Brain Development: Methods and Protocols [2nd ed. 2020] 978-1-4939-9731-2, 978-1-4939-9732-9

Table of contents :
Front Matter ....Pages i-xiv
Front Matter ....Pages 1-1
Combining BrdU-Labeling to Detection of Neuronal Markers to Monitor Adult Neurogenesis in Hydra (Wanda Buzgariu, Marie-Laure Curchod, Chrystelle Perruchoud, Brigitte Galliot)....Pages 3-24
Reverse Genetic Approaches to Investigate the Neurobiology of the Cnidarian Sea Anemone Nematostella vectensis (Jamie A. Havrilak, Michael J. Layden)....Pages 25-43
Generating Transgenic Reporter Lines for Studying Nervous System Development in the Cnidarian Nematostella vectensis (Fabian Rentzsch, Eduard Renfer, Ulrich Technau)....Pages 45-57
Immunostaining and In Situ Hybridization of the Developing Acoel Nervous System (Elena Perea-Atienza, Brenda Gavilán, Simon G. Sprecher, Pedro Martinez)....Pages 59-80
Immunostaining of the Embryonic and Larval Drosophila Brain (Frank Hirth, Danielle C. Diaper)....Pages 81-96
Nonfluorescent RNA In Situ Hybridization Combined with Antibody Staining to Visualize Multiple Gene Expression Patterns in the Embryonic Brain of Drosophila (David Jussen, Rolf Urbach)....Pages 97-113
Analysis of Complete Neuroblast Cell Lineages in the Drosophila Embryonic Brain via DiI Labeling (Karoline F. Kraft, Rolf Urbach)....Pages 115-135
Flybow to Dissect Circuit Assembly in the Drosophila Brain: An Update (Emma L. Powell, Iris Salecker)....Pages 137-152
Live Cell Imaging of Neural Stem Cells in the Drosophila Larval Brain (Karolina Miszczak, Boris Egger)....Pages 153-160
CRISPR/Cas9 Genome Editing to Study Nervous System Development in Drosophila (Cornelia Fritsch, Simon G. Sprecher)....Pages 161-189
The Red Flour Beetle as Model for Comparative Neural Development: Genome Editing to Mark Neural Cells in Tribolium Brain Development (Max S. Farnworth, Kolja N. Eckermann, Hassan M. M. Ahmed, Dominik S. Mühlen, Bicheng He, Gregor Bucher)....Pages 191-217
A Protocol for Double Fluorescent In Situ Hybridization and Immunohistochemistry for the Study of Embryonic Brain Development in Tribolium castaneum (Marita Buescher, Georg Oberhofer, Natalia Carolina Garcia-Perez, Gregor Bucher)....Pages 219-232
Immunohistochemistry and Fluorescent Whole Mount RNA In Situ Hybridization in Larval and Adult Brains of Tribolium (Vera S. Hunnekuhl, Janna Siemanowski, Max S. Farnworth, Bicheng He, Gregor Bucher)....Pages 233-251
X-Ray Microscopy of the Larval Crustacean Brain (Jakob Krieger, Franziska Spitzner)....Pages 253-270
Immunolocalization of Neurotransmitters and Neuromodulators in the Developing Crayfish Brain (Steffen Harzsch, Caroline Viertel)....Pages 271-291
Immunostainings in Nervous System Development of the Nematode C. elegans (Janet S. Duerr)....Pages 293-310
Methods in Brain Development of Molluscs (Andreas Wanninger, Tim Wollesen)....Pages 311-324
A Simple Method to Identify Ascidian Brain Lineage Cells at Neural Plate Stages Following In Situ Hybridization (Clare Hudson)....Pages 325-345
Spawning Induction and Embryo Micromanipulation Protocols in the Amphioxus Branchiostoma lanceolatum (Yann Le Petillon, Stéphanie Bertrand, Héctor Escrivà)....Pages 347-359
Front Matter ....Pages 361-361
In Situ Hybridization and Immunostaining of Xenopus Brain (Kai-li Liu, Xiu-mei Wang, Zi-long Li, Ying Liu, Rong-qiao He)....Pages 363-375
Morpholino Studies in Xenopus Brain Development (Jennifer E. Bestman, Hollis T. Cline)....Pages 377-395
Sensitive Multiplexed Fluorescent In Situ Hybridization Using Enhanced Tyramide Signal Amplification and Its Combination with Immunofluorescent Protein Visualization in Zebrafish (Gilbert Lauter, Iris Söll, Giselbert Hauptmann)....Pages 397-409
Live Morphometric Classification of Sensory Neurons in Larval Zebrafish (Gema Valera, Hernán López-Schier)....Pages 411-419
Immunohistochemistry and In Situ Hybridization in the Developing Chicken Brain (Richard P. Tucker, Tatsuto Ishimaru, Qizhi Gong)....Pages 421-437
Gene Silencing in Chicken Brain Development (Georgia Tsapara, Irwin Andermatt, Esther T. Stoeckli)....Pages 439-456
Transplantation of Neural Tissue: Quail–Chick Chimeras (Andrea Streit, Claudio D. Stern)....Pages 457-473
Immunohistochemistry and RNA In Situ Hybridization in Mouse Brain Development (Jinling Liu, Aimin Liu)....Pages 475-489
The Cre/Lox System to Assess the Development of the Mouse Brain (Claudius F. Kratochwil, Filippo M. Rijli)....Pages 491-512
In Utero Electroporation to Study Mouse Brain Development (Emilie Pacary, François Guillemot)....Pages 513-523
Back Matter ....Pages 525-527

Citation preview

Methods in Molecular Biology 2047

Simon G. Sprecher Editor

Brain Development Methods and Protocols Second Edition

Methods

in

M o l e c u l a r B i o lo g y

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, UK

For further volumes: http://www.springer.com/series/7651

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible stepby-step fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of theMethods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.

Brain Development Methods and Protocols Second Edition

Edited by

Simon G. Sprecher Department of Biology, University of Fribourg, Fribourg, Switzerland

Editor Simon G. Sprecher Department of Biology University of Fribourg Fribourg, Switzerland

ISSN 1064-3745     ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-4939-9731-2    ISBN 978-1-4939-9732-9 (eBook) https://doi.org/10.1007/978-1-4939-9732-9 © Springer Science+Business Media, LLC, part of Springer Nature 2020 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, express or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 233 Spring Street, New York, NY 10013, U.S.A.

Preface How the brain works remains one of the largest enigmatic questions in modern biology. Particularly, its cellular diversity, connectivity among neurons, formation of neuronal networks, use of distinct neurotransmitter systems, and underlying function for behavior raise the question of how this highly interconnected organ develops. It is therefore not surprising that the intersection between developmental biology and neuroscience provides an exceptional field to address and investigate impacting biological questions. Complementing findings of an array of distinct animal model systems provide the basis of brain development research. Our current understanding is based on widely used genetic model systems, including the fruit fly, zebra fish, chicken, and mouse. These animal models are particularly impacting since they allow elaborate genetic manipulations including transgenic expression systems and conditional knockout or knockdown of developmental genes. However, the advent of genome editing techniques makes it possible to extend investigations towards other animals that may thus be used to answer specific questions of molecular neuroscience. These noncanonical experimental systems further substantiate general principles and mechanisms in brain development. Questions that can be investigated often depend on the methodological accessibility. Therefore, progress and developments in constantly improving laboratory technologies provide an essential ground for the advancement in the field. This book aims to provide a comprehensive overview and introduction to widely used leading-edge techniques on a representative range of animals. A particular focus lies on recent technical advances in molecular genetics. Fribourg, Switzerland

Simon G. Sprecher

v

Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . v Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . xi

Part I Invertebrate Models 1 Combining BrdU-Labeling to Detection of Neuronal Markers to Monitor Adult Neurogenesis in Hydra . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .   3 Wanda Buzgariu, Marie-Laure Curchod, Chrystelle Perruchoud, and Brigitte Galliot 2 Reverse Genetic Approaches to Investigate the Neurobiology of the Cnidarian Sea Anemone Nematostella vectensis . . . . . . . . . . . . . . . . . . . . . . .  25 Jamie A. Havrilak and Michael J. Layden 3 Generating Transgenic Reporter Lines for Studying Nervous System Development in the Cnidarian Nematostella vectensis . . . . . . . . . . . . . . . . . . . . . . .  45 Fabian Rentzsch, Eduard Renfer, and Ulrich Technau 4 Immunostaining and In Situ Hybridization of the Developing Acoel Nervous System . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .  59 Elena Perea-Atienza, Brenda Gavilán, Simon G. Sprecher, and Pedro Martinez 5 Immunostaining of the Embryonic and Larval Drosophila Brain . . . . . . . . . . . . . . .  81 Frank Hirth and Danielle C. Diaper 6 Nonfluorescent RNA In Situ Hybridization Combined with Antibody Staining to Visualize Multiple Gene Expression Patterns in the Embryonic Brain of Drosophila . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .  97 David Jussen and Rolf Urbach 7 Analysis of Complete Neuroblast Cell Lineages in the Drosophila Embryonic Brain via DiI Labeling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115 Karoline F. Kraft and Rolf Urbach 8 Flybow to Dissect Circuit Assembly in the Drosophila Brain: An Update . . . . . . . . . 137 Emma L. Powell and Iris Salecker 9 Live Cell Imaging of Neural Stem Cells in the Drosophila Larval Brain . . . . . . . . . 153 Karolina Miszczak and Boris Egger 10 CRISPR/Cas9 Genome Editing to Study Nervous System Development in Drosophila . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 161 Cornelia Fritsch and Simon G. Sprecher 11 The Red Flour Beetle as Model for Comparative Neural Development: Genome Editing to Mark Neural Cells in Tribolium Brain Development . . . . . . . . 191 Max S. Farnworth, Kolja N. Eckermann, Hassan M. M. Ahmed, Dominik S. Mühlen, Bicheng He, and Gregor Bucher

vii

viii

Contents

12 A Protocol for Double Fluorescent In Situ Hybridization and Immunohistochemistry for the Study of Embryonic Brain Development in Tribolium castaneum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 219 Marita Buescher, Georg Oberhofer, Natalia Carolina Garcia-Perez, and Gregor Bucher 13 Immunohistochemistry and Fluorescent Whole Mount RNA In Situ Hybridization in Larval and Adult Brains of Tribolium . . . . . . . . . . . . . . . . . . . . . . 233 Vera S. Hunnekuhl, Janna Siemanowski, Max S. Farnworth, Bicheng He, and Gregor Bucher 14 X-Ray Microscopy of the Larval Crustacean Brain . . . . . . . . . . . . . . . . . . . . . . . . . 253 Jakob Krieger and Franziska Spitzner 15 Immunolocalization of Neurotransmitters and Neuromodulators in the Developing Crayfish Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271 Steffen Harzsch and Caroline Viertel 16 Immunostainings in Nervous System Development of the Nematode C. elegans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 293 Janet S. Duerr 17 Methods in Brain Development of Molluscs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 311 Andreas Wanninger and Tim Wollesen 18 A Simple Method to Identify Ascidian Brain Lineage Cells at Neural Plate Stages Following In Situ Hybridization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 325 Clare Hudson 19 Spawning Induction and Embryo Micromanipulation Protocols in the Amphioxus Branchiostoma lanceolatum . . . . . . . . . . . . . . . . . . . . . . . . . . . . 347 Yann Le Petillon, Stéphanie Bertrand, and Héctor Escrivà

Part II Vertebrate Models 20 In Situ Hybridization and Immunostaining of Xenopus Brain . . . . . . . . . . . . . . . . . 363 Kai-li Liu, Xiu-mei Wang, Zi-long Li, Ying Liu, and Rong-qiao He 21 Morpholino Studies in Xenopus Brain Development . . . . . . . . . . . . . . . . . . . . . . . . 377 Jennifer E. Bestman and Hollis T. Cline 22 Sensitive Multiplexed Fluorescent In Situ Hybridization Using Enhanced Tyramide Signal Amplification and Its Combination with Immunofluorescent Protein Visualization in Zebrafish . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 397 Gilbert Lauter, Iris Söll, and Giselbert Hauptmann 23 Live Morphometric Classification of Sensory Neurons in Larval Zebrafish . . . . . . . 411 Gema Valera and Hernán López-Schier 24 Immunohistochemistry and In Situ Hybridization in the Developing Chicken Brain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 421 Richard P. Tucker, Tatsuto Ishimaru, and Qizhi Gong 25 Gene Silencing in Chicken Brain Development . . . . . . . . . . . . . . . . . . . . . . . . . . . 439 Georgia Tsapara, Irwin Andermatt, and Esther T. Stoeckli

Contents

ix

26 Transplantation of Neural Tissue: Quail–Chick Chimeras . . . . . . . . . . . . . . . . . . . . 457 Andrea Streit and Claudio D. Stern 27 Immunohistochemistry and RNA In Situ Hybridization in Mouse Brain Development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 475 Jinling Liu and Aimin Liu 28 The Cre/Lox System to Assess the Development of the Mouse Brain . . . . . . . . . . . 491 Claudius F. Kratochwil and Filippo M. Rijli 29 In Utero Electroporation to Study Mouse Brain Development . . . . . . . . . . . . . . . . 513 Emilie Pacary and François Guillemot Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 525

Contributors Hassan M. M. Ahmed  •  Department of Developmental Biology, Johann-Friedrich-­ Blumenbach Institute, GZMB, University of Göttingen, Göttingen, Germany; Department of Crop Protection, Faculty of Agriculture, University of Khartoum, Khartoum-North, Khartoum, Sudan Irwin Andermatt  •  Neuroscience Center Zurich, Institute of Molecular Life Sciences, University of Zurich, Zurich, Switzerland Stéphanie Bertrand  •  Sorbonne Université, CNRS, Biologie Intégrative des Organismes Marins (BIOM), Observatoire Océanologique, Banyuls-sur-Mer, France Jennifer E. Bestman  •  Biology Department, William and Mary, Williamsburg, VA, USA Gregor Bucher  •  Department of Evolutionary Developmental Genetics, Johann-­ Friedrich-­Blumenbach Institute, GZMB, University of Göttingen, Göttingen, Germany; Department of Evolutionary Developmental Genetics, Georg-August-University Göttingen, Göttingen, Germany Marita Buescher  •  Department of Evolutionary Developmental Genetics, Johann-­ Friedrich-­Blumenbach Institute, GZMB, University of Göttingen, Göttingen, Germany Wanda Buzgariu  •  Department of Genetics and Evolution, iGE3, Faculty of Sciences, University of Geneva, Geneva, Switzerland Hollis T. Cline  •  The Dorris Neuroscience Center, The Scripps Research Institute, La Jolla, CA, USA Marie-Laure Curchod  •  Department of Genetics and Evolution, iGE3, Faculty of Sciences, University of Geneva, Geneva, Switzerland Danielle C. Diaper  •  Department of Basic and Clinical Neuroscience, Maurice Wohl Clinical Neuroscience Institute, Institute of Psychiatry, Psychology and Neuroscience, King’s College London, London, UK Janet S. Duerr  •  Department of Biological Sciences, Ohio University, Athens, OH, USA Kolja N. Eckermann  •  Göttingen Graduate Center for Molecular Biosciences, Neurosciences and Biophysics, Göttingen, Germany; Department of Developmental Biology Johann-Friedrich-Blumenbach Institute, GZMB, University of Göttingen, Göttingen, Germany Boris Egger  •  Department of Biology, University of Fribourg, Fribourg, Switzerland Héctor Escrivà  •  Sorbonne Université, CNRS, Biologie Intégrative des Organismes Marins (BIOM), Observatoire Océanologique, Banyuls-sur-Mer, France Max S. Farnworth  •  Department of Evolutionary Developmental Genetics, Johann-­ Friedrich-­Blumenbach Institute, GZMB, University of Göttingen, Göttingen, Germany; Göttingen Graduate Center for Molecular Biosciences, Neurosciences and Biophysics, Göttingen, Germany; Department of Evolutionary, Developmental Genetics, GeorgAugust-University Göttingen, Göttingen, Germany Cornelia Fritsch  •  Department of Biology, University of Fribourg, Fribourg, Switzerland Brigitte Galliot  •  Department of Genetics and Evolution, iGE3, Faculty of Sciences, University of Geneva, Geneva, Switzerland

xi

xii

Contributors

Natalia Carolina Garcia-Perez  •  Department of Evolutionary Developmental Genetics, Johann-Friedrich-Blumenbach Institute, GZMB, University of Göttingen, Göttingen, Germany Brenda Gavilán  •  Departament de Genètica, Universitat de Barcelona, Barcelona, Spain Qizhi Gong  •  Department of Cell Biology and Human Anatomy, University of California, Davis, Davis, CA, USA François Guillemot  •  The Francis Crick Institute, London, UK Steffen Harzsch  •  Department of Cytology and Evolutionary Biology, Zoological Institute and Museum, University of Greifswald, Greifswald, Germany Giselbert Hauptmann  •  Department of Molecular Biosciences, The Wenner-Gren Institute, MBW, Stockholm University, Stockholm, Sweden Jamie A. Havrilak  •  Department of Biological Sciences, Lehigh University, Bethlehem, PA, USA Bicheng He  •  Department of Evolutionary Developmental Genetics, Johann-Friedrich-­Blumenbach Institute, GZMB, University of Göttingen, Göttingen, Germany; Department of Evolutionary Developmental Genetics, Georg-August-University Göttingen, Göttingen, Germany Rong-qiao He  •  State Key Laboratory of Brain and Cognitive Science, Institute of Biophysics, Chinese Academy of Sciences, Beijing, China Frank Hirth  •  Department of Basic and Clinical Neuroscience, Maurice Wohl Clinical Neuroscience Institute, Institute of Psychiatry, Psychology and Neuroscience, King's College London, London, UK Clare Hudson  •  Laboratoire de Biologie du Développement de Villefranche-sur-mer (LBDV), Sorbonne Université, CNRS, Villefranche-sur-mer, France Vera S. Hunnekuhl  •  Department of Evolutionary Developmental Genetics, , Georg-August-University Göttingen, Göttingen, Germany Tatsuto Ishimaru  •  Department of Cell Biology and Human Anatomy, University of California, Davis, CA, USA David Jussen  •  Institute of Genetics, University of Mainz, Mainz, Germany Karoline F. Kraft  •  Institute of Genetics, University of Mainz, Mainz, Germany Claudius F. Kratochwil  •  Department of Biology, Zoology and Evolutionary Biology, University of Konstanz, Konstanz, Germany Jakob Krieger  •  Department of Cytology and Evolutionary Biology, Zoological Institute and Museum, University of Greifswald, Greifswald, Germany Gilbert Lauter  •  Department of Biosciences and Nutrition, Neo, Karolinska Institutet, Huddinge, Sweden Michael J. Layden  •  Department of Biological Sciences, Lehigh University, Bethlehem, PA, USA Yann Le Petillon  •  Sorbonne Université, CNRS, Biologie Intégrative des Organismes Marins (BIOM), Observatoire Océanologique, Banyuls-sur-Mer, Banyuls-sur-Mer, France Zi-long Li  •  State Key Laboratory of Brain and Cognitive Science, Institute of Biophysics, Chinese Academy of Sciences, Beijing, China Aimin Liu  •  Department of Biology, Center for Cellular Dynamics, Eberly College of Science, Huck Institute of Life Sciences, The Penn State University, Pennsylvania, PA, USA

Contributors

xiii

Jinling Liu  •  Department of Biology, Center for Cellular Dynamics, Eberly College of Science, Huck Institute of Life Sciences, The Penn State University, Pennsylvania, PA, USA Kai-li Liu  •  State Key Laboratory of Brain and Cognitive Science, Institute of Biophysics, Chinese Academy of Sciences, Beijing, China Ying Liu  •  State Key Laboratory of Brain and Cognitive Science, Institute of Biophysics, Chinese Academy of Sciences, Beijing, China Hernán López-Schier  •  Sensory Biology and Organogenesis, Helmholtz Zentrum Munich, Munich, Germany Pedro Martinez  •  Departament de Genètica, Universitat de Barcelona, Barcelona, Spain; Institut Català de Recerca i Estudis Avancats (ICREA), Barcelona, Spain Karolina Miszczak  •  Department of Biology, University of Fribourg, Fribourg, Switzerland Dominik S. Mühlen  •  Department of Evolutionary Developmental Genetics, Johann-­ Friedrich-­Blumenbach Institute, GZMB, University of Göttingen, Göttingen, Germany; Göttingen Graduate Center for Molecular Biosciences, Neurosciences and Biophysics, Göttingen, Germany Georg Oberhofer  •  Department of Evolutionary Developmental Genetics, Johann-­ Friedrich-­Blumenbach Institute, GZMB, University of Göttingen, Göttingen, Germany Emilie Pacary  •  INSERM U1215, Neurocentre Magendie, Bordeaux, France; Université de Bordeaux, Bordeaux, France Elena Perea-Atienza  •  Departament de Genètica, Universitat de Barcelona, Barcelona, Spain Chrystelle Perruchoud  •  Department of Genetics and Evolution, iGE3, Faculty of Sciences, University of Geneva, Geneva, Switzerland Emma L. Powell  •  Visual Circuit Assembly Laboratory, The Francis Crick Institute, London, UK Eduard Renfer  •  Department for Molecular Evolution and Development, Faculty of Life Sciences, Center of Organismal Systems Biology, University of Vienna, Vienna, Austria Fabian Rentzsch  •  Sars International Centre for Marine Molecular Biology, University of Bergen, Bergen, Norway; Department of Biological Sciences, University of Bergen, Bergen, Norway Filippo M. Rijli  •  Friedrich Miescher Institute for Biomedical Research, Basel, Switzerland Iris Salecker  •  Visual Circuit Assembly Laboratory, The Francis Crick Institute, London, UK Janna Siemanowski  •  Department of Evolutionary Developmental Genetics, Georg-August-University Göttingen, Göttingen, Germany Iris Söll  •  Department of Molecular Biosciences, The Wenner-Gren Institute, Stockholm University, Stockholm, Sweden Franziska Spitzner  •  Department of Cytology and Evolutionary Biology, Zoological Institute and Museum, University of Greifswald, Greifswald, Germany Simon G. Sprecher  •  Department of Biology, University of Fribourg, Fribourg, Switzerland Claudio D. Stern  •  Department of Cell and Developmental Biology, University College London, London, UK Esther T. Stoeckli  •  Neuroscience Center Zurich, Institute of Molecular Life Sciences, University of Zurich, Zurich, Switzerland

xiv

Contributors

Andrea Streit  •  Faculty of Dental, Oral and Craniofacial Sciences, Centre for Craniofacial and Regenerative Biology, King’s College London, London, UK Ulrich Technau  •  Department for Molecular Evolution and Development, Faculty of Life Sciences, Center of Organismal Systems Biology, University of Vienna, Vienna, Austria Georgia Tsapara  •  Neuroscience Center Zurich, Institute of Molecular Life Sciences, University of Zurich, Zurich, Switzerland Richard P. Tucker  •  Department of Cell Biology and Human Anatomy, University of California, Davis, CA, USA Rolf Urbach  •  Institute of Genetics, University of Mainz, Mainz, Germany Gema Valera  •  Sensory Biology and Organogenesis, Helmholtz Zentrum Munich, Munich, Germany Caroline Viertel  •  Department of Cytology and Evolutionary Biology, Zoological Institute and Museum, University of Greifswald, Greifswald, Germany Xiu-mei Wang  •  State Key Laboratory of Brain and Cognitive Science, Institute of Biophysics, Chinese Academy of Sciences, Beijing, China Andreas Wanninger  •  Department of Integrative Zoology, Faculty of Life Sciences, University of Vienna, Vienna, Austria Tim Wollesen  •  European Molecular Biology Laboratory, Heidelberg, Germany

Part I Invertebrate Models

Chapter 1 Combining BrdU-Labeling to Detection of Neuronal Markers to Monitor Adult Neurogenesis in Hydra Wanda Buzgariu, Marie-Laure Curchod, Chrystelle Perruchoud, and Brigitte Galliot Abstract The nervous system is produced and maintained in adult Hydra through the continuous production of nerve cells and mechanosensory cells (nematocytes or cnidocytes). De novo neurogenesis occurs slowly in intact animals that replace their dying nerve cells, at a faster rate in animals regenerating their head as a complete apical nervous system is built in few days. To dissect the molecular mechanisms that underlie these properties, a precise monitoring of the markers of neurogenesis and nematogenesis is required. Here we describe the conditions for an efficient BrdU-labeling coupled to an immunodetection of neuronal markers, either regulators of neurogenesis, here the homeoprotein prdl-a, or neuropeptides such as RFamide or Hym-355. This method can be performed on whole-mount animals as well as on macerated tissues when cells retain their morphology. Moreover, when antibodies are not available, BrdU-labeling can be combined with the analysis of gene expression by whole-mount in situ hybridization. This co-­ immunodetection procedure is well adapted to visualize and quantify the dynamics of de novo neurogenesis. Upon continuous BrdU labeling, the repeated measurements of BrdU-labeling indexes in specific cellular populations provide a precise monitoring of nematogenesis as well as neurogenesis, in homeostatic or developmental conditions. Key words Hydra nervous system, Interstitial stem cells, Neurogenesis, Nematogenesis, In situ hybridization, Immunofluorescence, Hydroxyurea, BrdU, prdl-a, Hym-355, RFamide

1  Introduction 1.1  Anatomy of the Nervous System, Neurogenesis, and Nematogenesis in Hydra

The freshwater hydrozoan Hydra belongs to Cnidaria, an early-­ branched eumetazoan phylum, sister group to bilaterians. Hydra is an attractive model for biologists not only for its outstanding regenerative properties but also for its highly dynamic neurogenesis. Indeed the nervous system is continuously renewed with nerve precursors produced in the body column, then displaced towards the extremities where they terminally differentiate to replace the nerve cells that die [1] (Fig. 1a). In addition, animals bisected at any level along the body column replace the missing part in few

Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_1, © Springer Science+Business Media, LLC, part of Springer Nature 2020

3

4

A

Wanda Buzgariu et al.

C

INTACT HYDRA POLYP apical differentiated nerve cells

nerve cell precursors

neurogenesis

interstitial stem cells

interstitial stem cells migration

nerve cell precursors

4x

8x

basal differentiated nerve cells

B

sensory neurons

sensory-motor bipolar neurons

HEAD-REGENERATING HYDRA

16 a.p

a.p

nematoblasts

4 hpa

24 hpa

36 hpa

a.p=amputation plane

nematocytes

ganglia neurons

Fig. 1 Adult neurogenesis in Hydra and cell types that form its nervous system. (a, b) Neurogenesis in adult intact (a) and in head-regenerating (b) Hydra polyp (modified from ref. 2). Only nerve cells are shown, denser in the apical (top) and basal (bottom) regions than in the body column where neurogenesis takes place. The interstitial stem cells (ISCs) provide precursors for all nerve cells and nematocytes, which migrate towards the extremities [3]. Terminal differentiation of nerve cells predominantly takes place at the extremities. In head-­ regenerating Hydra (b) interstitial cells and derivatives are first eliminated upon injury-induced cell death after mid-gastric bisection [4] and de novo neurogenesis takes place at the tip where the apical nervous system get built within 2 days, with precursors detected after 24 h [5, 6]. (c) ISCs divide every 24–30 h and frequently appear as pairs (top-left panel). Nematogenesis starts with a series of synchronous syncytial divisions that lead to the formation of nematoblast clusters that contain 4×, 8×, 16× or even 32× cells. At any stage, a cluster can stop proliferating to enter differentiation, i.e. each nematoblast forms a venom capsule named nematocyst (arrows) and a cnidocil (not visible), both fully mature in nematocytes [7, 8]. Note the moon-shaped nucleus in nematocytes, compressed by the nematocyst. Hydra differentiates different classes of neurons: sensory, sensory-motor bipolar, and multipolar ganglia neurons (right panels). Cells obtained after tissue maceration [9], were immunodetected with an anti α-tubulin antibody followed by an anti-mouse secondary antibody coupled with Alexa-488 (green). The nuclei were counterstained with DAPI (light blue). Pictures were taken on a SP8 Leica confocal microscope and optimized on Photoshop with channelmixer to lighten the dark blue color and M-curves to increase the contrast. Scale bar: 10 μm

days, e.g. a new head equipped with a complete nervous system (Fig. 1b). The Hydra nervous system is made of nerve cells and mechanosensory cells (named nematocytes or cnidocytes) that form a nerve net much denser at the apical and basal extremities of the animal. There it controls a series of more or less complex behaviors such as peristaltic movements, walking, prey capture, food ingestion, and reaction to light. Cnidarian neurons are equipped with neurites but no axons, therefore not polarized and often named “nerve cells.” Every nerve cell is able to function as sensory, sensory-motor, or inter-

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neuron. Nevertheless, nerve cells show diverse anatomies, sensory, bipolar, or multipolar as ganglia neurons (Fig. 1c), and the neuronal phenotype of a given cell can change according to its position along the body axis [10]. At the base of the apex, ganglia neurons can form a nerve ring that allows coordinated behaviors [11]. Although cnidarian nervous systems are highly peptidergic with peptide-gated ion channels as receptors [12, 13], they share with bilaterians all the basic properties of synaptic conduction and chemical neurotransmission [14, 15], even though the neuromuscular junction might have evolved independently in cnidarians and bilaterians [16]. Still, the transcriptional factors that control neurogenesis are largely evolutionarily conserved [2, 5, 9, 17–21] and cnidarian nervous systems might represent the first evolutionary attempt of centralized nervous system in eumetazoans [11]. The two myoepithelial layers of the body wall, the epidermis and the gastrodermis, are made of cells that derive from three not interchangeable stem cell populations: the two unipotent ectodermal and endodermal epithelial stem cells (ESCs) and the multipotent interstitial stem cells (ISCs). ISCs, predominantly located in the central body column, give rise to a variety of cell types, including the two cell lineages that build up the nervous system (Figs. 1a and 2): the rare nerve cells and the abundant nematocytes, which represent 3% and 35% of the total cell number, respectively [41, 42]. The nerve cells directly differentiate from migrating precursors and then get displaced towards the extremities where they form a dense diffuse nerve net [6, 42–44]. In intact animals (homeostasis), neurogenesis is a slow process, leading to the replacement of the nerve cells that get sloughed off at the extremities, while after bisection a faster de novo neurogenesis process takes place in the regenerating structure (Fig. 3a). In contrast to neurogenesis, nematogenesis is a multiple step process where interstitial progenitors synchronously divide up to five times, providing clusters that contain 4, 8, 16, or 32 nematoblasts (Figs. 1b and 2) [7]. Cells from a given cluster can exit the cell cycle at any time from the 4-cell stage to differentiate as nematocytes equipped with a sensory cnidocil and a vacuole named cnidocyst that contains a paralyzing venom, the latter structure being the hallmark of cnidarians [8]. The turnover of nematocytes is highly dynamic as once the venom capsule is discharged, the ­nematocyte is eliminated and replaced. Interestingly, ISCs cycle three to four times faster than ESCs and can thus be easily eliminated upon pulse treatments of antimitotic drugs [45–47]. Elimination of the cycling interstitial cells leads to the loss of the nervous system, after several weeks in intact animals, within few days in regenerating ones, as such animals regenerate a missing part that lacks a nervous system (Figs. 3b and 4). Nerve-free animals actually maintain their developmental properties if they are

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Fig. 2 Molecular markers of neurogenesis and nematogenesis in Hydra. A partial list of genes expressed during neurogenesis (upper) and/or nematogenesis (lower). Gene products detected with antibodies are written plain. ISC interstitial stem cells, pc precursor cell, nb nematoblasts. The spatial and cell-type expression pattern of genes presumably involved in neurogenesis and neurotransmission (103 and 156, respectively) was identified by RNAseq transcriptomics [22], and for any Hydra gene of interest the spatial and cell-type expression patterns are now available on the HydrAtlas server (https://hydratlas.unige.ch/blast/blast_link.cgi) [23]. Specific signatures were obtained by analyzing the expression patterns of (1) neuropeptides such as RGamide, RFamide, KVamide, or LWamide [24–26], the neuropeptide Hym-355 enhancing neuronal differentiation [3], (2) 86 genes identified in a microarray screened with cRNAs from nerve-less animals [27], (3) neurogenic genes that regulate the proliferation and/or the differentiation of progenitors such as CnASH [17, 28], prdl-a [23], COUP-TF, prdl-b [29], hyZic [30], cnox-2 [22], CREB [29], Myc1 [31, 32], FoxN1, PaxA, PaxB, Pou4F2 [22], (4) the two Piwi proteins (HyWi, HyLi) strongly expressed in ISCs and nematoblasts [33], (5) some gap junction proteins, innexin-2 being involved in neurotransmission [34]. No pan-neuronal marker was identified in Hydra yet. As early markers of nematocyst differentiation, the minicollagens N-COL1 and NOWA [35, 36] are components of the wall, spinalin of the spines [37], N-COL15 of the tubule [38], nematogalectins A and B of the tubule from distinct types of nematocysts [39]. Finally, a proteomic analysis of the venom identified 410 secreted proteins in nematocysts [40] (not shown here)

Fig. 3 The homeoprotein prdl-a as a marker of apical neurogenesis. (a) Double immunostaining of BrdU (green) and prdl-a (red) expressed in apical neuronal progenitors or nerve cells of homeostatic animals (upper panel) or animals having regenerated their head (lower panel). All animals were incubated with BrdU for 4 h and then washed out to remain intact or to undergo mid-gastric bisection (red arrow). White arrow heads indicate the prdl-a+/BrdU+ cells, which are more numerous in the regenerated than in the homeostatic head. (b) De novo neurogenesis evidenced by prdl-a (red) immunostaining in the presence (top panel) or the absence (lowerpanel) of interstitial progenitors after hydroxyurea (HU) treatment. The animals were exposed or not to 10 mM HU in three rounds (2× 24 h and 1× 32 h). After the third HU treatment, animals were transferred to HM, bisected (red arrow) and let to regenerate for seven days. Note the absence of prdl-a+ cells in HU-treated animals. Nuclei were counterstained with DAPI (blue). Image acquisition was done on a Leica LSM700 confocal microscope. Scale bar: 100 μm

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Fig. 4 Loss of terminal neuronal differentiation in Hydra undergoing apical or basal regeneration in the absence of ISCs or interstitial progenitors. HU treatment and regeneration were performed as in Fig. 3. The apical (a) and basal (b) nerve nets were identified after RFamide immunostaining (green) and nuclear DAPI counterstaining (purple). Note the sparse RFamide+ nerve cells in apical (a) and basal (b) regions regenerated after HU treatment. Image acquisition was done on a Leica LSM700 confocal microscope. Scale bars: 100 μm

force-fed, pointing to an important morphogenetic role played by the epithelial cells [46]. The Hydra Peptide Project identified about 500 peptides, half neuropeptides, half epitheliopeptides, with specific roles in neurotransmission, morphogenesis, and differentiation [48, 49]. However, not much is known concerning the role and the regulation of the neuronal-epithelial cross talk on the slow and fast modes of neurogenesis in intact and regenerating animals, respectively. These questions require the monitoring of the expression of neuronal markers in proliferating and differentiating progenitors as well as in fully mature nerve cells (Fig. 5). 1.2  Principles of Immunodetection on Whole-Mount and Macerated Tissues in Hydra

The Hydra nervous system is well visualized with the immunostaining procedures applied either on whole animals or on cells that keep their morphology after tissue maceration [41]. In both contexts, the procedure is based on the specific antigen–antibody recognition followed by different detection techniques to visualize the cells that express or overexpress a known antigen. The immunodetection procedure follows the classical steps: fixation and permeabilization of the tissue, saturation of unspecific sites, and antigen recognition by a specific antibody followed by detection and visualization. The expression patterns obtained in intact or regenerating Hydra as well as in specific cell types can also be verified at the transcriptomic level on the HydrAtlas server (https:// hydratlas.unige.ch) [23].

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Fig. 5 BrdU immunostaining combined with Hym-355 detection in homeostatic animals. The neuropeptide Hym-355 is expressed in a subset of apical and basal neurons. The animals were exposed to BrdU for 4 h, washed and subsequently maintained in HM for 2 days and then fixed. After the in situ hybridization procedure to detect Hym-355 expressing nerve cells (a, purple), the animals were immunolabeled for BrdU (b, c, green). In homeostatic condition, only few apical neurons do express Hym-355 and are positive for BrdU (arrows). The bright field (a) and fluorescence (b) images were acquired with a Leica DM5500 fluorescence microscope. Box in (b) corresponds to panel (c). Scale bars: 100 μm (a, b) and 20 μm (c)

1.2.1  Immunodetection on Whole-Mounts

The whole-mount immunodetection procedure is a robust method to detect with specific antibodies the spatial or temporal expression pattern of neurogenic proteins or neuropeptides. In complement the same procedure can be used to detect with antibodies raised against tagged material incorporated into macromolecules such as Bromodeoxyuridine (BrdU) incorporated into newly synthesized DNA [50] or digoxygenin (DIG)-labeled riboprobes that bind transcripts [51] (Fig. 5). As a consequence, immunodetection allows the visualization of proteins, DNA, and transcripts in tissues that show a preserved architecture.

1.2.2  Immunodetection on Cells Obtained After Tissue Maceration

To get a more accurate analysis at the cellular level, the immunodetection procedure can be performed on histological sections [18, 52] or on cells directly fixed on slides after tissue maceration [41]: In the presence of acetic acid and glycerol, the tissues are dissociated into single cells or small clusters, which are subsequently fixed with paraformaldehyde (pFA) that preserves their typical cellular architecture. Hydra-specific or exogenous epitopes as BrdU can next be immunodetected. The maceration technique can be adapted for small amount of tissue such as head- or foot-­ regenerating tips [4]. The strengths of this method are multiple: it identifies cell types and specific subcellular localization, it allows the precise quantification of each expressing cell type, including neurons, interstitial cells or clusters of nematoblasts (Fig. 1b).

1.2.3  Antibodies

As mentioned above, the Hydra nervous system produces numerous neuropeptides. Since 1982, their immunodetection on intact animals proved to be extremely useful to identify the anatomical organization and localization of specific subsets of neurons in Hydra [53, 54]. The vasopressin-like or RFamide sera were critical to monitor the phenotypic conversion of neurons as they get displaced along the body axis [10]. With the

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development of bioinformatic tools and genomics, evolutionarily conserved regions can be easily mapped and commercially available antibodies raised against such mammalian epitopes can be used to cross-react against Hydra proteins. Alternatively monoclonal [55] or polyclonal antibodies can be raised against Hydra proteins, either lab-made such as the sera against the RFamide neuropeptide [53], the transcription factor CREB [56], and the homeoprotein prdl-a [18] or commercially produced. The most efficient way to validate the specificity of a given antibody is to perform gene silencing, usually through RNA interference in Hydra, to assess the decrease in protein abundancy by Western analysis or immunofluorescence [57]. 1.2.4  Tissue Fixation and Tissue Permeabilization

Fixation and permeabilization are crucial steps as they allow the preservation of cells and tissue architecture. Paraformaldehyde (pFA) and formaldehyde (FA) are two cross-linking agents commonly used as fixatives; they ensure a good penetrability and preserve the antigens by forming chemical bonds between the proteins. In contrast, alcohols such as ethanol and methanol, or acetic acid are precipitating fixatives that disrupt the hydrophobic bonds and thus denature the tertiary structure of proteins. In fact, ethanol or methanol allow a good permeabilization by precipitating membrane proteins, leading to the formation of pores in the membrane, thus contributing to the leakage of RNA or cytoplasmic proteins and to a poor cellular preservation. Therefore, the commonly used method to fix Hydra polyps for whole-mount immunodetection or in situ hybridization (ISH) combines pFA and alcohol fixatives. As an alternative the Lavdowsky fixative that combines ethanol, FA, acetic acid and water (50/3.7/4/42), also takes advantage of the properties of cross-linking and precipitating agents [55]. A variant of the Lavdowsky fixative that does not contain acetic acid is often used to detect nuclear antigens. Finally, the mercury-containing “Helly” fixative or the Zinc–FA fixative used on insect brains to preserve morphology and improves immunodetection at synapses [58], can be used efficiently when the other fixatives fail. For each new epitope/antibody several fixatives need to be tested together with the fixing conditions such as temperature (4 °C, 18 °C, 37 °C) or duration of fixation. Tissue permeabilization is usually completed by adding a detergent such as Triton X-100 or Tween 20, again applied for a period of time that needs to be adapted to each antigen of interest.

1.2.5  Antigen Detection

The detection of an antigen can be made directly with the primary antibody or indirectly through a secondary antibody that binds to the primary one. The indirect method is preferred as Hydra specific antibodies conjugated with fluorochrome are rare and immunodetection is more sensitive when indirect as the signal is amplified. A large choice of secondary antibodies coupled to fluorochromes or enzymes are available. Amplification is optimal

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when a Tyramide detection system is used with conditions adapted for each antibody (see supplemental figure 1 in ref. 21). Unspecific binding of antibodies to reactive sites of proteins is blocked by BSA (bovine serum albumin) or normal serum from the species used to raise the secondary antibody. A common counterstaining is made by nuclear staining, which allows a better cell-type and tissue-layer identification through the shape and size of nuclei.

2  Materials All stock solutions are prepared with MilliQ water in sterilized bottles with screw-cup, then autoclaved or filtered through a 0.22 μm Steritop filter, and finally stored at 4 °C or at room temperature (RT) depending on the buffers. After opening, the stock solution should be checked regularly to be discarded before they become viscous or turbid, with visible signs of contamination. Experiments are always performed with fresh solutions, i.e. 10× stock solutions freshly diluted in sterile dishware with MilliQ water to prepare the requested amount of 1× solution for a single experiment. 2.1  Hydra Maceration

1. Hydra medium (HM): 1 mM NaCl, 1 mM CaCl2, 0.1 mM KCl, 0.1 mM MgSO4, 1 mM Tris pH 7.6 [59]. HM is prepared from three stock solutions that can be stored several weeks at RT once autoclaved: For the stock solution A (0.5 M Tris, 500×) dissolve 60.57 g Tris-base in 900 ml H2O, adjust the pH to 7.7 and complete the volume to 1000 ml. For the stock solution B (1 M MgSO4, 10,000×) dissolve 61.6 g MgSO4⋅7H2O in 250 ml H2O. For the stock solution C (0.5 M CaCl2, 0.5 M NaCl, 0.05 M KCl, 500×) dissolve 54.77 g CaCl2⋅6H2O, 14.6 g NaCl, 1.85 g KCl in 500 ml H2O. Autoclave the solutions A, B, and C. To prepare 1 l HM solution, dilute 2 ml solution A, 100 μl solution B and 2 ml solution C in 1000 ml H2O. 2. Macerating solution (MS): 7% acetic acid, 7% glycerol in H2O. Add 0.7 ml glycerol and 0.7 ml glacial acetic acid to 8.6 ml H2O. Use the fume hood for preparation. Do not autoclave. Store at RT no longer than 2 weeks. 3. 10% Tween 80: Add 1 ml Tween 80 to 9 ml H2O (see Note 1). Do not autoclave. Keep the solution for 1 month at RT.

2.2  Fixation

1. 4% Urethane: Dissolve 1 g urethane in 25 ml HM. Store at 4 °C. Always wear gloves as urethane is carcinogenic. 2. Lavdowsky fixative: 50% ethanol, 3.7% FA, 4% acetic acid. Add 1 ml 37% FA to 5 ml ethanol, 0.4 ml acetic acid and 4 ml H2O. Prepare it fresh, do not store.

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2.3  BrdU Immunodetection



(a) Lavdowsky fixative without acetic acid: 50% ethanol, 3.7% FA. Add 1 ml 37% FA to 5 ml ethanol and 4 ml H2O. Prepare it fresh and do not store.



(b) 8% Paraformaldehyde (PFA): Dissolve 8 g pFA in 80 ml HM preheated at 65–70 °C in a glass bottle and stir the mixture on a water bath maintained at 70 °C under the hood. Add 100 μl 1 N NAOH and stir the solution until the solution becomes clear. After cooling to RT, adjust the volume to 100 ml with HM and adjust the pH to 7.5 with 0.1 N NaOH. Filter the solution using a steriflip filter. Store at 4 °C for 2 weeks maximum (see Note 2).



(c) 4% PFA: Dilute 50 ml 8% pFA with 50 ml H2O and readjust the pH to 7.5 with 0.1 N NaOH.



(d) 3.7% Formaldehyde (FA): Add 1 ml 37% FA to 9 ml H2O.

BrdU (5-Bromo-2'deoxyuridine is an analog of thymidine that is rapidly incorporated into DNA during the replication phase of the cell cycle. 1. BrdU solution (10 mM): Add 154 mg BrdU to 50 ml HM in a 50 ml centrifugation tube, protect the tube from light and stir the solution that can be stored at 4 °C for a few days (see Note 3). 2. 10× PBS buffer: 1.37 M NaCl, 27 mM KCl, 81 mM Na2HPO4, 11 mM KH2PO4. Dissolve 80 g NaCl, 2 g KCl, 29 g Na2HPO4 12H2O and 2 g KH2PO4 in 1000 ml H2O. Verify that pH is 6.8. Autoclave and keep at RT (see Note 4). 3. PBS: 137 mM NaCl, 2.7 mM KCl, 8.1 mM Na2HPO4, 1.1 mM KH2PO4. Dilute 50 ml 10× PBS to 500 ml (see Note 4). 4. PBST: 0.5% (v/v) Triton X-100 in PBS. Add 0.5 ml Triton X-100 to 100 ml PBS (see Note 1). 5. Blocking solution: 2% BSA. Dissolve 1 g BSA in 50 ml PBS and stir until the solution becomes clear. Filter through a Steriflip and store at 4 °C (see Note 5). 6. 2 N HCl: Add 2 ml 37% HCl to 10 ml of H2O (see Note 6). 7. DAPI (4,6-diamidino-2-phenylindole) stock solution (1 mg/ ml): Dissolve 10 mg DAPI in 10 ml H2O. Aliquot and store at −20 °C. 8. DAPI working solution (1 μg/ml): Dilute 1 μl DAPI 1 mg/ml into 1 ml PBS.

2.4  Antibodies

1. The polyclonal anti-prdla antibody was produced in rabbit after immunization with the N-terminal fragment of prdl-a fused with 6HIS [18]. The polyclonal anti-RFamide antibody was produced in rabbits and kindly provided by Cok

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Grimmelikhuijzen [53]. For BrdU immunodetection the BrdU labeling and detection kit from Roche (ref 11444611001) provides the most reliable results. Dilute the antibody 1:20 in the labeling solution found in the kit (see Note 7). 2. The optimal titer for each primary antibody should be established after testing a series of dilutions; too high concentrations increase the background staining as the antibody accumulate at the cell or tissue surface, too low concentrations provide low-­intensity signals due to a reduced antigen–antibody interaction. 3. In the indirect immunostaining procedure, the secondary antibody needs to be chosen according to the available filters and lasers that equip the fluorescent/confocal microscopes. If a double antigen detection is required, select the secondary antibodies considering the possible cross talk between the excitation and emission wavelengths of each fluorochrome. 2.5  Mounting Medium

The samples should be mounted in a medium adapted for fluorescence, i.e. characterized by a high refractory index, no autofluorescence and protecting against photobleaching. The MOWIOL mounting medium (polyvinyl alcohol) fulfills these criteria. It is a widely used lab-made preparation, less expensive than the commercially available ones. 1. 0.2 M Tris pH 8.5: Dissolve 2.42 g Tris base in 100 ml of H2O. Adjust in a fume hood the pH to 8.5 with approximately 710 μl of 37% HCl solution. 2. Add 24 g MOWIOL 4–88 to 60 g glycerol and stir well. Subsequently add 60 g H2O and mix well for another 2 h at RT. 3. Add 100 ml 0.2 M Tris pH 8.5 and continuously stir at 52 °C on a heating plate until the powder is completely dissolved. This might take 4–5 h. 4. Dispatch the mix in 50 ml centrifugation tubes and centrifuge at 5000 × g for 15 min. 5. Carefully collect the supernatant and aliquot into 15 ml conical tubes. 6. Store the Mowiol mounting medium at −20 °C up to 1 year. For current use, keep aliquots at 4 °C.

2.6  Equipment

1. Stereomicroscope. 2. Pasteur glass pipette. 3. 360° vertical rotator. 4. Shaker. 5. Superfrost Plus microscope slides (Thermo Scientific, Gerhard Menzel).

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6. Coverslips: 22 × 40 mm, 0.13–0.17 thickness. 7. Surgical scalpel N°3. 8. Surgical blades N°10 (Ruttgers Solingen). 9. Plastic Petri dishes (6 and 9 cm diameter). 10. Silicon bulbs, 5 mm diameter for Pasteur pipette. 11. Steriflip and steritope for filtration (Millipore). 12. Hydrophobic pen (DAKO Pen). 13. Slide staining tray, Simport Scientific.

3  Methods 3.1  Hydra Tissue Maceration Protocol and Fixation

1. Collect five intact animals in a 1.5 ml Eppendorf tube with the help of a Pasteur pipette. Remove the medium and replace  it with 400 μl of fresh HM, repeat the washing 2–3× times (see Note 8). 2. After the last wash, eliminate carefully all the liquid and add 100 μl MS. Let the tube on a rack and mix gently, from time to time, until the tissue dissociates into a homogenous cell suspension (see Note 9). 3. After 30–60 min, fix the cell suspension by adding 100 μl 8% pFA, mix gently and let incubate for 1 h at RT. 4. Add 10 μl of 10% Tween 80 and mix gently. 5. With the DAKO PEN, label a square of about 25 × 20 mm on a Superfrost Plus slide. Let it dry for about 10 min. 6. Add the cell suspension drop by drop on the surface delimitated by the hydrophobic marker lines. 7. Let the slides dry for about 2 days at RT. 8. The slides are ready to be processed or can be stored into a box at −20 °C.

3.2  Hydra Fixation for ISH and Whole-­ Mount Immunostaining

1. Collect 15–20 intact animals with a glass Pasteur pipette in a graded 2 ml Eppendorf tube and adjust the final volume to 1 ml HM. 2. Add 1 ml 10 mM BrdU solution and transfer the animals in a plastic Petri dish (6 cm diameter) that contains 10 ml 5 mM BrdU solution. Protect from light and incubate for the desired period of time (see Note 3). 3. To wash out BrdU, collect the animals in 2 ml tube and wash them several times in HM by gently aspirating the liquid and replacing it with fresh medium.

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4. To initiate regeneration, transfer the animals in a plastic Petri dish (9 cm diameter) in 50 ml HM and place the dish under the stereomicroscope. Let the animals relax for a few minutes. 5. Bisect the animals with a scalpel at half of the body column length and transfer the head regenerating halves to a new Petri dish filled with 50 ml HM. Let the animals to regenerate (see Note 10). 6. Collect 15–20 intact or regenerated animals in a graded 2 ml Eppendorf tube and wash them several times with HM as described at step 3. 7. Adjust the final volume to 1 ml. Let the animals to relax a few minutes. 8. Immediately before fixation, add 1 ml 4% urethane to reach a 2% final concentration. Mix gently and wait for 60 s until the animals are completely relaxed (see Note 11). 9. Remove 1 ml urethane solution and immediately add 1 ml 8% pFA to reach a 4% final concentration. Mix well and let the animals fall to the bottom of the tube and aspirate the maximum amount of the liquid. 10. Wash 3–4 times with 4% PFA by eliminating each time the liquid and replacing it with fresh fixative. 11. Place the tubes on a shaker/rotator and fix the animals for 4 h at RT (see Note 12). 12. Aspirate the PFA and replace it with ethanol 100%; replace several times the ethanol until the brown pigment is solubilized. 13. Store the fixed samples at −20 °C in ethanol until the ISH procedure. 3.3  Hydra Fixation for BrdU Immunostaining on Whole-Mount

1. For BrdU incubation and animal fixation, proceed as in Subheading 3.2 from steps 1 to 8. 2. Remove the maximum amount of the urethane solution and replace it with Lavdowsky fixative without acetic acid. Wash several times with the fixative. 3. Place the tubes on a shaker or rotator and let the animals fix either for 1 h at 37 °C or overnight at 4 °C (see Note 12). Proceed immediately after fixation with the immunodetection.

3.4  Immuno­ detection of Hydra Cells After Tissue Maceration

The immunodetection procedure is performed in a dark humidity chamber (see for example the StainTray slide staining system proposed by Sigma). During antibody incubation, the atmosphere is kept humid by adding PBS in the bottom of the chamber.

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1. Rehydrate the cells by washing 3× 10 min with PBS (see Note 13). 2. Incubate the cells with 0.1% PBST for 30 min (see Note 14). 3. Add the blocking solution for 1 h at RT (see Note 15). 4. Dilute the primary antibody as desired in blocking solution. 5. Gently remove the blocking solution and add 100 μl of ­primary antibody solution. Incubate overnight at 4 °C or alternatively 3–4 h at RT (see Note 16). 6. Wash the slides 4× 10 min with PBS. 7. Dilute the secondary antibody in PBS as recommended by the supplier. 8. Remove PBS, add the secondary antibody solution and incubate for 2–4 h protected from light. 9. Wash the slides 4× 10 min with PBS. 10. Incubate with 1 μg/ml DAPI for 10 min (see Note 17). 11. Wash the slides 2× 5 min with PBS. 12. Wash the slides fast with H2O and let them dry at RT for 15–20 min protected from light. 13. Once dried, add one drop of mounting medium and apply the coverslip by holding it at 45 °C. Gently descend the coverslip allowing the mounting medium to cover the surface delimitated by the square. Avoid producing air bubbles (see Note 18). 3.5  Immunostaining on Whole-Mount Hydra

The procedure presented below is a general protocol, which has to be adapted for each antibody. Unless specified, the incubation steps are performed at RT in 1.5 ml Eppendorf tubes. The protocol should be adjusted according to the fixative. If samples are freshly fixed with Lavdowsky fixative, with or without acetic acid, start with step 1. If samples are stored in ethanol after pFA fixation, directly proceed to step 2. 1. Remove the Lavdowsky fixative and replace it with PBS. Repeat quickly the washing step twice. Proceed to step 3. 2. Rehydrate the pFA-fixed animals stored in ethanol by washing successively in 75%, 50% and 25% ethanol, each step for 5–10 min (see Note 8). 3. Wash 4× 10 min with PBS. 4. Remove PBS, add PBST to permeabilize for 30 min (see Note 14). 5. Optional for nuclear antigen detection. Otherwise proceed to step 7. Remove PBST and incubate in 2 N HCl for 30 min (see Note 19).

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6. Remove HCl and wash quickly 6× in PBS to eliminate all HCl traces. Complete with 2× 5 min washes in PBS (see Note 20). 7. Add the blocking solution for 60 min at RT (see Note 15). 8. Dilute the primary antibody as desired in blocking solution. 9. Remove the blocking solution, add at least 100 μl primary antibody solution. Incubate overnight at 4 °C or alternatively 3–4 h at RT (see Note 21). 10. Remove the primary antibody, wash quickly twice in PBS, then 4× 10 min. 11. Dilute the secondary antibody in PBS as recommended by the supplier. 12. Remove PBS, add the secondary antibody solution and incubate for 2–4 h protected from light (see Note 22). 13. Remove the secondary antibody, wash quickly twice in PBS, then 4× 10 min. 14. Remove PBS and add 300 μl 1 μg/ml DAPI for 10 min (see Note 23). 15. Wash quickly twice with PBS, then 2× 5 min. 16. Wash quickly with H2O. 17. Mount the animals on glass slide with the mounting medium. Gently descend the coverslip allowing the mounting medium to cover the surface. Avoid producing air bubbles (see Note 24). 3.6  Immunostaining on Whole-Mount Hydra After In Situ Hybridization

The immunodetection protocol described above can be applied to samples previously processed for in situ hybridization (ISH) to combine the detection of both protein and gene expression. Since the 1990s, gene expression patterns can be investigated in whole-­ mount Hydra. The classical procedure comprises fixation of the samples, hybridization of Digoxygenin (DIG) or Fluorescein-­ labeled riboprobes to the targeted transcript, and immunodetection of these hybridized riboprobes with an anti-DIG or an anti-Fluorescein antibody coupled to alkaline phosphatase, followed by a colorimetric detection of the coupled enzyme with NBT/BCIP substrate (for a detailed protocol, see ref. 53). The presentation of the complete ISH procedure is omitted here and only the steps that link the two methods are detailed. 1. At the end of the ISH procedure, stain the samples with NBT/ BCIP as in [51] and follow carefully the development of the staining. Once the pattern is obtained, wash out as indicated the NBT/BCIP substrate to block the staining. 2. Post-fix the samples in 3.7% FA for 30 min at RT. 3. Wash out FA by rinsing two to three times with methanol.

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4. Incubate the samples for 10 min in methanol. 5. Rinse three-four times with PBST and proceed with step 4 from Subheading 3.5.

4  Notes 1. Tween 80 and Triton X-100 are viscous detergents and, therefore, the pipetting should be done gently in order to take up the requested volume. The solution should be stirred at low speed to ensure a proper solubilization and to circumvent foam and bubbles formation. 2. Wear gloves and mask when preparing the PFA solution as pFA is a hazardous chemical. All manipulations should be performed in a fume hood. PFA wastes should be discarded in accordance with the safety procedure established at your institution. Given the low osmolarity of Hydra cells the pFA is diluted in HM and not in PBS as for mammalian tissues [60]. As HM has a lower buffering capacity than PBS, a critical step is pH adjustment, which should be done precisely. 3. BrdU (5-bromo-2′-deoxyurdine) is light sensitive and mutagenic; chemical and solutions should be manipulated with gloves. During incubation of live animals in the BrdU solution, protect the dishes from light by covering them with aluminum foil. 4. The pH of the 10× PBS stock solution is approximately 6.8; it does not need to be adjusted if the components were correctly weighted. After dilution to 1×, the pH will increase to 7.4. 5. When preparing the BSA solution, add first the powder into a 50 ml centrifugation tube and then carefully pour the buffer. Stir at low speed to avoid foam and formation of bubbles. After filtration, the BSA solution can be stored at 4 °C for several months. 6. 2 N HCl should always be diluted freshly from a 12 N HCl stock solution that is stable at RT. Do not use old 2 N HCl solution as it is unstable, hence less efficient to denature DNA in nuclei. 7. The most effective anti-BrdU antibody for Hydra contains nucleases that cleave DNA and generate single-stranded DNA, thus enhancing the exposure of the sites that have incorporated BrdU. Always use the buffer supplied by the manufacturer when diluting the anti-BrdU antibody. If multiple antigens are detected, dilute the second primary antibody in the buffer supplied for BrdU. To avoid unspecific binding caused by the second primary antibody, pre-incubate it with fixed Hydra polyps for few hours at 4 °C.

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8. To avoid animal loss during manipulations: (1) always wait few seconds after each washing step to let the animals settle at the bottom of the tube, then aspirate carefully the washing solution and replace it with the next solution; (2) cover part of the bench with a black sticky back plastic. 9. During the maceration procedure, dissociate the tissues by gently pipetting up and down every 10 min and by flicking the tube. A too strong pipetting might break the large epithelial cells as well as the dendrites that are fragile, or disrupt the large nematoblast clusters. In contrast, interstitial cells, nematoblasts, and nematocytes are resistant to mechanical pressure. 10. To correctly estimate the amputation level when bisecting Hydra for regeneration, the animals should be relaxed to reach their maximal length. This is easily obtained by placing them in a Petri dish under the stereomicroscope and letting them stretching out for a few minutes under light. Orientate the scalpel perpendicular to the body axis and cut fast. If the animals are floating in the dish, transfer them with a low amount of liquid in the cover lid of the Petri dish and cut them there under the light. After bisection, transfer back the halves into dishes prefilled with HM, usually 1 ml/animal as when animal density is too high, regeneration is delayed [52]. 11. Urethane added for 1 min is used to relax the myoepithelial layers [61]. When animals are not sufficiently relaxed at the time the fixative is added, permeabilization is reduced, antibodies accumulate at the animal surface and no specific signal is obtained. However, a too long exposure to urethane damages the cell membranes; therefore, this step should be limited to 2 min. Always wear gloves, urethane is highly carcinogenic. 12. Alternatively, the samples can be fixed overnight at 4 °C. The fixation time should not exceed 16 h for PFA, 24 h for modified Lavdowsky (no acetic acid). 13. To wash the cells attached to the slide, add the liquid by gently pipetting within the surface delimitated by the hydrophobic square. Reverse the slide to get the liquid and replace it with fresh PBS. Wipe the excess of liquid found outside of the square limits with a tissue paper and avoid touching the hydrophobic rim. 14. Triton X-100 concentration is indicative, as the optimal concentration depends on the antigen localization in the cell. In case of a poor permeabilization or reduced antigen accessibility, a concentration range varying between 0.1% and 2% might be tested. Alternatively, the incubation time with the detergent can be increased. Tissue permeabilization also depends on the thickness of the mucus and cuticle layer that protect the animals. Finally, Triton concentration should be adapted to the

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Hydra strain, as the cuticle layer varies between different strains/species. 15. BSA is a widely used blocking agent to prevent the potential unspecific binding of antibodies. As an alternative, the serum from the species where the secondary antibody was raised, would be preferred. However normal serum from any other sources can be used as it contains albumin and other proteins that bind nonspecifically to the reactive sites of proteins. 16. When the signal to noise ratio is inappropriate meaning a high background is noticed, the detection might be improved by pre-adsorbing the primary antibody on Hydra tissue (see Note 7) or by varying the environmental conditions such as lowering the temperature or shortening the antibody incubation time. The primary antibody dilution can be reused several times over several weeks, if stored at 4 °C. 17. DAPI is a widely used fluorescent DNA dye that is excited by UV and has a broad emission spectrum. Alternatively, other DNA stains such as Hoechst 33258 (bis-benzimidazole), TOPRO 3 can be selected. The option has to be taken according to the availability of the filters/lasers, and to the selected fluorophores used for the antigen detection. 18. Once dried, the slides can be stored at −20 °C for long, although the fluorescence preservation depends on the nature of the selected fluorochromes. Alexa dyes are very stable and samples can be stored for years at −20 °C with fluorescence preservation. 19. When antigens are nuclear, i.e. transcription factors, DNA-­ tagged molecules, a DNA denaturation step is required to expose the reactive sites to the antibody. The most common DNA denaturation agent is HCl (2 N), usually applied 30 min at RT. However, depending on the environmental conditions (season, temperature), the denaturation process can be faster (20 min) or slower (40 min). The duration of the denaturation step is critical, as a too long treatment destroys and fragments the DNA, while a too short exposure is not sufficient to reveal the antibody binding sites. Examples of insufficient denaturation and permeabilization are given in Fig. 6. 20. HCl should be removed fast and the washing step should not take more than 15 min, as longer washing time might allow renaturation of the DNA and thus reduce the accessibility of antibody to the incorporated BrdU (Fig. 6). 21. When double immunostaining is done, the primary antibodies can be incubated simultaneously if the species in which the antibodies were raised are not closely related and do not

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Fig. 6 Pitfalls of the BrdU labeling procedure on whole-mount Hydra. Detection of BrdU-labeled cells (green) on whole-mount animals requires a strong DNA denaturation obtained with HCl treatment (a). It can fail after a too partial DNA renaturation followed by an extensive washing step (b), or after an insufficient permeabilization of the tissues, which leads to the lack of penetration of the anti-BrdU antibody (c). Images were acquired with a Leica DM5500 fluorescence microscope. Scale bars: 50 μm

cross-­react. For instance, a combo of rat and mouse primary antibodies should be avoided, as both the anti-mouse and the anti-rat secondary antibodies recognize the antibodies from the other species. 22. When selecting the secondary antibodies, consider those that recognize specifically the species in which the primary antibody was raised. Moreover, as a large choice of antibodies coupled with different fluorochromes are commercially available, select the antibodies labeled with fluorochromes that ensure a bright and stable fluorescence with a reduced background, according to the available equipment for image acquisition. Fluorochromes are sensitive to light, to avoid photobleaching, protect from light the samples at all steps following incubation with the secondary antibody by covering the tubes/slides with aluminum foil, or hiding them in a drawer. 23. When BrdU immunodetection is performed, DAPI is a convenient nuclear stain. In contrast, Hoechst-33258 or Hoechst-33342 should be avoided as the intensity of the fluorescence staining is quenched by the thymidine analogs such as BrdU incorporated into DNA [62]. 24. When mounting, collect the animals in the minimum amount of liquid and place them onto the slide. Remove all excess of the liquid by pipetting up or by draining with a tissue paper. Add one or two drops of mounting medium and place the animals by softly pushing them with a pincer. When applying the coverslip, avoid the formation of air bubbles. Let the slides dry for a few hours at RT protected from light and transfer them at −20 °C.

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Acknowledgments This work was supported by the Swiss National Science Foundation (SNF grants 31003A_149630, 31003_169930), the Claraz donation, and the Canton of Geneva. References 1. Koizumi O (2016) Origin and evolution of the nervous system considered from the diffuse nervous system of cnidarians. In: Goffredo S, Dubinsky Z (eds) The cnidaria, past, present and future. Springer International Publishing, Cham, pp 73–91 2. Galliot B, Quiquand M, Ghila L, de Rosa R, Miljkovic-Licina M, Chera S (2009) Origins of neurogenesis, a cnidarian view. Dev Biol 332:2–24 3. Takahashi T, Koizumi O, Ariura Y, Romanovitch A, Bosch TC, Kobayakawa Y, Mohri S, Bode HR, Yum S, Hatta M et al (2000) A novel neuropeptide, Hym-355, positively regulates neuron differentiation in Hydra. Development 127:997–1005 4. Chera S, Ghila L, Dobretz K, Wenger Y, Bauer C, Buzgariu W, Martinou JC, Galliot B (2009) Apoptotic cells provide an unexpected source of Wnt3 signaling to drive hydra head regeneration. Dev Cell 17:279–289 5. Richards GS, Simionato E, Perron M, Adamska M, Vervoort M, Degnan BM (2008) Sponge genes provide new insight into the evolutionary origin of the neurogenic circuit. Curr Biol 18:1156–1161 6. Teragawa CK, Bode HR (1995) Migrating interstitial cells differentiate into neurons in hydra. Dev Biol 171:286–293 7. Fujisawa T, Nishimiya C, Sugiyama T (1986) Nematocyte differentiation in hydra. Curr Top Dev Biol 20:281–290 8. Tardent P (1995) The cnidarian cnidocyte, a high-tech cellular weaponry. BioEssays 17:351–362 9. Galliot B, Quiquand M (2011) A two-step process in the emergence of neurogenesis. Eur J Neurosci 34:847–862 10. Bode HR (1992) Continuous conversion of neuron phenotype in hydra. Trends Genet 8:279–284 11. Koizumi O (2007) Nerve ring of the hypostome in hydra: is it an origin of the central nervous system of bilaterian animals? Brain Behav Evol 69:151–159 12. Grimmelikhuijzen CJP, Westfall JA (1995) The nervous systems of Cnidarians. In: Breidbach

O, Kutsch W (eds) The nervous systems of invertebrates: an evolutionary and comparative approach. Birkhaüser Verlag, Basel, pp 7–24 13. Grunder S, Assmann M (2015) Peptide-gated ion channels and the simple nervous system of Hydra. J Exp Biol 218:551–561 14. Anderson PA, Spencer AN (1989) The importance of cnidarian synapses for neurobiology. J Neurobiol 20:435–457 15. Kass-Simon G, Pierobon P (2007) Cnidarian chemical neurotransmission, an updated overview. Comp Biochem Physiol A Mol Integr Physiol 146:9–25 16. Steinmetz PRH, Kraus JEM, Larroux C, Hammel JU, Amon-Hassenzahl A, Houliston E, Wörheide G, Nickel M, Degnan BM, Technau U (2012) Independent evolution of striated muscles in cnidarians and bilaterians. Nature 487:231–234 17. Grens A, Mason E, Marsh JL, Bode HR (1995) Evolutionary conservation of a cell fate specification gene: the Hydra achaete-scute homolog has proneural activity in Drosophila. Development 121:4027–4035 18. Gauchat D, Kreger S, Holstein T, Galliot B (1998) prdl-a, a gene marker for hydra apical differentiation related to triploblastic pairedlike head-specific genes. Development 125:1637–1645 19. Gauchat D, Escriva H, Miljkovic-Licina M, Chera S, Langlois MC, Begue A, Laudet V, Galliot B (2004) The orphan COUP-TF nuclear receptors are markers for neurogenesis from cnidarians to vertebrates. Dev Biol 275:104–123 20. Miljkovic-Licina M, Gauchat D, Galliot B (2004) Neuronal evolution: analysis of regulatory genes in a first-evolved nervous system, the hydra nervous system. Biosystems 76:75–87 21. Miljkovic-Licina M, Chera S, Ghila L, Galliot B (2007) Head regeneration in wild-type hydra requires de novo neurogenesis. Development 134:1191–1201 22. Wenger Y, Buzgariu W, Galliot B (2016) Loss of neurogenesis in Hydra leads to compensatory regulation of neurogenic and neurotrans-

Methods to Monitor Adult Neurogenesis in Hydra mission genes in epithelial cells. Philos Trans R Soc Lond Ser B Biol Sci 371:20150040 23. Wenger Y, Buzgariu W, Perruchoud C, Loichot G, Galliot B (2019) Generic and contextdependent gene modulations during Hydra whole body regeneration. BioRXiv 587147. https://doi.org/10.1101/587147 24. Darmer D, Hauser F, Nothacker HP, Bosch TC, Williamson M, Grimmelikhuijzen CJ (1998) Three different prohormones yield a variety of Hydra-RFamide (Arg-Phe- NH2) neuropeptides in Hydra magnipapillata. Biochem J 332:403–412 25. Hansen GN, Williamson M, Grimmelikhuijzen CJ (2000) Two-color double-labeling in situ hybridization of whole-mount Hydra using RNA probes for five different Hydra neuropeptide preprohormones: evidence for colocalization. Cell Tissue Res 301:245–253 26. Hansen GN, Williamson M, Grimmelikhuijzen CJ (2002) A new case of neuropeptide coexpression (RGamide and LWamides) in Hydra, found by whole-mount, two-color doublelabeling in situ hybridization. Cell Tissue Res 308:157–165 27. Hwang JS, Ohyanagi H, Hayakawa S, Osato N, Nishimiya-Fujisawa C, Ikeo K, David CN, Fujisawa T, Gojobori T (2007) The evolutionary emergence of cell type-specific genes inferred from the gene expression analysis of Hydra. Proc Natl Acad Sci U S A 104:14735–14740 28. Hayakawa E, Fujisawa C, Fujisawa T (2004) Involvement of Hydra achaete-scute gene CnASH in the differentiation pathway of sensory neurons in the tentacles. Dev Genes Evol 214:486–492 29. Chera S, Kaloulis K, Galliot B (2007) The cAMP response element binding protein (CREB) as an integrative HUB selector in metazoans: clues from the hydra model system. Biosystems 87:191–203 30. Lindgens D, Holstein TW, Technau U (2004) Hyzic, the Hydra homolog of the zic/oddpaired gene, is involved in the early specification of the sensory nematocytes. Development 131:191–201 31. Hartl M, Mitterstiller AM, Valovka T, Breuker K, Hobmayer B, Bister K (2010) Stem cellspecific activation of an ancestral myc protooncogene with conserved basic functions in the early metazoan Hydra. Proc Natl Acad Sci U S A 107:4051–4056 32. Ambrosone A, Marchesano V, Tino A, Hobmayer B, Tortiglione C (2012) Hymyc1 downregulation promotes stem cell proliferation in Hydra vulgaris. PLoS One 7:e30660

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33. Juliano CE, Reich A, Liu N, Götzfried J, Zhong M, Uman S, Reenan RA, Wessel GM, Steele RE, Lin H (2014) PIWI proteins and PIWI-interacting RNAs function in Hydra somatic stem cells. Proc Natl Acad Sci U S A 111:337–342 34. Takaku Y, Hwang JS, Wolf A, Bottger A, Shimizu H, David CN, Gojobori T (2014) Innexin gap junctions in nerve cells coordinate spontaneous contractile behavior in Hydra polyps. Sci Rep 4:3573 35. Engel U (2001) A switch in disulfide linkage during minicollagen assembly in Hydra nematocysts. EMBO J 20:3063–3073 36. Engel U, Ozbek S, Streitwolf-Engel R, Petri B, Lottspeich F, Holstein TW (2002) Nowa, a novel protein with minicollagen Cys-rich domains, is involved in nematocyst formation in Hydra. J Cell Sci 115:3923–3934 37. Koch AW, Holstein TW, Mala C, Kurz E, Engel J, David CN (1998) Spinalin, a new glycineand histidine-rich protein in spines of Hydra nematocysts. J Cell Sci 111:1545–1554 38. Adamczyk P, Meier S, Gross T, Hobmayer B, Grzesiek S, Bachinger HP, Holstein TW, Ozbek S (2008) Minicollagen-15, a novel minicollagen isolated from Hydra, forms tubule structures in nematocysts. J Mol Biol 376:1008–1020 39. Hwang JS, Takaku Y, Momose T, Adamczyk P, Ozbek S, Ikeo K, Khalturin K, Hemmrich G, Bosch TC, Holstein TW et al (2010) Nematogalectin, a nematocyst protein with GlyXY and galectin domains, demonstrates nematocyte-specific alternative splicing in Hydra. Proc Natl Acad Sci U S A 107:18539–18544 40. Balasubramanian PG, Beckmann A, Warnken U, Schnolzer M, Schuler A, Bornberg-Bauer E, Holstein TW, Ozbek S (2012) Proteome of Hydra nematocyst. J Biol Chem 287:9672–9681 41. David CN (1973) A quantitative method for maceration of hydra tissue. Wilhelm Roux Arch Dev Biol 171:259–268 42. Bode HR, Berking S, David C, Gierer A, Schaller H, Trenker E (1973) Quantitative analysis of cell types during growth and regeneration in hydra. Wilhelm Roux Arch Entw Mech Org 171:269–285 43. Fujisawa T (1989) Role of interstitial cell migration in generating position-dependent patterns of nerve cell differentiation in Hydra. Dev Biol 133:77–82 44. Technau U, Holstein TW (1996) Phenotypic maturation of neurons and continuous precursor migration in the formation of the

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peduncle nerve net in Hydra. Dev Biol 177:599–615 45. Campbell RD (1976) Elimination by Hydra interstitial and nerve cells by means of colchicine. J Cell Sci 21:1–13 46. Marcum BA, Campbell RD (1978) Development of Hydra lacking nerve and interstitial cells. J Cell Sci 29:17–33 47. Marcum BA, Fujisawa T, Sugiyama T (1980) A mutant hydra strain (sf-1) containing temperature-sensitive interstitial cells. In: Tardent P, Tardent R (eds) Developmental and cellular biology of coelenterates. Elsevier, Amsterdam, pp 429–434 48. Takahashi T, Fujisawa T (2009) Important roles for epithelial cell peptides in hydra development. BioEssays 49. Takahashi T, Takeda N (2015) Insight into the molecular and functional diversity of cnidarian neuropeptides. Int J Mol Sci 16:2610–2625 50. Plickert G, Kroiher M (1988) Proliferation kinetics and cell lineages can be studied in whole mounts and macerates by means of BrdU/antiBrdU technique. Development 103:791–794 51. Bode H, Lengfeld T, Hobmayer B, Holstein TW (2008) Detection of expression patterns in Hydra pattern formation. Methods Mol Biol 469:69–84 52. Kaloulis K, Chera S, Hassel M, Gauchat D, Galliot B (2004) Reactivation of developmental programs: the cAMP-response elementbinding protein pathway is involved in hydra head regeneration. Proc Natl Acad Sci U S A 101:2363–2368 53. Grimmelikhuijzen CJ (1985) Antisera to the sequence Arg-Phe-amide visualize neuronal centralization in hydroid polyps. Cell Tissue Res 241:171–182

54. Koizumi O, Bode HR (1991) Plasticity in the nervous system of adult hydra. III. Conversion of neurons to expression of a vasopressin-like immunoreactivity depends on axial location. J Neurosci 11:2011–2020 55. Dunne JF, Javois LC, Huang LW, Bode HR (1985) A subset of cells in the nerve net of Hydra oligactis defined by a monoclonal antibody: its arrangement and development. Dev Biol 109:41–53 56. Galliot B, Welschof M, Schuckert O, Hoffmeister S, Schaller HC (1995) The cAMP response element binding protein is involved in hydra regeneration. Development 121:1205–1216 57. Chera S, Ghila L, Wenger Y, Galliot B (2011) Injury-induced activation of the MAPK/CREB pathway triggers apoptosis-induced compensatory proliferation in hydra head regeneration. Develop Growth Differ 53:186–201 58. Ott SR (2008) Confocal microscopy in large insect brains: zinc-formaldehyde fixation improves synapsin immunostaining and preservation of morphology in whole-mounts. J Neurosci Methods 172:220–230 59. Loomis WF (1956) Growth and sexual differentiation of hydra in mass culture. J Exp Zool 132 60. Gierer A, Berking S, Bode H, David CN, Flick K, Hansmann G, Schaller H, Trenkner E (1972) Regeneration of hydra from reaggregated cells. Nat New Biol 239:98–101 61. Macklin M (1976) The effect of urethan on hydra. Biol Bull 150:442–452 62. Latt SA (1973) Microfluorometric detection of deoxyribonucleic acid replication in human metaphase chromosomes. Proc Natl Acad Sci U S A 70:3395–3399

Chapter 2 Reverse Genetic Approaches to Investigate the Neurobiology of the Cnidarian Sea Anemone Nematostella vectensis Jamie A. Havrilak and Michael J. Layden Abstract The cnidarian sea anemone Nematostella vectensis has grown in popularity as a model system to complement the ongoing work in traditional bilaterian model species (e.g. Drosophila, C. elegans, vertebrate). The driving force behind developing cnidarian model systems is the potential of this group of animals to impact EvoDevo studies aimed at better determining the origin and evolution of bilaterian traits, such as centralized nervous systems. However, it is becoming apparent that cnidarians have the potential to impact our understanding of regenerative neurogenesis and systems neuroscience. Next-generation sequencing and the development of reverse genetic approaches led to functional genetics becoming routine in the Nematostella system. As a result, researchers are beginning to understand how cnidarian nerve nets are related to the bilaterian nervous systems. This chapter describes the methods for morpholino and mRNA injections to knockdown or overexpress genes of interest, respectively. Carrying out these techniques in Nematostella requires obtaining and preparing embryos for microinjection, designing and generating effective morpholino and mRNA molecules with controls for injection, and optimizing injection conditions. Key words Nematostella, Morpholino, mRNA, Embryo microinjection, Gene knockdown, Overexpression

1  Introduction The desire to investigate neuronal development and function in cnidarians (sea anemones, corals, jellyfish, hydras, etc.) is driven by the potential that they have to impact our understanding of the origin and evolution of bilaterian nervous systems, including centralization into a brain and nerve cords. Cnidarians have nerve net nervous systems (Fig. 1a, b), and are thought to represent the morphology of the ancestral nervous systems that gave rise to centralized nervous systems. Functional experiments and conserved expression of neurogenic genes in progenitors and differentiating neurons provide strong arguments that cnidarian and bilaterian Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_2, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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Fig. 1 Introduction to Nematostella vectensis. (a) Image of an adult Nematostella. (b) Nvelav1::mOrange transgene shows the nerve net of a juvenile polyp. The longitudinal tracts of bundled neurites are detectable. (c) Phylogeny showing relative position of the five major metazoan clades. (d) Schematic representation of Nematostella during development. Orange indicates endoderm. White indicates ectoderm. Green bar indicates stages where neurogenesis occurs. In all images, oral is up. Images (a), (b), and (d) are adapted from [4]

nervous systems arose from a common ancestor [1–10]. Because Cnidaria is the closest phylum to the Bilateria (Fig. 1c) [11, 12], it is the critical out group to study in order to reconstruct the ancestral neurobiology from which complex brains and centralized nervous systems emerged. Reconstruction of the ancestral state provides researchers with the necessary knowledge to infer what key changes in nervous system development and neurobiology led to the emergence of centralized nervous systems and impacted the evolution of nervous systems. In addition to their ability to impact our understanding of nervous system evolution, many cnidarian species are relatively small and optically clear animals that are highly regenerative. These properties make them well suited for whole animal imaging of neuronal activity at single cell resolution, suggesting that this group of animals will also impact our understanding of systems neuroscience and neuronal regeneration [8, 13–16]. Recent advances in low-­ cost next-generation sequencing technology and reverse genetic

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approaches have made investigating cnidarian biology much more accessible than even a decade ago [4, 17–23]. As a result, gene disruption using reverse genetic approaches ranging from CRISPR/Cas9-mediated gene editing, transgenesis, RNA interference, and mRNA misexpression have been reported in multiple cnidarian species [4, 10, 18, 19, 24–26]. To date the anthozoan sea anemone Nematostella vectensis is the most established cnidarian to carry out functional genetic approaches because it has a variety of techniques available to disrupt gene function, assay neurogenic phenotypes, and it has a rapidly growing list of publically available data sets using a variety of next-generation sequencing approaches [4, 5, 16, 27, 28]. Nematostella is an anthozoan cnidarian and has the cnidarian polyp bauplan (Fig. 1a, d). They are diploblastic animals possessing an endoderm (sometimes referred to as the endomesoderm/ mesendoderm) and ectoderm, which makes them effectively two cell layers thick. The polyp possesses a single oral opening derived from the blastopore that leads to a sac-like gut. The mouth is surrounded by tentacles that are used to capture prey and transport it to the oral opening. There are eight mesenteries that run the length of the oral–aboral axis that contain contractile muscles, cnidoglandular tracts, the gonads, and bundles of neurons. The Nematostella nerve net is comprised of both endodermal and ectodermal nerve nets formed from neurons that are likely generated in the tissue layer in which they reside [29]. As a result, the nerve nets are essentially a two-dimensional array of interconnected neurons. Coupled with the fact that it is optically clear at all life stages, this makes the Nematostella nerve net one of the most accessible and easily visible nervous systems for in vivo investigation. Neuronal development initiates at blastula stages in the future ectoderm as Notch-Delta signaling acts to select which cells will respond to MAPK signaling and become NvsoxB [2] and Nvatonal-­ like (Nvath-like) positive neuronal progenitor cells [2, 3, 30, 31]. Post-mitotic daughter cells express differentiation markers that include Nvachaete-scute homolog A (NvashA). Neuronal progenitors and differentiating neurons are detected in early planula larval stages shortly after gastrulation is completed, and neurogenesis continues until the juvenile polyp stages [1, 2, 32]. Until recently, nerve nets were thought to be minimally organized and to lack stereotypy. However, transgenic analysis of individual neuronal subtypes suggests that patterning individual neuronal fates throughout development is a highly stereotyped process [31, 32]. The exact molecular mechanisms that pattern individual neuronal fates are not yet known, but axial patterning programs along the oral–aboral axis and the directive axis (perpendicular to the oral– aboral axis), as well as temporal cues, are thought to contribute to neural subtype specification [32–36]. The next generation of functional studies will be to better understand how individual neuronal

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subtypes are patterned and to better determine how regenerative neurogenesis occurs. A yet untapped area of cnidarian neurobiology will be to deploy functional gene disruptions for the purpose of investigating neuronal physiology and systems neuroscience to better understand how similar cnidarian neuronal physiology is to the physiology of bilaterian neurons. Additional efforts should focus on the use of functional genetics to query how cnidarian nerve nets perceive and interpret inputs to generate appropriate responses. Marrying insights from developmental neurobiology to those of neurophysiology will undoubtedly provide key data about how similar or dissimilar cnidarian and bilaterian nervous systems are, and ultimately provide insights about how nervous systems evolve. Morpholino-mediated gene knockdown and mRNA misexpression are the most widely used methods of gene disruption in Nematostella. mRNA injections date back to 2003 and morpholino injections were first reported in Nematostella in 2007 [37, 38]. TALEN and CRISPR/Cas9-mediated gene editing have also been reported as methods for gene knockdown in Nematostella [19]. Each of these approaches is still in its infancy in this system, and as of yet are not widely used. It is likely that any or all of the alternative gene knockdown approaches will one day be standard practice in most Nematostella laboratories. The availability of multiple methods is crucial, as it provides an opportunity to choose the approach that best fits a particular experimental design. Other than mRNA injection, the only alternative gain of function approach that hyperactivates a single gene is protein soaking, but this is limited to proteins that act as morphogens or signaling molecules. Transgenics offer alternative methods to introduce or misexpress genes of interest, but the number of available enhancers is currently limited. To date, mRNA injections remain the most reliable gain of function approach available in the Nematostella system. To perform morpholino and mRNA injections in Nematostella requires designing the mRNA and/or morpholino and planning appropriate controls. The exact design of individual mRNAs will depend on the particular biology of each protein of interest. In general, we observe 100% of injected animals displaying expression (Fig. 2a–g). However, cloning fluorescent reporters in frame with the gene of interest offers an effective strategy to ensure each mRNA is properly translated [1, 18]. Morpholinos can be designed to act as splice blocking or translation blocking morpholinos (Fig. 3). Design of morpholinos is most effective if done in collaboration with the design team at Gene Tools (http://www.genetools.com). Controls for mRNA injection should include the injection of mRNA encoding just the fluorescent tag to ensure phenotypes are not the result of a load on the translation machinery resulting from the injected mRNA. It is also recommended to perform a rescue experiment by co-injecting a morpholino that

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Fig. 2 mRNA injections in Nematostella. (a–c) Close-up of a single embryo injected with an mRNA encoding the NvashA transcription factor with a 3′ venus tag (NvashA:venus). The nuclear Venus localization is indicated by colocalization with the Hoechst DNA stain. (d–g) Embryos 24 h post injection (hpi) that were injected with NvashA:venus and the Rhodamine Dextran tracer dye. (h–k) Control embryos 24 hpi after injection with only the Rhodamine Dextran tracer dye. Images (d)–(k) are adapted from [18]

targets the endogenous mRNA and/or the injected mRNA. The rescue experiment ensures that all observed phenotypes are specific to the overexpressed gene and not an artifact of the injection. Controls for morpholino injections should include injecting mismatch morpholinos, which confirms that resulting phenotypes are specific for the targeted reduction in gene activity and not the morpholino injection itself. Similar to mRNA injection, a rescue experiment is a recommended additional control. The simplest experiment is to co-inject the morpholino with an mRNA that encodes the target gene but cannot be recognized by the morpholino (Fig. 3a). Phenotypes due to knockdown of the target gene should be rescued by co-injection of the morpholino and mRNA, whereas potential off-target effect phenotypes will remain. Additional requirements for morpholino injections are efforts to ensure the morpholino is effectively impacting the intended target. The activity of splice blocking morpholinos can be measured by PCR (see Fig. 3b, c) [18, 30]. Translation blocking morpholinos can be measured for activity by observing reduced antibody

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5’

TTTTACTCACATGAGACGGTAGT(ATG)CCTCAGCTTCCTAGGAA 3’

Nv-tcf:Venus Nv-tcf5’:Venus

5’ 5’

(ATG)CCTCAGCTTCCTAGGAA 3’ CACATGAGACGGTAGT(ATG)CCTCAGCTTCCTAGGAA 3’

MoTcf_trans

5’

Nv-tcf:Venus

CACATGAGACGGTAGT(ATG)CCTCAG

Nv-tcf:Venus + MoTCF

3’

Nv-tcf5’:Venus + MoTCF

Nv-tcf5’:Venus

B F1

R2

MoEtsA1

F1:R2 - unspliced:

~ 800 bp -

F1:R2 - spliced:

~ 440 bp -

Control

MoEtsA1

C Ladder

Control

Morpholino

Ladder

Control

Morpholino

Fig. 3 Morpholino injections in Nematostella. (a) Sequence of the 5′ UTR and ATG start site (in parentheses) of the Nv-tcf gene, as well as the sequence covered by the Nv-tcf translation blocking morpholino (red). Sequences for the beginning of two Nv-tcf:Venus mRNA constructs are also shown. One encoding a construct that cannot be targeted by the morpholino (Nv-tcf:Venus) and one that can be targeted by the morpholino (Nv-tcf5′:Venus). Below are images of animals injected with the indicated combinations. The Nv-tcf morpholino is effective at inhibiting translation of only the Nv-tcf5′:Venus construct containing the morpholino binding site. (b) Schematic of the PCR primers (F1 and R2) used to amplify the region that contains an intron targeted by the MoEtsA1 morpholino. Below is an agarose gel showing miss-splicing that results in an increase in the size of the amplified Ets fragment in MoEtsA1 but not control-injected animals. (c) Examples of splice blocking morpholinos that show the expected increased size of the PCR product (left and right). However, the morpholino injection shown on the left also resulted in a splicing at a cryptic splice site that resulted in a fragment that is actually smaller than the control fragment. Images in (a) and (b) are adapted from [18]

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staining against the protein product of the targeted gene, carrying out in vitro translation assays [2, 39], or co-injecting the morpholino and a fluorescently tagged mRNA that can be targeted by the morpholino of interest. The activity of the morpholino can be measured by observing reduced fluorescence resulting from translation of the injected mRNA. For each mRNA and morpholino, the optimized concentration for injection must be empirically determined by injecting a range of concentrations and assessing toxicity, as well as testing the effectiveness of each reagent (see above and Figs. 2 and 3) [18]. Prior to injection, eggs must be obtained, fertilized, and de-­ jellied to allow access for injection. To obtain embryos for injection, adult animals are placed on a light box or in a 25 °C incubator with a light timed to turn on at the appropriate hour to induce spawning within the desired timeframe the next morning [40]. After spawning, eggs from female-only bowls are placed in water containing males for ~10 min to allow fertilization to occur. Following fertilization, eggs are de-jellied in a cysteine solution for ~20 min and then washed in filtered 1/3× artificial seawater. Eggs are then placed in injection dishes as described below and are ready to inject. Both mRNA and morpholinos are introduced by microinjection into oocytes using forced air injections. At 17 °C there is a 170-min window to inject between fertilization and first cleavage [40]. Following injection, embryos are aged at the desired temperature and to the desired stage for analysis. Previous reports have indicated that morpholinos are effective up to a week post injection, whereas mRNAs will vary based on the rate at which each individual mRNA is targeted for degradation.

2  Materials 2.1  Embryo Preparation

1. 1/3× Artificial Sea Water (ASW): To 900 ml dH2O, add 12 g sea salt (Instant Ocean). Adjust to pH 8.1–8.2. Mix thoroughly and bring volume to 1 L with dH2O. The seawater should have a salinity of 12 parts per thousand. Sterile-filter the 1/3× ASW through a 0.2-μm Nalgene filter and aerate by shaking bottle following filtration. Store indefinitely at 17–22 °C. 2. 4% l-Cysteine (de-jellying solution): Dissolve 0.4 g of l- Cysteine (Sigma-Aldrich, see Note 1) in 10 ml 1/3× ASW (4% wt/vol with cysteine), adjust pH to 7.4–7.6 by adding 2 to 3 drops of 5 M NaOH. Make fresh for each use [40, 41].

2.2  Morpholino and mRNA Preparation

1. 100× Alexa Fluor dye stock solution: Make a 20 mg/ml concentration by dissolving the Alexa Fluor dye (Dextran, Alexa Fluor 488 or 555) in Non-DEPC-treated nuclease-free water (see Note 2). Assay for RNase activity by incubating the solu-

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tion with RNA at 37 °C for 1 h, then running it on an agarose gel to test for degradation. Aliquot and store at −20 °C for up to 2 years. 2. 5× Alexa Fluor dye stock solution: Dilute 100× stock 1:20 with sterile non-DEPC-treated nuclease-free water. 3. Ambion mMessage mMachine T3 Transcription kit. 4. Ambion MEGAclear purification kit. 5. 1% (wt/vol) agarose gel. 6. Nuclease-free water. 7. Phenol:chloroform:isoamyl purification).

alcohol

solution

(for

DNA

8. Isopropanol. 9. Glycogen. 10. RNase-free 70% (wt/vol) ethanol. 11. RNase-free 75% (wt/vol) ethanol. 12. RNase-free 100% ethanol. 13. Nuclease-free water. 14. RNase-free DNase. 15. MEGAclear purification kit. 16. Tris saturated phenol:chloroform:isoamyl alcohol (pH 6.8). 2.3  Equipment

1. Dissecting microscope (microinjection/injection apparatus). 2. Fluorescent apparatus).

lamp

3. Micromanipulator apparatus).

and

filters

apparatus

(microinjection/injection (microinjection/injection

4. Model PLI-OHN output hose (1-mm tubing) (injection apparatus). 5. Wall Superthane ester-based tubing 1/8 in, 130 PSI at 70 F (injection apparatus). 6. Picospritzer with external air source (injection apparatus). 7. Stainless steel pipette holder, 130 mm for 1–1.5 mm outer diameter glass pipettes (injection apparatus). 8. Glass needles with filament, 1 mm, thin-walled glass capillaries with filament (microinjection). 9. Electrode storage jar (microinjection). 10. Glass micropipette puller (injection prep). 11. Falcon disposable Petri dishes, 60 × 15 mm (Corning, 351007) (see Note 3) (microinjection). 12. Pasteur pipettes (embryo prep and microinjection).

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13. 15 mL conical tubes (embryo prep). 14. RNase-free 1.5 mL microcentrifuge tubes (mRNA prep). 15. Orbital shaker or rocker (embryo prep). 16. Microcentrifuge (injection prep). 17. Nalgene vacuum filter (embryo prep). 18. Incubator (animal care). 19. Thermocycler.

3  Methods 3.1  Morpholino Preparation

1. Design control and splice- or translation- blocking morpholinos against your gene of interest in collaboration with the GeneTools design team (see Note 4) (Gene Tools, LLC) (http://www.gene-tools.com). 2. Make 3 mM stock solution of morpholino by resuspending it in the appropriate volume of nuclease-free water. Aliquot and store stock solutions at −22 °C (see Note 5). 3. Prepare the morpholino injection mix by adding 3 μl of the 5× Alexa Fluor of choice, x μl of the morpholino stock to achieve desired concentration (see Note 6) and x μl nuclease-free water for a total volume of 15 μl. 4. Microcentrifuge the morpholino preparation at 13,000 × g at 4 °C to pellet any debris (see Note 7). 5. Optimize morpholino concentration for microinjection (see Note 6). 6. Assay for morpholino efficiency (see Notes 8–10) (Fig. 3). 7. Assay for toxicity (see Note 11).

3.2  mRNA Preparation

1. Linearize 5–10 μg of the transcription vector containing the coding sequence for your gene of interest (see Notes 12 and 13). 2. Ensure complete digestion by running 100–200 ng of digested DNA on a 1% (wt/vol) agarose gel. 3. Add nuclease-free water to the remaining digested sample for a total volume of 200 μl. 4. Add 200 μl phenol:chloroform:isoamyl alcohol to the digested sample and shake vigorously for 15 s. Incubate at room temperature for 5 min. 5. Centrifuge at 12,500 × g for 15 min at 4 °C. 6. Transfer the aqueous phase to an RNase-free microcentrifuge tube.

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7. Add 200 μl isopropanol and 0.5 μl of 20 mg/ml glycogen to the solution and mix well. Incubate the solution for 10 min at room temperature. 8. Centrifuge at 12,500 × g for 15 min at 4 °C. 9. Remove liquid, being careful not to disturb the pellet. 10. Add 1 ml of RNase-free 70% (wt/vol) ethanol and vortex briefly to wash the pellet (see Note 14). Centrifuge the DNA at 7500 × g for 5 min at room temperature. 11. Remove the ethanol with a pipette and air-dry the pellet in a hood for ~10–20 min, or dry the pellet with a SpeedVac. 12. Resuspend the pellet in 20 μl of nuclease-free water. 13. Determine DNA concentration with a spectrophotometer by measuring the OD at 260 nm (OD260) (see Note 15). 14. Set up the transcription reaction as per manufacturer instructions. We describe here the protocol for Ambion mMessage mMachine T3 Transcription kit and MEGAclear purification kit. 15. Use up to 1 μg of linearized DNA template (up to 6 μl), with a final reaction volume of 20 μL (see Note 16). 16. Use a thermocycler to incubate the reaction mixture for 2 h at 37 °C (see Note 17). 17. Add 1 μl of RNase-free DNase to the reaction and mix gently by pipetting. Incubate the mixture again for 15 min at 37 °C. 18. Run 0.5  μl of the reaction mixture on a 1% (wt/vol) agarose gel to verify transcription. 19. Purify per manufacturer’s instructions using MEGAclear purification kit. 20. To 100  μl of eluted RNA from step 19, add 1:1 volume (100 μl) of Tris saturated phenol:chloroform:isoamyl alcohol (pH 6.8) and shake vigorously for 15 s (see Note 18). Incubate for 5 min at room temperature. 21. Spin the tube at 12,500 × g in a microcentrifuge for 15 min at 4 °C. 22. Remove the aqueous phase with a pipette and transfer it to a new RNase-free microcentrifuge tube. 23. Add an equal volume of chloroform. Shake vigorously and incubate as in step 20 and then repeat steps 21 and 22. 24. To precipitate the mRNA, add 10 μl of 5 M ammonium acetate (provided in the MEGAclear kit) and 275 μl RNase-free 100% ethanol. Precipitate the mRNA overnight at −20 °C (see Note 19).

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25. Pellet the mRNA by spinning in a microcentrifuge at 12,500 × g for 15 min at 4 °C. Remove the supernatant with a pipette (see Note 20). 26. Add 1 ml of ice-cold RNase-free 75% (wt/vol) ethanol to wash the mRNA pellet and vortex briefly. Spin at 12,500 × g for 5 min at 4 °C. Remove the ethanol with a pipette. 27. Repeat the wash step by adding 1 ml of ice-cold RNase-free 75% (wt/vol) ethanol to wash the mRNA pellet and vortex briefly. Spin at 12,500 × g for 5 min at 4 °C. 28. Remove the ethanol and air-dry the pellet for ~10–20 min at room temperature in a hood (see Note 21). 29. Add 30  μl of nuclease-free water and pipette gently to resuspend the pellet. 30. Determine mRNA concentration with a spectrophotometer (see Note 22). 31. To ensure that the mRNA is not degraded, run 1 μg of the mRNA sample on a 1% (wt/vol) agarose gel and ensure only a single band with no lower molecular weight smearing is present. The mRNA can be stored at −80 °C for up to a year. 32. Determine optimal mRNA concentration (see Note 6). 33. Score for effectiveness of the mRNA (Fig. 2) (see Notes 23 and 24). 34. Assay for toxicity (see Note 11). 35. To prepare the injection mixture, combine 3 μl of 5× Alexa Fluor, x μl of the mRNA sample to obtain the desired concentration, and x μl nuclease-free water for a total volume of 15 μl. 36. Microcentrifuge the mRNA preparation mixture at 13,000 × g at 4 °C to pellet any debris (see Note 7). 3.3  Equipment Setup

1. Pulling glass needles: There are many variables that can affect how the glass pipettes are pulled for injections (e.g., the puller machine/model being used, environmental factors, and personal preference for thinness of the needle) (see Note 25). We recommend using these parameters as a starting point when using the Sutter Instrument Co Model P-97 Flaming/Brown Micropipette Puller and then adjusting for preferences: Heat = 545; P = 200; Pull = 90; Vel = 80; Del = 80. 2. Microinjection setup: The basic injection setup requires a picospritzer, micromanipulators, dissecting scope with sufficient working distance, and external white and fluorescent light sources. There are multiple brands and models for each component. In addition to our setup there is an alternative injection rig setup that is described in [42]. Both are effective, and your preference will determine which system is best for you.

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Jamie A. Havrilak and Michael J. Layden

An example of our setup is shown in Fig. 4a. Features to consider are: (1) Having a foot pedal to initiate air pulses with the picospritzer. Having both hands available during injections improves efficiency. (2) If you do not have house air to feed into the picospritzer, either choose a model that has a selfcontained bladder or purchase an external compressor. (3) Whether using an external compressor or house air, attach an air filter to the lines to avoid oil and debris from damaging your picospritzer. (4) Having both course and fine micromanipulators to control the needle position. It is possible to inject with only a course manipulator, but it is slow and inefficient. (5) If possible, set up the injection rig in a cool (17 °C) climate-controlled room (see Note 26). (6) When beginning the injection, place embryos in a dish under the scope. Use the course manipulators to bring the needle into position and roughly center it in the field of view. 3.4  Embryo Preparation

1. Induce spawning by placing adult Nematostella on a light box or in a 25 °C incubator with a light controlled by a timer the evening before the day of injection. Gametes will be released between 13–15 h following the light and temperature increase [40, 41] (see Note 27). Before the 2-h spawning window remove animals from the light box and change the water, being careful to remove any night-spawned egg masses. Having bowls of female-only animals will enable you to control the timing of fertilization by placing those egg masses into spermcontaining bowls. 2. Place unfertilized spawned egg masses with sperm-containing 1/3× ASW and incubate for 10 min [40]. 3. Transfer fertilized embryos into a 15 ml conical tube containing 10 ml of freshly prepared 4% l-cysteine de-jellying solution [40] (see Note 1). 4. Gently rock the embryos at room temperature until the jelly is dissolved and the individual embryos are free (~20 min) (see Note 28). 5. Wash the embryos 3× with ~10 ml of 1/3× ASW, allowing embryos to settle between washes. 6. Wash the embryos with 10 ml of sterile-filtered 1/3× ASW, then transfer them to a glass dish containing sterile-filtered 1/3× ASW.

3.5  Microinjection

1. Pull glass micropipette needles [18] (see Note 29) and store them upright in an electrode storage jar. 2. Load needles with 0.5–0.75 μl of injection mixture (see Note 30). Let the needles sit for 3 min to allow for the mixture to move to the bottom of the needle.

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37

Fig. 4 Example of injection rig setup. (a) Injection setup around the dissecting scope. The numbers are as follows: 1—The micropipette holder that is attached by tubing to the picospritzer. 2—The course manipulator, which manipulates the micropipette holder. 3—The micromanipulator injector used during injection. 4—The single source oblique lighting used to illuminate with transmitted light. 5—The fluorescent light source. 6— The Picospritzer. 7—The foot pedal used to activate a pulse of air from the picospritzer during injection. 8— The electrode storage jar that holds the micropipettes prior to use. 9—The house air used to provide pressure to the picospritzer. (b) Example of embryos lined up in between the scratches in the injection dish prior to injection. (c) Examples of needles used for injection. Lines on scale indicate millimeters. (d) Schematic of the needle at a 45° angle and relative volume to inject

3. Scratch parallel lines (~5 mm apart) into the bottom of a plastic petri dish with forceps (see Notes 3 and 31) (Fig. 4b). 4. Add sterile-filtered 1/3× ASW to cover the bottom of the dish (see Note 32). 5. Use a Pasteur pipette to transfer embryos into the dish, being careful to line them up between the parallel scratches in the dish (see Note 33) (Fig. 4b). 6. Insert the pulled needle loaded with the injection mixture into a picospritzer needle holder and adjust it so that it is at a steep angle (>45°) (see Note 34) (Fig. 4d).

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Jamie A. Havrilak and Michael J. Layden

7. With the needle in the middle of the field of view at the lowest magnification, slowly lower it down through the water, increasing the magnification as the needle reaches the bottom of the dish. 8. Use the micromanipulator to break the tip of the needle by pushing the tip of the needle into the sides of the scratches or by tapping it on the bottom of the dish. Alternatively, a diamond pen can be used to break the tip of the needle (see Note 35). 9. Adjust the balance pressure on the picospritzer so that there is a visible but small amount of dye flowing out of the needle. 10. Estimating by eye, adjust the time and pressure on the picospritzer so that a single pulse fills ~5–10% of the embryo volume (see Note 36) (Fig. 4d). 11. Use the micromanipulator to insert the needle into the top of the embryo, stopping just below the surface, and inject the embryo with a single pulse. Inject all the embryos in the dish (see Note 33). 12. Remove any uninjected embryos (see Note 37). 13. Set up a second dish and inject with the appropriate control. 14. After completion of all injections, transfer the embryos to the desired temperature and age until the desired stage (see Note 38). 15. Score for phenotype.

4  Notes 1. It is important to use the l-cysteine recommended here, as we have found l-cysteines vary in their impurities, which can cause lethality in some cases. 2. Use non-DEPC-treated H2O, as DEPC interferes with the efficiency of the morpholino. 3. We have found the Corning Falcon Petri dishes (cat 351007 and 351008) work the best for adhering embryos to the bottom. However, it is also possible to coat injection dishes with polylysine to improve adhesion. 4. Due to the potential lack of availability of antibodies against Nematostella proteins, the splice-blocking morpholino may offer an advantage to the translation-blocking morpholino, because it is easier to quantify its effectiveness (see Notes 8–10). 5. If the morpholino precipitates out of solution, heat the aliquot of morpholino at 65 °C for 5 min [18].

Reverse Genetics in Nematostella

39

6. To determine the optimal morpholino or mRNA concentration, inject 100–300 embryos with a series of concentrations and test for toxicity. Effective morpholino concentrations range from 300–900 μm and effective mRNA concentrations range from 150–700 ng/μl [18]. 7. Debris in the injection cocktail can clog the micropipette needle. It is best to pipette from the top of the liquid opposite the side of any potential pellet. 8. The best method to determine effective morpholino injection is western blot analysis with an antibody that recognizes the endogenous target protein, allowing for the visualization of a reduction in protein levels [18, 43]. If no antibody exists against the protein target of interest, see Notes 9 and 10 to test for the effectiveness of translation-blocking and splice-­blocking morpholinos, respectively. 9. Test the effectiveness of a translation-blocking morpholino in the absence of an antibody against the target protein by co-­ injecting an in vitro-transcribed mRNA containing the morpholino binding site that encodes the target gene fused to a fluorescent reporter. Effective knock down can be visualized by a decrease in the fluorescence of the reporter with the co-­ injection compared to mRNA injected alone [18]. Alternatively, perform an in vitro translation assay with and without the morpholino present [39]. 10. Assay splice-blocking morpholino effectiveness by performing RT-PCR on cDNA from control- and morpholino- injected animals. Use PCR primers that will amplify a DNA fragment that includes the intron targeted by the splice-blocking morpholino. Run the product on an agarose gel to visualize size difference in PCR products. Defects in splicing can range from cryptic splice sites being activated, to a complete failure in splicing. The resulting defectively spliced RNAs will result in smaller or longer PCR fragments relative to those amplified from control animals (Fig. 3b, c). The efficiency of the morpholino can be estimated by comparing the relative strength of wild-type bands to mis-spliced bands [18, 30, 39, 44]. It is strongly recommended to subclone and sequence the mis-­ spliced band in order to reconstruct the potential protein products in silico to ensure that any potential protein generated in the morphant animals does not in any way recapitulate functional domains of the target protein. 11. Assay for toxicity by scoring the injected embryos for failed cleavage and/or death at ~4 h and 24 h. Concentrations that are too high are toxic to the embryos, which is often discernible by observing blastomeres disassociating in early cleavage stages. Additionally, death within the first 24 h post injection is

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uncommon and likely indicates toxicity rather than a specific phenotype. However, rescue experiments are necessary to definitively distinguish between the two possibilities. High mortality at 24 h post injection is indicated by the loss of intact embryos and excessive organic debris in the injection dish. 12. Before starting this protocol, cDNA for your gene of interest should be subcloned into a vector appropriate for in vitro transcription. This can be done using standard recombinant DNA techniques or by commercial gene synthesis. We have successfully used pCS2 and Gateway vectors, but any standard vector for in vitro mRNA translation should be effective. We also recommend, if possible, generating mRNAs encoding fluorescently tagged fusion proteins to make visualization of effective translation easy to quantify (see Notes 23 and 24). 13. Use the appropriate restriction enzyme for the destination vector of choice. 14. The DNA precipitate can be stored at 4 °C (for up to a week) or − 20 to −80 °C (for a year or more). 15. The precipitated DNA can be stored at 4 °C (for a week) or − 20 to −80 °C (for a year or more). 16. The Ambion (Invitrogen) mMessage mMachine T3 transcription kit can accommodate up to 6 μl linearized DNA for a final volume of 20 μl. Check the specific manufacturer’s instructions if using a different transcription kit. 17. We recommend carrying out the incubation using a thermocycler because it will keep a constant temperature and result in more consistent yields compared to a heat block or incubator. 18. Do not vortex. 19. If necessary, the sample can be precipitated for as little as 1 h at −20 °C. 20. Be careful to not disturb the pellet while removing the supernatant. 21. Do not allow the pellet to dry completely or it may become difficult to resuspend. 22. This protocol will typically yield 30–40 μg mRNA. 23. Score for effectiveness of mRNA by cloning the gene in frame with a fluorescently tagged protein (e.g. Venus, GFP) and observe directly with fluorescent imaging or by ­immunofluorescence (Fig.  2). We define optimal concentrations of mRNA as being able to observe the fluorescence from the reporter tag encoded within the injected mRNA and none to minimal observed mortality/toxicity at 24 h post injection. 24. Co-injecting the in vitro-transcribed mRNA with a fused fluorescent reporter protein with a translation-blocking morpho-

Reverse Genetics in Nematostella

41

lino targeting the same gene can be used as a control to test for induction of mRNA-induced nonspecific toxicity. Co-­injection with the morpholino should rescue any lethal phenotype induced by the overexpression of mRNA (Layden et al. [18]). 25. Nematostella embryos are tolerant of a wide range of needle shapes, but thinner needles will afford more control over injection volume/pulse. Additionally, thinner needles tend to be sharper and accumulate less debris, making it easier to inject more embryos. 26. A climate-controlled room is not necessary, but it allows for longer injection sessions yielding more injected animals. 27. The timer can be adjusted accordingly for individual injection timing needs. For example, a timer set to turn the light on at 10:30 pm will often result in spawns between 10:30–11:30 am the following morning. 28. Rock the embryos slowly. Rocking the embryos too hard will cause them to become elongated. 29. Nematostella embryos are robust and can handle different shaped needles. For optimal microinjections, needles should be as fine as possible. Pulling long and thin needles and then cutting them back until the desired flow is achieved (see Note 35 below) is easier than pulling the preferred size. 30. If the injection cocktail is evaporating from the needle, only load needles as needed. 31. The lines have several uses. (1) To act as guides to line up embryos in the dish, (2) to break the needle, and (3) for cleaning the tip of the needle during the injection. 32. Add sterile-filtered 1/3× ASW to the Petri dish immediately before adding the embryos. Adding water too soon will decrease adherence of the embryos to the dish. 33. Start loading the embryos at one point in the dish, following the lines, and then move up or down the line as you inject. If the animals start to cleave before they are all injected one can easily remove the uninjected animals and minimize the amount of sorting required. 34. An angle >45° is ideal for injecting the embryos because it generates a downward force on the embryo, which maintains the adhesion of the embryo to the dish. Lower angles can dislodge the embryo, making it difficult to pierce the cell membrane. 35. If a diamond pen is used to break the tip of the needle, it is important to break it as close to the end as possible. Breaking it too far back may result in a blunted needle with a large bore size that creates large holes, effectively killing the injected animals and/or resulting in too high volume being injected.

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36. Estimate volume by examining the sphere present after a single pulse before the die diffuses (Fig. 4d). This balance of pressure and time may be utilized to adjust the volume throughout the injection, as issues such as a clogged needle or broken tip may decrease or increase the flow, respectively. 37. Removing any uninjected embryos at this stage will minimize the amount of sorting that will be necessary before future analysis. 38. We suggest leaving the embryos at a cooler temperature (17 °C) for 2–4 h following injection then moving them to a warmer temperature if necessary (22–25 °C). References 1. Layden MJ, Boekhout M, Martindale MQ (2012) Nematostella vectensis achaete-scute homolog NvashA regulates embryonic ectodermal neurogenesis and represents an ancient component of the metazoan neural specification pathway. Development 139:1013–1022 2. Richards GS, Rentzsch F (2014) Transgenic analysis of a SoxB gene reveals neural progenitor cells in the cnidarian Nematostella vectensis. Development 141:4681–4689 3. Richards GS, Rentzsch F (2015) Regulation of Nematostella neural progenitors by SoxB, Notch and bHLH genes. Development 142:3332–3342 4. Layden MJ, Rentzsch F, Röttinger E (2016) The rise of the starlet sea anemone Nematostella vectensis as a model system to investigate development and regeneration. Wiley Interdiscip Rev Dev Biol:1–21 5. Rentzsch F, Layden M, Manuel M (2016) The cellular and molecular basis of cnidarian neurogenesis. Wiley Interdiscip Rev Dev Biol:1–19 6. Hayakawa E, Fujisawa C, Fujisawa T (2004) Involvement of Hydra achaete?scute gene CnASH in the differentiation pathway of sensory neurons in the tentacles. Dev Genes Evol 7. Käsbauer T, Towb P, Alexandrova O et al (2007) The Notch signaling pathway in the cnidarian Hydra. Dev Biol 303:376–390 8. Münder S, Tischer S, Grundhuber M et al (2013) Dev Biol 383:146–157 9. Jager M, Quéinnec E, Le Guyader H et al (2011) Multiple Sox genes are expressed in stem cells or in differentiating neuro-sensory cells in the hydrozoan Clytia hemisphaerica. EvoDevo 2:12 10. Gahan JM, Schnitzler CE, DuBuc TQ et al (2017) Functional studies on the role of Notch signaling in Hydractinia development. Dev Biol 428:224–231

11. Dunn CW, Hejnol A, Matus DQ et al (2008) Broad phylogenomic sampling improves resolution of the animal tree of life. Nature 452:745–749 12. Hejnol A, Obst M, Stamatakis A et al (2009) Assessing the root of bilaterian animals with scalable phylogenomic methods. Proc R Soc B Biol Sci 276:4261–4270 13. Dupre C, Yuste R (2017) Non-overlapping neural networks in Hydra vulgaris. Curbio 27:1085–1097 14. Amiel AR, Johnston HT, Nedoncelle K et al (2015) Characterization of morphological and cellular events underlying oral regeneration in the sea anemone, Nematostella vectensis. Int J Mol Sci 16:28449–28471 15. DuBuc TQ, Traylor-Knowles N, Martindale MQ (2014) Initiating a regenerative response; cellular and molecular features of wound healing in the cnidarian Nematostella vectensis. BMC Biol 12:24–138 16. Schaffer AA, Bazarsky M, Levy K et al (2016) A transcriptional time-course analysis of oral vs. aboral whole-body regeneration in the sea anemone Nematostella vectensis. BMC Genomics:1–22 17. Sebe-Pedros A, Chomsky E, Saudemont B et al (2017) Cnidarian cell type diversity revealed by whole-organism single-cell RNA-seq analysis, pp 1–29 18. Layden MJ, Röttinger E, Wolenski FS et al (2013) Microinjection of mRNA or morpholinos for reverse genetic analysis in the starlet sea anemone, Nematostella vectensis. Nat Protoc 8:924–934 19. Ikmi A, McKinney SA, Delventhal KM et al (2014) TALEN and CRISPR/Cas9-mediated genome editing in the early-branching metazoan Nematostella vectensis. Nat Commun 5:5486–5488

Reverse Genetics in Nematostella 20. Putnam NH, Srivastava M, Hellsten U et al (2007) Sea anemone genome reveals ancestral eumetazoan gene repertoire and genomic organization. Science 317:86–94 21. Genikhovich G, Technau U (2009) The starlet sea anemone Nematostella vectensis: an anthozoan model organism for studies in comparative genomics and functional evolutionary developmental biology. Cold Spring Harb Protoc 2009:pdb.emo129–pdb.emo129 22. Tulin S, Aguiar D, Istrail S et al (2013) A quantitative reference transcriptome for Nematostella vectensis early embryonic development: a pipeline for de novo assembly in emerging model systems. EvoDevo 4:16–11 23. Helm RR, Siebert S, Tulin S et al (2013) Characterization of differential transcript abundance through time during Nematostellavectensis development. BMC Genomics 14:1–1 24. Wittlieb J, Khalturin K, Lohmann JU et al (2006) Transgenic Hydra allow in vivo tracking of individual stem cells during morphogenesis. PNAS 103:6208–6211 25. Galliot B, Miljkovic-Licina M, Ghila L et al (2007) RNAi gene silencing affects cell and developmental plasticity in hydra. C R Biol 330:491–497 26. Dunn SR, Phillips WS, Green DR et al (2007) Knockdown of actin and caspase gene expression by RNA interference in the symbiotic anemone. Biol Bull 212:250–258 27. Warner J, Guerlais V, Amiel A et al (2017) NvERTx: a gene expression database to compare embryogenesis and regeneration in the sea anemone Nematostella vectensis, pp 1–18 28. Sebe-Pedros A, Chomsky E, Saudemont B et al (2017) Cnidarian cell type diversity revealed by whole-organism single-cell RNA-seq analysis, bioRxiv, pp 1–29 29. Nakanishi N, Renfer E, Technau U et al (2012) Nervous systems of the sea anemone Nematostella vectensis are generated by ectoderm and endoderm and shaped by distinct mechanisms. Development 139:347–357 30. Layden MJ, Martindale MQ (2014) Non-­ canonical Notch signaling represents an ancestral mechanism to regulate neural differentiation. EvoDevo 5:30–14 31. Layden MJ, Johnston H, Amiel AR et al (2016) MAPK signaling is necessary for neurogenesis in Nematostella vectensis. BMC Biol:1–19 32. Havrilak JA, Faltine-Gonzalez D, Wen Y et al (2017) Characterization of NvLWamide-like

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neurons reveals stereotypy in Nematostella nerve net development. Dev Biol. 431(2):336–346 33. Watanabe H, Kuhn A, Fushiki M et al (2014) Sequential actions of β-catenin and Bmp pattern the oral nerve net in Nematostella vectensis. Nat Commun 5:5536–5514 34. Leclère L, Bause M, Sinigaglia C et al (2016) Development of the aboral domain in Nematostella requires β-catenin and the opposing activities of Six3/6 and Frizzled5/8. Development 143:1766–1777 35. Sinigaglia C, Busengdal H, Leclère L et al (2013) The bilaterian head patterning gene six3/6 controls aboral domain development in a cnidarian. PLoS Biol 11:e1001488 36. Marlow H, Matus DQ, Martindale MQ (2013) Ectopic activation of the canonical wnt signaling pathway affects ectodermal patterning along the primary axis during larval development in the anthozoan Nematostella vectensis. Dev Biol 380:324–334 37. Wikramanayake AH, Hong M, Lee PN et al (2003) An ancient role for nuclear beta-catenin in the evolution of axial polarity and germ layder segregation. Nature 426:446–450 38. Magie CR, Daly M, Martindale MQ (2007) Gastrulation in the cnidarian Nematostella vectensis occurs via invagination not ingression. Dev Biol 305:483–497 39. Rentzsch F, Fritzenwanker JH, Scholz CB et al (2008) FGF signalling controls formation of the apical sensory organ in the cnidarian Nematostella vectensis. Development 135:1761–1769 40. Fritzenwanker JH, Technau U (2002) Induction of gametogenesis in the basal cnidarian Nematostella vectensis (Anthozoa). ­ Dev Genes Evol 212:99–103 41. Hand C, Uhlinger KR (1992) The culture, sexual and asexual reproduction, and growth of the sea anemone Nematostella vectensis. Biol Bull 182:169–176 42. Renfer E, Technau U Meganuclease-assisted generation of stable transgenics in the sea anemone Nematostella vectensis. Nat Protoc 12:1844 EP 43. Wolenski FS, Bradham CA, Finnerty JR et al (2012) B is required for cnidocyte development in the sea anemoneNematostella vectensis. Dev Biol:1–11 44. Marlow H,Roettinger E,Boekhout M et al. (2012) Developmental biology. Dev Biol:1–14

Chapter 3 Generating Transgenic Reporter Lines for Studying Nervous System Development in the Cnidarian Nematostella vectensis Fabian Rentzsch, Eduard Renfer, and Ulrich Technau Abstract Neurons often display complex morphologies with long and fine processes that can be difficult to visualize, in particular in living animals. Transgenic reporter lines in which fluorescent proteins are expressed in defined populations of neurons are important tools that can overcome these difficulties. By using membrane-­attached fluorescent proteins, such reporter transgenes can identify the complete outline of subsets of neurons or they can highlight the subcellular localization of fusion proteins, for example at preor postsynaptic sites. The relative stability of fluorescent proteins furthermore allows the tracing of the progeny of cells over time and can therefore provide information about potential roles of the gene whose regulatory elements are controlling the expression of the fluorescent protein. Here we describe the generation of transgenic reporter lines in the sea anemone Nematostella vectensis, a cnidarian model organism for studying the evolution of developmental processes. We also provide an overview of existing transgenic Nematostella lines that have been used to study conserved and derived aspects of nervous system development. Key words Transgenic reporter, Nematostella, Cnidaria, Fluorescent protein, Neurogenesis, Microinjection

1  Introduction Cnidarians are a group of morphologically diverse animals that includes corals, sea anemones, and various types of jellyfish. They are the sister group to bilaterians [1] and as such occupy an informative position in the tree of life for understanding the early evolution of nervous systems. The relevance of cnidarians for the evolution of nervous systems has long been recognized, but only with the advent of molecular tools in the last 10–20 years it became possible to identify common and divergent features of cnidarian and bilaterian nervous system development [2–5].

Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_3, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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Cnidarians are grouped into anthozoans and medusozoans, the latter typically being characterized by the presence of a pelagic medusa in their life cycle, which is generated asexually from polyps [6]. Anthozoans, in contrast, lack a medusa stage and exist only as sexually reproducing, sessile polyps. The nervous system of polyps contains areas of lower and higher density of neurons and can feature tracts of longitudinal neurites, but its overall organization is best characterized as a nerve net [2–5]. Free-swimming medusae display a much higher degree of concentration of neural elements [7, 8]. This is particularly evident in the rhopalia, light and gravity sensing structures located at the margin of the bell in scyphozoan and cubozoan jellyfish. Rhopalia can contain thousands of neural cells and contribute to the control of swimming behavior [9, 10]. Cells of the nervous system in cnidarians have traditionally been grouped into sensory cells (including sensory-motor cells), ganglion cells (morphologically equivalent to interneurons), and cnidocytes (“stinging cells”). Morphological, immunohistochemical, and gene expression analyses have shown that these three main groups consist of many subgroups, but the integration of morphological and molecular data to define these subgroups more precisely is still in its early stages [4, 11]. The development of the nervous system has been studied mainly in two groups of cnidarians, hydrozoans (part of the medusozoans) and anthozoans. In both groups, nerve cells develop in a spatially distributed manner throughout the tissue. In hydrozoans, most neural cells are generated from interstitial stem cells (i-cells), a heterogeneous population of cells with mesenchymal appearance that includes multipotent cells [12–14]. Embryonically, i-cells originate from the endoderm before they can migrate to the ectodermal layer [15–17]. Anthozoan embryos do not possess cells that resemble interstitial cells at the morphological level. Neural cells first appear in the ectoderm and after gastrulation also in the endoderm [18]. Epithelial neural progenitor cells are located in both germ layers and give rise to sensory cells and ganglion cells [4, 19]. Cnidocytes (stinging cells), a sensory cell type used for predation and defense are formed exclusively in ectodermal tissue. Despite this uncommon, broad neurogenic potential, cnidarian neurogenesis shares many molecular features with bilaterian neurogenesis, e.g. the involvement of SoxB and bHLH transcription factors and the Notch and Wnt signaling pathways (though a role for Notch in early neurogenesis seems to be absent in hydrozoans, [20–24]). These observations now allow us to move beyond candidate gene approaches to obtain a detailed understanding of the molecular basis for the broad neurogenic potential of cnidarians during embryonic development and regeneration. A key technological progress for studying neural development and neural function in cnidarians was the establishment of stable transgenic lines [25–27].

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Fig. 1 Transgenic Nematostella reporter lines. (a–c) Elav1::mOrange, detected by an anti-dsRed antibody and pseudocolored. (a) Primary polyp (8 dpf), with aboral pole to the left, F-actin is stained with Phalloidin (green). (b, c) Body wall of 3-week old polyps. (b) is a F1 fully transgenic animal, note the two tracts of neurites that run along the oral–aboral axis (left to right). Nuclei are stained with DAPI (blue). (c) is a mosaic F0 transgenic animal that has been injected with the reporter plasmid. Fewer cells are labeled, allowing better resolution of individual neurons. (d) Elav1::cerulean (cyan), FoxQ2d::mOrange (magenta) double positive planula with aboral pole to the left. There is no overlap of the two neural cell poulations

The distributed organization of the nervous system in cnidarians makes it difficult or impossible to use the position of cells expressing particular genes (detected by in situ hybridization) as an indicator of their neural identity. Transgenic reporter lines overcome this problem by visualizing the morphology of cells expressing a transgene, for example, a membrane-tethered fluorescent proteins (Fig. 1). Moreover, they allow tracing the progeny of the cells after they terminated the expression of a gene of interest and they have been used to record neural activity via genetically encoded calcium sensors [28]. Successful use of transgenic animals has been reported for the hydrozoans Hydra and Hydractinia, and for the anthozoan Nematostella [25, 27, 29]. Here we focus on Nematostella vectensis to summarize features of published “neural” Nematostella reporter lines and provide a protocol for the generation of transgenic lines. This protocol is based on the one published by Renfer and Technau [26] and uses the I-SceI Meganuclease to obtain transgenic lines with good efficiency.

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1.1  Available Nematostella Reporter Lines for the Nervous System

NvSoxB(2) [19]: The HMG box transcription factor NvSoxB(2) is expressed from blastula stages on in mitotic and non-mitotic cells scattered throughout the ectoderm and (after gastrulation) the endoderm. The transgenic reporter line revealed that these cells develop into sensory cells, ganglion cells, and cnidocytes. Mitotic memOrange-positive cells can also be detected, but their number is significantly lower than that of mitotic cells expressing NvSoxB(2) mRNA. The NvSoxB(2) mRNA expressing cells thus include neural progenitor cells that generate a large fraction of the larval nervous system. Individual memOrange positive cells can be detected by immunohistochemistry (using anti-dsRed antibodies) from gastrula stage on; in living or unstained animals, however, individual cells can be distinguished only at planula stage. NvElav1 [18]: Orthologs of the RNA binding protein Elav are often considered pan-neural markers in bilaterians. In Nematostella, the NvElav1::memOrange reporter line labels a population of sensory cells and ganglion cells in both germ layers, starting from gastrula stage and persisting in adult polyps. Double transgenic animals suggest that NvElav1::Cerulean-positive cells constitute a subpopulation of the NvSoxB(2)::memOrange-positive cells. While the NvElav1 transgenics label a large number of morphologically heterogeneous neurons, co-labeling with antibodies against neuropeptides has shown that neither the transgenic line nor the elav1 transcript are pan-neural markers in Nematostella. Notably, the NvElav1 transcript and the transgenic line do not label cnidocytes. The Nematostella genome encodes a second elav gene, which is not part of the “neural” group of elav genes. It has so far not been possible to determine the mRNA expression pattern of NvElav2. NvLWamide-like [30]: Transcripts for the neuropeptide NvLWamide-like are first detected in the aboral half of gastrula stage embryos and from planula stage on in ecto- and endodermal cells, including the pharynx and (at later stages) the tentacles. Fluorescence derived from the NvLWamide-like::mCherry transgene is visible in neurons from mid-planula stage on in the ectoderm and pharynx and subsequently in the endoderm. The number of NvLWamide-like::mCherry-positive cells is small compared to the number of NvElav1::memOrange cells, which allowed the identification of specific subsets of these cells. Interestingly, some of these subsets display only little variability in their neurite projection patterns and positions in the body column. This high level of stereotypy will likely be beneficial for studying the mechanisms that control the positioning of these neurons and the outgrowth of their neurites. NvFoxQ2d [31]: The transcription factor NvFoxQ2d is expressed in both mitotic and non-mitotic cells in the ectoderm, starting at gastrula stage. Mitotic cells are not found in the NvFoxQ2d::memOrange transgenic line. One explanation for this observation can be that the NvFoxQ2d mRNA expressing cells

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undergo a terminal division and that the maturation time of the fluorescent protein precludes their identification in the transgenic line. The NvFoxQ2d::memOrange-positive cells are excluded from the oral area of the planula and polyp and they display a rather uniform morphology that resembles sensory cells. While they have only short neurite-like processes, they can be co-labeled with an antibody against FMRFamide neuropeptides, supporting their neural nature. NEP3 [32]: NEP3 encodes a toxin whose mRNA is expressed in scattered cells of the ectoderm and in the tentacles. As suspected, a NvNEP3::mOrange2 transgene encoding a fusion protein of the predicted signal peptide of NvNEP3 and the fluorescent reporter identified the NvNEP3-expressing cells as a large subpopulation of cnidocytes.

2  Materials 2.1  Animal Maintenance and Preparation of Eggs

1. Nematostella Medium—sea water (artificial of filtered natural) diluted to 14–16 ppt. 2. 3% Cysteine solution (l-cysteine, Sigma) in Nematostella medium, adjust pH with NaOH to 7.4. 3. Pasteur pipettes with smooth opening (see Note 1). 4. Petri dishes (see Note 1). 5. Injection dish with a low wall (see Note 2).

2.2  Injection Solution

1. I-SceI meganuclease (5 U/μl, New England Biolabs). 2. Alexa-conjugated Dextran MW 10,000 (Life Technologies, see Note 3). 3. Nuclease-free water for molecular biology.

2.3  Capillaries

1. Holding capillary (custom-made Vacu Tip, Eppendorf, outer diameter 110 μm, inner diameter 60 μm, angle 25°, limb length 500 μm, round front surface). 2. Injection capillaries (made from GB100TF-10 borosilicate capillaries with filament, Science Products, Germany, see Note 4).

2.4  Equipment

1. Heating block. 2. Puller for injection capillaries (Sutter P-97). 3. Inverted microscope with fluorescence lamp (Fig. 2). 4. Coarse and fine micromanipulators (MN-4 and MMO-­ 202ND, Narishige). 5. CellTram vario (Eppendorf, see Note 5). 6. Microinjection pump (FemtoJet, Eppendorf).

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Fig. 2 Microscope setup for Nematostella microinjections. (a) Overview; the injection setup is mounted on an inverted microscope equipped with a mercury lamp. The injection pump is connected to the injection capillary (on the right side of the microscope), the Cell Tram to the holding capillary (on the left side). A joystick operates the micromanipulator. Injection is triggered with a foot pedal or a by mouse click (not on the image). (b) The coarse manipulator is mounted on the microscope

3  Methods 3.1  Design and Cloning of Constructs

3.2  Preparation of Embryos

The cloning cassette of the vector that has been used for the generation of transgenic Nematostella is flanked by sites for the I-SceI homing endonuclease and it contains an SV40 polyadenylation signal following the ORF of a fluorescent reporter protein of choice [25, 26]. The use of Gibson Assembly® [33] for cloning is a good option to keep the distance between the regulatory elements and the sequence encoding the fluorescent protein minimal. The cloning is performed by standard procedures; plasmids can be prepared with midi-prep kits to achieve high purity (see Note 6). 1. To induce spawning, expose separate boxes with male and female polyps to bright light and a temperature of 24–25 °C for 12 h (overnight, see Note 7). 2. In the following morning, discard egg packages that have been laid overnight. Keep the boxes at the standard culture temperature (18–19 °C) and collect freshly laid egg packages after two hours. Fertilize the egg packages by incubating them for 20 min with an excess of medium from the box containing the male polyps. 3. Transfer the fertilized egg packages to a petri dish and incubate them for 20 min in the 3% Cysteine solution (pH 7.4) on a rotating shaker (see Notes 8 and 9). 4. Collect the fertilized eggs and wash them in Nematostella medium 5× for 2 min, 1× 5 min. 5. Place the de-jellied eggs in the injection room (see Note 10).

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3.3  Preparation of Injection Solution

3.4  Transfer of Zygotes to Injection Plate and Preparation of Capillaries

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Prepare the solution for injection in a 1.5 ml tube. Incubate at 37 °C for 20 min, keep dark. Store on ice. 1. I-SceI buffer (10×)

1.0 μl

2. Dextran MW 10,000 (200 ng/μl)

2.0 μl

3. Plasmid (100 ng/μl, see Note 11)

2.0 μl

4. I-SceI (5 U/μl)

2.0 μl

5. Nuclease-free water

to 10 μl

1. Fill the injection dish with Nematostella medium and place it on the stage of the injection microscope. Use a Pasteur pipette with smooth opening to place the fertilized eggs in the injection dish. Attempt to place them as a column (see Note 12). 2. Insert the holding capillary in the capillary holder connected to the Cell Tram. The distal part of the capillary should be parallel to the surface of the injection dish. 3. Use a microloader pipette tip to fill the injection capillary from the back end. Insert the pipette tip as deep as possible and release the injection solution close to the tip of the capillary. 4. Insert the injection capillary in the capillary holder (see Note 13). 5. Connect the capillary holder to the FemtoJet injection pump via the attached tube (see Note 14). 6. Use the microscope stage to move the embryos into the field of view, then use the coarse and the fine micromanipulators to move the two capillaries into focus. 7. Lower the injection capillary until the site at which you want to break it touches the bottom of the dish. Move the tip of the holding capillary on top of the injection capillary and gently move it down to open the injection capillary (Fig. 3, see Note 15).

3.5  Injection

1. Open the shutter for the fluorescent light and use the “inject” or the “clean” button of the injection pump to release air from the tip of the injection capillary. Adjust the injection volume via the injection pressure and the injection time (see Note 16). 2. Adjust the “hold pressure” to avoid that medium from the dish is aspired into the injection capillary.

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Fig. 3 Opening of the injection capillary. Schematic depiction of the opening of the injection capillary with the holding capillary. The injection capillary is lowered to touch the bottom of the injection dish (point of contact is indicated by the bend of the injection capillary). The holding capillary is lowered to break the injection capillary (middle panel)

3. Use the CellTram to hold a zygote and the micromanipulator to move the needle (Fig. 2). Inject and remove the needle before releasing the zygote (see Note 17). 4. Move the microscope stage to place the next zygote in front of the holding capillary. 3.6  Identification of Transgenic Animals

1. Transfer the animals with clearly visible fluorescent dextran from the injection dish to a petri dish. 2. Use a stereomicroscope or an inverted compound microscope to monitor the animals for expression of the fluorescent protein at the expected stage of development. Separate animals into groups according to the strength/breadth of expression (see Note 18). 3. Raise the animals according to standard culture protocols. 4. Identify the sex of the animals (see Note 19) and cross them individually to wild-type polyps (see Note 20) to identify carriers of the transgene.

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4  Notes 1. Nematostella eggs and embryos stick to fresh plasticware and glass pipettes. To reduce stickiness, we fill fresh pipettes and dishes with Nematostella medium for at least one night before first use and reuse them when possible. 2. A low wall allows mounting the injection capillary with a low angle in relation to the surface of the injection dish. With a low angle, the egg is pressed towards the holding capillary during penetration of the injection capillary. As injection dishes we use the lids of Nunc 4-well plates. 3. The color of the fluorescent dextran should be chosen according to the fluorescent protein of the construct. 4. The quality of the injection capillary is important for the survival of the injected embryos and the number of embryos that can be injected in one session. Capillaries that are too wide will damage the embryo and render the control of the injection volume difficult. If the capillary is too thin, it will bend when it comes into contact with the egg and it gets clogged more easily. Parameters for pulling the injection needles need to be determined for each heating filament of the puller. The Pipette Cookbook is a good source of information available on the website of Sutter Instruments. 5. Note that simpler microinjection setups can be used for Nematostella [34]. 6. The identification of putative regulatory elements is greatly aided by the availability of genome-wide maps of chromatin modifications [35] that can be used to predict promoters and enhancers. When using upstream sequences only, we place the reverse primer immediately upstream of the start codon to include the 5′UTR. For many genes, however, the first intron contains putative regulatory elements [35] and should be included in the construct. We have so far used putative regulatory regions of 1.4–6 kb for generating stable transgenic lines (unpublished data). Congruence of endogenous and transgene-­ derived expression can be tested by fluorescent double in situ hybridization with probes for the ORF of the fluorescent protein and the gene of interest. 7. In our feeding regime (Artemia nauplii 5× per week), spawning of Nematostella polyps can be induced every 14–21 days for years. Shorter intervals (7 days) can be sustained for a few weeks, but fecundity tends to decrease thereafter. Protocols for Nematostella culture conditions and the induction of spawning have been published [36–38].

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8. After fertilization, use a stereomicroscope to select egg packages in which the eggs are round and have a rather uniform diameter. Differences in egg size require repeated adjustment of the position of the capillaries during injection. 9. De-jellying by cysteine is an important step. If the eggs are not sufficiently de-jellied, both the holding and injection capillary will stick to the jelly and get clogged. If the eggs are exposed to the cysteine solution too long, the survival rate decreases and more embryos display developmental abnormalities. 10. Injection can be done until the blastomeres are separated, this is often the 4-cell stage. Lower temperature delays development and allows more time for injection. We cool the injection room to 16–18 °C. 11. The plasmid concentration that can be used for injection varies with the injection volume and with the quality of the plasmid preparation. A concentration of 20 ng/μl is a good starting point, but the concentration can be increased as long as developmental defects and mortality remain limited. 12. To align the eggs in a column, they need to be released slowly into the injection dish. This, however, increases the risk of them sticking to the glass or plastic of the pipette. 13. Move the capillary into the holder carefully until it touches the O-ring. 14. Refer to the user manual of the injection pump for detailed description. 15. The opening of the injection capillary with the holding capillary can be time-consuming and the exact point of breakage is difficult to predict. The part of the holding capillary that is furthest down has to be used for breaking; otherwise, the holding capillary itself might be damaged. Depending on the angle in which the holding capillary is mounted, the bending point (the “knee”) of the holding capillary might be furthest down. In this case, the desired break point has to be placed under the “knee.” Alternatively, fine forceps can be used to cut the injection capillary. 16. Different labs use different volumes for the injection. The exact volume is difficult to determine because the fluorescent dextran diffuses very quickly throughout the egg. Injection into drops of oil to determine the injected volume is also difficult, because good injection capillaries tend to be too thin and flexible to be inserted into the oil. We are trying to inject a drop of about 1/4 of the diameter of an egg. With an average egg diameter of 230 μm, this injection volume corresponds to approximately 100 pl.

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17. During the injection session, we keep both the shutters for the fluorescent and the transmitted light open. This allows control of the injected volume throughout the injection session. We have not observed any aversive effects of the excitation of fluorescent tracer. 18. The best time point for the screening of the injected animals depends on the injected construct and the expected pattern. In F0 planulae, we often observe expression that does not match the expected pattern, but this is no longer the case in primary polyps and it is not present in F1 planulae. When possible, we sort the F0 animals at primary polyp stage (10 dpf), when they don’t swim anymore. However, for genes that are expressed only at very early developmental stages, the fluorescent signal might no longer be detectable at primary polyp stage. The F0 animals display mosaic expression and we sort them according to the area in which expression is visible, with the assumption that a bigger area of expression increases the likelihood of germline transmission. Ideally, we raise ca 100 F0 polyps that show expression of the transgene. It should be noted, though, that for genes that are expressed distant from the mesenteries (e.g. in tentacles) transgene expression in F0 animals may not be a good selection criteria, as nonfluorescent, but transgenic patches may well reside in the germline, while fluorophore-expressing animals may not harbor transgenic patches in the germline. In such cases, it is best to simply raise all or a random selection of embryos to the next generation. 19. The sex of the adult polyps can only be determined by inducing them to spawn individually.  20. With the described method, integration of the transgene can occur at different positions in the genome. Spatial and temporal aspects of the expression of the transgene can thus be affected by the integration site. It is therefore desirable to generate transgenic lines from more than one F0 animal to allow comparison of the observed expression.

Acknowledgements This work is supported by funding from the University of Bergen and the Research Council of Norway (to F.R.) and by the Austrian Science Fund (FWF) (P27353) to U.T.

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References 1. Telford MJ, Budd GE, Philippe H (2015) Phylogenomic Insights into Animal Evolution. Curr Biol 25(19):R876–R887 2. Galliot B, Quiquand M (2011) A two-step process in the emergence of neurogenesis. Eur J Neurosci 34(6):847–862 3. Watanabe H, Fujisawa T, Holstein TW (2009) Cnidarians and the evolutionary origin of the nervous system. Dev Growth Differ 51(3):167–183 4. Rentzsch F, Layden M, Manuel M (2017) The cellular and molecular basis of cnidarian neurogenesis. Wiley Interdiscip Rev Dev Biol 6(1) 5. Kelava I, Rentzsch F, Technau U (2015) Evolution of eumetazoan nervous systems: insights from cnidarians. Philos Trans R Soc Lond B Biol Sci 370(1684):20150065 6. Zapata F, Goetz FE, Smith SA, Howison M, Siebert S, Church SH, Sanders SM, Ames CL, McFadden CS, France SC, Daly M, Collins AG, Haddock SH, Dunn CW, Cartwright P (2015) Phylogenomic Analyses Support Traditional Relationships within Cnidaria. PLoS One 10(10):e0139068 7. Katsuki T, Greenspan RJ (2013) Jellyfish nervous systems. Curr Biol 23(14):R592–R594 8. Mackie GO (2004) Central neural circuitry in the jellyfish Aglantha: a model simple nervous system. Neurosignals 13(1-2):5–19 9. Garm A, Ekstrom P, Boudes M, Nilsson DE (2006) Rhopalia are integrated parts of the central nervous system in box jellyfish. Cell Tissue Res 325(2):333–343 10. Petie R, Garm A, Nilsson DE (2011) Visual control of steering in the box jellyfish Tripedalia cystophora. J Exp Biol 214(Pt 17):2809–2815 11. Sebe-Pedros A, Saudemont B, Chomsky E, Plessier F, Mailhe MP, Renno J, Loe-Mie Y, Lifshitz A, Mukamel Z, Schmutz S, Novault S, Steinmetz PRH, Spitz F, Tanay A, Marlow H (2018) Cnidarian cell type diversity and regulation revealed by whole-organism single-cell RNA-seq. Cell 173(6):1520–1534 e20 12. Bosch TC, Anton-Erxleben F, Hemmrich G, Khalturin K (2010) The Hydra polyp: nothing but an active stem cell community. Dev Growth Differ 52(1):15–25 13. Watanabe H, Hoang VT, Mattner R, Holstein TW (2009) Immortality and the base of multicellular life: Lessons from cnidarian stem cells. Semin Cell Dev Biol 20(9):1114–1125 14. Frank U, Plickert G, Muller WA (2009) Cnidarian interstitial cells: The dawn of stem cell research. In: Rinkevich B, Matranga V (eds) Stem cells in marine organisms. Springer, New York, pp 33–59

15. Martin VJ (1990) Development of Nerve-Cells in Hydrozoan Planulae. 3. Some Interstitial-­ Cells Traverse the Ganglionic Pathway in the Endoderm. Biol Bull 178(1):10–20 16. Martin VJ, Archer WE (1986) Migration of interstitial cells and their derivatives in a hydrozoan planula. Dev Biol 116:486–496 17. Martin VJ, Littlefield CL, Archer WE, Bode HR (1997) Embryogenesis in hydra. Biol Bull 192(3):345–363 18. Nakanishi N, Renfer E, Technau U, Rentzsch F (2012) Nervous systems of the sea anemone Nematostella vectensis are generated by ectoderm and endoderm and shaped by distinct mechanisms. Development 139(2):347–357 19. Richards GS, Rentzsch F (2014) Transgenic analysis of a SoxB gene reveals neural progenitor cells in the cnidarian Nematostella vectensis. Development 141(24):4681–4689 20. Grens A, Mason E, Marsh JL, Bode HR (1995) Evolutionary conservation of a cell fate specification gene: the Hydra achaete-scute homolog has proneural activity in Drosophila. Development 121(12):4027–4035 21. Richards GS, Rentzsch F (2015) Regulation of Nematostella neural progenitors by SoxB, Notch and bHLH genes. Development 142(19):3332–3342 22. Watanabe H, Kuhn A, Fushiki M, Agata K, Ozbek S, Fujisawa T, Holstein TW (2014) Sequential actions of beta-catenin and Bmp pattern the oral nerve net in Nematostella vectensis. Nat Commun 5:5536 23. Layden MJ, Boekhout M, Martindale MQ (2012) Nematostella vectensis achaete-scute homolog NvashA regulates embryonic ectodermal neurogenesis and represents an ancient component of the metazoan neural specification pathway. Development 139(5):1013–1022 24. Gahan JM, Schnitzler CE, DuBuc TQ, Doonan LB, Kanska J, Gornik SG, Barreira S, Thompson K, Schiffer P, Baxevanis AD, Frank U (2017) Functional studies on the role of Notch signaling in Hydractinia development. Dev Biol 428(1):224–231 25. Renfer E, Amon-Hassenzahl A, Steinmetz PR, Technau U (2009) A muscle-specific transgenic reporter line of the sea anemone, Nematostella vectensis. Proc Natl Acad Sci U S A 107(1):104–108 26. Renfer E, Technau U (2017) Meganuclease-­ assisted generation of stable transgenics in the sea anemone Nematostella vectensis. Nat Protoc 12(9):1844–1854 27. Wittlieb J, Khalturin K, Lohmann JU, Anton-­ Erxleben F, Bosch TC (2006) Transgenic

Transgenic Reporter in Nematostella Hydra allow in vivo tracking of individual stem cells during morphogenesis. Proc Natl Acad Sci U S A 103(16):6208–6211 28. Dupre C, Yuste R (2017) Non-overlapping Neural Networks in Hydra vulgaris. Curr Biol 27(8):1085–1097 29. Kunzel T, Heiermann R, Frank U, Muller W, Tilmann W, Bause M, Nonn A, Helling M, Schwarz RS, Plickert G (2010) Migration and differentiation potential of stem cells in the cnidarian Hydractinia analysed in eGFP-­transgenic animals and chimeras. Dev Biol 348(1):120–129 30. Havrilak JA, Faltine-Gonzalez D, Wen Y, Fodera D, Simpson AC, Magie CR, Layden MJ (2017) Characterization of NvLWamide-like neurons reveals stereotypy in Nematostella nerve net development. Dev Biol 431(2):336–346 31. Busengdal H, Rentzsch F (2017) Unipotent progenitors contribute to the generation of sensory cell types in the nervous system of the cnidarian Nematostella vectensis. Dev Biol 431(1):59–68 32. Columbus-Shenkar YY, Sachkova MY, Macrander J, Fridrich A, Modepalli V, Reitzel AM, Sunagar K, Moran Y (2018) Dynamics of venom composition across a complex life cycle. Elife 7:e35014

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33. Gibson DG, Young L, Chuang RY, Venter JC, Hutchison CA 3rd, Smith HO (2009) Enzymatic assembly of DNA molecules up to several hundred kilobases. Nat Methods 6(5):343–345 34. Layden MJ, Rottinger E, Wolenski FS, Gilmore TD, Martindale MQ (2013) Microinjection of mRNA or morpholinos for reverse genetic analysis in the starlet sea anemone, Nematostella vectensis. Nat Protoc 8(5):924–934 35. Schwaiger M, Schonauer A, Rendeiro AF, Pribitzer C, Schauer A, Gilles AF, Schinko JB, Renfer E, Fredman D, Technau U (2014) Evolutionary conservation of the eumetazoan gene regulatory landscape. Genome Res 24(4):639–650 36. Fritzenwanker JH, Technau U (2002) Induction of gametogenesis in the basal cnidarian Nematostella vectensis(Anthozoa). Dev Genes Evol 212(2):99–103 37. Genikhovich G, Technau U (2009) Induction of spawning in the starlet sea anemone Nematostella vectensis, in vitro fertilization of gametes, and dejellying of zygotes. Cold Spring Harb Protoc 2009(9):pdb prot5281 38. Stefanik DJ, Friedman LE, Finnerty JR (2013) Collecting, rearing, spawning and inducing regeneration of the starlet sea anemone, Nematostella vectensis. Nat Protoc 8(5):916–923

Chapter 4 Immunostaining and In Situ Hybridization of the Developing Acoel Nervous System Elena Perea-Atienza, Brenda Gavilán, Simon G. Sprecher, and Pedro Martinez Abstract The study of acoel morphologies has been recently stimulated by the knowledge that this group of animals represents an early offshoot of the Bilateria. Understanding how organ systems and tissues develop and the molecular underpinnings of the processes involved has become an area of new research. The microscopic anatomy of these organisms is best understood through the systematic use of immunochemistry and in situ hybridization procedures. These methods allow us to map, in precise detail, the expression patterns of genes and proteins, in space and time. With the additional use of genomic resources, they provide us with insights on how a group of “early” bilaterians have diversified over time. As these animals are new to the world of molecular studies, the protocols have involved a lot of new and specific adaptations to their specific anatomical-histological characteristics. Here we explain some of these protocols in detail, with the aim that should prove useful in our much-needed understanding of the origins of bilaterian animals. An anatomical sketch is provided at the beginning as a necessary guide for those not familiar with the Acoela. Key words Acoela, Embryos, Juveniles, Immunochemistry, In situ hybridization, FISH methodology, Nervous system

1  Introduction Acoel worms are (mostly) marine animals belonging to the phylum Xenacoelomorpha. The phylum contains three major clades: Xenoturbellida, Nemertodermatida, and Acoela. It is to this last clade that our worms belong. Acoela is nowadays represented by more than 400 nominal species, all with relatively simple morphologies and sizes that are in the millimeter range. These animals share a series of characteristics, they are bilaterally symmetric, have a multiciliated epithelia, an underlying musculature with a few overlapping muscle sheets, a relatively simple nervous system with anteElena Perea-Atienza and Brenda Gavilán contributed equally to the development of the described methodology. Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_4, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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rior condensations, a single opening of the gut, are hermaphroditic, with assacate gonads, and possess a posteriorly located copulatory apparatuses (more on the anatomy, below). The importance of studying xenacoelomoprhs, or their constitutive clades, resides on the key phylogenetic position that this phylum occupies. In most recent phylogenomic studies it has been shown that Xenacoelomorpha represent, most probably, the sister group of the remaining Bilateria (the so-called Nephrozoa [1]). Some studies, though, still place the group in different positions within the Deuterostomia [2]. Given the fact of the solid support for the early split of Xenacoelomorpha and Nephorozoa, xenacoelomorphs became good proxies for understanding the morphological complexity of the ancestral bilateral animal [3, 4]. In this context, any hypothesis regarding the events leading to the origin of Bilateria has to contemplate the organization of xenacoelomorph morphologies. And here we find the most powerful argument to study this group of animals. The richest clade within the Xenacoelomorpha is the Acoela, with the 400 species living in a range of environments, from intertidal flats to the deep sea [5, 6]. The internal relationships of the acoel clades have been the subject of some historical debate, though Jondelius and collaborators, using molecular and morphological characters have proposed, recently, a well-supported phylogeny [4]. However, and before dealing with the specific protocols developed to study the morphology and development of these animals, let’s spend, as stated above, a few lines describing the major characteristics of the acoel morphology. 1.1  A Quick Guide to the Acoel Morphology

The Acoela are small, flat, soft-bodied, aquatic (predominantly marine) worm-like organisms. They lack a gut cavity, an oviduct, and an excretory system. The Acoela are hermaphrodites, and the same animal has both male and female reproductive organs (Fig. 1a). The epidermis is completely ciliated and richly glandular. The cilia are mostly involved in locomotion, but some of them act as sensory receptors. A thick mucus layer covers the epidermis. Several other types of glands are also distinguished between the epidermal cells all along the body. The frontal organ consists of a collection of two or more large mucus-secreting glands whose necks emerge together through a frontal pore at the apical pole of the body [7]. Their function as a sensory organ is still controversial. The statocyst [8] is a sensory organ informing the animal about its position in the water column. This gravity-sensitive organ consists of a round capsule surrounding a hemispherical concretion. It is located at the anterior tip of the body and embedded in the central nervous system.

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A

Brain Commisure Longitudinal nerves Neuropile

Commisure

B

Statocyst

Wrapping cells

Central syncytium

Food vacuoles

Nuclei

C

Frontal pore

Frontal glands Germ cells

Mouth Developing Sperm spermatids tract Bursa

D

Ovary

Mature egg

E Longitudinal muscles

Foreign sperm

Sperm

Seminal vesicle

Bursal Penis Male nozzle Male Sphincter Female copulatory genital pore genital pore organ Rhaboid gland Circular muscle

Epidermis

Longitudinal muscle Parenchyma cells

Circular muscles

Circular muscles

Longitudinal muscles

Epidermis

Fig. 1 General morphology of the Acoela adapted from different diagrams made by Kathryn Apse and Prof. Seth Tyler, University of Maine

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The ocelli correspond to the eyes of Acoela [9]. Unlike most of turbellarians the ocelli of Acoela lack rhabdomeres and cilia. Instead the ocellus of Acoela consists of several sensory cells and a concave pigment cell containing spindle-shaped platelets. The structure of the nervous system (Fig. 1b) in Acoela is very variable, and this partially reflects the different approaches used to study it. Indeed, there are two types of nerve concentration that have been called “brain” in Acoela [10]. The first is a thickening of the single plexus at the base of the epidermis. This thickening may form distinct rings and longitudinal cords. This type of nervous concentration was also called the “orthogonal brain,” and later on renamed “commissural brain,” [11] because the “thickening of transverse commissures” forms it. The second is a small concentration of nerve cells around or caudal to the statocyst. It was also called the “endonal brain” [12]. The recent, thorough, analysis by TEM of the S. roscoffensis brain has led us to abandon this nomenclature [13]. Acoela are characterized by the absence of a central cavity, as evoked by the name of this phylum (“a” = without, “coel” = cavity). Like the nervous system, the structure of the digestive system is poorly understood, because in previous studies two very different organizations of the digestive system were described. Most studies described a syncytial digestive system, and this is considered to be the common pattern for all acoels [14, 15], but others have suggested that it is, actually, a cellularized structure, difficult to identify in histological sections [16]. Our recent studies suggest that the gut is clearly syncytial (Gavilán et al., unpublished) (Fig. 1c). The digestive system opens at the surface of the body through a mouth. In Acoela, the anterior part of the digestive system shows variable morphology. The morphology of the mouth varies from a simple opening on the ventral surface of the body to a complex pharynx open at either the anterior or the posterior tip of the body. This important difference provides the basis for distinguishing between the acoel families. The structure of the copulatory organs (Fig. 1d) is variable and was used originally to establish the internal phylogeny of the Acoela. The male copulatory apparatus is generally located posterior to the female copulatory apparatus. Both apparatuses open on the ventral side of the body. The male copulatory apparatus [17] consists of a paired channel (vasa deferentia or false seminal vesicle) in the parenchyma for the transport of the sperm. The false seminal vesicle unites to a common terminal duct, in whose wall the penis lies. In Acoela the structure of the penis varies among the families from a muscular thickening of the body wall to a conical penis composed of parenchyma, muscle fibers and gland cells. The female copulatory apparatus [17] in Acoela is characterized by a lack of ducts. Instead the female reproductive system consists of a seminal

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bursa that is a rounded parenchymatous organ. The latter may be connected to the exterior by its own female gonopore or through the male gonopore, or may not be so connected. The seminal bursa is indirectly connected to the eggs through a bursal nozzle that is a strong structure arranged in stacked rings (previously called “mouthpieces”). The body wall musculature (Fig. 1e) has also been used also as a phylogenetic character, because the great internal variability within the Acoela [18–20]. All acoel species possess three main types of muscle: longitudinal, circular, and diagonal. They differ mainly in the presence of accessory muscle types (V-shape, cross-­ over, parenchymal) and in the relative position of the main muscle types (outer circular and inner longitudinal). The development of Acoela has not been studies in many species. Our best knowledge derives from the study carried out by Henry and collaborators [21]. Acoels fertilize internally and zygotes are extruded through the body wall in packages of several embryos (cocoons), all included within a thick, translucent, mucilaginous egg capsule, the jelly layer. Within these cocoons the development of embryos tends to be well synchronized. Only the details of the early embryogenesis of the species Neochildia fusca are known [21]. Organogenesis proceeds later on, giving rise to the different tissues (further details in: [22]). 1.2  Tools: Antibodies and RNA Probes

In laboratories working with acoels, most of the antibodies used to date are commercial, and recognize large groups of neurons, like those synthesizing serotonin [23–25] or those producing the FMRF-amide (or FMFR-amide like) peptide [25, 26]. However, recently, and thanks to the transcriptomic sequences produced in some acoel species, a number of species-specific antibodies have been produced (i.e.: anti-synaptotagmin or anti-ELAV; [27, 28]). Though these represent a very small cohort, more antibodies are being produced in different laboratories that will further widen the catalog of useful markers. The in situ methodologies rely on availability of RNA probes, labeled with colorimetric or fluorescent tags. RNA probes are synthesized routinely from clones identified in transcriptomes. In the following paragraphs we detail the methodologies used in acoels, with an emphasis on those specific modifications of standard protocols necessary to deal with this group of animals.

2  Materials All stock solutions must be made using sterile material and kept in sealable bottles to avoid contamination. Prepare the solutions with either sterilized distilled milli-Q water or filtered sea water (FSW)

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(artificial or natural) according to the protocol. The correct preparation and use of the solutions should keep them uncontaminated and in good conditions, at least for a month. Make aliquots of every stock solution, for instance in Falcon tubes, to avoid introducing pipettes or manipulating the original solution directly. For any of the in situ hybridization protocols remember to use, in all steps (including fixation and pretreatment), sterilized and RNAse-free solutions and material. 2.1  Embryo Fixation and Preparation

1. Filtered sea water (FSW); artificial (any of the available protocols will do) or natural (if available). 2. Thioglycolate-Pronase solution (use it freshly made): 0.2 g of Thioglycolate in 20 ml of FSW. Adjust to pH 8 with 1 M NaOH. Add 0.01 g of Pronase. 3. 4% formaldehyde methanol-free, in FSW. Use it freshly made. Alternatively, a 4% paraformaldehyde solution, from powder, can be used. 4. 1× PBS. 5. 100% methanol. 6. 75% MeOH in PBS. 7. 50% MeOH in PBS. 8. 25% MeOH in PBS. 9. 20× SSC stock: 0.3 M Na citrate and 3 M NaCl in water and pH adjusted to 7. 10. Bleaching solution (use it freshly made): 1.5% H2O2, 5% formamide, 0.5× SSC in milliQ RNase-free water (add the water and the formamide first).

2.2  Fixation and Preparation of Adults and Juveniles

1. Filtered sea water (FSW) artificial (any of the available protocols will do) or natural (if available). 2. 7% MgCl2 in FSW (the concentration needed may vary depending on the species, if it is necessary use the alternative dilution of 3.5% MgCl). 3. 4% formaldehyde, methanol-free for juveniles, in FSW (use it freshly made). For adults it does not have to be necessarily methanol-free. Alternatively, a 4% paraformaldehyde solution, from powder, can be used. 4. 1× PBS. 5. 100% methanol. 6. 75% MeOH in PBS. 7. 50% MeOH in PBS. 8. 25% MeOH in PBS. 9. 20× SSC stock: 0.3 M Na citrate and 3 M NaCl in water and pH adjusted to 7.

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10. Bleaching solution (use it freshly made): 1.5% H2O2, 5% formamide, 0.5× SSC in milliQ RNase-free water (add water and formamide first). 2.3  Immunostaining

1. 1× PBS. 2. 0.1% PBTx: 0.1% Triton X-100 in 1× PBS. 3. Blocking solution: 6% Normal Goat Serum (NGS) or 0.5% Bovine Serum Albumin (BSA) in PBTx. 4. Primary antibodies: their working concentration in PBTx should be determined experimentally, given that different species and different antibodies will require specific concentrations. 5. Secondary antibodies: use a specific one, taking into consideration in which animal the primary antibody was raised, and the emission fluorescence you will need in your experiment. Dilute it in PBTx as specified for each product. 6. Molecular marker of interest (if any) diluted in PBTx (for instance, DAPI or phalloidin). 7. Fluoromount™ Aqueous Mounting Medium (SIGMA-­ ALDRICH, Cat. No. F4680-25ML) or any other similar mounting medium specific for fluorescence samples.

2.4  Colorimetric and Fluorescence Whole Mount In Situ Hybridization (ISH and FISH)

All the components used in these reactions should be prepared in sterile conditions, and using filter tips to prepare the solutions. Use sterile water or DEPC-treated water to prepare them or, alternatively, filter the stock solutions i.e.: PBTx, EDTA or SSC. It is highly recommended that every person prepare their own stocks and pipette from them to avoid contaminating the main stocks. It is important that the formamide used in this protocol is deionized. 1. 1× PBS. 2. 0.1% PBTx: 0.1% Triton X-100 in 1× PBS. 3. 100% methanol. 4. 75% MeOH in PBTx. 5. 50% MeOH in PBTx. 6. 25% MeOH in PBTx. 7. Mucolytic reagent: 5% N-acetylcysteine in PBTx (use it freshly made). 8. Fixative: 5% formaldehyde in PBTx (use it freshly made). 9. Reduction solution: 50 mM Dithiothreitol (DTT), 1% NP40, 0.5% Sodium Dodecyl Sulphate (SDS) in PBS (use it freshly made).

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10. 0.1 M Triethanolamine in PBTx (use it freshly made). Be sure that the pH is adjusted to 8 by adding enough HCl 1 M. 11. 2.5  μl/ml acetic anhydride in 0.1 M triethanolamine (use it freshly made, mix it right before its use, do not shake). 12. 5  μl/ml acetic anhydride in 0.1 M triethanolamine (use it freshly made, mix it right before its use, do not shake). 13. 20× SSC stock: 0.3 M Na citrate and 3 M NaCl in water and pH adjusted to 7. 14. 0.05 M EDTA stock: dissolve the correct amount of ethylenediaminetetraacetic acid in water. It is important to bring the pH to 8, with 10 M NaOH, for complete dissolution. 15. Hybridization Buffer (HybBuffer): 50% deionized formamide, 5× SSC buffer pH 7, 0.005 M EDTA, 0.1% tween-20, 0.1 mg/ ml heparin, 1× Dendhart’s solution, 1 mg/ml yeast RNA (or 0.1 mg/ml salmon sperm DNA) in sterilized or DEPC-treated water. Add the components in this order, with the exception of the water, that should be added in the third place, after the SSC. Keep the solution at −20 °C. 16. Wash Solution: same as HybBuffer, but without heparin, Dendhart’s solution and the yeast RNA (or salmon sperm DNA). Keep it at −20 °C. 17. RNA probe of your gene of interest, diluted in HybBuffer. Probes are synthesized according to the instructions provided by the company of your selected kit. The optimal concentration of the probe has to be determined empirically, it is recommended to start at 1 ng/μl, and depending on the intensity of the signal (and the signal to noise ratio) modify the concentration, usually in a range between 0.5 and 2 ng/μl. (It is important to remember that the probe has to be synthesized using, apart from the regular dNTPS, with also modified UTP, a labeling compatible with the conjugated antibody that will be used later on, for instance Digoxigenin-11-UTP if a Digoxigenin antibody will be used.) Keep the probes stored at −20 °C. 18. 2×SSC-Formamide: 2× SSC and 50% formamide in water. 19. 0.2×SSC-Tx: 0.2× SSC and 0.1% Triton X-100 in water. 20. 0.1% MAB-TrX: Prepare Maleic Acid Buffer with 100 mM maleic acid and 150 mM NaCl in water. Adjust the pH to 7.5 with NaOH. Add 1 μl Triton 100-X for every ml of MAB. 21. Blocking solution: Use a 2% blocking reagent (from Roche) and 10% horse serum, heat inactivated (H.I.), in MAB-TrX. 2.4.1  Only for ISH (Colorimetric In Situ Hybridization)

1. Anti-DIG-AP Fab fragments (Roche, Cat. No. 000000011093274910): antigen-binding fragments from polyclonal anti-digoxigenin antibodies, conjugated to alkaline

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phosphatase (we use regularly alkaline phosphatase, AP, reaction to detect the signal of our probe). 2. AP buffer: 100 mM Tris–HCl pH 9.5, 100 mM NaCl, 50 mM MgCl2, 0.1% Tween-20, adjust the volume with water. 3. AP substrate solution: add to the AP buffer: 20 μl/ml of NBT-BCIP. 4. 80% Glycerol in PBTx. 2.4.2  Only for FISH (Fluorescent In Situ Hybridization)

1. Anti-DIG-POD Fab fragments (from Roche, Cat. No. 11207733910): antigen binding fragments from polyclonal anti-digoxigenin antibodies, conjugated to horseradish peroxidase. 2. Anti-DNP-HRP (from Perkin Elmer): anti-Dinitrophenyl conjugated to horseradish peroxidase. If we perform double FISH experiments, each probe will be detected with a different antibody, so both probes should be synthesized with their corresponding labeled dUTPs. 3. Tyramide Signal Amplification (TSA) red and/or green. 4. H2O2 30% stock. 5. 2×SSC-Formamide-Tx: 2× SSC, 50% formamide and 0.1% Triton X-100 in water. 6. TSA Buffer: 2 M NaCl and 100 mM Borate buffer in water, pH 8.5. 7. Fluoromount™ Aqueous Mounting Medium (SIGMA-­ ALDRICH, Cat. No. F4680-25ML) or any similar mounting medium specific for fluorescence samples.

2.5  Equipment

1. Stereoscope. 2. Bright field microscope (ISH samples). 3. Fluorescence microscope with mercury or LED lamps (fluorescent samples). 4. Confocal microscope (fluorescent samples). 5. Horizontal shaker. 6. Timer. 7. Petri dishes. 8. Pasteur pipettes. 9. 15 ml Falcon tubes. 10. Filter with a mesh of pore size 300 um (embryo sampling). 11. Glass embryo dishes (embryo preparation). 12. Forceps (embryo preparation). 13. Pipettes set 1000 μl; 200 μl; 20 μl and 2 μl.

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14. Sterile filter tips (ISH and FISH first steps). 15. Multiple-wells sterile tissue culture plates (immunostaining, ISH and FISH). 16. Baskets of 0.2 μm mesh pore size (embryo and juvenile ISH and FISH). 17. Oven (for ISH and FISH). 18. Glass microscope slides, StarFrost. 19. Cover slips: we use 18 × 18 and 24 × 32 mm, 0.13–0.17 mm thick.

3  Methods All steps are carried out at room temperature, unless otherwise stated (see Note 1). 3.1  Sampling and Fixation 3.1.1  Embryo Sampling and Fixation

1. Obtain cocoons from the total population (kept in glass trays) by filtering through a fine mesh of 300 μm. Juveniles and adults will pass through the filter due to their slender body shape and to another tray. Keep the filter in close contact to the sea water of the tray while pouring all the media with the acoel sample population through the mesh. Most of the cocoons stay in the filter because of its volumetric size. Transfer them to a dish by inverting the filter and washing it with FSW to detach the cocoons (Fig. 2a). They can be fixed at different times of their development if necessary (see Notes 1 and 2). 2. Collect the cocoons with a pipette (see Note 3) and introduce them in a 15 ml Falcon tube (see Note 4). 3. Suck up the FSW as much as possible, but be careful not to dry the embryo sample completely (see Note 5). Fill the Falcon tube with a Thioglycolate-Pronase solution, freshly made, and incubate the embryos for 30 min, on the shaker. 4. Wash three times × 5 min each in FSW. 5. Fix with 4% formaldehyde, methanol-free, in FSW during 1 h at room temperature, or overnight at 4 °C. 6. Wash three times × 5 min each in PBS. 7. For ISH and most procedures involving the immunostaining of embryos (see also Note 6): dehydrate the samples progressively with the following three washes: 25% MeOH in PBS; 50% MeOH in PBS; 75% MeOH in PBS; use at least 30 min per wash. 8. Keep samples at −20 °C in 100% MeOH.

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Fig. 2 (a) Typical glass tray in which acoels are kept. The content of a tray with sea water and acoels is poured through the cylindrical filter. This way, cocoons will stay in the filter and the adults and juveniles will swim through the mesh to a clean tray. Put the filter upside down on a Petri dish and detach the cocoons from the mesh applying pressure with some clean seawater. (b) Multiwell dish typically used to ISH and FISH protocols. Small baskets (arrow) are used to keep the animals. This way, liquid can be aspired in each wash without risk of taking the specimens 3.1.2  Juveniles’ Sampling and Fixation

1. Separate an amount of cocoons of approximately the same developmental stage in a Petri dish until they hatch (see step 1 for Subheading 3.1.1). After hatchling, remove the cocoon wrapping membranes by filtering and proceed to fixation (juveniles pass through the filter). Animals can be kept in FSW for the necessary time to obtain juveniles of the desired age. 2. Wash three times × 5 min in FSW to remove any cocoon debris (see Note 5). 3. Relax the worms by adding some drops of 7% MgCl2 to the FSW dish (see Note 7).

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4. Collect the juveniles with a Pasteur pipette (see Note 3) and introduce them in a 15 ml Falcon tube (see Note 4). 5. Fix them with 4% formaldehyde, methanol-free, in FSW, during 1 h at room temperature or overnight at 4 °C. 6. Wash three times × 5 min in PBS (see Note 5). 7. For ISH and, as before, most procedures involving the immunostaining of juveniles (see Note 6): dehydrate the samples progressively through the following three solutions: 25% MeOH in PBS; 50% MeOH in PBS; 75% MeOH in PBS; using at least 30 min per wash. 8. Keep samples at −20 °C in 100% MeOH. 3.1.3  Adult Sampling and Fixation

3.2  Sample Preparation and Photobleaching for Embryos, Juveniles and Adults

1. Collect some adults using a Pasteur pipette. 2. Follow the steps 2–8 of the protocol for Subheading 3.1.2. The only difference for adults is that the formaldehyde for the fixation step does not need to be methanol-free, unless they will be used in in situ hybridization protocols (the specifics for the protocol with adults are not included in this chapter). Although a photobleaching treatment is not necessary for all immunostaining and FISH experiments, it always helps to improve the quality of the results, given that it reduces the specimens’ autofluorescence. Moreover, in the case of Symsagittifera roscoffensis, for adults we highly recommend to photobleach the samples, due to the presence of pigment from the symbiotic algae. Skip the photobleaching step if the downstream applications are not fluorescence-related. We recommend carrying out the next steps in embryo dishes. If you are going to perform an in situ hybridization protocol later, remember to use sterilized and RNAse-free material, clean gloves, etc. in all the steps. 1. Transfer to an embryo dish. 2. (Only if the sample was kept in MeOH.) Rehydrate the samples progressively with the following three incubations: 75% MeOH in PBS; 50% MeOH in PBS; 25% MeOH in PBS; use, at least, 10 min per wash. 3. Wash three times × 5 min each in PBS. 4. Optional step, only for embryos (depending on the species, though it is always necessary for S. roscoffensis): in the stereoscope, open the cocoons using forceps, breaking the external membrane. It is not necessary to remove the membrane completely, just leave it open (see Note 8). 5. Remove the PBS (see Note 5) and add 600 μm of the bleaching solution to each embryo dish. Put them under a direct white light (see Note 9) and keep the animal samples during

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15 min for embryos and juveniles. For adults, keep them, instead, during 30 min or more (see Note 10). 6. Suck up the bleaching solution and wash the samples three times × at least 5 min each in PBS. 7. Transfer the animals to a clean embryo dish or a multiwell plate and carry on with the downstream applications. Alternatively, keep specimens stored at 4 °C in PBS. 3.3  Whole Mount Immunostaining for Juveniles and Adults

1. Suck up the PBS (as much as you can, see Note 5) and add the blocking solution NGS/PBTx, which performs permeabilization and blocking, 1 h RT on the shaker (see Note 11). 2. Remove the blocking solution. Add the primary antibody or antibodies (if a double or triple immunostaining is desired) diluted in PBTx (see Note 12). Keep shaking the samples at 4 °C overnight (see Note 13). 3. Wash 3 × 5 min each in PBS. 4. Change the PBS solution for the secondary antibody diluted in PBTx (see Note 14). Keep the samples shaking for 2 h at room temperature, or overnight 4 °C. 5. Remove the liquid (as much as you can). Wash with PBS three times × 5 min each. Sometimes, other specific structures can be easily revealed through fluorescent chemical molecules that bind them. Some of those additional markers are: phalloidin [muscular tissue marker (see Note 15)], WGA (Wheat Germ Agglutinin, as plasmatic membrane marker) or DAPI (cells nuclei marker). Usually they work easily in most of the samples. As an optional step, samples can be also incubated with some of them just before mounting. 6. Incubate in the chemical marker diluted in PBTx for 20 min to 1 h at RT while shaking. 7. Wash 3 × 5 min each PBS. 8. Mounting (if this step is not carried out immediately after the protocol, keep the sample stored at 4 °C). Transfer the worms to microscope slides and add about 15 μl of Fluoromount. Cover with 18 × 18 or 24 × 32 cover slips (fix the corners with nail polish) and let the preparation dry over night at 4 °C. To avoid that the animals look smashed, add some modeling clay to the corners of the cover slips before mounting. 9. A preliminary check of the results of the above protocol can be carried out with a fluorescence microscope with mercury or LED lamp. However, for more detailed analysis, it is advised to acquire images using a confocal microscope (Fig. 3c, pink).

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Fig. 3 Expression of synaptotagmin gene orthologue in embryo and juvenile in of the acoel S. roscoffensis. (Reproduced from EvoDevo 2018 with permission from BioMed Central) [34]. (a) ISH expression pattern of the gene in a 24 h post-fertilization (hpf) embryo. Asterisk indicates the anterior pole. (b) ISH expression of the same gene in a juvenile from 12 to 24 h post hatching (hph). Arrowhead points the position of the statocyst, and therefore the anterior part. C. FISH expression pattern (in green) of the same gene combined with anti-­ SrStg antibody (in pink) on a 12–24 hph juvenile. Embryo scale bar: 60 μm; Juveniles scale bar: 100 μm

In order to develop the protocol explained above, we started with the one previously used in our lab described in [25], and then make the necessary changes for improving it. Moreover, there are other available immunochemistry protocols for acoels that have been used in other laboratories like for instance [26, 29, 30]. 3.4  Colorimetric and Fluorescence Whole Mount In Situ Hybridization (ISH and FISH) for Embryos and Juveniles

The first steps of the protocol are identical for both. In the following text, it will be specified when the steps or reactions are specific for ISH or FISH. Always use filter tips until it is indicated and handle the samples with clean gloves. Preform all steps at room temperature unless otherwise stated. Embryos and juveniles previously fixed are kept stored at −20 °C in 100% MeOH. Take the desired amount of them to be analyzed, of the specific developmental time selected, and place them in a multiple-wells sterile tissue-culture plate. We perform the washes with 500 μl volume, but that will depend on the size of each well (keep in mind Note 5). The procedure is now developed in a more condensed form, since the details of most steps have been described before. DAY 1 1. Rehydrate the animals using the following steps:

(a) 5 min wash with 75% MetOH in PBTx.



(b) 5 min wash with 50% MetOH in PBTx.



(c) 5 min wash with 25% MetOH in PBTx.



(d) 2× 5 min wash with PBTx. After rehydration, we can take the animals from the wells, put them in small baskets (Fig. 2b) and then transfer them into other clean wells. This way it is easier to perform the washes through aspiration, without taking accidentally the specimens. For each change, work with vacuum pipette, taking out the

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liquid with one hand (from outside the basket) while at the same time you are adding the next solution with the other (remember Note 5). 2. Preform the mucolytic reagent treatment during 10 min with 5% N-acetylcysteine in PBTx (see Note 16). 3. Refix the animals during 30 min with 5% formaldehyde in PBTx. After this step, wash the animals 2× 5 min in PBTx. 4. Preform the chemical reduction treatment during 5 min in “reduction solution” (50 mM DTT, 1% NP40 and 0.5% SDS in PBS) (see Note 16). 5. Carry out the triethanolamine and acetic anhydride treatments following the steps:

(a) 2× 5 min wash with 0.1 M triethanolamine (pH 8).

(b)  5 min in 2.5 μl/ml acetic anhydride in 0.1 M triethanolamine. (c)  5 min in 5 μl/ml acetic anhydride in 0.1 M triethanolamine.

(d) 2× 5 min wash PBTx.

6. Rinse the sample in HybBuffer and pre-hybridize during 1–2 h, also in HybBuffer and at 70 °C. 7. Hybridization:

(a) The probes should be stored at −20 °C. Before hybridizing, denature the probe at 70 °C, during 10 min. If we perform double FISH, prepare a mix of the 2 probes, elaborated with differently labeled nucleotides (UTPs), which will react later with their corresponding antibodies (see Subheading 2 in the FISH probe part).



(b)  Replace the pre-hybridization solution with a solution containing the denatured probe.



(c) 2 or 3 days hybridization at hybridization temperature (58–62 °C) in a shaking hybridization oven (see Note 17). Use a wet chamber to avoid evaporation (see Note 18).

DAY 2 8. Do the post-hybridization washings (do at the hybridization temperature while shaking):

(a) 10 min wash with preheated HybBuffer.



(b) 4× 15 min wash with preheated Wash Solution.



(c) 2× 10 min wash with 2×SSC-Formamide.



(d) 2× 5–10 min wash with 0.2×SSC-Tx. In the second wash, change to room temperature and continue at room temperature until something else stated (see Note 19).



(e) 2× 5–10 min in 0.1% MAB-TrX.

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9. Proceed with the blocking during 30 min, using 2% of Blocking reagent plus 10% Horse serum H.I. (heat inactivated) in MABTrX (shake slowly). From now on keep the samples in dark, in order to avoid the fading of the signal. 10. Carry out the antibody incubation:

(a) 4 h at RT or ON at 4 °C incubation with the appropriate antibody, diluted in blocking reagent, and on a rotating shaker (check the ISH and FISH material sections for more details about the antibodies that should be used, according to how the probe was synthesized). We use, regularly, the following dilutions: ••

Anti-DIG-AP 1:5000 for colorimetric ISH.

••

Anti-DIG-POD 1:500 for FISH (see Note 20).

Anti-DNP-HRP 1:200 for FISH. DAY 3 From now on, there is no need of using filter tips. ••

11. Wash the samples during 2 h in 0.1% MAB-TrX, Change to new solution every 15 min. Take the animals from the baskets and transfer them (with the pipette) to new wells. After this step is completed, there are two options for continuing the protocol: following a colorimetric ISH or a fluorescent alternative, FISH. 3.4.1  Only for Colorimetric ISH (NBT-BCIP Labeling)

1. Start the labeling process by incubating 2× 5 min in AP buffer. Let it develop in AP substrate solution, at room temperature and in dark (see Note 21). 2. Stop the reaction’s development using the following washes:

(a) 1× 5 min in AP buffer.



(b) 2× 5 min PBTx rinses and then wash several times consecutively in PBTx.



(c) 30 min post-fix incubation in 5% formaldehyde in PBTx.



(d) Wash thoroughly with PBTx, at least 3 × 15min.

3. Proceed to mount the samples:

(a) In 80% Glycerol in PBTx. Add modeling clay to the corners of the cover slip if you consider it necessary, as explained in the mounting of the immunostaining section.

4. Check the samples under a Bright field Microscope. It is recommended to take pictures of several different animals for each probe, in order to see if there is individual variability in the staining patterns (Fig. 3a for embryo example and B for juvenile).

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1. Wash 2× during 5 min PBTx. 2. Label the samples following the next steps:

(a)  Incubate 30 min with the preferred Tyramide Signal Amplificaton (TSA green or red) at 1:300 in TSA Buffer.



(b) First Spike (for reamplification): adding H2O2 to a final concentration of 0.002–0.015% and let it develop for 30 min.



(c) ReSpike 2 times and let develop for 30 min each. ReSpike once more for 1 h. If necessary, perform extra ReSpike steps for 1 h development each. For doing so, add 2.5 μl/ well from the 1:10 dilution of the 30% H2O2 stock.

3. Rinse 2× 5 min in PBTx and then perform several washes with the same solution. If we are preforming FISH with only one probe: STOP here the protocol (see Note 22) and mount the samples in Fluoromount and put the slides at 4 °C. Acquire images using a confocal microscope. If we are preforming double FISH: carry on with the protocol as stated below. Quench the first antibody (conjugated to the first probe) before incubating the second one (see Note 23). 4. Preform the quenching steps:

(a) First quenching: 45 min in 1% H2O2 in PBTx shacking.



(b) 2× 5 min PBTx wash.

(c)  Second quenching: 10 min at 56 °C in 2×SSCFormamide-­Tx, shaking.

(d) 2× 5 min PBTx.

Go back to step 10 (Subheading 3.4), and chose the second antibody this time (Anti-DNP-HRP 1:200). Carry on steps 10 and 11 (Subheading 3.4) for the general protocol, then steps 1–3 (Subheading 3.4.2) specific for FISH, choosing a different TSA from the one used previously. To finish, mount the samples in Fluoromount as explained in the immunostaining section, and keep the slides at 4 °C. Acquire images using a confocal microscope (see example of a juvenile single FISH plus immunostaining in Fig. 3c). For developing these ISH and FISH protocols described above, we started using the ones described in [24, 31, 32], making the needed modifications. Other ISH available protocols also used for acoels are described in [30, 33].

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4  Notes 1. Please take into account that the protocols detailed here are optimized for our model species Symsagittifera roscoffensis. Other species may need slight variations, due to their different development, size, or color, among others. 2. To obtain cocoons from a certain time window (developmental time), clean the tray from all (pre-laid) cocoons through filtering. Keep the adults under observation until they lay the first cocoons and leave these cocoons for a desired period of time, to obtain different animals of different developmental stages. For instance, filtering again 6 h after the laying of the first cocoons, allows obtaining embryos from 0 to 6 h (the animals lay asynchronously the cocoons in the dish). Also, filtered cocoons can be kept in Petri dishes longer time to obtain later developmental stages embryos and eventually the juveniles. 3. To take embryos or juveniles from a Petri dish or a tray and transfer them to a Falcon tube (or Eppendorf tubes, plates, etc.) utilize the centrifuge force, moving the plate with your hands in the horizontal plane in circles or, alternatively, in a shaker, trying to concentrate the specimens in the middle of the plate, all together. Transferring is now easier through pipetting. For that, it is necessary to move manually the plate slowly making circles while with the other hand we pipette the embryos. This step could be done in the stereoscope because it is easier to follow the sample during the process. 4. The following steps are done in 15 ml Falcon tubes and in a horizontal shaker. Sometimes keeping them vertical for a few seconds before changing to the following solution helps the precipitation of the embryos (a similar phenomenon also happens in juveniles and adults). This step makes easier to throw away the liquid by decantation or pipetting. 5. It is recommended, in all the protocols, to keep some solution left in the dish, or well, or Falcon tube, etc., between all the changes, otherwise the sample tends to stick to the walls of the container and becomes damaged. 6. Samples stored in MeOH can be used for both in situ hybridization and immunostaining. However, some immunostaining reactions may not work as well in MeOH stored samples. In those cases they can also, alternatively, be kept in PBS (for short time periods) or PBS + 0.1% NaN3 (for longer time periods; NaN3 avoids fungi growth) at 4 °C. 7. MgCl2 should be added slowly, and constantly, observing the animals’ movements, until all of them are completely relaxed.

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Not using enough MgCl2 can lead to muscle contractions when the animals are fixed. Importantly, using too much MgCl2 or too high concentration could damage the specimens or even disintegrate them completely. 8. The following steps of the protocol will remove the rests of the membrane left. 9. Covering the base and the sides of the recipient with aluminum paper helps with the photobleaching process, because the light is reflected directly to the sample. It is also necessary to remove any lid or cover to the dish. 10. For S. roscoffensis, due to the presence of a symbiotic algae, it is necessary to keep adults in the solution longer than 30 min, until we see them turning almost white. But, careful, too much time in the bleaching solution can easily break the samples. 11. Alternatively, a mix of BSA in PBTx also blocks the unspecific binding sites, while PBTx (because the presence of detergent) permeabilizes the sample. However, sometimes we have observed that the result looks cleaner when using NGS. 12. The working concentration of the primary antibody may vary in each particular case, due to the own properties of the antibody, the animal species and also the developmental stage of the animals in the sample. It is necessary to try several dilutions to determine the most appropriate concentration. At too low concentration, the antibody will not penetrate properly and at too high concentration it would tend to aggregate on the surface of the animal. A good concentration to start with is usually 1:10. From there, further dilutions can be tried. In our experience, antibodies designed specifically for our species (e.g. synaptotagmin antibody) need to be used at higher concentration (1:100 at least) than commercial ones [i.e. tyrosinated tubulin (1:200) and serotonin (1:500 to 1:1000) antibodies]. Moreover, some antibodies need to be preabsorbed in order to work. Whereas others do not need to, we observe, in general, that better results are obtained if they go through a preabsorption step. 13. The incubation time may vary, depending on the antibody, the developmental stage and the animal species used. Some antibodies in S. roscoffensis need longer incubations. For instance, in adult’s synaptotagmin antibody needs more than 72 h at 4 °C to acquire a good quality staining of the nervous system. The same antibody in juveniles can be detected readily after 24 h incubation). 14. A series of concentrations should be checked for each secondary antibody used [i.e. anti-mouse (1:200) and anti-rabbit

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(1:1000)]. Too high concentration of the second antibody could produce unspecific fluorescence on the whole organism. 15. We use phalloidin at 1:100 concentration (diluted in PBTx). It is important to fix samples that will be reacted with phalloidin, with MeOH-free formaldehyde (alternatively, fresh prepared 4%PFA can be used as well) and instead of keeping them in MeOH, better use PBS + NaN3, as explained in Note 6. 16. Mucolytic and reduction treatments are preformed to eliminate, as much as possible, any sticky material present in the epidermis of the animals, to which the probe could get stuck. In this way, the background is reduced for most probe reactions. Acoels, as Platyhelminthes, secrete a lot of mucus, needed for mobility and bacterial protection. 17. The hybridization temperature should be determined empirically for each probe (it depends of their Tm). As a rule of thumb we always start trying 60 °C. If too much background is observed increase the hybridization temperature to a more restrictive one (max 62 °C). If no signal is observed, then proceed to the converse strategy, decrease the temperature to facilitate hybridization (but no less than 58 °C). 18. For instance, we can use a hermetic box and cover the bottom with wet cotton wool. The multiwell dish is placed inside this box, so the evaporation is reduced. 19. If you experience that your probe gives too much background, increase the number of washes. 20. If you are doing a double FISH protocol, with Anti-DIG-­ POD and Anti-DNP-HRP, in this step ONLY put Anti-DIG-­ POD. It allows more washes without losing the signal quality. Anti-DNP-HRP to detect the other probe will be used later. 21. The developing time can vary considerably between probes (according to Tm, probe concentration, prevalence of the endogenous mRNA, etc.), from some minutes to several days. It is strongly recommended to check every 5–10 min during the first hour, just to gauge the speed of signal development. 22. If we want to mark the sample with another molecule (DAPI, phalloidin) this labeling can be done at the end of the protocol. Also, if we want to perform a single FISH and immunostaining, at this point, go to Subheading 3.3 and perform the immunostaining protocol. 23. When double FISH is performed using TSA requires 2 rounds of amplification (one for each probe) with reagents conjugated to peroxidase. In order to avoid residual peroxidase activity from the previous reaction (that would generate false signal) it is necessary to quench (to stop) peroxidase activity between the use of one TSA and the other.

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Acknowledgments The research in P. Martínez laboratory was carried out with the support of the Spanish Ministry of Science, Grants BFU2006-­00898, BFU2009-07383, and BFU2012-32806. E. Perea-Atienza and B. Gavilán were supported by PhD fellowships from the Universitat de Barcelona (APIF). S.G. Sprecher acknowledges the Swiss National Science Foundation 31003A_169993. Elena Perea-­ Atienza and Brenda Gavilán contributed equally to the development of the described methodology. The authors would also like to thank Kathryn Apse and Prof. Seth Tyler (University of Maine) for letting us publish the acoel morphology diagrams in Fig. 1 of this chapter. References 1. Cannon JT, Vellutini BC, Smith J, Ronquist F, Jondelius U, Hejnol A (2016) Xenacoelomorpha is the sister group to Nephrozoa. Nature 530:89–93. https://doi. org/10.1038/nature16520 2. Philippe H, Brinkmann H, Copley RR, Moroz LL, Nakano H, Poustka AJ, Wallberg A, Peterson KJ, Telford MJ (2011) Acoelomorph flatworms are deuterostomes related to Xenoturbella. Nature 470:255–260. https:// doi.org/10.1038/nature09676 3. Baguñà J, Riutort M (2004) The dawn of bilaterian animals: the case of acoelomorph flatworms. BioEssays 26:1046–1057 4. Jondelius U, Wallberg A, Hooge M, Raikova OI (2011) How the worm got its pharynx: phylogeny, classification and bayesian assessment of character evolution in acoela. Syst Biol 60:845–871. https://doi.org/10.1093/sysbio/syr073 5. Achatz JG, Chiodin M, Salvenmoser W, Tyler S, Martinez P (2012) The Acoela: on their kind and kinships, especially with nemertodermatids and xenoturbellids (Bilateria incertae sedis). Org Divers Evol 13:267–286 6. Arroyo AS, López-Escardó D, de Vargas C, Ruiz-Trillo I (2016) Hidden diversity of Acoelomorpha revealed through metabarcoding. Biol Lett 12:20160674. https://doi. org/10.1098/rsbl.2016.0674 7. Smith J, Tyler S (1986) Frontal organs in the Acoelomorpha (Turbellaria): ultrastructure and phylogenetic significance. Hydrobiologia 132:71–78 8. Ehlers U (1991) Comparative morphology of the statocysts in the Plathelmithes and the Xenoturbellida. Hydrobiologia 227:263–271

9. Yamasu T (1991) Fine structure and function of ocelli and sagittocysts of acoel flatworms. Hydrobiologia 227:273–282. https://doi. org/10.1007/BF00027612 10. Rieger RM, Tyler S, Smith JPS, Rieger GE (1991) Platyhelminthes: Turbellaria. In: Harrsion FW, Bogitsch BJ (eds) Microscopic anatomy of invertebrates. Wiley, New York 11. Raikova OI, Reuter M, Kotikova EA, Gustafsson MKS (1998) A commissural brain! The pattern of 5-HT immunoreactivity in acoela (Plathelminthes). Zoomorphology 118:69–77. https://doi.org/10.1007/ s004350050058 12. Reisinger E (1925) Ein landbewohnender Archiannelide. (Zugleich ein Beitrag zur Systematik der Archianneliden). Z Morphol Tiere 3:197–254 13. Martínez P, Hartenstein V, Sprecher SG (2017) Xenacoelomorpha nervous systems. In Oxford Research Encyclopedia of Neuroscience. Ed. S. Murray Sherman. New York: Oxford University Press 14. Jennings JB (1957) Studies on feeding, digestion, and food storage in free-living flatworms (Platyhelminthes: Turbellaria). Biol Bull 112:63–80 15. Achatz J, Gschwentner R, Rieger R (2005) Symsagittifera smaragdina n. spec., a new acoel (Acoela, Acoelomorpha) of the Mediterranean Sea. Zootaxa 1085:33–45 16. Pedersen KJ (1964) The cellular organization of Convoluta convoluta, an Acoel Turbellarian: a cytological, histochemical and fine structural study. Z Zellforsch 64:655–687 17. Hyman LH (1951) The invertebrates: Platyhelminthes and Rhynchocoela. The acoe-

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lomate bilateria vol II. McGraw-Hill Book Company, Inc, New York 18. Hooge M (2001) Evolution of body-­wall musculature in the Platyhelminthes (Acoelomorpha, Catenulida, Rhabditophora). J Morphol 249:171–194 19. Hooge M, Tyler S (2006) Concordance of molecular and morphological data: The example of the Acoela. Integr Comp Biol 46: 118–124 20. Semmler H, Bailly X, Wanninger A (2008) Myogenesis in the basal bilaterian Symsagittifera roscoffensis (Acoela). Front Zool 5:1–15. https://doi.org/10.1186/1742-9994-5-14 21. Henry JQ, Martindale MQ, Boyer BC (2000) The unique developmental program of the acoel flatworm, Neochildia fusca. Dev Biol 220:285–295. https://doi.org/10.1006/ dbio.2000.9628 22. Hejnol A (2015) Acoelomorpha and Xenoturbellida. In: Evolutionary developmental biology of invertebrates, vol 1. Springer Verlag, New York, pp 203–214 23. Raikova O, Reuter M, Gustafsson MKS, Maule AG, Halton DW, Jondelius U (2004) Evolution of the nervous system in Paraphanostoma (Acoela). Zool Scr:71–88 24. Semmler H, Chiodin M, Bailly X, Martinez P, Wanninger A (2010) Steps towards a centralized nervous system in basal bilaterians: Insights from neurogenesis of the acoel Symsagittifera roscoffensis. Develop Growth Differ 52:701–713. https://doi. org/10.1111/j.1440-169X.2010.01207.x 25. Achatz JG, Martinez P (2012) The nervous system of Isodiametra pulchra (Acoela) with a discussion on the neuroanatomy of the Xenacoelomorpha and its evolutionary implications. Front Zool 9:27 26. Raikova OI (2004) Neuroanatomy of basal bilaterians (Xenoturbellida, Nemertodermatida, Acoela) and its phylogenetic implications (PhD thesis). Åbo Akademi University 27. Perea-Atienza E, Gavilan B, Chiodin M, Abril JF, Hoff KJ, Poustka AJ, Martinez P (2015) The nervous system of Xenacoelomorpha: a

genomic perspective. J Exp Biol 218:618–628. https://doi.org/10.1242/jeb.110379 28. Gavilán B, Perea-Atienza E, Martínez P (2016) Xenacoelomorpha: a case of independent nervous system centralization? Philos Trans R Soc B Biol Sci 371. https://doi.org/10.1098/ rstb.2015.0039 29. De Mulder K, Kuales G, Pfister D, Willems M, Egger B, Salvenmoser W, Thaler M, Gorny AK, Hrouda M, Borgonie G, Ladurner P (2009) Characterization of the stem cell system of the acoel Isodiametra pulchra. BMC Dev Biol 9: 1–17. https://doi.org/10.1186/1471-213X9-69 30. Srivastava M, Mazza-Curll KL, Van Wolfswinkel JC, Reddien PW (2014) Whole-body acoel regeneration is controlled by Wnt and Bmp-­ Admp signaling. Curr Biol 24:1107–1113. https://doi.org/10.1016/j.cub.2014.03.042 31. Chiodin M, Børve A, Berezikov E, Ladurner P, Martinez P, Hejnol A (2013) Mesodermal gene expression in the acoel Isodiametra pulchra indicates a low number of mesodermal cell types and the endomesodermal origin of the gonads. PLoS One 8:e55499. https://doi. org/10.1371/journal.pone.0055499 32. Albuixech-Crespo B, López-Blanch L, Burguera D, Maeso I, Sánchez-Arrones L, Moreno-Bravo JA, Somorjai I, Pascual-Anaya J, Puelles E, Bovolenta P, Garcia-Fernàndez J, Puelles L, Irimia M, Ferran JL (2017) Molecular regionalization of the developing amphioxus neural tube challenges major partitions of the vertebrate brain. PLoS Biol 15. https://doi.org/10.1371/journal.pbio.2001 573 33. Hejnol A, Martindale MQ (2008) Acoel development indicates the independent evolution of the bilaterian mouth and anus. Nature 456:382–386. https://doi.org/10.1038/ nature07309 34. Perea-Atienza E, Sprecher SG, Martínez P (2018) Characterization of the bHLH family of transcriptional regulators in the acoel S. roscoffensis and their putative role in neurogenesis. EvoDevo 9:1–16. https://doi. org/10.1186/s13227-018-0097-y

Chapter 5 Immunostaining of the Embryonic and Larval Drosophila Brain Frank Hirth and Danielle C. Diaper Abstract Immunostaining is used to visualize the spatiotemporal expression pattern of developmental control genes that regulate the genesis and specification of the embryonic and larval brain of Drosophila. It is also used to visualize the effects of targeted misexpression or inactivation of disease-related genes. Immunostaining uses specific antibodies to mark expressed proteins and allows their localization to be traced. This method reveals insights into gene regulation, cell type specification, neuron and glial differentiation, axonal and synaptic scaffolding and posttranslational protein modifications underlying the patterning and specification of the maturing brain. Depending on the targeted protein, it is possible to visualize a multitude of regions of the Drosophila brain, such as small groups of neurons or glia, defined subcomponents of the brain’s axon scaffold, or pre- and postsynaptic structures of neurons. Thus, antibody probes that recognize defined tissues, cells, or subcellular structures like axons or synaptic terminals can be used as markers to identify and analyze phenotypes in embryos and larvae. Several antibodies, combined with different labels can be used concurrently to examine protein colocalization. This protocol spans over 3–4 days. Key words Drosophila, Embryo, Larva, Brain, Immunostaining, Fluorescence immunocytochemistry, Dissection, Antibody

1  Introduction Similar to mammalian brain development, the Drosophila brain derives from a monolayered epithelium called the neuroectoderm. Subsequent neurogenesis is characterized by two neurogenic periods: one during embryogenesis and another during larval and pupal stages. The precursor cells of the developing brain, termed neuroblasts (NBs), derive from the embryonic procephalic neurogenic region to form proliferative clusters; they divide repeatedly and asymmetrically in a stem cell mode to generate a new NB and a smaller daughter cell, called ganglion mother cell (GMC). Each GMC is a transient intermediate progenitor cell that generally divides once to produce two lineage-specific postmitotic cells, either neuron or glia. Both neuron and glia subsequently initiate

Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_5, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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their differentiation processes, finally shaping neural segments and circuits along the major body axes of the embryo [1, 2]. The central brain of Drosophila derives from 106 embryonic brain NBs that can be identified based on their positional relationships and NB-specific gene expression [3]. Towards the end of embryogenesis, most NBs stop proliferating and enter a period of quiescence. During the first larval instar stage, quiescent NBs reenter the cell cycle in a characteristic spatiotemporal pattern to perform a second round of neurogenesis, resulting in postembryonic neural progeny. These two phases of brain development are followed by an extensive morphological transformation during metamorphosis that ultimately leads to the adult fly brain [1–3]. Although approximately 95% of the 200,000 neurons in the adult Drosophila brain are generated post-embryonically, the main architecture of the adult brain is already laid down during embryonic neurogenesis [3, 4]. Thus, the Drosophila brain is composed of an anterior (supraesophageal) and a posterior (subesophageal) part, both of which are interconnected by an axon scaffold surrounding the gut; the anterior part comprises the protocerebral, deutocerebral, and tritocerebral neuromeres, whereas the posterior part comprises the mandibular, maxillary, and labial neuromeres (Fig. 1). Immunostaining of the embryonic and larval Drosophila brain is used for two primary purposes. First, once a gene of interest has been cloned and antibodies raised against its encoded protein product(s), immunostaining can be used to visualize the spatiotemporal pattern of protein expression during embryonic and larval brain development. Second, antibody probes that recognize defined tissues, cells, or subcellular structures like axons or synaptic terminals can be used as markers to identify and analyze phenotypes in embryos and larvae of germline mutants or where a disease-­related gene is misexpressed. Thus, following protein localization throughout Drosophila development reveals insights into gene regulation, cell type specification, neuron and glial differentiation, and posttranslational protein modifications underlying the patterning, specification, and neuronal connectivity of the maturing brain in a wild-type or disease-related condition. In this respect, whole-mount immunohistochemistry (IHC), which is the process of using antibody probes to detect antigens (i.e. proteins), is among the most valuable of tools for analysis. It is a relatively cheap and reliable way to visualize discrete structures or cell types and neuronal structures, including synapses, in the embryonic and larval brain of Drosophila. With the use of multiple antibodies, or when used in conjunction with genetic labeling, such as lineage analysis or synaptic tagging [5], it is possible to follow several different proteins, examining colocalization, or identify specific cells and follow their neuronal projections from the cell body to its terminal dendritic arborizations [6–8]. Immunostaining

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Fig. 1 The embryonic brain of Drosophila. (a–d) Laser confocal microscopy, superimposition of optical sections of stage 14 embryo immunolabeled with anti-HRP-FITC. (a) frontal view of brain; (b) ventral view of ventral nerve cord; (c) lateral view of brain; (d) optical section along the midline of (c); (e) cartoon summarizing major neural segments and axon scaffolding which together constitute the embryonic Drosophila brain. Abbreviations: sec supraesophageal commissure, fc frontal connective, cnn circumesophageal connective, sco subesophageal commissure, an antennal nerve, ac anterior commissure, pc posterior commissure, pcn protocerebral connective, lc longitudinal connective, PC protocerebrum, DC deutocerebrum, TC tritocerebrum, MD mandibular neuromere, MX maxillary neuromere, LB labral neuromere, es esophagus. Scale bar: 25 μm

of Drosophila brain tissue is often used to help deconstruct the complexities of neural circuit development or neurodegeneration [9, 10], in many cases with the aim of understanding human disease pathogenesis (e.g. [11, 12]). The availability of Drosophila antibodies to human homologues can pave the way for follow-up studies in mammals or human tissue [13, 14]. There is a growing supply of Drosophila-specific antibodies available that bind to all manner of cellular proteins, from organelle components to synaptic vesicle markers [15]. Immunostaining can be done in basically two ways, which are defined by way of visualization method. One option visualizes proteins/antigens with secondary antibodies that carry labels, which, upon enzymatic reactions, lead to precipitates that are visible under the light microscope. Typical examples are alkaline phosphatase or horseradish peroxidase reactions that lead to brown, black, or purple precipitates (for details and references, see [16]). These histochemical staining methods have obvious limitations: enzymatic reactions do not penetrate well into tissue and resulting precipitates do not

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allow 3D reconstructions unless the tissue is embedded in plastic, micro-dissected, and subsequently scanned, which is laborious and time-consuming. The advent of laser confocal and super-resolution microscopy together with computer algorithms nowadays allows rapid and reproducible 3D reconstructions of optical sections derived from fluorescence immunostaining, which has thus become the method of choice. It is based on secondary antibodies that are conjugated with various fluorochromes that, following excitation, emit at specific wavelengths that are detected by the various optical imaging techniques. Our protocol outlines the basics for this method. First the tissue must be carefully dissected from the whole organism and fixed; the fixation process described here uses paraformaldehyde; however, other fixation processes may be used depending on the antibody used or the structures being visualized (see Subheading 1.1). In the case of the embryo, the two protective membranes, an outer shell called the chorion and a thick inner vitelline membrane [17], must first be removed. This vitelline barrier becomes permeable to the fixative agent when treated with a fixative–heptane mix. The membrane is then removed by methanol to allow the diffusion of the antibodies. Following fixation, the tissue is blocked, usually with normal serum. This step reduces binding of the antibody to nonspecific reactive sites of Drosophila proteins. Blocking with normal serum from the species used to generate the secondary antibodies is preferred. Either monoclonal or polyclonal primary antibodies can be used for immunohistological staining of Drosophila tissue. Antibodies are raised against specific antigens and their most effective concentration should be determined by first carrying out a dilution series on embryos. Occasionally, primary antibodies are already conjugated to a fluorophore meaning that secondary antibodies are not needed to visualize the antigen’s location. When choosing fluorophores for multiple antibody stainings, you should bear in mind what filters/lasers are available for imaging the brains, and any cross-over in the excitation or emission wavelengths [18]. Separating each incubation step is a series of washes. Thorough washing is essential to prevent antibodies interacting with surplus fixative and to clear the sample of residual antibodies. This immunohistological protocol can also be applied to adult brain tissue as well as to larval imaginal discs. 1.1  Trouble Shooting

Although now a relatively standard technique, elements of this protocol may benefit from tweaking, depending on what you are hoping to visualize and the antibodies you use. If you have persistent trouble in visualizing your proteins, you may want to try the following suggestions.

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1.1.1  Alter the Fixation Technique

Formaldehyde is suitable for deep penetration of the tissue; it forms strong cross-links between proteins and is suitable for long-­ term storage [19] and preserving chromosome morphology [20]. You may want to consider replacing the paraformaldehyde with freshly prepared formaldehyde solution. Gluteraldehyde may also be used [21, 22]; however, as it is a larger molecule it does not diffuse as well through deep tissue. Its cross-links span a larger distance so it can stably fix proteins that are further apart. Gluteraldehyde fixation is not ideal for immunohistochemistry as further treatment of the tissue is necessary to avoid aldehyde groups binding to the antibody [23]. Embryos may be fixed with methanol; while suitable for studying embryo morphology, this method shows poor preservation of cytoplasmic proteins [24]. A more complex technique is cryo-fixation [25]. With all these techniques, inadequate or over-fixation may occur so it may be useful to slightly increase or decrease the fixation period.

1.1.2  Antibody Optimization

Too high or low concentrations of antibody may either cause antibody aggregation on the surface of the tissue or poor penetration of the tissue [26]; therefore, carrying out a dilution series will help to determine the most effective titer. To improve the signal –to-­ noise ratio try using affinity purified antibodies or altering the temperature, e.g., room temperature instead of 4 °C, or length of antibody incubation, e.g., 3 h instead of overnight [22].

2  Materials Make all stock buffers in a sterile, sealable bottle. Prepare solutions with distilled H2O (dH2O). To avoid contamination of the stock solutions, keep individual 50 ml aliquots of PBS, PBL, and PBT and use these for dissections. These solutions are prone to contamination, check for wispy or cloudy cultures that may form after around 1 month of storage and dispose of contaminated solutions. All solutions should be stored at 4 °C unless stated otherwise. 2.1  Embryo Preparations

1. PEM—100 mM PIPES, 2 mM EGTA, 1 mM MgSO4. To 800 ml dH2O add 34.63 g PIPES, 0.76 g EGTA and 0.12 g MgSO4. Adjust to pH 7 with HCl. Make up to final volume of 1 l with dH2O. 2. PEM-FA: Add 1 ml 37% formaldehyde solution to 9 ml PEM. Make fresh. Do not store. 3. PBT 0.1%—for embryo and early larval stages (L1 and L2): Add 0.2 g BSA and 0.2 ml Triton X-100 to 200 ml PBS. Store at 4 °C (see Note 1). 4. 5% PBT-NGS: Add 1 ml normal goat serum (Invitrogen) to 19 ml PBT 0.1%.

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5. 50% sodium hypochlorite. 6. Heptane. 7. Methanol. 2.2  Larval Preparations

1. PBL: Dissolve 1.8 g of Lysine HCl in 50 ml dH2O, adjust pH to 7.4 by adding X ml of 0.1 M NaH2PO4 (see Note 2), adjust volume to 100 ml 0.1 M PBS. Store at 4 °C. 2. PBS 0.1 M buffer: Make 0.1 M Na2HPO by dissolving 8.52 g in 600 ml dH2O. Make 0.1 M NaH2PO4 by dissolving 2.4 g in 200 ml dH2O. Take 500 ml of 0.1 M Na2HPO and adjust pH to 7.4 by adding X ml of 0.1 M NaH2PO4 (see Note 2). Store at 4 °C. 3. PBT 0.5%—late larval stage (L3): Add 1 ml Triton X-100 to 200 ml PBS. Store at 4 °C (see Note 1). 4. 10% PBT-NGS: Add 1 ml normal goat serum (Invitrogen) to 9 ml PBT 0.5%. 5. 8% PFA: Caution—make in a fume hood. In a sealable 50 ml container dissolve 1.6 g paraformaldehyde in 20 ml dH2O, add 140 μl 1 M NaOH. Place in 37 °C water bath, vortexing occasionally until completely dissolved. Aliquot 400 μl into 2 ml tubes and store at −20 °C (see Note 3). 6. PLP: Mix 3 parts PBL (1.2 ml) to 1 part 8% PFA (400 μl). Make fresh. Do not store (see Note 3).

2.3  Mounting Medium and Antibodies

1. VECTASHIELD mounting medium with or without DAPI (see Note 4) (Vector Laboratories, Burlingame, CA).

2.4  Equipment

1. Microscope (larval preps).

2. Antibodies—Developmental Studies Hybridoma Bank supplies a large-range of Drosophila-specific primary antibodies. Secondary antibodies should be selected based on the animal in which the primary antibody was raised, i.e., if you are using rat-Elav, then you will need something like a goat anti-rat 488 secondary antibody.

2. Nylon mesh (embryo preps). 3. Vertical rotator. 4. Glass pipette (embryo preps). 5. Small paint brush (embryo preps). 6. Rocker. 7. Fine tip forceps and sharpening stone (Dumont no. 5 tweezers, super fine, straight tip) (larval preps). 8. Glass microscope slides, SuperFrost (Thermo Scientific Gerhard Menzel).

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9. Large cover slips: 22 × 50 mm, 0.13–0.17 thickness (embryo preps). 10. Small cover slips: 22 × 22 mm, 0.13–0.17 thickness (larval preps). 11. Silicone grease or petroleum jelly (larval preps). 12. Modeling clay (such as Plasticine) (embryo preps). 13. Dissection watch glass (larval preps). 14. Pin holder and stainless steel minutien pins 0.2 mm diameter (InterFocus Ltd., 15. Fine Science Tools) (larval preps). 15. Timer.

3  Methods To be carried out at room temperature unless otherwise stated. 3.1  Embryo Fixation

1. Recover plates from egg collection and remove any remaining yeast or dead flies with a spatula or brush being careful not to damage the agar (see Note 5). 2. Dechorionate the embryos by adding 50% hypochlorite and agitate for 2–5 min until dechorionated embryos float to the surface (see Note 6). 3. Pour the embryos and hypochlorite through the nylon mesh and rinse thoroughly with dH2O (see Note 7) (see Fig. 2). 4. Transfer embryos to a 2 ml tube containing 1 ml heptane and 1 ml PEM-FA (see Note 8). 5. Agitate on a rotator at high speed for 10–30 min (no longer than 30 min). 6. Allow embryos to settle—they should be at the interface between the two phases. 7. Remove the lower phase, then the upper phase (see Note 9). 8. Replace with another 1 ml heptane. Then add 1 ml 100% methanol and shake vigorously for 1–2 min. The devitellinized embryos will fall to the bottom of the tube. 9. Remove both phases making sure to remove all vitelline debris (see Note 10). 10. Quickly add 1.5 ml of 100% methanol and agitate on rotator for 2 × 5 min, then 1 × 30 min (see Note 11). 11. Replace with fresh methanol and store at −20 °C or rehydrate for immunostaining.

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Fig. 2 Embryo collection apparatus. An open-bottom container (a) has a hole cut in the lid (b); the hole is closed by a fine mesh (c), which is included into the lid when screwed on top of the container. A 50  ml plastic container can also be adapted in this way 3.2  Embryo Immunostaining

1. Transfer embryos into a 1.5 ml tube (see Note 12). 2. Rehydrate the embryos by removing the methanol and washing 2 × 5 min, then 1 × 30 min with PBT 0.1%. 3. Incubate embryos for 30 min in 5% PBT-NGS. 4. Remove PBT-NGS and add appropriate amount of primary antibody diluted in 5% PBT-NGS to a reaction volume of 100 μl (see Note 13). Incubate overnight at 4 °C. 5. Remove antibody solution and wash 1 × 1 min, 3 × 5 min and 4 × 30 min with PBT 0.1% (see Note 11). 6. Incubate embryos for 30 min in 5% PBT-NGS. 7. Remove PBT-NGS and add appropriate amount of secondary antibody diluted in 5% PBT-NGS to a reaction volume of 100 μl. Incubate at 4 °C overnight in the dark (see Note 14). 8. Remove antibody solution and wash 3 × 5 min and 4 × 30 min with PBT 0.1% (see Note 11). 9. Add 1 drop of VECTASHIELD mounting medium and incubate at 4 °C overnight in the dark. 10. Mount on glass slide with 22 × 50 mm cover slip (see Note 15). 11. Your samples are now ready for image acquisition (see Note 16).

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1. Collect larvae and place into a watch glass containing cold PBS. 2. Fill a 0.5 ml tube with cold PBS and keep on ice with the lid open. 3. Roughly remove the larval CNS (see Note 17), placing dissected brains into the PBS-containing 0.5 ml tube. 4. After 30 min of dissection, remove PBS and add 500 μl of PLP. Agitate on a rotator for 1 hour at room temperature. 5. Remove PLP and wash 3 × 10 min in PBT (see Note 18). 6. Block by incubating the brains in 5% PBT-NGS (for L1 and L2) or 10% PBT-NGS (for L3) on the rotator for 15 min. 7. Remove PBT-NGS and add appropriate amount of primary antibody diluted in 5 or 10% PBT-NGS to a reaction volume of 100 μl (see Note 13). Incubate overnight (unless alternative duration is specified by antibody manufacturer) at 4 °C. 8. Remove antibody solution and wash 1 × 1 min, then 3 × 20 min in PBT. 9. Add appropriate amount of secondary antibody diluted in 5 or 10% PBT-NGS to a reaction volume of 200 μl. Incubate for 2–3 h on a rocker in the dark (see Note 14). 10. Wash brains 2 × 15 min in PBT, then 2 × 15 min in PBS (see Note 19). 11. Add ~2 drops of VECTASHIELD mounting medium and incubate overnight at 4 °C in the dark. 12. Fine dissect in mounting medium on microscope slide under microscope (see Note 20). Remove debris, arrange brains, and make a small dab of silicone grease or petroleum jelly at four corners where the cover slip will go. Gently lower the 22 × 22 mm cover slip in to place and seal with nail varnish (see Note 21). 13. Store in a microscope slide box in the dark at 4 °C (see Note 22). 14. Your samples are now ready for image acquisition (see Note 16). Examples of successful fixation, labeling, and imaging under a standard fluorescence microscope are given in Fig. 3.

4  Notes 1. Triton X-100 is very viscous, so pipette slowly to make sure you take up the right amount. Mix by using a slow setting on a magnetic stirrer to avoid an abundance of bubbles.

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Fig. 3 Immunostaining of whole mount embryonic CNS and third instar larval CNS. Top row: The embryonic nervous system is staining with Horse Radish Peroxidase Cy3 (HRP-Cy3; red) and visualized under a standard fluorescence microscope. The anterior brain is visible (arrow) as well as the ladder-like axon tracts of the ventral nerve cord (arrowhead); compare to Fig.  1. The nuclei of all embryonic cells are visualized with DAPI. Merge shows co-labeling of HRP-cy3 and DAPI. Bottom row: third instar larval CNS immunostained with anti-­Bruchpilot counterlabeled with goat anti-mouse-568, and nuclei are highlighted with DAPI; merge shows co-labeling of nc82 and DAPI.  The nc82 antibody recognizes the Bruchpilot protein which is specifically enriched in the active zone of synapses

2. Place the solution on a magnetic stirrer while measuring the pH and use a 10 ml pipette to add small amounts of NaH2PO4 to the solution. 3. Before you start the dissections, place a frozen aliquot of PFA in a 37 °C water bath. By the time you have finished the dissection and are ready to fix the tissue, the PFA should be fully dissolved and you can add the PBL directly to it. 4. DAPI (4,6-diamindino-2-phenylindole) is a nuclear stain that binds to double stranded DNA and fluoresces when excited with a mercury arc lamp or UV. Even when not examining the nuclei, it is useful to use the consistent DAPI staining pattern to help orientate around the embryonic and larval brains. 5. Embryos can be collected by placing fruit agar plates (12.5 g sugar, 21.25 g agar, 750 ml dH2O, autoclave in large flask, add 250 ml apple juice, 2 g nipagen, stir and pour into 55 mm Petri dishes, allow to cool, store at 4 °C for 1 month), that have a smear of yeast paste in the middle, on top of a bottle of

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flies. Keep the bottle, which should have holes pierced in the bottom for air circulation, inverted in the dark and replace the agar plate as necessary. 6. You can use 50% household bleach instead of hypochlorite but make sure that it hasn’t gone off—yellow bleach is still in date. 7. Wash agar plates with water and pour this through the mesh × 2. Hold the nylon mesh over a waste beaker and use a water bottle to rinse thoroughly. You can test for remaining bleach by dabbing it on a colored paper towels (e.g., blue roll), if any bleach is present it will change the paper color. Residual bleach will interfere with the fixation process. If you plan on doing a lot of embryo stainings, it is worth trying to make a straining tool out of a small glass jar with a screw lid and the bottom removed (see Fig. 2). 8. Hold the mesh with forceps and dip into the 2 ml tube to transfer the embryos, use the brush to transfer any embryos stuck to side of straining tool (if you have one). 9. Use a glass pipette as embryos are less likely to adhere to glass than to a plastic pipette. Remove as much of the solutions as possible—this may mean pipetting off some of your embryos too. 10. Remove the layers by pipetting from the interface of the two phases. 11. For each wash, remove the old solution and add the same amount of fresh solution. Washes should be agitated on a rotator at full speed. 12. Pipette as fast as you can, as soon as the liquid stops being turbulent, the embryos tend to stick to the sides of the pipette and are very difficult to remove. This means you should have your two tubes ready, with lids open and suck up the liquid and embryos and transfer as quickly as possible. 13. Depending on what you are attempting to visualize, you can add several primary antibodies at once. Take care to avoid using similar species that may cross react, for example, if you use a mouse and rat primary antibody, the secondary anti-­ mouse will bind to the rat antibody and the secondary anti-rat will bind to the mouse antibody. If you are visualizing an endogenous GFP signal (e.g., using the GAL4>UAS system), you may get a better signal by also using an antibody against GFP and a secondary with a 488 (green) fluorophore. Some primary antibodies are already conjugated to a fluorophore and do not need a secondary antibody, such as Horse Radish Peroxidase Cy3 (HRP-Cy3), which specifically labels the Drosophila nervous system.

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14. You will need a secondary antibody that targets the animal species in which the primary antibody was raised. You can use up to four separate channels by picking fluorophores that are unlikely to interfere with each other, e.g., Dapi—UV, 488— green, 568—red, 647—far red. Of course, the fluorophore you use will depend on what filters, lamps or lasers are available for image capture. 15. Suck up VECTASHIELD and embryos and pipette out in a wide zig-zag on the labeled microscope slide (see Fig. 4a). Roll 4 small balls of modeling clay and place in the corners where the cover slip will go. Lower one short end of the cover slip in place and gently lower the cover slip, avoiding air bubbles. Gently press cover slip at the four corners. 16. To avoid bleaching the brains, first use a fluorescent microscope to select your best brains or embryos. When imaging using a laser scanning microscope scan each channel sequentially to avoid interference between fluorophores. If you have issues with the noise–signal ratio check that you are using the antibody at the appropriate dilution, ensure you are completing all wash steps and check the pH of the buffer solutions.

Fig. 4 Cartoon illustrating slide preparation. (a) Pipette embryos in VECTASHIELD mounting medium in a wide zig-zag line onto labeled microscope slide, place small balls of modeling clay where the corners of the cover slip will be and gently lower the cover slip (arrow) on to the embryos in VECTASHIELD; make sure to avoid air inclusions. (b) Arrange the larval brains/CNS’ in a rosette onto labeled microscope slide, place small dabs of silicone grease (or petroleum jelly) at the corners of where the cover slip will be, and gently lower the cover slip (arrow); make sure to avoid air inclusions

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17. This bit takes practice! There are several ways to pull the CNS from the larvae but the thing that will help the most is being able to differentiate the CNS from the imaginal discs and fat bodies (see Figs. 5 and 6). I find it easier to gently clamp the body of the larvae about two thirds the way towards the mouth hooks, then when the larvae extends its mouth hooks, grab them with the forceps (see Fig. 5). Gently pull the mouth

Fig. 5 Cartoon illustrating a rough larval CNS dissection (1). First grab the extended mouth hooks and make a gentle tear in the cuticle (2). Rip the larvae in half (3). Holding the mouth hooks again, slowly pull off the remaining cuticle until (4) you are left with the larval brain still attached to the head, imaginal discs and fat bodies

Fig. 6 Identifying the third instar larval brain and CNS. (a) Several imaginal discs (filled arrow), fat bodies (arrowhead) and digestive components (empty arrow) will be attached to the third instar larval brain and CNS. Identifying the brain among these other tissues will aid successful dissections. The larval brain and CNS are outlined with a black line. (b) DAPI labeled third instar larval brain and CNS visualized under the fluorescence microscope after successful dissection; note that wing and leg imaginal discs are still attached

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hooks until the cuticle rips and the internal matter is exposed. The round optic lobes of the CNS may be visible. Place both forceps on the body and rip the lower half of the larvae off. Grab the mouth hooks and body again and gently pull until the mouth hooks and CNS come away from the rest of the larvae. Holding on to the mouth hooks, transfer to the 0.5 ml tube that is sat on ice. If you do not have much luck this way, or find that the ventral nerve cord is lost using this method, you can also pinch both forceps just below the mouth hooks (by the scruff of its neck!) and rip the cuticle open then bit by bit remove the lower part of the larvae, the digestive tract, fat and some imaginal discs. 18. When washing it is best to leave brains to settle for 15 s then pipetting off the solution. Eject the pipetted solution into a clean watch glass, check under the microscope and rescue any brains by pipetting them back into the tube. If carrying out several dissections, you can leave samples washing for an extended period of time until all preparations have reached the same stage. 19. As the secondary antibodies are photosensitive you should keep your preparation in the dark as much as possible to avoid bleaching the fluorophores. Cover the tubes with tin foil or re-purpose a small container to hold your tubes and attach to the rotator using Velcro strips. 20. Cut around 2 cm off the end of a P200 pipette tip and transfer brains and VECTASHIELD to a microscope slide. Label the microscope slide with a dissection code (for cross-reference with your records e.g., Initials_01), genotype, the primary antibody and secondary fluorophore used and the date. Remove the majority of the VECTASHIELD, so that the brains are no longer floating, and pipette back into your ­dissection tube. Use the pin holder tools to remove unwanted imaginal discs, fat bodies, and mouth hooks. You can use them like a knife and fork to slice away tissue, or pin an imaginal disc into place with one needle and cut the connecting tissue with the other (see Fig. 7). Take care not to pin or damage the CNS itself. 21. Arrange brains in concentric circles with the ventral nerve cord pointing outwards (see Fig. 4b). Orientate the brains in a mixture of dorsal or ventral side up. Add a small amount of VECTASHIELD in a circle around the brains—not too much or they will float out of position. Once the cover slip is in place, gently press on the four corners to flatten the brains slightly, be careful as they are easily ruptured by squashing. Add more VECTASHIELD if necessary by pipetting small amounts at the edge of the cover slip. Remove excess

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Fig. 7 Cartoon illustrating fine dissection of the larval brain and CNS (1). Using the dissecting pins like a knife and fork, remove the mouth hooks from the brain, then (2) gently remove all other tissue taking care (3) not to damage or puncture the brain

VECTASHIELD with cotton buds or tissues dipped in ethanol—again, be very careful not to dislodge the cover slip as this will destroy your brains! 22. A standard 4 °C fridge is a good place to store your slide box. Cold rooms often have a damp atmosphere, which can cause the nail varnish to peel off.

Acknowledgments This work was supported by the UK Medical Research Council (G0701498; MR/L010666/1), the Biotechnology and Biological Sciences Research Council (BB/N001230/1), the MND Association (Hirth/Nov15/914-793; Hirth/Oct13/6202; Hirth/Mar12/6085; Hirth/Oct07/6233), and Alzheimer’s Research UK (Hirth/ARUK/2012) to F.H. References 1. Skeath JB, Thor S (2003) Genetic control of Drosophila nerve cord development. Curr Opin Neurobiol 13:8–15 2. Homem CC, Repic M, Knoblich JA (2015) Proliferation control in neural stem and progenitor cells. Nat Rev Neurosci 16:647–659 3. Urbach R, Technau GM (2004) Neuroblast formation and patterning during early brain development in Drosophila. BioEssays 26:739–751 4. Hirth F, Reichert H (1999) Conserved genetic programs in insect and mammalian brain development. BioEssays 21:677–684 5. Venken KJ, Simpson JH, Bellen HJ (2011) Genetic manipulation of genes and cells in the nervous system of the fruit fly. Neuron 72:202–230

6. Shaw RE, Kottler B, Ludlow ZN, Buhl E, Kim D, Morais da Silva S, Miedzik A, Coum A, Hodge JJ, Hirth F, Sousa-Nunes R (2018) In vivo expansion of functionally integrated GABAergic interneurons by targeted increase in neural progenitors. EMBO J 37:e98163 7. Diaper DC, Adachi Y, Lazarou L, Greenstein M, Simoes FA, Di Domenico A, Solomon DA, Lowe S, Alsubaie R, Cheng D, Buckley S, Humphrey DM, Shaw CE, Hirth F (2013) Drosophila TDP-43 dysfunction in glia and muscle cells cause cytological and behavioural phenotypes that characterize ALS and FTLD. Hum Mol Genet 22:3883–3893 8. White KE, Humphrey DM, Hirth F (2010) The dopaminergic system in the aging brain of Drosophila. Front Neurosci 4:205

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9. Muqit MK, Feany MB (2002) Modelling neurodegenerative diseases in Drosophila: a fruitful approach? Nat Rev Neurosci 3:237–243 10. Koizumi K, Higashida H, Yoo S et  al (2007) RNA interference screen to identify genes required for Drosophila embryonic nervous system development. Proc Natl Acad Sci U S A 104:5626–5631 11. Hirth F (2010) Drosophila melanogaster in the study of human neurodegeneration. CNS Neurol Disord Drug Targets 9:504–523 12. Li T, Bellen HJ, Groves AK (2018) Using Drosophila to study mechanisms of hereditary hearing loss. Dis Model Mech 11:dmm031492 13. Tsuji T, Higashida C, Yoshida Y et  al (2011) Ect2, an ortholog of Drosophila’s pebble, negatively regulates neurite outgrowth in neuroblastoma × glioma hybrid NG108-15 cells. Cell Mol Neurobiol 31:663–668 14. Venderova K, Kabbach G, Abdel-Messih E, Zhang Y, Parks RJ, Imai Y, Gehrke S, Ngsee J, Lavoie MJ, Slack RS, Rao Y, Zhang Z, Lu B, Haque ME, Park DS (2009) Leucine-Rich Repeat Kinase 2 interacts with Parkin, DJ-1 and PINK-1  in a Drosophila melanogaster model of Parkinson’s disease. Hum Mol Genet 18:4390–4404 15. Developmental Studies Hybridoma Bank developed under the auspices of the NICHD and maintained by The University of Iowa, Department of Biology, Iowa City, IA 52242. http://dshb.biology.uiowa.edu/ 16. Patel N (1994) Imaging neuronal subsets and other cell types in whole mount Drosophila emrbyos and larvae using antibody probes. In: Goldstein LSB, Fryberg E (eds) Methods in cell biology, vol 44. Drosophila melanogaster: practical uses in cell biology. Academic, New York. For an amended and updated version, follow the link: http://patelweb.berkeley.edu/Images/Protocols/pdf%20files/ Antibody%20Methods%202006.pdf 17. Ashburner M (1989) Drosophila: a laboratory manual. Cold Spring Harbor Laboratory Press, New York

18. Hoffman, G. (2008) Seeing is believing: use of antibodies in immunohistochemistry and in situ hybridization. In: Short course II of SfN’s 38 annual meeting: 15–19 November 2008. Society for Neuroscience, Washington, DC 19. Rothwell WF, Sullivan W (2000) Fluorescent analysis of Drosophila embryos. In: Sullivan W, Ashburner M, Hawley RS (eds) Drosophila protocols. Cold Spring Harbor Laboratory Press, New York, p 141 20. Bonaccorsi S, Giansanti MG, Cenci G, Gatti M (2012) Formaldehyde fixation of Drosophila testes. Cold Spring Harb Protoc:10.1101 21. Heimbeck G, Bugnon V, Gendre N, Häberlin C, Stocker RF (1999) Smell and taste perception in Drosophila melanogaster larva: toxin expression studies in chemosensory neurons. J Neurosci 19:6599–6609 22. Stocker RF, Heimbeck G, Gendre N, de Belle JS (1997) Neuroblast ablation in Drosophila P[GAL4] lines reveals origins of olfactory interneurons. J  Neurobiol 32:443–456 23. Hassell J, Hand AR (1974) Tissue fixation with diimidoesters as an alternative to aldehydes. I.  Comparison of cross-linking and ultrastructure obtained with dimethylsuberimidate and glutaraldehyde. J  Histochem Cytochem 22:223–229 24. Wieschaus E, Nüsslein-Volhard C (1998) Looking at embryos. In: Roberts DB (ed) Drosophila, a practical approach. Oxford University Press, New York, p 205 25. Ripper D, Schwarz H, Stierhof YD (2008) Cryo-section immunolabelling of difficult to preserve specimens: advantages of cryofixation, freeze-substitution and rehydration. Biol Cell 100:109–123 26. Rebay I, Fehon R (2000) Generating antibodies against Drosophila proteins. In: Sullivan W, Ashburner M, Hawley RS (eds) Drosophila protocols. Cold Spring Harbor Laboratory Press, New York, p 400

Chapter 6 Nonfluorescent RNA In Situ Hybridization Combined with Antibody Staining to Visualize Multiple Gene Expression Patterns in the Embryonic Brain of Drosophila David Jussen and Rolf Urbach Abstract In Drosophila, the brain arises from about 100 neural stem cells (called neuroblasts) per hemisphere which originate from the neuroectoderm. Products of developmental control genes are expressed in spatially restricted domains in the neuroectoderm and provide positional cues that determine the formation and identity of neuroblasts. Here, we present a protocol for nonfluorescent double in situ hybridization combined with antibody staining which allows the simultaneous representation of gene expression patterns in Drosophila embryos in up to three different colors. Such visible multiple stainings are especially useful to analyze the expression and regulatory interactions of developmental control genes during early embryonic brain development. We also provide protocols for wholemount and flat preparations of Drosophila embryos, which allow a more detailed analysis of gene expression patterns in relation to the cellular context of the early brain (and facilitate the identification of individual brain neuroblasts) using conventional light microscopy. Key words Drosophila, Embryonic brain, Neuroectoderm, Neuroblast identification, In situ hybridization, Antibody staining

1  Introduction In situ hybridization and immunohistochemistry are fundamental and widely used methods in the field of developmental biology. They serve multiple purposes, such as the identification of a specific cell type or tissue by the visualization of marker genes, or the study of genetic interactions by the comparative analysis of gene expression patterns in different genetic backgrounds. Huge progress has been made in that field in the last decades. The invention of fluorescence imaging techniques allows the detection of gene products in deeper layers of tissue, at consistently increasing resolutions. This also led to the development of staining protocols involving different fluorescent dyes, which allow the detection of multiple gene products at once [1]. However, fluorescence ­imaging methods have Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_6, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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limitations. Those concern, for example, the relatively low stability of fluorescence dyes hampering the long-­term storage of specimens, or the need of costly documentation systems. Here, we present a protocol for nonfluorescent double in situ hybridization combined with antibody staining on Drosophila wholemount embryos, which allows the simultaneous visualization of transcripts and proteins in up to three different colors. This method combines the benefits of simultaneously labeling multiple factors at once, like it is popular in fluorescence microscopy, while allowing the easy handling of traditional, nonfluorescent labeling techniques. That way, a high amount of specimens can be analyzed under a conventional light microscope, without any need of fluorescent detection systems. Using differential interference contrast (i.e. Nomarski), histochemically labeled cells are visualized in context of the surrounding (uncolored) tissue which often simplifies interpretation of gene expression patterns and their relation to specific cells without the need of additional markers (as e.g. membrane markers used in fluorescence microscopy to visualize cell boundaries). This protocol is based on the sequential detection of differentially labeled RNA probes via the alkaline phosphatase (AP)catalyzed turnover of different chromogenic compounds. The precipitates formed in those reactions are insoluble in aqueous solution, which allows the elution of AP-bound antibodies after the chromogenic reaction without affecting the staining per se. By that, the AP-coupled antibody directed against the first probe can be eluted after the first chromogenic reaction, with the subsequent introduction of an AP-coupled antibody directed against the second probe. Here, we use NBT/BCIP and Vector Red as chromogenic compounds to sequentially detect DIG- and FITC-labeled probes, in combination with a subsequent antibody staining which is visualized by the peroxidase-catalyzed turnover of DAB. That way, three different gene products are visualized by blue, red, and brown precipitates, respectively. Such stainings are particularly useful to analyze the expression and regulatory interactions of developmental control genes during the early period of embryonic brain development. In organisms as diverse as insects and mammals, the brain arises from multipotent neural stem cells (in insects called neuroblasts) which originate from the neuroectoderm. Products of developmental control genes are expressed in spatially restricted domains in the neuroectoderm and provide positional cues that regulate the development of neuroblasts [2, 3]. Therefore, the expression of these genes has to be precisely controlled. Recently, we uncovered a novel gene regulatory network in Drosophila, in which ‘DorsoVentral’ and ‘AnterioPosterior’ patterning genes interact to precisely control their spatially restricted expression in the brain neuroectoderm, which is essential for the correct formation and fate specification of

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neuroblasts [4–7]. These patterning genes display not only a high degree of intersegmental modulation, but also considerable changes in their expression patterns over time. The staining technique presented here has been used to visualize multiple patterning genes at once in order to analyze their dynamic expression patterns in relation to each other, and to uncover their genetic interactions and cellular functions. Nonfluorescent multiple stainings are also suitable to identify embryonic brain neuroblasts. The brain of Drosophila is built up by about 100 neuroblasts, which develop in a stereotypic pattern during the first third of embryonic development [8]. Brain neuroblasts can be identified individually by position, time point of formation, morphology, and the specific combination of marker genes they express [9]. As the identified marker genes are usually not selective enough, it is often necessary to stain for a combination of those in order to identify individual brain neuroblasts. Using this protocol, marker gene expression can be analyzed using conventional Nomarski optics. That way, labeled as well as unlabeled neuroblasts are visible alongside each other, which allows the identification of individual neuroblasts with the use of only a few molecular markers.

2  Materials 2.1  Reagents

1. Anti-digoxigenin (DIG)-AP, Fab fragments from sheep (Roche Applied Science). 2. Anti-fluorescein (FITC)-AP, Fab fragments from sheep (Roche Applied Science). 3. Methanol (≥99.9%, p.a.). 4. ssDNA (10 mg/μl, MB grade, from fish sperm) (Roche Applied Science).

2.2  Buffers

1. AP buffer (50 ml): 41.2 ddH2O, 1 ml 5 M NaCl, 2.5 ml 1 M MgCl2, 5 ml 1 M Tris–HCl. Adjust pH to 9.5. Add 50 μl Tween®20. 2. Blocking buffer (100 ml): 10 ml 1 M Tris–HCl, 3 ml 5 M NaCl. Adjust pH to 7.5. Add 0.5 g blocking reagent (delivered with TSA Biotin System, Perkin Elmer). Store aliquots at −20 °C. 3. Glycine buffer (50 ml): 50 ml ddH2O, 0.038 g glycine, 1.46 g NaCl. Adjust pH to 2.3. Add 0.05 g BSA and 50 μl Triton™ X-100. 4. Hybridization buffer (50 ml): 12.5 ml SSC (20×), 12.5 ml DEPC-H2O, 25 ml Formamide, 50 μl Tween®20. 5. 20× PBS (500 ml stock solution): 500 ml ddH2O, 75.97 g NaCl, 9.94 g Na2HPO4, 4.14 g NaH2PO4. Adjust pH to 7.4. Dilute with ddH2O to obtain 1× working solution.

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6. 0.3%, PBT: for 50 ml mix 47.5 ml ddH2O, 2.5 ml PBS (20×), 150 μl Triton™ X-100. 7. 0.1% PBTween: for 50 ml mix 2.5 ml PBS (20×), 47.5 ml ddH2O, 50 μl Tween®20 (use DEPC-treated PBS and ddH2O for PBTween-DEPC). 8. Vector Red buffer: for 100 ml mix 100 ml distilled ddH2O, 1.21 g Tris–HCl. Adjust pH to 8.2–8.5. Add 100 μl Tween®20. 9. Washing buffer: for 100 ml mix 10 ml 1 M Tris–HCl, 7.5 ml 2 M NaCl, 82.5 ml ddH2O. Adjust pH to 7.5. Add 50 μl Tween®20. 2.3  Solutions and Media

1. ABC solution (avidin-biotinylated peroxidase-complex): 1 ml PBT, 4 μl solution A, 4 μl solution B (both are delivered with the Vectastain ABC Kit). Prepare solution 1 h before use and keep on a shaker (100 rpm) at room temperature (RT). 2. AP staining solution: 1 ml AP-Buffer, 3 μl NBT stock solution, 1.5 μl BCIP stock solution. Prepare freshly before use. 3. Apple juice agar: 1000 ml apple (or grape) juice, 28 g agar. Heat until the apple juice agar solution starts boiling and becomes clear. Dispense in fly culture vials or petri dishes (the latter requires additional fly cages). Once the agar has cooled and hardened, store at −4 °C. 4. BCIP stock solution: 10 ml dimethylformamide (100%), 500 mg BCIP. Store 1 ml aliquots at −20 °C. 5. Chlorine bleach (6%): Dilute sodium hypochlorite solution (12% Cl) 1:1 with H2O. 6. 3,3′-Diaminobenzidine tetrahydrochloride (DAB) stock solution: Dissolve 1 DAB tablet (10 mg; Sigma-Aldrich) in 35 ml PBT. Store 400 μl aliquots at −20 °C. 7. DAB staining solution: 400 μl DAB stock solution, 600 μl PBT, 2 μl H2O2. Prepare freshly before use. 8. DEPC-H2O: 1000 ml H2O, 1 ml DEPC. Autoclave before use. 9. Fixative: 450 μl PBT, 70 μl formaldehyde (37%), 600 μl n-heptane. 10. Glycerol (70%/90%). Dilute glycerol with PBS to obtain the particular concentration. 11. NBT stock solution: 10 ml dimethylformamide (70%), 500 mg NBT. Store 1 ml aliquots at −20 °C. 12. Vector Red (VR) staining solution: 1 ml VR buffer, 16 μl VR substrate solution 1, 16 μl VR substrate solution 2, 16 μl VR substrate solution 3 (VR substrate solutions are delivered with the VectorRed alkaline substrate kit I). Vortex ~5 s after addition of each VR substrate solution. Prepare freshly before use.

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1. TSA™ Biotin System (PerkinElmer). 2. Vectastain ABC Kit (Standard) (Vector Labs). 3. VectorRed Alkaline Phosphatase Substrate Kit I (Vector Labs).

2.5  Equipment for Staining, Preparation, and Mounting

1. Aluminum foil (thickness 0.02–0.03 mm). 2. Preparation forceps (e.g. Dumont no. 5, Fine Science Tools). 3. Concavity slide. 4. Cover slips (18 × 18 mm, 22 × 22 mm and 24 × 60 mm). 5. Microscope slides. 6. Minutien pins (stainless steel, 0.1 mm diameter, Fine Science Tools). 7. Nail polish (transparent). 8. 2 Nickel plated pinholders (Fine Science Tools). 9. Nylon mesh (120 μm, e.g. from Merck Millipore). 10. Small scalpel. 11. Spot plate (white). 12. Weighing dishes (white, 41 × 41 mm). 13. Whetstone (e.g. Sharpening stone for Dumont forceps, Fine Science Tools; alternatively abrasive paper, grain size 600–1000).

2.6  Equipment for Microscopy

1. Cold light source (halogen or LED) equipped with fiber light guides (e.g. KL1500 by Schott). 2. Digital microscope camera (e.g. ProgRes® series, Jenoptik). 3. Upright light microscope (equipped with differential interference contrast and 40–100× objectives, e.g. Axioscope Zeiss). 4. Stereo microscope (e.g. MZ series by Leica Microsystems).

3  Methods General remarks: Wash steps and incubations are carried out at RT upon shaking (100 rpm) with 1 ml of solution used, unless stated differently. Reactions are carried out in standard 1.5 ml reaction tubes (eppendorf tubes). 3.1  Embryo Collection and Dechorionization

1. Place flies on apple juice agar for egg laying (see Note 1). 2. Dechorionize embryos after egg laying by covering the apple juice agar with chlorine bleach (6%) for 3 min. Slightly rotate once in a while. Dechorionized embryos will float up. 3. Collect embryos by transferring the chlorine bleach into an egg basket (see Note 2). 4. Wash embryos with water.

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3.2  Fixation

1. Open the egg basket. Take embryos from the nylon mesh with a small scalpel and transfer them into an eppendorf tube with fixative. 2. Fix embryos by vigorously shaking for 20 min (900 rpm). 3. Let the two phases of the fixative separate. 4. Remove the lower phase of the fixative without removing too many embryos. 5. Add 500 μl methanol. 6. Remove the vitelline membrane by vortexing at maximum speed for 2 min. 7. Let the devitellinized embryos sink down and remove lower phase of the solution. 8. Add 500 μl methanol. 9. Vortex for 1 min. 10. Let the embryos sink down and remove as much of the solution as possible without removing too many embryos. 11. Rinse 4× with methanol. Embryos may be stored in methanol at −20 °C (see Note 3).

3.3  (Double-) In Situ Hybridization

Use gloves, filtered tips and DEPC-treated PBTween for the following in situ hybridization procedure. 1. Rinse 5× with PBTween. 2. Incubate 5 min in PBTween/hybridization buffer (1:1). 3. Incubate 5 min in hybridization buffer. 4. Perform prehybridization by incubating 1 h in hybridization buffer + ssDNA (1:100) at 55 °C upon shaking (300 rpm). 5. Perform hybridization by incubating in hybridization buffer + ssDNA (1:100) containing your probe (both of your probes for double in situ hybridization) at the appropriate working dilution(s) (see Note 4). 6. Incubate overnight at 55 °C upon shaking (300 rpm). 7. Incubate 30 min in hybridization buffer at 65 °C upon shaking (350 rpm). 8. Incubate 30 min in PBTween/hybridization buffer (1:1) upon shaking (350 rpm). 9. Wash 4× 20 min with PBTween at 65 °C upon shaking (350 rpm). Standard tips and untreated PBTween may be used from now on. 10. Wash 10 min with PBTween at RT.

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1. Incubate 1.5 h in anti-DIG-AP or anti-FITC-AP (1:1000 in PBTween) (see Note 5). (a) The antibody can alternatively be incubated overnight at 4 °C upon slight shaking. 2. Rinse 3× and wash 3× 10 min with PBTween. 3. Incubate 2× 5 min in AP buffer. (a) Prepare NBT/BCIP staining solution during that step. 4. Incubate in AP-staining solution and transfer the solution containing the embryos into a white weighing dish (41 × 41 mm). Keep it cool and dark. 5. Judge the chromogenic reaction from time to time under a stereo microscope (see Note 6). 6. When staining intensity has reached the desired level, transfer the embryos back into the tube (see Note 7). 7. Rinse 3× with PBTween and terminate the reaction by incubating 10 min in methanol. 8. Rinse 3× with PBTween. (a) For double in situ hybridization combined with antibody staining, continue with Subheading 3.5. (b) For single in situ hybridization combined with antibody staining, continue with Subheading 3.7. (c) For double in situ hybridization without further antibody staining, continue with Subheading 3.9.

3.5  Antibody Elution and Inactivation of Residual AP

1. Rinse 1× and incubate 3 × 10 min in glycine Buffer.

3.6  Detection of the Second Probe (via VectorRed)

1. Rinse 3× and wash 3 × 10 min with PBT.

2. Rinse 3× with PBT. 3. Incubate 5 min in PBT/formaldehyde (10:1).

2. Incubate overnight in anti-FITC-AP or anti-DIG-AP (1:1000 in PBT; depending on the choice of the first antibody) at 4 °C. 3. Rinse 3× and wash 3 × 10 min with PBT. 4. Incubate 10 min in Vector Red buffer. (a) Prepare Vector Red staining solution during that step. 5. Incubate in Vector Red staining solution. Transfer the solution containing the embryos into a white weighing dish (41 × 41 mm). Keep it cool and dark. Judge the chromogenic reaction (in this case a red colored staining) from time to time under a stereo microscope (see Note 8). 6. Transfer embryos back into the tube. 7. Rinse 3× and wash 3 × 10 min with PBT. 8. Incubate 5 min in PBT/formaldehyde (10:1) (see Note 9).

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3.7  Incubation with Primary Antibody

1. Rinse and wash 3 × 10 min in PBT. 2. Incubate overnight with primary antibody (at the appropriate working dilution in PBT) at 4 °C (see Notes 10 and 11). Depending on the quality of the antiserum, 2–4 h incubation at RT may be suitable, too. 3. Rinse 3× and wash 3 × 10 min with PBT.

3.8  Detection of Primary Antibody (via DAB)

1. Incubate 2–3 h with Biotin-coupled secondary antibody (1:500 in PBT) RT. 2. Optional: Continue with Subheading 3.10 for Tyramide signal amplification (see Note 12). If not, prepare ABC before the following step. ABC has to incubate for at least 1 h before use upon shaking (100 rpm). 3. Rinse 3× and wash 3 × 10 min with PBT. 4. Incubate 1 h in ABC. 5. Rinse 3× and wash 3 × 10 min with PBT. Prepare DAB-staining solution during the last wash step. 6. Incubate in DAB-staining solution. Transfer the solution containing the embryos into a white weighing dish (41 × 41 mm). Judge the chromogenic reaction under a stereomicroscope. The staining should emerge quickly (~1–15 min). 7. When the intensity of the brown DAB staining has reached the desired level, transfer the embryos back into the tube.

3.9  Final Washing Steps

1. Rinse 3× and wash 10 min with PBT. 2. Rinse 2× with PBS. 3. Store in 70% glycerol.

3.10  Optional: Tyramide Signal Amplification

1. Rinse 1× and wash 1 × 10 min with PBT. 2. Incubate 1 × 5 min and 1 × 30 min in blocking buffer. 3. Incubate 30 min in Streptavidin-HRP (1:500 in blocking buffer). 4. Wash 3 × 10 min with TSA washing buffer. (a) Prepare ABC before the following step. ABC has to incubate for at least 1 h upon shaking (100 rpm) before use (see step 4 of Subheading 3.8). 5. Incubate 5 min in 140 μl TSA-Reagent (1:70 in Amplification Diluent) (see Note 13). 6. Continue with step 3 of Subheading 3.8.

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Two different kinds of needles are needed for the preparation of embryos. One will be used for handling the embryos (i.e. moving in glycerol, transferring onto microscope slide, holding during preparation; termed “handling needle”), while the other is used for cutting the tissue during preparation (termed “cutting needle”) (Fig. 1a). For that, the pinpoint of both needles will be processed differently. 1. Lock the dull end of a minutien pin tightly in a pinholder. 2. Process the tip of the needle under a stereomicroscope. 3. Place the needle horizontally on a solid steel ground (e.g. on the flat end of a forceps). 4. Flatten the tip of the needle by placing the flat end of another forceps on the tip of the needle, slowly streaking away from it with some pressure (see Note 14). 5. Adjust both needles according to Fig. 1a with the aid of a whetting stone or fine abrasive paper (see Note 15).

3.12  Initial Examination of Embryos

1. Transfer embryos into a well of a white spot plate. 2. Examine embryos under a stereo microscope using maximum magnification. 3. Sort embryos with the handling needle according to staining quality, stage and genotype of interest. 4. Examine selected embryos in more detail by making wholemounts (Subheading 3.13) or filet preparations (Subheading 3.14) for microscopy.

3.13  Wholemount Preparation of Embryos

1. Fix two cover slips (22 × 22 mm) side by side on a microscope slide with a drop of nail polish. The distance between both should be approx. 15 mm. 2. Transfer selected embryos between the cover slips with a pipette. 3. Place another cover slip (22 × 22 mm) on the embryos. Make sure that it stays on top of both spacing cover slips. 4. Examine under a microscope (see Note 16; Fig. 2a, d). 5. Seal for documentation and storage by applying nail polish to the edges of the cover slip.

3.14  Filet Preparation of Early to Mid-stage Embryos

The following filet preparation is recommended for embryos until developmental stage 12, and is performed under a stereo microscope (see Note 17). 1. Fill the pit of a concavity slide with 70% glycerol. 2. Transfer the selected embryo into the pit with the handling needle.

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Fig. 1 Filet preparation of Drosophila embryos. (a) The cutting needle has a flattened tip with sharp edges, while the tip of the handling needle is rounded. (b) Main steps performing filet preparation of embryos at different stages. The posterior part of the abdomen is removed by cutting the embryo transversally. Then, the remaining ectoderm is cut along the dorsal midline toward the anterior tip of the embryo to open the head capsule. Finally, the ectoderm has to be flattened. (c–e) Instructions for flat preparations. Black and white dots indicate corresponding positions of the head capsule. Arrowheads indicate how to move the upper (small) cover slip. Arrows indicate the flow of glycerol caused by moving the cover slip. (c) An embryo with opened head capsule is positioned in glycerol between a large (24 × 60 mm) and a small (18 × 18 mm) cover slip (separated by aluminum spacers) with anterior facing forward and ventral facing down. (d) The small cover slip is carefully moved in posterior direction (regarding the body axis of the embryo), which causes the hemispheres to erect. (e) The small cover slip is carefully moved back until the hemispheres are unfurled flatly on the cover slip. a anterior, p posterior, d dorsal, vML ventral midline, pNE procephalic neuroectoderm

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Fig. 2 Nonfluorescent multiple stainings of the head neuroectoderm and brain neuroblasts in wholemounts and flat preparations. (a) Ventral view on the head region of a stage 8 wholemount embryo (anterior is up) stained for Nkx6-mRNA (via NBT/BCIP, blue) and single-minded-protein (via DAB, brown). (b) Schematic representation of the details presented in c/d′. (c) View on the head region (left hemisphere) of a filet prepared stage 6 embryo with almost all neuroectodermal cells in one plane: blue is ind-mRNA (via NBT/BCIP), magenta is mshmRNA (via Vector Red), and brown is Vnd-Protein (via DAB). (d/d′) Early brain neuroblast pattern of a late stage 9 embryo as wholemount (d) and after filet preparation of the same embryo (d′), stained against engrailed (via NBT/BCIP, blue) and svp-LacZ (via DAB, brown). Note that filet preparation allows a more clear view, and thus the identification of individual neuroblasts by marker expression and relative position (as illustrated in d″) according to [8, 9]. a anterior, p posterior, v ventral, d dorsal, CF cephalic furrow, vML ventral midline, hs engrailed head spot, as engrailed antennal stripe, is engrailed intercalary stripe. (d–d″) Adapted from [8]

3. Place the embryo on the lateral side and hold it carefully with the handling needle. 4. Remove the posterior part of the abdomen and cut along the dorsal midline from posterior to anterior with a cutting needle to open the head capsule (Fig. 1b). 5. Spread the hemispheres slightly with both the needles or a Dumont forceps. 6. Remove excess yolk carefully with the cutting needle while holding the embryo with the handling needle. 7. Arrange two spacers (i.e. strips of aluminum foil, approx. 2 × 15 mm) in parallel at a distance of 10–12 mm in the middle of a large cover slip (24 × 60 mm). 8. Add 45 μl glycerol (90%) between the aluminum spacers.

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9. Transfer the prepared embryo with the handling needle into the glycerol drop and push it carefully to the ground, with anterior forward and ventral downward. 10. Place a small cover slip (18 × 18 mm) on the glycerol drop. And wait until the glycerol is spread between the two cover slips (Fig. 1c). 11. Carefully pull the small cover slip (e.g., with a forceps) backwards (i.e., toward yourself). This will cause the hemispheres to erect (Fig. 1d). 12. Push the small cover slip forward (i.e., away from yourself). By that, glycerol will float toward the erected hemispheres causing them to unfurl flatly on the cover slip (Fig. 1e; see Note 18). 13. When the embryo is appropriately flattened, fix the small cover slip by applying a drop of nail polish to every corner. Let the nail polish dry, remove excess glycerol (e.g., with small stripes of tissue paper), and seal the cover slip by applying nail polish to the edges (two times). 14. Clean the cover slip from glycerol and fix it on a microscope slide with small stickers. 15. Under a microscope, the filet preparation can be examined from both sides (with a view from ventral or dorsal) by turning the cover slides including the filet preparation (Fig. 2c, d′; see Note 19). 3.15  CNS Preparation of Stage 17 Embryos

1. Fill the pit of a concavity slide with 70% glycerol. 2. Transfer the selected embryo into the pit. 3. Place the embryo with the dorsal side up and hold it carefully at the anterior end with the handling needle. 4. Cut the epidermis from the posterior tip of the embryo along the dorsal midline toward the anterior tip with the cutting needle. 5. Open the embryo by moving the epidermis aside with the preparation needles or a Dumont forceps. 6. Remove the nonneural tissue that lies on top of the CNS (i.e. guts and fat body). 7. Carefully detach the CNS from the epidermis. 8. Continue as in steps 7–10 of Subheading 3.14. 9. Push the small cover slip slightly forward. The glycerol flow should push the brain hemispheres forward in front of the ventral nerve cord (see Note 20). 10. When the CNS is appropriately aligned, fix the small cover slip by applying a drop of nail polish to each corner. Let the nail

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polish dry and seal the cover slip by applying nail polish to the edges (two times). 11. Clean the cover slip from glycerol and fix it on a microscope slide with small stickers. 12. Under a microscope, the CNS can be examined from both sides (with a view from ventral or dorsal) by turning the cover slips including the filet preparation. 3.16  Analysis of the Developing Pattern of Brain Neuroblasts

The development of brain neuroblasts occurs in a stereotypic spatiotemporal pattern, with every neuroblast expressing a unique combination of marker genes. Both have been described in detail for Drosophila [8, 9], which allows the identification of individual brain neuroblasts. The brain neuroblast pattern is best examined in filet preparation of embryos from late stage 8 to late stage 11 using 100× magnification and differential interference contrast. We recommend (combinatorial) antibody stainings for distinct molecular markers (particularly against Deadpan, svp-LacZ, and engrailed protein) to analyze the patterns of brain neuroblasts in more detail. Deadpan (Dpn) is a general neuroblast marker. In early developmental stages (i.e. late stage 8-stage 9), Dpn antibody may even be sufficient to identify single neuroblasts according to their position within the entire brain neuroblast pattern [8]. From stage 9 onwards identification of single brain neuroblasts becomes increasingly difficult with general markers. We recommend the use of svp-­ LacZ enhancer trap line (available at Bloomington Stock Center, Bloomington, Indiana, USA). The expression pattern of svp-LacZ lines is stable, well described and persists throughout embryonic brain development. Svp-LacZ marks a subset of (about 40) brain neuroblasts in a characteristic pattern, which covers all three main subdivisions of the brain (i.e., proto-, deuto-, and tritocerebrum). By that, it can be used to identify the svp-LacZ-labeled neuroblasts within the entire pattern of brain neuroblasts, but also the adjacent (unlabeled) ones (Fig. 2d–d″) according to ref. [8]. In any case, it is helpful to combine stainings with engrailed, which is segmentally expressed in the posterior part of each subdivision of the brain neuroectoderm (engrailed head spot—protocerebrum, engrailed antennal stripe—deutocerebrum, engrailed intercalary stripe—tritocerebrum) and in distinct subsets of brain neuroblasts that emerge from the corresponding engrailed neuroectodermal domains (Fig. 2d–d″). These engrailed-positive brain neuroblasts additionally serve as reference points for orientation within the Dpn- or svp-LacZ-labeled neuroblast patterns [8, 9]. The (combinatorial) use of the markers mentioned above may in most cases be sufficient to identify single brain neuroblasts. However, further markers are listed in [9].

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4  Notes 1. The desired embryonic stage, and thus time window of egg collections, depends on the process to be examined and has to be chosen according to [10]. For example, the analysis of patterning in the neuroectoderm requires early developmental stages (starting at stage 5) while the analysis of NB formation requires stages of mid-embryogenesis (stages 8–11). For the former, we suggest egg collections of 3–4 h time windows, while for the latter, 8–16 h time windows are suitable (at 25 °C). 2. Egg baskets can easily be made by cutting a 50 ml falcon tube in half, cutting a hole into the cap and screwing a piece of nylon mesh between tube and cap (see ref. 11). 3. Note that long-term storage of fixed embryos may affect the staining quality. Some antibodies yield best stainings with freshly fixed embryos. 4. For double in situ hybridization, one probe has to be labeled with DIG and the other one with FITC (the use of different labels like biotin or dinitrophenol may be suitable, too). The dilution at which the probes work best must be tested before. We suggest testing probes by performing a standard in situ hybridization procedure (as described above) for a serial dilution of probes (e.g., 1:50, 1:500, 1:2500) on wildtype embryos. Probes should be detected via NBT/BCIP, since this method is very sensitive and easy. The optimal dilution is reached when the specific signal develops clearly without background staining (which may take up to 2 h or longer). If this is not the case, dilute the probe further. Higher dilutions may prolong the staining reaction. However, a clear signal should be preferred, since background staining of the different colorimetric reactions adds up and can decrease the overall staining quality significantly. Probes may be reused multiple times when stored at −20 °C. Often, the best results are reached when probes have been used before, since less unspecific bindings occur. 5. The choice of the first antibody (anti-DIG-AP or anti-FITC­AP) depends on the probe to be detected first. The first detection step is the generally less demanding and more sensitive NBT/BCIP reaction. It results in a deep blue precipitate, which is per se richer in contrast alongside the whitish tissue of the embryo. For those reasons, the inferior probe should be detected first. Here, only AP-dependent reactions are used for probe detection. Other enzymes used for colorimetric detection are peroxidases (using DAB as chromogen) or beta-­ galactosidase (using X-beta-d-Gal, Salmon-beta-d-Gal or

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Magenta-beta-d-Gal as chromogen). However, we found that the peroxidase-dependent reactions produce relatively poor signal/background ratio with this protocol, while the beta-­ Gal-­dependent reactions have been reported to be little sensitive (see ref. 12; this report also contains information on the use of further AP substrates). 6. The AP reaction may take from ~5 min up to several hours, depending on the probe used and the amount of transcript present. The staining solution should be replaced every hour. Usually, staining patterns start to emerge during the first 15 min. The reaction should be terminated as soon as (a) the relative amount of background vs. specific staining rises markedly or (b) the specific signal becomes too strong. The optimal staining intensity also depends on the context. When co-­ labeling of certain cells is expected (e.g., when a general NB marker is combined with a NB subset marker), the staining should be terminated earlier, since strong stainings may mask the presence of further dyes. Apart from that, it is recommended to terminate stainings relatively late to assure the representation of low transcript levels. Note that the NBT/BCIP staining will appear slightly lighter after the incubation in methanol and changes to a blue color since the red component of the initially purple reaction product becomes washed out. 7. Embryos can be transferred into the tube by slightly tilting the dish and flushing the embryos into one corner with PBTween. From there, they can be sucked up with a pipette. 8. Vector Red staining reactions may take up to several hours. The staining solution should be replaced every hour. The expression pattern usually starts to emerge during the first 30 min. The reaction can be slowed down by incubating at 4 °C. This may allow overnight incubation. Note that none of the dye will be washed away during subsequent steps of this protocol. Therefore, the staining reaction should be terminated as soon as the staining intensity is judged optimal. 9. This fixation step inactivates AP. Methanol is not suitable for terminating the Vector Red reaction, because the red precipitate is (at least partially) soluble in methanol. Efficient AP inactivation is especially important if further ­ NBT/BCIP-­ stainings are planned (e.g., for the detection of balancer expression) to prevent cross-reactions. 10. Many primary antibodies can be used in solutions containing 1% sodium azide; this prevents contamination with fungi or bacteria and is advantageous as an antibody can be reused several times often with increasingly better staining results. However, keep in mind that some antibodies can be impaired

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or do not work at all when using sodium azide; therefore, we recommend comparing stainings in which the antibody has been incubated in PBT and in PBT with sodium azide. 11. Additional antibodies for balancer detection can be added to the primary antibody solution. Balancers can be detected via DAB in parallel to the actual primary antibody. However, we recommend detecting balancers via AP and NBT/BCIP, since this method is more sensitive. 12. Tyramide signal amplification increases the staining intensity of DAB-stainings. This amplification step may not be necessary with good antisera. 13. The parameters of this signal amplification step are particularly critical and may have to be adapted. Prolonged incubation and inappropriately high concentrations of TSA reagent quickly lead to high background levels during the DAB staining reaction. 14. The cutting needle has to be flattened more intensely than the handling needle to obtain a sharp edge for preparation. 15. Add a drop of glycerol on the whetstone/abrasive paper. Prepare the needle under the stereo microscope by gently streaking it over the glycerol-covered surface of the stone/ paper. The cutting needle has to be sharpened to obtain an even blade with somewhat rectangular edges, while the handling needle should be whetted in a way to obtain a thicker tip with rounded edges to prevent damaging of embryos during preparation. 16. Embryos can be examined from different angles by carefully moving the upper cover slide over the spacers, which will cause the embryos to rotate in the glycerol. By that, different regions of the embryonic brain can be examined in one specimen. 17. Flat preparations, though time-consuming in production, allow the most detailed examination of gene expression pattern, even on the level of single cells (e.g. neuroblasts) [8, 9]. Yolk and other tissue which often accumulate unspecific staining are removed, and the whole neuroectoderm is presented in one plane, which significantly improves differential interference contrast. That way, neuroectodermal cells and neuroblasts, including the unstained, can be easily detected. 18. This step may need some exercise. The position of the embryo may be corrected by sliding the small cover slip in different directions. Avoid squeezing the embryo by pushing the cover slip down. 19. We recommend 40× or 63× objectives for the examination of gene expression in the neuroectoderm and 63× or 100× objectives for the analysis of the NB pattern. Stainings can be docu-

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mented on a microscope equipped with a standard digital (CCD/CMOS) camera. For digital documentation, we recommend recording selected regions of flat preparations. Depending on the magnification used, a stack of five to ten pictures along the z-axis is usually enough to cover all cells of the embryonic brain (i.e. neuroectoderm and all underlying neuroblasts of one hemisphere). 20. The hemispheres of the late embryonic brain lie on top of the ventral nerve cord, which hampers the view during microscopy. After this step, the brain hemispheres should lie in front of the ventral nerve cord, so that the whole embryonic CNS is arranged in a straight line.

Acknowledgments The authors thank Janina Seibert and Dagmar Volland for sharing protocols and considerable expertise, Janina Seibert for Fig. 2c, and Karoline F. Kraft for critically reading the manuscript. This work was supported by grants from the Deutsche Forschungsgemeinschaft (UR163/2-1 and UR163/3-1). References 1. Kosman D, Mizutani CM, Lemons D, Cox WG, McGinnis W, Bier E (2004) Multiplex detection of RNA expression in Drosophila embryos. Science 305:846 2. Skeath JB, Thor S (2003) Genetic control of Drosophila nerve cord development. Curr Opin Neurobiol 13:8–15 3. Urbach R, Technau GM (2004) Neuroblast formation and patterning during early brain development in Drosophila. BioEssays 26:739–751 4. Urbach R, Volland D, Seibert J, Technau GM (2006) Segment-specific requirements for dorsoventral patterning genes during early brain development in Drosophila. Development 133:4315–4330 5. Seibert J, Volland D, Urbach R (2009) Ems and Nkx6 are central regulators in dorsoventral patterning of the Drosohila brain. Development 136:3937–3947 6. Seibert J, Urbach R (2010) Role of en and novel interactions between msh, ind, and vnd in dorsoventral patterning of the Drosophila brain and ventral nerve cord. Dev Biol 346:332–345

7. Jussen D, von Hilchen J, Urbach R (2016) Genetic regulation and function of epidermal growth factor receptor signalling in patterning of the embryonic Drosophila brain. Open Biol 6:160202 8. Urbach R, Schnabel R, Technau GM (2003) The pattern of neuroblast formation, mitotic domains and proneural gene expression during early brain development in Drosophila. Development 103:3589–3606 9. Urbach R, Technau GM (2003) Molecular markers for identified neuroblasts in the developing brain of Drosophila. Development 103:3621–3637 10. Campos-Ortega J, Hartenstein V (1997) The embryonic development of Drosophila melanogaster. Springer, Berlin 11. Rothwell WF, Sullivan W (2000) Fluorescent analysis of Drosophila embryos. In: Sullivan W, Ashburner M, Hawley RS (eds) Drosophila protocols, 1st edn. CSHL, Cold Spring Harbor, pp 143–145 12. Hauptmann G (2001) One-, two-, and three-­ color whole-mount in situ hybridization to Drosophila embryos. Methods 23:359–372

Chapter 7 Analysis of Complete Neuroblast Cell Lineages in the Drosophila Embryonic Brain via DiI Labeling Karoline F. Kraft and Rolf Urbach Abstract Proper functioning of the brain relies on an enormous diversity of neural cells generated by neural stem cell-like neuroblasts (NBs). Each of the about 100 NBs in each side of brain generates a nearly invariant and unique cell lineage, consisting of specific neural cell types that develop in defined time periods. In this chapter we describe a method that labels entire NB lineages in the embryonic brain. Clonal DiI labeling allows us to follow the development of an NB lineage starting from the neuroectodermal precursor cell up to the fully developed cell clone in the first larval instar brain. We also show how to ablate individual cells within an NB clone, which reveals information about the temporal succession in which daughter cells are generated. Finally, we describe how to combine clonal DiI labeling with fluorescent antibody staining that permits relating protein expression to individual cells within a labeled NB lineage. These protocols make it feasible to uncover precise lineage relationships between a brain NB and its daughter cells, and to assign gene expression to individual clonal cells. Such lineage-based information is a critical key for understanding the cellular and molecular mechanisms that underlie specification of cell fates in spatial and temporal dimension in the embryonic brain. Key words Drosophila, Embryonic brain, Neural stem cell, Neuroblast lineage, DiI labeling, Antibody staining

1  Introduction Proper functioning of the brain relies on an enormous diversity of neural cells generated by neural stem cells called neuroblasts (NBs) in insects. In Drosophila, the central nervous system (CNS) including the brain develops from a bilaterally symmetrical sheet of neuroectodermal cells. It gives rise to a fixed number of NBs that segregate to the interior of the embryo. Upon segregation, an NB typically undergoes rounds of asymmetric cell division, budding off smaller intermediate precursor cells, which usually divide once to produce two postmitotic cells. Each NB produces specific neural cell types in defined time periods, and by that, generates a nearly invariant and unique cell lineage, suggesting stereotyped patterns of lineage development [1]. The fate of an individual Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_7, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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NB, and accordingly, the main features of its developing cell lineage, are specified by positional cues within the neuroectoderm [2, 3], and thereafter, by temporal cues as well as by a combination of further developmental control genes that it expresses [1, 4–8]. Many studies have shown that knowledge about the precise lineage relationships between an NB and its daughter cells is a critical key to investigate the molecular mechanisms that underlie specification of cell fates in spatial and temporal dimension [5, 8– 16]. Such lineage-­based information on the developmental origin of an NB and the neuron types it develops will not only decipher the sophisticated circuitry of the brain but also elucidate how a complex brain develops. Therefore, analysis of NB lineages with high resolution is needed, permitting a systematic identification of neural cell types by resolving the development of each single cell in an NB lineage. To unravel NB lineages in the fly brain, especially in the postembryonic period of development, very straightforward genetic clonal labeling techniques have been established, such as FLP/ FRT-based (e.g., refs. 17, 18), Gal4-based G-TRACE [19], or MARCM-based methods [20–23]. However, due to technical limitations, these genetic labeling techniques seem to be less applicable to elucidation of entire NB lineages in the embryo. As the production of FLP/FRT-clones depends on heat-shock flippase necessary to induce recombination, during embryogenesis critical levels of heat-shock flippase may not become enriched before the early-born part of a lineage has developed [24]. Also, MARCM fails to disclose entire NB lineages in the embryo, since after recombination (which depends also on a critical level of heat-shock flippase) the clonal reporter expression additionally relies on the loss of the GAL80 repressor, which seems to persist over the entire embryonic development [25]. To our knowledge, only clonal DiI labeling undoubtedly reveals the entire lineage of embryonic brain NBs. Clonal DiI labeling is highly selective and noninvasive. It is based on the application of a lipophilic fluorescent dye onto early neuroectodermal cells. The dye is easily absorbed by the membrane of the cell and diffuses through the lipids quickly without being transmitted to neighboring cells (see also ref. 26). There are several different versions of carbocyanine dyes available (e.g., the most commonly used DiI, DiO, and DiD) with different fluorescent properties. DiI for example has similar excitation properties to rhodamine; excited by green light it fluoresces red. Depending on requirement a number of alternative versions of the classical lipophilic carbocyanine dyes have been developed over the last few years, e.g., a chloromethylbenzamino derivate of DiI (CM-DiI), which shows a better staining persistence after standard tissue fixation procedures. As lipophilic carbocyanine dyes are entirely atoxic [27] the labeled cell can develop completely unaffected by the dye according to its posi-

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tion-dependent fate. Clonal DiI labeling allows in vivo to study the development of all the progeny cells derived from a single labeled neuroectodermal precursor cell (e.g., processes of cell proliferation and differentiation) and, after fixation, to analyze the cellular composition (i.e., glial or neuronal cells) and overall morphology of the fully differentiated cell clone. By using this technique the embryonic lineage of all neuroblasts in the ventral nerve cord (VNC) has been described previously [28–32]. Recently, clonal DiI labeling in combination with molecular markers and cell ablation experiments allowed us to analyze the embryonic development of a prominent central brain structure, the mushroom bodies (MBs). We unraveled the origin of the four mushroom body neuroblasts (MBNBs), their mode of formation, and could clarify the spatiotemporal development and individual cellular composition of their embryonic lineages [33]. Here we describe a protocol that was basically developed by Bossing and Technau [34], and modified by us to allow targeted DiI labeling of brain NB clones. Our methodical modifications also describe how to selectively ablate cells within a labeled NB clone, which, for example, reveals information about the temporal succession at which daughter cells are generated. Finally, we describe how to combine clonal DiI labeling with fluorescent antibody staining that permits relating protein expression to individual cells within a DiI-labeled NB clone.

2  Materials 2.1  Reagents

1. DiI (1,1′-dioctadecyl-3,3,3′,3′-tetramethylindocarbocyanineperchlorate; DiI, CM-DiI, DiD and other carbocyanine tracers can be obtained at Molecular Probes). 2. DiI/oil (5 mg/ml). 3. CM-DiI/oil (3 mg/ml). 4. DiD/oil (10 mg/ml). 5. Apple juice agar: Boil 29 g agar in 1 l apple juice until mixture is translucent. Decant into Petri dishes as long as the mixture is hot; let cool. 6. PBS (1 l 20× stock solution): Solve 151.92 g NaCl, 19.88 g Na2HPO4 (dehydrated) and 8.28 g NaH2PO4 in ddH2O, pH 7.4. 7. PBTween (0.1% Tween20 in PBS). 8. Heptane glue: Mix Cello 31.39 (Tesa, Beiersdorf) with heptane in a 2:1 ratio. 9. DAB solution: Dissolve 2–3 mg/ml DAB in 100 mM Tris– HCl, pH 7.4.

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10. Fixatives: For antibody staining after DiI labeling: Mix 200 μl 37% formaldehyde in 900 μl 1× PBS. For photoconversion: Mix 260 μl 37% formaldehyde in 900 μl 1× PBS. 11. 70% Glycerol: Dissolve 70 ml 100% glycerol in 30 ml 1× PBS. 12. Fluorocarbon oil Voltalef 10S. 13. Fetal calf serum. 2.2  Equipment

1. Inverse microscope (e.g., Leitz Fluovert FU) equipped with different fluorescent filters (Fluorescein FITC, Rhodamin, etc.). 2. 100 W halogen lamp. 3. 100 W mercury lamp. 4. Stereomicroscope. 5. Cold light source. 6. Capillary grinder (e.g., Bachofer Type 462). 7. Capillary puller (e.g., Sutter P97). 8. Micromanipulator (e.g., Leitz M). 9. Bunsen burner. 10. Cover slips (22 × 22 mm, 24 × 60 mm). 11. Glass slide (76 × 26 × 1 mm). 12. Plasticine. 13. Borosilicate glass “Meterware”).

microcapillaries

(Hilgenberg

normal

14. Glass cutter. 15. Wet chamber (e.g., Petri dish containing wet pieces of filter paper). 16. Petri dish (60 × 20 mm). 17. Scalpel. 18. Preparation needle. 19. Fine forceps (e.g., Dumont 5SF). 20. Black block dish. 21. Single-use syringe (5 or 10 ml). 22. Polyethylene tube (6 × 4 × 1 mm). 23. Transparent nail polish. 24. Tape (single sided, double sided). 25. Dissecting knife. 26. Cage (60 × 70 mm). 27. Weighing tray (41 × 41 × 8 mm).

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3  Methods 3.1  The Workplace

1. We recommend performing clonal DiI labelings, if possible, on a microscopic stage with controllable thermocouple element, or alternatively, in a temperature-controlled room (18–20 °C), so that the development of embryos is decelerated. This prolongs the time period in which successful labeling can be obtained. At warmer temperatures the viability of treated embryos declines and precise timing of DiI labeling becomes more difficult. 2. Labeling and observation are carried out on an inverted microscope. 3. Ideally, to minimize vibration, the microscope and micromanipulator carrying the DiI filled capillary are posed on a heavyweight balance table or comparable equipment. 4. The microscope needs to be equipped with exchangeable halogen and mercury lamps and appropriate filter settings capable of exciting the respective dye (for detailed information on different dyes and filter settings see “Molecular Probes Handbook of Fluorescent Probes and Research Chemicals”) and GFP.

3.2  Preparation of Fluorescent Dye/Oil Mixture

1. The fluorescent, lipophilic dye is mixed at a given concentration (see Subheading 2.1) with any commercially available vegetable oil (e.g., canola or soya oil). 2. Shake slowly (ca. 200 rpm) for 3–5 h in the dark at room temperature. The aliquots (~10 μl) can be stored at −20 °C for several years.

3.3  Preparation of Polyethylene Tube

1. Take a piece of polyethylene tube (ca. 8 cm) and heat it briefly over a small flame of a Bunsen burner (until the polyethylene starts “bubbling”). 2. Pull apart slowly from both ends until the tube reaches a length of about 50 cm. 3. Cut one end of the tube at appropriate position so that the inner diameter fits the capillary (Fig. 1c).

3.4  Manufacturing Labeling Capillaries

1. Capillaries most suitable for DiI labeling have the following characteristics: (1) a relatively long shank and (2) a relatively small tip diameter. The outer diameter at the distal tip of the capillary should be around 3–5 μm (Fig. 1d) (see Note 1). 2. Grind the capillary on a commercial capillary grinder in an angle of ca. 30°; the resulting capillary will be sharp enough to easily penetrate the vitelline membrane. For best results the capillary should be wet-ground. In the following this capillary type will be called “labeling capillary,” whereas a pulled but

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Fig. 1 Set up for DiI labeling. (a) Gastrula-stage embryo (at stage 7) glued to the cover slip for DiI labeling in the brain neuroectoderm. cf. cephalic furrow, dML dorsal midline. (b) Schematic cross-section through the head region boxed in (a); the embryo is fixed on a cover slip in appropriate orientation to label cells in the ventral or intermediate neuroectoderm (NE) under an inverse microscope. vML, ventral midline. Modified from [34]. (c) Micromanipulator equipped with a labeling capillary coupled with a polyethylene tube which is connected to a syringe. (d) The angle of the bevel of the labeling capillary should be around 30°, and the outer diameter at the distal-most tip between 3–5 μm. (e) Labeling of two neighboring neuroectodermal cells with DiI (magenta) and DiD (green). Dashed line indicates the adhesive border

unground capillary, needed for transfer and preparation of embryos, will be called “flat capillary.” 3. Under visual control through a stereomicroscope, slowly lower the capillary until its tip touches the abrasive wheel of the grinder. The tip is open when you see water climbing up into the capillary. Note, grinding does not take longer than a few seconds. 4. Mark the untreated, upper end of the labeling capillary with a water-resistant pen while it is still clamped in the grinder holder. This will help you later to fix the capillary in the micromanipulator in the right orientation. 5. To evaluate the inner diameter of the capillary tip couple a polyethylene tube (as prepared above, see Subheading 3.3) to the capillary. Insert the syringe at the other end of the tube and

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apply pressure while the tip of the capillary is under distillated water filled in a black block dish: the capillary is useful for labeling if you can hardly detect the escaping small air bubbles under intense light. For ablation of cells (see Subheading 3.16) we recommend using capillaries with a slightly larger tip opening. 6. Using the syringe, draw acetone into the capillary and eject it. The labeling capillary is then dry and ready for storage. 7. Store the labeling capillaries dust-free in a Petri dish; fix each capillary on a thin strip of plasticine. After labeling or ablation the capillary can be restored when before rinsed with acetone. 3.5  Preparing Heptane Glue Covered Cover Slips

1. In order to prepare a glue-coated cover slip (24 × 60 mm), spread a drop of heptane glue with the help of a small cover slip (22 × 22 mm) so that the cover slip becomes coated homogenously with a thin film of glue and let it dry for at least 10 min (Fig. 2a) (see Note 2). 2. Cut frames out of single-sided tape (width 25 mm). The border of a frame should be ~5 mm and the opening ~15 mm wide (Fig. 2a). Stick the frame onto the glue-coated cover slip (see Note 3).

3.6  Egg Collection and Staging

1. At least 2 days before you start collecting embryos for DiI labelings: Place the flies (between 4 and 14 days old) in a fly cage on an apple juice plate with yeast. Change the plate every 24 h (see Note 4). 2. For DiI labeling, precisely staged embryos within a range of 1 h can be obtained when changing the plate every hour at 25 °C.

3.7  Dechorionization and Mounting of Embryos for DiI Labeling

1. Fix a piece of double-sided tape to a cover slip (22 × 22 mm) (Fig. 2b). 2. Take cooled, fresh apple juice agar plates, and cut the agar into blocks of 2 × 2 cm. Place two blocks of agar in a distance of about 1.5 cm on a glass slide (76 × 26 mm), and the cover slip on top, leaving a part of each agar block exposed (Fig. 2b1). 3. Make three lines of five to seven holes each, by pressing a preparation needle into the agar blocks at an angle of about 20° (Fig. 2b2). 4. Transfer and scatter a few embryos onto the tape with a dissecting knife (Fig. 2b3). 5. Wait about 3 min to let the chorion dry. 6. Collect eggs at the blastoderm stage and dechorionate by rotating the embryo slightly along the dorsoventral axis with

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Fig. 2 Preparation of embryos for DiI labeling. (a) Spread a drop of heptane glue on a cover slip. Let the heptane glue dry for at least 10 min. Stick a single-sided tape frame on the glue-coated cover slip. (b) (1) Fix a piece of double-sided tape on a cover slip. Place the cover slip on top of two blocks of agar. (2) Make holes into the agar. (3) Transfer embryos onto the tape. Dechorionate embryos. (4) Transfer each embryo on an agar block and orientate them. (5) Cut up the agar blocks so that every embryo is on an individual block. (6) Check for optimal orientation. (7) Fix the embryo on the prepared cover slip. (8) When embryos are properly dried, cover them with Voltalef 10S oil. a anterior, p posterior, d dorsal, v ventral

the tip of a preparation needle. The chorion breaks and releases the embryo (see Note 5). 7. Carefully pick up each dechorionated embryo and transfer it to one of the agar blocks to prevent further drying (Fig. 2b4).

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8. For DiI labelings in the brain neuroectoderm, the anterioposterior axis of the embryo should be orientated perpendicular to the injection capillary. Transfer embryos into the prepared holes in the agar block, so that the posterior half of the embryo is sticking in the hole, and the procephalon is facing you (Fig. 2b6). Each embryo has to be orientated accurately with respect to the dorsoventral axis depending on the area of the brain neuroectoderm you want to label in (Fig. 2a) (see Note 6). 9. Using a scalpel, separate embryos on smaller agar blocks, each embryo on a single block (Fig. 2b5). 10. Now, stick the part of the procephalon facing you on the cover slip. Carefully lower a prepared cover slip (see below, and Subheading 3.5) onto a single block and press slightly in order to fix the embryo in proper orientation (Fig. 2b7) (see Note 7). 11. Repeat this process until all embryos are stuck next to each other (head-to-tail) to the prepared cover slip. 12. Let embryos desiccate for 5–12 min at room temperature (see Note 8). 13. As soon as the embryos are dried properly, cover them with a few drops of fluorocarbon oil Voltalef 10S (Fig. 2b8). Fluorocarbon oil (like halocarbon oil) allows oxygen to permeate to the embryo but prevents water from evaporating. 14. Until the beginning of injection the cover slip is kept in a wet chamber at room temperature. 3.8  Filling the Capillary with Dye Solution

1. Put the unbeveled end of the capillary (as indicated by the pen mark, see Subheading 3.4) into the polyethylene tube, and a single-use syringe on the other end of the tube. 2. Fix the capillary with the topside up (as indicated by the pen mark) in the holder of a micromanipulator (Fig. 1c). 3. Place a drop of 1 μl dye/oil on a cover slip (24 × 60 mm) and bring both the dye drop and the capillary tip under the microscope in focus using a 10× objective. 4. Now change to a 50× objective, and slowly draw up the dye into the capillary. 5. As soon as the tip is filled with dye, the syringe has to be removed from the polyethylene tube, the tip of the capillary to be lifted up, and the syringe, with the plunger pulled out, to be inserted again into the tube. 6. To reuse the drop of dye later you can store the cover slip in a dark, dry, and dust-free box.

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3.9  Labeling of Individual Neuroectodermal Cells and Determination of Their Position

1. Place the cover slip with the embryos under the microscope. 2. If the embryos are properly desiccated there should be a relatively sharp adhesive border around the area where each embryo is stacked to the glue and several rows of neuroectodermal cells should lie in one focal plane (Fig. 1b). 3. To identify the optimal time point for labeling and the position of the cell to be labeled it is useful to have at least a 600× or higher magnification (e.g., using a 50× immersion oil objective and a 12.5× ocular magnification). Optionally, the labeling can be controlled on a monitor using a standard digital camera (CCD/CMOS). 4. Check under the microscope the developmental stage of the embryos. By stage 7 (staging according to [35]) you can detect, for example, the dorsal and ventral midline, the cephalic furrow (see Note 9). The latest time point for successful labeling seems to be around stage 9. 5. Determine the neuroectodermal cell to be labeled by making use of the above-mentioned, morphological landmarks that help to recognize the position of individual cells (see Note 9). 6. Via the micromanipulator, lower the tip of the capillary until it is in focus, but still in a certain distance to the embryo. From now on, do not work with the micromanipulator anymore, as all movements should be effected via the microscope table. 7. Penetrate the vitelline membrane slowly and carefully with the tip of the capillary in the position where the neuroectodermal cell of interest is located, and approach the tip as closely as possible toward its cell membrane (see Note 10). 8. By slowly pushing the plunger of the syringe deposit a small drop of dye next to this cell. The diameter of the applied drop is critical for successful labeling: about 1/2 to 1/5 of the diameter of a neuroectodermal cell seems to be most promising (Fig. 1e). Immediately after depositing the drop, the dye diffuses into the cell membrane leaving behind the nonfluorescent drop of oil solvent. 9. Slowly, pull the capillary out of the embryo. 10. For double labeling in the same embryo, place two capillaries filled with different dyes (each on a different micromanipulator) as close as possible to the embryo before labeling. If no additional micromanipulator is available you can also quickly exchange the two capillaries between the labeling. 11. After labeling, briefly inspect the quality of the labeling under fluorescent light using a halogen lamp (see Note 11). 12. Finally, control the position of the labeled cell via DIC optics.

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13. Take notes of each labeling concerning position and behavior of the labeled cell (see Subheading 3.10). 14. Store the labeled embryos on the cover slip in a wet chamber at 18–20 °C. 3.10  Control of Cell Behavior After Labeling

1. About 1 h after labeling (around late stage 8/early stage 9), judge the behavior of the labeled cell using the same fluorescence and magnification settings (see Note 12). 2. If the neuroectodermal cell has delaminated, the NB was successfully labeled. To follow the development of the NB lineage in more detail, it is possible to inspect the labeling in shorter time intervals (e.g., every 30 min) without disturbing the labeled cells. 3. The identification of a labeled NB clone is facilitated when labeling is done in embryos of appropriate Gal4 strains (most can be ordered in the Bloomington or Kyoto Stock Center) in which the Gal4 pattern is visualized via a GFP reporter. Alternatively, you can combine your labeling with an antibody staining (see Subheading 3.15). 4. For further development the embryos are kept under fluorocarbon oil Voltalef 10S at 18 °C until they reach the desired stage. To obtain fully developed stage 17 NB clones, embryos are kept at 18 °C overnight. 5. Before further preparation, monitor the embryos and eliminate all those, which do not show properly labeled cell clones.

3.11  Preparation of Embryos at Developmental Stage 17

1. For better handling, we recommend removing the parts of the cover slip outside of the single-sided tape frame using a glass cutter. 2. Tip the cover slip at an ~80° angle for several minutes so that the fluorocarbon oil can drain off. 3. Then, place the cover slip in a weighing tray (41 × 41 × 8 mm) and spread about 1 ml heptane on the leftover fluorocarbon oil in order to remove it. 4. Rock the cover slip slowly for about 10–20 s. Do not rock too long since the heptane will also solve the glue and the embryos will detach. 5. Quickly place the cover slip on a glass slide for better handling. 6. Pour away the remaining heptane carefully. Wait a few seconds until the heptane is almost completely evaporated and cover the embryos with PBS immediately to prevent drying. 7. Get the embryos, one by one, out of the vitelline membrane using a flat capillary.

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8. Dissect the CNS out of the embryo by placing one fine forceps at the anterior end of the embryo and another one in its middle/posterior part. Carefully lacerate the embryo to make the CNS visible. Remove all remaining tissue from the CNS. 9. Use the flat capillary to transfer each CNS to a glue-coated cover slip (24 × 60 mm) prepared with a tape frame (as explained in Subheading 3.5) that is filled with PBS. 10. Stick the ventral nerve cord with its ventral side on the glue so that the brain hemispheres are orientated upwards. 11. The preparation can be stored in a wet chamber at 4 °C until fixation and documentation. 3.12  Preparation of First Instar Larvae

After about 30 h at 18 °C embryos are checked for eclosion under a stereomicroscope in intervals of about 15 min. Take notes on which larva hatches at which position on the cover slip, as this information is necessary to make correlations to the notes you took during the labeling (see Note 13). 1. Immediately after eclosion, take the larvae out of the oil by using a flat capillary. Transfer each larva separately to a glue-­ coated cover slip prepared with a tape frame (as explained in Subheading 3.5) filled in with PBS. 2. To prepare the CNS, tear the larva in half with a pair of fine forceps. Then turn the cuticle “inside-out” and the CNS will appear attached to the cuticle. Remove the CNS entirely from the tissue, transfer it with a capillary to another glue-coated cover slip prepared with a tape frame (as explained in Subheading 3.5) filled with PBS, and stick the ventral nerve cord with its ventral side on the glue. 3. The preparation can be stored in a wet chamber at 4 °C. If you want to photoconvert the clone continue as described in Subheading 3.17, otherwise go on with fixation (see Subheading 3.13).

3.13  Fixation

1. Remove the PBS under visual control using a stereomicroscope and replace it immediately with fixation solution (see Subheading 2.1). Note, as fixatives are toxic, take precautions (e.g., place the stereomicroscope on a portable fume hood). 2. Fix 20 min for subsequent antibody staining (as described in Subheading 3.15), otherwise 10 min. 3. Thereafter, carefully replace the fixative with PBS. 4. Change the PBS 4–5 times.

3.14  Documentation

1. Fix the cover slip (24 × 60 mm) carrying the preparation on a glass slide with tape for better handling.

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2. Remove excessive PBS so that the border of the liquid slightly overtops the single-sided tape frame (see Note 14). 3. Now carefully place a cover slip (22 × 22 mm) onto the single-­ sided tape frame. 4. By slightly moving the upper cover slip, the brain hemispheres that get in contact with this cover slip (as explained in Subheadings 3.11 and 3.12) can be brought into the best possible position for exposure. 5. Fix the cover slip to the underlying cover slip by applying nail polish at the corners. 6. Immediately document the labeled NB clone using a confocal microscope (Fig. 3a, b). Transport the specimen in a wet chamber (see Note 15). Optionally, 3D reconstructions can be generated from the LSM stacks using, for example, Amira or comparable software (Fig. 3c). 3.15  Optional: DiI Labeling Combined with Fluorescent Antibody Stainings

As far it was only possible to combine photoconverted DiI labelings with nonfluorescent histochemical antibody stainings (visualized e.g., via the alkaline phosphatase reaction) with the disadvantage that the photoconverted cells themselves, which have accumulated strong levels of diamino benzidine (DAB), could not be antibody stained. Now this limitation can be avoided by combining labeling, performed with the improved fixable DiI derivatives (e.g., CM-DiI), with fluorescent antibody stainings. To carry out fluorescent antibody stainings with fluorescent DiI-labeled clones, another critical issue is the proper usage of detergents, needed for antibodies to penetrate the membranes of cell tissues. As the carbocyanine dyes are lipophilic, conventional detergents in widely used concentrations (such as 0.1–0.3% Triton X-100 or 0.05% saponin) result in extensive diffusion of the DiI label out of the tissue (own observations; see also [36]). Presented below is a protocol in which we apply a low concentration of 0.1% Tween20, and in addition, mechanically perforate the neurilemma that envelops the brain. 1. After fixation, to facilitate the penetration of the antibodies into the brain tissue, carefully perforate the neurilemma with a flat capillary. Perforate generously but do not damage the area containing the labeled clones. 2. Thereafter, cover the specimen for 48 h with primary antibody/antibodies dissolved in 0.1% PBTween (~200 μl). Put the cover slip in a wet closed chamber at 4 °C. 3. Then take off the primary antibody/antibodies and replace with 0.1% PBTween twice. 4. Cover three times for 20 min with 0.1% PBTween.

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Fig. 3 Different applications of clonal DiI labeling. (a) Combined confocal images of a DiI-labeled mushroom body NB clone (a) in a Ok107 (ey)-Gal4; UAS-CD8::GFP brain (a′) at mid/late stage 17. Note that some cells of the DiI-labeled clone express GFP, indicating that this is a mushroom body NB clone. (b) DiI-labeled mushroom body NB clone at late stage 17 after ablation of the NB at stage 11. Accordingly, only the early-born part of its lineage has developed. (c) 3D reconstruction of a DiI-labeled mushroom body NB clone (magenta) in the early first larval instar in comparison to the structures of the entire mushroom body (green) as revealed by Ok107 (ey)-Gal4; UAS-CD8::GFP expression. (d) A mushroom body NB clone (magenta) and another protocerebral NB clone (green) at late stage 17 obtained after labeling of two neighboring cells with DiI (magenta) and DiD (green) (see inset). (a–d) Adapted from [33]. (e) Photoconverted protocerebral NB clone at early stage 17. (f) DiI-labeled protocerebral NB clone (at early first larval instar) before (f) and after (f′) fluorescent antibody staining against the gene products Eyes absent (f″a) and Retinal homeobox (f″b). Note that antibody staining leads to partial loss of DiI signal. (f″) Merge

5. Dissolve the secondary antibody/antibodies (take in consideration that you use appropriate fluorescent conjugates which do not interfere with applicated carbocyanine dyes) in 0.1% PBTween and incubate for 18–20 h at 4 °C. In case of weak antibody stainings, a Tyramide Signal Amplification system (TSA) is recommended (according to the manufacturer’s protocol of PerkinElmer, Waltham, USA).

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6. After taking off the secondary antibody/antibodies, replace with 0.1% PBTween twice and then cover for 15 min with 0.1% PBTween twice. 7. Subsequently, wash twice with PBS. 8. Finally, put a cover slip on the tape frame, fix with nail polish, and follow up with documentation immediately as the DiI signal fades relatively fast (see Subheading 3.14). 3.16  Optional: Ablation of DiI-Labeled NBs and/or Daughter Cells

1. For cell ablations, use embryos that have been labeled as described in Subheading 3.9 and have developed into the desired stage. Use a microscope equipped with a 100 W halogen lamp and appropriate filter settings. Bring the specimen in which cell ablations will be done under a sufficient magnification (600× or higher) in focus. Connect the ablation capillary (see Subheading 3.4) with the tube, then clamp the capillary into the micromanipulator and lower it into the Voltalef oil. Finally, connect the syringe to the other end of the tube to prevent Voltalef oil entering the tip. 2. Identify the labeled cell(s) to be ablated under the halogen lamp. 3. Using the micromanipulator, move the capillary toward the embryo under transmitted light. 4. Slowly and carefully move the cross table of the microscope toward the capillary tip until the capillary penetrates the vitelline membrane, and then carefully direct it to the cell(s) designated for ablation. 5. Once the tip of the capillary reaches the cell to be ablated, cautiously pull the syringe plunger in order to suck off the cell(s) (see Note 16). 6. Remove the capillary from the embryo by moving the cross table. 7. Store the treated embryos in a wet chamber at 18 °C until they have reached the desired developmental stage. 8. Then prepare, fix and document the specimen as described in Subheadings 3.11–3.14.

3.17  Optional: Photoconversion of DiI-Labeled Preparations

Photoconversion transforms the DiI label into a permanent reaction product via the Maranto reaction in which illumination releases a singlet oxygen that oxidizes DAB [37] (see also ref. 38). DiI-labeled photoconverted clones can be investigated using conventional light microscopy, and preserved for many years. 1. Incubate for 1 h in calf serum. 2. Exchange the drop of PBS with a drop of DAB solution (that has been briefly centrifuged before). Note, DAB is a hazardous chemical—handle with appropriate precautions!

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3. Photoconvert the labeled NB clone under the microscope with a 100 W mercury lamp and rhodamine filter using a water or oil immersion objective with about 50× magnification. Photoconversion is completed when all fluorescent signal is gone, this takes usually 10–20 min. Check regularly to make sure that the preparation is not over dyed. 4. After photoconversion, remove the DAB. 5. Rinse the preparation with PBS 3–4 times. 6. Fix in formaldehyde solution for 15 min. 7. Rinse 3–4 times with PBS. 8. Place a drop (20–25 μl) of 70% glycerol on another cover slip (24 × 60 mm). 9. Using a flat capillary, transfer the preparation to the glycerol drop and bring it in appropriate orientation. 10. Carefully cover the glycerol drop containing the CNS with a small cover slip (18 × 18 mm). Control the orientation of the preparation under a stereomicroscope while the glycerol spreads between both glass slides. Excess glycerol can be removed with tissue paper. 11. Fix the cover slip by applying a drop of nail polish to all four corners. Let those drops dry and seal all sides two times with nail polish. The photoconverted NB clones can now be documented using Nomarski optics and a 63× (or 100×) immersion oil objective, as camera lucida drawings, or can be digitized. For example, with the help of a standard digital camera (CCD/ CMOS), a motorized microscope table and appropriate documentation software, it is useful to take a sequence of individual pictures in z-axis (the distance between different focal planes should be 1 μm). Saving each focal plane in Tiff format, you can merge all files to a film sequence using, e.g., QuickTime Player; this often facilitates understanding the spatial complexity of brain NB clones.

4  Notes 1. Glass capillaries for DiI labeling can be produced by any commercial pipette puller, e.g., Sutter P97, which allows both the heat and the strength to be varied. Change the settings of the puller according to the operating manual until you get ideal needles (see, e.g., the comprehensive guide to pulling capillaries on the website of Sutter instruments http://www.sutter. com/contact/faqs/pipette_cookbook.pdf). 2. Spread the heptane-glue out carefully so that you have one thin but continuous layer of glue. If there is not enough glue, the embryos will not stick to the cover slip. However, if you apply

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too much glue, the embryo sinks into the glue and does also not stick properly. In addition, the capillary will be clogged by glue. 3. Make sure the frame is big enough so that there is a distance of at least 4 mm between the embryos and the inner edges of the frame. Otherwise the capillary cannot be placed near enough to the embryo. Fix the frame carefully without producing air bubbles. Otherwise the oil or PBS will later flow off the cover slip. After all embryos are fixed on the cover slip, place it onto a glass slide for better handling. 4. You do not need a vast amount of individuals, but as you will only work with embryos from a 1 h collection at 25 °C you need to make sure that your flies lay enough eggs. In order to guarantee efficient labeling it is important that you have sufficient numbers of precisely staged embryos. Of course, the collection schedule is temperature dependent. 5. We do not recommend chemical dechorionization as according to our experience this seems to cause misdevelopment or damage of embryos at higher rates. 6. The orientation of the embryo on the agar block is critical for the accessibility of the capillary to the neuroectodermal area one wants to label in. Orientate the embryos on the agar block by slightly moving them with the tip of the preparation needle. Consider that the adhesive border of the attached part of the neuroectoderm is close to the cell you want to label when you stick the embryo to the cover slip (Fig. 1b). For example, if you plan to label cells in the ventral neuroectoderm of the brain, the embryo should be placed sagitally so that the adhesive border is close to the ventral head midline (Fig. 1a). Accordingly, if you want to label in the dorsal neuroectoderm, orientate the embryo in the opposite direction. It is also important for the viability of the embryo that the ectodermal region that sticks to the glass surface is not too large, as otherwise the embryo does not develop any further. 7. While mounting the embryos on the one hand it is important to press carefully and vertically so that the embryos are neither damaged nor altered in position, on the other hand you need to press slightly onto the cover slip so that several rows of neuroectodermal cells come to lie in one focal plane. 8. The average time for desiccation depends highly on temperature and humidity. Control the drying of the embryos under a stereo microscope. Observe the embryos under a cold light source until you note two reflecting stripes on each side of the ventral midline. These stripes will disclose the level of desiccation: at the beginning these stripes are well defined and smooth, while after a few minutes they show tiny folds; once they are visible, the right level of desiccation has been achieved.

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9. To perform targeted labeling of neuroectodermal cells in the brain, it is necessary to make use of morphological landmarks which help to recognize the position of individual neuroectodermal cells: easily identifiable are the cephalic furrow and the dorsal and ventral midline. It is also helpful to use the stereotypical pattern of procephalic mitotic domains as morphological landmarks, in addition to the above-mentioned landmarks. The procephalic ectoderm can be subdivided into several mitotic domains, which are characterized as discrete groups of cells that synchronously enter mitosis [39]. Each mitotic domain has a distinct time of entry into mitosis, and discloses a specific shape. Mitotic domains (except domain B) are only recognizable during their period of mitosis but not before or thereafter. Almost all procephalic mitotic domains have already completed mitosis (by early/mid-stage 8) before they give rise to first NBs. Recently, we were able to establish mitotic domain B, positioned in the central area of the brain neuroectoderm, as a useful landmark for a systematical DiI labeling of neuroectodermal cells, and could show that, among others, the NBs of the mushroom body originate there [33]. Mitotic domains can be visualized in vivo, for example, in ubi-GFPnls embryos: cells of the mitotically inactive domain B (continuously showing nuclear GFP) can be distinguished from the surrounding mitotically active domains which transiently lose nuclear GFP [33]. 10. Once the tip of the dye-filled capillary has entered the embryo you must completely avoid movements along the anterioposterior axis of the embryo. Otherwise you will destroy tissue or the labeled cell, and risk misdevelopment of the cell clone. 11. You must use a halogen lamp for the inspection. Using a mercury lamp, due to the problem of phototoxicity, will lead to apoptosis of the labeled cells! 12. Check the behavior of the cell after labeling. The mode of brain NB formation reveals some differences depending on the neuroectodermal region an NB originates. For example, except in mitotic domain B, cells in all other parts of the brain neuroectoderm undergo mitosis before first NBs emerge. If a neuroectodermal cell divides in parallel to the ectodermal surface (typical for mitotic domain 1 or 5), then it may produce an NB and an epidermoblast, each generating a subclone, one becoming located in the later brain, the other in the epidermis (perhaps mapping in a sensory organ). If the neuroectodermal cell divides perpendicular to the ectodermal surface, then it may belong to mitotic domain 9. If the cell divides perpendicular (and asymmetrical) but in subectodermal position, then this is likely to be an NB producing a ganglion mother cell. It is also helpful to make notes on the time point of NB formation

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(indicating if it is an early or late developing NBs). For further details on different modes of brain NB formation see [40]. 13. According to our experience, DiI labelings can be followed until the first hours of the first larval instar since the lipophilic dye gradually dilutes, and thus fades, in the growing membrane of developing neuronal/glial cells. 14. Do not embed the preparations in pure glycerol, nor in other glycerol-based mounting mediums (e.g., Vectashield). Lipophilic dyes, such as DiI, dissolve in glycerol. Embedding can only be done in PBS without glycerol. However, due to the evaporation of PBS, every supplement should be prepared individually and immediately prior to the exposure under microscope. 15. As DiI signal may become weaker over time, it is often useful to document a labeled NB clone before and after the antibody staining. Because of the relatively long procedure you risk to lose the signal. 16. For cell ablations, halogen lamps with rhodamine filters and transmitted light at once are recommended. In order to recognize the DiI fluorescent signal of the cell(s) to be ablated, and simultaneously, the surrounding tissue, reduce the intensity of the transmitted light to an appropriate low level.

Acknowledgments The authors thank Thomas Kunz, Martin Steimel, and Christof Rickert for sharing protocols and considerable expertise, and David Jussen for critically reading the manuscript. We are grateful to Gerd Technau for his general support. This work was supported by grants from the Deutsche Forschungsgemeinschaft (UR163/2-1 and UR163/3-1) and by a research stipend to K.F.K. from the FTN of the University Mainz. References 1. Technau GM, Berger C, Urbach R (2006) Generation of cell diversity and segmental pattern in the embryonic central nervous system of Drosophila. Dev Dyn 235:861–869 2. Skeath JB (1999) At the nexus between pattern formation and cell-type specification: the generation of individual neuroblast fates in the Drosophila embryonic central nervous system. BioEssays 21:922–931

3. Urbach R, Technau GM (2004) Neuroblast formation and patterning during early brain development in Drosophila. BioEssays 26:739–751 4. Jacob J, Maurange C, Gould AP (2008) Temporal control of neuronal diversity: common regulatory principles in insects and vertebrates? Development 135:3481–3489 5. Kao C-F, Lee T (2010) Birth time/order-­ dependent neuron type specification. Curr Opin Neurobiol 20:14–21

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6. Lin S, Lee T (2012) Generating neuronal diversity in the Drosophila central nervous system. Dev Dyn 241:57–68 7. Skeath JB, Thor S (2003) Genetic control of Drosophila nerve cord development. Curr Opin Neurobiol 13:8–15 8. Doe CQ (2017) Temporal Patterning in the Drosophila CNS. Annu Rev Cell Dev Biol 33:219–240 9. Baumgardt M, Karlsson D, Terriente J et al (2009) Neuronal subtype specification within a lineage by opposing temporal feed-forward loops. Cell 139:969–982 10. Jefferis GS, Marin EC, Stocker RF et al (2001) Target neuron prespecification in the olfactory map of Drosophila. Nature 414:204–208 11. Lai S-L, Awasaki T, Ito K et al (2008) Clonal analysis of Drosophila antennal lobe neurons: diverse neuronal architectures in the lateral neuroblast lineage. Development 135:2883–2893 12. Pearson BJ, Doe CQ (2004) Specification of temporal identity in the developing nervous system. Annu Rev Cell Dev Biol 20:619–647 13. Yu H-H, Lee T (2007) Neuronal temporal identity in post-embryonic Drosophila brain. Trends Neurosci 30:520–526 14. Yu H-H, Kao C-F, He Y et al (2010) A complete developmental sequence of a Drosophila neuronal lineage as revealed by twin-spot MARCM. PLoS Biol 8 15. Bahrampour S, Gunnar E, Jonsson C et al (2017) Neural lineage progression controlled by a temporal proliferation program. Dev Cell 43:332–348 16. Walsh KT, Doe CQ (2017) Drosophila embryonic type II neuroblasts: origin, temporal patterning, and contribution to the adult central complex. Development 144:4552–4562 17. Golic KG, Lindquist S (1989) The FLP recombinase of yeast catalyzes site-specific recombination in the Drosophila genome. Cell 59:499–509 18. Xu T, Rubin GM (1993) Analysis of genetic mosaics in developing and adult Drosophila tissues. Development 117:1223–1237 19. Evans CJ, Olson JM, Ngo KT et al (2009) G-TRACE: rapid Gal4-based cell lineage analysis in Drosophila. Nat Methods 6:603–605 20. Lai S-L, Lee T (2006) Genetic mosaic with dual binary transcriptional systems in Drosophila. Nat Neurosci 9:703–709 21. Lee T, Luo L (1999) Mosaic analysis with a repressible cell marker for studies of gene function in neuronal morphogenesis. Neuron 22:451–461

22. Lee T (2009) New genetic tools for cell lineage analysis in Drosophila. Nat Methods 6:566–568 23. Yu HH, Chen CH, Shi L et al (2009) Twin-­ spot MARCM to reveal the developmental origin and identity of neurons. Nat Neurosci 12:947–953 24. Larsen C, Shy D, Spindler SR et al (2009) Patterns of growth, axonal extension and axonal arborization of neuronal lineages in the developing Drosophila brain. Dev Biol 335:289–304 25. Luo L (2005) A practical guide: single-neuron labeling using genetic methods. In: Yuste R, Konnerth A (eds) Imaging in neuroscience and development. A laboratory manual. Cold Spring Harbor Laboratory Press, New York, NY, pp 99–110 26. Honig MG, Hume RI (1986) Fluorescent carbocyanine dyes allow living neurons of identified origin to be studied in long-term cultures. J Cell Biol 103:171–187 27. Honig MG, Hume RI (1989) Dil and DiO: versatile fluorescent dyes for neuronal labelling and pathway tracing. Trends Neurosci 12:333–340 28. Bossing T, Udolph G, Doe CQ et al (1996) The embryonic central nervous system lineages of Drosophila melanogaster. I Neuroblast lineages derived from the ventral half of the neuroectoderm. Dev Biol 179:41–64 29. Schmidt H, Rickert C, Bossing T et al (1997) The embryonic central nervous system lineages of Drosophila melanogaster. II Neuroblast lineages derived from the dorsal part of the neuroectoderm. Dev Biol 189:186–204 30. Schmid A, Chiba A, Doe CQ (1999) Clonal analysis of Drosophila embryonic neuroblasts: neural cell types, axon projections and muscle targets. Development 126:4653–4689 31. Birkholz O, Rickert C, Nowak J et al (2015) Bridging the gap between postembryonic cell lineages and identified embryonic neuroblasts in the ventral nerve cord of Drosophila melanogaster. Biol Open 4:420–434 32. Rickert C, Lüer K, Vef O, Technau GM (2018) Progressive derivation of serially homologous neuroblast lineages in the gnathal CNS of Drosophila. PLoS One 13:e0191453 33. Kunz T, Kraft KF, Technau GM, Urbach R (2012) Origin of Drosophila mushroom body neuroblasts and generation of divergent embryonic lineages. Development 139:2510–2522 34. Bossing T, Technau GM (1994) The fate of the CNS midline progenitors in Drosophila as revealed by a new method for single cell labelling. Development 120:1895–1906

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tion, cryosectioning, and photoconversion for 35. Campos-Ortega JA, Hartenstein V (1997) The light and electron microscopic analysis. embryonic development of Drosophila melaJ Histochem Cytochem 38:725–733 nogaster. Springer, Berlin 36. Elberger AJ, Honig MG (1990) Double-­ 39. Foe VE (1989) Mitotic domains reveal early commitment of cells in Drosophila embryos. labeling of tissue containing the carbocyanine Development 107:1–22 dye DiI for immunocytochemistry. J Histochem Cytochem 38:735–739 40. Urbach R, Schnabel R, Technau GM (2003) The pattern of neuroblast formation, mitotic 37. Maranto AR (1982) Neuronal mapping: a phodomains and proneural gene expression during tooxidation reaction makes Lucifer yellow useful early brain development in Drosophila. for electron microscopy. Science 217:953–955 Development 130:3589–3606 38. von Bartheld CS, Cunningham DE, Rubel EW (1990) Neuronal tracing with DiI: decalcifica-

Chapter 8 Flybow to Dissect Circuit Assembly in the Drosophila Brain: An Update Emma L. Powell and Iris Salecker Abstract Visualization of single neurons and glia, as well as neural lineages within their complex environment is a pivotal step towards uncovering the mechanisms that control neural circuit development and function. This chapter provides detailed technical information on how to use Drosophila variants of the mouse Brainbow-2 system, called Flybow, for stochastic labeling of individual cells or lineages with different fluorescent proteins in one sample. We describe the genetic strategies and the heat shock regime required for induction of recombination events. Furthermore, we explain how Flybow and the mosaic analysis with a repressible cell marker (MARCM) approach can be combined to generate wild-type or homozygous mutant clones that are positively labeled in multiple colors. This is followed by a detailed protocol as to how to prepare samples for imaging. Finally, we provide specifications to facilitate multichannel image acquisition using confocal microscopy. Key words Drosophila, Brainbow, Confocal laser scanning microscopy, Genetics, Immunostaining, Mosaic analysis with a repressible cell marker, Multicolor celllabeling

1  Introduction Neuron and glial subtypes display elaborate branching patterns with distinct shapes that are indicative of their specific functions within neural circuits. Genetic approaches labeling single neurons or glial cells within the context of their complex surroundings are thus much needed additions to the neurobiologist’s technical repertoire. Furthermore, the ability to follow several neural lineages in parallel is instrumental for determining the relative contributions and interactions of different clones during brain development. Multicolor cell labeling techniques offer important opportunities for advancing both types of studies. In 2007, Livet et al. [1] pioneered the Brainbow system, a genetic multicolor cell labeling approach for mice, which makes it possible to visualize neurons and glia in different hues by the stochastic and combinatorial expression of three spectral variants of fluorescent proteins (FPs). Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_8, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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The Drosophila Brainbow [2] and Flybow (FB) [3] systems were the first adaptations of this approach for use in flies, containing distinct features that take advantage of genetic methods available in this model organism. During the past years numerous multicolor cell labeling techniques with extended functionalities have been steadily added to the genetic toolbox of mice, chicken, zebrafish, and Drosophila (reviewed in ref. 4;  5). Here, we provide detailed information about how to use one of these approaches, Flybow, for studies in the brain. FB is based on the Brainbow-2 strategy (see Fig. 1a). It relies on the Gal4/UAS system [6] to control transgene expression, while a FLP recombinase with altered specificity (mFLP5) promotes inversions and excisions of cassettes flanked by mFRT71 sites [7]. Each cassette contains two FP-encoding cDNAs in opposing orientations. FPs are membrane-tethered either by a Cd8a (cd8) [8] or the myr/palm (mp) sequence of Lyn kinase [9]. FP sequences are followed by SV40 and hsp70Ab polyadenylation (pA) signals. mFLP5 is expressed under the control of the heat shock promoter (hs-mFLP5) and induces inversions of DNA cassettes by recombining mFRT71 sites in opposing orientations, or excisions (FLP-out) by recombining mFRT71 sites in the same orientation. Cassettes have been subcloned into a vector containing 10 UAS sites [10]; the FP closest to these sites will be expressed. FB1.0 consists of one cassette encoding two FPs (mCherry [11] and Cerulean-V5 [12]). FB1.1 contains two cassettes, each encoding two FPs (EGFP [13] and mCitrine [14]; mCherry and Cerulean-V5). FB2.0 features an additional stop cassette, flanked by canonical FRT sites facing in the same orientation, which can be excised by FLP recombinase. The stop cassette consists of lamin cDNA, followed by two hemagglutinin (HA) epitope-tag sequences and hsp70Aa and hsp27 pA signals. Since expression is only induced in cells, in which Gal4 and canonical FLP expression overlap, sparse labeling can be achieved. Lineage tracing relies on the fact that daughter cells maintain expression of the genetic marker they inherited from their progenitor cell. Thus, the timing of site-­ specific recombination events can be adjusted to promote multicolor labeling of single cells or lineages. The first set of FB constructs [3] relied on the cyan FP Cerulean, which requires detection with a V5 antibody because of its weak endogenous fluorescent signal in flies. To eliminate the necessity for antibody detection, we generated a second set of transgenic lines, i.e. FB1.0B, FB1.1B, and FB2.0B [15], in which cd8-tethered Cerulean was replaced by myr/palm-tethered mTurquoise, a brighter cyan FP variant [16] (see Figs. 1b and 2). In the third set of transgenes, i.e. FB1.0C, FB1.1C, and FB2.0CB [20], cd8-mCherry was substituted by myr/palm-tethered orange-­ shifted TagRFP-T [21], while mTurquoise was replaced by mTurquoise 2 [22] to enable imaging with a two-photon microscope

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Fig. 1 Schematic representation of Flybow (FB) transgenes and color outcomes. Expression of the modified FLP recombinase mFLP5 induces inversions and excisions of cassettes each flanked by inward facing mFRT71 recognition sites (black triangles). Cassettes consist of two FP cDNAs with opposite orientations. In FB2.0, an additional FLP-out cassette flanked by canonical FRT sites facing in the same orientation (grey triangles) precedes the FP-containing cassettes. (a) A detailed description of FB1.0, FB1.1, and FB2.0 transgenes is provided in [3]. (b) In FB1.0B, FB1.1B, and FB2.0B transgenes, Cerulean-V5 has been replaced by mTurquoise (mTq). In FB1.0C, FB1.1C, and FB2.0C transgenes, mCherry was replaced by TagRFP-T and mTurquoise (mTq) by mTurquoise 2 (mTq 2)

(see Fig. 1c). As FB tools solely rely on endogenous fluorescence signals and not immunolabeling, they can be used for live imaging. As a further improvement, we generated additional hs-mFLP5 stocks for our toolkit [15]. Three are homozygous viable transgene insertions on the X, 2nd, and 3rd chromosomes, while one line contains a homozygous lethal insertion on the 2nd chromosome showing higher recombination efficiency (see Table 1). As FB relies on a modified FLP-FRT system and only one UAS-transgene [3], it is readily compatible with available genetic loss- and gain-of-function approaches, including mosaic analysis

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Fig. 2 Expression of B set of FB transgenes in the adult Drosophila optic lobe. Examples for original transgenes can be found in [3]. Photoreceptor cells (R1–R8) visualized with mAb24B10 (blue) extend axons into the optic lobe: R1–R6 axons innervate the lamina; R8 and R7 axons terminate in the medulla [17]. NP4151-Gal4, an enhancer trap insertion into the Netrin B locus [18, 19] drives expression of FB transgenes in lamina neurons L3 and medulla neuron subtypes, which extend axons into the lobula and lobula plate. (a, a′) Heat shock application induces expression of mFLP5 to trigger inversion events in FB1.0B260b, labeling individual neurons with mTurquoise instead of mCherry. (b, b′) In FB1.1B260b, excision and inversion events lead to expression of mCitrine, mCherry, or mTurquoise instead of the default marker EGFP. (c, c′) FB2.0B260b facilitates sparse labeling of single neurons in a population because it relies on heat shock-induced expression of FLP for excision of the stop cassette flanked by FRT sites, and of mFLP5 for stochastic expression of the four FPs. Confocal images represent single optic sections. Images in b, b′, c, and c′ were processed using channel separation software. Heat shock exposure: 3 × 45 min (a–b′) and 3× 30 min (c, c′) at 48, 72, and 96 h after puparium formation. Scale bars: 20 μm

with a repressible cell marker (MARCM), a landmark mosaic analysis tool developed by Lee and Luo in 1999 [25]. This technique enables Drosophila geneticists to generate homozygous –wild-type or mutant– clones of single cells that are labeled with a reporter (see Fig. 3a). When combining FB with MARCM, FLP mediates recombination events between FRT sites on homologous chromosomes in trans, whereas mFLP5 is responsible for in cis recombination events of mFRT71 sites within FB1.1 transgenes (see Fig. 3b). Finally, in MARCMbow [26], mFLP5 recombinase expression is not induced by heat shock but controlled by Gal4 expression of a UAS-mFLP5 transgene to diversify FP choice in lineages.

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Table 1 Basic Flybow transgene toolkit Flybowa

Chr.

Insertion

FB1.0260b

2

Viable

[3]

FB1.049b

3

Viable

[3]

FB1.0B

260b

2

Viable

[15]

FB1.0B

49b

3

Viable

[15]

FB1.0C260b

2

Viable

[20]

FB1.0C49b

3

Viable

[20]

FB1.1

260b

2

Viable

[3]

FB1.1

49b

3

Viable

[3]

FB1.1B260b

2

Viable

[15]

FB1.1B49b

3

Viable

[15]

FB1.1C

2

Viable

[20]

FB1.1C49b

3

Viable

[20]

FB2.0260b

2

Viable

hs-FLP1; FB2.0260b

[3]

FB2.0

3

Viable

hs-FLP ; FB2.0

[3]

FB2.0B

2

Viable

hs-FLP ; FB2.0B

[15]

FB2.0B49b

3

Viable

hs-FLP1; FB2.0B49b

[15]

FB2.0C260b

2

Viable

[20]

FB2.0C

3

Viable

[20]

hs-mFLP

Chr.

Insertion

Related parental stocks

Efficiencyb

hs-mFLP5

2

Lethal

hs-mFLP5/Gla Bc; TM2/TM6b

∗∗

[3]

hs-mFLP5

3

Lethal

Gla Bc/CyO; hs-mFLP5/TM2

∗∗

[3]

hs-mFLP5MH15

X

Viable

∗∗

[15]

hs-mFLP5MH1

2

Lethal

∗∗∗

[15]

hs-mFLP5

2

Viable

∗∗

[15]

hs-mFLP5

3

Viable

∗∗

[15]

260b

49b 260b

49b

MH12 MH3

Related parental stocks

1 1

References

49b 260b

∗∗ good efficiency, ∗∗∗ very good efficiency a 260b and 49b indicate the attP site-containing loci used for FB transgene insertion on the 2nd and 3rd chromosomes, respectively [3]. For additional information regarding the 260b attP landing site, see [23, 24] b Recombination efficiencies were estimated by monitoring excision events of a stop cassette flanked by mFRT71 sites in 3rd instar larval eye imaginal discs

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Fig. 3 Schematic representation of MARCM and combined MARCM and Flybow (FB) approaches. (a) The MARCM approach relies on two FRT recombination sites inserted at the same location close to the centromeres (open circles) of homologous chromosomes. The Gal80 transgene controlled by the ubiquitous enhancer tubP is located distal to the FRT site of one of the chromosomes. Expression of the repressive Gal80 protein prevents Gal4-mediated activation of any UAS-transgene in the cell. FLP recombinase and Gal4 transgenes can be positioned on any chromosome except on the FRT-containing chromosome arm. The homologous chromosome can carry a mutation (red asterisks). Site-specific somatic recombination between FRT sites mediated by FLP results in daughter cells, which are homozygous either for the chromosome carrying the mutation or the chromosome carrying the Gal80 transgene. The former is positively labeled with a marker, such as GFP, because Gal80 is absent following chromosome segregation during mitosis. (b) When combining MARCM and FB, an additional mFLP5 transgene e.g. under the control of a heat shock (hs) promoter is introduced to mediate inversions and excisions within the FB1.1 reporter. Positively labeled progeny can express one of four FPs

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FB transgenes can support anatomical and functional studies of any genetically accessible cell population in the developing and adult nervous system and other tissues. While FB also can be applied to studies in embryos, in this updated chapter, we focus on its use in the larval, pupal, and adult nervous system.

2  Materials Prepare all solutions with ultrapure deionized water (18.2 MΩ cm resistivity) from a water purification system. 2.1  Genetic Crosses and Clone Induction

1. Plastic vials containing standard cornmeal/agar medium. 2. Virgin female and male flies with the proper genotype (see Table 1). 3. Water bath at 37 °C.

2.2  Buffers

1. Phosphate buffered saline (PBS, 130 mM NaCl, 7 mM Na2HPO4 7H2O, 3 mM NaH2PO4 H2O): Prepare 1 L of 10× PBS stock solution by weighing 75.97 g NaCl, 18.76 g Na2HPO4·7H2O and 4.14 g NaH2PO4·H2O into a glass beaker. Add water to a volume of 990 mL. Dissolve crystals by stirring with a magnetic stirrer and adjust pH to 7.4 with drops of concentrated 12 N HCl and 10 M NaOH, and subsequently 1 M dilutions. Fill up to 1 L with water. Store this solution at room temperature in a glass bottle (see Note 1). To prepare 1 L PBS solution, mix 100 mL 10× PBS stock solution with 900 mL water and readjust pH using 1 N HCl or 1 M NaOH if necessary. Store at 4 °C. 2. 0.5% Phosphate-buffered saline with Triton (PBT): 100 mL PBS, 0.5 mL Triton®-X-100 (Sigma). Add 0.5 mL Triton®-X-100 (Sigma) by using a 1 mL syringe and mix vigorously with a magnetic stirrer to dissolve the detergent. Store solution at 4 °C. 3. 0.1 M Phosphate buffer (PB): Weigh 1.73 g NaH2PO4·H2O into a glass beaker, add 125 mL distilled water, and stir until crystals are dissolved to make a 0.1 M NaH2PO4 solution. Weigh 13.4 g Na2HPO4·7H2O into a 2nd larger glass beaker, add 500 mL water and stir to make a 0.1 M Na2HPO4 solution. Store 100 mL of 0.1 M Na2HPO4 solution in a separate glass bottle. Pour the 0.1 M NaH2PO4 solution into the remaining 0.1 M Na2HPO4 solution until pH reaches 7.4 to make 0.1 M PB. Store solutions at 4 °C. 4. Phosphate buffer with lysine (PBL): Dissolve 3.6 g L-Lysine-­ HCl (Sigma) in 100 mL water. Add approximately 40 mL 0.1 M Na2HPO4 solution until pH 7.4 is reached. Add

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0.1 M PB to make up 200 mL. Filter-sterilize using a 250 mL filter unit (0.2 μm pore size, 50 mm membrane diameter, Nalgene). Store solution at 4 °C for up to three months. 5. Blocking buffer: Add 1 mL normal goat serum (NGS, Sigma) to 9 mL PBT to get 10% NGS-PBT (see Note 2). 2.3  Fixative

1. 8% Paraformaldehyde stock solution: 0.8 g paraformaldehyde (PFA, EM grade powder), 10 mL water, 70 μL of 1 M NaOH. Weigh PFA powder in 50 mL self-standing centrifuge tube (Corning). Wear a mask and gloves when weighing paraformaldehyde to minimize exposure, as the powder is classified as carcinogenic. Add 10 mL water and 70 μL of 1 M NaOH. Shake gently and microwave for 10 s with the cap slightly loosened. Fully dissolve in a 37 °C water bath for 30–60 min with occasional shaking of the tube. Filter into a fresh 15 mL centrifuge tube using a sterile hydrophilic syringe filter with 0.2 μm pore size (Sartorius). Store solution for up to 7 days at 4 °C (see Note 3). 2. Phosphate buffer with lysine and 2% paraformaldehyde (PLP): 3 mL PBL, 1 mL 8% PFA stock solution. Make up freshly prior to use.

2.4  Antibodies

1. Primary antibodies: mouse monoclonal antibody anti-V5 (1:500 dilution in blocking buffer; Invitrogen) to visualize Cerulean expression (see Note 4). 2. Secondary antibodies: goat anti-mouse F(ab′)2 fragments conjugated with Cy5 or Alexa Fluor®647 (Jackson ImmunoResearch Laboratories) used as a dilution of 1:200 in blocking buffer (see Note 5).

2.5  Dissections, Immunostaining and Mounting

1. Stainless steel No. 5 forceps. 2. Glass embryo dish. 3. Mesh baskets. 4. 24-well multidish (e.g., Nunclon™ Δ surface). 5. 5. Terasaki plate (e.g., Nunc 60-well, MicroWell™ MiniTray). 6. Parafilm®. 7. Plastic box serving as a humidity chamber. 8. Sarstedt microtubes with O-ring screw caps. 9. Fine nylon mesh. 10. Rotating titer plate shaker (e.g., IKA® MTS 2/4 digital microtiter plate shaker). 11. Aluminum foil. 12. Vectashield (Vector Laboratories).

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13. Microscope slide (SuperFrost®). 14. Cover slip (18 × 18 mm, No. 1.5). 15. Soft modeling mass (e.g., Fimo). 16. Nail polish (clear). 17. Medical wipes. 18. Glass pipettes. 19. Sylgard (Dow Corning, 184 silicone elastomer kit). 20. Dissecting and fluorescence-dissecting microscopes. 2.6  Image Acquisition and Analysis

1. Confocal laser scanning microscope. 2. Long-range objectives (e.g., Leica 20× (0.7 NA) air, 40× (1.25 NA) and 100× (1.46 NA) oil immersion objectives). 3. Immersion oil (e.g., Zeiss Immersol™ 518F). 4. Confocal image acquisition software (e.g. Leica LAS software). 5. Image analysis software (e.g. Volocity Improvision PerkinElmer; Image J; Fiji including Simple Neurite Tracer plug-in).

3  Methods 3.1  Genetic Crosses and Clone Induction Protocol

1. Build and expand the driver stock, which contains both an hs-­ mFLP5 and a Gal4 transgene active in a cell type, brain area, or other tissue of interest (see Note 6). 2. Collect male and unfertilized female flies from parental stocks for about four days (see Note 7). Set up crosses in plastic vials containing standard cornmeal/agar medium with about 10–12 females and 5–6 males per vial. The crosses should be designed so that progeny will have one Gal4, one hs-mFLP5 and one FB1.0, FB1.1, or FB2.0 transgene. Note that for FB2.0, an additional transgene is required that expresses canonical FLP recombinase, such as hs-FLP1 to remove the stop cassette (see Figs.  1 and 4). FB and hs-mFLP5 transgenes are available as insertions on different chromosomes to facilitate genetic crosses (see Table 1). We usually set up three to five parallel crosses for a given experiment. After a period of 24-h of egg laying, transfer parents into fresh vials. Repeat this process for maximally 1 week. All vials are kept in an incubator at 25 °C. 3. To induce recombination events (see Fig. 4), progeny of crosses are heat shocked by placing the vials into a water bath with the temperature set to 37 °C (see Note 8). The water level should be sufficiently high, so that vials, when weighed down, are immersed well above the food level to ensure that also larvae leaving the medium are exposed to heat.

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Fig. 4 Example of experimental strategy to induce recombination events. After setting up or transferring the cross into a fresh vial (t = 0), flies are left to lay eggs for 1 day (t = 24 h after egg laying (AEL)). Subsequently, parents are transferred into a fresh vial or removed. Progeny in vials are heat shocked up to three times at 48, 72, and 96-h AEL in a 37 °C water bath. Brains can be dissected from larvae, staged pupae or adults. APF after puparium formation

3.2  Immunolabeling

1. Dissect 3rd instar larval, pupal, or adult brains (see Note 9) in drops of cold PBS using forceps on a dissecting pad. To maximize the number of dissected samples with recombination events, some Gal4 drivers make it possible to preselect flies with clones under a fluorescence-dissecting microscope. Transfer the brains into a glass embryo dish filled with PBS using forceps or a glass pipette. Keep the dish on ice until all samples of a given genotype are dissected (see Notes 10 and 11). 2. Transfer the brains into a mesh basket (see Note 12) positioned in one well of a 24-well multidish filled with 1 mL PLP. Fix for 1 h at room temperature (see Note 13). 3. Wash the brains by transferring the mesh baskets sequentially into three wells filled with 1 mL PBT each. Keep baskets in the first two wells for a few seconds each (short washes) and in the third well for at least 15 min (long wash) (see Note 14). 4. Block tissue for at least 15 min by transferring the baskets into a new well containing blocking buffer (10% NGS in PBT). 5. For incubation in primary antibody (see Note 15), transfer brains into the wells of a Terasaki plate, each filled with 10 μL of antibody diluted in blocking buffer. Distribute brains over multiple wells, with each well not containing more than ten brains (see Note 16). Add a stripe of moist filter paper on one side of the plate, close and seal tightly with Parafilm®. Place the plate into a humid chamber, such as a small closable box with a moist paper towel at the bottom. Keep on a gently horizontally rotating titer plate shaker overnight at 4 °C in a cold room.

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6. Return the brains into mesh baskets placed into 24-well plates filled with PBT. Wash for 15 min by transferring the baskets three times into fresh wells for two short and one long washes. 7. For incubation in secondary antibodies, transfer the baskets into a new well containing the antibody diluted in blocking buffer. Wrap the 24-well plate with aluminum foil and keep on a titer plate shaker for 2.5 h at room temperature. 8. Transfer baskets into fresh wells: two filled with PBT for two short washes, and two filled with PBS for one short and one long wash. In total, wash at least for 1 h. Immunolabeled brains should be stored in PBS at 4 °C, not in mounting media, to avoid softening of the tissue prior to mounting on slides. 3.3  Mounting Samples

1. Take immunolabeled brains with clones individually out of PBS and place in a drop of about 30 μL cold Vectashield (Vector Laboratories) on a precleaned microscope slide. 2. Take a cover slip (18 × 18 mm, No. 1.5) and place small pieces of soft modeling mass on the four corners of the cover slip to prevent squashing of brains. 3. Using forceps, position the cover slip with the modeling mass pieces facing down on the sample. Gently press the corners. Then place the slide under a fluorescence-dissecting microscope. 4. By pressing the forceps against the cover slip edges, move the cover slip gently up and down or left and right to roll at least one brain hemisphere into the correct orientation. Subsequently, brains can be gently flattened by pressing down the cover slip corners. Seal slide with clear nail polish. Store slides at 4 °C in the dark.

3.4  Image Acquisition and Analysis

1. Collect images using a confocal laser scanning microscope equipped with high-quality long-range objectives. For oil objectives, use immersion oil suitable for immunofluorescence microscopy recommended by the confocal microscope provider. Set up a confocal microscope imaging method that combines sequential and simultaneous scan modes (see Note 17). For each FP or dye, select a laser line suitable for optimal excitation, as well as detection windows or acousto-optical beam splitter (AOBS) settings for collecting specific emission signals. An example of such a method is provided in Table 2. For the original set of FB transgenes, mCitrine and Cy5/Alexa Fluor®647 signals are collected using a simultaneous scan mode, followed by sequential scans of EGFP and mCherry signals. For set B and C of FB transgenes, a fourth sequential scan is added to image mTurquoise/mTurquoise2. To obtain optimal signals and minimal cross talk between channels, adjust the

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Table 2 Example of a confocal microscopy scanning method for imaging FB transgenes

Scan

Fluorescent proteina/dye

PMTb (e.g.)

Excitation maximum (nm)

Emission maximum (nm)

Excitation laser line (nm)

Emission range imaged (nm)

1

mCitrine

2

516

529

514

525–565

no line above, one scan

Alexa Fluor®647

4

651

667

633

675–735

2

mCherry or Tag-RFP-T

3 (HyD) 587 555

610 584

561

572–630

3

EGFP

1

488

507

488

497–515

4

mTurquoise or mTurqouise 2

3 (HyD) 434

475

458

460–495

a Detailed information about the FPs used in the Flybow (FB) approach can be found in these references: EGFP [13], mCitrine [14], mCherry [11], and mTurquoise and mTurquoise 2 [16, 22] b PMT Photomultiplier tube as with the other table the last two lines should not be part of the table, they are comments

power of individual laser lines for each sample. The detection windows may also require fine-tuning depending on the experiment or microscope used. Assign a color to each channel, e.g., EGFP, green; mCitrine, yellow; mCherry or TagRFP-T, red; Cerulean-­V5/Cy5, medium blue; and mTurquoise or mTurquoise 2, cyan. 2. For samples displaying extensive bleed-through between channels, images can be processed using channel separation software. We use the Leica LAS AF suite channel separation tool for this purpose. In representative images of each detection channel, regions of interest with unambiguous strong but not saturated signals are manually selected for each FP or dye. Using these obtained values, the software algorithms subtract unspecific proportions of detected signals. 3. Confocal images can be further analyzed and processed using Volocity (Improvision Perkin Elmer) and Image J (Fiji) software to perform z projections of selected sections. Fiji can be used for brightness and contrast adjustments of the four or five channels in color composite images, as well as for subsequent conversion into RGB images. The Simple Neurite Tracer plugin is highly useful for the reconstruction of individual neurons from stacks.

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4  Notes 1. We store most of our solutions in autoclaved standard screw cap laboratory glass bottles. 2. We use higher-quality goat serum to extend the lifetime of solutions. 3. For optimal fixation of tissue, we use EM grade paraformaldehyde powder (Polysciences) in our protocol. Some i­ nvestigators freeze paraformaldehyde aliquots. However, we prepare fresh solutions at a weekly basis to obtain best staining results. 4. Although Cerulean had been described as the best CFP derivative with respect to brightness, quantum yield, and oligomerization properties [12], consistent with other studies [2] we observed that this FP is less suitable for studies in Drosophila because of low endogenous emission signals. Cerulean is therefore visualized using anti-V5 primary and Cy5 or Alexa Fluor®647 conjugated secondary antisera. 5. F(ab′)2 fragment-based secondary antibodies ensure even immunolabeling throughout the entire brain, because their smaller size facilitates penetration deeper into the tissue. 6. To build stable stocks, we prefer to combine hs-mFLP5 transgenes with the Gal4 driver instead of FB transgenes, as continuous low levels of FLP recombinase expression may cause transmittable recombination events in the germline. Stocks containing both hs-FLP1 and FB2.0 transgenes (see Table 1) should be continuously monitored. Note that the two hs-­ mFLP5 transgenes described in [3] are homozygous lethal insertions and are therefore kept over a balancer. 7. As the occurrence of hs-mFLP5-induced recombination events decreases with the parental age of flies, maximally 4-day old adult flies should be used for genetic crosses. The first emerging progeny of a cross should be prepared for analysis to maximize the number of samples with recombination events. In a 24-h egg collection, these correspond most closely to the progeny, at which the heat shock protocols were aimed. 8. Color outcomes are influenced by the time points, duration, and number of heat shocks. These parameters can be adjusted for each Gal4 line, cell or brain area of interest, and experimental aim. While early heat shocks lead to recombination events in dividing progenitors and thus tend to label larger lineagerelated groups of cells with the same FP, later or shorter heat shocks facilitate labeling of single cells. Repeated heat shocks further promote labeling of neurons or glia with different FPs. We observed that lineage-related cells born in a narrow time window are difficult to separate by the expression of different

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FPs using the FB approach. For studies in the optic lobe, the earliest time point at which larvae are subjected to heat shock is 48-h after egg laying (AEL). The time point at which crosses are transferred into a fresh vial is defined as t = 0. In a regime requiring multiple heat shocks, this procedure was repeated at 72 and 96 h. The length of heat shocks ranges from 30 to 45 min. When combining the FB approach with MARCM [25], it is useful to extend heat shocks to 90 min [3]. 9. For the visual system, large late 3rd instar larvae, which stopped wandering on the side of vials, are selected for dissections. To stage pupae, white pre-pupae are collected from vials at 1-h intervals, placed on grape-juice agar plates to avoid desiccation, stored in a 25 °C incubator and dissected at specific time points after puparium formation. Pharate adult flies are dissected shortly before eclosion to avoid tracheal filling. 10. Brains are dissected on plates coated with Sylgard® (Dow Corning, 184 silicone elastomer kit). We use the lids of Terasaki plates for this purpose. When transferring brains with a glass or plastic pipette, keep samples in the tip of the pipette, filled with PBS, to avoid losing them during the process. 11. Keep unfixed brains for no longer than 30 min on ice, as extended storage can cause connectivity defects. Fixed brains can be stored in PBT while completing all dissections for the day. 12. Mesh baskets are handmade using conical 1.5 mL Sarstedt microtubes with O-Ring Screw Caps and a fine nylon mesh. Cut microtubes at the bottom border of the ridged part using a pair of sharp scissors, so that the 24-well plate can be still closed with the lid. Carefully, cut off the top of the screw cap below the O-Ring using a sharp razor blade. Cut out a 2 × 2 cm square of nylon mesh. Place the mesh on top of the cut tube, and screw the cap ring in place while tightening the mesh. Remove all fabric on the outside of baskets using a razor blade. Newly made baskets should be extensively washed in PBT followed by deionized water to avoid sticking of brains. After each use, baskets should be thoroughly washed by soaking and rinsing them in deionized water. They can be reused for years. 13. Fixation for 1 h in PLP at room temperature is central to obtaining good signals both from FPs and fluorophore conjugated secondary antibodies. We observed that fixation using 4% PFA even for 30 min significantly quenched FP signals. 14. Even if solely endogenous FP signals are collected and immunolabeling steps are not required, brains should still be washed extensively in PBT after fixation before storage in PBS. Otherwise the tissue will shrink when transferred into mounting media.

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15. For the 2nd and 3rd sets of FB transgenes, immunolabeling with anti-V5 is not required. Instead, additional antibody stainings, e.g. mAb24B10 (1:75 dilution in blocking buffer; Developmental Studies Hybridoma Bank) can be performed and visualized with a Cy5 or Alexa Fluor®647-coupled secondary antibody. 16. To avoid damaging brains during the transfer into the Terasaki plate wells, position brains in the remaining liquid between slightly held apart or curved forceps tips. To minimize the transfer of excessive PBT, gently blot up the liquid held between the forceps arms using a rolled up corner of a medical wipe (e.g. Kimberly-Clark Professional). 17. We optimized imaging conditions for a Leica TCS SP5 II upright laser confocal microscope equipped with a resonant scanner and four photomultiplier tubes (PMTs). However, all confocal microscope models are suitable, which have the five listed laser lines, and whose detection windows can be adjusted. We typically collect image stacks using 1024 × 1024 pixels image size, 200 Hz line speed, and 5-line or 4-frame averages. To accelerate image acquisition, a bidirectional scan mode is used. Although not essential, the use of the resonant scanner provides an alternative way to increase the speed of image acquisition of large z stacks and thus to minimize photobleaching. When using the resonant scanner, images are collected at the fixed speed of 8 kHz and averaged 96 times. If the microscope is equipped with hybrid detection (HyD) GaAsP technology, scan the weakest fluorescent signals using the HyD PMT(s).

Acknowledgments We thank J. Goedhart for sharing the mTurquoise and mTurquoise2 cDNA. D. Hadjieconomou developed the original and C sets of FB transgenes. N. Shimosako generated the B set of transgenes and validated the hs-mFLP5MH insertions. The original FB approach was developed in collaboration with B.J. Dickson, S. Rotkopf, C. Alexandre, and D.M. Bell. This work was supported by the Francis Crick Institute, which receives its core funding from Cancer Research UK (FC001151), the UK Medical Research Council (FC001151), and the Wellcome Trust (FC001151), and by the UK Medical Research Council (U117581332). References 1. Livet J, Weissman TA, Kang H, Draft RW, Lu J, Bennis RA et al (2007) Transgenic strategies for combinatorial expression of fluorescent proteins in the nervous system. Nature 450(7166):56–62

2. Hampel S, Chung P, McKellar CE, Hall D, Looger LL, Simpson JH (2011) Drosophila Brainbow: a recombinase-based fluorescence labeling technique to subdivide neural expression patterns. Nat Methods 8(3):253–259

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3. Hadjieconomou D, Rotkopf S, Alexandre C, Bell DM, Dickson BJ, Salecker I (2011) Flybow: genetic multicolor cell labeling for neural circuit analysis in Drosophila melanogaster. Nat Methods 8(3):260–266 4. Richier B, Salecker I (2015) Versatile genetic paintbrushes: Brainbow technologies. Wiley Interdiscip Rev Dev Biol 4(2):161–180 5. Nern A, Pfeiffer BD, Rubin GM (2015) Optimized tools for multicolor stochastic labeling reveal diverse stereotyped cell arrangements in the fly visual system. Proc Natl Acad Sci U S A 112(22):E2967–E2976 6. Brand AH, Perrimon N (1993) Targeted gene expression as a means of altering cell fates and generating dominant phenotypes. Development 118(2):401–415 7. Voziyanov Y, Konieczka JH, Stewart AF, Jayaram M (2003) Stepwise manipulation of DNA specificity in Flp recombinase: progressively adapting Flp to individual and combinatorial mutations in its target site. J Mol Biol 326(1):65–76 8. Liaw CW, Zamoyska R, Parnes JR (1986) Structure, sequence, and polymorphism of the Lyt-2 T cell differentiation antigen gene. J Immunol 137(3):1037–1043 9. Zacharias DA, Violin JD, Newton AC, Tsien RY (2002) Partitioning of lipid-modified monomeric GFPs into membrane microdomains of live cells. Science 296(5569):913–916 10. Dietzl G, Chen D, Schnorrer F, Su KC, Barinova Y, Fellner M et al (2007) A genome-­ wide transgenic RNAi library for conditional gene inactivation in Drosophila. Nature 448(7150):151–156 11. Shaner NC, Campbell RE, Steinbach PA, Giepmans BN, Palmer AE, Tsien RY (2004) Improved monomeric red, orange and yellow fluorescent proteins derived from Discosoma sp. red fluorescent protein. Nat Biotechnol 22(12):1567–1572 12. Rizzo MA, Springer GH, Granada B, Piston DW (2004) An improved cyan fluorescent protein variant useful for FRET. Nat Biotechnol 22(4):445–449 13. Shaner NC, Steinbach PA, Tsien RY (2005) A guide to choosing fluorescent proteins. Nat Methods 2(12):905–909 14. Griesbeck O, Baird GS, Campbell RE, Zacharias DA, Tsien RY (2001) Reducing the environmental sensitivity of yellow fluorescent protein. Mechanism and applications. J Biol Chem 276(31):29188–29194

15. Shimosako N, Hadjieconomou D, Salecker I (2014) Flybow to dissect circuit assembly in the Drosophila brain. Methods Mol Biol 1082:57–69 16. Goedhart J, van Weeren L, Hink MA, Vischer NO, Jalink K, Gadella TW Jr (2010) Bright cyan fluorescent protein variants identified by fluorescence lifetime screening. Nat Methods 7(2):137–139 17. Hadjieconomou D, Timofeev K, Salecker I (2011) A step-by-step guide to visual circuit assembly in Drosophila. Curr Opin Neurobiol 21(1):76–84 18. Hayashi S, Ito K, Sado Y, Taniguchi M, Akimoto A, Takeuchi H et al (2002) GETDB, a database compiling expression patterns and molecular locations of a collection of Gal4 enhancer traps. Genesis 34(1-2):58–61 19. Timofeev K, Joly W, Hadjieconomou D, Salecker I (2012) Localized netrins act as positional cues to control layer-specific targeting of photoreceptor axons in Drosophila. Neuron 75(1):80–93 20. Apitz H, Salecker I (2018) Spatio-temporal relays control layer identity of direction-­ selective neuron subtypes in Drosophila. Nat Commun 9(1):2295 21. Shaner NC, Lin MZ, McKeown MR, Steinbach PA, Hazelwood KL, Davidson MW et al (2008) Improving the photostability of bright monomeric orange and red fluorescent proteins. Nat Methods 5(6):545–551 22. Goedhart J, von Stetten D, Noirclerc-Savoye M, Lelimousin M, Joosen L, Hink MA et al (2012) Structure-guided evolution of cyan fluorescent proteins towards a quantum yield of 93%. Nat Commun 3:751 23. Green EW, Fedele G, Giorgini F, Kyriacou CP (2014) A Drosophila RNAi collection is subject to dominant phenotypic effects. Nat Methods 11(3):222–223 24. Vissers JH, Manning SA, Kulkarni A, Harvey KF (2016) A Drosophila RNAi library modulates Hippo pathway-dependent tissue growth. Nat Commun 7:10368 25. Lee T, Luo L (1999) Mosaic analysis with a repressible cell marker for studies of gene function in neuronal morphogenesis. Neuron 22(3):451–461 26. Enriquez J, Venkatasubramanian L, Baek M, Peterson M, Aghayeva U, Mann RS (2015) Specification of individual adult motor neuron morphologies by combinatorial transcription factor codes. Neuron 86(4):955–970

Chapter 9 Live Cell Imaging of Neural Stem Cells in the Drosophila Larval Brain Karolina Miszczak and Boris Egger Abstract Live cell imaging gives valuable insights into the dynamic biological processes within and between cells. An important aspect of live cell imaging is to keep the cells under best physiological condition and to prevent abnormal cellular behavior, which might be caused by phototoxicity during microscopy. In this chapter we describe a protocol to visualize division patterns of neural stem cells in live whole mount brains of Drosophila larvae. We also present a newly developed live cell chamber that allows us to control the environmental air during live cell imaging. The protocol can be adapted to look at a wide range of cellular and tissue behavior in the Drosophila model system. Key words Neural stem cells, Drosophila, Brain, Live cell imaging

1  Introduction The Drosophila larval brain is a well-proven and informative model system to study the division pattern of neural stem cells. In the central brain, neural stem cells or neuroblasts undergo several rounds of asymmetric divisions thereby self-renewing and producing ganglion mother cells (GMCs). GMCs, which are intermediate progenitor cells, usually only divide once more to generate two sibling neurons or a neuron/glial cell pair [1]. Lateral to the central brain are the optic lobe anlagen that give rise to the visual system of the adult fly [2]. The developing optic lobe harbors symmetrically dividing neuroepithelial cells that progressively transform to asymmetrically dividing neuroblasts [3, 4]. Fixed tissue and live cell studies in Drosophila have greatly contributed to the understanding of the mechanisms regulating proliferation, self-­ renewing asymmetric cell division and differentiation of neural stem cells. Here we present a protocol to visualize live cell division patterns in the developing larval brain. Our methods are based on two main pieces of previous work [5, 6]. We further developed a live Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_9, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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cell chamber that allows us to look at dividing cells under different environmental air conditions, using small volumes of air flow. The chamber can for example be used to expose larval brains to different oxygen tensions and thereby allows to study the effects of hypoxia or hyperoxia on neural stem cell behavior.

2  Materials 2.1  Staged Larval Collection

1. Fly cages. 2. Apple juice embryo collection plates with wet yeast. 3. Petri dishes 35 mm filled with fly food and wet yeast.

2.2  Larval Brain Dissection

1. Binocular. 2. Fine forceps (Dumont no. 5, Fine Science Tools). 3. 200 μl pipette and sterile tips. 4. Paper tissue. 5. Petri dishes (plastic). 6. 1× Phosphate-buffered saline (PBS): 10 mM Na2HPO4, 2.68 mM KCl, 140 mM NaCl (PBS Tablets, Gibco, Thermo Fisher Scientific). pH 7.45. Sterilize by autoclaving.

2.3  Preparation of Conditioned Culture Medium

1. Schneider’s Drosophila medium (Gibco, Thermo Fisher Scientific). 2. Fetal bovine serum. 3. Third instar larval fat body tissue. 4. Phosphate-buffered saline (1× PBS). 5. Optional: 0.5 M ascorbic acid (Sigma-Aldrich).

2.4  Live Brain Sample Preparation

1. Low melting agarose (Sigma-Aldrich). 2. Schneider’s Drosophila Medium 1× (Gibco, Thermo Fisher Scientific). 3. Thermoblocks (2×). 4. Live cell microscopy chamber made from aluminum with plexiglass lid (see Fig. 1), further developed from [5, 7]. 5. Coverslips (22 × 22 mm, no.1.5, thickness 0.17 mm, Carl Roth GmbH). 6. Paint brush. 7. Vaseline.

2.5  Microscopy and Image Analysis

1. Spinning disk or single point scanning laser confocal microscope with inverted stand. 2. High-performance computer. 3. Image analysis software such as Fiji [8] or Bitplane Imaris.

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3  Methods 3.1  Staging of Larvae

1. Collect embryos on apple juice plates containing a drop of wet yeast for 4–6 h and let embryos develop at 25 °C (see Note 1). 2. 24 h after midpoint of egg collection, pick about 80–100 freshly hatched larvae and place them in a food plate containing a drop of wet yeast. 3. Let larvae develop at 25 °C to the appropriate larval stage (e.g. 72–96 h ALH, after larval hatching).

3.2  Dissection of Larval Brains

1. Pick the larvae from food plates and place them on a paper tissue soaked with PBS (see Note 2).

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2. Place several drops of 1× PBS in a circle on the inside of a plastic petri dish lid. Add one larva to each drop (see Fig. 2a). 3. To dissect larvae (72–96 h ALH), use fine forceps and the so-­ called “inverted sock” technique. Pull the larva in the middle apart and discard the posterior part. Grab the larval mouth hooks at the anterior half with one forceps and invert the larval cuticle over the tip of the forceps in a manner like to invert a sock. The interior organs of the larva are now on the outside and the brain can be further dissected. 3.3  Preparation of Conditioned Media

1. Dissect 10–20 third instar larvae as described above to collect fat body tissue (see Note 3). 2. Add fat body tissue to an Eppendorf tube containing 1 ml of sterile Schneider’s Drosophila medium. 3. Add 10 μl of fetal bovine serum (FBS) to the media. 4. Optional: add 1 μl of 0.5 M ascorbic acid to the media (see Note 4).

3.4  Preparation of Low Melting Agarose

1. Weight in 10 mg of low melting agarose to a 2 ml Eppendorf tube. 2. Add 1 ml of 1× PBS to obtain a 1% agarose stock solution. 3. Mix and heat up to 80 °C on a heat block (see Note 5). 4. Add 50 μl stock solution to 50 μl conditioned media to obtain a 0.5% agarose solution. 5. Mix and cool down to 37 °C on a heat block (see Note 6).

3.5  Mounting of Larval Brains

1. Place a drop of 1× PBS on a clean 22 × 22 mm cover slip. 2. Transfer three to four dissected brains to the drop and place the brain in the desired orientation on the cover slip (see Note 7). 3. Gently remove 1× PBS using a 200 μl pipette. 4. Gently pipette 100 μl of 0.5% melted agarose onto the brains. Quickly correct orientation of brains if necessary (see Fig. 2b). 5. Wait 5–10 min for the agarose to get solid. 6. Turn cover slip and place it onto the downside of the live cell chamber (see Fig. 2c). 7. Use a paint brush to seal the cover slip with preheated liquid Vaseline (see Fig. 2c and Note 8). 8. Turn chamber and add 0.5–1 ml conditioned medium. 9. Optional: if altered environmental conditioned are required close lid and flood chamber with desired gas mixture (see Note 9).

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Fig. 2 (a) Larvae are placed in a drop of 1× PBS in a petri dish lid and dissected using fine forceps. (b) Larval brains are transferred and positioned on a cover slip containing a drop of PBS. The liquid is pipetted off and replaced by low melting agarose. (c) After the low melting agarose has solidified the cover slip is inverted and placed on the opening of the live cell imaging chamber. The edges of the cover slip are sealed with liquefied Vaseline using a paint brush. (d) Conditioned Schneider’s medium is added to the live cell imaging chamber and the lid may be closed to control the air environment 3.6  Live Cell Imaging

1. Turn on and prepare microscope well in advance. 2. Place live cell chamber onto microscopy stage (see Fig. 2d). 3. Wait 15–30 min before imaging (see Note 10). 4. Adjust laser power, exposure time, frame rate, and z-stack (see Note 11). 5. Process and analyze movies on high-performance computer (see Fig. 3, and Note 12). 6. See Movies: Movie 1: Live whole mount larval brain lobe at third instar expressing insc-GAL4-driven UAS-mCD8-GFP (green) and UAS-­H2B-­mRFP1 (red). Shown is a single section with large central neuroblast cells and smaller progeny cells. Several neuroblasts undergo multiple rounds of asymmetric mitotic divisions. https://tube.switch.ch/videos/ fc51d03f Movie 2: Live whole mount larval brain lobe at third instar expressing insc-GAL4-driven UAS-mCD8-GFP (green) and UAS-­H2B-­mRFP1 (red). Single central brain neuro-

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Fig. 3 (a) Live whole mount larval brain lobe at third instar expressing insc-GAL4-driven UAS-mCD8-GFP (green) and UAS-H2B-mRFP1 (red). Shown is a single section with large central brain neuroblasts and smaller progeny cells. (b) Close up of a central brain neuroblast (asterisk) and progeny cells. At 15 min a GMC undergoes a mitosis (arrows). The neuroblast undergoes mitosis with prometaphase at 40 min, metaphase at 42 min, anaphase at 44 min, and beginning telophase at 45 min. At 1 h a new GMC is present (arrow). Scale bars are 10 μm

blasts that undergothree asymmetric divisions in the course of 3 h. https://tube.switch.ch/videos/0ca94b83 Live whole mount larval brain lobe at third instar expressing insc-­GAL4-­driven UAS-mCD8-GFP (green) and UASH2B-mRFP1 (red). Single central brain neuroblasts as shown in Fig. 3. https://tube.switch.ch/videos/b87cc0a6 (see Note 13).

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4  Notes 1. For more accurate staging make a 1 h pre-collection for eggs that have been retained and developed in the female abdomen before starting the experimental collection. 2. Larvae clean themselves of yeast and fly food by moving around on a wet paper tissue. 3. Fat body tissue is the source of a mitogen that stimulates divisions of neural stem and progenitor cells in the developing larval brain, possibly via the insulin signaling pathway [9–11]. 4. Ascorbic acid (Vitamin C) is a known antioxidant and can help to reduce phototoxicity by scavenging reactive oxygen species (ROS) [12]. 5. 1% stock solution of low melting agarose in 1× PBS can be stored for several days at 4 °C. Before use, melt agarose again at 80 °C and proceed with step 4 under Subheading 3.4. 6. 0.5% agarose solution should be cooled down to 37 °C or lower. Larval tissue might be harmed at higher temperature. 7. To visualize and investigate central brain neuroblasts orient the dorsal side of the brain lobes towards the cover slip. To visualize optic lobe neuroepithelial cells orient the lateral side of brain lobes towards the cover slip. 8. Heat up Vaseline well in advance to 80 °C until it becomes completely liquid. Carefully, seal the edges of the cover glass to the live cell chamber using a paint brush. 9. Use minimal gas flow to fill the lid closed chamber. If pressure is too high the cover slip will be moved or pushed away. 10. Wait several minutes before imaging to let the tissue acclimatize and to reduce drift. We use a spinning disk confocal microscope assembled by Visitron and equipped with solid-state lasers 488 nm and 561 nm, a CSU-W1 disk unit (Yokogawa), a Nikon Ti-2 stand with 40× oil immersion objective and an Evolve 512 Delta EMCCD camera (Photometrics). 11. A frame stack of 18 sections (z = 1 μm) were taken at 1 min time intervals for 6 h. Nikon’s Perfect Focus was used to correct for drift of the cover slip in z. 12. Movies were processed with Imaris Bitplane on a high-­ performance computer. Processor: AMD Threadripper 1920X (12 cores, 3.5 GHz), Memory: 128 GB DDR4, Storage Fast: 2× 1 TB Samsung 960 Pro (read 3.5 GB/s, write (2.5 GB/s), Graphics Card: MSI GeForce GTX 1070 AERO (8 GB), Display: BenQ PD3200U 32″ (16:9, 100% sRGB).

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13. Movies can be found online: Movie 1 https://tube.switch.ch/videos/fc51d03f Movie 2 https://tube.switch.ch/videos/0ca94b83 Movie 3 https://tube.switch.ch/videos/b87cc0a6 The movies show dividing central brain neuroblasts labeled by inscuteable-­GAL4 (insc-GAL4) [13]-driven UAS-mCD8-GFP and UAS-Histone2B-mRFP1 [14]. mCD8-GFP reveals cell membranes, whereas Histone2B-mRFP1 localizes to the DNA and reveals nuclei.

Acknowledgments We thank Jean-Daniel Niederhäuser for the construction of the live cell chamber and we thank Clemens Cabernard for advising us on live cell imaging protocols. References 1. Doe CQ (2008) Neural stem cells: balancing self-renewal with differentiation. Development 135:1575–1587 2. Apitz H, Salecker I (2014) A challenge of numbers and diversity: neurogenesis in the Drosophila optic lobe. J Neurogenet 28:233–249. https:// doi.org/10.3109/01677063.2014.922558 3. Egger B et al (2007) Regulation of spindle orientation and neural stem cell fate in the Drosophila optic lobe. Neural Develop 2:1 4. Yasugi T et al (2008) Drosophila optic lobe neuroblasts triggered by a wave of proneural gene expression that is negatively regulated by JAK/STAT. Development 135:1471–1480 5. Cabernard C, Doe CQ (2013) Live imaging of neuroblast lineages within intact larval brains in Drosophila. Cold Spring Harb Protoc 2013:970–977. https://doi.org/10.1101/ pdb.prot078162 6. Tsao CK et al (2017) Long-term live imaging of Drosophila eye. Disc J Vis Exp. https://doi. org/10.3791/55748 7. Kiehart DP et al (1994) High-resolution microscopic methods for the analysis of cellular movements in Drosophila embryos. Methods Cell Biol 44:507–532 8. Schindelin J et al (2012) Fiji: an open-­source platform for biological-image analysis. Nat Methods 9:676–682. https://doi. org/10.1038/nmeth.2019

9. Britton JS, Edgar BA (1998) Environmental control of the cell cycle in Drosophila: nutrition activates mitotic and endoreplicative cells by distinct mechanisms. Development 125:2149–2158 10. Chell JM, Brand AH (2010) Nutrition-­ responsive glia control exit of neural stem cells from quiescence. Cell 143:1161–1173. https://doi.org/10.1016/j. cell.2010.12.007 11. Sousa-Nunes R et al (2011) Fat cells reactivate quiescent neuroblasts via TOR and glial insulin relays in Drosophila. Nature 471:508– 512. https://doi.org/10.1038/ nature09867 12. Knight MM et al (2003) Live cell imaging using confocal microscopy induces intracellular calcium transients and cell death. Am J Physiol Cell Physiol 284:C1083–C1089. https://doi.org/10.1152/ ajpcell.00276.2002 13. Luo L et al (1994) Distinct morphogenetic functions of similar small GTPases: Drosophila Drac1 is involved in axonal outgrowth and myoblast fusion. Genes Dev 8:1787–1802 14. Langevin J et al (2005) Lethal giant larvae controls the localization of notch-signaling regulators numb, neuralized, and Sanpodo in Drosophila sensory-organ precursor cells. Current Biol 15:955–962. https://doi. org/10.1016/j.cub.2005.04.054

Chapter 10 CRISPR/Cas9 Genome Editing to Study Nervous System Development in Drosophila Cornelia Fritsch and Simon G. Sprecher Abstract Continuous implementation of new techniques allowing increasingly precise genetic manipulations makes the fruit fly Drosophila melanogaster an impacting model to study the nervous system. While transgenic approaches have been heavily used to investigate how the brain develops, genome editing has been notoriously hard in the fruit fly. The advent of versatile CRISPR/Cas9-based genome editing techniques allow the generation of engineered loci using homologous repair to replace the endogenous genome sequence with a designed template of interest. We here provide a protocol to generate an FRT/FLP-based conditional GFP or HA-flagged gene knockout. Key words Drosophila, Brain, Genome editing, CRISPR, Conditional alleles

1  Introduction Conditional knockout experiments allow to study the function of a gene in a specific organ, tissue, or cell type at a defined time-point [1–4]. Several techniques have been well established in Drosophila that allow controlled misexpression or knockdown of a gene of interest [5]. Here we use a combination of the CRISPR/Cas9 technique [6–17], the UAS/Gal4 system [18, 19], and FRT-­ mediated recombination [20, 21] to tag an endogenous gene with either GFP [22] or HA, at the same time flanking it with FRT sites for a cell- or tissue-specific knockout mediated by the UAS/Gal4 system and temporarily controlled by temperature sensitive Gal80. Any gene of interest can be modified due to the presence of CRISPR sites in its exons and introns. We have generated three cloning vectors in which GFP or 3xHA is flanked by two FRT sites (Fig. 1a–c). Most Drosophila genes have several exons separated by small introns. Our cloning vectors allow tagging of the gene of interest at the N-terminus, the C-terminus or internally. Figure 1a shows an example where the first three coding exons of a gene are cloned into pBsFGF in frame with GFP generating an N-terminal Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_10, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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Fig. 1 Cloning, genomic insertion, and deletion strategies for CRISPR-mediated introduction of a genomic knockout construct. (a–c) Cloning of different fragments of a gene into the template plasmids. The coding sequence is shown in red, noncoding exonic sequences are shown in purple. Two CRISPR sites (orange triangles) should be selected flanking the region that will be knocked-out. These CRISPR sites will be replaced by FRT sites (blue triangles in the plasmids). Genomic fragments that should be in frame with the reporter (GFP or HA) are shown in green;other genomic fragments are shown in blue. (a) Generation of an N-terminal GFP fusion in the pBsFGF plasmid. The first CRISPR site should be located within an intron upstream of the first coding exon and the

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GFP-fusion directly flanked by FRT sites. In this case the Start codon of the gene of interest is replaced by the GFP coding sequence directly followed by an FRT site. Upon Flp recombination only the GFP coding sequence will be removed. This means that the gene of interest could still be translated from an AUG codon further downstream generating a truncated version of the protein. Figure 1b shows an example where the GFP-tag gets inserted into the coding sequence. In this case, the sequence inserted between the first FRT-site and GFP, as well as the sequence inserted between GFP and the second FRT-site have to be in frame with GFP. To allow more flexibility, we added some restriction sites between GFP and the second FRT site in pBsFGF∗ (Fig. 1b). In this case the coding sequence of the gene of interest located between the two FRT-sites will be excised along with the GFP coding sequence upon FRT-mediated recombination. Both vectors could also be used for C-terminal tagging as shown for the pBsF3xHAF vector (Fig. 1c). In this case the last coding exon(s) are cloned between the first FRT-site and the tag, replacing the stop codon of the gene of interest with the tag cloned in frame with the

Fig. 1 (continued) second CRISPR site should be located near the start codon. The upstream homology arm should be cloned before the first FRT-site. The sequence to be excised should be cloned between the upstream FRT-site and GFP The downstream homology arm should be cloned after the second FRT-site in frame with GFP and lacking the start codon. (b) Generation of an internal GFP fusion in the pBsGFG∗ plasmid. The first CRISPR site should be located within an intron upstream of the first coding exon and the second CRISPR site should be located in and intron downstream of the first coding exon. The upstream homology arm should be cloned before the first FRT-site. One part of the sequence to be excised should be cloned between the upstream FRT-site and GFP in frame with GFP; the second part should be cloned between GFP and the second FRT-site in frame with GFP. The downstream homology arm should be cloned after the second FRT-site. (c) Generation of a C-terminal HA fusion in the pBsF3xHAF plasmid. The first CRISPR site should be located in an intron; the second CRISPR site should be located near the stop codon. The upstream homology arm should be cloned before the first FRT-site. The sequence to be excised should be cloned between the upstream FRT-site and the HA-tag in frame with the tag. The downstream homology arm should be cloned after the second FRT-site. (d) CRISPR mediated insertion of the template. The template plasmid shown here has an internal GFP-fusion within the first coding exon with FRT sites flanking the first and second coding exons. It is injected into nosCas9 flies along with a plasmid expressing gRNAs for both CRISPR sites. Cas9 nuclease is guided to the two sites to generate two double-­strand breaks. DNA repair by homologous recombination using the injected plasmid as template will lead to insertion of the GFP-tagged sequence flanked by the two FRT-sites. Since the CRISPR sites have been replaced by FRT, Cas9 will no longer cut in this region. (e) UAS-Gal4 driven Flp expression to delete the FRT-flanked sequence. Transgenic flies expressing Gal4 (green) under the control of a celltype-specific enhancer (orange), Gal80ts (red) under the control of a tubulin promoter (cyan), Flp (blue) under the control of an upstream activating sequence (UAS) element (green dashed line), and containing the knockout construct, are kept at low temperature (18 °C) during development to keep Gal4 inactive and prevent Flp activation. Adult flies are shifted to 29 °C resulting in Gal4 activation by Gal80 degeneration and expression of Flp. Flp expression leads to FRT recombination and deletion of the FRT-flanked sequence removing the tag and a part of the coding sequence of the gene of interest

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coding sequence. Upon Flp recombination the last coding exons will be removed along with the tag generating a truncated version of the protein. For adding the tagged sequences into the fly genome, the CRISPR sites should not be present in the resulting template plasmid, since that would lead to Cas9 cutting the template. Therefore, the selected CRISPR sites should be located in noncoding parts of the gene of interest where they can be replaced with the sequence of the two FRT-sites. For an N-terminal tagging the first CRISPR site should be located within 1 kb upstream of the start codon of the gene of interest, and the second CRISPR site should be located within 1–40 bp upstream of the start codon (Fig. 1a). For an internal tagging, both CRISPR sites should be located around 1 kb apart from each other within two introns flanking the coding exon in which the tag will be inserted (Fig. 1b). For a C-terminal tagging the first CRISPR site should be located about 1 kb upstream of the Stop codon, and the second CRISPR site should be located 1–40 bp downstream of the stop codon (Fig. 1c). Three genomic fragments should be cloned into the vector: the first fragment spanning 1–2 kb of sequence upstream of the first CRISPR site to provide the upstream homology arm for homologous recombination; the second fragment containing the sequence between the two CRISPR sites that will be flanked by FRT sites; and the third fragment spanning 1–2 kb of sequence downstream of the second CRISPR site to provide the downstream homology arm for homologous recombination. When using the pBsFGF∗ vector, the central fragment can be divide into two fragments due to the presence of additional restriction sites between GFP and the second FRT-site (Fig. 1b). The resulting plasmids can be used as a template for homologous repair after DNA double-strand break (Fig. 1d). Here we show integration of the template plasmid generated in Fig. 1b as an example. The two guide RNAs (gRNA1 and gRNA2) direct Cas9 endonuclease to the two selected CRISPR sites within the gene of interest. Two double-strand breaks are introduced by Cas9. The plasmid serves as a template for the cell-endogenous repair mechanism replacing the sequence located between the two CRISPR sites with the GFP-tagged and FRT-flanked version provided by the plasmid that can no longer be cut due to the replacement of the CRISPR sites by FRT sites (Fig. 1d). The resulting flies will now express a GFP-tagged version of the gene of interest. The FRT sites can be used to remove the FRT-flanked and tagged fragment in a specific cell type and at a specific time during or after development (Fig. 1e). Gal4 is a transcription factor binding specifically to an upstream activating sequence (UAS). In Drosophila many Gal4 lines have been generated expressing Gal4 under the control of various cell type-specific enhancers, and thus, providing spatial regulation of Gal4 production [23, 24]. Temporal

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regulation is achieved by combination with a transgenic line ubiquitously expressing a temperature sensitive version of the Gal4 inhibitor Gal80 (Gal80ts). At low temperature (18 °C) Gal80 represses Gal4 activity preventing the expression of UAS-regulated genes. When the flies are shifted to high temperature (29 °C), Gal80 is degraded allowing Gal4 to activate the expression of UAS-regulated genes. Thus, Gal4 can regulate the expression of a UAS-Flp transgene in a cell type-specific manner during a defined time interval. Flp is a recombinase that recognizes FRT sites in the genome. The sequence flanked by these FRT sites will be deleted upon FRT activation. Thus, when all four constructs are combined in a fly, Flp activity will lead to deletion of the FRT-flanked fragment of the gene of interest in a specific cell type (regulated by the Gal4 enhancer) and at a specific time (determined by the Gal80 inactivation upon shifting the flies to 29 °C). Successful knockout of the gene of interest will lead to a loss of the inserted tag, which can be verified by antibody staining.

2  Materials 2.1  General Material for Molecular Biology

1. 1.5 ml microtubes, lab glass ware, 15 ml round bottom tubes. 2. 0.2 ml PCR tubes. 3. Micropipettes and pipette tips (20 μl, 200 μl, 1000 μl). 4. Table top centrifuge. 5. Spectrophotometer/nanodrop. 6. PCR machine. 7. Water baths (37 °C, 42 °C). 8. 37°C bacteria shaker. 9. 37 °C incubator 10. Heat block. 11. Vortex. 12. Agarose-gel electrophoresis set-up (chambers, combs, power-­ pack, documentation system). 13. Magnetic stirrer. 14. Microwave oven. 15. Ice. 16. Kits for PCR-purification, Midipreps (e.g. QIAGEN)

Gel-purification,

Mini-

and

17. Gibson Assembly kit. 18. Different restriction enzymes, Taq polymerase, Pfu polymerase, T4 DNA-ligase, dNTP mix. 19. Agarose.

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20. 0.5× TBE buffer (pH 8.0): Tris–HCl 44.5 mM, Boric acid 44.5 mM, EDTA 1 mM. 21. Ethidiumbromide. 22. DNA ladder. 23. Competent E.coli cells. 24. LB medium. 25. Ampicillin. 26. LB-ampicillin plates. 2.2  Genomic DNA of Cas9 Flies

1. Transgenic flies expressing Cas9 in their germline are available at the “Bloomington Drosophila Stock Center” (bdsc. indiana.edu). 2. In our lab we are using M{nos-Cas9.P}ZH-2A flies (Bloomington stock number 58491) expressing Cas9 under the control of the nanos promoter. 3. Solution A: made of Tris–HCl 0.1 M (pH 9.0) EDTA 0.1 M and SDS 1%. 4. Phenol-Chloroform: 1:1 shake and spin for 10 min at 4000 rpm (1700 × g). 5. Potassium acetate (KAc) 8 M. 6. Isopropanol. 7. 70% Ethanol 8. Tris-EDTA buffer (TE): Tris–HCl 10 mM (pH 8.0), EDTA 1 mM. 9. Micropestle.

2.3  Plasmids

1. We generated different plasmids in our lab, which allow cloning of genomic sequences between two FRT sites, fusing the coding sequence with a GFP- or HA-tag, and flanking it with up- and downstream genomic sequence (see Note 1). (a) pBsFGF (Fig.  2a) (see Note 2). (b) pBsFGF∗ (Fig. 2b) (see Note 3). (c) pBsFHAF (Fig.  2c) (see Note 4). 2. For the guide RNAs preparation use “pCFD4-U6:1_ U6:3tandemgRNAs” from Simon Bullock’s lab; addgene number 49411 [13].

Fig. 2 (continued) depicted as a green box with the first and last two codons written, the 3×HA sequence is shown in magenta. Various restriction enzymes can be used for cloning genomic DNA fragments upstream of the first FRT site, between the first FRT site and GFP, between GFP and the second FRT site (only for plasmid pBsFGF∗), after the second FRT site. The restriction sites are indicated above the sequence at the position of the cut in the coding strand. Unique restriction sites are shown in bold letters. The binding sites for standard primers are depicted as arrows; or dashed arrows if their sequence is not fully shown in this figure

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a

pBsFGF M13 rev

SacI

T3 promoter

SacII

NotI

XbaI

ACAGGAAACAGCTATGACCATGATTACGCCAAGCTCGAAATTAACCCTCACTAAAGGGAACAAAAGCTGGAGCTCCACCGCGGTGGCGGCCGCTCT SpeI BamHI XmaI SmaI

PstI

BglII

StuI

XbaI

PmeI

AGAACTAGTGGATCCCCCGGGCTGCAGCAGATCTGAAGTTCCTATACTTTCTAGAGAATAGGAACTTCTTCGAGAG GCC TGT TTA AAC GAT A C L N D AgeI

NcoI

XbaI

GFP

CCA CCG GTC GCC ACC ATG GTG..........TAC AAG GAA TTG AAG TTC CTA TAC TTT CTA GAG AAT AGG AAC TTC P P V A T M V Y K E L K F L Y F L E N R N F HindIII

ClaI

SalI

XhoI

Acc65I

ApaI

KpnI

AAG CTT ATC GAT ACC GTC GAC CTC GAG GGG GGG CCC GGT ACC CAA TTC GCC CTA TAG TGAGTCGTATTACAATTCACT K L I D T V D L E G G P G T Q F A L * M13 for T7 promoter

b

pBsFGF*

M13 rev

SacI

T3 promoter

SacII

NotI

XbaI

ACAGGAAACAGCTATGACCATGATTACGCCAAGCTCGAAATTAACCCTCACTAAAGGGAACAAAAGCTGGAGCTCCACCGCGGTGGCGGCCGCTCT SpeI BamHI XmaI SmaI

PstI

BglII

StuI

XbaI

PmeI

AGAACTAGTGGATCCCCCGGGCTGCAGCAGATCTGAAGTTCCTATACTTTCTAGAGAATAGGAACTTCTTCGAGAG GCC TGT TTA AAC GAT A C L N D AgeI

NcoI

EcoRI

EcoRV

HindIII

XbaI

GFP

CCA CCG GTC GCC ACC ATG GTG..........TAC AAG GAA TTC GAT ATC AAG CTT GAA GTT CCT ATA CTT TCT AGA P P V A T M V Y K E F D I K L E V P I L S R SalI

XhoI

ApaI

Acc65I KpnI

GAA TAG GAACTTCGTCGACCTCGAGGGGGGGCCCGGTACCCAATTCGCCCTATAGTGAGTCGTATTACAATTCACTGGCCGTCGTTTTACAACGT E * T7 promoter M13 for

pBsFHAF

c M13 rev

SacI

T3 promoter

SacII

NotI

XbaI

CAGGAAACAGCTATGACCATGATTACGCCAAGCTCGAAATTAACCCTCACTAAAGGGAACAAAAGCTGGAGCTCCACCGCGGTGGCGGCCGCTCT SpeI BamHI XmaI SmaI

PstI

BglII

StuI

XbaI

PmeI

AGAACTAGTGGATCCCCCGGGCTGCAGCAGATCTGAAGTTCCTATACTTTCTAGAGAATAGGAACTTCTTCGAGA GGC CTG TTT AAA CGA G L F K R AgeI

NcoI

TCC ACC GGT CGC CAC CAT GGT TAC CCA TAC GAT GTT CCT GAC TAT GCG GGC TAT CCC TAT GAC GTC CCG GAC S T G R H H G Y P Y D V P D Y A G Y P Y D V P D HindIII

BamHI

XbaI

TAT GCA GGA TCC TAT CCA TAT GAC GTT CCA GAT TAC GCT GCA AGC TTG AAG TTC CTA TAC TTT CTA GAG AAT Y A G S Y P Y D V P D Y A A S L K F L Y F L E N SalI

XhoI

ApaI

Acc65I

KpnI

AGG AAC TTC GTC GAC CTC GAG GGG GGG CCC GGT ACC CAA TTC GCC CTA TAG TGAGTCGTATTACAATTCACTGGCCGTC R N F V D L E G G P G T Q F A L * M13 for T7 promoter

Fig. 2 Multiple cloning sites (mcs) of the three plasmids for the cloning of CRISPR templates for tagging and FRT mediated deletion. (a) mcs sequence of pBsFGF, (b) mcs sequence of pBsFGF∗, (c) mcs sequence of pBsF3xHAF. The DNA sequence of the coding strand flanking the tags is separated into triplets in the same frame as the tag with the encoded amino acids shown below each triplet. FRT sites are shown in blue, GFP is

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2.4  PCR Primers

1. One primer pair to insert the two gRNA sequences into the pU6 expression vector (see Subheading 2.4) by Gibson Assembly [25] (see Note 5). 2. One primer pair to amplify 1–2 kb of flanking genomic sequence upstream of the insertion site for the first FRT sequence (see Notes 6 and 7). 3. One primer pair to amplify the sequence that should be excisable (flanked by FRT sites and fused in frame with GFP (or HA)) (see Notes 6 and 8). 4. One primer pair to amplify 1–2 kb of flanking genomic sequence downstream of the insertion site for the second FRT sequence (see Notes 6 and 9). 5. Two primer pairs to test the insertion of the FRT sites and the tag into the genome (see Note 10).

2.5  Injection

1. Stereomicroscope. 2. Microscope with 10× objective, movable stage (Fig. 3a). 3. Micromanipulator (Fig. 3a). 4. Needle holder connected to a 50 ml syringe via a rubber tube (Fig. 3a, b). 5. Cage (plexiglass, 9 cm diameter, 13 cm height, covered with metal grid, Fig. 3c). 6. Plastic petri dishes (94 mm diameter). 7. Bleach. 8. Dechorionation cage: 50 ml Falcon tube with a 2 cm hole in the cap, cut open at the 40 ml mark, 3 × 3 cm square of nylon mesh (sefar 03–100/44) screwed with the cap onto the tube (Fig. 3d). 9. Cobalt(II)chloride crystals. 10. Voltalef 10S oil. 11. Borosilicate thin wall capillaries with filament (Harvard apparatus 30-0039). 12. Needle puller. 13. Tape (SCOTCH or TESA) dissolved in heptane. 14. Fine brush, preparation needle, Pasteur pipettes, cover slips (24 × 24), microscope slides, plastic tube.

2.6  Apple Juice Plates

1. 1.5 l H2O. 2. 40 g agar. 3. 20% Nipagin (dissolved in Ethanol). 4. Magnetic stir bar.

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a

c

b

d

e embryo micropyle (anterior end)

cover slip glue agar slice

lined up embryos

Fig. 3 Injection devices. (a) Microscope with 10× objective (middle), micromanipulator (right), syringe (left) connected to the needle holder (arrow). (b) syringe connected to the needle holder. (c) Fly cage to collect eggs on an apple juice plate. (d) Dechorionation cage consisting of a fine mesh and a cut Falcon tube and lid with hole. (e) Dechorionated embryos are lined up on an agar slice with the anterior end (micropyle) pointing to the edge of the slice. A cover slip with glue is used to pick up the lined-up embryos so that their posterior ends are pointing towards the edge of the cover slip

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5. 2 l glass bottle. 6. 500 ml apple juice. 7. Dry yeast. 8. Plastic petri dishes (5 cm diameter). 2.7  Establishment of Lines

1. Balancer stocks for the different chromosomes. 2. Vials with fly food. 3. Fly-station (Stereomicroscope, CO2-pad). 4. Brush. 5. Marker pens. 6. Fly incubators (18 °C, 25 °C, 29 °C).

2.8  PCR-Screening and Sequence Verification

1. Primers flanking the inserted FRT sites. 2. Squish buffer: Tris–HCl 10 mM pH 8.0, EDTA 1 mM, NaCl 25 mM. 3. ProteinaseK. 4. PCR reaction mix: 32 μl water, 5 μl 10× buffer 5 μl forward primer (10 mM stock), 5 μl reverse primer (10 mM stock), 1 μl dNTP mix (10 mM each), 1 μl Cas9 genomic DNA (see Note 11), 1 μl Taq polymerase (see Note 12).

2.9  Conditional Knockout

1. UAS-Flp flies. w[1118]; P{y[+t7.7] w[+mC]=20XUAS-FLPG5.PEST} attP40 (BDSC stock 55806) w[1118]; P{y[+t7.7] w[+mC]=20XUAS-FLPD5.PEST} attP2 (BDSC stock 55804) 2. Appropriate Gal4 line(s).

3. Gal80ts flies. w[∗]; P{w[+mC]=tubP-GAL80[ts]}D20; TM2/TM6B, Tb [1] (BDSC stock 7019) w[∗]; sna[Sco]/CyO; P{w[+mC]=tubP-GAL80[ts]}7 (BDSC stock 7018)

3  Methods 3.1  Preparation of Apple Juice Plates

1. Add magnetic stir bar to 1.5 l H2O and 40 g agar in glass bottle. 2. Autoclave. 3. Let cool down to 60 °C on magnetic stirrer.

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4. Add 500 ml apple juice. 5. Slowly add 20 ml of Nipagin. 6. Pour into plastic petri dishes. 7. Solidified plates can be stored at 4 °C. 8. Dissolve dry yeast in water until it has the consistency of mayonnaise. 9. Streak yeast in a crescent shape onto some of the plates. 3.2  Preparation of Genomic DNA

1. Collect 25 nos-Cas9 flies in a 1.5 ml tube—keep on ice (see Note 11). 2. Add 250 μl of solution A. 3. Homogenize the flies with a micropestle—put back on ice. 4. Incubate for 30 min at 70 °C. 5. Add 35 μl of KAc shake (no vortexing). 6. Incubate for 30 min on ice. 7. Spin for 15 min at 13,000 rpm (18,000 × g). 8. Move supernatant to a new tube (leave back any precipitate or interphase). 9. Add 1 volume of Phenol-Chloroform to the supernatant (ca. 250 μl) shake thoroughly (no vortexing). 10. Spin for 5 min at 13,000 rpm (18,000 × g). 11. Repeat steps 8–10. 12. Move supernatant to a new tube. 13. Add 150 μl of Isopropanol and shake. 14. Spin for 5 min at 10,000 rpm (10,000 × g). 15. Suck off supernatant (don’t lose pellet!) 16. Wash the pellet with 1 ml 70% Ethanol. 17. Spin for 5 min at 13,000 rpm (18,000 × g). 18. Dry the pellet. 19. Resuspend the pellet in 100 μl of TE.

3.3  PCR on Genomic DNA of Cas9 Flies

1. Set up three PCR reactions: PCR amplification for the upstream homology arm. PCR amplification for the downstream homology arm. PCR amplification for the sequence that will be put between the FRT sites.

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We use the following PCR program: 92 °C

2 min

92 °C

30 s

57 °C

30 s

72 °C

1–5 min

72 °C

10 min

30 cycles  

2. After the PCR, run 5 μl of the reaction on a 1% agarose gel (in TBE, with 2–3 μl of Ethidiumbromide; at 100-130 V). 3. Purify the PCR products using a PCR purification kit (the DNA should be eluted in 45 μl elution buffer). 3.4  Cloning into Knockout Plasmid

1. Digest the PCR products with the appropriate restriction enzymes, along with pBluescript or another cloning vector (see Note 13): 45 μl purified PCR product or plasmid DNA (0.1  μg/μl in ddH2O), 5 μl 10× restriction buffer, 1 μl enzyme1, 1 μl enzyme2, incubate for 2 h at 37 °C. 2. Optional: run digested bands on a 1% agarose gel and cut them out. 3. Purify the digested DNA using PCR purification kit (or a gel extraction kit), 4. Ligate the PCR products into the cloning vector digested with the same enzyme combination, respectively: 1–7 μl digested PCR product (use a molar ratio of PCR product to vector of 3:1), 1–2 μl digested vector (see Note 14 for control), 1 μl 10× ligation buffer, 1 μl T4 ligase, fill to 10 μl with ddH2O, incubate for 2 h at room temperature (or overnight at 15 °C), 5. Transform the ligations into competent E. coli cells: 100 μl competent cells (thawed on ice) plus 5 μl ligation reaction. 6. Incubate in ice for 20–30 min. 7. Heatshock for 60 seconds in a 42 °C waterbath. 8. Add 1 ml LB medium (without antibiotics). 9. Incubate for 30–60 min at 37 °C. 10. Spread onto LB-agar plates (with antibiotic, see Note 15). 11. Incubate over night at 37 °C. 12. Pick 4–12 single colonies per plate using a toothpick or a pipette tip and inoculate each colony in a 14 ml round bottom tube containing 2 ml of LB medium (with antibiotic, see Note 15). 13. Incubate overnight in a 37 °C shaker. 14. Purify plasmids using a Miniprep kit (see Note 16).

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15. Test digest each Miniprep with the same combination of restriction enzymes that were used for cloning: 7 μl ddH2O, 1 μl 10x reaction buffer, 2 μl miniprep DNA, 0.1 μl enzyme1, 0.1 μl enzyme 2, incubate for 1 hour at 37 °C. 16. Run on a 1% agarose gel. 17. Send for sequencing with standard primers matching the cloning vector used (for pBluescript use for example M13 and M13r). 18. Compare your sequencing results with the database sequence (see Notes 11 and 12). 19. Assemble all three fragments in one of the template plasmids following the same procedure as for cloning into the cloning vector (steps 1–8 of this section). Make sure that the coding sequence of your gene of interest is in frame with GFP (or 3×HA). 3.5  Gibson Assembly of gRNA Plasmid

1. Digest the “pCFD4-U6:1_U6:3tandemgRNAs” plasmid with BbsI (see Note 17): 30–40 μl ddH2O, 5 μl 10× restriction buffer, 5–15 μl plasmid (use 10 μg of DNA), 1 μl enzyme1, 1 μl enzyme2. 2. Incubate for 2 h at 37 °C. 3. Load the entire digestion reaction on a 1% agarose gel (in TBE, with 2-3 μl of Ethidiumbromide; at 100–130 V) to separate the bands. 4. Cut out the bands and purify them from the gel (using a gel extraction kit). 5. Use the smaller (493 bp) fragment containing the dU6–3 promoter as template for PCR with the gRNA primers (see Note 18). 6. PCR reaction mix: 32 μl water, 5 μl 10× buffer, 5 μl forward primer (10 mM stock), 5 μl reverse primer (10 mM stock), 1 μl dNTP mix (10 mM each), 1 μl pU6–3 promoter fragment (see Note 18), 1 μl Pfu polymerase. 7. PCR program: 95 °C

2 min

95 °C

30 s

57 °C

30 s

72 °C

1 min

72 °C

10 min

30 cycles  

8. After the PCR, run 5 μl of the reaction on a 1% agarose gel (in TBE, with 2–3 μl of Ethidiumbromide; at 100–130 V).

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9. Purify the PCR product using a PCR purification kit (the DNA should be eluted in 30 μl elution buffer). 10. Determine the concentration of the purified PCR product and the BbsI-digested plasmid fragment (6718 bp). 11. Set up Gibson Assembly reaction according to manufacturer’s instructions. 12. Incubate samples on a heating block at 50 °C for 15 min. 13. Transform into bacteria and make minipreps according to steps 5–8 of Subheading 3.3. 14. Test digest the assembled plasmid with EcoRI: 7 μl ddH2O, 1 μl 10× reaction buffer, 2 μl miniprep DNA, 0.1 μl EcoRI. 15. Incubate for 1 h at 37 °C. 16. Run on a 1% agarose gel; you should get a 7.2 kb and a 1.0 kb band. 17. Send for sequencing with standard primer M13r (see Note 19). 3.6  Midiprep

1. Retransform your plasmids (template and gRNA) into competent E. coli cells (see Note 20): 50 μl competent cells (thawed on ice) plus 1 μl plasmid. 2. Incubate on ice for 20–30 min. 3. Heatshock for 60 s in a 42 °C waterbath. 4. Add 1 ml LB medium (without antibiotics). 5. Incubate for 30–60 min at 37 °C. 6. Spread 200 μl onto LB-agar plates (with antibiotic, see Note 15). 7. Incubate over night at 37 °C. 8. Pick a single colony per plate using a toothpick or a pipette tip and inoculate each colony in a sterile Erlenmeyer bottle containing 100 ml of LB medium with ampicillin. 9. Incubate overnight in a 37 °C shaker. 10. Purify plasmids using a Midiprep kit. 11. Test digest each Midiprep along with the corresponding Miniprep using appropriate restriction enzymes (see Note 21): 8 μl ddH2O, 1 μl 10× reaction buffer, 1 μl midiprep or 2 μl miniprep DNA, 0.1 μl of each enzyme. 12. Incubate for 1 h at 37 °C. 13. Run on a 1% agarose gel (the pattern of the Midiprep should be identical to that of the Miniprep). 14. Determine the DNA concentration using a spectrophotometer (or nanodrop devise).

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15. Set up an injection mix: 0.4 μg/μl template DNA, 0.2 μg/μl gRNA plasmid in 100 μl total volume (ddH2O). 3.7  Injection

1. Set up a cage with Cas9 flies by combining flies from eight to ten big vials. Tape a yeasted apple juice plate to the bottom of the cage. 2. Incubate over night at 25 °C. 3. Change apple juice plate every 30–60 min (see Note 22). 4. Pull injection needles from glass capillaries using a needle puller (see Note 23). 5. Pull Pasteur pipettes on a Bunsen burner and break off the end of the tips (see Note 24). 6. Spin the injection mix in a table top centrifuge at full speed for 20–30 min. 7. Glue a cover slip onto a microscope slide using a drop of heptane with dissolved tape leaving a 2–3 mm gap between the edge of the cover slip and the edge of the slide. Put a drop of 10S oil on the slide covering the edge of the cover slip. Put it on the microscope and focus on the edge of the cover slip using a 10× objective. 8. Connect a rubber tube on one end with a blue pipette tip and on the other end with a drawn Pasteur pipette. Put the blue pipette in your mouth and suck a drop of injection mix into the Pasteur pipette. 9. Insert the Pasteur pipette into an injection needle to fill in the injection mix from the back of the needle by blowing into the blue pipette tip. 10. Tightly attach the injection needle to the needle holder, fasten it to the micromanipulator and bring the tip close to the edge of the coverslip on the microscope. When the tip of the needle is in the same focal plane as the edge of the cover slip, move the microscope stage so that the coverslip gently touches the tip of the needle to break it open. With the syringe attached to the needle holder put some pressure onto the needle until you see small droplets coming out of the tip (see Note 25). 11. Prepare 12–20 coverslips by pipetting a thin line of glue (tape in heptane) along one edge and labeling them with numbers. Cut out a slice of apple juice agar (around 30 mm × 8 mm). Put it onto the lid of a plastic petri dish. 12. Change the apple juice plate of the fly cage. Remove the yeast from the plate with the back of a brush, add some tap water on the plate and streak off the eggs with the brush. Transfer the eggs into the dechorionation cage. Put the cage into a jar with bleach and dechorionate the eggs for 1–2 min. Rinse the eggs

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thoroughly with water and transfer them onto the agar slice using the brush (see Note 26). 13. Line up the embryos along the edge of the agar slice with the anterior side (micropyle) pointing towards the edge. Transfer the lined up embryos onto a cover slip by gently touching them with the side that has glue on it (Fig. 3e). Repeat this for another two to four cover slips (see Note 27). 14. Desiccate the embryos for 10–15 min by putting them in a petri dish with cobalt(II) chloride crystals. 15. Cover the embryos with 10S oil, put the cover slip with the embryos on top on a microscope slide and put the slide under the microscope in the same focal plane as the tip of the needle (focus on the embryos and adjust the needle with the micromanipulator if necessary). 16. Using the stage control of the microscope move the first embryo towards the tip of the needle until the needle penetrates the vitelline membrane. While pulling back the embryo, put some pressure onto the needle using the syringe to release a drop of injection mix. Repeat this for all embryos on the cove slip (see Note 28). 17. When the embryos on the next cover slip are sufficiently desiccated, exchange the cover slip, putting the first one onto a plate with apple juice agar (see Note 29). Repeat this until all lined up embryos are injected. Keep the plates with the embryos at 18 °C. 18. After 2 days collect the larvae from the plates using a preparation needle and transfer them into a vial with food. Keep the vial at 25 °C. 3.8  Balancer Crosses

1. Within 10–12 days at 25 °C the injected embryos will develop into adult flies. The number of adults after CRISPR is usually rather small: of 500 injected embryos 100–150 larvae might hatch. 20–30 of those will develop into adults half of which might be sterile. 2. Collect every adult G0 fly, separate males and females, and cross each male and female individually with 3–5 balancer flies for the chromosome on which your gene is located. Keep the crosses at 25 °C. 3. After 10–14 days the F1 flies will start to eclose. Separate the balanced males and females. For each fertile G0 cross, set up 10–20 crosses of their offspring again crossing each male (and female) individually with 3–5 balancer flies. In total you should set up 100–200 F1 crosses (see Note 30). Keep the crosses at 25 °C.

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1. After a week, your F1 flies should have produced some larvae. Take the F1 male (or female) out of each vial and transfer each of them into a 1.5 ml reaction tube. Put the tubes on ice (see Note 31). 2. We usually squish 12–16 flies at a time. Suck 50 μl of squish buffer in into a pipette tip, use the tip to squish the fly in the first reaction tube, release the squish buffer, add 1 μl of proteinaseK and incubate the tube at 37 °C in a waterbath. Continue with the next fly in the next tube. Incubate the squished flies for 20–30 min (see Note 32). 3. Inactivate proteinaseK by incubating the tubes for 2 min at 95 °C on a heatblock. 4. Prepare a master mix for 12–16 test reactions (use the test primers flanking the tag insertion site (see Note 10): 264 μl water, 40 μl 10× buffer, 40 μl forward primer (10 mM stock), 40 μl reverse primer (10 mM stock), 8 μl dNTP mix (10 mM each), 3 μl Taq polymerase. 5. Distribute the master mix into 12–16 PCR tubes (24 μl each). Add 1 μl of supernatant from the squished flies to the PCR tubes (one test reaction per F1 fly). 6. PCR program: 92 °C

2 min

92 °C

30 sec

57 °C

30 sec

72 °C

1 min

72 °C

10 min

30 cycles  

7. Run 5 μl of each PCR reaction on a 1% agarose gel (see Note 33). 8. Discard any F1 crosses that produced only one PCR band (this will be the majority 90–100%). Keep the crosses that produced two bands and cross the balanced F2 flies inter se (see Note 34). Keep the crosses at 25 °C. 3.10  Sequence Verification

1. Take a single homozygous fly of the F3 generation and perform a squish prep as described in steps 2 and 3 of Subheading 3.9. Do this for all candidate lines you have established (see Note 35). 2. Set up two 50 μl PCR reactions for each candidate line: PCR reaction mix: 32 μl water, 5 μl 10× buffer, 5 μl forward primer (10 mM stock), 5 μl reverse primer (10 mM stock), 1 μl dNTP mix (10 mM each), 1 μl squish prep, 1 μl Taq polymerase.

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3. PCR program: 92 °C

2 min

92 °C

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57 °C

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72 °C

1 min

72 °C

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30 cycles  

4. Run 5 μl of each PCR reaction on a 1% agarose gel (see Note 36). 5. Pool the two remaining 45 μl of each candidate line and purify them using a PCR purification kit. Elute the columns with 30 μl of elution buffer. 6. Split the sample(s) into two tubes, add the forward primer used for the PCR to one tube, and the reverse primer to the other tube and send them for sequencing. 3.11  Conditional Knockout

1. Cross your genomic insertion line(s) and the UAS-Flp line inserted on another chromosome with double balancer flies. 2. Cross the F1 generation flies of the insertion line with the F1 generation of the UAS-Flp line to combine both features. Establish a stock. 3. Optional: Combine the Gal4 driver line(s) with Gal80ts using the same approach. 4. Cross the UAS-Flp; FRT-GFP-FRT insertion flies with the Gal4 (+ Gal80ts) flies. If you are using the Gal80 repressor, keep the cross at 18 °C until the offspring has reached the stage at which you want to knock out the FRT-flanked gene. Then switch the temperature to 29 °C to inactivate Gal80 thereby activating Gal4 mediated expression of Flippase and excision of your FRT-­flanked sequence (Fig. 1e). 5. Perform antibody stainings against GFP and a marker for the cells in which the Gal4 driver is expressed to check for the absence of GFP. Use uncrossed UAS-Flp; FRT-GFP-FRT insertion flies as controls. 6. Perform behavior experiments or check for other phenotypes caused by the knock out of the FRT-flanked sequence.

4  Notes 1. We have designed three plasmids in which any gene of interest can be tagged by GFP (or 3xHA) and flanked by FRT sites (Figs. 1a–c and 2). The tags can be added to the N-terminus or the C-terminus, or somewhere in the middle of your coding

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sequence, depending on the presence of alternative ­splicing and/or any functional domains at the N- or C-terminus. For small, intronless genes the entire coding sequence can be flipped out. For larger genes containing big introns only the start codon, or one or a few exons will be removed. 2. pBsFGF (Fig. 2a): With this plasmid you can put a GFP-tag to the N-terminus of your protein replacing the start codon of your gene. After FRT recombination the GFP sequence and thus, the start codon of the fusion protein will be removed. Note that in this case truncated versions of your protein starting from a second AUG codon might be produced. For the removal of more than just the start codon use plasmid pBsGFG∗. You can also use this plasmid for tagging the C-­terminus of you protein with GFP. For genes that have alternative splicing of the last exon, not all isoforms will be tagged. After FRT recombination the last exon(s) of the fusion protein including the GFP-tag will be removed. Note that in this case a truncated version of you protein will be produced. For an N-terminal GFP fusion, the first FRT site should be located upstream of the Start codon either within the first coding exon, or in the genomic sequence preceding the first coding exon (especially for genes that have more than one upstream noncoding exons and alternative transcription start sites). The coding sequence of your gene should be cloned after the second FRT site in frame with the GFP sequence, starting with the second amino acid of your gene. For a C-terminal fusion, the first FRT site should be located in an intron around 1–2 kb upstream of the Stop codon. The last coding exon(s) should be cloned without stop codon in frame wi.th GFP between the first FRT site and the GFP-tag. 3. pBsFGF∗ (Fig. 2b): In contrast to the pBsFGF plasmid, this plasmid does not only generate a sequence from which the GFP-tag including the translation start codon can be excised but also allows to add some additional coding sequence between GFP and the second FRT site. In this case the coding sequence of your gene should be cloned in frame with the GFP sequence between the GFP tag and the second FRT site. Note that there is a stop codon in the second FRT site. The second FRT site should be placed in the intron following the first coding exon (or the second coding exon, if the total length of this fragment will not exceed 1 kb). After FRT recombination the GFP and the sequence encoded by the first coding exon will be removed. For a C-terminal fusion the same rules apply as for pBsFGF. 4. pBs3xHA (Fig. 2c): The same rules apply as for the pBsFGF plasmid, except that for an N-terminal fusion the start codon has to be added upstream of the 3xHA tag fusing it in frame

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with the tag. After FRT recombination the Start codon and the HA-tag will be removed. This plasmid is also suitable for tagging your gene of interest internally. In this case, clone the 5′ part of the coding sequence between the first FRT site and the 3xHA-tag (in frame with the tag), and the 3′ part of the coding sequence after the second FRT site (in frame with the sequence coming from the 3xHA-tag. After FRT recombination, the 5′ part including the tag will be removed. 5. For the insertion of FRT sites into your gene of interest, you need to find two CRISPR sites. They should be flanking the sequence you want to excise in your final experiment (Fig. 1a– c). They will be replaced by the FRT sites in the CRISPR template sequence to prevent Cas9 cutting the template. To select two CRISPR sites in your gene of interest, paste the genomic sequence of your gene on the CRISPR target finder website (http://tools.flycrispr.molbio.wisc.edu/targetFinder/) [10]. You can also just paste around 1 kb of sequence in the region, in which you want to insert the FRT sites. Check the “All CRISPR targets” box and press the “Find CRISPR Targets” button. The program will give you a list of CRISPR target sites located in your region of interest. Check the boxes for “High” stringency and for “NGG Only” PAM sequences and press the “Evaluate” button. The program will give you a map with the positions of all CRISPR sites in your region of interest and a list of these sites with potential off-target sites. If possible choose unique sites. Note that the CRISPR sites can be oriented in + or – direction relative to your sequence. Include the sequence of your first CRISPR site into the forward primer (in 5′ to 3′ orientation without the PAM sequence): CRISPR fw primer: 5′-TATATAGGAAAGATATCCG-GGTG A A C T T C G - N 1 9 / 2 0 G T T T TA G A G C TA G A A A TAGCAAG-­3′. If your gRNA sequence starts with a G, N will be 19, otherwise N will be 20. Include the sequence of your second CRISPR site into the reverse primer (in the reverse complementary orientation without the PAM sequence): CRISPR re primer: 5′-ATTTTAACTTGCTATTTCTAGC TCTAAAAC-N19/-20revcomp-CGACGTTAAATTG AAAATAGGTC-3′. If your gRNA sequence starts with a G, N will be 19, otherwise N will be 20. See also this website for information about the primers: h t t p : / / w w w. c r i s p r f l y d e s i g n . o r g / w p - c o n t e n t / uploads/2014/06/Cloning-with-pCFD4.pdf.

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6. The primers should have a 20–30 nucleotide overlap with the template sequence and a GC content of 50–60% giving a melting temperature of around 62 °C. They should have a G or C at their 3′-end. To the 5′-end a restriction site can be added (if possible, choose one that is partially overlapping with the genomic sequence at this position). Two additional nucleotides should be added at the 5′-end of the restriction site. 7. Clone 1–2 kb genomic sequence upstream of the first CRISPR site into the chosen template assembly plasmid, upstream of the first FRT site using a pair of the following restriction sites: SacI, SacII, Not, SpeI, BamHI (not for the 3xHA plasmid), SmaI, PstI, BglII. Choose enzymes that do not cut in this fragment and if possible also do not cut elsewhere in your gene of interest (see Note 6). Design the forward primer so that it binds 1–2 kb upstream of your first CRISPR site and the reverse primer as close as possible (within 50 bp) 5′ of the first CRISPR site. 8. Clone the sequence that should be excised between the first CRISPR site and the tag using a pair of the following restriction sites: StuI, PmeI, AgeI or NcoI. Note that none of these enzymes cuts in pBluescript. In this case a given fragment can also be cloned directly into the final plasmid, or can be TOPO-­ cloned into a pCRII plasmid, or a second restriction site can be added to the primers. The second restriction sites can then be used for cloning into pBluescript and the first sites for cloning into the final template plasmid. Choose enzymes that do not cut in this fragment and if possible also do not cut elsewhere in your gene of interest (see Note 6). For an N-terminal fusion this sequence should contain the noncoding part of the first coding exon (plus a start codon for the pBsF3xHAF plasmid. Note that in this plasmid the AUG of the NcoI site is not in frame with the 3xHA-tag!). For a C-terminal (or internal) fusion the sequence should start in an intron an end in the last coding exon (or an internal one) in frame with the tag. Design the forward primer so that it binds as close as possible (within 50 bp) to the 3′-end of the first CRISPR site and the reverse primer at the end of the noncoding sequence (for an N-­terminal fusion), or the coding sequence (for a C-terminal fusion) (or internal sequence) in frame with the tag. When using the pBsFGF∗ plasmid additional coding sequence (for example the coding part of the first coding exon plus some additional intronic sequence in case of an N-­terminal fusion, or, in case of an internal fusion, the downstream coding sequence, can be cloned between the GFP tag and the second FRT site using a pair of the following restriction sites: EcoRI, EcoRV, HindIII (see Note 6). This will require an additional PCR reaction with the forward primer in frame

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with the tag and the reverse primer as close as possible (within 50 bp) 5′ of the second CRISPR site in an intron or the 3′UTR. 9. Clone 1–2 kb genomic sequence downstream of the second CRISPR site into the chosen template assembly plasmid, downstream of the second FRT site using a pair of the following restriction sites: HindIII (only for pBsFGF), SalI, XhoI, ApaI, Acc65I, or KpnI. Choose enzymes that do not cut in this fragment and if possible also do not cut elsewhere in your gene of interest (see Note 6). Design the forward primer so that it binds as close as possible (within 50 bp) 3′ of the second CRISPR site and the reverse primer 1–2 kb downstream of your second CRISPR site. 10. The test primers should be 20–30 nucleotides long. The forward primer should bind the genomic DNA in a region 200– 1000 bp upstream of the inserted tag, the reverse primer should bind the genomic DNA in a region 200–1000 bp downstream of the second inserted FRT site. For a GFP-tag insertion, the distance to the tag on both sides can be up to 1 kb, for a 3xHA-tag insertion the distance of both primers should not be more than 300 bp on each site. A second primer pair flanking the first FRT site insertion at a distance of 300– 500 bp to each side will be required for sequence verification. 11. Since the CRISPR reagents will be directly injected into nos-­ Cas9 expressing flies, the template for the genomic changes should be cloned from the genomic DNA of nos-Cas9 flies that can be quite different from the database sequence. We recommend to PCR amplify and sequence your genomic region including the selected CRISPR sites to confirm that the CRISPR sites, including the PAM sequences are present in the nos-Cas9 genome. For this you can use the test PCR primers (see Note 6). If you want to inject your CRISPR reagents (including a plasmid encoding Cas9) into other flies, you should use genomic DNA of those flies as PCR template. 12. Ideally one should use a proof-reading polymerase like Pfu for this step. However, to our experience Pfu does not perform very well on genomic DNA templates. Therefore, we usually use Taq polymerase at this step. For the later sequence verification this means, that there will not only be the sequence variations between the Cas9 genome and the database, but also PCR errors coming from the Taq polymerase. SNPs will usually be identical between different clones, while PCR errors are usually unique to one clone. Small differences in the noncoding regions of your gene of interest should not be a problem; however, the coding sequence should be identical. If you have sequenced your gene of interest in the genome of nos-­Cas9

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flies (see Note 6), you should compare the sequences of your fragments with the nos-Cas9 sequence. 13. The different fragments should first be cloned into a cloning vector like pBluescript. This will allow sequencing of the cloned fragments from both ends using standard primers. From there the different parts can be assembled in the final FRT-tag-FRT template plasmid. If an enzyme that was chosen for cloning of one fragment should cut in another fragment, this latter fragment has to be inserted in the final plasmid after the one that requires the enzyme cutting in the latter fragment. Note that not all restriction enzymes might cut in your cloning vector. In this case a given fragment can also be cloned directly into the final plasmid, or TOPO cloned into a pCRII plasmid, or a second restriction site can be added to the primer containing the site that does not cut in the vector. The second site can then be used for cloning into the vector and the first site for cloning into the final template plasmid. 14. Incomplete digestion of the vector can result in a high ratio of self-ligation yielding an empty plasmid. Therefore, we recommend performing a control ligation for every digested vector, using the same amount of purified vector DNA but adding more water instead of the insert. This control ligation should be transformed and plated in the same concentration as the actual ligation. If significantly less colonies are growing on the control plate than on the actual cloning plate, six to eight clones of the actual cloning plate should be sufficient for successful minipreps. If the amount of colonies on the control plate is as high or even higher than on the actual cloning plate, the vector should be redigested and dephosphorylated before the ligation. 15. For pBluescript, pBsFGF, pBsFGF∗, and pBsF3xHAF ampicillin should be used at 100 μg/ml. Other plasmids might require other antibiotics. To concentrate the bacteria and reduce the amount of liquid added to the plate, spin the bacteria briefly (10 s), remove about 900 μl of the supernatant and resuspend the pellet in the remaining 200 μl by gently pipetting it up and down. Add the entire suspension to the plate. 16. As an alternative to using the spin columns of the kit, after addition of the neutralization buffer and centrifugation, the supernatant can also be transferred into a fresh 1.5 ml tube and the DNA precipitated by adding 650 μl of isopropanol. Mix well, centrifuge for 7 min, discard the supernatant, wash the pellet with 1 ml of 70% ethanol, centrifuge for 1 min, remove the supernatant, dry the pellet until it becomes transparent, and resuspend it in 50 μl of ddH2O.

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17. BbsI gradually loses activity when stored at −20 °C. A fresh tube of BbsI enzyme can be aliquoted and stored at −70 °C. The BbsI digested and purified fragments can be stored at −20 °C to be used for other experiments. 18. Both, the forward and reverse primer used for this PCR amplification contain sequences matching the dU6 promoter and gRNA scaffold sequences occurring twice in the plasmid. Therefore, we recommend performing the PCR on the excised 493 bp fragment rather than the intact plasmid. 19. M13r binds to the pCFD4 plasmid about 1 kb downstream of the first gRNA insertion site. Thus, the resulting sequence will be reverse to the plasmid sequence. The quality of the DNA has to be good enough for a read of more than 1 kb. 20. To avoid this retransformation step, you can also keep a backup of your overnight cultures. Divide an LB-plate (with antibiotic) into as many sectors as you have overnight cultures. Label the sectors so, that you can correlate each of them to the corresponding clone. Before doing the minipreps, streak a bit of each overnight culture onto a sector of the plate with an inoculation loop (or the tip used for picking the colony), and incubate the plate over night at 37 °C. The plates can be kept at 4 °C for 2–4 weeks. 21. You can use the same enzyme combinations as for cloning of the different fragments. However, you can use one or two enzymes that cut within the amplified fragments, resulting in a characteristic pattern of 4–12 bands that should be identical between the midiprep and the corresponding miniprep. For example, the pCFD4 plasmid can be digested with PstI and EcoRI resulting in the following bands: 3929 bp, 1624 bp, 991 bp, 711 bp. 22. The flies usually start laying eggs in the afternoon. However, to get them into a good rhythm and to prevent the accumulation of older embryos and larvae on the plates, the plates should be changed regularly during the morning. Observe the number of eggs on the plates after each change. Once the number increases, you can start preparing the cover slips and loading the needle. If only one person is injecting and the flies are not laying so well, the plates can be changed every hour. If one person is injecting while another person is lining up the embryos, the plates can be changed every 30 min, provided that the flies lay enough eggs. 23. Pulling needles for injection can be quite tricky. They should be long and thin, but stiff enough so that they do not bend. Every needle puller requires different settings which might also have to be adjusted from time to time.

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24. The thin end of the Pasteur pipette should be pulled so that it will fit into the end of the injection needle. Hold the base of the thin end of a Pasteur pipette into the flame of a Bunsen burner until it gets soft. Take the pipette out of the flame and slowly pull both ends apart so that the thin part of the pipette reaches a length of 12–15 cm. 25. Usually there is some air in the injection needle. Keep the pressure of the syringe up until no more air bubbles are coming out of the needle. Small drops of injection mix should appear whenever you move the coverslip with the oil away from the needle even if you do not put extra pressure on the syringe. If the needle gets blocked, it can sometimes be reopened by touching the cover slip again. But if the opening gets too big, the embryos can no longer be injected and you have to prepare a fresh needle. After breaking the tip of the needle it will be sharp enough to penetrate the vitellin membrane of the embryos. After some time, however, it will become rather blunt, especially if the embryos are not well dechorionated. Then you also have to prepare a fresh needle. 26. The embryos have to be well dechorionated to be injected. Should you still see parts of the chorion on the embryos, repeat the dechorionation step for another 30 s. The efficiency of the bleach might decrease over time, so that longer dechorionation times will be required. 27. If one person is lining up the embryos, while the other person is injecting, up to six cover slips can be prepared within half an hour provided that the flies lay enough eggs. For one person alone, lining up three coverslips, then injecting them and then getting the next plate can also be quite efficient. Use only those embryos that do not show any structures (like furrows) inside. Place them with about 1.5–2 embryo widths apart from each other in a straight line. 28. Only inject embryos that have not yet started to cellularize. After cellularization the DNA will not be able to enter the germ cells. To avoid getting too many larvae that do not have their genomes of their germ cells altered, all embryos that are too old have to be destroyed by putting the needle into them and then moving them so that their vitellin membranes are ripped apart. Any embryos that were not injected at all for example because they were not sticking well to the cover slip also need to be removed. This can be done once the cover slips have been transferred to an apple juice plate using a preparation needle. 29. The injection needle should always be covered with oil. Therefore, after injection of a cover slip it should remain on

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the microscope until the next cover slip is ready and covered with oil. 30. If you injected a template with your gene of interest fused to GFP, you might try to select F1 flies expressing GFP. However, we found that depending on the expression levels of your gene, the GFP signal might be too weak to distinguish it from background fluorescence. However, you can preselect for example larvae that show high levels of fluorescence in the tissue in which your gene of interest should be expressed. Nevertheless, just blindly setting up the F1 crosses and screening later by PCR has turned out to be more efficient with our GFP tagged genes. For the F1 crosses we used mainly males, because in this case we could use both the males that were over the balancer and the males that were over the dominant marker of the balancer stock (for example Sp for the second or Dr for the third chromosome). Some G0 flies produced only very few offspring, in this case we also crossed their F1 daughters that had the balancer chromosome. 31. If the F1 animal is no longer alive, you can also wait for its offspring to eclose and use an F2 animal for the test PCR. 32. You can squish about 12–16 flies within 10 min. So, for a proteinaseK digestion of 20–30 min, the first squished fly will be incubated for 30 min and the last squished fly for 20 min. During those 20 min you can prepare the PCR mix. This will allow you to test 12–16 flies at a time. Alternatively, you can perform squish preps for up to 96 F1 flies within 90 min putting them into the waterbath in eight groups of 12 tubes and then test PCR them all together. 33. Since the F1 (or F2) flies are heterozygous, the test PCR should yield 2 bands. The shorter band should be the size of the wildtype sequence without insertions coming from the balancer chromosome; the longer band should be the sequence with the tag and FRT insertions. After the GFP insertion the resulting PCR product should be 700 bp longer than the wildtype sequence. Insertion of the 3xHA-tag should increase the size of the PCR product by 90 bp (+34 bp for the FRT site). Therefore the test primers for a 3xHA-tagged gene should be relatively close to the place of insertion, while those for a GFP-­ tagged gene can be further away (see Note 6). 34. In some cases you might also get an excision of the genomic sequence around one of the two CRISPR sites, or even between both sites. If you are interested in such mutants, you could also screen for PCR products shorter than the wildtype band, or use primers flanking both CRISPR sites.

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35. The nos-Cas9 stock might carry some lethal mutations on the chromosome that contains your gene of interest. Therefore it is possible that after balancing this chromosome the resulting flies are homozygous lethal even when the inserted tag should not interfere with the function of your gene of interest. Therefore, we recommend to establish stable lines for all candidates that were tested positive to see, if any of them yields homozygous flies; or if you get homozygous flies when crossing two established lines that came from different G0 animals and are homozygous lethal. For sequencing across a GFPinsertion one can also use heterozygous flies and separate both bands on an agarose gel. For the 3x-HA tagged flies and the first FRT insertion, separation of the inserted and uninserted PCR products may not be possible. In this case one could digest the PCR reactions, the one for the FRT site with XbaI and the one for the 3xHA site with BamHI, run them on a gel, purify the two digested bands (they will be shorter than the wildtype band), and send them for sequencing. The sequences will then end at the XbaI or BamHI sites located in the inserted sequence. 36. If you used homozygous flies, you should get only one band the size of the genomic region that was amplified, including the inserted sequence. However, since you are using genomic DNA as a template, additional bands might appear that could interfere with the sequencing reaction. In this case the whole PCR reaction should be loaded on an agarose gel. Cut out the band that has the expected size and gel purify it.

Acknowledgments We thank S. Bullock, M. Harrison, K. O’Connor-Giles, and J. Wildoger for plasmids. Special thanks to Jenifer Kaldun for generating the pBsF3xHAF plasmid. We thank the Bloomington Drosophila Stock center for fly strains. This work was funded by the Swiss National Science Foundation (31003A_149499 to S.G.S.). References 1. Consortium IMK, Collins F, Rossant J, Wurst W (2007) A mouse for all reasons. Cell 128(1):9–13. . Epub 2007/01/16. https:// doi.org/10.1016/j.cell.2006.12.018 2. Guan C, Ye C, Yang X, Gao J (2010) A review of current large-scale mouse knockout efforts. Genesis 48(2):73–85. . Epub 2010/01/23. https://doi.org/10.1002/dvg.20594

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with transgenic Cas9 in Drosophila. G3 (Bethesda) 4(5):925–929. https://doi. org/10.1534/g3.114.010496. Epub 2014/03/25 5. Schertel C, Albacara M, Rockel-Bauer C, Kelley N, Bischof J, Hens K et al (2015) A large-scale, in vivo transcription factor screen defines bivalent chromatin as a key property of regulatory factors mediating Drosophila wing development. Genome Res 25(4):514–523. https://doi.org/10.1101/gr.181305.114. Epub 2015/01/09 6. Jinek M, Chylinski K, Fonfara I, Hauer M, Doudna J, Charpentier E (2012) A programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity. Science 337(6096):816–821. Epub 2012/06/30. https://doi.org/10.1126/science.1225829 7. Bassett A, Tibbit C, Ponting C, Liu J (2013) Highly efficient targeted mutagenesis of Drosophila with the CRISPR/Cas9 system. Cell Rep 4(1):220–228. https://doi. org/10.1016/j.celrep.2013.06.020. Epub 2013/07/06 8. Bier E, Harrison M, O’Connor-Giles K, Wildonger J (2018) Advances in engineering the fly genome with the CRISPR-Cas system. Genetics 208(1):1–18. https://doi. org/10.1534/genetics.117.1113. Epub 2018/01/06 9. Gratz S, Cummings A, Nguyen J, Hamm D, Donohue L, Harrison M et al (2013) Genome engineering of Drosophila with the CRISPR RNA-guided Cas9 nuclease. Genetics 194(4): 1029–1035. https://doi.org/10.1534/genetics.113.152710. Epub 2013/05/28 10. Gratz S, Ukken F, Rubinstein C, Thiede G, Donohue L, Cummings A et al (2014) Highly specific and efficient CRISPR/Cas9-catalyzed homology-directed repair in Drosophila. Genetics 196(4):961–971. https://doi. org/10.1534/genetics.113.160713. Epub 2014/01/31 11. Kondo S, Ueda R (2013) Highly improved gene targeting by germline-specific Cas9 expression in Drosophila. Genetics 195(3): 715–721. https://doi.org/10.1534/genetics.113.156737. Epub 2013/09/05 12. Korona D, Koestler S, Russell S (2017) Engineering the Drosophila genome for developmental biology. J Dev Biol 5(4). https:// doi.org/10.3390/jdb5040016. Epub 2018/04/05 13. Port F, Chen H, Lee T, Bullock S (2014) Optimized CRISPR/Cas tools for efficient germline and somatic genome engineering in Drosophila. Proc Natl Acad Sci U S A

111(29):E2967–E2976. https://doi. org/10.1073/pnas.1405500111. Epub 2014/07/09 14. Ren X, Sun J, Housden B, Hu Y, Roesel C, Lin S et al (2013) Optimized gene editing technology for Drosophila melanogaster using germ line-specific Cas9. Proc Natl Acad Sci U S A 110(47):19012–19017. https://doi. org/10.1073/pnas.1318481110. Epub 2013/11/06 15. Xu J, Ren X, Sun J, Wang X, Qiao H-H, Xu B-W et al (2015) A toolkit of CRISPR-based genome editing systems in Drosophila. J Genet Genomics 42(4):141–149. https://doi. org/10.1016/j.jgg.2015.02.007 16. Yu Z, Chen H, Liu J, Zhang H, Yan Y, Zhu N et al (2014) Various applications of TALENand CRISPR/Cas9-mediated homologous recombination to modify the Drosophila genome. Biol Open 3(4):271–280. https:// doi.org/10.1242/bio.20147682. Epub 2014/03/25 17. Yu Z, Ren M, Wang Z, Zhang B, Rong Y, Jiao R et al (2013) Highly efficient genome modifications mediated by CRISPR/Cas9 in Drosophila. Genetics 195(1):289–291. https://doi.org/10.1534/genetics.113.153825. Epub 2013/07/09 18. Elliott D, Brand A (2008) The GAL4 system : a versatile system for the expression of genes. Methods Mol Biol 420:79–95. Epub 2008/07/22. https://doi.org/10.1007/ 978-1-59745-583-1_5 19. Brand A, Perrimon N (1993) Targeted gene expression as a means of altering cell fates and generating dominant phenotypes. Development 118(2):401–415 20. Golic K, Lindquist S (1989) The FLP recombinase of yeast catalyzes site-specific recombination in the Drosophila genome. Cell 59(3):499–509. Epub 1989/11/03. doi:. https://doi.org/10.1016/0092-8674(89) 90033-0 21. Duffy J, Harrison D, Perrimon N (1998) Identifying loci required for follicular patterning using directed mosaics. Development 125(12):2263–2271 22. Chalfie M, Tu Y, Euskirchen G, Ward W, Prasher D (1994) Targeted gene expression as a means of altering cell fates and generating dominant phenotypes. Science 263(5148):802–805 23. Jennett A, Rubin G, Ngo T, Shepherd D, Murphy C, Dionne H et al (2012) A GAL4-­ driver line resource for Drosophila neurobiology. Cell Rep 2(4):991–1001. https://doi. org/10.1016/j.celrep.2012.09.011

CRISPR/Cas9 Genome Editing to Study Nervous System Development in Drosophila 24. Pfeiffer B, Jenett A, Hammonds A, Ngo T, Misra S, Murphy C et al (2008) Tools for neuroanatomy and neurogenetics in Drosophila. Proc Natl Acad Sci U S A 105(28):9715–9720. https://doi.org/10.1073/pnas.0803697105. Epub 2008/07/16

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Chapter 11 The Red Flour Beetle as Model for Comparative Neural Development: Genome Editing to Mark Neural Cells in Tribolium Brain Development Max S. Farnworth, Kolja N. Eckermann, Hassan M. M. Ahmed, Dominik S. Mühlen, Bicheng He, and Gregor Bucher Abstract With CRISPR/Cas (Clustered Regularly Interspaced Short Palindromic Repeats/CRISPR-associated) scientists working with Tribolium castaneum can now generate transgenic lines with site-specific insertions at their region of interest. We present two methods to generate in vivo imaging lines suitable for marking subsets of neurons with fluorescent proteins. The first method relies on homologous recombination and uses a 2A peptide to create a bicistronic mRNA. In such lines, the target and the marker proteins are not fused but produced at equal amounts. This work-intensive method is compared with creating gene-specific enhancer traps that do not rely on homologous recombination. These are faster to generate but reflect the expression of the target gene less precisely. Which method to choose, strongly depends on the aims of each research project and in turn impacts of how neural cells and their development are marked. We describe the necessary steps from designing constructs and guide RNAs to embryonic injection and making homozygous stocks. Key words CRISPR/Cas9, Genome engineering, HDR, NHEJ, Tribolium, Neural lineage, Brain development, 2A peptide

1  Introduction 1.1  The Red Flour Beetle as Model for Brain Development and Evolution

The brain is among the most complex structures of an organism and understanding its development has been a major challenge in developmental biology. Highly advanced model systems with their plethora of tools and resources are spearheading this research and these ongoing efforts have been revealing basic principles of neural development of both deuterostomes and protostomes, with the dipteran Drosophila melanogaster being the model for arthropods [1–3]. Another major enigma in the field is the developmental

Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_11, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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basis of the evolution of brain diversity [4, 5]. In insects, for instance, the basic neuropil structure is highly conserved, but there is great variety of absolute brain size, relative size, and shape of homologous neuropils [6], in addition to heterochronic development [7]. In order to study evolutionary differences on a genetic and developmental level within insects a comparison of Drosophila to another insect species is required. Such a species should provide the tools for functional genetic work and transgenic approaches and as many resources as possible. While no insect species is in sight that will match the resources and the tool kit of Drosophila melanogaster, the red flour beetle Tribolium castaneum fulfills all necessary requirements. RNAi is strong and systemic such that all cells are targeted and the effect is transmitted to the offspring [8– 10]. A genome-wide RNAi screen is being performed [11], transgenesis is well established [12], and enhancer trap screens have been performed [13, 14]. Recently, genome editing via CRISPR/ Cas9 has been established in this species [15, 16]. Given this availability of robust tools for functional genetics in Tribolium, we think that this beetle has the potential to become the main comparative organism for studying mechanisms of brain diversification in insects. One approach is to mark homologous cells in both species and compare similarities and differences throughout development [7]. We want to elaborate here on the possibilities of CRISPR/Cas to extend the Tribolium toolkit for brain development and evolution research. For protocols about immunostaining and in situ hybridization of embryonic, larval and adult brains in Tribolium please refer to respective chapters in this book. 1.2  Using Transgenic Lines to Study Tribolium castaneum Brain Development

In order to study the development of the Tribolium brain, subsets of neural cells need to be visualized. Of particular use is the visualization of whole neural cells including soma, projections, and fine arborizations. In Drosophila, extensive enhancer trap screens [17–19] have created collections of transgenic lines where the expression of genes and the respective anatomical structures are marked with expression of fluorescent proteins. In addition, large collections of lines have been generated where markers are under the control of specific enhancers and promotors [20]. Subsequent immunostaining (e.g., [21]) or live-imaging (e.g., [22]) can generate fascinating insights into neural morphology and development. An enhancer trap screen has been performed in Tribolium castaneum and some transgenic brain imaging lines have become available [7, 13]. However, the collection is comparably small such that transgenic lines suited for individual research projects will in many cases have to be generated. Before the advent of CRISPR/Cas Tribolium transgenic lines were generated by enhancer trapping [13] or by insertion of constructs containing a gene’s regulatory regions [7]. In both cases the genetic construct is randomly integrated into the genome by

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transposon-mediated mutagenesis [12]. In case of enhancer trap experiments, a construct devoid of enhancers, but comprising a basal promoter and reporter gene jumps randomly into a gene locus. Because of the proximity, enhancers that control expression of that gene start to regulate the reporter gene as well. In case of genes involved in brain development, neural expression patterns are observed [13]. An alternative approach is to identify a gene of interest (GOI) and include parts of the regulatory region of this GOI in front of a reporter gene and let it integrate randomly [7, 23]. If functional enhancers are included in the construct, expression is at least partially similar to the one of the GOI. In both approaches, position effects are a major issue [24, 25]. Depending on where the construct is inserted, reporter gene expression can be influenced by additional enhancer elements and/or loose regulation by others. In case of reporter constructs, only parts of the regulatory region are usually included leading to loss of important enhancers besides position effects. As consequence, marker gene expression usually does not precisely reflect the GOI’s expression. CRISPR/Cas-mediated genome editing can be used to in two ways to mark neural cells in Tribolium: First, enhancer trap constructs can specifically be targeted to loci of genes with interesting neural expression obviating the need of time-consuming random screens. Second, homologous recombination allows engineering transgenic lines where the reporter is encoded by the same mRNA as the GOI. It therefore exactly mirrors GOI expression. 1.3  CRISPR/Cas

CRISPR (Clustered Regularly Interspaced Short Palindromic Repeats/CRISPR-associated) loci are repetitive elements that are part of the bacterial and archaeal adaptive immune system [26]. They contain CRISPR-associated (Cas) genes that encode for endonucleases. One of those genes encodes for Cas9 which is part of the type II adaptive immune response. This system is most widely exploited in genome editing, because it only requires one Cas protein, making the system more easy to use than type I and III [27]. The second major part of CRISPR loci are CRISPR arrays. They consist of repeat sequences and spacers that vary in sequence and correspond to foreign genetic elements (protospacers) of, e.g., phages [26]. These arrays are transcribed as single RNA and further processed to CRISPR RNAs (crRNAs) [27]. Associated transactivating CRISPR RNAs (tracrRNAs) hybridize with the repeat sequences, and are cleaved to include only one spacer sequence per duplex, which subsequently forms a complex with the Cas9 protein [27]. This active complex is also referred to as chimeric RNA or guide RNA, which uses the spacer sequence to bind to complementary DNA, and the Cas9 protein cuts the foreign DNA three base pairs upstream of the PAM (protospacer adjacent motif), which is a nucleotide triplet specific for each Cas protein. The resulting double-strand breaks (DSBs) can be repaired by two cellular repair mechanisms. The first mechanism is the

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error-prone nonhomologous end joining (NHEJ). Here, free DNA ends are fused, often resulting in indels (insertion-deletions) that frequently cause frameshift mutations and gene disruption. Importantly, besides indels, available linear DNA fragments can be inserted during NHEJ repair as well. The second mechanism is homology-directed repair (HDR), where the DSB is repaired using homologous sequences as template such that indels are avoided. The CRISPR/Cas9 system has been modified to edit genomes of diverse species at specific locations [26, 28, 29] including Tribolium [15]. Genome editing can on the one hand be used to generate mutations of a specific gene or to remove whole genes or other DNA elements. On the other hand, directed insertion of linear DNA fragments at specific loci by NHEJ is feasible. In addition, HDR allows tailoring genetic modifications with single base precision. Hence, gene loci can be modified in multiple ways including imaging lines suitable for neurodevelopmental research and other purposes [15, 26–28]. 1.4  Two Major Strategies to Generate Imaging Lines Using CRISPR/ Cas9

In this chapter, we highlight two strategies to make transgenic lines suitable for neurodevelopmental research in Tribolium. The first approach is the generation of enhancer traps in selected loci by NHEJ. Technically, this is the simpler approach, but the resulting reporter expression may lack precision. The second approach consists of the generation of bicistronic reporter lines using HDR. While the design is more demanding, the reporter will reflect the expression of the targeted gene with high precision. The transformation efficiencies appear to be in a similar range in both approaches (Johannes Schinko, TriGenes, personal communication). Specifically for HDR, observed rates (i.e., number of positive G0 founders of fertile injected animals) are 0.5% (Johannes Schinko, TriGenes, personal communication) and 0.65% [30]. Hence, we strongly advise to inject high numbers of embryos to counteract the relatively low rates of integration.

1.5  Prerequisite: Selection of the Gene of Interest

Depending on the cells that one would like to mark, the GOI could for instance be a neural differentiation gene that marks certain cell types, e.g., an enzyme involved in the production of certain neurotransmitters. When using such differentiation markers, expression of the respective reporter is expected to emerge rather late in development, i.e., during cell differentiation. Alternative GOIs are neural patterning genes that mark certain neuroblasts and/or neuroectodermal regions and later subsets of neural cells [7]. Here, reporter expression is expected both early in embryogenesis and later in the differentiated brain reflecting both early and late functions of such transcription factors. Due to the dynamics of patterning gene expression, loss of expression in subsets of cells at certain stages might be observed as well.

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1.6  Gene-Specific Enhancer Traps via NHEJ 1.6.1  Selecting Insertion Sites Within the GOI Locus

1.6.2  Generation of the Enhancer Trap Construct

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The first strategy exploits NHEJ to generate enhancer traps in the regulatory region of a gene of interest [13]. Sites of insertion can greatly influence reporter gene expression. Therefore, mindful choice of target sites is crucial. The exact location of the insertion within the locus of the GOI is defined by the DSBs which depends on appropriate guide RNAs. A mutation of the GOI might interfere with proper brain development or morphology, so that the insertion should not interrupt exons. Likewise, disruption of regulatory elements should be avoided, because these may lead to a mutant phenotype as well. As regulatory elements are difficult to predict based on DNA sequence, data on putative enhancers by Faire or ATACseq experiments might be used to further exclude insertion sites [31–33]. Unfortunately, it is unpredictable what location of an insertion will produce a good enhancer trap. In previous screens a large portion of enhancer traps identified was located upstream of the transcription start site (TSS) or in the first intron [13, 34] reflecting the predominant location of enhancers around the TSS [35]. However, insertions downstream of the polyA signal are producing enhancer trap patterns as well. Given these uncertainties, it is recommended to design guide RNAs for at least two to three different insertion sites per GOI and produce independent lines for each. Note that gene annotations found on genome browsers (e.g., J-Browse at iBeetle-Base http://ibeetlebase.uni-goettingen.de/jbrowse/) were generated automatically and may contain errors. Hence, the annotation needs to be checked manually and in case of doubt, the TSS needs to be identified, e.g., by RACE reactions. Depending on needs, enhancer trap constructs to be inserted may have different designs. For imaging, the construct must contain a reporter gene, e.g., encoding EGFP (or a transactivator like Gal4 [36]) under the control of a basal promotor. A basal promoter is a 300–400 bp region around the TSS which enables the binding of the polymerase, but does not initiate transcription on its own (Fig. 1a). Upon insertion into the genome, close-by enhancers will activate transcription from that promoter. In Drosophila, the basal heat shock promoter (bhsp) has been widely used in constructs and enhancer trap screens and a respective Tribolium bhsp has been successfully tested for that purpose [37]. Sequences of core promoters influence the interaction with enhancer elements modulating the expression of the reporter [20, 38]. Hence, using respective core promoters from neural genes might be an option. However, each new core promoter under consideration has to be functionally tested first. Next, the construct needs a transformation marker (Fig. 1a) allowing for the identification of the few transformants in between hundreds or thousands of nontransformed animals. The best option currently for brain imaging is the use of the ­3XP3-­Tc-­vermilion marker which rescues the white eyes of the

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Tribolium vermilion white mutation to black eyes [39]. This marker is easy to screen without epifluorescence and does not interfere with fluorescent brain imaging. Note that the artificial 3XP3-EGFP marker often used as transformation marker in beetles does not only drive expression in the eyes but also in glia cells [7]. Hence, if a fluorescent eye marker is used one has to choose a fluorophore such that overlap with the reporter protein is minimized. Importantly, the construct needs to be linearized in order to be inserted into a double-strand break (Fig. 1a). This can be facilitated by flanking the construct with sequences that are targeted by additional guide RNAs that do not match the genome of Tribolium. Previously tested guide RNA sequences from other species are a good option. A guide RNA targeting the ebony locus from Drosophila was successfully tested (Klingler, personal ­communication

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and own experience), although an alternative target site to Dm-yellow was also used previously and is an alternative (Klingler, personal communication). Such a construct has been cloned and used successfully to generate a number of enhancer traps of neural cells in Tribolium and is available from Addgene [plasmid #124068]. It contains the abovementioned components, but in addition to EGFP drives the Cre recombinase in the same pattern. Judging from our still limited experience we tend to believe that simply linearizing the plasmid close to the basal promotor of the construct may be more efficient than cutting out the construct at both sides. The disadvantage of simple linearization is that the plasmid-backbone is still in the genome and might generate an artificial situation at the locus. Comparing different insertion sites in one GOI locus, we found that the reporter patterns were in most cases related to the GOI but also showed quite some differences, probably depending on insertion site and orientation. 1.6.3  When to Use This Strategy

The enhancer trap approach has the advantage that the same construct can be used for all loci and does not need to be adopted specifically for each gene, substantially reducing the effort for cloning. A disadvantage can be that not the entire expression is captured by reporter expression and/or ectopic expression may occur. Hence, the resulting EGFP pattern may not be an exact copy of the GOI’s expression. This can actually turn into an advantage when only a small subset of cells is marked such that following neural projections can be easier.

1.7  Bicistronic Lines via HDR

If a precise copy of GOI expression is required, the more laborious generation of bicistronic lines is the option to choose. Essentially, the genome is edited such that an mRNA encoding both GOI and reporter gene is transcribed. The consequence of the fused mRNA is that both GOI and reporter are regulated from the same promoter in identical patterns and dynamics. This can be realized on the one hand by constructing fusion proteins [40]. Here, a reporter gene is inserted in frame with the GOI extending its ORF. However, the resulting fusion protein may have a strongly modified 3D structure and thus, may not function properly. Further, the signal of the reporter will only be present at the target protein’s cellular location. In case of transcription factors, the nucleus would be marked, but not the projections. To avoid these restrictions, 2A peptides can be used to generate two separate proteins from one mRNA. 2A peptides are short, approximately 20 amino acid long peptides that cause “ribosomal skipping” [41]. An mRNA where the sequence of the GOI is separated from the reporter by a 2A sequence will lead to the translation of the GOI, interruption of the polypeptide chain within the 2A sequence and to subsequent translation of the reporter by the same ribosome [42, 43]. Hence,

1.7.1  Principle of Bicistronic Expression by Using 2A Peptides

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a reporter gene such as EGFP is expressed alongside the GOI in the same cells, in the same amount and without affecting GOI function. Importantly, it is expected that EGFP will be located in the cytoplasm resulting in marking of the whole neural cells including projections. 1.7.2  Design of the Repair Template

In the construct for the repair template, parts of the GOI, the 2A peptide and a reporter gene, e.g., EGFP, need to be cloned in-­ frame before a STOP codon (Fig. 2). The original STOP codon of the GOI needs to be removed. The new STOP is followed by the 3′UTR containing the polyA signal of the endogenous gene. Upon integration, such a construct leads to the production of one mRNA which apart from its extension by the reporter is identical to the endogenous GOI mRNA. Downstream of the 3′UTR, a transformation marker needs to be included (see above for criteria for selection). The purpose of putting the artificial eye promotor 3XP3 [12] downstream of the bicistronic gene locus (Fig. 2) is to reduce the chance of interference of these two transcription units. In order to allow for homology-dependent integration, sequences homologous to the DNA up- and downstream of the sequence to be edited, need to be included. One sequence contains parts of the GOI sequence upstream of the STOP codon (5′ homology arm) and the other is identical to sequences downstream of the 3′UTR (3′ homology arm) (Fig. 2b). Hence, these homology arms flank the construct and serve for alignment of the construct to the chromosome during HDR. The construct does not need to be linearized for integration.

1.7.3  Selection of the guide RNAs

Homology-dependent repair only occurs efficiently after DSBs initiate the repair process. For efficient insertion of the construct, the sequence to be replaced is cut out by two guide RNAs via CRISPR/ Cas9: guide RNA 1 induces a DSB as close to the STOP codon of the GOI as possible (Fig. 2a). The second guide RNA 2 introduces a DSB in the intergenic region downstream of but close to the 3′UTR. Using both guide RNAs at the same time will delete the entire 3′UTR (Fig. 2a) and substitute it with the cassette including the 2A peptide, the reporter, the endogenous 3′UTR and the transformation marker (Fig. 2c). It is important to make sure that the guide RNAs do not target the homology arms or any other part of the repair template, because this would lead to fragmentation of the repair template, prohibiting proper integration. Hence, it is necessary to modify the PAM sequence in the repair template plasmid for both guide RNAs.

1.7.4  When to Use This Strategy

The strategy of a bicistronic line should be chosen when the expression dynamics of the GOI have to be copied as exactly and comprehensively as possible. It also is a method with a minimal chance of disrupting a gene’s function. Disadvantages are that the strategy is

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much more work-intensive, because repair templates need to be cloned individually for each gene. Also, the size of the gene’s 3′UTR might be a limiting factor, since larger constructs are more difficult to be inserted via HDR. To generate transgenic lines, the following steps need to be performed: 1. Sequencing of insertion locus. 2. guide RNA design. 3. guide RNA cloning. 4. guide RNA efficiency test. 5. Repair template and enhancer trap construct cloning. 6. Embryonic injection. 7. (Back-) Crossings of G0 to wildtype. 8. Screening for transgenics in F1. 9. Characterization of the integration event. 10. Generating homozygous stocks.

2  Materials 2.1  Tribolium Husbandry

2.2  Genomic DNA Extraction (See Note 1)

Standard equipment and supplies for molecular work as well as knowledge of Tribolium castaneum husbandry [44] are implied and not listed. For experiments, beetles of vermilionwhite (vw) strain should be used [39]. Double-deionized water should be used at all steps, as well as analysis-grade Ethanol. 1. DNA extraction buffer (80 mM EDTA pH = 8, 100 mM Tris-­Cl pH = 8, 0.5% SDS, 100 μg/ml Proteinase K, added freshly). 2. Micro-Pestle suited for 1.5 ml tubes. 3. Squishing buffer (10 mM Tris-Cl pH = 8.2, 1 mM EDTA, 25 mM NaCl, 200 μg/ml Proteinase K, added freshly). 4. 10 mg/ml BSA. 5. Wing buffer (10 mM Tris-Cl pH = 8.2, 1 mM EDTA, 25 mM NaCl, 500 μg/ml Proteinase K, added freshly). 6. 24-well plates or similar. 7. Dumont No. 5 forceps.

2.3  DNA Plasmid Vectors and Cloning

1. p(U6b-BsaI-gRNA), Addgene plasmid #65956. 2. p(bhsp68-Cas9), Addgene plasmid # 65959. 3. pBac[3xP3 g Tc’ v], Addgene plasmid #86446. 4. pCR™II vector or pJET1.2/blunt (Thermo Fisher Scientific, Waltham, USA).

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5. BsaI restriction enzyme. 6. T4 DNA ligase and buffer. 7. Transfection-grade plasmid-prep kits for purification. 8. T7 Endonuclease I. 9. DNA Assembly kit [e.g., Gibson Assembly® Cloning Kit (New England Biolabs, Ipswich, USA) or In-Fusion Cloning kit (Takara Bio Inc., Kusatsu, Japan)]. 10. DpnI restriction enzyme. 2.4  Embryonic Injections

1. Borosilicate capillaries (e.g., from Ligenberg GmbH, Malsfeld, Germany) 10 mm × 1 mm. 2. P-2000 micropipette puller (Sutter Instrument, Novato, USA) or similar. 3. Optional: Microelectrode beveler (e.g., from Bachofer GmbH, Reutlingen, Germany). 4. FemtoJet® Microinjector (Eppendorf, Hamburg, Germany). 5. 0.45 μm (for 0.5 ml tubes) and 0.22 μm filters. 6. 10× injection buffer (14 mM NaCl, 0.7 mM Na2HPO4 × 2H2O, 0.3 mM KH2PO4, 40 mM KCl, filter-sterilize with 0.22 μm filter, aliquot and store at −20 °C). 7. Phenol red. 8. Apple agar plates. 9. Bleach (DanKlorix, CP GABA GmbH, Hamburg, Germany). 10. Voltalef 10S oil (Lehmann & Voss & Co., Hamburg, Germany).

3  Methods 3.1  Sequencing of Insertion Locus

Even though the genome of Tribolium is available [45] it is still necessary to sequence the regions to be targeted by Cas9, since beetle strains and laboratory stocks may differ in sequence. Single nucleotide polymorphisms (SNPs) can occur in a potential target sequence and a different nucleotide in the PAM usually abolishes Cas9 function. In addition, differences in the rest of the target sequence can drastically reduce Cas9 efficiency [27]. For gene-­ specific enhancer traps, the regions to be sequenced should comprise the potential insertion sites, thus mainly the region upstream of the transcription start site and first intron (Fig. 1). In the case of bicistronic lines, we advise to sequence around 1 kB upstream of the STOP codon reaching 250 bp into the 3′UTR after the STOP for guide RNA 1, and 250 bp of the 3′UTR end as well as 1 kB of the downstream intergenic region for guide RNA 2 (Fig. 2). First, extract genomic DNA of adult vw beetles (i.e., the strain in which you later want to integrate your CRISPR construct):

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1. Put ~10 cold-anaesthetized adult beetles in a 1.5 ml tube, add 200 μl DNA extraction buffer. 2. Homogenize using a pestle suited for 1.5 ml tubes. 3. Put the homogenate in a heat block at 50 °C for 1 h, mix by pipetting every 20 min. 4. Add 200 μl 5 M NaCl, mix, add 300 μl Chloroform and mix by inverting. 5. Spin down at 16,000 × g for 15 min at RT. 6. Transfer 300 μl of the upper (aqueous) phase to a 1.5 ml tube. 7. Add 30 μl 7.8 M ammonium acetate, mix by pipetting and add ice-cold 600 μl 100% Ethanol, invert 5–10×. 8. Keep at −20 °C for at least 1 h. 9. Spin down at 16,000 × g for 20 min at 4 °C. 10. Discard supernatant. 11. Wash pellet with 300 μl 70% Ethanol. 12. Spin pellet down at 16,000 × g for 5 min at 4 °C. 13. Repeat steps 10–12. 14. Remove as much of the supernatant as possible and air-dry the pellet for ~10 min. 15. Resuspend DNA in 20 μl double-deionized H2O. Design suitable sets of primers to cover the areas to be sequenced (see Note 2). These should include primers to amplify the regions of interest (see Note 3) and sequencing primers that bind within this amplicon. Sequencing results should be compared with the Tribolium reference genome (currently, GCF_000002335.3 Tcas5.2; please check http://ibeetle-base.uni-goettingen.de) (see Note 4). 3.2  Guide RNA Design

Guide RNA sequences can be determined with the CRISPR Optimal Target Finder (see Note 5) at http://tools.flycrispr.molbio.wisc.edu/targetFinder/ [46]. The aim is to find target sites at the location where DSBs should be induced and that occur only at this location in order to avoid off-target effects (see Notes 6–9). For the generation of bicistronic lines, two guide RNAs are used to excise the 3′UTR and substitute it with the construct. Hence, guide RNA 1 should cut as close to the gene’s STOP codon as possible and guide RNA 2 downstream of but as near as possible to the end of the 3′UTR (Fig. 2). These guide RNAs will likely bind to both the genome and the repair template. In order to avoid destruction of the latter, the respective sequence in the repair template should be modified (see Subheading 3.5), but without affecting the encoded amino acid sequence. Unfortunately, this can further restrict suitable guide RNAs. In case of gene-specific enhancer traps, any target sites outside the exons and apart from putative regulatory elements can be chosen, but we recommend targets not too far from the transcription start site (Fig. 1).

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We recommend following settings for the target finder tool: 1. Set to finding all CRISPR targets. The U6 promotors need a 5′ Guanine for proper transcription [47] and a respective G is contained in the plasmid vector the guide RNA will be cloned in (see purple G in Fig. 3). It is advisable to search for guide sequences that have a G at the 5′ end in order to match the U6 promoter requirement. However, one mismatch at the 5′ end of the guide has been shown to not much reduce efficiency. Hence, this is not an absolute requirement, especially because a G is available in the plasmid vector. 2. Guide RNA length of 20 bp is suitable in our experience. 3. Check for and use the latest released Tribolium genome release (check http://ibeetle-base.uni-goettingen.de). 4. It is advisable to restrict the search to target sites with only “NGG” PAMs, as “NAG” PAMs have reduced efficiency [48]. 5. We use high stringency settings since maximum stringency criteria are based on cleavage effects in cell lines only. The “Design Experiment” button is helpful to extract the actual oligonucleotide sequences to be ordered. It automatically adds Gs if necessary. However, the overhangs generated will not be the correct ones for the plasmid containing the Tribolium U6 promotor (p(U6bBsaI-gRNA, see Fig. 3). To generate the correct overhangs, we recommend using the TriGenes guide RNA oligo design tool at https://trigenes.com/crispr/grna-oligo-design-tool/. We recommend using Golden Gate reactions to clone guide RNAs (Fig. 3). An alternative protocol can be found under https://trigenes.com/crispr/grna-oligo-design-tool/.

3.3  Guide RNA Cloning

1. Anneal oligonucleotides by mixing 10 μl of the forward and reverse oligonucleotide (100 μM) each with 80 μl double-­ deionized H2O. Heat to 98 °C for 5 min on a heat block and let cool down slowly to approximately 40 °C by switching off the heat block and leaving the tubes in the block for 45–60 min (see Note 10). Monitor the temperature. 2. Set up following Golden Gate reaction for one guide RNA (scale up as master mix for all guide RNAs):

(a) 50 ng p(U6b-BsaI-gRNA) vector (see Note 11). (b) 1  μl annealed oligonucleotides (10 μM).



(c) 1  μl ATP (10 mM).



(d) 1× Enzyme buffer.



(e) 0.3  μl BsaI.



(f) 0.3  μl T4 DNA Ligase.



(g) X μl double-deionized H2O up to 10 μl total volume.

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}

...GTCGATGGCATGTCATTCCA NGG...

1. target sequence

20 bp long

GTCGATGGCATGTCATTCCA

2. guide sequence

5’ AGTGGTCGATGGCATGTCATTCCA 3’ 5’ CAGCTACCGTACAGTAAGGTCAAA 3’

3. annealed oligos with BsaI overhangs

4. Cloning of annealed gRNA oligos into p(U6b-BsaI-gRNA) TTCGGTCGATGGCATGTCATTCCA CAGCTACCGTACAGTAAGGTCAAA Tc-U6b promotor

... ...AAGC

GTTT... ...

Fig. 3 Guide RNA design and cloning (figure partially modified from [15]). First, a suitable target sequence is selected containing an “NGG” PAM and with a seed sequence that does not occur elsewhere in the genome and, hence, cannot mediate off-target DSBs. The guide RNA sequence should be 20 bp long and does not contain the PAM which is already present in the construct. This guide RNA sequence will be synthesized by annealing two oligonucleotides that include suitable overhangs for the restriction enzyme BsaI. Annealed oligonucleotides (oligos) will be cloned into the plasmid p(U6b-BsaI-gRNA) containing a Tc-U6b promotor sequence and a downstream chiRNA scaffold (crRNA and tracrRNA) by Golden Gate cloning or regular ligation. The U6 promoter needs a G as the first base for transcription. As the backbone already contains a G 5′ of the target sequence (purple G), a G does not need to be part of the original target sequence in order to facilitate proper transcription by the U6 promotor

3. Perform Golden Gate reaction in thermocycler:





(a) heat lid to 40 °C.



(b) 37 °C for 5 min.



(c) 20 °C for 10 min. (d) Repeat steps b and c 10–15 times.

4. Transform 5 μl of the reaction in chemically competent bacterial cells. 5. Sequence guide RNA sequence of each guide RNA plasmid including the U6 promotor and the whole chiRNA scaffold (guide RNA and tracrRNA) (Fig. 3). 6. Prepare transfection-grade mini-preps of all guide RNA plasmids (midi-preps are not necessary for the testing stage) (see Note 12).

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Efficiency to guide the Cas9 nuclease can differ for each guide RNA. Hence, before moving on to the next steps, especially for the HDR approach, efficiency needs to be tested (see Note 13). There are different methods to test guide RNA efficiency. However, in this chapter we focus on the T7 Endonuclease I (T7 Endo I) assay, since it only requires standard lab equipment (see Note 14). The assay involves embryonic injection of guide RNAs individually, along with Cas9, which will lead to different indels in a variety of cells of the developing embryo. A PCR reaction is used to amplify the sequence encompassing the target site(s). Upon denaturing the PCR product and reannealing the single strands, heteroduplexes are formed. At the site of an indel, the base pairing is disturbed such that a T7 Endo I digest will cut both strands at those non-annealed regions. This can be visualized by gel-­ electrophoresis: If two fragments smaller than the amplicon are visible, the guide RNA has successfully mediated Cas9 targeting. The protocol is very similar to the one at www.crisprflydesign.org. Perform following steps:

3.4  Guide RNA Efficiency Test

1. Co-inject the combination of guide RNA (400 ng/μl) and Cas9 plasmid (500 ng/μl) into 100 embryos (needs to be done for each potential guide RNA) (see Subheading 3.6 for injection procedure). 2. Incubate them for three days at 32 °C (see Subheading 3.6). Alongside the injected embryos, uninjected embryos of the same strain should be incubated and DNA isolated identically to the injected condition, as control for potential differences in the sequence of individuals that will also result in heteroduplexes without a Cas9 cut. 3. Extract genomic DNA from injected embryos.



(a) Collect 15–20 L1 larvae that survived and transfer to a 1.5 ml tube.



(b) Add 100  μl squishing buffer and homogenize with a yellow tip at the sides of the tube. Make sure to not use the same tip for a different guide RNA batch.



(c) Incubate the homogenate for 1 h at 55 °C.



(d) Inactivate the Proteinase K at 95 °C for 6–8 min.



(e) Spin the homogenate down for 15 min at 16,000 × g at 4 °C. (f) Use 5  μl supernatant as template for a 50 μl PCR reaction. It is recommended to add 2.5 μl of a 10 mg/ml BSA solution.

4. PCR amplify sequence encompassing target site and perform T7 endo I assay (see Note 15).

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(a) Design primers to amplify ~700 bp, flanking target sites asymmetrically, resulting in two distinguishable bands after T7 endo I treatment.



(b) Run two 50 μl PCR reactions to generate enough material and gel-purify the kit of choice.



(c) Denature and reanneal 400 ng of the PCR product in 1× T7 endo I buffer (total volume 19 μL) as described under Subheading 3.3 with a heat block.



(d) Add 0.7  μl (7 U) T7 Endo I.

(e) Incubate at 37 °C for 20 min.



(f) Stop the reaction immediately by adding 2 μl of 0.25 M EDTA.



(g) Run the assay on a 1.5% agarose gel alongside the same amount of DNA of uncut DNA as control.

If the guide RNA is highly efficient, bands corresponding to the sequence upstream and downstream of the target will be seen. An uncut fraction will be visible as well. Successful genome editing has been performed with guide RNAs where the cut bands were much weaker than the main band at the size of the PCR product. The intensity of uncut and cut bands can thus vary, but do not directly relate to whether the guide RNA works in effect. 3.5  Repair Template and Enhancer Trap Construct Cloning

The construct will be provided in form of a plasmid. Construct cloning greatly differs between the two strategies, and in themselves strategies can be adapted to personal needs and situations (see Notes 16–22). We recommend assembling the repair template not with classic restriction digest/ligation cloning, but using DNA assembly cloning kits [e.g., Gibson Assembly® Cloning Kit (New England Biolabs, Ipswich, USA) or In-Fusion Cloning kit (Takara Bio Inc., Kusatsu, Japan)]. This is especially beneficial when cloning repair templates for bicistronic lines.

3.6  Generation of Enhancer Trap Construct

To generate a universally usable enhancer trap construct, following parts should be assembled: 1. 3XP3-Tc’v-SV40: The recommended eye marker consists of the eye-specific promotor 3XP3 [12], the rescue genomic Tc-vermillion gene and the termination sequence SV40 [49]. This can be retrieved from plasmid #86446 from Addgene. SV40 will work bidirectionally and thus, does not need to be included in the enhancer trap cassette. 2. bhsp68-EGFP: The enhancer trap cassette consists of EGFP and the basal promoter of the heat shock protein 68 (bhsp68) which acts as basal promotor. This cassette should be oriented

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in the opposite direction to the eye marker, so that SV40 can act as terminal sequence for both parts. Sequences can be retrieved from plasmids at Addgene. 3. Exogenous guide RNA target site (Fig. 1): Using megapriming cloning [50], a guide RNA target site should be included that is not found in the Tribolium genome. We used a guide RNA sequence from the Drosophila ebony gene, specifically (GAACCGGGCAGCCCGCCTCC TGG). This sequence should be located near the ends of the construct to be inserted and will lead to linearization of the plasmid thereby facilitating the integration of the repair template by NHEJ. Such a construct has been assembled and successfully tested by the authors. In addition to the abovementioned components, it contains a bicistronic mRNA encoding for EGFP and the Cre recombinase. The plasmid can be obtained from Addgene  [plasmid #124068] and contains the Dm-ebony target site. Prepare transfection-grade midi-preps of all constructs. Check for integrity of these preparations by sequencing at least the coding regions of the repair template or enhancer trap construct. 3.7  Generation of Bicistronic Repair Template

The generation of a bicistronic repair template depends on the guide RNAs of choice and its design needs to await the successful testing and selection of the guide RNAs to be used. See above and Fig. 2 for basic design. Following parts need to be assembled: 1. Backbone: We recommend using pCR™II vector or pJET1.2/ blunt (Thermo Fisher Scientific, Waltham, U.S.A.) as these are small vectors. The size of the complete repair template can be quite large. 2. 5′ homology arm: The ~1 kB 5′ homology arm (see [51] for a detailed analysis on the efficiency of differently long homology arms) should always end right before the gene’s STOP codon irrespective of which guide RNA is used (see Note 23), so that in the repair template sequence you will have 1 kB of ORF sequence fused to the 2A sequence (instead of the STOP) followed by the reporter sequence—all in frame (Fig. 2). 3. 2A peptide: We recommend using the P2A peptide sequence, as this was shown to be most efficient [43]. You can introduce this ~70 bp sequence either similar to the others by amplifying it from a donor plasmid or by having the sequence as part of the primer/oligonucleotide for the assembly reaction. Both have proven to work (see Note 24). 4. EGFP: This sequence can be retrieved from numerous sources. Make sure that the STOP codon is included or that you include it as part of the assembly primer.

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5. 3′UTR: This part also includes parts of the intergenic region. It starts after the gene’s STOP and ends at the cut position of guide RNA 2. Cut positions are always 3 bp upstream of the PAM [52]. 6. 3XP3-Tc’v-SV40: The recommended eye marker consists of the eye-specific promotor 3XP3 [12], the rescue genomic Tc-­ vermillion gene and the termination sequence SV40 [49]. This can be retrieved from plasmid #86446 from Addgene. 7. 3′ homology arm: The ~1 kB 3′ homology arm will start at the guide RNA 2 cut site (Fig. 2). To avoid that the repair template itself will be targeted by Cas9, PAMs must be mutated in the repair template, removing the PAM sequence “NAG” or “NGG.” If the PAM is located in a coding sequence, mutations must be chosen such that the amino acid sequence is not changed. This mutation can be introduced during the amplification processes for the homology arms and 3′ UTR by simply including these modified sequences in the primer sequence. This can be also achieved with a separate PCR mutagenesis or megapriming (see, e.g., [50] for details). Both processes involve amplification of the whole plasmid and a subsequent DpnI digestion to remove the original methylated plasmid which does not contain the intended modification. 3.8  Embryonic Injection

The procedure for embryonic injections is based on standard protocols [53–55]. 1. Prepare injection needles (Use Borosiliate capillaries for injections). Use a P-2000 micropipette puller to form the needle applying the following settings: Heat = 350, Fil = 4, Vel = 50, Del = 225, PUL = 150 or other pullers with respective settings. The capillaries should be similar to those used for Drosophila injection. Open and sharpen the needle either manually using a tweezer or a scissor or by using a microelectrode beveler and check after each sharpening step (see Note 25). 2. Prepare injection mix: Mix all purified plasmids to following concentrations and a volume of 10–20 μl: Repair template and p(bhsp68-Cas9) 500 ng/μl; Inject individual guide RNAs with 400 ng/μl and guide RNA 1 and 2 simultaneously with a concentration of 250 ng/μl each. Mix 8 μl plasmid mix with 1 μl 10× injection buffer and 1 μl Phenol red. Filter-sterilize with a 0.45 μm filter for 0.5 ml tubes by putting mixtures on the filter inside a tube and centrifuge for 5 min at 11,000 × g. 3. Place vw beetles on white flour and let them lay eggs for 1 h at 28 °C. 4. Remove embryos and let them develop further for 1 h at 28 °C.

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5. This can be repeated to get fresh embryos as often as necessary. 6. Wash embryos in 1% bleach (equivalent to 0.05% sodium hypochlorite) for up to 3 min in a 150 μm gaze sieve. Extended bleaching can lead to mortality. 7. Wash embryos thoroughly in room temperature water. 8. Make sure that all flour is removed. 9. Moisten an object slide with water. 10. Transfer embryos on the object slide with a fine brush, arrange them in a line near the long edge of the slide (90 embryos can fit on one side). The more pointed posterior side of the egg should point towards the outside (see Note 26). 11. Place slide on an apple agar plate. 12. Load needle with 4 μl of injection mix. 13. Place needle in the microinjector. 14. Test the needle position and opening by placing it into a drop of Voltalef® oil (VWR) on a slide. 15. The droplet size should be roughly a fifth of the embryo size. 16. Constant pressure should be adjusted so that no liquid is leaking. 17. Inject into first posterior third so that you see either movement in the embryo or red stain. 18. Do not move the needle inside the embryo, do not inject too deeply and not too much; there should be no leakage. 19. Put the slide back on an apple agar plate and collect in an airtight box and keep them at 32 °C for 72 h. High humidity is required for the injected embryos to survive. However, drops of water are deleterious for hatched larvae. Hence, it is important to keep the embryos as long as possible under high humidity, but upon hatching of the first animal they should be dried. 20. Transfer larvae to whole grain flour. 3.9  (Back-) Crossings of G0 to Wildtype

1. Rear injected animals at 32 °C until they pupate, then sex them and keep them separately. 2. Set up single crosses by crossing each injected adult to three vw wildtype animals of the opposite sex. 3. Rear single crosses at 32 °C and remove the parental generation when the next generation starts to pupate.

3.10  Screening for Transgenics in G1

1. Using a suitable stereomicroscope, screen G1 animals for black eyes. The pupal stage is best as these do not move. When screening adults, they have to be anesthetized either by CO2 or by placing them on ice.

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2. For both strategies, keep all positive animals and subsequent offspring as founder lines, as they can differ in quality and strength of the signal. This is especially true for gene-specific enhancer traps. 3. Cross each positive G1 separately to vw to generate offspring that can be screened for the expression in the (developing) brain. 4. Depending on your GOI, its expression amount, and timing, choose a time point where individuals can be easily prepared in a larger scale. 5. Prepare embryos for anti-GFP immunostaining (see Chapter 12), larvae or adults accordingly to detect native GFP expression (after preparation, directly put in mounting medium and imaged), or anti-GFP immunostaining (see Chapter 13). 6. Overlap to the GOI can be tested with an antibody against the protein of interest, if available, or alternatively with in situ hybridization (see Chapter 13). 7. Select three to five founder lines which have the strongest expression, with the highest degree of overlap to the GOI. 3.11  Characterization of the Integration Event

1. Check for proper integration at the expected location by designing primers for amplicons spanning at least the regions surrounding the DSBs or the whole inserted construct. 2. Extract genomic DNA similar to Subheading 3.1. 3. Perform standard PCR. 4. Make homozygous stocks of all of the selected.

3.12  Generating Homozygous Stocks

Having homozygous stocks makes subsequent experiments much easier, be it live-imaging or subsequent genetics. The following protocol is based on genotyping individuals using DNA extracted from wing tissue [56]: 1. Sex individuals as pupae and rear sexes separately until adult stage. 2. On the experiment day, prepare ice, wing buffer, glass slides, forceps, as well as a 24-well plate with small amounts of flour in the wells. 3. Put 0.5 ml tubes on ice, put beetles in a vial on ice for cold anesthetization. 4. Prepare glass side for preparation by wrapping it in parafilm. 5. Put a beetle with one forceps on its right side, head to the left, left elytron up. 6. Hold at the thorax and try to enter under elytra with other forceps carefully.

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7. If the elytron lifts, take out hindwing, and rip and cut first at thicker more distal dark part of the wing and then rip carefully at more proximal parts. 8. Be careful to not remove the whole hindwing as this might leave a wound in the thorax. 9. Put the wing in the tube keeping it cool at all times, put the beetle in a well, with both well and tube marked with a unique identifier for each beetle. 10. Repeat preparation for all. 11. Put wings on −80 °C for 15 min. 12. Add 10  μl of buffer to the wing, and crushing it on the tube wall, rolling it and pushing tissue up and down with a thin pipette tip (this can take 2 min per wing). 13. Spin down shortly. 14. Put tubes on 37 °C for 1 h. 15. Put wings on 75 °C for 20 min. 16. Spin down the evaporated water. 17. Perform PCR.

(a)  The wildtype (wt) amplicon (hence the insertion area without inserted construct) should be always smaller so that you always see the wt band at least if it is heterozygous (since smaller amplicons will outcompete bigger ones in PCR); this makes the PCR less error-prone.



(b) Amplicon size difference should be at least 200 bp and amplicon size between 300 and 800 bp. We recommend ~40 cycles to promote the amplification of the smaller wt band.

18. Put beetles, that are homozygous for the transformation marker, all together and raise homozygous stocks.

4  Notes 1. Any method for genomic DNA isolation will suffice (e.g., kits). These are the protocols we used. 2. Primers should be designed carefully so that unique sites are amplified, the melting temperature is high, and no secondary structures are likely to form. For sequencing primers and the sequencing reactions themselves, please consult the company you are using. If recommendations are followed, mistakes or suboptimal sequencing results may be avoided. 3. The areas to be sequenced can also be subcloned into a blunt cloning vector, such as pJET1.2, if sequencing from an amplicon does not yield good results.

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4. It is highly recommended to use a suitable software that allows in silico cloning, analysis of sequencing data, etc. Use of such will greatly aid the cloning work. 5. There are other design tools available. We have used the one mentioned above nearly exclusively, but we recommend considering other tool websites as well. 6. If a region is particularly low in GC-content, an alternative to Cas9 is CPf1 where the PAM is needs to be T-rich (“TTN”) [57]. 7. If there is an otherwise suitable guide RNA sequence which would mediate cuts in one off target, a T7 Endo I assay can be performed with two regions to be amplified for this guide RNA, one for the aimed target region, one for the off-target region. If the guide RNA mediates DSBs also in the off-target, this will be seen in the T7 endo I assay and the guide RNA can be discarded. 8. Guide RNAs can also be chosen by investigating whether the sequence can form hairpins, including the tracrRNA as this can greatly influence efficiency of the guide RNA. This can be done by the tool http://chopchop.cbu.uib.no/ [58, 59]. So far, the Tribolium genome cannot be used as reference, but you can ask the website administrators to include the genome of choice. 9. It might be advantageous to restrictively target the template strand with a guide RNA of choice, see [60]. 10. Oligo annealing can be alternatively achieved by programming a thermocycler to ramp down to 25 °C with a rate of −0.1 °C/s. This follows the advice found at http://flycrispr.molbio.wisc. edu/protocols/gRNA. 11. It would be worthwhile to test which of the three U6 promotors in the Tribolium genome is the most effective in driving Cas9 transcription, similar to [61]. Two were cloned and no pronounced difference was observed [15]. We have only used p(U6b-BsaI-gRNA). 12. In case transfection-grade kits are not available, use normal kits, but add a precipitation step to increase DNA purity and remove salts:

(a) Mix ~50  μg DNA with 10 μl 3 M Sodium acetate pH = 5.2 (NaOAc) and fill up with H2O to a total volume of 100 μl.



(b) Add 800  μl 100% Ethanol (analysis grade).

(c) Keep at −80 °C for 2 h or o/n.

(d) Centrifuge at 4 °C at maximum speed for 30 min.



(e) Remove supernatant.

(f) Wash with 500 μl precooled 70% Ethanol.

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(g) Centrifuge at 4 °C at maximum speed for 15 min.



(h) Repeat steps e–g once.

(i) Remove as much of the supernatant as possible and air-dry the pellet for ~10 min.



(j) Resuspend in 15–30 μl H2O.

13. If you design gene-specific enhancer traps, you could also skip this part and inject mixes with all available guide RNAs. You would need to inject more embryos, but save time potentially by avoiding this assay. The same is true for when guide RNAs of choice are really close to each other or even overlapping, this is even true for building a bicistronic line. For example, if three guide RNAs for cut-site A and two for cut-site B are strongly overlapping to each other, it might be a good alternative to prepare six different injection mixes instead of testing them beforehand. This is an issue of personal preference and should not be considered as a rule, as it is also dependent on each individual experiment. 14. An alternative to T7 Endonuclease I assays is high resolution melting analysis [62] which requires a qPCR machine, but is more sensitive to detect guide RNA functionality. Both methods can be used. 15. The T7 Endonuclease I assay is very sensitive to the amount of enzyme added, the temperature, as well as the length of incubation. So make sure that the amounts as well as the timing are exact. 16. It could be advantageous to design a way to remove the eye marker after stocks are made homozygous. In Drosophila this is usually achieved by introducing loxP sites [28]. In Tribolium the option is to introduce exogenous sequences from Drosophila such as guide RNAs of ebony on both sides of the eye marker via PCR mutagenesis. Then after the Tribolium stock was made homozygous using the eye marker, it can be removed by injecting the line with the suitable guide RNAs for the introduced target sequences. A fraction of injected embryos will lose the eye marker and those can be used for further experiments. This strategy reduces the potential influence of the 3XP3 promotor on the gene’s and marker gene’s expression. We note, however, that we have so far not observed strong influence of the eye marker. 17. An alternative strategy could be using only guide RNA 1 and putting the eye marker directly after a SV40 sequence following the fluorescent protein. This is especially useful if the 3′UTR of your gene of interest is too big to be considered as part of the construct. The eye marker influence can then be reduced by cutting the eye marker out (as in Note 16).

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18. The repair templates can be modified in multiple fashions. One could substitute EGFP with other fluorescent proteins of other colors or brightness, or with Gal4 to cross with suitable UAS lines. Also tricistronic lines with the gene, a fluorescent protein and Gal4 are possible. However, the size can be a constraint for cloning and HDR and can be a potential disadvantage. Ideas for multiple variations can be potentially gathered in [28]. 19. Repair template size can be a constraint, as rates of integration might decrease. We have successfully integrated 2.5 kB, but the homologous recombination mechanism itself was employed integrating constructs as big as 13 kB [63]. 20. You can also provide Cas9 as protein which is more efficient than providing Cas9 via a plasmid [64]. 21. Cas9 expression is driven by bhsp68. It would be better to have Cas9 under control of a germ-line-specific promoter. 22. Providing guide RNA and a repair template as single-stranded DNA can increase CRISPR efficiency drastically [28]. In most cases, single-stranded repair template had strong size restrictions, so that our strategies could not have worked. However, using a production system especially for long single-stranded DNAs can circumvent these limitations (e.g., https://www. takarabio.com/products/gene-function/gene-editing/ crispr-cas9/long-ssdna-for-knockins). 23. For bicistronic construct cloning, you can already start amplifying the 5′ homology arm as this always ends with the last codon before the STOP and is, hence, independent of guide RNA choice. The design of 3′ UTR and 3′ homology arm depend on the chosen guide RNA, so this needs to be postponed to after guide RNA tests (but see Note 13). 24. The sequence for the P2A peptide is 95 °C for 5 min. Cool rapidly. 8. Change solution to 100 μl hybridization solution containing 50 ng of RNA probe (see Note 8). Incubate at 50–55 °C for at least one night and up to three nights. Longer incubations give better results but are not necessary for many probes used. Day 2 9. Carefully remove probe solution and keep it at −20 °C. Probes can be reused many times and results become cleaner with repeated use of probe solution. 10. Wash embryos in Wash solution 1 for 20 min at 50–55 °C. Repeat. 11. Wash embryos in Wash solution 2 for 20 min at 50–55 °C. Repeat (see Note 10). 12. Wash embryos in Wash solution 3 for 20 min at 50–55 °C. Repeat. 13. Wash embryos in Wash solution 4 for 20 min at 50–55 °C. Repeat. 14. Wash in PBST for 5 min. 15. Incubate in 100 μl blocking buffer for 30 min.

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16. Replace with fresh 100 μl blocking buffer containing 1/2000 anti-digoxigenin-AP antibody and incubate overnight at 4 °C (see Note 14). Day 3 17. Wash 3× 5 min in PBST. 18. Wash 3× 1 h in PBST. 19. Wash in 100 μl TMNT solution, 2× 5 min. 20. Replace solution with 100 μl TMNT solution containing 6.6 μl NBT and 3.5 μl BCIP per ml of TMNT. 21. Incubate in the dark until the desired staining intensity has been reached (see Note 15). 22. Postfix embryos in 100 μl of 4% paraformaldehyde in PBS for 1 h at room temperature or overnight at 4 °C. If embryos are to be processed for nuclear staining, embryos should be stored in fix until further processing. Parafilm is placed around the top of the microcentrifuge tube and embryos can be stored for up to a few weeks at 4 °C (see Note 16). 3.4  Mounting and Photography

1. Fixed embryos are rinsed in PBST (3 washes) to remove fixative. 2. Remove as much PBST as possible and replace with a drop of Vectashield-DAPI (from the supplier’s pipette). 3. Transfer embryos in Vectashield-DAPI into a depression slide containing a small pool of Vectashield-DAPI. When not under stereomicroscope protect from light with an aluminum foil-­ covered petri-dish lid. 4. Take one embryo and place in a tiny drop of Vectashield-DAPI onto a glass slide. 5. Take a coverslip and, using a Vaseline dispenser (Fig. 2c), place one dot of Vaseline on each corner of the coverslip. 6. Place the coverslip over the embryo so that the embryo is in the center (Fig. 2d). 7. Under a stereomicroscope carefully press down until the coverslip touches the Vectashield liquid (without squashing the embryo). Gently adjust the position of the coverslip until the embryo is in the desired orientation for photography. Pushing on the side of the coverslip with a 200 μl micropipette cone is my preferred method. 8. Photograph the embryo first under bright field, then UV fluorescence for nuclear stain. The embryo should be fixed in position for this process. It is important to obtain a good brightfield image first, as recent lots of Vectashield-DAPI occasionally cause the ISH stained embryos to darken after UV exposure (e.g. Fig. 4). While this effect only takes place from time to

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Fig. 4 Vectashield-DAPI problem. Embryo stained with DllB probe prior to (a) and after (b) UV exposure

Fig. 5 Adobe Photoshop DAPI-ISH overlay. (a) Screen shot of Gsx ISH in Photoshop. (b) Screenshot of DAPI image after conversion to grayscale. (c) Overlay of DAPI shot (50% transparency) and ISH image. (d) Contrast, light-adjusted, rotated, and cropped image of Gsx ISH with DAPI stain

time, photographing the bright field first ensures the sample is not lost (see Notes 17 and 18). 9. Alternatively, nuclei can be stained with Hoechst. After fixation and washing in PBST, embryos are placed in 100 μl PBST with 1/1000 dilution of the Hoechst stock solution for 15 min in the dark. Embryos are then washed 2× 10 min in PBST, then washed overnight at 4 °C in PBST. After the overnight wash, embryos are placed in 100 μl 80% glycerol for 30 min and mounted as described above in 80% glycerol (see Notes 17 and 18). 10. To make ISH plus DAPI images, an overlay is made in Photoshop (Fig. 5). Open both images in Adobe Photoshop or similar program. 11. Change the DAPI image to gray-scale. 12. Adjust contrast and light for both images. 13. Copy and paste gray-scale DAPI image on top of the bright field image. 14. Using the “calques” window, adjust the transparency of the gray-scale image (to around 50%) to see at the same time both the nuclei and the ISH stain.

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A

B

C

V/VI

V/VI III/IV

III/IV I/II

3-row

II I

V/VI

V/VI

III/IV

III/IV

I/II

II I

row I/II dividing

D

E

VI V III/IV II I

II I

VI V IV III II I

row V/VI dividing III/IV starts to divide

VI V IV III II I

VI V III/IV

VI V IV III II I

VI V IV III II I

row III/IV dividing early 6-row stage

6-row

Fig. 6 Stages 11–13 DAPI images (3-row stage to 6-row stage). Top: DAPI-stained images. Embryo is outlined in blue, brain precursors are outlined in red. Middle: drawings of neural plate and surrounding nuclei of the corresponding embryo (top). The embryo is outlined in blue, the brain precursors are outlined in red and rows of cells are indicated. Dividing cell nuclei are connected by black bars. Bottom: schematic drawings of neural plates representing the embryo above. (a) 3-row neural plate. (b) The medial three pairs of cells (columns 1–3) of row I/II are dividing. (c) Row V/VI is dividing, one cell in row III/IV is dividing. (d) Row III/IV is dividing. (e) 6-row neural plate stage

15. Fuse visible calques to make one image. 16. Adjust brightness, contrast, rotate and crop image to desired size. 3.5  Staging

Comparing a new ISH plus DAPI-stained embryo series to the documented pattern of cell divisions should enable the precise identification of which cells express a given gene. The pattern of cell divisions is illustrated in Figs. 6, 7, 8 and 9 and described below. In Figs. 6, 7, 8 and 9 I have provided, as a guide, DAPI-­ stained images showing the progression of neural lineage divisions from approximately stage 11 (early gastrula) to stage 15 (mid ­neurula) of development [32]. In Figs. 6 and 8, at the top, are DAPI ISH-stained embryos, beneath is a drawing highlighting the nuclei in the neural plate and surrounding cells, with the corresponding rows of neural plate cells labeled, the brain precursors outlined in red and the embryo outlined in blue. At the bottom is a schematic drawing of the neural plate with the cell divisions shown and brain precursors outlined in red. Figure 6 covers stages 11–13 (“3-row” to “6-row” neural plate stages). Figure 7 shows the same embryos with ISH signals, detailing expression patterns in the neural plate cells on the schematic drawing (bottom) by gray

In Situ Hybridization in Ascidian Brain Precursors A- Fgf9/16/20

B- Fgf9/16/20

V/VI

V/VI III/IV

III/IV I/II

3-row

C- Snail

II I

V/VI

V/VI

III/IV

III/IV

I/II

II I

row I/II dividing

D- Fgf9/16/20

E- ZnF266

VI V III/IV

VI V IV III

VI V III/IV

II I

II I

II I

VI V IV III II I

VI V IV III II I

VI V IV III II I

row V/VI dividing III/IV starts to divide

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row III/IV dividing early 6-row stage

6-row

Fig. 7 Stages 11–13 ISH images. Same embryos as Fig. 6, but showing ISH-DAPI merged images. On the schematic drawings (bottom), the cells expressing each gene are colored in gray

filled boxes. Similarly Fig. 8 covers stages 13–15 (“6-row” to “12row” neural plate stages) with Fig. 9 illustrating the same embryos with ISH signals. At the early gastrula stage, the neural cells are arranged in three rows, row I/II (precursors to rows I and II), row III/IV, and row V/VI (Figs. 1b and 6a). Cells within each row of the neural plate divide in a stereotypical fashion (Fig. 6) [24]. Firstly, cells in row I/II divide (with the medial columns of cells starting to divide before the lateral columns of cells) (Fig. 6b). Then cells in row V/ VI and finally cells in row III/IV divide (Fig. 6c, d). This gives rise to the 6-row neural plate with the six rows (I–VI) of cells aligned along the posterior-anterior axis, respectively (Figs. 1c and 6e). Between the 6-row and 12-row neural plate stages, row II cells start to divide first followed by row I (Fig. 8a–c). Next, rows III, V and VI divide approximately at the same time, but variably, depending on the embryo (Fig. 8d–g). Row VI is often hard to see as it is situated close to the anterior tip of the embryo (gray boxes in Fig. 8f, h, i). The variability in the exact temporal sequence of divisions of rows III, V and VI is made clear in Fig. 8. In Fig. 8d–g, rows I and II have divided. In Fig. 8d, row VI cells are dividing before rows III and V. The embryo in Fig. 8e on the other hand shows division of row V before rows III and VI. In Fig. 8f, cells in rows III and V are dividing (row VI is not visible). In Fig. 8g, cells in rows V and VI are dividing before row III. Nonetheless, it is still relatively easy to identify each cell nuclei. The divided row I nuclei

A

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C

VI V IV III II I

VI V IV III II I

VI V IV III II

VI V IV III II

I

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row II dividing

F

II I

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row V, VI dividing

I

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I

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VI V IV III II I

row IV dividing 12-row neural plate

row V dividing

J

V IV III

II

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VI V IV III II I

II

row VI dividing row V compacted nuclei

H

V IV III

VI V IV III

VI V IV III II I

row I dividing

G

E

VI V IV III II

VI V IV III II I

row II starts to divide

row III, V dividing

D

II

VI V IV III II I

all cells divided once row II starts to divide again

Fig. 8 Stages 13–15 DAPI images (6-row to 12 row stage). Top: DAPI-stained images. Embryo is outlined in blue, brain precursors are outlined in red. Middle: drawings of neural plate and surrounding nuclei of the corresponding embryo (top). The embryo is outlined in blue, the brain precursors are outlined in red and rows of cells are indicated. Nuclei of cells which have divided from the 6-row neural plate stage are connected by black bars. Bottom: schematic drawings of neural plates representing the embryo above. Cells shown in light gray could not be clearly identified on the DAPI image. (a) Row II cells start to divide. (b) Row II cells are dividing. (c) Row II cells have divided, row I cells are dividing. (d–j) Row I and II cells are divided. (d–g) Embryos are at a similar stage, but show cell divisions in different orders between rows III, V and VI. (d) Cells in row VI are dividing. (e) Cells in row V are dividing. (f) Cells in rows III and V are dividing (row VI not visible). (g) Cells in rows V and VI are dividing. (h) Cells in rows III and V have divided and cells in row IV are dividing. (i) Cells in all rows have divided, row II cells have divided again. (j) Top: close up of embryo and nuclei drawing in (i) showing characteristic organization of row III cells. Bottom two panels: the characteristic organization of row III cells is even clearer in this embryo, in which there is no ISH stain. Brain precursors are outlined in red and row III cell nuclei are colored in red in the nuclei drawing

In Situ Hybridization in Ascidian Brain Precursors A Delta-like

B Msx

C Foxb

VI V IV III II I

VI V IV III II I

row II dividing

F Foxc

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I

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V IV III II

II

row IV dividing 12-row neural plate

I

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VI V IV III II I

II

VI V IV III II I

row VI dividing row V compacted nuclei

I

VI V IV III

II

H Msx

V IV III

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row I dividing

G Foxc

E Efna.b

VI V IV III II

VI V IV III II I

row II starts to divide

D Efna.b

339

VI V IV III II I

all cells divided once row II starts to divide again

Fig. 9 Stages 11–13 ISH images. Same embryos as Fig. 8, but showing ISH-DAPI merged images. On the schematic drawings (bottom), the cells expressing each gene are colored in gray. In (j) the close up of (i) is shown with the stained cells indicated in purple on the nuclei drawing

are small and bright and the medial four columns of cells are curved along the dorsal marginal lip (e.g. Fig. 8d, e). Divided row II cell nuclei are large and diffuse also providing good landmarks for orientation. Row IV is the last row of cells to divide (Fig. 8h) rapidly followed by another division of row II cells (Fig. 8i). At the mid

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neurula stage (Fig. 8i) the brain precursors are in “12-row” stage. At this stage, row III cells serve as good landmarks: the nuclei of column 3 sister cell pairs are separated, while the nuclei of the anterior column 1 sister cells are more compacted and closer together than those of the posterior sister cells (Fig. 8j, row III in red). At later stages, the exact identification of lateral row I and II cells becomes difficult with this technique due to their intercalation (gray boxes). While using the terms 3-row, 6-row and 12-row stages are useful for illustration purposes, it should be noted that the cells are dividing continuously and there is no “pause” at these stages.

4  Notes 1. BSA is added to ASW to help prevent the embryos from sticking to the sides of the microcentrifuge tubes prior to fixation. Low adhesion 1.5 ml tubes should be prerinsed in 1 ml of 0.5% BSA in ASW. Leave some BSA-ASW (100–500 μl) in the tube to add the embryos to. Add embryos near the bottom of the tube with the homemade micropipette, they will sink rapidly. Remove as much liquid as possible prior to fixation. 2. Paraformaldehyde is prepared as a 40% solution with heating at 65 °C and stirring to dissolve with addition of NaOH (to pH 7.5) and frozen in small aliquots. Alternatively, it can be bought commercially. NaCl is prepared as a 5 M stock solution and MOPS buffer is as a 1 M pH 7.5 stock solution. 3. Tween20 is viscous, so a 20% solution is prepared, which is much easier to pipette accurately. 4. PBS is made as a 20× solution (160 g/l NaCl, 28.84 g/l Na2HPO4.2H2O, 4 g/l KCl, 4 g/1 KH2PO4) filtered using a Nalgene Supor machV filtration unit and diluted as required. We do not adjust the pH. 5. Proteinase K treatment makes embryos stick to each other. Therefore we opted for a light proteinase K treatment at room temperature instead of 37 °C. If embryos appear “sticky” after ISH, try reducing the proteinase K concentration or incubation time. 6. We buy commercially prepared 20× SSC solution. We prepare 100× Denhardt’s solution (2% Ficoll 400, 2% polyvinylpyrollidone, 2% BSA), which is stored frozen in aliquots. Torula yeast RNA is prepared as a 100 mg/ml stock and stored frozen according to the published protocol [33]. 7. We add dextran sulfate to our hybridization solution following [34]. After testing several probes (“easy” and “difficult”) we found that adding dextran sulfate improved some probes tested

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but had little effect on others. Since it did not appear to have any detrimental effects, we systematically add it to our hybridization solution. 8. Antisense RNA probe synthesis is carried out using standard procedures (e.g. [35]). We use ProbeQuant G-50 micro-­ columns to purify probes, according to the manufacturer’s instructions (Table 1). While we use 50 ng probe in 100 μl of hybridization solution as standard, this is sometimes modified during trouble-shooting. If a probe gives rise to too much “background” then it is diluted further (i.e. 25 μg, 10 μg, 5 μg per 100 μl hybridization solution) and tested to determine the best probe concentration. Frequently we found that dilution of the probe concentration resulted in dramatic improvement for “difficult probes.” However, this results in longer color reaction times. 9. We reduced the amount of SDS in the original protocol for the washes from 1% to 0.2%. This decreased the time taken for the color reaction without significantly increasing background for several probes tested. 10. Optionally RNase treatment can be carried out between wash solutions 2 and 3. However, we found that for standard ISH for several genes tested, RNaseA treatment reduced Table 1 Unique identifiers for Ciona robusta genes. The ISHs presented here were carried out on Ciona intestinalis (Roscoff, France). Gene models for Ciona intestinalis are not currently available. Therefore, here are listed the common gene name used and the unique gene identity number for Ciona robusta

Gene name

Ciona robusta (unique gene identity)

Fgf9/16/20

Cirobu.g00004295

Snail

Cirobu.g00005955

ZnF266

Cirobu.g00001049

Delta-like

Cirobu.g00012743

Msx

Cirobu.g00005203

Foxb

Cirobu.g00006425

EfnA.b (ephrin Ab)

Cirobu.g00005364

Foxc

Cirobu.g00012813

Gsx

Cirobu.g00005160

DllB

Cirobu.g00008660

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foreground without significantly decreasing background. However, when we use ISH to detect lacZ driven by upstream regulatory sequences, I found that RNase treatment was essential. If background is high, it is worth testing if RNase treatment improves the result. RNase treatment is carried out after wash solution 2 washes. Embryos are rinsed in wash solution 3 at room temperature three times. Then 100 μl RNase A solution is added (20 μg/ml RNase A in 2× SSC) for 20 min at 37 °C. Embryos are then rinsed 3× 5 min in room temperature in wash solution 3 before rejoining the basic protocol with the 2× 20 min wash solution 3 washes at 50–55 °C. 11. Roche blocking reagent is made as a 10% stock solution in maleic acid buffer (0.1 M Maleic acid pH 7.5; 0.15 M NaCl) with heating and vortexing. Solution is stored in frozen aliquots. 12. Whole mount ISH is performed on embryos developed from dechorionated eggs. However, it is known that the chorion is essential for the later left-right asymmetry of the ascidian larval brain [36]. This technical caveat should certainly be considered when analyzing embryos developed from dechorionated eggs, particularly if one is interested in left-right patterning/ asymmetry. This issue can be avoided by dechorionating embryos just prior to fixation [21, 36]. 13. Embryos can alternatively be fixed for 90 min at room temperature. 14. For the standard protocol the embryos are incubated in the antibody solution overnight at 4 °C. This incubation can be extended to two nights or even three and good results can be obtained. It is also possible to incubate for 1.5–2 h at room temperature but I would not recommend this for difficult probes. 15. During NBT-BCIP staining, embryos are checked approximately every 30 min. Incubations can take as little as 30 min or up to 3–6 days. When long incubations are required, embryos are rinsed and stored in PBST at 4 °C overnight and then steps 19 and 20 followed the next morning. After a few days, if there is no background staining, embryos can be left overnight in the color reaction solution. However, the solution should be changed regularly (minimum morning and evening). For difficult probes if background levels start to rise, it is best to immediately wash embryos in PBST overnight at 4 °C and then resume staining. Once high background levels are reached, it is much more difficult to remove the background, but washing overnight (or several nights) in PBST at 4 °C

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or room temperature with agitation can sometimes rescue an experiment. 16. If nuclear staining is not required, embryos can be washed with PBST to remove fix and stored in 80% glycerol. Embryos stored in this way remain stable and can be rephotographed even after several months of storage. 17. The choice between Vectasheild-DAPI and Hoechst is difficult. The quality of the nuclear staining is similar. The advantage of Vectasheild is that embryos can be mounted immediately without the overnight wash and it is easier to handle than glycerol. The disadvantage is that, sometimes, for unknown reasons, the embryos go dark after UV exposure. This problem will affect all embryos in a batch or none and only happens from time to time. Each manipulator should test which of these techniques is best suited to him/her. For Hoechst stained embryos, adjusting the amount of glycerol between 50 and 80% is also possible as decreased transparency (with 50% glycerol) is sometimes better for probes with staining in internal tissues. 18. The “going black” issue was also noted by another ascidian researcher (Hiroki Nishida and Shih Yu, Osaka University, Japan; personal communication). Their current method of DAPI staining and mounting embryos to avoid this problem is as follows. Embryos are first stained with DAPI (Final concentration 0.1–0.2 μg/ml PBS) for 20 min and then washed with PBS once for 10 min. Embryos are mounted in 80% glycerol (in PBS) 0.5% n-Propyl gallate. Embryos treated by this method usually show a strong background just after mounting in 80% glycerol, but the background decreases in one day after mounting.

Acknowledgments The author is a CNRS researcher in the laboratory of Hitoyoshi Yasuo. I would like to thank H. Yasuo and Cathy Sirour for their help and for critical reading of the manuscript. Many thanks to Hiroki Nishida and Shih Yu (Osaka University) for kindly sharing their DAPI staining and mounting protocol and agreeing to publish it in this chapter. This work was supported by Central National de la Recherche Scientifique (CNRS), Sorbonne Université, Sorbonne Université Emegence project (2016), and the Agence Nationale de la Recherche (ANR-09-BLAN-0013-01, ANR-17CE13-0003).

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32. Hotta K, Mitsuhara K, Takahashi H et al (2007) A web-based interactive developmental table for the ascidian Ciona intestinalis, including 3D real-image embryo reconstructions: I. From fertilized egg to hatching larva. Dev Dyn 236:1790–1805 33. Jing L (2012) Preparation of Torula yeast RNA for Hybe solutions. BIO-Protoc 2 34. Lauter G, Söll I, Hauptmann G (2011) Two-­ color fluorescent in situ hybridization in the embryonic zebrafish brain using differential detection systems. BMC Dev Biol 11:43 35. Thisse B, Thisse C (2014) In situ hybridization on whole-mount zebrafish embryos and young larvae. In: Nielsen BS (ed) In situ hybridization protocols. Springer New York, New York, NY, pp 53–67 36. Shimeld SM, Levin M (2006) Evidence for the regulation of left-right asymmetry in Ciona intestinalis by ion flux. Dev Dyn 235: 1543–1553 37. Conklin EG (1905) The organisation and cell lineage of the ascidian egg. Science 23: 340–344

Chapter 19 Spawning Induction and Embryo Micromanipulation Protocols in the Amphioxus Branchiostoma lanceolatum Yann Le Petillon, Stéphanie Bertrand, and Héctor Escrivà Abstract In the last decades, the cephalochordate amphioxus has reached a peculiar place in research laboratories as an excellent animal model to answer Evo/Devo questions. Nevertheless, mainly due to its restricted spawning season and to the small size of its embryos, only a few basic techniques in developmental biology could be used until recently. Fortunately, these last years, and thanks to the development of high-­ throughput techniques, new technical approaches have been possible, such as comparative transcriptomics and/or genomics. However, classic micromanipulation techniques are still difficult to apply. Here we present simple protocols for the manipulation of amphioxus embryos. First, we present the spawning induction method used with the European amphioxus species Branchiostoma lanceolatum. Second, we explain simple methods to manipulate the developing amphioxus embryo during the first steps of its development (before the hatching stage). These methods open many technical possibilities for future functional studies. Thus, we present here a simple technique to efficiently dechorionate a large number of embryos, we detail a protocol for the dissociation of cells during the first steps of the embryonic development and, finally, we describe micromanipulation approaches for tissue isolation during the gastrula stage. Key words Amphioxus, EvoDevo, Spawning induction, Micromanipulation, Dissection, Graft experiment

1  Introduction The cephalochordate amphioxus is a marine animal that occupies an extremely interesting phylogenetic position for researchers interested in the origin and evolution of vertebrates since its lineage diverged very early during the evolution of chordates [1, 2]. In addition, unlike the third chordate phylum, the urochordates, cephalochordate morphology, anatomy, and genomic characteristics have diverged very little from those assumed to have existed in the ancestor of chordates. In the last decades, different laboratories around the world have promoted amphioxus as a key and amenable organism to answer different Evo/Devo questions in research teams interested in the non-vertebrate chordates to vertebrates

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evolutionary transition [3]. Nevertheless, this animal model still suffers from technical limitations, mainly due to two aspects: 1. The limited access to living embryos. All the different amphioxus species around the world (i.e. about 30 different species are described [4]) have a restricted spawning season. Depending on the species, the natural spawning season is limited to 2–5 months [3], thus hampering the possibility to develop new techniques for embryological studies. Moreover, a given animal might spawn once or twice per month during this season and the time of natural spawning cannot be predicted. Hence, efforts have been made to try to control the spawning behavior of the animals in captivity. These efforts include changes of different laboratory conditions (control of the sea water temperature, feeding rhythm and diet, day/night cycle, etc.). Thus, spawning induction in captivity of the Mediterranean amphioxus, Branchiostoma lanceolatum, on a daily basis during the spawning season was one of the most important advances in the field in recent years [5]. This method, or small variations of it, is also applied today in other species around the world (B. belcheri in Asia and B. floridae in USA) showing its reliability regardless of the amphioxus species. 2. The difficulty to work on individual embryos for microinjection or micromanipulation. Even if it is easy to obtain important amounts of eggs during a spawning event (e.g. an adult female may spawn around 2000 eggs), it remains difficult to work on individual eggs/embryos due to their small size and fragility. The size of the egg ranges from 80 to 110 μm and is unchanged until the gastrula stage. This makes the manipulation of the embryo extremely difficult, for example, in approaches using grafts. Moreover, in addition to the size of the egg, its physical properties, such as the strength of the vitelline membrane or the intracellular pressure, make the egg extremely fragile and also complicates microinjection approaches. In this chapter, we present some techniques that we have developed over the last few years and that allowed us to obtain some interesting results in different studies despite the difficulties involved in working with amphioxus embryos. We first detail the technique allowing the spawning induction of amphioxus. Second, we present a very simple but efficient way to get important amounts of dechorionated embryos from the 1-cell stage. This technique offers the possibility of developing new microinjection or electroporation approaches. Finally, we present a protocol for the manipulation of the embryos and ectodermal explants allowing cell/tissue excision and graft experiments during embryonic development. Taken together, these protocols offer new tools to improve classical embryology as well as genetic manipulation experiments in amphioxus. As an example, these methods have recently been used in the reproduction of the classic Spemann-Mangold graft experiments for the demonstration of the presence of an organizer in amphioxus [6].

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2  Materials 1. Open circuit of natural sea water. 2. Tank of 5 l with 3–4 cm of sand thickness. 3. Algae (Isochrysis galbana, Tisochrysis sp., Dunalliela sp.). 4. Day/night cycle and temperature control systems. 5. Incubator dedicated to the thermal shock, with day/night cycle and temperature control systems. 6. Transparent plastic cups of 4–5 cm diameter. 7. Black background. 8. Portable red light. 9. 0.2 μm filtered sea water (FSW). 10. Agarose (molecular biology grade). 11. 5.5 cm diameter Petri dishes. 12. Microscalpel (Micro Feather Ophthalmic Scalpel with Aluminum Handle, 45°). 13. Eyelash. 14. Pipette type P20. 15. Pipette type P200. 16. Tips 0.1–20  μl (Tips 20 μl, neutral, Sarstedt). 17. Tips 20–200  μl (Tips 200 μl, Sarstedt). 18. Binocular microscope.

3  Methods 3.1  Preparation of Petri Dishes for Dechorionation and Micromanipulation

1. Petri dishes (5.5 cm) are rinsed with FSW. 2. Fill the Petri dishes with 0.8% agarose/FSW in order to cover the bottom. Replace the lid immediately after pouring the agarose/FSW (see Note 1). 3. As soon as the agarose/FSW is solidified in the Petri dishes, add enough FSW to cover the whole layer of agarose/FSW. 4. Conserve at 4 °C (see Note 2).

3.2  Eyelash and Tips Preparation and Conservation

1. Stick the root of an eyelash to the interior of the extremity of a 200 μl tip with nail polish, and wash it in distilled water for at least 24 h before use. 2. During the experiments, conserve both the tips dedicated to the transfer of explanted tissue/cells and the eyelash fixed to the tip plunged in FSW. 3. Replace the FSW every 2–3 days.

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4. The eyelash is delicately wiped with a clean paper before being used in order to remove any impurity and to avoid it to stick to the embryos. 5. After use, the eyelash is washed with fresh water, rinsed with FSW and conserved as mentioned above. 3.3  Spawning Induction Protocol

All experiments described here detail the protocol for the amphioxus species Branchiostoma lanceolatum. For other amphioxus ­species, the protocol should be adapted (particularly water temperature). The amphioxus used here are collected near the Racou beach (Argelès-sur-Mer, France) from May to July, during their natural spawning season when the gonads are mature [5]. 1. Keep ripe animals at 17 °C in tanks with sand in a sea water open circuit (see Note 3). Feed the animals 2–3 times per day and with a 14/10 h day/night cycle for a period of time ranging from 2 weeks for animals collected early during the season (early May) to 3–5 days for animals collected later (June and July). 2. In order to induce the spawning, place the amphioxus at 23 °C in a FSW tank without sand, keeping the day/night cycle during the afternoon (2–3 h before the light is off). 3. 24 h later (2–3 h before the light is off), separate the animals individually in small plastic cups with 20–30 ml of FSW (without sand) (see Note 4). Place the plastic cups on the black background in a quiet room at 19 °C with the light on and turn off the light when the night is supposed to start in the previous day/night cycle (see Note 5) (Fig. 1).

Fig. 1 Collection and preparation of the spawning induction of the amphioxus B. lanceolatum. The animals are collected by dragging the sand with a Charcot-Picard dredge (a) and manual sieving (b). Following the thermal shock from 17 °C to 23 °C during 24 h, the amphioxus are brought to a quiet room at 19 °C (c) and separated in individual transparent plastic cups (d). The animals begin to release their gametes 1 h after the light is turned off

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4. Around 1 h after the light is turned off, the first animals start to spawn [5]. Use a portable red light to check for spawning. Then, transport the plastic cups with the animals and their gametes in another room and leave the animals that did not spawn yet in the dark. Most likely, they will spawn latter. 5. Recover the sperm with a pipette and keep it diluted in the sea water at 4 °C to use it for several hours (see Note 6). 6. Transfer gently the eggs in a scratched Petri dish (see Note 7). 7. Fertilize the eggs with 60–70 μl of sperm/FSW per Petri dish (see Note 8) and check the efficiency of the fertilization by looking a few minutes later if the fertilization membrane has swelled up. 3.4  Dechorionation Protocol

1. Concentrate in the middle of the Petri dish the fertilized eggs after the fertilization membranes reach their maximum size (see Note 9). 2. Between 30 min after fertilization until the first cell division (around 1 h at room temperature, 22/23 °C), fill a tip by pipetting 20–30 μl of FSW then 80 μl of FSW with concentrated embryos and finally 20–30 μl of FSW again (see Note 10) (Fig. 2). 3. Pipette the eggs mightily toward the border of an agarose Petri dish (Fig. 2). 4. Remove the non-dechorionated embryos from the agarose Petri dish (see Note 11). 5. Remove the fertilization membranes floating. 6. Separate dechorionated embryos from each other by moving laterally the Petri dish strongly enough in order to avoid embryos sticking to each other during their development.

Fig. 2 Representations of the dechorionation technique. Schemes represent: (a) the proportion of FSW/FSW+ embryos/FSW in the tip during the pipetting step; (b) lateral and (c) top views of the orientation of the tip in the FSW in an agarose Petri dish in order to maximize the amount of dechorionated embryos during the expulsion of the embryos from the tip

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3.5  Dissociation of Cells and Transfer of Dissociated Cells to a Different Petri Dish

This manipulation is designed for experiments using dechorionated embryos. Thus, the manipulation takes place in agarose Petri dishes as in Subheading 3.1. The example below describes the ­dissociation and the transfer of dissociated cells from an 8-cell stage embryo. Dissociation of cells 1. Position the embryo using the eyelash in the expected orientation by weakly moving it or creating a flow of FSW around it. 2. Just after the cell division from 4- to 8-cell stage, wait until the developmental timing when the eight cells are almost entirely separated (see Note 12). 3. Induce a weak pressure perpendicular to the agarose with the eyelash at the level of the separation zone between the cells (Fig. 3) (see Note 13). Transfer of dissociated cells 4. Mount a 0.2–20 μl tip on a 2–200 μl tip and use a pipette type 20 calibrated for a volume of 15 μl (see Note 14). 5. Fill the tip with 5 μl of FSW, then 5 μl of FSW with the dissociated cells and then 5 μl of FSW (see Note 15). 6. Gently drop the dissociated cells in the new Petri dish directly in the FSW (see Note 16).

Fig. 3 Scheme of the animal/vegetal dissociation technique at the 8-cell stage. (a) Schematic representation of the position of the 8-cell stage embryo over the agarose plate in order to dissociate animal (green cells) from vegetal cells (blue cells). The most stable position is shown here, allowing the most perpendicular movement to separate the cells. Gray arrows show the direction of the eyelash movement to dissociate the cells. (b) Picture of the extremity of an eyelash and blastomeres after the separation of the animal (right) and the vegetal cells (left) at the 8-cell stage. Note the obvious difference of size between the blastomeres. Scale bar: 100 μm

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3.6  Dissection During Gastrulation

Depending on the experimental need, we will divide this section in two distinct parts. These protocols are designed to recover part of the mesendoderm, of the ectoderm, or of the blastoporal lips during gastrulation (see [6]).

3.6.1  Dissection of Non-dechorionated Gastrula Embryos

This technique of ectodermal dissection relies on the constant position of the embryo, which is immobile inside the fertilization membrane until the activation of the cilia movement, which occurs at the end of gastrulation. The gastrula stage starts when the vegetal pole, which will develop into the mesendoderm, begins to flatten (5.5–6 h post-fertilization at 19 °C in B. lanceolatum). Then the mesendoderm invaginates into the blastocoel (6–7 h post-­ fertilization at 19 °C in B. lanceolatum) (Fig. 5). Dissection is undertaken before the mesendoderm touches the ectoderm. 1. In a scratched Petri dish filled with FSW, use the microscalpel to create micro-grooves (many weak stripes) or make three strips separated of 50 μm (i.e. a third of the fertilization membrane diameter) at the bottom of the Petri dish. 2. Roll the embryo on the micro-grooves or on the stripes in order to place it in the expected orientation (the dorsoventral axis perpendicular to the bottom of the plate) (Fig. 4). 3. Cut the embryo and the fertilization membrane with the microscalpel, with a movement from the top to the bottom (see Note 17). 4. Transfer the obtained ectodermal explants as described in the Subheading 3.5, steps 5 and 6.

Fig. 4 Scheme showing the ectodermal explantation in a non-dechorionated embryo at the early gastrula stage. The pink plan represents the orientation of the cutting movement with the microscalpel through the fertilization membrane and the early gastrula in order to separate the most anterior part of the ectoderm from the rest of the embryo

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3.6.2  Dissections of Dechorionated Gastrula Embryos

The gastrula stage used in this experiment corresponds to the early gastrula in which the mesendoderm starts to get flat before its invagination into the blastocoel (Fig. 5).

Explantation of Ectodermal Tissue (Corresponding to the Most Anterior Part of the Animal Pole)

1. In an agarose Petri dish, dig a hole in the agarose (approximately the size of the embryo) with the extremity of the microscalpel (see Note 18). 2. Using the morphology of the embryo, define the orientation of the gastrula (see Note 19) (Fig. 5) and position the embryo the dorsoventral axis perpendicular to the bottom of the plate. 3. Cut the embryo with the microscalpel, applying a movement from the top to the bottom (see Note 20). 4. Transfer the obtained ectodermal explant as described in the Subheading 3.5, steps 5 and 6.

Explantation of Blastoporal Lips, Ectodermal or Mesendodermal Components

The gastrula stage used in this experiment corresponds to the gastrula in which the invaginated mesendoderm is about to contact the ectoderm through the blastocoel (7 h post-fertilization at 19 °C in B. lanceolatum) (Fig. 5).

Fig. 5 Morphology of the embryo at the early-mid gastrula stage and examples of tissue graft. (a) Blastoporal view of an early-mid gastrula embryo (7 hpf). The embryo presents a left/right symmetry and is larger in the dorsal than in the ventral region. (b) Blastoporal view of an early-mid gastrula embryo (7 hpf). During the invagination of the mesendoderm, a depression of the mesendoderm shape in the dorsal region (arrows) can be observed, helping to distinguish the dorsal from the ventral part of the gastrulae. (c) Scheme representing graft experiments of the dorsal blastoporal lip (in green) on ectodermal explant or on early-mid gastrula embryo (6 hpf and 7 hpf respectively) represented in lateral view. Ectoderm is in orange, mesendoderm is in blue. D dorsal, L left, R right, V ventral, hpf hours post-fertilization. Scale bars: 50 μm

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1. In an agarose Petri dish, position the blastopore in contact with the agarose (the ectoderm to the top). 2. Using the morphology of the embryo, define the orientation of the embryo to delimitate the explantation region (see Note 19) (Fig. 5). 3. Apply a weak pressure with the eyelash to cut the two layers (ectoderm plus mesendoderm). 4. To separate the two layers of the blastoporal lip (if necessary) (see Note 20):

(a) Slip the eyelash between the two layers to separate them.



(b) Cut the expected zone of tissue and separate it from any unwanted additional tissue. Or



(c) Cut the region linking the ectoderm and the mesendoderm (in the case of the blastoporal lip explantation).



(d) Push the ectodermal tissue sliding it on the mesendoderm applying a weak and oblique pressure on it with the eyelash (see Notes 21 and 22).

5. Transfer the obtained explant as described in the Subheading 3.5, steps 5 and 6. 3.7  Graft Experiments 3.7.1  Grafting of a Blastoporal Lip into a Host Gastrula

1. From the step 4 in Subheading “Explantation of Blastoporal Lips, Ectodermal or Mesendodermal Components”: transfer the blastoporal lip explant close to a dechorionated host gastrula embryo (7–8 h post-fertilization stage at 19 °C in B. lanceolatum). 2. Position the host gastrula with the blastopore to the top. 3. With the extremity of the eyelash, press slightly the mesendodermal part of the host gastrula to create a tiny hole (see Note 23). 4. With the eyelash, create a flow around the blastoporal lip explant with delicate movements, suspend it in the FSW and move it above the gastrula. 5. Let the blastoporal lip explant drop by gravity into the archenteron, still making tiny movements with the eyelash in the FSW to control the exact position of the explant. 6. Once the blastoporal lip explant is in the archenteron, position it with the eyelash into the hole previously created in the mesendoderm (Subheading 3.7.1, step 3). 7. The grafted embryo is ready to develop (see Note 24).

3.7.2  Grafting of a Blastoporal Lip Over a Host Ectodermal Explant (See Note 25)

1. Use the ectodermal explants from Subheading “Explantation of Ectodermal Tissue (Corresponding to the Most Anterior Part of the Animal Pole)”. 2. Graft experiments can be performed in agarose Petri dish or in scratched Petri dish (see Note 26).

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3. Position the ectodermal explant with the convex part to the top. 4. Slightly push the ectodermal explant to create an incurved shape in order to receive the blastoporal lip (or any other tissue) (see Note 27). 5. With the eyelash, make a flow on the FSW around the blastoporal lip explant with delicate movements, suspend it in the FSW and move it above the ectodermal explant. 6. Let the blastoporal lip explant drop by gravity in the incurved part of the host ectodermal explant, and control the exact position of the explant with tiny movements of the eyelash in the FSW. 7. Slightly push on the blastoporal lip to ensure its adhesion to the ectodermal explant. 8. Keep the Petri dish immobile during at least 1 h to ensure that the blastoporal lip remains fixed to the ectodermal explant. 9. The grafted explant is ready to develop.

4  Notes 1. No difference is observed between embryonic development in scratched Petri dishes and in agarose Petri dishes covered with 0.8% or 1% agarose/FSW. 2. Agarose/FSW Petri dishes can be conserved 2–3 days at 4 °C without formation of a biofilm on the agarose. This time of conservation can be increased preparing the agarose/FSW Petri dishes in a laminar hood. It remains possible to avoid biofilm formation during embryonic development adding antibiotic in the FSW or in the agarose/FSW during its preparation [7, 8]. 3. If it is not possible to keep the animals in an open system, a close system with sea water filtration can be used [5], with a regular or continuous change of sea water in the tank. It is also recommended to wash the sand every 4–5 days, especially when an individual dies in the tank. 4. Ideally, using transparent plastic cups (diameter: 4–5 cm) placed over a black background helps to visualize which animals have already spawned. A low volume of FSW (20–30 ml) in the transparent plastic cups helps concentrating the gametes. 5. The animals are extremely sensitive to any source of light. As soon as the light is turned off be careful to avoid any light in the room where the amphioxus are kept. A weak red light can be used to check if the animals spawn, avoiding a direct and sustained illumination on the amphioxus.

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6. Amphioxus sperm in FSW can be kept for 2 days at 4 °C. It is recommended to keep an aliquot of sperm from one spawning day to another. 7. Do not put too many eggs in a single Petri dish. Use several Petri dishes to separate the eggs with approximately 300–500 eggs per dish. Indeed, a high concentration of eggs per Petri dish results in a high rate of malformed embryos. 8. Adding too much sperm/FSW results in abnormal development of some embryos due to a high amount of spermatozoids sticking to the fertilization membrane. 9. Just after fertilization, it is recommended to scatter the eggs in order to increase the space for the fertilization membrane formation. This will increase the tension of the fertilization membrane during its development and increase the efficiency of dechorionation. 10. Some embryos can explode or can stick to the tip. To minimize the number of eggs lost during the pipetting step, it is recommended to first pipette FSW without embryos to humidify the interior of the tip before the manipulation. In addition, the 20–30 μl of FSW pipetted before pipetting the embryos help to remove almost all of the embryos from the tip and to minimize the amount of exploding embryos in the step 3 in Subheading 3.4. Moreover, the additional 20–30 μl of FSW pipetted helps to avoid surface tension at the extremity of the tip, again minimizing the amount of exploding embryos in the step 3, Subheading 3.4. 11. Part of the non-dechorionated embryos float at the surface of the FSW and are easy to remove. Among the embryos that drop down, non-dechorionated embryos are easily distinguishable from the dechorionated embryos by weakly moving the Petri dish. Indeed, the nondechorionated embryos follow the movement of the surrounding FSW. In addition, playing with light orientation of the binocular microscope also allows to identify embryos with the fertilization membrane. 12. Just after every cell division during early development, cells are joined together by a part of the membrane they still share. At this cell division timing, it is possible to dissociate the cells, with a higher risk to break them. Nevertheless, as the embryo development progresses to the next cell division, each cell becomes individualized and is only slightly connected to the adjacent cells. At this moment, cells can be easily separated. 13. To separate cells or a group of cells within a dechorionated embryo, a thin needle of glass can also be used instead of the eyelash [9].

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14. Dechorionated embryos/cells tend to stick to the tips. In order to avoid this phenomenon, the tips used to transfer the cells are kept in FSW during at least 2 days before being used. 15. To avoid explosion or separation of the cells or groups of cells, it is necessary to be careful not to expose them to the water surface tension. To do so, it is recommended to use thin tips of 0.2–20 μl with a thin extremity. This will allow to reduce the movement of the cells in the tip. In addition, pipetting FSW before and after the volume of FSW containing the cells limits the exposition to surface tension. The use of a total of 15 μl volume of pipetted FSW allows controlling the position of the cells in the tip as well as limiting the pressure of the FSW flow around the cells when pipetting. 16. Depending on the experimental need, the dissociated cells can be dropped in an agarose or in a scratched Petri dish. Note that in the scratched Petri dish, the cells in contact with the plastic will stick to the plastic very tightly and movements of the Petri dish in these conditions can induce the separation of the dividing cells during cleavage stages. 17. The movement of the microscalpel should be weakly oblique in order to maintain the embryo in the fertilization membrane correctly oriented during the microdissection. 18. Presence of biofilm on the agarose will increase the difficulty to recover clean explants. It is recommended to use agarose Petri dishes of one-day old maximum. 19. It is possible to distinctly observe that the shape of the invaginating mesendoderm is larger in the dorsal than in the ventral part, thus helping to identify the orientation of the embryo (Fig. 5). 20. Apply a movement slightly oblique in order to maintain the dechorionated embryo correctly oriented during the microdissection. 21. Ectodermal and mesendodermal regions can be easily distinguished during the gastrulation process since the ectodermal tissue is constituted by cells of smaller size. Moreover, these cells reflect the light more than mesendodermal cells. 22. We recommend to push and slip the ectodermal part rather than the mesendodermal part since the ectoderm is more robust and resist micromanipulation better than the mesendoderm. 23. If the aim of the experiment is to put in contact the blastoporal lip with the ectoderm, create a hole in the mesendoderm. Thus the contact between the tissues will take place in the disappearing blastocoel.

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24. Avoiding any movement of the Petri dish for at least half an hour after the graft manipulation increases the chance to keep the blastoporal lip explant stuck to the mesendoderm of the host gastrula. 25. The ectodermal explants tend to close during their development generating ball shaped explants. We describe in the Subheading 3.7.2 the most difficult graft manipulation which is the external graft (graft over the external part of the host ectodermal explant) but it is possible to simply position the tissue to be grafted (blastoporal lip or any other embryonic tissue) into the closing host ectodermal explant before it acquires the ball shape (internal graft). 26. Ectodermal tissue dissected from dechorionated gastrula will stick to the scratched plate. The cells from this ectodermal explant will develop cilia 3–4 h at 19 °C after the dissection, and as soon as these cilia develop, the explant will detach from the dish. 27. It is possible to create a tiny hole in the ectodermal explants with the extremity of the eyelash in order to facilitate the blastoporal lip attachment on the ectoderm. Nevertheless, the ­cellular content liberated in the FSW by the few cells that will explode during this manipulation increases the difficulty to position the blastoporal lip correctly. References 1. Bourlat SJ, Juliusdottir T, Lowe CJ, Freeman R, Aronowicz J, Kirschner M et al (2006) Deuterostome phylogeny reveals monophyletic chordates and the new phylum Xenoturbellida. Nature 444(7115):85–88 2. Delsuc F, Brinkmann H, Chourrout D, Philippe H (2006) Tunicates and not cephalochordates are the closest living relatives of vertebrates. Nature 439(7079):965–968 3. Bertrand S, Escriva H (2011) Evolutionary crossroads in developmental biology: amphioxus. Development 138(22):4819–4830 4. Poss SG, Boschung HT (1996) Lancelets (Cephalochordata: Branchiostomatidae): how many species are valid? Isr J Zool 42(sup1):S13–S66 5. Fuentes M, Benito E, Bertrand S, Paris M, Mignardot A, Godoy L et al (2007) Insights into spawning behavior and development of the

European amphioxus (Branchiostoma lanceolatum). J Exp Zool B Mol Dev Evol 308B(4):484–493 6. Le Petillon Y, Luxardi G, Scerbo P, Cibois M, Leon A, Subirana L et al (2017) Nodal– activin pathway is a conserved neural induction signal in chordates. Nat Ecol Evol 1(8):1192–1200 7. Holland LZ, Yu J-K (2004) Cephalochordate (amphioxus) embryos: procurement, culture, and basic methods. Methods Cell Biol 74:195–215 8. Tung TC, Wu SC, Tung YY (1960) The developmental potencies of the blastomere layers in Amphioxus egg at the 32-cell stage. Sci Sin 9:119–141 9. Tung TC, Wu SC, Tung YF (1958) The development of isolated blastomeres of Amphioxus. Sci Sin 7(12):1280–1320

Part II Vertebrate Models

Chapter 20 In Situ Hybridization and Immunostaining of Xenopus Brain Kai-li Liu, Xiu-mei Wang, Zi-long Li, Ying Liu, and Rong-qiao He Abstract The dynamic expression pattern analysis provides the primary information of gene function. Differences of the RNA and/or protein location will provide valuable information for gene expression regulation. Generally, in situ hybridization (ISH) and immunohistochemistry (IHC) are two main techniques to visualize the locations of gene transcripts and protein products in situ, respectively. Here we describe the protocol for the whole brain dissection, the in situ hybridization, and the immunostaining of the developing Xenopus brain sections. Additionally, we point out the modification of in situ hybridization for microRNA expression detection. Key words Expression pattern, Brain dissection, Section, Immunostaining, In situ hybridization, Xenopus, MicroRNA

1  Introduction In situ hybridization and immunostaining are widely used practical techniques to detect the locations of gene transcript and protein product in situ, respectively. Moreover, the co-application of in situ hybridization and immunostaining in one section is increasingly employed for simultaneously observing location of the gene transcripts of interest as well as the spatial distribution of another gene product at the biochemical level. For the gene expression analysis in early developing Xenopus brain, usually earlier than stage 35/36 (st.35/36), in situ hybridization and immunostaining are often conducted with fixed whole mount embryo followed by sectioning for following analysis. Whole-mount in situ hybridization (WISH) was adapted according to Harland [1], with modifications [2–6]. Whole-mount immunohistochemistry (WIHC) was carried out as described in [3, 4]. WISH for detecting microRNA (miRNA) expression was performed according to our previous work [7–10]. However, as embryonic development proceeds, it gradually encounters difficulty for probe and antibody to penetrate into the Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_20, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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brain and other tissues completely if the embryos or tissues are too large. This low permeability can cause poor signal-to-background ratio in ISH and IHC, although whole-brain clearing methods have been reported compatible with whole-mount immunohistochemical analysis [11]. Therefore, it is necessary to dissect brain and other tissues at later embryonic (st.37/38 and later) and adult stages. Here, we describe the method for dissecting and preparing the brain for in situ hybridization and immunostaining. Particularly, we will present the updated approach for examining miRNA expression.

2  Materials 2.1  Tools for Xenopus Embryo Manipulation, Brain Dissection

1. A stereotype microscope is required for observation during embryo or tissue dissection. If the embryos were injected with gfp mRNA, a stereotype fluorescent microscope is needed. 2. Forceps with blunt or sharp tips (e.g., Sigma Tweezers style #5) are used to hold and dissect embryos or tissues, which will be placed on agarose plate for operation. 3. Small sharp surgical scissors that are for eye operation are used to cut skin and nerve. 4. Stainless steel bone clamp is adequate for opening adult head skull to obtain the intact brain and separate nerve fiber. 5. Superfrost® Plus slides or other Poly-l-Lysine-coated slides. 6. Hairloop is used for moving embryos gently. 7. Whatman No. 2 filter paper.

2.2  Chemical Reagents

Representative chemical reagents for ISH and IHC are listed as follows: 1. HEPES, tricaine methanesulfonate (Sigma). 2. Formaldehyde. 3. Paraformaldehyde (PFA). 4. Tween-20. 5. Methanol. 6. Ethanol. 7. Sucrose. 8. OCT. 9. Paraffin. 10. Xylene. 11. Proteinase K. 12. BSA (BSA V; Sigma).

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13. Polyvinylpyrrolidone (PVP-40). 14. Ficoll 400 (phamacia). 15. Formamide (redistilled). 16. Torula RNA (Type IX, Sigma). 17. Heparin. 18. CHAPS. 19. Blocking reagent (Roche). 20. Lamb serum. 21. Anti-digoxigenin–alkaline-phosphatase antibody (Sigma cat. No. 11093274910). 22. Tetramisole (Sigma cat. No. L9756-5G). 23. NBT/BCIP (Sigma cat. No. 11681451001). 24. BM purple (Sigma cat. No. 1144207400). 25. Fast Red Tablets (Sigma cat. No. F4648). 26. DAB substrate kit (zsbio). 27. AEC substrate kit (zsbio). 28. Hoechst 33258 (Sigma). 2.3  Solutions for In Situ Hybridization and Immunostaining

Solutions for in situ hybridization should be prepared using ultrapure water (18.2 MΩ at 25 °C). For IHC, double distilled water (ddH2O) is feasible. Chemical reagents are the analysis pure: 1. MMR: 0.1 M NaCl, 2.0 mM KCl, 1 mM MgSO4, 2 mM CaCl2, 5 mM HEPES (pH 7.8), 0.1 mM EDTA. For 1 L 10× MMR, add NaCl 58.44 g, KCl 1.49 g, MgCl2 ·6H2O 2.03 g, CaCl2 (anhyd) 2.22 g, and HEPES 11.92 g into 800 mL water, mix dissolve and adjust pH to 7.5 with NaOH, and bring to 1 L with water. Autoclave and store at room temperature (RT). 2. MS222 anesthetic solution: MS222 0.2 mg/mL in 0.1× MMR, pH 7.5–7.8. 3. 1× PBS: For 1 L 10× PBS, dissolve NaH2PO4·2H2O 2.89 g, Na2HPO4·12H2O 26.73 g, and NaCl 102.2 g in 800 mL ultrapure water, and then adjust pH to 7.4 with NaOH. Bring volume to 1 L and autoclave. Store this solution at RT. Dilute the stock solution with water before use and adjust pH if necessary. 4. MEMFA: 0.1 M MOPS (pH 7.4), 2 mM EGTA, 1 mM MgSO4, 3.7% formaldehyde (see Note 1). 5. 4% PFA/1× PBS: For 50 mL, dilute 10 mL 20% PFA with 5 mL 10× PBS with water prior to use (see Note 2). 6. 20% Sucrose/1× PBS: For 50 mL, dissolve 10 g RNase-free sucrose with 1× PBS, sterilize the solution through 0.45 μM filter, and store at 4 °C.

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7. 50× Denhardt’s solution: 1% BSA, 1% PVP-40, 1% Ficoll 400. For 50 mL, dissolve 0.5 g BSA, 0.5 g PVP-40, and 0.5 g Ficoll 400 in water, sterilize the solution through 0.22 μM filter, and store at −20 °C. 8. 20× SSC: Dissolve 175.3 g NaCl and 88.2 g sodium citrate in 800 mL of water, adjust to pH 7.0 with 1 N HCl. Make up to 1000 mL of water. Autoclave and store the solution at RT. 9. Hybridization buffer (for ordinary RNA probe): 50% formamide, 5× SSC, 0.1% Tween-20, 100 μg/mL Heparin, 1 mg/mL torula RNA, 1× Denhardt’s solution, 0.1% Tween-­20, 10 mM EDTA, 0.1% CHAPS. Sterilize the solution with 0.22 μM filter and store it at −20 °C. 10. Hybridization buffer (for LNA probe for miRNA): 50% Formamide, 5× SSC, 0.1% Tween-20, 50 μg/mL Heparin, 500 μg/mL yeast tRNA or torula RNA, 10 mM Citric acid (pH 6.0). Filter the solution with 0.22 μM filter and store it at −20 °C. 11. Washing solution (post hybridization): 50% formamide, 2× SSC, 0.1% Tween-20 for ordinary probe; 2× SSC for LNA probe. 12. PBT: 1× PBS, 0.1% Tween-20. 13. MABT: 100 mM maleic acid, 150 mM NaCl, 0.1% Tween-20, pH 7.5 (see Note 3). 14. Blocking buffer: 2% blocking reagent, 20% heat inactivated sheep serum in MABT for ordinary probe; 1% blocking reagent, 1% heat inactivated sheep serum in MABT for LNA probe (see Note 4). 15. Alkaline-phosphatase buffer: 100 mM NaCl, 100 mM Tris– HCl pH 9.5, 50 mM MgCl2, 0.1% Tween-20, 2 mM tetramisole (see Note 5). 16. NBT/BCIP staining solution: For every mL of alkaline phosphatase buffer, add 1 μL of NBT and 3.5 μL of BCIP (see Note 6). 17. Fast Red staining solution: Add one Roche tablet in 2 mL 0.1 M Tris–HCl, pH 8.2, vortex 2 min and centrifuge 30 s to discard the red precipitates.

3  Methods 3.1  Brain Dissection and Fixation

Induction of ovulation in females, in vitro fertilization, embryo culture, and staging are carried out as described [12, 13]: 1. For tadpoles (st.37/38 and later), transfer the embryos in a small dish (diameter 35 mm) and make them anesthetized by replacing the culture solution with MS222 solution (see Note 7).

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2. When the embryos stop moving, place them into agarose plate with precooled 1× PBS. Use the forceps with blunt tips to hold the embryo at the trunk; use another forceps with sharp tips to make a cut through the spinal cord at the dorsal truck just behind the hindbrain. Tip up the skin of the front cut with forceps and carefully peel off the skin of the dorsal head to expose the brain. Then insert the tips of the forceps from the cut to the bottom of the brain, lift up the brain by a careful forward movement of the forceps, and shear the nerve bundles with forceps tips or scissors. Transfer the dissected brain immediately into fixation solution with a pipette or forceps. 3. For the brain dissection of developing frogs to adult, make the frog anesthetized in the MS222 for around 20 min and then place it on the ice. Cut off the spine and tear the skin from foramen magnum toward the head with a sharp scissors and then use bone clamp to open the skull and expose the brain (see Note 8). The procedure of dissecting brain follows as above (step 2). If this step takes a long time, to prevent the decay of brain tissue, add several drops of fixation solution onto the brain to fix it in situ for 1 h before taking it out (see Note 9). 4. Fix the dissected brain in MEMFA or 4% PFA at 4 °C overnight (see Note 10). 5. For WISH or paraffin section preparation, the brain could be dehydrated gradually by replacing the fixation solution with 25% ethanol/PBS, 50% ethanol/50% PBS, 75% ethanol, 100% ethanol sequentially, 5–10 min each, and stored in 100% ethanol at −20 °C (see Note 11). For WIHC, ethanol should be replaced with methanol. For cryosection section preparation, go on as per Subheading 3.2 directly without dehydration. 3.2  Cryosection Preparation and Pretreatment Prior to Hybridization

1. Pipette off the fixation solution and add precooled 1× PBS to wash for three times. 2. Pipette off the PBS and add 20% sucrose/1× PBS to cryoprotect the brains at 4 °C for 4 h to overnight until the brains sink to the bottom. 3. Label molds the name and the direction of specimens. Transfer the brain(s) into the mold and remove the sucrose solution as much as possible. Add appropriate volume (enough to immerse the whole specimen) of OCT in the molds. Orientate the brains in right directions with a long syringe needle, and then carefully move the mold on the iron plate upon dry ice or into the 70% ethanol precooling at −80 °C. After the OCT has been completely frozen, store the block at −80 °C before sectioning. 4. Slightly tickle or press the mold to take out the frozen block, mount the block on the specimen stand of cryostat at right direction, and cut the edges to modify it into right shape before

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sectioning. Cut 10–15-μm sections of the embedded brain at −24 °C. 5. Collect sections one by one on ready-to-use Superfrost® Plus slides or other Poly-l-Lysine-coated slides, air-dry the sections, and store the slides at −80 °C. 6. The stored sections should be defrosted at RT for at least 30 min prior to use for hybridization or immunostaining. 3.3  Paraffin Section Preparation and Pretreatment Prior to Hybridization

1. Dissolve paraffin at 60 °C before the day of embedding and keep it at 60 °C. 2. Wash the stored dehydrated brains with 100% ethanol, and then transfer specimens to xylene by gradually replacing the solution with 75% ethanol/25% xylene, 50% ethanol/50% xylene, 25% ethanol/75% xylene, and 100% xylene, and shake at RT for 5–10 min each time. 3. Wash the brains with new 100% xylene, pour the specimens with xylene into a small beaker (specimens should be immersed in xylene), and incubate at 60 °C for 20 min (see Note 12). 4. Embedding: Add the same volume of prewarmed paraffin, incubate at 60 °C for 45 min, and then replace with 100% paraffin, incubated at 60 °C for 20 min. Replace with 100% new paraffin, incubated at 60 °C for 3 h; wash specimens with 100% new paraffin, incubate at 60 °C for 20 min; replace with new paraffin, pour specimens with paraffin into prewarmed mold or plastic disc, and set the specimens at the right orientation and positions with prewarmed needle. Turn off the oven, and leave the paraffin block with specimens to be solidified slowly in the oven overnight (see Note 13). 5. Modify the embedded block and cut 5–10 μm sections in a microtome. Transfer sections on the surface of 0.2% ethanol on slides prewarmed at 37 °C (see Note 14). Discard the solution when sections have been completely extended, dry the sections at 37 °C overnight, and then store the slides at 4 °C. 6. For hybridization or immunostaining, dewax the section in 100% xylene by three washes, 10 min each time, 100% ethanol wash twice, and then rehydrate the section by washes with graded alcohol sequentially reverse to the dehydration, 2 min each wash.

3.4  In Situ Hybridization of Sections 3.4.1  Pretreatment of Sections and Hybridization

1. Digoxigenin (DIG)- or fluorescein-labeled antisense RNA probes should be generated prior to hybridization step. The labeled LNA probe for miRNA is available from EXIQON Company. 2. Wash the sections with PBT twice at RT, 2–5 min each wash. 3. For ordinary RNA probe, the sections could be applied for hybridization (step 4) directly. For the LNA probe for miRNA,

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rinse sections in PBT with 1 μg/mL proteinase K at RT for 10 min, followed with two washes in PBT with 2 mg/mL glycine, 5 min each wash, and three washes in PBT, 3 min each wash. Then refix the sections in the 4% PFA solution for 15 min, followed with three washes in PBT, 3 min each time (see Note 15). 4. Place slides horizontally in humid hybridization chamber bottomed with filter paper immersed in 1× SSC/50% formamide. Prewarm hybridization buffer with 1 μg/mL probe at 60–65 °C for ordinary probe or at the temperature suggested by the manufacturer for LNA probe for 5–10 min. Add 120– 150  μL probe mix per slide, and carefully cover slide with a coverslip. Seal the box with plastic film and transfer the box into oven prewarmed at the right temperature to hybridize overnight (see Note 16). 3.4.2  Washing Steps and Antibody Visualization

1. Set a glass trough with washing solution stand in water bath. Warm up the solution and the glass trough to the hybridization temperature. Add 100 mL washing solution to a glass trough for ten slides. 2. Transfer slides to the glass trough with prewarmed washing solution (see Note 17), and take away the coverslips after they detach off the slides (usually in 1–2 min). Then, for ordinary probe, make two washes at 65 °C with stirring or shaking, 30 min each; for LNA probe, make two washes in 2× SSC at RT, 10 min each, followed with another wash in 0.2× SSC at RT for 10 min. 3. Wash the slides two times in MABT at RT, 30 min each for ordinary probe, or five times in PBT, 3 min each, for LNA probe. 4. Take the slide out one by one with forceps, wipe away the excessive solution with filter paper, and then lay the slides on the holder of the humid chamber with filter paper soaked in 1× PBS or water. Add around 1 mL blocking buffer per slide to cover all the sections (without coverslip), and block for a half to 1 h at RT. 5. Discard the blocking buffer, add 150 μL antibody mix (antibody diluted 1:2000 to 1:8000 in blocking buffer) per slide, cover with a piece of parafilm cut as the same size of coverslip, and incubate the slide in a humid chamber at RT overnight. 6. Discard the antibody solution, transfer the slides in a glass trough with MABT, and make five washes on a shaker, 30 min each wash at RT (see Note 18), followed with another five washes in PBT, 5 min each at RT. 7. Staining: Wash the slides three times in alkaline-phosphatase buffer for 5 min each at RT. Take out the slides and set them in a humid chamber, add 150 μL NBT/BCIP or Fast Red

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staining buffer per slide, cover with parafilm, and then develop in the dark for usually 1 h to overnight with NBT/BCIP or 20 min to 6 h with Fast Red, depending on the abundance of the RNA. 8. Stop the staining reaction by two washes in PBT, 1× PBS sequentially. Then the sections could be mounted for observation (go step 13) and go on with subsequent immunostaining (see Subheading 3.5) or the following counterstaining (e.g., Hoechst nuclear staining) for histological examination. 9. Hoechst staining for counterstaining of cell nuclei: Add several drops of freshly prepared 1 μg/mL Hoechst 33258/1× PBS to the slides, cover with a piece of parafilm to spread the staining solution to all section, incubate for around 10 min at RT to stop the staining reaction, and wash sections with 1× PBS twice, 3 min each. 10. Mounting: For fluorescent staining, mount in a water-soluble, nonfluorescing mounting medium, e.g., Aqua-Poly/Mount (Polysciences) for observation. For chemical staining, dehydrate and clear sections by graded alcohols two washes in 100% ethanol and two washes in xylene, 2–3 min each, and then mount in Histomount or Canadian gel solution (see Fig. 1). 3.5  Immunohistochemistry of Sections

1. For paraffin sections, antigen should be retrieved in 0.01 M sodium citrate pH 6.0 at 80 °C for 20 min. For cryosections, this step can be omitted. When HRP-conjugated second antibody is applied, 0.3% H2O2 in 1× PBS treatment for 10–30 min would be needed to inactivate the endogenous peroxidase. 2. Rinse sections with PBS for three times, then PBT for three times, 5 min for each time. 3. Add around 1 mL 10% goat serum/2% BSA/PBT per slide to cover all the sections, and block for 30 min. 4. Discard the blocking buffer, and overlay with 150 μL primary antibody solution (antibody diluted in blocking solution as recommended by the manufacturer), covered with coverslips and incubated for 6 h at RT or overnight at 4 °C in a humidified chamber. 5. Rinse the sections with PBT for three times, 5 min each. 6. Re-block sections with 10% goat serum/2% BSA/PBT for 30 min. Then incubate sections with second antibody (dilute antibody in the blocking solution as recommended by the ­manufacturer) in the wet chamber for 2 h at RT or overnight at 4 °C. 7. Wash sections with PBT for three times, with PBS for three times, 5 min.

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Fig. 1 In situ hybridization (ISH) of st.46 Xenopus brain sections for detecting transcripts of zic2 (a, b), ath3/neurod4 (c), microRNA-181 (d). ISH for Zic2 was carried out in cryosection, stained by using AP substrate Fast Red (red), which shows expressed in the dorsal region of the developing brain (b), the nuclei of which were counterstained with Hoechst (a). ISH for ath3/neurod4 and microRNA-181 were conducted in paraffin sections and stained by applying AP substrate BM purple (c, d). Ath3/neurod4 was expressed higher in ventricular zone (proximal) and lower in marginal zone (distal) of the developing brain/neural tube (c). microRNA-181 is rarely detectable in the developing brain and relatively highly expressed in the retina inner nuclear layer (d). Br brain, Re retina, Vz ventricular zone, Mz marginal zone

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Fig. 2 Immunostaining of Xenopus brain sections for detecting protein locations of N-tubulin (a, b) and phosphohistone-­H3 (pH 3) (c–e). IHC for n-tubulin was carried out in cryosection in (a, b), stained by TRITC-­ conjugated secondary antibody showing red fluorescence and expressed in the whole developing brain/neural tube and otic vesicle (b), the nuclei of which were counterstained with Hoechst (a). IHC for pH 3 was conducted in paraffin sections and stained with TRITC-conjugated secondary antibody (c, d) and HRP substrate AEC (e). pH 3 is specifically expressed in the proliferating cells in the midline of the developing brain as shown by the arrows. Br brain, Ov otic vesicle

8. If the second antibody is conjugated with fluorescent dye like FITC and TRITC, sections can be directly mounted and observed under fluorescent microscope. If the second antibody is AP or HRP conjugated, the substrate such as BM purple/FastRed for AP and DAB or AEC for HRP should be added on the slides for staining as described in Subheading 3.4 or the kit instruction (see Fig. 2). The post staining washes and mount are as described above.

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4  Notes Generally, the volume of solution for fixation and washing should be no less than 10 volumes of the specimens unless indicated. During the process of ISH or IHC, tissue sections should not be dry after initiating PBS washing, especially when the antibody is incubated. Otherwise, the background will appear to be too high. Sense probe control and blank control should be set in in situ hybridization. For immunohistochemistry, the blank control without adding the primary antibody is also required. The signal-to-­ background ratio of in situ hybridization or immunohistochemistry depends on the abundance of transcript/antigen of interest and also the quality of the antibody. Therefore, the final quality of the transcript/protein detection may be optimized in modulating the corresponding step when necessary: 1. The solution with formaldehyde is all right when stored at 4 °C and used within a week at most of the cases. But it is better to make the ten-times solution without formaldehyde (10× MEM) as stock solution and add fresh formaldehyde prior to use. For 100 mL 10× MEM, add MOPS 20.93 g, 0.2 M EGTA 10 mL, and 1 M MgSO4 1 mL, adjust pH to 7.4 with NaOH, and filter sterilized and aliquoted with 50 mL plastic tube, stored at −20 °C. 2. 20% PFA is commercially available. For self-preparation of 100 mL solution, heat 80 mL distilled water at 60–65 °C. Add 20 g paraformaldehyde and slowly add drops (1 mm in diameter. Cut the tip of the pulled syringe until you find it is not sealed and is sufficiently small to fit inside the micropipettes (see Fig. 3 in ref. 22). This syringe can be kept and reused to load micropipettes with plasmid and MO reagents. 19. Xenopus laevis tadpoles. Fertilized eggs are acquired from hormone-­induced matings of albino Xenopus laevis frogs in our colony. Tadpoles raised at 23 °C with a 12 h light: 12 h dark diurnal cycle until used for experiments. Alternatively, tadpoles can also be purchased as fertilized eggs/young embryos from commercial sources (Nasco, Fort Atkinson, WI), Xenopus Express (Brooksville, FL), or Xenopus One (Dexter, MI). 20. Aquarium air pump. Not necessary, but recommended to add to the bowl to speed recovery from anesthesia. E.g., Tetra 77851 “Whisper” Air Pumps for 10 gallon tank. 21. Compound microscope equipped with epifluorescence and appropriate filter cubes and a low power (20×) air objective, optional. E.g., Nikon Optiphot-2. This microscope is used to screen quickly tadpoles in order to determine whether sufficient cells are transfected and MOs are taken up into cells. 22. A multiphoton or confocal microscope system equipped with filters and/or lasers appropriate for the fluorescent molecules in the experiment. E.g., Perkin-Elmer Ultraview spinning disc confocal system using a Nikon Eclipse FN1 with a 25× 1.1 NA water immersion objective. 23. Incubator for tadpole housing. Set to 23 °C with a 12 h light: 12 h dark diurnal cycle.

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3  Methods 3.1  Animal Husbandry

3.2  Preparing the Electroporation Equipment

Tadpole armies are reared as groups of ~100 in large bowls at 23 °C with a 12 h light: 12 h dark diurnal cycle. For our experiments we typically use animals that are between Nieuwkoop and Faber stage 46 (5 days post fertilization, dpf) and stage 49 (~12 dpf) [32], though electroporation is effective on younger and old animals. Once experiments begin, single tadpoles are kept in 6-well tissue culture plates so that time-lapse images from identified individual tadpoles can be acquired. All procedures are conducted with Xenopus tadpoles anesthetized with 0.02% MS-222 solution. Before electroporation or imaging protocols, the tadpoles are transferred into the MS-222 solution and within minutes, they are immobile, unresponsive to touch and ready for the procedure. Afterward, they are revived within minutes of placing them back to their rearing containers containing Steinberg’s solution. 1. Configuring the electroporation equipment. We use an equipment configuration shown in Fig. 1a, so that the Grass Stimulator and Picospritzer are within arm’s reach while looking through the microscope. The wiring diagram is shown in Fig. 1b. What is not visible in the photo in Fig. 1a, is the 3 μF capacitor shown in the wiring diagram in Fig. 1b. Before you begin the electroporation procedure, practice adjusting the micropipette and electrode so that they both can reach where the stage/tadpole in the field of view. Minimizing the time that the tadpoles are anesthetized and on the stage of the microscope by preparing the equipment will help their recovery from the procedure. 2. Fabricating the electrode. Two platinum foil plates are fabricated from new or discarded filaments from Sutter puller that are folded and cut to a shape that fits the contour of the tadpole head/brain (Fig. 2). Typical platinum electrodes are approximately 0.5–1 mm wide and 5–10 mm long, and are soldered to wire leads. These plates must remain electrically isolated but secured so that the tips of the plates are ~0.5 mm apart and can straddle the area of the brain to be electroporated. We use put heat shrink tubing around the soldered connection of one electrode (bracket, Fig. 2b) and then shave it down to decrease its diameter so that the second plate can be positioned very close to it. Next, position the plates so they are parallel and use heat shrink tubing to secure them together. Lastly, secure the plates with shrink-wrap along a rod that fits in the micromanipulator. Connect the wire leads as shown in Fig. 1b.

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1. Preparation of transfer materials for electroporation. Using molecular biology-grade, endotoxin-free water prepare 10–20 μl working solutions of plasmid, MO and fast green in microcentrifuge tube. Recommended solution concentrations are: 1–5 μg/μl for CMV-promoter plasmids; 0.1–1 μg/μl for Gal4-UAS plasmids; 100–500 μM MO (see Note 2); 0.01– 0.1% fast green. Though there are reports that some MOs become unstable over time [33], we have had success keeping these working solutions at 4 °C for weeks and repeatedly drawn from them. 2. Loading micropipettes. Using the disposable 1 ml syringe that has been melted and pulled to a 100 μm into the distal processes of the cells. Arrows indicate the distal tips of tectal cells expressing turboGFP and show the lissamine MO signal. (i–vi) Panels are 50 μm square

electroporation, the MO concentration injected into the ventricle is known, but the diluted MO concentration within the ventricle is hard to estimate because it is not a closed system. More importantly, the efficiency of MO transfer from the ventricle into cells by electroporation is difficult to estimate and whether transfer is linearly related to ventricular MO concentration is unknown. Therefore, electroporation may effectively concentrate MOs in target cells, so injecting a range of MO concentrations into the ventricle may not result in a corresponding range of MO concentrations in target cells in a complex tissue like the brain. Figure 2 gives an indication of the

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heterogeneity of MO distribution in electroporated optic tecta and suggests that MO distribution is uneven across the tissue. The specificity of a MO on the knockdown of a target gene can be analyzed through a variety of experiments. First, the results of experimental MO electroporation should be compared the results of electroporation of control MOs. MO knockdown can also be compared to other the results of knockdown methods, such as the electroporation of dominant-negative plasmid DNA constructs or other interfering reagents [39–41]. Electroporation also works well to coadminister a “rescue” construct to replace the gene targeted by MO knockdown along with the MO [35, 42]. Several authorities recommend using multiple MOs against the same target transcript to validate specificity of knockdown. For translation blocking MOs, the MO is most effective when it overlaps with the start site, which limits the region against which MOs can be designed. This is a particular issue for neural genes, which have many different splice isoforms in the 5′UTR, which govern promoter specificity. Similarly, recommendations to validate knockdown by comparing effects of translation blocking MOs and MOs that interfere with splicing are valid in systems in which genomic information is complete, but cannot be applied where this is not the case. The central nervous system has the highest density of splice variants of an organ. Even as this feature makes studying the molecular genetic control of brain development particularly exciting, it also means that the challenges unique to studying nervous system development and plasticity must be recognized. When antibodies to the target protein are available, it is quite common to test whether MOs decrease levels of protein expression with immunocytochemistry or western blots. However, western blot analysis may not detect protein knockdown in the brain when MOs are administered with electroporation. Unlike MO injection into the fertilized egg or blastomere, where protein knockdown can be widespread, targeted electroporation of the tectum, retina or other areas of the CNS, may result in fewer cells that take up the MO. The sensitivity of the western blot to reveal knockdown will be hindered by the relatively few MO-containing cells in the tissue that will be homogenized with the surrounding cells that have not taken up the MO. Despite these caveats, this method has been shown to verify MO knockdown [39, 43]. Lastly, alternative methods to evaluate knockdown of genes of interest in the development of the CNS include functional analysis, such as electrophysiological recordings from neurons electroporated with MOs against neurotransmitter receptor subunits [36, 41].

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4  Notes 1. Design of experimental MOs. The MO vendor, Gene Tools, provides a free design service. Discussion of MO design can be found here [44]. 2. MO concentration: The MO vendor, GeneTools, provides detailed methods for storing and reconstituting the MO. Briefly, MOs are stable when reconstituted in pure water and stored at room temperature. Fluorescently tagged MOs should be stored in the dark. As stated on the GeneTools website, the lissamine tag decreases the solubility of the MO and lissaminetagged MOs in particular should be heated to 65 °C for 10 min and vortexed for 30 min until the MO solution is fully resuspended. MO concentration can be measured using the protocol on the GeneTools website. Considerations for long-term storage and dilution of MOs are also discussed in [27]. 3. Control MOs, choices and their design. Though the knockdown efficacy will be attenuated, MOs are capable of complementing a target sequence that shares 21/25 bases [16, 45]. “Specific” control MOs that are incapable of interacting with the target sequence consist of a 5/25 base mispair, and can be designed for each experimental MO. 4. There are two signs to look for as you electroporate as a way to verify that the settings are correct and the equipment is wired correctly. The first is the formation of small electrolysis induced bubbles (>10 bubbles). These will appear where the electrode makes contact with the moist skin. Large violently rolling bubbles are an indication of a problem with the settings/equipment. The second indicator is a slight twitch of the ocular muscles and movement of the eyes. 5. Positioning tadpoles in Sylgard chamber. For prescreening, this can be done quickly with little regard to the position of the coverslip and tadpole. When z-stacks are to be acquired, take care that there are no bubbles are transferred with the tadpoles inside of the Sylgard chamber because they might move and shift the tadpole during the image acquisition. Bubbles can usually be brushed out with the fine paintbrush. It is also important that the coverslip gently press on the head of the tadpole. The best image is acquired when there is no space between the tadpole and the coverslip. The coverslip must be secure, as it will have the water droplet for the immersion objective on it. If the coverslip becomes loose during the acquisition, it will shift the tadpole and disrupt the image mid-acquisition. 6. Inspecting tadpoles for damage as a result of the electroporation procedure. We occasionally find tadpoles that do not recover, or do not recover properly, from electroporation. This

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may result from the tadpole drying out during the procedure, and with experience, this can be minimized. Tadpoles that sustain damage can be spotted because they fail to swim properly, and inspection of the brain may show herniation of cells into the ventricle or signs of bleeding. The evidence of bleeding usually clears up within 48 h, but these are signs that indicate that the electrode pressed too hard on the tadpole, the volume injected into the ventricle was too great, and/or voltage level and number of electroporation pulses should be scaled back. References 1. Staton AA, Giraldez AJ (2011) Use of target protector morpholinos to analyze the physiological roles of specific miRNA-mRNA pairs in vivo. Nat Protoc 6(12):2035–2049. https:// doi.org/10.1038/nprot.2011.423 2. Draper BW, Morcos PA, Kimmel CB (2001) Inhibition of zebrafish fgf8 pre-mRNA splicing with morpholino oligos: a quantifiable method for gene knockdown. Genesis 30(3):154–156. Epub 2001/07/31 3. Morcos PA (2007) Achieving targeted and quantifiable alteration of mRNA splicing with Morpholino oligos. Biochem Biophys Res Commun 358(2):521–527. https://doi. org/10.1016/j.bbrc.2007.04.172. Epub 2007/05/12 4. Choi WY, Giraldez AJ, Schier AF (2007) Target protectors reveal dampening and balancing of Nodal agonist and antagonist by miR-430. Science 318(5848):271–274. https://doi.org/10.1126/science.1147535 5. Kloosterman WP, Lagendijk AK, Ketting RF, Moulton JD, Plasterk RH (2007) Targeted inhibition of miRNA maturation with morpholinos reveals a role for miR-375 in pancreatic islet development. PLoS Biol 5(8):e203. https://doi.org/10.1371/journal. pbio.0050203. Epub 2007/08/07 6. Bruno IG, Jin W, Cote GJ (2004) Correction of aberrant FGFR1 alternative RNA splicing through targeting of intronic regulatory elements. Hum Mol Genet 13(20):2409–2420. https://doi.org/10.1093/hmg/ddh272. Epub 2004/08/31 7. Kimmel CB, Law RD (1985) Cell lineage of zebrafish blastomeres. I. Cleavage pattern and cytoplasmic bridges between cells. Dev Biol 108(1):78–85 8. Heasman J, Kofron M, Wylie C (2000) Beta-­ catenin signaling activity dissected in the early Xenopus embryo: a novel antisense approach. Dev Biol 222(1):124–134. https://doi.

org/10.1006/dbio.2000.9720. [pii] S0012-­1606(00)99720-3. Epub 2000/07/08 9. Tandon P, Showell C, Christine K, Conlon FL (2012) Morpholino injection in Xenopus. Methods Mol Biol 843:29–46. https://doi. org/10.1007/978-1-61779-523-7_4. Epub 2012/01/10 10. Robu ME, Larson JD, Nasevicius A, Beiraghi S, Brenner C, Farber SA et al (2007) p53 activation by knockdown technologies. PLoS Genet 3(5):e78. https://doi.org/10.1371/ journal.pgen.0030078. Epub 2007/05/29 11. Hardy S, Legagneux V, Audic Y, Paillard L (2010) Reverse genetics in eukaryotes. Biol Cell 102(10):561–580. https://doi. org/10.1042/BC20100038. Epub 2010/09/04 12. Akcakaya P, Bobbin ML, Guo JA, Malagon-­ Lopez J, Clement K, Garcia SP et al (2018) In vivo CRISPR editing with no detectable genome-wide off-target mutations. Nature 561(7723):416–419. https://doi. org/10.1038/s41586-018-0500-9 13. Summerton JE (2007) Morpholino, siRNA, and S-DNA compared: impact of structure and mechanism of action on off-target effects adn sequence specificity. Curr Top Med Chem 7:651–660 14. Chen CM, Chiu SL, Shen W, Cline HT (2009) Co-expression of Argonaute2 enhances short hairpin RNA-induced RNA interference in Xenopus CNS neurons in vivo. Front Neurosci 3:63. https://doi.org/10.3389/ neuro.17.001.2009. Epub 2009/01/01 15. Lund E, Sheets MD, Imboden SB, Dahlberg JE (2011) Limiting ago protein restricts RNAi and microRNA biogenesis during early development in Xenopus laevis. Genes Dev 25(11):1121– 1131. https://doi.org/10.1101/gad.2038811. Epub 2011/05/18 16. Eisen JS, Smith JC (2008) Controlling morpholino experiments: don’t stop making antisense. Development 135(10):1735–1743.

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28. Mende M, Christophorou NA, Streit A (2008) Specific and effective gene knock-down in early chick embryos using morpholinos but not pRFPRNAi vectors. Mech Dev 125(11– 12):947–962. https://doi.org/10.1016/j. mod.2008.08.005 29. Bestman JE, Huang LC, Lee-Osbourne J, Cheung P, Cline HT (2015) An in vivo screen to identify candidate neurogenic genes in the developing Xenopus visual system. Dev Biol 408(2):269–291. https://doi.org/10.1016/j. ydbio.2015.03.010 30. Osterele A. Pipette cookbook 2018 P-97 & P-1000 micropipette pullers [pdf]. Sutter instruments; 2018 [cited 2018 14 October]. Rev. F. https://www.sutter.com/PDFs/ pipette_cookbook.pdf 31. Koster RW, Fraser SE (2001) Tracing transgene expression in living zebrafish embryos. Dev Biol 233(2):329–346. https://doi. org/10.1006/dbio.2001.0242. [pii] S0012-­1606(01)90242-8. Epub 2001/05/05 32. Nieuwkoop PD, Faber J (1994) Normal table of Xenopus Laevis (Daudin): a systematical & chronological survey of the development from the fertilized egg till the end of metamorphosis, 1st edn. Garland Science, New York 33. Bedell VM, Westcot SE, Ekker SC (2011) Lessons from morpholino-based screening in zebrafish. Brief Funct Genomics 10(4):181– 188. https://doi.org/10.1093/bfgp/elr021. Epub 2011/07/13 34. Kos R, Tucker RP, Hall R, Duong TD, Erickson CA (2003) Methods for introducing morpholinos into the chicken embryo. Dev Dyn 226(3):470–477. https://doi.org/10.1002/ dvdy.10254. Epub 2003/03/06 35. Faulkner RL, Wishard TJ, Thompson CK, Liu HH, Cline HT (2015) FMRP regulates neurogenesis in vivo in Xenopus laevis tadpoles. eNeuro 2(1):e0055. https://doi. org/10.1523/ENEURO.0055-14.2014 36. Ewald RC, Van Keuren-Jensen KR, Aizenman CD, Cline HT (2008) Roles of NR2A and NR2B in the development of dendritic arbor morphology in vivo. J Neurosci 28(4):850– 861.https://doi.org/10.1523/JNEUROSCI.507807.2008. Epub 2008/01/25 37. Bestman JE, Lee-Osbourne J, Cline HT (2012) In vivo time-lapse imaging of cell proliferation and differentiation in the optic tectum of Xenopus laevis tadpoles. J Comp Neurol 520(2):401–433. https://doi.org/10.1002/ cne.22795. Epub 2011/11/25 38. Sauka-Spengler T, Barembaum M (2008) Gain- and loss-of-function approaches in the chick embryo. Methods Cell Biol 87:237–256.

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Chapter 22 Sensitive Multiplexed Fluorescent In Situ Hybridization Using Enhanced Tyramide Signal Amplification and Its Combination with Immunofluorescent Protein Visualization in Zebrafish Gilbert Lauter, Iris Söll, and Giselbert Hauptmann Abstract Fluorescent in situ hybridization (FISH) provides sensitive detection and visualization of RNA transcripts in tissues and cells with high resolution. We present here a multiplex RNA FISH method using enhanced tyramide signal amplification (TSA) for colocalization analysis of three different transcripts in intact zebrafish brains. To achieve enhancement of fluorescent signals, essential steps of the FISH procedure are optimized including embryo permeability, hybridization efficacy, and fluorogenic TSA-reaction conditions. Critical to this protocol, the enzymatic peroxidase (PO) reactivity is significantly improved by the application of viscosity-increasing polymers, PO accelerators, and highly effective bench-made tyramide substrates. These advancements lead to an optimized TSA–FISH protocol with dramatically increased signal intensity and signal-to-background ratio allowing for visualization of three mRNA transcript patterns simultaneously. The TSA–FISH procedure can be combined with immunofluorescence (IF) to compare mRNA transcript and protein expression patterns. Key words Fluorescent in situ hybridization, FISH, Tyramide signal amplification, TSA, Peroxidase, Zebrafish, Transcript, RNA, Immunofluorescence

1  Introduction Fluorescent in situ hybridization (FISH) provides extremely sensitive detection and visualization of RNA transcripts in embryos, organs, tissues, and cells up to subcellular and even single molecule resolution [1, 2]. To achieve such incredible sensitivity nonenzymatic and enzymatic amplification systems have been developed yielding dramatically enhanced signal intensity and signal-to-noise ratio [3]. Nonenzymatic amplification strategies include branched DNA-based technology [4] such as in situ hybridization chain reaction (HCR) [5, 6], while enzymatic amplification makes use of alkaline phosphatase (AP)- and horseradish peroxidase (PO)-based

Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_22, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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immunoassays [7, 8]. Substrate turnover is continuously catalyzed by these enzymes leading to multiplied chromogenic or fluorescent signals depending on the specific substrate applied to the reaction. For the PO detection procedure, the catalyzed reporter deposition (CARD) technique [9] can be implemented to further enhance signal intensity. The PO activity results in the deposition of labeled tyramide molecules, which are bound to electron-rich moieties of nearby proteins, so that fluorochromes are multiplied at the target site. This results in tyramide signal amplification (TSA) by up to 100-fold as compared to nonamplified FISH methods [7, 10]. The enhanced TSA–FISH signals allow for multiplex mRNA detection in nonmodel and model species including Drosophila [11], Platynereis [12], zebrafish [13, 14], frog [15], and chicken [16]. Thus, TSA–FISH has been applied to interrogate the tissueand cell type-specific expression profiles of a large number of mRNAs in Drosophila [17] and used to generate molecular maps of the developing zebrafish brain [18, 19]. Extending earlier chromogenic expression data [20, 21], the use of multiplex TSA–FISH allowed a detailed and precise characterization of neuronal cell types and brain subdivisions to potentially reveal novel aspects of the underlying basic brain organization [18, 19, 22]. The zebrafish multiplex TSA–FISH protocol is based on the original whole-mount in situ hybridization (WISH) protocol developed first in Drosophila [23] and extended later for chromogenic detection of up to three transcripts in multiple colors [24– 30]. The chromogenic WISH procedures were based on simultaneous hybridization of differently hapten-labeled probes followed by AP-based immunohistochemical chromogenic detection of up to three different transcript patterns [31]. For fluorescent mRNA visualization, AP enzymatic reactions were exchanged by the PO detection system with application of fluorochrome-­ coupled TSA substrates. One problem with initial TSA protocols was increased background in parallel to specific signal enhancement. In addition, strong autofluorescence of zebrafish embryos [13] further lowered the signal-to-noise ratio making multiplexing a difficult task. Therefore optimization of embryo preparation, hybridization, and detection as described in this chapter was necessary to achieve optimal sensitivity and specificity of fluorescent signals in zebrafish embryos [14, 32]. For the second book edition we included in this follow-up chapter [33] the colocalization analysis of three different mRNA transcripts in whole zebrafish brains and added immunofluorescence (IF) protein detection to the TSA–FISH protocol [3]. For multiplex RNA FISH, dinitrophenol-, digoxigenin-, and fluorescein-­labeled antisense RNA probes are hybridized simultaneously and detected in three consecutive rounds of probe detection using the respective anti-hapten antibodies conjugated to horseradish peroxidase (PO) (Fig. 1). After each detection round

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Fig. 1 Examples of the methods: 28-hpf embryonic zebrafish brains with anterior to the left are shown. Scale bars = 50 μm. (a–c) Two-color RNA-FISH using enhanced TSA: Lateral views of dlx2a and lhx1a forebrain expression. Transcript signals are visualized in cyan and magenta as indicated in the panels. Single-channel detection and the overlay of both channels in the same confocal plane are shown. (a) Expression of dlx2a was visualized by using a DNP-labeled RNA probe together with the DyLight633-tyramide. The TSA reaction was allowed to proceed for 20 min in the presence of 0.15 mg/ml 4-iodophenol. (b) Expression of lhx1a was visualized by using a DIG-labeled RNA probe together with the FAM-tyramide. The TSA reaction was allowed to proceed for 30 min in the presence of 0.15 mg/ml 4-iodophenol. (c) Overlay of both single channels shows the expression of dlx2a and lhx1a in the developing forebrain with areas of overlap indicated in yellow. (d–f) Tri-­ color RNA-FISH: Lateral views of nkx2.1, pax6a, and eomes expression in the fore- and midbrain. Overlay of two (d, e) or all three (f) different channels of the same confocal plane. The distribution of axonal tracts (h) in relation to ephA4a rhombomere expression (g) was visualized by combining RNA-FISH with anti-acetylated α-tubulin IF (i)

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consisting of antibody-PO conjugate incubation and TSA reaction, the PO is inactivated by acidic pH treatment. Visualization of transcript distribution and colocalization is performed through ­confocal microscopy. Key steps of the procedure were optimized previously [14, 32, 34]: Tissue permeability is improved by hydrogen peroxide/methanol treatment in addition to proteinase K digestion. Hybridization efficacy is enhanced by adding the polymer dextran sulfate to the hybridization buffer to increase viscosity and local probe concentration. Inclusion of dextran sulfate in the TSA reaction and addition of substituted phenol compounds as PO accelerators strongly enhance fluorescent signal output. A short protocol for synthesis of tyramide substrates is provided, since bench-made substrates are highly efficient in producing high signal strength [14]. The RNA FISH protocol can be followed by immunofluorescence to visualize and directly compare the distributions of mRNAs and proteins.

2  Materials 2.1  Tyramide Synthesis

1. Succinimidyl esters: 5-(and-6)-carboxyfluorescein succinimidyl ester (FAM-SE; Molecular Probes C-1311), DyLight 633N-hydroxysuccinimide ester (DyLight633-SE; Pierce: Rockford, IL, USA 46414), 5-(and-6)-carboxytetramethylrhodamine succinimidyl ester (TAMRA-SE; Molecular Probes C-1171). Prepare a 10 mg/ml stock solution of each succinimidyl ester in dimethylformamide (Sigma D4551) just before use. 2. Tyramine hydrochloride: Prepare a 10× stock solution at a concentration of 100 mg/ml in dimethylformamide just before use. Dilute with dimethylformamide to yield the 1× working solution and add 10 μl triethylamine (Sigma T0886) per 1 ml solution. 3. Absolute ethanol (EtOH).

2.2  Embryo Preparation

1. Paraformaldehyde (PFA): 4% paraformaldehyde in 1× PBS, pH 7.3. Dissolve 4 g PFA in 100 ml PBS and stir in a fume hood at about 65 °C until everything has dissolved (1–2 h). Let solution cool to RT and adjust to pH 7.3 with NaOH. Store the fixative in 5–10 ml aliquots at −20 °C. 2. Phosphate buffered saline (1× PBS): 8% (w/v) NaCl, 0.2% (w/v) KCl, 16 mM Na2HPO4, 4 mM NaH2PO4, pH 7.3. 3. Phosphate buffered saline plus Tween (PBST): 1× PBS, 0.1% (v/v) Tween-20. 4. Methanol (MeOH). 5. 30% hydrogen peroxide (H2O2), stabilized (Sigma31642).

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6. Rehydration series: 75%, 50%, 25% (v/v) of methanol in PBST. 7. Proteinase K: 20 mg/ml stock in TE (10 mM Tris–HCl pH 8.0, 1 mM EDTA), store in aliquots at −20 °C. Prepare working solution of 50 μg/ml proteinase K in PBST just prior to use. 8. Glycine buffer: 2 mg/ml glycine (Sigma G7126) in 1× PBST. 2.3  Hybridization

1. Prehybridization buffer (HYB): 50% deionized formamide, 5× SSC, 5 mg/ml torula RNA (Sigma R6625), 50 μg/ml, heparin sodium salt, 0.1% Tween-20. Store in aliquots at −20 °C. Store heparin as a 50 mg/ml stock in ddH2O at −20 °C. 2. Dextran sulfate (Sigma D6001): Prepare a 50% (w/v) stock solution in water. When used for hybridization, autoclave stock solution for 30 min at 110 °C, otherwise store in aliquots at −20 °C. 3. Hybridization buffer (HYBD5): HYB including 5% (v/v) dextran sulfate. 4. Hybridization wash: 50% deionized formamide (AppliChem 2156), 2× SSC, 0.1% Tween-20. Store in aliquots at −20 °C. 5. 20× SSC: 3 M NaCl, 300 mM trisodium citrate, pH 7.0. 6. 2× SSCT: 2× SSC, 0.1% Tween-20. 7. 0.2× SSCT: 0.2× SSC, 0.1% Tween-20.

2.4  Immunohistochemical Detection

1. Blocking solution: 8% normal sheep serum in PBST. Heat inactivate sheep serum (Sigma S-2263) at 56 °C for 30 min and store in aliquots at −20 °C. 2. Rabbit-anti-fluorescein/Oregon Green 488-POD (Molecular Probes A21253) prepare 1:500 working solution in blocking solution prior to use. 3. Sheep-anti-digoxigenin-POD Fab fragments (Roche 11207733910) prepare 1:500 working dilution in blocking solution prior to use. 4. Anti-dinitrophenyl-POD (PerkinElmer TSA Plus DNP System NEL747A001KT) prepare 1:100 working dilution in blocking solution prior to use.

2.5  Fluorogenic Reaction

1. 4-Iodophenol (Fluka 58020): prepare a 150 mg/ml stock in DMSO. Store tightly sealed at 4 °C (see Note 1). 2. Vanillin (Sigma V110-4): prepare a 150 mg/ml stock in DMSO. Store tightly sealed at 4 °C. 3. TSA reaction buffer: 100 mM borate buffer pH 8.5, 2% dextran sulfate, 0.1% Tween-20, 0.003% H2O2. 4. Borate Buffer: 100 mM borate pH 8.5 plus 0.1% Tween-20.

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5. Bench-made FAM, DyLight633 and TAMRA tyramide substrates: 1 mg/ml stocks in ethanol. 6. POD inactivation: 100 mM glycine pH 2.0 plus 0.1% Tween-20. 2.6  Immunofluorescence (IF)

1. Phosphate buffered saline plus Triton (PBSX): 1× PBS, 0.3% (v/v) TritonX-100. 2. Western blocking reagent (Roche 11921673001) prepare 1% working solution in PBSX plus 1% DMSO, aliquot and store at −20 °C. 3. E.g. Monoclonal mouse anti-acetylated α-tubulin (Sigma T-6793) diluted 1:500 in Western blocking working solution. 4. Appropriate fluorochrome coupled anti-mouse secondary antibody.

2.7  Mounting

1. Glycerol series: 25%, 50%, 75% (v/v) of glycerol in PBST, 40 mM NaHCO3. 2. Mounting gel: 1% low melting agarose in 75% (v/v) glycerol in PBST, 40 mM NaHCO3. Keep the solution stirring at 50 °C, as it will turn yellow after repeated heating. 3. Mounting slides: Two stacks of coverslips are clued to a microscope slide with Eukitt (Fluka 03989) leaving a gap, which can be easily bridged by a 24 × 32 mm coverslip. To mount samples of varying thickness, prepare a series of mounting slides using one to several coverslips for each stack.

3  Methods 3.1  Tyramide Synthesis

Despite the advantage of being cost-efficient, bench-made tyramides are also highly concentrated and offer the possibility to specifically match the requirements of the microscope in use. 1. Mix the freshly prepared tyramine working solution and the respective succinimidyl esters at a 1:1.1 equimolar ratio. 2. Allow the reaction to proceed for 2 h in the dark without agitation. 3. The resulting tyramide product is diluted with absolute ethanol to a concentration of 1 mg/ml and stored protected from light at −20 °C (see Note 2).

3.2  Embryo Pretreatment

1. Transfer dechorionated embryos of the desired developmental stage to a 2 ml microcentrifuge tube. Aspirate the supernatant while taking care that the embryos remain immersed in liquid. Fixate embryos in 1 ml of 4% paraformaldehyde (PFA) for 24 h at 4 °C (see Note 3).

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2. Rinse embryos four times for 5 min with phosphate-buffered saline containing 0.1% Tween (PBST). Transfer embryos into 100% methanol (MeOH). Exchange the 100% MeOH after 5 min and incubate the embryos at −20 °C for at least 30 min. Alternatively embryos can be kept in 100% MeOH at −20 °C for long-term storage. 3. Incubate the embryos in a 2% hydrogen peroxide solution in 100%MeOH for 20 min. Gradually rehydrate by going through a series of 5 min washing steps of 75%, 50%, 25% MeOH in PBST followed by two PBST washes (see Note 4). 4. Embryos developed to tailbud stage or further have to be digested with proteinase K in order to increase permeability. The optimal digestion times for different stages have to be determined experimentally. After incubation in proteinase K solution (10 μg/ml) at RT and under gentle agitation, stop the reaction by rinsing twice with glycine buffer. Postfix the embryos for 20 min in 4% PFA. Afterwards wash four times for 5 min in PBST. Transfer Embryos into 0.5 ml prehybridization buffer (HYB) and exchange with 1 ml HYB after 5 min. Embryos are now ready to use or can be stored in HYB at −20 °C. 5. Transfer up to 25 embryos into 200 μl of HYB in a 2-ml tube. Incubate in a water-bath at 60 °C for 1 h for prehybridization. 3.3  Hybridization

1. Prepare the probe-mix by adding fluorescein (FAM)-, dinitrophenol (DNP)-, and digoxigenin (DIG)-labeled RNA probes in 150 μl HYBD5 (see Note 5). Denature the probe-mix for 5 min at 80 °C and equilibrate it afterwards to 60 °C. 2. Carefully aspirate the prehybridization solution from the embryos and add the prewarmed probe-mix instead. Incubate in a water-bath at 60 °C overnight (min. 15 h). 3. For all posthybridization washes the solutions have to be prewarmed to 60 °C. 4. Wash embryos twice with 1 ml of 50% formamide-2× SSCT for 30 min. 5. Wash once with 1.5 ml 2× SSCT for 15 min. 6. Wash twice with 1.5 ml 0.2× SSCT for 30 min. 7. Add fresh 0.2× SSC and let cool down to RT. 8. Rinse twice with PBST.

3.4  Immunohistochemical Detection

The differently hapten-labeled RNA probes are sequentially detected using peroxidase (POD)-conjugated antibodies directed against the respective hapten. Typically FAM-labeled probes are detected first using an anti-FAM-POD antibody followed by detec-

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tion of DNP- and DIG-labeled probes with anti-DNP-POD (see Note 6) and sheep-anti-DIG-POD, respectively (see Note 7). 1. Incubate embryos with 100 μl of 8% blocking solution for 30 min on a gently rocking table at RT. 2. Carefully remove the blocking solution and add the appropriate antibody solution depending on the intended order of probe detection. In each round only one antibody solution is used. For detection of FAM-labeled probes apply rabbit antifluorescein/Oregon Green 488-POD (Molecular Probes) diluted 1:500 in blocking solution. For detection of DNPlabeled probes apply anti-DNP-POD (Perkin Elmer) diluted 1:100 in blocking solution. For detection of DIG-labeled probes apply sheep anti-­ DIG-­ POD Fab fragments (Roche) diluted 1:500 in blocking solution. 3. Incubate overnight at 4 °C without agitation. 4. To remove unbound antibody wash six times with PBST for 20 min at RT with gentle agitation. 3.5  Fluorogenic Reaction

The TSA reaction can be significantly enhanced by the use of a POD accelerator. Use 4-iodophenol or vanillin at a concentration of 0.15 mg/ml and 0.45 mg/ml respectively (see Note 8). 1. For 1 ml reaction buffer combine 500 μl Borate-buffer pH 8.5 (200 mM), 40 μl 50% dextran, 10 μl of 10% Tween-20, 6 μl of 0.5% H2O2 and an appropriate accelerator, adjust the volume with water to 1 ml (see Note 9). 2. The addition of 4-iodophenol will result in the appearance of a cloudy smear. Mix the reaction buffer well by pipetting up and down until the dextran sulfate has dispersed and the solution appears opaque. 3. Dilute the desired tyramide with the reaction buffer and mix well by pipetting. Use either 6 μl of DyLight633-tyramide or 4 μl of FAM tyramide or 4 μl of TAMRA tyramide stock solution per 1 ml reaction buffer (see Note 10). 4. Rinse embryos twice with 100 mM borate buffer pH 8.5 containing 0.1% Tween-20. Remove supernatant borate buffer as closely as possible and add 90 μl of the tyramide solutions to the embryos. Gently mix by pipetting using a cut tip. Incubate for the desired length of time in the dark at RT without agitation (see Note 11). 5. To stop the TSA reaction, wash the tubes for four times with PBST. For each washing step fill the entire tube with PBST. After inverting the tubes for several times, wait until the embryos have sank down to the bottom and then remove the

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excess buffer carefully. From this step onward embryos should be protected from light. 6. To inactivate POD activity of the applied antibody, incubate samples in 100 mM glycine–HCl pH 2.0 for 10 min followed by four 5 min washes in PBST under agitation (see Note 12). 3.6  Second and Third Detection Rounds

1. For the second antibody detection round repeat the steps described under subheadings 3.4 and 3.5 using a different anti-­hapten-­POD antibody and fluorogenic substrate. 2. For the third antibody detection round repeat the steps described under subheadings 3.4 and 3.5 using the antihapten-­POD antibody and the fluorogenic substrate, that has not been used in the previous detection rounds (see Note 13).

3.7  Immunofluorescence (IF) (See Note 14)

1. Wash the embryos 2 × 5 min in PBSX. 2. Incubate embryos in 1% Western blocking reagent in PBSX/1% DMSO for 30 min at RT with gentle agitation (see Note 15). 3. Dilute the primary antibody to the appropriate concentration in Western blocking working solution and apply to the embryos overnight at 4 °C without agitation. 4. Wash embryos in PBSX for 6 × 15 min at RT with gentle agitation (see Note 16). 5. Incubate embryos in 1% Western blocking reagent in PBSX/1% DMSO for 30 min at RT with gentle agitation. 6. Dilute the secondary fluorochrome-labeled antibody to the appropriate working concentration in Western blocking working solution and apply to the embryos overnight at 4 °C. 7. Wash embryos in PBSX for 6 × 15 min at RT with gentle agitation.

3.8  Mounting

1. To avoid shrinkage gradually transfer the embryos through 5 min washing steps in 25%, 50%, and 75% glycerol in PBST, 40 mM NaHCO3 (see Note 17). 2. After preparation of the embryo under a dissecting microscope, immerse your sample in the mounting gel solution (see preparation of mounting gel under subheading 2.7). 3. Transfer the sample onto a microscope slide between spacers of respective heights and apply cover slip. 4. By gently moving the coverslip the sample can be rotated into the desired orientation. 5. Put the mounted sample into the fridge until the agarose has solidified.

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4  Notes 1. Be careful when working with 4-iodophenol, as it is a highly aggressive substance and should ONLY be handled under a fume hood wearing appropriate protections at all time (even when highly diluted). 2. Tyramide reagents diluted in absolute ethanol and stored at −20 °C are at least stable for 3 years. 3. The optimal fixing conditions for different stages have to be determined experimentally by adjusting fixation time and temperature. Embryos that are older than 1 day are usually fixed for 24 h at 4 °C. 4. Hydrogen peroxide treatment promotes embryo permeabilization properties resulting in improved signal detection. 5. As a rule of thumb, use comparable concentration of RNA probes as with chromogenic BCIP/NBT detection. Exaggerated probe concentrations will result in a decreased signal-to-­noise ratio. In case just two different mRNA transcript patterns are compared, combination of DIG- and DNPlabeled RNA probes are the best choice, as these yield stronger signals than when using fluorescein as hapten-label. Labeling probes with biotin is not recommended because biotin may produce high background signals in zebrafish embryos [24]. However, biotin-­labeled probes work well in Drosophila [27]. For a detailed descriptions of RNA probe preparation see [35]. 6. The anti-DNP-POD antibody (PerkinElmer) shows cross-­ reactivity with the periderm, which surrounds the zebrafish embryo as a thin cell layer during early stages of development. This usually results in visualization of the outline of the embryo. As the first round of detection usually requires only short staining times resulting in low background, this effect is minimized when detecting the DNP RNA first. 7. Please note that in each detection round only one POD-­ conjugated antibody is used. Be careful to apply the appropriate antibody according to the desired sequence of probe detection. In a dual FISH experiment we routinely apply first the anti-DNP-POD antibody and in the second round the anti-DIG-POD Fab fragments. 8. In general, 4-iodophenol is a more potent enhancer than vanillin. Higher concentrations of accelerator maybe used to further increase signal intensity, although adverse effects on the signal-to-noise ratio should be kept in mind. 9. Always make fresh TSA reaction buffer just before use. Preparing 1 ml reaction buffer in a 2 ml tube helps to minimize ­eventual spill of aggressive 4-iodophenol-rich solution during mixing.

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10. Since acidic POD inactivation greatly diminishes the signal intensity of FAM-tyramide staining [36], DyLight633-­ tyramide is usually used in the first and FAM-tyramide in the second round of detection. In case of three-color detection, the third transcript is visualized with carboxytetramethylrhodamine (TAMRA) tyramide [14]. 11. Since the 50% dextran sulfate stock solution is very viscose, use a cut 200 μl tip for pipetting. Take special care not to transfer embryos by accident from one tube to another. In order to avoid a decline in the signal-to-noise ratio the reaction time should not exceed 30 min. 12. The acidic inactivation step is pivotal to avoid that the first detected probe is visible in two fluorescence detection channels. Incubation in hydrogen peroxide solutions bears the risk of partial POD inactivation [14, 37]. This may cause false-­ positive colocalization signals. 13. We found that best results in a three-color experiment are obtained when the fluorescein-label is detected first with DyLight633 followed by DNP with TAMRA and DIG with FAM. 14. Immunofluorescence (IF) can be done before or after the FISH procedure. If the IF procedure is done before FISH, it is advisable to reduce the hybridization temperature to 55 °C. 15. If the IF procedure is done after FISH 8% sheep serum can be applied instead of Western blocking reagent in all blocking and antibody steps. If the IF is done before FISH, Western blocking reagent is obligatory to avoid RNase, which is contained in normal sheep serum. 16. If the IF procedure is done after FISH 1% DMSO is usually added to all buffers. 17. Using 40 mM NaHCO3 ensures that the pH is above 8 (see also Note 10). Stained embryos stored in 75% glycerol in PBST, 40 mM NaHCO3 are stable for several months and the use of special anti-fading agents is usually not necessary.

Acknowledgments Imaging was in part performed at the Live Cell Imaging unit, Department of Biosciences and Nutrition, Karolinska Institutet, Huddinge, Sweden, supported by grants from the Knut and Alice Wallenberg Foundation, the Swedish Research Council, and the Centre for Biosciences.

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References 1. Femino A, Fay FS, Fogarty K, Singer RH (1998) Visualization of single RNA transcripts in situ. Science 280(5363):585–590 2. Raj A, van den Bogaard P, Rifkin SA, van Oudenaarden A, Tyagi S (2008) Imaging individual mRNA molecules using multiple singly labeled probes. Nat Methods 5(10):877–879 3. Hauptmann G, Lauter G, Söll I (2016) Detection and signal amplification in zebrafish RNA FISH. Methods 98:50–59. https://doi. org/10.1016/j.ymeth.2016.01.012 4. Player AN, Shen LP, Kenny D, Antao VP, Kolberg JA (2001) Single-copy gene detection using branched DNA (bDNA) in situ hybridization. J Histochem Cytochem 49(5): 603–611 5. Choi HM, Chang JY, Trinh le A, Padilla JE, Fraser SE, Pierce NA (2010) Programmable in situ amplification for multiplexed imaging of mRNA expression. Nat Biotechnol 28(11):1208–1212. https://doi. org/10.1038/nbt.1692. [pii] nbt.1692 6. Choi HMT, Beck VA, Pierce NA (2014) Next-­ generation in situ hybridization chain reaction: higher gain, lower cost, greater durability. ACS Nano 8(5):4284–4294 7. Speel EJ (1999) Robert Feulgen Prize Lecture 1999. Detection and amplification systems for sensitive, multiple-target DNA and RNA in situ hybridization: looking inside cells with a spectrum of colors. Histochem Cell Biol 112(2): 89–113 8. Speel EJ, Ramaekers FC, Hopman AH (1995) Cytochemical detection systems for in situ hybridization, and the combination with immunocytochemistry, ‘who is still afraid of red, green and blue?’. Histochem J 27(11): 833–858 9. Bobrow MN, Harris TD, Shaughnessy KJ, Litt GJ (1989) Catalyzed reporter deposition, a novel method of signal amplification. Application to immunoassays. J Immunol Methods 125(1–2):279–285. [pii] 0022-1759 (89)90104-X 10. Speel EJM, Hopman AHN, Komminoth P (1999) Amplification methods to increase the sensitivity of in situ hybridization: play CARD(S). J Histochem Cytochem 47(3): 281–288 11. Kosman D, Mizutani CM, Lemons D, Cox WG, McGinnis W, Bier E (2004) Multiplex detection of RNA expression in Drosophila embryos. Science 305(5685):846. https://

doi.org/10.1126/science.1099247. [pii] 305/5685/846 12. Tessmar-Raible K, Steinmetz PR, Snyman H, Hassel M, Arendt D (2005) Fluorescent two-­ color whole mount in situ hybridization in Platynereis dumerilii (Polychaeta, Annelida), an emerging marine molecular model for evolution and development. BioTechniques 39(4):460, 462, 464. doi:000112023 [pii] 13. Clay H, Ramakrishnan L (2005) Multiplex fluorescent in situ hybridization in zebrafish embryos using tyramide signal amplification. Zebrafish 2(2):105–111. https://doi. org/10.1089/zeb.2005.2.105 14. Lauter G, Söll I, Hauptmann G (2011) Multicolor fluorescent in situ hybridization to define abutting and overlapping gene expression in the embryonic zebrafish brain. Neural Dev 6(1):10. https://doi.org/10.1186/17498104-6-10. [pii] 1749-8104-6-10 15. Vize PD, McCoy KE, Zhou X (2009) Multichannel wholemount fluorescent and fluorescent/chromogenic in situ hybridization in Xenopus embryos. Nat Protoc 4(6):975–983. https://doi.org/10.1038/nprot.2009.69. [pii] nprot.2009.69 16. Denkers N, Garcia-Villalba P, Rodesch CK, Nielson KR, Mauch TJ (2004) FISHing for chick genes: triple-label whole-mount fluorescence in situ hybridization detects simultaneous and overlapping gene expression in avian embryos. Dev Dyn 229(3):651–657. https:// doi.org/10.1002/dvdy.20005 17. Lecuyer E, Yoshida H, Parthasarathy N, Alm C, Babak T, Cerovina T, Hughes TR, Tomancak P, Krause HM (2007) Global analysis of mRNA localization reveals a prominent role in organizing cellular architecture and function. Cell 131(1):174–187. https://doi.org/10.1016/j. cell.2007.08.003. [pii] S0092-8674 (07)01022-7 18. Lauter G, Söll I, Hauptmann G (2013) Molecular characterization of prosomeric and intraprosomeric subdivisions of the embryonic zebrafish diencephalon. J Comp Neurol 521(5):1093–1118. https://doi.org/ 10.1002/cne.23221 19. Herget U, Ryu S (2015) Coexpression analysis of nine neuropeptides in the neurosecretory preoptic area of larval zebrafish. Front Neuroanat 9:2. https://doi.org/10.3389/ fnana.2015.00002 20. Hauptmann G, Gerster T (2000) Regulatory gene expression patterns reveal transverse and

Zebrafish Multiplex RNA FISH longitudinal subdivisions of the embryonic zebrafish forebrain. Mech Dev 91(1–2):105– 118. [pii] S0925-4773(99)00277-4 21. Hauptmann G, Söll I, Gerster T (2002) The early embryonic zebrafish forebrain is subdivided into molecularly distinct transverse and longitudinal domains. Brain Res Bull 57(3– 4):371–375. S0361923001006918 [pii] 22. Affaticati P, Yamamoto K, Rizzi B, Bureau C, Peyrieras N, Pasqualini C, Demarque M, Vernier P (2015) Identification of the optic recess region as a morphogenetic entity in the zebrafish forebrain. Sci Rep 5:8738. https:// doi.org/10.1038/srep08738 23. Tautz D, Pfeifle C (1989) A non-radioactive in situ hybridization method for the localization of specific RNAs in Drosophila embryos reveals translational control of the segmentation gene hunchback. Chromosoma 98(2):81–85 24. Hauptmann G (1999) Two-color detection of mRNA transcript localizations in fish and fly embryos using alkaline phosphatase and betagalactosidase conjugated antibodies. Dev Genes Evol 209(5):317–321 25. Hauptmann G (2001) One-, two-, and three-­ color whole-mount in situ hybridization to Drosophila embryos. Methods 23(4):359–372. https://doi.org/10.1006/meth.2000.1148. [pii] S1046-2023(00)91148-4 26. Hauptmann G, Gerster T (1994) Two-color whole-mount in situ hybridization to vertebrate and Drosophila embryos. Trends Genet 10(8):266 27. Hauptmann G, Gerster T (1996) Multicolour whole-mount in situ hybridization to Drosophila embryos. Dev Genes Evol 206(4):292–295. https://doi.org/10.1007/ s004270050055 28. Jowett T, Lettice L (1994) Whole-mount in situ hybridizations on zebrafish embryos using a mixture of digoxigenin- and fluorescein-­ labelled probes. Trends Genet 10(3):73–74 29. O’Neill JW, Bier E (1994) Double-label in situ hybridization using biotin and digoxigenin-­

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Chapter 23 Live Morphometric Classification of Sensory Neurons in Larval Zebrafish Gema Valera and Hernán López-Schier Abstract Sensory systems convey environmental information to the brain. A comprehensive description of neuronal anatomy and connectivity is essential to understand how sensory information is acquired, transmitted, and processed. Here we describe a high-resolution live imaging technique to quantify the architecture of sensory neurons in larval zebrafish. This approach is ideal to assess neuronal-circuit plasticity and regeneration. Key words Sensory system, Lateral line, Zebrafish, Neuronal morphometry

1  Introduction Sensory reception allows organisms to sample the environment and to react appropriately [1–3]. Sensory organs communicate environmental cues to the central nervous system through neuronal afferent circuits. The assembly of a coarse sensory circuit during embryogenesis is often governed by intrinsic genetic programs, and is subsequently refined by evoked activity. One widely conserved property of sensory systems is that the spatial distribution of the peripheral receptors is represented in the central nervous system by layered neuronal projections [4]. This type of central mapping is used by the visual system, where it receives the name of retinotopy, the olfactory system (rhinotopy), and in body mechanoreception (somatotopy). In this chapter, we explain how to trace neurons and describe their central architecture at submicrometer resolution. As an example, we will use the mechanosensory lateral-line afferent neurons (LANs) in 5-days-old larval zebrafish, and address their central and peripheral arborization and connectivity [5–9]. Fishes and amphibians rely on this system to detect hydromechanic variations around their bodies [10, 11]. The lateral-line system offers a simple model of sensorineural circuits whose dynamics can be visualized over Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_23, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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long periods and under controlled conditions [12, 13]. The method that we present here is based on transient expression of engineered fluorescent proteins in individual neurons after microinjection of DNA into fertilized eggs [6, 7, 12, 13]. It can be implemented in most biology laboratories. It requires minimal technical expertise, a small zebrafish colony, and off-the-shelf inexpensive equipment and materials to permit DNA microinjection, screening of specimens by fluorescent microscopy, general molecular biology tools, a confocal microscope for intravital imaging, and a computer equipped with appropriate software for three-­ dimensional data visualization and analysis.

2  Materials 2.1  Fish Care and Breeding

1. Equipment and facilities needed to raise and cross fish and collect eggs: breeding tanks, strainers, and 80 mL plastic Petri dishes. 2. Egg water: 1.5 mL of 40 mg/mL stock salts added to 1 L distilled water. 3. E3 water stock solution (60×): mix 172 g NaCl, 7.6 g KCl, 29 g CaCl2·2H2O, and 49 g MgSO4·7H2O in 10 L of distilled water. Store at 4 °C. 4. Transgenic zebrafish lines: Tg[HGn39D] and Tg[Myo6b:β-­ actin-­GFP] [6, 14] (see Note 1).

2.2  DNA Injection and Screening

1. A stereomicroscope and material needed for injection: glass capillary, puller, microinjector, micropipette holder, sharp forceps, 1% agarose dish with lanes, plastic Pasteur pipette, and 10 μL pipette tip. 2. DNA plasmid cloned with Tol2Kit® (or similar) in RNase-free water. Here we use SILL1 (hsp70:mCherry-SILL) which specifically labels single Lateralis Afferent Neurons (LANs). 3. Tricaine (MS222) stock solution (25×): 0.4 g tricaine, 97.9 mL distilled water. Adjust pH to 7 by adding 9.1 mL of 1 M Tris– HCl (pH 9). Store at 4 °C. 4. Fluorescence stereomicroscope to see GFP and mCherry.

2.3  Mounting and Live Imaging

1. Stereomicroscope and material needed for mounting: tricaine 1× in E3 water, cover-glass bottomed mini dish, 1% low-­ melting-­point agarose in E3 water, 1-hair brush, and micropipette p1000. 2. Confocal microscope equipped with 20×-dry, 40×-dry, and 63×-water-immersion objectives and lasers for excitation at 488 nm and 587 nm.

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1. Fiji software. 2. Amira 5.3.3 software.

3  Methods 3.1  Labeling of LANs

1. One day before the DNA injection, cross adult Tg[HGn39D] and Tg[Myo6b:β-actin-GFP] zebrafish in pairs, separating the male and the female by a transparent plastic divider. The resulting double transgenic progeny carrying HGn39D and Myo6b:β-­actin-­GFP will reveal the planar polarity of the hair cells and the LANs. 2. Pull glass microcapillaries. 3. The next morning open the barrier, wait for the oviposition and fertilization, collect the eggs, and place them in a Petri dish. 4. Rinse the embryos in egg water at 28.5 °C in Petri dishes with a density of approximately 50 embryos per plate. 5. Place eggs over a 1% agarose lawn containing depressed lanes that will hold the eggs in place. 6. Fill microcapillaries with a 20 ng/μL dilution of DNA plasmid SILL1. Inject this solution into the eggs at 2-cells stage, releasing into the cell a solution bolus of approximately 1/6 of the cell’s volume (see Fig. 1). This will amount to around 20 pg of DNA.

Fig. 1 Scheme of the injection of SILL1 in 2-cells eggs, the monomeric mCherry driven by a minimal promoter hsp70 and an enhancer specific of lateral-line afferent neurons (LANs). Injected eggs are raised in egg water at 28.5 °C and embryos liberated from the chorion are screened to select samples with a single LAN labeled in red. Neuromasts are named with an “L” followed by a number that progressively increases from the head until the end of the trunk (L1, L2, L3, L4, and L5). The cluster of neuromasts at the tip of the tail is known as terminal neuromasts (T). In this example, the neuron innervates the neuromast L1 of the posterior lateral line

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7. One or two days later, use a fluorescence stereomicoscope screen anesthetized embryos to select those ones with one single LAN labeled with mCherry. Change the water on a daily basis keeping the water clean (see Note 2). Look for a single neuronal somata in the lateral-line ganglia, located at either side of the otic vesicle (see Notes 3 and 4). 8. Ensure that specimens are Tg[HGn39D] and Tg[Myo6b:β-­ actin-­GFP] by checking that, respectively, all LANs and hair cells are GFP positive. 3.2  Imaging of Central and Peripheral Projections of Single LANs

1. Anesthetize selected larvae and mount them sideways on a glass-bottom mini dish by embedding them into a drop of 1% low melting-point agarose in E3 solution, right at the point of agarose solidification. This will avoid overheating the fish. After agarose hardening, cover the agarose dome with tricaine 1× in 3 solution (see Notes 5 and 6). 2. Image embryos with a confocal microscope equipped with a 20× dry objective to image the central arbor including the ganglion (see Note 7). Image GFP (488 nm) and mCherry (587 nm) to acquire z-stacks with a z-step of 1 μm. Adjust the exposure according to the intensity of the fluorescent signal and acquire enough focal planes over the whole volume of the lateralis central column by using the GFP signal from Tg[HGn39D] as reference (see Note 8 and Fig. 2). 3. Next, image specimens with a 63× water-immersion objective to assess hair-cell innervation. Acquire z-stacks with a z-step of 0.6 μm of the whole volume of the peripheral arborization as well as the entire apicobasal length of hair cells suing the GFP signal from Tg[Myo6b:β-actin-GFP] as reference.

3.3  Single LANs Tracing

1. Make a composite of the two fluorescent channels using the Fiji software (see Fig. 2c). 2. Open the composite with the Amira 5.3.3 software, and choose All channels in order to access GFP and mCherry. On the resulting tab, introduce the voxel size of the image for a correct 3D visualization of the neuron. 3. Use the Filament Editor to draw the neuron in a semi manual manner. Select the red channel and start bringing to the maximum the Thickness to work with the maximal projection of the sample. 4. Choose the center of the soma as starting point of the tracing schedule, by clicking on it and then drawing lines along the central axon until the first bifurcation. From the bifurcation point, draw lines along each branch until the following bifurcation (see Notes 9–11 and Fig. 3a). Repeat this work until ­reaching the end of each neurite revealed by a terminal varicosity (see Note 12).

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Fig. 2 Maximal projection of a single LAN z-stack in a 6-days-old zebrafish larva of Tg[Hgn39D]. (a) Central arbor of a L1 neuron genetically labeled in mCherry by injection of SILL1 (hsp70:mCherry_sill enhancer). (b) Entire column of central projections of all LANs of one half of the lateral-line system labeled in GFP in Tg[HGn39D]. (c) Merge of both channels

5. Finish work by removing intermediate nodes to leave only the ending point and branch points. 6. Run the action Identify Graphs found in Tools and after that select Identified_Graphs, both for node coloring and segment coloring, this will highlight all the branches and varicosities in different colors. Select the desired size for the axon and the varicosity points (see Notes 13 and 14 and Fig. 3b, c). 7. Save the skeleton in the same folder where the composite is located. 3.4  Quantification of Central-Arbor Complexity

1. Amira displays the number of points marked on the skeleton, which is the number of synaptic buttons. 2. Count and write down the number of secondary and subsequent branches. 3. Use the bifurcations as landmarks to split regions of desired size (see Note 15).

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Fig. 3 Progression of semi-manual skeletonization of the L1 neuron in Fig. 2 with Amira 5.3.3. (a) Early status of the skeletonization in which only the main branches have been traced. (b) Intermediate status of the skeletonization including secondary branches. (c) Final skeleton with all the branches traced and all the varicosities marked with spheres. (a–c) Colors indicate individual branches. Varicosities are highlighted under the same color code of the branch which they belong to. (d) Summary table of the number of secondary branches and varicosities. The bifurcation of the neuron serves to define anterior and posterior sides of the arbor in the longitudinal axis of the fish 3.5  Quantification of Peripheral-Arbor Complexity and Connectivity

1. Open the z-stack of a neuromast innervated by a LAN and make a composite of the two channels with Fiji (see Fig. 4). 2. Move back and forth through the planes in the composite z-stack and write down which hair cells are innervated by the singly labeled LAN. Connectivity can be assigned based on swelling in the neuronal terminals adjacent to the basal aspect of the hair cell.

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Fig. 4 Planar polarity hair cells and their innervation, as in Figs. 2 and 3. (a) Yellow arrows indicate innervated hair cells. (b) Maximal-intensity projection of selected z-planes where innervation is visible (enlarged red spots of the neuronal arbor). (c) Detail of the hair bundles showing the planar polarity of each hair cell. The innervated hair cells are rostrocaudal in this sample. (d) Maximal intensity projection of a neuromast innervated by the single LAN

3. Follow the innervated hair cells from their base to the apical hair bundle, and write down the polarity of the hair cell according to the position of the kinocilium, seen as a dark spot (see Note 16 and Fig. 4).

4  Notes 1. Ideally, the genetic background of the animals should be such that it prevents melanization. For instance, using the casper or crystal lines. This will facilitate imaging by avoiding dark epithelial cells obscuring the underlying neurons. 2. It is recommended to provide Ca2+ to embryos by starting using E3 water instead of egg water after the hatching of the eggs. 3. It is recommended to screen the fish when they are 3 or 4 days old, the expression of mCherry begins about the second day but it is still too weak. 4. Exclude those samples with more than one neuron given that it is very difficult to distinguish them during the description of the central complexity of the arbors. 5. The side of the fish where the single LAN is located should remain the closest to the objective, flip the fish if needed.

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6. The best position to image the central projection is with the fish slightly tilted to leave the dorsal side of the otic vesicle more accessible to the objective. However, the trunk of the fish must be completely flat and parallel to the bottom of the dish in order to assure a good image of the neuromast to determine the planar polarity of the hair cell. 7. Neurons were imaged in 2 z-stacks and subsequently they were stitched with Fiji software. Nevertheless, we have realized that imaging the same neurons with 20× is faster and changes in resolution are not appreciated. 8. The central column is the result of central projections of all the LANs from the ALL, DLL, and PLL. 9. It is possible to work on single- or double-panel view. 10. The Amira software will draw lines from the last point highlighted in red. This can be changed it by clicking on q different point under the option Select a single node, edge or point. 11. It is helpful to draw the neuron in stretches without covering long distances. We have discovered that the Fiji software is faster using this approach if the fluorescent signal is weak. 12. The varicosities are seen as enlarged spots at the end of the axon terminals. They will be drawn as small spheres in the final skeleton to facilitate analysis. 13. It is advisable to compare the skeleton with the original confocal image to correct potential errors, by manually removing or adding branches or varicosities on the skeleton. 14. Change the channel to GFP in order to see the location of the neuron, represented by its skeleton, within the column. 15. For a more specific analysis of branches and varicosities distribution, one can divide the entire column in boxes and count the number of varicosities by them fitting inside each box. For this purpose, built a grid of 1 row × 8 columns with Photoshop and placed it over the composite image with the skeleton. 16. The planar polarity of hair cell has a direction determined by the location of the kinocilium within the hair bundle.

Acknowledgments We thank the Central Animal Facility of the HMGU (Munich, Germany) for expert animal care. This work was supported by the Helmholtz Gemeinschaft. Ethical considerations: All procedures on live specimens should be performed under guidelines and approved protocols by local and general/federal agencies. The method and experiments described here were performed according to EU Directive 2010/63/EU,

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under which an ethics committee-approved animal protocol is required for freely feeding zebrafish (older than 5.5 days). Work in our laboratory employs zebrafish of 5 days or younger. References 1. Schwander M, Kachar B, Muller U (2010) The cell biology of hearing. J Cell Biol 190:9–20 2. Sung CH, Chuang JZ (2010) The cell biology of vision. J Cell Biol 190:953–963 3. Lumpkin EA, Marshall KL, Nelson AM (2010) The cell biology of touch. J Cell Biol 191:237–248 4. Luo L, Flanagan JG (2007) Development of continuous and discrete neural maps. Neuron 56:284–300 5. Rouse GW, Pickles JO (1991) Paired development of hair cells in neuromasts of the teleost lateral line. Proc Biol Sci 246:123–128 6. Faucherre A, Pujol-Martí J, Kawakami K, López-Schier H (2009) Afferent neurons of the zebrafish lateral line are strict selectors of hair-cell orientation. PLoS One 4:e4477 7. Lozano-Ortega M, Valera G, Xiao Y, Faucherre A, López-Schier H (2018) Hair cell identity establishes labeled lines of directional mechanosensation. PLoS Biol 16(7):e2004404 8. Ji YR, Warrier S, Jiang T, Wu DK, Kindt KS (2018) Directional selectivity of afferent neurons in zebrafish neuromasts is regulated by Emx2 in presynaptic hair cells. elife 7:pii: e35796 9. Dow E, Jacobo A, Hossain S, Siletti K, Hudspeth AJ (2018) Connectomics of the

zebrafishs lateral-line neuromast reveals wiring and miswiring in a simple microcircuit. Elife 7:pii: e33988 10. Pujol-Martí J, López-Schier H (2013) Developmental and architectural principles of the lateral-line neural map. Front Neural Circuits 7:47 11. Oteíza P, Odstrcil I, Lauder G, Portugués R, Engert F (2017) A novel mechanism for mechanosensory-­ based rheotaxis in larval zebrafish. Nature 547(7664):445–448 12. Xiao Y, Faucherre A, Pola-Morell L, Heddleston JM, Liu TL, Chew TL, Sato F, Sehara-Fujisawa A, Kawakami K, López-Schier H (2015) Highresolution live imaging reveals axon-glia interactions during peripheral nerve injury and repair in zebrafish. Dis Model Mech 8(6):553–564 13. Pujol-Martí J, Faucherre A, Aziz-Bose R, Asgharsharghi A, Colombelli J, Trapani JG, López-Schier H (2014) Converging axons collectively initiate and maintain synaptic selectivity in a constantly remodeling sensory organ. Curr Biol 24(24):2968–2974 14. Kindt KS, Finch G, Nicolson T (2012) Kinocilia mediate mechanosensitivity in developing zebrafish hair cells. Dev Cell 23(2):329–341

Chapter 24 Immunohistochemistry and In Situ Hybridization in the Developing Chicken Brain Richard P. Tucker, Tatsuto Ishimaru, and Qizhi Gong Abstract One of the first steps in studies of gene function is the spatiotemporal analysis of patterns of gene expression. Indirect immunohistochemistry is a method that allows the detection of a protein of interest by incubating a histological section with an antibody or antiserum raised against the protein, and then localizing this primary antibody with a tagged secondary antibody. To determine the cellular source of a protein of interest, or if a specific antibody is not available, specific transcripts can be localized using in situ hybridization. A histological section is incubated with a labeled RNA probe that is complementary to the target transcript; after hybridization with the target transcript the labeled RNA probe can be identified with an antibody. Here we describe materials and methods used to perform basic indirect immunohistochemistry and in situ hybridization on frozen sections through the developing chicken brain, emphasizing controls and potential problems that may be encountered. Key words Immunohistochemistry, In situ hybridization, Riboprobe, Antibody, Fluorescence, Protocol, Technique, Cryosection

1  Introduction Immunohistochemistry was first used in the 1940s to identify bacterial antigens in mouse tissues by applying fluorescein-labeled antibodies to frozen histological sections [1]. The technique was modified in the following decades to amplify the signal by “indirectly” identifying the so-called primary antibodies with labeled secondary antibodies [2]. The basic method of indirect immunohistochemistry is unchanged to this day: a histological section is incubated with a primary antibody (e.g., a mouse monoclonal antibody specific to an antigen of interest), then the section is incubated with a tagged secondary antibody (e.g., fluorescently tagged rabbit polyclonal antibodies against mouse antibodies) and the protein of interest is then observed using a microscope fitted with special illumination and optics. The technique is widely used in the neurosciences for determining patterns of expression of novel gene Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_24, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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products as well as for identifying specific types of neurons and supporting cells in brain sections. In situ hybridization is used to detect transcripts in their native tissue environment. The technique was first used in the early 1980s to study gene expression in Drosophila embryos [3, 4], but it was quickly adapted to studies of vertebrate brain [5]. In short, a tissue section is treated to make it amenable to hybridization, then it is incubated with a labeled nucleic acid probe with sequences complementary to the target transcript. Excess probe is removed and the label is detected. There are several established methods for tissue in situ hybridization [6]. One of the major differences between the published methods is the choice of hybridization probes; in general, RNA probes provide more sensitive detection and cleaner background [7]. Probe detection has improved over the years, making it easier to detect rarer transcripts and to perform double-­ label techniques to identify two different transcripts in the same section [8]. Here we present detailed protocols for fixing and cryosectioning embryonic chicken brains as well as methods for performing indirect immunohistochemistry and in situ hybridization with RNA probes on these frozen sections. The methods are easily adapted to other tissues and to tissues from other species.

2  Materials The required materials are listed below in three sections: Cryosectioning (Subheading 2.1), Immunohistochemistry (Subheading 2.2), and In Situ Hybridization (Subheading 2.3). 2.1  Cryosectioning Components

1. Buffer: Phosphate buffered saline, pH 7.4 (PBS). To make 10× PBS, dissolve 10 g NaCl, 2 g KCl, 11.5 g Na2HPO4 7H2O, and 2 g KH2PO4 in 800 mL of ddH2O. Bring volume to 1 L and adjust pH. 2. Fixative: 4% paraformaldehyde in PBS, pH 7.4, made fresh (same day). Add 2 g of reagent grade paraformaldehyde powder (Millipore-Sigma, Burlington, MA, USA) to 40 mL of PBS in a 50 mL conical tube. Add two NaOH pellets (MilliporeSigma), cap, and allow the pellets to dissolve with gentle shaking. When the paraformaldehyde is in solution, adjust the pH (using pH sensitive paper) with 2 N HCl (Millipore-Sigma), then top off with PBS to a final volume of 50 mL. Store on ice (see Note 1). 3. Cryoprotection: Sucrose (Fisher Scientific, Waltham, MA, USA). 4. Embedding medium: Tissue-Tek O.C.T. Compound (Sakura Finetek, Torrance, CA, USA).

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5. Embedding molds: Disposable base molds (various sizes; e.g., 15 × 15 × 5 mm; Ted Pella, Redding, CA, USA). 6. Freezing solution: 2-methylbutane (Fisher Scientific). Just before use, break up a 10 cm × 10 cm × 3 cm block of dry ice into powder and chunks approximately 1 cm3. Put the dry ice into a glass dish and add the 2-methylbutane to make a slurry 1–2 cm deep. 7. Slides: Superfrost Plus precleaned (25 × 75 × 1 mm; Fisher Scientific).

microscope

slides

8. Cryostat microtome: For example, Leica CM3050. 2.2  Immunohistochemistry Components

1. Buffer: PBS (see above). 2. Blocking agent: 0.5% bovine serum albumin (BSA) (Fisher Scientific) in PBS. 3. Coplin jars: Polypropylene Coplin staining jar (01-816-21, Fisher Scientific). 4. Staining tray: Any flat-bottomed glass or plastic tray, approximately 5 cm deep. Place a layer of paper towels on the bottom and dampen with water, pushing out excess water and air bubbles to keep the surface flat. Cover with sticky plastic wrap (Fig. 1). 5. Antibodies: Secondary antibodies should be against the animal source of the primary antibody, and the fluorescent marker should be compatible with local fluorescent microscopy filter sets.

Fig. 1 Schematic illustration of an immunohistochemistry incubation chamber, viewed from above (top) and the side (bottom). The chamber should have a flat bottom and either a securely fitting lid or a lid fashioned from plastic wrap. As the slides rest directly on the moist paper towel care should be taken not to let the slides touch each other, and the “puddle” of antibody should not extend to the very edge of the slide

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6. Coverslips: Various sizes (e.g., 22 × 30 mm, Corning, Corning, NY, USA). 7. Nuclear stain: Hoechst 33342 (Millipore-Sigma); Make a 2 mg/mL 100× stock and store in a foil-wrapped tube at 4 °C. 8. Wet coverslip mounting medium: 5 mL PBS/5 mL glycerol. 2.3  In Situ Hybridization Components

1. cDNA clones in an RNA polymerase binding site containing plasmid.

2.3.1  For Riboprobes

3. QIAquick gel extraction kit (Qiagen, Hilden, Germany).

2. Desired restriction enzymes to linearize the plasmid. 4. 100% ethanol (RNase-free). 5. T7 and Sp6 RNA polymerase (or appropriate RNA polymerase) 20 U/μL (Promega, Madison, WI, USA). 5× transcription buffer and 100 mM DTT are provided with the enzyme. 6. Fluorescein (FLU) and digoxigenin (DIG) RNA labeling mix (Millipore-Sigma). 7. RNase-free DNase 10 U/μL (Promega). 8. RNase inhibitor 20 U/μL (New England Biolabs, Ipswich, MA, USA). 9. Micro Bio-Spin 30 Columns (Bio-Rad, Hercules, CA, USA).

2.3.2  For Hybridization

1. Hair dryer with cool air setting. 2. Slide mailer (Ted Pella). 3. Plastic storage box with a tight-fitting lid. 4. Microscope cover glass (e.g., 40 × 22 or 50 × 22 mm; Fisher Scientific or Corning). 5. DEPC-treated H2O: Add 1 mL of DEPC to 1 L of ddH2O, mix well. Let stand at room temperature overnight or 37 °C for 2 h. Autoclave at least 15 min to inactivate the DEPC. 6. DEPC-treated PBS: For DEPC-treated 1× PBS, combine 100 mL of 10× PBS (see above) and 900 mL of ddH2O, add 1 mL of DEPC, mix well by shaking, let stand in room temperature overnight. Autoclave to inactivate the DEPC. 7. 1 M Tris–Cl, pH 8.0: Dissolve 60.5 g Tris base in 400 mL DEPC-treated H2O, adjust pH to 8.0 with concentrated HCl, bring volume up to 500 mL with DEPC-treated H2O. Filter-­ sterilize (see Note 2). 8. 1 M Tris–Cl, pH 7.5 and pH 9.5: Same as above; adjust pH to 7.5 and 9.5, respectively. 9. 0.5 M Ethylenediaminetetraacetic acid (EDTA), pH 8.0: 19 g of EDTA in 80 mL DEPC-treated H2O, add 10 N NaOH (about 4 mL) and mix to dissolve, adjust pH to 8.0 with NaOH, bring volume to 100 mL with DEPC-treated H2O. Filter-sterilize.

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10. TE buffer, pH 8.0: 5 mL of 1 M Tris–Cl, pH 8.0 and 1 mL of 0.5 M EDTA. Bring to 500 mL with DEPC-treated H2O. 11. Proteinase K: 12.8 μL of 15.6 mg/mL Proteinase K (539480, Millipore-Sigma) in 20 mL of TE buffer, pH 8.0. 12. 2 N HCl. 13. 1 M Triethanolamine-HCl stock: Combine 66.25 mL of triethanolamine and 11.25 mL of HCl into 500 mL of DEPC H2O. Dilute with DEPC-treated H2O to make 0.1 M Triethanolamine-HCl. 14. Acetic anhydride (Millipore-Sigma). 15. Ethanol series: Dilute ethanol with DEPC-treated H2O to make 50%, 75%, 90% ethanol. 16. Hybridization solution (all reagents from Millipore-Sigma): Stock

For 10 mL solution

Final concentration

Formamide

5 mL

50%

Yeast tRNA, 10 mg/mL

200 μL

0.2 mg/mL

Dextran sulfate, 50%

2 mL

10%

1 M Tris–Cl, pH 8.0

100 μL

10 mM

0.5 M EDTA, pH 8.0

20 μL

1 mM

Denhardt’s, 50×

100 μL

0.5×

5 M NaCl

1.2 mL

600 mM

10% SDS

250 μL

0.25%

DEPC-treated H2O

To 10 mL

17. 20× SSC: Dissolve 175.3 g of NaCl, 88.2 g of sodium citrate (Na3C6H5O7∙2H2O) in 800 mL of DEPC-treated H2O, adjust pH to 7.0 with 1 M HCl, add DEPC-treated H2O to i1 L. 18. RNase A (Millipore-Sigma) stock solution 10 mg/mL. 19. 5 M NaCl: Dissolve 146.1 g NaCl in 450 ddH2O, mix by stirring; add ddH2O to 500 mL. 20. TNE buffer: Combine 5 mL of 1 M Tris–Cl, pH 7.5, 50 mL of 5 M NaCl, 1 mL of 0.5 M EDTA pH 8.0 and 444 mL of ddH2O. 21. Buffer 1: 100 mM Tris–HCl pH 7.5, 150 mM NaCl. 22. Blocking reagent (Roche Applied Science, Mannheim, Germany). 23. Alkaline phosphatase (AP) conjugated anti-DIG antibody (Roche Applied Science). 24. Buffer 2: 10 mM Tris pH 8.0, 100 mM NaCl, 10 mM MgCl2).

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25. NHPP Fluorescent detection set (Millipore-Sigma). 26. POD conjugated (Millipore-Sigma).

anti-FLU

antibody,

Fab

fragment

27. TSA Biotin system (PerkinElmer, Waltham, MA, USA). 28. Alexa488 conjugated Streptavidin (Jackson ImmunoResearch, West Grove, PA, USA). 29. Fluoromount-G (SouthernBiotech, Birmingham, AL, USA).

3  Methods If studying a range of developmental stages it can be convenient to put the fertilized chicken eggs into the incubator at different intervals and then collect and process all of the embryos at once. Be sure to follow appropriate institutional and regional ethical guidelines for the treatment of research animals. The fixation and sectioning protocol described below is used to make frozen section suitable for both immunohistochemistry and in situ hybridization. 3.1  Cryosectioning

1. Remove the embryo from the egg by cracking the egg into a small bowl containing a small volume of PBS (room temperature). After trimming put the whole embryo (embryonic day [E]4–E7) or the head (E8–E16) into a 50 mL conical centrifuge tube with ice cold 4% paraformaldehyde in PBS (pH 7.6). Cap the tube and place it on ice on a rotary shaker for 1–6 h, then store the tube at 4 °C overnight (see Note 3). 2. Pour off and dispose of the fixative appropriately and add ice-­ cold PBS to the tube. Cap the tube and place on a rotary shaker in an ice bath for 30 min. Pour off the PBS and repeat twice with fresh PBS. 3. Add sucrose to the third PBS rinse to a final concentration of 20–25%. Cap the tube and place on rotating shaker until the sucrose has gone into solution. The tissues should ‘float’ on the sucrose. Store the tube with floating tissues at 4 °C overnight (see Note 4). 4. Put the sucrose infiltrated (cryoprotected) tissues into a petri dish or onto a piece of Parafilm© and trim them with a fresh razor blade such that the trimmed surface will match the orientation of the sections to be cut in the cryostat. Transfer the trimmed tissues to a “puddle” of embedding compound to minimize the presence of any sucrose solution, then transfer the tissue to an embedding mold partially filled with embedding medium. With forceps orient the tissue so that the cut surface is resting along the bottom of the mold (see Note 5). Float the mold in a shallow slurry of 2-methylbutane and

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crushed dry ice, taking care not to let the 2-methylbutane come into direct contact with the embedding medium. When the embedding medium is solid white, transfer the mold with the frozen tissue to dry ice (see Note 6). 5. Cut sections with a cryostat using appropriate protocols and collect the sections on pre-subbed slides (see Note 7). Allow the sections to air dry for 4–6 h, then either proceed with the protocol or store the slides at −20 °C to −70 °C (see Note 8). 3.2  Immunohistochemistry

All immunohistochemistry procedures described below are carried out at room temperature. 1. Allow slides to come to room temperature (if stored in the freezer), then place them into a Coplin jar or slide staining rack (depending on the number of slides being processed) with PBS for 10–30 min. Pour off the PBS and add PBS with BSA blocking solution (see Note 9). Incubate the slides in blocking solution for 15–30 min (longer blocking can reduce background, but typically is not necessary). 2. Dilute the primary antibody in blocking solution in a 1.5 mL snap cap tube. It may be necessary to use a range of dilutions (e.g., 1:10, 1:100 and 1:1000) with antibodies that have not been characterized before. Spin the diluted antibodies in a table top microfuge at 9000–13,000 rpm for 5 min. Reserve a few mL of blocking solution for the secondary antibody (store at 4 °C until used). 3. As the primary antibody is spinning remove the slides from the blocking solution. Wipe away excess solution using a KimWipe© taking care not to disturb the section and taking care not to let the section dry out (see Note 10). Label the slide and place it on the moist paper towel in the incubation chamber; add an appropriate volume of diluted and centrifuged primary antibody until the section is completely covered (about 200 μL). The antibody should stay on the region that is wet and not spread onto the parts of the slide that were dried with the KimWipe©. 4. Cover the chamber (e.g., with a clingy plastic wrap) and leave undisturbed on the bench top overnight (Fig. 1). 5. Gently tap the primary antibody from the slides onto a KimWipe© and rinse the slides three times for 10 min in PBS. 6. Dilute the secondary antibody (following the manufacturer’s instructions) in the reserve blocking solution in a 1.5 mL snap cap tube and centrifuge in a table top microfuge at 9–13K rpm for 5 min. 7. As the secondary antibody is spinning, remove the slides from the PBS rinse and carefully dry them (as above), leaving the

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area where the section is found wet. Place the slides back into the incubation chamber and add the secondary antibody until it covers the section. Take the same precautions noted above for adding the primary antibody. 8. Cover the chamber and incubate the slides with the secondary antibody for 1–3 h (longer incubations are possible, but usually not necessary). 9. Gently tap the secondary antibody off each slide onto a KimWipe© and rinse the slides three times for 10 min in PBS. Optional: Sections can be counterstained with a fluorescent nuclear dye at this stage. Between the second and third PBS rinse put the slides into a Coplin jar with nuclear stain (Hoechst 33342) diluted to the 1× working concentration in PBS for 1 min. 10. Coverslip carefully, without introducing air bubbles, using either the PBS:glycerol wet mount or a permanent mounting medium suited for fluorescence microscopy (e.g., Fluoromount-­G) and appropriately sized glass coverslips (see Note 11). 11. One or more control incubations should also be run. If the antibody is well characterized, it may be sufficient to incubate a section in the blocking solution overnight and then treat it with the secondary antibody the following day. Such “secondary antibody only” controls will reveal background fluorescence or nonspecific binding of the secondary antibody. When illustrating this type of control be sure to use the same camera settings that were used when imaging the experimental sections. If a polyclonal antiserum is used one should run a control that includes similarly diluted pre-immune serum. 3.3  In Situ Hybridization

3.3.1  Preparation of Labeled Riboprobes

In situ hybridization is a multistep procedure that can take up to 3 days. For better planning, make all solutions ahead of time. Frozen sections can be prepared days to weeks ahead and stored at −20 °C. DIG and FLU labeled RNA probes need to be made and the quality of the probes evaluated before starting in situ hybridization. The method below describes double-label in situ ­hybridization, which allows two different transcripts to be identified in the same tissue section, as well as single-label in situ hybridization. Typical results with two probes are illustrated in Fig. 2. 1. cDNAs were cloned into an appropriate vector. Orientation was confirmed by sequencing (see Note 12). 2. 10 μg of cDNA-containing plasmids were linearized at either 5′ end for an antisense probe, or at the 3′ end of the cDNA for a control sense probe, with appropriate restriction endonucle-

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Fig. 2 An example of double-label in situ hybridization. (a) A section through the olfactory bulb following in situ localization with a FLU-labeled probe to Tenm1. (b) To identify the cell type expressing Tenm1, the section was also incubated with a DIG-labeled probe to vGlut1, a marker of mitral and tufted cells. (c) Color images can be merged to show co-expression. Hoechst 33342 was used as a nuclear counterstain (blue). GL glomerular layer, MCL mitral cell layer

Fig. 3 Steps for making riboprobes. An appropriate cDNA fragment is cloned into the pCRII-TOPO plasmid (for example) with an Sp6 promoter at its 5′ end. Restriction enzyme Spe I and EcoRV sites are located at different ends of the cloned cDNA. To make antisense probes, plasmids are digested with Spe I and transcribed with T7 RNA polymerase. To make control sense probes, plasmids are digested with EcoR V and transcribed with Sp6 RNA polymerase. DIG or FLU-labeled UTPs are used to incorporate label (indicated by black dots) into the riboprobes

ases (Fig. 3). The restriction digest should be done overnight to ensure completeness (see Notes 13 and 14). 3. Take out 1/20 of the reaction to run on an agarose gel to confirm that the restriction digest is complete. 4. Purify the restriction digest reactions with QIAquick gel extraction kit: add 300 μL of QG, mix by vortexing, then add 100 μL of isopropanol to the sample, mix, load to the column,

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wash with PE, spin additional 1 min then elute with 30 μL of EB. Steps are described in detail on the product sheet. 5. Measure the concentration of linearized DNA using spectrophotometer or Nanodrop spectrophotometer. 6. To make labeled RNA probes, assemble the reaction as follows: Template DNA

1 μg

5× transcription buffer

4 μL

DIG or FITC RNA labeling mix

2 μL

0.1 M DTT

1 μL

RNA polymerase

2 μL

RNase inhibitor

0.5 μL

RNase-free water to 20 μL

7. Incubate the reaction at 37 °C for 2 h. 8. Add the following and incubate at 37 °C for 15 min: RNase inhibitor

1 μL

DNase

1 μL

9. Purify riboprobes with Micro Bio-Spin 30 Columns by following manufacture’s instruction (see Note 15). 3.3.2  In Situ Hybridization: Day 1. Prehybridization and Hybridization— RNase-Free

1. Dry sections with cold air using a hair dryer for up to 1 min (see Note 16). 2. All prehybridization and subsequent washing steps are done in a vertical slide mailer that holds five slides. Complete i­ mmersion of tissue sections requires 15 mL of solution for each mailer. 3. Place slides into a slide mailer and postfix with 4% paraformaldehyde in PBS for 15 min. 4. Rinse in DEPC-treated PBS three times, 1 min each. 5. Digest sections with Proteinase K at room temperature for 6–12 min (see Note 17). 6. Inactivate the Proteinase K by incubating the sections in 4% paraformaldehyde in PBS for 10 min at room temperature. 7. Rinse in DEPC-treated PBS three times, 1 min each. 8. Treat the sections with 0.2 N HCl for 10 min at room temperature (see Note 18). 9. Rinse in DEPC-treated PBS three times, 1 min each.

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10. Measure 15 mL of 0.1 M Triethanolamine-HCl into a 15 mL tube. Add 37.5 μL of acetic anhydride drop by drop. Mix by inverting and pour into the mailer immediately. 11. Transfer slides into the mailer and incubate sections for 10 min at room temperature. 12. Wash in DEPC-treated PBS three times, 1 min each. 13. Dehydrate sections in increasing concentrations of ethanol (50%, 75%, 95%, 100%, 100%) for 2 min each. Sections are subsequently air dried for 2 min. 14. Add 200  μL of hybridization solution to each 1.5 mL tube. Heat the hybridization solution at 85 °C for 10 min. 15. Add labeled sense or antisense probes (~1 μg/mL) to the preheated hybridization solution and incubate at 85 °C for 3 min (for double-labeling experiment, mix DIG and FITC labeled probes) (see Note 19). 16. Add 200  μL of probe solution onto each slide. Cover the slide with an RNase-free plastic coverslip. Be sure there are no bubbles and that the hybridization solution covers the entire coverslip area. 17. Place the slides into a sealed, humid chamber. We line the bottom of the plastic box with a piece of paper towel soaked with 50% formamide in 5× SSC. Instead of placing the slides directly on the paper towel place a 1.5 mL tube rack up-side down to build a platform for the slides. Cover the box with a tight lid. 18. Hybridize at 60 °C overnight (see Notes 20 and 21). 3.3.3  In Situ Hybridization: Day 2. Posthybridization Washes and Immunological Detection—Non-RNase-­ Free

1. Carefully remove the coverslips by dipping the slides in and out of 5× SSC at 60 °C (see Note 22). 2. Incubate in 2× SSC and 50% formamide at 60 °C for 30 min. 3. Transfer sections to TNE buffer at 37 °C for 10 min. 4. Replace the TNE buffer with TNE plus RNaseA (20 μg/mL) and incubate at 37 °C for 30 min. 5. Wash with TNE for 10 min at 37 °C. 6. Further wash the sections in 2× SSC at 60 °C for 30 min. 7. Proceed with high stringency washes in 0.2× SSC twice for 30 min at 60 °C (see Note 23). 8. Let the sections stay cool at room temperature for 5 min. 9. Wash with Buffer 1 for 5 min at room temperature. 10. Incubate with 1% blocking reagent for 1 h at room temperature. 11. Briefly rinse the sections with Buffer 1 plus 0.1% Tween 20 (B1-T). Put 300 μL of alkaline phosphatase conjugated anti-­ DIG antibody (1:1000 with B1-T) onto each slide. Cover the

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sections with coverslips and place the slides into a humidified chamber (e.g., a flat-bottomed box with a moist paper towel as in Fig. 1, but with the sections coverslipped to reduce the volume of antibody solution). For double-label in situ hybridization, add POD conjugated anti-FLU antibody (1:100) with the anti-DIG antibody. 12. Incubate overnight at 4 °C. 3.3.4  In Situ Hybridization: Day 3. Signal Detection—Non-RNase-­ Free

1. Remove the coverslips by dipping the slides in TBS-T. Wash in TBS-T three times 10 min each at room temperature. 2. Precondition with Buffer 2 (B2) for 10 min. 3. Make HNPP/Fast Red solution by mixing 10 μL of HNPP and 10 μL of Fast Red TR (abcam, Burlingame, CA, USA) into 1 mL of B2. Filter with 0.2 μm Nylon Syringe filter (see Note 24). 4. Put 400 μL of HNPP/Fast Red on each slide. Incubate for 30 min in the dark at room temperature. Check for staining. If necessary, repeat steps 3 and 4. 5. Wash sections and mount with Fluoromount-G for single probe in situ (see Note 25). 6. To visualize the FLU-labeled probe, slides are incubated with POD conjugated anti-FLU (1:100) overnight. 7. Wash in B1-T at room temperature for 5 min, three times. 8. Dilute TSA-biotin with Amplification Diluent (1:50, provided in the kit). Use 100 μL per slide. Cover with a coverslip size parafilm. Incubate for 10 min at room temperature. 9. Wash with B1-T for 5 min, three times. 10. Apply 300 μL of Alexa488 conjugated Streptavidin (1:300) to each slide and incubate at room temperature in the dark. 11. Wash in water for 10 min and mount with Fluoromount G. Sections can be counterstained with Hoechst 33342 diluted to 1× working solution prior to mounting.

4  Notes 1. Paraformaldehyde causes skin, eye, and respiratory tract irritation, and is a suspect carcinogen. All handling of paraformaldehyde should be done in the fume hood. Read and follow precautions provided by the manufacturer. 2. DEPC inactivates RNase at 0.1% concentration and is therefore used to treat solutions used for in situ hybridization. It is important to remember that DEPC should not be used to treat Tris buffer. When making RNase-free Tris buffer, dissolve Tris base into DEPC-treated water and filter-sterilize.

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3. It is important to use freshly prepared formaldehyde as fixative made more than a day or 2 before use can increase background fluorescence. Do not over fix. Tissues left in fixative for more than 24 h can become so cross-linked that in situ hybridization will not work, and over-fixation can also adversely affect epitopes. Some antibodies will not work on formaldehyde-­ fixed tissues. It may be necessary to experiment with different fixatives, like methanol, ethanol or acetone-based fixatives. 4. The sucrose will prevent tissue disruption from the expansion of water in tissues as it freezes. Tissues that have taken up sufficient sucrose to withstand freezing will sink into the solution over time. Smaller tissues will sink sooner and can be processed the same day. Larger tissues will take more time, but most will sink after 24 h. Air bubbles trapped in the tissues may prevent them from sinking even when they have taken up the sucrose, so avoid introducing air bubbles into the specimens. 5. It is important to orient the material properly before it is frozen. Chose traditional orientations that correspond to orientations found in atlases to help interpret your results. 6. Tissues frozen in embedding medium can typically be stored in zip-lock storage bags at −20 °C for several weeks or at −70 °C for several months. Tissues can usually be stored for longer periods if the sections will be used for immunohistochemistry than if they are to be used for in situ hybridization. 7. Wear gloves when handling slides that will be used for in situ hybridization. It is easier to cut and collect thicker sections, but try to collect sections in the 12–16 μm range. The ideal temperature for cutting sections through chicken brain is usually colder than the ideal temperature for cutting murine tissues; precise block, knife, and chamber tissues will vary from cryostat to cryostat and require practice before working with precious specimens. 8. Store the slides in a clean box and wrap the edges of the box with Parafilm© and place it in a zip-lock bag before refreezing. It is always best to work with fresh slides, but not always practical. Slides kept in the freezer for several years are often suitable for immunohistochemistry, but slides used for in situ hybridization should be used within a few weeks. 9. The BSA-based blocking solution is easy to make and use and works well for most antibodies, but with some antibodies it may be necessary to use a fetal calf serum or condensed milk-­ based blocking solution. 10. The moisture left behind after drying around the section will help keep the primary antibody from spreading away from the section during the antibody incubation step. There is an

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increased chance of auto-fluorescence if the section dries out during this step. To avoid this, work with small batches of slides. 11. The wet mounting medium (PBS:glycerol) is easy to use, but care must be taken not to use too much or the medium may spread onto the microscope stage and/or objectives. Dab at the edge of the coverslip after mounting to remove excess medium and let the coverslipped slides air dry for several hours before use. Another advantage of this medium is that it is easy to “float off” the coverslip after photography, allowing the section to be counterstained or stained with another primary and secondary antibody (double label immunohistochemistry). 12. The most important factor for probe selection is its sequence specificity to the target gene. When choosing cDNA fragments as template, perform a blast search to find out whether or not they have homology to other transcripts. The optimal probe size is ~500 bp. If the probe is too long, it can inhibit tissue penetration. 13. We often use the vector pCRII-TOPO (Fig. 3). In this vector, T7 and Sp6 polymerase binding sites are located on different sides of the cloned cDNA. When selecting restriction enzymes, it is preferable to use enzymes that result in either a 5′ overhang or a blunt end DNA template. We used Spe I to linearize the template for the antisense probe and EcoR V for the sense probe. Selected restriction enzymes will cut on either end of the cDNA respectively to allow the RNA polymerase to transcribe through the cDNA, but relatively little of the plasmid sequences. 14. When performing in situ hybridization, it is important to incorporate controls. The sense probe often serves as a negative control for nonspecific hybridization signals. Under high stringency condition, the sense probe should give no in situ signal while the antisense probe hopefully hybridizes specifically to its target. Others suggest to evaluate tissue transcript quality by using labeled poly T probes [9]. 15. The yield of the labeled RNA probe is usually 10–15 μg. To evaluate the quantity of the probe, measure the concentration using a nanodrop or conventional spectrophotometer. To determine the quality and the size of the RNA probe, run 0.5–1 μg of the probe on a 2% agarose formaldehyde gel. To assemble a 2% agarose formaldehyde gel, combine 2 g of agarose and 73.3 mL of DEPC-treated ddH2O, heat to boil; in the fumehood add 16.7 mL formalin (37% formaldehyde), 10 mL of 10× MOPS (200 mM MOPS, pH 7.0, 20 mM sodium acetate, 10 mM EDTA, pH 8.0); mix and pour into

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the tray to set. The probes are diluted with 1× MOPS, 20% formaldehyde and 50% formamide; heat denature at 70 °C for 10 min before loading onto the gel. The gel is run in 1× MOPS. 16. RNases are long lived and difficult to inactivate. Small amount of contamination can destroy the transcripts in the tissue section, so it is important to prevent RNase contamination. On day 1 of in situ hybridization, all solutions should be RNase-­ free. Use DEPC-treated water to make all solutions. Wear gloves during day 1. It is important to make sure that the slide mailers for prehybridization steps are brand new and free of RNase. 17. Proteinase K treatment is used to allow better access of the probe to its target transcripts in tissue sections. Excessive digestion will result in damage to tissue morphology. The concentration, duration, and the temperature of the treatment should be determined empirically. Proteinase K is commonly used at 10 μg/mL. Dependent upon the tissue type, digestion can be done between 25 and 37 °C for 5–15 min. A good starting point is 10 min at room temperature. When treating embryonic tissue, it is suggested to start with lower concentration of proteinase K (2 μg/mL) or a shorter digestion time. 18. The function of HCl is not entirely known. It is believed that HCl extracts protein and hydrolyzes the target sequence. The treatment may allow better permeability and also appears to reduce background. 19. Dependent upon the abundance of the target transcript, the optimal concentration of the labeled probe should be determined empirically. A good starting range is 0.5–1 μg/mL. 20. The theoretical melting temperature (Tm), which is the temperature at which 50% of the probe is dissociated from the target, is determined by a number of factors including monovalent cation concentration, the presence of formamide, probe length, and GC content. Tm can be calculated as follows: Tm = 79.8 + 18.8× log [Na] + 0.58 × GC% + 11.8 × [G C%]2 − 0.35 × % formamide-820/length [10]. Hybridization is often carried out at 20 °C below the Tm. 21. The Tm on tissue (tissue Tm) is different from the theoretical Tm. While theoretical Tm is determined by DNA or RNA behavior in solution, tissue Tm is determined by chromogenic detection of a specific hybridization signal. In general, tissue Tm is around 10 °C higher than the theoretical Tm. It is important to keep this mind when designing hybridization and washing conditions [11–14]. 22. During posthybridization washes and all subsequent steps, it is not necessary to maintain an RNase-free environment. In gen-

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eral, RNases cleave single-stranded but not double-stranded RNAs. After hybridization, probes are hybridized with their transcript targets and are resistant to degradation by RNase contamination. 23. The stringency of the washes needs to be determined empirically for each probe. Washing stringency is determined by both the salt concentration and the washing temperature. For RNA probes larger than 300 bp, washing temperatures are generally between 55–65 °C and the salt concentrations vary between 1× SSC to 0.1× SSC. As discussed in Note 21, tissue Tm is generally 10–1560 °C higher than theoretical Tm. In this experiment, washing with 0.2× SSC at 60 °C yields the most specific in situ signal for TNC. 24. Fast Red is an AP substrate-chromagen. The reaction results in a red precipitate where hybridization has occurred. If the tissue has endogenous AP activity, it will yield a false positive signal. 25. Mounting should be done with a quick wash with water and using an aqueous mount. Do not dehydrate with xylene as the treatment will result in the formation of crystals. References 1. Coons AH, Creech HJ Jr, Jones RN, Berliner E (1942) The demonstration of pneumococcal antigen in tissues by the use of fluorescent antibody. J Immunol 45:159–170 2. Beutner EH, Witebsky E (1963) Studies on organ specificity. XV. Immunohistologic evaluation of reactions produced by thyroid autoantibodies. J Immunol 91:204–209 3. Akam ME (1983) The location of ultrabithorax transcripts in Drosophila tissue sections. EMBO J 2:2075–2084 4. Hafen E, Levine M, Garber RL, Gehring WJ (1983) An improved in situ hybridization method for the detection of cellular RNAs in Drosophila tissue sections and its application for localizing transcripts of the homeotic Antennapedia gene complex. EMBO J 2:617–623 5. Garner CC, Tucker RP, Matus A (1988) Selective localization of messenger RNA for cytoskeletal protein MAP2 in dendrites. Nature 336:674–677 6. Wilkinson DG (ed) (1998) In situ hybridization: a practical approach. Oxford University Press, Oxford, p 224 7. Zeller R, Rogers M, Harami AG, Carrasceo AS (2001) In situ hybridization to cellular

RNA. Curr Protoc Mol Biol Chapter 14:Unit 14.3 8. Ishii T, Hirota J, Mombaerts P (2003) Combinatorial coexpression of neural and immune multigene families in mouse vomeronasal sensory neurons. Curr Biol 13:394–400 9. Iezzoni JC, Kang JH, Bucana CD, Reed JA, Brigati DJ (1993) Rapid colorimetric detection of epidermal growth factor receptor mRNA by in situ hybridization. J Clin Lab Anal 7:247–251 10. Bodkin DK, Knudson DL (1985) Assessment of sequence relatedness of double-stranded RNA genes by RNA-RNA blot hybridization. J Virol Methods 10:45–52 11. Evans MF, Aliesky HA, Cooper K (2003) Opimization of biotinyl-tyramide-based in situ hybridization for sensitive background-­ free applications on formalin-fixed, paraffin-­ embedded tissue specimens. BMC Clin Pathol 3:2 12. Herrington CS, Graham AK, Flannery DM, Burns J, McGee JO (1990) Discrimination of closely homologous HPV types by nonisotopic in situ hybridization: definition and derivation of tissue melting temperatures. Histochem J 22:545–554

Immunohistochemistry and In Situ Hybridization 13. Herrington CS, Anderson SM, Graham AK, McGee JO (1993) The discrimination of high-­ risk HPV types by in situ hybridization and the polymerase chain reaction. Histochem J 25: 191–198

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14. Herrington CS, McGee JO (1994) Discrimination of closely homologous human genomic and viral sequences in cells and tissues: further characterization of Tmt. Histochem J 26:545–552

Chapter 25 Gene Silencing in Chicken Brain Development Georgia Tsapara, Irwin Andermatt, and Esther T. Stoeckli Abstract Despite the development of brain organoids and neural cultures derived from iPSCs (induced pluripotent stem cells), brain development can only be studied in an animal. The mouse is the most commonly used vertebrate model for the analysis of gene function because of the well-established genetic tools that are available for loss-of-function studies. However, studies of gene function during development can be problematic in mammals. Many genes are active during different stages of development. Absence of gene function during early development may cause aberrant neurogenesis or even embryonic lethality and thus prevent analysis of later stages of development. To avoid these problems, precise temporal control of gene silencing is required. In contrast to mammals, oviparous animals are accessible for experimental manipulations during embryonic development. The combination of accessibility and RNAi- or Crispr/Cas9-based gene silencing makes the chicken embryo a powerful model for developmental studies. Depending on the time window during which gene silencing is attempted, chicken embryos can be used in ovo or ex ovo in a domed dish for easier access during later stages of development. Both techniques allow for precise temporal control of gene silencing during embryonic development. Key words Neural development, Cerebellum, Chicken embryo, Artificial miRNA, RNA interference, Electroporation, Gene silencing

1  Introduction The chicken embryo has been used to study developmental processes for a long time. The advantage of the chicken as a model is its easy accessibility for experimental manipulation during embryonic development. However, compared to the mouse, the chicken cannot be manipulated with the same powerful genetic tools. Furthermore, size of the adult animals and long generation time are unfavorable features of the chicken with respect to genetic approaches. The development of RNAi-based loss-of-function approaches has overcome these problems, and turned the chicken embryo into a very useful model organism for developmental studies [1–4]. In fact, the precise temporal control and the cell-type specificity of Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_25, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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gene silencing provide unique opportunities for the analysis of gene function during development. In principle, RNAi-based gene silencing can be replaced by Crispr/Cas9-mediated genome editing, but because chickens may not be bred in a lab setting, there is not really an advantage of Crispr/Cas9 compared to RNAi. Tools have been developed however, and were successfully used in chicken embryos [5, 6]. Because many genes involved in neural development are also involved in the development of other tissues, such as the heart, classical genetic knockout strategies can lead to early lethality. One approach to overcome this restriction in the mouse is a conditional, tissue-specific knockout of a gene of interest, provided that suitable Cre-lines are available to excise the floxed target gene in mouse. Conversely, in chicken, organ- or even cell-type specificity is easy to achieve and can be combined with temporal control of gene silencing [4]. We have developed approaches for temporally and spatially controlled gene silencing based on RNAi in chicken embryos [1, 3, 4, 7]. Temporal control can be achieved with both in ovo and ex ovo RNAi by using long dsRNA or plasmid-based miRNA/shRNA for gene silencing [1, 4]. The latter, miRNA-based RNAi comes with the additional advantage of cell-type specificity of gene silencing. Both dsRNA- and miRNA-based RNAi can be used to silence several genes at the same time [8]. For gene analysis during early stages, when embryos can easily be accessed through a window in the eggshell, in ovo RNAi is the method of choice, as it has higher survival rates compared to ex ovo RNAi. To analyze brain function during late developmental stages, embryos can be cultured ex ovo (shell-less) in a dish which enables direct access to the desired brain areas for injection and electroporation [3, 9]. Timing is important! Keep in mind that RNAi does not remove the preexisting protein. Electroporation has to be done before the protein of interest has accumulated. Therefore, it is necessary to carefully analyze the temporal expression of the target gene. We routinely use in situ hybridization to analyze gene expression during embryonic development [10]. Timing of injection and electroporation of dsRNA or miRNA is determined by the temporal expression of the target gene, not by the time point of analysis or the developmental milestones of the part of the nervous system that is analyzed. In general, injection and electroporation are easier at younger stages. However, cell proliferation will dilute the active RNA-induced silencing complex (RISC) loaded with the specific siRNA produced from the injected dsRNA or the miRNA. Thus, gene silencing will be very effective over long time periods in neurons but will be less efficient in cells that keep proliferating after electroporation.

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Once manual skills for handling embryos and efficient injection and electroporation have been acquired, RNAi will provide results on gene function very rapidly. For a first approach, it is often easiest to simply use long dsRNA prepared by in vitro transcription from cDNA fragments [1]. For thorough functional analysis of a target gene, miRNA-based plasmids can be used in a cell type-specific manner [4]. Since these miRNAs are coupled to a fluorescent protein, efficiency of electroporation is directly visualized [4]. These vectors can be used in combination to knock-down several genes with different tissue-specific promoters or together with constructs that are resistant to the miRNA to perform rescue experiments [8]. Examples demonstrating the power of precise temporal control of gene silencing are provided by studies that identify a role of the morphogens Shh and Wnts in axon guidance [11–14]. Morphogens are required for cell differentiation and patterning of the nervous system during early stages of development [15–18]. Thus, precise temporal control of gene silencing was key to our finding of a direct and an indirect effect of Shh signaling as well as Wnt signaling on post-crossing commissural axon guidance [11– 14]. Loss of Shh or Wnt function during the morphogenesis phase of neural development would have prevented these findings as cell types in the neural tube would not have been induced properly. Here, we describe how to use chicken embryos in ovo or ex ovo for electroporation of miRNA-constructs (described in [4]) or long dsRNA [1, 3] to analyze neural development. In ovo RNAi is well-suited for manipulation of embryos at young stages and limited to about the fourth day of embryonic development. After these stages the brain of the embryo is no longer easily accessible. For manipulations at older stages, chicken embryos can be transferred from the egg to a plastic dish enabling injection and electroporation of diverse brain regions at late stages [3, 9]. To study neural crest derivatives, injection and electroporation have to be carried out in the first 2.5 days of development. Injections into the eye are easier in the first 3–4 days, as the poor elasticity of the sclera makes good injections without leakiness more difficult at later stages. The cerebellum, for instance, starts to emerge very late, only after about one week of embryonic development. At this stage, the head needs to be fixed for injection in a more upright position and the blood vessels at the back of the head can be used as landmarks to guide the injection needle [3]. We include two protocols in this chapter, one for injections and electroporation of the neural tube within the first 3–3.5 days of development, and the second one for electroporation of the cerebellum at late stages (HH34-36; Hamburger and Hamilton stages 34–36 [19]). Furthermore, we describe an alternative method to study genes in brain development using electroporation of slices.

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2  Materials 2.1  Windowing the Eggs

1. Fertilized eggs from a local hatchery. 2. Incubator set at 38.5 °C and at least 45% humidity. We use two incubators, one to incubate eggs before they are windowed, and a second one for incubation during the experiment (e.g. Heraeus/Kendro Model B12, Kendro Laboratory Products, Germany, or Juppiter 576 Setter+Hatcher; FIEM, Italy). 3. Facial tissues. 4. 70% ethanol. 5. Paraffin wax (Paraplast tissue embedding medium). 6. Heating plate set at 80 °C to melt paraffin. 7. Paint brush. 8. Scalpel. 9. Scotch tape or coverslips (24 × 24 mm). 10. Soldering iron (when eggs are closed with coverslips, alternatively Scotch tape can be used to seal the window). 11. Sterile syringe with 18 G needle. 12. Fine scissors (e.g. Fine Science Tools, 14090-09).

2.2  Ex Ovo Culture

1. Fertilized eggs from a local hatchery. 2. Domed dish with a diameter of 80 mm and a depth of 40 mm. (These dishes are produced for the food industry from oriented polystyrene (OPS; Bellaplast, Altstaetten, Switzerland).) 3. Lid for domed dish (we use the lid of a 10 cm Ø petri dish). 4. Incubator set at 38.5 °C and at least 45% humidity. We use two incubators, one to incubate eggs, and a second one for incubation of embryos in the dishes (e.g. Heraeus/Kendro Model B12, Kendro Laboratory Products, Germany, for dishes; Juppiter 576 Setter+Hatcher; FIEM, Italy, for eggs). 5. Facial tissues. 6. 70% ethanol.

2.3  Electroporation

1. Borosilicate glass capillaries (outer Ø/inner Ø: 1.2 mm/0.68 mm; World Precision Instruments, 1B120F-4). 2. Glass needle puller (Narishige, PC-10). 3. Square wave electroporator (BTX ECM 830). 4. Spring scissors (Fine Science Tools, 15003-08). 5. Dumont #5 forceps (Fine Science Tools, 11252-20). 6. For in ovo electroporation: Platinum electrodes (4 mm length, 4 mm distance between cathode and anode).

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7. For ex ovo electroporation: Platelet electrode of 7 mm diameter (Tweezertrodes Model #520, BTX Instrument Division, Harvard Apparatus, Holliston, MA, USA, also see http:// www.btxonline.com/tweezertrodes/). 8. For slice electroporation: electroporation chamber and electrodes (7 × 7 mm; Sonidel, CUY701P7E). 9. Spritz bottle filled with ddH2O. 10. Polyethylene tubing (Ø 1.24 mm). 11. 0.2-μm filter (Sarstedt, Switzerland). 12. Trypan blue solution, 0.4% (Invitrogen). 13. 20× phosphate buffered saline (PBS). 14. Sterile PBS. 15. Spatula bent to a hook (ex ovo electroporation). 2.4  Brain Slices

1. Fertilized eggs from a local hatchery. 2. Incubator set at 38.5 °C and at least 45% humidity. We use two incubators, one to incubate eggs before they are windowed, and a second one for incubation during the experiment (e.g. Heraeus/Kendro Model B12, Kendro Laboratory Products, Germany, or Juppiter 576 Setter+Hatcher; FIEM, Italy). 3. Cell culture incubator (10% CO2, 37 °C). 4. Facial tissues. 5. 70% ethanol. 6. Tissue chopper and blades (McIlwain Tissue Chopper). 7. 35 mm tissue culture dishes (Corning). 8. 6-well plates (Nunclon Delta Si, ThermoScientific). 9. MillicellR cell culture inserts with 0.4 μm pore size, 30 mm diameter (Merck, PIC03050). 10. Spatulae straight and round. 11. Two fine brushes. 12. Two spring scissors (Fine Science Tools, 15003.08). 13. Two forceps Dumont #5 (Fine Science Tools, 11252-30). 14. Two forceps Dumont #5 (Fine Science Tools, 11254-20). 15. Fine scissors (e.g. Fine Science Tools, 14090-09). 16. Heavy duty scissors to decapitate old embryos. 17. DMEM with Glutamax (Gibco, 31966). 18. DMEM/Glutamax with 10% (for older stages: HH40) or 25% horse serum (for younger stages: HH38). 19. Prepare 3% agarose gels (low gelling agarose, for instance FMC 50103).

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3  Methods 3.1  Windowing Eggs

To obtain access to the developing embryo a window is cut into the eggshell (Fig. 1). Independent of the time of electroporation, eggs are windowed at the second or third day of incubation (E2 and E3, respectively; see Note 1). 1. Incubate eggs at 38.5 °C and 45% humidity (see Notes 2–4). 2. Let embryos develop until they have reached the desired stage for the experimental manipulation. Staging of the embryo is done according to Hamburger and Hamilton [19] (see Note 5). 3. About 20 min before windowing, place the egg on its long side to allow the embryo to reposition on top of the egg yolk. Maintain the orientation of the egg throughout the following steps. 4. Wipe the eggshell using facial tissue and 70% ethanol to avoid contamination (see Note 6).

Fig. 1 Egg windowing. (a) After the egg is cleaned with 70% ethanol, a strip of sticky tape is put on the top of the egg and two holes are drilled into the edge of the presumptive window and into the blunt end of the egg (arrowheads). (b) The needle is inserted at an angle greater than 45° in order not to damage the egg yolk. Removal of about 3 mL of albumen will detach the embryo and the blood vessels from the eggshell. (c) The hole at the blunt end is sealed (arrowhead) and a window is cut into the shell starting at the previously drilled hole. (d) With a brush melted paraffin is applied to the edges of the window and immediately covered with a coverslip (open arrowheads)

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5. Place a strip of Scotch tape to cover the area of the planned window. This will prevent pieces of eggshell from falling onto the embryo when cutting the window. 6. Use a scalpel to drill a hole into the corner of the intended window and into the blunt end of the egg (see Fig. 1 and Note 7). 7. Remove ~3 mL of albumen by pushing the needle of the syringe at an angle greater than 45° into the hole at the blunt end of the egg. This will avoid damage to the egg yolk (see Note 8). 8. Use a paint brush to seal the hole at the blunt end of the egg with melted paraffin. 9. Cut a window into the eggshell. To avoid damaging the embryo carefully hold scissors horizontally. 10. Seal the window with a coverslip and paraffin. Use a brush to apply melted paraffin to the edges of the window and carefully press a coverslip onto the hot paraffin (alternatively, use Scotch tape to seal the window, see Note 9). As paraffin cools down quickly, carefully pressing a soldering iron on the coverslip will re-melt the paraffin below the glass and lead to proper sealing. 11. Put the windowed egg in the incubator until further use (see Note 3 and 4). 3.2  In Ovo Electroporation

1. Clean the working space with 70% ethanol and autoclave your tools (see Note 6). 2. Prepare capillaries to make injection needles using the glass needle puller (see Note 10). 3. Remove the cover slip or the tape covering the window. To remove the coverslip briefly press the hot soldering iron onto the glass. 4. To get direct access to the embryo, carefully remove the extra-­ embryonic membranes with forceps and spring scissors (Fig. 2). 5. Break off the tip of the previously pulled needles to obtain a diameter of 5–7 μm. Plug the needle into the polyethylene tubing and fill the tip of the needle with your injection mix (see Note 10). 6. Inject the mix containing miRNA-plasmids (see Notes 11 and 12) or long dsRNA (see Notes 11 and 13) into the central canal of the neural tube and control injection volume by mouth (Fig. 2e, f). The maximum volume is reached when the blue solution extends from the ventricle to the tail of the embryo.

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Fig. 2 (a, b) Schematic drawing of in ovo electroporation of the caudal neural tube. (a) Injection of dsRNA or miRNA into the central canal of the spinal cord. (b) Electroporation with anode and cathode positioned parallel to the body axis. (c) Electroporation can be spatially controlled by the position of the electrodes. (d) Successful unilateral transfection of EGFP in the right hemi-segment is visualized on a cryosection of a HH25 embryo. (e) Unilateral in ovo injection and electroporation of an E3 chicken embryo. Using forceps, the extraembryonic membranes are removed to obtain access to the spinal cord. The injection mix is injected into the central canal of the lumbar region of the spinal cord. The injection volume is controlled by mouth. The maximal injection volume is achieved when the blue solution reaches both the ventricle and the tail (arrowheads). For unilateral electroporation the electrodes are placed laterally in parallel to the body axis of the embryo. Make sure not to touch major blood vessels to avoid fatal bleeding. (f) For dorsal or ventral electroporation HH19 or older embryos are easy to handle because their body is more detached from the egg yolk and slightly tilted to the side compared to the younger embryo shown in (e, HH18). Note that the lumbar region is tilted to the side and therefore you see more lateral parts of the spinal cord. Use the forceps to pull the extraembryonic membranes sidewards to position the embryo and stabilize it in the desired position for injection and electroporation with anode and cathode positioned dorsally and ventrally respectively. For dorsal electroporation the polarity would be reversed. Scale bar: 2 mm

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7. Before applying an electric field, add a few drops of PBS to the embryo to prevent overheating and high electrical resistance during subsequent electroporation. 8. Place the electrodes parallel to the anterior-posterior axis of the spinal cord. Do not touch major blood vessels while applying current to avoid bleeding. Use five pulses of 50 ms duration at 18 V for E2 or at 25 V for E3 embryos for electroporation (Fig. 2, see Notes 14–16). 9. Put the resealed egg back into the incubator until the embryo reaches the desired developmental stage for analysis (see Note 4). 3.3  Ex Ovo Culture of Chicken Embryos

1. Incubate fertilized eggs for 2.5 days at 38.5 °C and at least 45% humidity (see Notes 4 and 17). 2. Position the eggs on the long side for 20 min to allow for the embryo to position on top of the egg yolk. 3. Wipe the egg with 70% ethanol and crack it on a sharp edge. Transfer the whole egg content into the domed dish without destroying the egg yolk (Fig. 3). 4. Cover the culture with a lid from a petri dish to minimize evaporation and keep the ex ovo cultures in the incubator until further use.

3.4  Ex Ovo Injection and Electroporation

Here we describe injection and electroporation of the developing cerebellum of embryos at E8 as an example and as previously described [3]. Other areas of the brain can be targeted with the same parameters as a starting point. 1. Use autoclaved tools and clean the workspace with 70% ethanol. 2. Stage the embryos according to Hamburger and Hamilton [19]. 3. Remove the lid and cut a small hole of 3–4 mm diameter into the extraembryonic membranes above the head, where the injection is planned (Fig. 4). 4. Depending on the age of the embryo and the injection site, it may be required to fix the head of the embryo with a spatula bent to a hook by placing it underneath the neck. 5. Use a glass capillary connected to a piece of tubing to inject the mix (see Notes 10–13) into the cerebellum (Fig. 4c). To target all the cerebellar layers insert the glass capillary first into the ventricular zone and apply constant pressure while pulling the needle out (see Note 18). 6. Put a few drops of sterile PBS on the injection site and place the platelet electrodes on either side of the head of the embryo.

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Fig. 3 Ex ovo culture. (a) The embryo is transferred from the egg into a domed dish at E2.5 and kept at 38.5 °C. (b, c) Ex ovo culture of a HH34 embryo (E8) ready for cerebellar injection and electroporation

Fig. 4 Ex ovo electroporation. (a, b) Using spring scissors and forceps a small hole is cut into the extraembryonic membranes in order to get access to the desired injection site. In the example shown here, the injection was planned into the eye. (c) For injections into the developing cerebellum, the blood vessels on the back of the head can be used as landmarks. (d) The tweezer electrodes are placed parallel to the head for electroporation. Importantly, electrodes should not touch the embryo. Scale bar in (d): 2 mm. (e, f) Two examples of successful electroporation of the cerebellum. Coronal sections are shown with EGFP-positive cells (green) indicating successful transfection. Axonin-1/Contactin2 is a marker for parallel fibers, the axons of granule cells (red). V ventricle, PS pial surface

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Avoid touching embryonic tissue with the electrodes to prevent damage. Use six pulses of 40 V and 99 ms duration with 1 s interpulse intervals (Fig. 4d; see Note 16). 7. Cover the dish and put it back to the incubator until the desired stage for analysis is reached. 3.5  Verification of Gene Silencing

First, electroporation efficiency has to be high to get strong downregulation of the gene of interest. The best way to monitor successful targeting of the tissue is to check for the expression of fluorescent proteins (Figs. 2d and 4e, f; see Note 19). A first impression can be obtained by simply looking at the embryo in ovo or ex ovo under a fluorescent stereomicroscope. The embryos expressing the reporter protein can then be further analyzed for verification of gene knockdown. The efficacy of electroporation at late stages in ex ovo cultures has to be verified after dissection [3]. There are several possibilities to analyze gene silencing. Immunohistochemistry as described previously is certainly the best way to verify the downregulation of protein expression [1, 4, 20]. Lysates from spinal cord or brain tissue can be used for analysis by Western blot. Alternatively, if there are no antibodies available, in situ hybridization on cryosections [4, 12] or maybe on whole-­ mount embryos at very young stages [21, 22] can monitor expression levels of the targeted mRNA. Also RT-PCR can serve to demonstrate efficient downregulation, although spatial information is lost [23]. The nontargeted half of the spinal cord or the cerebellum may serve as internal control when compared to the electroporated hemisegment (see Note 20).

3.6  Phenotype Analysis

Depending on the target gene and its function the methods of phenotype analysis will vary. It is not within the scope of this chapter to list methods for immunohistochemistry, axonal tracing, or analysis of cell migration. No matter what kind of analysis you will use, keep in mind that phenotypes can only be assessed by comparison of your experimental group with two control groups (see Note 20).

3.7  Cerebellar Slice Cultures

1. Incubate fertilized eggs for 12 days (until stage HH38 [19]) at 38.5 °C and at least 45% humidity (see Note 5). 2. Use autoclaved tools and clean workspace with 70% ethanol. 3. Decapitate the embryo and transfer head to sterile dish. 4. Use forceps and fine scissors to cut off and remove skin. 5. Remove skull cartilage with forceps and scissors. 6. Carefully remove the meninges with forceps and spring scissors. 7. Remove brain from brainstem and cerebellum.

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8. Put cerebellum with brainstem attached in a dish filled with DMEM. 9. Use binoculars to remove the remaining meninges and remove brainstem from cerebellum. 10. Transfer the cerebellum with spatula and put it on the stage of the chopper. Align it by moving it with spatula. Remove all liquid. 11. Cut sagittal slices of 350 μm thickness (Fig. 5a; see Note 21). 12. Add a few drops of cold DMEM to the slices and transfer them to 35 mm tissue culture dish filled with DMEM (see Note 22). 13. Carefully separate the slices with brushes or forceps. (Do not grab slices with forceps!) 14. Wet the membrane of the Millicell insert with DMEM. 15. Remove liquid and place slices on filter with spatula and forceps (see Note 23). 16. Use forceps to put the insert into a well of the 6-well plate filled with 750 μL DMEM and horse serum (25% for slices taken from HH38 brain; 10% for HH40). 17. Put cultures in cell culture incubator for desired time, depending on the experimental question (see Note 24). 18. Fix slices for staining depending on immunohistochemistry protocol. 3.8  Electroporation of Cerebellar Slices

1. Follow steps 1–15 from Subheading 3.7. 2. Cut membrane to small pieces with one slice per piece. 3. Carefully place the membrane with the cerebellar slice above the electrode (prewetted with PBS) in the electroporation dish (Fig. 5b; see Note 25). 4. Pipet the solution containing your construct of interest or dsRNA on top of the slice. 5. Carefully place a piece of 3% agarose on top of the slice (Fig. 5b; see Note 26). 6. Place anode on top of the agarose slab and electroporate with 80 V and five pulses of 50 ms duration with 1 s interpulse interval. 7. Put slice in 35 mm culture dish with cold DMEM on ice for 10 min. 8. Separate the slice from the membrane and transfer it to a Millicell insert by following steps 14–17 of Subheading 3.7.

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Fig. 5 Electroporation of cerebellar slices. The cerebellum is dissected from the rest of the brain and cut into sagittal slices as indicated in (a). The slices are separated from each other with fine brushes or forceps, and are placed on Millicell membranes. (b) The membrane with the slice is transferred to the electroporation chamber, the transfection mix is pipetted onto the slice. The electrode is positioned on top of the agarose slab that protects the slice during electroporation. (c) Example of a cerebellar slice, cut at HH40, after 14 days in vitro stained with anti-Calbindin antibodies to visualize Purkinje cells

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4  Notes 1. Eggs have to be windowed no later than the third day of incubation even when the injection and electroporation is made at later stages. After 3 days of development, the blood vessels no longer detach from the eggshell and will therefore be damaged during windowing. 2. Eggs can be stored at 15 °C for up to 1 week before incubation. Longer storage will impair development and viability of the embryos. 3. After windowing, the eggs need to be sealed properly. Loss of humidity from the egg will strongly decrease the viability of the embryo. Furthermore, high humidity in the incubator is crucial (at least 45%). Place a tray of water containing 0.1 g/L of copper sulfate into the incubator. Copper sulfate prevents contamination of the water. 4. We routinely use two incubators, one to incubate eggs before they are windowed or used for ex ovo cultures and another one for windowed eggs or dishes. Avoid opening incubators too many times during an experiment, as temperature fluctuations, especially going up to ~40 °C and more, will have negative effects on embryo development and strongly decreases survival rates. 5. Staging of the embryos at the beginning and at the time of analysis is very important. Careful comparison of the developmental progress between experimental and control embryos that have not been handled can give important information about potential interference of embryo handling and/or injection and electroporation procedure with normal development. 6. To reduce the risk of contamination always clean your work space with 70% ethanol, use autoclaved tools and sterile solutions. Additionally, keep the time during which the egg is unsealed to a minimum. Working in a laminar flow hood is not necessary, however. 7. Drilling a hole into the corner of the intended window is necessary for allowing airflow during removal of albumen, which will result in the detachment of the embryo from the eggshell. 8. Damaging the yolk will result in death of the embryo and therefore has to be avoided both for in ovo and ex ovo development. 9. If the eggs need to be reopened several times, sealing with a coverslip is advantageous since the paraffin can be quickly melted by placing a soldering iron on the coverslip. No matter

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whether you seal the egg with Scotch tape or a coverslip, make sure the window is properly closed, because dehydration will dramatically lower survival rate. 10. The diameter of the injection needle should be kept at 10–15 μm to minimize tissue damage and to prevent leakage of the injection mix upon retraction of the needle. It is important to have no leakage in order to get efficient and reproducible gene silencing. 11. Make sure that salt concentrations and pH of the injection mix are in the physiological range. Plasmids used in the mix should be purified carefully by making sure that there is no alcohol remaining from previous precipitation steps. Tris buffers should be avoided as they tend to cause unspecific, toxic effects. 12. DNA injection mix: RNAi plasmid at a concentration that must be determined by the user and Trypan blue diluted in PBS (for detailed designing of RNAi plasmids and concentration of ingredients of the injection mix, see [4]). We design artificial miRNAs using Genscript’s siRNA Target Finder http://www.genscript.com/siRNA_target_finder.html. 13. dsRNA injection mix: sterile PBS containing dsRNA derived from the gene of interest (200–400 ng/μL), EGFP reporter plasmid (20 ng/μL) and 0.04% (vol/vol) Trypan Blue (for detailed description of dsRNA synthesis and troubleshooting advice see [1, 9]). Long dsRNA is easily produced by in vitro transcription from a cDNA template. Expressed sequence tags (ESTs) are available from Ark Genomics or Source BioScience LifeSciences. 14. For targeting dorsal or ventral cell types electrodes can be positioned accordingly along the body of the embryo (see Fig. 2c). Make sure not to touch any major blood vessels during electroporation because this will lead to fatal bleeding. Keep the electrodes away from the heart. 15. For in ovo spinal cord electroporation we use custom-made wire electrodes. There are also commercially available ones from BTX, Harvard Apparatus (http://www.btxonline.com/ genetrodes/). 16. Make sure to clean the electrodes from denatured proteins after each electroporation to maintain a proper electric field between the electrodes for the following embryos. 17. For best survival rate the eggs should be cracked at E2.5. E2 works also but older stages will not give good survival rates. 18. Since injection depth and volume cannot be seen easily at older stages of development, we recommend using injection of a mix of a dye (DiI, CFSE (carboxyfluorescein succinimidyl

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ester)), or fluorescent dextran beads together with an EGFP reporter plasmid to establish the landmarks that can be used to guide injection. For instance, for the cerebellar anlage, the blood vessels on the back of the head can be used as landmarks [3]. 19. If you have low transfection rates consider the following points:

(a) Make sure the electrodes are positioned close enough to the embryo without actually touching it.



(b) The injection volume should be maximized but kept small enough to avoid tissue damage or leakage. It is absolutely important to prevent leakage of the injection mix before electroporation in order to get reproducible and effective gene silencing.



(c) Take into account that in highly proliferating cells the electroporated plasmid will be diluted strongly over time. In this case, reconsider the time point of injection and electroporation.



(d)  Finally, injection and electroporation require extensive practice to obtain adequate manual skills for an efficient downregulation of the targeted mRNA.

20. Note depending on time of electroporation and position of the electrodes it is not possible to compare the two halves of the embryo, as both sides would be affected by the manipulation. Therefore it is important to have adequate controls. At least two control groups are always required. One group consists of untreated embryos taken out of unwindowed eggs at the time of analysis. The second control group consists of mocktreated embryos. These are handled exactly the same way as the experimental group(s) but without dsRNA derived from the gene of interest. In these embryos either only a reporter plasmid or dsRNA from a gene that is not expressed in the nervous system is injected. The best control would be dsRNA or miRNA-based constructs targeting a family member of the gene of interest. This is of course not always available. The treated control group and the embryos from the untreated group have to be indistinguishable to make sure that the handling of the embryos did not cause any artifacts that may be mistaken as a phenotype caused by silencing the target gene. The comparison between the control-treated and the experimental group will provide the desired result indicating the function of the gene of interest. 21. The blade and the stage of the tissue chopper need to be sterile. Mount the blade and tighten screws to have it aligned with the stage.

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22. Slices have to be kept in cold DMEM until they are cultured. 23. Arrange slices on the insert so that they do not touch the edge or each other. 24. Cultures can be kept for at least 14 days (up to months) but medium has to be changed every 2 days. 25. Use as little PBS as possible for electroporation to avoid movement of the slice during electroporation procedure. 26. Use a new piece of agarose for each slice.

Acknowledgements We thank Evelyn Avilés and Nicole Wilson for their help in Figs. 2 and 4. Work in the laboratory of E.S. is supported by a grant from the Swiss National Science Foundation. References 1. Pekarik V, Bourikas D, Miglino N et al (2003) Screening for gene function in chicken embryo using RNAi and electroporation. Nat Biotechnol 21:93–96. https://doi. org/10.1038/nbt770 2. Chesnutt C, Niswander L (2004) Plasmid-­ based short-hairpin RNA interference in the chicken embryo. Genesis 39:73–78. https:// doi.org/10.1002/gene.20028 3. Baeriswyl T, Stoeckli ET (2008) Axonin-1/ TAG-1 is required for pathfinding of granule cell axons in the developing cerebellum. Neural Dev 3:7. https://doi. org/10.1186/1749-8104-3-7 4. Wilson NH, Stoeckli ET (2011) Cell type specific, traceable gene silencing for functional gene analysis during vertebrate neural development. Nucleic Acids Res 39:e133. https://doi. org/10.1093/nar/gkr628 5. Abu-Bonsrah KD, Zhang D, Newgreen DF (2016) CRISPR/Cas9 Targets Chicken Embryonic Somatic Cells In Vitro and In Vivo and generates Phenotypic Abnormalities. Sci Rep 6:34524. https://doi.org/10.1038/ srep34524 6. Morin V, Véron N, Marcelle C (2017) CRISPR/Cas9 in the Chicken Embryo. Methods Mol Biol 1650:113–123. https:// doi.org/10.1007/978-1-4939-7216-6_7 7. Baeriswyl T, Mauti O, Stoeckli ET (2008) Temporal control of gene silencing by in ovo electroporation. Methods Mol Biol 442:231– 244. https://doi. org/10.1007/978-1-59745-191-8_16

8. Alther TA, Domanitskaya E, Stoeckli ET (2016) Calsyntenin 1-mediated trafficking of axon guidance receptors regulates the switch in axonal responsiveness at a choice point. Development 143:994–1004. https://doi. org/10.1242/dev.127449 9. Luo J, Redies C (2005) Ex ovo electroporation for gene transfer into older chicken embryos. Dev Dyn 233:1470–1477. https://doi. org/10.1002/dvdy.20454 10. Mauti O, Sadhu R, Gemayel J et al (2006) Expression patterns of plexins and neuropilins are consistent with cooperative and separate functions during neural development. BMC Dev Biol 6:32. https://doi. org/10.1186/1471-213X-6-32 11. Bourikas D, Pekarik V, Baeriswyl T et al (2005) Sonic hedgehog guides commissural axons along the longitudinal axis of the spinal cord. Nat Neurosci 8:297–304. https://doi. org/10.1038/nn1396 12. Domanitskaya E, Wacker A, Mauti O et al (2010) Sonic hedgehog guides post-crossing commissural axons both directly and indirectly by regulating Wnt activity. J Neurosci 30:11167–11176. https://doi.org/10.1523/ JNEUROSCI.1488-10.2010 13. Wilson NH, Stoeckli ET (2013) Sonic hedgehog regulates its own receptor on postcrossing commissural axons in a glypican1-dependent manner. Neuron 79:478–491. https://doi. org/10.1016/j.neuron.2013.05.025 14. Avilés EC, Stoeckli ET (2016) Canonical wnt signaling is required for commissural axon

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guidance. Dev Neurobiol 76:190–208. https://doi.org/10.1002/dneu.22307 15. Avilés EC, Wilson NH, Stoeckli ET (2013) Sonic hedgehog and Wnt: Antagonists in morphogenesis but collaborators in axon guidance. Front Cell Neurosci 7:86. https://doi. org/10.3389/fncel.2013.00086 16. Gouti M, Metzis V, Briscoe J (2015) The route to spinal cord cell types: A tale of signals and switches. Trends Genet 31:282–289. https:// doi.org/10.1016/j.tig.2015.03.001 17. Briscoe J, Small S (2015) Morphogen rules: Design principles of gradient-mediated embryo patterning. Development 142:3996–4009. https://doi.org/10.1242/dev.129452 18. Sokol SY (2015) Spatial and temporal aspects of Wnt signaling and planar cell polarity during vertebrate embryonic development. Semin Cell Dev Biol 42:78–85. https://doi. org/10.1016/j.semcdb.2015.05.002 19. Hamburger V, Hamilton HL (1992) A series of normal stages in the development of the chick

embryo. 1951. Dev Dyn 195:231–272. https://doi.org/10.1002/aja.1001950404 20. Rao M, Baraban JH, Rajaii F et al (2004) In vivo comparative study of RNAi methodologies by in ovo electroporation in the chick embryo. Dev Dyn 231:592–600. https://doi. org/10.1002/dvdy.20161 21. Katahira T, Nakamura H (2003) Gene silencing in chick embryos with a vector-based small interfering RNA system. Develop Growth Differ 45:361–367 22. Dai F, Yusuf F, Farjah GH et al (2005) RNAi-­ induced targeted silencing of developmental control genes during chicken embryogenesis. Dev Biol 285:80–90. https://doi. org/10.1016/j.ydbio.2005.06.005 23. Sato F, Nakagawa T, Ito M et al (2004) Application of RNA interference to chicken embryos using small interfering RNA. J Exp Zool A Comp Exp Biol 301:820–827. https:// doi.org/10.1002/jez.a.99

Chapter 26 Transplantation of Neural Tissue: Quail–Chick Chimeras Andrea Streit and Claudio D. Stern Abstract Tissue transplantation is an important approach in developmental neurobiology to determine cell fate, to uncover inductive interactions required for tissue specification and patterning as well as to establish tissue competence and commitment. Combined with state-of-the-art molecular approaches, transplantation assays have been instrumental for the discovery of gene regulatory networks controlling cell fate choices and how such networks change over time. Avian species are among the favorite model systems for these approaches because of their accessibility and relatively large size. Here we describe two culture techniques used to generate quail–chick chimeras at different embryonic stages and methods to distinguish graft and donor tissue. Key words Chick, Brain, Central nervous system, Neural plate, Neural tube, Organizer, Peripheral nervous system, Quail, Transplantation

1  Introduction During development, the entire central nervous system arises from the neural plate, which is induced in the ectoderm by signals from the organizer [1–3]. Shortly thereafter, precursors for the forebrain, midbrain, hindbrain, and spinal cord occupy different, albeit overlapping, territories [4–7]. As the neural plate folds to form the neural tube, anterior-posterior and dorsoventral patterning is established through signals from surrounding tissues, and also through the action of local organizers like the midbrain–hindbrain boundary, the floor plate, and roof plate [8]. Many of these paradigms were originally established by transplantation experiments using various ways to distinguish host and donor tissue. In particular, grafting experiments established fate maps of the neural plate and brain [4, 5, 9–12], the location and action of organizers [8, 13], and also the time of competence during which tissue can respond to organizer signals and the time when cells become committed to a particular fate [14–16]. Although many of the signaling mechanisms that control these processes have been identified,

Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_26, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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the downstream transcriptional networks largely remain elusive. In this context transplantation experiments turned out to be critical tools to dissect inductive processes over time and to compare tissue interactions and organizer functions. For example, molecular screens allowed the discovery of new organizers in early development [17], while the comparison of different inductive processes revealed an unexpected similarity between the induction of the central and peripheral nervous system and how both diverge [18, 19]. Thus, although transplantation assays are generally considered to be classical embryological approaches, combined with modern molecular techniques like transcriptional and epigenomic profiling they have the power to uncover gene regulatory networks that control cell fate choices. Avian model systems have been very popular for such studies, because the development of their nervous system parallels that of mammalian embryos in many aspects and has been described in great detail. Unlike mammals, however, avian embryos are easily accessible, relatively cheap to obtain, and little specialized equipment is needed for operations and for growing the embryos. Most, if not all transplantation experiments require a reliable system to distinguish host and donor tissue to locate the graft, to follow its progeny or examine cell behavior like axonal projections, and also to establish whether, e.g., changes in gene expression or neuronal morphology occur in the cell autonomously (e.g., within the graft) or are induced in surrounding cells (e.g., in neighboring tissues). Many studies have used transplantation of tissues labeled with fluorescent dyes (e.g. DiI or DiO [20]), infected with retroviral vectors (transplanted into resistant hosts) [21], and more recently tissues from GFP-transgenic chickens [22, 23]. However, one of the most extensively used techniques is cross-species transplantations generating chick–quail chimeras to provide permanent cell tracing [9– 12]. Quail and chick are closely related species; their early development is fairly similar, but they differ very slightly in timing. The chimeras generated by transplantation of neural tissue and neural crest cells develop normally and are even able to hatch. Early experiments to distinguish quail and chick tissue made use of the fact that quail nucleoli are associated with a fair amount of heterochromatin, which is absent in most other species including the chick. Therefore, histological staining for DNA can differentiate quail and chick tissue [24]. More recently, however, quail-specific antibodies have become available, which either recognize all quail cells or quail neurites [25, 26]. These are now frequently used and their detection can be combined with other techniques like in situ hybridization [27]. This chapter focuses on quail–chick transplantation of neural tissue at early neural plate and at later neural tube stages. After embryonic day 3–4, the brain becomes less accessible due to the formation of blood vessels and extraembryonic membranes, while the spinal cord remains accessible. The procedures described are

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used to replace neural tissue from chick with the identical tissue from quail (orthotopic) at the same stage of development (isochronic). However, similar strategies can be used for heterotopic or heterochronic grafts as well as to any other tissue.

2  Materials All procedures described below require two pairs of watchmakers’ forceps (number 5), one pair of coarse forceps, about 15 cm long, one pair of small, fine scissors, with straight blades about 2 cm long, Pasteur pipettes (short form), end lightly flamed to remove sharp edges and rubber teats, container for egg waste and small beakers (50–100 ml). Instruments should be cleaned with lightly soapy water, rinsed in distilled water and washed in 70% ethanol before drying on a tissue. You need a good stereo-microscope with transmitted light base and for in ovo work a cold light source (fiber optics) for illumination from the top. Fertile hens’ or quails’ eggs are incubated in a humidified incubator at 38 °C until they have reached the stage desired; staging of host and donor embryos is performed according to Hamburger and Hamilton [28]. All solutions are diluted from autoclaved stock solutions in distilled water immediately before use; beakers for salines are autoclaved before use. 2.1  Materials for Preparing Chick Hosts for New Culture

Operations on primitive streak to early somite stage (HH3+-8) chick host embryos are performed in modified New culture [29, 30]; at this stage embryos are fragile, difficult to manipulate in ovo and survival rate in ovo is poor. On the other hand, in New culture embryos can only be grown for 24–36 h even in an expert’s hands. In addition to the above materials, this method requires a Pyrex baking dish about 5 cm deep with 2 l capacity, watch glasses about 5–7 cm diameter, rings cut from glass tubing (approx. 27 mm outer diameter, 24 mm inner diameter and 3–4 mm deep; obtained from a local glass blower), 35 mm plastic dishes with lids (bacteriological grade) and a plastic box with lid for incubating culture dishes. Pannett-Compton saline is prepared from two stock solutions, which can be kept at 4 °C if autoclaved; solution A: 121 g NaCl, 15.5 g KCl, 10.42 g CaCl2·2H2O, 12.7 g MgCl2∙6H2O, H2O to 1 l and solution B: 2.365 g Na2HPO4·2H2O, 0.188 g NaH2PO4·2H2O, H2O to 1 l. To prepare working solution just before use, mix (in order) 120 ml A, 2700 ml H2O and 180 ml B. Do not mix concentrated stocks of A and B.

2.2  Materials for Preparing Chick Hosts for In Ovo Operation

After HH8-9, operations are performed in ovo; under perfect conditions embryos can be grown for a long time, even until hatching. Collect the following materials in addition to those listed at the beginning of Subheading 3. A scalpel with No. 3 handle and

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No. 11 blades, plasticine (Playdo) or foam (from packaging) to make a ring for resting eggs on their side, PVC tape to seal the eggs, 5 ml syringe with 21 G needle (for removing albumen), 1 ml syringe with 27 G (or finer) needle (for ink injection), 1 ml syringe with 21 G needle (for antibiotics), paper tissues, Indian ink (Pelikan Fount India or Windsor and Newton; diluted 1:10 in saline), silicone grease in a 10 ml syringe (no needle). 10× stock Tyrode’s saline (80 g NaCl, 2 g KCl, 0.5 g NaH2PO4∙2H2O and 10 g glucose in 1 l H2O) is prepared in advance, autoclaved and kept at 4 °C after opening. Dilute this to 1× working solution with autoclaved water (about 100 ml are needed). 100× penicillin/streptomycin solution (Sigma A9909) and 70% ethanol are also required. 2.3  Materials for Preparing Quail Donors

The following materials are required to harvest quail embryos: one glass Petri dish (10 or 15 cm diameter, depending on the number of embryos to be collected), one spoon-spatula for collecting embryos, a glass Pasteur pipette cut at the shoulder and fire polished (for embryo transfer), rubber teats, 500 ml Tyrode’s saline (see above), dissecting microscope with transmitted light base.

2.4  Materials for Grafting

To dissect quail and chick tissue for grafting the following materials are required: 35 mm Sylgard-coated dish for dissecting (this should never come into contact with fixative), entomological insect pins (A1; steel) for pinning out embryos on the Sylgard dish, insect pins mounted on Pasteur pipettes or Tungsten needles (Goodfellow; 100  μm diameter, mounted on aluminum holders or glass rods using sealing wax; sharpen by repeated exposure of the tip to a very hot Bunsen flame), 30 G needles mounted on 1 ml syringes, P20 Gilson pipette, and yellow tips. In addition, for in ovo transplantation in older embryos the following materials are needed: microknife (e.g., micro-feather microsurgery knives for eye surgery 15E blade angle), aspirator tube (Sigma A5177), 50 μl borsilicate glass capillaries (for trypsin injection) pulled to fine injection needles using an electrode puller, tips broken off (puller settings need to be determined; needles should be fine enough to avoid fluid uptake by capillary forces, but large enough to deliver small amounts of trypsin by air pressure); about 50 ml 0.12% trypsin (Difco) in Tyrode’s saline, 5% serum (any species) in Tyrode’s saline for stopping the Trypsin.

2.5  Materials for Fixing Embryos and Analyzing Results

Embryos are fixed several hours or days after transplantation; they can be analyzed by in situ hybridization or immunostaining to label specific tissues or cell types, followed by labeling with quail specific antibodies to detect the graft, by in situ hybridization using chick and quail-specific probes or by histological sectioning followed by Feulgen and Rossenbeck staining to reveal nucleoli [24]. Depending on the analysis, different fixatives are used.

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Fixing requires a Petri dish for collecting embryos (for in ovo), 3.5 or 10 mm Sylgard dish for pinning out embryos, insect pins (see above), 7 ml glass vials and phosphate buffered saline (PBS). For whole mount in situ hybridization embryos are fixed in 4% PFA, 1 mM EGTA in PBS, 0.1% Tween-20 and stored in methanol. For Feulgen staining and in situ hybridization on sections [31], embryos are fixed in Zenker’s (50 g HgCL2, 25 g K2Cr2O7, 10 g of Na2SO4 ×10 H2O in 1 l distilled water; before use, add 5 ml glacial acetic acid to 100 ml of the solution) or Carnoy’s fixative (50% ethanol, 11.1% formaldehyde, 10% glacial acetic acid), respectively. Further analysis requires materials for in situ hybridization; see detailed descriptions of these procedures in [27, 31]. The most important point for further analysis is the detection of quail tissue. While traditionally Feulgen staining has been used to reveal the difference between chick and quail nucleoli [25, 32, 33], now quail-specific antibodies are the favorite method. The monoclonal mouse antibody QCPN (Developmental Studies Hybridoma Bank) labels all quail cells, while QN is specific for neurites [26]. We generally perform QCPN staining after whole mount in situ hybridization; this procedure requires: glass vials, Pasteur pipettes and rubber teats, a rocking platform, PBS, blocking buffer (1% goat serum, 0.5% Triton X100 in PBS), anti-mouse IgG-HRP coupled (Jackson), 100 mM Tris pH 7.4, 50 mg/ml 3,3′-Diaminobenzidine (DAB) in 100 mM Tris pH 7.4, 0.3% H2O2 in 100 mM Tris pH 7.4. It is also possible to perform in situ hybridization with a probe directed against sequences that differ between chick and quail (most likely the 3′ UTR of a specific gene of interest) to distinguish transcripts produced by cells of the graft and donor [34].

3  Methods 3.1  Preparing Quail Donor Embryos for Grafting

Collect the materials listed under Subheading 2.3; to dissect the donor tissue, quail embryos are first removed from the egg and cleaned using the following steps. 1. Remove quail eggs from incubator. Using fine scissors, gently tap the shell near the blunt end of the egg to penetrate the shell. Use the tip of the scissors to cut off a small cap of shell; avoid damaging the yolk. 2. Pour thin egg white into waste; use the scissors to help and cut through the rather thick albumen if required (see Note 1). 3. Once most albumen is removed, turn the yolk by stroking it very gently with the sides of the scissors to make the embryo become visible on top of the yolk.

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4. Using the scissors make four cuts into the vitelline membrane around the embryo; make sure that all the cuts meet (see Note 2). 5. Using the spoon, pick up the square of embryo and membrane including a little yolk; try to collect as little yolk as possible. 6. Transfer the embryo including yolk and membrane into the large Petri dish containing Tyrode’s saline under a dissecting microscope by sliding it carefully off the spoon. Using fine forceps, turn the square of yolk/membrane/embryo so you can see the embryo. 7. Once all donor embryos have been collected in the Petri dish, separate the embryos from adhering yolk. Work at low magnification; use two pairs of forceps to pick up a corner of the vitelline membrane with one and slowly but steadily fold it back, steadying the yolk with the other. Make sure that the membrane and embryo remain submerged in saline. The embryo will remain attached to the membrane. If not, peel off the membrane completely and then use the forceps gently to remove the embryo from the underlying yolk. 8. Using the wide-mouth Pasteur pipette pick up the embryo, with (better) or without membrane, and transfer to a 10 cm dish with clean saline. To clean the embryo use a fire-polished Pasteur pipette and gently blow saline over it; this will remove yolk particles. The embryos are now ready for dissection and grafting and can be kept for 1–2 h before proceeding further. 3.2  Preparing Chick Hosts for New Culture

At primitive streak and neural plate stages, operations in avian embryos are most easily performed in New culture. The method described below is based on New’s original technique [30] modified by Stern and Ireland [29]. This modified culture method uses rings cut from glass tubes, instead of rings bent from glass rods and 35 mm plastic dishes instead of glass watch glasses resting inside a large glass Petri dish. The rings cut from tubing generate a slightly rough surface that grips the vitelline membrane and therefore allows easy transfer of the culture into the plastic dish. Collect materials and solutions described in Subheading 2.1; to set up the cultures proceed as follows (Fig. 1): 1. Remove eggs from the incubator. 2. Fill the large Pyrex dish with about 1.5 l of Pannett-Compton saline; the volume should be large enough that eggs yolks are submerged completely. 3. To open an egg, tap its blunt egg with the coarse forceps and carefully remove pieces of the shell. Discard the thick albumen into the waste bucket, assisted with the coarse forceps. Collect the thin albumen in a small beaker (see Note 3).

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Fig. 1 Setting up New culture. (a) Setup. (b) Instruments. (c) Opening eggs. (d) Removing albumen. (e) Yolks in Pyrex dish. (f) Cutting the vitelline membrane at the equator. (g) Removing membrane and embryo from the yolk. (h) Membrane with embryo facing upwards on watch glass. (i) Placing glass ring on the membrane. (j) Culture assembled on ring and watch glass. (k) Removing assembly form Pyrex dish. (l) Cleaning the culture. (m) Primitive streak stage embryo on watch glass. (n) Embryo after removal of a piece of neural plate. (o) Setting up the culture in a Petri dish with albumen. (p) Finished New culture

4. Carefully tip the yolk into the Pyrex dish containing saline, taking care not to damage the membrane on the edges of the shell. Carefully turn the yolk with the side of the coarse forceps so the embryo is facing upwards. Now place a watch glass and a glass ring into the dish.

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5. Cut the vitelline membrane enveloping the yolk just below the equator using small scissors; you can use one pair of forceps to push the yolk around gently, while continuing to cut all the way around its circumference (see Note 4). 6. Using both pairs of fine forceps peel the vitelline membrane with the embryo attached slowly but very steadily off the yolk. Use one pair of forceps to pull the edge of the membrane slightly upwards (about 25–30° angle from the yolk surface) and the other to hold the yolk down. Do not pull tangentially along the yolk: this may detach the embryo from the membrane. Do not stop during this process. The embryo should come off with the membrane (see Note 5). 7. Turn the vitelline membrane with the inner face containing the embryo pointing upwards and slide it, preserving its orientation, onto the watch glass. Place the ring over it so that membrane protrudes around the ring and the embryo sits in its center. Remove the watchglass, ring, and embryo from the dish; tilt the assembly gently to pour off some saline while steadying the ring with one finger (see Note 6). 8. Dry the bottom of the watch glass on some tissue. Using fine forceps, carefully wrap the lose edges of the vitelline membrane over the edge of the ring, all the way around its circumference. Pull the membrane slightly so its bottom is smooth and free from wrinkles, but be careful not to pull so tight that it breaks (see Note 7). 9. Using the fire-polished Pasteur pipette rinse the outside of the ring to remove yolk particles. If there is a lot of egg albumen remaining under the membrane, lift the ring gently and use the Pasteur pipette to remove it. Clean the yolk over and around the embryo using clean saline; be careful not to dislodge the embryo form the membrane. Damaged embryos do not grow well or normal. If there is a lot of vitelline membrane inside the ring, trim off the excess with the fine scissors while lifting the edges with fine forceps. At this stage embryos are ready for transplantation and can be kept on the bench for some time; make sure they remain well submerged under saline and there is sufficient saline on the watch glass. If keeping them for a few hours, place them on a wet tissue and cover with a large glass or plastic plate/dish. 10. To finalize the cultures after transplantations work under the microscope; carefully remove any remaining saline, both inside and outside the ring. Drying helps the graft and host tissue to heal faster. During culture the embryo and the inside of the ring must remain dry. 11. Pour some thin albumen (about 2–3 mm thick layer) on the bottom of a 35 mm Petri dish. Using fine forceps, slide the

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ring with vitelline membrane off the watch glass, and transfer it to the dish; lower it onto the albumen making sure that no air is trapped underneath. Press the ring lightly onto the bottom of the dish using two forceps to allow it to adhere. 12. Remove the excess albumen if its level comes close to the edge of the ring. At this point, remove any remaining liquid from inside the ring using a fire-polished Pasteur pipette. The vitelline membrane should be slightly dome-shaped; this will help to drain off any fluid that accumulates during culture (see Note 8). 13. To seal the Petri dish, wet the inside of the lid with a thin film of albumen all around the egde, discard the excess and place onto the bottom part. Press lid down slightly to seal (see Note 9). 14. Place the dish in a plastic box containing a piece of wet tissue, seal the box, and place it into an incubator at 38 °C. 3.3  Preparing Chick Hosts for In Ovo Culture

Later stage embryos are generally grown in ovo, which allows embryos to grow for long periods, even until hatching. Eggs must be incubated lying on their side, so the yolk turns with the embryo facing upwards to make it accessible for manipulation. Collect the materials described in Subheading 2.2; to prepare hosts for in ovo operations use the following procedure (Fig. 2): 1. Remove the eggs from the incubator, place one egg onto the egg rest and clean with 70% ethanol; be careful not to rotate the egg. 2. Hold the 5 ml syringe with 21 G needle nearly vertical and insert the needle into the blunt end of egg until you feel the shell at the bottom. Remove about 1 ml egg albumen and discard. This lowers the embryo, away from the top of the shell. 3. Using the scalpel score a 1 cm × 1 cm square on the top of the shell and lift it up using the blade or a pair of forceps. 4. Moisten the white membrane under the shell with a little Tyrode’s saline and remove it with fine forceps, to the edge of the window. Be careful to avoid damage to the embryo underneath. 5. Add saline to the egg so that the embryo floats up to the level of the window. 6. Take up diluted ink into a 1 ml syringe with a 27 G needle; make sure there are no air bubbles in the syringe. Insert the needle under the vitelline membrane almost parallel to the yolk surface; choose a position away from the embryo proper and point the needle towards and underneath the embryo. Inject about 50–100 μl ink; the amount should be kept as little as

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Fig. 2 In ovo culture. (a) Eggs are incubated on their side and placed on egg rest. Circle labels the blunt end of the egg. (b) Blunt end is used to remove albumen with 5 ml syringe. (c) Inserting syringe. (d) Scoring the window. (e) Windowed egg. (f) Windowed egg with vaseline border surrounding the window. (g) Embryo before ink injection. (h) Ink injection using a 1 ml syringe. (i) Ink injection: embryo is clearly visible. (j) Embryo after ink injection. (k) Removing albumen to lower the embryo. (l) Eggs sealed with tape

possible. Avoid moving the needle after initial penetration; otherwise the hole will become too big and yolk and/or ink will leak out. The embryo should now be clearly visible on a dark background (see Note 10). 7. Line the shell window with a shallow edge of Vaseline by ejecting it from the syringe. This will allow you to cover the embryo with a drop of saline during the operation for moisture and good optics. Fill the chamber with saline until there is a good dome of fluid. Adjust the fiber optic light so the light shines tangentially onto the embryo, which is now ready for manipulation (see Note 11). 8. Once the embryo has received the graft and it is in the correct position, remove the saline very carefully from above the embryo using a Pasteur pipette. Watch under low magnification to ensure the graft does not move. If required, reposition it using a pin mounted on a Pasteur pipette.

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9. Carefully insert the 5 ml syringe with 21 G needle into the original hole in the blunt end of the shell. Carefully remove about 3 ml egg albumen to lower the embryo to its original position. 10. Add 1–2 drops (50–100 μl) of antibiotic solution (see Note 12). 11. Use a tissue with 70% ethanol to wipe the Vaseline off the shell; dry the shell thoroughly with another tissue. 12. Cut a piece of PVC tape about 3–4 cm long, stretch it slightly and allow it to relax again. To seal the egg, place the tape over the window and carefully smoothen any wrinkles without putting too much pressure on the shell. Make sure the edges of the tape are firmly attached to the shell; if not they will roll up and expose the embryo. 13. Incubate the egg in a well-humidified incubator at 38 °C; next day you can turn the egg window side down which helps to keep the embryo moist and improves their development. Incubate for the desired period; generally the 2–4 day survival rate should be 80–90% (see Note 13). 3.4  Grafting Procedure: Quail Neural Plate into Chick Hosts in New Culture

The procedure below describes orthotopic, isochronic neural plate grafts from quail donors into chick hosts at HH3+/4. The same method can be applied for heterotopic and heterochronic grafts or transplantation of other tissues. 1. To prepare the host follow the procedure described under Subheading 3.1 until step 9. 2. Replace one of the eyepieces of the microscope with an eyepiece containing a graticule; a protractor, to measure angles, is particularly useful. 3. Place a host embryo from Subheading 3.1, step 9 (kept on a watch glass) under the microscope and center the graticule on the node. 4. Define the area to be replaced using the graticule coordinates. New culture embryos face ventral side up; to reach the ectoderm lower layers need to be removed. Fold back the endoderm and mesoderm above the area to be replaced by quail tissue using 30 G needles on a 1 ml syringe serving as a holder. Use the sharp side of the needle gently to score both layers on three sides (e.g., anterior, posterior and lateral leaving them attached medially); then using the back of the needle carefully peel away the endoderm and mesoderm overlying the area to be grafted. 5. Use tungsten needles, mounted insect pins or 30 G needles to cut out the region of the neural plate to be replaced by quail

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tissue (see Note 14; Fig. 1n). Set aside the host embryo and turn to the quail donor. 6. Pin out a quail embryo of the same stage, ventral side up, on a Sylgard dish containing Tyrode’s saline; use insect pins through the extraembryonic region to stretch the embryo slightly. 7. Locate the area to be grafted using the graticule, remove the endoderm and mesoderm as described in step 4 and excise the underlying neural plate as described in step 5. 8. Working under low magnification, use a Gilson pipette to pick up the graft in 3–5 μl saline. 9. Move the host embryo back on the stage and, working under low magnification, place the graft close to the target site. 10. Use mounted insect pins to move the graft into the hole cut previously. It is crucial to maintain the apical-basal orientation of the graft; after excision the ectoderm generally contracts basally and the tissue curves slightly. 11. Carefully remove all the liquid outside the ring and most of the fluid inside the ring. 12. Flip back the mesoderm and endoderm to secure the graft in its position and carefully remove all remaining liquid inside the ring. Excess liquid around the grafted area should be removed using pulled capillaries on aspirator tubes (see Note 15). 13. Now finish setting up the cultures by following steps 10–13 in Subheading 3.2. Make sure that the dome of albumen is rather flat; a high dome causes too much tension and the grafts do not integrate properly. Leave embryos on the bench for 30–60 min to let the grafts heal and then proceed to step 14 in Subheading 3.2. 3.5  Grafting Procedures: Quail Neuroectoderm into Chick Host In Ovo

This section describes orthotopic and isochronic neural tube grafts in embryos older than HH9; as with transplantations in younger embryos described above, the same techniques can be used for heterotopic and heterochronic experiments. First prepare the donor embryo and then turn to the host. 1. To prepare the quail donor, pin out the embryo dorsal side up on a Sylgard dish in Tyrode’s saline and place under a dissecting microscope with transmitted light base. 2. Using tungsten needles or a micro-knife, make a longitudinal incision into the ectoderm dorsal to the neural tube on both of its sides. 3. Replace the Tyrode’s with trypsin solution; working at high magnification peel the ectoderm away from the neural tube using the back of a 30 G needle. In the same way, gently scrape any loosely attached cells (neural crest depending on stage)

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off the neural tube and free it from the adjacent tissues (see Note 16). 4. Progressively separate the neural tube from the underlying notochord using a micro-knife to push it from side to side, allowing the Trypsin to penetrate; when completely detached, cut the neural tube transversely at its anterior and posterior ends to free it. 5. Remove the excised neural tube using a using a Gilson pipette set to 3–5 μl and place into a 35 mm Petri dish containing saline with 5% serum (see Note 17). Graft can be kept on ice until use. 6. Prepare the chick host by following steps 1–7 in Subheading 3.3. Using a mounted insect pin make a small hole into the vitelline membrane just over the area to be operated. The hole should be as small as possible. 7. Replace the drop of Tyrode’s saline with trypsin solution and follow steps 2–4 above to excise the same section of the neural tube as in the quail donor. 8. Remove the excised neural tube using a Gilson pipette set to 3–5  μl and replace the trypsin solution with fresh Tyrode’s saline twice. 9. Pick up the graft using a Gilson pipette, rinse in Tyrode’s saline without serum before transferring it to the host. 10. Using a Gilson pipette and working under low magnification, transfer the graft into the saline bubble over the host embryo. 11. Use a mounted insect pin or 30 G needle to place the graft into the hole made by removal of the host neural tube. Preserve anterior-posterior and dorsoventral orientation (see Note 18). 12. Once the transplant is in position, carefully remove the saline using a Pasteur pipette while observing under low magnification. If needed, reposition the graft using a mounted pin. 13. Finish the egg by following steps 8–13 in Subheading 3.3. 3.6  Detecting Quail Tissue

As outlined above, grafted embryos can be analyzed in various ways depending on the question; these include whole mount or section in situ hybridization and tissue- or cell-specific immunohistochemistry. All of these techniques can be combined with the antibody staining using the quail-specific antibody QCPN. We generally perform whole mount in situ hybridization for embryos up to embryonic day 3 and section in situ for older embryos followed by QCPN staining. The protocol below describes the whole mount procedure; for other applications see [22, 23, 31]. For embryos incubated to HH13 or older, it is a good idea to treat them with 6% H2O2 in PBS, 0.1% Tween after fixation or rehydra-

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tion after storing in methanol (see [27]); this reduces background for the in situ hybridization and immunostaining signal. 1. After developing the in situ hybridization color reaction, wash and fix embryos as normal. Remove fixative by washing in PBS three times for 10–30 min depending on the age of the embryo. 2. Block embryos for 1–3 h in blocking buffer at room temperature on a rocking platform. 3. Replace blocking solution with QCPN antibody solution (dilute antibody in blocking buffer; determine concentration for each batch of antibody) and incubate at 4 °C on a rocking platform for two nights. 4. Remove antibody and wash embryos in PBS for 5–7 times 1 h each; for older embryos leave the final wash overnight at 4 °C. 5. Incubate embryos in secondary antibody (generally 1:1000 in blocking buffer, but may need titration) over one or two nights at 4 °C. 6. Wash as in step 4. 7. Wash twice for 15 min in 100 mM Tris pH 7.4 (see Note 19); in the second wash measure the volume (generally 1 ml is sufficient). 8. Add DAB from the stock to vial with embryos to a final concentration of 0.5 mg/ml; incubate for 10–15 min rocking in the dark (see Note 20). 9. Add the appropriate amount of H2O2 from the 0.3% stock to the embryos to make a final concentration of 0.003%. 10. Incubate in the dark until brown color develops normally within 5–10 min; check occasionally using illumination from the top on a white background. 11. Stop reaction by rinsing several times in distilled H2O to remove residual substrate and post-fix embryos in 4% formaldehyde. 12. Embryos can now be cleared, photographed, embedded for paraffin or vibratome sectioning as required.

4  Notes 1. Remove as much albumen as possible; the yolks move less in the next step. 2. Make sure scissors are cleaned after each egg; crusts of egg yolk make the vitelline membrane stick to the scissors and the embryos tend to sink into the yolk. Do not hesitate when making the cuts, work rapidly so the embryos do not move.

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3. Try to remove as much albumen as possible; albumen adhering to the vitelline membrane makes the following steps more difficult. 4. Make sure to cut at or slightly below the equator; otherwise it will be difficult to fit the membrane around the ring in the next step. 5. Occasionally embryos remain attached to the yolk particularly at early primitive streak stages. Keep the membrane because it can be used for other embryos, in case you accidentally punctured a membrane. The embryo can also be retrieved from the yolk, but requires thorough cleaning: use a pair of fine forceps, close them and gently push the edges of the extraembryonic region away from the yolk. Work all the way around the edge of the embryo. Transfer the embryo using a wide-mouthed pipette into a dish with fresh saline, ventral side up gently blow the attached yolk plug off the embryo using a Pasteur pipette. The embryo can now be returned to the membrane; make sure to keep its orientation: embryos do not grow with the ventral side on the membrane. 6. Make sure you do not turn the membrane inside out; embryos do not grow on the outer surface. 7. Be careful not to make any holes into the membrane; this will allow albumen to accumulate inside the ring and prevent the embryo from growing. 8. Be careful not to use too much albumen for grafted embryos; this will increase the tension and prevent healing. 9. Sealing is important to prevent condensation on the lid during incubation. 10. Too much or too high concentration of ink is toxic for the embryos. Recently Pelikan Indian ink has become difficult to obtain; if you are using other brands they need to be tested for toxicity. 11. A dome of liquid considerably improves the optics and also prevents the embryo from drying. The latter is critical as drying out reduces the survival rate. 12. Antibiotics are generally only required for long culture periods. Normally embryos survive well for 2–3 days without this. 13. Low survival rates can be due to a number of factors. The most common problems are dehydration, damage to critical blood vessels and infection. Working in a drop of saline helps to alleviate dehydration. To avoid infection ensure you use clean solutions; instruments can be cleaned periodically while working in distilled H2O and ethanol (make sure it evaporates before using to operate), while tungsten needles are flamed periodically to keep them sharp.

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14. Be careful not to make any holes in the vitelline membrane. 15. Drying the area surrounding the graft greatly improves healing; be careful not to suck up the graft into the capillary. 16. Be patient; the trypsin works almost by itself and it generally sufficient to push the adjacent tissues away using your instruments. Avoid going too deep and cut a hole into the endoderm; this will make ink and yolk leak out. 17. This inactivates trypsin; this is important because the tissue should be exposed to proteolytic enzymes for as short as possible to avoid disintegration. 18. If you encounter difficulties to preserve orientation, mark one end of the neural tube with a small crystal of carmine powder. 19. Make sure the pH is properly adjusted to 7.4; the reaction is pH sensitive. 20. DAB is carcinogenic; make sure to wear appropriate protective clothing (lab coat, gloves) and consult the local health and safety regulations for inactivation and disposal.

Acknowledgments This work is supported by the BBSRC, NIH, and ERC. We thank Anneliese Norris for assistance with New culture photography. References 1. Stern CD (2005) Neural induction: Old problem, new findings, yet more questions. Development 132:2007–2021 2. Waddington CH (1934) Experiments on embryonic induction. Part I: the competence of the extra-embryonic ectoderm. Part II: experiments on coagulated organisers in the chick. Part III: a note on inductions by chick primitive streak transplanted to the rabbit embryo. J Exp Biol 11:211–227 3. Spemann H, Mangold H (1924) Über induktion von embryonalanlagen durch implantation artfremder organisatoren. Arch Mikroskop Anat Entwicklungsmech 100:599–638 4. Cobos I, Shimamura K, Rubenstein JL, Martinez S, Puelles L (2001) Fate map of the avian anterior forebrain at the four-somite stage, based on the analysis of quail-chick chimeras. Dev Biol 239:46–67 5. Fernandez-Garre P, Rodriguez-Gallardo L, Gallego-Diaz V, Alvarez IS, Puelles L (2002)

Fate map of the chicken neural plate at stage 4. Development 129:2807–2822 6. Garcia-Martinez V, Alvarez IS, Schoenwolf GC (1993) Locations of the ectodermal and nonectodermal subdivisions of the epiblast at stages 3 and 4 of avian gastrulation and neurulation. J Exp Zool 267:431–446 7. Eagleson GW, Harris WA (1990) Mapping of the presumptive brain regions in the neural plate of xenopus laevis. J Neurobiol 21:427–440 8. Vieira C, Pombero A, Garcia-Lopez R, Gimeno L, Echevarria D, Martinez S (2010) Molecular mechanisms controlling brain development: an overview of neuroepithelial secondary organizers. Int J Dev Biol 54:7–20 9. Couly G, Le Douarin NM (1988) The fate map of the cephalic neural primordium at the presomitic to the 3-somite stage in the avian embryo. Development 103:101–113

Quail–Chick Chimeras 10. Couly G, Le Douarin NM (1990) Head morphogenesis in embryonic avian chimeras: evidence for a segmental pattern in the ectoderm corresponding to the neuromeres. Development 108:543–558 11. Couly GF, Le Douarin NM (1985) Mapping of the early neural primordium in quail-chick chimeras. I Developmental relationships between placodes, facial ectoderm, and prosencephalon. Dev Biol 110:422–439 12. Couly GF, Le Douarin NM (1987) Mapping of the early neural primordium in quail-chick chimeras. II The prosencephalic neural plate and neural folds: Implications for the genesis of cephalic human congenital abnormalities. Dev Biol 120:198–214 13. Martinez S, Wassef M, Alvarado-Mallart RM (1991) Induction of a mesencephalic phenotype in the 2-day-old chick prosencephalon is preceded by the early expression of the homeobox gene en. Neuron 6:971–981 14. Baker CV, Stark MR, Marcelle C, Bronner-­ Fraser M (1999) Competence, specification and induction of pax-3 in the trigeminal placode. Development 126:147–156 15. Bhattacharyya S, Bronner-Fraser M (2008) Competence, specification and commitment to an olfactory placode fate. Development 135:4165–4177 16. Groves AK, Bronner-Fraser M (2000) Competence, specification and commitment in otic placode induction. Development 127:3489–3499 17. Anderson C, Khan MA, Wong F, Solovieva T, Oliveira NM, Baldock RA, Tickle C, Burt DW, Stern CD (2016) A strategy to discover new organizers identifies a putative heart organizer. Nat Commun 7:12656 18. Trevers KE, Prajapati RS, Hintze M, Stower MJ, Strobl AC, Tambalo M, Ranganathan R, Moncaut N, Khan MAF, Stern CD, Streit A (2017) Neural induction by the node and placode induction by head mesoderm share an initial state resembling neural plate border and es cells. Proc Natl Acad Sci U S A 115:355–360 19. Hintze M, Prajapati RS, Tambalo M, Christophorou NAD, Anwar M, Grocott T, Streit A (2017) Cell interactions, signals and transcriptional hierarchy governing placode progenitor induction. Development 144:2810–2823 20. Guthrie S, Prince V, Lumsden A (1993) Selective dispersal of avian rhombomere cells in orthotopic and heterotopic grafts. Development 118:527–538

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21. Fekete DM, Cepko CL (1993) Retroviral infection coupled with tissue transplantation limits gene transfer in the chicken embryo. Proc Natl Acad Sci U S A 90:2350–2354 22. Barraud P, Seferiadis AA, Tyson LD, Zwart MF, Szabo-Rogers HL, Ruhrberg C, Liu KJ, Baker CV (2010) Neural crest origin of olfactory ensheathing glia. Proc Natl Acad Sci U S A 107:21040–21045 23. Sabado V, Barraud P, Baker CV, Streit A (2012) Specification of gnrh-1 neurons by antagonistic fgf and retinoic acid signaling. Dev Biol 362:254–262 24. Feulgen R, Rossenbeck H (1924) Mikroskopisch-­chemischer nachweis einer nukleinsaeure vom typus der thymonukleinsaeure und die daruf beruhende elecktive faerbung von zellkernen in miroskopischen praeparaten. Hoppe Seylers Z Physiol Chem 135:203–248 25. Teillet MA, Ziller C, Le Douarin NM (2008) Quail-chick chimeras. Methods Mol Biol 461:337–350 26. Tanaka H (1990) Selective motoneuron outgrowth from the cord in the avian embryo. Neurosci Res Suppl 13:S147–S151 27. Streit A, Stern CD (2001) Combined wholemount in situ hybridization and immunohistochemistry in avian embryos. Methods 23:339–344 28. Hamburger V, Hamilton HL (1951) A series of normal stages in the development of the chick embryo. J Morph 88:49–92 29. Stern CD, Ireland GW (1981) An integrated experimental study of endoderm formation in avian embryos. Anat Embryol (Berl) 163:245–263 30. New DAT (1955) A new technique for the cultivation of the chick embryo in vitro. J Embryol Exp Morph 3:326–331 31. Lassiter RN, Dude CM, Reynolds SB, Winters NI, Baker CV, Stark MR (2007) Canonical wnt signaling is required for ophthalmic trigeminal placode cell fate determination and maintenance. Dev Biol 308:392–406 32. Storey KG, Selleck MA, Stern CD (1995) Neural induction and regionalisation by different subpopulations of cells in hensen's node. Development 121:417–428 33. Le Douarin N, Dieterlen-Lievre F, Creuzet S, Teillet MA (2008) Quail-chick transplantations. Methods Cell Biol 87:19–58 34. Izpisua-Belmonte JC, De Robertis EM, Storey KG, Stern CD (1993) The homeobox gene goosecoid and the origin of organizer cells in the early chick blastoderm. Cell 74:645–659

Chapter 27 Immunohistochemistry and RNA In Situ Hybridization in Mouse Brain Development Jinling Liu and Aimin Liu Abstract During development, the mouse brain is progressively divided into functionally distinct compartments. Numerous neuronal and glial cell types are subsequently generated in response to various inductive signals. Each cell expresses a unique combination of genes encoding proteins from transcription factors to neurotransmitters that define its role in brain function. To understand these important and highly sophisticated processes, it is critical to accurately locate the various proteins and cells that produce them. In this chapter, we introduce the techniques of Immunohistochemistry, which detects the localization of specific proteins, and RNA in situ hybridization, which enables the visualization of specific mRNAs. Key words Immunohistochemistry, RNA in situ hybridization, Cryosection, Antibody, Digoxigenin, Fluorescent

1  Introduction The mouse brain consists of multiple divisions (cerebrum, epithalamus, thalamus, hypothalamus, cerebellum, and brain stem), and more than 100 million neurons and glia [1]. Extrinsic inductive signals and intrinsic cellular programs both play key roles in the compartmentalization of the brain as well as cellular behaviors such as proliferation, differentiation, migration, and cell death. To better understand the developmental processes involved in mouse brain development, it is important to obtain information regarding the spatial and temporal patterns of gene expression. In this chapter, we describe methods for the detection of protein (immunohistochemistry, IHC) and mRNA (RNA in situ hybridization, RISH) in brain sections. IHC detects particular proteins present in tissues. The principle underlying this technique is the specific bindings between antibodies and antigens. To visualize the antibody–antigen interaction, an antibody is tagged with a fluorophore, which can be conveniently detected with a fluorescent microscope. Alternatively, the Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_27, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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antibody can be conjugated to an enzyme that catalyzes a color-­ producing reaction, which can be visualized under a regular microscope. In this chapter we describe the method of IHC with the fluorophore-labeled antibody. The procedure comprises tissue preparation, blocking, primary and secondary antibody incubation, mounting, and visualization. The success of IHC heavily depends on the availability of high-­ quality antibodies. In addition, secreted signaling proteins as well as proteins of the extracellular matrix are not restricted to the cells producing them, preventing a high resolution identification of the signaling centers. On the other hand, RISH allows the detection of the expression of virtually all genes in the cells, even genes that do not encode proteins, providing more flexibility compared with the antibody-based IHC method. Traditionally, RISH depends on the hybridization of the specific RNA sequence in situ to radiolabeled probes [2]. Currently, Digoxigenin-labeled probes are more commonly used in RISH, which can be recognized with antibodies coupled with fluorophore or enzymes such as alkaline phosphatase or peroxidase [3]. RISH can be performed on both frozen sections and paraffin sections, with frozen sections allowing more sensitive detection of weak signals [4]. The RISH method we introduce in this chapter uses Digoxigenin-labeled riboprobes (complementary RNA probes) to detect specific mRNA on frozen brain sections. The procedure includes synthesis of riboprobes, hybridization of sections with Digoxigenin-labeled riboprobes, post-hybridization washes, incubation with alkaline phosphatase (AP)-conjugated anti-Digoxigenin antibody, and a color reaction using the phosphatase substrate BM purple solution.

2  Materials The materials should be stored at room temperature unless otherwise specified. 2.1  Materials for Cryosection Preparation

1. Dissection tools: student quality iris scissors (Fine Science Tools), Dumont forceps (Fine Science Tools), spoon (Fine Science Tools). 2. 6 cm petri dishes. 3. 24-well tissue culture plates (see Note 1). 4. Stereomicroscope, such as Nikon SMZ645. 5. Phosphate buffered saline (PBS): Prepare 10× stock by dissolving 80 g NaCl, 2 g KCl, 14.4 g Na2HPO4 and 2.4 g KH2PO4 in 1 l of distilled, de-ionized water (ddH2O). Adjust pH to 7.4 and autoclave at 121 °C for 25 min. Dilute to 1× solution with ddH2O for use.

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6. 4% Paraformaldehyde (PFA): prepare 16% stock by adding 32 g PFA powder and 100 μl 5 N NaOH in 150 ml ddH2O prewarmed at ~65 °C (see Note 2). Once PFA is dissolved, add 20 ml 10× PBS and adjust the volume to 200 ml with ddH2O. Filter through Whatman paper and make 10 ml aliquot in 50 ml centrifuge tubes. Store at −20 °C. Thaw one tube at ~65 °C and dilute to 4% PFA with PBS before use. Four percentage PFA can be stored at 4 °C for up to 1 week (see Note 3). 7. Nutator mixer (VWR). 8. 30% sucrose: Dissolve 12 g sucrose powder in 40 ml PBS. Store at 4 °C. 9. Tissue-Tek® O.C.T. compound (VWR): Store at 4 °C. 10. Disposable embedding molds. 11. Dry ice or liquid nitrogen in appropriate containers. 12. Cryostat, such as Leica CM1900. 13. Superfrost Plus microscopic slides (see Note 4). 14. Slide boxes. 15. Micro Slide trays. 2.2  Materials for Immunohistochemistry

1. Coplin Jars. 2. Blocking buffer: 1% normal goat serum, 0.1% Triton X-100, in PBS. Keep at 4 °C (see Note 5). 3. A humidified slide incubation chamber (Fig. 1): Cut two 5 ml serological pipettes and tape them to the bottom of a flat-­ bottom plastic box with lid, such that the two pipettes are parallel and 5 cm apart. Place paper towels soaked with ddH2O on the bottom to keep a moist environment. 4. Paper towels. 5. Fluorophore-conjugated secondary −20 °C in 50% Glycerol.

antibodies:

Store

at

6. Micro Slide trays. 7. Coverslips, 24 × 50 mm. 8. Forceps: Dumont forceps (Fine Science Tools). 9. Dabco® 33-LV: Aldrich. Store at 4 °C (see Note 6). 10. Nail polish. 2.3  Materials for the Synthesis of RNA Probes

1. A plasmid with the promoters for viral RNA polymerases (T3, T7 and Sp6) flanking the multiple cloning sites, Store at −20 °C. 2. Restriction endonucleases: Store at −20 °C. 3. Horizontal DNA electrophoresis apparatus and power supply.

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Fig. 1 A homemade humidified slide incubation chamber. This chamber is made by taping two 5 ml serological pipettes, broken to appropriate length, to the bottom of a plastic container with lid. Paper towels soaked in ddH2O provide humidity during the incubation. For RISH, a microwavable box or a container that can withstand 55 °C temperature is needed

4. Agarose. 5. 10 mg/ml Ethidium Bromide solution: keep away from light (see Note 7). 6. TBE buffer: Dissolve 54 g Tris base, 27.5 g Boric Acid in ddH2O, add 20 ml 0.5 M EDTA (PH 8.0), bring the volume to 5 l with ddH2O. 7. 1 kb DNA ladder. 8. 3 M sodium acetate (NaOAc), pH 5.2: Dissolve 408.3 g of NaOAc∙3H2O in 800 ml ddH2O. Adjust pH to 5.2 with acetic acid and adjust the volume to 1 l with ddH2O. Aliquot and autoclave. 9. Phenol saturated with Tris–HCl, pH 8.0: Store at 4 °C. 10. Chloroform. 11. 70% and 100% ethanol. 12. Tabletop microcentrifuge, such as Eppendorf 5415D. 13. Magnetic stir plate. 14. Diethylpyrocarbonate (DEPC): Store at 4 °C (see Note 8). 15. DEPC-H2O: Add 0.1% v/v DEPC to ddH2O and mix overnight on a magnetic stir plate at room temperature, autoclave at 121 °C for 25 min (see Note 9).

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16. 10× DIG labeling mix (Millipore Sigma): Store at −20 °C (see Note 10). 17. RNase inhibitor (Promega, 40 U/μl): Store at −20 °C. 18. T3 (Millipore Sigma), T7 (Millipore Sigma) and Sp6 (Millipore Sigma) RNA polymerases: Store at −20 °C. 19. DNaseI (RNase-free, Millipore Sigma): Store at −20 °C. 2.4  Materials for RNA In Situ Hybridization

The amount of reagents in this section is enough for processing five slides in a 50 ml Coplin jar, and should be adjusted according to the number of slides and size of the container used.

2.4.1  For Day 1: Hybridization of Cryosections on the Slides

All containers and reagents need to be RNase-free on Day 1. 1. Coplin jars: 50 ml. 2. Diethylpyrocarbonate (DEPC): Store at 4 °C (see Note 8). 3. Magnetic stir plate. 4. DEPC-H2O (400 ml): add 400 μl DEPC to 400 ml ddH2O. Mix overnight on a magnetic stir plate at room temperature, and autoclave at 121 °C for 25 min (see Note 9). 5. DEPC-PBS (400 ml): add 400 μl DEPC to 400 ml PBS. Mix overnight on a magnetic stir plate at room temperature, and autoclave at 121 °C for 25 min (see Note 9). 6. 4% PFA in DEPC-PBS (80 ml): Thaw two 10 ml aliquots of 16% PFA (see Subheading 2.1 for 16% PFA preparation) and dilute each with 30 ml DEPC-PBS. Store at 4 °C. 7. 0.25% acetic anhydride in 0.1 M TEA-HCl (40 ml): add 0.742 g Triethanolamine-HCl and 360 μl 5 N NaOH to 39.8 ml DEPC-H2O to make TEA-HCl. Add 100 μl acetic anhydride right before use (see Note 11). 8. 20 μg/ml Proteinase K (proK; Millipore Sigma): Dissolve one vial of proK (100 mg) in 5 ml DEPC-H2O to make 20 mg/ml stock. Aliquot and store at −20 °C. Thaw an aliquot before use and add 40 μl into 40 ml DEPC-PBS (see Note 12). 9. 70% ethanol in DEPC-H2O (40 ml): 28 ml 100% ethanol, 12 ml DEPC-H2O. 10. 95% ethanol in DEPC-H2O (40 ml): 38 ml 100% ethanol, 2 ml DEPC-H2O. 11. Hybridization solution (40 ml): mix the following in a clean 50 ml tube, aliquot and store at −20 °C (see Note 13).

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Formamide (deionized, aliquot in 50 ml tubes and store at −20 °C)

8 ml

50% dextran sulfate

400 μl

100× Denhardt’s (VWR, aliquot in 1.5 ml tubes and store at −20 °C)

1 ml

tRNA (10 mg/ml, aliquot in 1.5 ml tubes, store at −20 °C)

2.4 ml

5 M NaCl

800 μl

1 M Tris–HCl (PH 8.0)

400 μl

0.5 M EDTA

400 μl

1 M NaPO4

4 ml

10% Sarcosyl

2.6 ml

DEPC-H2O

12. Parafilm (cut into 24 mm × 50 mm pieces) or RNase-free plastic coverslips. 13. A hybridization oven. 14. A humidified slide incubation chamber: see Subheading 2.2. 2.4.2  For Day 2: Post-hybridization Washes and Antibody Incubation

Starting from day 2, reagents do not need to be RNase-free. 1. 20× saline-sodium citrate (SSC) buffer: mix the following in an appropriate container. 800 ml ddH2O 175.3 g NaCl 88.2 g Sodium citrate

Adjust PH to 7.0 with a few drops of 12 N HCl. Adjust the volume to 1 l with ddH2O. Aliquot and autoclave at 121 °C for 25 min. 2. 5× SSC (40 ml): 10 ml 20× SSC, 30 ml ddH2O. 3. 2× SSC (40 ml): 4 ml 20× SSC, 36 ml ddH2O. 4. 0.1× SSC (40 ml): 0.2 ml 20× SSC, 39.8 ml ddH2O. 5. High-Stringency Wash Buffer (120 ml) (Make fresh): 60 ml formamide, 12 ml 20× SSC, 48 ml ddH2O. 6. PBT (350 ml, enough for both day 2 and 3): add 350 μl Tween 20 to 350 ml PBS in a 500 ml bottle. Mix well by shaking vigorously. Alternatively, 10% Tween 20 stock can be made in ddH2O in advance and stored at room temperature. 7. RNase buffer (400 ml) (see Note 14): mix the following.

In Situ and Immunostaining in Mouse 40 ml

5 M NaCl

4 ml

1 M Tris–HCl, pH 7.5

4 ml

0.5 M EDTA, pH 8.0

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8. 10 mg/ml RNase A stock (DNase-free): add 10 ml 0.01 M NaOAc (pH 5.2) to a vial of 100 mg RNase A. Heat at 100 °C for 15 min. Cool to room temperature. Add 1 ml 1 M Tris– HCl (pH 7.5) and aliquot. Store at −20 °C (see Note 15). On the day of experiment, thaw one aliquot and make 20 μg/ml RNase A in RNase buffer by adding 80 μl 10 mg/ml RNase A stock to 40 ml RNase buffer. 9. Alkaline Phosphatase-conjugated anti-Digoxigenin antibody (Fab fragments from sheep, Millipore Sigma): Store at 4 °C. 10. A humidified slide incubation chamber: see Subheading 2.2. 2.4.3  For Day 3: Color Reaction

1. NTMT (80 ml) (Make fresh): see Note 16. 8 ml

1 M Tris 9.5

4 ml

1 M MgCl2

1.6 ml

5 M NaCl

800 μl

10% Tween20

40 mg

Levamisol

Adjust to 80 ml with ddH2O

2. Levamisol: make 50 mg/ml stock in ddH2O, aliquot and store in 1.5 ml tubes at −20 °C. 3. BM Purple (Millipore Sigma): Store at 4 °C. 4. Water-based mounting medium, such as Mount Quick (VWR). Store at 4 °C. 5. Coverslips, such as VWR, 24 × 50 mm. 6. Nuclear fast red solution (100 ml): Dissolve 5 g aluminum sulfate in 100 ml ddH2O, then add 0.1 g nuclear fast red. Boil and stir on a heated magnetic stir plate to dissolve Nuclear fast red. Filter the solution right before use.

3  Methods Conduct all procedures at room temperature unless otherwise specified.

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3.1  Mouse Brain Cryosection Preparation

1. Dissect embryos in ice-cold PBS in a 6 cm dish (see Note 17). 2. Cut the embryos at the shoulder level and transfer the heads into a 24-well plate with a spoon. 3. Fix the embryos in 4% PFA on a nutator at 4 °C for 1 h for IHC or overnight for RISH. 4. Rinse the embryos with PBS, and then wash the embryos in PBS on a nutator overnight at 4 °C. 5. Immerse the embryos in 30% sucrose overnight at 4 °C, on a nutator (see Note 18). 6. Change the 30% sucrose and incubate for another 2–3 h for further infiltration at 4 °C, on a nutator. 7. Transfer the brain samples into a disposable embedding mold and immerse the embryos in O.C.T. compound for 1 h at 4 °C (see Note 19). 8. Position the samples at desired orientation and freeze them on dry ice, or alternatively, in a paperboard box floating on the surface of liquid nitrogen. Wait for 5 min (see Note 20). 9. Transfer the frozen O.C.T. block containing the brain samples to the cryostat and wait at least 1 h so that the temperature of the block can reach the optimal cutting temperature (see Note 21). 10. Cut 10 μm sections using a cryostat (see Note 22). Collect sections on Superfrost Plus slides. 11. Dry the slides in a micro slide holder for 1 h at room temperature. Store them in a slide box at −80 °C (see Note 23).

3.2  Immunostaining on Mouse Brain Sections

1. Remove sections from −80 °C freezer and dry slides in a micro slide holder at room temperature for about 45 min (see Note 24). 2. Place the slides in a Coplin jar and incubate the sections with PBS plus 0.1% Triton X-100 for 1 h. 3. Take slides out, wipe the backside (the one without sections) and edges with paper towel. Place them with the front side (the one with sections) up on the pipettes of the humidified slide incubation chamber (Fig. 1; see Note 25). 4. Apply primary antibodies to the sections (diluted in blocking buffer, 300 μl per slide), and incubate overnight at 4 °C (see Note 26). 5. Pour primary antibody onto a paper towel and place the slides in a Coplin jar. Wash the slides with PBS plus 0.1% Triton X-100, 3 × 10 min. 6. Similar to step 3, place slides in the humidified chamber and incubate in the dark with appropriate fluorophore-conjugated secondary antibody (diluted in blocking buffer, 300 μl per slide) for 2 h (see Note 27).

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Fig. 2 Immunostaining of brain sections. Shown are two coronal sections of an E12.5 brain incubated with a monoclonal antibody against Pax6 and a Cy3-­conjugated secondary antibody. (a) Pax6 is weakly expressed in the dorsal region and strongly expressed in the intermediate region in the hindbrain. (b) Pax6 is expressed in the diencephalon (Di), but not the Mesencephalon (Mes). Dashed line demarcates the boundary between the Diencephalon and Mesencephalon

7. Pour secondary antibody onto a paper towel and place the slides in a Coplin jar. Wash with PBS plus 0.1% Triton X-100, 3 × 10 min. 8. Take slides out and wipe the backside and edges with paper towel. Place them into a micro slide tray. 9. Apply 30 μl Dabco evenly onto each slide and mount with a coverslip (see Notes 28 and 29); seal the slides with nail polish. 10. Observe the sections under a fluorescence microscope and take photos with a cooled CCD camera. Some examples are shown in Fig. 2. 11. Slides can be stored at 4 °C protected from light (see Note 30). 3.3  Synthesis of RNA Probes for RISH

1. Clone the cDNA (or part of the cDNA if the full-length cDNA is longer than 1500 bps; see Note 31) of the gene of interest into a plasmid containing the promoters for the viral RNA polymerases (T3, T7 and Sp6; such as pBluescript). 2. Linearize the template: Choose a unique restriction site in the multiple cloning sites on the 5′ end of the cDNA. Cut ~10 μg DNA in 20–50 μl reaction with the corresponding restriction endonuclease overnight using at least 20 IU enzyme. Run small amount (0.1–0.5 μg) in a 0.8% agarose gel to check the efficiency of restriction enzyme digestion.

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3. Purify the linearized DNA with a DNA clean up or PCR purification kit (such as The Nucleospin Gel and PCR clean up kit from Macherey-Nagel) and elute in 30 μl elution buffer (final concentration is ~0.3 μg/μl). The linearized template can be stored at −20 °C. 4. In vitro transcription: 2 μl

10× transcription buffer

2 μl

DIG labeling mix

6 μl

linearized template DNA

0.5 μl

RNase inhibitor

2 μl

RNA polymerase (T3, T7 or Sp6; use the one whose promoter is at the 3′ end of the cDNA)

7.5 μl

DEPC-H2O

Incubate at 37 °C for 2 h

5. Run 1 μl in a 1% agarose gel for 20 min to 1 h to check the yield of the probe. 6. DNase I treatment (optional): Add 1 μl DNase I and 1 μl RNase inhibitor to the RNA probe, incubate at 37 °C for 15 min to remove the template. 7. Add 180 μl DEPC-H2O, 20 μl 3 M NaOAc, mix well. Then add 600 μl 100% ethanol, mix well, and leave at −80 °C for 30 min. 8. Centrifuge at top speed (>14,000 × g) in a tabletop centrifuge for 15 min at 4 °C (see Note 32). 9. Discard the supernatant and rinse the pellet with 70% ethanol once without disturbing the pellet. 10. Discard 70% ethanol and dry the pellet for 5 min. Dissolve the RNA probe in 40–50 μl DEPC-H2O and store at −80 °C. 3.4  RNA In Situ Hybridization of Mouse Brain Sections 3.4.1  Day 1: Hybridization of Cryosections

All steps are carried out in Coplin jars unless otherwise specified; avoid RNase contamination. 1. Dry slides at room temperature for about 45 min (see Note 24). 2. Post-fix slides in 4% PFA in DEPC-PBS for 10 min (see Note 33). 3. Wash with DEPC-PBS, 2 × 5 min. 4. Drain excess DEPC-PBS and incubate for 6 min in 20 μg/ml proteinase K in DEPC-PBS (see Note 34). 5. Drain and wash with DEPC-PBS for 5 min. 6. Refix in 4% PFA for 5 min, then wash 5 min in DEPC-PBS (see Note 35).

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7. Acetylate sections with acetic anhydride in 0.1 M TEA-HCl for 10 min (see Note 36). 8. Wash in DEPC-PBS for 5 min, dehydrate in 70% ethanol for 5 min and 95% ethanol for 2 min. Air dry for 30 min to 2 h. 9. Add 2 μl RNA probe (approx. 1 μg) to 1 ml hybridization solution and heat at 80 °C for 2 min (see Note 37). 10. Place slides horizontally in a humidified slide incubation chamber. Cover sections with 200 μl of hybridization solution with the probe and lower parafilm coverslips over sections avoiding bubbles (see Note 38). 11. Seal the slide incubation chamber carefully and hybridize at 55 °C in a hybridization oven overnight (16–18 h). 3.4.2  Day 2: Post-­ hybridization Washes and Antibody Incubation

All steps are carried out in Coplin jars; RNase-free environment is not required. 1. Dip slides gently in a Coplin jar filled with 5× SSC to let the coverslips float off the slides (see Note 39). 2. Incubate the sections in high-stringency wash at 65 °C in a Coplin jar for 30 min (see Note 40). 3. Wash in RNase Buffer at 37 °C, 3 × 10 min. 4. Wash in RNase Buffer with 20 μg/ml RNase A at 37 °C for 30 min (see Note 41). 5. Wash in RNase buffer at 37 °C for 15 min. 6. Repeat high-stringency wash (as in step 2) at 65 °C, 2 × 20 min. 7. Wash in 2× SSC, then 0.1× SSC for 15 min each at 37 °C. 8. Wash with PBT for 15 min. 9. Take the slide out of PBT, wipe the backside and edges of the slide with paper towel. Place slides horizontally in a humidified slide incubation chamber and block for 1 h with 10% goat serum in PBT (300 μl per slide). 10. Pour the blocking buffer onto paper towels. Wipe the backside and edges of the slide with paper towel. Incubate with AP-conjugated anti-Digoxigenin antibody (diluted 1/5000 in PBT with 1% goat serum, 300 μl per slide) at 4 °C overnight in the same humidified chamber.

3.4.3  Day 3: Color Reaction

1. Pour the antibody onto paper towels. Place slides in a Coplin jar and wash in PBT for five times, 1 h each. 2. Wash 2 × 10 min in freshly prepared NTMT buffer. 3. Wipe the backside and edges of the slide with paper towel. Place slides horizontally in a humidified slide incubation chamber and incubate overnight to several days in BM purple solution (300 μl per slide) supplemented with 0.5 mg/ml Levamisol in the dark (see Note 42).

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Fig. 3 RNA in situ hybridization of brain sections. Shown are two sections of E12.5 brains hybridized with a Digoxigenin-labeled RNA probe against Otx2. (a) A coronal section through the forebrain region shows Otx2 expression in ventral telencephalon (tel) and the retinal pigment epithelium of the eye, but not in ventral Diencephalon (di). (b) A sagittal section shows that Otx2 is expressed in the Mesencephalon (mes), Diencephalon (di), but not in the Metencephalon (met) and Pons. The arrowhead points to the boundary between Diencephalon and Mesencephalon. Dashed lines demarcate the boundary between the midbrain and hindbrain. is isthmus

4. Observe periodically the progress of the color reaction under a microscope. If the staining is not ready, reapply BM purple solution and incubate for longer time (see Note 43). 5. When the signal is strong and the background staining just begins to show, place slides back into a Coplin jar. Wash slides in PBS for 2–5 min and dip briefly in ddH2O (see Note 18). 6. (Optional) Counterstain the sections with Nuclear fast red until the sections turn slightly pink. Usually it takes 2–3 min. 7. Wash excess Nuclear fast red in slow-running tap water. 8. Wipe the backside and edges of the slides with paper towel. Apply mounting medium to the slides and put coverslips on (see Note 44). 9. Observe the staining under a microscope and take photos with a color camera. Some examples are shown in Fig. 3. 10. The mounted sections can be stored at 4 °C.

4  Notes 1. The 24-well plates are for the convenient storage of individual small sample, such as the brains of E12.5 embryos or younger. Vials of appropriate size should be used for older/ bigger brain samples.

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2. Distilled, de-ionized water (ddH2O) used in this protocol is ultrapurified water with a resistance of 18.2 MΩ. 3. 16% PFA should be prepared in a fume hood because PFA is highly volatile and irritant. 4. It is critical to use Superfrost Plus slides as they contain a special coating to prevent the sections from falling off the slides during the incubation and washing steps. 5. Make fresh blocking buffer for each experiment. Do not store for more than a week. 6. Dabco is irritant. Avoid contact with skin and eyes. Avoid inhalation of vapor or mist. 7. Ethidium is highly carcinogenic. Avoid direct contact and dispose Ethidium-containing waste properly. 8. DEPC is toxic. Avoid direct contact. 9. All DEPC-treated solutions need to be autoclaved to degrade DEPC, which can react with RNA. 10. When thawing the DIG-labeling mix, avoid prolonged incubation at 37 °C to reduce the chance of NTP degradation. 11. Keep acetic anhydride from moisture (keep the container tightly closed all the time) and add acetic anhydride immediately before use. 12. Do not refreeze proK for RNA in situ hybridization. The activity of proK should be tested every time a new batch is introduced. 13. To make 50% dextran sulfate, mix 10 ml dextran sulfate and 10 ml ddH2O in a 50 ml tube by inverting, shaking, vortexing, and heating at 60 °C. When dextran sulfate starts to get into water, the total volume will decrease. Add more ddH2O to keep the final volume at 20 ml. 14. Sterile RNase buffer can be stored at room temperature. 15. Take caution to avoid contaminating the bench and other lab materials with RNase A. 16. NTMT should be freshly prepared. 17. At this step, we usually remove the extra-embryonic membranes or the tips of the tails to genotype the embryos. 18. The tissue should float on the surface of the 30% sucrose initially, and sink to the bottom of the well after sucrose has fully infiltrated the tissue. Make sure the tissue is completely immersed in the sucrose solution, as the morphology of tissues staying at the air/liquid interface can be distorted by surface tension. 19. The time the embryos immersed in OCT depends on the size and density of the tissue. For large and/or dense tissue, longer time is needed.

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20. Do not freeze samples by directly placing the embedding mold in liquid nitrogen. 21. If not cutting sections right away, wrap the embedding mold with parafilm and keep it at −80 °C for up to 1 week. 22. See manufacturer’s manual for how to cut cryosections. 23. Cryosections can be stored for months at −80 °C without noticeable degradation of proteins and mRNAs. 24. Dried sections do not fall off the slides in subsequent experiments. In addition, drying creates holes in the subcellular structure, permeabilizing the cells for further experiments. 25. Take caution not to let the sections dry completely. If necessary, process slides one at a time. This applies to all steps that involve taking slides out of the solution. 26. The antibody solution stays on the top of slides only if the edges and the bottom of the slides are dry. Therefore, add the antibody solution to the center of the slide and avoid moving the chamber once the antibody is added. Place a sign on the chamber to warn others not to move it during incubation. 27. Covering the humidified slide incubation chamber with foil or perform the incubation in a cabinet. 28. Cut the pipet tip to make a large orifice because Dabco is sticky. 29. To avoid bubbles, lower the coverslip slowly. 30. The fluorescent signal decreases over time, so try to observe the fluorescence as early as you can. 31. In general, shorter probes (1500 bps) have difficulty penetrating the cell membranes. 32. Take caution when removing the supernatant because the pellet may not be visible. 33. Post-fixation ensures the tissue is fixed equally with cross-­ linked RNA molecules. It also improves the retention of the tissue on the slide. 34. To drain excess DEPC-PBS, hold the slides and tilt the Coplin jar onto the paper towels. Proteinase K treatment improves the signal intensity by allowing greater access of the target mRNA for the probes. 35. Refixation improves the section stability after proteolytic digestion. 36. Acetylation chemically modifies proteins and reduces their nonspecific bindings. 37. Preheat the hybridization solution at 80 °C before use.

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38. To avoid bubbles, lower the parafilm coverslips slowly from one side to the other. Take caution when transporting the slide chamber to the oven such that the coverslips do not fall off the slides. 39. Don’t force the coverslips off the slides with forceps, or the sections may tear. 40. Prewarm solution for this step and steps 3–7. During the high-stringency wash, low salt concentration and high temperature inhibits nonspecific bindings. 41. RNase A digests single-stranded RNA to reduce the background signal. 42. Wrap the slide chamber in foil or place the chamber in a dark cabinet. 43. Before checking the staining status of the sections, prepare BM purple solution with 0.5 mg/ml levamisol in case the staining is not ready and more incubation with BM purple is needed. Otherwise, the sections may become dry before the BM purple solution is ready. 44. The mounting medium is very sticky and solidifies quickly. It is better to apply the mounting medium before the slides dry and put coverslip on immediately.

Acknowledgments We would like to thank Dr. Simeone for providing the RNA in situ probe for Otx2. The monoclonal antibody against Pax6 developed by Dr. Jessell was obtained from the Developmental Studies Hybridoma Bank developed under the auspices of the NICHD and maintained by Department of Biological Sciences, The University of Iowa, Iowa City, IA 52242, USA. Research in the Liu lab has been supported by US NSF (IOS-0949877 and IOS-­ 1257540), US NIH (HD083625) and a Penn State University new faculty start-up fund. References 1. Herculano-Houzel S, Mota B, Lent R (2006) Cellular scaling rules for rodent brains. Proc Natl Acad Sci U S A 103(32): 12138–12143. https://doi.org/10.1073/ pnas.0604911103 2. Gall JG, Pardue ML (1969) Formation and detection of RNA-DNA hybrid molecules in cytological preparations. Proc Natl Acad Sci U S A 63(2):378–383

3. Komminoth P, Merk FB, Leav I, Wolfe HJ, Roth J (1992) Comparison of 35S- and digoxigenin-­labeled RNA and oligonucleotide probes for in situ hybridization. Expression of mRNA of the seminal vesicle secretion protein II and androgen receptor genes in the rat prostate. Histochemistry 98(4):217–228 4. Wilcox JN (1993) Fundamental principles of in situ hybridization. J Histochem Cytochem 41(12):1725–1733

Chapter 28 The Cre/Lox System to Assess the Development of the Mouse Brain Claudius F. Kratochwil and Filippo M. Rijli Abstract Cre-mediated recombination has become a powerful tool to confine gene deletions (conditional knockouts) or overexpression of genes (conditional knockin/overexpression). By spatiotemporal restriction of genetic manipulations, major problems of classical knockouts such as embryonic lethality or pleiotropy can be circumvented. Furthermore, Cre-mediated recombination has broad applications in the analysis of the cellular behavior of subpopulations and cell types as well as for genetic fate mapping. This chapter gives an overview about applications for the Cre/LoxP system and their execution. Key words Cre recombinase, CRISPR-Cas9, Transgenesis, Conditional knockout, Conditional knockin, CreERT2, Flpe recombinase, MADM, Split-Cre, Brainbow

1  Introduction After the first gene knockout (KO) in the mouse was obtained by Thomas and Capecchi using site-directed mutagenesis of the HPRT gene in 1987 [1], the functions of a multitude of genes have been analyzed using this technique. Still the “simple” KO approach has two main restrictions. First, genes whose inactivation is embryonically lethal cannot be analyzed for their function in late development and adulthood. Secondly, KOs of pleiotropic genes (i.e. genes with function in multiple tissues and/or cell types) are difficult to analyze, as phenotypes might be combinations of multiple distinct defects and therefore quite complex to disentangle. The use of the Cre/Lox recombination system to induce gene knockouts in mice has been described for the first time by Gu et al. in 1994 [2]. The technology made it possible to—conditionally— knockout genes solely in subsets of cells (i.e., in a cell-type- or tissue-­specific manner), where Cre recombinase is expressed. Two years back, the technique had been already used to conditionally overexpress the SV40 large tumor antigen in mice [3]. The key principle of Cre-mediated recombination is that the recombinase Simon G. Sprecher (ed.), Brain Development: Methods and Protocols, Methods in Molecular Biology, vol. 2047, https://doi.org/10.1007/978-1-4939-9732-9_28, © Springer Science+Business Media, LLC, part of Springer Nature 2020

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enzyme can catalyze the deletion or inversion of a genomic fragment. If a sequence is deleted or inverted depends on the orientation of small flanking recognition sequences called Lox or LoxP sites. Mirrored LoxP sites lead to an inversion, LoxP sequences with the same direction to a deletion. Since the 1990s, Cre-mediated recombination has been used successfully for many applications, tissues, and model organisms including not only mice [4–7] but also Drosophila [8], Xenopus [9], zebrafish [10], and plants [11] Also the recombination mechanism has been elucidated by the analysis of the crystal structure of Cre and the Cre/LoxP interface [12]. In mammalian cells, Cre is the predominantly used recombinase for site-specific recombination and has been shown to be more effective than another (in Drosophila widely used) recombination system, the Flp/Frt system from Saccharomyces cerevisiae [13]. Similar to Cre, Flp deletes or inverts DNA fragments (and therefore was named Flp—pronounced “Flip”—recombinase), though using different target sequences. Later on a more efficient version, Flpe has been created, which works reliably in mice [14, 15] and offers a suitable alternative for some applications. To further restrict the knockout in time, inducible variants of Cre were designed. The most widely used version is CreERT2 [16]. Hereby Cre is fused to a mutated ligand-binding domain of the estrogen receptor (ER) [17]. The fusion protein is normally confined to the cytoplasm, while in the presence of the synthetic ligand tamoxifen or 4-hydroxytamoxifen, it translocates to the nucleus, where it can trigger recombination events (see Subheading 3.5). A light-activatable Cre recombinase to control activity in time and space has also been generated [18]. Another concept to restrict Cre or CreERT2 further in space came from Hirrlinger et al. [19]. By splitting Cre into two inactive fragments, which regain Cre activity when co-expressed, recombination could be restricted to the intersection of two expression domains (by using different enhancers/promoters for the two fragments). A similar system was established by Farago et al. [20], using a combination of the two recombination systems Cre/Lox and Flpe/Frt (see Subheading 3.6). Both systems have been also tested in vivo in transgenic animals. A powerful tool to construct a fate map of cells as well as to analyze gene function on a single-cell resolution was introduced with the MADM (mosaic analysis with double markers) system [21, 22], an adaptation of the MARCM (mosaic analysis with a repressible cell marker) system from Drosophila [23]. MADM uses Cre-mediated interchromosomal recombination. The approach combines two fluorescent markers (e.g. GFP and RFP) and thereby allows tracing of single-cell progenies. Moreover, if one of the markers is linked to a mutation, homoyzgous mutant, heterozygous, and wild-type cells can be obtained by interchromosomal recombination and distinguished by their fluorescent markers [24]

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(see in detail in Subheading 3.7). Another approach to achieve single-­cell resolution was given by the brainbow system from Livet et al. [25]. Here, recombination leads to a stochastic choice of expression of a fluorescent protein. Multiple integrations result in a combinatorial color code, creating dozens of distinguishable colors (see Subheading 3.8). Lastly, we report on further ideas to introduce and target Cre recombination (see Subheading 3.9) and discuss novel strategies that take advantage of CRISPR-Cas9 (see Subheading 3.10). In this chapter, we focus on the basic principles and applications of the Cre/LoxP and CreERT2/LoxP systems and their variations (including Split-Cre, MADM, brainbow) and will provide strategies and protocols for their use.

2  Materials 2.1  Reporter Mice

1. Rosa26 LacZ reporter line [26]; available at Jackson Laboratories: # 003504 B6.129S4-Gt(ROSA)26Sortm1Sor/J. 2. Rosa26 tdTomato reporter line [27]; available at Jackson Laboratories: # 007914 B6.Cg-Gt(ROSA)26Sortm14(CAG-tdTomato)Hze/J. 3. Rosa26 ZsGreen reporter line [27]; available at Jackson Laboratories: # 007906 B6.Cg-Gt(ROSA)26Sortm6(CAG-ZsGreen1)Hze/J. 4. Brainbow Reporter lines [25]; six transgenic lines available at Jackson Laboratories: # 007901 B6.Cg-Tg(Thy1-Brainbow1.0)HLich/J. # 007910 B6.CBA-Tg(Thy1-Brainbow1.0)LLich/J. # 007911 B6.Cg-Tg(Thy1-Brainbow1.1)MLich/J. # 007921 B6.Cg-Tg(Thy1-Brainbow2.1)RLich/J. # 013731 STOCK Gt(ROSA)26Sor/J. # 017492 B6.129P2-Gt(ROSA)26Sor /J. 5. MADM mice [21, 22, 28, 29]; several transgenic lines are available at Jackson Laboratories, the following are the most widely used: # 013749 STOCK Tg(ACTB-EGFP,-tdTomato)11Luo/J. # 013751 STOCK Tg(ACTB-tdTomato,-EGFP)11Luo/J. # 017932 STOCK Tg(ACTB-EGFP∗)10Luo/J. # 017923 STOCK Tg(ACTB-EGFP∗,-tdTomato)10Luo/J.

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# 017912 STOCK Gt(ROSA)26Sor/J. # 017921 STOCK Gt(ROSA)26Sor/J. 2.2  Generation of Genetically Modified Mice

1. Genomic DNA (for amplifying enhancers). 2. Cloning plasmid(s) (for Cre, CreERT2, Lox sites, minimal promoter, fluorescent proteins, resistance cassettes (e.g., Addgene)). 3. Standard reagents for molecular biology (restriction enzymes (NEB), ligase (NEB), competent E. coli (e.g., DH5alpha, Top10 (Invitrogen)), primers, antibiotics). 4. Modified recombinogenic bacterial strains for recombination [30] (see Note 1), alternatively reagents for CRISPR/Cas9 (see Subheading 3.10).

2.3  Tamoxifen Treatment for CreERT2-­ Mediated Recombination

1. Gavage/feeding needle to administer tamoxifen (Fine Science Tools). 2. Tamoxifen (SIGMA). 3. Corn oil (SIGMA) or sunflower oil. 4. Syringe (graded in 100 μl intervals).

2.4  Genotyping

1. Standard reagents for DNA extraction (1 M Tris–HCl pH 8.5; 5 M NaCl; 0.5 M EDTA; 20% SDS; proteinase K; isopropanol; ethanol). 2. Standard reagents and primers for polymerase chain reaction (PCR) or GoTaq® Green Master Mix (Promega). 3. Standard reagents for gel electrophoresis.

3  Methods 3.1  Applications of the Cre/LoxP System

To create a conditional knockout, the gene of interest is flanked with recognition sites (LoxP sites; Locus of crossing [x-ing]-over of bacteriophage P1) for the bacteriophage P1 Cre recombinase [31] (Fig.  1). If Cre recombinase is present, the sequence flanked by two LoxP sites (the “floxed” sequence) is excised (if Lox sites have the same orientation) or inverted (if Lox sites have opposing orientation) (Fig. 1). It is also possible to recombine between two different plasmids or chromosomes (interchromosomal recombination), which is, for example, used in the MADM system (see Subheading 3.7). The excision of genomic fragments gives the possibility to perturb gene function either (I) completely, by removing the whole gene or the start codon, (II) partly, by removing certain parts/exons of the gene or by truncating it, or by (III)

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Fig. 1 Distinct genomic outcomes of Cre/LoxP-mediated recombination. (a) If two Lox sites have opposing directionality, the flanked sequence is inversed upon Cre-mediated recombination. This is reversible via the same process. (b) If Lox sites have the same orientation, the flanked sequence is excised, resulting in a circular fragment and the sequence lacking this fragment. (c) If LoxP sites are present on two chromosomes, recombination can also occur interchromosomally, albeit at a lower frequency than if the LoxP sites are on the same chromosome. (d) The LoxP site is 34 base pairs (bp) long. Two palindromic 13 bp sequences, containing the Cre binding site (boxed), flank an asymmetric 8 bp sequence. Arrows indicate the sites of cleavage during recombination. The underlined base pairs are mutated in the most commonly used variants LoxN and Lox2722

changing its expression pattern and levels by excising, replacing, or modifying regulatory elements as enhancers or promoters. Similarly, Cre-mediated recombination can be used to overexpress genes, by excision of an intervening transcriptional termination sequence, flanked by LoxP sites, that prevents the transcription of the target gene [3] (Fig. 2b). As these constructs might result in low levels of leaking readthrough transcription, especially if many copies are present (e.g., when using gene transfer by viral vectors or electroporation), a different approach (also called flip-excision (FLEX) switch) was created, using LoxP sites, which are put in inverted orientation (Fig. 2c), causing an inversion of the intervening DNA (Fig. 1). Thereby the transcription of a gene can be initiated by inverting the open-reading frame. Because here none of the LoxP sites is removed, the inversion would continue forth and back. To interfere with that, a second LoxP-incompatible site pair is introduced (e.g., Lox2272) which results in the termination of the ongoing recombination by recombining out the specific recognition

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Fig. 2 Three examples for applications using the Cre/LoxP system. (a) The classic approach to generate a conditional knockout is to place LoxP sites in introns either flanking the exon, which contains the ATG, or exons that contain functionally important domains. Using this strategy, the probability to interfere with gene expression before Cre recombination is reasonably smaller than if LoxP sites are placed in promoter regions. (b) To overexpress genes an intervening sequence is placed between promoter and target gene in turn blocking transcription. Upon Cre-mediated recombination this sequence is removed, and the gene starts to be expressed under the control of the upstream promoter. (c) To reduce the risk of leaking, the use of double-inverted Lox sites might be preferred (flip-excision switch, or FLEX). Here, the gene of interest is placed in inverted or antisense orientation. Cre mediates the inversion of the sequence in the presence of a pair of Lox sites in inverted orientation. The use of a second pair of Lox sites also in reverse orientation, though incompatible with the first pair, eventually results in an irreversible sense configuration, as both sites eventually lack its partner site, allowing stable transcription

partners of both Lox sites (Fig. 2) [32]. It should be mentioned that the efficiency of Cre/LoxP-mediated recombination decreases in general with increasing genetic distance, but in principle any desired rearrangement can be made with the Cre/LoxP system [33]. 3.2  Designing Constructs

Cre/LoxP is a binary system. First, the gene of interest has to be flanked by LoxP sites (“floxed”) (Fig. 1). Second, recombinase expression in the cell or tissue of interest must be provided either by a knockin (KI) of Cre, a transgene in which Cre expression is

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Fig. 3 Two approaches to obtain tissue-/cell-type-specific Cre expression. (a) A tissue-specific enhancer is combined with a minimal promoter to drive tissue-specific Cre expression, where the enhancer is active. (b) Another approach is site-directed recombination into a bacterial artificial chromosome (BAC) or into the genome of embryonic stem cells

driven by a specific promoter/enhancer, a bacterial artificial chromosome (BAC) in which Cre is inserted in-frame at a specific locus, or gene transfer using, e.g., viruses or in utero electroporation (Fig.  3). Many tissue- or cell-type-specific enhancers have been described in the literature. Others can be found in enhancer databases such as the VISTA Enhancer Browser [34] or can be identified by selecting evolutionarily conserved sequences in proximity of genes with expression patterns of interest [35] and by cloning a fragment of a few kilobases of core and proximal promoter in front of the Cre recombinase gene. Finally, relevant enhancers can be also identified by genome-wide search, e.g., by ChIP-Seq or ATAC-Seq [36, 37]. Such distal regulatory elements that lack core promoter elements are usually combined with the minimal/core promoter (minP) of the human ß-globin promoter [38] to be able to drive efficient transcription (see examples of this approach in [39]). Also, several national and international programs have been launched that generate Cre lines and make them available to the community including the “Cre-driver mice Network” (www.credrivermice.org) or “The Jackson Laboratory’s Cre Repository” (www.jax.org). 3.3  Mouse Mating Schemes

Once Cre KI (i.e., inserted at specific loci) or Cre-expressing transgenic mice have been created, they need to be crossed to mice carrying LoxP site-bearing conditional alleles, in order to generate double heterozygotes for Cre and the floxed locus. As most laboratories dispose of multiple Cre driver lines, it is more space efficient to generate a few double heterozygous males for the Cre-expressing line and the conditional allele and mate them to homozygous conditional mutant females, which can be readily maintained as a pool. However, it should be noted that in this type of crossings, double heterozygous Cre/conditional allele and/or homozygous conditional mutant specimen in the absence of Cre-­ mediated excision usually displays a wild-type-like phenotype, thus serving as controls. It may be useful to additionally cross a floxed conditional reporter line into the background of Cre/conditional

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allele double heterozygotes. In many projects this will ease the analysis, as it will allow to first have a direct readout of the cells in which the gene has been knocked out, second directly analyze cellular behavior between heterozygotes and homozygotes, and third select tissue or cells using methods such as fluorescence-activated cell sorting (FACS). Conditional Cre-inducible reporter mice in which the reporter genes are inserted in housekeeping gene loci are available from the Jackson Laboratories. These lines provide stable and constitutive reporter expression once activated by Cre-­mediated excision. In particular, three KIs in the ROSA26 locus are the most frequently used reporter lines as they are highly sensitive to Cre-mediated activation, which express for either ß-­galactosidase (LacZ) [26], ZsGreen, or tdTomato [27], but additional reporter lines expressing other fluorescent proteins or genes for functional manipulations are also available [40] (see Subheading 3.9). The choice depends on the application. ß-Galactosidase catalyzes the transformation of X-gal into an insoluble blue enzymatic product and is preferentially used for nonfluorescent histochemistry or whole mount stainings of embryos or organs. The strongly fluorescent proteins ZsGreen (see Note 2) or tdTomato are a better choice for fluorescence histochemistry, live imaging, or cell sorting. The crossing scheme (Fig.  4) will give litters with all needed controls: 25% homozygotes (+Cre), 25% homozygotes (−Cre), and 25% heterozygotes

Fig. 4 Mating scheme to analyze gene function by conditional knockout. The use of pools for the conditional knockout (with or without reporter) eases the analysis if multiple Cre drivers are used to test gene function in different tissues. The offspring contains animals displaying mutant (Cre tg/+, Gene fl/fl) and control (Cre tg/+, Gene fl/+, and Gene fl/fl) phenotypes

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(−Cre). The controls can be used to exclude an effect of the Cre (heterozygotes +Cre) or of the floxed locus (homozygotes −Cre) when analyzing the phentoype of the homozygous mutants. 3.4  Genotyping

To genotype mice for maintenance and experiments, polymerase chain reaction (PCR) is the method of choice. Primers have to be designed to simultaneously detect (I) Cre, (II) floxed locus, (III) locus after Cre-mediated deletion, and (IV) wild-type locus of the respective gene. For (I), specific primers for Cre can be used or, alternatively, primers which span the Cre extremities and its flanking regions. For (II–IV), usually three primers are needed (Fig. 5). One primer pair is chosen on each side of one or the other of the LoxP sites to detect the small size difference, as compared to the wild-type fragment, due to the presence of the LoxP site (Fig. 5b). Another pair of primers is designed 5′ and 3′ to the two LoxP sites, respectively, to be able to detect the Cre-mediated deletion of the locus and confirm the efficiency of the recombination (Fig. 5c). PCRs for genotyping can be done using standard PCR protocols for genotyping on clipped toes, ears, or tails or any other tissue, where recombination has to be tested. We recommend the use of the GoTaq® Green Master Mix (Promega), because it is faster, reduces pipetting mistakes and contaminations, and increases reproducibility.

Fig. 5 Genotyping by PCR to detect gene deletion. (a) The targeted allele can be detected by primer pair 1 (F1 and R1). (b) The floxed allele can be detected by the same primers, yielding a longer amplicon. (c) A distinct primer pair (F2 and R1) detects the Cre-mediated recombination. The nonrecombined locus usually cannot be detected, as the amplicon will be too long. (d) Theoretical result of a PCR with different genotypes in Cre-­ positive (Cre+) and Cre-negative (Cre−) tissues

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3.4.1  DNA Extraction

1. Prepare Tail Buffer (500 ml Stock): 100 mM Tris–HCl pH 8.5

50 ml of 1 M

200 mM NaCl

20 ml of 5 M

5 mM EDTA

5 ml of 0.5 M

0.2% SDS

5 ml of 20% SDS

ddH2O

420 ml

2. Clip the tails or ears and collect them in 1.5 ml Eppendorf tubes. 3. Add 500 μl tail buffer and 10 μl proteinase K (20 mg/ml). 4. Digest overnight in a water incubator at 55 °C or 2 h on a shaking dry incubator at 55 °C. 5. Vortex tubes and centrifuge for 5 min at 12,000–16,000 × g. 6. Collect supernatant in new tube (to remove undigested tissue) with 500 μl isopropanol (2-propanol). 7. Shake vigorously and 12,000–16,000 × g.

centrifuge

for

10–15 min

at

8. Remove supernatant, add 500 μl 70% ethanol to wash pellet, and remove supernatant again. 9. Add 500 μl ddH2O and vortex vigorously. 3.4.2  Genotyping Using GoTaq® Green Master Mix

1. Aliquot 2× Master Mix into aliquots between 200 μl and 1 ml. 2. Prepare Primer Mix for each genotype (200 μl stock): Upstream primer, 100 μM

20 μl

Downstream primer, 100 μM

20 μl

ddH2O

160 μl

If two Up- or Downstream Primers are needed, use 10 μl of each. 3. Prepare Master Mix for each primer set: 1 Reaction 20 Reactions GoTaq® Green Master Mix, 2×

4 μl

Primer Mix, 10 μM for 0.4 μl each primer

80 μl 8 μl

4. Pipet 3.2 μl genomic DNA template and 4 μl Master Mix (for many samples a dispenser can be used) per reaction tube. Also half reactions can be done.

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5. Annealing conditions should be optimized. It is recommended to design all primers for approximately the same annealing temperature (e.g. 58 °C) because PCRs can be pooled in the same block and different primers can be combined. The following program works for many primer sets: Initial denaturation 30–35 cycles:

3 min

94 °C

Denaturation

40 s

94 °C

Annealing

40 s

52–62 °C

Extension

1 min

72 °C

Final extension

5 min

72 °C

Soak/refrigeration cycle

–

4 °C

6. Load DNA samples on a 1.5% Agarose Gel (one band) 2% Agarose Gel (multiple bands with