSPECTS PSCs are a promising new tool for potential future clinical application (Fig. 4). Exogenous cell replacement therapy represents an obvious approach in cases where retinal cells, such as photoreceptors, have already degenerated. However, with the advent of more sophisticated culture methodologies that allow retinal organoids to develop in vitro and with similar properties to retinae in vivo, future studies should see an increase in drug screens and evaluation of treatments in vitro. Even in cases where disease etiology and pathobiology are unknown, in vitro retinal differentiation of PSCs will likely contribute to a better understanding of the disease process, through disease modeling.

FIG. 4 Utility of directed differentiation of pluripotent stem cells in retinal research. Pluripotent stem cells (including ESCs and iPSCs; A) can be differentiated in vitro into neural retina containing physiologically relevant populations of retinal cell types (B) capable of modeling disease mechanisms in culture (C), be used to screen different treatment options including small molecule compound libraries for potential therapeutic use (D), and provide a source of cells for developing cell therapies (E).

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6.1 CELL TRANSPLANTATION Photoreceptor cell replacement studies initially used donor-derived photoreceptors (Bartsch et al., 2008; MacLaren et al., 2006; Pearson et al., 2012; Radtke et al., 2008). While the most convenient source for proof-of-concept studies, this source of cells is of very limited potential for clinical application. This is because humans reach a developmental stage equivalent to that of the early postnatal donor mouse within the second trimester, which thus imposes significant ethical and logistical barriers. In transitioning toward PSC-derived photoreceptor transplantation strategies, Homma et al. (2013) showed that miPSC-derived Nrl.GFP + cells showed rod-like functional properties in vitro. An initial study on the transplantation of mESC-derived photoreceptors from 2D cultures did not result in tumor formation, an inherent risk with stem cell-derived cellular therapies; it did, however, fail to detect photoreceptor integration (West et al., 2012). Improved transplantation outcomes were observed upon using 3D differentiation to generate mESCs-derived photoreceptors (Gonzalez-Cordero et al., 2013). Retinal organoids allow photoreceptors to mature in a more in vivo-like manner, which likely enables them to acquire more of the characteristics required to achieve good transplantation outcomes compared with 2D differentiation protocols (Gonzalez-Cordero et al., 2013). Of clinical relevance, we, and others, have demonstrated that photoreceptors can be isolated (and transplanted) from retinal organoids based on a cell surface biomarker panel (CD73 +, CD24 +, CD133 +, CD47 +, CD15
) rather than via the transgenic expression of fluorescent proteins (Eberle et al., 2011; Lakowski et al., 2011, 2015). A number of studies have reported improvements in various measures of visual function following the transplantation of photoreceptor-like cells derived from mouse or human PSCs into various mouse models of RP, including some of advanced retinal degeneration (Barnea-Cramer et al., 2016; Homma et al., 2013; Lamba et al., 2009; Neves et al., 2016; Santos-Ferreira et al., 2016; Tucker et al., 2011a). Often, studies have used a single measure of function and it will be important to validate the extent to which visual function can be restored by PSC-derived donor cells using a variety of measures. As described earlier, we and others have recently shown that donor-derived photoreceptors appear to rescue nondegenerative cells by cytoplasmic material transfer from donor to host cells (Pearson et al., 2016; Santos-Ferreira et al., 2016). It is, therefore, of paramount importance to determine the relative contributions made by cytoplasmic material transfer and cell integration to rescue by PSC-derived photoreceptors in the diseased—as opposed to the intact—retina. The first clinical trials for photoreceptor replacement are likely to involve patients with few or no photoreceptors remaining. For this reason, there have been an increasing number of studies focusing on transplantation into murine models of advanced disease. Barnea-Cramer et al. (2016) transplanted hESC- and hiPSC-derived retinal cells into the severely degenerated, end-stage PDEbrd1/ rd1 mouse. The transplanted cells expressed mature photoreceptor markers,

6 Clinical prospects

exhibited signs of synaptic connectivity with cells of the recipient INL and appeared to contribute to improvements in vision behavior. Human vision relies heavily on cone photoreceptors and their transplantation should thus be a major research focus. Recent work from our group has focused on the transplantation of mESC- and hESC-derived cone photoreceptors into the severely degenerated, end-stage Aipl1
/
mouse, where the transplanted cells differentiated and matured within the recipient eye (A. Gonzalez-Cordero and K. Kruczek, unpublished data). Using a different approach, the Takahashi group transplanted mESC- and miPSC-derived retinal organoid sheets into the ONL-lacking PDEbrd1/rd1 mouse (Assawachananont et al., 2014). More recently, they transplanted hESC-derived retinal sheets into primates with induced focal photoreceptor degeneration (cobalt chloride and laser photocoagulation) (Shirai et al., 2015). In both studies, the retinal sheet grafts developed elaborate photoreceptor outer segments, despite the cells being at an immature stage at the time of transplantation. Contact between graft ONL (containing graft photoreceptors) and host INL is the most desirable transplantation outcome, but graft ONL and host INL may also be bridged by graft INL and GCL. It was suggested that, in addition to graft INL/GCL remodeling, graft photoreceptors may synaptically “switch” from graft to some host bipolar cells, forming putative synaptic connections (Shirai et al., 2015).

6.2 DISEASE MODELING PSC-derived retinal cells/organoids are now being used to try to understand disease etiology and pathobiology. For instance, a recent study has shown that an Alu insertion in the male germ cell-associated kinase (MAK) gene causes dysregulation in its developmentally regulated, retina-specific alternative splicing program resulting in RP (Tucker et al., 2011b). In another study, Tucker et al. (2013) identified disease-causing USH2A mutations in an adult patient with autosomal recessive RP, resulting in the exonification of intron 40, a translation frameshift and a premature stop codon. Ultimately, this caused ER stress in hiPSC-derived retinal organoids, the long-term effects of which were likely to eventually lead to photoreceptor degeneration. Understanding disease pathobiology with hiPSCs derived from patients with previously unknown mutations, as was the case with these two studies, could thus facilitate the development of suitable therapeutic strategies.

6.3 DRUG SCREENING/EVALUATION OF POTENTIAL TREATMENTS Our ability to generate retinal organoids from patient-derived hiPSCs that are increasingly similar to endogenous retinal tissue offers opportunities for the in vitro evaluation of therapies in human tissue as well as high-throughput drug screens, which could lead to an acceleration in the identification and evaluation of treatment

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options, a notoriously time-consuming process (Tucker et al., 2014). This approach is still in its infancy, however, and needs to be fully validated before it can translate into the clinic. Readers are referred to the review by Inoue and Yamanaka (2011) for further discussion on this topic. An example of its potential, however, was provided in a recent study by Cheetham and colleagues (Parfitt et al., 2016): A common, deep intronic mutation (c.2991+1665A>G) in the cilia-related CEP290 gene results in aberrant splicing and premature termination. This leads to photoreceptor degeneration due to defects in ciliogenesis and ciliary transport (den Hollander et al., 2008; Drivas et al., 2015). The amount of mis-splicing, and thus truncated gene product, was found to be especially pronounced in hiPSC-derived photoreceptors vs hiPSC-derived RPE. In the latter, sufficient amounts of full-length gene product were still produced (Drivas et al., 2015; Parfitt et al., 2016). This explains why photoreceptors appear to be particularly affected by this splice site mutation. Additionally, splice donor site targeting antisense morpholinos were demonstrated to reduce aberrant splicing in hiPSC-derived optic cups and to rescue ciliogenesis and ciliary transport in photoreceptors.

7 CONCLUSION The last decade has seen considerable progress in our ability to generate PSC-derived retinal cells and tissue that increasingly resemble their endogenous counterparts. This new technology is likely to be of increasing importance for studying developmental biology and disease processes as well as developing new therapies, including cell and gene therapies and conventional drugs. The study of PSC-derived retina may even facilitate the development of methods to activate endogenous repair mechanisms that are dormant in the retina of higher vertebrate species. The ability to generate human PSC-derived photoreceptors provides a major boost for retinal repair through cell therapy. Although photoreceptor transplantation is still some way from effective clinical application, with further optimisation, evaluation and validation, we may see significant clinical advance in the next decade.

ACKNOWLEDGMENTS This work was supported by grants from Fight for Sight (1448/1449), Moorfields Eye Charity, Macular Vision Research Foundation, and the Medical Research Council UK (mr/j004553/1). N.A. is a UCL Grand Challenge PhD student. R.A.P. is a Royal Society University Research Fellow (UF120046). R.R.A. is supported by the National Institute for Health Research (NIHR) Biomedical Research Centre for Ophthalmology at Moorfields Eye Hospital, and UCL Institute of Ophthalmology. R.A.P. is part-funded by the Alcon Research Institute.

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Taylor, C.J., Peacock, S., Chaudhry, A.N., Bradley, J.A., Bolton, E.M., 2012. Generating an iPSC bank for HLA-matched tissue transplantation based on known donor and recipient HLA types. Cell Stem Cell 11, 147–152. http://dx.doi.org/10.1016/j.stem.2012.07.014. Terada, N., Hamazaki, T., Oka, M., Hoki, M., Mastalerz, D.M., Nakano, Y., et al., 2002. Bone marrow cells adopt the phenotype of other cells by spontaneous cell fusion. Nature 416, 542–545. http://dx.doi.org/10.1038/nature730. Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S., et al., 1998. Embryonic stem cell lines derived from human blastocysts. Science 282, 1145–1147. http://dx.doi.org/10.1126/science.282.5391.1145. Trapani, I., Colella, P., Sommella, A., Iodice, C., Cesi, G., de Simone, S., et al., 2014. Effective delivery of large genes to the retina by dual AAV vectors. EMBO Mol. Med. 6, 194–211. http://dx.doi.org/10.1002/emmm.201302948. Tucker, B.A., Park, I.-H., Qi, S.D., Klassen, H.J., Jiang, C., Yao, J., et al., 2011a. Transplantation of adult mouse iPS cell-derived photoreceptor precursors restores retinal structure and function in degenerative mice. PLoS One 6, e18992. http://dx.doi.org/10.1371/ journal.pone.0018992. Tucker, B.A., Scheetz, T.E., Mullins, R.F., DeLuca, A.P., Hoffmann, J.M., Johnston, R.M., et al., 2011b. Exome sequencing and analysis of induced pluripotent stem cells identify the cilia-related gene male germ cell-associated kinase (MAK) as a cause of retinitis pigmentosa. Proc. Natl. Acad. Sci. U.S.A. 108, E569–E576. http://dx.doi.org/10.1073/ pnas.1108918108. Tucker, B.A., Mullins, R.F., Streb, L.M., Anfnson, K., Eyestone, M.E., Kaalberg, E., et al., 2013. Patient-specific iPSC-derived photoreceptor precursor cells as a means to investigate retinitis pigmentosa. Elife 2013, e00824. http://dx.doi.org/10.7554/eLife.00824.001. Tucker, B.A., Mullins, R.F., Stone, E.M., 2014. Stem cells for investigation and treatment of inherited retinal disease. Hum. Mol. Genet. 23, 9–16. http://dx.doi.org/10.1093/hmg/ ddu124. Wahlin, K.J., Maruotti, J., Zack, D.J., 2014. Modeling retinal dystrophies using patient-derived induced pluripotent stem cells. Adv. Exp. Med. Biol. 801, 157–164. http://dx.doi.org/10.1007/978-1-4614-0631-0. Wang, X., Willenbring, H., Akkari, Y., Torimaru, Y., Foster, M., Al-Dhalimy, M., et al., 2003. Cell fusion is the principal source of bone-marrow-derived hepatocytes. Nature 422, 897–901. http://dx.doi.org/10.1038/nature01564.1. Warre-Cornish, K., Barber, A.C., Sowden, J.C., Ali, R.R., Pearson, R.A., 2014. Migration, integration and maturation of photoreceptor precursors following transplantation in the mouse retina. Stem Cells Dev. 23, 941–954. http://dx.doi.org/10.1089/scd.2013.0471. Watanabe, K., Kamiya, D., Nishiyama, A., Katayama, T., Nozaki, S., Kawasaki, H., et al., 2005. Directed differentiation of telencephalic precursors from embryonic stem cells. Nat. Neurosci. 8, 288–296. http://dx.doi.org/10.1038/nn1402. West, E.L., Pearson, R.A., Tschernutter, M., Sowden, J.C., MacLaren, R.E., Ali, R.R., 2008. Pharmacological disruption of the outer limiting membrane leads to increased retinal integration of transplanted photoreceptor precursors. Exp. Eye Res. 86, 601–611. http:// dx.doi.org/10.1016/j.exer.2008.01.004. West, E.L., Gonzalez-Cordero, A., Hippert, C., Osakada, F., Martinez-Barbera, J.P., Pearson, R.A., et al., 2012. Defining the integration capacity of embryonic stem cellderived photoreceptor precursors. Stem Cells 30, 1424–1435. http://dx.doi.org/10.1002/ stem.1123.

References

Young, R.W., 1985. Cell differentiation in the retina of the mouse. Anat. Rec. 212, 199–205. http://dx.doi.org/10.1002/ar.1092120215. Zarbin, M., 2016. Cell-based therapy for degenerative retinal disease. Trends Mol. Med. 22, 115–134. http://dx.doi.org/10.1016/j.molmed.2015.12.007. Zhao, X., Liu, J., Ahmad, I., 2002. Differentiation of embryonic stem cells into retinal neurons. Biochem. Biophys. Res. Commun. 297, 177–187. http://dx.doi.org/10.1002/9780470151808. sc01f01s2. Zhong, X., Gutierrez, C., Xue, T., Hampton, C., Vergara, M.N., Cao, L.-H., et al., 2014. Generation of three dimensional retinal tissue with functional photoreceptors from human iPSCs. Nat. Commun. 5, 4047. http://dx.doi.org/10.3109/10641955.2015. 1046604.Association. Zhou, S., Flamier, A., Abdouh, M., T
etreault, N., Barabino, A., Wadhwa, S., et al., 2015. Differentiation of human embryonic stem cells into cone photoreceptors through simultaneous inhibition of BMP, TGFb and Wnt signaling. Development 142, 3294–3306. http://dx.doi.org/10.1242/dev.125385.

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Stem cell-derived retinal pigment epithelium transplantation for treatment of retinal disease

9

Britta Nommiste*, Kate Fynes*, Victoria E. Tovell*, Conor Ramsden*,†, Lyndon da Cruz*,†,‡, Peter Coffey*,†,§,1 *Institute of Ophthalmology, London, United Kingdom NIHR Biomedical Research Centre, Moorfields Eye Hospital NHS Foundation Trust, London, United Kingdom ‡ Moorfields Eye Hospital NHS Foundation Trust, London, United Kingdom § Center for Stem Cell Biology and Engineering, Neuroscience Research Institute, University of California, Santa Barbara, Santa Barbara, CA, United States 1 Corresponding author: Tel.: 20-7608-4037; Fax: 20-7608-4015, e-mail address: [email protected] †

Abstract Age-related macular degeneration remains the most common cause of blindness in the western world, severely comprising patients’ and carers’ quality of life and presenting a great cost to the healthcare system. As the disease progresses, the retinal pigmented epithelium (RPE) layer at the back of the eye degenerates, contributing to a series of events resulting in visual impairment. The easy accessibility of the eye has allowed for in-depth study of disease progression in patients, while in vivo studies have facilitated investigations into healthy and diseased RPE. Consequently, a number of research groups are examining different approaches for the replacement of RPE cells in age-related macular degeneration (AMD) patients. This chapter examines some of these initial proof-of-principle studies and goes on to review the use of pluripotent stem cells as a source for RPE replacement in a number of current AMD clinical trials. Finally, we consider just some of the regulatory and manufacturing challenges presented in taking a promising AMD treatment from the research bench into clinical trials in patients, and how to mitigate potential risks early in process development.

Keywords Retinal pigmented epithelium, Retinal disease, Age-related macular degeneration

Progress in Brain Research, Volume 231, ISSN 0079-6123, http://dx.doi.org/10.1016/bs.pbr.2017.03.003 © 2017 Elsevier B.V. All rights reserved.

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1 AGE-RELATED MACULAR DEGENERATION Age-related macular degeneration (AMD) is the most common irreversible blinding condition in the western world in the over 60 age group (Buchholz et al., 2009, 2013; Carr et al., 2009a; Du et al., 2011; Kokkinaki et al., 2011; Ramsden et al., 2016; Vaajasaari et al., 2011). It is estimated that AMD will affect over 3 million people in the United States by 2030 and around 700,000 people in the United Kingdom by 2020 (Owen et al., 2012). The exact etiology of AMD is still unclear but is influenced by genetic (e.g., apolipoprotein E, complement factors H and B, and complement components C2 and C3) and environmental factors (e.g., smoking and diet) (Katta et al., 2009; Zeiss, 2010). Susceptibility to AMD is determined by gene–gene and gene–environment interactions (Katta et al., 2009). There are two forms of AMD—dry and wet (Whiting et al., 2015). The latter, affecting about 10% of the overall AMD cases, is acute and presents with neovascularization from the choroid that penetrates under or into the retina (Du et al., 2011). The growth of new fragile blood vessels can result in bleeds underneath the retina, secondary scarring, and eventually leads to sudden loss of vision due to retinal pigment epithelium (RPE) and photoreceptor loss (Ramsden et al., 2013). While there are costly and laborintensive treatments for wet AMD with regular intraocular injections of vascular endothelial growth factor-targeting antiangiogenic drugs (e.g., Lucentis), these are only effective in a limited population and are not a curative therapy (Buchholz et al., 2013; Du et al., 2011; Ramsden et al., 2016; Vaajasaari et al., 2011). The majority of AMD patients (approximately 90%) suffer from dry AMD, which presents with the development of fatty protein deposits, known as drusen, which form between the RPE and Bruch’s membrane, causing RPE and photoreceptor layer degeneration, ultimately leading to loss of vision (Whiting et al., 2015). The formation of these soft drusen interferes with the essential role of the RPE, preventing them from delivering nutrients and removing waste to and from the retina. With age, development of small, hard drusen that is not pathologic is common. However, these hard drusen can increase in size and soften and cause detachment of RPE from the photoreceptor cells, leading to AMD (Zeiss, 2010). The macular region of the retina, an area that is crucial for central vision and important for daily tasks including reading, driving, and recognizing faces, is most vulnerable to the buildup of drusen (Fig. 1). While there has been progress in developing treatments to stabilize, but not reverse, wet AMD, there are currently no treatment options for dry AMD. Hence, patients suffering from AMD require urgent curative clinical therapy options.

2 RPE, ITS FUNCTIONS, AND ROLE IN AMD Age-related degenerative changes in the RPE, resulting in the disruption and failure of RPE, choroid, and Bruch’s membrane function, are a major factor in the development of AMD. The RPE is a polarized monolayer of pigmented, cobblestone-like cells that lies at the back of the eye and maintains close interaction with the overlying

2 RPE, its functions, and role in AMD

Normal vision

Retina Ciliary body Lens Iris

Cornea Anterior chamber

Macula

Amd vision

Damaged macula with drusen

FIG. 1 Schematic presentation of the visual field through a healthy eye, and the eye of an age-related macular degeneration (AMD) patient. The aging RPE undergoes several pathological changes—including RPE injury, oxidative stress, complement activation, mitochondrial dysfunction, and extracellular matrix remodeling—that all cause RPE injury and are a major contributor to AMD pathogenesis (Katta et al., 2009).

photoreceptor cells and the underlying choroid (Ramsden et al., 2013). Interaction with the photoreceptor layer is essential for vision as the RPE is responsible for many critical functions. These include recycling compounds associated with the visual cycle (all-trans retinal reisomerization), phagocytosis of photoreceptor outer segments (POS), secretion of growth factors and nutrients, transport of ions and water, removal of oxidative waste, and providing a barrier to create the immune-privileged posterior chamber of the eye (see Fig. 2; Ramsden et al., 2016; Strauss, 2005; Whiting et al., 2015). During a lifetime, the apical microvilli of the RPE phagocytose millions of outer segment discs. The components of these discs are degraded, recycled, or redelivered to the photoreceptor layer for normal visual cycle function (e.g., all-trans retinal to 11-cis retinal). With age, a derivative from this process, bis-retinoid N-retinyl-Nretinylidene ethanolamine, can accumulate at the base of the RPE, where it may

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FIG. 2 Schematic presentation of the functions of the retinal pigment epithelium (RPE). AMV, apical microvilli; POS, photoreceptor outer segments; RPE, retinal pigment epithelium; TJ, tight junctions.

contribute to drusen formation and potential AMD later in life (Zeiss, 2010). In addition, the photoreceptor cells that transform light into electrical energy release free radicals as a by-product which are also toxic and eventually contribute to the degeneration of the RPE (Ramsden et al., 2016). The basal side of the RPE is in close proximity to the choroidal blood supply, which delivers key nutrients for the normal visual cycle (glucose and vitamin A) and phagocytosis functions (Kokkinaki et al., 2011; Zeiss, 2010). RPE dysfunction contributes to buildup of soft drusen causing RPE detachment from the photoreceptor cell layer which, over time, leads to the failure and loss of the photoreceptor layer and eventual blindness, as seen in AMD (Carr et al., 2013; Zeiss, 2010). During normal aging, accumulation of some hard drusen occurs at the back of the eye accompanied by thickening of the Bruch’s membrane. In AMD Bruch’s membrane becomes fragmented and calcified, and potentially contributes toward accumulation of large soft drusen (Zeiss, 2010). For dry AMD, the geographic atrophy associated with the disease is in general slowly progressing, providing a wide window of opportunity for therapeutic intervention to prevent further degradation. In the case of wet AMD, the timing for intervention is critical. The newly formed blood vessels associated with this disease

3 Proof-of-principle studies

are fragile and have the tendency to leak, resulting in bleeds underneath the retina which can lead to irreversible photoreceptor layer death within months of the onset of neovascularization. Thus, if we are to stem the progression of these debilitating conditions and indeed to develop a curative treatment, it is crucial that we establish a means of replacing the degenerated RPE cells under the macula. Establishing an effective cure for AMD would not only vastly improve the quality of life that patients can enjoy, but would also have significant economic benefit (Shah and Williams, 2016; Velez-Montoya et al., 2014).

3 PROOF-OF-PRINCIPLE STUDIES Many research groups use animal model to perform proof-of-principle studies for testing the efficacy of potential therapeutics. One of the most commonly used models for testing AMD cellular therapies is the Royal College of Surgeons (RCS) rat. This retinal degenerative animal model has a mutation in the Mertk gene, which encodes a protein (met proto-oncogene tyrosine kinase) essential for the phagocytosis of POS discs. Defects in the phagocytosis pathway lead to degeneration of the photoreceptor cell layer and subsequent blindness within months after birth (D’Cruz et al., 2000). Transplantation of donor RPE from nondystrophic rats, or of fetal, neonatal, adult, and stem cell-derived human RPE, has been shown to delay the onset, and prevent or reverse the degeneration of RPE in the RCS rat (Alvarez Palomo et al., 2015; Carr et al., 2009a; Coffey et al., 2002; Lavail et al., 1992; Vugler et al., 2008). These experiments established the paradigm that transplantation of healthy donor RPE could prevent the degeneration if not completely reverse it. Experimental surgeries, such as autologous donor RPE transplants, where an RPE patch is harvested from the periphery and immediately transplanted under the macular region, or macular translocation surgery, where the retina is detached and repositioned over a healthy area of RPE, provide proof of principle for RPE cellreplacement therapy in the human eye. However, although these treatments can maintain visual function in some patients, these approaches may have limited long-term success (MacLaren et al., 2007; Treumer et al., 2007; van Meurs and Van Den Biesen, 2003; Whiting et al., 2015), the procedures themselves are lengthy, involve follow-up surgery, and have a high rate of postoperative complication (Eckardt et al., 1999; Machemer and Steinhorst, 1993; MacLaren et al., 2007; Treumer et al., 2007). Transplantation of postmortem RPE tissue into degenerative eyes has had mixed results; this may be due to changes in RPE cell physiology and function after harvest (Blenkinsop et al., 2015; Buchholz et al., 2009). Cadaver RPE transplants have led to chronic allograft rejection and complete encapsulation of the patch with white fibrotic tissue without immunosuppression (Del Priore et al., 2001). The transplantation of cadaver and/or donor RPE as well as translocation of RPE surgeries is an option for AMD treatment; however, due to the large number of patients, these approaches cannot provide the urgent, limitless source of RPE that is required for a viable cellular therapeutic. Hence, the need for an alternative source

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of RPE cells. Focus has now switched to developing treatments using pluripotent stem cell-derived RPE that could in theory be produced in limitless amounts. A body of literature has shown that pluripotent stem cell-derived RPE cells are capable of recapitulating the same functions in vivo as the native healthy human RPE (Carr et al., 2013; Coffey et al., 2000; Kokkinaki et al., 2011; Vugler et al., 2008). Pluripotent stem cells can be generated from few cells of a human embryo (human embryonic stem cells, hESCs) or skin cells that have been reprogrammed back to pluripotency (induced pluripotent stem cells, iPSCs). hESCs as well as iPSCs can be differentiated into RPE by withdrawal of basic fibroblast growth factor (bFGF) from the tissue culture media, a process known as spontaneous differentiation (Carr et al., 2009a,b; Vaajasaari et al., 2011; Vugler et al., 2008). Pigmented foci can be identified between weeks 3 and 5 after bFGF withdrawal. These foci can then be manually isolated to purify the RPE from the differentiated cultures. The isolated RPE cells can then be cultured as a monolayer and used for downstream assays in research facilities or for clinical use as stem cell-derived donor RPE. In research, RPE cells derived from patient iPSCs can be used to resolve the molecular pathology of inherited mutations that contribute toward RPE diseases, an essential step in identifying alternative novel and viable therapeutics for these patient groups (Schwarz et al., 2015). Our group has previously demonstrated that RPE can be produced from hESCs and iPSCs, and is now working toward using these cells as a clinical therapeutic. The hESC-derived RPE project has reached clinical trial, while iPSC-derived RPE manufacture is still in process development. However, the latter has and will continue to provide a limitless source of RPE to model various patient diseases, including AMD, in a dish. iPSC-derived RPE could, therefore, not only provide autologous or even human leukocyte antigen-matched RPE transplants in the future, but also contribute to the understanding of disease (e.g., AMD) progression. In clinical studies, the source of cells as well as the purity and maturity of these cells plays a crucial part in ensuring the recipient of the cells will gain the best outcome. The successful transition of stem cell-derived RPE to clinical applications will have many challenges, involving safety as well as financial and regulatory concerns. In order to surmount this rigorous process, the first step must be to ensure that the RPE cells derived in vitro are as comparable to the native RPE tissue as possible. Ideally, stem cell-derived RPE should have the correct polarized morphology in vitro, possess a gene signature that reflects that of native RPE, and be capable of all of the cardinal functions of RPE, which have been shown by many research groups (Alvarez Palomo et al., 2015; Carr et al., 2009a; Coffey et al., 2002; Lavail et al., 1992; Lu et al., 2009; Sugino et al., 2011; Vugler et al., 2008). Second, advances in medical retina imaging and functional testing techniques have enabled researchers to track the survival and measure the functional capacity of transplanted cells in vivo. These techniques will be essential to evaluate the efficacy and safety of cells in human trials (Schwartz et al., 2012, 2015). A number of other concerns should be taken into account when considering using pluripotent stem cell-derived RPE as a cellular therapy. Although the eye is an immune-privileged site, there may be issues with immunocompatibility of the donor

4 Clinical results and considerations

RPE, particularly when using hESC-derived RPE cells. Due to its autologous nature, iPSC-derived RPE holds great promise for overcoming immune rejection since RPE for transplantation could be created from the patient’s own cells (Blenkinsop et al., 2015; Buchholz et al., 2009; Kokkinaki et al., 2011). Autologous iPSC-derived RPE cells have already been used in a Japanese clinical trial for AMD. Given the degenerative nature and slow progression of AMD, patient-derived cells could provide a viable source of donor tissue which could circumvent rejection issues. Genetic predispositions toward AMD contained within a patient’s own cells should not be of concern since reintroduction of cells with genetic defect will probably take years to reveal pathology (Buchholz et al., 2009). However, due to cost issues, it would not be economically viable to use this approach for all patients. To overcome this, banking enough hESC and iPSC from various healthy donors with a range of human leukocyte antigens will potentially have a better chance in overcoming the graft-host disease and would be a more cost-effective source of cells. Another major issue when using stem cell-derived cells for transplantation is the possible formation of teratomas. Unlike other organs within the body, the eye has the benefit of having a transparent window through which light travels; this window can also be used to track the progress of transplanted cells. Using relatively simple ophthalmic instruments, surgeons will be able to track cells within the eye over the long term. In the case of a serious adverse incident, such as teratoma formation, the transplanted tissue could be treated with laser ablation or, as a last resort, the eye could be removed from a patient who, without the transplant, would have been rendered blind by the disease (Whiting et al., 2015).

4 CLINICAL RESULTS AND CONSIDERATIONS Owing to the complexity and risks associated with the translocation and cadaveric/ autologous RPE transplantation surgeries, recent research has focused on preparing novel sources of donor RPE. With the success and current advances in stem cell research, scientists now have the opportunity to generate hESC- or iPSC-derived RPE which demonstrates huge potential for the treatment of AMD, once safety and cell survival issues are addressed. The means by which these cells should be delivered to the back of the eye in order to achieve maximum efficacy is still under investigation. To date, there are two types of clinical studies involving cell therapies for AMD in progress: a bolus injection of RPE cell suspension to the back of the eye and the transplantation of an RPE monolayer graft patch between the retina and existing RPE, either on a substrate or not. Injection of RPE cells offers a convenient, low cost, and speedy treatment due to required culture time and ease of transplantation. At the forefront of hESC-based therapy is Astellas Pharma US (IL, USA, formerly Ocata Therapeutics/Advanced Cell Technology) who were the first to transplant RPE cells derived from hESC. In this trial, hESC-derived RPE cell suspensions were injected into the subretinal space of patients with advanced dry AMD, as well as patients with Stargardt’s

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disease. This was a small early-stage clinical trial involving 18 patients in total (9 with dry AMD and 9 with Stargardt’s disease) with the primary aim of assessing the safety of transplanted hESC-derived RPE (Schwartz et al., 2012). A 22-month follow-up showed no indication of adverse effects or immune reactions, providing the first evidence that hESC-derived RPE has the potential to be a safe source of cells for the treatment of AMD (Schwartz et al., 2015). Alongside this, a more recent trial by Cell Cure Neurosciences (Jerusalem, Israel) is moving into its second patient cohort of a phase I/IIa dose-escalation study. A total of 15 patients with dry AMD will be treated with injections of hESC-derived RPE (OpRegen therapy). Although no data have been released describing detailed results, this trial was supported by preclinical studies using the RCS rat model where the authors demonstrate the potential of this treatment to rescue visual acuity (unpublished preclinical work). In contrast to the bolus injection, simultaneous clinical trials have been investigating the safety of transplanting monolayers of polarized RPE, which have been prepared in vitro. This method provides an alternative to injecting RPE in suspension, and aims to replace the existing damaged RPE/basement membrane and reestablish the support and maintenance of the patients’ photoreceptor cells. Developing an RPE monolayer for transplantation requires a scaffold on which to grow the RPE and to act as an artificial basement membrane. The scaffold must, therefore, be permeable to allow for the free movement of water, nutrients, and small molecules without hindering the normal RPE function. Several ongoing clinical trials, including our own (The London Project to Cure Blindness), are investigating the use of different scaffolds and support membranes for transplanting RPE monolayers. The London Project to Cure Blindness collaborates with Pfizer (London, UK) and so far has treated 2 patients out of a planned 10 in a phase I clinical trial (Carr et al., 2013). Patients with severe exudative wet AMD were treated with a “patch” of hESC–RPE immobilized on a polyester membrane with the aim to provide a permanent replacement for the existing RPE. The RPE “patch” measures approximately 6 mm  3 mm and covers the majority of the macula. The clinical study aims to investigate the long-term survival and functionality of hESC–RPE as well as any safety issues. A similar trial is taking place in California whereby hESC–RPE cultured on an ultrathin parylene membrane will be transplanted as part of a phase I/IIa clinical trial (Lu et al., 2012). This work is led by the University of Southern California Eye Institute (CA, USA) and funded by the California Institute for Regenerative Medicine (CIRM), and involves the collaborative work of a number of different institutions. The trial aims to treat two cohorts, each of 10 patients with advanced dry AMD, and to assess the safety and tolerability of the transplanted monolayer. The scaffolds used in the above trials are nondegradable and aim to replace the existing basement membrane and facilitate in surgical delivery. However, more recently, a clinical study is being planned by the National Eye Institute and the National Institutes of Health (Bethesda, MD, USA) which will use a biodegradable scaffold made of poly-lactic-co-glycolic acid (PLGA) with the aim of providing a

4 Clinical results and considerations

temporary substrate for implantation but ultimately allowing the RPE monolayer to interact with patient’s existing Bruch’s membrane (Liu et al., 2014). The study aims to use iPSC-derived RPE in conjunction with PLGA membranes for an autologous cell therapy. This project, however, is in its early stages and performing investigational new drug studies before a clinical trial can begin (Song and Bharti, 2016). Another technique using RPE as a monolayer involves the transplantation of RPE that is cut into strips and, more importantly, is the only study involving the use of iPSC-derived RPE (Reardon and Cyranoski, 2014). This clinical study was the result of a collaboration between RIKEN and the Foundation for Biomedical Research and Innovation (FBRI; Kobe, Japan) and after being given approval by Japan’s health minister in 2013, the first patient was treated early in 2014. However, the Act on Safety of Regenerative Medicine came into effect late in 2014 leading to that particular trial being suspended until a new appropriately revised clinical study is proposed (Mandai et al., 2017). The patient underwent an autologous surgery with iPSC-derived RPE, which had been reprogrammed from the patient’s own skin fibroblasts. Similar to preclinical studies, strips of RPE measuring approximately 1.3 mm  3 mm were transplanted subretinally into the back of the patient’s eye by injection (Kamao et al., 2014). These strips were generated by culturing a monolayer of RPE on a collagen-coated membrane before enzymatically dissolving the collagen gel, leaving an RPE sheet without any artificial scaffold which can be cut into appropriately sized strips. The 1-year follow-up of the patient showed no signs of recurring neovascularization and the morphology of the macula revealed signs of improvement. Further to this, identification of tumorigenesis or other major abnormalities was not observed (http://www.riken-ibri.jp). Although initial progress and results are encouraging, there are a number of considerations to take into account with the methods of delivery under investigation. Injection of an RPE cell suspension has shown great promise in terms of safety. However, there are several issues to be considered, including cell survival, migration from the transplantation site, and the propensity of RPE cells to dedifferentiate after dissociation from a monolayer. Indeed, there is a lack of sufficient data to suggest that injected cells form a polarized monolayer in vivo (Diniz et al., 2013), and hence, the challenge remains. The use of polarized RPE monolayer on scaffolds provides an alternative method for transplantation and may overcome the need for cells to migrate to the right location, polarize, and function properly once in vivo. This RPE scaffold method, combined with advanced retinal imaging, will allow surgeons to easily observe any loss of cells from the implant. However, to develop a fully polarized RPE scaffold in culture may prolong the product development timeline and be costly, since a custom-made delivery device accommodating the RPE scaffold has to be designed and the transplantation surgery is more complicated in comparison to the simple injection method. Ultimately, the RPE on an artificial scaffold holds a better promise in terms of RPE transplant survival in vivo (Diniz et al., 2013). However, the type of substrate (e.g., biodegradable or not) used must support the growth of RPE

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monolayer, be compatible with host immune system as well as vision, and be easy to transplant (da Cruz et al., 2007). Emerging data from current clinical trials using various transplantation approaches will enable researchers to make a more informed decision on the best route forward to drive AMD therapies from the research bench to the clinic. The recent advances in technology and speed at which stem cell therapies are developing put us in an exciting era for the treatment of AMD, and indeed many other diseases. However, as a therapeutic product is being designed and developed, even at the research stage, there are a multitude of essential regulations to consider and implement before embarking on the manufacturing process.

5 PRODUCTION OF CELL THERAPIES FOR AMD 5.1 REGULATIONS Where more traditional therapeutics such as small molecules and biologics have proved ineffective, we are now increasingly looking to novel cell replacement-based approaches for treatment options for these refractory conditions. With this rapid shift in focus comes considerable challenges for regulatory bodies to ensure patients are ultimately treated with a consistent, safe, and effective cell product. Within Europe, the production of medicines and pharmaceuticals used for regenerative medicine purposes must comply with European regulations on the production of human medicines (Directive 2001/83/EC), and is overseen by the European Medicines Agency (EMA) (for a discussion of the regulatory environment in the USA, see Carpenter, 2017). In the past, conventional medicinal products have been manufactured to good manufacturing practice (GMP) guidelines. These guidelines ensure products are consistently produced and controlled according to quality standards throughout the manufacturing process, minimizing any risk to the final product (World Health Organisation (WHO), 2014). In 2004, the European Commission sanctioned the Tissues and Cells Directive (EUTCD) (Directive 2004/23/EC) which reclassified gene therapy, somatic cell therapies (including stem cell therapies), and tissue-engineered products as advanced therapy medicinal products (ATMPs) and stipulated that they should be manufactured to the same GMP standards as conventional medicinal products (Commission Directive 2003/94/EC). Safely regulating the manufacture of these ATMPs has presented a significant challenge to regulatory bodies (Pearce et al., 2014). They have had to adapt quickly to produce and implement appropriate legislation to cover their manufacture and quality assurance (Commission Directive 2009/120/EC). In the United Kingdom, the Medicines and Healthcare products Regulatory Agency (MHRA) are the competent authority regulating ATMPs and their safe manufacture to GMP standards. It is also the MHRA that authorizes their use as investigational medicinal products

5 Production of cell therapies for AMD

(IMPs) in clinical studies. Facilities hoping to manufacture IMPs must also obtain a manufacturing license for IMPs (an “MIA IMP” license) from the MHRA. Navigating the often complex regulatory pathways for taking your product into clinical trials and to market can seem daunting and tough at times. However, in light of the end goal of treating patients with safe, effective medicines, it is both necessary and appropriately stringent. Furthermore, regulatory bodies, in our experience, have been extremely helpful in facilitating this process.

5.2 PRECLINICAL CONSIDERATIONS The translation of a discovery at the research bench to production of a therapeutic product in a GMP-compliant clinical trial setting is incredibly complex. As such, it requires an extensive network of interconnecting areas of expertise (HartmannFritsch et al., 2016; Trounson and DeWitt, 2016; Whiting et al., 2015). Further to the regulatory issues mentioned previously, these span engineering, toxicology, cell biology, manufacturing, quality assurance, as well as clinical operations to name just a few (Whiting et al., 2015). These areas all require extensive early consideration and continuous attention to ensure the safety and efficacy of a candidate therapeutic. Critical to a successful cell therapy product is extensive basic research and understanding of the mechanism of action at the molecular level. In addition, characterization of the manufacturing process and of the target cell population throughout process development is essential and will facilitate the production of quality specifications for intermediate process checkpoints. This will also aid in the development of in-process quality assays used to ensure the product is within appropriate functional parameters. The establishment of these parameters will be critical in confirming the quality and lot-to-lot consistency of the final product, and will reduce the risk of shifts in cell function that could be overlooked. In this way, you can effectively anticipate suboptimal manufacturing runs in real time (Campbell et al., 2015). With any new medicinal product approaching clinical trials, safety of the patient is the primary concern. With a novel stem cell-derived ATMP such as our hESCderived RPE therapeutic, however, safety is preeminent due to the largely unchartered territory of introducing PSC-derived products into patients. Some of the main risks identified with PSC-derived products such as ours include: contaminant tumorigenic or residual hESCs in the final product that could cause tumor or teratoma, respectively; the presence of any contaminating cell types where target cell purity has not been achieved that could disrupt the therapeutic benefit of the product; or finally, the presence of any adventitious agents that could have contaminated the hESCs originating from cell banks, or human or animal-derived components (Whiting et al., 2015). Significant time, effort, and financial investment help to derisk these factors to an acceptable level, and these steps should be built into a product development timeline from the start. Again, with relevance to our manufacturing experience we will describe just a few of the actions we took to mitigate some of our perceived risks:

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•

•

•

Ensuring production of a pure therapeutic cell population. Here, we developed stringent protein-based assays to support various stages of the manufacturing process. We were able to confirm the identity and purity of our RPE cell population, through the analysis of markers such as PMEL17 (RPE marker) and TRA-1-60 (identifies residual hESCs). Analysis of chromosomal/genetic stability. It is widely known that hESCs maintained for extended time in culture can incur chromosomal changes, an effect known as adaptation (Narva et al., 2010). We implemented a strategic sampling method for regular metaphase karyology checks to detect any chromosomal abnormalities throughout the process. In vitro and in vivo validation of the safety of the manufacturing process. It is critical to be able to demonstrate safety of the product in animal models in order to achieve clinical trial authorization. We approached this in two ways: first, we spiked cell cultures at different stages of the RPE differentiation process with hESCs and showed that they were unable to survive the culture conditions (unpublished data). Additionally as a further part of manufacturing process validation, the scaffolds with hESC-derived RPE were implanted into the eyes of pigs. These studies not only confirmed safety, but also revealed evidence of rescue of retinal degeneration incurred by implantation of the control scaffold without any hESC-derived RPE (unpublished data).

5.3 MANUFACTURING Having completed the required preclinical work and successfully obtained the authorization to carry out a clinical trial, manufacturing of the ATMP can begin. This must be performed within a GMP-compliant cleanroom, which must hold an IMP MIA license as mentioned previously. Manufacturing under GMP conditions brings with it its own set of challenges when compared to working in a conventional research laboratory. The cleanroom environment is precisely controlled to ensure levels of particulates and microbial contamination are kept to an appropriate minimum level. Different areas are graded on their stipulated particle limits (European Union, 2003) with most products, including production of our RPE cells, being carried out in Grade A. Grade A is considered the most critical area where the product can be open to the environment during processing and as such has the lowest particle limits which are 3520 and 20 for 0.5- and 5-mm particles, both in use and at rest, respectively (MHRA). These levels are controlled by a sophisticated air-handling unit that maintains temperature, humidity, and differential pressures between the rooms. This produces a positive pressure gradient throughout the facility, which effectively removes particles or contaminants with the outward flow of air (Sheth-Shah et al., 2016). To ensure manufacturing adheres to the strict regulations and guidelines set out by the directives mentioned previously, a quality management system (QMS) is developed by each facility, which ensures product specification and quality is

5 Production of cell therapies for AMD

maintained throughout the manufacturing process. This is achieved through an integrated system of activities involving planning, training, implementation, documentation, assessment, and improvement to ensure that the product is fit for purpose (Sheth-Shah et al., 2016). We will highlight here just two key quality assurance considerations with respect to our manufacturing experience that will affect all manufacturing processes: •

•

Validations: these are required for all aspects of manufacturing, from validating the function of the cleanroom itself, to equipment, reagents, and processes to ensure the item or process is fit for purpose and that the particular parameter is functioning within specified limits that ensure product safety and function. For example, following the installation of a new incubator within the facility, a validation was carried out before it could be used to house cells. In the case of an incubator, the critical factor is temperature, and so temperature mapping was carried out for an appropriate length of time to ensure all regions were maintained within predetermined temperature set points. Service-level agreements: it is imperative for the quality assurance of the final clinical-grade product that there is total, traceable knowledge of the quality of the materials and reagents used throughout the manufacturing process. In the case of RPE differentiation media, prior to purchasing the item, an audit of the provider was carried out by the quality manager to ensure they were satisfied with the quality and sourcing of their products. Once this was confirmed, an agreement was put in place to ensure the requirements of both parties were maintained and the quality and specification of the media was consistent every time. Quality certificates such as Certificates of Analysis are included with each delivery of a material or reagent and can be matched up against original agreed specifications. Once the item has been determined to be of the appropriate quality, it is released from quarantine and can be used within the facility.

As highlighted in the examples given here, manufacturing within a cleanroom, with its associated activities and considerations, requires considerably more time and effort than producing the equivalent product in the research laboratory. Adding to these pressures, the manufacturing timelines of PSC-derived therapies are often relatively long compared to conventional medicines. This is usually due to either a considerable expansion phase, depending on the required clinical dosage, or an extended differentiation phase depending on the maturity of the therapeutic cells. In the case of our product, the manufacturing timeline is a minimum of 7 months but can be longer due to the flexibility built into the process (Fig. 3A). This is a long period of continual processing, and inevitably issues can arise. These can take the form of contamination events or, for example, an unplanned facility shutdown. The employment of the QMS should minimize the likelihood of these events occurring due to continual training of operators and preventative facility and equipment maintenance checks. However, it is clear that breakpoints in these extended protocols would be desirable. To this end, we would advise the inclusion of a cryopreservation

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FIG. 3 Generation and characterization of stem cell-derived retinal pigment epithelium (RPE) for research and the manufacture of an advanced therapeutic medicinal product (ATMP) to treat age-related macular degeneration (AMD). (A) Schematic representation and comparison of the RPE derivation process from iPSC and hESC in research and in GMP. (B) Derivation of iPSCs from fibroblasts and initial identification of stem cell colonies—(i) merging iPSC colonies, (ii) higher magnification of the merging colonies, (iii) confirmation of pluripotency using StainAlive TRA-1-60 marker, (iv) early passage images of the isolated clonal lines, (v and vi) confirmation of pluripotency using StainAlive TRA-1-60 and TRA-1-81 markers on the expanded lines, and (vii) normal karyotype of derived iPSC. (C) Confirmation of pluripotency in higher passage stem cells. (D) Confirmation of RPE polarity by

5 Production of cell therapies for AMD

step(s) at appropriate stages of the process. Being able to cryopreserve cells would additionally facilitate more efficient distribution of product to other facilities for multicenter trials, as well as reduce cost of goods during manufacture where cells could be thawed as and when required instead of ongoing culture and possible loss of material in the event a patient is not recruited in time. Evidence of the benefits of a step such as this can be observed in the Astellas AMD clinical trial mentioned earlier in the chapter. Here, the differentiated RPE suspension is cryopreserved and the product then transported to various clinics, where it is thawed, washed, and reconstituted before being injected into the patient’s eye (Schwartz et al., 2012). However, to obtain regulatory approval, we would advise introducing cryopreservation reagents into the system as far away from the final dosing of the product as possible. In this way, there is adequate time to ensure that thawed cell populations are of the required specification and pass in-process quality checks in advance of administrating the product to patients. As is demonstrated in this discussion, the translation of a cell-replacement therapy for clinical usage is multifaceted and not without its challenges. This is only amplified when the cell product is derived from PSCs, as demonstrated currently by the relatively fewer PSC trials in the clinic. It is certainly feasible though, as long as product development is approached in a logical manner, with a comprehensive understanding and characterization of the process and therapeutic cell population, and with the appropriate expertise and experience available. Fig. 3 depicts our manufacturing timeline for production of a scaffold covered with RPE cells from either iPSCs or hESCs. The PSCs are expanded prior to their spontaneous differentiation into RPE. After approximately 6 weeks, pigmented foci can be purified by manual dissection and seeded to encourage monolayer formation. The comparative timings and culture requirements of these stages when carried out in research (blue) or in the cleanroom (black) are indicated to highlight the differences. The main disparity being that in research there are minimal constraints to timings or materials used and activity is very much down to operator discretion, within reason. Manufacturing in the cleanroom, however, requires operators to strictly follow standard operating procedures and production protocols that have been agreed with the quality assurance manager, and from which users cannot deviate. Production

immunostaining. (E) Confocal immunofluorescent micrographs depicting staining for typical regional RPE markers as part of the characterization of the monolayer as RPE. (F) Bright-field images depicting the chronological stages of RPE manufacture: (i) stem cell differentiation, (ii) cobblestone morphology, (iii) RPE on a transwell culture, and (iv) cellular therapeutic “patch.” (G) Internalization of POS by control stem cell-derived RPE cells. (H) ScoreCard assay confirming trilineage differentiation potential and self-renewal of control stem cell-derived RPE. (I) Quantification of PEDF secretion in spent culture medium during late expansion phase using an ELISA assay. The classical RPE PEDF secretion asymptote occurs around 3–6 weeks. (J) Assessment of control stem cell-derived RPE barrier function using transepithelial volt/ohm meter.

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protocols can depict anything from the cell substrate used, tissue culture vessel size, to number of times a trituration movement can be made to resuspend a cell pellet with a pipette. It is, however, crucial that flexibility is built into manufacturing production protocols at certain stages when dealing with living cells. It could be that the operator feels the cells need to expand for another day before transferring them to the next stage of the process. If the protocol had stipulated that this action must occur after an exact number of days, any deviation from this would initiate a chain of quality control alerts including risk analyses to establish if there is any risk to the product. Instead, during the process validation of each step, we would establish production “windows” in which safety and efficacy of the product is maintained, as specified by the well-developed in-process quality assays and characterization. Second, Fig. 3 provides examples of the detailed characterization profiles that our research group has established for the RPE and its functional capabilities. Using various functional (e.g., transepithelial resistance measurements, electrophysiology, phagocytosis, pigment epithelium-derived factor (PEDF) secretion assays) and characterization assays (e.g., immunocytochemistry, PCR, Western blot, pluripotency assays) have previously shown that the spontaneously differentiated control stem cell-derived RPE is consistently capable of recapitulating the native RPE phenotype as well as function in vitro and in animal models (Carr et al., 2009a,b; Carter et al., 2016; Vugler et al., 2008). It is this comprehensive knowledge of the therapeutic cells and how they function throughout production that has allowed us to develop a robust, highly characterized manufacturing process, with sufficient in-built quality assurance that gives us, and the regulators, confidence that our cell-replacement therapy is safe to trial in patients.

6 FUTURE The eye as indicated has a number of advantages in determining whether cell therapies for disease will be safe and efficacious. Equally, the pathway from bench to the clinic, especially regulatory, has led to a number of first-in-human studies for eye disease which have progressed the field of regenerative medicine as a whole. Safety concerns using hESC-derived therapeutics have been tested and shown to be safe in a number of clinical trials over an extended period of time. The initial trials were to ensure safety and were in patients with loss of central vision (end-stage disease). Subsequent trials are now exploring how effective such transplants are in saving vision.

ACKNOWLEDGMENTS This work was supported by The London Project to Cure Blindness.

References

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Transplantation of reprogrammed neurons for improved recovery after stroke

10

Zaal Kokaia1, Daniel Tornero, Olle Lindvall Laboratory of Stem Cells and Restorative Neurology, Lund Stem Cell Center, Lund, Sweden 1 Corresponding author: Tel.: +46-462220276; Fax: +46-462220560, e-mail address: [email protected]

Abstract Somatic cells such as fibroblasts, reprogrammed to induced pluripotent stem cells, can be used to generate neural stem/progenitor cells or neuroblasts for transplantation. In this review, we summarize recent studies demonstrating that when grafted intracerebrally in animal models of stroke, reprogrammed neurons improve function, probably by several different mechanisms, e.g., trophic actions, modulation of inflammation, promotion of angiogenesis, cellular and synaptic plasticity, and neuroprotection. In our own work, we have shown that human skinderived reprogrammed neurons, fated to cortical progeny, integrate in stroke-injured neuronal network and form functional afferent synapses with host neurons, responding to peripheral sensory stimulation. However, whether neuronal replacement plays a role for the improvement of sensory, motor, and cognitive deficits after transplantation of reprogrammed neurons is still unclear. We conclude that further preclinical studies are needed to understand the therapeutic potential of grafted reprogrammed neurons and to define a road map for their clinical translation in stroke.

Keywords Fibroblasts, Reprogramming, Stroke, Regeneration, Functional recovery, Brain repair

1 INTRODUCTION Stroke is the second most common cause of death worldwide and a major cause of disability (Donnan et al., 2008). In ischemic stroke, occlusion of a cerebral artery due to thromboembolism causes focal brain ischemia, cell death and sensory, motor, and cognitive impairments. More than half of patients suffer significant residual deficits, causing huge economic and societal burden. Apart from thrombolysis Progress in Brain Research, Volume 231, ISSN 0079-6123, http://dx.doi.org/10.1016/bs.pbr.2016.11.013 © 2017 Elsevier B.V. All rights reserved.

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and thrombectomy during the first hours after an ischemic stroke, which can be given to only a fraction of patients, no effective treatment to improve functional recovery exists in the postischemic phase. Therefore, efforts at bringing the latest advances in science and technology a step closer to the development of therapies against stroke should be vigorously promoted. It should be emphasized that some degree of spontaneous recovery occurs in virtually all patients surviving stroke but varies from modest improvement to almost complete restoration. This reflects the extremely complex nature of poststroke recovery which involves plastic changes in surviving neurons and neurons on the contralateral side, redistribution of brain representation, release of growth factors and antiinflammatory factors from immune cells, synaptogenesis and changes in synaptic strength, changes in dendritic arborization and spines, as well as generation of new neurons, and glial and endothelial cells from endogenous stem cells (Moskowitz et al., 2010). Research has previously focused on the acute and subacute phases after stroke, assuming that the therapeutic window for rescuing neurons from secondary degeneration is limited to this period. However, it is now clear that the recovery phase is longer, allowing for the application of treatments that affect the environment and promote repair processes, e.g., modulate brain plasticity and replace injured or dead cells (Dihne et al., 2011; Lindvall and Kokaia, 2010). Stem cell-based approaches hold much promise as potential novel therapies to restore brain function after ischemic stroke (George and Steinberg, 2015). Transplantation of various types of stem cells or their progeny has been reported to improve behavioral impairments in animal models of stroke (Lindvall and Kokaia, 2011). Several possible mechanisms underlying these improvements have been proposed such as neuronal replacement, modulation of inflammatory responses, trophic action, and stimulation of plasticity in host brain neuronal circuitries (Kokaia et al., orklund, 2017). Ongoing or completed clinical trials using in2012; Dunnett and Bj€ tracerebral, intravenous, or intraarterial transplantation of mesenchymal or bone marrow-derived stem cells provide evidence for the safety of these treatments and also suggest some clinical improvements (Lee et al., 2010; Moniche et al., 2015; San Roman et al., 2015; Steinberg et al., 2016; Taguchi et al., 2015), but the underlying mechanisms are unclear. When delivered systemically in animal models, these cells infiltrate the damaged brain in limited numbers and hardly engraft in the brain parenchyma or transdifferentiate to neurons (reviewed in Doeppner and Hermann, 2010). Possibly, these cells act by modulating the inflammatory environment, stimulating endogenous neurogenesis and angiogenesis, and reducing the formation of the glial scar. Recently, the outcome of a Phase 1 clinical trial using allogeneic transplantation of a human fetal brain-derived immortalized neural stem cell line was reported (Kalladka et al., 2016). The trial demonstrated feasibility and safety of intracerebral transplantation of up to 20 million cells in stroke patients with no immunological or cell-related adverse events and also suggested some neurological improvements. In our view, the ultimate goal for stem cell research in stroke, which would give rise to optimum, long-term recovery, should be to provide the stroke-injured brain with new cellular elements, including specific neurons for reconstruction of

2 Improving functional recovery in stroke

damaged neural circuitry. It is inconceivable that stroke patients, who are mainly elderly people, would exhibit major improvements due to just plastic changes in surviving neurons, especially because such processes are limited in the aging brain. Neuronal replacement has been shown to be feasible in the diseased human brain. Thus, clinical trials in Parkinson’s patients have provided proof of principle for the cell replacement strategy by demonstrating that transplanted neurons can survive for many years, become integrated in host circuitry, and improve motor function (Lindvall, 2015). Reprogramming of somatic cells (Takahashi and Yamanaka, 2006) seems particularly attractive for the generation of neurons for cell replacement in stroke. This strategy allows for the production of patient-specific cells, without the need for immunosuppression after autologous transplantation, and also avoids the ethical issues associated with the use of human embryonic stem cells. In this chapter, we summarize recent data supporting the idea that transplantation of reprogrammed neurons to the stroke-injured brain can lead to functional improvements. We consider possible underlying mechanisms and present evidence that these neurons can become integrated into host neuronal circuitries. Finally, we discuss the further development of this research and how it can be moved toward the clinic in a responsible way.

2 IMPROVING FUNCTIONAL RECOVERY IN STROKE BY TRANSPLANTATION OF REPROGRAMMED NEURONS Somatic cells such as fibroblasts can be reprogrammed to pluripotent stem cells by introduction of transcription factors (Fig. 1; Takahashi and Yamanaka, 2006). These induced pluripotent stem cells (iPSCs) can then be transplanted into the injured brain, either directly or after they have been fated to neural progenitor cells (NPCs) or to specific neuron types (Fig. 1). Fibroblasts can also used to produce induced neural stem cells (iNSCs), avoiding the pluripotent stage. Similar to embryonic stem cells, iPSCs can generate tumors (Kawai et al., 2010), but if iPSCs are fated to neural phenotype, tumorigenicity is virtually abolished. From a clinical perspective, it is important to note that whether iPSC-derived neurons can express the complete functional phenotype of the neurons they are going to replace has been unclear. In recent years, several studies have demonstrated the feasibility of the reprogramming approach to produce cells for transplantation in experimental stroke models. Mouse (Chau et al., 2014; Chen et al., 2010; Kawai et al., 2010; Liu et al., 2014; Qin et al., 2015; Yamashita et al., 2016; Yao et al., 2015) and human (Chang et al., 2013; Eckert et al., 2015; Jensen et al., 2013; Jiang et al., 2011; Lam et al., 2014; Mohamad et al., 2013; Oki et al., 2012; Tatarishvili et al., 2014; Tornero et al., 2013; Yuan et al., 2013) skin-derived iPSCs have been implanted in mouse (Chen et al., 2010; Eckert et al., 2015; Kawai et al., 2010; Lam et al., 2014; Liu et al., 2014; Mohamad et al., 2013; Oki et al., 2012; Yamashita et al., 2016) and rat (Chang et al., 2013; Chau et al., 2014; Jensen et al., 2013; Jiang et al., 2011; Qin et al., 2015; Tatarishvili et al., 2014; Tornero et al., 2013; Yao

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Oct4, Sox2, Klf4, c-myc

Intracerebral transplantation Induced neural stem cells (iNSCs)

Expansion/ modification Neural progenitor cells (NPCs) Somatic (skin) cells/ fibroblasts

Stroke-subjected rodent

Induced pluripotent stem cells (iPSCs) BMP4, Wnt3A, cyclopamine Cortical Long-term progenitors neuroepithelial-like stem cells (lt-NESCs)

FIG. 1 Current approaches for reprogramming somatic cells for application in stroke models. (i) Skin-derived fibroblasts are reprogrammed to induced pluripotent stem cells (iPSCs) or induced neural stem cells (iNSCs), which are transplanted directly into stroke-injured brain. (ii) Through expansion and modification, the iPSCs are transformed to neural progenitor cells (NPCs) or long-term neuroepithelial-like stem cells (lt-NESCs), which are then transplanted into stroke-injured brain. (iii) Using BMP4, Wnt3A, and cyclopamine, the lt-NESCs are first fated to cortical progenitors in vitro and then transplanted into stroke-injured brain.

et al., 2015; Yuan et al., 2013) brains subjected to different types of ischemic stroke (Table 1). The reprogrammed cells have been transplanted directly as iPSCs or, more often, after they have been predifferentiated or transformed into neural lineage cell lines such as NPCs or long-term neuroepithelial-like stem cells (lt-NESCs). One of these studies detected hypercellularity and rosette formation (Jensen et al., 2013), and two other studies described tumorigenicity with high c-myc activity (Kawai et al., 2010) and formation of tumors in all iPSC-grafted animals (Chen et al., 2010). However, in the other studies, iPSCs directly transplanted in stroke-injured brain did not cause tumor formation (Jiang et al., 2011; Qin et al., 2015), which suggests that factors other than pluripotency also influence tumorigenicity. The relative safety of reprogrammed cells generated from lt-NESCs and iNSCs is supported by findings that their implantation in immune-deficient mice and rats revealed no transplant overgrowth or tumor formation (Tornero et al., 2013; Yao et al., 2015). In our study (Tornero et al., 2013), we detected a threefold increase in the number of cells in the graft at 2 months after transplantation of iPSC-derived lt-NESCs in the cortex of

Table 1 Design and Results of Studies Using Intracerebral Transplantation of Stem/Progenitor Cells Derived by Reprogramming of Somatic Cells in Rodent Stroke Models Implantation: When/Where/ Number of Cells

Improvements After Transplantation

Possible Mechanisms of Improvements

1. Rotarod 2. Grasping power test/ 1, 2, and 4 weeks

1. Decreased latency @1, 2, and 4 weeks 2. Increased grasping power @1, 2, and 4 weeks

Decreased IL-1b, IL-2, IL-6, TNFa, iNOS mRNA; Increased IL-4, IL-10 mRNA in ipsilateral hemisphere

Chen et al. (2010)

Yes. @2 and 4 weeks, high c-myc level

1. mNSS 2. Rotarod/ 1 day, 2 and 4 weeks

No

ND

Kawai et al. (2010)

GFAP+, nestin +, vimentin + cells, NQ

No

Cylinder test/4, 8, 1, and 16 days

Decreased curve of score

ND

Jiang et al. (2011)

HuNu+ cells, 10%/11 weeks

HuD+ (79%) and DCX+ (13%) cells

No

1. Staircase 2. Corridor/ 2, 5, and 9 weeks

1. Improvements @9 weeks 2. No improvements

Increased VEGF immunoreactivity in ipsilateral hemisphere

Oki et al. (2012)

ND

hNu + and hMito+ cells, NQ/8 weeks

Nestin +, MAP2+, NeuN +, DARPP32 +, GAD65/67+, TH +, GFAP+, O4+ cells, NQ

No

1. Rotarod 2. Stepping 3. mNSS 4. Aporotation/ weekly from 0 to 8 weeks

1. Improvements @2–8 weeks 2. Improvements @1–8 weeks 3. Improvements @2–8 weeks 4. Improvements @2 and 4 weeks

1. TUNEL+ cells decreased 2. Decreased inflammation (less Iba1+, ED1 +, GFAP+ cells) 3. Increased neurogenesis (more BrdU +, DCX+, PSANCAM+ cells) in ipsilateral hemisphere

Chang et al. (2013)

ND

HuNu+ cells, about 500,000/5 weeks

b-Tubulin + and MAP2+ (41%), GFAP+ (5%), MBP+ (0%), nestin + (32%) cells

No; hypercellularity, rosettes (50% of rats). Ki67 + cells (13%)

1. Elevated body swing 2. Cylinder 3. Adhesive removal/1, 3, and 5 weeks

No

ND

Jensen et al. (2013)

Survival/ Time After Stroke

Phenotype of Cells

1. Decreased @1 week 2. Decreased @1 week

ND

ND

100% with pure iPSCs, less with glue

1 day after MCAO/ipsi striatum and cortex/ 500,000 cells

ND

GFP+ cells, NQ/2 weeks

DCX+, NeuN +, Nanog + cells/NQ

MCAO/70 min

1 week after MCAO/ipsi striatum/ 800,000 cells

Decreased @4 weeks

GFP+ cells, NQ/4 weeks

Mouse/ C57BL/ male/ adult

MCAO/30 min

1 week after MCAO/ipsi striatum/ 100,000 cells

No

iPSCNPCs

Rat/SD/ male/ adult

MCAO/90 min

1 week after MCAO/contra striatum/ 200,000 cells

iPSCNPCs

Rat/ Wistar/ male/ adult

MCAO/30 min

1 week after MCAO/ipsi cortex/ 250,000 cells

Species/ Strain/ Sex/Age

Cell Origin

Cell Type

Stroke Model/ Duration

Mouse fibroblasts

1. Pure iPSCs 2. iPSCs mixed with fibrin glue

Rat/Long Evans/ male/ adult

MCAO/60 min

1. Just before MCAO/ipsi cortex/ 1,000,000 cells 2. Immediately after MCAO/ ipsi subdural/ 1,000,000 cells

Mouse fibroblasts

iPSCsNPCs (hydrogel coated)

Mouse/ C57BL/ 6N/male/ adult

MCAO/30 min

Human fibroblasts

iPSCs

Rat/SD/ female/ adult

Human fibroblasts

iPSC-ltNESCs

Human fibroblasts

Human fibroblasts

Behavioral Test: Which/ When After Stroke

Effect on Infarct Size

Tumorigenicity

References

Continued

Table 1 Design and Results of Studies Using Intracerebral Transplantation of Stem/Progenitor Cells Derived by Reprogramming of Somatic Cells in Rodent Stroke Models—cont’d Species/ Strain/ Sex/Age

Implantation: When/Where/ Number of Cells

Improvements After Transplantation

Possible Mechanisms of Improvements

Adhesive removal/1, 2, 3, and 4 weeks

Improvement @2 and 3 weeks, not @4 weeks

Increased BDNF mRNA in penumbra of the stroke

Mohamad et al. (2013)

No

1. Cylinder 2. Stepping/ 1, 4, and 8 weeks

1. No Improvements 2. Improvements @8 weeks

ND

Tornero et al. (2013)

Nestin +, b-tubulin +, GFAP+ cells, NQ

ND

1. Rope grabbing 2. Beam walking 3. Morris water maze/ 1, 2, and 3 weeks

Improvement @3 weeks

ND

Yuan et al. (2013)

mCherry+ cells, NQ/2 weeks

NeuN+, neurofilament +, GFAP+ cells, NQ

ND

Vibrissaeelicited forelimb placement/ 3 weeks

Improvements @3 weeks

Increased SDF-1 and VEGF protein levels @2 days after transplantation Increased neurogenesis and angiogenesis in periinfarct area

Chau et al. (2014)

ND

GFP+ cells, NQ/2 weeks

Sox2+, DCX+, NF200 cells, NQ

ND

ND

ND

ND

Lam et al. (2014)

During MCAO/ ipsi cortex/ 1,000,000 cells

ND

Bisbenzimide+ cells, NQ/3 weeks

Tuj1 +, NeuN+, GFAP+ cells, NQ

No

1. Locomotor activity 2. Beam walking 3. Rotarod/ 1, 2, and 3 weeks

1. Improved @2 and 3 weeks 2. Improved @1, 2, and 3 weeks 3. Improved @2 and 3 weeks

ND

Liu et al. (2014)

2 days after MCAO/ipsi cortex/ 300,000 cells

Decreased

HuNu+ cells 49%/8 weeks

HuD + (91%), DCX+ (30%), TBR1 + (3%), GFAP+ (1%) cells

No

1. Cylinder 2. Stepping/ 1, 4, and 7 weeks

1. Improvements @4 and 7 weeks 2. No improvements

Increased VEGF immunoreactivity and decreased inflammation in ipsilateral hemisphere

Tatarishvili et al. (2014)

Effect on Infarct Size

Survival/ Time After Stroke

Phenotype of Cells

Tumorigenicity

Hoechst33342 + cells, NQ; 15% TUNEL + cells/4 weeks

NeuN+ cells (26%)

No

No

HuNu+ cells 270%/8 weeks

Fox3+ (13%), DCX+ (77%), GABA + (20%) cells

Immediately after MCAO/ ipsi striatum/ 1,000,000 cells

ND

CM-Dil + cells, NQ/1 and 2 weeks

MCAO/ permanent

7 days after MCAO/ipsi cortex/ 400,000 cells

ND

Mouse/ C57BL/ 6/male/ adult

Photothrombotic/ permanent

1 week after stroke/ipsi cortex/ 200,000 cells

iPSCNPCs

Mouse/ C57BL/ 6/male/ adult

MCAO/120 min

iPSC-ltNESCs

Rat/SD/ male/ aged

Distal MCAO/ 30 min

Stroke Model/ Duration

Behavioral Test: Which/ When After Stroke

Cell Origin

Cell Type

Human fibroblasts

iPSCNPCs

Mouse/ C57BL/ 6/male/ adult

MCAO/ permanent

1 week after MCAO/ipsi cortex/ 400,000 cells

ND

Human fibroblasts

iPSC-ltNESCs (cortically fated)

Rat/ nude/ male/ adult

Distal MCAO/ 30 min

2 days after MCAO/ipsi cortex/ 300,000 cells

Human fibroblasts

iPSCNPCs

Rat/SD/ male/ adult

MCAO/120 min

Mouse fibroblasts

iPSCNPCs

Rat/ Wistar/ neonate (P7)

Human fibroblasts

iPSCNPCs (hydrogel coated)

Mouse fibroblasts

Human fibroblasts

References

Human fibroblasts

iPSCNPCs

Mouse/ C57BL/ 6J/male/ adult

MCAO/60 min

1 days after MCAO/ipsi hippocampus/ 100,000 cells

No

STEM121+ cells, NQ/30 days

Nestin + cells (90%), Tuj1 +, and S100b+ cells (small %)

No

1. Adhesive removal 2. Beam walk 3. Rotarod/ 2, 4, 6, 8, 12, 16, 20, and 24 days

1. Improvements @2, 4, 6, and 8 days 2. Improvements @2, 4, 6, and 8 days 3. Improvements @ all days

Reduced TNFa, IL-6, IL-1b mRNA, and MCP-1 and MIP-1a mRNA; decreased MMP-9 activity; decreased number of Iba1+ microglia; ameliorated BBB damage in ipsilateral hemisphere

Eckert et al. (2015)

Mouse fibroblasts

iPSCs

Rat/SD/ male/ adult

Intracerebral hemorrhage/ collagenase VII injection into striatum

6 h after stroke/ipsi striatum/ 1,000,000 cells

ND

CM-Dil+ cells, NQ/6 weeks

GFAP+ cells, NQ

No

Limb placement/ 1 and 3 days and 1, 2, 4, and 6 weeks

Improvements @2, 4, and 6 weeks

Decreased MPO+ neutrophils and CD11b+ microglia @3 days; decreased IL-1b, IL-6, TNFa protein, and mRNA; Increased IL-10 mRNA. Decrease of glial scar (GFAP +) and apoptosis (Caspase3+) in perihematomal area

Qin et al. (2015)

Mouse fibroblasts

iNSCs

Rat/SD/ male/ adult

MCAO/ permanent

2 days after MCAO/ipsi cortex/ 1,000,000 cells

Decreased @2 weeks

GFP+ cells, NQ/2 weeks

GFAP+ cells (@2 weeks), Tuj1+ cells (@6 weeks)

No

1. mNSS 2. Adhesive removal/2, 4, 7, 10, and 14 days

Improvements @4, 7, 10, and 14 days

Change in metabolism status in ipsilateral hemisphere

Yao et al. (2015)

Mouse fibroblasts

iNSCs

Mouse/ C57BL/ 6N/male/ adult

MCAO/30 min

1 day after MCAO/ipsi striatum and cortex/ 500,000 cells

No; increased animal survival

GFP+ cells, NQ/4 weeks

GFAP+ (90%), O4+ (3%) cells, No Tuj1+, DCX+, MAP2+ cells

No

1. mNSS 2. Rotarod 3. Corner/1 day, 1, 2, 3, and 4 weeks

1. Improvements @1 week 2. No improvements 3. Improvements @2 weeks

ND

Yamashita et al. (2016)

Apo-rotation, apomorphine-induced rotation; BBB, blood–brain barrier; iNSCs, induced neural stem cells; iPSCs, induced pluripotent stem cells; lt-NESCs, long-term neuroepithelial-like stem cells; MCAO, middle cerebral artery occlusion; mNSS, modified neurological severity score; ND, not determined; NPCs, neural progenitor cells; NQ, not quantified; SD, Sprague-Dawley.

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CHAPTER 10 Reprogrammed neurons in stroke

rats subjected to stroke. At this time point, the cellular proliferation rate in these cortically fated grafts was about 10% and we did not detect any tumors or cellular overgrowth. Our findings indicate that the initial proliferation of the reprogrammed cells might counteract the cell death after implantation, thereby maintaining a high number of cells in the transplants. However, if this is the case, gradual spontaneous downregulation of proliferative activity will be necessary in order to avoid tumor formation and tissue overgrowth. The effect of intracerebral transplantation of reprogrammed, skin-derived iPSCs, iPSC-derived NPCs, iPSC-derived lt-NESCs, and iNSCs on behavioral recovery after stroke has been evaluated in a large number of studies (Table 1). Multiple tests for the assessment of motor, sensory, and cognitive functions have been used at different time points from 1 day up to 9 weeks after stroke. Most studies have used two or more behavioral tests. Interestingly, in the two studies which did not report any improvement (Jensen et al., 2013; Kawai et al., 2010), transplantation of human and mouse fibroblast-derived iPSC-NSCs gave rise to hypercellularity and rosette formation (Jensen et al., 2013) or tumor formation (Kawai et al., 2010), respectively. It is conceivable that this brain pathology had interfered with any beneficial effect of reprogrammed cells on functional recovery. Numerous studies have shown, although with different dynamics, improvement in all performed behavioral tests, e.g., rotarod, stepping, modified neurological severity scores (mNSS) and apomorphine-induced rotation (Chang et al., 2013), rope grabbing, beam walking and Morris water maze (Yuan et al., 2013), mNSS and adhesive removal (Yao et al., 2015). In other studies, only some of the applied tests showed improvements (Oki et al., 2012; Tatarishvili et al., 2014; Tornero et al., 2013; Yamashita et al., 2016). These data indicate that grafted reprogrammed cells might have differential effects on the restoration of specific types of impaired function. In future studies, it will be important to use behavioral tests to target specific brain functions, and also to discriminate between impairments, which can spontaneously recover over time or remain permanent. Data from many studies indicate that the grafted reprogrammed cells induce functional improvement in stroke models by mechanisms other than neuronal replacement. In most of these studies, the beneficial effect of transplantation was first observed within 1–2 weeks, i.e., before any functional neurons could have developed from the grafted reprogrammed cells. It is inconceivable that this early improvement is due to neuronal replacement. In support of this interpretation, we found that the enhancement of behavioral recovery early after transplantation of iPSC-derived lt-NESCs was observed irrespective of degree of graft survival and generation of neurons at later time points (Oki et al., 2012; Tatarishvili et al., 2014; Tornero et al., 2013). It is well established that cellular and synaptic plasticity in the injured brain is heavily involved in regenerative processes accompanying poststroke recovery (for review, see Wieloch and Nikolich, 2006). Although the underlying mechanisms have not been clarified in detail, available data indicate that the beneficial effects on behavior seen early after transplantation are due to direct or indirect influences on these poststroke processes mediated, e.g., by trophic action,

2 Improving functional recovery in stroke

neuroprotection, modulation of inflammation, and stimulation of angiogenesis. In support, using mouse-derived iPSCs, Chen et al. (2010) demonstrated decreased levels of the proinflammatory cytokines IL-1b, IL-2, IL-6, TNFa, and iNOS mRNA concomitant with increased level of the antiinflammatory cytokine IL-4 and increased expression of IL-10 mRNA in the hemisphere ipsilateral to stroke. Similarly, decreased inflammation was observed when human-derived iPSCs were transplanted in stroke-damaged brain (Chang et al., 2013; Tatarishvili et al., 2014). Chang et al. (2013) reported promotion of neurogenesis in subventricular zone, as evidenced by increased number of BrdU +, DCX +, and PSA-NCAM+ cells, and Mohamad et al. (2013) described increased BDNF mRNA levels in the stroke-injured brain of mice grafted with iPSC-NPCs. In our own studies, we observed that transplanted reprogrammed iPSC-derived lt-NESCs expressed high levels of VEGF immunoreactivity and gave rise to increased VEGF immunoreactivity in host brain astrocytes (Oki et al., 2012; Tatarishvili et al., 2014). Secretion of VEGF might be important for suppression of inflammation and neovascularization in the periinfarct region, leading to improved recovery. Interestingly, iPSC-derived lt-NESCs cells have been shown to survive transplantation also into the stroke-damaged brain of aged rats. More than 90% of reprogrammed cells differentiated to mature neurons expressing the marker HuD. Moreover, a population of transplanted cells (19%) expressed GABA (Tatarishvili et al., 2014). Since the majority of stroke patients are elderly people, the ability of reprogrammed somatic cells to give rise to neurons also in the aged brain tissue environment is important from a translational perspective. Several studies have attempted to characterize the phenotype of the reprogrammed cells after implantation into the stroke-injured brain. When human and mouse iPSCs were used directly without any priming or fating, they generated GFAP +, nestin +, and vimentin + cells (Jiang et al., 2011) or only GFAP+ (Qin et al., 2015) cells, respectively. However, in virtually all other studies using either iPSC-derived NPCs (Chang et al., 2013; Eckert et al., 2015; Jensen et al., 2013; Liu et al., 2014; Mohamad et al., 2013; Yuan et al., 2013) or iPSC-derived lt-NESCs (Oki et al., 2012; Tatarishvili et al., 2014; Tornero et al., 2013), cells expressing either early (Eckert et al., 2015; Lam et al., 2014; Yao et al., 2015; Yuan et al., 2013) or mature (Chau et al., 2014; Kawai et al., 2010; Liu et al., 2014; Oki et al., 2012; Tatarishvili et al., 2014; Tornero et al., 2013; Yamashita et al., 2016) neuronal markers were observed. We have explored whether reprogrammed cells give rise to functional neurons after transplantation. We showed that human iPSC-derived lt-NESCs (Falk et al., 2012), implanted into the stroke-injured, adult rodent brain (Oki et al., 2012), stop to proliferate, and survive for many months. The vast majority of grafted cells had become mature neurons at 11 weeks after transplantation, and some of them sent efferent projections to appropriate brain structures such as globus pallidus. Using patch-clamp recordings on acute brain slices, we investigated the electrophysiological properties of grafted cells at 5 months after transplantation and showed that they had the capacity to generate action potentials and to receive afferent inputs.

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In virtually all studies with intracerebral transplantation of reprogrammed cells in stroke-damaged brain performed so far, no efforts have been made to fate the differentiation of the grafted cells toward a specific neuronal phenotype. Importantly, in our clinical study (Kokaia et al., 2012; Delavaran et al., 2013), we found that predominantly subcortical lesions are rare in stroke patients and tend to give only mild neurological deficits. Therefore, we believe that if the neuronal replacement strategy should be translated to the clinic, it should primarily aim for patients with cortical and/or combined cortical–subcortical lesions. Consequently, generation of cortical neurons and reconstruction of cortical neuronal networks are major issues for the development of cell-based therapies in stroke. We have fated iPSC-derived lt-NESCs to cortical neuronal progenitors in culture and then transplanted them into strokeinjured cortex (Tornero et al., 2013). The cortically fated cells survived, differentiated to functional neurons, and improved neurological outcome in stroke-subjected nude rats (Tornero et al., 2013). In support of their cortical phenotype, we also demonstrated pyramidal morphology of the grafted, fated cells and expression of the cortex-specific marker TBR1 in a certain layered pattern. Patch-clamp recordings in acute brain slices obtained at 5 months after transplantation demonstrated their neuronal maturity and functionality.

3 EVIDENCE FOR RECONSTRUCTION OF NEURONAL CIRCUITRY AFTER IMPLANTATION OF REPROGRAMMED CELLS IN STROKE-INJURED BRAIN Although functional integration of grafted reprogrammed neurons in injured host neural circuitry will most likely lead to optimum functional recovery after stroke, evidence that neuronal replacement really occurs is virtually lacking. It requires the formation not only of electrophysiologically and morphologically competent neurons but also of efferent projections to the appropriate targets and establishment of functional synaptic inputs on the grafted neurons from host brain. We have previously shown that intracortically grafted neurons, generated from human iPSC-derived lt-NESCs and differentiated to a cortical phenotype, provide efferent projections to cortical and subcortical areas and exhibit spontaneous excitatory postsynaptic currents (Tornero et al., 2013). The only evidence that the grafted reprogrammed cells can project axons to the appropriate target structure comes from our study (Oki et al., 2012), showing that injection of retrograde tracer in globus pallidus leads to labeling of iPSC-derived lt-NESCs grafted in stroke-injured striatum. Direct experimental data demonstrating that transplanted stem cell- or reprogrammed cell-derived neurons receive synaptic inputs from stroke-injured host brain are scarce. Neurons, derived from transplanted mouse or human NSCs or human ES cells, have been shown to be surrounded by structures expressing synaptic markers and to exhibit spontaneous excitatory postsynaptic currents (Buhnemann et al., 2006; Daadi et al., 2009). Some ultrastructural data indicate synaptic connections between

4 Direct in vitro and in vivo reprogramming of somatic cells to neurons

host axon terminals and grafted neurons (Daadi et al., 2009; Muneton-Gomez et al., 2012). We have shown that stimulation of the intact cortical region adjacent to the transplant triggered monosynaptic evoked responses from the intracortically grafted neurons, generated from human iPSC-derived lt-NESCs, which indirectly suggested that the host neurons had established afferent synaptic contacts on the grafted cells (Tornero et al., 2013). We have now demonstrated (Tornero et al., 2017) for the first time that the host brain regulates the activity of grafted, cortically fated human lt-NESC-derived neurons, providing strong evidence that these transplanted cortical neurons can become incorporated into injured cortical circuitry. In this study, we used the rabies virus-based trans-synaptic tracing method and showed that the grafted neurons receive direct synaptic inputs from neurons in different host brain areas located in a pattern similar to that of neurons projecting to the corresponding endogenous cortical neurons in the intact brain. Immunoelectron microscopy confirmed that neurons of stroke-injured host brain establish excitatory axodendritic synaptic contacts with the grafted human cortical neurons. Electrophysiological in vivo recordings from the cortical implants showed that mechanical stimulation of nose and paw evokes spikes or inhibits spontaneous activity in the grafted neurons, indicating that some of the afferent inputs to the these neurons are functional. Confirming the functionality of the synaptic inputs, using patch-clamp recordings, we showed that the grafted neurons respond to photostimulation of virally transfected, channelrhodopsin-2-expressing thalamocortical axons in acute brain slices. Although our data clearly demonstrate that the reprogrammed grafted neurons can become part of injured host neuronal circuitry, it remains to be demonstrated that this repair mechanism contributes to long-term amelioration of functional impairments after stroke.

4 DIRECT IN VITRO AND IN VIVO REPROGRAMMING OF SOMATIC CELLS TO NEURONS In addition to neurons derived from iPSCs, reprogrammed neurons suitable for transplantation into stroke-injured brain could potentially also be produced by direct in vitro conversion of somatic cells. Functional neurons, so-called induced neurons (iNs), can be generated by directly reprogramming mouse somatic cells (Fig. 1; Vierbuchen et al., 2010). This conversion does not occur through a pluripotent stem cell stage and thereby eliminates the risk for tumor formation. Using forced expression of lineage-specific transcription factors that act during brain development, specific subtypes of neurons, including dopaminergic (Caiazzo et al., 2011; Pfisterer et al., 2011), spinal motor (Son et al., 2011), cholinergic (Liu et al., 2013), and striatal medium spiny neurons (Victor et al., 2014), have been generated directly from human fibroblasts. For the use of directly converted cells in stroke, it has to be shown that specific neuronal subtypes can be produced and survive transplantation after this insult. So far, generation of cortical neurons from fibroblasts or other somatic cells by direct in vitro conversion has not been reported. Moreover, there is no published

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study using transplantation of iNs in stroke-subjected animals. Thus, it is presently unclear whether this approach can become a viable strategy for functional restoration in stroke. Another strategy to replace neurons in the stroke-injured brain could be by direct in vivo reprogramming. A number of studies have demonstrated the neurogenic potential of glia (Heins et al., 2002; Magnusson et al., 2014) and the ability to reprogram astrocytes, NG2 glia, and pericytes into neurons in vitro (Berninger et al., 2007; Guo et al., 2014; Karow et al., 2012). These studies open up the possibility to directly reprogram glia to neurons within the brain parenchyma. Zhang and colleagues have developed a strategy to convert resident astrocytes to proliferative NPCs and functionally mature neurons in the adult brain and spinal cord (Niu et al., 2013, 2015; Su et al., 2014). When the transcription factor NeuroD1 was ectopically expressed in reactive astrocytes at stab injury sites in adult mouse cortex, they could be directly reprogrammed to functional neurons (Guo et al., 2014). However, it is not clear whether these neurons expressed a specific cortical phenotype or could functionally integrate in injured neuronal networks and contribute to repair. A shortcoming of the in vivo reprogrammed neurons reported to date is that they do not adopt the same subtype-specific phenotype as observed in parallel in vitro experiments. Thus, unlike direct neural conversion in vitro, it has so far not been possible to direct the formation of dopaminergic neurons via direct conversion in vivo (Torper et al., 2015). To our knowledge, no study has so far addressed the issue of direct in vivo reprogramming of cortical glia cells to cortical neurons. Moreover, nothing is known about the possibilities to reprogram brain cells to specific neuron types in strokedamaged animals, nor whether this approach can be used to promote recovery of impaired motor, sensory, and cognitive functions. These major challenges need to be addressed in future studies.

5 RESEARCH CHALLENGES AND PROSPECTS FOR CLINICAL TRANSLATION Several studies in animal models have demonstrated that transplantation of reprogrammed cells can reverse functional deficits in ischemic stroke without tumorigenicity or other side effects. Although these studies suggest a therapeutic potential of reprogrammed cells in stroke, available data are definitely not sufficient to move this new strategy to clinical application. Further advancements in both experimental and clinical research are needed. Much better understanding of the mechanisms by which transplanted, reprogrammed cells exert their recovery-promoting action is required for choosing the optimum cell, for inducing maximum recovery, and for selecting the most suitable stroke patients based on the location and size of their ischemic lesion. Most often, grafts of nonsorted reprogrammed cells are heterogeneous and include neurons of several phenotypes at different stages of differentiation as well as astrocytes and undifferentiated cells. Which cell types are important for functional

5 Research challenges and prospects for clinical translation

recovery needs to be clarified. Moreover, the efficient reconstruction of strokeinjured neuronal networks will require generation of neurons with specific phenotypes, and we must also identify mechanisms affecting their integration into host neural circuitries. In addition, how much the “bystander effect” of grafted reprogrammed cells, mediated through release of trophic factors or immunomodulation, stimulation of cell and angiogenesis, and other plasticity-promoting events, contributes to behavioral improvements needs to be characterized in detail. Understanding the dynamics of these various processes during the recovery phase might be crucial for optimization of the functional restoration after stroke. It should be pointed out, though, that the properties and mode of action over time of intracerebrally grafted, reprogrammed cells in animal models of stroke may only partly reflect how these cells will behave in patients. Since stroke patients are in most cases elderly people, age-related events and comorbid conditions such as diabetes, atherosclerosis, hypertension, or dementia might influence not only the fate of the transplanted cells but also their ability to exert the recovery-promoting effect observed in animal models (Wechsler, 2009). Therefore, data obtained in animal models that do not mimic the pathology of human stroke may not translate into meaningful effects in a clinical setting. For successful clinical translation, it will be necessary to demonstrate in animal models that grafted reprogrammed cells induce substantial improvement of functional deficits that resemble the debilitating symptoms in stroke patients. Many experimental studies have reported improved behavioral outcome after stroke using tests which do not reflect the clinical situation and have a strong learning component (Hicks et al., 2009). Behavioral tests that assess somatosensory, motor, and cognitive impairments should be used for quantifying possible beneficial effects. Moreover, tests assessing permanent as well as transient functional impairments of different severity should be used to determine the ability of the grafted cells to either promote spontaneous recovery or restore functions that are not spontaneously improving. One fundamental research question is which stroke patients could potentially benefit from transplantation of neurons generated by reprogramming somatic cells. As mentioned earlier, comorbid diseases and overall health status of aged patients will probably be important factors for the patient’s suitability for such a therapeutic approach. Size and location of the ischemic lesion, as determined by noninvasive imaging, as well as magnitude of somatosensory, motor, and cognitive impairment and prognosis for potential spontaneous recovery will be crucial for determining of type and number cells as well as location of the transplant. After implantation, it will be important to be able to trace the grafted cells and monitor their survival, migration, function, and effect in the patient’s brain. Several studies have demonstrated feasibility of magnetic resonance imaging, bioluminescence imaging, and positron emission tomography for tracing transplanted cells in vivo (Daadi et al., 2009; Guzman et al., 2007; Hoehn et al., 2005; Love et al., 2007; Modo et al., 2004; Ramos-Cabrer et al., 2010). Unfortunately, the current resolution of these techniques does not allow tracing single or even few cells. Development of new approaches, probably based on the combination of several imaging techniques

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for monitoring with high resolution the proliferation, long-term survival, and functionality of intracerebrally grafted cells in the stroke-damaged brain, is highly warranted. Surprisingly, it has still not been convincingly demonstrated whether neuronal replacement and integration in injured circuitry of grafted stem cells or reprogrammed cells contribute to the recovery of impaired functions following stroke. To clarify this issue, several fundamental issues regarding the reconstruction of stroke-damaged neuronal networks need to be addressed. First, it has to be shown not only whether the grafted cells can receive functional afferent inputs from the host brain but also if they send functional efferent projections to the appropriate brain structures. Second, it should be analyzed if host oligodendrocytes can myelinate the axons of grafted neurons, which will be important for the functionality of the efferent projections. Third, and most importantly, it has to be determined whether integration of reprogrammed neurons and their neuronal function contribute to sensory, motor, and cognitive recovery after stroke. Optogenetic approaches, which can be used to inhibit or stimulate the activity of grafted neurons at different stages of poststroke recovery while animals are performing various behavioral tasks, will be instrumental in determining the mechanisms underlying functional recovery and the significance of integration of grafted cells in host neural circuitry (Carter et al., 2012). If neuronal replacement using reprogrammed cells is shown to be a viable strategy for brain repair and functional restoration in stroke, it will be important in the next step to develop protocols for fating the reprogrammed cells toward different neuronal subtypes, including various cortical projection neurons and interneurons (Gaspard et al., 2008), and for producing the neuronal progenitors for intracerebral transplantation in large numbers.

6 CONCLUSIONS The results from studies in animal models are promising and provide supportive evidence that the use of reprogrammed cell-based approaches has a potential to be developed into a new, clinically useful strategy to promote recovery after stroke. However, there is no fast track for reprogrammed cells to reach the stage of clinical application. Only with a long-term commitment to high-quality basic and clinical research, which addresses all the crucial issues, some of which are discussed earlier, will it eventually be possible to offer reprogrammed cell-based treatments to stroke patients, leading to substantial improvements of their quality of life.

ACKNOWLEDGMENTS Our own work was supported by the Swedish Research Council, EU FP7 grant TargetBraIn, S€oderberg Foundation, and Swedish Brain Foundation.

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Combined Index The index covers both Parts A and Part B of Functional Neural Transplantation IV, Progress in Brain Research Volumes 230 and 231, respectively. The relevant part is indicated by ‘A:’ and ‘B:’, as appropriate.

A A2B5 expression, B: 137, 167, 168, 172 ABCA4 gene, B: 194 Aberrant splicing, B: 212 Abortion, A: 171, 228, 229, 239, 247, 267. B: 34, 37 Accelerometer, A: 318 Acetylcholine, A: 2, 17–22, 31, 57, 87, 105. B: 255 Acetylcholinesterase (AChE), A: 17, 19, 21, 31 Activin, A: 139, 195, 215, 271, 272, 275. B: 204 Activities of daily living, A: 315, 316, 322. B: 195, 226 Addenbrooke’s Cognitive Examination, A: 183 Adeno-associated virus (AAV), A: 62, 83, 86, 100, 102, 103, 108, 115. B: 45, 194, 195 Adenovirus, A: 102, 103 Adenylate cyclase, B: 119 Adenylyl cyclase, B: 110, 114, 115 Adhesive removal, B: 249–252 Adipocytes, B: 4, 40 Adipose (fat) tissue, A: 287. B: 119 Adipose (fat) tissue stem cells, A: 101. B: 3, 111 Adrenal grafts, A: 8, 9, 167. B: 88 Adrenal medulla, A: 3, 8, 9, 167, 193. B: 88 Adrenoleukodystrophy, B: 170 Adult neurogenesis, A: 74, 273, 308. B: 7, 246, 249 Adult stem cells, A: 102, 151. B: 135–153, 202 Advanced Cell Technologies, A: 137, 159–161. B: 231 Advanced disease, A: 170, 182, 233. B: 210 Advanced therapeutic medicinal product (ATMP), A: 173. B: 124, 234–236, 238 Adventitious agents, A: 156, 158, 160, 161. B: 235 Adverse effects. See Adverse events Adverse events, A: 75, 76, 78, 88, 100, 101, 106–108, 114–116, 134, 142, 167, 169, 170, 184, 214, 220, 228, 232, 234–237, 241, 244, 248, 254, 286, 298–300. B: 88, 126, 169, 177, 246 Aerobic exercise, A: 319–321 Aetiology, A: 105, 107, 110, 268, 312. B: 74, 87, 100, 178, 193, 209, 211, 226 Age, A: xxvii, 62, 73, 105, 137, 166, 168, 182, 232, 235, 238, 244, 247, 266, 275, 316. B: xxvii, 65–67, 70, 100, 137, 140, 142–144, 147, 149, 174–176, 180, 193, 226–228, 257 Age of onset, A: 112, 232, 314, 316. B: 193 Age related macular degeneration, B: 193, 198, 200, 225–240

Aged hosts, A: 2, 9, 220, 315. B: 73, 141, 142, 227, 247, 253, 257 Age-related macular degeneration (AMD), A: xxvii, 34, 137, 159. B: 191–194, 225–227, 238. B, xxvii Aggregate suspension culture, A: 216, 220. B: 203–206 Aggression, A: 236 Aipl1 mouse, B: 211 Akt/mTOR pathway, B: 146 Alemtuzumab, B: 175 Alexander’s disease, B: 170, 171 Alkaline phosphatase, B: 12 Allodynia, B: 21, 73, 88, 90–92, 97 Allografts, A: 116, 152, 156, 167, 168, 175, 213–215, 221, 222, 228, 266. B: 165, 168, 169, 171, 174, 182, 208, 210, 229, 246 Alloimmunisation, A: 228, 238, 241, 242, 249, 253 All-trans retinal re-isomerisation, B: 227 Alzheimer’s disease, A: 63, 99, 105. B: 58, 67, 72, 73, 166. See also Dementia Amacrine cells, B: 195 Amblyopia, B: 66, 67, 69 4-Aminopyridine (4-AP), B: 71 Amniotic fluid stem cells, A: 101. B: 3 AMPA receptors, A: 310. B: 145 Amphetamine, A: 13, 22, 30, 173, 179, 181, 200 Amygdala, A: 10, 12, 22, 26. B: 60, 61, 69 Amyloid, A: 63, 74, 105. B: 73 b-Amyloid. See Amyloid Ab protein Amyloid Ab protein, A: 63, 105, 106. B: 73 Amyloid plaques, A: 74, 105, 106. B: 73 Amyotrophic lateral sclerosis (ALS), A: 34, 63, 99, 103, 104, 109–112. B: 165, 166, 178 Anaerobic glycolysis, A: 84, 289 Anaesthesia, A: 248. B: 88 Analgesia, B: 88, 90, 93, 97 Angiogenesis, A: 101. B: 39, 42, 46, 245, 246, 253, 257 Animal models, A: 8–10, 32, 34, 76, 78, 102, 106, 108–112, 116, 117, 135, 136, 141, 167, 173, 178, 179, 181, 191–205, 213, 228, 246, 247, 264, 265, 276, 283, 286, 292, 306, 319. B: 7, 17, 34, 47, 67, 69, 70, 72, 73, 90, 111, 150, 167, 172, 181, 229, 245, 246, 256–258 Animal retroviruses, A: 156. B: 208

265

266

Combined Index

Anterograde degeneration, A: 13 Anterograde tracing, A: 12, 17, 23, 27. B: 12, 17, 255 Antibody rHlgM22, B: 152 Antibody vx15-2503, B: 153 Anticonvulsant drugs, B: 89 Antidepressant drugs, B: 69 Antiepileptic drugs, B: 71 Antihistamine drugs, B: 88, 150 Antimitotic agents, B: 113, 115, 122 Antimuscarinic drugs, B: 150 Antioxidants, A: 8, 108, 113 Antipsychotic drugs, A: 252 Antisense morpholinos, B: 212 Anxiety, B: 67, 69 Apical microvilli, B: 227, 228 ApoE4 knock-in mouse, B: 73 Apolipoprotein ApoE4 allele, B: 73 Apomorphine, A: 166. B: 249, 252 Apoptosis, A: 235, 249, 289, 290. B: 35, 65, 151, 206 Aromatic amino acid decarboxylase (AADC), A: 292, 295, 299 Artifact, B: 4, 12 Arylsulfatase, B: 171 Ascl1, A: 56, 57, 60, 61, 71–73, 83, 85–87, 89, 270, 271. B: 9 Astellas Pharma, A: 137. B: 231, 239 Asterias, A: 137, 151, 152, 159–161. B: 11 Astrocytes, A: 30, 59–63, 70, 73, 75–79, 81–85, 87, 88, 100–102, 107, 108, 140, 194, 195, 235, 290, 292, 294. B: 3, 5, 9, 10, 35–39, 41, 42, 44, 46, 137, 143, 144, 165–82, 253, 256 Atherosclerosis, B: 257 Atrial natriuretic peptide (ANP), A: 216 Atrophy, A: 100, 112, 228, 233–235, 241, 247, 248, 288, 291. B: 38, 150, 194, 228 Atropine, A: 18 Australia, A: 159, 161. B: 87 Autism, B: 58 Autoantibodies, B: 150, 151 Autocrine signalling, A: 269. B: 47, 116 Autoimmune disorders, A: 9. B: 139, 140, 175 Autologous grafts, A: 29, 99–103, 106, 111, 116, 117, 140, 152–154, 161, 214, 220, 221, 232, 249, 267, 273. B: 8, 14, 40, 107–109, 111–114, 116, 123–127, 168–170, 208, 229–231, 233, 247 Autonomic function, A: 166. B: 1, 72 Autosomal dominant disorders, B: 194 Availability, A: 33, 172, 239, 248, 267, 319. B: 112 Axin2, B: 145 Axon growth, A: 1, 22–27, 30, 76, 167, 168, 180, 192, 197, 202, 205, 222, 308. B: 1–22, 33–48, 109, 113, 117, 119, 127, 144, 148

Axon guidance, A: 6, 8, 27, 30. B: 21, 144 Axon regeneration, B: 2, 33–48, 107–128 Axons, A: xxvi, 2, 4, 6–8, 13, 14, 18, 20–31, 61, 62, 75, 77, 88, 89, 110, 285, 289, 308. B: xxvi, 1–3, 8, 10–21, 33–47, 72, 98, 108–119, 123, 135, 136, 139, 140, 143–145, 151, 166, 172, 173, 175, 177, 192, 254, 258 Azathioprine, A: 172, 184, 204, 239

B B11B061, B: 153 Baclofen, B: 90 Bands of Bungner, B: 109 Basal ganglia, A: 3, 7, 10, 11, 13, 72, 108, 134, 228, 263, 286, 306, 317, 322. B: 72. See also Striatum; Caudate nucleus; Putamen; Globus Pallidus; Substantia nigra Basal lamina, A: 27. B: 204 Basiliximab, A: 204 Basso mouse scale (BMS), B: 38, 39 Batten disease, A: 103 B-cells, A: 233. B: 151 BCL11B. See CTIP2 Bcl-2, A: 84, 142 Beam walking test, B: 250–252 Beck Depression Inventory, A: 183 Behavioural rescue, B: 69 Belgium, A: 227 Benzodiazepine receptor, A: 296 Benztropine, B: 150 Best medical therapy, A: 182 Bexarotene, B: 149, 153 Bhlhb5, B: 98–100 Bicuculline, B: 94, 95 BIIB033, B: 148 Biodegradable materials, B: 19, 232, 233 Biodistribution, A: 157, 158, 160, 179, 200 Bioluminescence, A: 115. B: 257 Biomarkers, A: 204, 244, 290, 292, 297. B: 210 Biopsy, A: 29, 72, 75, 275, 284. B: 109, 127 Bioreactor, B: 127, 207 Biotin, B: 152 Biotinylated dextran amine (BDA), A: 23. B: 18 Bipolar cells, B: 195–197, 200, 211 bis-Retinoid N-retinyl-N-retinylidene ethanolamine (A2E), B: 227 Blastocyst, A: 134, 137, 143, 161, 268. B: 5, 7, 37, 201, 202 Blastomere, A: 137, 161 Blind assessment, A: 243, 244, 254 Blindness, B: 191–212, 225–240

Combined Index

Blood vessels, A: 105, 241. B: 3, 117, 119, 193, 226, 228 Blood-brain barrier, A: 7, 75, 88, 101, 105, 172, 204, 248, 249. B: 147, 151 Blood-derived stem cells, B: 201 Bone bank, B: 122 Bone marrow stem cell grafts, A: 109, 111, 215. B: 1–4, 40, 111, 199, 246 Bone marrow stem cells, A: 101, 109, 111, 215. B: 1–4, 40, 111, 199, 246 Bone morphogenetic protein (BMP), A: 61. B: 6, 37, 248 Bone tissue, A: 101. B: 122 Bovine spongiform encephalopathy (BSE), A: 158, 175, 176 Bovine viruses. See Animal retroviruses Brain bank, A: 182 Brain morphogenic protein (BMP), A: 24, 61, 139, 195, 215. B: 145, 205, 248 Brain repair, A: 1–34, 69–89, 307, 311. B: 74, 192, 245–258 Brain-derived neurotrophic factor (BDNF), A: 8, 80, 81, 85, 106, 108, 110–113, 249, 276, 311–313. B: 3, 11, 18, 20, 35, 41, 45, 46, 90, 113, 253 Brainstem, A: 18, 23, 24, 31. B: 3, 13, 16, 94, 124, 172. See also Midbrain; Hindbrain Bromodeoxyuridine (BrdU), A: 85. B: 249, 253 Bridge grafts, A: 6, 7, 26–32. B: 1, 2, 9, 17, 19–21, 33–39, 42–48, 107–128 Brn2, A: 56, 57, 61, 73, 83, 89. B: 9 Bruch’s membrane, B: 193, 226, 228, 233 Bupivacaine, A: 111 Bystander effect, A: 6, 9. B: 171, 257

C C3Hrd1/rd1 mouse, B: 197 C9orf72, A: 110 CA1, A: 81. B: 61, 71 Cajal Retzius cells, B: 65 Calbindin, A: 22 Calretinin, B: 60 cAMP, A: 57. B: 3, 108, 110–115, 119 Canada, A: 101, 214 Canavan’s disease, B: 170 Cancer, A: 34, 104, 141, 142. B: 40, 88, 91, 137, 149 Cancer therapy, A: 104. B: 149 Cancerous cells, A: 34, 141, 173 CANTAB, A: 236 CANTAB test, A: 236 CAPIT-HD, A: 228, 232, 234, 235, 251–253, 315 Capsaicin, B: 94

Cardiac disease, A: 9, 159, 161, 201 Cardiomyocytes, A: 56 Carnegie stage, A: 266 Carotid grafts, A: 167, 193 Caspase 3, B: 251 Catechol-o-methyl-transferase (COMT), A: 248 b-Catenin, A: 139. B: 145, 146 Cats, B: 176 Caudal ganglionic eminence (CGE), B: 59–64, 68, 89 Caudalisation, A: 196. B: 8, 18, 203 Caudate nucleus, A: 4, 22, 107, 109, 232, 234, 235, 237, 239, 241 CDn surface markers. See also Human Leukocyte Antigens (HLA) CD3. A: 233 CD4 T cells, A: 235, 250 CD8 T cells, A: 235, 250 CD9, B: 168 CD11b, B: 40, 251 CD14 T cells, B: 40 CD15, B: 210 CD19 T cells, B: 40 CD20, A: 233 CD24, B: 210 CD34, A: 138. B: 40 CD44, B: 168, 179, 180 CD45, B: 40 CD47, B: 210 CD68, A: 233, 296, 300. B: 249 CD73, B: 40, 210 CD79a, B: 40 CD90, B: 40 CD105, B: 40 CD133, B: 6, 210 CD140a, B: 168, 172 Cell adhesion molecules, A: 27. B: 41, 42, 144 Cell autonomous effects, B: 46, 178 Cell bank, A: 100, 172, 178, 197, 204. B: 207, 235 Cell Cure Neurosciences, A: 159–161. B: 232 Cell cycle, A: 77. B: 111, 137, 203 Cell fate, A: xxv, xxvi, 57–59, 71, 79, 85, 87, 274, 291, 299. B: xxv, xxvi, 35, 41, 60, 63, 64, 137, 247, 248, 258 Cell lineage, A: 57, 59, 60, 63, 76, 85, 141, 142, 192, 193, 195, 196, 200, 215, 216, 266, 268, 273–275. B: 4, 5, 8, 37, 39, 40, 64, 109–112, 139, 146, 148, 149, 166, 169, 170, 201, 202, 238, 248, 254, 255 Cell lysis, B: 115 Cell manufacturing facility, A: 154, 197, 199, 201, 205. B: 121–125, 127, 236, 237 Cell phenotype, B: 240, 254

267

268

Combined Index

Cell product, A: xxvi, 100, 104, 133, 135–137, 140–142, 144, 145, 152, 155, 157, 158, 161, 162, 165, 173–181, 192, 197–201. B: xxvi, 126, 234, 235, 239, 240 Cell proliferation, A: 33, 76, 77, 79, 82, 84, 181, 194, 198, 218, 220, 275, 283, 289. B: 9, 107, 108, 110, 111, 114, 116, 118, 120, 151, 252, 258 Cell settling, A: 247 Cell sorting, A: 140, 205, 213–222. B: 6, 8, 166–168, 172, 196, 198 Cell source, A: xxv, xxvi, 2, 22, 29, 30, 32, 33, 53, 69–71, 73, 99–102, 133, 134, 136, 139, 153, 165, 167, 172–175, 178, 179, 191, 193, 195, 200, 205, 214, 229, 246, 263, 267, 268, 275, 276, 306. B: xxv, xxvi, 1–3, 5, 20, 37, 39, 40, 108, 111, 121, 147, 167, 168, 170, 191, 201, 202, 208–210, 225, 229–231 Cell spiking, A: 201. B: 236 Cell suspension culture, B: 205 Cell suspension grafts, A: 11, 19, 172, 178, 229, 235, 236, 242, 247, 287, 291, 306. B: 5, 12, 120, 121, 196, 200, 231–233 Cell therapy, A: xxv–xxvii, 133–137, 139–146, 167, 173, 193–207, 215–224, 229–257, 265–278, 285–302, 307, 308, 318, 324. B: xxv–xxvii, 7, 33–48, 87–101, 169, 170, 174, 175, 179, 181, 182, 192, 196, 202, 207, 209, 210, 229, 230, 234, 235, 239, 240, 247 CellTracker CM-DiI, B: 251 Center for Biologics Evaluation and Research, A: 153 Central vision, B: 226, 240 CERE-120, A: 108 Cerebellum, A: 3, 20, 306. B: 150, 172 Cerebral palsy, B: 165, 170 Cerebrospinal fluid (CSF), A: 101, 240, 249, 287. B: 89, 93, 151 c-Fos, A: 13, 14. B: 20, 94 Chandelier cells, B: 64 Channelrhodopsin, B: 94, 95, 255 Chemotherapy, B: 74, 91, 92, 100 Childhood disorders. See Paediatric disorders Chimeric mice, B: 177–180 China, A: 104. B: 108 Chloride gradient, B: 90 Cholecystokinin, A: 14 Cholera toxin, B: 114–121 Choline, A: 288–290, 292 Choline acetyltransferase (ChAT), A: 61. B: 16 Chondrocytes, B: 4, 40 Chondroitin sulphate proteoglycan (CSPG), B: 19, 42, 47, 144 Chondroitinase, A: 218. B: 19, 47, 48, 113, 144

Chorea, A: 232, 236, 248 Choroid cyst, A: 228, 246 Choroid plexus, A: 239, 246, 247. B: 193, 226 Chromaffin cells, A: 8, 193. B: 88 Chromatin, A: 56–58, 143, 273 Chromosomal abnormalities, B: 122 Chromosomal spreads. See Karyotyping Chromosomal stability, A: 133–146 Chromosomes, A: 103, 142 Chromosome 12, A: 142 Chromosome 17, A: 142 Chromosome 19, A: 103 Chromosome 20, A: 143 Chronic pain, B: 67, 87–89, 93, 100, 101 Cilia, B: 212 Ciliary body, B: 202 Ciliary neurotrophic factor (CNTF), A: 253. B: 19, 41 Ciliary transport, B: 212 Ciliogenesis, B: 206, 212 Circuit reconstruction, A: xxvi, 1, 3, 4, 6, 10–13, 27, 36, 264, 267, 294, 312, 314, 319, 321, 322. B: xxvi, 1–22, 33–48, 74, 107–128, 247, 255, 258 Cleanroom, B: 236, 237, 239 Clemastine, B: 150 Clinch token transfer test (C3T), A: 317–319 Clinical assessment, A: 181, 182, 184, 192, 228, 232, 240, 241, 247, 252, 315–318. B: 123–126, 237, 252 Clinical grade cells, A: 62, 103, 111, 117, 133, 135–139, 142–144, 161, 221. B: 119–127, 237. See also Good manufacturing practice Clinical Laboratory Improvement Amendments (CLIA), A: 155 Clinical trials, A: 26, 29, 32–34, 83, 99, 100, 102–104, 106, 108–113, 134, 135, 137, 141, 151, 152, 155–158, 160–162, 165–185, 191–205, 213–222, 227–255, 283, 292, 314, 317, 321, 322. B: 4, 7, 11, 41, 88, 107–128, 147–150, 152, 153, 171, 177, 195, 199, 207, 209–212, 225, 230–236, 239, 246, 247 Clobetasole, B: 150 c-Myc. See Myc proto-oncogene Cobalt chloride, B: 211 Cobblestone morphology, B: 226, 238 Co-culture, A: 27, 61, 156–158, 194, 217, 269, 270. B: 108, 148, 205 Cognitive assessment, A: xxvi, 252, 316, 317 Cognitive deficits, A: 2, 5, 182, 232, 238, 251, 266, 286, 306–308, 315, 316. B: 69, 70, 73, 180, 182, 245, 257

Combined Index

Cognitive function, A: 3, 106, 113, 233, 234, 236–238, 251, 317, 319. B: 72, 178, 180, 252, 256, 258 Cognitive training, A: 305–322 Co-grafts, A: 8. B: 19, 42 Collagen, A: 75. B: 42, 114, 116, 120, 233 Collagenase, B: 115, 117–120 Collagenase VII lesion, B: 251 Collateral sprouting, A: 7, 308 Colour vision, B: 198 Community setting, A: 320 Comorbid conditions, B: 149, 257 Compensation, A: xxvi, 89, 307 Complement, B: 115, 194, 226, 227 Complete Freund’s Adjuvant (CFA), B: 141 Conduction velocity, B: 95, 172 Cone dystrophy, B: 194 Cones, B: 193–195, 198, 199, 207, 211 Congenital hypomyelination, B: 139, 167, 172, 181 Contactin, B: 42 Contamination, A: 140, 141, 155, 158, 170, 178, 180, 181, 198, 200, 201, 215, 216, 239, 247. B: 9, 115, 119, 126, 235–237 Controls, A: 26, 75, 112, 117, 172, 201, 202, 220, 242, 244, 245, 290, 309, 313, 321. B: 4, 19, 20, 35–37, 39, 42, 44, 45, 71, 88, 98–100, 125, 126, 143, 145, 179, 180, 200 Contusion injury, B: 5, 6, 10, 43, 45, 112, 113, 124, 177 Copy number variations (CNVs), A: 54, 143 Cord blood stem cells, A: 101, 139. B: 3, 172 Core assessment protocol for intracerebral transplantation in HD. See CAPIT-HD Corin, A: 141, 213, 216–218 Corpus callosum, B: 62 Corridor test, B: 249 Cortex, A: 2, 14, 21, 23–26, 57, 59, 60, 73, 74, 76, 77, 79, 81, 82, 84–88, 104, 237, 238, 263, 265, 269, 271, 286, 306, 310, 312. B: 41, 45, 57–60, 62, 63, 65, 66, 69–71, 89, 90, 93, 248, 254, 256 Cortical grafts, A: 17–21. B: 57–74, 245–258 Cortical Interneurons, B: 57–74 Cortical lamina, A: 12, 19, 24. B: 64, 65, 254 Cortical phenotype, B: 256 Cortical silent period (CSP), A: 253 Corticospinal tract (CST), A: 29. B: 17–20, 41 Corticostriatal pathway, A: 5, 12–17, 286, 311, 314 Cranial nerves, B: 172 Creatine, A: 289, 290 CREB1, A: 57 Cre-Lox inducible expression, A: 83, 86. B: 66, 94, 145

Cresyl violet, A: 293 CRISPR/Cas9, A: 53–63, 89, 100, 103, 104, 114, 274. B: 195, 208 Critical period, B: 66–69, 74 Cross-sectional studies, A: 251 Crown-rump length, A: 247 Cryopreservation, A: 133–145, 152, 156, 177–179, 191–205, 266, 267, 274. B: 237, 239 CTIP2, A: 24, 270–272, 274, 275 Culture (2D / monolayer), A: 244, 268. B: 6, 201, 205, 206, 210, 226, 230, 239 Culture (3D), B: 202–204, 210 Culture medium, A: 17, 55, 140, 197, 198, 217, 269, 273, 311. B: 37, 38, 91, 98, 109, 110, 114–116, 118–121, 125, 126, 203, 205, 230, 237, 238 Cuprizone, B: 141, 148, 150 Cyclosporine, A: 168, 172, 184, 204, 232, 236, 237, 239, 243 Cylinder test, A: 196. B: 249, 250 Cynomolgous monkeys, A: 217 Cyst, A: 228, 232–234, 241, 246. B: 109, 113 Cytokines, A: 78, 101, 108, 113, 296. B: 6, 10, 34, 35, 46, 99, 194, 253 Cytoplasmic material transfer, B: 199, 210 Cytosine arabinoside, B: 115

D DARPP-32, A: 11, 14, 33, 82, 87, 217, 270–272, 274, 275, 309 Database of Genomic Variants, A: 142 Databases, A: 142, 239 Daylight vision, B: 198 Dedifferentiation, B: 41, 109 Deep brain stimulation, A: 6, 7, 134, 166, 182, 183, 203 Delayed alternation, A: 11, 14, 21, 266 Delayed onset, A: 308. B: 229 Delayed start design, A: 239, 244 Dementia, A: 105, 171, 182, 234. B: 257. See also Alzheimer’s disease Demyelinating disease, B: 135–153, 165–182 Demyelination, A: 9. B: 11, 135–153, 165–182 Dendrites, A: 16, 23, 87, 116, 310, 319. B: 16, 73, 93, 95, 96, 197, 246, 255 Dendritic spines, A: 13, 16, 310. B: 246 Denmark, B: 165 Dentate gyrus, A: 18, 19, 85. B: 72, 73 Depolarisation block, A: 7 Depression, A: 25, 232, 236, 246 Dermal tissue stem cells, A: 101

269

270

Combined Index

Dermatitis, B: 100 Designer receptors exclusively activated by designer drugs (DREADDs), A: 17, 114, 205, 218 Desomorphine, A: 195 Development. See Embryonic development Developmental gradients, A: 269 Diabetes, A: 2, 139, 159, 161. B: 36, 174, 175, 257 Diet, B: 226 Dietary restriction, B: 147 Differentiation, A: xxv, xxvi, 7, 8, 33, 34, 53, 54, 56, 58, 60, 61, 63, 72, 84, 85, 133, 135, 139, 140, 145, 153, 155, 160, 173, 175–177, 181, 184, 191–205, 213–222, 236, 263–276, 283–286, 290, 292–294, 299, 313. B: xxv, xxvi, 3–7, 9–11, 13, 14, 17, 19, 21, 34, 35, 38, 40, 44, 46, 57, 72, 100, 111, 112, 114, 142–146, 148–151, 167, 170, 173, 192, 201–210, 230, 236–239, 254, 256 Dimethylfumarate, B: 175 Diphtheria toxin, B: 36, 45, 66 Direct conversion, A: 56, 60. B: 20, 255–258 Disease duration, A: 171, 182, 237 Disease modelling, A: 53–63, 73, 192, 275. B: 192, 201, 202, 208, 209, 211 Disease modification, A: 110. B: 58, 67, 87, 100, 149, 151 Disease progression, A: 1, 105, 107, 110, 115, 228, 236, 246, 251, 254, 288, 295, 306, 315. B: 179, 180, 193, 194, 225, 230 Disinhibition, B: 70, 73 Dispase, B: 117–119 Dispersal, A: 10. B: 58, 59, 61, 63, 68, 72–74, 166, 171, 172, 176, 177 Dissection, A: 265 Dissociation, A: 154. B: 111, 115–121, 196, 233 Distribution, A: 247. B: 208 DKK1, A: 270. B: 203–205 Dlx2, A: 72, 73, 271–274 DNA, A: 54, 79, 85, 86, 100, 143, 144, 272. B: 115, 195 Domperidone, B: 153 Donald Hebb, A: 307 Donor age, A: 71, 168. B: 201 Donor eligibility requirements, A: 155, 156, 158, 174 Dopamine (DA), A: 2–5, 8, 12–17, 20–23, 25, 26, 30, 32, 34, 60, 69, 71, 73, 83, 87, 107–109, 114–116, 133–145, 158, 165–185, 191–205, 213–222, 233, 235, 236, 252, 263, 264, 269, 273, 286, 292–296, 299. B: 13, 72, 255, 256

Dopamine D1 receptors, A: 294, 295 Dopamine D2 receptors, A: 60, 234, 236–238, 252, 293–295, 297, 299. B: 70 Dopamine D3 receptors, A: 299 Dopamine release, A: 167 Dopamine transporter (DAT), A: 293, 295, 299 Dorsal horn, B: 73, 87–101 Dorsal root, A: 26, 27. B: 41, 95, 108 Dorsal root entry zone, A: 27 Dorsal root ganglia, A: 26. B: 108, 148 Dorsal striatum, A: 8, 169, 171 Dorsalisation, A: 269. B: 37 Dorsomorphin, A: 57 Dose escalation studies, A: 104, 110, 160, 161, 179. B: 125, 232 Doublecortin (DCX), A: 81, 85. B: 249–251, 253 Doxycycline, A: 78 Driving, B: 226 Drug cocktail, A: 57–59. B: 4, 117, 118, 121 Drug dose, A: 8, 104, 115, 239, 243, 249, 270. B: 19, 88 Drug screening, A: 73, 268, 275. B: 148, 150, 192, 201, 208, 209, 211 Drug therapy, A: 5, 6, 12, 141, 203, 319. B: 66, 74, 87, 88, 90, 97, 100, 201 Drug withdrawal, A: 204, 236. B: 113, 230 Drusen, B: 226–228 Dual task tests, A: 316, 317 Dual-SMAD inhibition, A: 139, 195, 272 DuoDopa, A: 166, 167, 182 Dyskinesias, A: 107, 134, 168, 169, 180, 216, 220 Dystrophic rats, B: 229

E Economic issues, A: 166, 175. B: 229–231, 245 Ectoderm, A: 73, 195. B: 201, 205, 207 Efficacy, A: xxvi, 3, 5, 7, 9, 29, 30, 34, 103, 107, 108, 112, 113, 116, 117, 133–136, 140, 141, 145, 151, 152, 157, 158, 162, 167–170, 173–181, 192, 199, 201, 202, 204, 205, 218, 220, 227–229, 236, 238–240, 243, 248, 252, 254, 265, 274, 283, 285, 286, 295, 297, 300, 307, 321. B: xxvi, 4, 34, 38, 45, 71, 74, 108, 112, 113, 148, 150, 174, 229–231, 235, 240 Efficiency, A: 53–57, 61–63, 69, 82, 83, 104, 175, 195, 196, 215, 250, 270, 272. B: 10, 21, 37, 142–144, 148, 172, 176, 178, 201 Electroconvulsive shock, B: 71 Electroencephalography (EEG), B: 70, 71 Electron microscopy, A: 23. B: 16, 36, 42, 45, 63, 94–96, 120, 255

Combined Index

Electrophysiology, A: 2, 5, 13, 14, 26, 56, 62, 73, 83, 86, 89, 228, 233, 234, 243, 253, 254, 266, 311. B: 8, 20, 39, 41, 43–45, 63, 71, 90–95, 97, 125, 126, 137, 138, 148, 179, 180, 206, 240, 253–255 Electroretinography, B: 198, 200 Elevated body swing test, B: 249 ELISA, A: 250. B: 238 Embryoid body, A: 177, 194, 195, 270. B: 37, 38, 203–205, 207 Embryonic age, A: 19, 23, 247, 264, 266. B: 17, 59, 60, 64, 66, 167 Embryonic development, A: xxvi, 2, 8, 10, 24, 25, 57–61, 71, 73, 74, 193–195, 198, 199, 205, 215, 263–276, 306, 308. B: xxvi, 5, 6, 17, 21, 37, 46, 57–67, 70, 89, 110, 136, 137, 139, 144, 146, 151, 191, 192, 196, 201, 203, 207, 210–212, 255 Embryonic donors, A: 19, 102, 134, 143, 156, 216, 306. B: 38 Embryonic stem cell grafts, A: 178, 263–276. B: 168–170. See also hESC grafts Embryonic stem cells (ESCs), A: 24–26, 70, 102, 107, 133–145, 151–159, 161, 162, 165–185, 191–205, 214, 216, 217, 263–276, 292. B: 2, 7–11, 13, 14, 16–18, 21, 37, 38, 42, 168–170, 191–212, 225–240, 247. See also hESCs Endoderm, A: 138, 195. B: 201 Endogenous cues, A: 114, 218, 274, 308. B: 64 Endogenous glial cells, A: 71, 72, 74, 76, 79, 101. B: 9, 41, 167 Endogenous neurogenesis, A: 79, 81, 82, 85. B: 246 Endogenous remyelination, B: 146–153 Endogenous stem cells, B: 33, 40, 113, 135, 136, 146, 147, 212, 246 Endothelial cells, B: 246 ENGAGE-HD, A: 321 Engineered cells, A: 30, 31, 99, 102, 103, 106, 108–116, 167, 205, 263, 276. B: 10 Enkephalin, B: 89 ENROLL trial, A: 244 Environmental enrichment, A: 305–322 Ependymal cells, A: 74, 233, 246 Ependymal cyst, A: 233 Epiblast stem cells, B: 150 Epidermal growth factor (EGF), A: 81, 101, 273. B: 5, 110, 204 Epigenetic factors, B: 143, 147, 170, 201 Epigenetic markers, A: 275. B: 43, 170 Epigenetic memory, B: 201, 208

Epilepsy, A: 10, 34, 114. B: 58, 67, 70, 71, 73, 89 Episodic memory, A: 16 ErbB, B: 110, 111, 148 ES Cell International (ESI-n) cell lines, A: 137, 138 Essential-8, A: 197 Estradiol, A: 218, 219 Estradiol-2-benzoate (E2B), A: 219 Ethical issues, A: 69–71, 102, 133, 134, 165, 168, 172, 174, 193, 197, 202, 203, 242–245, 250, 267. B: 3, 8, 34, 37–39, 147, 182, 208, 210, 247 Ethical review board, A: 244, 250 Europe, A: 32, 107, 139, 174, 185, 214 European Huntington’s Disease Network (EHDN), A: 251 European Medicines Agency (EMA), A: 135, 142, 158. B: 234 European Union, A: 135, 138, 322 European Union Tissue and Cells Directive (EUTCD), A: 138. B: 234 Evolution, B: 60, 136, 178, 182 Ex vivo gene therapy, A: 99–117. B: 19 Excitatory amino acids, B: 34 Excitatory Interneurons, B: 93, 96 Excitotoxic amino acids (EAAs), A: 4, 11, 21, 25, 291, 309. B: 45 Excitotoxic lesions, A: 3–5, 11, 13, 75, 265, 266 Exclusion criteria, A: 182, 183, 202, 203, 220, 245, 254. B: 25 Exercise, A: 305–322. B: 27 Expansion, A: 54, 55, 140, 143, 152, 154, 155, 172, 173, 194, 197, 198, 216, 274. B: 9, 21, 60, 100, 109–111, 114–117, 119–121, 167, 169, 170, 237, 238, 248 Experimental allergic encephalomyelitis (EAE), A: 9. B: 141, 144, 148–150 Extracellular matrix, A: 235. B: 41, 57, 108, 109, 144, 201, 204, 227 Eye, B: 66, 191–212, 225–240

F Facial recognition, B: 226 Fascicles, B: 117, 119–121, 126, 192 Fate specification. See Specification F-DOPA, A: 183–184, 204, 218, 220 Fear conditioning, B: 69 Feasibility, A: 78, 89, 106, 107, 181, 184, 227, 229, 235, 243, 244, 321. B: 11, 112, 126, 172, 246, 247, 257 Feeder cells, A: 137, 139, 141, 157, 158, 194, 195, 198, 213, 215, 217

271

272

Combined Index

Fetal calf serum, B: 203–206 Fetal tissue, A: 3, 18, 19, 21, 25, 26, 31, 33, 69, 70, 83, 101, 102, 106–108, 110, 112, 116, 134, 135, 139–141, 165–185, 192, 193, 202–205, 214, 216, 220, 222, 227–229, 232, 234, 246, 249, 254, 264, 266, 267, 273, 276, 286, 288–290, 294. B: 2, 3, 5–9, 11, 12, 14, 17, 19, 21, 34, 36, 37, 45, 57, 100, 121, 126, 167, 172, 201, 203, 229, 246 FGF1 (acidic FGF), B: 203 FGF2 (basic FGF), A: 61, 80, 81, 109. B: 5, 35, 37, 203–205, 230 FGF8, A: 194, 215. B: 11 Fibrin glue, B: 249–251 Fibrin matrix, A: 23, 161. B: 13, 14, 21 Fibroblast growth factor (FGF), A: 8, 24, 25, 30, 61, 81, 101, 109, 194, 196, 215, 269, 273, 274. B: 5, 11, 35, 37, 111, 203–205 Fibroblasts, A: 21, 31, 56–63, 72, 73, 75, 78, 83, 84, 87–89, 101, 105, 106, 108, 138, 143, 156, 161, 215, 274, 275. B: 4, 9, 10, 38, 39, 108, 112, 115, 117–119, 122, 126, 169, 170, 201, 233, 238, 245–258 Fibrosis, B: 229 Fimbria-fornix lesions, A: 18, 20, 21, 31 Fingolimod, B: 149, 150, 175 First-in-man trials, A: 32, 175, 181, 184, 283. B: 177 Fischer 344 strain, B: 13 FK506 (Tacrolimus), A: 184, 204, 250 Floor plate, A: 25, 60, 139, 140, 144, 173, 178, 195, 196, 215, 216, 218, 269. B: 192, 203 Flow cytometry, A: 179 Fluorescence-activated cell sorting, A: 140, 216. B: 6, 8, 166, 196, 198 FluoroDOPA. See Radioligand Fluoro-Gold, A: 12. B: 12 Food and Drug Administration (FDA), A: 103, 105, 110, 117, 135, 139, 141, 151–162, 175, 176, 196, 199. B: 107, 108, 113, 119, 121–126, 128, 149, 150, 169 Forebrain, A: 4, 10, 17, 20–22, 25, 73, 195, 264, 269–272, 275. B: 5, 13, 37, 166, 171, 204 Forebrain identity, A: 195. B: 204 Forelimb placement, B: 250, 251 Formalin, B: 92–94, 98 Forskolin, A: 57, 58, 61, 84. B: 112, 115, 116, 118–121, 126 Foundation of Accreditation for Cellular Therapy, B: 124 FoxA1, A: 60 FoxA2, A: 25, 60, 140, 144, 195, 196, 200, 216

FoxG1, A: 270 FoxP1, A: 271, 274, 276 FoxP2, A: 271, 272 France, A: 159, 161, 227, 233, 238, 283 Free radicals, B: 34, 35, 228 Frizzled receptors, B: 145 Frontal cortex, A: 5, 20, 237, 253, 266, 294. B: 15. See also Prefrontal cortex Frontoparietal cortex, A: 12, 14, 21 Fronto-striatal pathway, A: 5, 228, 233, 266, 294, 317–319 Functional Neural Transplantation, A: xxv–xxvii. B: xxv–xxvii Functional recovery, A: 1–34, 69, 89, 109, 111–113, 167, 168, 179, 196, 219, 220, 236, 245, 246, 264, 267, 270, 294, 307, 309, 311–316, 319. B: 1, 4, 8, 10, 19–21, 34–47, 67, 72, 100, 126, 137, 171, 172, 177, 200, 232, 245–258 Functional rescue, A: 60, 201. B: 70, 73, 74, 171, 179, 181, 199, 232

G GABA, A: 10, 11, 13, 14, 18, 20, 23, 60, 72, 73, 82, 85, 88, 114, 270, 272, 274, 289, 290, 296, 299. B: 9, 39, 57, 58, 61, 66, 69, 71–74, 87–101, 253 GABA receptors, B: 89, 90 GABAergic grafts, B: 57–74, 87–101 Gabapentinoids, B: 88 GABAstat, B: 96, 97 GAD expression, B: 73, 90, 96, 98 GAD staining, A: 13, 23 GAD65, A: 23, 272. B: 96, 249, 250 GAD67, A: 272. B: 96, 249, 250 GAD67 promoter, B: 91 Gain of function toxicity, A: 263 Gait analysis, A: 237, 319. B: 39 Galactocerebrosidase, B: 172 Galectin-1, A: 113 Ganglion cell layer, B: 192, 193, 195, 211 Ganglionic eminence, B: 57–74, 80–101. See also CGE; LGE; MGE; WGE Ganglionic eminence grafts, B: 60, 61, 87–101. See also Striatal grafts Gangliosides, B: 168 Gangliosidosis, B: 170 G-banding. See Karyotyping Gene editing, A: 55 Gene therapy, A: 55, 99–117, 244, 249. B: 168, 192, 194, 234 Genetic diagnosis, A: 228

Combined Index

Genetic disorders, B: 139, 168, 193–195 Genetic engineering, A: 8, 21, 30, 31, 55, 99, 102, 103, 106, 108–116, 167, 205, 263, 276. B: 10, 171 Genetic manipulation, B: 113 Genetic mutation, A: 54, 55, 100, 104, 110, 144, 175, 176, 203, 215, 235, 263, 265. B: 11, 43, 65, 70, 98–100, 139, 172, 178–180, 193, 195, 200, 201, 208, 211, 212, 229, 230 Genome editing, A: 53–63 Genomic stability, A: 133, 136, 141–143. B: 9 Gentamicin, B: 126 Geographic origin, A: 175 Germ layers, A: 70, 71, 178. B: 37, 201 Germany, A: 69, 227, 241, 250, 305 Geron. See Asterias GForce-PD, A: 32, 142, 185 Girk2, A: 22 GLI3, A: 271 Glial cell line-derived neurotrophic factor (GDNF), A: 8, 30, 103, 108–115, 218. B: 11, 42, 46 Glial cell replacement, A: xxvi, xxvii, 6, 62. B: xxvi, xxvii, 47, 170, 174, 175, 179, 181, 182, 229, 234 Glial fibrillary acid protein (GFAP), A: 73–76, 82, 83, 85, 233, 291, 296, 298. B: 5, 42, 112, 180, 249–251, 253 Glial grafts, A: 29. B: 20, 39–42, 107–128, 147, 165–182 Glial growth factor (GGF), B: 114–116, 118 Glial progenitor cells, A: 102 Glial progenitor cells (GPCs). See Glial progenitors Glial progenitors, A: 62, 76, 108, 289. B: 145, 165–182 Glial scar, A: 6, 9, 76, 77. B: 3, 9, 16, 33, 34, 36, 47, 246 Glial support, A: 6 Gliomas, A: 77, 141, 290 Gliosis, A: 75, 83, 250. B: 193 Globus pallidus, A: 4, 5, 11–14, 21, 22, 166, 286, 294, 306. B: 61, 253, 254 Glucose, A: 84, 294–297. B: 228 Glucose metabolism, A: 233, 234, 237–239, 241, 289, 294–297, 319 b-Glucuronidase, B: 171 Glutamate, A: 16, 23, 26, 57, 59, 72–76, 82, 88, 290, 310. B: 9, 57, 178 Glutamate synthase, A: 74 Glutamate transport, B: 178 Glutamic acid decarboxylase (GAD), A: 13, 23, 108, 272. B: 73, 90, 91, 96, 98 Glutamine, A: 74. B: 126 Glycolysis, A: 84, 289, 290

Gnat1, B: 197 GNbAC1, B: 153 Gold standard, A: 70, 173, 174, 251, 252, 296, 315, 318. B: 47, 201 Golgi staining, A: 13 Good clinical practice (GCP), A: 242 Good manufacturing practice (GMP), A: 54, 70, 133–145, 151–162, 165–185, 191–205, 213–222, 250, 283. B: 121–125, 127, 205, 208, 234–238 Good Tissue Practice (GTP), A: 151–162 Graft connections, A: 1, 12, 13, 219, 232, 233, 251, 266, 276, 294, 307, 309. B: 2–4, 6, 10–13, 18–20, 34, 44, 64, 196, 200, 211, 254, 255 Graft dose, A: 104, 105, 110, 117, 141, 157, 160–162, 180–184, 192, 196, 199, 201–203, 284. B: 124, 125, 174, 232, 239 Graft growth, A: 77, 81, 84, 109, 112, 290, 300. B: 9, 15, 16, 21, 248, 253 Graft integration, A: xxvi, 1, 3, 6, 10, 13, 16, 20, 23, 25, 31, 33, 79, 87–89, 102, 103, 138, 158, 162, 169, 172, 173, 228, 263, 266, 275, 285, 286, 294, 296, 305, 306, 309. B: xxvi, 6, 8, 12, 57, 58, 61, 66, 68, 72–74, 87–89, 93, 94, 97, 100, 180, 196, 199, 210, 237, 245, 247, 255, 256, 258 Graft metabolism, A: 314 Graft misplacement, A: 237, 238, 248, 249 Graft overgrowth, A: 6, 7, 25, 26, 34, 181, 184, 232, 233, 238, 247, 248, 270, 271, 273, 287–290, 298. B: 9, 21, 118, 248, 252 Graft placement, A: 3, 5, 30, 168, 169, 180, 181, 184, 247, 248, 251, 267, 285, 309, 314. B: 44, 172, 196 Graft rejection, A: 29, 71, 72, 101, 116, 168, 172, 228, 239, 241, 249, 250, 253, 266, 267, 283, 288–290, 296, 297, 300. B: 3, 8, 21, 169, 208, 229, 231 Graft removal, B: 20, 36, 68, 231 Graft survival, A: xxv, xxvi, 2, 6–9, 17, 24, 30, 60, 61, 78, 104–117, 139, 168, 169, 171–173, 184, 193, 194, 196, 198, 200, 202–204, 215–222, 235, 236, 241, 266, 283–287, 291, 292, 299, 307–311, 313, 314, 319, 321. B: xxv, xxvi, 6, 8, 10–14, 19, 21, 34, 35, 41, 42, 45–47, 64, 65, 70, 71, 100, 109, 111, 112, 123, 124, 151, 172, 179, 180, 198, 200, 230–233, 252, 257, 258 Graft tracking, A: 99, 115, 283–297, 299, 300, 314. B: 231, 257 Graft vascularisation, A: 233–235, 245 Graft-induced dyskinesia (GID), A: 168–170, 180, 181

273

274

Combined Index

Grafts, A: 14–16, 18, 20–26, 29, 31, 71, 84, 100, 101, 110, 116, 134, 139, 168, 169, 172, 174, 178–182, 184, 215, 216, 228, 232–234, 236–245, 248–250, 253, 283–288, 290, 292–294, 296–300, 305, 309–314, 316, 319, 321. B: 3, 5, 9, 11, 13–21, 33–48, 57–74, 87–101, 113 Grasping, B: 249 Gray matter, A: 22, 73, 76, 77. B: 1, 14, 15, 166, 177 Green fluorescent protein (GFP), A: 22–24, 79, 81–83, 86, 312. B: 12–16, 18, 91, 93–96, 124, 196–199, 210, 249–251 Growth cone, A: 8, 26 Growth factors, A: 9, 24, 81, 82, 99–117, 155, 194, 198, 269. B: 11, 14, 21, 46, 109–111, 113, 119, 142, 201, 227, 246 GSK239512, B: 153 Gsx2, A: 271 GTP cyclohydrolase 1 (GCH1), A: 115 Gurdon, Sir John, A: 71

H H9 cell line, A: 137, 139, 156, 158, 197 Hadassah Research Centre (HADCn) cell lines, A: 137 Haematoma, A: 232 Haematopoiesis, A: 71 Haematopoietic stem cells, B: 201 Haemorrhage, A: 220. B: 193 Haplotype, A: 215, 221, 267. B: 208 Hb9, A: 61 Heart, A: 9, 71, 161, 201, 216 Helix-loop-helix factors, A: 57 Hemiparkinsonism, A: 200, 293 Heparin, B: 35 Hepatocyte growth factor (HGF), B: 11, 46 Hepatocytes, A: 56 Heregulin, B: 108, 110–112, 118–122, 126 hESC grafts, B: 177. B: 225–240 hESCs, A: 25, 26, 102, 133–145, 151, 152, 155–158, 160–162, 165, 173–175, 178, 180, 181, 184, 191–205, 269, 270, 272, 275, 293. B: 167–170, 177, 179, 201, 203, 205, 206, 208, 210, 211, 230–232, 235, 236, 238, 239 Heterochromatin, B: 120 Heterochronic transplantation, B: 58, 61–63, 66–69, 74 Hexokinase, A: 295, 297 b-Hexosaminidase, B: 171 Hibernation, A: 231, 267 High-frequency stimulation, A: 312 Hindbrain, A: 171, 181, 195, 196, 216, 269

Hippocampal grafts, A: 17–21. B: 251 Hippocampus, A: 1, 2, 10, 16–21, 28, 31, 75, 81, 106, 114, 290, 311. B: 7, 13, 60, 61, 67, 70, 71, 73, 89 hiPSC grafts, B: 173, 176, 196, 247, 249–251 hiPSCs, A: 54, 55, 60, 173–175, 191–205, 272. B: 39, 44–46, 167–170, 173, 205, 206, 208, 210–212 Histamine, B: 88, 100 Histamine H3 receptor agonists, B: 153 Histone deacetylase (HDAC), B: 143, 146 HLA antibodies, A: 228, 237–239, 241, 246, 250 HLA matching, A: 213, 215, 246, 267. B: 208, 230 HLA monitoring, A: 228 HLA-DR, B: 40 HLA-matching, B: 208 hMito, B: 247, 248 Hoechst stain, B: 250 Hoehn & Yahr scale, A: 220 Homeostasis, A: 74, 77, 88, 273. B: 167, 178 Homing, A: 9, 104 Homotopic grafts, A: 4, 5. B: 61, 64, 65 Horizontal cells, B: 195–197 Horseradish peroxidase (HRP), A: 12, 13, 27. B: 11–13, 45, 94 Host brain environment, A: 1, 30, 101, 213, 218, 219, 221, 222, 266, 284, 319. B: 7, 12, 33, 36, 47, 87, 89, 90, 97, 109, 176, 246, 253 HuD, B: 253 Human immunodeficiency virus (HIV), A: 104, 140 Human leukocyte antigens (HLA), A: 213–222. B: 208, 230, 231 Human nuclear antigen (HuNu), B: 44, 173, 180, 249, 250 Human tissue, A: 76, 142, 154, 155, 162, 168, 193. B: 167 Huntingtin (Htt), A: 112, 235, 263, 265. B: 179 Huntington’s disease, A: 2, 34, 99, 104, 111, 112, 227–255, 263–276, 283, 284, 286, 287, 291, 294, 295, 297, 300, 305, 306, 315, 318. B: 69, 165, 166, 178, 180, 181 Hyaluronan, B: 144, 179 Hydrogel, B: 249–251 6-Hydroxydopamine (6-OHDA), A: 3, 4, 15, 25, 26, 30, 60, 107–109, 135, 141, 173, 179, 216, 219, 293, 294. B: 72 Hyperactivation, B: 70 Hyperactivity, A: 3, 11. B: 73 Hyperalgesia, B: 73, 90–92 Hypercellularity, B: 248, 252 Hyperexcitability, A: 10. B: 57, 58, 64, 69, 70, 74, 87, 89 Hyperoxia, B: 206

Combined Index

Hypersensitivity, B: 73, 74, 88, 89, 91, 92, 97, 100 Hypertension, A: 240. B: 174, 175, 257 Hypertrophy, A: 75, 77 Hypometabolism, A: 233, 234, 239, 294, 296 Hypomyelination, B: 167, 172, 173, 176, 177, 181 Hypothalamus, A: 2, 3, 31 Hypoxia, A: 290

I Iba-1, A: 291, 298. B: 249, 251 Ibotenic acid, A: 11 IGF-1, A: 106, 109, 110. B: 204–206 192 IgG-saporin. See Immunotoxin Immediate early gene (IEG) expression, B: 70 Immune privilege, A: 204, 249, 266. B: 227, 230 Immune rejection. See Graft rejection Immunogold, B: 96 Immunosuppression, A: 12, 32, 101, 116, 168, 172, 174, 175, 184, 197, 202, 203, 205, 220, 227, 232, 234, 235, 238, 239, 241–243, 246, 249, 250. B: 13, 14, 16, 18, 168, 169, 174, 229, 247. See also Azathioprine; Cyclosporine; FK506; Prednisolone; Triple regime Immunotherapy, A: 168, 172, 184. B: 192, 194 Immunotoxin, 192 IgG-saporin, A: 20, 21. B: 71 Implantation instrument, A: 105, 155, 158, 171, 180, 202, 204, 222, 247–249. B: 122, 124, 233 iN cells. See Induced neurons (iN cells) In utero transplantation, B: 61, 64, 65 In vitro fertilisation (IVF), A: 156 In vivo gene therapy, A: 99, 100 In vivo imaging, A: 204, 283–286, 288–300, 314 Inclusion criteria, A: 202, 203, 220, 245, 246, 254. B: 125 Induced glia (iGlia), A: 61, 62. B: 170 Induced neural stem cells (iNSCs), B: 2–4, 9, 10, 247, 248, 252. See also Neural stem cells (NSCs) Induced neurons (iN cells), A: 53–63, 84. B: 2, 9, 10, 255–258 Induced pluripotent stem cells (iPSCs), A: 24, 25, 32, 53–63, 70, 84, 100, 102, 104, 107, 116, 117, 133–145, 152–162, 173–175, 191, 213–222, 249, 268. B: 2, 4, 8–11, 14, 20, 21, 34, 38–40, 43–46, 111, 167–170, 173, 191–212, 230, 245–258. See also hiPSCs Induced plurpotent stem cell grafts, A: 263–276. B: 100, 168–170, 247–252, 254, 255. See also hIPSC grafts

Infection, A: 75, 78, 82, 116, 232 Inflammation, A: 9, 62, 75, 76, 78, 84, 88, 108, 113, 234, 242, 249, 250, 283, 285, 290, 296–298, 300. B: 3, 6, 10, 13, 34–36, 40, 42, 87, 88, 90, 92, 93, 98, 100, 137, 139–141, 150, 165, 170, 175, 177, 193, 245, 246, 253 Informed consent, A: 156, 175, 197, 203, 228, 240, 244, 245, 253, 254, 267. B: 125 Inherited retinal degeneration, B: 191–212 Inhibition, A: 14, 58, 60, 110, 143, 215, 270, 272. B: 6, 57, 64, 68–73, 89, 90, 96–98, 141, 144–146, 148 Inhibitory Interneurons, B: 57–74, 87–101 Injection device. See Implantation instrument Innate immune response, B: 141, 143 Inner nuclear layer, B: 192, 193, 195, 197, 200, 211 iNOS, B: 249, 253 Insertional mutagenesis, A: 55, 100, 102 Integrin, A: 219. B: 42, 88, 151, 175 Interleukins, B: 35 IL-1a, B: 35 IL-1b, B: 35, 249, 251, 253 IL-2, B: 249, 253 IL-4, B: 40, 249, 253 IL-6, B: 35, 40, 249, 251, 253 IL-10, A: 113. B: 35, 249, 251, 253 IL-13, B: 40 IL-31, B: 99, 100 International Stem Cell Corporation, 159–161 Interneurons, A: 3, 10, 13, 14, 20, 23, 33, 60, 72, 75, 85, 87, 159–161, 233, 235, 241, 246, 308. B: 2, 10, 34, 35, 44, 45, 57–74, 87–101, 192, 258 Interneuron grafts, B: 57–74, 87–101 Interneuropathy, B: 58, 69 Intraarterial delivery, B: 246 Intracerebral haemorrhage, B: 251 Intraparenchymal delivery, A: 105, 110. B: 246 Intrathecal delivery, A: 105, 110, 111. B: 3, 4, 88, 89, 100 Intravascular delivery, A: 104, 105 Intravenous delivery, A: 239, 249. B: 3, 246 Intraventricular delivery, A: 20, 101, 105, 106, 201 Investigational new drug (IND) application, A: 196. B: 124, 125 Iodobenzamide, A: 237 Ion transport, B: 227 IPBZ, A: 237, 238, 252 Iran, B: 108 Iris, B: 202 Irradiation, A: 104, 284, 285

275

276

Combined Index

Ischaemia, A: 70, 75, 81, 113, 288. B: 140, 245, 246, 248, 256, 257 Isl1, A: 61 Isochronic grafts, B: 61–65 Isogenic grafts, B: 13 Israel, A: 111, 139, 159, 161. B: 232 Italy, A: 53, 237, 250 Itch, B: 67, 87–101

J Japan, A: 32, 34, 139, 152, 161, 175, 185, 213–215, 221. B: 33, 34, 37, 48, 231, 233

K Karyotyping, A: 54, 142, 197, 198, 233. B: 127, 236, 238 Kearns-Sayre syndrome, B: 139 Keratinocyte stem cells, B: 201 Ki67, B: 203, 249, 250 Kidney, A: 141, 201, 214, 215, 235 Kings College London (KCL) cell lines, A: 137, 138 KLF4, A: 215, 268. B: 38–40, 201 Knock-in mouse, A: 103. B: 73 Knockout mouse, A: 269. B: 98, 145, 148, 149 Knockout serum replacement, A: 144, 269, 272. B: 203, 205 Korea, B: 7 Krabbe’s disease, B: 139, 170–172 Krebb’s cycle, A: 295 Kyoto trial, A: 213–222

L L1, B: 10, 42 Lactate, A: 288, 289 Laminin, A: 30, 177, 213, 215–217. B: 35, 42, 119–122, 126 Laser photocoagulation, B: 211 Lateral ganglionic eminence (LGE), A: 72, 242, 246, 264–266, 269–275. B: 59, 61–63 Lateral ventricle, A: 74, 264, 291 LDN193189, A: 195 L-dopa, A: 4, 5, 115, 134, 166, 203, 213, 214, 220, 292 L-dopa-induced dyskinesia (LID), A: 166, 170, 171, 181, 182 Learning, A: 11, 14–21, 71, 308–312. B: 69, 73, 136, 178, 257 Learning and memory. See Memory Learning to use the grafts, A: 15, 16, 305–322 Leber’s congenital amaurosis, B: 195 LeftyA, B: 203, 204 Lens, B: 205, 207

Lentivirus, A: 58, 62, 82–84, 103. B: 12, 14, 20 Leukemia inhibitory factor (LIF), B: 37 Leukocytes, A: 221 Leukocyte antigens. See Human Leukocyte antigens (HLA) Leukodystrophy, B: 165, 167–171, 175 Leukomalacia, B: 175 Lewy body pathology, A: 169 Life expectancy, A: 166. B: 140 Life span, B: 193 Lifespan, A: 110, 111, 117. B: 147, 172 Light avoidance test, B: 200 Light-detecting electrical devices, B: 195 Light-evoked activity, B: 200, 206 LIN28, A: 215 Lineage commitment, A: 266, 274, 275, 318. B: 109, 112, 198 Lingo1, B: 148 Liothyronine, B: 152 Lipids, A: 289, 290, 292. B: 136, 139 Lipopolysaccharides, A: 78 Lithium carbonate, B: 152 Liver, A: 201, 214 Lmx1a, A: 25, 60, 86, 140 Lmx1b, A: 60, 140 L-Myc, A: 215. See also Myc proto-oncogene Locomotor activity, A: 110, 111, 308. B: 43, 72, 118, 127, 250 Logistics, A: 69–71, 133, 134, 156, 165, 171, 172. B: 208, 210 London Project to Cure Blindness, A: 34. B: 232 Long-distance axon growth, A: 21–25, 27, 28, 181. B: 1, 12–14, 16, 20, 44, 45 Longitudinal studies, A: 183, 235, 244, 251, 283, 285, 286, 299, 300, 318, 319 Long-term depression (LTD), A: 15–17, 311–313 Long-term neuroepithelial stem cells (lt-NESCs), B: 248–255 Long-term potentiation (LTP), A: 15–17, 311–313 Lou Gehrig disease. See Amyotrophic lateral sclerosis LRTM1, A: 141 Lucentis, B: 226 Luciferase, A: 115 Luminex, A: 250 Lymphocytes, A: 104. B: 140, 150 Lymphoid organ, B: 150 Lyn kinase, B: 151 Lysolecithin, B: 141, 145, 148, 150, 176 Lysosomal storage disorder, B: 139, 170, 171 Lysosomes, B: 120, 171

Combined Index

M M6 (mouse-specific marker), A: 12. B: 13 MAb-O4, B: 168, 251 Macrophage, A: 78, 233. B: 3, 40, 47, 143, 144, 147, 149 Macrophage inflammatory protein (MIP-1a), B: 251 Macula, B: 193, 194, 226, 229, 232, 233 Macular atrophy, B: 194 Macular dystrophy, A: 137. See also Age-related macular degeneration (AMD) Macular translocation, B: 229, 231 Magnetic resonance imaging (MRI), A: 115, 182, 183, 204, 218, 232–235, 239–241, 243, 248, 252, 283–300, 314, 315. B: 257 Magnetic resonance spectroscopy (MRS), A: 283–300 Magnetic-activated cell sorting (MACS), B: 196 Male germ cell-associated kinase (MAK) gene, B: 211 Malignant cells, A: 34, 141 Manchester University (Mann) cell lines, A: 137, 138, 158, 159, 175 Mannose-6-phosphate, B: 171 Manufacture of Sterile Medicinal Products, B: 236 Market approval, A: 193, 205 Marmoset, A: 168. B: 37, 39 Master cell bank, A: 138, 139, 152, 153, 160, 177, 197 Master transcriptional regulators, A: 71 Matrigel, B: 61, 204, 205 Matrix metallopeptidase (MMP-9), B: 251 Maturation, A: 56, 57, 60, 63, 81-83, 85–88, 140, 168, 174, 216, 218, 243, 275, 290, 292, 294, 296, 299, 314. B: 6, 17, 62, 66, 67, 93, 144, 146, 192, 203, 206, 207, 230 Maze learning, A: 2, 11, 18–21. B: 198, 250, 252 mCherry, A: 83 Mechanism, A: 1–34, 53, 56, 58, 59, 63, 73, 85, 104, 105, 113, 228, 235, 249, 285, 311, 312, 319. B: 6, 10, 19, 20, 33, 34, 41–43, 46, 47, 64, 68, 71, 72, 74, 90, 94, 96, 97, 136, 140, 143, 149, 150, 192, 235, 245–247, 252, 255, 256, 258 Medial forebrain bundle, A: 22, 173 Medial ganglionic eminence (MGE), A: 10, 264, 265, 270–272. B: 59–74, 87–101 Medial striatum, A: 14 Median nerve, A: 253 Medical ethics. See Ethical issues Medical Research Council, A: 133, 165, 322. B: 135, 212 Medical termination of pregnancy (MTOP), A: 171, 267. See also Abortion

Medicines and Healthcare Products Regulatory Agency (MHRA), A: 135, 141. B: 234–236 Medium spiny neurons (MSNs), A: 12–14, 16, 17, 33, 71, 73, 87, 229, 263–276, 294, 311, 314. B: 61, 179, 180, 255 Memory, A: 11, 14, 16, 18, 21, 275, 308, 317. B: 69, 72, 73, 136, 201, 208 Memory deficits, B: 72 2-Mercaptoethanol, B: 205 Mertk defects, B: 193, 229 Mesencephalic DA (mDA) neurons, A: 3–5, 8, 12–15, 17, 20, 22, 23, 25, 26, 30, 60, 69, 71, 73, 133–145, 165–184, 191–205, 213–222 Mesenchymal stem cell grafts, A: 111, 167. B: 1–22, 33, 34, 40, 42, 43, 46–48, 246 Mesenchymal stem cells (MSCs), A: 9, 99, 101, 102, 105, 106, 109, 111–113. B: 1–22, 33, 34, 40, 42, 43, 46–48, 111 Mesoderm, A: 74. B: 4, 201 Metabolic storage disorders, B: 170, 174 Metabolic support, B: 136 Metachromatic leukodystrophy, B: 170, 171 Methyl-phenyl-tetrahydro-piridine (MPTP), A: 108, 202 Met proto-oncogene tyrosine kinase, B: 229 Miami Project to Cure Paralysis, B: 107–128 Miconasole, B: 150 Microarray, A: 142. B: 207 Microduplication, A: 143 Microglia, A: 74, 78, 82, 102, 106–108, 113, 116, 221, 241, 291, 296, 298, 300. B: 3, 90, 137, 143 MicroRNAs, A: 57, 274 Microspheres, B: 19 Microtubule-associated protein, MAP2, A: 73, 270, 274. B: 249, 250 Midbrain, A: 5, 22, 25, 30, 32, 60, 71, 81, 83, 87, 134, 136, 139–141, 165, 168, 170, 173, 176, 178, 179, 191–196, 198, 202, 204, 215, 216, 219, 269, 273, 286. B: 13 Midbrain identity, A: 140, 196 Middle cerebral artery occlusion (MCAO), B: 249–251 Mifepristone, A: 114 MIG-HD trial, A: 227–255 Migration, A: 10, 74, 77, 78, 101, 105, 115, 158, 162, 201, 237, 241, 242, 248, 284, 285. B: 19, 41, 47, 57–63, 68, 74, 89, 90, 111, 117, 123, 124, 136, 137, 142–144, 172, 176, 177, 200, 205, 233, 257 Mini mental state examination (MMSE), A: 236, 237 Minipig, A: 110, 265. B: 124 Mitochondria, A: 296. B: 35, 139, 227

277

278

Combined Index

Mitogens, A: 101. B: 110–112, 115–121, 137 Mitosis, A: 269. B: 166, 197, 201 Molecular signature, A: 25, 73, 274. B: 74, 230, 236 Moloney murine leukaemia virus (MMLV), A: 79 Monkey. See Primate Monoclonal antibodies, A: 184. B: 148, 151, 167, 168, 251 Monocyte, A: 78. B: 149 Monogenic disorders, B: 194 Monolayer grafts, A: 34. B: 231–234, 238 Monounsaturated fatty acids (MUFAs), A: 292 Montreal cognitive assessment, A: 183 Morphine, B: 97 Morphology, A: 2, 11, 56, 57, 74, 77, 81, 84, 86, 309, 310, 316, 319. B: 59, 62, 63, 93, 112, 117, 120, 127, 137, 192, 196–199, 230, 233, 254 Morris water maze, A: 11, 20, 21. B: 198, 250, 252 Motion detection, B: 195 Motor cortex, A: 23, 25 Motor deficits, A: 238, 251, 307, 308, 315, 316. B: 72, 245, 257 Motor function, A: 10, 26, 110, 111, 134, 192, 218, 321. B: 1, 11, 20, 35, 36, 38, 39, 45–47, 91, 118, 123, 126, 179, 247, 252 Motor learning, A: 3, 5, 309, 310, 312, 319. B: 179 Motor neurone disease. See Amyotrophic lateral sclerosis Motor neurons, A: 34, 61, 71, 73, 109–111, 269. B: 3, 8–10, 12, 16, 37, 91, 178 Motor tests, A: xxvi, 11, 182, 216, 218, 316. B: xxvi, 179 Mouse, A: 2, 10, 12, 15, 22–26, 56–63, 69, 70, 73–75, 77, 79, 81–84, 86–88, 106, 108, 112, 115, 137, 140, 141, 156–158, 161, 193, 194, 196, 198, 200, 201, 203, 215, 216, 264, 270, 271, 273–276, 306, 308. B: 4–11, 13, 19, 36–39, 42–47, 58–61, 65–67, 69–74, 87, 89, 91–94, 96–100, 118, 123, 141–143, 145, 146, 148, 150, 167, 169, 171–173, 177–180, 182, 191, 196–198, 200–203, 205, 206, 208, 210, 211, 247–256 mRNA, A: 54, 55, 138, 143. B: 96, 253 Msx1, A: 141 Mucopolysaccharidosis type VII, B: 171 M€ uller cells, B: 195, 202, 205 Multicentre clinical trials, A: 32, 33, 108, 227–255 Multidisciplinary team, A: 146, 185, 193, 197, 248, 319, 322. B: 123 Multikinase inhibitors, B: 112 Multiple sclerosis, A: 9, 34. B: 135, 144–153, 165–167, 174–176 Multipotency, A: 9, 101, 241, 268. B: 4–6, 40, 137, 202

Murine viruses, A: 157 Muscle, A: 71, 110, 111. B: 126 Mutant mouse, A: 2, 54. B: 11, 43, 70, 98–100, 172, 177, 180, 200 Mutant rat, B: 200, 229 Myc proto-oncogene, A: 84, 215, 268. B: 38, 39, 201, 248–250 Mycophenolate, A: 204, 250 Mycoplasm, A: 140 Myelin, A: 61, 62. B: 11, 13, 33, 41–44, 57, 108, 109, 112–116, 118, 120, 123, 135–153, 165–182 Myelin basic protein (MBP), B: 15, 43, 150, 172, 173, 249, 250 Myelin protein zero (MPZ) protein, A: 61 Myelin sheath, A: 61, 62. B: 42, 43, 109, 116–118, 136–137, 139, 140, 173 Myelination, A: 62, 89. B: 2, 7, 10, 17, 20, 21, 39, 44, 46, 119, 135–137, 139, 145, 146, 148–151 Myelinoclastic disorders, B: 170 Myeloperoxidase (MPO), B: 251 Myocytes, B: 4 MyoD, A: 57, 71 Myogenesis, A: 71 Myo-inositol, A: 290 Myt1L, A: 56, 57, 73, 83, 89, 274. B: 9

N N2B27 medium, A: 272 N-acetyl-aspartate (NAA), A: 288–290, 292 Nanog, A: 140, 195. B: 249, 250 Natalizumab, B: 175 National Institute for Health Research (NIHR), B: 212 National Institutes of Health (NIH), A: 32, 138, 170. B: 22, 74, 232 Natural killer (NK) cells, A: 235 N-cadherin, B: 41 N-CAM, A: 25, 78. B: 13, 42, 249 Necrosis, A: 234, 287 Negative effects, A: 6 Neocortex, A: 4–7, 10–13, 17–21, 24, 228. B: 57–74 Neonatal transplantation, A: 2, 22, 24. B: 61, 63–67, 70, 71, 89, 90, 141, 172, 173, 176, 178–180 Neostriatum. See Striatum; Basal ganglia; Caudate nucleus; Putamen Neovascularisation, B: 193, 194, 226, 229, 233, 253 Neprilysin, A: 106 Nerve conduction, B: 34, 95, 126, 136–138, 140, 172 Nerve growth factor (NGF), A: 8, 31, 105, 112, 249. B: 11, 35, 41, 45, 46

Combined Index

Nestin, B: 14, 249–251, 253 NEST-UK, A: 235–237, 246, 256 NeuN, A: 73, 82–85, 291. B: 5, 7, 8, 14, 15, 93, 180, 249, 250 Neural crest, A: 61, 195. B: 112, 137 Neural induction, A: 194, 196, 198, 215, 268–270. B: 204 Neural progenitors (NPCs), A: 23, 63, 72, 74, 75, 81, 82, 85, 101–103, 105, 108–112, 114, 115, 173, 193, 194, 216, 241, 269, 272–275, 290, 292. B: 2, 5, 7, 8, 12, 13, 141, 247–253, 256 Neural rosettes, A: 144, 216, 270, 291, 298. B: 248–250, 252 Neural stem cells (NSCs), A: 70, 72, 74, 76, 101, 105, 106, 108, 110, 112, 113, 193, 268, 273, 274. B: 1–22, 33–39, 42, 43, 45–48, 166, 171, 252, 254 Neurexophilin 3, A: 218 Neuroblasts, A: 13, 17, 21, 22, 24, 59, 70, 81, 82, 85, 86, 237, 242, 294. B: 61, 245 NeuroD, A: 57, 72, 82. B: 9, 256 Neurodegeneration, B: 195, 229, 245, 246 Neurodegenerative disease, A: xxvii, 1, 7, 34, 62, 70, 73, 75, 76, 78, 88, 89, 101, 102, 109, 111, 116, 117, 166, 173, 222, 228, 229, 255, 275, 283, 284, 305, 306, 313. B: xxvii, 9, 21, 140, 178, 179 Neurodevelopmental disorders, B: 69, 182 Neuroectoderm, A: 195, 268, 271. B: 192 Neuroepithelium, A: 200, 268–270. B: 5, 45, 136, 204, 205, 248 Neurofibrillary tangles, A: 105 Neurofilament, A: 12. B: 12, 14, 17, 46, 173 Neurofilament NF-kB, B: 35 Neurog2, A: 57, 59, 61 Neurogenesism, A: 71, 76, 79, 81, 85, 114, 194, 307, 308, 319. B: 46, 57, 60, 64, 246, 253 Neurogenin Ngn2, A: 60, 72, 73, 77, 81, 82, 84 Neurogliaform interneurons, B: 60, 62 Neuroinflammation, A: 9, 106, 107, 113, 114, 285, 287, 296–298, 300 Neuroleptics, A: 233, 252 Neurological disorders, A: xxvii, 69, 99–113, 115–117, 222, 235, 288, 296, 319. B: xxvii, 9, 21, 57, 58, 69, 70, 89, 135, 140, 165, 166, 177, 182, 246, 252, 254 Neuromyelitis optica, B: 176 Neuronal ceroid lipofuscinosis, B: 170, 171 Neuronal morphology, A: 56, 73, 79, 83 Neuronal phenotype, A: xxvi, 269, 283, 286. B: xxvi, 93, 166, 247, 254 Neuronal relays. See Bridge grafts

Neuronal replacement, A: xxvi, xxvii, 3, 7, 8, 11, 61, 62, 70, 102, 107, 112, 134, 143, 167, 173, 191, 192, 263, 264, 286, 305, 306, 322. B: xxvi, xxvii, 7, 11, 196, 202, 209, 210, 234, 245–247, 252, 254, 258 Neuronal rescue, A: 84, 133, 135. B: 10, 101, 171, 172, 179, 210, 212, 236, 246 Neuropathic pain, A: 10. B: 69, 73, 74, 87–101, 126 Neuropeptide Y, B: 60, 93 Neuroplasticity, A: 1–34, 321. B: 57–74, 87–101 Neuroprotection, A: xxvi, 6, 8, 99–13, 115–117. B: xxvi, 1–3, 7, 8, 10, 19–21, 35, 42, 46, 47, 136, 195, 198, 245, 253 Neuropsychiatric Inventory, A: 183 Neurorehabilitation, A: 305–322 Neurospheres, A: 72, 76. B: 6, 8, 36–38 NeuroStemCell, A: 34, 171, 300 NeuroStemcellRepair, A: 34, 171, 175, 300 Neurosurgical approach, A: 6, 29, 170, 241. B: 70 Neurotoxicity, A: 78, . B: 35, 139 Neurotrophic support, B: 33–39, 42, 43, 45–48 Neurotrophin, A: 107. B: 12, 18, 19, 21, 45, 46, 65, 113 Neutrophils, B: 251 NG2 glia, A: 69–89. B: 9, 137, 142, 166, 256 Niemann-Pick disease, B: 170, 171 Night vision, B: 193, 195, 197, 198 Nigral grafts, A: 3–5, 21–23, 26, 30, 133–145, 165–185, 191–205, 213–222, 295, 297. B: 247 Nigrostriatal pathway, A: 4, 14, 15, 21–23, 25, 28, 30, 134, 136, 145, 166, 167, 286, 292. B: 72 Nitric oxide synthase (NOS), A: 85 Nkx2.1, A: 270–272. B: 60, 64 NMDA receptors, A: 310. B: 69, 70 N-methyl-D-aspartic acid (NMDA), B: 45 Nociceptive C-fibres, B: 94 Nocifensive behaviour, B: 92. B: 93, 98 Nodal, A: 62, 139, 195. B: 203 Nodes of Ranvier, B: 172 Noggin, A: 85, 195. B: 6, 37, 204, 205 Nogo receptor, B: 148 Nogo66, B: 148 NOLZ1, A: 271, 272 Non-coding transcripts, A: 141 Non-specific effects, A: 1, 6–9, 21, 100, 103 Noradrenaline (NA), A: 2, 17, 18, 20, 22, 31. B: 19, 88, 89 Notch inhibitor (DAPT), B: 203 Notch signalling, A: 85. B: 145 Notch1, A: 85 Notochord, A: 269

279

280

Combined Index

NP-zones, A: 11 NT-3, A: 8, 108, 110. B: 18, 19, 45, 46, 113 NT-4, B: 18, 45, 46 Nucleus accumbens, A: 22, 26. B: 61 Nucleus basalis magnocellularis (NBM), A: 18–21 Nude mice, A: 103 Nude mouse, A: 24. B: 8, 36, 38, 39, 44, 123, 179, 248 Nude rat, A: 110, 200, 202, 291, 293, 298. B: 123, 248, 254 Nurr1, A: 60, 86, 140, 216 Nutrients, A: 74, 266. B: 201, 207, 226–228, 232 NYSTEM, A: 32, 33, 191–205. B: 182

O Observational studies, A: 172, 182, 183, 197, 218, 234, 235, 251, 253 Ocata. See Advanced Cell Technologies Occipital cortex, A: 25 Oct4, A: 84, 140, 179, 215, 268. B: 9, 38–40, 201 Ocular dominance, B: 66, 67 Ocular stem cells, B: 202 Oedema, A: 237, 239, 287 Off-the-shelf products, A: 152, 191, 198, 199. B: 205, 208 Olesoxime, B: 153 Olfactory bulb, A: 29. B: 15, 41, 62 Olfactory ensheathing cell grafts, A: 29. B: 41, 44 Olfactory ensheathing cells (OECs), A: 29. B: 41, 42, 44 Olfactory tubercle, A: 22, 26. B: 61 Olig2, A: 61, 77, 79, 81. B: 145, 180 Oligodendrocyte grafts, B: 165–182 Oligodendrocyte precursor cells (OPCs), A: 61, 74, 77. B: 7, 8, 11, 38, 42, 135–153, 166 Oligodendrocytes, A: 23, 61, 63, 74, 76, 77, 100–102, 195. B: 5, 6, 8, 9, 11, 14, 17, 35–44, 47, 72, 135–153, 165–182, 258 Oligodendrocytic progenitors, B: 165–182 Oncogenic mutation. See Genetic mutation Oocyte, A: 71, 156 Open-label studies, A: 134, 139, 169, 170, 180, 184, 220, 227, 266. B: 125 Opioids, B: 88, 98 Opsins, B: 203 Optic cup, B: 202, 204–206, 212 Optic nerve, B: 140, 150, 192 Optic neuritis, B: 148 Optic vesicle, B: 204–206 Optimisation, A: 3, 135, 139, 144, 181, 228, 267, 268, 271, 307. B: 74, 206, 207, 212, 256

Optogenetics, A: 17, 26, 205, 221, 288. B: 94, 95, 255, 258 Organ transplantation, A: 215, 250 Osteoblast, B: 4, 40 Outer nuclear layer, B: 192, 193, 197, 198, 211 Outer segment, B: 192, 197, 200, 205–207, 211, 227–229 b-Oxidation, A: 84 Oxidative metabolism, A: 84. B: 141, 227 Oxidative stress, A: 84. B: 227 Oxygen, A: 84, 266, 288. B: 201

P p53, A: 215 p75 receptor, B: 35, 122, 148 Paclitaxel, B: 74, 91, 92 Paediatric disorders, B: 167, 170–172, 174, 176, 182, 193 Pain, B: 21, 67, 73, 74, 87–101 Palmitic acid, A: 292 Parabiosis, B: 147 Paracrine signalling, A: 104, 269. B: 40, 47 Paranode, B: 136 Paraplegia, A: 29. B: 126 Parenchyma, A: 56, 160, 232, 249, 285, 287. B: 57, 59, 74, 171, 246, 256 Parietal cortex, A: 253 Parkinson’s disease, A: xxvi, 2, 3, 6, 8, 32, 60, 69, 99, 103, 104, 107, 109, 114, 115, 133–145, 158, 159, 162, 165–185, 191–205, 213–222, 228, 233, 235, 243–247, 249, 263, 283, 284, 286, 292–295, 297, 299, 305, 306, 314. B: xxvi, 58, 67, 69, 72, 150, 247 Parthenogenesis, A: 161 Parvalbumin, A: 87. B: 10, 59–62, 64–66, 68, 89, 93 Patch-clamp recording, B: 94, 206, 253–255 Patchy organisation, A: 11, 306 Pathogenesis, A: 1, 55, 86, 104, 155, 157, 160, 169, 175, 214, 267. B: 70, 122, 141, 179, 209, 211, 227 Pathophysiology, B: 72, 87, 97, 98, 191 Patient selection, A: 170, 181, 245 Patient stratification, A: 228, 246. B: 176 Patient understanding, A: 240, 244 Patient withdrawal, A: 244, 246 Pax2, B: 93 Pax6, A: 73, 79, 81, 200, 216, 270. B: 203 PDCD1, A: 104 PDE10A, A: 252, 294 PDEbrd1/rd1 mouse, B: 200, 210, 211 PDQ-39, A: 220 Pelizaeus-Merzbacher disease, B: 139, 170, 174

Combined Index

Pericyte, A: 86, 88. B: 256 Perineurial cells, B: 116, 119 Perineuronal net, B: 69 Peripheral blood lymphocytes, A: 104 Peripheral nerve, B: 108–112 Peripheral nerve grafts, A: 27, 30, 31, 232. B: 108–113, 123 Peripheral nerve injury, A: 30. B: 73, 89–91, 93, 97, 98, 100, 109, 110 Peripheral nerve-injury, B: 100 Peripheral nervous system (PNS), A: 27, 28, 195, 196. B: 41, 42, 46, 108, 109, 137 Peripheral vision, B: 193 Peripherin-2, B: 199 Periventricular leukomalacia, B: 170 Pfizer, A: 159, 160. B: 232 Phagocytosis, A: 88. B: 144, 147, 149, 227–229, 240 Pharmacological effects, A: 1, 6, 10, 294, 319. B: 87, 100 Phase I trials, A: 103, 104, 106, 108, 110, 113, 135–138, 155, 158, 160, 161, 175, 178, 181, 197, 199, 200, 220, 243. B: 122, 125, 126, 148, 151, 169, 171, 174, 195, 232, 246 Phase I/II trials, A: 139, 220. B: 195, 232 Phase II trials, A: 109, 110, 152, 155, 182, 192. B: 148 Phase II/III trials, A: 181 Phase III trials, A: 155, 176, 193, 205. B: 127, 150 Phase IV trials, A: 176 Phaseolus vulgaris leucoagglutinin (PhAL), B: 12 Phencyclidine, B: 69, 70 Phenotype, A: 25, 62, 157, 283, 284, 290, 292, 294, 299. B: 110, 150, 169, 170, 172, 203 Phosphocholine, A: 289 Phosphocreatine, A: 289, 290 Photoreceptor degeneration, A: 104. B: 191–212, 226, 229 Photoreceptors, A: xxvii, 104. B: xxvii, 191–212, 226–229, 232 Photoreceptor grafts. See Retinal grafts Photothrombotic lesion, B: 250 Phrenic nerve, B: 12 Phylogeny, B: 182 Physical training, A: 305–322 Physiotherapy, A: 305–322 PI3K-Akt pathway, B: 35 Pig, A: 180. B: 236. See also Minipig Pigment epithelium derived factor (PEDF), B: 238, 240 Pilocarpine, B: 71 Pilot trials, A: 227–229, 239, 243, 245–247, 249, 251–253 Pituitary extract, B: 114–116

Pitx3, A: 60, 140 PK11195, A: 220, 296 Placebo, A: 172, 234, 243. B: 123, 125 Placenta stem cells, A: 101. B: 3 Plasticity, A: 1, 2, 6–9, 15–17, 70, 76, 81, 106, 113, 114, 308, 310–313. B: 4, 19, 33–48, 57–74, 109, 110, 136, 178, 198, 245, 246, 252, 257. See also Neuroplasticity Platelet-derived growth factor (PDGF), B: 35, 111, 168, 194, 198 Platelet-derived growth factor receptor a, A: 77. B: 137, 151, 172, 180 Pluripotency, A: 56, 177, 195, 198. B: 169, 176, 201–203, 230, 238, 240, 248 Pluripotent stem cells (PSCs), A: 24, 53–63, 70, 99, 100, 102, 104, 134, 140, 142, 151–159, 161, 162, 165, 191–205, 213–222, 263–276, 308. B: 2, 4, 34, 37, 38, 74, 109, 111, 165, 167, 191–212, 225–40, 245–258 Pneumonia, A: 236, 238 Point-to-point systems, A: 3 Polarisation, A: 22. B: 226, 232, 233 Polyglutamine (CAG) repeats, A: 112, 246, 263, 265. B: 179, 180 Poly-lactic-co-glycolic acid (PLGA), B: 232 Poly-L-lysine, B: 115, 118–120 Polymerase chain reaction (RT-PCR), A: 140, 179, 201, 250. B: 46, 240 Positron emission tomography (PET), A: 171, 172, 182, 184, 204, 218, 220, 233–241, 243, 252, 283–300, 314, 315. B: 257. See also Radioligand Post mortem analysis, A: 27, 75, 168, 169, 184, 192, 220, 229, 232–234, 236, 241, 250, 252, 290 Post-mitotic cells, A: xxv, 24, 58, 59, 63, 73, 77, 79, 83, 84, 87, 274, 275, 308. B: xxv, 170 Post-traumatic stress disorder, B: 69 Potassium (K+) levels, B: 180 Potassium channels, B: 71 Potassium electrodes, B: 180 Potassium homeostasis, B: 178 Potency assays, A: 26, 155, 174, 176, 180. B: 127 Preclinical studies, A: 112, 185, 217, 300. B: 74 Precursors, A: 9–11, 25, 135, 139, 140, 193, 194, 241, 264, 270, 272, 306. B: 5, 6, 20, 41, 58, 63, 71, 87–89, 98, 100, 111, 196–198, 203. See also Progenitors PREDICT trial, A: 244 Prednisolone, A: 239 Prednisone, A: 204 Prefrontal cortex, A: 11, 14. B: 70. See also Frontal cortex Pre-manifest disease, A: 228. B: 172

281

282

Combined Index

Preoptic area, B: 60 Primary end point, A: 169, 172, 184. B: 125 Primate, A: 18, 105, 107, 108, 112, 116, 117, 141, 161, 168, 180, 191, 194, 196, 197, 202, 213, 214, 217, 218, 220, 221, 228, 265, 286, 306. B: 7, 9, 17, 18, 36, 37, 39, 59, 109, 124, 178, 211 Primitive neural stem cells, A: 161. B: 16 Prion, A: 169 PrKACA kinase, A: 57 Product manufacture, A: 135, 141, 151–155, 157, 160, 162, 171, 173, 176, 192, 198, 199, 205, 213, 215, 218, 221. B: 113, 121, 122, 124–127, 203, 208, 225, 234–237, 239, 240 Product release, A: 154, 155 Progenitors, A: 269. B: 70, 89, 135, 165–182. See also Precursors Programmed cell death, A: 104. B: 62, 65, 66 Programming, A: 274. See also Reprogramming; Specification Proliferation, B: 201, 253 Proof-of-concept studies, A: 63, 105, 110, 116, 117, 134, 173, 192, 194, 288. B: 36, 199, 210 Proof-of-principle, A: 30, 70, 71, 81, 83, 88, 115, 165, 266, 286, 307. B: 225, 229, 247 Prophylaxis, A: 249. B: 70, 73, 93 Proprio-spinal pathway, B: 17 Protein aggregate, A: 75, 235 Protein kinases, A: 57. B: 110 Proxies, A: 240, 245, 254 Pruning, A: 13, 75, 78. B: 21 Pruritogens, B: 88 PsA-NCAM, A: 205. B: 144, 145, 253 Pseudorabies virus, B: 12, 94 Psychiatric disorders, A: 112, 171, 182, 203, 232, 236, 251, 252, 315. B: 58, 73, 165, 178, 181, 182 Psychogenics, B: 180 Psychotic disorders, A: 236. B: 70 PTEN/mTOR pathway, B: 19 Pump, A: 26, 109, 166. B: 88, 89, 93, 97, 98 Pupillary reflex, A: 309. B: 200 Purification, B: 196 Purity, A: 135, 136, 140, 165, 175, 176, 179, 218. B: 117, 120, 121, 126, 167, 169, 230, 235, 236 Putamen, A: 4, 22, 109, 167, 169, 174, 181, 232, 234, 235, 237, 239 Pyramidal neurons, A: 24, 25, 84, 88. B: 57–59, 64, 65, 71, 178, 254 Pyramidal tract, A: 23 Pyruvate, B: 205 P-zones, A: 11, 12, 14, 232, 233, 275

Q Quality assurance, A: 135, 136. B: 125, 127, 234, 235, 237, 239, 240 Quality control, A: xxv, 33, 135, 145, 171, 174, 177–179, 198, 267, 274. B: xxv, 235, 237, 240 Quality of life, A: 252, 306, 315, 316, 322. B: 229, 258 Quality system, A: 135, 155 Questionnaires, A: 240, 254, 315, 317 Quietapine, A: 236 Quinolinic acid, A: 33, 265, 270, 271, 291, 296, 298

R R6/1 mouse, A: 308 R6/2 mouse, A: 308. B: 179, 180 Rabies virus, A: 17, 87, 218. B: 55 Radial maze, A: 11, 21 Radial-glia, B: 58, 59, 61 Radiation therapy, A: 104. B: 176 Radioligand, A: 292, 294, 296 [11C]-DPA713, A: 298 [11C]-Raclopride, A: 236, 237, 252, 293–295, 297 [11C]-SCH23390, A: 295 [11C]-SSR180575, A: 298 [18F]-DPA714, A: 297 [18F]-Fallypride, A: 293–295, 297 [18F]-FECNT, A: 295 [18F]-FluoroDOPA, A: 294, 297 [18F]-Fluorodeoxyglucose (FDG), A: 233, 234, 237–239, 241, 252, 294, 295, 297 [18F]-Fluorothymidine (FLT), A: 222 [18F]-Fluoro-m-tyrosine, A: 292, 295 [18F]-FP-CIT, A: 295 [18F]-LBT, A: 293, 295 [18F]-PBR28, A: 297 [123I]-DaTscan, A: 295 Ramon y Cajal, Santiago, A: 26. B: 1, 33 Randomisation, A: 228, 239–241, 244 Rapamycin, A: 114 Raphe nucleus, A: 5, 11, 12, 18. B: 16 Raphe-spinal pathway, B: 17 Rat, A: 2, 5, 11, 12, 15, 16, 18–29, 31, 56, 78, 81, 82, 104, 107–111, 113–115, 135, 140, 142, 161, 173, 174, 180, 194, 196, 198, 200, 202, 216, 217, 219, 247, 264, 266, 270–272, 286, 287, 289–291, 293, 294, 296, 298, 305–307, 309, 311. B: 5–8, 10–16, 18–20, 35, 36, 38, 41, 72, 107–109, 112–118, 123, 124, 128, 150, 176, 177, 197, 229, 232, 247–254 Reaching test, B: 36

Combined Index

Reactive astrocytes, A: 69–89. B: 208 Reactive gliosis, A: 71, 74, 78, 79 Reactive oxygen species (ROS), A: 78, 84 Reading, B: 226 Recessive disorders, B: 194, 211 Recruitment, B: 135–153 Reelin (RLN), B: 60, 62–65 Regeneration, A: xxvi, xxvii, 2, 4, 6, 7, 9, 24, 26–31, 61, 70, 75, 165, 192, 205, 222, 263, 275, 276, 284, 285, 300, 308, 319–321,. B: xxvi, xxvii, 1, 3, 7, 10, 11, 17–21, 33–37, 39, 41–48, 109, 112, 118, 123, 135–137, 141, 142, 148, 149, 151, 192, 234, 240, 252 Regenerative medicine, A: xxvi, xxvii, 146, 192, 205, 222, 263, 275, 276, 285, 300, 319, 321. B: xxvi, xxvii, 39, 234, 240 Regenerative sprouting, A: 4, 8 Registry, A: 138, 244 Regulation, A: 151–162, 173. B: 230, 234, 235 Rehabilitation, A: 29, 213, 222, 254, 305–322. B: 21, 47, 48, 123, 125 Reinnervation, A: 1–34 Rejection. See Graft rejection Rejuvenation, B: 147 Relapse rate, B: 175 Relevant Communicable Disease Agents and Diseases (RCDADs), A: 155 Reliability, A: xxv, 246, 267, 315 Remyelination, A: 29. B: 1, 7, 11, 14, 19, 33–48, 109, 113, 135–153, 176, 258 Repair-HD, A: 33, 34, 252, 274, 277, 300 Reprogramming, A: 53–63, 69–89, 138, 143, 144, 154, 250, 268. B: 4, 9, 10, 38, 39, 169, 245–258 Reticular system, A: 3, 23. B: 3 Reticulo-spinal pathway, B: 3, 17, 20 Retina, A: 75. B: 191–212, 225–240 Retinal degeneration, A: 104. B: 192–194, 197–200, 210, 236 Retinal detachment, B: 193, 226, 228 Retinal dystrophies, B: 191–212 Retinal ganglion cell (RGC), A: 74. B: 192, 195, 197 Retinal grafts, B: xxvi, xxvii, 191–212 Retinal imaging, B: 230 Retinal organoids, B: 196, 205–211 Retinal pigment epithelium (RPE), A: xxvi, xxvii, 34, 161, 193. B: xxvi, xxvii, 191–212, 225–240 Retinal pigment epithelium grafts, A: xxvii, 167. B: xxvii, 225–240 Retinal progenitor cells (RPCs), B: 197. B: 198, 203, 205, 207 Retinal prosthesis, B: 192. B: 195

Retinitis pigmentosa, A: 104. B: 193 Retinogenesis, B: 192, 205, 206 Retinoic acid, A: 196, 269. B: 37, 149, 203–206 Retinoid isomerohydrolase, B: 195 Retinoid X nuclear receptors (RXR), B: 148, 149 Retrograde degereration, B: 33 Retrograde tracing, A: 12, 13, 17, 22, 24, 27, 87. B: 12, 13, 94, 254 Retrovirus, A: 79, 81, 82, 84, 86, 102, 103, 139, 143, 215. B: 12 Rhesus monkey, A: 196 Rhodopsin, A: 26, 104. B: 199 Risk factor, B: 21, 73, 125, 140, 193, 235 Risk of falling, A: 232, 239, 248, 316 Rituximab, B: 175 RNAseq, A: 179. B: 35 ROCK inhibitor, A: 177 Rod dystrophy, B: 194 Rod sensitivity, B: 195 Rods, B: 193–195, 197–200, 207, 210 Rope-climbing test, B: 250, 252 Roslin Cells, A: 137, 139 Roslin Cells (RCn) cell lines, A: 137, 138, 158, 159, 175 Rostralisation, B: 203, 205, 206 Rotarod, A: 270. B: 39, 180, 249–252 Rotation, A: 8, 11, 22, 30, 173, 179, 196, 200, 247, 272. B: 249, 252 Royal College of Surgeons (RCS) rat, B: 229, 232 RPE65 gene, B: 195 Rubro-spinal pathway, B: 17 RXR agonists, B: 148, 149, 153

S S100 protein, B: 112, 122, 251 Safety, A: xxv, xxvi, 7, 29, 54, 55, 58, 62, 88, 103, 104, 108, 110–113, 115–117, 133, 135, 136, 140–145, 151–162, 169, 174–179, 181–185, 191, 192, 199–205, 216, 218, 220, 227, 229, 232, 235, 240, 243, 244, 250, 254, 266, 273, 284, 286, 288, 321. B: xxv, xxvi, 4, 8, 10, 21, 34, 38–40, 74, 100, 110, 113, 116, 122, 124–127, 148–151, 167, 169, 171, 174, 177, 230–237, 240, 246, 248 Saltatory conduction, B: 136–138 Sandhoff strain, B: 171 Saturated fatty acids (SFAs), A: 292 Scaffolds and matrices, A: 34, 161. B: 13, 14, 21, 42, 109, 232, 233, 236, 239 Scar formation, A: 75. B: 226 Schizophrenia, B: 58, 64, 67, 69, 70, 166, 182 Schwann cell grafts, B: 41, 42, 44, 107–128

283

284

Combined Index

Schwann cells, A: 27–30, 61, 62. B: 19, 41, 42, 44, 107–128, 137, 151 Sciatic nerve, A: 30, 111. B: 20, 74, 91, 94, 114–116, 118, 123 Sciatic nerve lesion, B: 74. B: 91, 94, 123 SCID mouse. See Nude mouse SCOPE, the Spinal Cord Outcomes Partnership Endeavor, B: 122 ScoreCard assay, B: 238 Scratching, B: 97–100 Secondary degeneration, B: 246 Seizure, A: 114. B: 70–72, 89, 90 Self-renewal, A: 55, 77, 101, 141, 143, 269. B: 36, 109, 111, 137, 167, 238 Semaphorins (SEM3A, SEM3F), B: 144 Senescence, A: 275. B: 118 Sensitisation, B: 88 Sensorimotor function, A: 16, 113. B: 41 Sensorimotor neglect, A: 16 Sensory deficits, B: 245, 257 Sensory function, B: 252, 258 Sensory ganglia, B: 20, 113, 114 Sensory neglect, A: 16 Sensory pathways, A: 26. B: 17, 19, 20, 46, 47, 113 Sensory placode, A: 195 Septal grafts, A: 19, 20 Serious Adverse Events. See Adverse events Serotonin, A: 2, 17, 18, 20, 22, 23, 107, 141, 170, 171, 179–181, 216, 294. B: 8, 16, 17, 19, 59, 88 Serum, A: 198, 250, 269. B: 37, 110, 115, 117–119, 121, 126, 147, 150, 151, 201, 203 Service Level Agreement, B: 237 Seural nerve grafts, A: 232 Sham surgery, A: 108, 172, 243–245 Sheep, A: 265 Shiverer mouse, B: 11, 43, 143, 172, 173 Side effects, B: 88, 90. See Adverse events Simulect, A: 204 Singapore, A: 138 Single nucleotide polymorphisms, A: 133–145, 177 Single-photon emission computed tomography (SPECT), A: 237, 314 Skin dermatomes, B: 99 Slice preparation, A: 13, 15, 16, 26, 311–313. B: 41, 60, 63, 94, 95, 150, 253–255 SMAD signaling, A: 58, 140, 195, 215, 270, 272, 273, 276 Small molecules, A: 57, 58, 274. B: 4, 232, 234 SmartCube test, B: 180 Smoking, B: 193, 226 Societal issues, A: 166, 168, 172. B: 245 Solid grafts, A: 242. B: 5

Somatic cell reprogramming, A: 53–63, 71, 134, 138, 143, 153, 274, 275. B: 2, 4, 8–10, 20, 38, 169, 245–258 Somatic stem cells, A: 61. B: 142, 234, 246 Somatosensory function, A: 26. B: 1, 47, 123 Somatostatin, A: 241. B: 59–68, 73, 89, 93 Sonic hedgehog (Shh), A: 24, 25, 139, 194, 196, 217, 271–274, 277. B: 19, 37, 203, 204 SOX gene family Sox1, A: 216 Sox2, A: 61, 84–86, 215, 268. B: 9, 14, 38, 39, 201 Sox6, A: 140. B: 60 Sox9, A: 62 Sox10, A: 61, 62, 77 Sox11, A: 61, 77 Spasticity, B: 67, 126 Specification, A: xxv, 25, 59–61, 63, 71, 89, 195, 215, 216, 248, 254, 272–276. B: xxv, 57, 111, 112, 145, 201, 236, 237, 239 Sperm, A: 156 Spheramine, A: 167 Sphingomyelinase, B: 171 Sphingosine-1-phosphate receptor, B: 150, 175 Spinal cord, A: 3, 7, 9, 10, 17, 18, 22–24, 27–31, 34, 61, 75, 77, 81, 84, 85, 88, 102, 104, 105, 110, 111, 137, 159–161, 181, 196, 201, 216, 269. B: 1–22, 33–48, 63, 67, 73, 74, 87–101, 107–128, 140, 144, 145, 166, 167, 172, 174, 176, 256 Spinal cord grafts, A: 30. B: 1–22, 33–48, 74, 87–101, 107–128, 177 Spinal cord injury, A: 7, 9, 27–29, 31, 34, 75, 81, 137, 159, 160, 181. B: 1–22, 33–48, 107–128, 140, 144, 166, 169, 176, 177 Spinal cord repair, B: 3–22, 33–48 Spinal cord transection, A: 23, 27–29, 31. B: 6, 8, 13, 14, 17, 19, 20, 91, 112, 118, 123 Spin-lattice weighting, A: 286 Spinomuscular atrophy (SMA), A: 100, 103, 104 Spontaneous activity, A: 13, 20, 85. B: 63, 70, 71, 90, 97, 98, 254, 255 Spontaneous recovery, A: 200, 202. B: 123, 246, 252, 257 Sprouting, A: 4, 6–8, 26, 29, 308. B: 19, 46, 112. See also Collateral sprouting; Regenerative sprouting SSEA-1 progenitor cells, A: 161 Stab wound, A: 79, 82, 84, 86. B: 256 Staircase test, B: 249 Standard operating procedure (SOP), A: 135, 136, 154, 192, 199–201. B: 121, 124, 125, 239 Stargardt’s disease, A: 137, 159, 161. B: 193, 231, 232

Combined Index

Stem cell bank, A: 133, 136, 139, 144, 153, 158, 161, 205, 215, 266, 267, 273. B: 208, 231, 235 Stem cell differentiation protocol, A: 24–26, 32–34, 133, 135–140, 144, 173, 174, 176–178, 181, 192, 194–-199, 213, 216, 218, 220, 268–272, 275, 283. B: 11, 173, 201, 203–208, 210, 211, 239, 240, 245–258 Stem cell grafts, A: xxv–xxviii, 9, 24–26, 29, 31–34, 101–117, 158–162, 173–175, 180–183, 191–205, 213–222, 267, 268, 283, 284. B: xxv–xxviii, 1–22, 33–48, 165–182, 232–240, 245–258 Stem cell lines, A: xxv, xxvi, 70, 133, 137, 138, 142, 156, 158, 159, 174, 175, 197. B: xxv, xxvi, 246 Stem cell tourism, A: xxvii, 9, 33 Stem121, B: 251 STEM-PD, A: 165, 171, 174, 175, 178–180, 183 Stepping test, A: 196. B: 249, 250, 252 Stereotactic surgery, A: 105, 204, 220, 248, 249, 306. B: 124 Sterility, A: 140, 155, 178, 179, 198. B: 121 Steroids, A: 172, 184, 239, 254 Storage, A: 135, 168, 179, 245, 267. B: 207, 208 Stress, A: 100, 117, 317. B: 69, 211 Striatal grafts, A: 1–5, 7, 8, 10–17, 20, 22, 23, 25, 26, 30, 31, 33, 34, 72, 85, 87, 107, 109, 112, 115, 134, 139, 165, 168, 170, 171, 180, 183, 196, 197, 218, 219, 227–255, 263–276, 286, 287, 291, 294–299, 305–307, 309–316, 322. B: 72, 179, 180, 249–251, 255. See also Ganglionic eminence grafts Striatal lesions, A: 2, 11, 14–16, 109, 291, 296, 298, 309, 311 Striatal topography, A: 170 Striatum, A: 2–5, 10–16, 18, 22–26, 30, 78, 81–87, 104, 107–109, 112, 114, 115, 133, 134, 139, 144, 161, 165–170, 180, 203, 219, 228, 229, 233, 235, 237, 247–254, 263–276, 286, 287, 289–291, 293, 294, 298–300, 305, 306, 309–313, 315, 316. B: 60, 61, 67, 72, 179, 180, 254. See also Basal ganglia; Caudate nucleus; Putamen Stroke, A: xxvii, 9, 34, 70, 81, 82, 85, 99, 113, 114, 309, 313, 315. B: xxvii, 70, 166, 175, 245–258 Stromal cells, A: 111, 195, 217, 269, 270. B: 2, 40, 111, 172 Subdural haematoma, A: 228, 233, 234, 237, 240, 245, 248 Subpallium, A: 272, 275. B: 59, 60, 64 Substantia nigra, A: 4, 11, 12, 22, 24, 25, 30, 61, 107, 108, 139, 162, 166, 306. B: 72

Subthalamic nucleus, A: 166, 306 Subventricular zone (SVZ), A: 82, 85. B: 7, 36, 59, 137, 139, 166, 253 Suicide, A: 197, 232, 245 Sulfatide clearance, B: 171 Superior colliculus, B: 200 Superoxide dismutase (SOD1), A: 110, 113 Sural nerve, B: 122 Surgical termination of pregnancy (STOP), A: 171, 267. See also Abortion Survival. See Graft survival Sweden, A: xxvii, 1, 34, 165. B: 245 Synapsin, A: 73 Synaptic connections, A: 2, 5, 6, 13, 15–18, 21, 23, 26, 29, 31, 56, 57, 73, 75–79, 83, 85, 88, 89, 106, 114, 167, 218, 219, 221, 222, 273, 276, 293–295, 297, 299, 307, 308, 310–313, 319. B: 1–22, 33–48, 57, 63–66, 68, 71, 73, 74, 89, 94–96, 178, 192, 193, 196, 200, 211, 245, 246, 252, 254, 255 Synaptic remodelling, A: 78, 96, 190, 191, 209 Synaptic transmission, A: 13, 20, 26, 75, 89, 311. B: 20, 69, 73, 136, 137, 140 Synaptogenesis, A: 309, 310. B: 17, 57, 65, 244 Synaptophysin, B: 16, 18 Synovial fluid stem cells, A: 101 Synthetic mRNA, A: 54, 55, 138, 143 a-Synuclein, A: 169 Systematic review, A: 134. B: 40

T T cell leukemia, A: 103, 104 T1-weighting, A: 285–287 T2-weighting, A: 285–287, 291, 298, 300 Target support, A: 6 Tau, A: 105 Taurine, B: 203–206 TBR1, B: 254 T-cells, A: 103, 104, 116, 233, 249. B: 149 Tcf4, B: 145, 146 TDP43, A: 110. B: 178 Temperature control, A: 247 Temporal factors, A: 11, 69, 75, 85, 114, 195, 284. B: 37, 59 Temporal lobe epilepsy, B: 71 Teratoma, A: 141, 200, 201, 273. B: 8, 231, 235 Terflunamide, B: 175 Terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL), B: 249, 250 Tetracycline, A: 114, 115 Tetraplegia, B: 122 Thalamo-cortical connections, B: 255

285

286

Combined Index

Thalamus, A: 3, 4, 7, 11–13, 23, 24, 306. B: 66 Theiler’s murine encephalomyelitis virus (TMV), B: 141, 151 Therapeutic time window, A: 113, 267, 308. B: 34–36, 228, 246 Thrombectomy, B: 245 Thromboembolism, B: 245 Thrombolysis, B: 245 Thy1, A: 106. B: 115 Thymidine, A: 85 Tibial nerve, B: 91 Tight junctions, B: 206, 228 Tissue banking, A: See Bone bank; Brain bank; Cell bank; Master cell bank; Stem cell bank; Working cell bank Tissue dissection, A: 11, 171, 242, 246, 254, 264, 266. B: 117, 239 Tissue donor, A: 151–159, 161, 162, 168, 175, 307. B: 231 Tissue engineering, B: 234, 235 TLR2 receptors, B: 144 T-maze tasks, A: 19 a-Tocotrienol, A: 80 Tolerability, A: 181, 184. B: 148, 232 Tolerance (immunological), B: 151 Toluidine blue, B: 138 Tooth pulp stem cells, A: 101 Topographic organisation, B: 91 Toxicity, A: 108, 110, 157, 158, 160, 200, 201, 290. B: 73, 123, 124, 170, 171, 194, 228 Toxicology, A: 179, 201, 218. B: 124, 235 TRACK-HD trial, A: 244 Tracking. See Graft tracking Training, A: 16, 135, 240, 246, 251, 305, 307–310, 313–316, 319–321. B: 21, 47, 69, 124, 127, 147, 237 Transcranial magnetic stimulation (TMS), A: 253 Transcription activator-like effector nucleases (TALENs), A: 55, 103, 104 Transcription factors, A: 25, 53, 55, 56, 58, 59, 69, 71–73, 77, 79, 82–84, 86, 87, 89, 103, 106, 140, 215, 268–271, 274–276. B: 2, 4, 9, 35, 45, 60, 98, 99, 143, 145, 148, 149, 169, 176, 197, 201, 208, 247, 255, 256. See also individual factors by name Transepithelial resistance, B: 240 TRANSEURO, A: 34, 134, 139, 165–185 Transfer-of-training, A: 16 Transforming growth factor (TGF-b), A: 107, 113, 139, 195, 215, 271–273. B: 35, 46 Transgenic mouse, A: 23, 80, 83, 86, 106, 265, 308. B: 12, 70, 99, 100, 179, 180, 182, 196–198, 210

Transgenic rat, A: 23. B: 13 Translocator protein (TSPO), A: 296–298, 300 Trans-neuronal labelling, B: 12 Transplant overgrowth. See Graft overgrowth Transplants (various). See Grafts Transportation, A: 135, 144. B: 126, 239 Transynaptic tracing, B: 255 Traumatic injury, A: 6, 7, 9, 34, 70, 75, 77, 78, 299, 308. B: 7, 40, 69, 70, 90, 91, 98, 122, 139, 140, 170, 174, 175, 193 Treadmill, B: 39, 47 Trial design, A: xxvi, 32, 134, 169, 170, 172, 180–182, 205, 229, 239, 252 Trial safety committee, A: 250 Trigeminal neuralgia, B: 90 Trimethoprim (TMP), A: 115 Triple labelling, B: 15, 16 Triple therapy, A: 172, 233, 250. See also Immunosuppression Trisomy, A: 142 TrkB receptors, A: 311. B: 65 TrkC receptors, B: 19 Trophectoderm, A: 195 Trophic effects, A: 6. B: 120 Trophic factors, A: 6, 8, 30, 108–110, 112, 228, 251. B: 21, 36, 42, 43, 173, 257 Trophic grafts, A: 6–8, 10, 27, 29, 30, 102, 103, 106, 108–115. B: 33–43, 45–48, 246, 252 Trophic support, B: 72, 73, 136 Trypan blue, A: 247 Trypsinisation, B: 120 b-Tubulin, A: 73, 270. B: 44, 249, 250 Tuj1, B: 6, 15, 249 Tumorigenesis, B: 115, 169, 170, 173 Tumorigenicity, A: 117, 157, 158, 160, 179, 200, 201, 218. B: 34, 39, 231, 235, 247, 248, 256 Tumour necrosis factor (TNF)a, B: 35, 40, 249, 251, 253 Tumours, A: 25, 77, 86, 140, 141, 157, 158, 160, 161, 178, 200, 201, 213, 215–218, 220, 235, 288, 300. B: 9, 10, 21, 38–40, 70, 100, 116, 123, 124, 210, 247, 248, 252, 255 Tyrosine hydroxylase (TH), A: 21–23, 108, 114, 115, 180, 184, 194, 200, 220, 232, 233, 242. B: 249, 250 Tyrosine kinase, B: 110, 229 Tyrosine phosphorylation, B: 111

U UK Stem Cell Bank, A: 138 Ultrasound guidance, B: 61 Umbilical cord stem cells, A: 101, 139. B: 3, 172

Combined Index

Unified Huntington’s disease rating scale (UHDRS), A: 229, 236, 253, 315, 318 Unified Parkinson’s disease rating scale (UPDRS), A: 108, 172, 182, 183, 220 United Kingdom, A: 1, 34, 104, 133, 159, 165, 230, 231, 235–237. B: 191, 225, 226 United States, A: 34, 104, 110, 135, 151, 154, 159, 175, 185, 191, 214, 229–231, 234. B: 1, 57, 87, 107, 122, 165, 170, 175, 226, 232, 234 Sheffield University (Shef and MasterShef ) cell lines, A: 137, 138, 158, 159, 175

V Vacuolation, B: 140 Validation, A: 33, 34, 54, 135, 173, 179, 192, 199, 200, 240, 283, 284, 290, 298, 319. B: 148, 212, 236, 237, 240 Valproic acid, B: 6, 19, 45 Vanishing white matter disease, B: 171 Vascular damage, A: 241 Vascular endothelial growth factor (VEGF), A: 8, 110, 111, 113. B: 46, 194, 226, 249, 253 Vascularisation, A: 233–235, 245. B: 193, 194, 226, 229, 233, 253 Vasoactive intestinal polypeptide (VIP), B: 62 Vasopressin, A: 2 Ventral forebrain, A: 11, 269, 272. B: 59, 62–64, 180 Ventral horn, B: 45, 91 Ventral mesencephalic grafts. See Nigral grafts Ventral mesencephalon (VM), A: 4, 25, 26, 30, 165–185 Ventralisation, A: 215, 269–271, 275. B: 37 Vesicular GABA transporter (VGAT), B: 74, 91, 92 Vesicular transporter (VAT), A: 293 Viability, A: 116, 140, 144, 155, 171, 179, 191, 200, 247, 252, 266, 289, 291, 292. B: 121, 126, 127, 175 Viacyte (CyTn) cell lines, A: 137, 138, 158–161 Vimentin, A: 75, 76. B: 253 VIP, B: 60, 63 Viral integration, A: 215. B: 208 Visual acuity, B: 66, 191, 194, 198, 200, 232 Visual cortex, A: 25. B: 66, 67, 69, 70, 197 Visual deprivation, B: 69 Visual field, B: 192

Visual function, A: 104. B: 191, 196, 197, 199, 200, 210, 211, 227, 229, 230 Visual system, A: 3. B: 66 Visually guided mobility, B: 195 Vitamin A, B: 228 Vitamin D, A: 84. B: 149 Vitamin E, A: 84 Vitronectin, A: 140

W

Waisman Biomanufacturing. See WiCell H9 cell line Wallerian degeneration, B: 139 Western blot, B: 240 WGA-HRP, A: 12, 13. B: 11, 12, 45, 94 White matter, A: 22, 23, 29, 288, 308. B: 1, 3, 12–15, 44, 136–138, 141, 143, 166, 170, 172, 174, 175, 177, 181, 182 Whole ganglionic eminence (WGE), A: 4, 11, 12, 30, 242, 246, 264–266, 306, 314 WiCell (WAn or Hn) cell lines, A: 137, 139, 144, 156, 158, 159, 197 Wisconsin Card Sorting Test, A: 236 Withdrawal reflex, B: 91, 97 Wnt signaling, A: 24, 25, 61, 139, 195, 196, 215, 269, 270. B: 37, 145, 146, 203, 205, 248 Wnt3a, B: 37, 248 Working cell bank, A: 152, 153, 160, 197–199, 204 Working memory, A: 11, 14, 317 Wound healing, A: 74, 76, 77

X Xeno-free reagents, A: 138, 140, 144, 161, 217. B: 206, 208 Xenogeneic pathogens, A: 157, 160. B: 235 Xenogeneic products, A: 158, 160 Xenografts, A: 12, 116, 157, 160, 174, 194, 201–203, 296. B: 12, 14, 177, 179 X-linked severe combined immunodeficiency (SCID). See Nude mice

Z Zinc finger nucleases (ZFNs), A: 55, 103, 104 Zonisamide, A: 218 Zoonosis, B: 208

287