Mouse Cell Culture: Methods and Protocols [1st ed.] 978-1-4939-9085-6, 978-1-4939-9086-3

Table of contents :
Front Matter ....Pages i-xii
Front Matter ....Pages 1-1
The Applicability of Mouse Models to the Study of Human Disease (Kristina Rydell-Törmänen, Jill R. Johnson)....Pages 3-22
Optimizing the Cell Culture Microenvironment (Ivan Bertoncello)....Pages 23-30
Front Matter ....Pages 31-31
Propagation and Maintenance of Mouse Embryonic Stem Cells (Jacob M. Paynter, Joseph Chen, Xiaodong Liu, Christian M. Nefzger)....Pages 33-45
Production of High-Titer Lentiviral Particles for Stable Genetic Modification of Mammalian Cells (Michael R. Larcombe, Jan Manent, Joseph Chen, Ketan Mishra, Xiaodong Liu, Christian M. Nefzger)....Pages 47-61
Generation of Mouse-Induced Pluripotent Stem Cells by Lentiviral Transduction (Xiaodong Liu, Joseph Chen, Jaber Firas, Jacob M. Paynter, Christian M. Nefzger, Jose M. Polo)....Pages 63-76
Gene Editing of Mouse Embryonic and Epiblast Stem Cells (Tennille Sibbritt, Pierre Osteil, Xiaochen Fan, Jane Sun, Nazmus Salehin, Hilary Knowles et al.)....Pages 77-95
Identification of Circulating Endothelial Colony-Forming Cells from Murine Embryonic Peripheral Blood (Yang Lin, Chang-Hyun Gil, Mervin C. Yoder)....Pages 97-107
Imaging and Analysis of Mouse Embryonic Whole Lung, Isolated Tissue, and Lineage-Labelled Cell Culture (Matthew Jones, Saverio Bellusci)....Pages 109-127
Mouse Hematopoietic Stem Cell Modification and Labelling by Transduction and Tracking Posttransplantation (Benjamin Cao, Songhui Li, Claire Pritchard, Brenda Williams, Susan K. Nilsson)....Pages 129-142
Genetic Manipulation and Selection of Mouse Mesenchymal Stem Cells for Delivery of Therapeutic Factors In Vivo (Donald S. Sakaguchi)....Pages 143-155
Isolation and Culture of Primary Mouse Middle Ear Epithelial Cells (Apoorva Mulay, Khondoker Akram, Lynne Bingle, Colin D. Bingle)....Pages 157-168
Isolation and Propagation of Lacrimal Gland Putative Epithelial Progenitor Cells (Helen P. Makarenkova, Robyn Meech)....Pages 169-180
Organotypic Culture of Adult Mouse Retina (Brigitte Müller)....Pages 181-191
Langendorff-Free Isolation and Propagation of Adult Mouse Cardiomyocytes (Matthew Ackers-Johnson, Roger S. Foo)....Pages 193-204
Isolation, Culture, and Characterization of Primary Mouse Epidermal Keratinocytes (Ling-Juan Zhang)....Pages 205-215
Isolation and Propagation of Mammary Epithelial Stem and Progenitor Cells (Julie M. Sheridan, Jane E. Visvader)....Pages 217-229
An Organoid Assay for Long-Term Maintenance and Propagation of Mouse Prostate Luminal Epithelial Progenitors and Cancer Cells (Yu Shu, Chee Wai Chua)....Pages 231-254
Isolation, Purification, and Culture of Mouse Pancreatic Islets of Langerhans (Youakim Saliba, Nassim Farès)....Pages 255-265
Identification and In Vitro Expansion of Adult Hepatocyte Progenitors from Chronically Injured Livers (Naoki Tanimizu)....Pages 267-273
The Preparation of Decellularized Mouse Lung Matrix Scaffolds for Analysis of Lung Regenerative Cell Potential (Deniz A. Bölükbas, Martina M. De Santis, Hani N. Alsafadi, Ali Doryab, Darcy E. Wagner)....Pages 275-295
Mouse Lung Tissue Slice Culture (Xinhui Wu, Eline M. van Dijk, I. Sophie T. Bos, Loes E. M. Kistemaker, Reinoud Gosens)....Pages 297-311
Back Matter ....Pages 313-315

Citation preview

Methods in Molecular Biology 1940

Ivan Bertoncello Editor

Mouse Cell Culture Methods and Protocols

METHODS

IN

MOLECULAR BIOLOGY

Series Editor John M. Walker School of Life and Medical Sciences University of Hertfordshire Hatfield, Hertfordshire, AL10 9AB, UK

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

Mouse Cell Culture Methods and Protocols

Edited by

Ivan Bertoncello Lung Health Research Centre, Department of Pharmacology and Therapeutics, University of Melbourne, Melbourne, Victoria, Australia

Editor Ivan Bertoncello Lung Health Research Centre Department of Pharmacology and Therapeutics University of Melbourne Melbourne, Victoria, Australia

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-4939-9085-6 ISBN 978-1-4939-9086-3 (eBook) https://doi.org/10.1007/978-1-4939-9086-3 Library of Congress Control Number: 2019930653 © Springer Science+Business Media, LLC, part of Springer Nature 2019 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 Press 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 Mouse models have long underpinned discovery in the biomedical sciences and the development of enabling technologies for the systematic analysis of cellular and molecular mechanisms of organ development, regeneration, and repair. In particular, mouse cell culture systems continue to play an important role in identifying key factors, genes, and pathways regulating cell lineage commitment and differentiation and validating potential cellular and molecular targets which could be exploited to develop novel therapies for intractable diseases. This compendium describes recently devised and refined best-practice cell culture protocols for the maintenance, propagation, manipulation, and analysis of primary explanted cells from various mouse organ systems commonly used in current research applications. Each chapter provides a step-by-step description of cell culture methodologies for specific mouse cell types and lineages, highlighting caveats and commonly encountered pitfalls. These protocols are preceded by two introductory chapters that review the applicability of mouse models as a discovery tool and describe factors and variables that influence cell culture endpoints and need to be considered and controlled in order to achieve optimal results. In conclusion, I would like to acknowledge and thank the many authors who have enthusiastically contributed their protocols to this volume. I also thank John Walker (series editor) for his invitation to edit the volume and for his advice and assistance in developing and preparing the volume for publication. Melbourne, Victoria, Australia

Ivan Bertoncello

v

Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

PART I

PRACTICAL CONSIDERATIONS

1 The Applicability of Mouse Models to the Study of Human Disease. . . . . . . . . . . Kristina Rydell-To¨rm€ a nen and Jill R. Johnson 2 Optimizing the Cell Culture Microenvironment . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ivan Bertoncello

PART II

v xi

3 23

METHODS AND PROTOCOLS

3 Propagation and Maintenance of Mouse Embryonic Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jacob M. Paynter, Joseph Chen, Xiaodong Liu, and Christian M. Nefzger 4 Production of High-Titer Lentiviral Particles for Stable Genetic Modification of Mammalian Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Michael R. Larcombe, Jan Manent, Joseph Chen, Ketan Mishra, Xiaodong Liu, and Christian M. Nefzger 5 Generation of Mouse-Induced Pluripotent Stem Cells by Lentiviral Transduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Xiaodong Liu, Joseph Chen, Jaber Firas, Jacob M. Paynter, Christian M. Nefzger, and Jose M. Polo 6 Gene Editing of Mouse Embryonic and Epiblast Stem Cells. . . . . . . . . . . . . . . . . . Tennille Sibbritt, Pierre Osteil, Xiaochen Fan, Jane Sun, Nazmus Salehin, Hilary Knowles, Joanne Shen, and Patrick P. L. Tam 7 Identification of Circulating Endothelial Colony-Forming Cells from Murine Embryonic Peripheral Blood . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yang Lin, Chang-Hyun Gil, and Mervin C. Yoder 8 Imaging and Analysis of Mouse Embryonic Whole Lung, Isolated Tissue, and Lineage-Labelled Cell Culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Matthew Jones and Saverio Bellusci 9 Mouse Hematopoietic Stem Cell Modification and Labelling by Transduction and Tracking Posttransplantation . . . . . . . . . . . . . . . . . . . . . . . . . . Benjamin Cao, Songhui Li, Claire Pritchard, Brenda Williams, and Susan K. Nilsson 10 Genetic Manipulation and Selection of Mouse Mesenchymal Stem Cells for Delivery of Therapeutic Factors In Vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . Donald S. Sakaguchi 11 Isolation and Culture of Primary Mouse Middle Ear Epithelial Cells . . . . . . . . . . Apoorva Mulay, Khondoker Akram, Lynne Bingle, and Colin D. Bingle

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47

63

77

97

109

129

143 157

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13 14

15

16

17

18

19

20

21

Contents

Isolation and Propagation of Lacrimal Gland Putative Epithelial Progenitor Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Helen P. Makarenkova and Robyn Meech Organotypic Culture of Adult Mouse Retina . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ¨ ller Brigitte Mu Langendorff-Free Isolation and Propagation of Adult Mouse Cardiomyocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Matthew Ackers-Johnson and Roger S. Foo Isolation, Culture, and Characterization of Primary Mouse Epidermal Keratinocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ling-Juan Zhang Isolation and Propagation of Mammary Epithelial Stem and Progenitor Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Julie M. Sheridan and Jane E. Visvader An Organoid Assay for Long-Term Maintenance and Propagation of Mouse Prostate Luminal Epithelial Progenitors and Cancer Cells. . . . . . . . . . . Yu Shu and Chee Wai Chua Isolation, Purification, and Culture of Mouse Pancreatic Islets of Langerhans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Youakim Saliba and Nassim Fare`s Identification and In Vitro Expansion of Adult Hepatocyte Progenitors from Chronically Injured Livers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Naoki Tanimizu The Preparation of Decellularized Mouse Lung Matrix Scaffolds for Analysis of Lung Regenerative Cell Potential . . . . . . . . . . . . . . . . . . . . . . . . . . . . ¨ kbas, Martina M. De Santis, Hani N. Alsafadi, Deniz A. Bo¨lu Ali Doryab, and Darcy E. Wagner Mouse Lung Tissue Slice Culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Xinhui Wu, Eline M. van Dijk, I. Sophie T. Bos, Loes E. M. Kistemaker, and Reinoud Gosens

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

169 181

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Contributors MATTHEW ACKERS-JOHNSON  Cardiovascular Research Institute, Centre for Translational Medicine MD6, National University Health System, Singapore, Singapore; Genome Institute of Singapore, Singapore, Singapore KHONDOKER AKRAM  Academic Unit of Respiratory Medicine, Department of Infection, Immunity and Cardiovascular Disease, University of Sheffield, Sheffield, UK HANI N. ALSAFADI  Department of Experimental Medical Sciences, Faculty of Medicine, Lund University, Lund, Sweden; Wallenberg Centre for Molecular Medicine, Faculty of Medicine, Lund University, Lund, Sweden; Stem Cell Centre, Lund University, Lund, Sweden SAVERIO BELLUSCI  Faculty of Medicine, Excellence Cluster Cardio-Pulmonary System (ECCPS), University of Giessen, Giessen, Germany IVAN BERTONCELLO  Lung Health Research Centre, Department of Pharmacology and Therapeutics, University of Melbourne, Melbourne, Victoria, Australia COLIN D. BINGLE  Academic Unit of Respiratory Medicine, Department of Infection, Immunity and Cardiovascular Disease, University of Sheffield, Sheffield, UK LYNNE BINGLE  Oral and Maxillofacial Pathology, Department of Clinical Dentistry, University of Sheffield, Sheffield, UK DENIZ A. BO¨LU¨KBAS  Department of Experimental Medical Sciences, Faculty of Medicine, Lund University, Lund, Sweden; Wallenberg Centre for Molecular Medicine, Faculty of Medicine, Lund University, Lund, Sweden; Stem Cell Centre, Lund University, Lund, Sweden I. SOPHIE T. BOS  Faculty of Science and Engineering, Department of Molecular Pharmacology, University of Groningen, Groningen, The Netherlands; Groningen Research Institute for Asthma and COPD, University Medical Center Groningen, University of Groningen, Groningen, The Netherlands BENJAMIN CAO  Biomedical Manufacturing, CSIRO Clayton, Clayton, VIC, Australia; Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia JOSEPH CHEN  Department of Anatomy and Developmental Biology, Monash University, Clayton, VIC, Australia; Development and Stem Cells Program, Monash Biomedicine Discovery Institute, Clayton, VIC, Australia; Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia CHEE WAI CHUA  State Key Laboratory of Oncogenes and Related Genes, Renji-Med X Clinical Stem Cell Research Center, Department of Urology, Ren Ji Hospital, School of Medicine, Shanghai Jiao Tong University, Shanghai, China MARTINA M. DE SANTIS  Department of Experimental Medical Sciences, Faculty of Medicine, Lund University, Lund, Sweden; Wallenberg Centre for Molecular Medicine, Faculty of Medicine, Lund University, Lund, Sweden; Stem Cell Centre, Lund University, Lund, Sweden ALI DORYAB  Helmholtz Zentrum Mu¨nchen, Member of the German Center for Lung Research (DZL), Institute of Lung Biology and Disease, Neuherberg, Germany XIAOCHEN FAN  Children’s Medical Research Institute, The University of Sydney, Westmead, NSW, Australia

ix

x

Contributors

NASSIM FARE`S  Laboratoire de Recherche en Physiologie et Physiopathologie LRPP, Poˆle Technologie Sante´, Faculte´ de Me´decine, Universite´ Saint Joseph, Beirut, Lebanon JABER FIRAS  Department of Anatomy and Developmental Biology, Monash University, Clayton, VIC, Australia; Development and Stem Cells Program, Monash Biomedicine Discovery Institute, Clayton, VIC, Australia; Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia ROGER S. FOO  Cardiovascular Research Institute, Centre for Translational Medicine MD6, National University Health System, Singapore, Singapore; Genome Institute of Singapore, Singapore, Singapore CHANG-HYUN GIL  Department of Pediatrics, Herman B. Wells Center for Pediatric Research, Indiana University School of Medicine, Indianapolis, IN, USA REINOUD GOSENS  Faculty of Science and Engineering, Department of Molecular Pharmacology, University of Groningen, Groningen, The Netherlands; Groningen Research Institute for Asthma and COPD, University Medical Center Groningen, University of Groningen, Groningen, The Netherlands JILL R. JOHNSON  School of Life and Health Sciences, Aston University, Birmingham, UK MATTHEW JONES  Faculty of Medicine, Excellence Cluster Cardio-Pulmonary System (ECCPS), University of Giessen, Giessen, Germany LOES E. M. KISTEMAKER  Faculty of Science and Engineering, Department of Molecular Pharmacology, University of Groningen, Groningen, The Netherlands; Groningen Research Institute for Asthma and COPD, University Medical Center Groningen, University of Groningen, Groningen, The Netherlands HILARY KNOWLES  Children’s Medical Research Institute, The University of Sydney, Westmead, NSW, Australia MICHAEL R. LARCOMBE  Department of Anatomy and Developmental Biology, Monash University, Clayton, VIC, Australia; Development and Stem Cells Program, Monash Biomedicine Discovery Institute, Clayton, VIC, Australia; Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia SONGHUI LI  Biomedical Manufacturing, CSIRO Clayton, Clayton, VIC, Australia; Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia YANG LIN  Department of Pediatrics, Herman B. Wells Center for Pediatric Research, Indiana University School of Medicine, Indianapolis, IN, USA; Department of Biochemistry and Molecular Biology, Indiana University School of Medicine, Indianapolis, IN, USA XIAODONG LIU  Department of Anatomy and Developmental Biology, Monash University, Clayton, VIC, Australia; Development and Stem Cells Program, Monash Biomedicine Discovery Institute, Clayton, VIC, Australia; Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia HELEN P. MAKARENKOVA  Department of Molecular Medicine, The Scripps Research Institute, La Jolla, CA, USA JAN MANENT  Department of Anatomy and Developmental Biology, Monash University, Clayton, VIC, Australia; Development and Stem Cells Program, Monash Biomedicine Discovery Institute, Clayton, VIC, Australia; Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia ROBYN MEECH  Discipline of Clinical Pharmacology, College of Medicine and Public Health, Flinders University, Bedford Park, SA, Australia KETAN MISHRA  Department of Anatomy and Developmental Biology, Monash University, Clayton, VIC, Australia; Development and Stem Cells Program, Monash Biomedicine

Contributors

xi

Discovery Institute, Clayton, VIC, Australia; Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia APOORVA MULAY  Academic Unit of Respiratory Medicine, Department of Infection, Immunity and Cardiovascular Disease, University of Sheffield, Sheffield, UK BRIGITTE MU¨LLER  Department of Ophthalmology, Justus-Liebig-University Gießen, Gießen, Germany CHRISTIAN M. NEFZGER  Department of Anatomy and Developmental Biology, Monash University, Clayton, VIC, Australia; Development and Stem Cells Program, Monash Biomedicine Discovery Institute, Clayton, VIC, Australia; Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia; Institute for Molecular Bioscience, The University of Queensland, St Lucia, QLD, Australia SUSAN K. NILSSON  Biomedical Manufacturing, CSIRO Clayton, Clayton, VIC, Australia; Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia PIERRE OSTEIL  Children’s Medical Research Institute, The University of Sydney, Westmead, NSW, Australia; School of Medical Sciences, The University of Sydney, Camperdown, NSW, Australia JACOB M. PAYNTER  Department of Anatomy and Developmental Biology, Monash University, Clayton, VIC, Australia; Development and Stem Cells Program, Monash Biomedicine Discovery Institute, Clayton, VIC, Australia; Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia JOSE M. POLO  Department of Anatomy and Developmental Biology, Monash University, Clayton, VIC, Australia; Development and Stem Cells Program, Monash Biomedicine Discovery Institute, Clayton, VIC, Australia; Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia CLAIRE PRITCHARD  Biomedical Manufacturing, CSIRO Clayton, Clayton, VIC, Australia; Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia € KRISTINA RYDELL-TO¨RMANEN  Lung Biology Group, Department of Experimental Medical Science, Lund University, Lund, Sweden DONALD S. SAKAGUCHI  Department of Genetics, Development and Cell Biology, Iowa State University, Ames, IA, USA; Neuroscience Program, Iowa State University, Ames, IA, USA NAZMUS SALEHIN  Children’s Medical Research Institute, The University of Sydney, Westmead, NSW, Australia YOUAKIM SALIBA  Laboratoire de Recherche en Physiologie et Physiopathologie LRPP, Poˆle Technologie Sante´, Faculte´ de Me´decine, Universite´ Saint Joseph, Beirut, Lebanon JOANNE SHEN  Children’s Medical Research Institute, The University of Sydney, Westmead, NSW, Australia JULIE M. SHERIDAN  Molecular Genetics of Cancer Division, Walter and Eliza Hall Institute of Medical Research, Parkville, VIC, Australia; Department of Medical Biology, University of Melbourne, Melbourne, VIC, Australia YU SHU  State Key Laboratory of Oncogenes and Related Genes, Renji-Med X Clinical Stem Cell Research Center, Department of Urology, Ren Ji Hospital, School of Medicine, Shanghai Jiao Tong University, Shanghai, China TENNILLE SIBBRITT  Children’s Medical Research Institute, The University of Sydney, Westmead, NSW, Australia; School of Medical Sciences, The University of Sydney, Camperdown, NSW, Australia JANE SUN  Children’s Medical Research Institute, The University of Sydney, Westmead, NSW, Australia

xii

Contributors

PATRICK P. L. TAM  Children’s Medical Research Institute, The University of Sydney, Westmead, NSW, Australia; School of Medical Sciences, The University of Sydney, Camperdown, NSW, Australia NAOKI TANIMIZU  Department of Tissue Development and Regeneration, Research Institute for Frontier Medicine, Sapporo Medical University School of Medicine, Sapporo, Japan ELINE M. VAN DIJK  Faculty of Science and Engineering, Department of Molecular Pharmacology, University of Groningen, Groningen, The Netherlands; Groningen Research Institute for Asthma and COPD, University Medical Center Groningen, University of Groningen, Groningen, The Netherlands JANE E. VISVADER  Department of Medical Biology, University of Melbourne, Melbourne, VIC, Australia; Stem Cells and Cancer Division, Walter and Eliza Hall Institute of Medical Research, Parkville, VIC, Australia DARCY E. WAGNER  Department of Experimental Medical Sciences, Faculty of Medicine, Lund University, Lund, Sweden; Wallenberg Centre for Molecular Medicine, Faculty of Medicine, Lund University, Lund, Sweden; Stem Cell Centre, Lund University, Lund, Sweden BRENDA WILLIAMS  Biomedical Manufacturing, CSIRO Clayton, Clayton, VIC, Australia; Australian Regenerative Medicine Institute, Monash University, Clayton, VIC, Australia XINHUI WU  Faculty of Science and Engineering, Department of Molecular Pharmacology, University of Groningen, Groningen, The Netherlands; Groningen Research Institute for Asthma and COPD, University Medical Center Groningen, University of Groningen, Groningen, The Netherlands MERVIN C. YODER  Department of Pediatrics, Herman B. Wells Center for Pediatric Research, Indiana University School of Medicine, Indianapolis, IN, USA; Department of Biochemistry and Molecular Biology, Indiana University School of Medicine, Indianapolis, IN, USA LING-JUAN ZHANG  School of Pharmaceutical Sciences, Xiamen University, Xiamen, China; Department of Dermatology, School of Medicine, University of California San Diego, La Jolla, CA, USA

Part I Practical Considerations

Chapter 1 The Applicability of Mouse Models to the Study of Human Disease Kristina Rydell-To¨rm€anen and Jill R. Johnson Abstract The laboratory mouse Mus musculus has long been used as a model organism to test hypotheses and treatments related to understanding the mechanisms of disease in humans; however, for these experiments to be relevant, it is important to know the complex ways in which mice are similar to humans and, crucially, the ways in which they differ. In this chapter, an in-depth analysis of these similarities and differences is provided to allow researchers to use mouse models of human disease and primary cells derived from these animal models under the most appropriate and meaningful conditions. Although there are considerable differences between mice and humans, particularly regarding genetics, physiology, and immunology, a more thorough understanding of these differences and their effects on the function of the whole organism will provide deeper insights into relevant disease mechanisms and potential drug targets for further clinical investigation. Using specific examples of mouse models of human lung disease, i.e., asthma, chronic obstructive pulmonary disease, and pulmonary fibrosis, this chapter explores the most salient features of mouse models of human disease and provides a full assessment of the advantages and limitations of these models, focusing on the relevance of disease induction and their ability to replicate critical features of human disease pathophysiology and response to treatment. The chapter concludes with a discussion on the future of using mice in medical research with regard to ethical and technological considerations. Key words Mouse, Model, Disease, Genetics, Physiology, Immunology, Ethics

1

The Mouse: From Pest, to Pet, to Predominant Tool in Medical Research Although the genetic lineages of mice and humans diverged around 75 million years ago, these two species have evolved to live together, particularly since the development of agriculture. For millennia, mice (Mus musculus) were considered to be pests due to their propensity to ravenously consume stored foodstuff (mush in ancient Sanskrit means “to steal” [1]) and their ability to adapt to a wide range of environmental conditions. Since the 1700s, domesticated mice have been bred and kept as companion animals, and in Victorian England, “fancy” mice were prized for their variations in coat color and comportment; these mouse strains were the

Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940, https://doi.org/10.1007/978-1-4939-9086-3_1, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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€nen and Jill R. Johnson Kristina Rydell-To¨rma

forerunners to the strains used in the laboratory today. Robert Hooke performed the first recorded inquiry-driven experiments on mice in 1664, when he investigated the effects of changes in air pressure on respiratory function [2]. More recently, with data from the Human Genome Project and sequencing of the Mus musculus genome showing remarkable genetic homology between these species, as well as the advent of biotechnology and the development of myriad knockout and transgenic mouse strains, it is clear why the mouse has become the most ubiquitous model organism used to study human disease. In addition, their small size, rapid breeding, and ease of handling are all important advantages to scientists for practical and financial reasons. However, keeping in mind that mice are fellow vertebrates and mammals, there are ethical issues inherent to using these animals in medical research. This chapter will provide an overview of the important similarities and differences between Mus musculus and Homo sapiens and their relevance to the use of the mouse as a model organism and provide specific examples of the quality of mouse models used to investigate the mechanisms, pathology, and treatment of human lung diseases. We will then conclude with an assessment of the future of mice in medical research considering ethical and technological advances. As a model organism used to test hypotheses and treatments related to human disease, it is important to understand the complex ways in which mice are similar to humans, and crucially, the ways in which they differ. A clear understanding of these aspects will allow researchers to use mouse models of human disease and primary cells derived from mice under the most appropriate and meaningful conditions.

2 2.1

Applicability of Mouse Models to Human Disease Genetics

In 2014, the Encyclopedia of DNA Elements (ENCODE) program published a comparative analysis of the genomes of Homo sapiens and Mus musculus [3], as well as an in-depth analysis of the differences in the regulatory landscape of the genomes of these species [4]. ENCODE, a follow-up to the Human Genome Project, was implemented by the National Human Genome Research Institute (NHGRI) at the National Institutes of Health in order to develop a comprehensive catalog of protein-encoding and nonproteincoding genes and the regulatory elements that control gene expression in a number of species. This was achieved using a number of genomic approaches (e.g., RNA-seq, DNase-seq, and ChIP-seq) to assess gene expression in over 100 mouse cell types and tissues; the data were then compared with the human genome. Overall, these studies showed that although gene expression is fairly similar between mice and humans, considerable differences were observed in the regulatory networks controlling the activity of

Mouse Models of Human Disease

5

the immune system, metabolic functions, and responses to stress, all of which have important implications when using mice to model human disease. In essence, mice and humans demonstrate genetic similarity with regulatory divergence. Specifically, there is a high degree of similarity in transcription factor networks but a great deal of divergence in the cis-regulatory elements that control gene transcription in the mouse and human genomes. Moreover, the chromatin landscape in cell types of similar lineages in mouse and human is both developmentally stable and evolutionarily conserved [3]. Of particular relevance regarding modeling human diseases involving the immune system, in its assessment of transcription factor networks, the Mouse ENCODE Consortium revealed potentially important differences in the activity of ETS1 in the mouse and human genome. Although conserved between the two species, divergence in ETS1 regulation may be responsible for discrepancies in the function of the immune system in mouse and human [4]. Certainly, the biological consequences of these differences in gene expression and regulation between human and mouse invite further investigation. 2.2 Anatomy and Physiology

The anatomical and physiological differences between model organisms and humans can have profound impacts on interpreting experimental results. Virtually every biological process under investigation in experimental studies involves at least one anatomical structure. To aid in interpretation, many anatomy compendia have been developed for model organisms; the most useful organize anatomical entities into hierarchies representing the structure of the human body, e.g., the Foundational Model of Anatomy developed by the Structural Informatics Group at the University of Washington [5]. Although an analysis of the myriad differences between mouse and human anatomy is beyond the scope of this chapter, a few of the most critical issues that have an impact on the interpretation of data from mouse experiments should be mentioned. The most obvious difference between mice and humans is size; the human body is about 2500 times larger than that of the mouse. Size influences many aspects of biology, particularly the metabolic rate, which is correlated to body size in placental mammals through the relationship BMR ¼ 70  mass (0.75), where BMR is the basal metabolic rate (in kcal/day). Thus, the mouse BMR is roughly seven times faster than that of an average-sized human [6]. This higher BMR has effects on thermoregulation, nutrient demand, and nutrient supply. As such, mice have greater amounts of metabolically active tissues (e.g., liver and kidney) and more extensive deposits of brown fat [6]. Furthermore, mice more readily produce reactive oxygen species than do humans, which is an important consideration when modeling human diseases involving the

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induction of oxidative stress (i.e., aging, inflammation, and neurodegeneration) [6]. The lung provides an excellent example of the similarities and differences between human and mouse anatomy. Similar to the human organ, the mouse lung is subdivided into lobes of lung parenchyma containing a branching bronchial tree and is vascularized by the pulmonary circulation originating from the right ventricle. There are a number of subtle variations in this general structure between species, i.e., the number of lobes on the right and left, the branching pattern, and the distribution of cartilage rings around the large airways, but the most important differences between the mouse and human lung are related to the organism’s size (airway diameter and alveolar size are naturally much smaller in the mouse) and respiratory rate. Moreover, there are important differences in the blood supply of the large airways in humans versus mice [7]. Specifically, the bronchial circulation (a branch of the high-pressure systemic circulation that arises from the aorta and intercostal arteries) supplies a miniscule proportion of the pulmonary tissue in mice (the trachea and bronchi) compared to humans; the majority of the lung parenchyma is supplied by the low-pressure, high-flow pulmonary circulation. In the mouse, these systemic blood vessels do not penetrate into the intraparenchymal airways, as they do in larger species [8]. This difference, although subtle, has important ramifications regarding the vascular supply of lung tumors which, in humans, is primarily derived from the systemic circulation [9]. These differences may also have profound consequences when modeling human diseases involving the lung vasculature. 2.3

Immunology

The adaptive immune system evolved in jawed fish about 500 million years ago, well before the evolution of mammals and the divergence of mouse and human ancestral species [10]. Many features of the adaptive immune system, including antigen recognition, clonal selection, antibody production, and immunological tolerance, have been maintained since they first arose in early vertebrates. However, the finer details of the mouse and human immune systems differ considerably, which is not surprising since these species diverged 75 million years ago [6]. While some have claimed that these differences mean that research into immunological phenomena in mice is not transferable to humans, as long as these differences are understood and acknowledged, the study of mouse immune responses can continue to be relevant. Research on mice has been vital to the discovery of key features of both innate and adaptive immune responses; for example, the first descriptions of the major histocompatibility complex, the T cell receptor, and antibody synthesis were derived from experiments performed on mice [6]. The general structure of the immune system is similar in mice and humans, with similar mediators and

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Table 1 A brief overview of the immunological differences between mice and humans Attribute

Mouse

Human

References

Proportion of leukocytes in the blood

75–90% lymphocytes 10–25% neutrophils

50–70% neutrophils 30–50% lymphocytes

[13]

Antigen presentation

Endothelial cells do not express Endothelial cells express MHC Class II and present antigen to MHC Class II, cannot activate CD4+ T cells CD4+ T cells

Costimulatory signaling

80% of CD4+ and 50% of CD8+ T cells express CD28 ICOS is not required for B cell maturation B7-H3 inhibits T cell activation

[14]

100% of CD4+ and CD8+ T cells [12] express CD28 ICOS is required for B cell [15, 16] maturation and IgM production B7-H3 promotes T cell activation [17]

Immunoglobulin IgD, IgM, IgA, IgE, IgG1, isotypes IgG2a/c, IgG2b, IgG3

IgD, IgM, IgA1, IgA2, IgE, IgG1, [12] IgG2, IgG3, IgG4

Immunoglobulin IL-4 induces IgG1 and IgE class switching

IL-4 induces IgG4 and IgE

[18]

Helper T cell differentiation

IFN-α does not activate STAT4 IFN-α induces Th1 polarization via [19] STAT4 and does not induce Th1 polarization Clear Th1/Th2 differentiation in Multiple T helper cell subsets occur [20] mice simultaneously

Responses to infection

Eradication of schistosomiasis requires a Th1 response and IFN-γ Low susceptibility to Mycobacterium tuberculosis; noncaseating granulomas; no latent infection

[21] Eradication of schistosomiasis requires a Th2 response and IgE Highly susceptible to Mycobacterium tuberculosis; caseating granulomas; latent infection is common

[22]

cell types involved in rapid, innate immune responses (complement, macrophages, neutrophils, and natural killer cells) as well as adaptive immune responses informed by antigen-presenting dendritic cells and executed by B and T cells. However, due to the anatomical and physiological differences between these species as described above, divergence in key features of the immune system, such as the maintenance of memory T cells (related to the life span of the organism) and the commensal microbiota (related to the lifestyle of the organism), has arisen [11]. Similar to what has been discovered regarding the genetics of mice and humans, i.e., broad similarities in structure but considerable differences in regulation, there are a number of known discrepancies in the regulation of innate and adaptive immunity in

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mice versus humans, including the balance of leukocyte subsets, T cell activation and costimulation, antibody subtypes and cellular responses to antibody, Th1/Th2 differentiation, and responses to pathogens (described in detail in Table 1). In addition to these differences in immune cell functions, the expression of specific genes involved in immune responses also differs, particularly those for Toll-like receptors, defensins, NK inhibitory receptors, Thy-1, and many components of chemokine and cytokine signaling; additionally, differences between mouse strains are known to exist for many of these mediators [12]. Another important consideration when using mice to perform immunological research (with a view to translating these findings to human medicine) is the availability of hundreds of strains of genetically modified mice that have enabled exquisitely detailed studies on immune cell function, regulation, and trafficking. Many of these strains involve the expression of inducible Cre or Cas9 that allow for targeted knockdown or overexpression of key immune function-related genes in specific cell types at specific moments in time. However, it is important to note that drift between mouse colonies has long been known to occur. In fact, a recent report described the fortuitous discovery of a point mutation in the natural cytotoxicity receptor 1 (NCR1) gene in the C57/Bl6 CD45.1 mouse strain, resulting in absent NCR1 expression. This mutation was found to have profound effects on the response of mice to viral infection, i.e., the mice were resistant to cytomegalovirus infection but more susceptible to influenza virus [23]. This cautionary tale highlights the importance of understanding the genetic evolution of laboratory strains of mice, the effect of these genetic and immunological changes on mouse biology, and the impact on the translation of these results to human medicine. In addition to the differences between mouse and human genetics, physiology, and immunology highlighted above, several factors must also be taken into account when performing in vitro assays using isolated mouse cells and applying these findings to our understanding of human disease. Particularly with regard to stem cell research, it should be noted that the telomeres of mouse cells are five- to tenfold longer than human telomeres, resulting in greater replicative capacity [24]. There are also important differences in the regulation of pluripotency and stem cell differentiation pathways in humans and mice [25]. Moreover, there are considerable species differences in the longevity of cultured cells; for example, mouse fibroblasts are capable of spontaneous immortalization in vitro, whereas human fibroblasts become senescent and ultimately fail to thrive in culture [26]. In summary, although there are considerable differences between mice and humans, constant improvement in the analytical techniques used to delineate these differences and their effects on whole organism and cell function have provided vital information

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and contributed to our understanding of both murine and human biology. Experimentation employing mouse models of human disease will continue to provide key insights into relevant disease mechanisms and potential drug targets for further clinical investigation. However, several important considerations must be taken into account when selecting a mouse model of human disease, as described in the following section, using mouse models of human lung disease to illustrate this point.

3

Mouse Models of Human Disease The two most salient features of a mouse model of human disease are the accuracy of its etiology (it employs a physiologically relevant method of disease induction) and its presentation (its ability to recapitulate the features of human disease). The relevance of any given mouse model can be judged on the basis of these two criteria, and there is considerable variation within mouse models of human disease in this regard. As a full assessment of the advantages and limitations of all currently available mouse models of human disease would be prohibitively long and complex, here we have elected to assess the accuracy of currently available models of human lung diseases, i.e., asthma, chronic obstructive pulmonary disease, and pulmonary fibrosis, focusing on the relevance of disease induction in these models and their ability to replicate critical features of human disease pathophysiology and response to treatment. The first and foremost notion when modeling human disease in mice is to acknowledge the species differences, which are significant [27]. As described above, genetics, anatomy, physiology, and immunology differ between mice and humans, but despite these differences, mouse models of human disease are useful and necessary, as long as data interpretation is performed appropriately.

4

Asthma An elegant example of differences between mice and humans that must be considered when designing a mouse model of human inflammatory lung disease is the key effector cell type in human asthma, i.e., mast cells. These leukocytes differ in granule composition as well as localization in the mouse and human airways [28]. Mice mostly lack mast cells in the peripheral lung [29], whereas humans have numerous mast cells of multiple subpopulations in the alveolar parenchyma [30]. Another example is anatomy: in contrast to humans, mice lack an extensive pulmonary circulation, which may have significant effects on leukocyte adhesion and migration, and subsequently inflammation [31]. Still, as long as these differences are taken into consideration, mouse models can be

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powerful tools in the discovery and exploitation of new targets for the treatment of human disease. The World Health Organization (WHO) defines asthma as a chronic disease characterized by recurrent attacks of breathlessness and wheezing, which may vary in severity and frequency from person to person. The disease is characterized by airway hyperresponsiveness, airway smooth muscle thickening, increased mucus secretion and collagen deposition, as well as prominent inflammation affecting both large and small airways [32]. Nowadays, it is recognized that asthma is not a single homogenous disease but rather several different phenotypes united by similar clinical symptoms [32, 33]. Only a few animal species develop asthma naturally, including cats and horses [34, 35], whereas mice do not [31]. However, mice can be manipulated to develop a type of allergic airway inflammation, which is similar in many ways to the human disease, in response to different aeroallergens [36]. Importantly, these models are capable of recapitulating only the allergic type of human asthma and have less relevance for other types of asthma (i.e., endotypes induced by medication, obesity, and air pollution). As with many human diseases, asthma has a complex and multifaceted etiology, where environmental factors, genetic susceptibility, and microbial colonization all contribute; thus, it is important to take strain differences into consideration. Generations of inbreeding have created mouse strains that differ not only in coat color and disposition but also from a physiological, immunological, and genetic perspective. Different strains may be more susceptible to allergic airway inflammation or pulmonary fibrosis, whereas others are more or less resistant. Choosing the right strain to model a specific disease or pathologic event is thus essential. The most widely used strains for models of allergic airway inflammation are BALB/c and C57BL/6. These strains differ regarding the type of immune response mounted to an inhaled allergen: C57BL/6 is generally considered a TH1-skewed strain, whereas BALB/c is regarded as a TH2-skewed strain [36]. Due to their strong TH2response, and subsequent development of robust asthmatic responses, BALB/c has been commonly used to model asthma [37]. However, most humans do not express such a strongly TH2-skewed immune system, suggesting this strain may not be the best model of human disease; instead, C57BL/6 may be more suitable as immune responses in this strain are more similar to those of atopic human subjects [37]. Furthermore, as C57BL/6 is the most commonly used strain for the development of genetically manipulated mice, using these mice allows for very specific investigations into disease pathology; thus, this strain is increasingly used in models of human lung disease.

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4.1

Ovalbumin

Besides the genetic differences in the mouse strains used in these models, the etiology (the method of disease induction) of commonly used models of asthma is highly variable. In humans with allergic asthma, environmental allergen exposure occurs at the airway mucosa; the immune response is coordinated in the bronchopulmonary lymph nodes, and the T cells, macrophages, and eosinophils recruited as part of this response travel to the lung where they mediate the cardinal features of asthma: airway inflammation, structural remodeling of the airway wall, and airway hyperreactivity [38]. Ideally, these features should be found in a physiologically relevant mouse model of asthma. However, for the sake of cost and convenience, early mouse models of asthma used the surrogate protein ovalbumin (OVA) [31] rather than an environmental allergen to induce an immune response, which also requires the use of a powerful TH2-polarizing adjuvant such as alum delivered via the intraperitoneal route, followed by OVA nebulization—a clear divergence from the etiology of human asthma [36]. In terms of disease presentation, mice develop some hallmarks of asthma, including airway eosinophilic inflammation, goblet cell metaplasia, and increased airway smooth muscle density [31]. After the cessation of OVA exposure, most of the remodeling resolves, although some structural alterations remain up to 1 month after the last challenge [39]. Based on these attributes, the OVA model is primarily a model to investigate the initiation of inflammation, rather than the chronic progression and maintenance of inflammation [31]. A clear advantage with the OVA model is the number of studies where it is used; both the pros and cons are familiar. It is easy to find a suitable protocol, and the model is readily accessible and flexible regarding the number of sensitizations and allergen doses. The model is relatively easy to reproduce, as OVA and different adjuvants are easily obtained. However, the resolution of remodeling following the cessation of allergen provocations is a disadvantage, as is the practical problem with the nebulization of an allergen—it ends up in the mouse’s coat and is ingested during grooming, potentially resulting in systemic exposure (this is particularly relevant in models employing systemic, intraperitoneal sensitization). In addition, concerns have been raised against the use of adjuvants to induce the immunological response, as well as the clinical relevance of OVA as an allergen, which have driven the development of more clinically relevant allergens and models [31].

4.2

House Dust Mite

The common environmental aeroallergen house dust mite (HDM) extract is increasingly used to initiate disease in mouse models of allergic airway inflammation, as it is a common human allergen (around 50% of asthmatics are sensitized to HDM [40]) that evokes asthma attacks and other allergic responses in susceptible individuals. In addition, HDM has inherent allergenic properties, likely

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due to components with protease activity [40], so there is no need to use an adjuvant, thus improving the etiological similarity of these models with the clinical situation [41]. In contrast to OVA, prolonged exposure of HDM (up to 7 weeks) induces asthma-like severe airway inflammation with prominent eosinophilia, severe hyperreactivity to methacholine, and robust remodeling of the airway wall [41], i.e., the presentation of chronic respiratory HDM exposure in mice effectively recapitulates the key features of human allergic asthma. Importantly, the airway structural changes induced by chronic HDM exposure, such as increased collagen deposition, airway smooth muscle thickening, and microvascular alterations, persist for at least 4 weeks after the cessation of HDM exposure [42], another commonality with human asthma in which airway remodeling is currently considered to be irreversible. Thus, the advantages of using HDM as the allergen in mouse models of asthma are the clinical relevance of the allergen [43] and the route of delivery via the respiratory tract. Moreover, studies have shown that the type of inflammation and characteristics of tissue remodeling are relatively similar to those seen in human asthmatics [35, 41, 43]. One disadvantage is the complexity of HDM extract; as a consequence of this complexity, variations exist in some components between batches, particularly regarding the content of lipopolysaccharide, so reproducibility in these studies may be problematic. 4.3 Cockroach, Aspergillus, and Other Model Allergens

With similarity to HDM, these models were developed to be as clinically relevant as possible, as many patients suffer from allergy toward cockroach allergen, molds, and other environmental irritants. A common feature of these allergens is their complex nature, as they commonly consist of a mix of different allergic epitopes and fragments. This complexity is most likely why the immunological reaction in mice is relatively similar to that seen in asthmatics [44]. Cockroach allergen (CRA) is a common allergen, known to induce asthma in susceptible individuals; thus, it shares with HDM the advantage of being highly clinically relevant [45]. CRA induces peribronchial inflammation with significant eosinophilic inflammation and transient airway hyperresponsiveness, both of which can be increased by repeated administrations of the allergen [45]. Colonization of the airways with Aspergillus fumigatus is the cause of allergic bronchopulmonary aspergillosis (ABPA), a disease where the lungs are colonized by the fungus, but allergens from Aspergillus fumigatus can also induce asthma similar to other allergens [46]. The reaction to Aspergillus allergens is robust, and often no adjuvants are needed to elicit inflammation [46]. In addition to Aspergillus, other fungi such as Penicillium and Alternaria can also induce asthma in humans and have been used to model disease in mice [47]. A common difficulty with these allergens is the method of administration, as the physiological route is believed to be the

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inhalation of dry allergens; mimicking this route with a nebulizer introduces the risk of the animals ingesting the allergen and thus causing systemic responses [47]. 4.4 Modeling Asthma Exacerbations

Exacerbations of asthma are defined as the worsening of symptoms, prompting an adjustment in treatment, and are believed to be associated with increased inflammation in the distal airways. Clinically, exacerbations are believed to be induced by infections (most common), allergen exposure, or pollutants, which can be modeled in different ways [48, 49]: 1. Infections with viruses and bacteria or exposure to proteins/ DNA/RNA derived from these microbes. 2. Administration of a high dose of allergen in a previously sensitized animal. 3. Exposure to environmental pollutants, such as diesel exhaust or ozone. Modeling exacerbations adds a layer of complexity, as robust ongoing allergic airway inflammation needs to be established first, before challenge with the exacerbating agent. Both the OVA and HDM models are used in this respect, and in both cases chronic protocols extending for several weeks before triggering an exacerbation have been used [48].

5

Chronic Obstructive Pulmonary Disease Chronic obstructive pulmonary disease (COPD) is characterized by chronic airway obstruction, in contrast to asthma where the obstruction is reversible (particularly in response to bronchodilator treatment). Clinically, in COPD, chronic bronchitis and emphysema can occur either separately or in combination. COPD is almost always associated with either first- or secondhand tobacco smoking or in rare cases with a deficiency in the production of α1antitrypsin (a serpin that prevents elastin breakdown as a result of neutrophil degranulation) [50]. The etiology of COPD is highly complex and is believed to develop after many years of smoking in combination with other known factors such as genetic susceptibility or environmental factors [51]. In similarity to asthma, inflammation is a major component in COPD, but the leukocyte profile is very different: the most prominent players in COPD-related inflammation are neutrophils and, to some degree, macrophages [51]. Due to the complex etiology of COPD, it is difficult to recapitulate all aspects of this disease in a single model, so in most cases, the aim is to induce COPD-like lesions by exposing mice to tissue-damaging substances (usually cigarette smoke) or to mimic emphysema by the administration of tissue-degrading enzymes [27, 51].

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5.1

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Cigarette Smoke

5.2 Protease Instillation

6

Clearly, mice do not smoke cigarettes on their own, so to model COPD by cigarette smoke (CS) inhalation, the mice need to be exposed to unfiltered CS in an induction chamber; moreover, in an attempt to better model the chronic aspects of COPD, this needs to be performed for a prolonged period of time. Mice are very tolerant to CS, but eventually (over a period of several weeks), CS induces pulmonary neutrophilic inflammation that is associated with some degree of tissue degradation and destruction [51]. An important advantage of this model is the fact that CS is the actual irritant responsible for disease in humans, and mice develop several features similar to the clinical disease, making this model highly clinically relevant [27]. A significant drawback is the self-limitation of the model—the pathological changes do not progress after the cessation of CS exposure [51]. Furthermore, the exposure time needed for mice to develop COPD-like pathology is extensive, i.e., studies have shown that an exposure protocol of 5 days per week for a minimum of 3 months is needed to generate robust structural changes to the lung [52]. The pathological image in COPD is complex and varies greatly between patients, commonly encompassing chronic bronchitis and bronchiolitis, emphysema, fibrosis, and airway obstruction. Although mice develop some of these symptoms when exposed to CS, they do not develop all the symptoms of human disease; thus, CS has advantages as a model but fails to mimic the complexity of the clinical situation and disease presentation [27]. Other models of COPD rely on the administration of proteases (protein-degrading enzymes) that are believed to be involved in the pathology of this disease in a subset of patients, such as elastindegrading elastase. This approach mimics the emphysematous changes seen in COPD, but the pathological process underlying tissue destruction is likely very different compared to the clinical situation [51], as very few patients show evidence of elastase dysregulation [27]. However, if the aim of the study is to investigate the general effect of protease-induced tissue destruction and regeneration, then this is a highly relevant method [51]. Some studies on COPD have also used genetically modified animals, such as mice overexpressing collagenase, which results in tissue destruction without inflammation or fibrosis with an end result fairly similar to the type of emphysema observed in COPD [53].

Pulmonary Fibrosis Pulmonary fibrosis, the accumulation of fibrotic tissue within the alveolar parenchyma, is merely a symptom of disease, and the etiology of this pathology in humans varies greatly [54]. The

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most enigmatic class is perhaps the idiopathic interstitial pneumonias, especially idiopathic pulmonary fibrosis (IPF). IPF is a debilitating and progressive disease with a grave prognosis, characterized by progressive fibrosis believed to reflect aberrant tissue regeneration [55]. As the reason behind this defective repair is unknown, although a combination of immunological, genetic, and environmental factors are suspected, it is very difficult to model disease in a clinically relevant fashion [56]. The most common method used to model pulmonary fibrosis in mice is administration of the chemotherapeutic agent bleomycin; this agent is known to cause pulmonary fibrosis in humans as well, but this may not accurately reflect the true etiology of most cases of human disease. The strain of choice is C57BL/6, as it is prone to developing pulmonary fibrosis, whereas BALB/c is relatively resistant, a feature believed to reflect the cytokine response following cellular stress and damage [57]. Bleomycin administration can be performed locally or systemically, producing very different results. 6.1 Local Bleomycin Administration

The most common model of pulmonary fibrosis is a single intranasal or intratracheal administration of bleomycin, with analysis 3 to 4 weeks later. During this time, the drug causes acute tissue damage in a restricted area of the lung (where the solution ends up during administration), followed by intense inflammation in this area and subsequent fibrosis, which gradually resolves within weeks. However, if older mice are used, the fibrosis will persist longer than in younger mice, which is in accordance with clinical IPF, where the majority of the patients are 65 years of age or older [56, 58]. A great advantage of this model is how well-characterized it is. In addition, local administration is labor-effective, as only one administration is required and the result is highly reproducible. The fibrosis is robust, only affects the lungs, and the accumulation of extracellular matrix can be easily measured using standard techniques [58]. Furthermore, as it is used throughout the world, studies performed in different labs and by different groups can be compared relatively easily. Unfortunately, the intense pulmonary inflammation may be lethal, and fatalities are to be expected with this model [59], representing an important ethical limitation. Furthermore, fibrosis is heterogeneous—it develops where the bleomycin solution is deposited. The solution usually deposits within the central lung, a localization that is not in agreement with the clinical situation where fibrosis is located in the more distal regions of the lung parenchyma. In addition, the fibrosis that develops as a result of severe tissue damage is self-limiting and reversible, unlike what is observed clinically [58]. The severe degree of tissue damage induced by bleomycin may in fact be more relevant for modeling acute lung injury (ALI) or acute respiratory distress syndrome (ARDS).

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6.2 Systemic Bleomycin Administration

Bleomycin can also be administered systemically, through intravenous or subcutaneous injection. In contrast to local administration, this route requires multiple administrations and is thus more laborintensive [58]. Some studies have described the usage of osmotic mini-pumps, where bleomycin is slowly administered over a short period of time, and then fibrosis continues to develop over subsequent weeks [60]. Irrespective of the route of delivery, systemic administration results in more homogenous fibrosis, affecting the entire lung through the pulmonary endothelium and persisting much longer than following local administration [61]. The main advantages of systemic administration are that inflammation is limited, while the fibrosis is more apparent and displays a more distal pattern, all of which mimics the clinical situation relatively well. The multiple administrations allow for lower doses with each injection; this is less stressful to the animals and results in little to no mortality [61] and is thus more ethically acceptable. A major disadvantage with this model is that it takes time for fibrosis to develop [58], which may be the reason it is used relatively scarcely, and thus the pathological development is less well-understood. In addition, as IPF is a local disease, local administration of the etiologic agent may better mimic the clinical reality [56].

6.3 Fluorescein Isothiocyanate Administration

The administration of fluorescein isothiocyanate (FITC) induces focal inflammation, primarily involving mononuclear cells and neutrophils, and localizes in areas where the FITC solution is deposited [58]. Antibodies against FITC can be detected after 1 week, and the fibrosis persists for up to 5 months after instillation [58]. The benefits of this model are mainly related to the persistent fibrosis that does not appear to be self-limiting, thus reflecting the clinical situation, and it is also very easy to determine which part of the lung has been exposed to FITC, as the molecule is fluorescent [58]. It is also an advantage that both C57BL/6 and BALB/c mice are susceptible and develop fibrosis following FITC administration [56]. The disadvantages of this model include profound variability due to differences between batches of FITC, as well as in the method used to prepare the solution before instillation. Importantly, given the characteristics of the etiologic agent used to induce this model of IPF, this model is considered a very artificial system with limited clinical relevance [56].

6.4 TGF-β Overexpression

Adenovirus vectors have been used to overexpress the pro-fibrotic cytokine transforming growth factor (TGF)-β, which results in pulmonary fibrosis. As TGF-β overexpression in the lungs is known to be crucial in the development of fibrosis in humans [62], this model mimics an important feature of disease etiology. However, the delivery system has some drawbacks, as the virus itself initiates an immune response. Moreover, adenoviruses display

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significant tropism for epithelial cells and rarely infect other cell types such as fibroblasts [58], which are the cells meant to be targeted in this model. As TGF-β has major effects on fibroblast biology, the main feature of this model is the effect of epitheliumderived TGF-β on fibroblasts and myofibroblasts, resulting in the deposition of ECM proteins and areas of dense fibrosis [63]. An advantage of this model is the relatively low degree of inflammation, as well as what appears to be a direct effect on fibroblasts/ myofibroblasts [63], which is in accordance with the clinical situation (as we understand it today). 6.5

7

Silica

Silica administration induces a similar pathology in mouse lungs as in humans exposed to silica, and as is also observed in human silicainduced fibrosis, structural remodeling persists when administration is halted [56]. Following the administration of silica particles, fibrotic nodules develop in mouse lungs, with considerable resemblance to the human lesions that develop after exposure to mineral fibers [56]. The fibrotic response is accompanied by a limited inflammatory response, and different pro-fibrotic cytokines such as TGF-β, platelet-derived growth factor, and IL-10 are involved in disease development, which is in accordance with the clinical situation [56]. Another advantage is that nodules develop around silica fibers, and these fibers are easy to identify by light microscopy. The response in this model is strain-dependent, with C57BL/6 mice being the most susceptible. The main drawbacks are the time required to establish disease, i.e., 30–60 days, and the need for special equipment to aerosolize the silica particles. However, since the route of administration, the driving etiologic agent, and the resulting pathobiology are all similar to the characteristics of this subtype of pulmonary fibrosis [56, 58], the silica exposure model can be considered to have very good clinical relevance.

What Does the Future Hold for Mouse Models of Human Disease? Medical research using experimental animals (not only mice but other animals including rats, guinea pigs, zebrafish, and fruit flies) has greatly contributed to many important scientific and medical advances in the past century and will continue to do so into the near future. These advances have contributed to the development of new medicines and treatments for human disease and have therefore played a vital role in increasing the human life span and improving quality of life. Despite the acknowledged benefits of performing research using experimental animals, a number of considerations must be made before embarking on this type of research. Of course, the financial aspects of conducting this type of work are an important limitation, as the costs of purchasing and housing mice can be prohibitive, especially when genetically modified mice and colony

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maintenance are required for the study. The practicalities of working with animals such as mice may also be an issue, as this type of work requires specialized facilities, equipment, and staff to ensure studies are carried out in a manner that is safe for both the researchers and the animals. Moreover, as discussed in detail in this chapter, the relevance of the selected animal model to human disease must be carefully evaluated to ensure that these experiments provide robust results that are translatable to human health and disease. Another important and demanding aspect of biomedical research using animals is the ethics of imposing pain and suffering on live animals. Although there has been a considerable reduction in the numbers of animals used in research in the last 30 years, animal research remains a vital part of biomedical research. However, no responsible scientist wants to cause unnecessary suffering in experimental animals if it can be avoided, so scientists have accepted controls on the use of animals for medical research. In the UK, this ethical framework has been enshrined in law, i.e., the Animals (Scientific Procedures) Act 1986. This legislation requires that applications for a project license to perform research involving the use of “protected” animals (including all vertebrates and cephalopods) must be fully assessed with regard to any harm imposed on the animals. This involves a detailed examination of the proposed procedures and experiments, and the numbers and types of animal used, with robust statistical calculations to support these numbers. The planned studies are then considered in light of the potential benefits of the project. Both within and outside the UK, approval for a study involving protected animals also requires an internal ethical review process, usually conducted by the research institution where the work is taking place, with the aim of promoting animal welfare by ensuring the work will be carried out in an ethical manner and that the use of animals is justified. Additionally, the UK has a national animal use reduction strategy supported by the National Centre for the Replacement, Refinement and Reduction of Animals in Research (NC3Rs; London, UK). This consortium was established in 2004 to promote and develop high-quality research that takes the principles of replacement, refinement, and reduction (the 3Rs) into account. 7.1

Replacement

Replacement strategies often involve the use of alternative, non-protected species (e.g., zebrafish, fruit flies, flatworms) and in vitro correlates (two-dimensional cell culture or threedimensional organoids containing multiple cell types) to test hypotheses and assess the effects of therapeutic interventions. The main obstacle with studies on non-protected animals is the difficulty of accurately mimicking the complex physiological systems involved in human health and disease, as described in detail above. For example, the fruit fly Drosophila melanogaster is an excellent

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model organism for studies on genetic diseases, aging, and pathogen-borne illnesses but may be less relevant for studies on complex lung diseases. Importantly, model organisms such as fruit flies, zebrafish, and flatworms do not possess lungs, which somewhat limits the translatability of research on these animals in the field of respiratory disease. As such, it is likely that rodents will remain the model organism of choice for studies into lung disease for some time to come. There has been considerable progress recently in imitating single organs such as the liver, lung, and brain in vitro using multiple cell types and a physical scaffold. As an important advantage, these in vitro tests have replaced a large number of rodents in initial drug discovery experiments, while also speeding up the process [64]. These studies still require further refinement and validation to establish them as suitable models for an entire organ; importantly, these in vitro organoids cannot take into account interactions between organ systems in complex, multisystem diseases such as COPD. 7.2

Refinement

Refinement involves selecting the most clinically relevant model for the disease available, informed by the discussion above on closely recapitulating the etiologic agent and disease pathobiology associated with clinical cases. Another important factor is refining the management of pain. An assessment of the procedures used and the effects of the substance on the animal, as well as the degree of handling, restraint, and analgesia, are other important aspects of refinement. This standard of animal care is achieved through strict regulations and controls on how personnel are trained to carry out experiments on live animals. Adequate training is an important aspect of refinement and should be reviewed and improved on an ongoing basis. Moreover, refinement can be achieved by improving animal housing by environmental enrichment, e.g., providing a place for mice to hide in the cage and housing social animals such as mice in appropriate-sized groups. These simple changes can improve the physiological and behavioral status of research animals; this not only increases animal well-being but also contributes to the quality of the experimental results by reducing stress levels.

7.3

Reduction

The 3Rs aspect of reduction focuses on the statistical power of experiments and by following the Animal Research: Reporting of In Vivo Experiments (ARRIVE) guidelines, originally published in PLOS Biology in 2010. These guidelines provide a framework to improve the reporting of research performed on live animals by maximizing the quality of the scientific data and by minimizing unnecessary studies. The ARRIVE guidelines provide a checklist of aspects that must be considered in good quality research using live animals. The guidelines are most appropriate for comparative studies involving two or more groups of experimental animals with at least one control group, but they also apply to studies involving

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drug dosing in which a single animal is used as its own control (within-subject experiments). The guidelines provide recommendations on what should be considered when preparing to report on the results of experiments involving live animals, i.e., by providing a concise but thorough background on the scientific theory and why and how animals were used to test a hypothesis, a statement on ethical approvals and study design including power and sample size calculations, a clear description of the methods used to ensure repeatability, objective measurements of outcomes and adverse effects, and interpretation of the results in light of the available literature and the limitations of the study. In addition to the positive impact of the ARRIVE guidelines on reducing the number of animals used in experiments, this checklist provides an easy-tofollow roadmap on what is required for good quality reporting of experimental results.

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Conclusion In conclusion, the use of animals in research will continue to be an important aspect of medical research, and these procedures can be ethically justified provided the proper controls are in place. The benefits of animal research have been vital to the progress of medical science; abandoning these studies would have severe negative consequences on human health. By considering aspects such as the 3Rs and the ARRIVE guidelines in planning experiments involving live animals, the number of animals used and suffering of these animals for the benefit of human health can be minimized. This requires a strong regulatory framework such as that found in the UK and many other countries, as well an ongoing public debate on the advantages and limitations of animal experimentation.

References 1. Vandenbergh J (2000) Use of house mice in biomedical research. ILAR J 41:133–135 2. Hedrich H (ed) (2012) The laboratory mouse. Academic Press, Cambridge 3. Yue F, Cheng Y, Breschi A, Vierstra J, Wu W et al (2014) A comparative encyclopedia of DNA elements in the mouse genome. Nature 515:355–364 4. Cheng Y, Ma Z, Kim BH, Wu W, Cayting P et al (2014) Principles of regulatory information conservation between mouse and human. Nature 515:371–375 5. Bodenreider O, Hayamizu TF, Ringwald M, De Coronado S, Zhang S (2005) Of mice and men: aligning mouse and human anatomies. AMIA Annu Symp Proc 2005:61–65

6. Perlman RL (2016) Mouse models of human disease: an evolutionary perspective. Evol Med Public Health 2016(1):170–176 7. Townsley MI (2012) Structure and composition of pulmonary arteries, capillaries, and veins. Compr Physiol 2:675–709 8. Mitzner W, Lee W, Georgakopoulos D, Wagner E (2000) Angiogenesis in the mouse lung. Am J Pathol 157:93–101 9. Yuan X, Zhang J, Ao G, Quan C, Tian Y, Li H (2012) Lung cancer perfusion: can we measure pulmonary and bronchial circulation simultaneously? Eur Radiol 22:1665–1671 10. Flajnik MF, Kasahara M (2010) Origin and evolution of the adaptive immune system: genetic events and selective pressures. Nat Rev Genet 11:47–59

Mouse Models of Human Disease 11. Bailey M, Christoforidou Z, Lewis MC (2013) The evolutionary basis for differences between the immune systems of man, mouse, pig and ruminants. Vet Immunol Immunopathol 152:13–19 12. Mestas J, Hughes CC (2004) Of mice and not men: differences between mouse and human immunology. J Immunol 172:2731–2738 13. Doeing DC, Borowicz JL, Crockett ET (2003) Gender dimorphism in differential peripheral blood leukocyte counts in mice using cardiac, tail, foot, and saphenous vein puncture methods. BMC Clin Pathol 3:3 14. Choo JK, Seebach JD, Nickeleit V, Shimizu A, Lei H, Sachs DH, Madsen JC (1997) Species differences in the expression of major histocompatibility complex class II antigens on coronary artery endothelium: implications for cellmediated xenoreactivity. Transplantation 64:1315–1322 15. McAdam AJ, Greenwald RJ, Levin MA, Chernova T, Malenkovich N et al (2001) ICOS is critical for CD40-mediated antibody class switching. Nature 409:102–105 16. Grimbacher B, Hutloff A, Schlesier M, Glocker E, Warnatz K et al (2003) Homozygous loss of ICOS is associated with adultonset common variable immunodeficiency. Nat Immunol 4:261–268 17. Chapoval AI, Ni J, Lau JS, Wilcox RA, Flies DB et al (2001) B7-H3: a costimulatory molecule for T cell activation and IFN-gamma production. Nat Immunol 2:269–274 18. Zhang Y, Fear DJ, Willis-Owen SA, Cookson WO, Moffatt MF (2016) Global gene regulation during activation of immunoglobulin class switching in human B cells. Sci Rep 6:37988 19. Farrar JD, Smith JD, Murphy TL, Leung S, Stark GR, Murphy KM (2000) Selective loss of type I interferon-induced STAT4 activation caused by a minisatellite insertion in mouse Stat2. Nat Immunol 1:65–69 20. Schmitt N, Ueno H (2015) Regulation of human helper T cell subset differentiation by cytokines. Curr Opin Immunol 34:130–136 21. Pearce EJ, Sher A (1991) Functional dichotomy in the CD4+ T cell response to Schistosoma mansoni. Exp Parasitol 73:110–116 22. Dharmadhikari AS, Nardell EA (2008) What animal models teach humans about tuberculosis. Am J Respir Cell Mol Biol 39:503–508 23. Jang Y, Gerbec ZJ, Won T, Choi B, Podsiad A, BM B, Malarkannan S, Laouar Y (2018) Cutting edge: check your mice-a point mutation in the Ncr1 locus identified in CD45.1 congenic mice with consequences in mouse susceptibility to infection. J Immunol 200:1982–1987

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24. Calado RT, Dumitriu B (2013) Telomere dynamics in mice and humans. Semin Hematol 50:165–174 25. Ernst M, Abu Dawud R, Kurtz A, Schotta G, Taher L, Fuellen G (2015) Comparative computational analysis of pluripotency in human and mouse stem cells. Sci Rep 5:7927 26. Macieira-Coelho A, Azzarone B (1988) The transition from primary culture to spontaneous immortalization in mouse fibroblast populations. Anticancer Res 8:669–676 27. Williams K, Roman J (2016) Studying human respiratory disease in animals – role of induced and naturally occurring models. J Pathol 238:220–232 28. Bischoff SC (2007) Role of mast cells in allergic and non-allergic immune responses: comparison of human and murine data. Nat Rev Immunol 7:93–104 29. Schmit D, Le DD, Heck S, Bischoff M, Tschernig T et al (2017) Allergic airway inflammation induces migration of mast cell populations into the mouse airway. Cell Tissue Res 369:331–340 30. Andersson CK, Mori M, Bjermer L, Lofdahl CG, Erjefalt JS (2009) Novel site-specific mast cell subpopulations in the human lung. Thorax 64:297–305 31. Mullane K, Williams M (2014) Animal models of asthma: reprise or reboot? Biochem Pharmacol 87:131–139 32. Holgate ST, Wenzel S, Postma DS, Weiss ST, Renz H, Sly PD (2015) Asthma. Nat Rev Dis Primers 1:15025 33. Ray A, Oriss TB, Wenzel SE (2015) Emerging molecular phenotypes of asthma. Am J Physiol Lung Cell Mol Physiol 308:L130–L140 34. Mueller RS, Janda J, Jensen-Jarolim E, Rhyner C, Marti E (2016) Allergens in veterinary medicine. Allergy 71:27–35 35. O’Brien R, Ooi MA, Clarke AH, Thomas WR (1996) Immunologic responses following respiratory sensitization to house dust mite allergens in mice. Immunol Cell Biol 74:174–179 36. Shin YS, Takeda K, Gelfand EW (2009) Understanding asthma using animal models. Allergy Asthma Immunol Res 1:10–18 37. Durham CG, Schwiebert LM, Lorenz RG (2013) Use of the cockroach antigen model of acute asthma to determine the immunomodulatory role of early exposure to gastrointestinal infection. Methods Mol Biol 1032:271–286 38. Islam SA, Luster AD (2012) T cell homing to epithelial barriers in allergic disease. Nat Med 18:705–715

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39. Rydell-Tormanen K, Uller L, Erjefalt JS (2009) Allergic airway inflammation initiates long-term vascular remodeling of the pulmonary circulation. Int Arch Allergy Immunol 149:251–258 40. Calderon MA, Linneberg A, Kleine-Tebbe J, De Blay F, Hernandez Fernandez de Rojas D, Virchow JC, Demoly P (2015) Respiratory allergy caused by house dust mites: what do we really know? J Allergy Clin Immunol 136:38–48 41. Johnson JR, Wiley RE, Fattouh R, Swirski FK, Gajewska BU et al (2004) Continuous exposure to house dust mite elicits chronic airway inflammation and structural remodeling. Am J Respir Crit Care Med 169:378–385 42. Rydell-Tormanen K, Johnson JR, Fattouh R, Jordana M, Erjefalt JS (2008) Induction of vascular remodeling in the lung by chronic house dust mite exposure. Am J Respir Cell Mol Biol 39:61–67 43. Doras C, Petak F, Bayat S, Baudat A, Von Garnier C, Eigenmann P, Habre W (2017) Lung responses in murine models of experimental asthma: value of house dust mite over ovalbumin sensitization. Respir Physiol Neurobiol 247:43–51 44. Sarpong SB, Zhang LY, Kleeberger SR (2003) A novel mouse model of experimental asthma. Int Arch Allergy Immunol 132:346–354 45. Campbell EM, Kunkel SL, Strieter RM, Lukacs NW (1998) Temporal role of chemokines in a murine model of cockroach allergen-induced airway hyperreactivity and eosinophilia. J Immunol 161:7047–7053 46. Kurup VP, Grunig G (2002) Animal models of allergic bronchopulmonary aspergillosis. Mycopathologia 153:165–177 47. Templeton SP, Buskirk AD, Green BJ, Beezhold DH, Schmechel D (2010) Murine models of airway fungal exposure and allergic sensitization. Med Mycol 48:217–228 48. Kumar RK, Herbert C, Foster PS (2016) Mouse models of acute exacerbations of allergic asthma. Respirology 21:842–849 49. Maltby S, Tay HL, Yang M, Foster PS (2017) Mouse models of severe asthma: understanding the mechanisms of steroid resistance, tissue remodelling and disease exacerbation. Respirology 22:874–885 50. Bashir A, Shah NN, Hazari YM, Habib M, Bashir S et al (2016) Novel variants of SERPIN1A gene: interplay between alpha1antitrypsin deficiency and chronic obstructive pulmonary disease. Respir Med 117:139–149 51. Groneberg DA, Chung KF (2004) Models of chronic obstructive pulmonary disease. Respir Res 5:18

52. Bartalesi B, Cavarra E, Fineschi S, Lucattelli M, Lunghi B et al (2005) Different lung responses to cigarette smoke in two strains of mice sensitive to oxidants. Eur Respir J 25:15–22 53. D’Armiento J, Dalal SS, Okada Y, Berg RA, Chada K (1992) Collagenase expression in the lungs of transgenic mice causes pulmonary emphysema. Cell 71:955–961 54. Knudsen L, Ruppert C, Ochs M (2017) Tissue remodelling in pulmonary fibrosis. Cell Tissue Res 36:607–626 55. Martinez FJ, Collard HR, Pardo A, Raghu G, Richeldi L et al (2017) Idiopathic pulmonary fibrosis. Nat Rev Dis Primers 3:17074 56. Tashiro J, Rubio GA, Limper AH, Williams K, Elliot SJ et al (2017) Exploring animal models that resemble idiopathic pulmonary fibrosis. Front Med (Lausanne) 4:118 57. Walkin L, Herrick SE, Summers A, Brenchley PE, Hoff CM et al (2013) The role of mouse strain differences in the susceptibility to fibrosis: a systematic review. Fibrogenesis Tissue Repair 6:18 58. Moore BB, Hogaboam CM (2008) Murine models of pulmonary fibrosis. Am J Physiol Lung Cell Mol Physiol 294:L152–L160 59. Manitsopoulos N, Nikitopoulou I, Maniatis NA, Magkou C, Kotanidou A, Orfanos SE (2018) Highly selective endothelin-1 receptor A inhibition prevents bleomycin-induced pulmonary inflammation and fibrosis in mice. Respiration 95:122–136 60. Yatomi M, Hisada T, Ishizuka T, Koga Y, Ono A et al (2015) 17(R)-resolvin D1 ameliorates bleomycin-induced pulmonary fibrosis in mice. Physiol Rep 3:e12628 61. Rydell-Tormanen K, Andreasson K, Hesselstrand R, Risteli J, Heinegard D et al (2012) Extracellular matrix alterations and acute inflammation; developing in parallel during early induction of pulmonary fibrosis. Lab Investig 92:917–925 62. Gauldie J, Kolb M, Ask K, Martin G, Bonniaud P, Warburton D (2006) Smad3 signaling involved in pulmonary fibrosis and emphysema. Proc Am Thorac Soc 3:696–702 63. Sime PJ, Xing Z, Graham FL, Csaky KG, Gauldie J (1997) Adenovector-mediated gene transfer of active transforming growth factorbeta1 induces prolonged severe fibrosis in rat lung. J Clin Invest 100:768–776 64. Festing S, Wilkinson R (2007) The ethics of animal research. Talking Point on the use of animals in scientific research. EMBO Rep 8:526–530

Chapter 2 Optimizing the Cell Culture Microenvironment Ivan Bertoncello Abstract The survival, proliferation, and differentiation of cells in culture are determined not only by their intrinsic potential but also by cues provided by the permissive or restrictive microenvironment in which they reside. The robustness and reproducibility of cell culture assays and endpoints relies on the stability of that microenvironment and vigilant attention to the control of variables that affect cell behavior during culture. These often underappreciated variables include, but are not limited to, medium pH and buffering, osmolarity, composition of the gas phase, the timing and periodicity of refeeding and subculture, and the impact of fluctuations in temperature and gas phase composition on frequent opening and closing of incubator doors. This chapter briefly describes the impact of these and other variables on the behavior of cultured cells. Key words Cell culture, Culture medium, Medium pH, Medium buffering, Oxygen tension, Incubation conditions

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Introduction The niche hypothesis first articulated by Schofield [1] to explain hematopoietic regulation posits that the regenerative potential of a cell population is defined in context: by its intrinsic potential and by its interaction with the microenvironment in which it resides [2, 3]. The dynamic interaction of regenerative cells with soluble and insoluble factors, signaling pathways, accessory cells, and matrix proteins comprising their microenvironment regulates their developmental potential, proliferation, and differentiation (Fig. 1). Conversely, reciprocal signaling mediated by proliferating and differentiating cells is able to modify their niche microenvironment, potentially leading to dysregulated cell growth [4–6]. The degree of difficulty encountered in deconstructing and elucidating these regulatory processes to develop informative and instructive in vitro cell culture models cannot be underestimated. In 1993, Quesenberry [7] estimated that there were at least 2.248 possible combinations of 40 known hemolymphopoietic cytokines with order being important and without allowing for dose-

Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940, https://doi.org/10.1007/978-1-4939-9086-3_2, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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Fig. 1 The developmental potential, proliferation, and differentiation of cultured cells are determined by their spatial orientation, their dynamic interaction, and their elaboration of soluble and insoluble stimulatory and inhibitory factors and matrix proteins that comprise their microenvironment

dependent differences in activity or target cell heterogeneity. Variables including cell adhesion and cell geometry [8], cell polarity [9], and extracellular matrix stiffness [3, 10] have also been identified as significant factors that influence the developmental fate, proliferation, and differentiation of different cell types. Over the years, reductionist experimentation exploiting powerful biochemical and molecular genetic technologies and cell separative strategies has enabled the progressive refinement of cell culture technologies. However, the optimization of cell culture assays and the identification and elimination of sources of experimental variability remain a work in progress [11]. Historically, the development of protocols for the maintenance and propagation of specific cell types has focused on the formulation and optimization of chemically defined tissue culture media that meet their requirement for unique combinations of nutrients, trace elements, growth factors, and hormones in order to thrive in vitro [11–13]. Often, optimal growth requires the addition of uncharacterized supplements such as fetal calf serum, conditioned media, tissue extracts, extracellular matrices, or hydrogels to chemically defined media. In addition to variable concentrations of known factors, these supplements contain a plethora of ill-defined peptides, proteins, and organic and inorganic molecules potentially capable of stimulating or inhibiting target cells. In one study [14], in-depth proteomic analysis identified a total of 14,060 unique peptides and 1851 unique proteins in different lots of standard and growth factor reduced (GFR) Matrigel preparations obtained from different suppliers, with only 53% batch-to-batch similarity in GFR Matrigel lots based on protein identification. These constituents will potentially affect the developmental potential,

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proliferation, and differentiation of cultured cells, by directly stimulating target cells synergistically or additively in a dosedependent fashion, or by indirectly activating or suppressing accessory cells inducing cytokine loops and cascades. Batch variation in these supplements is often a significant source of unexplained qualitative and quantitative differences in cell culture outcomes and assay readouts within laboratories or among different laboratories. Consequently, batch testing of these reagents is critical in order to ensure the stability and reproducibility of cell culture systems over time. Ideally aliquots of reagents with optimal growth supporting properties should be stored long term as a reference standard or as an aid in identifying confounding factors affecting reproducibility or replication of assay readouts. The commercial availability of highly defined cell culture media supplemented with unique combinations of essential nutrients, trace elements, hormones, and growth factors has enabled the development of cell culture systems for the maintenance, propagation, and manipulation of virtually all embryonic and adult cell types and cell lineages. However, cell-dependent variables, colligative properties of cell culture media, and the stability of the cell culture environment (Table 1) merit much greater attention when looking to optimize cell culture systems and improve the stability and reproducibility of cell culture assays.

Table 1 Variables affecting the proliferation and differentiation of cultured cells and the reproducibility of cell culture endpoints Heterogeneity of the initial cell inoculum Cell plating density Split ratio, growth phase, and cycling status of cultured cells Timing and frequency of refeeding Composition of the gas phase: pO2 and pCO2 Culture medium pH and buffering Volume and depth of cell culture medium Medium osmolarity Stability of tissue culture reagents Stability of incubation conditions—temperature, gas phase Period of incubation and criteria for culture endpoint (e.g., colony size)

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Cell-Dependent Variables The defined medium in which cells are propagated is only “defined” at the initiation of culture. From the moment cells are suspended in the defined medium, the properties of the culture system are instantly and progressively altered. The initial cell density and the heterogeneity of the cell inoculum are significant variables affecting the stability and reproducibility of cell culture outcomes. Cells interact in culture to secrete autocrine, juxtacrine, holocrine, and/or endocrine factors that modify their microenvironment to affect their survival, developmental potential, and rate of proliferation and differentiation (Fig. 1). Single cells or cells growing at clonal cell densities have more fastidious requirements than cells propagated at high cell densities [12]. The large volume of medium per cell in clonal cell cultures significantly compromises their ability to quickly generate optimal concentrations of secreted factors required for their survival and growth, potentially compromising their survival, proliferation, and differentiation. The replicative capacity, viability, and dynamic changes in the pattern of growth and differentiation of cells propagated in longterm culture also vary in response to the periodicity of refeeding, the phase of cell growth (exponential or stationary) at the time cultures are split, and the uniformity of split ratio. For example, hematopoietic stem cells secrete a large repertoire of stimulatory and inhibitory factors in the course of their proliferation and differentiation. Therefore, large fluctuations are observed in the repertoire and concentration of these factors each time medium is replenished during periodic refeeding of long-term cultures [15] markedly affecting the dynamics and heterogeneity of cell growth. The optimal performance and stability of cell culture assays, and the reproducibility of cell culture endpoints, relies on precise standardization of culture conditions. This includes number of cells, and the cell density of the cell inoculum, the ratio of the cell number to the volume of medium, and the timing and periodicity of subculture and refeeding.

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Medium pH and Buffering Early pioneers of cell culture recognized that the behavior of cultured cells is profoundly sensitive to changes in environmental pH, affecting parameters including protein synthesis, metabolism, cell growth rate [12, 16, 17], and cell differentiation and cloning efficiency [18]. Medium acidification as a result of catabolic and anabolic metabolism and the generation of inhibitory metabolites also affects the availability of nutrients due to complex interactions of medium constituents [17, 19] including sequestration of

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CO2 CO2 + H2O

H2CO3

HCO3- + H+

Fig. 2 The pH in cell culture is stabilized by the bicarbonate buffering system. Equilibrium is maintained by the relationship between the concentration of CO2 in the gas phase and the concentration of HCO3 in the cell culture medium. Acidification of culture medium drives the equation to the left raising CO2 concentration, whereas raising the concentration of CO2 in the gas phase, or in the medium due to the metabolic activity of cultured cells, drives the equation to the right lowering medium pH. For optimal buffering, the concentration of NaHCO3 in culture medium should be adjusted in line with the concentration of CO2 in the gas phase

essential nutrients by binding to albumin [20]. Optimal pH differs markedly for different cell types [16, 17] with some cell types exhibiting extreme sensitivity to medium acidification [21, 22] affecting cell cycling, cell growth, and differentiation and also causing DNA damage and genomic instability [23]. The regulation of pH in cell culture is primarily achieved by bicarbonate buffering as described by the Henderson-Hasselbalch equation (Fig. 2). Medium pH is maintained by the equilibrium between the CO2 concentration in the gas phase and the sodium bicarbonate (NaHCO3) concentration in the culture medium [11, 24]. An elevated concentration of CO2 in the gas phase will drive the equation to the right resulting in elaboration of an increased concentration of hydrogen ion (H+) and medium acidification. Conversely, medium acidification will drive the equation to the left increasing the elaboration of CO2. It is not uncommon for tissue culture protocols developed in different laboratories to specify different CO2 concentrations for the incubator gas phase: commonly 5% CO2 or 10% CO2. In my experience the fact that differences in the CO2 concentration of the incubator gas phase affects the buffering capacity of the medium is often overlooked, potentially affecting the optimal growth of cells and cell lines sensitive to medium acidification. Ideally, the CO2 tension in the incubator gas phase and/or the NaHCO3 concentration in different media should be adjusted accordingly [11, 24].

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The Gas Phase Cultured cells are most commonly maintained and propagated under normoxic conditions in a gas phase of 5% CO2 or 10% CO2 in air (i.e., 20% O2). However, there has been a growing awareness of the benefits of cell culture at low oxygen tension (5% O2), mimicking the hypoxic environment in which regenerative cells

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reside in tissues and organs in vivo [25, 26]. The reader is also referred to a comprehensive bibliography of historical studies in this field provided in supplementary information in the commentary by Wion et al. [26]. The beneficial effect of low O2 tension is less evident when analyzing the behavior of cell lines originally selected and propagated long term under normoxic conditions. Nor when cells or cell lines are grown at high cell density where O2 and CO2 tension in the pericellular microenvironment tends to be adjusted and regulated by cell metabolism. However, it is a different matter for many primary explanted cell types and cells grown at clonal or low cell densities where growth at low O2 tension often results in improved replicative capacity and cloning efficiency and qualitative and quantitative differences in differentiation potential or the elaboration of secreted factors. The expense of purchasing and maintaining triple gas incubators that regulate the delivery of CO2, O2, and nitrogen (N2) is often cited as an impediment to routine culturing of cells at low O2 tension. However, investigators should be aware that O2 toxicity and oxidative stress are detrimental to cells in culture [27–29] and that O2 toxicity can cause spontaneous genetic mutations [30]. A recent study has also demonstrated that low O2 tension is not only important during cell propagation but also during cell processing, noting that the incidence and recovery of hematopoietic stem cells (HSC) are significantly impaired in hematopoietic tissues processed in air [31].

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Osmolarity Medium osmolarity is another important variable affecting cell membrane transport and the metabolism, growth, and differentiation of cultured cells [32]. When evaluating the activity of cytokines, supplements, or reagents on cell proliferation and differentiation, cell metabolism, or gene expression, investigators should be aware of the possible contribution of these substances to changes in osmolarity that could affect culture outcome. Osmolarity of culture medium in open culture vessels can also be adversely affected by evaporation due to fluctuations in incubator temperature and humidity during injection of dry gases while purging to re-equilibrate the gas phase following opening and closing of the incubator. Frequent opening and closing of incubator doors to retrieve and examine cell cultures further exacerbate this problem creating a progressively hyperosmotic microenvironment in long-term cultures that will ultimately inhibit their proliferation and clonogenicity.

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Stability of Incubation Conditions The robustness of cell culture protocols, and the reproducibility of cell culture endpoints, relies on careful attention paid to the stability of incubation conditions during the period of culture: a factor underappreciated by many investigators. The previous section alluded to the impact of frequent opening and closing of incubator doors on the evaporation of medium and medium osmolarity due to loss of humidity. However, fluctuations in incubator temperature, and O2 and CO2 tension during examination of cultures, or following purging and re-equilibration of the incubator gas phase are equally significant, affecting medium pH and medium buffering. Investigators need to be aware that the equilibration of the gas phase in the incubator and gas phase inside a tissue culture vessel can take 30 min [33, 34]. Because equilibration of gas concentration in culture medium relies on diffusion, the rate of equilibration is also affected by the depth of the medium. Allen et al. [33] have shown that equilibration of O2 content in unstirred culture medium can take more than 3 h due to the low solubility and limited diffusion of O2 in aqueous solutions, potentially affecting the precision and reproducibility of cell culture endpoints over time.

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Conclusion Cell culture protocols for specific cell types will continue to evolve in lockstep with our understanding of the nature and function of the factors and signaling pathways that specify their fate and regulate their replicative capacity, proliferation, and differentiation. The impact of each of the variables discussed in this brief review on individual primary cell cultures or established cell lines will differ. But together, they are a source of experimental variation that if not appreciated and controlled potentially affect the reproducibility of cell culture endpoints within laboratories, as well as the ability of investigators to replicate assays and predictive models in different laboratories.

References 1. Schofield R (1978) The relationship between the spleen colony-forming cell and the haemopoietic stem cell. Blood Cells 4:7–25 2. Wagers AJ (2012) The stem cell niche in regenerative medicine. Cell Stem Cell 10:362–369 3. Muncie JM, Weaver VM (2018) The physical and biochemical properties of the extracellular matrix regulate cell fate. Curr Top Dev Biol 130:1–37 4. Nelson CM, Bissell MJ (2006) Of extracellular matrix, scaffolds, and signaling: tissue architecture regulates development, homeostasis, and cancer. Annu Rev Cell Dev Biol 22:287–309

5. Xu R, Boudreau A, Bissell MJ (2009) Tissue architecture and function: dynamic reciprocity via extra- and intra-cellular matrices. Cancer Metastasis Rev 28:167–176 6. Sneddon JB, Werb Z (2007) Location, location, location: the cancer stem cell niche. Cell Stem Cell 1:607–611 7. Quesenberry PJ (1993) Too much of a good thing. Reductionism run amok [editorial]. Exp Hematol 21:193–194 8. Folkman J, Greenspan HP (1975) Influence of geometry on control of cell growth. Biochim Biophys Acta 417:211–236

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9. Kaushik G, Ponnusamy MP, Batra SK (2018) Concise review: current status of threedimensional organoids as preclinical models. Stem Cells 36:1329–1340 10. Tharp KM, Weaver VM (2018) Modeling tissue polarity in context. J Mol Biol 430:3613–3628 11. Yao T, Asayama Y (2017) Animal-cell culture media: history, characteristics, and current issues. Reprod Med Biol 16:99–117 12. Ham RG (1981) Survival and growth requirements of nontransformed cells. In: Baserga R (ed) Handbook of experimental pharmacology, Tissue growth factors, vol 57. Springer-Verlag, Berlin, pp 13–88 13. Ham RG (1984) Formulation of basal nutrient media. In: Barnes DW, Sirbascu DA, Sato GH (eds) Methods for preparation of media, supplements, and substrata for serum-free animal cell culture, Cell culture methods for molecular and cell biology. Alan R. Liss, New York, pp 1–21 14. Hughes CS, Postovit LM, Lajoie GA (2010) Matrigel: a complex protein mixture required for optimal growth of cell culture. Proteomics 10:1886–1890 15. Csaszar E, Kirouac DC, Yu M, Wang W, Qiao W et al (2012) Rapid expansion of human hematopoietic stem cells by automated control of inhibitory feedback signaling. Cell Stem Cell 10:218–229 16. Ceccarini C, Eagle H (1971) pH as a determinant of cellular growth and contact inhibition. Proc Natl Acad Sci U S A 68:229–233 17. Eagle H (1974) Some effects of environmental pH on cellular metabolism and function. In: Clarkson B, Baserga R (eds) Control of proliferation in animal cells, vol 1. Cold Spring Harbor Laboratory, Cold Spring Harbor, pp 1–12 18. McAdams TA, Miller WM, Papoutsakis ET (1997) Variations in culture pH affect the cloning efficiency and differentiation of progenitor cells in ex vivo haemopoiesis. Br J Haematol 97:889–895 19. Waymouth C (1974) “Feeding the baby” – designing the culture milieu to enhance cell stability. J Natl Cancer Inst 53:1443–1448 20. Francis GL (2010) Albumin and mammalian cell culture: implications for biotechnology applications. Cytotechnology 62:1–16 21. Brodsky AN, Zhang J, Visconti RP, Harcum SW (2013) Expansion of mesenchymal stem cells under atmospheric carbon dioxide. Biotechnol Prog 29:1298–1306

22. Liu W, Ren Z, Lu K, Song C, Cheung ECW et al (2018) The suppression of medium acidosis improves the maintenance and differentiation of human pluripotent stem cells at high density in defined cell culture medium. Int J Biol Sci 14:485–496 23. Jacobs K, Zambelli F, Mertzanidou A, Smolders I, Geens M et al (2016) Higherdensity culture in human embryonic stem cells results in DNA damage and genome instability. Stem Cell Reports 6:330–341 24. Freshney RI (2016) Defined media and supplements. In: Culture of animal cells: a manual of basic technique and specialized applications, 7th edn. John Wiley & Sons Inc., Hoboken, pp 125–148 25. Toussaint O, Weemaels G, Debacq-ChainiauxF, Scharffetter-Kochanek K, Wlaschek M (2011) Artefactual effects of oxygen on cell culture models of cellular senescence and stem cell biology. J Cell Physiol 226:315–321 26. Wion D, Christen T, Barbier EL, Coles JA (2009) PO(2) matters in stem cell culture. Cell Stem Cell 5:242–243 27. Halliwell B (2003) Oxidative stress in cell culture: an under-appreciated problem? FEBS Lett 540:3–6 28. Halliwell B (2014) Cell culture, oxidative stress, and antioxidants: avoiding pitfalls. Biomed J 37:99–105 29. Ito K, Ito K (2018) Hematopoietic stem cell fate through metabolic control. Exp Hematol 64:1–11 30. Parshad R, Sanford KK, Jones GM, Price FM, Taylor WG (1977) Oxygen and light effects on chromosomal aberrations in mouse cells in vitro. Exp Cell Res 104:199–205 31. Mantel CR, O’Leary HA, Chitteti BR, Huang X, Cooper S et al (2015) Enhancing hematopoietic stem cell transplantation efficacy by mitigating oxygen shock. Cell 161:1553–1565 32. Waymouth C (1970) Osmolality of mammalian blood and of media for culture of mammalian cells. In Vitro 6:109–127 33. Allen CB, Schneider BK, White CW (2001) Limitations to oxygen diffusion and equilibration in in vitro cell exposure systems in hyperoxia and hypoxia. Am J Physiol Lung Cell Mol Physiol 281:L1021–L1027 34. Place TL, Domann FE, Case AJ (2017) Limitations of oxygen delivery to cells in culture: an underappreciated problem in basic and translational research. Free Radic Biol Med 113:311–322

Part II Methods and Protocols

Chapter 3 Propagation and Maintenance of Mouse Embryonic Stem Cells Jacob M. Paynter, Joseph Chen, Xiaodong Liu, and Christian M. Nefzger Abstract Mouse embryonic stem cells (mESCs) are pluripotent cells derived from preimplantation embryos that have the capacity to self-renew indefinitely in vitro. mESCs are an indispensable tool for studying cellular differentiation in vitro, generating disease in a dish models, and have been used extensively for the generation of transgenic animals. Therefore, maintaining their pluripotent state, even after extended culture, is crucial for their utility. Herein, we describe in detail a protocol for the culture of mESCs in the presence of fetal calf serum (FCS), leukemia inhibitory factor (LIF), and a layer of irradiated mouse embryonic fibroblasts (iMEFs). This culture system reliably sustains mESC pluripotency and self-renewal capacity, allowing their use in a wide range of experimental settings. Key words Mouse embryonic stem cells, Cell culture, Pluripotency, Mouse embryonic fibroblasts, Leukemia inhibitory factor

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Introduction Pluripotency is defined as the capacity of a cell to give rise to all somatic cell lineages and the germline of the embryo [1]. Our ability to maintain pluripotent stem cells in vitro was crucial to establish them as a platform for the study of early development in vitro and for the generation of transgenic animals [2]. Historically, pluripotent stem cell culture emerged from findings by Stevens and Little in 1954, who described a population of undifferentiated cells in mouse testicular teratocarcinomas termed embryonal carcinoma (EC) cells [3]. Most importantly, Kleinsmith and Pierce found that individual EC cells could give rise to bona fide teratocarcinomas when deposited into the peritoneum of secondary mice [4]. These tumors contained cells from each of the three germ layers, indicating that EC cells were pluripotent. In 1970, the groups of Sato and Ephrussi succeeded in maintaining EC cells

Jacob M. Paynter and Joseph Chen contributed equally to this work. Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940, https://doi.org/10.1007/978-1-4939-9086-3_3, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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in vitro as monolayer cultures in the presence of fetal calf serum (FCS) and a basal feeder layer of mitotically inactivated mouse fibroblasts [5, 6]. Over the next decade, biochemical and functional analyses showed that EC cells resemble cells of the early embryo [7–14]. The most stringent of these assays was the ability of EC cells to form chimeras upon blastocyst injection [15–17]. However, in the majority of cases, EC cells failed to contribute to the germline due to their excessive chromosomal abnormalities [18] which precluded their use for the generation of transgenic animals. Seminal work in 1981 culminated in two independent publications by Evans and Kaufman [19] and Martin [20] which described the direct in vitro derivation of pluripotent cells from preimplantation mouse embryos. These cells were named “embryonic stem cells (ESCs)” and could be maintained on a feeder layer of mouse embryonic fibroblasts (MEFs) in the presence of FCS. Mouse ESCs (mESCs) were found to readily give rise to germline chimeras [21] and, remarkably, whole mice when injected into a tetraploid blastocyst [22]. As mESCs are amenable to genetic modification in vitro, gene knockouts and knock-ins can be performed, including the insertion of conditional and reporter alleles, multiplexed gene targeting, and genome-wide mutagenesis. Transgenic animals derived via these routes have been instrumental for disease modeling and the study of gene function [23–26]. Subsequent work focused on characterizing the molecular pathways underpinning the maintenance of pluripotency in ESCs. In 1988, leukemia inhibitory factor (LIF) was identified as the principal component produced by feeder cells and can maintain mESCs in their absence [27, 28], albeit less efficiently, as feeder cells provide an additional attachment matrix as well as factors that support the maintenance of pluripotency in ESCs in addition to LIF [29, 30]. LIF activates STAT3 which feeds into the pluripotency network by upregulating the expression of pluripotency factors such as KLF4, Gbx2, and Tfcp2l1 [31–34]. Serum factors, particularly bone morphogenetic proteins, stimulate the SMAD signaling pathway and constrain lineage commitment by inducing expression of inhibitor of differentiation (ID) proteins [35, 36]. Building on these insights, in 2008, Ying et al. [37] established a culture system to maintain ESCs in the absence of LIF, serum, and feeders, with two small molecule inhibitors they termed “2i.” The components of 2i are PD03 (PD0325901) and CHIRON (CHIR99021) which modulate key pathways involved in lineage commitment and pluripotency. PD03 is a MEK inhibitor which blocks the auto-inductive effects of the FGF/ERK1/2 signaling cascade on differentiation [37], while CHIRON inhibits GSK-3 which mimics the effects of canonical Wnt signaling and thereby alleviates the repressive effects of TCF3 on pluripotency genes.

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Although providing a well-defined milieu, culture in 2i anchors mESCs in a so-called ground state of pluripotency [1], biasing lineage commitment under differentiation-inducing conditions [38]. Hence, most current mESC differentiation protocols still use cells cultured under serum/LIF conditions in the presence of feeder cells as a starting point. Albeit not chemically defined, this system remains the gold standard for mouse pluripotent stem cell culture. In this chapter, we describe the culture of mESCs under serum/LIF conditions on a feeder layer. We provide stepwise protocols for (I) the generation of high-quality growth-inactivated mouse embryonic fibroblast (iMEF) feeders, (II + III) the recovery of and routine culture of mESCs, and (IV) a protocol for their cryopreservation. The protocol described herein has broad applicability and is also compatible with the culture of induced pluripotent stem cells derived from a variety of different cell types [39].

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Materials 1. MEF culture medium: Dulbecco’s modified eagle medium (DMEM) containing 10% fetal calf serum (FBS) (v/v); 1% GlutaMAX™ Supplement (100
) (v/v); 1% MEM nonessential amino acid solution (100
) (v/v); 100 μM sodium pyruvate; 55 μM β-mercaptoethanol. 2. mESC culture medium: KnockOut™ DMEM containing 15% FBS (v/v) (see Note 1); 1% GlutaMAX™ Supplement (100
) (v/v); 1% MEM nonessential amino acid solution (100
) (v/v); 55 μM β-mercaptoethanol; 1000 units/mL recombinant murine LIF. 3. Cryopreservation medium: FBS containing 10% DMSO (v/v). 4. Dulbecco’s phosphate-buffered saline (DPBS). 5. 0.25% Trypsin-EDTA (1
), phenol red (380 mg/L EDTA, 2500 mg/L trypsin). 6. 175 cm2, angled neck, vented cap cell culture flasks. 7. 25 cm2, angled neck, vented cap cell culture flasks. 8. “Mr. Frosty” freezing containers. 9. 0.1% gelatine solution (w/v): Mix 1 g gelatine from porcine skin with 1 L ultrapure water (milli-Q). Autoclave to dissolve. 10. 15 mL centrifuge tubes. 11. 50 mL centrifuge tubes. 12. EmbryoMax® Primary Mouse Embryo Fibroblasts, Strain CF1, passage 1. 13. γ-Radiation source (e.g., Gammacell® 40 Exactor Low DoseRate Research Irradiator).

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Fig. 1 Generation of iMEFs. (a) Schematic overview of the process. (b–d) Morphology of MEFs prior splitting at the end of passages 1, 2, and 3, respectively. (e) Morphology of iMEFs 24 h after recovery from cryopreservation on a gelatine-coated surface. Scale bar: 25 μm

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Methods

3.1 Generation of iMEFs

3.1.1 Recovery of MEFs from Cryopreservation (Passage 1)

High-quality iMEFs are crucial to propagate mESC in a highly undifferentiated state. To generate feeder cells in large quantities, passage 1 (p1) MEFs are expanded by serial passaging, mitotically inactivated by irradiation, and cryopreserved for future use (Fig. 1a). While this protocol makes use of commercially available p1 MEFs, they can also be isolated de novo via dissection and homogenization of embryonic day 13.5 mouse embryos as described previously [40]. The procedure described in Subheading 3.1 is expected to yield between 2.4 and 3.6
108 iMEFs from the expansion of a single embryo. 1. Dispense 10 mL MEF medium into a 15 mL conical tube, and place in a water bath heated to 37  C. 2. Retrieve cryovials from liquid nitrogen storage collectively containing the MEFs from one embryo (~5
106) frozen down at the first passaging event (as described in Ref. 35), and place them in a 37  C water bath to thaw (see Note 2). 3. Once the contents of the vials have completely thawed, transfer the contents to the pre-warmed MEF medium prepared in step 1 of this section, and mix by gentle inversion. 4. Pellet the cells by centrifugation at 400
g for 5 min.

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5. Aspirate the supernatant from the pellet, and resuspend in 5 mL MEF medium by gentle pipetting. 6. Transfer the suspension in equal volumes to 2
175 cm2 culture flasks, and add additional MEF medium to achieve a working culture volume of 20 mL per flask. Mix by gentle pipetting. 7. Incubate the flasks at 37  C under low oxygen conditions (5% O2, 7% CO2) for 24 h (see Note 3). 8. Perform a media change to remove any dead cells and traces of DMSO: Aspirate culture medium and gently overlay the cells with 20 mL fresh MEF medium (see Note 3). 9. Incubate at 37  C under low oxygen conditions for a further 24–48 h. The cells will require passaging upon reaching ~90% confluence (Fig. 1b) (see Note 4). 3.1.2 Passage 2 (Expansion)

1. Aspirate culture media, and gently overlay the cells in each flask with 15 mL DPBS, and aspirate to remove traces of culture medium (see Note 5). 2. Dispense 3 mL Trypsin-EDTA solution into each flask, and rock back and forth to ensure the solution is distributed evenly over the cell layer. 3. Incubate the flasks at room temperature for 3–5 min. 4. Tap the flasks to dislodge the cells. Examine under a microscope to ensure all cells are fully detached (see Note 6). 5. Dispense 3 mL MEF medium into each flask to neutralize the trypsin. Gently pipette up and down several times to homogenize the cell suspension. 6. Pool the contents of each flask into a single 50 mL centrifuge tube. 7. Wash the surface of each flask with an additional 4 mL of MEF medium to recover any remaining cells, and pool into the 50 mL tube. 8. Pellet the cells by centrifugation at 400
g for 5 min. 9. Aspirate the supernatant from the pellet, and resuspend in 16 mL MEF medium by gentle pipetting. 10. Perform a 1:4 split: Transfer the suspension (20–30
106 cells total) in equal volumes into 8
175 cm2 culture flasks (see Note 7), and add additional MEF medium to achieve a working culture volume of 20 mL. Mix by gentle pipetting. 11. Incubate the flasks at 37  C under low oxygen conditions (5% O2, 7% CO2) for 48–72 h until the cells reach ~90% confluence (Fig. 1c).

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3.1.3 Passage 3 (Expansion)

1. Perform a 1:3 split: Passage cells as described previously in Subheading 3.1.2, effectively expanding the 8
p2 flasks into 24
175 cm2 culture flasks. 2. Incubate the flasks at 37  C under low oxygen conditions (5% O2, 7% CO2) for 48–96 h until the cells reach ~100% confluence (Fig. 1d) (see Note 8).

3.1.4 Harvesting, Irradiation, and Cryopreservation of MEFs

1. Trypsinize the cells as per Subheading 3.1.2, steps 2–5, and transfer the contents of each flask into 5
50 mL centrifuge tubes (see Note 9). 2. Wash the surface of each flask with an additional 4 mL of MEF medium to recover any remaining cells, and pool into the 5
50 mL tubes. 3. Pellet the cells by centrifugation at 400
g for 5 min. 4. Aspirate the supernatant from the pellets. Resuspend and pool the pellets in 30 mL MEF medium, and transfer to a single 50 mL tube. Determine cell number using an automated cell counter slide or hemocytometer (see Note 10). 5. Place the tube in a research irradiator, and expose to γ radiation at a dose rate of 1.0 Gy/min for 30 min. 6. Pellet the resulting iMEFs by centrifugation at 500
g for 5 min. 7. Aspirate the supernatant, and resuspend the pellet in cryopreservation medium at 1
107 cells/mL by gentle pipetting (see Note 11). 8. Using a serological pipette, dispense 500 μL aliquots (5
106 cells) into cryovials. 9. Place cryovials in a controlled rate freezer or freezing containers (“Mr. Frosty”) at 80  C for 24 h (see Note 12). After freezing, transfer the vials to liquid N2 cryo-storage (see Note 13).

3.2

mESC Culture

3.2.1 Gelatine Coating of Culture Vessels and Seeding of MEFs

In this protocol, mESCs (see Note 14) are maintained as monolayer cultures on gelatine-coated culture vessels that have been seeded with iMEF feeder cells (Fig. 2a). iMEFs can be obtained commercially (e.g., EmbryoMax® Primary Mouse Embryo Fibroblasts, Strain CF1, Irradiated, passage 3 (Merk Millipore)) or generated as described in the previous section. Feeder cells provide an additional attachment matrix as well as factors that support the maintenance of pluripotency in ESCs in addition to serum and LIF [29, 30]. 1. Overlay the surface of a 25 cm2 culture flask with 2 mL 0.1% gelatine solution. Incubate at room temperature for at least 20 min (see Note 15).

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Fig. 2 mESC culture. (a) Schematic overview depicting recovery of mESC from cryopreservation and their routine maintenance. (b–f) mESC recovery from cryopreservation and expected morphology. (b) iMEFs are seeded 12–24 h before thawing of mESCs. (c) Thawed mESCs are seeded at a density of 2.5
104 cells/cm2 and should reach 70% confluence after 3 days (d–f). Scale bar: 25 μm

2. Retrieve a vial of irradiated MEFs from cryo-storage. Thaw and resuspend as per Subheading 3.1.1, steps 1–5. Determine cell number using an automated cell counter slide or hemocytometer. 3. Aspirate the gelatine solution from the culture flask prepared in step 1 of this section. Transfer a volume of suspension containing 5
105 iMEF cells (2
104 cells/cm2) into the flask, and add additional MEF medium to achieve a working culture volume of 3 mL (see Note 16). 4. Incubate the flask at 37  C for 12–24 h to allow the feeders to attach. The cells should cover 70–100% of the flask (Fig. 2b). 3.2.2 Thawing of mESCs

1. Retrieve a vial containing ~1
106 mESCs from cryo-storage. Thaw and pellet as per Subheading 3.1.1, steps 1–4. 2. Aspirate the supernatant from the pellet, and resuspend in 1 mL mESC medium by gentle pipetting. Determine cell number using an automated cell counter slide or hemocytometer. 3. Aspirate MEF medium from the gelatine-coated flask with feeders prepared in Subheading 3.2.1. Transfer a volume of mESC suspension containing 6.25
105 cells

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(2.5
104 cells/cm2) into the flask (Fig. 2c), and add additional mESC medium to achieve a final culture volume of 3 mL. Mix by gentle pipetting. 4. Incubate the flask at 37  C for 24 h under atmospheric oxygen and 5% CO2. 5. Perform a media change to remove any dead cells and traces of DMSO: Aspirate culture medium and gently overlay the cells with 3 mL fresh mESC medium. 6. Incubate at 37  C for a further 48–72 h, changing medium after 48 h. The cells will require passaging upon reaching ~70% confluence (Fig. 2d–f) (see Note 17). 3.2.3 Routine Passaging of mESCs

1. Prepare a fresh gelatine-coated 25 cm2 flask seeded with iMEFs as per Subheading 3.2.1. 2. Aspirate culture media from the mESC culture vessel (25 cm2 flask), gently overlay with 2 mL DPBS, and aspirate to remove traces of culture medium. 3. Dispense 1 mL Trypsin-EDTA solution into each flask, and rock back and forth to ensure the solution is distributed evenly over the cell layer. 4. Incubate the flasks at room temperature for 3–5 min. 5. Tap the flasks to dislodge the cells. Examine under a microscope to ensure all cells are fully detached. 6. Dispense 1 mL mESC medium into each flask to neutralize the trypsin. Gently pipette up and down several times to homogenize the cell suspension. 7. Transfer the contents of the flask into a 15 mL centrifuge tube. 8. Pellet the cells by centrifugation at 400
g for 5 min. 9. Aspirate the supernatant from the pellet, and resuspend in 5 mL mESC medium by gentle pipetting (see Note 18). Determine cell number using an automated cell counter slide or hemocytometer. A 70% confluent 25 cm2 flask should yield between 7
106 and 1
107 cells. 10. Aspirate MEF medium from the gelatine-coated flask with feeders prepared in step 1 of this section. Transfer a volume of mESC suspension containing 1.25–2.5
105 cells (0.5–1
104 cells/cm2) into the flask, and add additional mESC medium to achieve a final culture volume of 3 mL (see Note 19). Mix by gentle pipetting. 11. Incubate at 37  C for 72 h under atmospheric oxygen and 5% CO2 until cultures reach ~70% confluence, changing medium after 48 h.

Mouse Embryonic Stem Cell Culture 3.2.4 Cryopreservation of mESCs

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1. After counting, pellet mESCs obtained in Subheading 3.2.3, step 9 by centrifugation at 500
g for 5 min. 2. Aspirate the supernatant, and resuspend the pellet in cryopreservation medium at 1
106 cells/mL by gentle pipetting. A 70% confluent 25 cm2 should yield 7–10 cryovials. 3. Using a serological pipette, dispense 1 mL aliquots (1
106 cells) into cryovials. 4. Place cryovials in a controlled rate freezer or (“Mr. Frosty”) freezing containers at 80  C for 24 h. After freezing, transfer the vials to liquid N2 cryo-storage.

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Notes 1. Certain batches of FCS can be detrimental to mESC cultures. Hence, we recommend using embryonic stem cell-qualified FCS which can be purchased from several vendors (e.g., Thermo Fisher, Merck, Applied StemCell). If this is undesirable due to financial constraints, batch testing of sera from other vendors can be performed. For batch testing, we recommend culturing mESCs for several passages (~5) in the alternative sera and to only use batches that are able to maintain the typical dome-shaped morphology and growth rate of mESCs. 2. Cells should be thawed as quickly as possible to maximize recovery. This can be facilitated by using a circulating water bath and periodically agitating the vial. 3. We recommend the use of low oxygen incubators for MEF expansions. MEFs cultured in a low oxygen environment show higher proliferation rates and delayed senescence compared to those cultured under atmospheric oxygen [41], thus giving rise to higher quality feeders. 4. MEF cultures should not be allowed to exceed 90% confluency, as this can induce growth inhibition and result in poor expansion. 5. When changing medium or washing with PBS, never dispense liquid directly onto the cells, as this can compromise the monolayer. Instead, eject gently down the side of the flask. 6. Prolonged trypsin exposure can compromise cell viability. We do not advise exposing cells to trypsin for much longer than 5 min. 7. When passaging MEFs, culture flasks may be reused to reduce financial costs. 8. The growth rate of MEFs slows down at later passages due to the onset of senescence, and cultures may take longer to reach confluence. Senescent MEFs have a flattened, spread out

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(“fried egg”) morphology. Cultures dominated by overtly senescent cells give rise to poor-quality feeders; hence we advise not to expand MEFs beyond passage 3. If culturing for longer than 72 h, cells should receive a media change every 48 h. 9. When handling high numbers of flasks, we recommend dividing the labor across subsets to avoid cells drying out during the washing steps. 10. Due to the high cell density of the suspension at this stage, we recommend taking a 20 μL aliquot of the concentrated suspension and diluting it 1:10 in MEF media before counting to determine the cell number more accurately. 11. It is advisable to minimize exposure of cells to DMSO at subfreezing temperatures due to its toxicity. Cell viability can be increased by working quickly and keeping the cells at a low temperature. Feeders should be resuspended in prechilled cryopreservation medium and aliquoted promptly into prechilled cryovials. 12. A controlled cooling rate of 1  C per minute is optimal for maximizing cell viability during freezing. While the use of “Mr. Frosty” freezing containers is acceptable for cryopreserving iMEFs, we recommend the use of a controlled rate freezer, as the freezing process induces a localized spike in temperature. A controlled rate freezer compensates for this and achieves a more uniform cooling rate resulting in superior cell viability after thawing. 13. High-quality feeders should have a high recovery rate after thawing (>70%) and adopt a spindle-shaped morphology (Fig. 1e). 14. The procedures outlined in this Methods chapter were established with R1 mESCs. While this protocol is compatible with mESC lines from other mouse strains, growth rates may vary slightly. Therefore, seeding densities may need to be optimized for other mESC lines. 15. Culture vessels can be scaled up or down as needed, e.g., use 6 mL of 0.1% gelatine solution for a 75 cm2 flask. 16. Although less cost-effective, feeders can also be cultured in mESC medium for convenience. 17. Culturing cells beyond 70% confluence may lead to spontaneous differentiation and poor cell viability. Always ensure that the morphology of the colonies is predominantly dome-shaped before using the cells for experiments. Immunostaining for pluripotency markers such as Oct4, Nanog, Sox2, and SSEA1 can be used to indicate the quality of mESC cultures (Fig. 3). Immunostaining can be performed as described previously [42] using primary antibodies against OCT4 (mouse IgG2b,

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Fig. 3 Assessment of differentiation status of mESC cultures. mESC colony stained for pluripotency markers (a) SSEA1 and (b) OCT4; (c) the DNA intercalating dye 40 ,6-diamidino-2-phenylindole (DAPI) was used to visualize cell nuclei; Panel d depicts a bright-field image of the mESC colony. Scale bar: 25 μm

1:100 dilution, Clone: C-10, Santa Cruz Biotechnology) and SSEA-1 (mouse IgM, 1:200 dilution, Clone: MC-480, DSHB); secondary conjugated antibodies goat anti-mouse IgG2b-AF 488 (1:400 dilution, Thermo Fisher Scientific) and goat anti-mouse IgM-AF 555 (1:400 dilution, Thermo Fisher Scientific); and nuclear stain 40 ,6-diamidino-2-phenylindole, dihydrochloride (DAPI) (1:1000 dilution, Invitrogen). Cultures with excessive spontaneous differentiation can be rescued by FACS isolation of SSEA-1 and EPCAM doublepositive cells and their use for re-culture [40, 43]. A more stringent assay for pluripotency is the teratoma formation assay entailing subcutaneous injection of mouse pluripotent stem cells into the flanks of immune compromised mice to assess their in vivo differentiation potential into derivates of all three germ layers [44, 45]. 18. Most differentiation protocols require the use of mESCs in the absence of contaminating feeders. mESC suspensions (after Trypsin/EDTA dissociation) can be depleted of feeders by transferring the 5 mL suspensions obtained in Subheading 3.2.1, step 9 onto a non-gelatine-coated 25 cm2 culture flask and incubating at 37  C for 30 min. After this period, the feeders will have attached to the plate, while the mESCs remain in suspension. The supernatant containing the mESCs can be collected and used for downstream differentiation applications as detailed previously [46]. 19. Experienced users can perform a 1:10–1:20 split which should approximate this seeding density. References 1. Hackett JA, Surani MA (2014) Regulatory principles of pluripotency: from the ground state up. Cell Stem Cell 15:416–430

2. Evans M (2011) Discovering pluripotency: 30 years of mouse embryonic stem cells. Nat Rev Mol Cell Biol 12:680–686

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40. Nefzger CM, Alaei S, Knaupp AS, Holmes ML, Polo JM (2014) Cell surface marker mediated purification of iPS cell intermediates from a reprogrammable mouse model. J Vis Exp (91):e51728. 41. Parrinello S, Samper E, Krtolica A, Goldstein J, Melov S, Campisi J (2003) Oxygen sensitivity severely limits the replicative lifespan of murine fibroblasts. Nat Cell Biol 5:741–747 42. Nefzger CM, Haynes JM, Pouton CW (2011) Directed expression of Gata2, Mash1, and Foxa2 synergize to induce the serotonergic neuron phenotype during in vitro differentiation of embryonic stem cells. Stem Cells 29:928–939 43. Nefzger CM, Alaei S, Polo JM (2015) Isolation of reprogramming intermediates during generation of induced pluripotent stem cells from mouse embryonic fibroblasts. Methods Mol Biol 1330:205–218 44. Alaei S, Knaupp A, Lim S, Chen J, Holmes M et al (2016) An improved reprogrammable mouse model harbouring the reverse tetracycline-controlled transcriptional transactivator 3. Stem Cell Res 17:49–53 45. Firas J, Liu X, Nefzger CM, Polo JM (2014) GM-CSF and MEF-conditioned media support feeder-free reprogramming of mouse granulocytes to iPS cells. Differentiation 87:193–199 46. Chen J, Nefzger CM, Rossello FJ, Sun YBY, Lim SM et al (2018) Fine tuning of canonical Wnt stimulation enhances differentiation of pluripotent stem cells independent of βcatenin-mediated T-Cell factor signaling. Stem Cells 36:822–833

Chapter 4 Production of High-Titer Lentiviral Particles for Stable Genetic Modification of Mammalian Cells Michael R. Larcombe, Jan Manent, Joseph Chen, Ketan Mishra, Xiaodong Liu, and Christian M. Nefzger Abstract Lentiviral gene transfer technologies exploit the natural efficiency of viral transduction to integrate exogenous genes into mammalian cells. This provides a simple research tool for inducing transgene expression or endogenous gene knockdown in both dividing and nondividing cells. This chapter describes an improved protocol for polyethylenimine (PEI)-mediated multi-plasmid transfection and polyethylene glycol (PEG) precipitation to generate and concentrate lentiviral vectors. Key words PEI transfection, PEG precipitation, Titration, Gene transfer

1

Introduction Effective delivery and expression of exogenous genes in mammalian cells is essential for the study of gene function. Viral transfer technologies are routinely used for transient and integrating gene delivery in vitro and, once biosafety concerns are addressed, have a vast potential for clinical applications [1–3]. Unlike the transient expression achieved with adenoviral vehicles, lentiviral and retroviral vectors allow stable transgene integration for sustained and heritable gene expression. This is particularly useful for generating transgenic animals, reprogramming fibroblasts into induced pluripotent stem cells (iPSCs), and creating stable cell lines overexpressing or silencing genes via RNA interference [4]. While both lentiviral and retroviral gene transfer methods integrate transgenes into the genome of targeted cells for continued expression, lentiviruses transduce both replicating and non-replicating cells, integrate away from cellular promoters, allow for a larger genomic payload, and maintain transgene

Michael R. Larcombe, Jan Manent, and Joseph Chen contributed equally to this work. Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940, https://doi.org/10.1007/978-1-4939-9086-3_4, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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expression in pluripotent cells [5–9]. Thus, lentiviruses are a particularly widely used and a popular tool for the stable genetic modification of mammalian cells. Furthermore, advances have been made to minimize risk and improve delivery efficiency via the design of new generation lentiviral vectors [10]. These systems prevent the generation of replication competent virus through deletion of nonessential viral genome components and separation of the remaining elements. As such the second-generation system used in this study requires three separate vectors encoding for the transgene (called “transfer vector”), replication genes (including the rev transactivator), and envelope genes. Omission of unnecessary virulence factors and isolation of the remaining components further reduce the risk of homologous recombination that could lead to the generation of unwanted replicating virus [11, 12]. Modern vector designs improve infectivity of the generated particles by substitution of viral coating proteins within the envelope vector for a broader (vesicular stomatitis Indiana virus G protein [VSV-G], as used in this protocol) [13, 14] or more specified tropism (human parainfluenza virus type 3 [HPIV3] for lung epithelial cells) [15, 16]. Numerous methods for producing lentiviral particles have been established, predominantly using transient co-transfection of the desired transfer vector with the accessory plasmids [17, 18]. This can be achieved with a variety of reagents utilizing lipid-based, polymer-based, or naked DNA delivery [19]. Nevertheless, price, efficiency, scalability, and simplicity must be accounted for when selecting transfection technique and reagents. Commercial products such as Lipofectamine and newer versions thereof offer a simple and efficient way to achieve effective plasmid transfection; however, especially for large-scale experiments, they can be cost intensive. Since viral particle production is generally achieved using highly transfectable human embryonic kidney 293T (HEK293T) cells, lipofection-based approaches can be substituted with the more cost-effective calcium phosphate (Ca-phosphate) or polyethylenimine (PEI) transfection systems. Ca-phosphate and PEI methods are also reasonably simple and capable of producing high viral titers from HEK293T cells [15, 20–22]. However transfecting with Ca-phosphate is very sensitive to fluctuations in pH [23], and therefore the resulting titers can differ largely between experiments. Since generation of high viral titers is dependent on efficient transfection, we consider the PEI-based method more reliable for consistent results. PEI transfection operates by condensing DNA into positively charged particles for delivery across the cell membrane. DNA is then released in the cytoplasm and incorporated into the nucleus during cell division for temporary expression [24], followed by the secretion of lentiviral particles into the growth medium. The resulting supernatant can then be collected and used for direct transduction of target cells or concentrated to achieve higher titer viral preparations. Protocols for lentiviral

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concentration typically require expensive ultrafiltration units and lengthy periods of ultracentrifugation [15, 25–27]. For laboratories without access to this high-end equipment and due to the ease of use, polyethylene glycol (PEG) precipitation offers a cheap and simple alternative to concentrate large volumes of supernatant and efficiently recover the viral particles [25, 28]. PEG is a highly hydrophilic polymer composed of repeating subunits of ethylene oxide and is commercially available in varying molecular weights. Generally, higher molecular weight PEGs are more efficient for precipitation compared to lower molecular weight PEGS [29]. Therefore we are using PEG (8000 Da) that has a relatively high molecular weight and is therefore very effective as a crowding agent in aqueous solution. PEG separates the viral particles from the aqueous medium and forces them to aggregate through a process called “steric exclusion” [30] which is supported by high levels of salt such as NaCl, through a “salting-out” mechanism [31]. This allows the virus to be pelleted with a simple benchtop centrifuge without the need for ultracentrifugation [32–35]. Therefore, in this chapter, we describe an improved protocol for the production and concentration of replication-deficient hightiter lentiviruses using PEI transfection and concentration by PEG precipitation. In the context of this protocol, we also describe the routine maintenance of the HEK293T producer cell line, a method for determining viral titers (for viral inserts with and without a fluorescent reporter gene) and the use of the viral concentrates to infect a target cell type of interest.

2

Materials

2.1 Lentiviral Production and Titration

1. Lenti-X™ 293T cell line (HEK293T) (Clontech, 632180) or 293T cell line (ATCC® CRL-3216™). 2. Complete growth medium (for 293T cells and MEFs): Dulbecco’s modified eagle medium (DMEM), containing 10% fetal calf serum (FBS) (v/v); 1% GlutaMAX™ Supplement (100
) (v/v); 1% MEM nonessential amino acid solution (100
) (v/v); 1% penicillin/streptomycin (100
) (v/v); 100 μM sodium pyruvate; 55 μM β-mercaptoethanol. 3. Viral production culture medium: Advanced DMEM containing (Gibco®, 12491015) 2% FBS (v/v) (see Note 1); 1% GlutaMAX™ Supplement (100
) (v/v); 1% MEM nonessential amino acid solution (100
) (v/v); 1% penicillin/streptomycin (100
) (v/v). 4. 15 mL centrifuge tubes. 5. Cryopreservation medium: FBS containing 10% DMSO (v/v). 6. Dulbecco’s phosphate-buffered saline (DPBS).

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7. 0.25% trypsin-EDTA (1
), phenol red (380 mg/L EDTA, 2500 mg/L trypsin). 8. 175 cm2, angled neck, vented cap cell culture flasks (T175). 9. 50 mL centrifuge tubes. 10. “Mr. Frosty” freezing container. 11. Linear polyethylenimine 25,000. 12. BSA fraction V (7.5%) (BSA). 13. Virkon® disinfectant cleaner. 14. UltraPure distilled water. 15. PAX2 plasmid (psPAX2 was a gift from Didier Trono (Addgene, #12260)). 16. MD2G plasmid (pMD2.G was a gift from Didier Trono (Addgene, #12259)). 17. OKSM plasmid (Merck Millipore, SCR512). 18. rtTA-GFP plasmid (designed and ordered from vector builder [Cyagen Biosciences]; lentiviral plasmid with a EF1A promoter driving the m2rtTA gene, followed by an internal ribosome entry site and an eGFP reporter (LV-EF1A-m2rtTA-IRESeGFP)). 19. 0.45 μm, HV Durapore® membrane filter (Merck Millipore). 20. Polyethylene glycol (PEG) 8000. 21. 5M NaCl in dH2O, filtered with 0.22 μm. 22. Doxycycline hyclate diluted to 2 mg/mL in dH2O (1000
stock). 23. Polybrene infection/transfection reagent 10 mg/mL stock (used at 1:1700 dilution). 2.2 Immunofluorescence and Viral Titration Analysis (See Note 18)

1. Paraformaldehyde (PFA). 2. Triton-X. 3. DAPI: 40 ,6-diamidino-2-phenylindole, dihydrochloride. 4. Mouse anti-Oct4 antibody (Santa Cruz Biotechnology, sc-5279). 5. Goat anti-mouse IgG Alexa Fluor 555 (Life Technologies®, A21422).

3

Methods

3.1 Recovery from Cryopreservation and Routine Culture of HEK293T Cells

1. Collect cryovial containing 4–5
106 HEK293T cells from liquid nitrogen storage, and thaw quickly in a 37  C water bath. 2. Transfer cells to a 15 mL conical tube along with 10 mL of pre-warmed complete growth medium.

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3. Centrifuge at 500
g for 5 min, and then aspirate supernatant containing toxic DMSO without disturbing cell pellet. 4. Resuspend cells in 4 mL of growth medium by gentle pipetting, and transfer 3.5
106 cells into a 175 cm2 flask (T175) made to 20 mL with growth medium. 5. Incubate at 37  C in a 5% CO2 incubator. Replace media every 2–3 days (as required) with 20 mL of fresh growth medium during routine culture until cells reach 80% confluence (see Note 1). 6. To harvest cells, firstly remove the spent growth medium, and then wash cells with 10 mL PBS. Apply DPBS to the side of the flask, and gently tilt to spread, trying not to disturb the weakly adherent cells. 7. Aspirate DPBS, and evenly distribute 5 mL trypsin-EDTA over the cells to dissociate. 8. Incubate the cells at room temperature for 3–5 min, using the microscope to verify when >90% of the cells have detached. 9. Neutralize the trypsin by adding 10 mL of growth medium. Pipet up and down several times over the cell culture surface to separate and collect remaining cells. 10. Transfer cells to a 50 mL centrifuge tube and pellet at 500
g for 5 min. 11. Resuspend the cells in 5 mL of complete growth medium, and remove a sample for counting. 12. In general, 80–90% confluent flasks can be passaged at a split ratio of 1:5 up to 1:20. Depending on the split ratio, the new flasks will become confluent in a 2–4-day time frame. 13. Create frozen stocks by resuspending the cells in cryopreservation medium at 2–6
106 cells/mL, and dispense 1 mL aliquots into cryovials. Use a controlled rate freezer or “Mr. Frosty” freezing container at 80  C for 24 h, and transfer frozen cells to liquid nitrogen for long-term storage. 3.2 PEI Transfection (See Note 2) and Collection of Primary Viral Supernatant

1. Thaw an aliquot of 1 mg/mL PEI (see Note 3). 2. The day before transfection, plate HEK293T cells (that have been passaged at least once after recovery from cryopreservation) in complete growth medium at a density of 1.5
107 to 2
107 cells per T175 flask (8.5
104 to 1.15
105 cells/ cm2). Depending on the growth rate of your HEK293T culture, adjust to have healthy, 80% confluent cultures for transfection the following day (Fig. 1) (see Note 4). 3. The next day in the late afternoon, 1 h prior to transfection, change complete growth medium to viral production culture medium (20 mL per T175 flask).

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Fig. 1 Schematic overview of viral particle production via PEI transfection Table 1 Transfection mix for second-generation lentivirus production in HEK293T cells T175

T75

T25

Vector

Concentration Ratios μg DNA Volume μg DNA Volume

Transfer DNA

100 ng/μL

3

15

150 μL 6.3

63 μL

2.1

21 μL

psPAX2

100 ng/μL

2

10

100 μL 4.2

42 μL

1.4

14 μL

pMD2.G

100 ng/μL

1

5

50 μL

21 μL

0.7

7 μL

120

120 μL 50.4

50.4 μL

16.8

16.8 μL

H2O

980 μL

423.6 μL

141.2 μL

Total volume

1.4 mL

0.6 mL

0.2 mL

PEI (4 μg/μg DNA) 1 mg/mL

2.1

μg DNA Volume

DNA concentrations are arbitrarily set at 100 ng/μL for the purpose of this example If transfecting more than one T175 flask, the values in this table can be scaled up accordingly

4. Prepare transfection mix as shown in Table 1: per T175 to transfect, mix 15 μg transfer DNA, 10 μg psPAX2, 5 μg pMD2.G, and 120 μg PEI in water to a final volume of 1.4 mL (see Note 5).

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5. Vortex solution for 10 s, and incubate at room temperature for 15 min. 6. Pipet up and down gently 2–3 times, and add the solution to the cells dropwise (Fig. 1). 7. Swirl flask gently to evenly distribute the DNA mix over the cells and return to incubator. 8. Incubate overnight at 37  C, 5% CO2. 9. Early the next morning, discard transfection medium into strong bleach or 1% Virkon® solution, and replace with fresh viral production culture medium (Fig. 1). 10. 24 h later, collect supernatant for the first time, and replace with fresh viral production culture medium. Keep supernatant at 4  C if not processed right away (Fig. 1) (see Note 6). 11. 24 h later (i.e., 48 h after removal of transfection complexes), collect supernatant for a second time. The flasks with any remaining cells can be discarded at this point (Fig. 1). 12. Proceed with concentration. 3.3 Viral Concentration

1. Filter the harvested medium through a 0.45 μm membrane, and take note of total volume (Fig. 2) (see Note 7). 2. Adjust NaCl concentration to 400 mM with a 5 M NaCl stock solution. This is achieved by adding a 1/17 volume of 5M NaCl to viral production culture medium (i.e for 10 mL of viral supernatant, add 588 μL of 5M NaCl). 3. Add 50% PEG solution to a final concentration of 8.5% (see Note 8). 4. Mix well and transfer mixture into 50 mL Falcon tubes. Make sure to have balanced volumes in the tubes for centrifugation in step 6 (Fig. 2). 5. Incubate the tubes at 4  C or on ice for 5 h, with regular agitation (see Note 9). 6. Spin samples for 1.5 h at >4000
g in a centrifuge pre-cooled to 4  C (see Note 10). 7. Gently discard supernatant (without disturbing viral pellet) in bleach or 1% Virkon® solution, and centrifuge empty tube again for 5 min (Fig. 2). 8. Gently remove any remaining supernatant with a P1000 pipette without disturbing pellet (see Fig. 2 for picture of the pellet). 9. We recommend to resuspend the pellet in DPBS containing 1% BSA (v/v) in a volume corresponding to 1/100th of the amount of primary supernatant used for concentration (e.g., resuspend the pellet resulting from the concentration of 30 mL supernatant in a volume of 300 μL; see Note 11).

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Fig. 2 Schematic overview of viral particle concentration with PEG

10. Aliquot into 20–30 μL aliquots (see Note 12). 11. Freeze at 80  C. 3.4

Titration

3.4.1 Viral Titration of a Lentiviral Vector Carrying a Fluorescent Reporter

1. Thaw and recover a vial of mouse embryonic fibroblasts (MEFs) from DMSO (see Note 13) into complete growth media. 2. Seed MEFs in 12 wells (9 for rtTA-GFP titering and 5 control wells (3 control wells and 2 wells for counting)) in a 24-well plate format at 1
104 cells/cm2 at least 12–24 h before viral transduction (Fig. 3a, c). 3. Count the cells of two control wells to obtain accurate cell count at the time of transduction. 4. Thaw and aliquot rtTA-GFP viral concentrate (at 1:1000, 1:10000, 1:100000 dilutions; see Note 14) into complete growth media containing polybrene (1:1700 dilution of stock) in 1000 μL volumes. 5. Remove media from the wells previously seeded with MEFs, and add 500 μL media of the respective serial dilutions onto cells (Fig. 3a, c). 6. Spin plates at 750
g for 60 min at room temperature to increase the efficiency of transduction. 7. The following day, replace media containing viral particles with fresh complete growth media. 8. 72 h after transduction, harvest each of the wells by dissociating cells with appropriate dissociation reagent (0.25% Trypsin, EDTA, etc.) for 5 min. Resuspend cells with FACS buffer (PBS w/2% FBS) supplemented with DAPI (1 μg/mL), and transfer cells into a FACS tube to determine the percentage of GFP

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Fig. 3 Titration of viral concentrates. (a) Schematic overview of rtTA-GFP viral titration. (b) FACS analysis of rtTA-GFP viral titration. (c) Schematic overview of OKSM viral titration. (d) Immunofluorescence images for GFP (green) and Oct4 (red) in MEFs transduced with OKSM and rtTA-GFP viruses after 48 h and quantification of Oct4þ cells within the GFPþ population. All nuclei were counterstained with DAPI (blue). Scale bar ¼ 25 μm

expressing cells by flow cytometry for the m2rtTA-GFP construct (Fig. 3b). 9. Calculate titers of viral concentrate using the following formula: (% of GFP+ cells [between 1% and 25%]
number of cells at the time of transduction
dilution factor)/volume of media for transduction (in mL). Titers are usually indicated as transducing units per mL (TU/mL) (see Notes 15 and 16 for example calculation).

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3.4.2 Viral Titration of a Lentiviral Vector Without a GFP Fluorescent Marker

The titration of the OKSM virus (which does not have a fluorescent reporter) follows a similar pattern as described in steps 3.4.1. steps 1–9 for the m2rtTA-GFP virus. However, there are noteworthy differences. In particular, for calculating OKSM viral titers, an immunofluorescence staining is required to determine the percentage of cells expressing exogenous Oct4, Klf4, Sox2, or C-Myc (Fig. 3c, d). Furthermore as the OKSM constructs is inducible, the titer determination will have to occur in the presence of excess m2rtTA-GFP virus and doxycycline (2 μg/mL) to enable OKSM expression (see Notes 17 and 18 for detailed information as well as Fig. 3c, d).

3.5 Viral Transduction of Target Cell Type

1. MEFs (or any other cell type that has ideally also been used to determine the viral titers) can be used for transduction in an experimental context. 2. For infection seed MEFs at 1
104 cells per cm2 as in Subheading 3.4.1., step 2, (albeit in a 6-well format in 2 mL of media or scale to other well formats) 12–24 h before transduction. 3. Prepare transduction mix as in Subheading 3.4.1., step 4 with outlined modifications: it is important for the experimenter to decide at what multiplicity of infection (MOI; see Note 17) to perform transduction. As a reference if the experimenter decides for an MOI ¼ 1, only ~60% of cells will become transduced as some cells become infected with more than one virus. Conversely infection with an MOI of 10 will result in infection of >85–90% of cells with an average of 10 viral integrants per cell. An example of how to calculate the amount of viral concentrate to add to a 6-well with a set number of cells for a specific target MOI is provided in Note 19. 4. Perform spin inoculation as described in Subheading 3.4.1. steps 5–7, using 2 mL of transduction mix per 6-well. Please note that it will take 48–72 h before gene products introduced with the viral inserts can be detected in the infected cells.

4

Notes 1. HEK293T cells undergo contact inhibition of growth if allowed to become confluent. This must be avoided as a key parameter for efficient viral production is healthy, exponentially growing cells. Gently move the freshly seeded flask back and forth, left and right to evenly distribute the cells throughout the flask. Try to recover low-passage stock (70% of HEK293T cells to express the fluorescent marker. 7. Using filters with a smaller pore size may result in retention of viral particles. 8. Resuspend PEG powder in water, 50% weight/volume, and filter (0.45 μm). The solution will be very viscous, so we recommend resuspending overnight at 37  C with agitation. 9. We recommend constant mixing using a rotating wheel or rolling table at 4  C; if this isn’t available, you can invert tubes on ice at half-hourly intervals. 10. We have successfully been using a Heraeus Multifuge X3R centrifuge (Thermo Fisher) at Vmax of 4700 rpm/4800
g.

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11. The PEG/viral pellet is quite sticky. It helps to resuspend the pellet and aliquot at room temperature but return to ice or freeze down immediately afterward. 12. Depending on your titer and experimental requirements, you may want to adjust the aliquot size in the next viral preparation. Avoid freeze thaw cycles as the amount of infectious particles drops dramatically with each additional freeze thaw cycle (ideally only have one freeze thaw cycle). 13. The transduction efficiency of a lentiviral vector varies between cell types; therefore it is necessary to titrate against the cell type intended for use in the planned experiments. Seed at least duplicate wells for each viral titration (for five concentrations, seed at least ten wells of cells), and include two control wells that are not transduced with virus. MEFs are only used as an example cell type in the context of this manuscript and should be substituted for the experimenters’ cell type of need. 14. You may prepare these solutions in an empty 24-well plate by adding viral concentrate into a 24 well at a 1:100 or 1:1000 dilution and then perform ten-fold serial dilutions into neighboring wells containing growth media supplemented with polybrene. 15. Use the dilutions that give values in the range of 1–20% GFP cells to calculate the MOI to avoid complications arising from cells receiving multiple insertions. 16. Example calculation of viral titer: (% of GFP+ cells [e.g., 5%, express as 0.05]
number of cells at the time of transduction [e.g., 2
104]
dilution factor [100000])/volume of media for transduction (0.5 mL) ¼ 2
108 transduction units (TU)/mL 17. The multiplicity of infection or mean occurrence of infection (MOI) is defined as the theoretical average number of viral integrants per target cell. 18. To determine the titer of OKSM viruses, seed MEFs in 24-well plates (Fig. 3c) at 1
104 cells/cm2 12–24 h before viral transduction. Aliquot rtTA-GFP virus into complete growth media containing polybrene (1:1700 dilution) at MOI ¼ 10. (The MOI of 10 for rtTA-GFP is chosen to ensure most of the cells are expressing the transactivator that is critical for doxycycline induction of OKSM expression (see Note 19 for example calculation). As an alternative, OKSM lentivirus can be titered using an rtTA-expressing stable cell line.) Separate out the rtTA-GFP containing media into 1000 μL volume aliquots, and establish 1:100, 1:1000, 1:10000, and 1:100000 dilutions of OKSM virus. We recommend using at least four 10
serial dilutions ranging from 1:100 to 1:100000 of OKSM virus to get a more accurate titration of the virus. In our experience, a

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59

dilution factor of up to 1:1000000 can be required to determine the transducing units of high-titer viral preparations. Add 500 μL of media into respective wells (as indicated in Fig. 3c) followed by spin inoculation as per Subheading 4.4.1.6. On the following day, remove media from wells, and replace with fresh media containing doxycycline (2 μg/mL). After 48–72 h, fix cells and perform immunofluorescence on cells to determine infective units of OKSM virus preparation. Further details on performing immunofluorescence can be referred from Nefzger et al. [37] and Chen et al. [38]. We recommend using an Alexa Fluor 555 secondary antibody as it does not interfere with the 488 nm (GFP) channel or the DAPI channel when analyzing expression of viral titration and a primary antibody against Oct4 (see Subheading 2). Calculate the subset of GFPþ cells that are positive for Oct4, Klf4, Sox2, or C-Myc (rtTA-GFP transduced cells) to determine the percentage of transduction (Fig. 3d). We perform cell quantification using the particle analysis option of the ImageJ software (http://rsb. info.nih.gov/ij/). We recommend counting cells from multiple images per replicate well to give a more accurate estimation of the transduction rate of the OKSM virus. It is recommended to take images from at least four fields of view per well for analysis. 19. Example calculation for transduction at MOI ¼ 10:   Number of cells to be infected 2
104
MOI ½10   ¼ TU required to transduce cells at MOI ¼ 10 2
105 TU Volume of viral aliquot to use for transduction of 2
104 cells at MOI ¼10   2
105 TU ðTU required to transduce cells at MOI ¼ 10Þ=   2
108 =mL ðTiter of viral concentrateÞ ¼ 0:001 mL or 1 μL Be advised that MOIs that are higher than 50 can lead to varying degrees of cell death due to cytotoxicity. References 1. Naldini L (2015) Gene therapy returns to centre stage. Nature 526:351–360 2. Bessis N, GarciaCozar FJ, Boissier MC (2004) Immune responses to gene therapy vectors: influence on vector function and effector mechanisms. Gene Ther 11(Suppl 1):S10–S17 3. Baum C, Kustikova O, Modlich U, Li Z, Fehse B (2006) Mutagenesis and oncogenesis by

chromosomal insertion of gene transfer vectors. Hum Gene Ther 17:253–263 4. Nefzger CM, Rossello FJ, Chen J, Liu X, Knaupp AS et al (2017) Cell type of origin dictates the route to pluripotency. Cell Rep. 21:2649–2660. 5. Naldini L, Blomer U, Gallay P, Ory D, Mulligan R et al (1996) In vivo gene delivery and

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stable transduction of nondividing cells by a lentiviral vector. Science 272:263–267 6. Vannucci L, Lai M, Chiuppesi F, CeccheriniNelli L, Pistello M (2013) Viral vectors: a look back and ahead on gene transfer technology. New Microbiol 36:1–22 7. Wu X, Burgess SM (2004) Integration target site selection for retroviruses and transposable elements. Cell Mol Life Sci 61:2588–2596 8. Montini E, Cesana D, Schmidt M, Sanvito F, Bartholomae CC et al (2009) The genotoxic potential of retroviral vectors is strongly modulated by vector design and integration site selection in a mouse model of HSC gene therapy. J Clin Invest 119:964–975 9. Pfeifer A, Ikawa M, Dayn Y, Verma IM (2002) Transgenesis by lentiviral vectors: lack of gene silencing in mammalian embryonic stem cells and preimplantation embryos. Proc Natl Acad Sci U S A 99:2140–2145 10. Dull T, Zufferey R, Kelly M, Mandel RJ, Nguyen M et al (1998) A third-generation lentivirus vector with a conditional packaging system. J Virol 72:8463–8471 11. Zufferey R, Dull T, Mandel RJ, Bukovsky A, Quiroz D et al (1998) Self-inactivating lentivirus vector for safe and efficient in vivo gene delivery. J Virol 72:9873–9880 12. Pacchia AL, Adelson ME, Kaul M, Ron Y, Dougherty JP (2001) An inducible packaging cell system for safe, efficient lentiviral vector production in the absence of HIV-1 accessory proteins. Virology 282:77–86 13. Akkina RK, Walton RM, Chen ML, Li QX, Planelles V, Chen IS (1996) High-efficiency gene transfer into CD34þ cells with a human immunodeficiency virus type 1-based retroviral vector pseudotyped with vesicular stomatitis virus envelope glycoprotein G. J Virol 70:2581–2585 14. Bartz SR, Vodicka MA (1997) Production of high-titer human immunodeficiency virus type 1 pseudotyped with vesicular stomatitis virus glycoprotein. Methods 12:337–342 15. Sena-Esteves M, Tebbets JC, Steffens S, Crombleholme T, Flake AW (2004) Optimized large-scale production of high titer lentivirus vector pseudotypes. J Virol Methods 122:131–139 16. Grzybowski B, Tong S, Compans RW, Le Doux JM Lentivirus vector pseudotyped with human parainfluenza virus type 3 glycoproteins. In: Proceedings of the Second Joint 24th Annual Conference and the Annual Fall Meeting of the Biomedical Engineering Society, Engineering in Medicine and Biology

17. Merten O-W, Hebben M, Bovolenta C (2016) Production of lentiviral vectors. Mol Ther Methods Clin Dev 3:16017 18. Kafri T, van Praag H, Ouyang L, Gage FH, Verma IM (1999) A packaging cell line for lentivirus vectors. J Virol 73:576–584 19. Kim TK, Eberwine JH (2010) Mammalian cell transfection: the present and the future. Anal Bioanal Chem 397:3173–3178 20. Boussif O, Lezoualc’h F, Zanta MA, Mergny MD, Scherman D et al (1995) A versatile vector for gene and oligonucleotide transfer into cells in culture and in vivo: polyethylenimine. Proc Natl Acad Sci U S A 92:7297–7301 21. Chen C, Okayama H (1987) High-efficiency transformation of mammalian cells by plasmid DNA. Mol Cell Biol 7:2745–2752 22. Baekelandt V, Eggermont K, Michiels M, Nuttin B, Debyser Z (2003) Optimized lentiviral vector production and purification procedure prevents immune response after transduction of mouse brain. Gene Ther 10:1933–1940 23. Tonini T, Claudio PP, Giordano A, Romano G (2004) Transient production of retroviral- and lentiviral-based vectors for the transduction of Mammalian cells. Methods Mol Biol 285:141–148 24. Longo PA, Kavran JM, Kim M-S, Leahy DJ (2013) Transient mammalian cell transfection with polyethylenimine (PEI). Methods Enzymol 529:227–240 25. Zhang B, Xia HQ, Cleghorn G, Gobe G, West M, Wei MQ (2001) A highly efficient and consistent method for harvesting large volumes of high-titre lentiviral vectors. Gene Ther 8:1745–1751 26. Reiser J (2000) Production and concentration of pseudotyped HIV-1-based gene transfer vectors. Gene Ther 7:910–913 27. Marino MP, Luce MJ, Reiser J (2003) Smallto large-scale production of lentivirus vectors. Methods Mol Biol 229:43–55 28. Nasri M, Karimi A, Allahbakhshian Farsani M (2014) Production, purification and titration of a lentivirus-based vector for gene delivery purposes. Cytotechnology 66:1031–1038 29. Yamamoto KR, Alberts BM, Benzinger R, Lawhorne L, Treiber G (1970) Rapid bacteriophage sedimentation in the presence of polyethylene glycol and its application to large-scale virus purification. Virology 40:734–744 30. Lee J, Gan HT, Latiff SMA, Chuah C, Lee WY et al (2012) Principles and applications of steric exclusion chromatography. J Chromatogr A 1270:162–170

Lentiviral Particle Production and Concentration ˚ , Frick G (1960) 31. Philipson L, Albertsson PA The purification and concentration of viruses by aqueous polymer phase systems. Virology 11:553–571 32. Vajda BP (1978) Concentration and purification of viruses and bacteriophages with polyethylene glycol. Folia Microbiol 23:88–96 33. Bhat R, Timasheff SN (1992) Steric exclusion is the principal source of the preferential hydration of proteins in the presence of polyethylene glycols. Protein Sci 1:1133–1143 34. Lewis GD, Metcalf TG (1988) Polyethylene glycol precipitation for recovery of pathogenic viruses, including hepatitis A virus and human rotavirus, from oyster, water, and sediment samples. Appl Environ Microbiol 54:1983–1988 35. Atha DH, Ingham KC (1981) Mechanism of precipitation of proteins by polyethylene

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Chapter 5 Generation of Mouse-Induced Pluripotent Stem Cells by Lentiviral Transduction Xiaodong Liu, Joseph Chen, Jaber Firas, Jacob M. Paynter, Christian M. Nefzger, and Jose M. Polo Abstract Terminally differentiated somatic cells can be reprogrammed into an embryonic stem cell-like state by the forced expression of four transcription factors: Oct4, Klf4, Sox2, and c-Myc (OKSM). These so-called induced pluripotent stem (iPS) cells can give rise to any cell type of the body and thus have tremendous potential for many applications in research and regenerative medicine. Herein, we describe (1) a protocol for the generation of iPS cells from mouse embryonic fibroblasts (MEFs) using a doxycycline (Dox)inducible lentiviral transduction system; (2) the derivation of clonal iPS cell lines; and (3) the characterization of the pluripotent potential of iPS cell lines using alkaline phosphatase staining, flow cytometry, and the teratoma formation assays. Key words Mouse-induced pluripotent stem cells, Fibroblasts, Reprogramming, Lentiviral transduction, OKSM, Teratoma assay

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Introduction In 2006, Shinya Yamanaka and Kazutoshi Takahashi reported that by overexpressing four transcription factors, namely, Oct4, Klf4, Sox2, and c-Myc (OKSM), mature differentiated cells such as mouse embryonic fibroblasts (MEFs) can be reprogrammed into embryonic stem (ES) cell-like cells, which they termed induced pluripotent stem (iPS) cells [1]. A year later, Yamanaka and colleagues generated human iPS cells using OKSM [2]. This discovery opened up a new research field and gave rise to a number of paradigms for the use of iPS technology in basic research and medicine. For example, iPS cells can be generated from patients with genetic disorders and then differentiated into a cell type of interest to model and study disease processes and progression

Xiaodong Liu and Joseph Chen contributed equally to this work. Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940, https://doi.org/10.1007/978-1-4939-9086-3_5, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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in vitro [3, 4]. In addition, these cells can be used as screening platforms for the identification and development of novel therapeutic compounds [5]. Furthermore, basic mechanistic studies during the reprogramming process are starting to provide us with a basic framework to understand transcription factor-based reprogramming [6–8]. However, more studies are required to complete this picture. For this purpose, mouse iPS cells remain a suitable and useful model because very stringent functional assays to test their potential such as gestational complementation, germline transmission, tetraploid complementation, and single-cell chimerism can be readily assessed in the mouse system [9]. In addition, cell types like MEFs are relatively easy to maintain and reprogram considerably faster than their human counterparts (~2 weeks vs ~4 weeks, respectively) [7, 10]. A variety of methods have been developed to generate iPS cells since their discovery, and each method has advantages and disadvantages as reviewed by Robinton and Daley [3]. Among these methods, using a lentiviral system is a cost-effective, robust, and efficient approach for transgene delivery since lentiviruses can transduce almost all mammalian cells, including dividing and nondividing cells [11]. Furthermore, using a doxycycline (Dox)inducible polycistronic cassette encoding the four reprogramming factors OKSM, in combination with a Tet-on transactivator (rtTA) [12], allows temporal control of the expression of the Yamanaka factors to obtain transgene-independent bona fide iPS cells. In order to determine if the cells that were reprogrammed are indeed pluripotent, further analyses to verify pluripotency are required. Analyses of iPS cell lines by flow cytometry and through the alkaline phosphatase assay are good and quick screening methods to discard aberrant or differentiated lines. As such pluripotent cell lines should be positive for pluripotent cell surface markers like SSEA1 and EpCAM [6] and express the cytoplasmic enzyme alkaline phosphatase at high levels [13]. A widely accepted and more stringent, albeit time-consuming, assay for a functional demonstration of pluripotency potential is the teratoma formation assay. This assay entails the injection of the iPS cells into flanks of immunocompromised mice to assess their in vivo differentiation potential [14]. The principle of this assay is to test whether the iPS or ES cell lines are capable of generating derivatives of all three germ layers, one of the crucial hallmarks of pluripotency [9]. In the context of this chapter, we describe (1) the generation of iPS cells from MEFs using a Dox-inducible lentiviral transduction system in the serum/LIF condition; (2) the subsequent isolation of clonal iPS cell lines for downstream experiments; and (3) characterization techniques like alkaline phosphatase staining, flow cytometry, and the teratoma formation assay to verify that the resulting iPS cell lines are pluripotent.

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Materials

2.1 Mouse Embryonic Fibroblasts

2.2 Reagents for Lentiviral Transduction

Mouse embryonic fibroblasts (MEFs), isolated from embryos of any genetic background of interest, can be used for reprogramming experiments. Isolation of MEFs from embryonic day (E) 13.5 mouse embryos is described in great detail in [15]. Alternatively, primary MEFs can be purchased commercially from various companies such as Merck Millipore (PMEF-CFL-P1). 1. Generation and titer determination of lentiviral particles harboring the OKSM construct (OKSM plasmid (Millipore, SCR513)) [12] and the m2rtTA (Ef1a-rtTA-GFP plasmid) construct are described in detail in a preceding chapter in this volume [16]. 2. Polybrene infection/transfection reagent 10 mg/mL stock (used at 1:1700 dilution).

2.3 General Cell Culture Reagents

1. MEF culture medium (MEF media): Dulbecco’s modified eagle medium (DMEM), containing 10% fetal bovine serum (FBS) (v/v); 1% GlutaMAX supplement (100
) (v/v); 1% MEM nonessential amino acid solution (100
) (v/v); 100 μM sodium pyruvate; 55 μM β-mercaptoethanol. 2. mESC/iPSC culture medium (iPSC media): KnockOut DMEM containing 15% FBS (v/v) (see Note 1); 1% GlutaMAX supplement (100
) (v/v); 1% MEM nonessential amino acid solution (100x) (v/v); 55 μM β-mercaptoethanol; 1000 unit/ mL recombinant murine LIF. 3. Dulbecco’s phosphate-buffered saline (DPBS) without calcium and magnesium. 4. 0.1% gelatin solution (w/v) is prepared by mixing 1 g of gelatin from porcine skin with 1 L ultrapure water (milli-Q). Autoclave to dissolve and sterilise. 5. Cryopreservation medium: 90% FBS (v/v) and 10% dimethyl sulfoxide (DMSO) (v/v). 6. 0.25% trypsin-EDTA. 7. Doxycycline hyclate diluted to 2 mg/mL in dH2O (1000
stock). 8. Mr. Frosty freezing containers.

2.4 Equipment for iPS Cell Colony Isolation

1. Dissection microscope. 2. Irradiated mouse embryonic fibroblasts (iMEFs) can be generated in house as described in a preceding chapter in this volume [15]. Alternatively, iMEFs can be purchased commercially from Merck Millipore (Merck Millipore, PMEF-CFX). 3. 24-well plates.

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2.5 Reagents and Equipment for Flow Cytometry

1. Anti-mouse BUV395 Thy1.2. 2. Anti-mouse BV421 EpCAM. 3. Anti-mouse SSEA1 Biotin. 4. Streptavidin Pe-Cy7. 5. DRAQ7 viability dye. 6. Tubes for flow cytometry analysis (5 mL Polystyrene RoundBottom Tube with Cell-Strainer Cap) and fluorescence-activated cell sorting (FACS) (5 mL Polypropylene RoundBottom Tube). 7. Flowcytometry instruments such as LSR-II analyzer (BD Biosciences) and BD Influx cell sorter.

2.6 Alkaline Phosphatase Assay

1. 100 mM Tris–HCl pH 9.5.

2.7 Teratoma Formation Assay

1. Dulbecco’s phosphate buffered saline (DPBS) with 1% bovine serum albumin (BSA) (1% BSA/DPBS).

2. Vector Black Alkaline Phosphatase Substrate Kit II (Vector Laboratories).

2. BD Luer-Lock Tip Syringe (1 mL). 3. 23G
1 1/4 in. needles. 4. Immunodeficient mice: NOD-SCID IL2Rgammanull (NSG). 5. Anesthesia unit. 6. Surgical station and tools. 7. Iodine solution. 8. 20, 70, 80, 90, and 100% ethanol. 9. Biopsy-processing cassettes and biopsy foam pads (see Note 2). 10. Chloroform. 11. 4% (w/v) paraformaldehyde (PFA) in PBS (4% PFA). 12. Tube rotator. 13. Shandon™ Paraffin (paraffin wax). 14. Paraffin bath. 15. Embedding molds. 16. Microtome (or cryostat). 17. Water bath at 56  C. 18. Superfrost™ Plus slides. 19. Hematoxylin and Eosin Stain Kit. 20. MIRAX scanner (or light microscope with camera).

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Methods

3.1 Reprogramming of MEFs into iPS Cells

1. Freshly derive MEFs, as described previously [15], or thaw low passage (p0–p2) cryopreserved MEFs for reprogramming experiments. 2. For thawing of cryopreserved MEFs, quickly transfer a cryovial of MEFs (~2–3 million cells) from liquid nitrogen into a 37  C water bath. 3. Once thawed, quickly transfer MEFs with a pipette into a 15 mL centrifuge tube containing 10 mL of pre-warmed MEF media. 4. Pellet the cells by centrifugation at 450
g for 3 min. 5. Remove the supernatant, resuspend the cell pellet (1.5–3
106 cells on average) in 12 mL MEF media, and transfer to a T75 cell culture flask with a vented cap to allow the cells to recover for 1–2 days before starting the reprogramming experiments. 6. One to two days after recovery, cellularize thawed or freshly derived cells as follows: remove MEF media, wash cells once with DPBS to remove traces of serum, and then add 3 mL of 0.25% trypsin-EDTA solution and incubate at 37  C for 3–5 min. Neutralize the enzymatic reaction by the addition of 3 mL MEF media, and pipette medium onto the surface of the flask 3–5 times to dissociate the MEFs. Transfer the cell suspension into a 15 mL tube. 7. Perform cell counting using a hemocytometer or automated cell counter. 8. It is recommended to seed cells at a range of 0.5–2.5
103 cells/cm2 in gelatin-coated 6-well plates (see Note 3) containing MEF media (see Note 4) (Fig. 1).

Fig. 1 Schematic depicting MEF to iPSC reprogramming protocol

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9. Twenty-four hours later, perform lentiviral transduction as follows: prepare viral mix by adding polybrene (1:1700), lentivirus-m2rtTA (mean occurance of infection [MOI] of 2), and lentivirus-OKSM (MOI of 2) (see Note 5 and Chap.4) in 2 mL iPSC media; following this aspirate culture media from wells to be infected, and replace with the iPSC media containing the viral mix (Fig. 1). 10. Perform spin inoculation by transferring the plate(s) into a centrifuge, and spin for 60 min at 750
g at room temperature. Afterward, transfer the plate(s) into a 37  C incubator with 20% O2 and 5% CO2 (see Note 6). 11. On the next day, remove virus-containing media, and replace with fresh iPSC media supplemented with doxycycline (2 μg/ mL) to initiate the reprogramming process (3 mL of media per well of 6-well plates). 12. Perform media changes every other day using doxycyclinesupplemented iPSC media for the first 6 days of reprogramming. 13. After 6 days, daily media changes are recommended due to increased cell densities. Alternatively, add 6 mL of media into one well of a 6-well plates if media changes can only be performed every other day. 14. Expected changes in cell morphology during reprogramming are shown in Fig. 2. iPS cell colonies should be identifiable after approximately 12 days, and it is recommended to transfer the cells into doxycycline-free iPSC media for another 4 days to remove aberrant iPS cell colonies that are still dependent on forced transgene expression. After this period proceed to isolate clonal lines by colony picking. 3.2 Isolation of Clonal iPS Cell Lines by Colony Picking

1. Six hours to 1 day before colony isolation, prepare recipient plates by seeding iMEFs onto gelatin-coated 24-well plates at a density of 2
104 cells/cm2 in 1 mL of iPSC media per well (see Note 3).

Fig. 2 Timeline of reprogramming from MEFs to iPSCs. Representative brightfield images of reprograming cultures on days 0, 3, 6, 12, and 16. Scale bar ¼ 25 μM

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Fig. 3 Establishment of clonal iPSC lines. (a) (i) Identify dome-shaped colony in culture (Day 16 of reprogramming). (ii) Isolate colony with a pipette by removing cells surrounding the colony. (iii) Lift colony with pipette, gently nudge side to detach cells from well plate. (iv) Dissociate cells by transferring colony into 1.5 mL tube containing 0.25% trypsin. Gently pipette to dissociate colony further. After 2–4 min, transfer contents of 1.5 mL tube into a 24-well with iMEFs. (v) Bright field image of cells after 3 days in culture after transfer. (b) Bright field image of iPSC colonies at passage 2. (c) FACS analysis of iPSC clonal line at passage 2. (d) Alkaline-phosphatase staining of iPSC colonies at passage 2. Scale bar ¼ 25 μM

2. Rinse the 6-well plate(s) containing the reprogrammed cultures with DPBS, and then add 1 mL of warm DPBS into each well. Colonies can be picked under an inverted light microscope or a dissection microscope (see Notes 7 and 8). 3. Identify a reasonably isolated (i.e., not fused to other colonies) iPS cell colony with characteristically dome-shaped morphology (Fig. 3a(i)). 4. Using a 20 μL pipette, draw a circle around the colony with a sterile tip to detach the colony from surrounding fibroblasts (Fig. 3a(ii)). 5. Nudge the colony with the tip to gently lift it from the underlying tissue culture plastic (Fig. 3a(iii)). 6. Aspirate the free-floating colony with the pipette in a 12.5 μL volume, and transfer into a 1.5 mL tube containing 50 μL of 0.25% trypsin-EDTA (Fig. 3a(iv)). 7. After 2–4 min, gently further dissociate the transferred colony by pipetting the medium within the tube several times. 8. Transfer the cell suspension from the tube directly into a well of the prepared 24-well plate with the iMEFs (Fig. 3a(iv)).

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9. Repeat steps 3–8 with other colonies to generate more potential clonal lines (see Note 9). 10. Change media with fresh iPSC media 24 h after colony picking. 11. New, dome-shaped colonies should form in the recipient 24-wells after 2 days (Fig. 3a). 12. Expand new clones through routine passaging to propagate the iPS cells (Fig. 3b) (see Notes 10, 11 and 12) as described for mouse embryonics stem cells in a preceding chapter in this volume [17]. 3.3 Characterization of Clonal iPS Cell Lines 3.3.1 Flow Cytometry

iPS cells can be analyzed by flow cytometry or purified through FACS using positive markers associated with pluripotent cells and a MEF marker they are negative for. For example, iPS cells can be FACS depleted from Thy1.2-positive feeder cells and enriched for SSEA1- and EpCAM-positive pluripotent cells (Fig. 3c). Only iPS lines that express SSEA1 and EpCAM should be considered as pluripotent. By extracting the SSEA1/EpCAM double-positive population by FACS, undifferentiated iPS cells can be effectively removed and the purified cells subsequently used for purposes such as differentiation assays. Preparation of single-colour compensation samples, antibody labelling process, and gating strategies were described in great detail previously [15, 18, 19]; setting up the voltages at the cell analyzer or sorter is to be performed by either an experienced user or a dedicated FACS operator as described previously [15, 19].

3.3.2 Alkaline Phosphatase Staining

Established clonal iPS cell lines (passaged 4–5 times to enrich for pluripotent cells) can be submitted to an alkaline phosphatase assay. For this assay, cells can be seeded in a 24-well or 12-well format. When cultures are confluent (roughly 70% confluent colonies), remove iPSC media from the wells, wash once with DPBS, and then stain with alkaline phosphatase assay (Vector Black Alkaline Phosphatase Substrate Kit II, SK-5200) according to manufacturer’s instructions. iPS cell colonies will stain black (Fig. 3d).

3.3.3 Teratoma Formation Assay

iPS cells that have been established for at least 4–5 passages are normally subjected to the teratoma assay. Upon injecting iPS cells subcutaneously, they should start proliferating and differentiate into the cell types of all three germ layers and thereby form a growth, the so-called teratoma. If the resulting teratoma indeed contains cells from the three germ layers (ectoderm, mesoderm, and endoderm), the iPS cell line is deemed to be pluripotent. In order to perform a teratoma formation assay, approval has to be obtained from the Animal Welfare Committee or other regulatory bodies before conducting any of these experiments.

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1. Expand subcloned iPS cell lines into one T25 flask in the presence of iMEF feeders. 2. When the cells are ready to be passaged, remove iPSC media, then wash once with DPBS, and add 3 mL trypsin-EDTA solution for 3–5 min at 37  C. 3. Neutralize the enzymatic reaction by the addition of 3 mL of MEF media, and pipette up and down three to five times to dissociate the iPS cells, and then transfer the cells to a 15 mL tube. 4. Pellet the cells by centrifugation at 450
g for 3 min, and then resuspend the cell pellet in 12 mL iPSC media. 5. To purify and enrich iPS cells prior to the assay, an iMEF feeder depletion step is performed. Transfer the cells onto a new gelatin-coated T75 flask (see Note 3), and place the flask into a 37  C incubator for 45–50 min. iMEF feeders and differentiated cells attach to the gelatin within 45 min of incubation time, whereas iPS cells require 2–4 h to attach. 6. Transfer the supernatant (containing the nonadherent iPS cells) to a 15 mL tube, pellet the cells by centrifugation at 450
g for 3 min, and then resuspend the cell pellet in 1–3 mL iPSC media to perform cell counting using a hemocytometer or automated cell counter. 7. Transfer ~1
106 cells into an Eppendorf tube, and pellet the cells by centrifugation at 450
g for 3 min. 8. Resuspend the cell pellet in 200 μL prepared injection mix containing 1% BSA/PBS or iPSC media, and keep on ice (see Note 13). 9. Set up the anesthesia apparatus and surgical station in the animal facility according to the instructions provided by the manufacturer (see Note 14). 10. Before anesthetizing the mice, fill the anesthetic apparatus’ induction chamber by setting the oxygen flow rate at 4 L/ min and isoflurane at 4–5% for 1–2 min. 11. Once the induction chamber is filled, decrease the oxygen flow rate and isoflurane to maintenance level (0.4 L/min flow rate and 2–3% isoflurane). 12. Anesthetize the NGS mice using 2–3% isoflurane and 0.4 L/ min oxygen flow rate for anesthesia and its maintenance once they are unconscious (Fig. 4a). 13. Remove one mouse from the induction chamber, and place a nose cone on it to provide consistent anesthetic air flow. 14. Wipe the skin around the dorsal flanks of the mouse with iodine solution and then 70% ethanol.

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Fig. 4 Overview of teratoma assay. (a) Schematic depicting injection of iPSCs into NGS mouse and resulting teratoma formation and isolation. (b) Hematoxylin and eosin (H&E) staining of teratoma sections and visualization of three representative tissue types per germ layer. Scale bar ¼ 25 μM

15. Using a 1 mL syringe with 23-gauge needle, slowly draw 200 μL of the prepared cell suspension, remove bubbles from the syringe, and proceed immediately to the next step. 16. Slowly inject 200 μL of the cell suspension subcutaneously into the dorsal flanks of the NGS mouse. This can be done by pinching a part of the mouse’s flank using the thumb and the index finger (Fig. 4a) followed by inserting the needle between

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the fingers. After ensuring that the needle is positioned subcutaneously, slowly inject the cell suspension. 17. After injection, keep the mouse anesthetized for ~10 min. 18. Monitor injected NGS mice at least once a week for 3–5 weeks to track growth at the injection site. 19. When teratoma formation is evident at a stage when the growth is still smaller than or around 1 cm3, it is recommended (and an ethical requirement) to cull the mouse and excise the teratomas (see Note 15). 20. Transfer the teratoma to a 50 mL tube, and submerge into an ample volume of 4% PFA. 21. Fix tissue overnight at 4  C on a tube rotator. 22. On the following day, wash the teratomas with DPBS, and cut into 2–5 slices using scalpels (the choice of the number of the slices depends on the size of the teratoma and the personal choice of the experimenter). 23. Place the teratoma slices into a labelled biopsy-processing cassettes with biopsy foam pads, and submerge in a container with 70% ethanol for sectioning. 24. Dehydrate tissue using graded alcohols (70, 80, 90, 100%) by successively incubating in each solution (starting with 70%) for at least 20 min, and then submerge in fresh chloroform solution twice for 1 h each. 25. Incubate cassettes twice in paraffin wax (molten at 56  C) for at least 1 h (see Note 16). 26. After the tissue has been infiltrated with paraffin wax (step 25), place cassette in a paraffin bath at 58  C for 15 min to melt away residual wax. 27. Open cassette and pick tissue out of the cassette with a pair of forceps. Transfer tissue onto embedding molds, and position it preferably in the center of the depression of the mold. When tissue is placed in the desired orientation, fill remaining portion of mold with hot paraffin to desired volume. Place mold in 20  C freezer for at least 3 h before separating the tissue block from the mold. 28. Section paraffin-embedded tissue block using a microtome (or cryostat). Cut sections at 2–5 μm according to the manufacturer’s instructions (see Note 17). 29. Using a pair of forceps, transfer sections onto a 20% ethanol bath (20% v/v ethanol in water), and then transfer sections onto a heated water bath of 56  C. Collect sections onto labelled slides, and leave to drain for 10 min (leave upright). Leave slides to dry overnight on a slide rack in an oven at 40  C.

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30. Hematoxylin and eosin staining should be performed according to histology facility’s or manufacturer’s instruction. Refer to Nelakanti et al. [20] for a detailed protocol of the staining technique. 31. To obtain high-quality images and identify tissues of all three germ layers, use a MIRAX scanner (or any other comparable slide scanners). 32. Score the images for the presence of derivatives of the three germ layers. Examples of representative tissues of each germ layer are provided in Fig. 4b.

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Notes 1. It is important to note that the batch and quality of FBS are crucial to support pluripotent stem cell culture and reprogramming. Not all FBS batches are suited for reprogramming. If batch testing is not possible, procure ES-qualified FBS (in general more expensive). 2. Contact local histology platforms in advance for submission of specimens for subsequent processing of teratoma assays. 3. Sterile 0.1% gelatin solution (w/v) is used to coat the plates or flasks to provide attachment support to the MEFs. It is recommended to add adequate 0.1% gelatin to cover the surface of the plates or flasks (e.g., 2 mL for a well of 6-well plate) and incubate for 30 min or more at 37  C to coat. 4. Starting cell density has a dramatic impact on the reprogramming efficiency. We recommend trying a range of cell densities to determine the optimal density for reprogramming experiments for that particular cell line. 5. MOI can be calculated based on the target cell number to be transduced multiplied by the number of infective viral particles per microliter of viral concentrate. When handling viruses, please ensure that transduction is performed in a class II hood and the user is double-gloved for safety and protection. Refer to the previous chapter [16] for further details. 6. This reprogramming protocol is optimized for culture in normoxic conditions. 7. It is recommended to pick iPSC colonies using a dissection microscope to derive clonal lines. Colony picking should be performed in a sterile condition (e.g. inside a hood; and the surface and the hood, and all tools and equipment should be cleaned thoroughly with 80% ethanol before colony picking). It is easiest to pick colonies a minimum format size of a 6-well plate.

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8. Before picking colonies, seed iMEFs in 24-well plates 24 h in advance. For better visualization of colonies (and to facilitate dissociation of colonies after picking later), remove iPSC media, and wash cells with PBS. Aspirate and add 2 mL of PBS in each 6-well plate (or 10 mL in a 10 cm dish). When selecting a colony to pick, avoid colonies with a flattened/ differentiated appearance. 9. In order to derive 3–5 clonal lines, we advise picking a minimum of 20 colonies as not all colonies will expand after this process. 10. Three to five days after establishing the clonal lines, cells can be expanded into larger formats (from 24-well into a 6-well plate) with iMEF feeders seeded 24 h prior to expansion. 11. It might be necessary to passage new clonal lines for a few times (at least 2–3 passages) to get rid of cells that are not fully reprogrammed/partially differentiated and enrich for true iPSC colonies with dome-shaped morphology. 12. For cryopreservation and routine passaging of iPSCs, it is recommended to follow the protocol described in a previous chapter in this volume [17]. 13. Matrigel diluted 1:3 in DMEM/F12 can be used to increase teratoma formation as it enhances cell engraftment after injection [21]. Keep thawed Matrigel on ice at all times to prevent solidification. 14. Depending on the equipment, delivery method and time of anesthetic exposure to the animal can vary. It is recommended that this part of the protocol be performed based on the facility’s preferences and equipment’s instruction manual. 15. Extraction of the formed teratomas is explained in detail by Nelakanti et al. [20]. 16. It is important to take note of the temperature of molten wax as high temperatures and prolonged exposure to molten wax may destroy antigens in tissue. Keep the wax at the lowest temperature possible in its molten state. Preferably, maintain this temperature at 2  C above the melting point of 56  C. 17. The operation of the microtome (or cryostat) and sectioning of slides should be performed by trained personnel in accordance to manufacturer’s instructions for the machines. References 1. Takahashi K, Yamanaka S (2006) Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell 126:663–676

2. Takahashi K, Tanabe K, Ohnuki M, Narita M, Ichisaka T et al (2007) Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell 131:861–872

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3. Robinton DA, Daley GQ (2012) The promise of induced pluripotent stem cells in research and therapy. Nature 481:295–305 4. Chen J, Nefzger CM, Rossello FJ, Sun YBY, Lim sm et al (2018) Fine tuning of canonical wnt stimulation enhances differentiation of pluripotent stem cells independent of β-catenin-mediated T-cell factor signaling. Stem Cells. 36:822–833. 5. Avior Y, Sagi I, Benvenisty N (2016) Pluripotent stem cells in disease modelling and drug discovery. Nat Rev Mol Cell Biol 17:170–182 6. Polo JM, Anderssen E, Walsh RM, Schwarz BA, Nefzger CM et al (2012) Molecular roadmap of reprogramming somatic cells into iPS cells. Cell 151:1617–1632 7. Nefzger CM, Rossello FJ, Chen J, Liu X, Knaupp AS et al (2017) Cell type of origin dictates the route to pluripotency. Cell Rep 21(10):2649–2660 8. Knaupp AS, Buckberry S, Pflueger J, Lim SM, Ford E et al (2017) Transient and permanent reconfiguration of chromatin and transcription factor occupancy drive reprogramming. Cell Stem Cell 21:834–845 9. De Los Angeles A, Ferrari F, Xi R, Fujiwara Y, Benvenisty N et al (2015) Hallmarks of pluripotency. Nature 525:469–478 10. Liu X, Nefzger CM, Rossello FJ, Chen J, Knaupp AS et al (2017) Comprehensive characterization of distinct states of human naive pluripotency generated by reprogramming. Nat Methods 14:1055–1062 11. Sakuma T, Barry Michael A, Ikeda Y (2012) Lentiviral vectors: basic to translational. Biochem J 443(3):603–618 12. Sommer CA, Stadtfeld M, Murphy GJ, Hochedlinger K, Kotton DN, Mostoslavsky G (2009) iPS cell generation using a single lentiviral stem cell cassette. Stem Cells 27:543–549

13. Singh U, Quintanilla RH, Grecian S, Gee KR, Rao MS, Lakshmipathy U (2012) Novel live alkaline phosphatase substrate for identification of pluripotent stem cells. Stem Cell Rev 8:1021–1029 14. Wesselschmidt RL (2011) The teratoma assay: an in vivo assessment of pluripotency. Methods Mol Biol 767:231–241 15. Nefzger CM, Alaei S, Knaupp AS, Holmes ML, Polo JM (2014) Cell surface marker mediated purification of iPS cell intermediates from a reprogrammable mouse model. J Vis Exp (91):e51728. 16. Larcombe MR, Manent J, Chen J, Mishra K, Liu X, Nefzger CM (2019) Production of high titer lentiviral particles for stable genetic modification of mammalian cells. Methods Mol Biol 1940:47–61 17. Paynter JM, Chen J, Liu X, Nefzger CM (2019) Propagation and maintenance of mouse embryonic stem cells. Methods Mol Biol 1940:33–45 18. Nefzger CM, Alaei S, Polo JM (2015) Isolation of reprogramming intermediates during generation of induced pluripotent stem cells from mouse embryonic fibroblasts. Methods Mol Biol 1330:205–218 19. Nefzger CM, Jarde T, Rossello FJ, Horvay K, Knaupp AS et al (2016) A versatile strategy for isolating a highly enriched population of intestinal stem cells. Stem Cell Reports. 6:321–329. 20. Nelakanti RV, Kooreman NG, Wu JC (2015) Teratoma formation: a tool for monitoring pluripotency in stem cell research. Curr Protoc Stem Cell Biol 32:4A.8.1–4A.817 21. Polanco JC, Ho MS, Wang B, Zhou Q, Wolvetang E et al (2013) Identification of unsafe human induced pluripotent stem cell lines using a robust surrogate assay for pluripotency. Stem Cells 31:1498–1510

Chapter 6 Gene Editing of Mouse Embryonic and Epiblast Stem Cells Tennille Sibbritt, Pierre Osteil, Xiaochen Fan, Jane Sun, Nazmus Salehin, Hilary Knowles, Joanne Shen, and Patrick P. L. Tam Abstract Efficient and reliable methods for gene editing are critical for the generation of loss-of-gene function stem cells and genetically modified mice. Here, we outline the application of CRISPR-Cas9 technology for gene editing in mouse embryonic stem cells (mESCs) to generate knockout ESC chimeras for the fast-tracked analysis of gene function. Furthermore, we describe the application of gene editing directly to mouse epiblast stem cells (mEpiSCs) for modelling germ layer differentiation in vitro. Key words CRISPR-Cas9, Embryonic stem cells, Epiblast stem cells

1

Introduction Conventional methods to perform genome editing in embryonic stem cells (ESCs) such as gene targeting by homologous recombination are inefficient and time-consuming, taking up to months to a year to generate a stock of genetically modified mice for experimental studies of gene function. Recently, advances in genetic manipulation technology have enabled the quick and efficient generation of edited genomes. Nucleases fused to specific DNA-binding domains, such as transcription activator-like effector nucleases (TALENs) and zinc-finger nucleases (ZFNs), have facilitated highly specific gene editing that can be achieved expeditiously, but the utility and the cost-effectiveness of these technologies remain challenging [1, 2]. CRISPR-Cas9 technology has recently arisen to the fore as the most amenable technique to perform genome editing [3]. In addition to producing the desired genetic modification within a shorter time frame, this technology is efficient, is relatively straightforward in design, and can be applied to both cell lines and whole organisms. Here, we describe the use of CRISPR-Cas9 for genome editing in mouse (m) ESCs and EpiSCs. Several resources are available to design the single-guide RNAs (sgRNAs) to target the gene of

Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940, https://doi.org/10.1007/978-1-4939-9086-3_6, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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interest, such as the Zhang Lab website (http://crispr.mit.edu/) or the Broad Institute’s Genetic Perturbation Platform (https:// portals.broadinstitute.org/gpp/public/). Commonly, a combination of these resources is used to select the optimal sgRNAs that target within the first 200 bp of the start codon in the coding region and have high specificity with low off-target effects. Furthermore, the sgRNA must be followed on the 30 end by the 3 bp NGG protospacer adjacent motif (PAM). The cloning of the sgRNA for targeting the gene of interest into plasmids containing the Cas9 nuclease, a selection marker (puromycin or GFP), and a sgRNA scaffold (pSpCas9(BB)-2A-Puro or pSpCas9(BB)-2A-GFP) can be ready within a week for use in the electroporation of stem cells [3]. Within another 3 weeks, mESCs or mEpiSCs harboring target mutations can be established, which can be taken further for clone selection by puromycin resistance or GFP expression. The subsequent expansion of positive clones for cryopreservation and genotyping usually takes at most another 4 weeks. We have previously generated chimeric embryos derived predominantly from the targeted genome-edited ESCs by microinjecting the stem cells into eight-cell preimplantation mouse embryos which are transferred to pseudopregnant mice for further intrauterine development [4, 5]. In addition to the capability to generate mid-gestation embryos of the edited genotype for the phenotypic analysis and elucidation of gene function, this approach has also been employed to derive genome-edited mEpiSCs from the chimeric embryos [4]. The generation of ESC-derived chimeras enables the transition of the stem cells through the development of the inner cell mass to an epiblast state from which authentic mEpiSCs can be derived. This approach bypasses the technical barrier that the transition from mESCs to mEpiSCs of a proper epiblast-like state has not been achieved in vitro. However, derivation of mEpiSCs with an edited genotype via chimeric embryo production is timeconsuming and laborious. Moreover, the efficiency of derivation of mEpiSCs is lower than that of mESCs, resulting in increased animal usage. This technically demanding protocol and the low efficiency of mEpiSC derivation could be replaced by a genome editing protocol that applies directly to mEpiSCs. Efforts to perform gene editing directly on mEpiSCs have been hampered by the technical difficulty in maintaining the mEpiSCs (1) without feeders (that interferes with the selection of edited cells) and (2) as single cells for clonal selection of editing events. These hurdles are reputed to be the different requirements for maintaining the mEpiSCs due to their primed pluripotent state and the poising of cell differentiation. We have overcome these issues with the use of conditioned mEpiSC medium obtained from culture with mouse embryonic fibroblasts (MEFs) and chemical reagents that enhance the viability of single mEpiSC cells during clonal selection in vitro.

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Gene editing of mESC and mEpiSCs by these approaches is efficient. In the case of puromycin selection, 80–100% of clones that successfully grow contain a mutation on at least one allele of the gene of interest. The success rate of generating biallelic frameshift mutations varies from 35% to 100%. With GFP selection, we generally find mutations in 60–80% of the sequenced clones, with 10–20% of these containing biallelic frameshift mutations. For several genes that have been edited by CRISPR-Cas9 technology, we were able to confirm the absence of protein for the biallelic frameshift mutations in mESC clones and the reduction of protein in some monoallelic mESC clones. For mEpiSCs, sequencing analysis of a mixed population of clones revealed that desirable editing events have taken place. In this chapter, we describe procedures for generating frameshift mutations in mESCs and directly in mEpiSCs (Fig. 1).

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Materials Prepare and store all reagents at room temperature unless indicated otherwise in protocol or packaging of reagent. Diligently follow all safety and waste disposal regulations when performing experiments.

2.1 CRISPR Plasmid Components

1. pSpCas9(BB)-2A-Puro (PX459) V2.0 (Addgene plasmid #62988) or pSpCas9(BB)-2A-GFP (PX458) (Addgene plasmid #48138) (gift from Feng Zhang) with the sgRNA ligated into the plasmid according to the protocol described in [3].

2.2 Cell Culture Components

1. 100 nucleosides: 0.16 g adenosine, 0.146 g cytidine, 0.17 g guanosine, 0.146 g uridine, 0.048 g thymidine in 200 mL H2O. Swirl at 37  C for several hours or overnight. Filter before use and store aliquots at 4  C. Warm to 37  C prior to use. 2. Mouse embryonic stem cell (mESC) culture medium: DMEM, 12.5% heat-inactivated fetal calf serum (FCS), 1000 U/mL leukemia inhibitory factor, 0.1 mM β-mercaptoethanol, 1 nonessential amino acids, 1 nucleosides. Store at 4  C. 3. Mouse embryonic fibroblast (MEF) culture medium: DMEM, 10% FCS, 0.1 mM β-mercaptoethanol. Store at 4  C. 4. Mouse epiblast stem cell (mEpiSC) medium: Knockout Serum Replacement, 1 nonessential amino acids, 1 GlutaMAX™, 0.1 mM β-mercaptoethanol. Store at 4  C. Media is supplemented with 10 ng/mL recombinant human FGF2 and 20 ng/mL recombinant Human/Mouse/Rat Activin A extemporaneously.

PX458

PX459 V2.0 2A

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Pu ro R

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20mer oligo cloning site

20mer oligo cloning site

Select GFP positive clones ~3-4 days after transfection

Rest for 24 h Puromycin select (48 h mESCs, 24 h mEpiSCs)

Select clones (add ROCKi to mEpiSCs for 24 h)

mESCs Freeze clones & grow on gelatin to extract genomic DNA for genotyping mEpiSCs (conditioned medium + ROCKi)

PCR amplify, Sanger sequence & analyse mutation using TIDE mEpiSCs

mESCs *

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Clone PCR products into pGEM®-T Easy Vector & Sanger sequence PAM

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Positive clone Allele 1: 7 bp deletion

Allele 2: 2 bp deletion

Thaw, expand and freeze positive clones

Validate clones Sanger sequence for final validation

mRNA expression analysis (e.g. RT-qPCR)

Protein expression analysis (e.g. western blot)

Fig. 1 Workflow for genome editing in mESCs and mEpiSCs using PX459 V2.0 or PX458. PX459 V2.0 and PX458 contain the S. pyogenes Cas9 nuclease ORF and a cloning backbone for the sgRNA. PX459 V2.0 contains the 2A-Puro ORF directly downstream of the Cas9 ORF, while PX458 contains the 2A-EGFP ORF

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5. Conditioned mEpiSC medium: mEpiSC medium is incubated overnight at 37  C on MEFs at a density of 9  104 cells/cm2. The following day, the medium is filtered. The conditioned medium can be stored at 20  C for up to 1 month. Use only for cell culture without MEFs, and supplement with Activin A and FGF2 as mentioned in step 4. 6. 2 freeze medium: 50% heat-inactivated FCS, 30% culture medium, 20% DMSO. Make up fresh each time and keep at 4  C. 7. TrypLE™ Select. 8. Collagenase IV: Make a stock of 2 mg/mL in mEpiSC medium. 9. ROCK inhibitor (Y-27632, TOCRIS): ROCKi is used only when mEpiSCs are seeded in a single cell suspension to improve cell viability. Add to mEpiSC media to a final concentration of 10 μM only for 24 h following TrypLE™ Select treatment. 10. Dulbecco’s (D)PBS. 11. Puromycin (10 mg/mL). 12. 0.1% gelatin: Mix 1 g gelatin from bovine skin with 1 L H2O. Autoclave and filter before use. 13. MicroTube Rack System™ Tubes. 2.3 Electroporation Components

1. Neon® Transfection System. 2. Neon® Transfection System 100 μL kit.

ä Fig. 1 (continued) directly downstream of the Cas9 ORF, allowing for puromycin or EGFP selection, respectively. After electroporation of the plasmids containing the sgRNA of interest, clones are left to grow. In the case of mEpiSCs, clones are grown in mEpiSC medium supplemented with 10 μM ROCKi for 24 h, while mESCs are grown in mESC medium. Once established, clones are selected and grown in a 96-well plate containing MEFs. One-third (mESCs) or one-half (mEpiSCs) of the cells are cryopreserved, while the remainder are grown on 0.1% gelatin for several passages to remove MEFs for genotyping of the mutation. In the case of mEpiSCs grown on gelatin, clones are passaged in conditioned medium with 10 μM ROCKi added after each passage for 24 h. Clones are lysed and subjected to PCR and Sanger sequencing, followed by analysis by TIDE to decompose the mutations on each allele. Examples of the chromatograms indicating gene editing are shown for mESCs and mEpiSCs; overlapping peaks in the sequence of several bases upstream of the PAM indicate mutations in at least one allele. PCR products are then cloned into pGEM®-T Easy Vector for confirmation of the gene editing event. An example of a confirmed biallelic frameshift mutation is shown for mESCs. Final validation of the mutation involves thawing out the cryopreserved clones of interest, expanding them from a 48-well plate to a 6-well plate over several weeks, and repeating the process of lysis, PCR, cloning, and Sanger sequencing. Final validation of the knockout is confirmed by mRNA expression analysis (RT-qPCR) using primers that overlap the indel or protein expression analysis (Western blot). * indicates the indel site

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2.4 Cell Lysis Components

1. Cell lysis buffer: 50 mM Tris–HCl, pH 8.0, 1 mM EDTA, 0.5% Tween-20. Freshly add 200 μg/mL Proteinase K.

2.5 PCR Amplification Components

1. BioMix™ (Bioline).

2.6 Agarose Gel Electrophoresis and PCR Purification Components

2. Forward and reverse primers spanning the mutation site (desired product size ~700 bp, with sufficient sequence flanking the mutation). 1. Agarose. 2. RedSafe™ Nucleic Biotechnology).

Acid

Staining

Solution

(iNtRON

3. HyperLadder™ 100 bp (Bioline). 4. 1 TAE buffer: Make up 50 TAE buffer by combining 424 g Tris base, 57.1 mL acetic acid, and 100 mL 0.5 M EDTA (pH 8.0), and make up to 1 L in H2O. To make 1 TAE buffer, combine 40 mL 50 TAE buffer with 1.96 L H2O. 5. Wizard SV Gel and PCR Clean-Up System (Promega).

2.7 Ligation, Bacterial Transformation, and Plasmid Purification Components

1. pGEM®-T Easy Vector System (Promega). 2. α-Select Silver Competent Cells (Bioline). 3. Luria-Broth (LB): Combine 4 g Tryptone (vegetable), 2 g Bacto™ Yeast Extract, and 4 g NaCl, and add 400 mL H2O. Autoclave. 4. LB ampicillin: As for LB, add ampicillin to a final concentration of 100 μg/mL once cooled enough to touch. 5. Combine 4 g Tryptone (vegetable), 2 g Bacto™ Yeast Extract, and 4 g NaCl, and add 400 mL H2O. Autoclave. 6. LB ampicillin agar plates: As for LB, add 6.5 g Bacto™ Agar. Autoclave. Cool to the point that you can touch. Add ampicillin to a final concentration of 100 μg/mL. Under sterile conditions pour ~20 mL into bacterial plates. Store at 4  C for up to 1 month. 7. LB ampicillin agar plates for bacterial transformations using pGEM®-T Easy Vector: As for LB ampicillin agar plates, add fresh 0.5 mM IPTG and 50 μg/mL X-Gal to the plates after the agar has set. Spread across the plate, and hold with the lid off in a 37  C incubator until plates have dried (~20 min). 8. ISOLATE II Plasmid Mini Kit (Bioline).

3

Methods Carry out all procedures at room temperature unless otherwise specified. Warm all cell culture media components to room

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temperature or 37  C before use. Unless otherwise stated, all cell culture is done under sterile conditions in a laminar flow hood. Follow all waste disposal regulations diligently when disposing waste materials. 3.1 Electroporation of mESCs with PX459 v2.0 or PX458

1. 24 h before electroporation, seed a 10 cm plate (two plates for PX458) with 1.5  106 MEFs in MEF medium, and place into the 37  C incubator. 2. On the day of electroporation, set up and save the following parameters on the Neon® Transfection System: Pulse voltage, 1200 V; pulse width, 20 ms; pulse number, 2 3. Set up the electroporation equipment according to the manufacturer’s instructions. 4. Remove 5 μg of PX459 V2.0 or 3 μg PX458 with the specific sgRNA ligated in from the stock tube, and aliquot into a 1.5 mL tube (see Note 1). Put aside. 5. Change the medium on the pre-seeded MEFs to 10 mL mESC medium. Label the plate with all the required details for the electroporation. Place back into the incubator until required. 6. Take the mESCs out of the 37  C incubator, aspirate the medium, and rinse with 4 mL DPBS. 7. Add 3 mL TrypLE™ Select to the mESC plate, and place back into the incubator for 5 min. 8. Take the plate out of the incubator and add 6 mL mESC medium. Inactivate and break into a single cell suspension (see Note 2). 9. Centrifuge cells at 1000 rpm (200  g) for 5 min. 10. Once spinning has completed, remove the medium leaving only the cell pellet, and then resuspend in 10 mL DPBS. Count the cells. 11. For electroporation with pX459 V2.0, remove 5  106 cells, and pipette into a new 15 mL tube. For electroporation with pX458, remove 1  106 cells, and pipette into a new 15 mL tube (see Note 3). 12. Centrifuge at 1000 rpm (200  g) for 5 min. During this time, add 3 mL E2 buffer into a Neon® Tube, and place into the Neon® Pipette Station. 13. When spinning has completed, remove all DPBS leaving only the cell pellet. 14. Resuspend cells in 120 μL R buffer until they are in a single cell suspension. 15. Transfer the cells to the 1.5 mL tube containing the plasmid from step 4. Mix well by gentle pipetting.

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16. Insert a 100 μL Neon® tip into the Neon® Pipette, and collect the cell and DNA mixture. Ensure there are no bubbles (see Note 4). 17. Move the Neon® Pipette containing the cells and DNA to the Neon® Pipette Station. 18. Load the saved parameters on the Neon® Transfection System from step 2. Press “start.” Once completed, the unit will display “complete.” 19. Transfer the electroporated cells onto the pre-seeded MEF plate. Evenly distribute the cells by rocking back and forth and side-to-side (not swirling). 20. If there are still substantial numbers of cells left in the 1.5 mL tube, repeat steps 14–19. 21. Return the plate to the incubator. 3.2 Electroporation of mEpiSCs with PX459 v2.0

1. 24 h before electroporation, seed 2  6 cm plates with 1  106 MEFs in MEF medium, and place into the 37  C incubator. 2. Repeat steps 2–4 from Subheading 3.1. 3. Rinse the plates containing pre-seeded MEFs with 2 mL DPBS, and then add 4 mL of mEpiSC medium supplemented with 10 μM ROCKi. Label the plate with all the required details for the electroporation. Place back into the incubator until required. 4. Add 2 mL of Collagenase IV to the plates containing mEpiSCs, and place back into the 37  C incubator for 10 min. 5. Take the mEpiSC plates out of the incubator, and add 2 mL mEpiSC medium to detach the colonies from the feeder layer. 6. Spin the clumped suspension at 1000 rpm (200  g) for 30 s. 7. Resuspend the cell pellet in 1 mL TrypLE™ Select, and incubate at room temperature for 2 min. 8. Break the cell clumps using a P1000 tip and add 2 mL mEpiSC medium. 9. Centrifuge cells at 1000 rpm (200  g) for 5 min. 10. Remove the medium leaving only the cell pellet, and then resuspend in 2 mL DPBS. Count the cells. 11. Proceed with step 11 from Subheading 3.1 for electroporation of mEpiSCs.

3.3 Puromycin Selection of mESCs and mEpiSCs Transfected with pX459 v2.0

1. 24 h post-electroporation, feed the electroporated mESC or mEpiSC media containing 2 μg/mL or 1 μg/mL puromycin, respectively (see Note 5). Repeat this the following day for mESCs with fresh mESC media (see Note 6).

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2. 72 h post-electroporation of the mESCs (48 h postelectroporation for mEpiSCs), wash the cells twice with DPBS, and feed with mESC or mEpiSC media. 3. For the electroporated mESCs, seed 1.5  106 MEFs in mESC medium onto the plate. For mEpiSCs, seed 1  106 MEFs in mEpiSC medium onto each plate (see Note 7). 4. Continue DPBS wash and feed daily. 3.4 Clone Picking of pX459 v2.0Transfected mESCs and mEpiSCs

1. The day before colonies are ready to be picked, seed 8.42  104 MEFs for mESCs or 6.24  104 MEFs for mEpiSCs into the wells of a 96-well plate (see Note 8). The number of wells to seed depends on the number of clones to be picked. If possible, try to pick 20–30 clones. Place the plate into the 37  C incubator. 2. On the day of picking the mESCs, change the media in the 96-well plate to 80 μL mESC media. For mEpiSCs, rinse once with 100 μL DPBS to remove any trace of FCS before adding 80 μL mEpiSC media with 10 μM ROCKi. Label the wells with the clone number (i.e., 1, 2, 3, etc.) and return to the incubator. 3. In another 96-well plate, add 30 μL TrypLE™ Select to the same series of wells seeded with MEFs on the plate prepared in step 2. Label the wells with the clone number (i.e., 1, 2, 3, etc.). 4. Take the electroporated cells out of the incubator, and examine the clones on the plate. Mark on the plate which clones are suitable for selection based on morphology (see Note 9). 5. Aspirate the medium from the plate, and rinse the mESCs or mEpiSCs with 4 mL or 2 mL DPBS, respectively. 6. Add 7 mL DPBS to the mESCs or 2 mL to the mEpiSCs (see Note 10), and under a microscope, start picking clones, with each being placed in a single well of the 96-well plate containing TrypLE™ Select (see Note 11). This can be done by setting a P20 pipette to 4 μL (see Note 12). 7. After 5 min, inactivate TrypLE™ Select by adding 70 μL mESC medium or mEpiSC medium supplemented with 10 μM ROCKi to each well. 8. Using a multichannel pipette, pipette each well up and down to dissociate into single cells. 9. Transfer the cells to the 96-well plate containing the MEFs, and place back into the incubator. 10. Feed the clones daily with 200 μL mESC medium or mEpiSC medium.

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3.5 Clone Picking of pX458-Transfected mESCs

1. Repeat steps 1–2 from Subheading 3.4. 2. Check growth of cells 3 days after electroporation for size and the presence of fluorescence to decide the best time for clone picking. A balance needs to be made between strong GFP expression and a sufficient number of cells per clone (see Note 13). 3. Under a fluorescence microscope, pick clones uniformly expressing GFP using a mouth pipette, and place directly into the 96-well plate containing pre-seeded MEFs (see Note 14). 4. Feed the clones daily with 200 μL mESC media until most clones are large enough to be passaged. As the clones that were picked were small, they will not grow to fill the entire well. 5. 24 h before passaging the clones, repeat step 1 from Subheading 3.4. On the day of passaging, repeat step 2 from Subheading 3.4. 6. Passage the entire clones onto the 96-well plate that was pre-seeded with MEFs 24 h earlier (see Note 15).

3.6 Passaging and Cryopreservation of mESC Clones

Once clones are almost confluent, it is necessary to cryopreserve a proportion of the cells as a stock as well as grow the remaining cells on gelatin to remove MEFs for genomic DNA (gDNA) extraction and genotyping of the mutation. Not all clones grow at the same rate, and some clones may not grow at all, so it is necessary to exercise a compromise between the expansion of cells by enhancing growth and preventing cell differentiation. 1. Remove mESC media from each well. 2. Rinse each well with 100 μL DPBS and add 30 μL TrypLE™ Select. Place in the incubator for 5 min. 3. During the incubation, coat wells of a fresh 96-well plate with 100 μL 0.1% gelatin, aspirate off gelatin after a few minutes, and immediately add 160 μL mESC media. The number of wells to coat is equivalent to the number of clones that has been picked. 4. Inactivate the TrypLE™ Select by adding 90 μL mESC media to each well. 5. Using a multichannel pipette, pipette each well up and down to dissociate into single cells. 6. Transfer 40 μL of the cells onto the plate containing gelatin. Label the plate accordingly and place into the incubator. 7. To the remaining 80 μL cells, slowly add 80 μL 2 freeze media, and gently pipette up and down. 8. Transfer each clone to a separate tube of the MicroTube Rack System™. Attach the lids and label the tubes with the necessary details.

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9. Place the MicroTube Rack System™ Tubes on ice, and transfer to a 80  C freezer. 10. Feed the cells on gelatin daily with 200 μL mESC media and passage when cells are confluent. Generally, three passages on gelatin are sufficient to remove all MEFs (see Note 16). 3.7 Passaging and Cryopreservation of mEpiSC Clones

1. Remove mEpiSC media from each well. 2. Rinse each well with 100 μL DPBS, add 30 μL Collagenase IV, and place back into the 37  C incubator for 10 min. 3. During the incubation, coat the wells of a new 96-well plate with 100 μL 0.1% gelatin for 20 min, then aspirate off gelatin, and immediately add 140 μL conditioned mEpiSC medium supplemented with 10 μM ROCKi. 4. Add 30 μL mEpiSC medium to the dissociated cells, and detach the colonies from the feeder layer. Transfer into 1.5 mL tubes. 5. Spin the clump suspension at 1000 rpm (200  g) for 30 s. 6. Resuspend the cell pellet in 120 μL conditioned mEpiSC medium supplemented with 10 μM ROCKi. Try to dissociate the clumps as much as possible. 7. Transfer 60 μL of the cell suspension to the plate containing gelatin. Label the plate accordingly, and place into the incubator. 8. To the remaining 60 μL cells, add 60 μL 2 freeze media, and gently pipette up and down. 9. Repeat steps 8 and 9 of Subheading 3.6. 10. Feed the cells daily with 200 μL conditioned mEpiSC medium, and, when cells are confluent, passage at a 1:2 ratio onto a new gelatin-coated 96-well plate. Add 10 μM ROCKi after each passage for 24 h. Three passages through gelatin-coated culture are sufficient to remove all MEFs; however, the mEpiSCs may take up to 2 weeks to recover.

3.8 Cell Lysis, PCR Amplification, and Sanger Sequencing for Genotyping

1. Once all MEFs are removed and cells are confluent, aspirate off media, and rinse with 100 μL DPBS. 2. Add 100 μL lysis buffer with Proteinase K to each of the clones, and incubate overnight at 56  C (see Note 17). 3. Transfer the lysates to eight-strip tubes, and inactivate Proteinase K by incubating at 95  C for 10 min in a thermocycler (see Note 18). 4. PCR amplify the region surrounding the mutation for each of the clones using the conditions in Tables 1 and 2. It will be necessary to perform the same PCR reaction on a sample that has not been edited in the same region (see Note 19).

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Table 1 PCR master mix for the amplification of gDNA surrounding the mutation site using BioMix™ Reagent

Volume for one reaction (μL)

BioMix™

10

Forward primer (10 μM)

1

Reverse primer (10 μM)

1

Template gDNA

2

Sterile deionized water

6

Table 2 PCR cycling conditions for the amplification of gDNA surrounding the mutation site using BioMix™ Stage

Temperature ( C)

Time

Initial denaturation

95

5 min

35 cycles

95 60 72

30 s 30 s 1 min/kb

Hold

Denaturation Annealing Extension

4

Forever

5. Make a 2% agarose gel in 1 TAE that is enough to fill a large casting tray (~200 mL). 6. Once cool, add 15 μL of RedSafe™ and gently swirl into the solution. Carefully pour the agarose solution into a gel cast tray with combs already inserted without introducing air bubbles to the gel. 7. Load 5 μL of the PCR products into each well. Add 6 μL of the appropriate ladder into a lane next to your samples (see Note 20). Electrophorese gel at 100 V for 2.5 h to allow for sufficient separation of the bands indicating potential biallelic mutations. 8. After imaging, select samples in which the fragment size is relatively close to the expected size, and purify samples from the remaining 15 μL PCR product using Wizard® SV Gel and PCR Clean-Up System according to manufacturer’s instructions (see Note 21). 9. Perform Sanger sequencing of the PCR products, including the unedited control, with the forward primer used for the PCR amplification. 10. Once the sequencing results have returned, use TIDE (https://tide-calculator.nki.nl/) [6] to predict the insertions/

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deletions (indels) of the samples compared to the unedited control sample (see Note 22). 11. Determine which clones are worth pursuing for further characterization based on the predicted indels, % of sequences with that indel, and total efficiency of the prediction. The desired indels should result in a biallelic frameshift mutation (number of base deletions not divisible by 3), with ~50% sequences with each indel at a high efficiency. 3.9 Decomposition of Mutations: Ligation, Transformation, and Extraction of Plasmid DNA

1. Ligate 2 μL of purified PCR product of the selected clones from Subheading 3.8 into the pGEM®-T Easy Vector according to manufacturer’s instructions (see Note 23). 2. Thaw a vial of the α-Select Silver Competent Cells on ice for ~5 min. Label a 1.5 mL tube for every ligation sample, and aliquot out 2 μL of the sample into the tubes, and pre-chill on ice, while competent cells are thawing. 3. Once completely thawed, gently mix the competent cells by pipetting up and down once before adding 50 μL of cells to each pre-chilled ligation mix. Pipette up and down gently once to mix. 4. Incubate mixture on ice for 30 min. 5. Heat shock the cells at 42  C for 30 s in a water bath. 6. Immediately place on ice for 2 min. 7. Add 350 μL of LB to the transformation mix. 8. Incubate transformation mix at 37  C for 1 h, shaking at 200 rpm. 9. Plate 200–300 μL of the transformation mix carefully onto a pre-warmed LB ampicillin agar plate with IPTG and X-gal (see Note 24). Incubate overnight at 37  C. 10. Seal transformation plates and place at 4  C the following morning (see Note 25). 11. Aliquot 4 mL of freshly made LB ampicillin into 8–12 labelled 14 mL Falcon™ Round-Bottom Polypropylene Tubes. 12. Pick 8–12 pure white colonies from the transformation plate using a yellow 200 μL pipette tip, suspend the colony into the LB ampicillin, and discard the tip within the tube (see Note 26). 13. Culture the picked subclones at 37  C overnight, shaking at 200 rpm. 14. Isolate the plasmid using the ISOLATE II Plasmid Mini Kit as per manufacturer’s instructions. Elute in 30 μL of H2O. 15. Send isolated plasmid samples off for Sanger sequencing using the M13 reverse primer.

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16. Use https://www.ebi.ac.uk/Tools/msa/muscle/ [7, 8] to compare sequenced results to the reference sequence (see Note 27). 3.10 Thawing and Validation of Confirmed mESC and mEpiSC Clones

1. Seed one well of a 48-well plate 1 day prior to thawing confirmed clones with MEFs. 2. On the day of thawing, remove MEF media, and replace with 400 μL mESC medium or mEpiSC medium supplemented with 10 μM ROCKi. 3. Cut the selected clones from the strips of the MicroTube Rack System™, and immediately put tube(s) on ice (see Note 28). 4. Hold the tube in the 37  C water bath until almost thawed. 5. Sterilize the outside of the tube with ethanol, and slowly add 200 μL of pre-warmed mESC or mEpiSC media into the tube. 6. Move the thawed cells to a 15 mL tube. Do not discard the original tube. 7. Add 500 μL of pre-warmed mESC or mEpiSC media into the original tube to clean up the leftover cells, and move all the contents into the 15 mL tube. 8. Add 1.5 mL of pre-warmed mESC or mEpiSC media slowly into the tube with the cells (see Note 29). 9. Centrifuge the cells at 1000 rpm (200  g) for 5 min. 10. Aspirate off the supernatant leaving about 200 μL of media. 11. Carefully resuspend the cells in another 200 μL of pre-warmed mESC or mEpiSC media supplemented with 10 μM ROCKi. 12. Transfer the cells into the single well of the 48-well plate pre-seeded with MEFs. 13. Add another 200 μL of pre-warmed mESC medium or mEpiSC medium supplemented with 10 μM ROCKi into the 15 mL tube to collect the remaining cells, and transfer them into the well of the 48-well plate (see Note 30). 14. Expand the clones, and freeze down vials of cells whenever possible until enough cells can be seeded onto a full 6-well plate (see Note 31). 15. After getting to the 6-well plate stage, also passage cells onto 0.1% gelatin in a 6-well plate for at least three passages. This plate will be used to extract gDNA from for validation of mutations. 16. Cryopreserve the 6-well plate of cells on MEFs once it becomes confluent. Each well is enough to cryopreserve into two vials that can be thawed onto a single well of a 6-well plate. 17. Once the cells on gelatin have been passaged three times and are confluent, rinse each well with 1 mL DPBS.

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18. Add 1 mL of DPBS into each well and scrape cells thoroughly (see Note 32). Resuspend cells and transfer each well to a separate 1.5 mL tube. 19. Rinse each well with another 0.5 mL of DPBS, and collect into the same 1.5 mL tube. 20. Centrifuge samples at 1000 rpm (200  g) for 5 min. 21. Aspirate excess DPBS, removing as much as possible without disrupting the cell pellet (see Note 33). 22. Snap freeze samples in liquid nitrogen, and either proceed to the next step or store at 80  C until required. 23. Extract gDNA from one cell pellet by adding 100 μL of lysis buffer with fresh Proteinase K and incubating overnight at 56  C. 24. Repeat steps 3–8 from Subheading 3.8, and then repeat Subheading 3.9 (see Note 34). 25. Confirm gene knockout by RT-qPCR and Western blotting (see Note 35).

4

Notes 1. It is best if ¼ 18.2 MΩ cm H2O Check preparation, remake buffers. Calibrate pH meter and check pH Reduce digestion time/enzyme concentration Use new FBS. Try new batch if still unsuccessful Use only ultrapure 18.2 MΩ cm H2O

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Fig. 2 Schematic illustrations of in situ heart flushing, removal, and ex vivo injection. (a) Chest cavity of anesthetized mouse is opened to below the diaphragm, which is then cut through to expose the heart. The descending aorta and inferior vena cava are cut (1), and the heart is immediately flushed with EDTA buffer by injection into the right ventricle (2). Reynolds hemostatic forceps then reach around the heart to clamp the emerging aorta (3), and hold the heart while it is removed by cutting around the forceps (4). (b) The excised heart, still held by the clamp, is transferred to 60 mm dishes for subsequent injection and digestion steps

the base of the RV, penetrating no more than a few mm, and the angle of entry may be carefully varied during injection. 3.5

Removal of Heart

1. The emerging aorta is then clamped. Any hemostatic clamp will suffice, but full-curved-ended Reynolds forceps are preferred. These can easily reach around the heart and clamp the emerging aorta in situ, which does not require high precision, and inclusion of additional emerging vessels does not matter, although clamping of atrial appendages should be avoided. 2. The heart is removed by simply cutting around the outside of the forceps and transferred, still held by the clamped forceps, to the 60 mm dish containing EDTA buffer, where it should be almost completely submerged (Fig. 2b).

3.6

Heart Digestion

1. Locate the left ventricle (LV), which is the larger of the ventricles and forms a pointed apex at the base of the heart. Using the second EDTA syringe, insert the needle into the base of the LV wall, 2 or 3 mm above the apex, penetrating no more than a few mm into the LV chamber, and inject the EDTA buffer

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starting at a flow rate of around 1 mL per 2 or 3 min (see Note 8). 2. After 6 min or application of all 10 mL EDTA buffer, whichever is first, the needle is removed, and the heart is transferred, still held by the clamped forceps, to the dish of perfusion buffer. 3 mL perfusion buffer is then similarly injected into the LV, if possible via the same perforation left by the previous injection. Inexperienced users may find a magnification lens beneficial for identification of the original injection point. 3. After 2 min or application of all 3 mL perfusion buffer, whichever is first, the heart is transferred to the dish containing 10 mL collagenase buffer, and the LV is injected sequentially with the five syringes of collagenase buffer (see Notes 6 and 9). 4. The clamp is removed, and scissors may be used to separate the heart into its constituent chambers, or other specific regions, as desired. The selected region is then transferred to the final 3 mL dish of collagenase buffer (multiple dishes can be used here in order to isolate cells from multiple regions). 5. Tissue is gently teased apart into roughly 1 mm  1 mm sized pieces using the round and sharp-end forceps, which requires very little force following a successful digestion. 6. Dissociation is completed by gentle trituration for 2 min using a 1 mL pipette, with a wide-bore tip (purchased or homemade using sterile scissors) to reduce shear stress. 7. To stop the enzymatic digestion, 5 mL of stop solution is added to the cell-tissue suspension, which may be gently pipetted for a further 2 min, and inspected under a microscope (see Note 10). 8. Cell suspension is then transferred to a 50 mL centrifuge tube, which should be stored on its side at room temperature to reduce clumping and hypoxic damage. Cells may be stored with little loss of viability for up to 2 h, in which time further isolations may be performed. However, such delays may not be suitable for sensitive applications. 3.7 Collection of Cardiomyocytes by Gravity Settling

If cells are to be cultured, subsequent steps are best undertaken in a laminar flow cabinet, to maintain sterility. 1. Cell suspension is passed through a 100 μm pore-size strainer, in order to remove undigested tissue debris. The filter is washed through with a further 5 mL stop buffer. 2. Total volume of cell suspension is now typically around 15 mL. This can be divided into two 15 mL centrifuge tubes, and cells are then allowed to settle by gravity for 20 min. Most myocytes will settle to a pellet, while most non-myocytes and cellular/ extracellular debris remain in suspension (see Note 11).

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3. Supernatant is removed. If cells are to be harvested immediately without further in vitro experiments, myocyte fractions are purified simply by three further rounds of sequential gravity settling for 10 min in 4 mL fresh perfusion buffer, retaining the myocyte-containing pellet each time. 3.8 Calcium Reintroduction and Culture of Cells

Where myocytes are to be returned to physiological extracellular calcium levels and/or plated, it is important to do so in gradual increments, in order to avoid spontaneous contraction and achieve healthy populations of calcium-tolerant cells, which may then be subjected to a wide range of experimental applications [4]. This can be easily incorporated into the gravity settling steps. 1. Similar to Subheading 3.7, step 3, myocyte pellets are instead resuspended sequentially in three calcium reintroduction buffers, containing increasing proportions of either Tyrode’s solution (for immediate calcium handling or electrophysiology experiments) or culture media (for plating and/or culturing of cells); see Subheading 2.2, item 3. 2. If required, the supernatant fractions, which contain non-myocyte cell populations as well as rounded myocytes and some viable myocytes, may be collected and combined from each round of gravity settling. Plating and fibroblast media can be warmed and equilibrated in a 37
C, 5% CO2, humidified tissue culture incubator during this process. 3. For plating of cardiomyocytes, laminin solution is aspirated from the prepared culture surfaces (see Subheading 3.1), which are then washed once with PBS. 4. The final cardiomyocyte pellet is resuspended in pre-equilibrated plating medium, and cells are plated at application-specific densities: Typically around 25,000 cells/ mL, or 5000 cells/cm2, but this may be substantially lowered for imaging studies. 5. Cardiomyocytes are transferred to the tissue culture incubator and shaken gently in a side-to-side (not swirling) motion to ensure even distribution. Adhesion of rod-shaped myocytes occurs rapidly, within 20 min for most cells. Culture medium may be pre-equilibrated in the incubator during this time. 6. Cells in the combined supernatant fraction may be collected by centrifugation at 300  g for 5 min, resuspended in pre-equilibrated fibroblast growth media, plated on tissue culture surfaces (area ~20 cm2 per LV), and transferred to the culture incubator (see Note 12). 7. After 1 h, plated cardiomyocytes are gently washed once with pre-equilibrated culture media and then incubated in culture media, for the required experimental duration. Rounded

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Fig. 3 Representative example of adult mouse cardiomyocytes after poor isolation procedure (a), containing many rounded, hypercontracted, and dying cells, and good quality procedure (b), showing a majority of healthy, rod-shaped cells. Healthy cardiomyocytes were plated and visualized at 40 (c) and 400 (d) magnification, whereby characteristic angular morphology and sarcomeric striations are clearly visible. Scale bars are 100 μm

myocytes do not adhere strongly and are removed by this process (see Note 13). 8. Culture and fibroblast media are changed after 24 h and every 48 h in culture thereafter. A successful isolation procedure yields up to one million cardiomyocytes per left ventricle, with 80% viable, healthy, rod-shaped cells [4]. Poor isolations yield high numbers of dying, round, hypercontracted cells, and troubleshooting is required; see Table 1 and Fig. 3a–d.

4

Notes 1. 100 collagenase and 1000 protease XIV (¼50 mg/mL) stocks may alternatively be prepared in ultrapure 18.2 MΩ cm H2O, filter-sterilized, and stored in aliquots at 80
C for at least 4 months. These can be added to perfusion buffer to produce collagenase buffer immediately before the

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isolation. We use collagenases 2 (LS004176) and 4 (LS004188) from Worthington Biochemical, Lakewood, USA, which exhibit high batch-to-batch reproducibility (collagenase 2, ~210 units/mg; collagenase 4, ~260 units/mg). Collagenase 2 is a less pure extract with more basal clostripain activity than collagenase 4, which can sometimes be advantageous, and in many cases, 2.5 mg/mL collagenase 2 alone is sufficient at 37
C to attain good yields of myocytes. However, using the described mixture as standard performs consistently. 2. 100 BDM (¼1 M) stocks can be prepared by dissolving 1.01 g BDM in 10 mL ultrapure 18.2 MΩ cm H2O, filtersterilized, and stored in aliquots at 20
C. Stock may require incubation at 37
C to redissolve BDM before adding to media as required. BDM is a myosin II ATPase inhibitor, used to reduce myocyte contractions and improve the yield of isolated cardiac myocytes. BDM must be removed from cultures before conducting contractility, calcium handling, or electrophysiology experiments. It is normal to see a number of cardiomyocytes becoming hypercontracted and dying 1–2 h after BDM removal. In culture media, blebbistatin can be used in place of BDM, which may have fewer off-target effects, and improve long-term survival and adenoviral transduction efficiency [6]. We find that 5 μM blebbistatin is optimal for this purpose. 3. 50 BSA (¼5% w/v) stocks can be prepared by dissolving 1 g BSA in 20 mL PBS, filter-sterilized, and stored at 4
C for at least 8 weeks, if kept sterile. 4. CD lipid mix is included to improve myocyte survival in longterm culture [4]. It is not typically required for short term culture. The lipid mix is susceptible to oxidation and to light damage and is therefore best stored in small aliquots containing minimal air space, at 4
C, in the dark. 5. Laminin-coated surfaces are best prepared fresh but may be sealed and stored at 4
C for up to 4 days. When using glass surfaces, extra volume may be required for complete coverage. Note that cells adhere less strongly to glass than plastic. 6. Following application of each syringe of collagenase buffer to the heart, 10 mL will need to be removed from the dish, to prevent overflow. To reduce consumption of enzyme, this buffer may be collected and recycled for subsequent injection. Care must be taken to prevent needle-prick injuries. However, to prevent cellular cross-contamination, fresh collagenase buffer is generally prepared for each heart. 7. Feedback from users suggests that the choice of euthanasia technique is one of the most common causes of problems encountered. Induction of rapid-onset anesthesia using oxygen-isoflurane ventilation is strongly recommended. This

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involves no injections and causes the mouse minimal stress. Furthermore, circulation is intact, and blood is well oxygenated up to the point of chest opening and introduction of EDTA buffer. Injected anesthetics such as pentobarbital and ketamine have a longer onset and significantly reduce respiration, increasing the risk of ischemia and subsequent cardiomyocyte calcium overload [1, 7]. Cervical dislocation carries the same risk, in addition to likely blood coagulation and thus blockage of coronary circulation in the time taken to open the chest and inject EDTA buffer, particularly for inexperienced users, leading to poor myocardial perfusion of dissociation buffers and low yields of healthy cardiomyocytes. If these techniques are necessary, heparin administration is recommended 30 min prior to euthanasia. It should be emphasized that euthanasia by CO2 inhalation causes ischemia and is not appropriate for myocyte isolation techniques. 8. The ideal flow rate when injecting buffers into the LV will vary between hearts, but the best measure of adequate perfusion is simply the minimum required to maintain full inflation of the heart. Initially, very little force is required for the heart to inflate, and flow rate may be only 1 mL per 2 or 3 min. As digestion progresses and the heart softens, flow rate typically reaches around 2 mL/min. A temptation is to over-apply, which can cause buffer to force into and perforate the left atrial appendage. This alone does not cause poor isolation results, and the researcher may proceed as normal, although such pressure is unnecessary. If desired, this protocol is compatible with automated infusion pump setups [4]. 9. The volume of collagenase buffer required for complete digestion varies between hearts. Small, young, healthy hearts can digest in as little as 25 mL, while larger, older, or fibrotic hearts may pass beyond 50 mL, necessitating the recycling of buffer (see Note 6). Signs of complete digestion include a noticeable reduction in resistance to injection pressure, loss of shape and rigidity, holes and/or extensive pale and fluffy appearance at the heart surface, and ejection of myocytes into the effluent buffer, which are just visible to the naked eye. The point of injection will often widen until significant buffer appears to be flowing directly backwards, but the researcher may proceed as necessary. 10. Myocytes may display contraction immediately after isolation due to mechanical stimulation but should quickly acquiesce. The presence of large numbers of rounded, hypercontracted myocytes (Fig. 3a) indicates a poor isolation and requires troubleshooting (see Table 1).

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11. Sequential gravity settling is a method to obtain a highly pure myocyte population and avoids damage caused by centrifugation. Viable rod-shaped myocytes tend also to settle faster than round hypercontracted and dying myocytes, so enriching the pellet for viable rod-shaped cells. Division of cell suspension into two 15 mL centrifuge tubes rather than one 50 mL tube aids the formation of a pellet due to the more steeply angled base. Sterile polystyrene round-bottom tubes are also good alternatives for this purpose. 12. Cardiac fibroblasts, and some other non-myocytes [4], adhere to untreated tissue culture plastic surfaces within 1–2 h. Remaining cardiomyocytes do not and can be washed off at this stage to effectively purify the (mostly) cardiac fibroblast population. 13. Cultured myocytes must be handled with great care. Avoid shocks, vibrations, and rapid aspiration/introduction of media. Always wash gently using warm culture media to reduce ionic fluctuations, and change media one well at a time to avoid prolonged exposure to air, particularly if culturing on glass surfaces. When fixing cells with formaldehyde for imaging, best results are obtained by adding 8% formaldehyde dissolved in culture medium slowly to an equal volume of culture medium already in the well and incubating for 15 min. Do not swirl or shake. References 1. Bers DM (2002) Cardiac Na/Ca exchange function in rabbit, mouse and man: what’s the difference? J Mol Cell Cardiol 34:369–373 2. Berry MN, Friend DS, Scheuer J (1970) Morphology and metabolism of intact muscle cells isolated from adult rat heart. Circ Res 26:679–687 3. Powell T, Twist VW (1976) A rapid technique for the isolation and purification of adult cardiac muscle cells having respiratory control and a tolerance to calcium. Biochem Biophys Res Commun 72:327–333 4. Ackers-Johnson M, Li PY, Holmes AP, O’Brien S-M, Pavlovic D, Foo RS (2016) A simplified, Langendorff-free method for concomitant

isolation of viable cardiac myocytes and nonmyocytes from the adult mouse heart. Circ Res 119:909–920 5. Chen X, O’Connell TD, Xiang YK (2016) With or without Langendorff: a new method for adult myocyte isolation to be tested with time. Circ Res 119:888–890 6. Kabaeva Z, Zhao M, Michele DE (2008) Blebbistatin extends culture life of adult mouse cardiac myocytes and allows efficient and stable transgene expression. Am J Physiol Heart Circ Physiol 294:H1667–H1674 7. O’Connell TD, Rodrigo MC, Simpson PC (2007) Isolation and culture of adult mouse cardiac myocytes. Methods Mol Biol 357:271–296

Chapter 15 Isolation, Culture, and Characterization of Primary Mouse Epidermal Keratinocytes Ling-Juan Zhang Abstract Epidermis, the outermost layer of the skin, plays a critical role as both a physical and immunological barrier protecting the internal tissues from external environmental insults, such as pathogenic bacteria, fungi, viruses, UV irradiation, and water loss. Epidermal keratinocytes (KC), the predominant cell type in the skin epidermis, are in the front line of skin defense. Here we describe methods to isolate and culture primary epidermal KC from neonatal and adult mouse skin and describe in vitro assays to study and characterize KC proliferation and differentiation and pro-inflammatory responses to viral products and UVB irradiation. These methods will be useful for researchers in the field of epidermal biology to set up in vitro assays to study the barrier and pro-inflammatory function of epidermal keratinocytes. Key words Skin epidermis, Keratinocyte, Skin barrier, Keratinocyte proliferation, Keratinocyte differentiation, Pro-inflammatory response, dsRNA, UVB irradiation, TNF release

1

Introduction Skin is the largest organ of the body, and epidermis is the outermost layer of the skin. At the front line of defense, epidermis plays a critical role in forming an intact barrier to protect the body from dehydration and external insults, such as pathogenic bacteria, virus, allergens, and UVB irradiation [1, 2]. This barrier function is mainly provided by keratinocytes (KC), the predominant cell type in the epidermis, and it is maintained by a tightly controlled balance between proliferation and differentiation of KC [3, 4]. KC located on the basal layer of the epidermis, including both epidermal stem cells and transit-amplifying cells, are proliferative. As these basal cells exit cell cycle, KC commit to terminal differentiation and gradually move upward toward the surface of the skin [5, 6]. During the differentiation process, KC undergo a series of biochemical and morphological changes that result in the formation of distinct layers of the epidermis. At the spinous layer, directly above the basal layer, KC express early differentiation markers, such as keratin 10 (K10).

Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940, https://doi.org/10.1007/978-1-4939-9086-3_15, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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As KC migrate to the granular layer, these cells become interconnected by tight junctions and express late differentiation markers, such as filaggrin (FLG), loricrin (LOR), and involucrin (INV). Eventually, when these cells reach the outer surface of the epidermis, KC become terminally differentiated corneocytes, which are enucleated and flattened and eventually sloughed into the environment as new cells replace them [1, 5, 6]. At the front line of host defense, epidermal KC are also an important component of the skin’s innate immune system. In response to PAMPs (pathogen-associated molecular patterns) released by invading pathogens or DAMPs (damage-associated molecular patterns) released by host cells during UVB irradiation or wounding, KC produce a variety of pro-inflammatory cytokines or chemokines, such as TNFα, IL6, IL8, CXCL10, and IFNβ [2, 7, 8]. These pro-inflammatory signals released from KC recruit or activate myeloid and resident immune cells, mounting a rapid host defense immune response leading to efficient pathogen clearance. However, uncontrolled inflammatory response may trigger the development of auto-inflammatory skin diseases, such as psoriasis and rosacea [9, 10]. Here we describe methods to isolate epidermal KC from neonatal or adult mouse skin. This is an extended protocol modified from our previous published protocol in Journal of Visualized Experiments [10]. While neonatal KC are collected from both whole body skin from neonates, adult KC are isolated from tail skin, which has thicker epidermis and lower hair follicle density compared to body skin in adult mice. Skin is first digested overnight with dispase, an enzyme to dissociate the epidermis from dermis. The separated epidermal sheet is then digested with a trypsin-like enzyme to release epidermal KC. Isolated KC are seeded on culture dishes coated with extracellular matrix and cultured in low calcium medium supplemented with defined growth supplements. Between days 2 and 5 after the initial seeding, dead cells or differentiated cells are washed away by daily medium changes; the remaining cells are proliferating and have cobblestone morphology, a characteristic morphology of basal KC. We also describe methods to characterize and study proliferation, growth factor starvation, and/or high calcium-induced differentiation as well as pro-inflammatory response of these primary KC triggered upon exposure to viral product or UVB irradiation.

2 2.1

Materials Animals

C57B/6 wild-type mice are bred and maintained in a specific pathogen-free (SPF) environment according to animal facility regulations. Neonates are used within 2 days of birth (postnatal days 0~2), and adult mice are used between 6 and 15 weeks of age and either female or male mice can be used.

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1. Sterile PBS, pH 7.4. 2. 10 cm Petri dish. 3. KC basal medium with 0.06 mM CaCl2. 4. Defined growth supplements include epidermal growth factor (EGF), bovine transferrin, insulin-like growth factor1 (IGF1), prostaglandin E2 (PGE2), bovine serum albumin (BSA), and hydrocortisone (Life Technologies, Carlsbad, CA, Catalog S0125). 5. Dispase. 6. Complete KC growth medium: Basal KC medium (0.06 mM CaCl2) supplemented with defined KC growth supplement and 1
antibiotic-antimycotic. 7. Dispase digestion buffer: 4 mg/mL dispase in complete KC growth medium. 8. Gelatin coating material. 9. Type 1 collagen coating material (either from bovine or rat tail or recombinant human protein). 10. Trypsin-like enzyme/TrypLE (Life Technologies, Carlsbad, CA, Catalog 12604-013). 11. 100 μm cell strainer.

2.3 Functional Cell Assay

1. Culture dishes: 24-well clear flat bottom TC-treated cell culture plates are used for phase contrast imaging and/or RTqPCR analyses. 96-well clear flat bottom TC-treated microplate is used for colorimetric cell counting assay. 8-well chamber slide is used for immunocytochemistry analysis. 2. Colorimetric cell counting kit: Colorimetric assays to measure metabolic activity of living cells, such as Cell Counting Kit-8 (CCK-8) or MTS assay or MTT assay, can be used for mouse KC. 3. 5-bromo-20 -deoxyuridine (BrdU) is dissolved as 20 mM stock in DMSO in aliquots. 4. Rat anti-BrdU antibody. 5. High molecular weight (HMW) poly(I:C). 6. Corded handheld UV lamps. 7. 8-watt UV tubes. 8. Mouse TNF ELISA kit. 9. Light inverted microscope for cell culture. 10. Fluorescent microscope for cover slides.

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Methods

3.1 Overnight Dispase Digestion of Neonatal or Adult Skin

1. Euthanize the postnatal day 0–day 2-old C57BL/6 wild-type neonatal pups by decapitation using scissors, and follow steps 2 and 3 to set up dispase digestion. For adult mice, euthanize adult mice according to animal facility regulations. Cut off the tail from the base, and follow steps 4 and5 to set up dispase digestion. 2. To peel neonatal skin off the body, first cut off limbs just above the wrist and joints, and cut off the tail from the base leaving a small hole. Insert sharp scissors through the hole from the tail and cut the skin along the dorsal midline through the neck. Next, use one forceps to gently lift a corner of skin off the neck and the other forceps to grasp the exposed neck and body, and then carefully peel the whole skin off the body and over the leg stumps in one continuous motion (see Note 1). 3. Rinse the peeled neonatal skin in a 10 cm Petri dish with 15 mL of sterile PBS, and then transfer the skin to a 2 mL tube prefilled with 2 mL ice cold dispase digestion buffer (see Note 2). Digest the skin overnight at 4  C on a rotator in a refrigerator. Next day, proceed to Subheading 3.2. 4. To peel adult tail skin off the bone, first use a sharp blade to cut through the tail base from the tail tip. Next, use one forceps to gently lift a corner of the skin off the tail bone at the base and the other forceps to grasp the exposed tail bone, and then carefully peel tail skin off the bone with one continuous motion. Cut each of the peeled skin into 2~3 pieces, each of which is ~2 cm in length. 5. Rinse the peeled adult tail skin in a 10 cm Petri dish with 15 mL of sterile PBS, and then transfer skin pieces from each tail to a 2 mL tube prefilled with 2 mL ice cold dispase digestion buffer (see Note 3). Digest the skin overnight at 4 refrigerator on a rotator. Next day, proceed to Subheading 3.2.

3.2 Isolation of Keratinocytes from Skin Epidermis

1. On the second day (within 12~18 h post dispase digestion), carefully transfer the skins together with the dispase solution to a Petri dish and then to a new Petri dish with 15 mL sterile PBS to wash away excess dispase. Using two pairs of forceps, carefully transfer each skin piece to a dry Petri dish with epidermis side down and dermis side up. Stretch the skin folds so that the skin is fully extended on the Petri dish. 2. Before separating the epidermal sheet from the dermis, place a drop of 500 μL TrypLE, a trypsin-like digestion solution (room temperature) in a new Petri dish (see Note 4). To remove the dermis, use one forceps to hold down a corner of the epidermis, and use the other forceps to gently lift the

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dermis (pink, opaque, gooey) away from the epidermal sheet (whitish, semitransparent; see Note 5). Dispose of the dermis as biohazardous material. 3. Use two pairs of forceps to grasp the cross corner of the separated epidermal sheet, and slowly transfer it onto the surface of the TrypLE digestion solution with the basal layer downward (see Note 6). 4. Cover skin on Petri dish with lid, and incubate for 20 min at room temperature on a horizontal shaker with gentle agitation. Basal KC become loosely attached to the epidermal sheet or are released from the epidermal sheet during this digestion process. 5. To stop digestion, add 2 mL ice cold complete KC growth medium per epidermis to the Petri dish. Using forceps vigorously rub the epidermal sheet, and the medium will become turbid as KC are released into solution from the epidermal sheet. Tilt the Petri dish to collect and transfer the cell suspension to a 50 mL centrifuge tube leaving the remaining epidermal sheet on the dish. Keep the collection tube on ice during the procedure. 6. Repeat step 5 two more times, and combine the cell suspensions into the same 50 mL tube. 7. Pipet the cell suspension up and down gently a few times to disperse cell clumps using a serological pipette, and then pass it through a 100 μm filter to a new 50 mL centrifuge tube. 8. Centrifuge the filtered cells at 180
g for 5 min. Aspirate the supernatant, and resuspend the cell pellet in 1 mL cold KC growth medium, and determine the cell number using a hematocytometer (see Note 7). 3.3 Primary Mouse KC Culture

1. Prior to cell seeding (see Note 8), culture dishes should be coated with appropriate ECM materials (see Note 9) according to the manufacturer’s instruction at 37  C for 30 min. Remove the coating material completely immediately before adding the cell suspension. 2. Seed the isolated neonatal KC at a density of 5
104/cm2, or adult KC at a density of 10
104/cm2, in KC growth medium in culture dishes coated with ECM material as described above. 3. Change the medium 24 h after the initial plating, and then change medium daily to remove unattached cells or cells that spontaneously differentiate and detach from culture dish. Between day 2 and day 5 of initial plating, cells should reach >70% confluency. Cells can then be used for experimentation. Representative phase contrast images for adult mouse KC from 8 h to 4 days after the initial plating are shown in Fig. 1a.

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Fig. 1 In vitro assays to measure KC growth and proliferation. (a) Phase contrast images at 10
magnification of primary adult mouse KC at 8 h, day 1, day 2, day 3, and day 4 post the initial seeding. Scale bar ¼ 100 μm. (b) The relative cell number of primary adult mouse KC at indicated day after the initial seeding was measured by the CCK-8 cell viability assay. All error bars indicate mean  s.e.m. *P < 0.05, **P < 0.01, ***P < 0.001 (ANOVA). (c) Subconfluent neonatal KC were pulse-labeled with BrdU prior to immunocytochemistry analyses using anti-BrdU antibody (red), and nuclei were counterstained with DAPI in blue. Scale bar ¼ 100 μm

3.4 In Vitro Assays to Study KC Proliferation (See Note 10)

1. Measurement of cell proliferation by colorimetric cell counting assay: Primary KC are seeded in 96-well flat bottom clear plate, and KC growth medium is changed daily during the assay (100 μL medium /well). To measure relative cell number, 10 μL of CCK-8 solution is added to each well and incubated for 1 h in a cell culture incubator (5% CO2 at 37  C), and then O.D. at 450 nm is measured by a spectrometer. Measurement of relative cell number over a time course of 4 days by CCK-8 assay is shown in Fig. 1b. 2. Labeling of S-phase cells by BrdU incorporation: Primary KC are grown on coverslips and incubated for 30 min with 10 μM BrdU, followed by fixation in 4% PFA/PBS. Fixed cells are treated with 0.2 M HCl for 30 min at room temperature followed by neutralization with a borate buffer. Cells are then permeabilized with 0.1% Triton X-100 and subjected to standard immunocytochemistry procedures using a rat anti-BrdU antibody and a Cy3-conjugated anti-rat secondary antibody. Nuclei are counterstained with DAPI. Representative image for BrdU labeling of primary neonatal KC in the S-phase of cell cycle is shown in Fig. 1c using fluorescence microscope.

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Fig. 2 Growth factor starvation and/or High calcium triggered KC differentiation. (a) Primary neonatal KC were cultured in growth medium (first lane), KC basal medium without growth factors (lanes 2 and 3), or growth medium with high calcium for 24 or 48 h as indicated. The expression of early differentiation marker K10 was measured by RTqPCR analysis. Fold induction of K10 compared to control cells (lane 1) was shown, and Hprt was used as housekeeping gene in the analysis. All error bars indicate mean  s.e.m. ***P < 0.001, ****P < 0.0001 (ANOVA). (b) Phase contrast images at 10
magnification of primary adult mouse KC treated with 0.2 mM CaCl2 at indicated time. Scale bar ¼ 100 μm 3.5 In Vitro Assays to Study KC Differentiation (See Note 11)

1. Early differentiation by growth factor starvation: Growth factor depletion alone is more efficient than high calcium to induce the expression of KC early differentiation markers, such as K10 (see Note 12). To starve the cells, remove growth medium and replace with basal medium without added growth supplements. K10 is induced (15-fold) as early as 24 h of GF removal, and this induction further increases by 48 h (~ 300-fold), whereas high calcium only leads to ~ten-fold of K10 induction by 48 h (Fig. 2a). 2. To trigger terminal differentiation by high calcium, proliferating KC are first starved overnight in basal medium without added growth supplements as described above (see Note 13). Next day, add CaCl2 to 0.2 mM in culture medium. As shown in Fig. 1a, b, within 8~12 h after high calcium switch, cells become flattened, and the distinct intercellular space becomes

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less apparent; by 24 h the cell-cell adhesion with tight junction becomes apparent, and the formation of corneocytes/cornified envelop and vertical cell stratification is observed around 48~72 h post high calcium switch. 3. In vitro assays to study KC differentiation: Cells can be harvested at desired time points for RTqPCR and/or western blot analyses to determine the mRNA or protein expression of KC differentiation markers, such as early differentiation marker K10 or late differentiation markers such as FLG, INV, and/or LOR [3]. 3.6 In Vitro Assays to Study Pro-inflammatory Response of KC to Viral Products

3.7 UVB IrradiationMediated Cell Death and Secretion of TNFa from KC

1. Culture the mouse KC in KC growth medium until cells reach ~80% confluency. Starve the cells for 6~16 h in basal medium without added growth supplement (see Note 14). 2. Add 1 μg/mL poly(I:C), the synthetic viral dsRNA, directly to the culture medium of the cells (see Note 15). Cells can be harvested 4~24 h posttreatment for RTqPCR analysis of the expression of pro-inflammatory cytokines, such as Ifnb1 as shown in Fig. 3a, or ELISA to measure cytokine secretion to conditioned medium. 1. Culture the mouse KC in KC growth medium until cells reach desired confluency. Immediately prior to UVB irradiation, remove growth medium and replace with sterile PBS warmed to 37  C. 2. Treat cells with 25 mJ/cm2 UVB using handheld UVB lamps. After UVB irradiation, change cells back to fresh growth medium. Representative images of cells treated with UVB for 12 h and 24 h are shown in Fig. 3b. 3. To measure and quantify cell viability, cells grown in 96-well flat bottom plate are first treated with UVB as described above, and then treated cells are subjected to CCK-8 cell assay at 12 and 24 h post-UVB treatment as described in Subheading 3.4, step 2. 4. To measure TNFα secretion (see Note 16), cells grown in 24-well flat bottom plate are first treated with UVB as described above; conditioned medium is then collected from UVB-treated cells at desired time point. The amount of TNFα in the conditioned medium is measured by the mouse TNFα ELISA kit following the manufacturer’s instructions. As shown in Fig. 3c, TNFα was abundantly secreted from UVB-treated adult KC compared to untreated control cells.

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Fig. 3 Pro-inflammatory response of primary mouse KC to viral dsRNA or UVB irradiation. (a) Primary adult KC were treated with 1 μg/mL poly(I:C) or vehicle control for 4 h, and cells were subjected to RTqPCR analysis. Fold induction of Ifnb1 in poly(I:C)-treated cells compared to control cells was shown, and Hprt was used as housekeeping gene in the analysis. (b, c) Primary adult mouse KC were grown to confluency and then exposed to 25 mJ/cm2 UVB irradiation. (b) Phase contrast images at 10
magnification 12 h or 24 h after UVB irradiation compared to untreated control cells. Scale bar, 200 μm. (c) Secretion of TNFα was measured by ELISA in conditioned medium from control cells or cells treated with UVB for 24 h. All error bars indicate mean  s.e.m. ***P < 0.001 (ANOVA)

4

Notes 1. Peel the skin off the whole body slowly as one piece, and be careful not to break skin into pieces as this will result in cell loss during the dispase digestion step. 2. Make sure the skin is extended and not folded in the tube to allow efficient digestion of the whole skin. 3. Tail skin pieces from mice with the same genetic background can be combined and incubated in one 15 mL tube (up to five tails per tube) filled with 15 mL dispase solution. 4. Compared to trypsin, the TrypLE digestion solution is gentler on cells and can be inactivated by dilution alone without the need for trypsin inhibitors, such as FBS. Each 10 cm Petri dish should fit up to five drops of the TrypLE solution.

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5. Because the epidermal sheet is very fragile, the dermis should be lifted away from the epidermis slowly to prevent tearing of the epidermal sheet. 6. Use forceps to carefully stretch and unfold the epidermal sheet so that it is fully extended and floats on the digestion solution. 7. KC spontaneously differentiate in suspension, so prior to cell seeding, the cell suspension should be kept on ice and plated onto ECM-coated culture dishes as soon as possible (preferably within 1 h). 8. Coating of culture dishes with the appropriate extracellular matrix (ECM) should be done immediately after the cell count and as soon as possible prior to cell seeding. 9. A gelatin-based coating matrix works well for neonatal KC, whereas a collagen-based coating matrix is preferable for adult KC due to the decreased ability of adult cells to adhere compared to their neonatal counterparts. Either rat tail, bovine, or recombinant human type 1 collagen can be used here. 10. Cell proliferation can be measured by either a colorimetric cell counting assay using a dye that measures the metabolic activity from living cells or 5-bromo-20 -deoxyuridine (BrdU) incorporation assay to measure BrdU incorporated into the newly synthesized DNA during cell proliferation. 11. Calcium is considered the most physiological agent to trigger epidermal KC differentiation in vitro and in vivo in a similar manner. In vitro, low calcium (0.02 mM~0.1 mM) maintains the proliferation of basal KC as a monolayer, whereas high calcium (>0.2 mM) rapidly triggers a terminal differentiation process converting KC from basal cell morphology to stratified corneocyte morphology. 12. We show there that growth factor starvation is more efficient than high calcium to induce genes that are associated with KC early differentiation, such as K10 (Fig. 2a). While high calcium weakly induces the expression of early differentiation genes, it strongly induces the expression of KC late differentiation process, such as FLG, INV, and LOR [3]. These observations are in line with the in vivo observation that calcium concentration is actually low in both basal and the spinous layer (in which K10 is expressed) but rises in the granular layer (where late differentiation markers express) [11–13]. Together these evidences suggest that calcium is unlikely the key factor that drives basal cells to commit to early differentiation process. Instead cell cycle arrest (which can be triggered by growth factor starvation in vitro) is likely the key factor that drives basal cells to commit to early differentiation stage.

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13. The growth factor removal step may enhance but is not required for the high calcium triggered cellular changes associated with the late differentiation processes, including tight junction formation and vertical cell stratification. 14. We always include a growth factor starvation step so that cells are synchronized and more consistent results can be obtained. Starvation can also lower basal inflammatory signal. Medium change should be done at least 6 h prior to treatment as medium change alone induces stress and inflammatory response from the cells. 15. Poly (I:C) is diluted and added in a small volume (10 μL) directly to culture well to minimize disturbance to the cells. 16. TNFα is an important pro-inflammatory cytokine that is induced by UVB irradiation, and it drives KC apoptosis following UVB irradiation [14]. References 1. Fuchs E, Raghavan S (2002) Getting under the skin of epidermal morphogenesis. Nat Rev Genet 3:199–209 2. Bernard JJ, Cowing-Zitron C, Nakatsuji T, Muehleisen B, Muto J et al (2012) Ultraviolet radiation damages self noncoding RNA and is detected by TLR3. Nat Med 18:1286–1290 3. Zhang LJ, Bhattacharya S, Leid M, GanguliIndra G, Indra AK (2012) Ctip2 is a dynamic regulator of epidermal proliferation and differentiation by integrating EGFR and Notch signaling. J Cell Sci 125:5733–5744 4. Sambandam SAT, Kasetti RB, Xue L, Dean DC, Lu Q, Li Q (2015) 14-3-3sigma regulates keratinocyte proliferation and differentiation by modulating Yap1 cellular localization. J Invest Dermatol 135(6):1621–1628 5. Yuspa SH, Hennings H, Tucker RW, Jaken S, Kilkenny AE, Roop DR (1988) Signal transduction for proliferation and differentiation in keratinocytes. Ann N Y Acad Sci 548:191–196 6. Mack JA, Anand S, Maytin EV (2005) Proliferation and cornification during development of the mammalian epidermis. Birth Defects Res C Embryo Today 75:314–329 7. Borkowski AW, Kuo IH, Bernard JJ, Yoshida T, Williams MR et al (2015) Toll-like receptor 3 activation is required for normal skin barrier repair following UV damage. J Invest Dermatol 135:569–578 8. Zhang LJ, Sen GL, Ward NL, Johnston A, Chun K et al (2016) Antimicrobial peptide

LL37 and MAVS signaling drive interferonbeta production by epidermal keratinocytes during skin injury. Immunity 45:119–130 9. Yamasaki K, Di Nardo A, Bardan A, Murakami M, Ohtake T et al (2007) Increased serine protease activity and cathelicidin promotes skin inflammation in rosacea. Nat Med 13:975–980 10. Li FW, Adase CA, Zhang LJ (2017) Isolation and culture of primary mouse keratinocytes from neonatal and adult mouse skin. J Vis Exp (125):56027 11. Menon GK, Grayson S, Elias PM (1985) Ionic calcium reservoirs in mammalian epidermis: ultrastructural localization by ion-capture cytochemistry. J Invest Dermatol 84:508–512 12. Elias PM, Nau P, Hanley K, Cullander C, Crumrine D et al (1998) Formation of the epidermal calcium gradient coincides with key milestones of barrier ontogenesis in the rodent. J Invest Dermatol 110:399–404 13. Mauro T, Bench G, Sidderas-Haddad E, Feingold K, Elias P, Cullander C (1998) Acute barrier perturbation abolishes the Ca2+ and K+ gradients in murine epidermis: quantitative measurement using PIXE. J Invest Dermatol 111:1198–1201 14. Zhuang L, Wang B, Shinder GA, Shivji GM, Mak TW, Sauder DN (1999) TNF receptor p55 plays a pivotal role in murine keratinocyte apoptosis induced by ultraviolet B irradiation. J Immunol 162:1440–1447

Chapter 16 Isolation and Propagation of Mammary Epithelial Stem and Progenitor Cells Julie M. Sheridan and Jane E. Visvader Abstract Several methods of mammary gland dissociation have been described that utilize a combined strategy of mechanical and enzymatic dissociation to isolate mammary epithelial cells (MECs) from intact tissue (Smalley et al., J Mammary Gland Biol Neoplasia 17:91–97, 2012). Here we detail a robust method that enables the isolation of all major stem and progenitor MEC populations, which has been successfully used to study stem cell behavior when coupled with transplantation and in vitro assays (Shackleton et al., Nature 439:84–88, 2006; Bouras et al., Cell Stem Cell 3:429–441, 2008; Sheridan et al., BMC Cancer 15:221, 2015; Jamieson et al., Development 144:1065–1071, 2017). Furthermore, we outline two prominent methods for culturing MECs for the purposes of ex vivo manipulation or study: 2D feeder layer cultures and 3D Matrigel colony assays. Importantly, all outlined methods retain stem and progenitor cell behaviors and can be used in combination with downstream in vivo, in vitro, or in silico analyses. Key words Mammary gland, Epithelial stem cell, Progenitor cell, Single cell suspension, Stem cell culture

1

Introduction A large body of evidence suggests that a stem cell-based mammary epithelial differentiation hierarchy establishes the mammary gland and maintains function through rounds of differentiation and regression that accompany pregnancy and weaning [1]. Several mammary epithelial stem cell (MaSC) and progenitor cell populations have been isolated including transplantable bipotent MaSCs that demonstrate luminal and basal/myoepithelial differentiation capacity as well as a range of more restricted cell types that contribute to limited luminal or basal cell subtypes [2–6]. The characterization of these populations has provided significant insights into the processes that guide normal mammary development and homeostasis and tumorigenesis [12]. Beyond phenotyping, the isolation of MaSC and progenitor cell populations provides a valuable tool with which to study or manipulate mammary epithelial

Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940, https://doi.org/10.1007/978-1-4939-9086-3_16, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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cell behavior ex vivo, and robust purification and culture protocols are key to these endeavors [7–10]. Herein we describe a MEC isolation protocol that reliably isolates and maintains the viability of MaSC and progenitor cell populations [2, 11]. Additionally, we outline two methods of MaSC and progenitor cell culture: (1) a tractable 2D feeder layer system that readily expands MaSC and progenitor cells ex vivo, providing a means to chemically or genetically manipulate cells prior to downstream analyses [2], and (2) a Matrigel-based assay that is permissive for the differentiation of different stem/progenitor cells into morphologically distinguishable colony types, a feature that makes this system suitable for studies of cell behavior or function [2].

2

Materials Where possible, materials are prepared using aseptic technique and solutions are filter sterilized with 0.2 μm filters.

2.1 Materials for the Dissociation of Mammary Gland Tissue to a Single Cell Suspension

1. Sterile Dulbecco’s phosphate buffered saline solution, without calcium and magnesium (DPBS). 2. Sterile wash buffer: DPBS with 2% fetal calf serum (FCS). 3. Sterile MEC medium supplemented with 1% FCS (1% MEC medium): DMEM/Ham’s F12 containing GlutaMAX, 5 μg/ mL insulin, 500 ng/mL hydrocortisone, 10 ng/mL epidermal growth factor, 20 ng/mL cholera toxin plus 1% FCS (Table 1). 4. 10 concentrated digestion buffer I: 150,000 U collagenase, 50,000 U hyaluronidase, 50 mL DPBS. Mix, filter sterilize, and use immediately or freeze in single use aliquots at 20  C. 5. Digestion buffer II: Dissolve 40 mg EGTA and 10 mg polyvinyl alcohol in 90 mL DPBS on a low heat with stirring, allow to cool, and then add 10 mL 2.5% trypsin. pH to 7.4, filter sterilize, and freeze in single use aliquots at 20  C.

Table 1 Recommended volumes of dissociation reagents for virgin mammary glands Number of virgin mice (BL6)

1–2

3–5

6–8

Number of virgin mice (FVB/N)

1

2–4

5–7

Digestion buffer I

5 mL

10 mL

20 mL

Digestion buffer II

0.5–1 mL

2 mL

3 mL

Digestion buffer III

1 mL

2 mL

5 mL

DNase I volume

100

200

400

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6. Digestion buffer II: Dissolve 250 mg dispase in 50 mL DPBS. Filter sterilize, and use within 1 week of storage at 4  C or freeze in single use aliquots at 20  C. 7. 1 mg/mL DNase I solution: Dissolve 10 mg DNase I in 10 mL medium, filter sterilize, and use within 1 week of storage at 4  C or freeze in single use aliquots at 20  C. 8. Optional, 1.25 concentrated red blood cell lysis solution; 8 g NH4Cl in 1 L deionized water, filter sterilize, and store at 4  C. 9. Orbital shaking incubator. 10. McIlwain tissue chopper with standard table fitted with razor blade as per manufacturer’s instructions. 2.2 Materials for the Purification of MaSC and Progenitor Cells by Flow Cytometric Sorting

1. Single cell suspension of mammary gland cells (as obtained from Subheading 3.2). 2. Sterile DPBS. 3. Sterile wash buffer. 4. Sterile collection buffer: DPBS with 10% FCS. 5. Fluorescently conjugated antibodies, refer to Table 2. 6. Viability dye such as propidium iodide (PI) or 7-actinomycin D (7-AAD).

2.3 Materials for the Culture and Expansion of Primary MECs on a Feeder Layer

1. MEC as obtained from Subheadings 3.2 or 3.3 2. Sterile MEC medium supplemented with 1% FCS. 3. Sterile MEC medium supplemented with 5% FCS (5% MEC medium, modified version of 1% MEC medium made in Subheading 2.1). 4. Sterile collagen coated 6-well tissue culture plates (see Note 1). 5. NIH/3T3 cells, irradiated (i3T3) (see Note 2). 6. 37  C incubator maintained with 5% CO2 and 5% O2.

2.4 Materials for the Culture of MEC in a Matrigel Colony Assay

1. MECs as obtained from Subheadings 3.2 or 3.3. 2. Growth factor reduced Matrigel, thawed on ice. 3. 1% MEC medium 4. Glass chamber slides (Ibidi). 5. 37  C incubator maintained with a low O2 gas phase (5% CO2, 5% O2, and 90% N2) 6. Pipette tips, precooled to 4  C. 7. Harvest only: Cell recovery solution (BD Bioscience) and wash buffer.

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Table 2 Anti-mouse antibody clones that facilitate the identification and isolation of indicated mammary epithelial stem and progenitor cell populations

Cell population and marker criteria

Antibody clone (conjugate)

References

Blocks non-specific (FcγIII and FcγII) staining

CD16/CD32 (2.4G2)

Lineage cocktail (to deplete hematopoietic cells, red blood cells, and endothelial cells)

CD45 TER-199 CD31

Lin CD29lo CD24+ CD14+ luminal progenitor-enriched Lin CD29lo CD24+ CD14 mature luminal cell-enriched (pregnant/lactating)

CD14

[13]

Lin CD29hi CD24+ basal cells incl. MaSC; Lin CD29lo CD24+ luminal cells incl. Luminal progenitor

CD29

[2]

Lin CD29hi CD24+ basal cells incl. MaSC; Lin CD29lo CD24+ luminal cells incl. Luminal progenitor

CD24

[2]

Lin CD24+ CD29lo CD49b+ Sca-1 hormone-receptor luminal progenitor; Lin CD24+ CD29lo CD49b+ Sca-1+ hormonereceptor+ luminal progenitor

CD49b

[14]

Lin CD29lo CD24+ CD61+ LP-enriched; Lin CD29lo CD24+ CD61 mature luminal cell-enriched (virgin)

CD61

[6]

Lin CD24+ CD29lo CD49b+ Sca-1 hormone-receptor luminal progenitor; Lin CD24+ CD29lo CD49b+ Sca-1+ hormonereceptor+ luminal progenitor

SCA-1

[14]

Lin CD29hi CD24+ TSPAN8hi/lo MaSC subsets

TSPAN8

[9]

3

Methods Where possible, manipulations are performed using aseptic technique and/or in a sterile environment such as a tissue culture hood.

3.1 Harvesting Mouse Mammary Gland Tissue

1. Wipe the ventral midline of the euthanased mouse with 70% ethanol. 2. Make an incision along the ventral midline through the skin from the pubis to the neck taking care to avoid damage to the peritoneal membrane (Fig. 1). 3. At the base of the midline incision, make further oblique cuts toward and half of the way along the hind legs (Fig. 1). 4. Using forceps to hold the skin to one side of the junction of the midline and oblique cuts, gently peel it outward to separate the skin from the internal membrane to reveal the fourth and fifth

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3

4

LN

5

Fig. 1 Diagram of mammary gland dissection. Incisions (dashed lines) are made through the skin along the midline and hind legs. The skin is peeled away from the body and pinned in position to reveal the third, fourth, and fifth mammary glands (3, 4, and 5, respectively). The inguinal lymph node (LN) can be removed prior to excision of the fourth gland to minimize hematopoietic cell contamination of the harvested tissue

mammary glands. This is best achieved using a second pair of forceps to apply counter-pressure to the internal membrane to prevent it from being pulled in the same direction as the skin. 5. Repeat this motion on the opposite side and pin the flayed skin to the dissection board (Fig. 1). 6. Grasp the skin to one side of the midline cut at the level of the forelegs and peel the skin outward from the body to reveal the second and third mammary glands on the underside of the skin. 7. Repeat this motion on the opposite side and pin the flayed skin to the dissection board (Fig. 1). 8. Remove and discard the inguinal lymph node that is located at the intersection of the three prominent vessels on the surface of the fourth mammary gland. This is best achieved by pushing the skin upward from the outside, thus raising the area of the vessel junction. In this position, the relative firmness of the lymph node and its characteristically gray-white, circular appearance facilitate its identification and excision using forceps or scissors.

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9. Remove the mammary gland tissue by grasping and raising the outer edge of the attached gland away from the skin and membrane. Cut and peel the outer edges of the gland away from the serous membrane and repeat this process while moving toward the dorsal midline of the animal, releasing the whole gland in one piece. 10. Place the mammary glands into MEC medium on ice. 3.2 Dissociation of Mammary Gland Tissue to a Single Cell Suspension

1. Prepare the materials outlined in Subheading 2.1.

3.2.1 Preparation

3. Thaw a suitable amount of digestion buffers II and III and DNase I at room temperature (Table 1).

2. Thaw and then dilute a suitable volume (Table 1) of 10 stock of digestion buffer I in MEC medium to yield a 1 solution and warm to 37  C.

4. Pre-warm a shaking incubator to 37  C. 3.2.2 Mechanical Disruption

1. Prepare the McIlwain tissue chopper as per manufacturer’s instructions. Briefly, with the machine turned off, affix a clean plastic disc to the cutting table using the spring-loaded clips and position the blade so that it touches the plastic disc. Once ready, position the blade in the starting position at the righthand side of the plastic circle (see Notes 3 and 4). 2. Drain the mammary glands of excess buffer/medium and place on the plastic disc. Multiple runs may be necessary to chop all of the tissue depending upon the number of glands collected and the mass of tissue obtained (see Note 4). 3. Turn the chopper on and allow to run across the tissue until it stops. Raise the blade, return it to the starting position, and rotate the plastic disc one quarter turn to crosscut the sample. This process should be repeated for a total of four cutting runs or until the gland no longer presents as lumps when lifted with forceps (see Notes 4 and 5). 4. Place chopped tissue into a 50 mL conical tube for further processing.

3.2.3 Enzymatic Digestion to a Single Cell Suspension

1. With reference to Table 1, add an appropriate volume of pre-warmed 1 digestion buffer I, seal the tube, and place in a shaking incubator at 37  C for 30 min (see Note 5). 2. Triturate the sample ten times using a pipette and incubate for a further 20 min at 37  C. 3. Triturate the sample a further ten times and check for the absence of large fragments. If any remain, incubate for a further 10 min and then re-triturate until a relatively homogeneous organoid solution has been achieved.

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4. Add 20–40 mL wash buffer to the sample and centrifuge for 5 min at 1200 rpm (200 RCF) to collect the organoids. 5. Pre-warm digestion buffers II and III and DNase I to 37  C. 6. Using a vacuum pump or pipette, remove the supernatant. Care must be taken to remove the concentrated band of adipose flocculate that overlays the supernatant (see Note 6). 7. Add DNase I to the organoid pellet, tap to partially resuspend, and leave at room temperature for 2 min. 8. Add a suitable volume (Table 1) (see Note 5) of pre-warmed digestion buffer II to the organoids, pipette gently to fully suspend, and incubate in a water bath at 37  C for 2–3 min. 9. Inactivate and dilute the trypsin by adding 30 mL wash buffer. Centrifuge for 5 min at 1200 rpm and discard the supernatant (see Note 6). 10. Add DNase I to the organoid pellet, tap to partially resuspend, and wait for 2 min. 11. Add a suitable volume (Table 1) (see Note 5) of pre-warmed digestion buffer III to the organoids, pipette gently to resuspend the organoids, and incubate in a water bath at 37  C for 5 min. 12. Gently triturate the suspension to check for the presence of a single cell suspension; no clumps should remain, and the “grainy” appearance of the cell/organoid suspension that was evident before digestion buffer III should now be gone. 13. Add 30 mL wash buffer and centrifuge for 5 min at 1200 rpm (200 RCF) to collect the cells. Remove the supernatant to yield a pellet of single cells and proceed with assays or further purification. 3.3 Purification of Mammary Epithelial Stem and Progenitor Cell Populations

1. Prepare the materials outlined in Subheading 2.2. 2. Isolate mammary cells using the protocol outlined in Subheading 3.2.

3.3.1 Preparation 3.3.2 Red Blood Cell Lysis

1. Resuspend cells in 100 μL DNase I and 900 μL wash buffer. 2. Slowly add 4 mL of room temperature red blood cell lysis buffer while swirling tube to facilitate gentle mixing. 3. Incubate at room temperature for 3 min. 4. Add 20 mL wash buffer and pass through a 40, 70, or 100 μm cell strainer to remove any clumps. 5. Pass another 20 mL wash buffer through the cell strainer to ensure maximal cell recovery.

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6. Take a small aliquot to determine the cell concentration using a hemocytometer or similar. Together with the known volume, use the cell concentration to calculate the total cell count. 7. Centrifuge for 5 min at 1200 rpm to collect the cells. Discard the supernatant and place the tube of cells on ice. Form this point on, use chilled solutions and keep the cells on ice. 3.3.3 Staining a Single Cell Suspension for the Purpose of Mammary Stem and Progenitor Cell Identification

What follows is a brief overview of a basic staining protocol. It is by no means exhaustive, and other considerations not covered in this protocol include the type of sorter to be used, the aim of the sort (yield or purity), and antibody-fluorochrome panel design. 1. Prepare the materials outlined in Subheading 2.2. 2. Resuspend the cells in cold wash buffer at a concentration of 2.5  107 cells per mL. 3. [Optional] Incubate the cells with an appropriate volume of unconjugated anti-mouse CD16/CD32 antibody (Table 2) to minimize non-antigen-specific binding of fluorescently conjugated antibodies to Fc receptors. 4. Distribute the cells to staining vessels as required. Where necessary, include cells for single color and fluorescence-minusone controls. 5. Add primary antibodies to the cell suspensions (Table 2). Incubate on ice for 25 min. 6. Add wash buffer and centrifuge for 5 min at 1200 rpm (200 RCF) to collect the cells. Discard the supernatant. 7. [Optional] When necessary, steps 5 and 6 may be repeated with secondary antibodies. 8. Prepare collection tubes for cells post-sort containing a small amount of collection buffer. Vessel choice and volume depend on several factors including expected cell yield and sort nozzle (and thus droplet) size. For example, a small number of cells (10,000) might be sorted through a 100 μm nozzle into an Eppendorf tube. However, a larger number of cells will require a 5 mL FACS tube or multiple Eppendorf tubes. 9. Resuspend the cell pellet in a suitable volume of wash buffer containing a viability dye such as 2 μg/mL propidium iodide (PI) or 5 μg/mL 7-actinomycin D (7-AAD) in preparation for sorting. 10. Fluorescence-minus-one controls and population contours should be used to inform appropriate gate placement. Gates may be configured to exclude doublets and debris on the basis of light scatter properties, and viable cells can be selected on the basis of viability dye exclusion. Several mammary epithelial stem and progenitor cell populations lie within the Lineage (Lin ) CD29hi CD24+ fraction and suggested sort criteria for

Singlets

Cells

Viable

FSC-A Luminal

FSC-A

Basal/ MaSC

Mature Luminal Modal

CD24

CD45/TER-119/CD31

FSC-A

225

7-AAD

SSC-A

SSC-W

Mammary Stem Cell Isolation and Culture

Luminal Progenitor

Lineageneg CD29

CD29

CD61

Fig. 2 Representative plots showing a flow cytometric gating strategy designed to distinguish single, viable CD29hi CD24+ basal cells, Lin CD29lo CD24+ CD61+ luminal progenitors, and Lin CD29lo CD24+ CD61 mature luminal cells

specific mammary stem or progenitor cell populations are outlined in Table 2 and Fig. 2. 11. Cells may be sorted on any flow cytometer and each requires optimization. Consideration should be given to both nozzle size and sort pressures since the factors can greatly affect viability of large fragile cells such as MECs (see Note 7). 12. [Optional] Reanalysis may be performed on a small aliquot of collected cells to confirm the identity and establish the purity of sorted cells. 3.4 Ex Vivo Propagation of MEC on a Feeder Layer

1. Prepare materials as per Subheading 2.3 under aseptic conditions.

3.4.1 Preparation 3.4.2 Plating Cells for Expansion

1. All cell manipulations are performed using aseptic technique in a sterile TC hood. 2. For each well of a 6-well plate to be plated, mix 2.5 mL 5% MEC medium with 100,000 i3T3 (100 μL of thawed stock, final density approximately 14,000/cm2) and purified MEC.

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Fig. 3 Representative bright-field images of colonies generated from plating purified mammary epithelial stem and progenitor cell-enriched populations. (Top panel) 300 Lin CD29hi CD24+ basal/MaSCs or Lin CD29lo CD24+ CD61+ luminal cells were plated in 2D feeder layer-based cultures. After 6 days, cultures were briefly fixed and stained with Giemsa to visualize colony morphology. Scale bars, 250 μm. (Bottom panel) 1000 Lin CD29hi CD24+ basal cells or 1500 Lin CD29lo CD24+ luminal cells were grown for 14 days in the 3D Matrigel assay. Scale bars, 500 μm. Inset, magnified region showing typical solid (basal) or acinar (luminal) colony types. Scale bars, 250 μm

As a guide, 10,000–20,000 basal/MaSC or luminal cells from FVB/N or 20,000–50,000 basal/MaSC or luminal cells from C57BL/6 will give a MEC-dominated plated after 6 days in culture. 3. Place in a 37  C incubator maintained at 5% CO2 and 5% O2. 4. Once situated, to ensure that cells are evenly distributed across the plate, gently slide the plate forward and backward five times and then repeat using a side-to-side motion. 5. 24 h later, viable cells will have attached. Discard the medium in the well and immediately replace with fresh 1% MEC. 6. Maintain the culture by changing the MEC medium every 3 days. Cells may be manipulated at any time during culture using transduction, transfection, or application of medium additives as described elsewhere.

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7. Culture until the colonies of epithelial cells have expanded and dominate the plate (typically less than 1 week with purified basal or luminal cells). Several different colony morphologies will contribute to epithelial outgrowth. These are most obvious when cells are plated at lower densities as shown in Fig. 3. 3.4.3 Harvesting Expanded MECs

1. Remove medium and wash cells gently with DPBS. 2. Add 0.5 mL pre-warmed trypsin-EDTA to each well and incubate at 37  C for 10–15 min. 3. Using a microscope, check to see that cells are detaching from the plate. Luminal cells will detach more easily. 4. Using a P1000 pipette, gently triturate cells to aid detachment from the plate and re-incubate if necessary. 5. Repeat steps 3 and 4 until cells are in a single cell suspension. 6. Collect cells into a falcon tube and add several times the volume of cold wash buffer (containing FCS) to quench the trypsin activity. 7. Centrifuge for 5 min at 1200 rpm to collect the cells for downstream analyses.

3.5 Culture of MECs as Colonies in Matrigel 3.5.1 Preparation

3.5.2 Organoid Culture Initiation and Maintenance

1. Prepare materials as in Subheading 2.4. Due to the small volumes involved with Matrigel culture initiation, manipulations should be done in bulk with master mixes, where possible. Always prepare extra to accommodate unavoidable losses, and keep plates, tubes, and pipette tips cold to facilitate the handling of Matrigel, which polymerizes quickly when warmed. 1. Calculate the cell number and Matrigel volume to be plated. As a starting point, 1000 single cells can be plated in one 20 μL drop of Matrigel with 1 drop per well of an 8-chamber slide. 2. Prepare materials as per Subheading 2.4, taking into account the quantity of material to be plated. 3. Cells to be plated are centrifuged in an Eppendorf tube and medium aspirated to leave a minimum volume (not greater than 10–15 μL). 4. Resuspend cell pellet in Matrigel by gentle trituration with a pre-chilled pipette tip, taking care not to introduce any bubbles (see Note 8). 5. Pipette 20 μL drop into each chamber of the slide (see Note 8). 6. Transfer to 37  C incubator for 10–15 min to allow Matrigel to polymerize. 7. Gently pipette 400 μL of pre-warmed 1% MEC medium into each well.

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8. Return to incubator and maintain with medium changes every 3–4 days intervals. 9. After a total of 12–14 days, mature colonies may be imaged or harvested for further analysis. Characteristic colony morphologies will be identifiable following culture (Fig. 3). 10. Cell recovery solution used as per manufacturer’s instructions can be used to isolate colonies for downstream analyses (see Note 9).

4

Notes 1. The use of plates pre-coated with collagen is not necessary but shortens the time to confluency. 2. For production: Grow sufficient numbers of NIH3T3, e.g., 1–5 T225 flasks, and harvest when actively growing but nearing confluence (~85% confluent). Harvest cells using trypsin, quench with NIH3T3 growth medium containing 10% FCS, centrifuge and resuspend cells in NIH3T3 growth medium containing 10% FCS, and place on ice. Irradiate the cells with 50 Gy and count the cell suspension using an automated cell counter or hemocytometer to determine the total cell number. Spin and resuspend the cells at a density of 1  106 cells/mL in cold freezing medium such as 50% DMEM + 40% FCS + 10% DMSO. Aliquots of 1 mL (1  106 i3T3) can be frozen using a typical cell freezing protocol for long-term storage. 3. When cells will be cultured, ensure that the cutting station is clean and the plastic discs are clean and sterilized prior to use. 4. Slice thickness and strike force for the McIlwain tissue chopper should be determined empirically. The chopped gland should have only small lumps and have a thick slurry-like consistency. 5. This protocol generalizes the requirements to dissociate glands from virgin mice into single cells. Due to the increased mass of pregnant or lactating glands, volumes in Table 1 should be adjusted commensurate with size. 6. Large flocculates may be observed that consist of viable cells held together by DNA released from damaged cells. Although these will disperse when DNase I is added, care must be taken to retain them since they can be easily lost during supernatant aspiration. 7. We routinely use 70 or 100 μm nozzles and sort at low-tomedium sort pressures to achieve consistent viability and yields 8. Avoid the introduction of bubbles into the Matrigel by careful pipetting. If bubbles are introduced, they will be difficult to remove but, by preparing a little extra Matrigel/cell suspension, they may be excluded during future pipetting strokes.

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A single bubble that has been introduced while pipetting 20 μL drops into the chambers can be occasionally removed by simply drawing the Matrigel back up into the tip and re-pipetting it. 9. To mitigate the sticky nature of 3D colonies during harvesting, care should be taken to avoid unnecessary manipulations and minimize the surfaces that they contact. Additional safeguards include using pipette tips pre-wet with FCS to resuspend the organoids, since tapping can distribute organoids across the Eppendorf tube walls, which will result in cell losses. References 1. Visvader JE, Stingl J (2014) Mammary stem cells and the differentiation hierarchy: current status and perspectives. Genes & Dev 28:1143–1158. 2. Shackleton M, Vaillant F, Simpson KJ et al (2006) Generation of a functional mammary gland from a single stem cell. Nature 439:84–88. 3. Stingl J, Eirew P, Ricketson I et al (2006) Purification and unique properties of mammary epithelial stem cells. Nature 439:993–997. 4. Sleeman KE, Kendrick H, Robertson D et al (2007) Dissociation of estrogen receptor expression and in vivo stem cell activity in the mammary gland. J Cell Biol 176:19–26. 5. Fu NY, Rios AC, Pal B et al (2017) Identification of quiescent and spatially restricted mammary stem cells that are hormone responsive. Nat Cell Biol 19:164–176. 6. Asselin-Labat M-L, Sutherland KD, Barker H et al (2007) Gata-3 is an essential regulator of mammary-gland morphogenesis and luminalcell differentiation. Nat Cell Biol 9:201–209. 7. Bouras T, Pal B, Vaillant F et al (2008) Notch Signaling Regulates Mammary Stem Cell Function and Luminal Cell-Fate Commitment. Cell Stem Cell 3:429–441. 8. Sheridan JM, Ritchie ME, Best SA et al (2015) A pooled shRNA screen for regulators of primary mammary stem and progenitor cells

identifies roles for Asap1 and Prox1. BMC Cancer 15, 221. 9. Jamieson PR, Dekkers JF, Rios AC et al (2017) Derivation of a robust mouse mammary organoid system for studying tissue dynamics. Development 144:1065–1071. 10. Smalley MJ, Kendrick H, Sheridan JM et al (2012) Isolation of Mouse Mammary Epithelial Subpopulations: A Comparison of Leading Methods. J Mammary Gland Biol Neoplasia 17:91–97. 11. Pal B, Bouras T, Shi W et al (2013) Global changes in the mammary epigenome are induced by hormonal cues and coordinated by Ezh2. Cell Rep 3:411–426. 12. Asselin-Labat M-L, Sutherland KD, Vaillant F et al (2011) Gata-3 negatively regulates the tumor-initiating capacity of mammary luminal progenitor cells and targets the putative tumor suppressor caspase-14. Mol Cell Biol 31:4609–4622. 13. Li W, Ferguson BJ, Khaled WT et al (2009) PML depletion disrupts normal mammary gland development and skews the composition of the mammary luminal cell progenitor pool. Proc Natl Acad Sci USA 106:4725–4730. 14. Barcellos-Hoff MH, Aggeler J, Ram TG, Bissell MJ (1989) Functional differentiation and alveolar morphogenesis of primary mammary cultures on reconstituted basement membrane. Development 105:223–235

Chapter 17 An Organoid Assay for Long-Term Maintenance and Propagation of Mouse Prostate Luminal Epithelial Progenitors and Cancer Cells Yu Shu and Chee Wai Chua Abstract Historically, prostate luminal epithelial progenitors and cancer cells have been difficult to culture, thus hampering the generation of representative models for the study of prostate homeostasis, epithelial lineage hierarchy relationship and cancer drug efficacy assessment. Here, we describe a newly developed culture methodology that can efficiently grow prostate luminal epithelial progenitors and cancer cells as organoids. Notably, the organoid assay favors prostate luminal cell growth, thus minimizing basal cell dominance upon the establishment and continuous propagation of prostate epithelial cells. Importantly, organoids cultured under this condition have demonstrated preservation of androgen responsiveness and intact androgen receptor signaling, providing a representative system to study castration resistance and androgen receptor independence. Key words Organoid culture, Prostate luminal progenitors, Prostate cancer, Androgen receptor, Genetically engineered mouse, Castration resistance

1

Introduction Recent advances in the three-dimensional culture system have enabled the maintenance and propagation of various cell lineages from different organ systems under defined in vitro conditions [1–3]. In normal prostate, the epithelium consists of three major cell lineages, including luminal cells, basal cells as well as an extremely rare neuroendocrine cell population [4]. The majority of prostate cancers in comparison are adenocarcinomas, which are characterized by an exclusively luminal phenotype [4]. In the past decades, there have been many attempts to establish culture conditions for prostate luminal and cancer cells, but most of these studies have yielded very little promising results [5]. However, these culture conditions are neither optimized for prostate luminal and/or cancer cell survival nor favor long-term propagation of the luminal

Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940, https://doi.org/10.1007/978-1-4939-9086-3_17, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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population. With prolonged culture, basal cells will eventually outcompete luminal cells and become the dominant cell population in culture. When using primary mouse or human prostate specimens to isolate different prostate epithelial cell lineages, introduction of tissue dissociation procedures may lead to changes in lineage marker expression (unpublished observation). Consequently, it is extremely difficult to ascertain the lineage origin of sorted cell populations to be used for subsequent optimization of culture conditions. The use of genetically engineered mouse (GEM) models has provided excellent opportunities to study prostate homeostasis, regeneration, tumor initiation, and progression [6]. In particular, lineage tracing using GEM models has identified the first prostate luminal epithelial progenitor population, termed castrationresistant Nkx3.1-expressing cells (CARNs) that can generate basal and luminal cell populations and are capable of serving as a cell of origin for prostate cancer [7]. Using CARNs as a starting population, we have developed and optimized a novel organoid assay that can maintain and propagate the prostate luminal progenitor population as well as other prostate epithelial cells in the long term (Figs. 1, 2 and 3) [8, 9]. Notably, using lineage-marked prostate luminal and basal cells derived from GEM models, we have convincingly demonstrated that the culture method favors luminal cell growth. Moreover, under this culture condition, prostate luminal progenitors can generate basal progeny, implying a model that recapitulates in vivo condition. While organoids derived from basal cells are generally much smaller in size, co-culture of basal and luminal cells can promote organoid growth (Fig. 4). These results have indicated that our organoid methodology minimizes the basal cell dominance issue in prostate epithelial culture and can serve as a representative model for the study of prostate homeostasis. More importantly, the organoid assay has facilitated the establishment of various tumor organoid lines from GEM models of prostate cancer (Fig. 5). Notably, derived organoids demonstrate preservation of androgen responsiveness and intact androgen receptor signaling, which are highly crucial and relevant for the study of prostate homeostasis as well as cancer initiation and progression (Fig. 6). Lastly, it is worth noting that another study has also established an ENR (EGF, Noggin, R-spondin)-based organoid culture condition, which enables the growth of both prostate basal and luminal epithelial cells [10]. Because this assay favors the growth of basal over luminal cells [10], it remains unclear whether basal and luminal cells in the derived organoids can be maintained equally after serial passaging. Here, we describe a comprehensive protocol, which involves the use of hepatocyte-based medium that is cost-effective and easy to prepare for the isolation, maintenence and propagation of mouse prostate luminal progenitors and cancer cells as organoids. We also

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present detailed protocols for the characterization of various organoid types as well as growth and gene expression assessment of prostate epithelial organoids in response to androgen withdrawal.

2

Materials We have included suppliers and catalogue numbers for certain reagents because in our hands, we have found that similar reagents from other suppliers do not perform as well.

2.1

Mouse Models

See Table 1 for primers used to genotype major GEM models listed below. 1. To lineage-mark the prostate luminal progenitor population, CARNs, castrate 8–12 weeks old Nkx3.1CreERT2/+; R26R-YFP/ + mice and treat the mice with tamoxifen (Sigma #T5648) (9 mg per 40 g body weight in corn oil) by daily oral gavage for 4 consecutive days after a month of castration. Dissect and analyze the mice after a month of tamoxifen induction (see Note 1). 2. Use the following Cre- or inducible Cre-expressing mouse models to isolate different prostate epithelial and cancer cells (see Note 2): (a) Nkx3.1Cre/+; R26R-YFP/+: Lineage-marks all prostate epithelial cells during embryonic stage (b) Nkx3.1CreERT2/+; R26R-YFP/+: Lineage-marks mainly prostate luminal cells as well as a small prostate epithelial population that co-expresses basal and luminal markers upon tamoxifen induction at adult stage (c) CK8-CreERT2; R26R-YFP/+ or CK18-CreERT2; R26RYFP/+: Lineage-marks prostate luminal cells upon tamoxifen induction at adult stage (d) Various types of GEM model of prostate cancer that carry R26R-YFP allele, such as Nkx3.1CreERT2/+; Pten flox/flox; R26R-YFP/+ (NP), Nkx3.1CreERT2/+; Pten flox/flox; R26R-YFP/+ (NPK), and KrasLSL-G12D/+; Nkx3.1CreERT2/+; Pten flox/flox; p53 flox/flox; R26R-YFP/+ (NPP53): Lineage-marks and oncogenic transforms prostate epithelial cells upon tamoxifen induction at adult stage 3. Prepare the following mice for isolation of unlabeled prostate epithelial and cancer cells: (a) Wild-type C56BL/6 mice—for isolation of prostate epithelial cells

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Table 1 Primers used for mouse genotyping

Allele

Amplicon size Forward 0

Reverse

Nkx3.1CreERT2 500 bp

5 -CAG ATG GCG CGG CAA CAC C-30

50 -GCG CGG TCT GGC AGT AAA AAC-30

Nkx3.1 wildtype

500 bp

50 -CTC CGC TAC CCT AAG CAT CC-30

50 -GAC ACT GTC ATA TTA CTT GGA CC-30

Nkx3.1 null

232 bp

50 -TTC CAC ATA CAC TTC ATT CTC AGT-30

50 -GCC AAC CTG CCT CAA TCA CTA AGG-30

Nkx3.1 wildtype

707 bp

50 -GTC TTG GAG AAG AAC TCA CCA TTG-30

50 -GCC AAC CTG CCT CAA TCA CTA AGG-30

CreERT2

500 bp

50 -CAG ATG GCG CGG CAA CAC C-30

50 -GCG CGG TCT GGC AGT AAA AAC-30

R26R-YFP

320 bp

50 -AAA GTC GCT CTG AGT TGT TAT-30

50 -AAG ACC GCG AAG AGT TTG TC-30

R26R wild-type 600 bp

50 -AAA GTC GCT CTG AGT TGT TAT-30

50 -GGA GCG GGA GAA ATG GAT ATG-30

R26R-Tomato

196 bp

50 -CTG TTC CTG TAC GGC ATG G-30

50 -GGC ATT AAA GCA GCG TAT CC-30

R26R-CAGYFP

212 bp

50 -ACA TGG TCC TGC TGG AGT TC-30

50 -GGC ATT AAA GCA GCG TAT CC-30

R26R wild-type 297 bp

50 -AAG GGA GCT GCA GTG GAG TA-30

50 -CCG AAA ATC TGT GGG AAG TC-30

Pten flox

320 bp

50 -CAA GCA CTC TGC GAA CTG AG-30

50 -AAG TTT TTG AAG GCA AGA TGC-30

Pten wild-type

156 bp

50 -CAA GCA CTC TGC GAA CTG AG-30

50 -AAG TTT TTG AAG GCA AGA TGC-30

Pten null

320 bp

50 -TTG CAC AGT ATC CTT TTG AAG-30

50 -ACG AGA CTA GTG AGA CGT GC-30

Pten wild-type

240 bp

50 -TTG CAC AGT ATC CTT TTG AAG-30

50 -GTC TCT GGT CCT TAC TTC C-30

KrasLSL-G12D

550 bp

50 -AGC TAG CCA CCA TGG CTT GAG TAA GTC TGC A-30

50 -CCT TTA CAA GCG CAC GCA GAC TGT AGA-30

Kras wild-type

500 bp

50 -GTC GAC AAG CTC ATG CGG GTG-30

50 -CCT TTA CAA GCG CAC GCA GAC TGT AGA-30

TRAMP

650 bp

50 -GCG CTG CTG ACT TTC TAA ACA TAA G-30

50 -GAG CTC ACG TTA AGT TTT GAT GTG T-30

p53 flox

370 bp

50 -CAC AAA AAC AGG TTA AAC CCA G-30

50 -AGC ACA TAG GAG GCA GAG AC-30 (continued)

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Table 1 (continued)

Allele

Amplicon size Forward

p53 wild-type

288 bp

50 -CAC AAA AAC AGG TTA AAC CCA G-30

50 -AGC ACA TAG GAG GCA GAG AC-30

Hi-Myc

177 bp

50 -AAA CAT GAT GAC TAC CAA GCT TGG C-30

50 -ATG ATA GCA TCT TGT TCT TAG TCT TTT TCT TAA TAG GG-30

Reverse

(b) Various types of GEM model of prostate cancer that are not lineage-marked, such as TRAMP, Nkx3.1/, Hi-Myc and Nkx3.1+/; Pten+/ mice (see Note 3). 2.2 General Equipment and Consumables

1. CO2 euthanasia chamber. 2. Laminar flow hood or biological safety cabinet. 3. Dissecting microscope. 4. Micro-dissecting instruments. 5. P20, P200, and P1000 pipettes and pipette tips. 6. Water baths set at 37  C and 55  C. 7. CO2 incubator. 8. Centrifuges (for Eppendorf and Falcon tubes). 9. Orbital shaker. 10. Hemocytometer/automated cell counter. 11. BD FACSAria cell sorter (or similar). 12. Olympus IX51 inverted microscope with fluorescent lamp, camera, and computer (or similar). 13. Tissue processor and embedder. 14. Leica microtomes, histology water bath, and slide warmer (or similar). 15. Leica cryostats (or similar). 16. Histology microscope. 17. Leica SP5 confocal microscope (or similar). 18. Eppendorf Mastercycler Realplex2 (or similar). 19. Luminometer plate reader. 20. Food steamer. 21. DAKO pen. 22. Humidified chamber. 23. Insulated cryofreezing container.

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24. Sterile petri dishes. 25. 96-well low attachment plates (Corning #3474). 26. 1.5 mL Eppendorf tubes. 27. 15 and 50 mL Falcon tubes. 28. Cell strainer 40 and 70 μm. 29. 1.8 mL cryotubes. 30. Parafilm. 31. Cryomold for biopsy specimen 10  10  5 mm, 100/pcs. 32. Greiner 96-well CELLSTAR plate (or similar opaque-walled 96-well plate). 2.3 Prostate Dissection and Dissociation, and Preparation of Single Cell Suspension

1. Phosphate-buffered saline (PBS) for dissection. 2. 10 collagenase/hyaluronidase solution (STEMCELL Technologies #07912). 3. Dulbecco’s Modified Eagle Medium: Nutrient Mixture F12 supplemented with 5% fetal bovine serum (FBS). 4. 0.25% trypsin-EDTA (STEMCELL Technologies #07901). 5. Hanks’ balanced salt solution modified (HBSS) (STEMCELL Technologies #37150) supplemented with 2% FBS. 6. Dispase 5 U/mL (STEMCELL Technologies #07913). 7. DNase I solution 1 mg/mL (STEMCELL Technologies #07900). 8. 0.4% trypan blue solution. 9. Reconstitute ROCK inhibitor Y-27632 (STEMCELL Technologies #72302) in PBS to make 5 mM stock solution (see Note 4).

2.4 FluorescenceActivated Cell Sorting (FACS)

1. APC anti-mouse EpCAM antibody (BioLegend #118214). 2. PerCP-Efluor710 anti-mouse/human E-cadherin antibody (eBiosciences #46-3249-82). 3. Prepare 0.5 mg/mL DAPI solution by dissolving DAPI powder in Milli-Q water (see Note 5). 4. Prepare collecting medium consisting of HBSS supplemented with 2% FBS and 10 μM ROCK inhibitor Y-27632.

2.5 Prostate Epithelial Organoid Culture

1. Hepatocyte Culture Media Kit (Corning #355056) (see Note 6). 2. Prepare heat-inactivated charcoal-stripped FBS by heating the reagent in 55  C water bath for 60 min (see Note 7). 3. 100 Glutamax. 4. Matrigel (Corning #354234) (see Note 8).

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5. 5 mM stock solution of ROCK inhibitor Y-27632. 6. 105 M dihydrotestosterone (DHT, Sigma #A-8380) in absolute ethanol (see Note 9). 7. 100 antibiotic-antimycotic. 2.6 Organoid Passage and Preparation of Frozen Organoid Stocks

1. Cold PBS. 2. 0.25% trypsin-EDTA. 3. HBSS supplemented with 2% FBS. 4. Organoid media with complete supplements (see Subheading 3.3). 5. Dispase 5 U/mL. 6. DNase I solution 1 mg/mL. 7. FBS. 8. Dimethyl sulfoxide (DMSO).

2.7 Histology and Immunostaining of Organoids

1. 10% neutral buffered formalin. 2. 4% paraformaldehyde (PFA) solution in PBS. 3. 30% sucrose in PBS. 4. Tissue-Tek OCT compound. 5. Collagen I, high concentration, rat tail, 100 mg (Corning #354249). 6. Prepare setting solution for collagen I by combining 100 mL 10 RPMI (with phenol red), 7.5 mL 1 M NaOH, and 42.5 mL Milli-Q water (see Note 10). 7. Citrus Clearing Solvent (or xylene). 8. 70%, 95%, and 100% ethanol. Prepare 70% and 95% ethanol by diluting 100% ethanol with Milli-Q water. 9. Hematoxylin and eosin stains. 10. 0.5% acid alcohol. 11. Scott water. 12. ClearMount mounting medium. 13. Prepare 3% hydrogen peroxide solution by diluting 30% hydrogen peroxide solution with Milli-Q water. 14. Antigen unmasking solution, citrate acid based. 15. Normal goat serum. 16. Prepare 1 PBS from 10 PBS solution. 17. Prepare 0.1% and 0.5% PBST by adding 1 and 5 mL of Triton X-100 to 999 and 995 mL 1 PBS, respectively. Mix well the solution prior to use. 18. Primary antibodies (refer to Table 2).

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Table 2 Antibodies used for immunofluorescent staining Antigen

Supplier

Species Dilution Remarks

AR

Sigma #A9853

Rabbit

CK5

BioLegend #905901

Chicken 1:500

Basal marker (formerly Covance #SIG-3475)

CK5

BioLegend #905501

Rabbit

1:500

Basal marker (formerly Covance #PRB-160P)

CK8

Abcam #ab14053

Chicken 1:500

Luminal marker (discontinued)

CK8

Abcam #ab53280

Rabbit

1:200

Luminal marker

CK8/18 Developmental Studies Hybridoma Rat Bank #TROMA-1

1:100

Luminal marker

CK18

Abcam #ab668

Mouse

1:100

Luminal marker

FoxA1

Abcam #ab55178

Mouse

1:100

Epithelial lineage marker

GFP

Abcam #ab13970

Chicken 1:1000

GFP

Abcam #ab290

Rabbit

1:1000

GFP

Roche #11814460001

Mouse

1:100

p63

Santa Cruz #sc-8343

Rabbit

1:50

phospho- Cell Signaling #3787 Akt

Rabbit

1:100

phospho- Cell Signaling #4370 Erk

Rabbit

1:200

Ki67

Rat

1:1000 Proliferation marker

eBiosciences #14–5698

1:1000 With tyramide amplification (less background staining)

Use 1:500 with tyramide amplification

19. Alexa-fluor-conjugated secondary antibodies. 20. Tyramide signal amplification kit. 21. 0.5 mg/mL DAPI solution. 22. Antifade mounting medium. 2.8 Growth and Gene Expression Assessment of Organoids

1. CellTiter-Glo 3D Cell Viability Assay (Promega #G9681). 2. TRIzol reagent. 3. MagMAX 96 for Microarray Total RNA Isolation Kit (Ambion, Life Technologies #AM1839). 4. Superscript First-Strand Synthesis System for RT-PCR (Invitrogen #11904-018). 5. SYBR green master mix reagent. 6. Primers (refer to Table 3 for primer sequences).

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Table 3 Primers used for quantitative real-time PCR Amplicon Allele size Forward

Reverse

Fkbp5 172 bp

5 -TGA GGG CAC CAG TAA CAA 50 -CAA CAT CCC TTT GTA GTG GAC TGG-30 AT-30

Mme

198 bp

50 -CTC TCT GTG CTT GTC TTG 50 -GAC GTT GCG TTT CAA CCA GC-30 CTC-30

Psca

103 bp

50 -GGA CCA GCA CAG TTG CTT 50 -GTA GTT CTC CGA GTC ATC CTC TAC-30 A-30

Igfbp3 101 bp

50 -CCA GGA AAC ATC AGT GAG 50 -GGA TGG AAC TTG GAA TCG GTC TCC-30 A-30

3

0

Methods The volume of reagents detailed below is for dissociation of an intact prostate from an 8- to 12-week-old C57BL/6 wild-type mouse. For larger prostate specimens, such as those from GEM models of prostate cancer, adjust the volume of reagents proportionally (see Note 11).

3.1 Prostate Dissection, Dissociation, and Preparation of Single Cell Suspension

1. In tissue culture hood, prepare 1 collagenase/hyaluronidase solution by combining 200 μL 10 solution and 1.8 mL DMEM/F12 supplemented with 5% FBS. Warm solution in 37  C water batch for at least 30 min prior to use. 2. Harvest whole urogenital system and transfer to sterile petri dish containing cold PBS for prostate dissection. Using a dissecting microscope, fine forceps, and tweezers, remove residual fats, connective tissues, and unrelated organs such as the bladder, ureters, seminal vesicles, and urethra (see Note 12). 3. Fill 1.5 mL Eppendorf tube with 0.75 mL pre-warmed 1 collagenase/hyaluronidase solution and transfer prostate tissue into the tube. Using small and sharp sterile scissors, lacerate prostate tissue by rapidly opening and closing scissors inside the tube until no visible tissue chunks are seen. Fill up the tube by adding 0.75 mL pre-warmed 1 collagenase/hyaluronidase. Incubate in 37  C CO2 incubator for 3 h with periodic shaking of tube once every hour (see Note 13). 4. Spin down digested tissue at 350  g for 5 min in pre-cooled centrifuge and discard supernatant. Resuspend pellet in 1.5 mL cold 0.25% trypsin-EDTA and incubate in 4  C refrigerator for 1 h (see Note 14).

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5. During trypsin-EDTA incubation, warm 900 μL dispase in 37  C water bath for at least 10–15 min prior to use. At the same time, thaw DNase I solution by leaving the tube carrying the solution on ice. Immediately before use, warm 100 μL DNase I solution briefly prior to combining with dispase solution. 6. After trypsinization step, transfer digested tissue in trypsinEDTA into 15 mL Falcon tube containing 3 mL cold HBSS supplemented with 2% FBS (equal to 2 volume of trypsinEDTA) to quench trypsin reaction. Centrifuge at 350  g for 5 min and discard supernatant. 7. Resuspend pellet with 1 mL pre-warmed dispase/DNase I solution. Pipette vigorously for 1–2 min using P1000 pipette to dissociate cells from extracellular matrices (see Note 15). 8. Add 5 mL cold HBSS supplemented with 2% FBS (equal to 5 volume of dispase 5 U/mL) to dilute dispase to 200 μm in diameter. To passage the organoids, transfer them into 1.5 mL Eppendorf tube, centrifuge at 350  g for 5 min, and discard supernatant. Wash cell pellet in cold PBS and then perform another round of centrifugation at 350  g for 5 min to spin down the cells. Repeat washing step until Matrigel is no longer seen on top of pelleted cells (see Note 23). 2. After removal of supernatant, add 1 mL warm 0.25% trypsinEDTA to tube and incubate in a 37  C water bath for 5–10 min. During the incubation period, pipette up and down for 30 s occasionally with P200 pipette tip to facilitate cell dissociation. In the case of very large organoids or organoids that are difficult to dissociate, separate dissociated cells from large organoids by filtering cell suspension with a 70 μm cell strainer. Quench trypsin reaction on dissociated cells with addition of HBSS supplemented with 2% FBS (2 volume of trypsin-EDTA) while continuing trypsinization on large organoids (see Note 24). 3. For single cell dissociation, add dispase/DNase I, pipette up and down vigorously for 2 min, and filter cell suspension with 40 μm cell strainer. 4. Upon the completion of trypsinization step (with or without dispase/DNase I step), transfer cell suspension into a 15 mL Falcon tube prefilled with 2 mL cold HBSS supplemented with 2% FBS. Spin down the cells by centrifugation at 350  g for 5 min and discard supernatant. Resuspend cell pellet in fresh media and plate onto a new 96-well low attachment plate (see Note 25). 5. Freeze organoids at any point during a passage cycle by pooling 4–6 wells of organoids, centrifuging at 350  g for 5 min, and resuspending in 1 mL freezing media containing 80% FBS, 10% basal hepatocyte culture media, and 10% DMSO in 1.8 mL cryotubes. Use an insulated cryofreezing container to gradually

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freeze the cells to 80  C. Transfer frozen stocks to liquid nitrogen tank for long-term storage. 6. To thaw frozen organoid stock, warm the cryotube rapidly in a 37  C water bath until only a small piece of ice is seen in the tube. Immediately transfer 1 mL of organoids containing freezing media to a 15 mL Falcon tube prefilled with 10 mL cold HBSS supplemented with 2% FBS to dilute DMSO. Spin down thawed organoids at 350  g for 5 min, resuspend in organoid culture media, and plate onto a new 96-well low attachment plate (see Note 26). 3.5 Characterization and Analysis of Organoids 3.5.1 Histology and Immunostaining Organoid Specimens Processing and Sectioning

1. To perform histopathological evaluation and molecular characterization, fix organoids with either 10% neutral buffered formalin (for paraffin embedding) or cold 4% PFA (for OCT embedding) at room temperature (RT) for 1–4 h depending on organoid size. Wash the fixed organoids with three changes of cold PBS by repeatedly centrifuging at 250  g for 5 min, discarding supernatant, and resuspending in fresh PBS. 2. For paraffin embedding, after last washing step, mix spun down organoids with 20–50 μL cold 9:1 collagen I/setting solution mixture. Prepare the collagen/setting solution mixture on ice at all time. When resuspending organoids with collagen solution, use a P200 pipette tip that is cut at the tip region for wider opening to allow homogeneous mixing of collagen and organoids. Avoid generation of bubbles during pipetting and place the organoid and collagen solution mixture on parafilm as a button and leave on a petri dish. Allow the collagen button to solidify in 37  C incubator for 30 min (see Note 27). 3. Slowly transfer the solidified collagen button into 1.5 mL Eppendorf tube prefilled with 10% neutral buffered formalin and fix overnight at RT. On the following day, wash the fixed button with three changes of PBS and then leave the sample in a tissue processor for dehydration, clearing, and infiltration of paraffin. Embed the processed collagen button by pressing and leaving the wider surface of the button on a stainless steel base mold before filling up with paraffin and placing on a cold platform. 4. For OCT embedding, after washing and spinning down the fixed organoids, resuspend them in cold 30% sucrose solution. Incubate at 4  C for overnight or until all organoids sink to the bottom of tube. Remove sucrose solution and transfer organoids in residual sucrose solution to cryomold for biopsy specimen prefilled with a thin layer of OCT compound. Use forceps to mix organoid with OCT compound and then place them close to the center base of cryomold using a dissecting microscope. Once organoids are positioned correctly, fill up

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cryomold with additional OCT compound and place on dry ice to flash freeze the sample (see Note 28). 5. Cut paraffin- or OCT-embedded organoids at a thickness of 4 μm per section. Use freshly cut paraffin sections for best immunostaining result. When cutting OCT blocks, perform serial sectioning until organoids are no longer seen on the sections by checking through a histology microscope. Leave OCT sections at RT for 30 min before storing at 20 or 80  C freezer or proceeding with immunostaining (see Note 29). Hematoxylin and Eosin (H&E) and Immunofluorescent (IF) Stainings

1. Use paraffin sections for H&E staining. Perform paraffin clearing and rehydration of slides by going through two changes of Citrus Clearing Solvent step (9 min each), two changes of 100% ethanol (7 min each), two changes of 95% ethanol (3 min each), and one change of 70% ethanol (1 min) before transferring to Milli-Q water for 5 min. From this point onward, avoid drying out sections at all time. 2. Immerse the slides in hematoxylin solution for 2 min and 30 s, transfer to running tap water for 1 min to wash out excessive hematoxylin, and then immerse into 0.5% acid alcohol for 30 s for differentiation. Put the slide in running tap water again for 1 min prior to a bluing step using Scott water for 1 min. Immerse the slides again in tap water for 2 min before putting into 95% alcohol for 1 min and then eosin solution for 3 min. Prior to mounting, dehydrate the slide by going through four changes of absolute alcohol for 30 s, 1 min, 2 min, and 3 min, respectively (see Note 30). Refer to Figs. 2c, d, 3c, d, and 5b, k–o for representative H&E-stained images for organoids derived from lineagemarked CARNs, EpCAM+, and/or E-Cad+ prostate epithelial cells and GEM model-derived prostate cancer cells. 3. Use paraffin or OCT section for IF staining. When using paraffin sections, perform clearing and rehydration steps as described in step 1. For OCT sections, bring out the sections from freezer, leave at RT for 30 min to 1 h, and then wash with two changes of Milli-Q water for 5 min each. From this point onward, avoid drying out sections at all time. Block endogenous peroxidase activity of organoids with 3% hydrogen peroxide solution for 20 min and then wash in Milli-Q water for 5 min (see Note 31). 4. Perform antigen retrieval by immersing slides into citrate acidbased antigen unmasking solution and leave in a food steamer for 45 min. Upon the completion of antigen retrieval step, cool down slides by leaving in refrigerator or cold room for 15 min and then wash with PBS for 5 min. Subsequently, perform permeabilization step by incubating paraffin sections with 0.5% PBST for 15 min or OCT sections with 0.1% PBST for

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10 min and followed by another washing step with PBS for 5 min. 5. Circle sections with DAKO pen to confine reagents used in the following steps. Incubate in 10% normal goat serum in PBS (or serum of species, in which secondary antibody is raised) at RT for 1 h. Prepare primary antibody of choice (up to three primary antibodies of different species) in 5% normal goat serum in PBS. Apply antibody (or mixture of antibodies) to sections and incubate at 4  C for overnight in a humidified chamber (refer to Table 2 for information of primary antibodies). 6. Wash sections with one change of 0.1% PBST and two changes of PBS for 5 min in each change. Prepare fluoresceinconjugated secondary antibody that recognizes primary antibody used (up to three secondary antibodies conjugated with different fluorochromes) in 5% normal goat serum in PBS with addition of 1:500 0.5 mg/mL DAPI solution. Apply antibody (or mixture of antibodies) to sections and incubate at RT for 1 h (see Note 32). 7. In the case of tyramide signal amplification, prepare HRP-conjugated secondary antibody alone or together with other fluorochrome-conjugated secondary antibodies in 5% normal goat serum in PBS with addition of 1:500 0.5 mg/ mL DAPI solution. Apply antibody (or mixture of antibodies) to sections and incubate at RT for 1 h. 8. Wash sections with one change of 0.1% PBST and two changes of PBS for 5 min in each change (in the case of tyramide signal amplification, refer to step 9). Mount sections with antifade mounting medium and then cover with coverslip. Press on the coverslip to remove excessive mounting medium and bubbles, and seal coverslip with nail polish. 9. Prepare tyramide amplification solution as suggested by the manufacturer’s protocol. Apply tyramide amplification solution at RT for 6 min (see Note 33). Wash sections with 0.1% PBST/ PBS (5 min, 0.1% PBST—one change and followed by PBS— two changes). Repeat step 8. 10. Store slides in dark at 4  C. For consistent result, visualize stainings and take images within 1–2 weeks using a confocal microscope. Refer to Figs. 2e–h, 3e–h, 4b, d, f, 5c–e, and 6c, d for IF stainings of various markers in organoids derived from lineagemarked CARNs, EpCAM+, and/or E-Cad+ prostate epithelial cells, lineage-marked basal and luminal cells, GEM model-derived prostate cancer cells, as well as prostate epithelial organoids grown in the presence or absence of DHT.

Fig. 2 Assessment of organoids derived from CARNs. (a, b) Bright-field (a) and epifluorescent (b) views of CARN-derived organoids that are either partially filled or with hollow lumen (arrow). (c, d) Representative H&Estained CARN-derived organoids at low (c) and high (d) magnification views. (e–h) Organoid derived from lineage-marked CARNs demonstrates uniformed YFP expression and high cellular proliferation as shown by Ki67 immunostaining (arrows, e) and correct localization of basal cells at the outer region as marked by CK5 (arrowheads) and the luminal cells at the internal region as marked by CK8 (f). The organoid also expresses AR (arrows, g) and epithelial lineage marker, FoxA1 (h). Scale bars in a–c corresponding to 100 μm and in d–h to 50 μm. Figure is adapted from Fig. 1c–j in ref. 8

Fig. 3 Characterization of organoids derived from EpCAM+ and/or E-Cad+ prostate epithelium. (a, b) Low (a) and high (b) magnification views of organoids derived from EpCAM+ and/or E-Cad+ sorted prostate epithelial cells at 20 days of plating that demonstrate heterogeneous phenotype. (c, d) Representative H&E-stained EpCAM+ and/or E-Cad+ sorted prostate epithelial cell-derived organoids. (e–h) Organoid derived from prostate epithelial cells expresses proliferation marker, Ki67 (arrows, e), demonstrates localization of basal cells at the outer layer as marked by p63 (arrowheads) and the luminal cells at the internal layers as marked by CK18 (f). The organoid also contains cells that co-expresses AR and CK8 (arrows, g) as well as epithelial lineage marker, FoxA1 (arrows, h). Scale bars in a–c corresponding to 100 μm and in d–h to 50 μm. Figure is adapted from Fig. 2b–f and h–j in ref. 8

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Fig. 4 Assessment of organoid-forming ability of lineage-marked basal or luminal prostate epithelial cells. (a) Organoid derived from YFP-labeled prostate basal epithelial cells. (b) CK5-trace organoid contains mostly CK5-positive cells, including internal cells (arrowheads). (c) Organoid derived from YFP-labeled prostate luminal epithelial cells. (d) CK8-trace organoid contains some CK5-positive cells at the outer layer (arrowheads). (e) Comparison of organoid formation efficiency between CK5-trace (n ¼ 4 experiments), CK8-trace (n ¼ 3 experiments), and CK18-trace (n ¼ 2 experiments) prostate epithelial cells. (f) Organoid derived from mixing of red CK18-trace luminal cells and green CK5-trace basal cells demonstrates green cells at the outer layer (arrowheads), consistent with the localization of basal cells. Scale bars in a–d and f corresponding to 50 μm. Figure is adapted from Fig. 3c–e, g, h, and n in ref. 8

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Fig. 5 Generation of tumor organoids from GEM models of prostate cancer. (a–e) Organoids derived from transformed CARNs isolated from castrated and tamoxifen-induced Nkx3.1CreERT2/+; Pten flox/flox; KrasLSL-G12D/ + ; R26R-YFP/+ (NPK) mice. NPK-CARN tumor organoid is YFP positive with extensive budding (a) and mostly filled morphology as evidenced by H&E staining (b), highly proliferative as marked by Ki67 (c) and expresses pAKT (d) and patchy pERK (arrows, e), consistent with the Pten deletion and Kras-G12D activation phenotype. (f–o) Bright-field (f–j) and H&E-stained sections of tumor organoids derived from various GEM models of prostate cancer through FACS sorting of EpCAM+ and/or E-Cad+ population. Showings are organoids generated from 22-week-old TRAMP mice (f, k), tamoxifen-induced Nkx3.1CreERT2/+; Pten flox/flox; p53 flox/flox (NPP53) mice at 2 months old and assayed after 8 months (g, l), 14-month-old Nkx3.1/ mice (h, m), 9-month-old Hi-Myc transgenic mice (i, n) and 10-month-old Nkx3.1+/; Pten+/ mice (j, o). Scale bars in a, b, and f–o corresponding to 100 μm and in c–e to 50 μm. Figure is adapted from Figs. 4b–f and 5a–j in ref. 8

3.5.2 Growth and Gene Expression Assessment (See Note 34) CellTiter-Glo Assay for Growth Assessment

1. To assess cell growth in response to androgen withdrawal, passage prostate epithelial cell-derived organoids and plate equal amount of organoids onto 96-well low attachment plate in the presence or absence of 100 nM DHT (see Note 35). 2. Assay cell viability at days 1, 3, and 5 after plating using CellTiter-Glo 3D (Promega) with five technical replicates for each time point. Briefly, thaw CellTiter-Glo 3D reagent at 4  C and leave at RT for at least 15 min prior to use. Add 100 μL of the reagent into each well, which contains approximately 100 μL culture medium. 3. Shake the 96-well low attachment plate vigorously at RT for 10 min and then transfer the mixture to an opaque-walled 96-well plate. Incubate at RT for 10 min prior to measurement using a luminometer plate reader.

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Quantitative Real-Time PCR Analysis

1. For RNA extraction, transfer four to six wells of organoids to a 1.5 mL Eppendorf tube, centrifuge at 350  g for 5 min, and discard supernatant. Dissolve pelleted organoids in TRIzol reagent followed by processing using the MagMAX 96 for Microarray Total RNA Isolation Kit as suggested by manufacturer’s recommendation. 2. Use 100–200 ng of RNA as template for cDNA synthesis using the Superscript First-Strand Synthesis System according to manufacturer’s recommendation. Perform quantitative realtime PCR by mixing cDNA, primers (refer to Table 3 for primer sequences used) and SYBR green master mix reagent in the Eppendorf Realplex2 instrument in a triplicate manner for each culture condition. Use the ΔΔCT method to obtain expression values and normalize values to GAPDH expression. Repeat the experiment with additional biological replicates (see Note 36). Refer to Fig. 6e for difference of various androgen-responsive genes between prostate epithelial organoids grown in the presence or absence of DHT.

Fig. 6 Androgen responsiveness of organoid derived from EpCAM+ and/or E-Cad+ prostate epithelial cells. (a, b) Organoids were passaged and plated in the presence (a) or absence of (b) DHT. (c, d) In the presence of DHT, organoids demonstrate strong nuclear AR expression, (c) whereas in the absence of DHT, AR expression of the organoids is weak and localized at the cytoplasm (d). (e) Quantitative real-time PCR analysis of androgen-responsive genes, Fkbp5, Mme, Psca, and lgfbp3, in organoids cultured in the presence or absence of DHT. Scale bars in a and b corresponding to 100 μm and in c and d to 50 μm. Figure is adapted from Fig. 2l–p in ref. 8

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Notes 1. Make sure tamoxifen is fully dissolved in corn oil before filtration by repeatedly warming mixture of tamoxifen powder and corn oil in 37  C water bath for 10 min and then vortexing vigorously for 15 min. Filtered tamoxifen solution should be stored at 4  C and used within a month of preparation. 2. We use identical tamoxifen treatment schedule, route, and dose for different inducible Cre mice. 3. Different GEM models of prostate cancer require different durations and genetic backgrounds to progress into hyperplasia and/or cancer. 4. Aliquot and keep the 5 mM stock solution at 20  C until use. Avoid repeated freeze and thaw cycles for optimal results. We recommend using ROCK inhibitor Y-27632 from STEMCELL Technologies for consistent organoid formation. 5. After DAPI is fully dissolved in Milli-Q water, filter, aliquot, and keep the stock solution at 20  C. Once thawed, the working stock solution should be in good condition for at least 3 months if it is kept at 4  C and without extensive light exposure. 6. This reagent has been regularly on back order. Make sure sufficient reagent is available prior to the start of a new experiment. 7. The use of heat-inactivated charcoal-stripped FBS is crucial for efficient organoid formation from prostate luminal cells. After the completion of heat inactivation, aliquot and keep the reagent at 80  C prior to use. 8. Thaw Matrigel by placing the reagent on ice in a 4  C refrigerator or cold room for overnight. Once thawed, Matrigel must remain on ice at all time to prevent polymerization. For consistent results, we avoid using Matrigel that has undergone more than two freeze-thaw cycles. 9. Keep DHT solution at 20  C at all time. Avoid leaving working solution outside the freezer for prolonged period when preparing culture medium. 10. Make sure there is no precipitation when using setting solution to ensure proper collagen I neutralization and solidification. 11. For regressed prostate, we use identical volume of reagents that are used for intact mouse prostate dissociation because prostate epithelium in castrated mice is high in cell density and more collagenous. 12. For detailed description of urogenital system harvest and prostate dissection procedure, refer to Lukacs et al. [11].

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13. For optimal dissociation, place Eppendorf tube on its side on a sterile petri dish during incubation to maximize surface area of prostate tissue exposed to collagenase/hyaluronidase solution. In addition, periodic shaking of tube helps redistributing tissues. 14. Place a petri dish on ice and leave tube on the petri dish. Subsequently, place the whole ice bucket into the refrigerator or cold room. Avoid use of warm trypsin-EDTA as this will cause over-trypsinization, consequently leading to extensive cell death. 15. Vigorous pipetting is crucial for optimal cell dissociation. Pipette sample until the solution is translucent without visible tissue fragments. Do not exceed 2 min for this step as this will cause extensive cell death. 16. Using this protocol, we should expect 1.5–4 million of cells isolated from intact prostate of an 8- to 12-week-old C57BL/6 mouse. 17. We use DAPI to exclude dead cells because they can be mistakenly identified as YFP-positive cells during flow sorting. However, high concentration of DAPI staining is lethal to cells. Avoid adding >0.1 μg/mL DAPI in resuspension media. 18. Use sheath pressure not exceeding 12 psi for cell sorting as higher sheath pressure will cause luminal cell death. It is also important to include media supplemented with ROCK inhibitor Y-27632 in the collecting tube to prevent cell loss and death. 19. Basal organoid culture media should be finished within a month of preparation to ensure consistent results. 20. Avoid extensive warming in 37  C water bath because this may cause Matrigel to solidify on top of the media. 21. Use P1000 pipette tip to transfer organoids as smaller tips may damage structure of organoids. If there are too many wells for media change, pool multiple wells prior to centrifugation and redistribute evenly when plating. 22. Organoid growth should be evidenced as early as days 2–3 (CARNs or prostate cancer cells) or days 4–5 (prostate epithelial cells) of plating. Larger size organoids will occasionally fuse together. Therefore, quantification of organoid formation should be done by days 7 of plating for accurate estimation of organoid-forming efficiency. 23. Trypsinization step will not be optimal with residual Matrigel, consequently leading to incomplete cell dissociation. 24. Prostate epithelial cell-derived organoids that are solid and do not have obvious lumen are harder to dissociate. These

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organoids contain more basal cells and require longer trypsinization duration during dissociation. In addition, prolonged trypsin-EDTA incubation of more easily dissociated cells will cause extensive death in this population. Therefore, adoption of different trypsinization time for different cell lineages can ensure continuous propagation of different lineage populations in organoid culture. 25. Passage organoids in a ratio of 1:4–1:6, i.e., dissociated cells from 1 well can be passaged into 4–6 wells. 26. During thawing, do not over-warm the freezing media as warm DMSO is lethal to the cells. 27. After neutralization by setting solution, collagen solution will change color from yellow to pink or red once it is solidified. 28. Make sure organoids are mixed well with OCT compound and embedded into it to provide proper support during tissue sectioning. 29. Avoid over-trimming when sectioning and always check sections with a histology microscope to confirm the presence of organoids on sections. Serial sectioning of OCT blocks can minimize tissue loss as each block may only carry 20–30 4 μm sections with intact organoid structure. 30. Check quickly the slides using a histology microscope after bluing step to see whether hematoxylin staining provides great nuclear details as well as after eosin step to see whether the counterstaining gives good nuclear/cytoplasmic contrast. Avoid using aged hematoxylin and eosin stains for optimal staining results. 31. Exclude this step if HRP-enzymatic reaction is not involved in the subsequent staining procedures. 32. When using two to three fluorochrome-conjugated secondary antibodies, avoid selecting combination of fluorochromes that have overlapped emission spectrums. Use Alexa-fluor 488 and 555 for combination of two antibodies and Alexa-fluor 488, 555, and 647 for combination of three antibodies. 33. Do not exceed 6-min incubation time as longer incubation will lead to high staining background. 34. The protocols are also applicable for drug assessment on prostate luminal progenitors and cancer cells. 35. Instead of supplementing 5% Matrigel in culture medium, use hepatocyte medium supplemented with 2% Matrigel for this assay. With prolonged culture period, higher concentration of Matrigel can affect accuracy of luminescence reading. 36. Dilute cDNA samples 1:5–1:10 when necessary.

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Acknowledgments We thank M. Shibata, S. Irshad, H.H. Zhu, M. Shen, W.Q. Gao, and members of the Chua lab (M.Y. Liu, Y. Zhang, X. Cai, and C.P. Liang) for insightful comments on the manuscript. This work was supported by grants from the National Natural Science Foundation of China (81672548 and 81874098 to C.W. Chua), the Program for Professor of Special Appointment (Eastern Scholar), the Shanghai Institutions of Higher Learning (2016012 to C.W. Chua), the State Key Laboratory of Oncogenes and Related Genes (90-17-01 to C.W. Chua), and the Shanghai Municipal Education Commission-Gaofeng Clinical Medicine Grant Support (20171917 to C.W. Chua). Author Contributions: Together with M. Lei, C.W. Chua developed and optimized the organoid culture protocol in M. Shen’s lab at the Columbia University. C.W. Chua and Y. Shu wrote the book chapter manuscript, and prepared the tables as well as the figures that were adapted from original figures in Chua et al. [8]. References 1. Debnath J, Brugge JS (2005) Modelling glandular epithelial cancers in three-dimensional cultures. Nat Rev Cancer 5:675–688 2. Clevers H (2016) Modeling development and disease with organoids. Cell 165:1586–1597 3. Fatehullah A, Tan SH, Barker N (2016) Organoids as an in vitro model of human development and disease. Nat Cell Biol 18:246–254 4. Shen MM, Abate-Shen C (2010) Molecular genetics of prostate cancer: new prospects for old challenges. Genes Dev 24:1967–2000 5. Peehl DM (2005) Primary cell cultures as models of prostate cancer development. Endocr Relat Cancer 12:19–47 6. Irshad S, Abate-Shen C (2013) Modeling prostate cancer in mice: something old, something new, something premalignant, something metastatic. Cancer Metastasis Rev 32:109–122 7. Wang X, Kruithof-de Julio M, Economides KD, Walker D, Yu H et al (2009) A luminal

epithelial stem cell that is a cell of origin for prostate cancer. Nature 461:495–500 8. Chua CW, Shibata M, Lei M, Toivanen R, Barlow LJ et al (2014) Single luminal epithelial progenitors can generate prostate organoids in culture. Nat Cell Biol 16:951–961 9. Chua CW, Shibata M, Lei M, Toivanen R, Barlow LJ, Shen MM (2014) Culture of mouse prostate organoids. Protoc Exch. https://doi. org/10.1038/protex.2014.037 10. Karthaus WR, Iaquinta PJ, Drost J, Gracanin A, van Boxtel R et al (2014) Identification of multipotent luminal progenitor cells in human prostate organoid cultures. Cell 159:163–175 11. Lukacs RU, Goldstein AS, Lawson DA, Cheng D, Witte ON (2010) Isolation, cultivation and characterization of adult murine prostate stem cells. Nat Protoc 5:702–713

Chapter 18 Isolation, Purification, and Culture of Mouse Pancreatic Islets of Langerhans Youakim Saliba and Nassim Fare`s Abstract Pancreatic islets constitute an important tool for research and clinical applications in the field of diabetes. They are used for transplantation, unraveling new mechanisms in insulin secretion, studying pathophysiological pathways in diseased cells, and pharmacological research aimed at developing improved therapeutic strategies. Therefore, fine-tuning islet isolation protocols remains an important objective for reliable investigations. Here we describe a relatively simple mouse islet isolation protocol that relies on enzymatic digestion using low-activity collagenase and several sedimentation and Percoll gradient steps. Key words Islets of Langerhans, Pancreas, Isolation protocol, Mouse

1

Introduction Islets of Langerhans constitute an important experimental tool in the field of diabetic research, and achieving successful cell isolation and culture is the most important requisite for reliable studies. Since Lacy’s group described a new enzyme-based method for islet isolation in 1967 [1], different protocols from rodents to humans have been proposed and fuelled the advances in islet research [2–16]. The main difference between all these procedures resides in the enzyme blends and their delivery route, with the common bile duct as the most commonly used [10, 13, 17, 18]. Islet separation and purification are then performed by Percoll [19, 20] or Ficoll gradient separation [15], filtration [21], or magnetic retraction [4, 22] and handpicking under a microscope [8, 10, 12, 15, 18, 23–25]. However, many hurdles still persist in these isolation procedures rendering it difficult to achieve reproducible maximum yield of viable islets that retain their in vivo characteristics. Besides, the fine details to perform the isolation that usually requires delicate microsurgery maneuvers are often lacking, leading to sometimes conflicting data when it comes to yield and function [26, 27]. Therefore, refining and revisiting islet

Ivan Bertoncello (ed.), Mouse Cell Culture: Methods and Protocols, Methods in Molecular Biology, vol. 1940, https://doi.org/10.1007/978-1-4939-9086-3_18, © Springer Science+Business Media, LLC, part of Springer Nature 2019

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isolation protocols remain important for more and more reliable research. In the protocol described here, mouse pancreas was excised and digested with a low-activity collagenase eliminating the need for microsurgery expertise. Since Percoll is chemically inert with minimum cellular toxic effects as compared to other cell separation gradient media [9, 21, 28], it was used here to purify the islets from the isolated pancreatic tissue. This purification step was preceded by several sedimentations and repeated to further purify the islets without time-consuming handpicking [5, 10, 13]. Islets were cultured in RPMI 1640 medium that has been shown to be optimal for keeping islets into culture [2]. Finally, variations in the islet numbers have been shown to exist between inbred mice [29], and age differences have been also noted [30–34]. In the current protocol, we isolated islets from male and female mice of different ages and species and obtained similar yields ranging from 750 to 1200 healthy islets per pancreas.

2

Materials All chemicals used in the protocol are of reagent grade (>99%). Deionized water with a resistivity of 18.2 mΩ cm at 25
C was used.

2.1 Modified Tyrode Solution

1. Add approximately 500 mL water in 1 L autoclaved glass beaker. Weigh each of the following and add consecutively to the beaker while stirring with a magnetic stir bar: 8.18 g NaCl, 0.425 g KCl, 0.346 g MgCl2·6H2O, 0.37 g NaHCO3, 0.204 g KH2PO4, 2.383 g HEPES, 1.492 g creatine monohydrate, 2.5 g taurine, and 2.11 g D-glucose. Adjust the pH to 7.2 with 1 M NaOH, and then make up the solution to 1 L. This solution will have the following composition: 140 mM NaCl, 5.7 mM KCl, 1.7 mM MgCl2, 4.4 mM NaHCO3, 1.5 mM KH2PO4, 10 mM HEPES, 10 mM creatine monohydrate, 20 mM taurine, and 11.7 mM D-glucose. 2. Filter the Tyrode solution on a sterile glass filter beaker using 0.22 μm membrane filter (Millipore, Merck, Massachusetts, USA) under a laminar flow hood and store it at 4
C (see Note 1).

2.2 Collagenase Solution

1. Prepare 10 mL of collagenase solution per mouse pancreas. Weigh in a 15 mL conical tube using the following: 10 mg collagenase A (activity >0.15 U/mg) (Roche Diagnostics, Basel, Switzerland) kept at 20
C and 10 mg bovine serum albumin (BSA) kept at 4
C (see Note 2).

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2. Under a laminar flow hood, filter the enzyme solution using 0.22 μm membrane syringe filter, transfer it in a new sterile 15 mL conical tube, and keep it on ice until used (see Note 3). 2.3

Percoll Solution

1. Percoll is a colloidal PVP-coated silica for cell separation with a density of 1.13  0.005 g/mL. 2. Prepare 500 mL of a 10 concentrated NaCl solution (1.5 M) by diluting 43.83 g NaCl in water. Prepare 500 mL of a 1 concentrated NaCl solution (0.15 M) by either diluting the 10 concentrated NaCl or by formulating 1 concentrated NaCl solution directly from powder. 3. Prepare a stock isotonic Percoll (SIP) solution by adding 9 parts of Percoll to 1 part of 1.5 M NaCl (10 concentrated) (see Note 4). In the following calculations, we will use an example of 10 mL SIP solution, that is, 9 mL Percoll and 1 mL 1.5 M NaCl. 4. Calculate the density of the SIP solution from the following formula: V X ¼ V 0  ðρ0  ρi Þ=ðρi  ρ10 Þ so ρi ¼ ½ðV 0  ρ0 Þ þ ðV X  ρ10 Þ=ðV X þ V 0 Þ where VX is the volume of diluting medium (mL) V0 is the volume of undiluted Percoll (mL) ρ0 is the density of Percoll (1.13  0.005 g/mL) ρ10 is the density of 1.5 M NaCl (1.058 g/mL) ρi is the density of the SIP solution (g/mL) The density of SIP obtained is 1.123 g/mL 5. Dilute the SIP solution to a final density of 1.045 g/mL (see Note 5) by adding 0.15 M NaCl using the following formula:   V y ¼ V i  ðρi  ρÞ= ρ  ρy where Vy is the volume of diluting 0.15 M NaCl (mL) Vi is the volume of SIP (mL) ρi is the density of SIP (g/mL) ρy is the density of 0.15 M NaCl (1.0046 g/mL) ρ is the density of final diluted Percoll solution (g/mL) In order to dilute all the SIP solution (10 mL) to a final density of 1.045 g/mL, proceed by adding the following: Vy ¼ 10  (1.123–1.045)/(1.045–1.0046) ¼ 19.3 mL of 0.15 M NaCl (see Note 6).

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2.4

Youakim Saliba and Nassim Fare`s

Mouse Sacrifice

Male and female adult and aged C57BL/6 mice (3 and 10 months old) as well as male adult BALB/C mice (3 months old) are used in the protocol (Janvier Labs, Le Genest-Saint Isle, France). The animals are kept at a stable temperature (25
C) and humidity (50  5%) and are exposed to a 12:12-h light-dark cycle. They are fed ordinary rodent chow, have free access to tap water, and are acclimatized for at least 1 week under these conditions before the start of the experiments (see Note 7). 1. Anesthetics: Ketamine hydrochloride 50 mg/mL (RotexMedica, Trittau, Germany) and Xyla, xylazine 2% injection (Interchemie, Waalre, Holland) kept at 4
C 2. Sterile syringe for injections (1 mL) 3. Ethanol 100% 4. Betadine solution, 10% povidone iodine 5. Sterilized surgical scissors

2.5 Pancreas Removal and Islet Isolation

1. Three sterile petri dishes (60 mm) kept on ice 2. Serological pipettes (2 mL) 3. Water bath incubator (37
C) 4. One sterilized Erlenmeyer flask (50 mL) 5. Eight sterile conical tubes (4 of 15 mL and 4 of 50 mL) 6. Pure oxygen canister

2.6 Cell Viability Tests

1. Trypan blue stock solution (0.4%), prepared in 0.81% sodium chloride and 0.06% potassium phosphate, dibasic 2. Propidium iodide stock solution (1 mg/mL in water)

2.7

Islet Culture

1. RPMI 1640 medium (Lonza, Basel, Switzerland) supplemented with 2 mM L-glutamine. 2. Fetal bovine serum. 3. Prepare a penicillin/streptomycin stock solution (100 concentrated). This solution contains 10,000 penicillin units and 10,000 μg streptomycin per mL. For a 50 mL stock volume, weigh 33.33 mg of penicillin G sodium salt (1500 U/mg) and 50 mg of streptomycin sulfate and complete with water to the final volume. The solution is aliquoted and kept at 20
C (see Note 8). 4. Cell culture plates (6, 24, or 96 wells). 5. 5% CO2 incubator at 37
C.

Mouse Islet Isolation

3

259

Methods

3.1 Mouse Sacrifice and Pancreas Removal

1. Anesthetize the animals by an intraperitoneal injection of a mixture of ketamine (75 mg/kg) and xylazine (10 mg/kg). To make sure of adequate depth of anesthesia, pedal withdrawal reflex (footpad pinch, on two feet) is performed (see Note 9). 2. When animals are completely nonresponsive to toe pinching, apply betadine and ethanol on the abdomen in order to keep the conditions as sterile as possible and to reduce the chance of hair contamination in the peritoneal cavity during subsequent steps. 3. Perform a V-shaped abdominal incision starting from the lower abdomen and extending to the lateral parts of the diaphragm in order to expose all organs in the peritoneal cavity. 4. Locate the spleen (dark red) which is about 1 cm below the diaphragm (Fig. 1). The pancreas is surrounded by the stomach, the duodenum and proximal jejunum, and the spleen. In the mouse, the duodenum wraps around the head of the pancreas that is a soft and diffuse organ as compared with the human pancreas. 5. Remove the pancreas from the mouse and place it into a petri dish containing ice cold Tyrode solution. Removal begins by snipping the common bile duct by scissors and gently detaching the pancreas from the intestines. Continue removing the pancreas from the stomach and finally the spleen (see Note 10).

Fig. 1 Mouse pancreas anatomical location. The pancreas is surrounded by the stomach, the duodenum and proximal jejunum, and the spleen. The duodenum wraps around the head of the pancreas that is a soft and diffuse organ as compared with the human pancreas. (1) Pancreas, (2) duodenum, (3) stomach, (4) spleen, (5) liver

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The usual number of islets obtained per pancreas is around 1000; however, if a large number of islets are required for the assays, use three or four mice and repeat the above steps for each animal. 6. Gently cut the pancreas into small pieces (2 mm) in ice cold Tyrode solution. Wash the tissue pieces three times in the same solution by transferring them in consecutive 60 mm petri dishes containing 5 mL Tyrode solution. Repeat this three times or until all blood and fat tissue is removed (fat floats easily in solution). 3.2 Pancreas Digestion and Islet Purification

1. Transfer the pieces into the Erlenmeyer flask and discard the remaining Tyrode solution. Add the collagenase A solution to the Erlenmeyer flask (see Note 11) and incubate in a shaking water bath at 37
C for 1 h at 100 rpm (see Note 12). Supply the solution by pure oxygen (see Note 13). 2. Stop digestion by transferring the cell suspension into a 50 mL tube containing an equal volume of ice cold Tyrode solution to stop the activity of collagenase. Shake the tube vigorously by hand to mechanically dissociate the digested pancreas and liberate the islets (see Note 14). 3. Leave the cell suspension on ice for 3 min in order to sediment the islets. Discard the supernatant and add on another 20 mL ice cold Tyrode solution. Repeat this sedimentation step for three times (see Note 15). 4. Remove as much liquid as possible from the final pellet and then purify the cells on the Percoll solution (see Note 16). Aspirate the pellet with a 1 mL tip and carefully place it on top of the Percoll solution in a 15 mL tube (see Note 17). 5. Allow the islets to sediment for 5 min (see Note 18). Meanwhile, prepare another two 15 mL tubes with Percoll solution. 6. Collect the pellet from the bottom with a 2 mL pipette and place it on top of the second Percoll solution. Repeat this again with the final pellet. 7. Collect the final pellet with a 2 mL pipette and transfer it into a new 15 mL tube. 8. Add 10 mL RPMI 1640 supplemented with 10% fetal bovine serum and 2 mM L-glutamine and let islets sediment for 5 min. 9. Discard the supernatant and add another 10 mL RPMI 1640 medium (as above) on the pellet. 10. Gently resuspend the islets.

3.3 Islet Viability Assessment

Trypan blue exclusion test and propidium iodide are used to assess islet cell viability. Following the pancreas digestion, perform the following:

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261

1. Transfer 500 μL of islet suspension into a well of a 24-well plate to perform trypan blue labeling. 2. Add 500 μL trypan blue stock solution to the previous suspension, and incubate for 2 min at room temperature. 3. Count trypan blue-positive and trypan blue-negative islets under a microscope. Trypan blue-negative islets are regarded as viable ones. Calculate the percentage of viable islets according to the following formula: total viable cells (unstained)/ total cells (stained and unstained)  100. 4. Transfer another 500 μL of islet suspension into a new well to perform propidium iodide labeling. 5. Add appropriate volume of propidium iodide stock solution to reach a final 3 μM concentration in the well, and incubate for 15 min at room temperature. 6. Count fluorescent cells in three view fields under an epifluorescence microscope at 620 nm. Propidium iodide-negative islets are regarded as viable ones. Calculate the percentage of viable islets as previously described for trypan blue (see Note 19). 3.4

Islet Culture

1. Homogenize the cell suspension. 2. Remove 500 μL of the suspension and place in a petri dish to count the islets under a microscope (see Note 20). The total number of islets is then obtained by a simple rule of three calculator. 3. Distribute islets in equal number in 6-, 24-, or 96-well plates as required, in 2 mL, 500 μL, or 100 μL, respectively 4. Incubate in ambient air with 5% CO2 incubator at 37
C till the time of experiment (see Note 21).

4

Notes 1. It is important to check the Tyrode solution for any bacterial or fungi contamination prior to each use. Therefore, it is better to store sterile solutions in smaller volumes and for short-term use. Alternatively, add phenol red to the Tyrode solution prior to filtration, and discard the solution when its color turns yellow at ph