Embryonic Stem Cell Protocols (Methods in Molecular Biology, 2520) 1071624369, 9781071624364

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Methods in Molecular Biology 2520

Kursad Turksen Editors

Embryonic Stem Cell Protocols Fourth Edition

METHODS

IN

MOLECULAR BIOLOGY

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

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

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

Embryonic Stem Cell Protocols Fourth Edition

Edited by

Kursad Turksen Ottawa, ON, Canada

Editor Kursad Turksen Ottawa, ON, Canada

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-2436-4 ISBN 978-1-0716-2437-1 (eBook) https://doi.org/10.1007/978-1-0716-2437-1 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022 This work is subject to copyright. All rights are solely and exclusively licensed 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, expressed 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. Cover Illustration Caption: Artwork created by Kursad Turksen. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface Since their introduction as a model for cell lineage and differentiation studies, embryonic stem cells (ESCs) have become a critical workhorse. Over the years, many early protocols have been improved and extended to become more directed and consequently more informative. In the new edition of this volume, I have collected a series of protocols that are representative of such recent developments and improvements in the ESC field. Once again, the protocols gathered here are faithful to the mission statement of the Methods in Molecular Biology series: They are well-established and described in an easy-tofollow, step-by-step fashion so as to be valuable for not only experts but also novices in the stem cell field. That goal is achieved because of the generosity of the contributors who have carefully described their protocols in this volume, and I am very grateful for their efforts. My thanks as well go to Dr. John Walker, the Editor-in-Chief of the Methods in Molecular Biology series, for giving me the opportunity to create and now update this volume and for supporting me along the way. I am also grateful to Patrick Marton, the Executive Editor of Methods in Molecular Biology and the Springer Protocols collection, for his continuous support from idea to completion of this volume. A special thank you goes to Anna Rakovsky, Assistant Editor for Methods in Molecular Biology, for her support from the beginning to the end of this project. I would also like to thank David C. Casey, Senior Editor of Methods in Molecular Biology, for his outstanding editorial work during the production of this volume. Finally, I would like to thank Sativa Rockey Samuel, Daniel Ignatius Jagadisan, and the rest of the production crew for their work in putting together an outstanding volume. Ottawa, ON, Canada

Kursad Turksen

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

Dissecting Molecular Phenotypes Through FACS-Based Pooled CRISPR Screens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Oriana Genolet, Liat Ravid Lustig, and Edda G. Schulz Feeder-Free Human Embryonic Stem Cell Culture Under Defined Culture Conditions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ¨ l Dog˘an Taha Bartu Hayal and Ays¸egu Rat-Induced Pluripotent Stem Cells-Derived Cardiac Myocytes in a Cell Culture Dish. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fatima Bianca A. Dessouki, Pawan K. Singal, and Dinender K. Singla Transient Induction and Characterization of Mouse Epiblast-Like Cells from Mouse Embryonic Stem Cells. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Federico Pecori and Shoko Nishihara High Content Image Analysis of Spatiotemporal Proliferation and Differentiation Patterns in 3D Embryoid Body Differentiation Model . . . . . . . . . Olga Gordeeva Derivation of Multipotent Neural Progenitors from Human Embryonic Stem Cells for Cell Therapy and Biomedical Applications . . . . . . . . . . . . . . . . . . . . . . . . Loriana Vitillo and Ludovic Vallier Feeder-Dependent/Independent Mouse Embryonic Stem Cell Culture Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hatice Burcu S¸is¸li, Selinay S¸enkal, Derya Sag˘rac¸, ¨ l Dog˘an Taha Bartu Hayal, and Ays¸egu ChIP-qPCR for Polycomb Group Proteins During Neuronal Differentiation of Human Pluripotent Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Divya Desai and Prasad Pethe Directed Differentiation of Human Pluripotent Stem Cells into Inner Ear Organoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yoshitomo Ueda, Stephen T. Moore, and Eri Hashino Glycolytic Profiling of Mouse Embryonic Stem Cells (mESCs). . . . . . . . . . . . . . . . . . . . ˜ o Ramalho-Santos Bibiana Correia, Maria Ineˆs Sousa, and Joa Replating Protocol for Human Induced Pluripotent Stem Cell–Derived Cardiomyocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Arzuhan Koc and Esra Cagavi Hematopoietic Cell Isolation by Antibody-Free Flow Cytometry in the Zebrafish Embryo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Katsuhiro Konno, Jingjing Kobayashi-Sun, Fumio Arai, Isao Kobayashi, and Daisuke Sugiyama

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Human Trophectoderm Spheroid Derived from Human Embryonic Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Wen Huang, Sze Wan Fong, William Shu Biu Yeung, and Yin Lau Lee Embryoid Bodies–Based Multilineage Differentiation of Human Embryonic Stem Cells Grown on Feeder-Free Conditions. . . . . . . . . . . . . . . . . . . . . . . . Luciana Isaja, Sofı´a Luja´n Ferriol-Laffouillere, Sofı´a Mucci, Marı´a Soledad Rodrı´guez-Varela, and Leonardo Romorini A Simple, Rapid, and Cost-Effective Method for Loss-of-Function Assays in Pluripotent Cells. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Helena Covelo-Molares, Yara Souto, Miguel Fidalgo, and Diana Guallar Embryoid Body Formation from Mouse and Human Pluripotent Stem Cells for Transplantation to Study Brain Microenvironment and Cellular Differentiation. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Magdalena Guerra-Crespo, Omar Collazo-Navarrete, Rodrigo Ramos-Acevedo, Carmen Alejandra Morato-Torres, ¨ le and Birgitt Schu A Modified SMART-Seq Method for Single-Cell Transcriptomic Analysis of Embryoid Body Differentiation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jianqun Zheng, Ying Ye, Qiushi Xu, Wei Xu, Wensheng Zhang, and Xi Chen Osteogenic Differentiation from Mouse Embryonic Stem Cells . . . . . . . . . . . . . . . . . . . Zahra Alvandi and Michal Opas Murine Embryonic Stem Cell Culture, Self-Renewal, and Differentiation . . . . . . . . . . Manar Elkenani and Belal A. Mohamed Mouse Embryonic Stem Cell Culture in Serum-Containing or 2i Conditions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Emre Balbasi, Gozde Guven, and Nihal Terzi Cizmecioglu Directed Differentiation of Mouse Embryonic Stem Cells to Mesoderm, Endoderm, and Neuroectoderm Lineages . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Emre Balbasi, Dersu Sezginmert, Ceren Alganatay, and Nihal Terzi Cizmecioglu Accessing the Human Pluripotent Stem Cell Translatome by Polysome Profiling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rubens Gomes-Ju´nior, Patrı´cia Shigunov, Bruno Dallagiovanna, and Isabela Tiemy Pereira Genome Engineering Human ESCs or iPSCs with Cytosine and Adenine Base Editors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Giulia Pavani, Joshua G. Klein, Deborah L. French, and Paul Gadue Proteomic Analysis of Human Neural Stem Cell Differentiation by SWATH-MS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jirina Tyleckova, Jakub Cervenka, Ievgeniia Poliakh, Jaromir Novak, Katerina Vodickova Kepkova, Helena Kupcova Skalnikova, and Petr Vodicka Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors CEREN ALGANATAY • Department of Biological Sciences, Middle East Technical University, Ankara, Turkey ZAHRA ALVANDI • Vascular Biology Program and Department of Surgery, Boston Children’s Hospital, Boston, MA, USA; Department of Surgery, Harvard Medical School, Boston, MA, USA FUMIO ARAI • Department of Stem Cell Biology and Medicine, Graduate School of Medical Sciences, Kyushu University, Fukuoka, Japan EMRE BALBASI • Department of Biological Sciences, Middle East Technical University, Ankara, Turkey ESRA CAGAVI • Regenerative and Restorative Medicine Research Center (REMER), Research Institute for Health Sciences and Technologies (SABITA), Istanbul Medipol University, Istanbul, Turkey; Department of Medical Biology, School of Medicine, Istanbul Medipol University, Istanbul, Turkey; Medical Biology and Genetics Graduate Program, Health Sciences Institute, Istanbul Medipol University, Istanbul, Turkey JAKUB CERVENKA • Institute of Animal Physiology and Genetics, Czech Academy of Sciences, Libechov, Czech Republic; Department of Cell Biology, Faculty of Science, Charles University, Prague, Czech Republic XI CHEN • Shenzhen Key Laboratory of Gene Regulation and Systems Biology, School of Life Sciences, Southern University of Science and Technology, Shenzhen, China OMAR COLLAZO-NAVARRETE • Laboratorio Nacional de Recursos Genomicos, Instituto de Investigaciones Biome´dicas, Universidad Nacional Autonoma de Me´xico, Mexico City, Mexico BIBIANA CORREIA • Department of Life Sciences, University of Coimbra, Calc¸ada Martim de Freitas, Coimbra, Portugal; CNC-Center for Neuroscience and Cell Biology, CIBB, Azinhaga de Santa Comba, Polo 3, University of Coimbra, Coimbra, Portugal HELENA COVELO-MOLARES • Center for Research in Molecular Medicine and Chronic Diseases (CiMUS), Universidade de Santiago de Compostela (USC), Santiago de Compostela, Spain BRUNO DALLAGIOVANNA • Instituto Carlos Chagas—FIOCRUZ-PR, Parana´, Brazil DIVYA DESAI • Department of Biological Sciences, NMIMS Sunandan Divatia School of Science, SVKM’s NMIMS (Deemed to-Be University), Mumbai, Maharashtra, India FATIMA BIANCA A. DESSOUKI • Division of Metabolic and Cardiovascular Sciences, Burnett School of Biomedical Sciences, College of Medicine, University of Central Florida, Orlando, FL, USA AYS¸EGU¨L DOG˘AN • Department of Genetics and Bioengineering, Faculty of Engineering, Yeditepe University, Istanbul, Turkey MANAR ELKENANI • Department of Cardiology and Pneumology, Heart Centre, University Medical Centre Go¨ttingen, Go¨ttingen, Germany; DZHK (German Centre for Cardiovascular Research), Partner Site Go¨ttingen, Go¨ttingen, Germany; Institute of Human Genetics, University Medical Centre Go¨ttingen, Go¨ttingen, Germany SOFI´A LUJA´N FERRIOL-LAFFOUILLERE • Laboratorios de Investigacion Aplicada en Neurociencias (LIAN-CONICET), Fundacion para la Lucha contra las Enfermedades

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Neurologicas de la Infancia (Fleni), Bele´n de Escobar, Provincia de Buenos Aires, Argentina MIGUEL FIDALGO • Center for Research in Molecular Medicine and Chronic Diseases (CiMUS), Universidade de Santiago de Compostela (USC), Santiago de Compostela, Spain SZE WAN FONG • Department of Obstetrics and Gynaecology, University of Hong Kong, Hong Kong, SAR, People’s Republic of China DEBORAH L. FRENCH • Children’s Hospital of Philadelphia Research Institute, Philadelphia, PA, USA PAUL GADUE • Children’s Hospital of Philadelphia Research Institute, Philadelphia, PA, USA ORIANA GENOLET • Otto Warburg Laboratories, Max Planck Institute for Molecular Genetics, Berlin, Germany RUBENS GOMES-JU´NIOR • Instituto Carlos Chagas—FIOCRUZ-PR, Parana´, Brazil OLGA GORDEEVA • Laboratory of Cell and Molecular Mechanisms of Histogenesis, Kol’tsov Institute of Developmental Biology, Russian Academy of Sciences, Moscow, Russia DIANA GUALLAR • Center for Research in Molecular Medicine and Chronic Diseases (CiMUS), Universidade de Santiago de Compostela (USC), Santiago de Compostela, Spain MAGDALENA GUERRA-CRESPO • Instituto de Fisiologı´a Celular, Division de Neurociencias, Universidad Nacional Autonoma de Me´xico, Mexico City, Mexico; Laboratorio de Medicina Regenerativa, Departamento de Cirugı´a, Facultad de Medicina, Universidad Nacional Autonoma de Me´xico, Mexico City, Mexico GOZDE GUVEN • Department of Biological Sciences, Middle East Technical University, Ankara, Turkey ERI HASHINO • Department of Otolaryngology – Head and Neck Surgery, and Stark Neurosciences Research Institute, Indiana University School of Medicine, Indianapolis, IN, USA TAHA BARTU HAYAL • Department of Genetics and Bioengineering, Faculty of Engineering, Yeditepe University, Istanbul, Turkey WEN HUANG • Department of Obstetrics and Gynaecology, University of Hong Kong, Hong Kong, SAR, People’s Republic of China LUCIANA ISAJA • Laboratorios de Investigacion Aplicada en Neurociencias (LIANCONICET), Fundacion para la Lucha contra las Enfermedades Neurologicas de la Infancia (Fleni), Bele´n de Escobar, Provincia de Buenos Aires, Argentina KATERINA VODICKOVA KEPKOVA • Institute of Animal Physiology and Genetics, Czech Academy of Sciences, Libechov, Czech Republic JOSHUA G. KLEIN • Children’s Hospital of Philadelphia Research Institute, Philadelphia, PA, USA ISAO KOBAYASHI • Faculty of Biological Science and Technology, Institute of Science and Engineering, Kanazawa University, Ishikawa, Japan JINGJING KOBAYASHI-SUN • Division of Life Sciences, Graduate School of Natural Science and Technology, Kanazawa University, Ishikawa, Japan ARZUHAN KOC • Regenerative and Restorative Medicine Research Center (REMER), Research Institute for Health Sciences and Technologies (SABITA), Istanbul Medipol University, Istanbul, Turkey; Medical Microbiology Graduate Program, Health Sciences Institute, Istanbul Medipol University, Istanbul, Turkey

Contributors

xi

KATSUHIRO KONNO • Department of Stem Cell Biology and Medicine, Graduate School of Medical Sciences, Kyushu University, Fukuoka, Japan; Translational Research Center, Hiroshima University, Hiroshima, Japan YIN LAU LEE • Department of Obstetrics and Gynaecology, University of Hong Kong, Hong Kong, SAR, People’s Republic of China; Shenzhen Key Laboratory of Fertility Regulation, University of Hong Kong Shenzhen Hospital, Shenzhen, People’s Republic of China BELAL A. MOHAMED • Department of Cardiology and Pneumology, Heart Centre, University Medical Centre Go¨ttingen, Go¨ttingen, Germany; DZHK (German Centre for Cardiovascular Research), Partner Site Go¨ttingen, Go¨ttingen, Germany STEPHEN T. MOORE • Department of Otolaryngology – Head and Neck Surgery, and Stark Neurosciences Research Institute, Indiana University School of Medicine, Indianapolis, IN, USA CARMEN ALEJANDRA MORATO-TORRES • Instituto de Fisiologı´a Celular, Division de Neurociencias, Universidad Nacional Autonoma de Me´xico, Mexico City, Mexico; Department of Pathology, Stanford University School of Medicine, Stanford, CA, USA SOFI´A MUCCI • Laboratorios de Investigacion Aplicada en Neurociencias (LIANCONICET), Fundacion para la Lucha contra las Enfermedades Neurologicas de la Infancia (Fleni), Bele´n de Escobar, Provincia de Buenos Aires, Argentina SHOKO NISHIHARA • Laboratory of Cell Biology, Department of Bioinformatics, Graduate School of Engineering, Soka University, Tokyo, Japan; Glycan & Life System Integration Center (GaLSIC), Soka University, Tokyo, Japan JAROMIR NOVAK • Institute of Animal Physiology and Genetics, Czech Academy of Sciences, Libechov, Czech Republic; Department of Cell Biology, Faculty of Science, Charles University, Prague, Czech Republic MICHAL OPAS • Department of Lab Medicine & Pathobiology, University of Toronto, Toronto, ON, Canada GIULIA PAVANI • Children’s Hospital of Philadelphia Research Institute, Philadelphia, PA, USA FEDERICO PECORI • Laboratory of Cell Biology, Department of Bioinformatics, Graduate School of Engineering, Soka University, Tokyo, Japan ISABELA TIEMY PEREIRA • Instituto Carlos Chagas—FIOCRUZ-PR, Parana´, Brazil PRASAD PETHE • Symbiosis Centre for Stem Cell Research (SCSCR), Symbiosis International University (SIU), Pune, Maharashtra, India IEVGENIIA POLIAKH • Institute of Animal Physiology and Genetics, Czech Academy of Sciences, Libechov, Czech Republic; Department of Cell Biology, Faculty of Science, Charles University, Prague, Czech Republic JOA˜O RAMALHO-SANTOS • Department of Life Sciences, University of Coimbra, Calc¸ada Martim de Freitas, Coimbra, Portugal; CNC-Center for Neuroscience and Cell Biology, CIBB, Azinhaga de Santa Comba, Polo 3, University of Coimbra, Coimbra, Portugal RODRIGO RAMOS-ACEVEDO • Instituto de Fisiologı´a Celular, Division de Neurociencias, Universidad Nacional Autonoma de Me´xico, Mexico City, Mexico; Laboratorio de Medicina Regenerativa, Departamento de Cirugı´a, Facultad de Medicina, Universidad Nacional Autonoma de Me´xico, Mexico City, Mexico LIAT RAVID LUSTIG • Otto Warburg Laboratories, Max Planck Institute for Molecular Genetics, Berlin, Germany MARI´A SOLEDAD RODRI´GUEZ-VARELA • Laboratorios de Investigacion Aplicada en Neurociencias (LIAN-CONICET), Fundacion para la Lucha contra las Enfermedades

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Contributors

Neurologicas de la Infancia (Fleni), Bele´n de Escobar, Provincia de Buenos Aires, Argentina LEONARDO ROMORINI on Aplicada en Neurociencias • Laboratorios de Investigaci (LIAN-CONICET), Fundacion para la Lucha contra las Enfermedades Neurologicas de la Infancia (Fleni), Bele´n de Escobar, Provincia de Buenos Aires, Argentina DERYA SAG˘RAC¸ • Department of Genetics and Bioengineering, Faculty of Engineering, Yeditepe University, Istanbul, Turkey BIRGITT SCHU¨LE • Department of Pathology, Stanford University School of Medicine, Stanford, CA, USA EDDA G. SCHULZ • Otto Warburg Laboratories, Max Planck Institute for Molecular Genetics, Berlin, Germany SELINAY S¸ENKAL • Department of Genetics and Bioengineering, Faculty of Engineering, Yeditepe University, Istanbul, Turkey DERSU SEZGINMERT • Department of Biological Sciences, Middle East Technical University, Ankara, Turkey PATRI´CIA SHIGUNOV • Instituto Carlos Chagas—FIOCRUZ-PR, Parana´, Brazil PAWAN K. SINGAL • Institute of Cardiovascular Sciences, St. Boniface Hospital Albrechtsen Research Centre, University of Manitoba, Winnipeg, MB, Canada DINENDER K. SINGLA • Division of Metabolic and Cardiovascular Sciences, Burnett School of Biomedical Sciences, College of Medicine, University of Central Florida, Orlando, FL, USA HATICE BURCU S¸IS¸LI • Department of Genetics and Bioengineering, Faculty of Engineering, Yeditepe University, Istanbul, Turkey HELENA KUPCOVA SKALNIKOVA • Institute of Animal Physiology and Genetics, Czech Academy of Sciences, Libechov, Czech Republic MARIA INEˆS SOUSA • CNC-Center for Neuroscience and Cell Biology, CIBB, Azinhaga de Santa Comba, Polo 3, University of Coimbra, Coimbra, Portugal YARA SOUTO • Center for Research in Molecular Medicine and Chronic Diseases (CiMUS), Universidade de Santiago de Compostela (USC), Santiago de Compostela, Spain DAISUKE SUGIYAMA • Translational Research Center, Hiroshima University, Hiroshima, Japan NIHAL TERZI CIZMECIOGLU • Department of Biological Sciences, Middle East Technical University, Ankara, Turkey JIRINA TYLECKOVA • Institute of Animal Physiology and Genetics, Czech Academy of Sciences, Libechov, Czech Republic YOSHITOMO UEDA • Department of Otolaryngology – Head and Neck Surgery, and Stark Neurosciences Research Institute, Indiana University School of Medicine, Indianapolis, IN, USA LUDOVIC VALLIER • Jeffrey Cheah Biomedical Centre, Department of Surgery, WellcomeMRC Cambridge Stem Cell Institute, University of Cambridge, Cambridge, UK LORIANA VITILLO • Jeffrey Cheah Biomedical Centre, Department of Surgery, WellcomeMRC Cambridge Stem Cell Institute, University of Cambridge, Cambridge, UK; Institute of Ophthalmology, University College London, London, UK PETR VODICKA • Institute of Animal Physiology and Genetics, Czech Academy of Sciences, Libechov, Czech Republic QIUSHI XU • Shenzhen Key Laboratory of Gene Regulation and Systems Biology, School of Life Sciences, Southern University of Science and Technology, Shenzhen, China WEI XU • Shenzhen Key Laboratory of Gene Regulation and Systems Biology, School of Life Sciences, Southern University of Science and Technology, Shenzhen, China

Contributors

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YING YE • Cam-Su Genomic Resource Center, Soochow University, Suzhou, China WILLIAM SHU BIU YEUNG • Department of Obstetrics and Gynaecology, University of Hong Kong, Hong Kong, SAR, People’s Republic of China; Shenzhen Key Laboratory of Fertility Regulation, University of Hong Kong Shenzhen Hospital, Shenzhen, People’s Republic of China WENSHENG ZHANG • Department of Physiology, School of Basic Medical Sciences, Binzhou Medical University, Yantai, China; Cam-Su Genomic Resource Center, Soochow University, Suzhou, China JIANQUN ZHENG • Shenzhen Key Laboratory of Gene Regulation and Systems Biology, School of Life Sciences, Southern University of Science and Technology, Shenzhen, China

Methods in Molecular Biology (2022) 2520: 1–24 DOI 10.1007/7651_2021_457 © Springer Science+Business Media, LLC 2022 Published online: 27 February 2022

Dissecting Molecular Phenotypes Through FACS-Based Pooled CRISPR Screens Oriana Genolet, Liat Ravid Lustig, and Edda G. Schulz Abstract Pooled CRISPR screens are emerging as a powerful tool to dissect regulatory networks, by assessing how a protein responds to genetic perturbations in a highly multiplexed manner. A large number of genes are perturbed in a cell population through genomic integration of one single-guide RNA (sgRNA) per cell. A subset of cells with the phenotype of interest can then be enriched through fluorescence-activated cell sorting (FACS). SgRNAs with altered abundance after phenotypic enrichment allow identification of genes that either promote or attenuate the investigated phenotype. Here we provide detailed guidelines on how to design and execute a pooled CRISPR screen to investigate molecular phenotypes. We describe how to generate a custom sgRNA library and how to perform a FACS-based screen using readouts such as intracellular antibody staining or Flow-FISH to assess phosphorylation levels or RNA abundance. Through the variety of available perturbation systems and readout options many different molecular and cellular phenotypes can now be tackled with pooled CRISPR screens. Key words CRISPR screen, Embryonic stem cell, Signaling, MAPK pathway, Intracellular antibody staining, Flow-FISH, RNA, sgRNA library

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Introduction Pooled CRISPR screens are a powerful tool for the unbiased identification of genes mediating a certain phenotype. Prior to the advent of the CRISPR/Cas9 technology, short-hairpin RNA pools were used for mRNA downregulation in pooled RNAi screens, but high variability in knock-down efficiency and off-target activity restricted their use [1, 2]. More recently, the combination of the Cas9 endonuclease together with pooled sgRNA libraries led to a broader adoption of pooled screens as a discovery tool in forward genetics [3, 4]. The number of available perturbation systems and phenotypes that can be investigated has rapidly expanded since then. Gene overexpression (CRISPR activation) and downregulation (CRISPR inhibition) through coupling catalytically inactive Cas9 (dCas9) to transcriptional effector domains was successfully used in CRISPR screens [5, 6]. Moreover,

Oriana Genolet and Liat Ravid Lustig contributed equally with all other contributors.

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in addition to the investigation of viability phenotypes, a variety of molecular phenotypes can now be studied through fluorescenceactivated cell sorting (FACS) [7]. CRISPR screens have been used to identify protein-coding genes, noncoding RNAs, and more recently also cis-regulatory elements that control a specific phenotype or gene of interest [3, 7–11]. In addition, also single-cell RNA sequencing (scRNAseq) has been used as readout in CRISPR screens, which allows omission of the phenotypic enrichment step [12–14]. However, since the technical considerations for scRNA-seq screens are rather different and they, so far, cannot be applied on a large scale and require extensive computational efforts to limit background noise and reduce the number of false positives, they are not covered here. In this protocol we will provide detailed guidelines for screen design and execution of enrichment-based pooled CRISPR screens, which can be applied to numerous systems, cell types, and experimental questions. Moreover, general considerations are additionally given at various steps that allow the adaptation of the protocol to different experimental requirements. In a pooled CRISPR screen, a population of cells is transduced with one sgRNA per cell to induce perturbation of a single gene. After enrichment of cells with the phenotype of interest the relative abundance of each sgRNA is quantified by deep sequencing. Importantly, the sgRNA library is usually delivered by lentiviral transduction, which allows a precise control of how many sgRNAs are expressed in each cell (usually only one) and enables quantification of the relative number of cells carrying each sgRNA in a population, through sequencing of the genomically integrated sgRNA vector. In pooled CRISPR screens a sufficiently strong perturbation must be induced with a single sgRNA per cell. In knock-out screens this can be achieved with high efficiency by inducing frameshift mutations. However, due to the fact that one third of repair events will generate in-frame mutations, this strategy generates a mixture of wildtype, homozygous, and heterozygous mutant cells, thus potentially reducing screen sensitivity. Cas9 nevertheless remains the most potent loss-of-function perturbation system to date, but is usually limited to protein-coding genes. An alternative strategy for loss-of-function that also allows interrogation of noncoding genes, such as long noncoding RNAs (lncRNAs), is CRISPR interference (CRISPRi), where dCas9 is fused with a repressor domain, such as KRAB or the more recently developed KRAB-MeCP2 fusion protein [5, 15]. To interrogate the function of RNAs also RNA-targeting CRISPR/Cas systems, such as Cas13/CasRx, are being established, and might be promising screening tools in the future [16, 17]. Through the development of CRISPR activation (CRISPRa) systems, also gain-of-function screens have become possible. However, typically, overexpression with a single sgRNA recruiting a

CRISPR Screens

3

simple dCas9 effector fusion protein is not sufficiently strong. Therefore CRISPRa screens have made use of systems that recruit multiple effector domains with one sgRNA, such as the SunTag or SAM systems [5, 6]. Also other potent CRISPRa systems, such as VPR, might potentially be used in a screen setting [18]. A general limitation of CRISPRa and also CRISPRi screens is that perturbation strength varies significantly across sgRNAs and genes [19]. Pooled CRISPR screens rely on the ability to physically separate a subset of cells with the phenotype of interest. For phenotypes that affect proliferation or viability, cells can be enriched or depleted through prolonged culture (for example [20, 21]). This approach can be easily scaled up and is therefore used in most genome-wide screens. Here we focus on screens that investigate molecular phenotypes and thus use FACS-sorting for phenotypic enrichment. To investigate the regulation of a specific protein, it can be tagged with a fluorescent protein. If protein tagging is not possible, the endogenous protein can be detected through antibody-mediated fluorescence staining [22, 23]. If a suitable antibody exists, this approach also allows investigation of specific post-translational modifications. A second tagging-free approach which additionally allows investigation of noncoding RNAs is based on RNA detection through Flow-FISH and can in principle be applied to any sufficiently abundant RNA [24–26]. To ensure high screen sensitivity, sufficient library representation, also called coverage, must be maintained at all steps of a screen. Coverage is defined as the average number of cells or molecules per sgRNA present in a sample. Maintenance of sufficient coverage is important to ensure that all sgRNAs are detected in all samples, which is a prerequisite for the subsequent statistical analysis. Similar coverage should be maintained throughout the different steps of the screen and is defined as follows: 1. Library cloning: Number of independent ligation and transformation events per sgRNA. 2. Lentiviral transduction: Number of cells transduced with each sgRNA. 3. Cell splitting: Number of cells plated per sgRNA. 4. Phenotypic enrichment: Number of cells per sgRNA in the selected/sorted population. 5. Sequencing library preparation: Number of genomically integrated sgRNA cassettes (¼ number of cells) that are amplified in the PCR. 6. Sequencing: Number of reads per sgRNA. Typically, a coverage between 100 and 1000 is used [3, 5] and simulations have confirmed that this range supports high performance [27]. The required coverage strongly depends on the width

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of the distribution of the cloned sgRNA library [28]. The distribution width can be quantified as the fold change between the 90th and 10th percentile. Simulations have shown that a 200 coverage is sufficient for a distribution width of 2.5, while 400 is required for a tenfold distribution width [28]. Here we provide detailed instructions on how to perform a CRISPR-KO screen, using intracellular antibody staining of phospho-Mek for phenotypic readout. We have used this assay to identify genes that modulate the differentiation-promoting MAPK signaling pathway in murine embryonic stem cells [23]. We also provide instructions on how to perform phenotypic enrichment based on RNA levels quantified by Flow-FISH, which we have recently used to investigate the regulation of the Xist lncRNA [25].

2 2.1

Materials Cloning and PCR

1. Synthesized oligo pool (e.g., Genscript or Twist Bioscience). 2. lentiGuide-Puro sgRNA expression vector (Addgene, plasmid ID 52963). 3. BsmBI restriction enzyme (alternatively Esp3I, New England BioLabs). 4. Phusion Hot Start Flex DNA Polymerase (New England BioLabs). 5. KAPA HiFi HS Ready Mix (Roche). 6. dNTP Mix: 10 mM each. 7. PCR purification kit (e.g., Qiagen QIAquick PCR Purification Kit). 8. Gel extraction kit (e.g., QIAquick Gel extraction kit). 9. LE Agarose. 10. 10 TBE Buffer (Thermo Fisher Scientific). 11. DNA Ladder 100 bp (New England Biolabs). 12. SYBR Safe DNA Gel Stain-400 (Thermo Fisher Scientific). 13. Gel Loading Dye, Purple (6, New England BioLabs). 14. Gibson Assembly Master Mix (New England Biolabs). 15. Pellet Paint Co-precipitant (Merck KGaA). 16. NaOAc: 3 M solution in H2O (Invitrogen). 17. Ethanol 100%. 18. NaCl. 19. MegaX DH10B T1R Electrocomp. Cells (Thermo Fisher Scientific). 20. LB medium: 10 g NaCl, 10 g Tryptone, 5 g Yeast Extract, 1 ml 1 M NaOH dissolved in 1 l ddH2O.

CRISPR Screens

5

21. Ampicillin. 22. SOC medium (Thermo Fisher Scientific). 23. Plasmid Maxi prep (e.g., Nucleobond Xtra MidiPlus, Macherey-Nagel). 24. Agar Agar. 25. Agar plates (1 l LB medium +15 g Agar Agar). 26. Primer Sequences (see Table 1). 2.2

Cell Culture

1. HEK293FT (Thermo Fisher). 2. ViraPower™ Lentiviral Packaging Mix: pLP1, pLP2, VSVG (Thermo Fisher Scientific). 3. HEK293T culture medium: DMEM, 10% FBS. 4. Opti-MEM I Reduced Serum (Thermo Fisher Scientific). 5. Lipofectamine 2000 (Thermo Fisher Scientific). 6. Lenti-X-Concentrator (Takara). 7. Cell line stably expressing Cas9 (e.g., 1.8 ESCs, transduced with lentiCas9-Blast, Addgene #52962). 8. Puromycin (Sigma-Aldrich). 9. Polybrene Infection/Transfection Reagent (Merck KGaA). 10. 10 PBS (Sigma-Aldrich). 11. Trypsin (Thermo Fisher Scientific). 12. ESC Culture medium: DMEM, 15% ES-grade FBS, 0.1 mM β-mercaptoethanol, 1000 U/ml LIF.

2.3 Antibody Staining and FlowFISH

1. Staining media: PBS, 1% BSA. 2. Sorting buffer: PBS, 1% FBS, 1 mM EDTA. 3. Paraformaldehyde: 3% solution in H2O. 4. Methanol: 100%. 5. Bovine Serum Albumin (BSA, Sigma-Aldrich). 6. Secondary fluorescently labeled antibody (e.g., AlexaFluor-647 Goat anti-Rabbit, Thermo Fisher Scientific). 7. PrimeFlow RNA assay (ThermoFisher) and gene-specific target probes. 8. Conical bottom 96-well plate.

2.4 Genomic DNA Extraction

1. UltraPure SDS Solution (Thermo Fisher Scientific). 2. Tris-EDTA (TE) buffer solution (Sigma-Aldrich). 3. Lysis Buffer: 1% SDS, 0.2 M NaCl, 5 mM DTT in TE Buffer. 4. RNase A: 10 mg/ml (Thermo Fisher Scientific). 5. Proteinase K: 20 mg/ml solution in H2O (Sigma-Aldrich).

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Table 1 Primer sequences. Illumina barcodes are represented in bold Name

Sequence

OG113

TAACTTGAAAGTATTTCGATTTCTTGGCTTTATATATCTTGTGGAAAGGAC GAAACACCG

OG114

ACTTTTTCAAGTTGATAACGGACTAGCCTTATTTTAACTTGCTATTTCTAG CTCTAAAAC

OG115

AATGGACTATCATATGCTTACCGTAACTTGAAAGTATTTCG

OG116

CTTTAGTTTGTATGTCTGTTGCTATTATGTCTACTATTCTTTCC

OG125

AATGATACGGCGACCACCGAGATCTACACTCTTTCCCTACACGACGCTCT TCCGATCTTCTTGTGGAAAGGACGAAACACCG

OG126_bc01 CAAGCAGAAGACGGCATACGAGATATCACGGTGACTGGAGTTCAGACGTG TGCTCTTCCGATCTTCTACTATTCTTTCCCCTGCACTGT OG126_bc02 CAAGCAGAAGACGGCATACGAGATCGATGTGTGACTGGAGTTCAGACGTG TGCTCTTCCGATCTTCTACTATTCTTTCCCCTGCACTGT OG126_bc03 CAAGCAGAAGACGGCATACGAGATTTAGGCGTGACTGGAGTTCAGACGTG TGCTCTTCCGATCTTCTACTATTCTTTCCCCTGCACTGT OG126_bc04 CAAGCAGAAGACGGCATACGAGATTGACCAGTGACTGGAGTTCAGACGTG TGCTCTTCCGATCTTCTACTATTCTTTCCCCTGCACTGT OG126_bc05 CAAGCAGAAGACGGCATACGAGATACAGTGGTGACTGGAGTTCAGACGTGT GCTCTTCCGATCTTCTACTATTCTTTCCCCTGCACTGT OG126_bc06 CAAGCAGAAGACGGCATACGAGATGCCAATGTGACTGGAGTTCAGACGTGT GCTCTTCCGATCTTCTACTATTCTTTCCCCTGCACTGT OG126_bc07 CAAGCAGAAGACGGCATACGAGATCAGATCGTGACTGGAGTTCAGACGTG TGCTCTTCCGATCTTCTACTATTCTTTCCCCTGCACTGT OG126_bc08 CAAGCAGAAGACGGCATACGAGATACTTGAGTGACTGGAGTTCAGACGTG TGCTCTTCCGATCTTCTACTATTCTTTCCCCTGCACTGT OG126_bc09 CAAGCAGAAGACGGCATACGAGATGATCAGGTGACTGGAGTTCAGACGTGT GCTCTTCCGATCTTCTACTATTCTTTCCCCTGCACTGT OG126_bc10 CAAGCAGAAGACGGCATACGAGATTAGCTTGTGACTGGAGTTCAGACGTGT GCTCTTCCGATCTTCTACTATTCTTTCCCCTGCACTGT LR256

CTGTGAATAAGATGTCCATT

LR257

ACACTCTCGTTATTTGTCAT

LR21

ACTATCATATGCTTACCGTAAC

6. Phenol/chloroform/Isoamyl (Roth).

alcohol

mixture

125:24:1

7. Qubit dsDNA HS Assay Kit-100 (Thermo Fisher Scientific).

CRISPR Screens

2.5

Equipment

7

1. ECM 399 electroporator (BTX). 2. Spectralphotometer NanoDrop™ 2000 (Peqlab). 3. Safe Imager™ 2.0 Blue-Light Transilluminator (Thermo Fisher Scientific). 4. Hybridization oven capable of maintaining temperature at 40  C (for Flow-FISH). 5. EVE™ Automated Cell Counter, NanoEnTek (VWR). 6. Qubit Fluorometer (Thermo Fisher Scientific). 7. Illumina sequencer.

3

Methods In this section we describe every step of a FACS-based pooled lentiviral CRISPR screen, where molecular phenotypes are detected by intracellular staining of proteins or RNA, using the lentiGuidePuro sgRNA expression vector (Fig. 1). Subheadings 3.1 and 3.2 provide detailed instructions for design and cloning of the sgRNA library, respectively. Subheading 3.3 covers packaging of the sgRNA library in lentiviral particles and transduction with the library. Phenotypic enrichment using antibody staining or RNA detection by Flow-FISH is described in Subheading 3.4. Subheadings 3.5 and 3.6 explain the preparation of sequencing libraries and statistical data analysis, respectively.

3.1 SgRNA Library Design

1. Download a genome-wide CRISPR knock-out library from the Addgene website (https://www.addgene.org/crispr/ libraries/) (see Note 1). 2. Decide which genes should be targeted by the library and how many sgRNAs should be included for each gene (see Note 2). Extract the sequences from the genome-wide library accordingly. In addition, add at least 100–1000 non-targeting or safetargeting control sgRNAs (see Note 3). Make sure to also include positive control genes in the library, which are known to affect your phenotype of interest. 3. Add homology arms for library cloning by extending the sgRNA sequences as follows: TGGAAAGGACGAAACACCG[sgRNA]GTTTTAGAGC TAGAAATAGCAAGTTAAAATAAGGC 4. Synthesize the sequences as a custom oligo pool. Several sgRNA libraries can be synthesized in the same pool (see Note 4).

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3.1-3.2. SgRNA library design and cloning

sgRNAs

3.3. Lentiviral transduction

Cas9

3.4. Phenotypic enrichment Antibody staining

Nr. Cells

Flow-FISH

Protein

A RN

Fluorescence genomic DNA

3.5. NGS library preparation and sequencing

PCR

sgRNA cassette

Input

Count table

3

sgRNA

Gene

Input

Sorted

1 2 3 4 5 :

A A A B B :

331 287 347 266 367 :

256 103 87 867 789 :

FDR (−log10)

3.6. Data analysis

Sorted

2

1

0 −2

−1

0

1

2

Fold change Sorted/Input

Fig. 1 Protocol overview. The sgRNA library is designed, synthesized in a pooled fashion on a microarray, and cloned into the lentiviral sgRNA expression vector lentiGuide-Puro (Subheadings 3.1 and 3.2). The cloned sgRNA library is packaged in lentiviral particles and used to transduce cells expressing the perturbation system (Subheading 3.3). Cells with the phenotype of interest are enriched by FACS using intracellular antibody staining or RNA detection by Flow-FISH (Subheading 3.4). Genomic DNA is extracted from the sorted and input samples. The genomically integrated sgRNA cassette is amplified by PCR, tagged with sample barcodes and sequencing adaptors, and subjected to NGS (Subheading 3.5). Reads are mapped to the sgRNA library to produce a count table and screen hits are identified by statistical analysis (Subheading 3.6)

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1. To amplify the oligo pool and extend the overhangs with homology to the vector (Fig. 2), set up the following PCR reaction. In this step sufficient product must be generated for the subsequent Gibson reactions, while keeping the number of amplification cycles low. It is thus recommended to set up several PCR reactions (between 4 and 12 reactions depending on library size).

3.2 SgRNA Library Cloning

BsmBI

cPPT sgRNA scaffold

EF-1a

Filler Puro

BsmBI

WPRE hU6 promoter

lentiGuide-Puro 3‘ LTR SIN

10182 bp

SV40 PolyA

RRE gag

f1 Ori psi RSV/5‘LTR pUC Ori

AmpR

BsmBI site BsmBI site digested ...ATATATCTTGTGGAAAGGACGAAACACCGGAGACG.......CGTCTCTGTTTTAGAGCTAGAAATAGCAAGTTAAAATAAGGCTAGTCCG... ...TATATAGAACACCTTTCCTGCTTTGTGGCCTCTGC.......GCAGAGACAAAATCTCGATCTTTATCGTTCAATTTTATTCCGATCAGGC... vector PCR N31...ATATATCTTGTGGAAAGGACGAAACACCGNNNNNNNNNNNNNNNNNNNNGTTTTAGAGCTAGAAATAGCAAGTTAAAATAAGGCTAGTCCG...N18 product primer OG113

primer OG114

N31...ATATATCTTGTGGAAAGGACGAAACACCG

synthesized oligo

GTTTTAGAGCTAGAAATAGCAAGTTAAAATAAGGCTAGTCCG...N18

TGGAAAGGACGAAACACCGNNNNNNNNNNNNNNNNNNNNGTTTTAGAGCTAGAAATAGCAAGTTAAAATAAGGC

sgRNA sequence

Fig. 2 Cloning of sgRNA library. The sgRNA expression vector lentiGuide-Puro (orange: sgRNA expression, green: antibiotic resistance; blue: lentiviral elements; gray: vector backbone) is digested with BsmbI. The synthesized oligos (bottom) are amplified with primers OG113/114 and ligated in the digested vector with homology-directed Gibson cloning

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Components

Amount

Final concentration

5 Phusion HF Buffer

10 μl

1

10 mM dNTPs

1 μl

200 μM

DMSO

1.5 μl

3%

10 μM Forward Primer (OG113)

2.5 μl

0.5 μM

10 μM Reverse Primer (OG114)

2.5 μl

0.5 μM

Oligo Pool

1–5 ng

Phusion Hot Start Flex DNA Polymerase

1 μl

Nuclease-free water

Add to 50 μl

2 units

2. Run the PCR with the following cycle parameters. Cycle

Denature

Anneal

Extend

0

98  C 30 s

–

–





1–14

98 C 15 s

63 C 15 s

72  C 15 s

15

–

–

72  C 5 min

3. Pool PCR reactions and purify with a PCR Purification Kit to concentrate the PCR reactions. Elute in 30 μl Nuclease-free water. 4. Gel-purify the pooled PCR product (140 bp) from a 2% agarose gel (LE) to eliminate primer dimers (approx. 120 bp). 5. Digest 2.5 μg of lentiGuide-Puro plasmid (see Note 5) with 2.5 μl of BsmbI for 1 h at 55  C or overnight at 37  C in a 50 μl reaction. 6. Gel-purify the digested plasmid (approx. 8.3 kb, filler sequence is approx. 1.9 kb). 7. Determine the concentration of the digested plasmid and the PCR product via Nanodrop. 8. Set up a Gibson assembly reaction as follows using the gel-purified lentiGuide-Puro plasmid from step 6 and the purified PCR amplicon from step 4, and one negative control reaction without insert. Perform 1–2 Gibson reactions per 10,000 sgRNAs.

CRISPR Screens

11

Component

Amount

BsmBI digested and gel-purified lentiGuide-Puro

100 ng

PCR-amplified oligo pool

7 ng

2 Gibson assembly master mix

10 μl

Nuclease-free water

Add to 20 μl

9. Incubate in a thermocycler at 50  C for 1 h and quench on ice. 10. Pool Gibson reactions and bring to 200 μl with nuclease-free water. 11. Add 1.5 μl of pellet paint, 20 μl 3 M NaOAc, and 600 μl 100% EtOH. 12. Incubate samples at 80  C for 30 min (or overnight at 20  C). 13. Centrifuge for 30 min at 20,000  g and 4  C to pellet DNA, followed by two washes with ice-cold 80% EtOH. 14. Air-dry pellet and resuspend in 5 μl nuclease-free water per Gibson reaction. 15. Freeze the eluted DNA for 3 h at 80  C (this step might increase transformation efficiency). 16. Add maximally 9 μl of the Gibson assembly reaction to 20 μl of thawed electrocompetent MegaX DH10B T1R E.coli and incubate 30 min on ice. Perform enough electroporation reactions to transform the entire Gibson assembly reaction. 17. Pipette the bacterial cells with the Gibson reaction into a prechilled electroporation cuvette and tap gently on a surface to remove bubbles. Place the cuvette in an electroporator and electroporate the sample at 2.0 kV, 200 Ω, and 25 μF. 18. Add 1 ml of prewarmed recovery or SOC medium to the cuvette quickly after electroporation (make sure to pipette up and down to resuspend all bacterial cells from the bottom of the cuvette). 19. Transfer bacterial cells to a culture tube and shake at 250 rpm for 1 h at 37  C. Pool electroporation reactions at this step if multiple reactions were performed. 20. Bring sample to a volume of 10 ml with LB medium supplemented with ampicillin (0.1 mg/ml) and mix well. 21. To estimate library coverage plate 10 μl (1:1000 dilution) and 100 μl (1:100 dilution) of the mixture on pre-warmed Ampicillin-supplemented (0.1 mg/ml) agar plates. Additionally, make a 1:100 dilution of the mixture (10 μl in 990 μl LB medium) and plate 1 μl (1:1,000,000 dilution), 10 μl (1: 100,000 dilution), and 100 μl (1:10,000 dilution) on three

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additional ampicillin agar plates and incubate overnight at 37  C. 22. Inoculate 500 ml of LB medium plus ampicillin (0.1 mg/ml) with the remaining bacteria and incubate overnight on a shaker at 37  C 250 rpm. 23. The next day count the colonies in the agar plates and multiply by the dilution factor. The average number of colonies divided by the library size yields the coverage of the cloned sgRNA library. The coverage of library cloning should be 100–1000 (see Note 6 and example in Table 2). 24. Pick 10 colonies, carry out mini-scale plasmid preparations and sequence the insert through Sanger sequencing with primer LR21. At least eight out of the ten colonies should carry an identifiable sgRNA sequence from the cloned library. 25. Pellet bacterial cells from step 22 by centrifugation for 20 min at 20,000  g at 4  C and carry out a plasmid Maxiprep. Table 2 Example calculation for coverage. Here we give an example how sufficient coverage can be ensured at all steps of the screen. The calculations below are for an sgRNA library of 1000 sgRNAs (library size), where 15% of cells were sorted after antibody staining and a 300 coverage is maintained Step

Material required

PCR to amplify oligo pool (Subheading 3.2, step 1)

Amplify 1–5 ng (~6–30  109 molecules)

Gibson assembly and transformation (Subheading 3.2, step 21)

3  105 transformants

Lentiviral transduction (Subheading 3.3, step 13)

Treat 9  105 cells with viral supernatant (MOI ¼ 0.3) ! 3  105 cells transduced with one sgRNA

Selection and expansion (Subheading 3.3, step 15)

Always plate 3  105 cells

Cell harvesting (Subheading 3.4, step 1)

Use 1.2  107 cells for staining

Antibody staining (Subheading 3.4, steps 6–11)

~30–50% loss (6  106 cells left) take 6  105 cells as input control

Single-cell gating (Subheading 3.4, step 12)

~30% loss (4  106 cells left)

Cell sorting (15%) (Subheading 3.4, step 12)

~85% loss (6  105 cells left)

Genomic DNA extraction (Subheading 3.5, steps 5–9)

~20–50% loss (DNA from 3  105 cells left)

NGS library prep (Subheading 3.5, steps 12–17)

Amplify DNA from 3  105 cells  1.8 μg [a mouse cell contains 6 pg DNA, a human cell 7 pg DNA]

Sequencing (Subheading 3.5, step 18)

Sequence 4.3  105 reads [assuming 70% alignment rate]

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13

26. In order to confirm good library representation by NGS, add sequencing adaptors and samples barcodes by PCR as follows. Components

Amount

Final concentration

2 Ready Mix Kapa

25 μl

1

10 μM Forward Primer (OG125)

1.5 μl

0.3 μM

10 μM Reverse Primer (OG126)

1.5 μl

0.3 μM

Plasmid library

100 ng

Nuclease-free water

Add to 50 μl

27. Run the PCR with the following cycle parameters. Cycle

Denature 

Anneal

Extend

0

95 C 3 min

–

–

1–12

98  C 20 s

55  C 15 s

72  C 15 s

13

–

–

72  C 1 min

28. Gel-purify the PCR amplicon of 351 bp. 29. Subject the purified PCR amplicon to Illumina sequencing as outlined in Subheading 3.5, step 18. 30. For analysis follow steps 1–8 in Subheading 3.6. Calculate the distribution width given as the fold change between the 90th and the 10th percentiles (see Fig. 3 for an example). The distribution width should ideally be Stacks > Z Project. Choose your Start and Stop slices and Maximum projection type. Image of Maximum Intensity Projection appears in a new window. 4. Constructing rotated 3D projected stacks and movies available in LAS X Full version: Click on the tab and then on the tab. Click on the tab under the Visualization header. A 3D Projection window will appear with a grayscale graphic of your image series. Click in the box and select options for movie files. 5. 3D Reconstruction/Volume Rendering with Fiji/ImageJ Software: Improve image quality using Image > Adjust > Brightness/Contrast or increase signal with Process > Enhance contrast. To make the rotated 3D projection video, go to Image > Stacks >3D Project. Select the Projection method and Axis of rotation. Save as an .avi file with no compression and then click on it to watch the movie. 6. 3D Reconstruction/Volume and Surface Rendering with Fiji/ ImageJ Software: Go to Plugins >3D Viewer. Select Volume or Surface in the Display As option. Select the color (if needed) and set the threshold to 20–50, and confirm OK. The 3D reconstructed model will appear in the ImageJ 3D Viewer window. Use the mouse manipulator for arbitrary rotation and analysis of 3D reconstruction. To make a snapshot, go to View > Take a snapshot, which can be saved as a jpeg or tiff. Image. To make a movie, go to View > Record 360
rotation or choose other options for the video file. For saving the movie, go to View > Save as > AVI.

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3.6 Quantification and High Content Imaging Analysis 3.6.1 The 2D Automated Cell Counter Tool in ImageJ/Fiji

To analyze proliferation and differentiation patterns in the EBs, the proportions of cells expressing markers relative to the total number of cells can be calculated using the Automated Cell Counter tool in ImageJ (https://imagej.nih.gov/ij/plugins/cell-counter.html). The 2D Object Counter is based on the object identification in 2D images of stained cells in a z-stack (Fig. 3). 1. Pre-processing of a z-stack is required for further Automated Cell Counter analysis. For quantitative analysis, acquire a zstack with 5–10 μm z-steps. Extra images can be removed from the stack in ImageJ. Right Click on a z-stack window and select Duplicate, then, in the opened dialog window, check Duplicate stack and set Range for images in the stack. Individual optical slices can be deleted via Image > Stacks > Delete Slice (see Note 11). 2. Image stacks acquired from multiple channels must be split into separate stacks for each channel for further calculation (Image > Color > Split channels). 3. Check the color intensity of the stained objects expected for calculation (nuclei or other organelles, or whole cells) on different optical sections along the z-axis. In case of significant differences, it is necessary to split the z-stack into several substacks with similar color intensity, since this is a significant point for the correct setting of thresholds during the further calculation. On average, with a scan depth of 150 μm and a step of 5–10 μm, you can get 12–25 analyzable images in a z-stack. 4. Check the image quality for each substack and, if necessary, increase the signal with Process > Enhance contrast or improve image quality with Image > Adjust > Brightness/Contrast. 5. Convert substacks for each channel to grayscale mode. Go to Edit > Options > Conversions to set checkbox Scale when converting. Then use Image > Type >8-bit or 16-bit to convert to grayscale. 6. If too many noise or background pixels are highlighted, fix this issue by going to Process > Subtract background. 7. After image grayscale conversion, use Image > Adjust > Threshold to highlight all of the objects for counting with the sliders. Review all images in the stack. If the resulting images are of good quality, click Apply, and set checkbox Calculate thresholds for each image in open Convert Stack to Binary dialog window. Thus, a binary version of the image with only two-pixel intensities is created. 8. At the segmentation step, merged objects, which were on initial images or run together during the threshold set, can be divided using the watershed procedure: go to Process > Binary > Watershed. Process all images in the z-stack. It can help to divide merged objects.

Fig. 3 A workflow for high content imaging analysis (HCIA) in the 3D EB differentiation model. A workflow for quantitative HCIA of the 3D EB differentiation model with Fiji/ImageJ software that includes the following

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9. To set the parameters for counting objects, the minimum and maximum area of objects need to be measured using Measure tools before starting to count the objects. Outline selected objects with freehand selection tool from the ImageJ menu bar and go to Analyze > Measure. The Results window appears with statistic values for the selected objects (see Note 12). 10. For 2D Automated Counting use Analyze Particles tool (Analyze > Analyze Particles). This command counts and measures objects in binary images by scanning the image until it finds the edge of an object. It outlines the object, measures it, fills it to make it invisible, then resumes scanning until it reaches the end of each image in the z-stack. 11. Use the dialog window to configure the object analyzer: l

specify sizes in the Size field, objects outside the range are ignored;

l

select Outlines from the Show dropdown selection to open a window containing numbered outlines of the measured objects;

l

check Display results to display measurements for each particle in the Results window;

l

check Summarize to display the object count, total particle area, average particle size, and area fraction in a separate window;

l

check Exclude on Edges to ignore particles touching the edge of the image;

l

check Include Holes to include interior holes;

l

check Add to Manager and the measured objects will be added to the ROI Manager;

l

check Summarize to calculates and displays the mean, standard deviation, minimum and maximum of the values in each column in the Results table;

l

uncheck In Situ show to displays numbered outlines of the measured objects of each image in the z-stack in a separate window;

l

check Clear Results to erase any previous measurement results.

 Fig. 3 (continued) steps: step 1, acquisition of z-stack images using confocal microscopy; step 2, pre-processing of z-stack images (checking and improving image quality); step 3, fluorochrome channel splitting; step 4, adjustments and threshold (binary images) for single-channel z-stack images; step 5, segmentation of merged objects; step 6, object measuring and automated counting; and step 7, calculation and statistical analysis

High-Content Image Analysis of 3D EB Differentiation Model

77

12. After completion of the calculation, check the results in all the windows and tables. To save the data, select all content in the Summary table and save it as a file in cvs. format (File > Save as). 13. Use this data to further calculate the number of cells expressing the markers in relation to the total number of cells in each image. The average over the entire z-stack is calculated from the data on the percentage of cells in each optical section. 14. For further statistical analysis, at least 10 EBs must be tested for each experimental group. It is necessary to calculate inter- and intra-assay coefficients of variability. 3.6.2 3D Objects Counter in the Fiji/ImageJ

The 3D Objects Counter plugin for Fiji/ImageJ (Analyze >3D Objects Counter) is an alternative approach to analyzing the cellular pattern in 3D cell models. This plugin counts the number of 3D objects in a single-channel stack (after channel splitting) and displays the volume, the surface, the center of mass, and the center of intensity for each identified object. 1. All required settings can be selected under Analyze >3D OC Options. 2. Go to the 3D Objects Counter and set up the threshold value which can be adjusted using two sliders for levels and navigation through the stack. For correct counting of the objects, set the voxel size range, which should be determined by preliminary test counting. 3. The results are presented in the summary tables, and several new stacks for the full volume, the edges, the center of mass, and center of intensity of each particle. Each of these stacks can be merged with single-channel stacks to build a 3D projection for further image analysis (see Subheading 3.5.2) (see Note 13).

4

Notes 1. Per one Petri dish with 90–100 drops, 1500 μL of ESC suspension must be prepared. 2. There must be a distance of at least 3 mm between drops to prevent droplets from merging. 3. EB adhesion is a common problem in EB culture, even in dishes with a low adhesion surface. This is because, after attachments, EBs lose their spheroid shape and are not suitable for further analysis of the spatiotemporal differentiation pattern. This also affects the morphological heterogeneity of EBs and interferes with standardization for further analysis.

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4. It is important to maintain the same volume of medium in the wells of the plate for correct measurement of EB diameters. 5. EB diameters can be measured during confocal microscopy analysis (Fig. 2). 6. For better confocal imaging, clarify EB preparations in several dilutions of glycerol with anti-fade additives (DAPCO) or mounting medium in PBS (30%, 50%, 70%, 100%). Store EB preparations for several days at 4
C, protected from the light. 7. The plates with EBs may be placed on a shaker in a thermostat at 37
C for 1 h. 8. At all stages of immunofluorescence staining, it is necessary to carefully remove and add solutions checking EBs under a stereomicroscope to prevent their loss or damage. 9. Be careful to prevent damage to the glass bottom of the plates or μ-slides. 10. To study spatiotemporal patterns in EB at different stages of differentiation, the following parameters for the Leica TSC SPE and SP5 confocal microscopes confocal microscopy were used: the 405, 488, and 596 lasers; HCX PL APO CS 20.0  0.70 IMM UV and 40.0  1.25 OIL UV objectives; x/y resolution: 512  512, 1024  1024 or 2048  2048 pixels, z-resolution: 0.3–10 μm at 100–150 μm depth of specimen scanning, and xyz sequential mode. 11. In some cases, optical sections of surface EB layers (up to 10 μm from the surface) containing a small number of cells need to be excluded from z-stacks, since this introduces a large variability and further incorrect calculation of the percentage of stained objects. 12. The square range for the counting of mouse ESC nuclei needs to be set at 15–350 μm2 based on our manual cell measurements. 13. In our experience, the 3D Objects Counter of Fiji/ImageJ software works well for counting objects in loose cell clusters, where cell nuclei are located separately and do not merge on optical sections. For medium to large EBs with a large number of cells, this option may not be suitable.

Acknowledgments This work was supported by the Russian Foundation for Basic Research (grant number 06-04-08279). The author thanks A. Gordeev (NLM NIH, USA) for his help with a high content image analysis using the ImageJ/Fiji software.

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References 1. Ebert AD, Svendsen CN (2010) Human stem cells and drug screening: opportunities and challenges. Nat Rev Drug Discov 9:367–372. https://doi.org/10.1038/nrd3000 2. Avior Y, Sagi I, Benvenisty N (2016) Pluripotent stem cells in disease modelling and drug discovery. Nat Rev Mol Cell Biol 17:170–182 3. Rowe RG, Daley GQ (2019) Induced pluripotent stem cells in disease modelling and drug discovery. Nat Rev Genet 20:377–388 4. Lauschke K, Rosenmai AK, Meiser I et al (2020) A novel human pluripotent stem cellbased assay to predict developmental toxicity. Arch Toxicol 94:3831–3846. https://doi.org/ 10.1007/s00204-020-02856-6 5. Dutta D, Heo I, Clevers H (2017) Disease modeling in stem cell-derived 3D organoid systems. Trends Mol Med 23:393–410 6. Takahashi T (2019) Organoids for drug discovery and personalized medicine. Annu Rev Pharmacol Toxicol 59:447–462 7. O’Connell L, Winter DC (2020) Organoids: past learning and future directions. Stem Cells Dev 29:281–289 8. Clevers H (2016) Modeling development and disease with organoids. Cell 165:1586–1597 9. Flamier A, Singh S, Rasmussen TP (2017) A standardized human embryoid body platform for the detection and analysis of teratogens. PLoS One 12:e0171101. https://doi.org/10. 1371/journal.pone.0171101 10. Kim J, Koo BK, Knoblich JA (2020) Human organoids: model systems for human biology and medicine. Nat Rev Mol Cell Biol 21:571–584 11. Kang HY, Choi YK, Jo NR et al (2017) Advanced developmental toxicity test method based on embryoid body’s area. Reprod Toxicol 72:74–85. https://doi.org/10.1016/j. reprotox.2017.06.185 12. Warkus ELL, Yuen AAYQ, Lau CGY, Marikawa Y (2016) Use of in vitro morphogenesis

of mouse embryoid bodies to assess developmental toxicity of therapeutic drugs contraindicated in pregnancy. Toxicol Sci 149:15–30. https://doi.org/10.1093/toxsci/kfv209 13. Gordeeva O, Gordeev A (2021) Comparative assessment of toxic responses in 3D embryoid body differentiation model and mouse early embryos treated with 5-hydroxytryptophan. Arch Toxicol 95:253–269. https://doi.org/ 10.1007/s00204-020-02909-w 14. De Bernardi MA, Hewitt SM, Kriete A (2006) Automated confocal imaging and high-content screening for cytomics. In: Handbook of biological confocal microscopy, 3rd edn. Springer, New York, pp 809–817 15. Sirenko O, Hancock MK, Crittenden C et al (2017) Phenotypic assays for characterizing compound effects on induced pluripotent stem cell-derived cardiac spheroids. Assay Drug Dev Technol 15:280–296. https://doi. org/10.1089/adt.2017.792 16. Chandrasekaran SN, Ceulemans H, Boyd JD, Carpenter AE (2021) Image-based profiling for drug discovery: due for a machine-learning upgrade? Nat Rev Drug Discov 20:145–159 17. Nagy A, Rossant J, Nagy R et al (1993) Derivation of completely cell culture-derived mice from early-passage embryonic stem cells. Proc Natl Acad Sci U S A 90:8424–8428. https:// doi.org/10.1073/pnas.90.18.8424 18. Robertson E, Bradley A, Kuehn M, Evans M (1986) Germ-line transmission of genes introduced into cultured pluripotential cells by retroviral vector. Nature 323:445–448. https:// doi.org/10.1038/323445a0 19. Salic A, Mitchison TJ (2008) A chemical method for fast and sensitive detection of DNA synthesis in vivo. Proc Natl Acad Sci U S A 105:2415–2420. https://doi.org/10. 1073/pnas.0712168105

Methods in Molecular Biology (2022) 2520: 81–100 DOI 10.1007/7651_2021_401 © Springer Science+Business Media, LLC 2021 Published online: 05 May 2021

Derivation of Multipotent Neural Progenitors from Human Embryonic Stem Cells for Cell Therapy and Biomedical Applications Loriana Vitillo and Ludovic Vallier Abstract Long-term neuroepithelial-like stem cells (lt-NES) derived from human embryonic stem cells are a stable self-renewing progenitor population with high neurogenic potential and phenotypic plasticity. Lt-NES are amenable to regional patterning toward neurons and glia subtypes and thus represent a valuable source of cells for many biomedical applications. For use in regenerative medicine and cell therapy, lt-NES and their progeny require derivation with high-quality culture conditions suitable for clinical use. In this chapter, we describe a robust method to derive multipotent and expandable lt-NES based on good manufacturing practice and cell therapy-grade reagents. We further describe fully defined protocols to terminally differentiate lt-NES toward GABA-ergic, dopaminergic, and motor neurons. Key words Human embryonic stem cells, Cell therapy, Differentiation, GMP, Lt-NES, Neural progenitors, Regenerative medicine, Stem cells

1

Introduction In the decades since their first derivation [1], human embryonic stem cells (hESCs) have gone from the theoretical promise of a new unlimited source of human cell types [2] to the reality of the first in human trials based on their derivatives [3, 4]. HESCs have been successfully differentiated into a large collection of cells that are otherwise unavailable or difficult to isolate without ethical challenges, such as human neural progenitors [5–7]. Indeed, hESCbased cell therapy represents a game-changing strategy for neurological diseases involving tissues that are permanently damaged and unable to regenerate, as in the case of multiple sclerosis, Parkinson’s disease, or spinal cord injury [6, 8]. Furthermore, therapeutic advancements crucially depend on a reliable source of high-quality neurons and glia to model the pathophysiology of neurological disorders and drug responses. Long-term neuroepithelial-like stem cells (lt-NES) have been generated from pluripotent stem cells as a viable alternative to fetal tissue. Their extensive self-renewal capacity, stable karyotype, and

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phenotypic plasticity provide an advantageous platform to model a multitude of disease and for biomedical applications [9–14]. Lt-NES retain multipotency as a rosette-type population when in the presence of EGF and FGF growth factors while differentiate with a bias for GABAergic neurons upon growth factors withdrawal [9, 10]. Furthermore, upon addition of patterning cues, they can be directed toward midbrain dopaminergic, motoneuron, or astroglia phenotypes [9–11]. Despite the tremendous applications offered by lt-NES in vitro, fully defined sets of highly qualified good manufacturing practice (GMP)-grade protocols based on cell therapy-grade reagents are required for regulatory compliance and in order to deploy them for clinical use [15–17]. In this chapter, we describe a step-by-step method (Fig. 1) for the derivation, maintenance, and differentiation of lt-NES based on GMP-grade and cell therapy-grade reagents including substrates and growth factors [18]. In particular, we describe methods to terminally differentiate lt-NES into GABAergic neuron and into midbrain dopaminergic neurons or motoneurons phenotypes (Fig. 1). In essence, this protocol involves as a first step the formation of same-sized embryoid bodies from pluripotent hESCs (step 1), followed by neural induction resulting in the formation of neural rosettes (step 2). Lt-NES are then isolated from these neural rosettes to be passaged at high density in the presence of EGF and FGF to promote self-renewal (step 3). Once established, lt-NES lines are maintained in continuous presence of EGF and FGF (step 4) while spontaneous and terminal neural differentiation is induced by the withdrawal of EGF and FGF (step 5). Patterning cues involving addition of FGF8b and SHH direct lt-NES to a midbrain dopaminergic phenotype after 21 days (Fig. 1, step 6). Patterning cues involving retinoid acid and SHH direct lt-NES toward motoneuron phenotype after 21 days of treatment (step 7). Overall, this method provides a GMP and cell therapy-grade platform to generate multipotent and bankable lt-NES amenable for downstream differentiation into clinically useful cell types.

2

Materials All the cell culture reagents are stored, reconstituted, and used as specified in the manufacturer instruction unless stated. Procedures are performed at room temperature (RT) unless otherwise specified.

2.1 Reagents Grade and Quality Specifications

1. The reagents used in this protocol are of cGMP quality, unless stated, regardless of this being written in their product name. Refer to each manufacturer for specific details on documentation and quality specifications provided.

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Fig. 1 Schematic overview of the step-by-step protocol. HESCs are dissociated to generate embryoid bodies of the same size in a microwell format (step 1). After 5 days, the formed EBs are dislodged from the microwell and plated for neural induction (step 2). Neural induction will follow for up to 5 days when characteristic neural rosettes morphologies appear by microscopic inspection, at which point they are ready for purification (step 3). Scalable expansions of dissociated rosettes in the presence of EGF and FGF leads to the establishment of a pure population of lt-NES, which is self-renewing, bankable, and multipotent (step 4). The downstream differentiation of lt-NES depends on the chosen patterning conditions, with a high neurogenic potential toward GABAergic neurons upon EGF and FGF withdrawal (step 5). Specific patterning conditions including FGF8b and SHH lead lt-NES toward dopaminergic phenotypes (step 6). On the contrary, with patterning media containing retinoid acid and SHH, lt-NES are redirected toward motoneurons phenotypes (step 7). (Figure created with BioRender.com)

2. Cell Therapy Systems (CTS™) reagents are GMP-grade, serum-free, xeno-free, and animal origin-free formulations. Moreover, they have cell and gene therapy-specific intended use statements. https://www.thermofisher.com/uk/en/ home/clinical/cell-gene-therapy/cell-therapy/cell-therapysystems.html. 3. GMP growth factors from R&D are produced following cGMP guidelines “that allow for their use as ancillary materials in cell therapy or for further manufacturing processes. GMP proteins also come with extensive documentation and traceability, as well as additional quality control testing.” https://www. rndsystems.com/products/good-manufacturing-practicesgmp-grade-proteins.

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4. Laminin 521 substrate is available as cell therapy grade (CTG) from the Biolamina manufacturer. This product is a seamless transition from the Laminin 521 of the same company used in this protocol. Notably, the CTG grade was in the pipeline at the time of this protocol development and was chosen in view of the availability of this grade as well as its high performance. 5. StemPro Accutase and STEMdiff™ Neural Rosette Selection Reagent would need custom due diligence qualification for therapy usage. STEMdiff™ Neural Rosette Selection Reagent is enzyme and animal derivative-free. The manufacturer is committed to working with cell therapy developers to support the use of this product in cell therapy manufacture and can assist with performing due diligence for specific regulators. Overall, these reagents provide the best solution to date for effectiveness and clinical compliance. 2.2 Equipment and Consumables

1. Class II biological safety cabinet. 2. Inverted microscope. 3. Incubator set at 5% CO2, 5% O2, and 37  C. 4. Fridge (4  C)/freezers ( 20  C/ 80  C). 5. Centrifuge fitted with 15-ml tubes holders. 6. Centrifuge with swinging rotor fitted with plate holder (capacity up to 2000  g). 7. Thermostat filled with beads (i.e., LabArmor Beads, Thermo Scientific) at 37  C. 8. Pipettor. 9. Hemocytometer. 10. Sterile and individually plastic-wrapped serological stripettes. 11. Pipettes and sterile tips (suitable for p1000, p200, p10). 12. Corning® CoolCell™ (CLS432001, SDS).

LX

Cell

Freezing

Container

13. Sterile 6-, 12-, 24-, and 48-well cell culture plates of polystyrene. 14. Sterile 50- and 15-ml tubes. 15. Sterile polypropylene 0.5- and 1.5-ml tubes. 16. Sterile 2-ml cryovials. 17. AggreWell™ 800 (27865, StemCell Technologies). 18. 1000 μl Filter Tip, Wide Orifice, Sterile, Racked (E1011-9618, Starlab UK). 19. 37 μm Reversible Strainer (27215, StemCell Technologies).

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Cell Culture

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1. CTS™ VTN-N Vitronectin (A27940, Life Technologies). 2. CTS™ Essential 8™ (A2656101, Life Technologies). 3. UltraPure™ 0.5 M EDTA (15575020, Invitrogen). 4. CTS™ DPBS

/

(A1285601, Life Technologies).

5. StemPro Accutase (A1110501, Life Technologies). 6. Essential 6 (A1516401, Life Technologies). 7. CTS™ DPBS+/+ (A1285801, Life Technologies). 8. CTS™ DMEM-F12 (A1370801, Life Technologies). 9. CTS™ N2 (A1370701, Life Technologies). 10. CTS™ B27 (A1486701, Life Technologies). 11. CTS™ Glutamax (A12860-01, Life Technologies). 12. Laminin 521 (LN521, Biolamina). 13. STEMdiff™ Neural Rosette Selection Reagent (5832, StemCell Technologies). 14. GMP FGF (233-GMP, Bio-Techne). 15. GMP EGF (236-GMP, Bio-Techne). 16. CryoStem (K1-0640, Geneflow). 17. Revitacell supplement (A2644501, Life Technologies). 18. Sterile water (10114292, Gibco™). 2.4 Additional Reagents for Spontaneous lt-NES Differentiation

1. CTS™ Neurobasal (A1371001, Life Technologies).

2.5 Additional Reagents for lt-NES Differentiation to Dopaminergic Neurons

1. CTS™ Neurobasal (A1371001, Life Technologies).

2. cAMP (A9501, Sigma Aldrich).

2. GMP Sonic Hedgehog (SHH, 130-095-727, Miltenyi Biotec). 3. GMP FGF8b (130-095-740, Miltenyi Biotec). 4. GMP GDNF (212-GMP-010, Bio-Techne). 5. GMP BDNF (248-GMP-025, Bio-Techne). 6. Dibutyryl-cAMP (D0260-25 mg, Sigma Aldrich). 7. Ascorbic acid (95210-250G, Sigma Aldrich).

2.6 Additional Reagents for lt-NES Differentiation to Motoneurons

1. CTS™ Neurobasal (A1371001, Life Technologies). 2. GMP Sonic Hedgehog (SHH, 130-095-727, Miltenyi Biotec). 3. GMP GDNF (212-GMP-010, Bio-Techne). 4. GMP BDNF (248-GMP-025, Bio-Techne). 5. cAMP (A9501, Sigma Aldrich). 6. Retinoid Acid (R2625, Sigma Aldrich).

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2.7 Coating Cell Culture Plates with VTN-N

1. Thaw one VTN-N vial at room temperature and aliquot in 60 μl volume in sterile polypropylene tubes. Store aliquots at 80  C. 2. Dilute the appropriate volume of VTN-N stock in CTS™ DPBS / based on the number of plates required to a concentration of 0.5 μg/cm2. 3. Add 6 ml of CTS™ DPBS 60 μl of VTN-N.

/

to a 15-ml tube followed by

4. Gently resuspend the VTN-N solution with a 5-ml stripette. 5. Add 1 ml of VTN-N solution to each well of a 6-well plate. 6. Swirl plate to assure even coverage. 7. Incubate at room temperature for 1 h. 8. Aspirate the diluted VTN-N solution from the wells prior to use (see Note 1). 2.8 Coating Cell Culture Plates with Laminin 521

1. Thaw the laminin 521 stock at 2–8  C. Once thawed is stable at 2–8  C for up to 3 months. 2. Dilute the appropriate volume of laminin 521 stock in CTS™ DPBS+/+ based on the number of plates required to a concentration of 10 μg/ml (see Note 2). 3. Incubate for 3 h at 37  C or overnight at 4  C with the plate sealed with parafilm to avoid evaporation. Plates can be kept at 4  C up to 4 weeks. 4. Visualize if the coating solution still covers the entire surface, particularly the center of the well, since after prolonger storage some coating may have evaporated. 5. The plates are ready to use, prior removal of the laminin 521 coating.

2.9 Preparation of N2 Base Differentiation Medium (N2 Medium)

1. Thaw CTS™ N2 and CTS™ B27 at room temperature. 2. Remove 10.5 ml of media from the CTS™ DMEM/F12 500 ml bottle. 3. Add 5 ml of CTS™ N2. 4. Add 500 μl of CTS™ B27. 5. Add 5 ml of Glutamax. 6. Mix well the components by inverting the bottle upside down. 7. Store media at 4  C up to 3 weeks. It is recommended to make 50 ml aliquots for the weekly requirement. 8. Always warm the media at room temperature 20 min before use.

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All growth factors and molecules must be stored according to the manufacturer’s instructions. In this protocol, we recommend reconstituting with the following solvents and final stock concentrations: to 10 μg/ml

1. EGF: Reconstitute 200 μg in CTS™ DPBS final stock concentration.

/

2. FGF: Reconstitute 25 μg in CTS™ DPBS stock concentration.

to 4 μg/ml final

/

3. FGF8b: Reconstitute 25 μg in CTS™ DPBS / to 100 μg/ml final stock concentration. 4. BDNF: Reconstitute 25 μg in CTS™ DPBS final stock concentration.

/

to 100 μg/ml

5. GDNF: Reconstitute 10 μg in CTS™ DPBS final stock concentration.

/

to 100 μg/ml

6. SHH: Reconstitute 25 μg in sterile water to 250 μg/ml final stock concentration. 7. Ascorbic acid: Reconstitute 50 mg in sterile water to 200 mM final stock concentration. 8. cAMP: Reconstitute 1 g in sterile water to 300 μg/ml final stock concentration. 9. dy-cAMP: Reconstitute 25 mg in sterile water to 50 mM final stock concentration. 10. Retinoid acid: Reconstitute in DMSO to 3 mM final stock concentration.

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Methods Perform all work at room temperature unless specified.

3.1 HESCs Maintenance, Passaging, Freezing

1. HESCs are maintained in Essential 8™ (E8) medium on VTNN-coated plates with an EDTA passaging method, performed without centrifugation (see Note 3). 2. Passage the cells when they are 70–80% confluent. 3. Wash the cells once with DPBS

/

.

/

4. Aspirate the DPBS and replace with 0.5 mM EDTA solution (prepared in DPBS / ). 5. Incubate the cells at room temperature for 1–2 min. 6. Aspirate the EDTA solution entirely when the cells start to round up, and the colonies appear to have holes in them. 7. Add 1 ml/well of E8 with a p1000 tip to gently detach cells as small clumps. 8. Add cell suspension in a 15-ml tube.

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9. Repeat steps 7 and 8 until the majority of the cells are collected. 10. Gently mix the cell suspension with a 5 or 10-ml stripette. 11. Resuspend the cells in half of the final volume of media needed for a 1:6 passaging ratio. 12. Remove DPBS / from the VTN-N-coated plates and replace with 1 ml/well (for a 6-well plate format) of E8 medium. 13. Add 1 ml/well of cell suspension drop-by-drop across each well. 14. Move the plate in the incubator and assure an even distribution of the cells by rocking side to side and top to bottom. Do not move for 24 h. 15. Replace E8 medium after 24 h and feed daily afterwards. 16. Freezing: cells are frozen in CryoStem and thawed in E8 plus GMP ROCK inhibitor Revitacell added to the media for the first 24 h. 3.2 Embryoid Body Formation (Day 0)

1. HESCs should be checked for suitability for differentiation by ensuring a level of spontaneous differentiation that is less than 2–5% of the entire culture. 2. The protocol requires two wells of 80% confluent hESCs in a 6-well plate format. 3. Prepare 5 ml of E6 medium plus 50 μl of Revitacell. 4. Prepare Aggrewell for cell seeding: (a) Wash a single well of Aggrewell with 2 ml of E6 media. (b) Aspirate the media and add 500 μl of E6 plus Revitacell to the well. (c) Seal the plate with parafilm and centrifuge at 2000  g for 5 min in a centrifuge fitted with plate holder. (d) Check under an inverted microscope that the microwells inside the Aggrewell do not contain bubbles. 5. Aspirate the E8 medium from two wells of hESCs in a 6-well plate and proceed for each well with the following steps (6–11). /

6. Add 1 ml of DPBS coverage. 7. Aspirate the DPBS

/

and swirl gently to ensure even

.

8. Add 1 ml of Accutase and swirl gently to ensure even coverage. 9. Incubate the wells at 37  C for 1–2 min until the cells start to detach but are not all floating in suspension. 10. Neutralize the Accutase by adding 1 ml of E6 media to each well.

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11. Pipette with a p1000 tip to collect all cells and possibly attached colonies. 12. Pull together the cell suspension from the two 6-wells into a 15-ml tube containing 8 ml of E6. 13. Pipette up and down 5–7 times with a 5-ml stripette in order to produce single-cell suspension (see Note 4). 14. Remove 10 μl of cell suspension and place it in a hemocytometer for cell counting. 15. Centrifuge the 10 ml cell suspension at 300  g for 5 min. 16. Count cells by averaging the number of cells in the 16 squares of four quadrants of a hemocytometer and multiply by 104 and then 10 ml, to obtain the total cell number. It is expected to obtain at least 4  106 cells. 17. Aspirate the supernatant and resuspend the cell pellet into 1 ml of E6 plus Revitacell with a p1000 pipette and mix well. 18. Based on the total cell count previously calculated, remove the appropriate volume of cell suspension corresponding to 3  106 cells and place it into a separate 15-ml tube. 19. Top up to 1.5 ml with E6 plus Revitacell. 20. Add 1.5 ml of cell suspension to the Aggrewell containing 500 μl of E6 plus Revitacell with a p1000 tip by gently pipetting few times to disperse the cells. 21. Inspect the Aggrewell at the microscope to assure cells are evenly distributed. 22. Seal the plate carefully with parafilm and centrifuge at 100  g for 3 min with a corresponding plate balance. 23. Remove the parafilm and inspect under the microscope to confirm that the cells are concentrated at the bottom of the microwell. 24. Incubate at 37  C for 24 h. 25. Day 1. Remove all the media carefully with a p1000 tip and replace with 2 ml of fresh E6 for a full media change (see Note 5). 26. Incubate at 37  C for 24 h. 27. Day 2 till day 4. Feed the EBs with half-media change by gently replacing only 1 ml of E6 per well. Incubate at 37  C for 24 h. 3.3 Neural Induction (Day 5)

For each EB preparation (one well of Aggrewell plate), one well of a 6-well plate coated with laminin 521 needs to be ready at day 5. 1. On day 5 of EB formation, prepare a 15-ml tube fitted with a 37 μm reversible strainer with arrow pointing up. 2. Prepare N2 base medium.

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3. With a standard p1000 tip, remove 1 ml of medium from the Aggrewell containing EBs and expel it back into the Aggrewell with vigor in order to detach EBs (see Note 6). 4. Use a large bore p1000 tip to collect all the media containing floating, and visible, EBs, and slowly add this suspension to the top of the cell strainer. 5. Repeat steps 3 and 4 to collect all EBs (see Note 7). 6. Remove the Laminin 521 coating solution from a precoated 6-well plate and place the cell strainer upside-down (arrow pointing down) on top of the corner of the well by holding it with one hand. 7. Add 1 ml of N2 base media with a standard p1000 tip slowly around the top of the cell strainer to release the EBs into the well. 8. Repeat step 7 in order to release all EBs in the well. 9. Incubate at 37  C to start neural induction and do not disturb for 24 h (see Note 8). 10. Day 1 of neural induction: Using a stripette, replace media with 2 ml of fresh N2 base medium. 11. Day 2 of neural induction: Using a stripette, replace media with 2 ml of fresh N2 base medium. 3.4 Lt-NES Derivation (Day 8/10)

1. Neural rosettes should be derived between day 3 and day 5 of neural induction when the rosettes appear, at microscopic inspection, to be flat, round-shaped, and with lumens, while the confluence of the surrounding differentiated cells has not yet reached 80% of the entire plate. 2. Prepare, or have already, at least 4 wells of a 48-well plate coated with Laminin 521. 3. Remove the N2 base medium from the rosette well. 4. Add 1 ml of rosette selection solution and incubate for 45 min at 37  C (see Note 9). 5. Remove the rosette selection solution entirely. 6. Using a p1000 tip firmly release 1 ml of N2 media on top of the rosette clusters (visible cell clumps by naked eye). 7. With the same tip, aspirate the media containing the rosettes and transfer it to a 15-ml tube (see Note 10). 8. Repeat step 5 and 6 until 70% of rosettes have been removed (see Note 11). 9. Spin the rosette suspension at 300  g for 5 min. 10. Aspirate supernatant.

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11. Suspend the rosettes pellet in 400 μl of N2 media plus 10 ng/ ml of EGF and FGF (named N2 EF) supplemented also with Revitacell (see Note 12). 12. Distribute the rosette suspension equally in 4 wells of a 48-well plate coated with Laminin 521. 13. Top up each well to a final volume of 500 μl with N2 EF media plus Revitacell. 14. Monitor cell attachment after 10 min and around 5–6 h (see Note 13). 15. After 24 h, perform a media change with fresh N2 EF media (see Note 14). 16. Prepare, or have already, at least 4 wells of a 24-well plate coated with Laminin 521. 17. On day 2, if lt-NES are 100% confluent, proceed with a p0 to p1 expansion from a 48- to a 24-well format, for each of the wells, as follows: (a) Remove media from one well of the 48-well plate. (b) Add 300 μl of Accutase and incubate at 37  C for 1 min. (c) Once the cells have started to look round but are not floating, use a p1000 to aspirate Accutase and pipette back on the cells to physically detach them. (d) Transfer the lt-NES suspension to a 15-ml tube containing 10 ml N2 media. (e) Centrifuge at 300  g for 3 min. (f) Aspirate supernatant and resuspend the lt-NES into 600 μl of N2 EF plus Revitacell. (g) Plate lt-NES into a single well of a 24-well format coated with Laminin 521. 18. Daily feed p1 lt-NES with 0.6 ml of N2 EF until reaching 100% confluency (see Note 15). 19. Keep one well of p1 lt-NES cell as security or for early stock (see Note 16). 20. Prepare, or have already, at least three wells of a 12-well plate coated with Laminin 521. 21. Once p1 lt-NES are 100% confluent, proceed with a p1 to p2 expansion from a 24- to a 12-well format, for each of the wells, as follows: (a) Remove media from one 24-well vessel. (b) Add 500 μl of Accutase and incubate at 37  C for 1 min. (c) Once the cells have started to look round but are not floating, use a p1000 to aspirate Accutase and pipette back on the cells to physically detach them.

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(d) Transfer the lt-NES suspension to a 15-ml tube containing 10 ml N2 media. (e) Centrifuge at 300  g for 3 min. (f)

Aspirate supernatant and resuspend the lt-NES into 1 ml of N2 EF plus Revitacell.

(g) Plate lt-NES into a single well of a 12-well plate coated with Laminin 521. 22. Daily feed p2 lt-NES with 1 ml of N2 EF until reaching 100% confluency. 23. Prepare, or have already, at least six wells of a 12-well plate coated with Laminin 521. 24. Once p2 lt-NES are 100% confluent, proceed with a p2 to p3 1:2 expansion in a 12-well format, for each of the wells, as follows: (a) Remove media from one 12-well vessel. (b) Add 500 μl of Accutase and incubate at 37  C for 1 min. (c) Once the cells have started to look round but are not floating, use a p1000 to aspirate Accutase and pipette back on the cells to physically detach them. (d) Transfer the lt-NES suspension to a 15-ml tube containing 10 ml N2 media. (e) Centrifuge at 300  g for 3 min. (f) Aspirate supernatant and resuspend the lt-NES into 2 ml of N2 EF plus Revitacell. (g) Plate lt-NES into two wells of a 12-well format coated with Laminin 521. 25. Daily feed p3 lt-NES with 1 ml of N2 EF until reaching 100% confluency. 26. Prepare, or have already, at least three wells of a 6-well plate coated with Laminin 521. 27. Once p3 lt-NES are 100% confluent, proceed with a p3 to p4 expansion from a 12- to a 6-well format, for two of the wells, as follows: (a) Remove media from two 12-well vessels. (b) Add 500 μl of Accutase and incubate at 37  C for 1 min. (c) Once the cells have started to look round but are not floating, use a p1000 to aspirate Accutase and pipette back on the cells to physically detach them. (d) Transfer the lt-NES suspension to a 15-ml tube containing 10 ml N2 media. (e) Centrifuge at 300  g for 3 min.

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(f) Aspirate supernatant and resuspend the lt-NES into 2 ml of N2 EF plus Revitacell. (g) Plate lt-NES into one well of a 6-well format coated with Laminin 521. 28. Daily feed p4 lt-NES with 2 ml of N2 EF until reaching 100% confluency. 29. Once p4 lt-NES are 100% confluent, proceed with a 1:2 passage in a 6-well format (see Note 17). (a) Remove media from one 6-well vessel. (b) Add 1 ml of Accutase and incubate at 37  C for 1 min. (c) Once the cells have started to look round but are not floating, use a p1000 to aspirate Accutase and pipette back on the cells to physically detach them. (d) Transfer the lt-NES suspension to a 15-ml tube containing 10 ml N2 media. (e) Centrifuge at 300  g for 3 min. (f) Aspirate supernatant and resuspend the lt-NES into 4 ml of N2 EF plus Revitacell. (g) Plate lt-NES into two wells of a 6-well format coated with Laminin 521. 30. Daily feed p5 lt-NES in N2 EF until 100% confluency is reached. 31. Once lt-NES reach a comfortable expansion in a 6-well plate format, they are ready for characterization (see Note 18). 3.5 Lt-NES Maintenance

Passaging

1. Lt-NES are split at a ratio of 1:2 or 1:3 every 3 days when culture is 95–100% confluent. 2. Remove media from one 6-well vessel. 3. Add 1 ml of Accutase and incubate at 37  C for 1 min. 4. Once the cells have started to look round but are not floating, use a p1000 to aspirate Accutase and pipette back on the cells to physically detach them. 5. Transfer the lt-NES suspension to a 15-ml tube containing 10 ml N2 media. 6. Centrifuge at 300  g for 3 min. 7. Aspirate supernatant and resuspend the lt-NES into 4 ml (or 6 ml) of N2 EF. 8. Plate lt-NES into two (or three wells) of a 6-well format coated with Laminin 521.

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9. Perform a media change with N2 EF (10 ng/ml) every day from passage 0 to passage 10 and every other day from passage 10 onwards. Freezing

1. Remove media from one 6-well vessel. 2. Add 1 ml of Accutase and incubate at 37  C for 1 min. 3. Once the cells have started to look round but are not floating, use a p1000 to aspirate Accutase and pipette back on the cells to physically detach them. 4. Transfer the lt-NES suspension to a 15-ml tube containing 10 ml N2 media. 5. Centrifuge at 300  g for 3 min. 6. Aspirate supernatant and resuspend the lt-NES pellet into 1 ml Cryostem freezing medium. 7. Immediately transfer cell suspension into a labeled cryovial and place into a CoolCell freezing device at 80  C for 24 h. 8. After 24 h, cells can be removed from the CoolCell device and transferred to liquid or vapor phase nitrogen tanks for longterm storage. Thawing

1. Remove one cryovial from the liquid nitrogen tank and place immediately into a dry ice box. 2. Place the cryovial to thaw in a bead-filled thermostat. Keep the vial at 37  C until visibly there is only a 1/10 of the solution in the solid state, in the form of a small piece of ice. 3. Remove the cell solution from the cryovial with a 5-ml stripette and transfer it into a 15-ml tube containing 10 ml of pre-warmed N2 base media. 4. Centrifuge at 300  g for 3 min. 5. Aspirate supernatant and resuspend the lt-NES pellet into 2 ml of N2 EF plus Revitacell. 6. Plate lt-NES into one well of a 6-well format coated with Laminin 521. 7. After 24 h remove N2 media containing Revitacell and add fresh N2 EF medium. 8. Place the cell culture 37  C in the incubator. 3.6 Lt-NES Spontaneous Differentiation

1. Remove media from one 6-well vessel containing lt-NES. 2. Add 1 ml of Accutase and incubate at 37  C for 1 min. 3. Once the cells have started to look round but are not floating, use a p1000 to aspirate Accutase and pipette back on the cells to physically detach them.

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4. Transfer the lt-NES suspension to a 15-ml tube containing 10 ml N2 media. 5. Remove 10 μl of suspension and count the cells by using a hemocytometer. 6. Centrifuge at 300  g for 3 min to pellet the cells. 7. Calculate the volume of cell suspension required to seed lt-NES at 40,000 cells/cm2. 8. Seed cell on wells (48-well format or higher, depending on the experimental design) coated with Laminin 521 in N2 media without EF. 9. After 24 h, remove N2 media. 10. Prepare terminal differentiation media by mixing 50:50 parts of DMEM-F12 (with N2 1:100): Neurobasal (with B27 1:50). 11. Add terminal differentiation media plus freshly added 300 ng/ ml cAMP. This constitutes day 1 of the terminal differentiation. 12. Daily feed the cells with differentiation medium plus cAMP till day 21. 13. Cells are ready for characterization from day 21 (see Note 19). 3.7 Lt-NES-Directed Differentiation Toward Dopaminergic Neurons

1. Remove media from one 6-well vessel containing lt-NES. 2. Add 1 ml of Accutase and incubate at 37  C for 1 min. 3. Once the cells have started to look round but are not floating, use a p1000 to aspirate Accutase and pipette back on the cells to physically detach them. 4. Transfer the lt-NES suspension to a 15-ml tube containing 10 ml N2 media. 5. Remove 10 μl of suspension and count the cells by using a hemocytometer. 6. Centrifuge at 300  g for 3 min to pellet the cells. 7. Calculate the volume of cell suspension required to seed lt-NES at 40,000 cells/cm2. 8. Seed cell on wells (48-well format or higher, depending on the experimental design) coated with Laminin 521 in N2 media without EF. 9. After 24 h, remove N2 media. 10. Day 1: Change the medium to patterning medium composed of DMEM-F12 (with N2 1:100) medium plus freshly added 200 ng/ml SHH, 100 ng/ml FGF8, and 160 μM ascorbic acid. 11. Daily feed the cells with patterning medium till day 13. 12. Day 14: Change medium to dopaminergic terminal differentiation medium composed of DMEM-F12 (N2 1:100):

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Neurobasal (B27 1:50) plus freshly added 20 ng/ml BDNF, 10 ng/ml GDNF, 160 μM ascorbic acid, 500 μM dy-cAMP. 13. Daily feed neurons with dopaminergic terminal differentiation medium till day 21. 14. From day 21, the neurons are ready for immunofluorescence characterization, although differentiation can continue longer in the same medium for long-term functionality and maturity studies (see Note 20). 3.8 Lt-NES-Directed Differentiation Toward Motoneurons

1. Remove media from one 6-well vessel. 2. Add 1 ml of Accutase and incubate at 37  C for 1 min. 3. Once the cells have started to look round but are not floating, use a p1000 to aspirate Accutase and pipette back on the cells to physically detach them. 4. Transfer the lt-NES suspension to a 15-ml tube containing 10 ml N2 media. 5. Remove 10 μl of suspension and count the cells by using a hemocytometer. 6. Centrifuge at 300  g for 3 min to pellet the cells. 7. Calculate the volume of cell suspension required to seed lt-NES at 40,000 cells/cm2. 8. Seed cell on wells (48-well format or higher, depending on the experimental design) coated with Laminin 521 in N2 media without EF. 9. After 24 h, remove N2 media. 10. Day 1: Change the medium to patterning medium composed of DMEM F12 (With N2 1:100, B27 1:50) plus 10 ng/ml EGF, 10 ng/ml FGF, and 1 μM retinoid acid. 11. Daily feed cells with patterning medium till day 5. 12. Day 5: Supplement the patterning medium with 1 μg/ml GMP SHH. 13. Day 6: Feed cells with patterning medium plus 1 μg/ml GMP SHH. 14. Day 7: Feed cells with patterning medium modified by removing EGF and FGF and reducing the concentration of retinoid acid to 0.01 μM. 15. Day 8 till day 11: Feed cells with patterning medium as per step 9. 16. Day 12: Change feeding media to motoneuron terminal differentiation media composed of equal parts of DMEM-F12 (N2 1:100): neurobasal (B27 1:50) plus 20 ng/ml GMP BDNF, 20 ng/ml GMP GDNF, 50 ng/ml SHH, and 300 ng/ml cAMP.

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17. Daily feed neurons with motoneuron terminal differentiation medium till day 21. 18. From day 21, the neurons are ready for immunofluorescence characterization, although differentiation can continue longer in the same medium (see Note 21).

4

Notes 1. It is important that wells do not dry; therefore, it is recommended to add Essential 8™ Medium to the wells as soon as the Vitronectin solution is removed. Culture vessels coated with Vitronectin can be stored at 2–8  C wrapped in parafilm for up to 2 days. However, it is recommended that the plates are coated on the day of use. 2. Instruction for one well of several formats. 6-well: 100 μl L521 to 900 μl of CTS™ DPBS+/+. 12-well: 50 μl L521 to 450 μl of CTS™ DPBS+/+. 24-well: 30 μl L521 to 270 μl of CTS™ DPBS+/+. 48-well: 17.5 μl L521 to 157.5 μl of CTS™ DPBS+/+. 96-well: 6 μl L521 to 54 μl of CTS™ DPBS+/+. 3. Media should be changed every day; it is acceptable once a week to add double the quantity of media and leave the cells without media change for 48 h. 4. Do not overly stress cells as this will affect their survival, over pipetting will result in cell death and hamper the quality of EBs in the subsequent steps. 5. After 24 h, the cells should have aggregated in EB-like shape, and the first media change is performed. Remove media very gently, without touching the microwells. Likewise, addition of the media needs to be performed very slowly by touching the border of the well plastic with the top of the tip. Failure to be gentle at this step will dislodge EBs form the microwells with the result of having more than 1 EB per microwell or loss of EBs. 6. Do not aspirate back the suspension with the standard p1000. 7. Do not pipette EBs up and down in the Aggrewell, simply do many more rounds if not all the EBs appear to be detached once inspected under microscope. 8. Neural rosettes, characteristic morphologies indicating a successful neural induction, will start to emerge around day 2–3 in the center of the EBs. Sometimes rosettes are already visible after 24 h, but these are at the earliest stages and maturation generally takes up to 3 days. Neural rosette should be visible with a 2 to 10 magnification in an inverted microscope.

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9. The solution will promote detachment of neural rosettes. Time of incubation may require optimization based on the cell line used, from 45 min up to 1.15 h. 10. Do not reuse the media containing rosettes to target new areas. 11. It is important to check under the microscope that only the rosettes have been removed while the surrounding cells remain largely attached as these non-progenitors cells can affect the derivation of pure lt-NES. 12. Assess if there are large clusters of fully formed round rosettes, if so, pipette with a p200 firmly until producing clusters of around 5–20 cells and some single cells. Do not pipette to produce a 100% single-cell suspension, the ideal preparation will have small aggregates and also single cells. 13. One of the main challenges in the derivation of lt-NES is the proper attachment of the cells to the matrix. Improper coating can lead to poor attachment, visible already after a couple of hours from plating. Lt-NES cells generally attach to the laminin after 5–10 min from plating and within 6 h start to spread to the surface. If clumps or round cells are visible, remove the cells (with gentle pipetting or short addition of Accutase) and place in a new laminin plate. Monitor attachment of lt-NES and prepare extra reserve plates with laminin made on different days. Batch testing of laminin lots is highly recommended. 14. Cells should be between 90% and 100% confluent. Inspect under the microscope in phase contrast for the presence of typical neural rosette progenitors morphology. Due to high cell density, the right rosette morphology may also not be visible as cells are too compacted and rounded. If cells are on top of each other, they need passaging 1:2 into a 48-well format. 15. This can vary from 24 h to 2 days. Typical lt-NES morphology should be visible at this stage. 16. Plan ahead a freeze of passage 1 and following early passages of lt-NES, even from a 48-well format. This will save work if lt-NES do not attach to the laminin for various reasons and also guarantee an early stock. 17. It is highly recommended to freeze 1:1 one 6-well of confluent lt-NES at this passage, and split 1:2 the other well. 18. Immunofluorescence for rosette and lt-NES markers can be performed with the following antibodies: SOX2 (1:100, Bio-techne); Nestin (1:100, 10C2, Abcam); PAX6 (1:100, Cambridge Bioscience); DACH1 (1:100, Proteintech); PLZF (1:100, Life Technologies); ZO-1 (1:50, Bio-techne). Gene expression assessment of lt-NES markers can be performed

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with the following genes (Taq man assays from Life Technologies): SOX2 (Hs01053049_s1); PLZF (Hs00957433_m1); PLAGL1 (Hs00414677_m1); MMNR1 (Hs00201182_m1); DACH1 (Hs00362088_m1); NOTCH1 (Hs01062014_m1); PAX6 (Hs00240871_m1); SOX1 (Hs01057642_s1); NANOG (negative) (Hs04260366_g1); PBGD (housekeeping gene) (Hs00609296_g1). 19. Spontaneously differentiated lt-NES are positive for the following markers by immunofluorescence: GABA (1:500, Sigma), MAP2 (1:500, Sigma), Tubulin TuJ1 clone (1:1000, Biolegend). 20. After 21 days, lt-NES are positive for the following dopaminergic markers by immunofluorescence: Tyrosine Hydroxylase (1:200, Millipore); Tubulin β3 [Clone: TUJ1] (1:1000, Biolegend); FOXA2 (1:100, Santa Cruz); LMX1alfa (1:500, Millipore); GIRK2 (1:500, Millipore); Nurr1 (1:500, Santa Cruz). 21. Motoneurons immunofluorescence markers: Tubulin β3 [Clone: TUJ1] (1:1000, Biolegend), HB9 (Insight Biotechnology).

Acknowledgments This work was supported with a UK Regenerative Medicine Platform grant funded by the Medical Research Council, the Biotechnology and Biological Sciences Research Council, the Engineering and Physical Sciences Research Council, the European Research Council Grant New-Chol (L.V.), the Cambridge Hospitals National Institute for Health Research Biomedical Research Center (L.V.), and a core support grant from the Wellcome Trust and Medical Research Council to the Wellcome Trust—Medical Research Council Cambridge Stem Cell Institute. References 1. Thomson JA, Itskovitz-Eldor J, Shapiro SS, Waknitz MA, Swiergiel JJ, Marshall VS, Jones JM (1998) Embryonic stem cell lines derived from human blastocysts. Science 282 (5391):1145–1147. https://doi.org/10. 1126/science.282.5391.1145 2. Trounson A, Pera M (2001) Human embryonic stem cells. Fertil Steril 76(4):660–661. https://doi.org/10.1016/s0015-0282(01) 02880-1 3. Kobold S, Guhr A, Mah N, Bultjer N, Seltmann S, Seiler Wulczyn AEM, Stacey G, Jie H, Liu W, Loser P, Kurtz A (2020) A manually curated database on clinical studies involving cell products derived from human

pluripotent stem cells. Stem Cell Rep 15 (2):546–555. https://doi.org/10.1016/j. stemcr.2020.06.014 4. Deinsberger J, Reisinger D, Weber B (2020) Global trends in clinical trials involving pluripotent stem cells: a systematic multi-database analysis. NPJ Regen Med 5:15. https://doi. org/10.1038/s41536-020-00100-4 5. Rossi F, Cattaneo E (2002) Opinion: neural stem cell therapy for neurological diseases: dreams and reality. Nat Rev Neurosci 3 (5):401–409. https://doi.org/10.1038/ nrn809 6. De Luca M, Aiuti A, Cossu G, Parmar M, Pellegrini G, Robey PG (2019) Advances in

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stem cell research and therapeutic development. Nat Cell Biol 21(7):801–811. https:// doi.org/10.1038/s41556-019-0344-z 7. Bernal A, Arranz L (2018) Nestin-expressing progenitor cells: function, identity and therapeutic implications. Cell Mol Life Sci 75 (12):2177–2195. https://doi.org/10.1007/ s00018-018-2794-z 8. Barker RA, Parmar M, Studer L, Takahashi J (2017) Human trials of stem cell-derived dopamine neurons for Parkinson’s disease: dawn of a new era. Cell Stem Cell 21(5):569–573. https://doi.org/10.1016/j.stem.2017.09. 014 9. Koch P, Opitz T, Steinbeck JA, Ladewig J, Brustle O (2009) A rosette-type, self-renewing human ES cell-derived neural stem cell with potential for in vitro instruction and synaptic integration. Proc Natl Acad Sci U S A 106 (9):3225–3230. https://doi.org/10.1073/ pnas.0808387106 10. Falk A, Koch P, Kesavan J, Takashima Y, Ladewig J, Alexander M, Wiskow O, Tailor J, Trotter M, Pollard S, Smith A, Brustle O (2012) Capture of neuroepithelial-like stem cells from pluripotent stem cells provides a versatile system for in vitro production of human neurons. PLoS One 7(1):e29597. https://doi. org/10.1371/journal.pone.0029597 11. Lundin A, Delsing L, Clausen M, Ricchiuto P, Sanchez J, Sabirsh A, Ding M, Synnergren J, Zetterberg H, Brolen G, Hicks R, Herland A, Falk A (2018) Human iPS-derived Astroglia from a stable neural precursor state show improved functionality compared with conventional astrocytic models. Stem Cell Rep 10 (3):1030–1045. https://doi.org/10.1016/j. stemcr.2018.01.021 12. Tailor J, Kittappa R, Leto K, Gates M, Borel M, Paulsen O, Spitzer S, Karadottir RT, Rossi F, Falk A, Smith A (2013) Stem cells expanded from the human embryonic hindbrain stably retain regional specification and high neurogenic potency. J Neurosci 33

(30):12407–12422. https://doi.org/10. 1523/JNEUROSCI.0130-13.2013 13. Fujimoto Y, Abematsu M, Falk A, Tsujimura K, Sanosaka T, Juliandi B, Semi K, Namihira M, Komiya S, Smith A, Nakashima K (2012) Treatment of a mouse model of spinal cord injury by transplantation of human induced pluripotent stem cell-derived long-term selfrenewing neuroepithelial-like stem cells. Stem Cells 30(6):1163–1173. https://doi.org/10. 1002/stem.1083 14. Gronning Hansen M, Laterza C, PalmaTortosa S, Kvist G, Monni E, Tsupykov O, Tornero D, Uoshima N, Soriano J, Bengzon J, Martino G, Skibo G, Lindvall O, Kokaia Z (2020) Grafted human pluripotent stem cell-derived cortical neurons integrate into adult human cortical neural circuitry. Stem Cells Transl Med 9(11):1365–1377. https://doi.org/10.1002/sctm.20-0134 15. Solomon J, Csontos L, Clarke D, Bonyhadi M, Zylberberg C, McNiece I, Kurtzberg J, Bell R, Deans R (2016) Current perspectives on the use of ancillary materials for the manufacture of cellular therapies. Cytotherapy 18(1):1–12. https://doi.org/10.1016/j.jcyt.2015.09.010 16. Iancu EM, Kandalaft LE (2020) Challenges and advantages of cell therapy manufacturing under good manufacturing practices within the hospital setting. Curr Opin Biotechnol 65:233–241. https://doi.org/10.1016/j. copbio.2020.05.005 17. Henriques D, Moreira R, Schwamborn J, Pereira de Almeida L, Mendonca LS (2019) Successes and hurdles in stem cells application and production for brain transplantation. Front Neurosci 13:1194. https://doi.org/10. 3389/fnins.2019.01194 18. Vitillo L, Durance C, Hewitt Z, Moore H, Smith A, Vallier L (2020) GMP-grade neural progenitor derivation and differentiation from clinical-grade human embryonic stem cells. Stem Cell Res Ther 11(1):406. https://doi. org/10.1186/s13287-020-01915-0

Methods in Molecular Biology (2022) 2520: 101–115 DOI 10.1007/7651_2021_402 © Springer Science+Business Media, LLC 2021 Published online: 05 May 2021

Feeder-Dependent/Independent Mouse Embryonic Stem Cell Culture Protocol Hatice Burcu S¸is¸li, Selinay S¸enkal, Derya Sag˘rac¸, Taha Bartu Hayal, and Ays¸egu¨l Dog˘an Abstract Mouse embryonic stem cells (mESCs) were first derived and cultured nearly 30 years ago and have been beneficial tools to create transgenic mice and to study early mammalian development so far. Fibroblast feeder cell layers are often used at some stage in the culture protocol of mESCs. The feeder layer—often mouse embryonic fibroblasts (MEFs)—contribute to the mESC culture as a substrate to increase culture efficiency, maintain pluripotency, and facilitate survival and growth of the stem cells. Various feederdependent and feeder-independent culture and differentiation protocols have been established for mESCs. Here we describe the isolation, culture, and preparation feeder cell layers and establishment of feeder-dependent/independent protocol for mESC culture. In addition, basic mESC protocols for culture, storage, and differentiation were described. Key words Cell culture, Mouse embryonic fibroblasts, Mouse embryonic stem cells

1

Introduction Ever since the discovery of mouse ESC (mESC) in 1981, the main concern of the researchers was to establish an effective condition to proliferate the cells while maintaining the undifferentiated, pluripotent state with self-renew capacity [1, 2]. To keep mESCs in this state, scientists came up with various culture conditions which are categorized under feeder-dependent or feeder-free culture techniques. The presence of feeder cells provides mESCs a matrix to attach and growth factors to help proliferation as well as maintenance of pluripotency [3]. Currently, one of the most commonly used methods in laboratories, studying mESCs, is mouse embryonic fibroblast (MEF) feeder dependent method. In this method, MEFs are used as feeder cells and the medium contains FBS, which provides hormones as well as essential nutrients, and leukemia inhibitory factor (LIF), which is cytokine-inducing JAK-STAT pathway to retain pluripotency and self-renew of mESCs [4]. The other standard method is a feeder-free method, that does not involve feeder cells, and uses a well-defined medium supplemented

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with N2, B27, LIF, and two small inhibitors, called CHIR99021 and PD0325901 [5, 6]. While CHIR99021 is a GSK3 inhibitor, which maintains pluripotency [7], PD0325901 inhibits ERK1/ 2 to block differentiation [8]. Mouse embryonic fibroblasts (MEFs) are primary cell lines derived from mouse embryos which are acquired from a pregnant mouse generally at day 12.5 to day 14.5 [9, 10]. MEFs are mitotically inactivated to stop cell division while retaining the metabolic activity in order to be used as feeder cells for mESCs. The condition is commonly met by either Mitomycin C treatment or gammairradiation of MEFs [11, 12]. Pluripotent properties of mESCs enable differentiation into three germ layers: endoderm, mesoderm, and ectoderm. mESCs are aggregated to form three-dimensional structure in vitro, called embryoid bodies (EBs), and during this process, the cells acquire regional differentiations indicating linage markers of three germ layers. Formation of EBs from mESC let scientist to study embryonic development as well as effect of genetic modifications [13]. In this chapter, successful methods for MEF culture, which describe how to isolate, culture, and mitotically inactivate MEF were described. We also described mESC culture, which involves the protocols to grow and cryopreserve mESCs on MEF feeder layer and without feeder layer, EB formation from mESCs, and differentiation of ESCs.

2 2.1

Materials Mouse

2.2 Preparation of Media

1. A mated mouse should be pregnant for 11–14 days. 1. MEF medium: Add 10% fetal bovine serum (FBS, Invitrogen, Gibco, UK), 1% penicillin, streptomycin, and Amphotericin B (PSA, Gibco, UK), and 1% L-glutamine (Gibco, UK) to Dulbecco’s modified Eagle’s medium (DMEM, Invitrogen, Gibco, UK) with high glucose (see Note 1). 2. Freezing medium for MEF: Add 10% dimethyl sulfoxide (DMSO, Sigma) to FBS. 3. Freezing medium for mESCs: Add 10% DMSO to ES-qualified FBS (ATCC) (see Note 2). 4. mESC medium: Add 15% ES-qualified FBS, 1% PSA, 1% Lglutamine, and 1% non-essential amino acids (#11140050, Gibco, UK) to DMEM with high glucose. 5. N2B27 medium: Mix DMEM F12 (Gibco, Thermo Fisher) and Neurobasal medium (Gibco, Thermo Fisher) at a ratio of 1:1. Add 0.5% bovine serum albumin (10%) (BSA, #A1595,

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Sigma), 0.5% N2 supplement (#17502001, Gibco, Thermo Fisher), 1% B27 supplement (#17504001, Gibco, Thermo Fisher), 1% L-glutamine, and 1% PSA to the medium [14] (see Note 3). Store it at 4
C up to 1 month. 6. Feeder-free medium (FFM): Add 3 μM CHIR99021 (#72054, Stem Cell), 1 μM PD0325901, and 10 ng/ml LIF to N2B27 medium just before the use. Filter the solution through 0.22 μm filter. 7. EB formation medium: Add 12 ng/ml bFGF and 10 μM Y-27632 (#72304, Stem Cell) to N2B27 medium just before the use (see Note 4). Filter the solution through 0.22 μm filter. 2.3 Preparation of Mitomycin C

1. Dissolve 2 mg Mitomycin C (#m4287, Sigma) in 200 ml DMEM to prepare 10 μg/ml stock. Filter the solution through 0.22 μm filter and aliquot the solution 10 ml per falcon tube. Store the solutions at 20
C for long-term use.

2.4 LIF

1. Dilute 100 μg recombinant mouse LIF (#Z200485, Applied Biological Materials Inc., Canada) in 1 ml distilled water to prepare 100 μg/ml stock solution. The working solution of LIF is 10 ng/ml in mESC medium [15].

3 3.1

Preparation of

Methods MEF Culture

3.1.1 Isolation

1. Euthanize a pregnant mouse at day 11–14 post-coitum (see Notes 5 and 6) and disinfect the fur of the mouse using 70% ethanol. 2. Cut the skin and remove the abdominal wall to reveal the uterine horns and count the number of embryos (see Note 7). Use sterile forceps and scissors to hold uterine horns and to dissect them out (see Note 8). 3. Put the uterine horns in a petri dish filled with 15–20 ml PBS without Ca2+ and Mg2+ which is enough to cover the uterus and wash them (see Note 9). 4. Carry the petri to a hood and place the uterus in a sterile dish filled with sterile 10 ml PBS. Use a sterile scissor to cut the placenta between the embryos and use forceps to make a little pressure to take out the embryos. 5. Place the embryos in a sterile dish filled with sterile PBS. While using forceps to hold the embryo, cut the head with a scalpel, and remove red organs, including heart, liver, and kidney, from the body. 6. Place the embryo bodies in a new dish, add 1 ml of 0.05% trypsin/EDTA on to the embryos, and use a scalpel blade to mince the embryos finely until the tissues can pass through a

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1000-μl pipette tip. Then, complete trypsin to 7 ml to cover the tissues and pipette up and down the tissues about ten times before incubating the dish at 37
C for 15 min. During the incubation, pipette up and down the tissues after each 5 min and place the dish back to the incubator. 7. Take out the dish from the incubator, pipette up and down 10–15 times and transfer the tissues to a 50-ml falcon tube. Wash the dish with 7 ml fresh MEF medium and transfer the media to the falcon tube to inactivate the trypsin/EDTA. 8. Incubate for a few minutes to allow tissue pieces to settle down and transfer the supernatant containing the cells to a sterile 50-ml falcon tube (see Note 10). 9. Centrifuge the falcon tube at 500  g for 5 min. Discard the supernatant and resuspend the pellet in MEF medium. Divide the suspension equally as cells of three embryos will be seeded into a T150 flask (see Note 7). Add MEF medium to the T150 flasks to complete volume to 25 ml and label the flasks Passage 0 (Fig. 1). 10. Place the flasks in a 5% CO2 incubator at 37
C and allow the cells to reach to confluency in 3–4 days.

Fig. 1 Illustration of MEF isolation protocol. Embryos are taken out from the uterus of pregnant mouse at day 11–14. Head and red organs are removed, and the embryonic body is chopped finely in trypsin/EDTA. The tissues are incubated with trypsin at 37
C for 15 min. Trypsin/EDTA is inactivated by MEF media, and the suspension is centrifuged at 500  g for 5 min. After the supernatant is removed, the pellet is dissolved with MEF media and transferred to the flasks

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11. When the cells are confluent, detach the cells using trypsin and either freeze for cryopreservation using freezing medium or continue the culture until Passage 6–7 to inactivate them for the purpose of feeder layer for mESCs (see Note 11). 3.1.2 Culture

1. Remove the medium from the T150 flask. Wash the flask gently with 5 ml PBS and discard the PBS. 2. Add 5 ml 0.25% trypsin/EDTA and incubate the plate at 37
C for 3–5 min. 3. Inactivate the trypsin/EDTA by adding 5 ml MEF medium to the flask and transfer the cell suspension to falcon tube. 4. Centrifuge the cells at 300  g for 5 min. 5. Discard the supernatant and resuspend the pellet in MEF medium by pipetting. 6. Divide the cell suspension of each flask into three flasks and complete volume of the flasks to 25 ml with MEF medium. 7. Incubate the flasks in a 5% CO2 incubator at 37
C to allow them to grow confluency.

3.1.3 Treatment

1. Remove the medium of MEF cells from the T150 flask (see Note 12). 2. Add 10 ml of 10 μg/ml Mitomycin C solution to the flask and incubate the flask in a 5% CO2 incubator at 37
C for 2–3 h (see Note 13) (Fig. 2).

Fig. 2 Morphological pictures of MEF cells. MEF cells (a) before and (b) after mitomycin C treatment. (c) Morphology of mitomycin C-treated MEFs after 3 weeks with rarely media change (scale bar ¼ 200 μm)

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3. Remove the Mitomycin C solution from the flask and wash the flask with 8 ml PBS. 4. Add 5 ml 0.25% trypsin/EDTA and incubate the plate at 37
C for 3–5 min. 5. Inactivate the trypsin/EDTA by adding 5 ml MEF medium to the flask and transfer the cell suspension to a falcon tube. 6. Centrifuge the cells at 300  g for 5 min. 7. Discard the supernatant and resuspend the pellet in MEF medium by pipetting. 8. Seed the cells to 6-well plates with 3  105 cells per well in 2 ml MEF medium (see Notes 14 and 15). 9. Incubate the plates overnight in a 5% CO2 incubator at 37
C to allow MEFs to attach the plate. The treated cells can be incubated for a week with media changes every 2–3 days (see Note 16). 10. Discard the media of MEF cells just before ESCs are seeded on them in ESC media. 3.2 Mouse Embryonic Stem Cell Culture on MEFs

1. Prepare MEF feeder layer as explained in Subheading 3.1.3.

3.2.1 Thawing and Culturing mESCs on MEFs

3. Place the vial in the 37
C water bath until melting starts.

2. Take a frozen mESC vial from the liquid nitrogen storage tank and transport it immediately to the cell culture room. 4. Transport the vial and spray with 75% ethanol to put it in the sterile hood. 5. Add 9 ml of fresh mESC medium in a 15-ml falcon tube. 6. Transfer the mESCs in the vial into the 15-ml falcon tube using 1000-μl pipette and wash the vial carefully with mESC medium. 7. Centrifuge cells at 270  g for 5 min and then spray the falcon tube with 75% ethanol before putting it into the sterile hood. 8. Discard the supernatant into the waste and add 1 ml of mESC medium containing 10 ng/ml LIF onto mESC pellets in a falcon tube. 9. Disperse the mESC pellets carefully to avoid damaging cells using 1000-μl pipette. 10. Aspirate the MEF media out of the 6-well plate and add 1 ml of mESC medium containing LIF. 11. Add 1 ml of mESC suspension to the 6-well plate using 1000-μ l pipette and then rock the plate X–Y direction gently (see Note 17). 12. Observe the concentration of cells under an inverted microscope and be sure that the cells are at enough confluency.

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Fig. 3 Morphological pictures of pluripotent mESCs on MEF. (a) mESCs after seeding. (b) mESCs 6 h after seeding. (c) mESCs 48 h after seeding. (d) Differentiated colony of mESCs (scale bar ¼ 200 μm)

13. Incubate the plate at 37
C, %5 CO2 incubator and check the next day for attachment of mESCs onto MEFs (see Note 18). 14. Change the medium every day with fresh mESC medium containing LIF. 15. See the morphology of pluripotent mESCs in colonies (Fig. 3) and when colonies are enlarged and their sizes are increased, passage the mESCs (Fig. 3c) (see Note 19). 3.2.2 Passaging and Freezing mESCs on MEFs

1. Aspirate the mESCs media containing LIF out of the 6-well plate including mESCs. 2. Add 1 ml of DPBS to wash out of the 6-well plate. 3. Aspirate out of the DPBS and add 1 ml of the 0.25% trypsin/ EDTA to the mESCs in the 6-well plate. 4. Incubate the 6-well plate for 3 min at 37
C, %5 CO2 incubator (see Note 20). 5. Check the detached cells under the inverted microscope after the trypsinization step. 6. Add 2 ml of mESC medium to the 6-well plate and rock the plate back and forth (6-well plate includes a total of 3 ml). 7. Divide the solution into two 15-ml falcon tubes and label as freezing (2 ml) and culturing (1 ml). 8. Transfer 1 ml of cell suspension containing mESCs using 1000μl pipette into 15-ml falcon tube. 9. Transfer remaining 2 ml of suspension containing mESCs using 1000-μl pipette into 15-ml falcon tube. 10. Centrifuge two falcon tubes for 5 min at 300  g and then spray the falcon tubes with 75% ethanol before putting them into the sterile hood. 11. Discard the medium from falcon tube including 2 ml of suspension and add 1.25 ml of freezing medium and suspend the mESC pellet (see Note 21). 12. Transfer the cell suspension in freezing medium into cryovial and placed to 80
C freezer with alcohol-free

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cryopreservation container for 24 h and then transfer to the liquid nitrogen storage tank for long-term storage (see Note 22). 13. Discard the medium from falcon tube including 1 ml of solution and suspend the mESC pellet with 2 ml of mESC medium containing LIF. 14. Seed the cells onto MEF layers in 6-well plate using 1000-μl pipette and then rock the plate X–Y direction gently. 15. Observe the concentration of cells under an inverted microscope and be sure that the cells are enough confluency. 16. Incubate the plate at 37
C, %5 CO2 incubator and check the next day for attachment of mESCs onto MEFs (see Note 18). 17. Check and change the medium every day as explained in Subheading 3.1.2. 3.3 Feeder-Free Protocol of Mouse Embryonic Stem Cell Culture and Differentiation

1. Thaw Matrigel® at 4
C overnight prior to the coating and keep on ice during procedure. Mix 500 μl of 10 mg/ml stock Matrigel® with 50 ml of ice-cold serum-free DMEM/F12 (see Note 23). Add coating solution to the tissue culture well plates at an appropriate amount (see Note 24). 2. Incubate the plates at 37
C for at least 30 min. Remove the coating solution prior to cell culture by aspiration (see Note 25). 3. mESCs should be separated from MEFs before feederindependent culture. Detach the mESCs as described in Subheading 3.2.2. 4. After centrifugation, seed the cell mixture on non-coated wells with mESC medium for 30 min at 37
C (see Note 26). 5. At the end of incubation time, check the dishes under the microscope to confirm MEF attachment. Collect the cell suspension containing ES cells and centrifuge at 300  g for 3 min. 6. Seed the 2.0  105 cells per well of 6-well plate with FFM medium as described in Subheading 2.2 (see Note 27). Incubate the plates at 37
C with 5% CO2 in a humidified chamber (Fig. 4). 7. Monitor the mouse ESCs and replace the medium every day. When mouse ESC colonies reach passaging density, transfer the cells to the freshly coated surfaces.

3.4 Embryoid Body (EB) Formation from mESCs in Suspension Culture

1. Separate mESCs from MEF feeder cells for 30 min as described in Subheading 3.3. 2. Prepare EB formation medium as described in Subheading 2.2 and filter medium after supplementation using 0.22 μm filter during 30 min incubation.

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Fig. 4 Morphological picture of feeder-free pluripotent mESCs. mESC colonies cultured in 2i + LIF medium for 2 days (magnification: 4)

3. Apply EB formation protocol for 2 days in order to reach epi-like state (see Note 28). 4. Check plate under inverted microscope in order to observe MEF attachment after incubation. 5. Collect cell suspension including mESCs after 30 min of incubation and transfer into a 15-ml falcon tube. 6. Centrifuge the tube at 300  g for 5 min. 7. Discard the supernatant and resuspend the pellet with 3 ml of EB formation buffer. 8. Add cell solution into low attachment 6-well plate for formation of EBs in suspension culture (see Note 29) (Fig. 5). 9. Incubate the plate at 37
C with 5% CO2 in a humidified chamber for 3 days but check the EB formation each day.

4

Notes 1. Some of the protocols in the literature include addition β-mercaptoethanol to prepare MEF medium [16]. However, we do not use it, since it is a toxic reagent, and MEF cells grow quite healthy without using it. 2. Although there are commercially available cryopreservation media, such as PSC Cryopreservation Kit (#A2644601, Thermo), our freezing medium works fine. 3. Some combinations of N2B27 may contain β-mercaptoethanol [17] as a reducing agent or more commonly monothioglycerol (MTG) [14]. However, we did not observe any change at the

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Fig. 5 Schematic representation of EB formation protocols using (a) RT-PCR plate, (b) low attachment plate, and (c) hanging drop technique

efficiency of the medium when MTG was added. So, we do not use it anymore. 4. If N2B27 medium was not prepared freshly, then add 1% Lglutamine each time while preparing EB formation medium. 5. Day of the embryos are counted after the formation of the vaginal plug. Although ideal day for dissection of fetus is day 13–14, MEF isolation can be performed as early as day 8.5 [16]. 6. Ethical approval must be taken before using the use and humanized techniques should be used for Euthanasia, such as cervical dislocation or CO2 exposure. 7. Count the embryos for cell seeding at the end of the isolation. Generally, three embryos per T150 flask is enough to grow the cells to confluency after 3–4 days of incubation.

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8. Avoid touching fur or damaging the organs to prevent any potential contamination. 9. The first three steps can be performed under non-aseptic conditions. However, the remaining steps should be performed in a sterile environment such as biosafety hoods with sterile instruments and aseptic techniques should be used to prevent contamination risk. 10. Separating large tissue pieces will provide a clear cell culture. However, their presence is not a problem since fibroblasts from the tissues will also migrate and contribute to the MEF culture. 11. Morphology of MEF cells should spread at day 2, and at day 3–4, the elongated MEFs should reach confluency. If there is a space between the cells, wait for it to be closed before passage or cryopreservation. 12. MEFs should be tested for mycoplasma contamination before using as feeder layer. 13. We use 10 ml Mitomycin C to treat MEF in T150 flask. However, we have tried different volumes for the treatment and concluded that 8–15 ml Mitomycin C can be used to treat MEFs in T150 flask for 2–3 h. 14. Mitomycin C-treated MEFs can be used directly by seeding them as a feeder layer or can be cryopreserved to use in the future. 15. Variation at seeding density of Mitomycin C-treated MEF according to the size of the plates are listed in Table 1. 16. Mitomycin C-treated MEF cells can be incubated ideally for a week and maximum for 2 weeks with media changes every 2–3 days. However, as the time pass, MEFs become unhealthy and should not be used as a feeder layer (Fig. 2). 17. Be sure to disperse mESCs onto MEF properly after thawing cells. 18. Be sure to check the morphology of the MEFs. If MEFs start to detach, mESCs will not be properly cultured and grown. 19. Check cells each day to avoid differentiation of colonies due to over-confluency or detached MEFs. Table 1 Cell density of Mitomycin C-treated MEFs for seeding Plates

Number of cells

Required volume (ml)

6-Well plate

3  10

2

12-Well plate

1.5  105

1

24-Well plate

8  10

0.5

5

4

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20. Be sure to check the requirements of the incubator such as temperature, water, and CO2 level properly every week. 21. For the ideal freezing procedure, split colonies in Fig. 3c into two and add 1.25–1.5 ml freezing media for each vial. 22. Although it is customary to store MEFs and mESCs in liquid nitrogen, we have also stored cells at 80
C for up to 6 months without loss of cell viability after thawing. 23. The volume of Matrigel® required for the preparation of coating solution varies according to each lot as concentration of proteins might be different. Check the lot-specific certificate of analysis of Matrigel® to find out the exact protein concentration. Matrigel® has been used in different concentrations such as 2, 4, and 6 mg/ml [18], and 5 mg/ml [19] for ES cell culture. Moreover, use the serum-free medium for solution preparation to prevent possible effects of serum. In addition to Matrigel® coating, 0.1% gelatin coating is also widely used [20, 21]. 24. During the incubation at 37
C, the water in the medium might evaporate and results in the non-homogenous distribution of coating materials. Therefore, 1000 μl of the solution will be sufficient for a well of 6-well plate (Table 2). 25. 1 h incubation at room temperature might be applied to store coated plates at 4
C for further use. These plates can be wrapped carefully and stored at 4
C for 2 weeks. Plates should be incubated at 37
C for 30 min before use. Do not let plates dry out. A dried surface may reduce the attachment of cells. Add culture medium immediately before use. Table 2 The required volume of a coating solution to cover the surface Surface growth area (cm2)

Required volume

6-Well plate

9.8

800–1,000 μl

12-Well plate

3.8

350–500 μl

24-Well plate

1.9

100–250 μl

48-Well plate

0.95

40–150 μl

96-Well plate

0.32

10–50 μl

35-mm dish

8

500–800 μl

100-mm dish

44

2.5–4 ml

150-mm dish

148

5.5–7 ml

Plates

Dishes

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26. The period can be extended to 40 min. To facilitate the adhesion of MEFs, pre-plating of the cell mixture on 0.1% gelatincoated wells has been utilized [22], but not performed here. 27. A third inhibitor can be used in addition to CHIR99021 and PD0325901. 3i + LIF protocols can be conducted by adding a TGF-β RI Kinase Inhibitor VI, SB431542 [23], a tankyrase inhibitor, XAV939 [24], or an FGF receptor tyrosine kinase inhibitor, SU5402 [25] to the medium in the presence of LIF. 28. There are different protocols to generate EB suspension culture by inducing an epi-like state for further differentiation. 5–12 ng/ml FGF2 and 10–20 ng/ml Activin exposure for 2 days might trigger Epi-like state [26]. Standalone 5 ng/ml bFGF treatment for 3 days induced Epi-like state [27, 28]. 29. There are several EB formation methods for different experiments which are summarized in Fig. 5. Uniform size EBs are formed by using round bottom RT-PCR plate [29]. Different size EBs are produced using low attachment plate in suspension culture. This technique is preferred for the generation of high number of EBs [30]. In hanging drop method, EBs are formed in the lid of petri dish as hanging drops. The advantages of this method are inexpensive and easy culture procedures. Also, homogeneous EBs from predetermined number of mESCs can be reproduced [31].

Acknowledgments € ˙ TAK 2232 International FelThis study was supported by TUBI lowship for Outstanding Researchers Program (Project no: 118C186). The authors declare no conflict of interest. References 1. Martin GR (1981) Isolation of a pluripotent cell line from early mouse embryos cultured in medium conditioned by teratocarcinoma stem cells. Proc Natl Acad Sci U S A 78:7634–7638. https://doi.org/10.1073/pnas.78.12.7634 2. Evans MJ, Kaufman MH (1981) Establishment in culture of pluripotential cells from mouse embryos. Nature 292:154–156. https://doi.org/10.1038/292154a0 3. Robertson EJ (1997) Derivation and maintenance of embryonic stem cell cultures. Methods Mol Biol 75:173–184 4. Graf U, Casanova EA, Cinelli P (2011) The role of the leukemia inhibitory factor (LIF) — pathway in derivation and maintenance of murine pluripotent stem cells. Genes (Basel)

2:280–297. https://doi.org/10.3390/ genes2010280 5. Ying QL, Wray J, Nichols J et al (2008) The ground state of embryonic stem cell selfrenewal. Nature 453:519–523. https://doi. org/10.1038/nature06968 6. Tai CI, Ying QL (2013) Gbx2, a LIF/Stat3 target, promotes reprogramming to and retention of the pluripotent ground state. J Cell Sci 126:1093–1098. https://doi.org/10.1242/ jcs.118273 7. Wray J, Kalkan T, Gomez-Lopez S et al (2011) Inhibition of glycogen synthase kinase-3 alleviates Tcf3 repression of the pluripotency network and increases embryonic stem cell resistance to differentiation. Nat Cell Biol

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13:838–845. https://doi.org/10.1038/ ncb2267 8. Kunath T, Saba-El-Leil MK, Almousailleakh M et al (2007) FGF stimulation of the Erk1/ 2 signalling cascade triggers transition of pluripotent embryonic stem cells from selfrenewal to lineage commitment. Development 134:2895–2902. https://doi.org/10.1242/ dev.02880 9. Nagy A, Gertsenstein M, Vintersten K, Behringer R (2006) Preparing mouse embryo fibroblasts. Cold Spring Harb Protoc 2006:pdb. prot4398. https://doi.org/10.1101/pdb. prot4398 10. Durkin M, Qian X, Popescu N, Lowy D (2013) Isolation of mouse embryo fibroblasts. Bio-Protocol 3. https://doi.org/10.21769/ bioprotoc.908 11. Nagy A, Gertsenstein M, Vintersten K, Behringer R (2006) Preparing feeder cell layers from STO or mouse embryo fibroblast (MEF) cells: treatment with mitomycin C. Cold Spring Harb Protoc 2006:pdb.prot4399. https:// doi.org/10.1101/pdb.prot4399 12. Nagy A, Gertsenstein M, Vintersten K, Behringer R (2006) Preparing feeder cell layers from STO or mouse embryo fibroblast (MEF) cells: treatment with γ-irradiation. Cold Spring Harb Protoc 2006:pdb.prot4400. https://doi.org/ 10.1101/pdb.prot4400 13. Itskovitz-Eldor J, Schuldiner M, Karsenti D et al (2000) Differentiation of human embryonic stem cells into embryoid bodies compromising the three embryonic germ layers. Mol Med 6:88–95. https://doi.org/ 10.1007/bf03401776 14. Gouon-Evans V, Boussemart L, Gadue P et al (2006) BMP-4 is required for hepatic specification of mouse embryonic stem cell-derived definitive endoderm. Nat Biotechnol. https:// doi.org/10.1038/nbt1258 15. Ying QL, Nichols J, Chambers I, Smith A (2003) BMP induction of Id proteins suppresses differentiation and sustains embryonic stem cell self-renewal in collaboration with STAT3. Cell. https://doi.org/10.1016/ S0092-8674(03)00847-X 16. Smith CL (2006) Mammalian cell culture. Curr Protoc Mol Biol 73:28.0.1–28.0.2. https://doi.org/10.1002/0471142727. mb2800s73 17. Di Stefano B, Ueda M, Sabri S et al (2018) Reduced MEK inhibition preserves genomic stability in naive human embryonic stem cells. Nat Methods. https://doi.org/10.1038/ s41592-018-0104-1

18. Ozdil B, Gu¨ler G, Acikgoz E et al (2020) The effect of extracellular matrix on the differentiation of mouse embryonic stem cells. J Cell Biochem 121:269–283. https://doi.org/10. 1002/jcb.29159 19. Chang SY, Carpena NT, Mun S et al (2020) Enhanced inner-ear organoid formation from mouse embryonic stem cells by photobiomodulation. Mol Ther Methods Clin Dev 17:556–567. https://doi.org/10.1016/j. omtm.2020.03.010 20. Tamm C, Galito´ SP, Annere´n C (2013) A comparative study of protocols for mouse embryonic stem cell culturing. PLoS One 8:81156. https://doi.org/10.1371/journal.pone. 0081156 21. Bae YU, Sung HK, Kim JR (2017) Collection of serum- and feeder-free mouse embryonic stem cell-conditioned medium for a cell-free approach. J Vis Exp 2017:55035. https://doi. org/10.3791/55035 22. Millipore M (2004) Murine embryonic stem cell culture manual 2004. Leukemia 23. Ma Y, Yu T, Cai Y, Wang H (2018) Preserving self-renewal of porcine pluripotent stem cells in serum-free 3i culture condition and independent of LIF and b-FGF cytokines. Cell Death Discov 4:21. https://doi.org/10.1038/ s41420-017-0015-4 24. Park TS, Zimmerlin L, Evans-Moses R, Zambidis ET (2018) Chemical reversion of conventional human pluripotent stem cells to a naı¨velike state with improved multilineage differentiation potency. J Vis Exp 2018:57921. https://doi.org/10.3791/57921 25. Nishihara K, Shiga T, Nakamura E et al (2019) Induced pluripotent stem cells reprogrammed with three inhibitors show accelerated differentiation potentials with high levels of 2-cell stage marker expression. Stem Cell Rep 12:305–318. https://doi.org/10.1016/j.stemcr.2018.12. 018 26. Morgani S, Nichols J, Hadjantonakis AK (2017) The many faces of pluripotency: in vitro adaptations of a continuum of in vivo states. BMC Dev Biol 17:1–20 27. Bernemann C, Greber B, Ko K et al (2011) Distinct developmental ground states of epiblast stem cell lines determine different pluripotency features. Stem Cells 29:1496–1503. https://doi.org/10.1002/stem.709 28. Greber B, Wu G, Bernemann C et al (2010) Conserved and divergent roles of FGF signaling in mouse epiblast stem cells and human embryonic stem cells. Cell Stem Cell 6:215–226. https://doi.org/10.1016/j.stem. 2010.01.003

Feeder-Dependent/Independent Mouse Embryonic Stem Cell Culture Protocol 29. Koike M, Kurosawa H, Amano Y (2005) A round-bottom 96-well polystyrene plate coated with 2-methacryloyloxyethyl phosphorylcholine as an effective tool for embryoid body formation. Cytotechnology 47:3–10 30. Stover AE, Schwartz PH (2011) The generation of embryoid bodies from feeder-based or feeder-free human pluripotent stem cell

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Methods in Molecular Biology (2022) 2520: 117–133 DOI 10.1007/7651_2021_400 © Springer Science+Business Media, LLC 2021 Published online: 05 May 2021

ChIP-qPCR for Polycomb Group Proteins During Neuronal Differentiation of Human Pluripotent Stem Cells Divya Desai and Prasad Pethe Abstract Neuronal differentiation is an intricate and a complex process which involves crosstalk among various signaling pathways, growth factors, transcription factors, and epigenetic modifiers. During different stages of neuronal development, there are various histone modifiers which drive the expression of lineage-specific genes. Polycomb group proteins are one of the histone modifiers that control transcriptional repression of specific genes in development, differentiation, and functionality of various tissues. Chromatin immunoprecipitation (ChIP) is a widely used technique to investigate the interaction of proteins and DNA; ChIP combined with quantitative real-time PCR (qPCR) gives a quantitative data about the occupancy of specific protein on a particular stretch of DNA, and this can help us investigate how a protein regulates expression of a specific gene. In this chapter, we describe a protocol for ChIP coupled to qPCR during early neuronal differentiation to identify the specific genomic targets regulated by components of Polycomb repressive complex 1. Key words Chromatin immunoprecipitation, Neuronal differentiation, PcG, PRC1, qPCR, RING1B

Abbreviations BMI1/PCGF4 ChIP H2AK119ub1 hESC hiPSC NPC PcG PRC1 RING1B

1

B Lymphoma Mo-MLV insertion region 1 homolog, Polycomb ring finger protein 4 Chromatin immunoprecipitation Monoubiquitination of lysine 119 on histone H2A Human embryonic stem cells Human-induced pluripotent stem cells Neuronal progenitor cells Polycomb group protein(s) Polycomb repressive complex 1 Really interesting new gene 1B/RING-type E3 ubiquitin transferase RING1

Introduction Human pluripotent stem cells (embryonic stem cells and induced pluripotent stem cells) exhibit the properties of pluripotency and self-renewal. Their ability to differentiate into any of the three germ

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layers (endoderm, mesoderm, and ectoderm) makes them unique. The mammalian central nervous system (CNS) is composed of three major differentiated cell types: neurons, astrocytes, and oligodendrocytes. Neurogenesis in vertebrates begins after differentiation of ectodermal cells into the neuroepithelial cells [1, 2]. Neuroepithelial cells form the radial glial cells, which undergo a period of expansion, then differentiate into committed transit amplifying progenitors that eventually give rise to mature cells of the central nervous system (CNS). Polycomb group (PcG) proteins play an important role by selectively repressing the activity of a specific set of developmental regulatory genes [3]. They comprise several proteins which assemble into two broad complexes, viz. Polycomb Repressive Complex 1 (PRC1) and Polycomb Repressive Complex 2 (PRC2). The PRC1 and PRC2 complexes repress genes by catalyzing histone modifications such as H2AK119 mono-ubiquitination and H3K27 di and tri methylation, respectively [4]. Chromatin immunoprecipitation (ChIP) is a useful technique to understand how PcG protein binds to promoters of neuronalspecific genes and regulates their expression by catalyzing histone modifications. ChIP can be combined with real-time PCR, microarray, or sequencing to analyze these complex interactions in silico [5]. It is crucial to understand the method of analyzing the results after performing ChIP-qPCR. The two most commonly used methods for analysis are (1) fold enrichment method and (2) ChIP input method [6]. In this chapter, we have given a detailed protocol describing ChIP combined with qPCR to determine the localization of the PcGs at neural lineage-specific genes during neuronal differentiation. Figure 1 shows the typical schedule followed for neuronal differentiation from human pluripotent stem cells that include the timepoints used for the ChIP-qPCR assay. Selective immunoprecipitation of DNA-protein complexes using antibodies against RING1B, BMI1, H2AK119ub1, and EZH2 coupled to magnetic beads (Dyna beads) provides a quicker and easier method to study the interaction of these PcG components in a stage-specific manner during neural differentiation. We described the ChIP input method to quantitatively measure the occupancy of core PRC1 components: RING1B and BMI1 along with its histone modification H2AK119ub1 on various neural lineage-specific genes.

2

Materials

2.1 Cell Culture Reagents Preparation

1. Basic fibroblast growth factor (bFGF/FGF2) (Life technologies, Cat. no. PHG0264). 2. Fibroblast growth factor-4 (FGF4) (Life technologies, Cat. no. PHG0154).

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Fig. 1 Schedule of neuronal differentiation. Preparation of EBs from undifferentiated pluripotent stem cells using hanging drop method followed by propagation on geltrex-coated dishes with the addition of synthetic inhibitors, growth factors, and morphogens during various timepoints in differentiation regimen. The differentiated cells in DMEM supplemented with N2, B27, NEAA, and Glutamax for efficient growth. The timepoints for ChIP-qPCR assay mentioned at the bottom of the image indicates each stage of neuronal differentiation

3. SB431542 hydrate (Sigma Aldrich, Cat. no. S4317). 4. LDN193189 hydrochloride no. SML0559).

(Sigma

Aldrich,

Cat.

5. Sonic hedgehog (SHH) (Sigma Aldrich, Cat. no. SRP3156). 6. Retinoic acid (RA) (Sigma Aldrich, Cat. no. R2625). 7. DKK1 (Wnt inhibitor) (Gibco, Cat. no. PHC9214). 8. N2 supplement (Life Technologies, Cat. no. 17502048). 9. B27 supplement (Life Technologies, Cat. no. 12587). 10. Glutamax (Life Technologies, Cat. no. 35050061).

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11. Non-essential amino acids (NEAA) (Life Technologies, Cat. no. 11140050). 12. Essential 8 medium (Gibco, A1517001). 13. Dulbecco’s modified Eagle’s medium (DMEM) (Gibco, Cat. no. 11995-065). 14. Fetal bovine serum (FBS) (Gibco, Cat. no. 10270106). 15. Dulbecco’s phosphate-buffered saline (DPBS) Gibco, Cat. no. 14190144). 16. Insulin transferrin selenium (ITS) (Sigma Aldrich, Cat. no. I3146). 17. Geltrex (Gibco, Cat. no. A1413301). 18. Embryoid body formation (EB) medium: DMEM + 20% FBS + 1
ITS + 0.5
Geltrex. 19. Neural differentiation medium 1 (NDM1): DMEM + 1
N2 + 1
B27 + 1
NEAA + 1
Glutamax. 20. Neural differentiation medium 2 (NDM2): DMEM + 2
N2 + 2
B27 + 2
NEAA + 2
Glutamax. Note: Spin the non-reconstituted vials of growth factors, morphogens, and synthetic inhibitors so as to sediment all the lyophilized particles at the bottom and then vortex thoroughly after adding the solvent or reconstitution buffer. Always spin down the stock aliquots of reagents which are to be added in small amounts. Note: The stock and working stock of reagents need to be stored at 20  C/80  C. Avoid multiple freeze-thaw treatments of the growth factors, morphogens, and inhibitors. 2.2 Chromatin Immunoprecipitation Reagents/Buffers

1. Dyna beads (Life technologies, ThermoFisher Scientific, Cat. no. 10001D). 2. Dyna Mag2 magnet (Invitrogen, ThermoFisher Scientific, Cat. no. 12321D). 3. Wash buffer (WB): 0.01% Tween 20 (MP chemicals) in 1
PBS solution. 4. Blocking buffer: 0.5% BSA (HiMedia TC194) in 1
PBS solution. 5. 1
phosphate-buffered saline (PBS) solution. 6. 200 mM phenylmethylsulfonyl fluoride (PMSF) (Sigma Aldrich, P7626) in 100% isopropanol, vortex it to dissolve all the crystals. 7. ChIP lysis buffer: 50 mM HEPES KOH solution (Cell Clone), 140 mM sodium chloride (Molychem), 1 mM EDTA (Merck), 1% Triton X-100 (Merck), 0.1% SDS (MP chemicals), Protease Inhibitor Cocktail (Amresco, M221), and 1 mM PMSF (Sigma Aldrich).

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Note: Adjust the pH of cell lysis buffer to 7.5 prior in addition of Triton X-100. Store at 4  C. Add PIC-Protease Inhibitor Cocktail and PMSF just prior to use. Prepare a stock of PMSF separately. 8. Primary antibodies: RING1B (1:100) (Abcam, Cat. no. ab101273), H2AK119ub1 (1: 100) (CST, Cat. no. 8240), BMI1 (1:50) (CST, Cat. no. 6964), EZH2 (Abcam, Cat. no. ab191250), and rabbit isotype (Abcam, Cat. no. ab172730). 9. 1.25 M glycine (MP chemicals). 10. 37% formaldehyde solution (Merck). 11. Blocking buffer/antibody diluent for ChIP: 0.5% BSA in 1
PBS. Prepare the buffer fresh prior to usage. 12. IP1 buffer: 50 mM HEPES KOH (Cell Clone, New Delhi, India), 140 mM NaCl (Molychem), 1 mM EDTA (Merck), 1% Triton X-100 (Merck), 0.1% SDS (HiMedia), and 1 mM PMSF (Sigma Aldrich), adjust the pH to 7.5 then add 500 mM NaCl (Molychem). Store the buffer at 4  C. 13. IP2 buffer: 10 mM Tris (MP chemicals), 1 mM EDTA (Merck), 250 mM LiCl (Sigma Aldrich), and 0.5% sodium desoxycholate, adjust the pH to 8.0. Add the 0.5% NP40 detergent (Qualigens) in the end. Note: LiCl and sodium desoxycholate precipitate when added directly in buffer, hence prepare a stock solution of 2 M LiCl and 10% sodium desoxycholate and add the required volume to final buffer. Store the buffer at RT. Do not refrigerate the buffer since LiCl and sodium desoxycholate will precipitate under cold conditions. 14. Tris EDTA (TE) buffer: 10 mM Tris (MP chemicals) and 1 mM EDTA (Merck), adjust the pH to 8.0 with 1 N NaOH (Merck). Store the solution at 4  C. 15. Elution buffer: TE buffer along with 1% SDS (HiMedia). 2.3 Sonication and DNA Extraction

1. 5 M and 1 M NaCl (Molychem). 2. Proteinase K: 0.2 μg/mL working stock (Roche, Cat. no. 0311536001). 3. Ribonuclease A: 0.2 mg/mL working stock (Sigma Aldrich, Cat. no. R6513). 4. Phenol : chloroform : isoamyl alcohol [25:24:1] (Sigma Aldrich, Cat. no. 2069). 5. Absolute ethanol (MP chemicals). 6. 80% ethanol (ethanol in 0.1% DEPC-treated water).

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Real-Time PCR

1. SYBR Green no. A25742).

Master

mix

(Life

Technologies,

Cat.

2. ChIP-specific DNA primers (Sigma Aldrich). 3. DEPC water (Sigma Aldrich). 4. Extracted DNA samples. 2.5

Instruments

1. Probe Sonicator (QSonica Q125). 2. Ice bath (Labman). 3. Water bath (Technosys). 4. Refrigerated centrifuge (Eppendorf). 5. Biosafety cabinet class 2 (ESCO, Singapore). 6. Phase contrast fluorescent microscope (Carl Zeiss, Germany). 7. Step One Plus Real-time PCR machine (Applied Biosystems, USA). 8. CO2 incubator (Thermo Fisher, USA). 9. Gel documentation system (BioRad, USA).

3

Methods

3.1 Differentiation of hESC and hiPSC into Neuronal Lineage

1. Culture hESC and hiPSC in feeder-free conditions using vitronectin-coated 60 mm tissue culture dish and expand the cells to 80–85% confluency using E8 medium. Passage the cells in growing phase in 1:4 or 1:6 ratio. Do not prewarm E8 media at 37  C; instead warm the media at room temperature (RT) (see Notes 1–3). 2. To begin differentiation, remove the E8 media from the well and give a wash with DPBS. Discard the DPBS and add 500 μL of 0.5 mM EDTA solution to the cells and keep them at 37  C/5% CO2 for 3–4 min with gentle swirling every 2 min (see Note 4). 3. Discard the EDTA solution and collect the cells in spent E8 media. Spin these cells at 1000 rpm/5 min/RT. Discard the supernatant completely and for 6
60 mm dishes, add around 6 mL of EB medium containing 60 μL of ITS and 30 μL of geltex (requirement per 60 mm dish: 1 mL of EB medium + 10 μL ITS + 0.5 μL geltrex). 4. Triturate the cells in the media to make single-cell suspension (suspension containing around three to four million cells) and make drops of 15-μL drops of cell suspension on the inverted lid of 60 mm culture dish. Add 3–4 mL of DPBS in the culture dish and place the lids having cell suspension drops onto the culture dish having 2–3 mL of DPBS. Around 30–35 drops (containing almost 100 cells/per drop) per plate can be made.

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Make almost 6–8 plates in similar pattern. Incubate hanging drops at 37  C/5% CO2 for 2 days. Make sure that the drops do not dry as dehydration will lead to cell death, the DPBS inside the plate will ensure the drop do not dry. 5. After 48 h, propagate the hanging drops containing the embryoid bodies onto geltrex-coated dishes. Coat the 35-mm dish with 1 ml of 0.5
geltrex in cold DMEM and incubate at 37  C/5% CO2 for 1 h. After 1 h, remove the DMEM and add 25–30 EBs per 35-mm dish along with 2 mL of differentiation media (see Note 5). 6. For day 0 to day 3 of neuronal differentiation, culture the attached cells in 2 mL NDM1 media and add 10 ng/mL DKK1, 20 ng/mL bFGF, 2 μM LDN193189 hydrochloride, and 2.5 μM SB435142 hydrate (see Note 6). 7. Day 4 to day 7 of neuronal differentiation: To these proliferating neuroectodermal cells, add 50 ng/mL SHH, 5 μM RA, and 20 ng/mL of bFGF in the NDM1 media. 8. Day 8 to day 11 of neuronal differentiation: Allow the neuroepithelial cells to grow in reduced amounts of morphogens, add 25 ng/mL SHH, 2.5 μM RA, and 20 ng/mL of FGF-4 in the NDM1 media. 9. Day 12 to day 20 of neuronal differentiation: Change the culture medium to NDM2 and propagate the differentiated cells only in this media (see Fig. 1) (see Note 7). 3.2 Harvesting Differentiated Cells for ChIP Sonication and Preparation of the Sample 3.2.1 Harvesting the Cells for ChIP Assay

1. Keep the 60-mm culture plate containing the cells on ice and discard the spent media (see Note 8). 2. Wash the cells with 500 μL ice cold 1
PBS twice, add fresh 1 mL of 1
PBS, and scrape of the cells, collect all the cells in a conical plastic centrifuge tube. 3. Add 1% of formaldehyde (final concentration) to the conical centrifuge tube and make up the volume up to 9 mL with chilled 1
PBS solution. 4. Incubate the cells with formaldehyde at RT for 10 min under rotatory conditions. 5. Add 1 mL of 1.25 M glycine to this tube to quench the formaldehyde and incubate for 10 min at RT under rotatory conditions. 6. Centrifuge the fixed cells at 2500
g for 10 min at 4  C. 7. Wash the fixed cells twice with 4–5 mL of 1
PBS at 2500
g for 10 min at 4  C. 8. Discard the supernatant completely and add 500 μL to 1 mL of complete ChIP lysis buffer (ChIP lysis buffer + PIC + PMSF), transfer lysate to a 1.5-mL vial, and process the sample for sonication immediately (see Note 9).

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3.2.2 Sonication Using Probe Sonicator

1. Keep the sample tube in a container submerged in ice completely. 2. Gently place the probe inside the vial, so that it is just above the bottom of the tube and does not touch the walls of the vial. 3. Set the amplitude at 40% and keep the sonics/pulse on and off for 30 s each for a total of 10 min. Repeat this for 5–6 cycles with a time interval of 10 min (see Note 10). 4. Once the chromatin is sheared, spin the samples at maximum speed 21,130
g for 10 min at 4  C, collect the supernatant in a fresh vial, and label it as the sheared chromatin DNA sample. Aliquot 10% volume in a separate vial and label it as the ChIP input sample. 5. Check the sonicated chromatin on 1.5% agarose gel to ensure that you have generated DNA fragments between 100 and 500 bp; thereafter, use it for the pull down. If the fragment size is too small or too large, then you need to adjust your sonication settings or amplitude or incomplete cell lysis. 6. Process the sample for chromatin immunoprecipitation. You may use the sheared chromatin up to 6 months for ChIP if it was stored at 80  C. The longer the sample stays under refrigerated conditions, the sheared chromatin may not give efficient results. Store the samples at 80  C. We recommend using the sonicated sample for chromatin immunoprecipitation without delay.

3.3 Chromatin Immunoprecipitation

Bertani et al. were one of the few groups to initially publish protocol of ChIP for embryonic stem cells [7]. We have followed the same protocol with few modifications. Day 1: Antibody Complex-Conjugate Formation

1. Use a concentration of 1.5 mg/mL of Dyna beads per immunoprecipitation reaction. Take 50 μL of Dyna beads in a fresh 1.5 mL vial (see Note 11). 2. Keep the vials containing beads on a magnetic rack (DynaMag 2), the beads will be collected on one side within 10–15 s. Discard the solvent and add 200 μL of wash buffer (WB) to the beads. 3. Remove the vial from the magnetic rack and vortex it thoroughly. Keep the vial back onto the magnetic rack. Once the beads settle down, remove the wash buffer. 4. Wash the beads with 200 μL of WB for four times in the same pattern (see Note 12). 5. Add 100 μL of primary antibody solution (typically the concentration of each antibody varies and but use the final concentration between 5 and 10 μg/mL) prepared in 0.5%BSA

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solution to the washed beads and seal the vial thoroughly. Keep the vial on a rotator or a rocker to allow the antibody-beads complex to form for 6 h or overnight at 4  C. Follow a similar protocol for appropriate isotype antibody (we used rabbit IgG antibody). You need to prepare the antibody-bead complex depending on the number of samples (or time points or conditions). 6. After the incubation, add 30–35 μg of sheared chromatin to this complex and vortex the sample thoroughly. Add the sheared chromatin from different timepoints or treatments. Incubate at 4  C for overnight on a rocker or a rotator. The overnight incubation will allow efficient interaction between chromatin and antibody : bead complex. Day 2: Washing

1. Add the solutions given below to the chromatin and antibody: bead complex (each wash of approximately 30 s) and vortex. In order to wash, keep the vial on magnetic rack and discard the supernatant once the beads are collected on sides. Repeat this process for all our washes. Do not discard supernatant when the tube is not placed in magnetic rack. 2. Wash the beads with 200 μL of 0.5% BSA solution twice. 3. Wash the beads with 500 μL of ChIP lysis buffer four times. 4. Wash the beads with 500 μL of IP1 buffer four times. 5. Wash the beads with 500 μL of IP2 buffer four times. 6. Wash the beads with 500 μL of TE buffer four times (see Note 13). Day 2: Elution and Reverse Crosslinking

1. After the last wash, spin the sample at 4000
g for 10 min at 4  C to remove any residual TE buffer. 2. Add 210 μL of elution buffer to the sample and incubate at 65  C in water bath for a period of 30 min (see Note 14). 3. Centrifuge the sample at 21,130
g for 5 min at RT. 4. Transfer supernatant to a fresh 1.5-mL vial and discard the beads. 5. Keep the sample for reverse crosslinking at 65  C water bath for 6–15 h (see Note 15). 6. Thaw 50 μL of ChIP input DNA (kept aside after sonication) and add 150 μL (three volumes) of elution buffer and mix. Reverse crosslink this ChIP input along with the pull down sample in a water bath at 65  C.

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Day 3: RNase A, Proteinase K, and DNA extraction

1. Cool down the samples to RT, add 4 μL of RNase A (final concentration 0.2 mg/mL) and 3 μL of 1 M NaCl (final concentration 15 mM) per sample, incubate in water bath for 2 h at 37  C. 2. Next, add 2 μL of Proteinase K (final concentration 0.2 μg/ mL) per sample and incubate in water bath for 2 h at 55  C. 3. Bring down the temperature to RT and proceed with DNA extraction using phenol:chloroform:isoamyl alcohol method. 4. Add 400 μL of phenol:chloroform:isoamyl alcohol (25:24:1) which is adjusted to pH 8.0 for every 200 μL volume of sample and mix vigorously for about a minute. Allow the tube to stand at RT for 2 min. 5. Centrifuge at 21,130
g for 10 min at 4  C. Collect the aqueous upper layer in a fresh vial. Always keep the samples on ice from hereon. 6. To this add 800 μL of chilled 100% absolute ethanol and 8 μL of 5 M NaCl solution per sample. NaCl aids in better DNA precipitation. 7. Spin the DNA at 21,130
g for 10 min at 4  C. Incubate the samples at 20  C overnight or at 80  C for 1 h for better DNA precipitation. 8. Spin the DNA at 21,130
g for 10 min at 4  C, discard the supernatant and wash the pellet with 80% ethanol at 7500
g for 7 min at 4  C. 9. Repeat the washing steps again, if the pellet is too small, avoid the second washing step. 10. Remove the supernatant completely and allow the pellet to air dry. Add appropriate amounts of 10 mM Tris (typically ranges from 25 to 35 μL) buffer to the dried pellet. Quantify the DNA before proceeding to qPCR (see Note 16). Day 4: ChIP-qPCR

1. Quantify the DNA at 260 nm using UV spectrophotometer; use 10 ng of gDNA as the starting material for amplifying by PCR. 2. Prepare a set of reactions for the genes which you want to investigate are controlled by PcG proteins. We have given a table below for a single PCR:

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Components

Volume per reaction

dDNA (10 ng)

Variable for 10 ng

Forward primer (250 nM)

0.5 μL

Reverse primer (250 nM)

0.5 μL

SYBR green mix

5 μL

DEPC water

Variable

Total

10 μL

3. Add appropriate volumes of SYBR Green master mix, 0.25 μM sense and antisense primers each along with calculated volumes of DNA material (input or pull down) and DEPC water to make up the total volume. 4. Set up the thermal cycles consisting of the following conditions: initial denaturation at 95  C for 2 min followed by 50 cycles of denaturation at 95  C for 15 s, annealing (specific to each primer set) for 15 s, and extension at 72  C for 15 s and data acquisition, followed by melt curve to check the homogeneity of the PCR products. The primers were standardized for each gene (see Note 17). 5. Once the PCR run is complete, analyze the results using ChIP input method (see Note 18 for understanding the ChIP calculations using percent input method). 3.4

Results

Chromatin immunoprecipitation provides a valuable method for the examination of chromatin-based developments during cellular differentiation. Figures 2 and 3 show the stage-specific neuronal differentiation from human pluripotent stem cells. ChIP-qPCR data analysis by percent input method is a widely accepted method for data analysis, where the amount of pulldown by RING1B/ BMI1/H2AK119ub1 is compared against input (which is 100%) as well as isotype (which should be least). Figure 4 shows the dynamic PcG proteins RING1B-BMI1 occupancy on the promoter (1000 bp upstream of transcription start site) of HOXA2 during the course of differentiation from day 0 to day20. As the neuronal differentiation proceeds, the occupancy of PRCI proteins (RING1B-BMI1 and EZH2) and histone modification they catalyze (H2AK119ub1) increases at HOXA2 promoters. Similar to HOXA2, we can look at the promoter occupancy of your gene of interest by PcG proteins by ChIP-qPCR. When we couple the ChIP-qPCR data with gene expression or protein expression data, we can better understand the role of PcG proteins in regulating gene expression. Some of the prerequisites for successful ChIP include ChIP-validated antibodies, known targets, and sheared

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Fig. 2 Neural differentiation days (day 0 to day 15). The image depicts the cells transformed into neuroectodermal, neural epithelial cells, and neural progenitors on various days. The yellow-colored arrows indicate the formation of neuroepithelial morphology and neural rosettes. The formation of rosette-like arrangement is a typical indication of cells transforming into neuroepithelia. The cells observed under 100
magnification (scale: 100 μm)

chromatin. ChIP can be combined with multiple downstream processes like qPCR, microarray, and sequencing, thus giving huge amount of information which can help us in understanding the dynamic associations of proteins, DNA/RNA, and protein modifications which regulate processes such as proliferation and differentiation.

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Fig. 3 Neural differentiation days (day 16 to day 20). The image depicts the cells transformed early neural progenitors through day 16 to day 20. The yellow-colored arrows indicate the formation of different neural rosettes. The typical formation of classical rosettes is marked by yellow-colored arrow in (a), (b), and (c), and the images indicate smaller number of rosettes seen in one particular field. Images (d)–(h) indicate the neuronal projections; the arrows show projections emerging from various rosettes structure giving it an appearance of a typical neuron. The cells observed under 200
and 400
magnification (scale: 200 μm and 400 μm)

Fig. 4 Example of promoter occupancy of HOXA2 gene by RING1B, BMI1, H2AK119ub1, and EZH2 during neuronal differentiation. The Ct values for HOXA2 in undifferentiated hESC/hiPSC and differentiated pull down samples were normalized using ChIP input Ct. Statistical analysis performed using unpaired Student’s t-test (two-tailed), standard error of mean (SEM) indicated in error bars. Statistical significance is denoted as *, where *p < 0.05, **p < 0.01, ***p < 0.005. HOXA2 is the known genomic target whose promoter is bound by the Polycomb group proteins

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Notes 1. The vitronectin coating should be prepared in prewarmed DPBS for better attachment and incubate the plate with the coating solution for about 45 min to an hour at RT or 37  C. Monitor the hESC and hiPSC cultures as continuous cell growth may result in spontaneous differentiation and will not give rise to neuronal differentiated cells. If the population of differentiated cells is more than 10%, remove the differentiated cells or start with fresh undifferentiated culture. The cells should be maintained in undifferentiated state by providing media change daily. 2. Upon reaching 80% confluency, the E8 medium needs to be replaced with fresh 2.5 ml of E8 medium for half an hour before passaging. Then collect this spent media and wash the cells with 500 μL of DPBS solution twice. 3. Begin with six to eight million cells for differentiation since lot of cell death occurs during hanging drop method and initial 4 days of differentiation. 4. Prewarm the EDTA in a water bath set to 37  C, if the EDTA solution is cold the detachment of pluripotent stem cells will not occur from the vitronectin substrate. The EDTA treatment makes the cells round in shape and shiny in appearance when observed under microscope, the cells at the colony edges start to detach first. Remove the EDTA as soon as most of the cells in the colony start appearing rounded. Do not over treat the cells with EDTA otherwise the cells will die immediately. 5. Geltrex should always be diluted using cold media. Geltrex stock should always be kept in ice to avoid the jelly-like formation of the stock, if the geltrex solidifies, use another aliquot. Once you coat the plate with geltrex, immediately incubate the plates at 37  C. To collect the EBs, use a wide bore tip or cut the tip and collect the EBs individually to avoid disintegration of the EBs. Use NDM1 media to collect the hanging drops gently. 6. Initial days into differentiation are very crucial, the cells can easily come off the plates; hence, add the medium from the sides of the culture dish very gently, to prevent dislodging the attached cells from the surface. While changing the media, do not apply direct force onto the cells otherwise the cells will detach from the plate. Give media change every alternate day. 7. The success of generating successful EBs and robust neuronal differentiated cultures depends critically on the quality of hESC and hiPSC cell lines. The colonies should show hallmarks of pluripotency such as distinct colony borders, epithelial

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morphology, and compact colonies; these are phenotypic indicators that cells are in undifferentiated state. Some common tests for pluripotency and differentiation for all three germ lineage include immunostaining for pluripotent markers: OCT4, NANOG, and SOX2; embryoid body formation followed by Western blot or RT-PCR analysis for lineage-specific genes. 8. Approximately 50,000–100,000 cells are required to begin with ChIP assay. Depending on the downstream application of ChIP, you can adjust the amount of cells for the same. Typically, for ChIP Seq, you need more amount of cells as a starting material. 9. Precisely note the volume of ChIP lysis buffer, as this volume plays an important role in analysis of qPCR data. At the end of washes, the pellet becomes very sticky and tends to stick to the plastic walls of the conical tube. Be gentle while breaking the pellet or collecting it in the complete ChIP lysis buffer. If you want to store the pellet and plan to sonicate later, you can snap freeze the pellet in the buffer using liquid nitrogen. The stored samples has a shelf life of up to 3 months. It is advisable to process the sample immediately or day after for sonication. 10. Check the instruction manual of your probe sonicator before using the instrument. Check if your instrument allows you to set specific amplitude. After every cycle, vortex the sample and give a quick spin. Also ensure that the vial should be cold throughout the process. It becomes difficult when ice starts to melt, so keep refilling the ice and discard the ice-cold water in between the procedure. If 40% amplitude does not shear the chromatin, you need to standardize the amplitude which gives you sheared chromatin between 150 and 300 bp. 11. Prior to aliquoting Dyna beads, vortex the stock vial thoroughly for 1–2 min. 12. While adding and removing buffers, always keep the vial on the magnetic rack. While working with many vials, you can detach the rack from the magnet and invert it gently to mix the beads and reagents, then you can attach it to the magnet. Invert the vials on the rack repeatedly till all the beads settle at the end of the vial. A separate IP reaction needs to be set up for isotype which will follow the same set of steps as that of samples. 13. During washes, invert the rack 4–6 times to collect all the beads at the bottom. Sometimes during washes the beads tend to stick at the top or the cap of the vial, invert the rack few more times until all the beads come down. 14. During incubation, vortex the sample for 30 s every 2 min to improve the recovery of the eluate.

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15. Do not exceed the reverse crosslinking treatment for more than 15 h because longer times of reverse crosslinking results in increased noise in the microarray analysis. 16. Over drying of pellets can make it difficult to resuspend, or liable to flake and peel away from the side of the tube. The concentration of DNA after pull down decreases ten times than that of input DNA. You can start with 5–10 ng of gDNA for qPCR reactions. 17. The primers need to be standardized prior to its use for ChIPqPCR. The primer efficiency can be determined using standard curve method using serially dilution technique. We used fivefold serial dilution of gDNA of hESC sample. Given below is the primer sequence for HOXA2 gene used for ChIP-qPCR.

Gene

Primer Sequences (50 ! 30 )

HOXA2 AGGAAAGATTTTGGTTGGGAAG AAAAAGAGGGAAAGGGACAGAC

Size (bp)

Accession ID

138

NC_000007.14

18. We have used “Percent ChIP Input” method for analysis. With this method, the signals collected (Cts) from ChIP need to be subtracted by signals collected (Cts) from an input sample for normalization. The input sample indicates the total amount of chromatin used for the ChIP (typically 1–10% of starting cell lysate volume can be used as the input). The input DNA is defined as the aliquot of chopped chromatin prior to immunoprecipitation and will be used to normalize the sample to the amount of chromatin used for each ChIP. Further the next step involves adjusting the Ct value of the input sample by converting the dilution factor 10 into logarithmic quantification cycles. The value obtained needs to be subtracted from Ct input; this would be termed as adjusted Ct, and the promoter occupancy will be now determined with respect to the normalized input Ct [6].

Acknowledgments This work is supported by DST-SERB, India, and NMIMS (deemed-to-be) University. References 1. Dang L, Tropepe V (2006) Neural induction and neural stem cell development. Regen Med 1:635–652. https://doi.org/10.2217/ 17460751.1.5.635

2. Kriegstein A, Alvarez-Buylla A (2009) The glial nature of embryonic and adult neural stem cells. Annu Rev Neurosci 32:149–184. https://doi. org/10.1146/annurev.neuro.051508.135600

Neuronal Differentiation and ChIP-qPCR 3. Kondo T, Isono K, Kondo K et al (2014) Polycomb potentiates Meis2 activation in midbrain by mediating interaction of the promoter with a tissue-specific enhancer. Dev Cell 28:94–101. https://doi.org/10.1016/j.devcel.2013.11. 021 4. Luis NM, Morey L, Di Croce L, Benitah SA (2012) Polycomb in stem cells: PRC1 branches out. Cell Stem Cell 11:16–21. https://doi.org/ 10.1016/j.stem.2012.06.005 5. Yan Y, Chen H, Costa M (2009) Chromatin immunoprecipitation assays. Methods Mol

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Biol:567. https://doi.org/10.1007/978-160327-414-2 6. Haring M, Offermann S, Danker T et al (2007) Chromatin immunoprecipitation: optimization, quantitative analysis and data normalization. Plant Methods 3:1–16. https://doi.org/10. 1186/1746-4811-3-11 7. Bertani S, Kan A, Sauer F (2008) Chromatin immunoprecipitation from human embryonic stem cells. J Vis Exp:1–6. https://doi.org/10. 3791/780

Methods in Molecular Biology (2022) 2520: 135–150 DOI 10.1007/7651_2021_448 © Springer Science+Business Media, LLC 2021 Published online: 02 November 2021

Directed Differentiation of Human Pluripotent Stem Cells into Inner Ear Organoids Yoshitomo Ueda, Stephen T. Moore, and Eri Hashino Abstract The sensory epithelia of the inner ear contain mechanosensitive hair cells that detect sound and head acceleration. This protocol details a 3D differentiation method to generate inner ear organoids containing sensory epithelia with hair cells. Human pluripotent stem cells are aggregated in low-binding 96-well plates and treated in chemically defined media with extracellular matrix to promote epithelialization. Small molecules and recombinant proteins are applied in a stepwise manner to recapitulate the morphogenic cues (BMP, TGF-β, FGF, and WNT) present during inner ear development in vivo. These treatments induce the sequential formation of nonneural ectoderm, otic-epibranchial progenitor domain, and otic placodes. The derived otic placodes then undergo self-guided morphogenesis to form otic vesicles, which eventually give rise to sensory epithelia containing hair cells and supporting cells, as well as neurons with synaptic formations to hair cells. This human stem cell–derived inner ear organoid system provides an ideal platform to study human inner ear development and disease in vitro. Key words Inner ear, Human, Mechanosensitive hair cells, Organoid, Otic development, Pluripotent stem cells, Supporting cells

1

Introduction Sensory hair cells in the inner ear exhibit unique mechanosensitive characteristics with hair bundles on their apical surface to detect the inclination and acceleration of the head, and sound. In both vestibule and cochlea, hair cells are surrounded by supporting cells and transmit electric signals to afferent ganglion neurons (Fig. 1a). Damage to hair cells due to loud noise, ototoxic drugs, aging, and genetic mutations may lead to irreversible hearing loss and/or balance disorders because hair cells do not regenerate in humans [1, 2]. At the early stage of embryonic development following neural tube closure, bone morphogenic protein (BMP) signaling drives the lateral domain of the ectoderm epithelium to form nonneural ectoderm (NNE) [3–6]. Subsequent attenuation of BMP signaling and activation of fibroblast growth factor (FGF) signaling induces the formation of a thickened epithelium from the NNE. This region gives rise to both otic and epibranchial lineages and is

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Fig. 1 Schematic of the ear which is responsible for hearing and balance. (a) Each sensory epithelium has hair cells and surrounding supporting cells. The hair cells are innervated with the neurons forming the vestibulocochlear nerve. (b) The outer ear, the middle ear, and the inner ear. The excitation of the hair cells is transmitted to the brainstem through the vestibulocochlear nerve. (c) The membranous labyrinth in the inner ear shown with the sensory epithelia labeled orange. Each semicircular canal has a dilated portion at the base (ampulla) which contains the sensory organ termed crista ampullaris. The utricle and the saccule have different sensory epithelium termed macula statica. The cochlea contains spiral-shaped sensory epithelium called the organ of Corti. Hair cells in the cristae and the maculae detect angular and linear acceleration of the head, respectively, while hair cells in the organ of Corti detect sound

termed the otic-epibranchial progenitor domain (OEPD) [7– 11]. Within the OEPD, WNT signaling plays an important role; elevated WNT activity promotes an otic fate while reduced pathway activation leads to epibranchial induction [12–14]. Shortly after this specification, the otic placode invaginates to form the otic pit which progressively submerges into the surrounding mesenchyme. Eventually, the otic pit pinches off from the surface epithelium forming an enclosed vesicle termed the otocyst. The mature inner ear, with its great diversity of cell types [13, 15] and structural complexity (Fig. 1b, c), is derived almost entirely from the otocyst, a structure composed of relatively uniform progenitors [16, 17]. By recapitulating the morphogenic cues of inner ear development in vivo, we previously developed 3D organoid differentiation methods to generate inner ear sensory epithelia, including hair cells, supporting cells, and innervating neurons, from mouse pluripotent stem cells (PSCs) [18–21] and human PSCs [22]. Cultures begin by making aggregates of the mouse or human PSCs in low cell-adhesion 96-well U-bottom plates. Matrigel is added as a source of extracellular matrix to coat the surface of the aggregates

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to promote epithelization. Over the next several days, small molecules and recombinant proteins are applied to the aggregates in a stepwise manner to modulate BMP, TGF-β, FGF, and WNT signaling. These treatments result in the sequential formation of NNE, OEPD, and otic placodes (Fig. 2). The otic placodes are evaginated from the core of the aggregates and form otic-pit-like ruffling on the aggregates’ surface. These tissues subsequently pinch off and form otocyst-like structures embedded in the surrounding mesenchymal tissues (Fig. 3a). The majority, if not all, of cells in otic vesicles express canonical otic marker proteins such as PAX2, PAX8, ECAD, EPCAM, SOX2, and JAG1 (Fig. 3b–e) as well as FBXO2 at later stages. Following self-guided differentiation in a maturation medium, ATOH1+ hair cells appear on the lumen of the vesicles. These nascent hair cells first appear around day 14 in the mouse inner ear organoids, and around day 40 in human inner ear organoids. As the hair cells mature, canonical hair cell markers such as MYO7A, PCP4, POU4F3, ANXA4, ESPN, and SOX2 can be detected (Fig. 3e). Furthermore, hair bundles containing actinrich stereocilia and microtubule-rich kinocilium are found on their apical surface. As with hair cells in vivo, hair cells in inner ear organoids are surrounded by supporting cells and form synaptic connections around their basolateral regions. Electrophysiology experiments suggest that these hair cells exhibit mechanosensitivity and intrinsic electrical properties that resemble those of native vestibular hair cells in vivo [22, 23]. In this chapter, we describe a detailed protocol for generating inner ear organoids from hPSCs. The inner ear cell types in the 3D structure provide an alternative in vitro resource for the investigation of human inner ear development and diseases, as well as for drug screening and therapy development. Moreover, the organoid system has the potential to contribute as donor tissues in regenerative therapies for hearing loss and balance disorders.

2 2.1

Materials Reagent Setup

Aliquoting should be done in a sterile environment in the biosafety cabinet. Aliquots are intended to be prepared in 0.5-mL tubes, unless otherwise noted. Reagents should be stored at 4  C except as otherwise indicated (see Note 1). 1. Human pluripotent stem cells (hPSC): Embryonic stem cells (hESC) and human induced pluripotent stem cells (hiPSC) are applicable. We primarily use the WA25 hESC line (WiCell) and CRISPR-engineered cell lines using the WA25 as a parental cell line. Note that BMP-4 treatment on day 0 may be necessary depending on the cell lines and its concentration should be optimized for the other hPSC lines.

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Fig. 2 Scheme of the differentiation protocol for generation of inner ear organoid from hPSCs. (a) The overview of the human inner ear organoid culture highlighting relevant signaling pathway. (b) Experimental procedure of the culture. hPSCs human pluripotent stem cells, NNE nonneural ectoderm, OEPD otic-epibranchial progenitor domain, HC hair cells, SC supporting cells, E8 flex Essential 8 flex medium, E6 Essential 6 medium, OMM Organoid maturation medium

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Fig. 3 Morphology of aggregates during the culture and the immunostaining showing inner ear lineage markers in human inner ear organoids. (a) Phase-contrast images of aggregates at early stages (from day 2 to day 20). (b–e) Immunostaining with the otic progenitor markers at d20 (b–d) and d25 (e), and hair cells and supporting cells with neural fibers at d60 (f). Scale bars, 200 μm (a); 50 μm (b–f)

2. Essential 8 Flex Medium (E8f; Gibco #A2858501). 3. Essential 6 Medium (E6; Gibco #A1516401). 4. Advanced DMEM/F-12 #12634028).

(A-DMEM/F12;

Gibco

5. Neurobasal Medium (Gibco #21103049). 6. DMEM/F-12, HEPES (Gibco #11330032). 7. Dulbecco’s phosphate-buffered saline (DPBS) without calcium and magnesium (Gibco): Store at RT. 8. 0.5 M ethylenediaminetetraacetic acid (EDTA; Invitrogen): Prepare a 0.5 mM solution by diluting 50 μL of 0.5 M EDTA in 50 mL of DPBS. Store at RT.

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9. Vitronectin (VTN-N) Recombinant Human Protein, Truncated (Gibco #A14700): Aliquot 20–40 μL of 0.5 mg/ mL VTN-N. Store at 80  C for up to 2 years. 10. RevitaCell (RvC) supplement (100; Gibco #A2644501): Thaw a bottle at 4  C. Then aliquot the arbitrary volume (e.g., 250 μL for 25 mL E8f) in 1.5-mL tubes and store at 20  C for up to 1 year. 11. GlutaMAX supplement (Gibco #35050061): Store at RT. 12. B-27 Supplement (50), minus vitamin A (Gibco #12587010): Store at 20  C. Use in 2 weeks after thawing or make aliquots and store at 20  C. Avoid freeze–thaw cycle. 13. N2 Supplement (100) (Gibco #17502048): Store at 20  C. Use in 2 weeks after thawing or make aliquots and store at 20  C. Avoid freeze–thaw cycle. 14. Accutase (Gibco #A1110501): Aliquot in 2-mL tubes and store at 20  C for up to 1 year. After thawing, store the aliquot at 4  C for up to 1 month. 15. Matrigel, Growth Factor Reduced, LDEV-free (Corning #354230): Thaw a bottle of Matrigel overnight or for several hours at 4  C. Chill wide-mouth P1000 pipette tips at 20  C. Aliquot completely thawed Matrigel into 1.5-mL tubes on ice. Aliquot volume varies by protocol (i.e., for a 2% solution, aliquot 400 μL of Matrigel for 20 mL of the media on d0; 500 μL Matrigel for 50 mL of the media on d11). Keep stock bottle and tubes on ice when making aliquots to prevent warming above 4  C. Store the aliquots at 20  C for up to 1 year. 16. Basic fibroblast growth factor (bFGF; STEMCELL Technologies #78003): Prepare 200 ng/μL stock solution. Dissolve 50 μg of bFGF in 250 μL of DPBS and gently mix well with pipetting. Aliquot at 2 and 11 μL. Store at 80  C for up to 3 months. 17. BMP-4 (Reprocell #03-0007): Prepare 100 ng/μL stock solution. Dissolve 10 μg of BMP-4 with 100 μL of 4 mM HCl and mix well. Aliquot at 2 μL and store at 80  C for up to 1 year. 18. 10 mM Y27632 (Stemgent #04-0012-02): Prepare 25 μL aliquots. Store at 20  C for up to 1 year. 19. 10 mM SB431542 (SB; Stemgent #04-0010-05): Prepare 32 μL aliquots and store at 20  C for up to 1 year. 20. 10 mM LDN193189 (LDN; Reprocell #04-0074-02): Prepare 2 μL aliquots and store at 20  C for up to 1 year. 21. 10 mM CHIR99021 (CHIR; Reprocell #04-0004-02): Prepare 16 μL aliquots and store at 20  C for up to 1 year. 22. 2-Mercaptoethanol (2-ME, 55 mM; Gibco #21985023)

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23. Normocin (500; Invivogen #ant-nr-1): Store at 20  C. Use in 1 month after thawing. 24. 0.4% Trypan Blue: Store at RT. 2.2 Preparation of Culture Media

1. E8f: To prepare a complete Essential 8 flex (E8f) stock, thaw a bottle of Essential 8 flex supplement (50) at room temperature (RT) for ~1 h or overnight at 4  C. DO NOT thaw at 37  C. In a biosafety cabinet, combine Essential 8 flex supplement (50) into a bottle of Essential 8 flex Basal Medium (500 mL). Rinse the supplement bottle with the basal medium by pipetting several times. Label the stock bottle “Supplement Added” and the date or immediately prepare aliquots in 50-mL conical tubes with appropriate labels (recommended). Store the complete medium at 4  C for up to 3 weeks. 2. E8f + Normocin (E8fn): Add 100 μL of Normocin to 50 mL of E8f. Store medium at 4  C and warm at RT before use. DO NOT warm at 37  C. 3. E8fn + RevitaCell supplement (E8fn + RvC) Add RevitaCell supplement to E8fn at 1:100 dilution and mix well. Store medium at 4  C and warm at RT before use. DO NOT warm at 37  C. 4. E6 + Normocin (E6n): Add 100 μL of Normocin to 50 mL of E6. Store medium at 4  C and warm at RT before use. DO NOT warm at 37  C. 5. E6n + 2% Matrigel, 10 μM SB, and 4 ng/mL bFGF (E6n-MSF): To prepare 20 mL of E6n-MSF, aliquot 20 mL of E6n into a conical tube. Use a prechilled P1000 pipette tip to add chilled E6n directly to a 400 μL aliquot of frozen Matrigel. Triturate and repeat until the entire frozen aliquot is dissolved, then vortex to dissolve the Matrigel completely. Add 20 μL SB and 4 μL of ten-fold diluted bFGF to the solution and mix well. (Optional: For certain cell lines, addition of BMP-4 may be necessary and/or improve efficiency. The concentration of BMP-4 should be optimized and may vary from 10 1 to 10 ng/mL final concentration). The medium should be freshly prepared immediately prior to performing the d0 treatment and used promptly after warming to RT. DO NOT warm to 37  C. 6. Organoid Maturation Medium (OMM): To prepare 50 mL of OMM, mix the following reagents in a 50 mL conical tube: 24.5 mL A-DMEM/F12, 24.5 mL Neurobasal, 500 μL GlutaMAX, 500 μL B-27 supplement, 250 μL N-2 supplement, 90 μL of 55 mM 2-ME, and 100 μL Normocin. Store OMM at 4  C for up to 1 week. Warm OMM to RT or 37  C before use (see Note 2).

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Equipment

1. Nunc Delta Surface Cell Culture Treated 6-well plates (Thermo Scientific 140675) to maintain hPSC cultures. 2. 100-mm dishes: For stationary aggregate culture on d11 and later. Use 100-mm dishes with 20 mm height (e.g., Corning 353003) rather than 15 mm height. Deeper dishes can hold a larger volume of culture medium and are less prone to accidental medium/aggregate spilling. 3. 60-mm dishes. 4. Low cell-adhesion (also known as low binding) U-bottom 96-well plates (e.g., Nunclon Sphera Plates, Thermo Fisher Scientific 174932). 5. Low cell-adhesion 6-well plate (e.g., Corning 3471). 6. Liquid nitrogen tank for storage of cryopreserved hPSCs. 7. Biosafety cabinet: Make sure to handle all culture media, reagents, and aliquots in biosafety cabinets. Make sure to handle all hPSC and organoid culture in biosafety cabinets, except for procedures such as centrifuging and imaging. 8. A vacuum aspirator with a glass Pasteur pipette equipped in the biosafety cabinet. 9. CO2 incubator set at 37  C, 5% CO2, and humidified with sterile water. 10. Water bath. 11. A benchtop centrifuge for 2-mL tubes. 12. A large centrifuge for 96-well plates: set the temperature to RT. 13. Vortex machine. 14. Automated cell counter and compatible cell counting slides. (Optional: a hemocytometer slide cleaned with ultra-pure water and then 70% ethanol can be used.) 15. Inverted microscope. 16. Single-channel pipettes (20-μL, 200-μL, and 1000-μL) and multichannel pipettes (200-μL). 17. Autoclaved pipette tips (20-μL, 200-μL, and 1000-μL), autoclaved microcentrifuge tubes (0.5-mL, 1.5-mL, and 2-mL), and conical tubes (15-mL and 50-mL). 18. Disposable pipette basins. 19. Scissors: To cut the pipette tips to make their mouth wide. Keep in the biosafety hood and expose to ultraviolet rays to sterilize before use. 20. Sprayer containing 70% ethanol. 21. Sterile H2O.

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Methods All steps should be processed in the sterile environment in the biosafety cabinet unless otherwise noted. The medium should be warmed to room temperature (RT), not 37  C except as otherwise indicated.

3.1 hPSC Maintenance and Passaging in FeederFree Condition 3.1.1 Maintenance

3.1.2 Passaging

1. It is possible to skip feeding up to two consecutive days when culturing with E8fn, (e.g., Saturday and Sunday), and up to 3 days total per week (see Note 3). Feed cells or passage cells regardless of the date if the cells are highly confluent and the culture medium becomes amber to yellow. 2. To change medium, aspirate spent medium, and slowly add 2 mL (3–4 mL on Fridays) of E8fn medium to each well of the 6-well plate using a wide-mouth p1000 pipette tip. Passage hPSCs when either the confluency of the cells reaches 70–80%, the cell density becomes too high, or the size of the colony becomes too large. 1. Coat several wells of a Nunc Delta Surface 6-well plate for over 1 h with 1 mL of 5 μg/mL VTN-N (1:100 dilution with DPBS) to each well (see Note 4). 2. Warm E8fn + RvC to RT (see Note 5). 3. Aspirate the spent medium and gently wash cells three times with 1 mL of DPBS using a wide-mouth P1000 pipette tip. 4. Slowly add 1 mL of 0.5 mM EDTA and incubate at 37  C for 5–7 min until the cells start to separate and round up, checking under the microscope. 5. During EDTA incubation, aspirate the VTN-N coating solution from the new plate and add 2 mL of E8fn + RvC to each coated well. 6. Carefully aspirate EDTA without damaging the cells. 7. Gently wash the cells from the surface of the plate with 1 mL of E8fn + RvC. Slowly pipet up and down several times to dissociate cell clumps and create homogenous cell suspension. Avoid creating bubbles in the media. 8. Slowly add an arbitrary volume of the cell suspension into prepared wells of the new plate (see Note 6). 9. Gently shake the plate in all directions several times to disperse the cells in the well. Avoid swirling motions to prevent concentrating the cells to the center. 10. Place the plate in the CO2 incubator. 11. After 16–24 h, change medium with E8fn, without RvC.

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3.2 Generating Human Inner Ear Organoids 3.2.1 Day 2: hPSC Aggregation

All media should be warmed at RT prior to use, unless otherwise noted. The protocol below is intended to generate 96 aggregates. 1. Ensure hPSCs are 50–80% confluent. Refeed cells 2–3 h prior to starting the differentiation culture if the medium looks amber to yellow. 2. For each 96-well plate of organoid culture (see Note 7), prepare a tube of E8fn-Y20: add 22 μL of Y-27632 to 11 mL of E8fn in a 15-mL conical tube. 3. Prepare two tubes of E8fn-Y10: add 1 μL of Y-27632 to 1 mL of E8fn in each 1.5-mL tube. 4. Warm an aliquot of Accutase to RT. 5. Aspirate the medium from one well keeping hPSCs and wash three times with DPBS using a wide-mouth P1000 tip. 6. Gently add 500 μL of Accutase and briefly swirl the plate to expand. Incubate at 37  C for 5–7 min. Ensure the cells have detached from the plate using a microscope before proceeding. 7. Gently wash the cells from the plate using 1 mL of E8Fn-Y10. Collect the cells in a 2-mL tube and gently pipet up and down several times to dissociate cells into single cells (see Note 8). 8. Centrifuge at 100  g for 3 min. 9. Completely aspirate the supernatant. Make sure not to damage the cell pellet. 10. Resuspend the cell pellet with 1 mL of E8fn-Y10. Gently pipet up and down several times to dissociate cell clumps and distribute the cells evenly in E8fn-Y10. 11. In a 0.5-mL tube, mix 50 μL of the cell suspension with 50 μL of trypan blue. Load the mixture onto a cell counting slide and measure the cell concentration by the automatic cell counter or by hand under the microscope. 12. Add the cell suspension into the 15-mL tube containing 11 mL of E8fn-Y20 to acquire a final concentration of 35,000 cells/ mL (i.e., 385  103 cells in 11 mL). For example, if 2  106 cells/mL are counted, add 192.5 μL of the cell suspension in 11 mL of E8fn-Y20 medium. 13. Gently invert the 15-mL tube a couple of times to distribute the cells evenly. Pour the cell suspension into a disposable pipette basin and slowly add 100 μL of cell suspension into each well of a low-cell-binding 96-well plate using a multichannel pipette with wide-mouth pipette tips. Each well contains 3500 cells. 14. Centrifuge the 96-well plates at 120  g for 5 min at RT. 15. Place the plates in the CO2 incubator.

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16. Four hours after seeding, add 100 μL of E8fn (with no Y-27632) to each well using a multichannel pipette and a disposable basin. 3.2.2 Day 0: Transfer Aggregates to Differentiation E6

1. Prepare 20 mL of E6n containing 2% Matrigel, 10 μM SB, and 4 ng/mL bFGF (E6n-MSF). (Optional: add 2.5 ng/mL BMP-4, see Note 9) Warm the media to RT (see Notes 10–12). 2. Warm 6 mL of DMEM/F12 to RT. 3. Using P200 wide-mouth pipette tips and a P200 multichannel pipette, carefully transfer aggregates from the 96-well plates into a 100-mm dish. Discard the used 96-well plates. 4. Transfer all aggregates to a 2-mL tube using a wide-mouth P1000 pipette tip. 5. Gently wash the aggregates 3 times with DMEM/F12. 6. Gently wash the aggregates 3 times with E6n-MSF. 7. Resuspend the aggregates in 1 mL of E6n-MSF. 8. Transfer all aggregates to a 60-mm dish and add additional E6n-MSF to the dish to pool. 9. Using a P200 pipette with a wide-mouth pipette tip, individually transfer each aggregate with 100 μL of the media to the central 6  10 wells of new low-cell-binding 96-well plates (see Note 13). 10. Fill each peripheral well of the 96-well plate with 200 μL of sterile H2O or DPBS. (The peripheral 36 wells are not used and filled with water) (see Note 14). 11. Place the plates in the CO2 incubator for 3 days.

3.2.3 Day 3: bFGF and LDN-193189 Treatment

1. In a 15-mL tube, add 7.5 μL of 200 ng/μL bFGF and 0.6 μL of 10 mM LDN to 10 mL of E6n (E6n-FL). Invert the tube several times to mix. Note that E6n-FL contains 150 ng/mL bFGF and 600 nM LDN (see Note 15). 2. Add 50 μL of E6n-FL to each well with wide-mouth P200 pipette tips and the multichannel pipette. Gently pipet several times to mix. Note that each well contains 150 μL of the medium with 50 ng/mL bFGF and 200 nM LDN as a final concentration. 3. Return the plates to the CO2 incubator.

3.2.4 Day 5: Medium Change and CHIR-99021 Treatment

1. Using P200 wide-mouth pipette tips and a P200 multichannel pipette, carefully transfer aggregates from the 96-well plates into a 100 mm dish. The dish should be kept in the CO2 incubator while washing and preparing the medium.

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2. Wash the central wells of the 96-well plates. Aspirate the residual medium, add 200 μL of sterile H2O, and then completely aspirate. 3. In a 50-mL tube, add 15 μL of 10 mM CHIR, 7.5 μL of 200 ng/μL bFGF, and 0.625 μL of 10 mM LDN to 50 mL of E6n (E6n-CFL). Invert the tube several times to mix. Note that E6n-CFL contains 3 μM of CHIR, 30 ng/mL bFGF, and 125 nM LDN. 4. Transfer all aggregates to a 2-mL tube using a wide-mouth P1000 pipette tip. 5. Gently Wash the aggregates 3 times with E6n-CLF. 6. Transfer the aggregates to a low-cell-binding 6-well plate using a wide P1000 pipette tip. 7. Transfer the aggregates with 100 μL of E6n-CLF to the central wells on the original 96-well plate. 8. Add 150 μL of E6n-CLF to each well. The final volume in each well is 250 μL. 9. Return the plates to the CO2 incubator. 3.2.5 Day 8: Medium Change

1. Make 25-mL of E6n-CFL as mentioned at Subheading 3.2.4. 2. Aspirate 200 μL of the medium from each well using P200 wide-mouth pipette tips and a P200 multichannel pipette. Make sure not to disturb the aggregates. 3. Add 110 μL of E6n-CLF twice to each well. The final volume in each well is 270 μL. 4. Return the plates to the CO2 incubator.

3.2.6 Day 11: Transfer to OMM Containing Matrigel and CHIR

1. Warm 6 mL of DMEM/F12 to RT. 2. Prepare 25 mL of OMM in a 50-mL tube and place on ice or in a refrigerator for at least 30 min. 3. Dissolve 250 μL of frozen Matrigel in the cold 25 mL OMM using a chilled P1000 pipette tip by pipetting up and down. Vortex the tube to dissolve Matrigel completely (see Note 10). 4. Add 7.5 μL of 10 mM CHIR to the 25 mL of OMM + Matrigel (OMM-MC). Mix well and warm to RT. Note that the OMM-MC contains 1% Matrigel +3 μM CHIR. 5. Using wide-mouth pipette tips and a multichannel P200 pipette, transfer up to 60 aggregates from the 96-well plates to each 100-mm dish. 6. Transfer aggregates into a 2-mL tube using a wide-mouth P1000 pipette tip. 7. Wash the aggregates with 1–2 mL of DMEM/F12 three times and then with 1 mL of OMM-MC twice.

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8. Transfer up to 60 aggregates to a new 100-mm dish using a wide-mouth P1000 pipette tip, and then add 10–20 mL of OMM-MC. 9. Gently shake the dish in all directions several times to disperse the aggregates. 10. Place the dishes in the CO2 incubator. 3.2.7 Day 13 and 15: Medium Change with OMM + CHIR

1. Prepare 25 mL of OMM and warm to RT. 2. Add 7.5 μL of 10 mM CHIR to the 25 mL of OMM and mix well. 3. Aspirate the spent medium and pour 10–20 mL into each dish. The d15–18 interval may require additional media compared to d13–15. 4. Return the dishes to the CO2 incubator.

3.2.8 Day 18 and Thereafter: Transition to the Long-Term Culture

1. Transfer the aggregates into new 100-mm dishes using a widemouth P1000 pipette tip. Carefully aspirate the excess medium on the new dish and add 10–20 mL of OMM (without Matrigel or CHIR). Place the dishes in the CO2 incubator. 2. Completely change the medium with OMM once or twice every week, or when the medium looks amber or yellow. Transfer aggregates to a new 100 mm dish when the dishsurface becomes confluent with cells migrating out from the aggregates. 3. PAX2+ PAX8+ ECAD+ otic vesicles can be observed at around d20. ATOH1+ MYO7A+ POU4F3+ PCP4+ hair cells appear from around d40, and the number of hair cells keeps increasing until around d70.

4

Notes 1. Medium should be stored as aliquots in conical tubes to prevent pH change during its storage. 2. Although OMM can be stored for up to 2 weeks, freshly made OMM is encouraged for use during early differentiation (e.g., d11–20). 3. Regular E8 medium (Gibco A1517001) + Normocin (E8n) can be used instead of E8fn for keeping hPSCs, for this substitution, the medium must be changed daily. Human inner ear organoid culture can also be started with E8n instead of E8fn on d-2. For this substitution, add 100 μL of E8n on d-1 instead of 4 h. 4. VTN-N coated plates can be stored for 1 week at 4  C in a sealed plastic bag.

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5. It is recommended to warm E8f at RT to prevent degradation of bFGF. However, warming at 37  C can be applied for immediate use; for example, E8fn + RvC can be placed in the CO2 incubator just before thawing or passaging cells. 6. The split ratio can be determined by the confluency and the growth rate of the hPSC line, as well as the duration until the next passaging or organoid culture. We regularly passage twice per week like as in the example described below: (a) On Mondays, the 60% confluent cells are passaged with the split ratio at 1:10–1:20 into two wells. (b) On Thursday, the 60% confluent cells are passaged with the split ratio at 1:10–1:25 into four wells. (c) The following Monday, one well of hPSC is used for another round of passaging. (d) On the next day (Tuesday), one of the three residual wells are used for starting an organoid culture. 7. Using low-cell-binding 96-well plates is necessary to form 3D aggregates. Either U-bottom or V-bottom 96-well plates can be used from d-2 to d0. 8. Most of the cells are dissociated into single cells after 5 min incubation with Accutase, although it is normal that some clumps remain after pipetting. To promote dissociation, 0.5 mM EDTA can be used for washing the cells before treating Accutase. A longer treatment of Accutase or vigorous pipetting is not recommended. 9. The optimization of the BMP-4 concentration may be necessary for different hPSC lines. In our experience, the WA25 hESC line and CRISPR-engineered cell lines based on WA25 do not require additional BMP-4 as the endogenous BMP-4 level from these cell lines at d0 is sufficient for nonneural ectoderm induction. 10. The Matrigel-containing medium should be used immediately after warming. Storing the medium in a refrigerator or room temperature for prolonged periods is not recommended. Never warm the medium at 37  C. An unheated water bath (i.e., at RT) can quicken the warming. 11. The freezing point of DMSO is 19  C, therefore adding small molecules dissolved in DMSO (e.g., Y, SB, LDN, CHIR) after warming at RT is recommended. 12. It is recommended to dissolve bFGF in the medium immediately prior to use to maximize its effect due to the low stability of this protein. On d0, bFGF can be added to the E6 + Matrigel + SB after transferring all aggregates from 96-well plates to the 100-mm dish.

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13. The aggregates on d0 are quite small and can be difficult to identify. Placing the 60-mm dish on a darker object (e.g., a dark-colored pipette tip box) can help increase visibility. 14. We only use the central 60 wells to culture aggregates and fill the peripheral wells with sterile H2O to reduce the evaporation from the central wells from d0 to d11. The peripheral wells are unsuitable for culture as they lose some amount of medium and therefore reagents in them become concentrated. 15. Use DMSO rather than water or DPBS to dilute small molecules with poor aqueous solubility; especially LDN, as a quite small volume is required. Otherwise, some precipitation appears, and the concentration becomes inaccurate.

Acknowledgments We would like to thank Jing Nie for the protocol optimization. This work was supported by a National Institutes of Health grant R01 DC015788 and a Department of Defense U.S. Army Medical Research and Materiel Command Congressionally Directed Medical Research Program grant W81XWH-18-1-0062 (to E.H.). References 1. Geleoc GS, Holt JR (2014) Sound strategies for hearing restoration. Science 344 (6184):1241062. https://doi.org/10.1126/ science.1241062 2. Muller U, Barr-Gillespie PG (2015) New treatment options for hearing loss. Nat Rev Drug Discov 14(5):346–365. https://doi.org/10. 1038/nrd4533 3. Barth KA, Kishimoto Y, Rohr KB, Seydler C, Schulte-Merker S, Wilson SW (1999) Bmp activity establishes a gradient of positional information throughout the entire neural plate. Development 126(22):4977–4987 4. Grocott T, Tambalo M, Streit A (2012) The peripheral sensory nervous system in the vertebrate head: a gene regulatory perspective. Dev Biol 370(1):3–23. https://doi.org/10.1016/ j.ydbio.2012.06.028 5. Harvey NT, Hughes JN, Lonic A, Yap C, Long C, Rathjen PD, Rathjen J (2010) Response to BMP4 signalling during ES cell differentiation defines intermediates of the ectoderm lineage. J Cell Sci 123 (Pt 10):1796–1804. https://doi.org/10. 1242/jcs.047530 6. Wilson PA, Hemmati-Brivanlou A (1995) Induction of epidermis and inhibition of neural

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e1007552. https://doi.org/10.1371/journal. pgen.1007552 18. DeJonge RE, Liu XP, Deig CR, Heller S, Koehler KR, Hashino E (2016) Modulation of Wnt signaling enhances inner ear organoid development in 3D culture. PLoS One 11(9): e0162508. https://doi.org/10.1371/journal. pone.0162508 19. Koehler KR, Hashino E (2014) 3D mouse embryonic stem cell culture for generating inner ear organoids. Nat Protoc 9 (6):1229–1244. https://doi.org/10.1038/ nprot.2014.100 20. Longworth-Mills E, Koehler KR, Hashino E (2016) Generating inner ear organoids from mouse embryonic stem cells. Methods Mol Biol 1341:391–406. https://doi.org/10. 1007/7651_2015_215 21. Nie J, Koehler KR, Hashino E (2017) Directed differentiation of mouse embryonic stem cells into inner ear sensory epithelia in 3D culture. Methods Mol Biol 1597:67–83. https://doi. org/10.1007/978-1-4939-6949-4_6 22. Koehler KR, Nie J, Longworth-Mills E, Liu XP, Lee J, Holt JR, Hashino E (2017) Generation of inner ear organoids containing functional hair cells from human pluripotent stem cells. Nat Biotechnol 35(6):583–589. https:// doi.org/10.1038/nbt.3840 23. Liu XP, Koehler KR, Mikosz AM, Hashino E, Holt JR (2016) Functional development of mechanosensitive hair cells in stem cell-derived organoids parallels native vestibular hair cells. Nat Commun 7:11508. https://doi.org/10. 1038/ncomms11508

Methods in Molecular Biology (2022) 2520: 151–159 DOI 10.1007/7651_2021_449 © Springer Science+Business Media, LLC 2021 Published online: 02 November 2021

Glycolytic Profiling of Mouse Embryonic Stem Cells (mESCs) Bibiana Correia, Maria Ineˆs Sousa, and Joa˜o Ramalho-Santos Abstract Mouse embryonic stem cells (mESCs) can be captured in vitro in different pluripotency states through media modulation, mimicking their natural environment during early embryo development. As highly proliferative cells, mESCs prefer to use glycolysis to support the energetic and biosynthetic demands, even in the presence of oxygen. Indeed, glycolysis can not only supply ATP at a much faster rate, when compared to other catabolic pathways, but also provides biosynthetic substrates to meet anabolic requirements. Considering that ESCs cultured in different media conditions display distinct metabolic requirements, it is of utmost importance to have a robust metabolic characterization methodology to understand how subtle metabolic variations may be coupled to ESC identity. Here we describe how to profile the glycolytic activity of naive mouse ESC, using the established Seahorse XFe24 Live-cell Metabolic Assay. This may be a useful protocol for understanding how the glycolytic function of mESCs changes in certain circumstances and how is it coupled to diverse pluripotency/differentiation phenotypes. Key words Glycolysis, mESCs, Metabolism, Pluripotency, Seahorse

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Introduction Pluripotency refers to the potential that some cells have to differentiate and functionally contribute to all somatic lineages and the germline of an organism. Although these cells arise in the blastocyst in vivo, there are different states of pluripotency stablished in vitro, which are sustained with different media formulations [1–3]. Cells in each pluripotency state share similarities to their in vivo equivalents in the pre-/peri- and early implantation embryo, named “naive” ESCs and “primed” ESCs or Epiblast Stem Cells (EpiSCs), respectively [1, 4–7]. Among all the characteristics that confer ESCs their unique potential, the metabolic profile presented by the cells in different pluripotency states (or even in the same state cultured in different media) is deeply related to the function and identity of these cells [7–11]. For most cell types glucose is the preferred carbon source used for the production of ATP. The catabolism of a single glucose molecule into to two pyruvate molecules is coupled with the

Bibiana Correia and Maria Ineˆs Sousa contributed equally and should be considered co-first authors.

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production of two NADH molecules and a net gain of two ATP molecules, in the glycolytic pathway [12, 13]. Despite the low ATP net gain of glycolysis, in excess of glucose and low oxygen conditions, glycolysis can supply ATP at a much faster rate compared to other catabolic pathways. Therefore, highly proliferative cells, as ESCs, prefer to use glycolysis to support the energetic and biosynthetic demands, even in the presence of oxygen [9, 14]. Indeed, highly proliferative cells require not only reducing cofactors (NADPH) and energy (ATP), but also carbon, nitrogen, and hydrogen to support macromolecule biosynthesis. Thus, a partial breakdown of glucose through glycolysis and management of intermediate metabolites have to be tightly coordinated to obtain the right balance between the generation of ATP and reducing cofactors and the production of biosynthetic substrates to meet anabolic requirements [8, 15]. In addition, the use of glycolytic metabolism over mitochondrial oxidation prevents the production of reactive oxygen species that might damage the cellular building blocks needed for cell division [16]. Core pluripotency factors were already demonstrated to directly regulate glycolysis in ESCs. For instance, OCT4 was shown to regulate the transcription of genes encoding the glycolytic enzymes HXK2 and PKM2, boosting glycolytic flux [7, 15, 17, 18]. Additionally, it has recently been shown that glucose transporter 1 (GLUT1) enhancer presents binding sites for OCT4, SOX2 and NANOG, that drive its expression and consequently lead to an increased glycolytic flux [19]. Moreover, metabolism can in turn influence pluripotency status. For example, the forkhead box protein O1 (FOXO1), which mediates cell response to insulin or insulin growth factors, regulates the expression of OCT4 and SOX2 [20, 21]. Additionally, inhibition of mitochondrial ETC complex III, by Antimycin A, not only enhanced hESC pluripotency [22] but was also shown to suppress differentiation of mESCs into neuronal dopaminergic cell lineages, maintaining mESC pluripotency-associated gene expression during the differentiation process [23]. Furthermore, inhibition of glycolysis by either dichloroacetate or 3-bromopyruvate induced loss of pluripotency in mESCs [10, 11]. Considering that, not only different states of pluripotency have distinct metabolic requirements, but even different media conditions that sustain the same naı¨ve ESCs identity, can affect the metabolic profile of these cells, a robust metabolic characterization methodology is needed, in order to understand how subtle metabolic variations are coupled to ESC identity. Here, we describe how to profile the glycolytic activity of naive mouse ESC, using the established Seahorse XFe24 Live-cell Metabolic Assay, which monitors media acidification related to lactic acid production, the final step used to regenerate NAD+ in mammalian glycolysis. The protocol can be applied to assess functionally

ECAR (pMoles/min/103 cells)

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Fig. 1 Example of a real-time extracellular acidification rate (ECAR) measurement assay comparing more quiescent mESCs cultured in 2i media with mESCs cultured in FBS-based media. Both are naive mESCs but show clear metabolic differences

different aspects of glycolytic activity, taking approximately 2 days to perform. This protocol is divided into three steps: (1) Assay preparation, (2) Seahorse assay protocol, (3) Normalization of the Seahorse assay by cell count. Cell culture should be performed as usual before the assay preparation, when cells are plated into the XFe24 Seahorse Microplate. The first measurements allow us to determine the basal acidification rates of cells cultured in basal media in glucose starvation. Subsequently, the first injected compound is glucose, which enables the prediction of the normal capacity that cells hold to convert glucose into pyruvate (followed by conversion to lactate). Then, in the second injection, the addiction of Oligomycin, which inhibits mitochondrial ATP synthesis, forces cells to use glycolysis at its maximal potential to support the ATP levels necessary to fulfil their energetic demands for ATP production, through acceleration of glycolysis rate. Finally, the third and final injection is the addiction of 2-Deoxy-D-glucose (2DG), which by inhibiting Hexokinase compromises the whole pathway, since this enzyme is involved in the first glycolysis reaction, catalyzing glucose phosphorylation and irreversibly trapping it inside the cell. Therefore, the 2-DG injection ensures that the previous measurement signals result from glycolytic function (Fig. 1).

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Materials

2.1 Mouse Embryonic Stem Cell (mESC) Culture and Seahorse XFe24 LiveCell Metabolic Assay Normalization by Cell Count

1. mESC line of interest. 2. 2i mESC medium: 1:1 solution of DMEM/F12 and Neurobasal medium, supplemented with 100 U/mL penicillin–streptomycin, 2 mM L-glutamine, 0.1 mM 2-mercaptoethanol, 1:200 N2 and 1:100 B27, GSK3 inhibitor—3 μM CHIR99021, MEK inhibitor—1 μM PD184352 and 106 U/ L Leukemia inhibitory factor (LIF). 3. Serum (FBS)-based medium: DMEM, supplemented with 15% embryonic stem-cell qualified FBS, 100 U/mL penicillin/streptomycin, 0.1 mM 2-mercaptoethanol, 1% nonessential amino acids, 2 mM sodium pyruvate, and 106U/L LIF. 4. Phosphate-buffered saline (PBS): 137 mM NaCl, 2.67 mM KCl, 8.06 mM Na2HPO4, 1.47 mM KH2PO4, pH 7.4. 5. Gelatin: 0.1% gelatin in PBS. 6. StemPro™ Accutase™ Cell Dissociation Reagent: 1 Accutase enzymes in PBS. 7. Trypan Blue stain solution. 8. Hemocytometer.

2.2 Seahorse XFe24 Live-Cell Metabolic Assay

1. Seahorse XFe24 Analyzer (Agilent Technologies). 2. Seahorse XFe24 FluxPak containing XFe24 sensor cartridges, XF24 cell culture microplates and Seahorse XF Calibrant Solution (Agilent Technologies). 3. Seahorse XF Glycolysis Stress Test medium: DMEM medium, 2 mM L-Glutamine, pH 7.4. 4. Glycolytic modulator stock solutions: (a) Glucose—100 mM Glucose, DMEM medium, 2 mM LGlutamine, pH 7.4. (b) 2-Deoxy-D-glucose (2-DG)—1 M medium, 2 mM L-Glutamine, pH 7.4

2-DG,

DMEM

(c) Oligomycin—10 μM Oligomycin, DMEM medium, 25 mM Glucose, 2 mM L-Glutamine, 1 mM Sodium Pyruvate, pH 7.4.

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3.1 Seahorse XF Glycolysis Stress Test 3.1.1 Assay Preparation (Performed the Day Before) (See Notes 1 and 2)

1. Hydrate the XFp sensor cartridge in XF Calibrant Solution. The sensor cartridge has a Utility Plate associated. Carefully lift the Sensor lid and fill each well of the Utility plate with 1 mL Seahorse XF Calibrant Solution. Lower the sensor lid into the Utility Plate, submerging the sensors in the XF Calibrant. Place the Sensor Cartridge in a non-CO2 incubator at 37  C, overnight. 2. In a cell culture room, prepare the materials and reagents required for plating mESCs into the XF24 cell culture microplate. 3. Coat the XF24 cell culture microplate with a 0.1% gelatin solution, to allow for mESC adherence. 4. Wash the cultured mESCs with PBS. 5. Incubate the cells with StemPro™ Accutase™ solution for 5 min, at 37  C. 6. Inactivate the enzyme by dilution with mESC medium, and collect the harvested cells in a centrifuge tube. 7. Centrifuge the cell suspension at 300  g, for 5 min. 8. Resuspend the obtained cell pellet in mESC medium. 9. Take a small aliquot of cell suspension and add Trypan Blue stain solution at 1:1 ratio. Count the cells in a hemocytometer. 10. Perform the calculations required in order to plate 100,000 cells/100 μL in each well of the XF24 cell culture microplate. 11. Aspirate off the gelatin in each well of the XF24 cell culture microplate, before seeding. 12. Plate the cell suspension in the designated wells of the XF24 cell culture microplate (see Note 3). 13. Place the seeded XF24 cell culture microplate in a standard cell culture incubator, at 37  C and 5% CO2 conditions and allow cells to deposit at the bottom the well (1–2 h). 14. After cells deposition, carefully add an extra 200 μL of mESC medium, without disturbing the cells, to allow cell growth and colony assembly (see Note 4).

3.1.2 Seahorse Assay Protocol

1. Switch on the Seahorse XFe24 Analyzer equipment and software, preferentially 30 min–1 h before use to allow the temperature (37  C) to stabilize. 2. Warm the pre-made Seahorse XF Glycolysis Stress Test medium at 37  C. 3. Take the XF24 cell culture microplate from the CO2 incubator and look at the plated cells under the microscope, to confirm cell adhesion and colony morphology (see Note 5).

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4. Carefully wash the plated cells with the assay medium. 5. Add 450 μL of Seahorse XF Glycolysis Stress Test medium to each well and place the microplate in an incubator at 37  C without CO2, 1 h prior to the assay (see Note 6). 6. Prepare the 10 stocks of glycolytic modulators that will be used in the assay. Final concentrations after injections will be: 10 mM Glucose, 100 mM 2-DG and 1 μM Oligomycin. 7. Without removing the Utility Plate, load the mitochondrial modulators into the drug delivery injection ports that are on top of the sensor cartridge as following: (Port A) 50 μL of Glucose; (Port B) 55 μL of Oligomycin; (Port C) 60 μL of 2-DG (see Note 7). 8. Leave the sensor cartridge with the Utility plate in a non-CO2 incubator at 37  C, while designing the assay in the XFe Wave software. 9. In the XFe Wave software, set up the desired Seahorse assay’s program. 10. After designing the Seahorse assay, click Start Assay. 11. The Seahorse XFe24 Analyzer will then ask to place the utility plate with the loaded assay cartridge on the instrument tray. 12. The standard Seahorse run comprises the following steps: (a) Calibration. (b) Replacement of the Utility plate for XF24 cell culture microplate. (c) Equilibration of the Base line readings: (Mix—3 min, Wait—2 min, Measure—3 min) x3; (d) Port A injection: (Mix—3 min, Wait—2 min, Measure— 3 min) x3; (e) Port B injection: (Mix—3 min, Wait—2 min, Measure— 3 min) x3; (f) Port C injection: (Mix—3 min, Wait—2 min, Measure— 3 min) x3; (g) End program. 3.1.3 Normalization of the Seahorse Assay by Cell Count (see Note 8)

1. Right after the Seahorse assay run ends, gently aspirate off the Seahorse assay medium and wash the wells once with PBS (see Note 9). 2. Using the dissociation reagent StemPro™ Accutase™, lift the cells from the base of the well. 3. Stop the enzymatic reaction by diluting the enzyme. 4. Thoroughly resuspend the cell suspension in order to break the mESC colonies into a single-cell suspension.

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5. Take a small aliquot of cell suspension and add Trypan Blue stain solution at 1:1 ratio. 6. Count the cells in a hemocytometer. 7. Calculate the total number of cells in each well. 8. Normalize the Seahorse assay data results by the total number of cells in each well, using the XFe Wave software.

4

Notes 1. Prior to the assay, plan/design the XF24 cell culture microplate in order to leave four wells without plated cells for background correction (with only assay medium). Likewise, it is recommended to use 3 replicate wells per condition to increase measurement accuracy. 2. To allow for mESCs to adhere and acquire the proper phenotypical colony morphology in the microplate (6–8 h), it is preferable to seed cells, in mESC culture medium, the day before the assay. 3. When seeding the mESCs, thoroughly homogenate the cell suspension, since mESCs that do not adhere in a single-cell fashion are prone to differentiate, influencing the outcome of the assay. 4. When gently adding the extra 200 μL of mESC medium, place the pipette tip against the side of the well, at the top of the well, to decrease the medium’s flow impact on the adhering cells. 5. Due to the high cell density required for this assay, it is to be expected that the mESCs cultured in the FBS-medium; which is more prone for differentiation, to present a slightly higher differentiation percentage after overnight adhesion. 6. It is necessary for plated cells to be for 1 h in an incubator without CO2 to allow the de-gassing the plate, permitting CO2 diffusion from the cells, medium, and plate. 7. Inspect the drug delivery injection ports to verify if there are no air bubbles and if the liquid is all the way down at the bottom of the port. 8. Multiple/other normalization methodologies (such as protein concentration) can be performed, depending on the specificities of each experimental design. 9. After the Seahorse assay, be extremely gentle when aspirating off the Seahorse assay medium, in order to avoid possible colony detachment and their aspiration during the process.

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Acknowledgments The authors would like to acknowledge the members of the Biology of Reproduction and Stem Cells research group, at the Center from Neuroscience and Cell Biology, for the discussion and constructive feedback related to this work.

Funding This work was funded by Fundac¸˜ao para a Cieˆncia e Tecnologia (FCT) Portugal: PhD scholarship attributed to B.C. (SFRH/BD/ 144150/2019), the STEM@REST Project (CENTRO-01-0145FEDER-028871) and PAC CANCEL_STEM (POCI-01-0145FEDER-016390. M.I.S. was hired through the STEM@REST Project (CENTRO-01-0145-FEDER-028871). Additional funding was provided by the European Regional Development Fund (ERDF), through the Centro 2020 Regional Operational Programme: project CENTRO-01-0145-FEDER-000012HealthyAging2020, the COMPETE 2020—Operational Programme for Competitiveness and Internationalisation, and the Portuguese national funds via FCT—Fundac¸˜ao para a Cieˆncia e a Tecnologia, I.P.: project POCI-01-0145-FEDER-007440, that attributed a fellowship to B. C. (BIM—IN0828). References 1. Nichols J, Smith A (2012) Pluripotency in the embryo and in culture. Cold Spring Harbor Perspect Biol 4(8):1–154. https://doi.org/ 10.1101/cshperspect.a008128 2. De Los Angeles A, Ferrari F, Xi R et al (2015) Hallmarks of pluripotency. Nature 525:469–478. https://doi.org/10.1038/ nature15515 3. Weinberger L, Ayyash M, Novershtern N et al (2016) Dynamic stem cell states: naive to primed pluripotency in rodents and humans. Nat Rev Mol Cell Biol 17:155–169. https:// doi.org/10.1038/nrm.2015.28 4. Brons IGM, Smithers LE, Trotter MWB et al (2007) Derivation of pluripotent epiblast stem cells from mammalian embryos. Nature 448:191–195. https://doi.org/10.1038/ nature05950 5. Tesar PJ, Chenoweth JG, Brook FA et al (2007) New cell lines from mouse epiblast share defining features with human embryonic stem cells. Nature 448:196–199. https://doi. org/10.1038/nature05972

6. Mathieu J, Ruohola-Baker H (2017) Metabolic remodeling during the loss and acquisition of pluripotency. Development 144:541–551. https://doi.org/10.1242/dev. 128389 7. Tsogtbaatar E, Landin C, Minter-Dykhouse K et al (2020) Energy metabolism regulates stem cell pluripotency. Front Cell Dev Biol 8:1–16. https://doi.org/10.3389/fcell.2020.00087 8. Varum S, Rodrigues AS, Moura MB et al (2011) Energy metabolism in human pluripotent stem cells and their differentiated counterparts. PLoS One 6(6):e20914. https://doi. org/10.1371/journal.pone.0020914 9. Pereira SL, Rodrigues AS, Sousa MI et al (2014) From gametogenesis and stem cells to cancer: common metabolic themes. Hum Reprod Update 20:924–943. https://doi. org/10.1093/humupd/dmu034 10. Rodrigues AS, Correia M, Gomes A et al (2015) Dichloroacetate, the pyruvate dehydrogenase complex and the modulation of mESC pluripotency. PLoS One 10:e0131663. https://doi.org/10.1371/journal.pone. 0131663

Glycolytic Profiling of Mouse Embryonic Stem Cells (mESCs) 11. Rodrigues AS, Pereira SL, Correia M et al (2015) Differentiate or die: 3-Bromopyruvate and pluripotency in mouse embryonic stem cells. PLoS One 10:e0135617. https://doi. org/10.1371/journal.pone.0135617 12. Nelson DL, Cox MM (2017) Lehninger principles of biochemistry, 7th edn. W. H. Freeman and Company All, New York 13. Roosterman D, Meyerhof W, Cottrell GS (2018) Proton transport chains in glucose metabolism: mind the proton. Front Neurosci 12:1–15. https://doi.org/10.3389/fnins. 2018.00404 14. Sousa MI, Rodrigues AS, Pereira S et al (2015) Mitochondrial mechanisms of metabolic reprogramming in proliferating cells. Curr Med Chem 22:2493–2504. https://doi.org/10. 2174/0929867322666150514095718 15. Folmes CDL, Nelson TJ, Dzeja PP et al (2012) Energy metabolism plasticity enables stemness programs. Ann N Y Acad Sci 1254:82–89. https://doi.org/10.1111/j.1749-6632.2012. 06487.x 16. Zhu S, Li W, Zhou H et al (2010) Reprogramming of human primary somatic cells by OCT4 and chemical compounds. Cell Stem Cell 7:651–655. https://doi.org/10.1016/j.stem. 2010.11.015 17. Folmes CDL, Nelson TJ, Martinez-Fernandez A et al (2011) Somatic oxidative bioenergetics transitions into pluripotency-dependent glycolysis to facilitate nuclear reprogramming.

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Cell Metab 14:264–271. https://doi.org/10. 1016/j.cmet.2011.06.011 18. Kim H, Jang H, Kim TW et al (2015) Core pluripotency factors directly regulate metabolism in embryonic stem cell to maintain pluripotency. Stem Cells 33:2699–2711. https:// doi.org/10.1002/stem.2073 19. Yu L, Ji K, Zhang J et al (2019) Core pluripotency factors promote glycolysis of human embryonic stem cells by activating GLUT1 enhancer. Protein Cell 10:668–680. https:// doi.org/10.1007/s13238-019-0637-9 20. Zhang X, Yalcin S, Lee D-F et al (2011) FOXO1 is an essential regulator of pluripotency in human embryonic stem cells. Nat Cell Biol 13:1092–1099. https://doi.org/10. 1038/ncb2293 21. Lee S, Dong HH (2017) FoxO integration of insulin signaling with glucose and lipid metabolism. J Endocrinol 233:R67–R79. https:// doi.org/10.1530/JOE-17-0002 22. Varum S, Momcˇilovic´ O, Castro C et al (2009) Enhancement of human embryonic stem cell pluripotency through inhibition of the mitochondrial respiratory chain. Stem Cell Res 3:142–156. https://doi.org/10.1016/j.scr. 2009.07.002 23. Pereira SL, Gra˜os M, Rodrigues AS et al (2013) Inhibition of mitochondrial complex III blocks neuronal differentiation and maintains embryonic stem cell pluripotency. PLoS One 8: e82095. https://doi.org/10.1371/journal. pone.0082095

Methods in Molecular Biology (2022) 2520: 161–170 DOI 10.1007/7651_2021_450 © Springer Science+Business Media, LLC 2021 Published online: 30 November 2021

Replating Protocol for Human Induced Pluripotent Stem Cell–Derived Cardiomyocytes Arzuhan Koc and Esra Cagavi Abstract Human induced pluripotent stem cell–derived cardiomyocytes (hiPSC-CM) create an unlimited cell source for basic and translational cardiac research. Obtaining hiPSC-CM culture as a single-cell, monolayer or three-dimensional clusters for downstream applications can be challenging. Thus, it is critical to develop replating strategies for hiPSC-CMs by evaluating different enzymatic or nonenzymatic reagents for dissociation and seeding on different coating materials. To reseed hiPSC-CMs with high viability and at structures desirable for the downstream applications, here we defined optimized protocols to dissociate hiPSC-CMs by using collagenase A&B, Collagenase II, TrypLE, and EDTA and reseeding on various matrix materials including fibronectin, laminin, imatrix, Matrigel, and Geltrex. By the replating methods described here, a single cell or cluster-containing hiPSC-CM cultures can be generated effectively. Key words Human induced pluripotent stem cells, Cardiomyocytes, Cardiac differentiation, Replating cardiomyocytes, Cell dissociation

Abbreviations ECM EDTA hiPSC-CM

1

Extracellular matrix Ethylenediaminetetraacetic acid Human induced pluripotent stem cell-derived cardiomyocytes

Introduction The human induced pluripotent stem cell (hiPSC) technology has provided an invaluable source for in vitro modeling of cardiac disease mechanisms, developing translational and personalized therapies [1–3]. As it is challenging to reach the patient-sourced cardiomyocytes from cardiac biopsies, and also the count and types of the cardiac cells obtained can be limited, hiPSC-derived cardiomyocytes (hiPSC-CM) are highly preferred in these studies [2–8]. Fully differentiated hiPSC-CMs generally exhibit tissue clusters at early days of differentiation, and the clusters become more compact as the cultures mature [8]. Within this maturation period,

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the cell–cell and cell–extracellular matrix (ECM) interactions of the cardiac cultures strengthen. Specific integrin subtypes are known to bind selectively and tightly to fibronectins and laminins in cardiac microenvironment [6, 9–12]. Therefore, ECM selection becomes an important issue for cardiac cultures at different levels of maturity. Strategies for reseeding of iPSC-CMs for the downstream analysis have been developed through using different enzymes. From these enzymes, collagenases and trypsin derivatives are commonly preferred, and seeding of cardiac cells on different ECMs has been evaluated to obtain effective attachment, high viability, and functional recovery [2, 6–11]. Different ECM components are used for culturing stem cells, terminally differentiated cells, or primary cell cultures, some of which are commercially available, including fibronectin, Matrigel, Geltrex, laminin derivatives, and different collagen subtypes [8, 11, 13, 14]. In this context, choosing a suitable dissociation enzyme and coating material becomes an essential part of cardiac cultures, as ECM elements provide cell attachment, survival, and differentiation cues by modulating signaling pathways [12–15]. Therefore, in this protocol, we defined optimal strategies to dissociate hiPSC-CMs by using the Collagenase A-B, Collagenase type II, TrypLE enzymes or EDTA as a nonenzymatic methodology. The protocol was expanded to provide insights into replating iPSC-CMs onto different matrices, such as Matrigel, Geltrex, laminin, fibronectin, and iMatrix-511 for various applications [8]. Furthermore, we have highlighted the important points to be followed with common troubleshooting strategies. Prolonged enzymatic digestions can be preferred in studies that require single cells such as, microfluidic studies or single-cell RNA sequencing, by compromising on cell viability. The use of EDTA and collagenases results in mild dissociation in less time with high viability that would be preferable for obtaining cardiac clusters for tissue engineering applications or organoid research.

2

Materials

2.1 Differentiation of hiPSC-CMs

1. CHIR99021 (10 mM) (Sigma-Aldrich, SML1046): Add 1.074 mL DMSO to 5 mg CHIR99021. Aliquot samples and store at 80  C for up to 1 year. 2. IWP-4 (10 mM) (Tocris, 5214): Add 2 mL DMSO to 10 mg IWP-4. Sonicate the mixture for 30 min to dissolve the IWP-4. Aliquot samples and store at 80  C for up to 1 year. 3. IWR-1 (10 mM) (Tocris, 3532): Add 2.4 mL DMSO to 10 mg IWR-1. Shake gently to dissolve the IWR-1. Aliquot samples and store at 80  C for up to 1 year. 4. RPMI–B27 (differentiation medium with insulin) (50 mL): Mix 48 mL of RPMI1640 (Multicell, 350-000-

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EL), 1 mL of B27 supplement (Gibco, 17504-044), 2 mM GlutaMAX (Gibco, 35050-061), and 100 penicillin–streptomycin (Gibco, 15140-122). The medium can be stored at 4  C for up to 2 weeks. 5. RPMI–B27(without insulin) (50 mL): Mix 48 mL of RPMI1640 (Multicell, 350-000-EL), 1 mL B27 supplement lacking insulin (Gibco, A1895601), 2 mM GlutaMAX (Gibco, 35050-061), and 100 penicillin–streptomycin (Gibco, 15140-122). The medium can be stored at 4  C for up to 2 weeks. 6. Cardiomyocyte enrichment medium with 10 mM L-lactate (50 mL): Mix 49 mL of RPMI1640 (Multicell, 350-060-CL), 56 mg sodium L-lactate (Sigma-Aldrich, 71718), 2 mM GlutaMAX (Gibco, 35050-061), and 100 penicillin–streptomycin (Gibco, 15140-122). Filter the medium with a sterile 22 μm membrane and store at 4  C for up to 2 weeks. 7. CRITICAL POINT: All medium and solutions should be prepared at aseptic conditions under a class-II biological safety cabinet. Filtration through a low protein binding and sterile 22 μm filter after preparation is highly recommended. 2.2 Coating Tissue Culture Plates with ECM Proteins

1. Fibronectin-coated plates: Fibronectin (Sigma, F1141) was prepared at a concentration of 10 μg/cm2 in DPBS (without Ca2+/Mg2+, Gibco, 14190-094) and incubated for 4 h at 37  C for homogeneous coating of tissue culture plates. 2. Laminin-coated plates: Poly-L-lysine (Sigma-Aldrich, P6282) was prepared at 1:10 dilution in DPBS and incubated at 37  C for 2 h for coating culture plates. After washing the wells three times with dH2O, laminin, composed of A, B1 and B2 chains (Sigma-Aldrich, L2020), was added on poly-L-lysine–coated wells at the concentration of 10 μg/cm2 in dH2O and incubated for 24 h at 37  C. 3. Geltrex-coated plates: Add 12 mL of cold (4  C) DMEM– F12 to a 15 mL conical tube mix 120 μL Geltrex (hESCqualified, Gibco, A1413302). Immediately add 1 mL/well Geltrex in DMEM–F12 (Gibco, 11330-032) for tissue-treated 6-well plates. Allow the Geltrex to set in the wells for 30 min at RT before use. The Geltrex-coated plates can be stored at 4  C for up to 2 weeks. 4. iMatrix-511-coated plates: For iMatrix-511 (Takara, T304) coating, the solution was prepared at the concentration of 10 μg/mL in DPBS (without Ca2+/Mg2+, Gibco, 14190094), and incubated for 1 h at 37  C before plating cells. 5. Matrigel-coated plates: Add 12 mL of cold (+4  C) DMEM– F12 (Gibco, 11330-032) to a 15 mL conical tube mix 120 μL Matrigel (hESC-qualified, Corning 354277). Immediately add

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1 mL/well Matrigel in DMEM–F12 for 6-well plates. Allow the Matrigel to set for 1 h at RT before use. The Matrigelcoated plates can be stored at 4  C for up to 2 weeks. 6. CRITICAL POINT: Matrigel and Geltrex aliquots should be prepared by using prechilled tips and tubes. The solutions should be kept on ice at all times during aliquoting and preferably use cold blocks. 2.3 Preparation of Dissociation Reagents

1. Collagenase A and B enzyme solution: Add 7% (w/v) Collagenase A (Roche, 10103586001) and Collagenase B (Roche, 11088815001) to 5 mL RPMI/B27. Mix well till the solution becomes homogenous and sterilize by passing through a 22 μm filter. It is recommended to prepare and use fresh. Alternatively, one-use only aliquots can be prepared and stored at 80  C for up to 1 year. 2. Collagenase type II solution (70% w/v) (Gibco, 17101015): Add 3.5 mg Collagenase type II to 5 mL HBSS. Mix and sterilize collagenase solution by passing through a 22 μm filter. It is recommended to prepare and use fresh. 3. DNaseI (10 mg/mL) (Sigma, D4513): The working concentration is 10 μg/mL. Prepare 11 mg/1100 mL NaCl (0.15 M) with sterile dH2O and filter the NaCl solution. Add NaCl solution into the DNase vial with a syringe. Mix gently without shaking to dissolve lyophilized DNase pellet. Aliquot samples and store at 20  C for up to 1 year. 4. CRITICAL POINT: DNase I was added to the enzymatic dissociation mixtures except EDTA treatment, to clear the genomic DNA exposed from dead cells. 5. 0.5 mM EDTA (Invitrogen, 15575-038): Add 50 μL 0.5 M EDTA in 50 mL sterile DPBS (without Ca2+/Mg2+) in aseptic conditions and mix well. The solution can be stored at 4  C.

2.4 Preparation of Reseeding Reagents

1. Enzyme inactivation medium: Mix 5 mL FBS (10%) and 45 mL DMEM or RPMI1640 under the biosafety cabin. Filter the medium with a 0.22 membrane filter. The medium can be stored at 4  C for up to 2 weeks. 2. Y27632 (5 mM) (Millipore, SCM075): Add 2.956 mL sterile dH2O to 5 mg Y27632. Shake gently. Aliquot samples and store at 20  C for up to 1 year. 3. Seeding medium: Add 5 μM Y27632 to the RPMI/B27 (Differentiation medium with insulin).

Replating Protocol for Human Induced Pluripotent Stem Cell–-erived. . .

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Methods 1. Grow feeder-free iPSCs in any iPSC medium of choice in a 37  C, 5% CO2 incubator till they get 70–90% confluency. We culture the hiPSCs on Matrigel-coated 6-well plates in E8 medium to 85% confluence (see Note 1).

3.1 Differentiation of hiPSC-CMs

2. Day 0: Change the hiPSCs culture medium with 6 μM CHIR99021-containing prewarmed RPMI/B27(-insulin) medium (see Note 2). 3. Day 2: Change the medium with prewarmed RPMI/B27 (-insulin) medium. 4. Day 3: Change the medium supplemented with 10 μM IWP-4 or IWR-1 containing prewarmed RPMI/B27(-insulin) medium. 5. CRITICAL POINT: The concentration of small molecules (CHIR99021, IWR-1, IWP-4) should be optimized as the different iPSC lines can exhibit batch-to-batch variation in differentiation capacity (see Note 2). 6. Day 5: Change the medium with prewarmed RPMI/B27 (-insulin) medium. 7. Day 7: Change the medium with prewarmed RPMI/B27 differentiation medium containing insulin. After this day, change the hiPSC-CM culture medium every 2–3 days and closely examine the cultures with light microscope to monitor spontaneous beating clusters (see Notes 3 and 4). 3.2 Replating of hiPSC-CMs by Dissociation with Collagenase A + B

l

CRITICAL POINT: The replating procedures discussed here describe the experimental steps for cardiac cultures at differentiation days between 15 and 30. If the iPSC-CM cultures are more mature and at differentiation days 60 or above, the cell clumps can be dissociated by increasing the enzyme treatment duration and/or the trituration. 1. Aspirate the hiPSC-CM culture medium usually grown on 6-well plates and wash the wells with 2 mL DPBS (without Ca2+/Mg2+) at RT. Add 2 mL DPBS (without Ca2+/Mg2+) to the wells and wait for 5 min at RT (see Note 5). 2. Add 1–1.5 mL/1 of 6 wells with prewarmed Collagenase A + B mixture, place the plate in a 37  C incubator and wait for 15–20 min. For mature cardiac differentiation cultures, the incubation time may be increased up to 30 min. 3. Every 5 min, check the size of cell clumps by light microscopy, pipette the cell clumps to dissolve until the clusters are not observed (see Note 6).

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4. Add 2 mL enzyme inactivation medium to inhibit the enzymatic activity to the wells, then pipetting can be reperformed while the cells are still in the well or in the falcon (see Note 7). 5. Collect the cells in 15 mL falcon tube, and wash the wells with an extra 3–4 mL of enzyme inactivation medium to collect the remaining cells (see Notes 7 and 8). 6. Spin the cells at 300  g for 3 min at 4  C. If the cell pellet is not observed clearly, DNaseI can be added to digest the genomic DNA exposed from dead cells (see Note 8). 7. Aspirate the supernatant and resuspend the cellular pellet with the seeding medium that is half of culture volume. 8. Aspirate the coating material, and seed the cells drop by drop on wells. Put the plate in a 37  C incubator and wait for 30 min. 9. After confirming settling of the cells by observing under light microscopy, gently add the other half of the seeding medium through the walls of the plate (see Notes 9 and 10). 3.3 Replating of hiPSC-CMs by Dissociation with Collagenase II

1. After washing the cells with DPBS (without Ca2+/Mg2+) (see Note 5), add 1–1.5 mL prewarmed Collagenase II mixture, put the plate in a 37  C incubator and wait for 15–20 min. If the cardiac clusters are not dissociated, the incubation can be extended another 10–15 min. 2. After checking the cell clumps every 5 min by light microscopy, triturate the sample to further dissolve cell clumps. Add 2 mL enzyme inactivation medium to the wells, then pipetting can be reperformed in the well or in the falcon (see Note 6). 3. Collect the cells in a 15 mL falcon tube, and wash the wells with extra 3–4 mL enzyme inactivation medium to collect the remaining cells. After this step, follow the protocol described in Subheading 3.2 at step 6 through 9 (see Notes 7–10).

3.4 Replating of hiPSC-CMs by Dissociation with TrypLE Express

1. After washing the cells with DPBS (without Ca2+/Mg2+) (see Note 5), add 1–1.5 mL prewarmed TrypLE Express enzyme (Gibco, 12604-013), put the plate in a 37  C incubator and wait for 3–5 min for early day hiPSC-CMs. When late day hiPSC-CMs were replated, incubation time can be increased up to 7–10 min. 2. After confirming dissociation of cardiac clusters by light microscopy, triturate further the sample to dissolve clusters. Add 2 mL enzyme inactivation medium (see Note 6). 3. Collect the cells in a 15 mL falcon tube, and wash the wells with extra 3–4 mL enzyme inactivation medium to collect the remaining cells. After this step, follow the protocol described in Subheading 3.2 at step 6 through 9 (see Notes 7–10).

Replating Protocol for Human Induced Pluripotent Stem Cell–-erived. . .

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1. After washing the cells with DPBS (without Ca2+/Mg2+) (see Note 5), wash the cells quickly with 1 mL prewarmed 0.5 mM EDTA solution. Aspirate EDTA, add 1.5 mL prewarmed 0.5 mM EDTA solution, put the plate in a 37  C incubator, and wait for 5 min. 2. For nonenzymatic dissociation, hiPSC-CMs were incubated with 10 mM EDTA at 37  C for 3–5 min for cardiac cultures at differentiation before day 30. For iPSC-CMs in differentiation cultures above 30 days, the incubation period can be extended to 7–10 min by carefully examining cultures microscopically ever 1–2 min during the treatment (see Note 6). 3. After confirming the dissociation of cardiac clumps by light microscopy aspirate EDTA solution, add the half of the final volume of seeding medium into the well, triturate if needed and transfer the dissociated cellular mixture to the culture plate (see Notes 7 and 11). 4. Place the culture plate in a 37  C incubator and wait for 30 min (see Note 11). 5. After confirming settling of the cells by observing under light microscopy, gently add the other half of the seeding medium through the walls of the plate (see Notes 9 and 10). Overall, after following through these experimental steps, the highest cell viability, mostly single cell morphology, and beating frequencies were achieved in replated iPSC-CM cultures dissociated with the TrypLE enzyme [8]. Dissociation with different collagenase subtypes resulted in cardiac clusters at various sizes. Laminin, i-Matrix, and fibronectin provided replated hiPSC-CMs to adhere in a flatter and monolayer manner pattern compared to other matrices. The repleated hiPSC-CMs were observed to be more cluster-like phenotype when seeded on Geltrex or Matrigel than on other matrices [8]. Consequently, the selection of dissociation reagent and the ECM for coating for replated iPSC-CMs may need to be evaluated and adjusted for the desired outcome for the researcher due to user- or lab-related variables.

4

Notes 1. For the efficiency of differentiation, the iPSC confluency is critical and should be optimized for each iPSC line. Although 85–90% hiPSC confluency would be recommended to start the differentiation process in literature, 60–70% confluency has been reported to give the best results having highest percentage of functional cardiomyocytes [16]. As CHIR99021 provokes the cell proliferation, the initial cell number should not be higher than 85–90% to prevent the contact inhibition.

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2. The concentration of CHIR99021 should be optimized for each iPSC line. In the literature, the concentrations ranging from 6 to 10 μM CHIR has given efficient results. However, if CHIR99021 concentration is increased, the duration of the treatment should be optimized as this increased concentration may lead to higher toxicity. Another optimization area is the concentration and duration of small molecules for WNT inhibition used at differentiation days 3–5. 3. Human iPSC-CMs are differentiated as a heterogenous culture, and the cardiomyocytes need to be enriched for subsequent experimental evaluation. For this, one of the most commonly used methodology is to metabolically select iPSCCMs and to eliminate noncardiomyocytes by using lactate. Lactate provides a switch from glucose metabolism to fatty acid oxidation, the major source of energy in the heart [17]. The metabolic selection with lactate in glucose-depleted culture medium improves the efficacy of cardiac differentiation [17]. Secondly, Mitomycin-C selection can be also conducted to purify cardiomyocyte population in a heterogenous culture, as Mitomycin-C prevents proliferation of non-CMs, such as fibroblasts [18]. 4. When the spontaneous contractions initiate on Day 8 or forward, changing half of the medium is recommended not to lose the secreted factors that support the cardiac differentiation. The conditioned medium collected from the beating cells can be stored at 4  C. 5. Pretreatment of the cells with DPBS (without Ca2+/Mg2+) for a couple minute will ease the dissociation of the cardiac clumps. This pretreatment step should be applied in all enzyme treatments. 6. During the enzymatic dissociation of cardiac cultures, the cells should be checked by light microscopy every 2–3 min to evaluate the efficiency of treatment and when to terminate the reaction. If the cell clumps are large then the mixture can be triturated gently during the digestion period to achieve a homogeneous cell mixture with mostly single cells. 7. The digested cell clusters should be collected with the FBS-containing medium for inhibiting the enzymatic activity. 8. After dissociation and centrifugation, if cellular pellet cannot be seen in the falcon, the live cells might be unsettled because of the genomic DNA exposed from the dead cells. DNase enzyme can be readded after centrifugation to expose the live cells trapped in genomic DNA. 9. Using ROCK inhibitor Y27632 is highly recommended to increase adherence and cell viability after cellular stress induced by dissociation and replating. However, Y27632 should not

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remain in cell culture for more than 24 h as this molecule can also induce off-target signaling activities. Thus, within 24 h of reseeding the hiPSC-CM medium should be changed with prewarmed RPMI/B27 medium. It is preferred not to add Y27632 during reseeding, but less adhesion and survival of the cardiac cells may be observed. 10. After reseeding of iPSC-CMs, if low adherence and viability in the culture is observed, the freshness of the growth mediums and aliquots of small molecules as well as the homogeneous coating should be checked. The incubation time or concentration of ECM material or small molecules could be optimized based on the cell line. 11. After dissociation with EDTA, the iPSC-CMs can be directly seeded onto the culture wells. The centrifuge step can be also applied at 300  g for 3 min at 4  C.

Acknowledgments This work was supported by the Scientific and Technological € ˙ TAK) under grant number Research Council of Turkey (TUBI 213S192. References 1. Takahashi K, Tanabe K, Ohnuki M, Narita M, Ichisaka T, Tomoda K, Yamanaka S (2007) Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell 131 (5):861–872 2. Tertoolen LGJ, Braam SR, van Meer BJ, Passier R, Mummery CL (2018) Interpretation of field potentials measured on a multi electrode array in pharmacological toxicity screening on primary and human pluripotent stem cell-derived cardiomyocytes. Biochem Biophys Res Commun 497(4):1135–1141 3. Garg P, Garg V, Shrestha R, Sanguinetti MC, Kamp TJ, Wu JC (2018) Human induced pluripotent stem cell-derived cardiomyocytes as models for cardiac channelopathies: a primer for non-electrophysiologists. Circ Res 123 (2):224–243 4. Kitaguchi T, Moriyama Y, Taniguchi T, Maeda S, Ando H, Uda T, Otabe K, Oguchi M, Shimizu S, Saito H, Toratani A, Asayama M, Yamamoto W, Matsumoto E, Saji D, Ohnaka H, Miyamoto N (2017) CSAHi study: detection of drug-induced ion channel/receptor responses, QT prolongation, and arrhythmia using multi-electrode arrays in combination with human induced pluripotent

stem cell-derived cardiomyocytes. J Pharmacol Toxicol Methods 85:73–81 5. Giacomelli E, Bellin M, Sala L, van Meer BJ, Tertoolen LG, Orlova VV, Mummery CL (2017) Three-dimensional cardiac microtissues composed of cardiomyocytes and endothelial cells co-differentiated from human pluripotent stem cells. Development 144(6):1008–1017 6. Koivum€aki JT, Naumenko N, Tuomainen T, Takalo J, Oksanen M, Puttonen KA, Lehtonen Sˇ, Kuusisto J, Laakso M, Koistinaho J, Tavi P (2018) Structural immaturity of human iPSCderived cardiomyocytes: in silico investigation of effects on function and disease modeling. Front Physiol 9:80 7. Ellis BW, Acun A, Can UI, Zorlutuna P (2017) Human iPSC-derived myocardium-on-chip with capillary-like flow for personalized medicine. Biomicrofluidics 11(2):024105 8. Koc A, Sahoglu Goktas S, Akgul Caglar T, Cagavi E (2021) Defining optimal enzyme and matrix combination for replating of human induced pluripotent stem cell-derived cardiomyocytes at different levels of maturity. Exp Cell Res 403(2):112599 9. Santoro R, Perrucci GL, Gowran A, Pompilio G (2019) Unchain my heart: integrins at the

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basis of iPSC cardiomyocyte differentiation. Stem Cells Int 13:8203950 10. Oberwallner B, Brodarac A, Anic´ P, Sˇaric´ T, Wassilew K, Neef K, Choi YH, Stamm C (2015) Human cardiac extracellular matrix supports myocardial lineage commitment of pluripotent stem cells. Eur J Cardiothorac Surg 47(3):416–425 11. Wu X, Sun Z, Foskett A, Trzeciakowski JP, Meininger GA, Muthuchamy M (2010) Cardiomyocyte contractile status is associated with differences in fibronectin and integrin interactions. Am J Physiol Heart Circ Physiol 298(6): H2071–H2081 12. Trappmann B, Gautrot JE, Connelly JT, Strange DG, Li Y, Oyen ML, Cohen Stuart MA, Boehm H, Li B, Vogel V, Spatz JP, Watt FM, Huck WT (2012) Extracellular-matrix tethering regulates stem-cell fate. Nat Mater 11(7):642–649 13. Burridge PW, Matsa E, Shukla P, Lin ZC, Churko JM, Ebert AD, Lan F, Diecke S, Huber B, Mordwinkin NM, Plews JR, Abilez OJ, Cui B, Gold JD, Wu JC (2014) Chemically defined generation of human cardiomyocytes. Nat Methods 11(8):855–860 14. Miyazaki T, Futaki S, Suemori H, Taniguchi Y, Yamada M, Kawasaki M, Hayashi M, Kumagai H, Nakatsuji N, Sekiguchi K, Kawase E (2012) Laminin E8 fragments support efficient adhesion and expansion of dissociated

human pluripotent stem cells. Nat Commun 3:1236 15. Nishiuchi R, Takagi J, Hayashi M, Ido H, Yagi Y, Sanzen N, Tsuji T, Yamada M, Sekiguchi K (2006) Ligand-binding specificities of laminin-binding integrins: a comprehensive survey of laminin-integrin interactions using recombinant alpha3beta1, alpha6beta1, alpha7beta1 and alpha6beta4 integrins. Matrix Biol 25(3):189–197 16. Balafkan N, Mostafavi S, Schubert M, Siller R, Liang KX, Sullivan G, Bindoff LA (2020) A method for differentiating human induced pluripotent stem cells toward functional cardiomyocytes in 96-well microplates. Sci Rep 10 (1):18498 17. Tohyama S, Hattori F, Sano M, Hishiki T, Nagahata Y, Matsuura T, Hashimoto H, Suzuki T, Yamashita H, Satoh Y, Egashira T, Seki T, Muraoka N, Yamakawa H, Ohgino Y, Tanaka T, Yoichi M, Yuasa S, Murata M, Suematsu M, Fukuda K (2013) Distinct metabolic flow enables large-scale purification of mouse and human pluripotent stem cellderived cardiomyocytes. Cell Stem Cell 12 (1):127–137 18. Mummery CL, van Achterberg TA, van den Eijnden-van Raaij AJ, van Haaster L, Willemse A, de Laat SW, Piersma AH (1991) Visceral-endoderm-like cell lines induce differentiation of murine P19 embryonal carcinoma cells. Differentiation 46(1):51–60

Methods in Molecular Biology (2022) 2520: 171–180 DOI 10.1007/7651_2021_459 © Springer Science+Business Media, LLC 2022 Published online: 17 May 2022

Hematopoietic Cell Isolation by Antibody-Free Flow Cytometry in the Zebrafish Embryo Katsuhiro Konno, Jingjing Kobayashi-Sun, Fumio Arai, Isao Kobayashi, and Daisuke Sugiyama Abstract The zebrafish is a useful model to identify genes functioning in hematopoiesis, owing to high conservation of hematopoiesis. Flow cytometry is widely used to isolate and quantitatively characterize human and mouse hematopoietic cells, often using fluorescently labeled antibodies. However, such analysis is limited in zebrafish due to lack of antibodies that recognize zebrafish hematopoietic cells. We here describe methods for isolation of hematopoietic cells by antibody-free flow cytometry in the zebrafish embryo. Hematopoietic stem cells (HSCs) are specified from a specific subset of vascular endothelial cells, the hemogenic endothelial cell (HEC), by a high level of Notch signaling. In combination with a Notch reporter line (Tp1:GFP) and a vascular specific line ( fli1a:dsRed), HECs can be isolated as Tp1:GFPhigh fli1a:dsRed+ cells at 20–22 hours post-fertilization (hpf). Zebrafish erythrocytes can be purified using 1,5-bis{[2-(dimethylamino)ethyl]amino}-4, 8-dihydroxyanthracene-9,10-dione (DRAQ5), a DNA-staining fluorescent dye, and gata1:dsRed (erythroid marker line). DRAQ5high dsRed+ cells are morphologically erythrocyte-like, hemoglobin-stained positive, and express erythropoiesis-related genes. Zebrafish neutrophils can be also isolated using the lectin Phaseolus vulgaris erythroagglutinin (PHA-E) and DRAQ5. PHA-Elow DRAQ5low cells have myeloperoxidase activity, are Sudan Black B-positive, and express neutrophil-related genes. These methods will help to perform genetical and functional assays for various types of hematopoietic cells in zebrafish embryos. Key words Zebrafish, Hematopoietic stem cell, Hemogenic endothelial cell, Erythrocytes, Neutrophil

1

Introduction The zebrafish is an excellent model to study hematopoietic development because of a number of unique advantages. Its embryos are externally fertilized and transparent, enabling in vivo visualization of early embryonic processes including developmental process of hematopoietic cells. In addition, genome editing systems and chemical screening make it possible to analyze hematopoietic ontogeny and mechanisms of blood diseases [1, 2].

Katsuhiro Konno and Jingjing Kobayashi-Sun contributed equally with all other contributors.

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Hematopoietic stem cells (HSCs) are defined by their abilities to self-renew and differentiate into all types of blood lineages, including erythroid, myeloid, and lymphoid lineage. During development, HSCs are formed from a specific subset of vascular endothelial cells, termed the hemogenic endothelial cell (HEC), through endothelial-to-hematopoietic transition (EHT) in the dorsal aorta [3–6]. After arising from the dorsal aorta, HSCs move to the transient hematopoietic organ, fetal liver in mammals and caudal hematopoietic tissue in zebrafish, where HSCs proliferate and differentiate into multiple lineages of blood cells [7, 8]. HSCs finally colonize in the permanent hematopoietic organ, bone marrow in mammals and kidneys in zebrafish [1, 9]. It is important to isolate HECs/HSCs and their progenies to understand the molecular mechanisms underlying hematopoietic development as well as to perform genetic and chemical screening of blood diseases. Isolation of hematopoietic cells in the mouse and human is traditionally achieved by flow cytometry in combination of multiple fluorescently labeled antibodies against surface antigens. Due to lack of antibodies in zebrafish, however, fluorescent transgenic lines that label specific blood cell types have instead been developed. Here, we describe methods for isolation of hematopoietic cells by antibody-free flow cytometry in the zebrafish embryo. The fate of HSCs is established by Notch signaling in the shared vascular precursor, the angioblast. In zebrafish, angioblasts arise in the lateral plate mesoderm and migrate along the ventral surface of the somite toward the midline to form vascular endothelial cells and HECs. During axial migration, a part of angioblasts become HECs by receiving a high level of Notch signaling from the somite based on robust interaction by cell adhesion molecules [10, 11]. Therefore, a Notch reporter line, Tp1:GFP, can be used for isolation of HECs in zebrafish embryos. Tp1:GFP expresses GFP under the control of Notch regulatory elements and its expression reflects the Notch activation level [12]. In combination with a vascular specific dsRed line, fli1a:dsRed, HECs can be detected as Tp1:GFPhigh fli1a:dsRed+ cells at 20–22 hours postfertilization (hpf), which is, to date, the earliest timing to isolate HSC precursors in zebrafish embryos [10]. Erythrocytes synthesize hemoglobin to supply to all tissues and organs and are generated from bone marrow through erythropoiesis [13]. In the bone marrow, erythroids are obtained by differentiating from megakaryocyte/erythroid progenitors differentiated from HSC. Zebrafish erythropoiesis is continuously performed in the mesodermal germ layer by two waves: a primitive wave and a definitive wave. At 12 hpf, erythroid progenitor cells expressing gata1a which is zinc finger transcription factor differentiate into primitive erythrocytes [14]. Blood circulation begins at 24 hpf, and primitive hematopoietic cells circulate and mature. The red

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fluorescent agent, 1,5-bis{[2-(dimethylamino)ethyl] amino}-4, 8-dihydroxyanthracene-9,10-dione (DRAQ5) is a synthetic anthraquinone that enters living cells [15]. It specifically binds to DNA with high affinity. The fluorescence intensity of DRAQ5 depends on the DNA content and chromatin complexity and affects access to DRAQ5 [16]. Therefore, by combining gata1a:dsRed [17] and DRAQ5, zebrafish erythrocytes at 48 hpf can be detected as DRAQ5high gata1a:dsRed+ [18]. Neutrophils play important roles in the innate immune response. Neutrophils are formed by the differentiation of granulocyte/monocyte progenitors in the bone marrow. Zebrafish neutrophils have myeloperoxidase granules [19] and are produced by myelopoiesis, which is regulated by genes involved in neutrophil differentiation such as transcription factor Spi1 [20, 21], granulocyte-colony stimulating factor and its receptor [22]. That is similar to the characteristics of neutrophils in humans and mice. Lectins, oligosaccharide-binding proteins, have been used to classify hematopoietic cells [23]. Phaseolus vulgaris erythroagglutinin (PHA-E), a type of lectin, binds to Terminal galactose, which is widely present in neutrophils [24]. Thus, by using PHA-E and DRAQ5, it is possible to isolate neutrophils from wild-type larvae as DRAQ5lowPHA-Elow at 96 hpf [25]. Here, we show the methods for isolation of HECs, erythrocytes, and neutrophils in zebrafish embryos and larvae.

2 2.1

Materials Isolation of HECs

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Double-transgenic zebrafish embryo, Tp1:GFPum14; fli1a: dsRedum13.

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E3 medium: 5 mM NaCl, 0.17 mM KCl, 0.33 mM CaCl2, 0.33 mM MgSO4.

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Pronase solution: 20 mg/mL Pronase (Sigma, 10165921001) in E3 medium.

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Cell medium: 2% fetal bovine serum (FBS) in phosphate buffered saline (PBS).

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Collagenase solution: 5401119001) in PBS.

l

l

50

μg/mL

Liberase

(Sigma,

5 mL round bottom tube with 35 μm cell strainer (Corning, 352235). 5 μM Sytox red solution (Thermo Fisher, S34859).

l

Lysis buffer: 1% 2-mercaptoethanol (Wako, 135-07522), 500 μg/mL polyinosinic acid (Poly(I)) (Sigma, P4154) in Buffer RLT (RNeasy Mini Kit, Qiagen, 74104).

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Fluorescent stereomicroscope: Axiozoom V16 (Zeiss).

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Flow cytometer: FACS Aria III (BD).

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2.2 Isolation of Erythrocytes

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Transgenic zebrafish embryo, gata1:dsRed.

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Fluorescence microscope: BZ-X800 (Keyence).

l

Egg water: 1 L of RO water containing 0.03% NaCl per 1 drop of new methylene blue (Muto pure chemicals, 15861).

l l

2.3 Isolation of Neutrophils

5 mM DRAQ5 (BioStatus, DR50200).

l

1 mM TO-PRO-1 iodide (Invitrogen, T3602).

l

Flow cytometer: FACS Aria (BD).

l

Wild-type zebrafish embryo.

l

Egg water: 1 L of RO water containing 0.03% NaCl per 1 drop of new methylene blue (Muto pure chemicals, 15861).

l

Wash buffer: ice-cold 0.9
PBS.

l

Cell medium: 0.9
PBS containing 2% FBS.

l

3.1

5 mL round bottom tube with 35 μm cell strainer (Corning, 352235).

l

l

3

Cell medium: 0.9
PBS containing 2% FBS.

Collagenase solution: 0.9
PBS containing 2% FBS and 1 mg/ mL collagenase type IV (Sigma, C428). 5 mL round bottom tube with 35 μm cell strainer (Corning, 352235).

l

Biotin-labeled PHA-E (cosmobio, J211).

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DRAQ5 (BioStatus, DR50200).

l

1 mM SYTOX Blue (Invitrogen, S34857).

l

Flow cytometer: FACS Aria (BD).

l

May-Gru¨nwald Giemsa staining: CytoSpin™4 (Thermo Fisher Scientific), May-Gru¨nwald reagent (Muto pure chemicals, 15053), Giemsa solution (Muto pure chemicals, 15003).

Methods Isolation of HECs

1. Obtain double-transgenic embryos by crossing Tp1:GFP and fli1a:dsRed zebrafish. 2. Incubate embryos in E3 medium at 28.5  C until 18–19 hpf (see Note 1). 3. Treat embryos with Pronase solution at 28.5  C for 10–20 min (dechorionation). 4. Screen GFP and dsRed double-positive embryos using a fluorescent stereomicroscope (see Note 2). 5. Collect embryos (20–22 hpf) into a 1.5 mL tube (up to 50 embryos/tube) and treat with 500 μL of collagenase solution at 37  C for 1 h.

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6. Pipet to suspend cells from embryos. 7. Add 1 mL of cell medium and filter cells with a 35 μm cell strainer into a 5 mL round bottom tube. 8. Centrifuge cells at 1600 rpm (approx. 400
g) for 7 min at 4  C. 9. Resuspend cells with 1 mL of cell medium. 10. Filter cells again with a 35 μm cell strainer into a 5 mL round bottom tube. 11. Add 1 μL of Sytox red solution to exclude dead cells and debris on the flow cytometer. 12. Set gates on the flow cytometer to isolate Tp1:GFPhigh fli1a: dsRed+ cells (Fig. 1) (see Note 3). 13. Sort 5000–10,000 cells into 500 μL of lysis buffer (see Note 4).

Fig. 1 Isolation of hemogenic endothelial cells (HECs) from zebrafish embryos. (a–e) Representative result of flow cytometric analysis in Tp1:GFP; fli1a:dsRed embryos at 22 hpf. Sytox Red-stained dead cells and FSC-Alow debris were excluded using gate A (a). Single cells were selected using gate B in a FSC-A vs. FSC-W plot (b) and gate C in a FSC-A vs. SSC-W plot (c). Cells in the SSC-Alow fraction, which contains endothelial cells, were selected using gate D (d) and displayed in a Tp1:GFP vs. fli1a:dsRed plot (e). fli1a:dsRed+ cells (endothelial cells) were subdivided into three fractions based on expression of Tp1:GFP, “Tp1-neg,” “Tp1-lo,” and “Tp1-hi.” (f) Expression of HEC marker genes, runx1, gata2b, and gfi1aa, in Tp1-neg, Tp1-lo, or Tp1-hi cells at 22 hpf. HEC marker genes were highly expressed in Tp1-hi cells (Tp1: GFPhigh fli1a:dsRed+ cells). Error bars, s.d. (n ¼ 3)

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3.2 Isolation of Erythrocytes

1. Obtain transgenic embryos by crossing gata1:dsRed and wildtype zebrafish. 2. Incubate embryos in egg water at 28  C until 48 hpf. 3. Screen dsRed-positive embryos using a fluorescent microscope (see Note 5). 4. Wash embryos three times with ice-cold 0.9
PBS. 5. After removing 0.9
PBS, immerse the embryos in the cell medium, crushed with the plunger of 1-mL syringe and passed through a 35-μm nylon filter (see Note 6). 6. Wash the resulting cells three times with 0.9
PBS. 7. Centrifuge the cells at 200
g for 5 min at room temperature. 8. Resuspend in cell medium. 9. Count living cells hemocytometer.

stained

with

Trypan

blue

using

10. Suspend 1
105 cells in 1 mL of 5 μmol/L DRAQ5 in cell medium and incubate in the dark at room temperature for 15 min. 11. Add 1 μL of TO-PRO-1 iodide to exclude dead cells. 12. Set gates on the flow cytometer to isolate DRAQ5high gata1: dsRed+ cells (Fig. 2) (see Note 7). 3.3 Isolation of Neutrophils

1. Obtain embryos by crossing wild-type zebrafish. 2. Incubate embryos in egg water at 28  C until 96 hpf. 3. Wash larvae three times with ice-cold 0.9
PBS. 4. After removing 0.9
PBS, mince larvae with a microscissors, immerse in ice-cold collagenase solution (see Note 8). 5. Incubate at 37  C for 40 min with intermittent shaking every 5 min. 6. Pass the cell through 35-μm nylon filter (see Note 9). 7. Centrifuge the cells at 200
g for 5 min at room temperature and resuspend them in cell medium three times. 8. Count living cells hemocytometer.

stained

with

Trypan

blue

using

9. Incubate 5
105 cells in cell medium with 10 μg/mL biotinlabeled PHA-E for 30 min on ice (see Note 10). 10. Centrifuge the cells at 200
g for 5 min at room temperature and resuspend them in 500 μL of cell medium. 11. Add 1 μL of PE-labeled streptavidin and incubate 30 min on ice (see Note 10). 12. After one wash with cell medium, suspend the cells in the 1 mL of cell medium containing 5 μM DRAQ5.

Hematopoietic Cell Isolation by Antibody-Free Flow Cytometry in the. . .

177

Fig. 2 Isolation of erythrocytes from zebrafish embryos at 48 hpf. (a–c) Representative analysis of flow cytometry in gata1:dsRed embryos at 48 hpf. (a) Dead cells are removed using TO-PRO-1. (b) The fraction surrounded in the red line is selected that contains erythroids. (c) dsRed vs. DRAQ5 plot. dsRed is detected by PE and DRAQ5 is detected by PerCP-Cy5. Only TO-PRO-1, TO-PRO-1 + DRAQ5 (wild-type embryos), TO-PRO-1 + DRAQ5 + dsRed (gata1:dsRed + embryos) cases are shown

13. Keep the cells in the dark at room temperature for 15 min. 14. Add 1 μL of SYTOX Blue to exclude dead cells. 15. Set gates on the flow cytometer to isolate PHA-Elow DRAQ5low cells (Fig. 3) (see Note 11).

4

Notes 1. To obtain 18–19 hpf embryos in the morning hours, incubate embryos at 28.5  C for 8 h, then move to 23  C, and further incubate for 16–18 h.

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Fig. 3 Isolation of neutrophils from zebrafish larvae at 96 hpf. (a, b) Representative analysis of flow cytometry in wild-type larvae at 96 hpf. (a) Unstained samples are shown as negative controls. Dead cells are removed using SYTOX Blue (detected by Pacific Blue). (b) Analysis example of stained sample. Live cells are first selected as Pacific Blue- V500- and developed by FSC-A and SSC-A. Next, the fraction surrounded by the red line is selected and expanded it with DRAQ5 (detected by PerCP-Cy5) and PE. Neutrophils contain in the fraction of PHA-Elow DRAQ5low. (c) Example of May-Gru¨nwald Giemsa staining of PHA-Elow DRAQ5low fractionated cells. Scale bar: 20 μm

2. The expression of fli1a:dsRed at 18–19 hpf is very dim. If dsRed is not detectable, GFP-positive embryos should at least be screened by a fluorescent stereomicroscope before cell preparation. 3. The signal from dsRed may leak into the GFP channel, resulting in false-positive detection. To compensate properly, embryos having GFP only and dsRed only should be prepared as well. 4. We estimated that 50–80 Tp1:GFPhigh fli1a:dsRed+ cells can be isolated from a 22 hpf embryo, indicating that 100–200 double-positive embryos should be used to isolate 5000–10,000 Tp1:GFPhigh fli1a:dsRed+ cells. 5. DsRed-positive erythroid cells can be seen in the blood circulation. If you do not care about efficiency of sorting the target cells, you can skip this step. 6. In case of crushing a large amount of embryo, you can do it well by loading it into the cell strainer, removing as much water as possible, and then crushing it.

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7. Single-stained or only dsRed-positive sample as well needs to be prepared to compensate properly. 8. It is enough to mince several times with microscissors. Be careful not to let the larvae dry. 9. A few minutes after the shaking is finished, the residue settles on the bottom of the container. If you pass it through the cell strainer from the supernatant side first, clogging is unlikely to occur. 10. If more than 1
105 cells are put in one tube, it may cause aggregation in this step. 11. Single-stained sample as well needs to be prepared to compensate properly. References 1. de Jong JL, Zon LI (2005) Use of the zebrafish system to study primitive and definitive hematopoiesis. Annu Rev Genet 39:481–501 2. Wu RS, Lam II, Clay H, Duong DN, Deo RC, Coughlin SR (2018) A rapid method for directed gene knockout for screening in G0 zebrafish. Dev Cell 46:112–125 3. Zovein AC, Hofmann JJ, Lynch M, French WJ, Turlo KA, Yang Y, Becker MS, Zanetta L, Dejana E, Gasson JC et al (2008) Fate tracing reveals the endothelial origin of hematopoietic stem cells. Cell Stem Cell 3:625–636 4. Bertrand JY, Chi NC, Santoso B, Teng S, Stainier DY, Traver D (2010) Haematopoietic stem cells derive directly from aortic endothelium during development. Nature 464: 108–111 5. Kissa K, Herbomel P (2010) Blood stem cells emerge from aortic endothelium by a novel type of cell transition. Nature 464:112–115 6. Boisset JC, van Cappellen W, Andrieu-Soler C, Galjart N, Dzierzak E, Robin C (2010) In vivo imaging of hematopoietic cells emerging from the mouse aortic endothelium. Nature 464: 116–120 7. Murayama E, Kissa K, Zapata A, Mordelet E, Briolat V, Lin HF, Handin RI, Herbomel P (2006) Tracing hematopoietic precursor migration to successive hematopoietic organs during zebrafish development. Immunity 25: 963–975 8. Jin H, Xu J, Wen Z (2007) Migratory path of definitive hematopoietic stem/progenitor cells during zebrafish development. Blood 109: 5208–5214 9. Davidson AJ, Zon LI (2004) The ‘definitive’ (and ‘primitive’) guide to zebrafish hematopoiesis. Oncogene 23:7233–7246

10. Kobayashi I, Kobayashi-Sun J, Kim AD, Pouget C, Fujita N, Suda T, Traver D (2014) Jam1a-Jam2a interactions regulate hematopoietic stem cell fate through Notch signalling. Nature 512:319–323 11. Rho SS, Kobayashi I, Oguri-Nakamura E, Ando K, Fujiwara M, Kamimura N, Hirata H, Iida A, Iwai Y, Mochizuki N, Fukuhara S (2019) Rap1b promotes notch-signalmediated hematopoietic stem cell development by enhancing integrin-mediated cell adhesion. Dev Cell 49:681–696 12. Parsons MJ, Pisharath H, Yusuff S, Moore JC, Siekmann AF, Lawson N, Leach SD (2009) Notch-responsive cells initiate the secondary transition in larval zebrafish pancreas. Mech Dev 126:898–912 13. McGrath K, Palis J (2008) Ontogeny of erythropoiesis in the mammalian embryo. Curr Top Dev Biol 82:1–22 14. Berman JN, Kanki JP, Look AT (2005) Zebrafish as a model for myelopoiesis during embryogenesis. Exp Hematol 33(9):997–1006 15. Smith PJ, Blunt N, Wiltshire M, Hoy T, Teesdale-Spittle P, Craven MR, Watson JV, Amos WB, Errington RJ, Patterson LH (2000) Characteristics of a novel deep red/infrared fluorescent cell-permeant DNA probe, DRAQ5, in intact human cells analyzed by flow cytometry, confocal and multiphoton microscopy. Cytometry 40(4):280–291 16. Smith PJ, Wiltshire M, Davies S, Patterson LH, Hoy T (1999) A novel cell permeant and far red-fluorescing DNA probe, DRAQ5, for blood cell discrimination by flow cytometry. J Immunol Methods 229(1–2):131–139 17. Traver D, Paw BH, Poss KD, Penberthy WT, Lin S, Zon LI (2003) Transplantation and

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in vivo imaging of multilineage engraftment in zebrafish bloodless mutants. Nat Immunol 4: 1238–1246 18. Kulkeaw K, Inoue T, Ishitani T, Nakanishi Y, Zon LI, Sugiyama D (2018) Purification of zebrafish erythrocytes as a means of identifying a novel regulator of hematopoiesis. Br J Haematol 180(3):420–431 19. Bennett CM, Kanki JP, Rhodes J, Liu TX, Paw BH, Kieran MW, Langenau DM, DelahayeBrown A, Zon LI, Fleming MD, Look AT (2001) Myelopoiesis in the zebrafish, Danio rerio. Blood 98(3):643–651 20. Le Guyader D, Redd MJ, Colucci-Guyon E, Murayama E, Kissa K, Briolat V, Mordelet E, Zapata A, Shinomiya H, Herbomel P (2008) Origins and unconventional behavior of neutrophils in developing zebrafish. Blood 111(1): 132–141 21. Lieschke GJ, Oates AC, Paw BH, Thompson MA, Hall NE, Ward AC, Ho RK, Zon LI, Layton JE (2002) Zebrafish SPI-1 (PU.1)

marks a site of myeloid development independent of primitive erythropoiesis: implications for axial patterning. Dev Biol 246(2):274–295 22. Pazhakh V, Hock B, Fearnley D, McLellan A, Vuckovic S, Hart DN (2017) A GCSFR/ CSF3R zebrafish mutant models the persistent basal neutrophil deficiency of severe congenital neutropenia. Sci Rep 7:44455 23. El Sherbini H, Hock B, Fearnley D, McLellan A, Vuckovic S, Hart DN (2000) Lectin ligands on human dendritic cells and identification of a peanut agglutinin positive subset in blood. Cell Immunol 200(1):36–44 24. Madsen-Bouterse SA, Xu Y, Petty HR, Romero R (2008) Quantification of O-GlcNAc protein modification in neutrophils by flow cytometry. Cytometry A 73(7):667–672 25. Konno K, Kulkeaw K, Sasada M, Nii T, Kaneyuki A, Ishitani T, Arai F, Sugiyama D (2020) A novel method to purify neutrophils enables functional analysis of zebrafish hematopoiesis. Genes Cells 25(12):770–781

Methods in Molecular Biology (2022) 2520: 181–187 DOI 10.1007/7651_2021_460 © Springer Science+Business Media, LLC 2022 Published online: 27 February 2022

Human Trophectoderm Spheroid Derived from Human Embryonic Stem Cells Wen Huang, Sze Wan Fong, William Shu Biu Yeung, and Yin Lau Lee Abstract The use of human embryos for studying the early implantation processes and trophoblast is restricted by ethical concerns. The development of models mimicking the peri-implantation embryos is critical for understanding the physiology of human embryos and many pathophysiological disorders including recurrent implantation failure and miscarriage. Three-dimensional (3D) models of trophoblastic spheroids have been successfully derived from human embryonic stem cells (hESC). Simultaneous activation of the BMP pathway and blockage of the Activin/Nodal pathway favor the differentiation of hESC into trophoblast. Here we describe a 3D trophectoderm differentiation protocol with the use of BAP (BMP4, A83-01, and PD173074) to generate hESC-derived trophectoderm spheroids (BAP-EB). BAP-EB is highly reproducible and exhibits morphological and transcriptomic similarities to human early blastocysts. Key words Human embryonic stem cells, Trophectoderm spheroids, BAP-EB, 3-Dimensional culture, In vitro implantation model, Early implantation processes

1

Introduction Despite the use of assisted reproduction for treatment of infertility, many treatment cycles fail to finish with successful implantation. Due to ethical concerns, studies of human peri-implantation development and early placentation are difficult without an appropriate model. Although some in vitro models mimicking human embryo implantation have been generated from primary trophoblast or choriocarcinoma cell lines, they showed obvious shortcomings in terms of limited proliferation and cancerous characteristics, respectively [1]. Human embryonic stem cells (hESC) derived from human blastocysts [2] are able to differentiate into all three embryonic germ layers, namely ectoderm, mesoderm, and endoderm. Recently, bone morphogenic protein 4 (BMP4) or its homologues treatment was shown to drive the differentiation of hESC into extraembryonic lineages [3–5]. BMPs are members of the transforming growth factor β (TGF-β) family of cytokines, which play crucial roles in the loss of pluripotency and the acquisition of trophoblast characteristics [6]. A combination of BMP4 with

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Fig. 1 The morphologies of hESC-derived BAP-EB at different time points. (a) After 24 h aggregation in AggreWell™ plate, hESC form homogenous EB in microwells. (b) After 48 h of BAP treatment, BAP-EB form spheroids with similar sizes to that of human blastocysts and possess blastocoel-like cavities

inhibitors of ALK4/5/7 (A83-01) and FGF2 (PD173074) pathways was subsequently reported to induce trophoblast differentiation from hESC with higher efficiency and long-term proliferation ability [7, 8]. Conventional differentiation protocols of hESC-derived trophoblast are conducted in 2-dimensional Matrigel-coated plates. Here we describe a novel 3-dimensional trophoblast differentiation protocol to generate hESC-derived trophectoderm spheroids (BAP-EB) [9] mimicking the early implanting human embryos. Human ESC derived embryonic bodies (EBs) are first generated using AggreWell plates (Fig. 1a). The EBs formed are then treated with BMP4, A83-01, and PD173074 to induce trophoblast differentiation. Upon 48 h of BAP treatment (BAPEB48h), cystic structure similar to that in human blastocysts (Fig. 1b) is formed. Transcriptomic analysis reveals a trophectoderm-like signature of BAP-EB [10]. Together with the ability to selectively attach onto receptive endometrial epithelial cells, our BAP-EB model can be used as embryo surrogates for studying human early embryo implantation and trophoblast development.

Human Trophectoderm Spheroid Derived from Human Embryonic Stem Cells

2

183

Materials The hESC line, VAL3, is used to generate BAP-EBs in this protocol. All centrifugation processes are performed at room temperature.

2.1 Equipment Required for Cell Culture and Spheroid Formation

1. Vertical laminar flow hood of biosafety level II with stereomicroscope. 2. Incubator with temperature at 37  C, CO2 concentration at 5%, and humidity >95%. 3. Low speed Swing-Bucket centrifuge fitted with plate holder and tube holder. 4. Inverted microscope with 2, 4, and 10 phase objectives. 5. Hemacytometer. 6. Water bath.

2.2 Materials Required for hESC Culture

1. Matrigel-coated 6-well culture plate: Coat the desired number of culture wells with Matrigel for at least 1 h at room temperature (see Note 1). The coated plate can be stored at 4  C for up to 1 week. Prewarm the coated plate to room temperature (>1 h) and aspirate the coating solution just before use (see Note 2). 2. Complete mTeSR™ Plus medium: Thaw mTeSR™ Plus 5 Supplement at room temperature or overnight at 2–8  C (see Note 3). To prepare 500 ml of complete mTeSR™ Plus medium, add 100 ml of mTeSR™ Plus 5 Supplement to 400 mL of mTeSR™ Plus Basal medium (see Note 4). Mix thoroughly. 3. Accutase Cell Dissociation Reagent.

2.3 Materials Required for BAP-EB Formation

1. AggreWell™ 400 (24-well) plate.

2.4 Materials Required for Suspension Culture of BAP-EB

1. Ultra-low attachment 6-well plates.

2. Medium for spheroid formation: Complete mTeSR™ Plus medium supplemented with 10 μM Y-27632.

2. Mouse embryonic fibroblast (MEF) medium: DMEM (high glucose), 1% P/S, 10% FBS, 1% MEM Non-Essential Amino Acids, and 0.1% β-Mercaptoethanol (see Note 5). 3. MEF cells: Sacrifice pregnant mice at around 13.5 days postcoitus and dissect out the uterine horns followed by several washes with PBS in a Petri dish. Separate each embryo from its placenta and embryonic sac. Wash the embryos with PBS and then mince them into fragments (30 )

Comments See Table 2 for the well barcode

CTACACGACGCTCTTCCGATCT[8-bp well barcode]NNNNNNNNNNTTTTTTTTTTTTTTTTTTTTTTTTTVN

Template AAGCAGTGGTATCAACGCAGAGTACATrGrGrG Switching Oligo (TSO) cDNA PCR Fwd Primer

CTACACGACGCTCTTCCGATCT

cDNA PCR Rev. Primer

AAGCAGTGGTATCAACGCAGAG

rG indicates riboguanosine

Library PCR AATGATACGGCGACCACCGAGATCTACACTCTTTCCCTACACGACGCTC Universal Primer Library PCR Index Primer

See Table 3 for the plate index

CAAGCAGAAGACGGCATACGAGAT[8-bp plate barcode]GTCTCGTGGGCTCGG

3. FACS buffer: dissolve 1 g bovine serum albumin (BSA) powder in 100 mL DPBS. Pass through 0.22 μm filter and store in 4  C. 4. Lysis Buffer (freshly prepared). Stock

For one well

Final Concentration

50% PEG-8000

0.4 μL

6.67%

10% Triton X-100

0.03 μL

0.1%

RNase Inhibitor (40 U/μL)

0.04 μL

0.53 U/μL

dNTPs (25 mM/each)

0.08 μL

0.67 mM/each

ddH2O

1.45 μL

N/A

Total

2 μL

N/A

5. Reverse Transcription Mix (4) (freshly prepared).

Stock

For one Final well concentration

1 M Tris–HCl, pH 8.0

0.1 μL

1 M NaCl

0.12 μL 120 mM

100 mM

(continued)

238

Jianqun Zheng et al.

Stock

For one Final well concentration

100 mM MgCl2

0.1 μL

100 mM GTP

0.04 μL 4 mM

100 mM DTT

0.32 μL 32 mM

RNase Inhibitor (40 U/μL)

0.05 μL 2 U/μL

Template Switching Oligo (TSO) (100 μM)

0.08 μL 8 μM

Maxima H-minus RT Enzyme (200 U/μL) (Thermo Fisher, cat. mo. EP0751)

0.04 μL 8 U/μL

ddH2O

0.15 μL N/A

Total

1 μL

10 mM

N/A

6. cDNA Amplification Mix (freshly prepared). For one Final well concentration

Stock KAPA HiFi HotStart ReadyMix (2) (KAPA, cat. no. KK2601)

5 μL

1.67

cDNA PCR Fwd Primer (100 μM)

0.05 μL

0.83 μM

cDNA PCR Rev. Primer (100 μM)

0.05 μL

0.83 μM

ddH2O

0.9 μL

N/A

Total

6 μL

N/A

7. Column Wash Buffer. Stock

For 50 mL

Final concentration

1 M Tris–HCl, pH 7.5

0.5 mL

10 mM

100% Ethanol

40 mL

80%

ddH2O

9.5 mL

N/A

Total

50 mL

N/A

8. TD Buffer (2), store in 20  C and discard after 3 months. Stock

For 1 mL

Final concentration

1 M Tris–HCl, pH 7.5

20 μL

20 mM

1 M MgCl2

10 μL

10 mM

Dimethylformamide (DMF)

200 μL

20%

ddH2O

770 μL

N/A

Total

1 mL

N/A

B01

CCGTTTAG

J01

AGACTACT

B02

CTGAACGA

J02

CAACTATA

B03

GAGCCTCG

J03

ATGATCTT

B04

CTTCCGGA

J04

CCCGCACA

B05

TGCCCTGG

J05

ACCAAGGC

B06

TCGCTAGA

J06

A01

AGTAAACC

I01

CAGTACTG

A02

TCACCTAG

I02

ACCTCTGT

A03

CTTTAAGA

I03

GACCGGAC

A04

AAGATTAC

I04

TAATACAC

A05

CCATTATT

I05

TGAGGTTT

A06

AGTTACAG

I06

Table 2 Well barcodes

K06

CACACTCT

C06

GAGTCCAG

K05

ATGAGGCA

C05

GTTATGGT

K04

GCCGAACT

C04

CCTTCAGG

K03

AGAAGGAC

C03

TGGGACCG

K02

AATTGGCT

C02

TCTACTGC

K01

GACGCCGA

C01

L06

GTAGGGTC

D06

CTTCTACA

L05

GATGACAC

D05

AGGCGTTG

L04

TGATGCTG

D04

TGAGATCA

L03

TCCGTCTT

D03

GTTAGGAC

L02

GGCGTATC

D02

GTCGGGAA

L01

TTACGGTT

D01

M06

CATACGGT

E06

AGCACTTT

M05

GATTAACG

E05

TATGAAGT

M04

GAAGAGTA

E04

GTTCCACT

M03

AGGGTTTC

E03

GGAGATGC

M02

ACATAGCG

E02

GAATCATA

M01

TCATGAAA

E01

N06

ATCTAATC

F06

CAGGTAGC

N05

AGCGTGGC

F05

GTCATCTC

N04

CGCACAGG

F04

AAGGAGAC

N03

GACAGAAA

F03

ACCTTACA

N02

TGGACAGC

F02

ACTGTGGG

N01

AGGGACGT

F01

O06

TCAGTCAG

G06

TTACAGCA

O05

CTGAGTAA

G05

CCACGGCA

O04

ACGTGTCC

G04

TGATGTGA

O03

TCACACGG

G03

CATCGCAG

O02

GATCTCAA

G02

TTCAGCAT

O01

CACATGCG

G01

(continued)

P06

GGGCGTCA

H06

GCTTGCAG

P05

TCACCCTT

H05

AGGTCTAG

P04

TTTCTCAT

H04

CCCATCTG

P03

CTTTCGCT

H03

TTGACGTT

P02

CTCGGTTT

H02

CGGCATCC

P01

GTTCCTTC

H01

Single Cell Gene Expression Profiling of Embryoid Body 239

GCGTACCT

B07

CGCCAGTT

J07

GGCCTCTG

B08

AGTCTGTA

J08

GAGAGATT

B09

GGTAAGAT

J09

TATGGTCT

B10

AATTCAGC

J10

TACCCAAA

B11

TGCTGATC

J11

GGTGTCTA

B12

ACCACATC

J12

TGAGGAAC

A07

ATAAGTGG

I07

ACGGCTGA

A08

TCAAGTCG

I08

CGAGAGAC

A09

TTCTGCGC

I09

AGATCCTC

A10

TCCGAGTG

I10

AGTAATGT

A11

GTGACGAG

I11

TAGCAAAT

A12

CATCTCCT

I12

Table 2 (continued)

K12

GGATGTAG

C12

CCCTGTGG

K11

CCTGTTGT

C11

GTGGGCTG

K10

GTACTTCA

C10

GCGCAGGA

K09

ACACCATA

C09

TCCCTCCG

K08

CACGCAAT

C08

TATTGACC

K07

TATGCCAA

C07

CATACTTA

L12

TTGGAGGA

D12

ATAACGCC

L11

AAACACCA

D11

CCATTGCC

L10

CGGAGCAT

D10

CTCATAAG

L09

CAGGTTCG

D09

ATTTCTGA

L08

GTGTACGC

D08

CTAAAGAT

L07

GCGTTACC

D07

ATCCTGGG

M12

GGGTATTG

E12

TTTGTTGC

M11

ATCGCATG

E11

TGTTAGGG

M10

CCTTGTTA

E10

AAACGAGA

M09

CACTGAGA

E09

TTCTCAAC

M08

GAAGTAAT

E08

TGAATTGT

M07

GGTCTAGG

E07

TGGTGGTA

N12

CATAGACA

F12

GACAACTA

N11

GAATATCA

F11

ATCGGAAT

N10

GAGGAACC

F10

CGCTAGCC

N09

ATAGTCTC

F09

CCAGATCT

N08

AGTTCGCA

F08

GCTGCGCA

N07

ACAGAGAT

F07

AAACTTAG

O12

TTCGTGGT

G12

CGATGGCG

O11

TCGATCAC

G11

GCAATCCA

O10

TGCATGAT

G10

GCGGTCAT

O09

GCTAAGCG

G09

AGTATCGA

O08

CCGAATGC

G08

AAGTGATG

O07

TAGAGCTA

G07

CTTGACCC

P12

ACACCCAC

H12

ACGCCAAT

P11

CGTCGGGT

H11

CAGCCTTC

P10

ATACCCGG

H10

TTTACTTG

P09

TGGCCTAT

H09

GAGCGGTG

P08

TTCCGCTG

H08

CTCCACAC

P07

CTCTCTCC

H07

GCCACAGT

240 Jianqun Zheng et al.

TTTGTGCA

B13

ACCCTTTC

J13

ACTAGGAG

B14

TCAGCGAA

J14

CTATATAC

B15

CCTCCTCC

J15

ATCAAACG

B16

TAATTCCA

J16

ATGTCCGT

B17

CTCGCTAG

J17

ATCTTGTT

B18

CAGTGAAC

J18

TGAACGTA

CCGCAAAG

A13

GTTTACCA

I13

CAATACCT

A14

CGCTAATG

I14

AAGGCGTG

A15

GGAAACAA

I15

CAGCGCTT

A16

GCTGCAAC

I16

CACCTGAA

A17

ACGTAGTC

I17

TCTGCACG

A18

GGAGCTTG

I18

ACTCTCGC

GTGGGACT

K18

TTTCACGA

C18

CGAAGCGA

K17

GATATAGA

C17

TCAAGATC

K16

CTCCATTG

C16

TGTGCTAC

K15

TTGTTAGG

C15

GCCCGAGA

K14

ATTCTTGT

C14

GGCCTTGA

K13

CGAGCAGT

C13

GACTGCTT

CACTATAG

L18

ACCATGCT

D18

GAGCATAC

L17

TGACGCCT

D17

GGTGATCG

L16

AGGAGGGT

D16

GCATTGGA

L15

AACGGGTT

D15

TGTATCCT

L14

GAGAGCCC

D14

TTGGCATC

L13

TAGAGGAG

D13

AGAACTGC

ATGCATCA

M18

AGCCAAGC

E18

TGTTCATG

M17

GTGCTCCG

E17

CTATTGGG

M16

CATGTAAA

E16

GAGCCGGT

M15

GCACATTT

E15

GTAACTAT

M14

AGCCGCAG

E14

AGGGCGAA

M13

GGTACCAC

E13

CATTGTGC

TCAGTAGC

N18

GTTTGTTT

F18

GCGAGTGA

N17

AATAGGTC

F17

AGCGACAC

N16

GTATCGCG

F16

ACTAACTC

N15

ATTTGCCC

F15

ACTCAGCA

N14

TTTGAGTC

F14

GCCCATCC

N13

CTATAGTA

F13

GTACCAAG

GGCAGGTG

O18

TAAGCCCG

G18

ATCCAGCT

O17

TCCTAAGT

G17

GCGACTCT

O16

ACCCGCGT

G16

CTAGGACG

O15

CGGATAGA

G15

TACGTAGG

O14

GAGTTTCT

G14

CATTGATT

O13

TACCTTGT

G13

ACGGTGCA

(continued)

CATTCCAT

P18

CCGATGAA

H18

CAAGTCAC

P17

CGAGCTAA

H17

TATCGATA

P16

TGGAATTC

H16

TGCTTTAA

P15

TACGCGAG

H15

CGGTGCTC

P14

CCAACAGA

H14

TTAATCGG

P13

ACGGGACG

H13

TGCAACTT

Single Cell Gene Expression Profiling of Embryoid Body 241

B19

CTGAGATC

J19

CGGATGCT

B20

ATCACAGA

J20

ATTTCGAG

B21

TGTGGGAT

J21

GCTCTGCT

B22

GTCTGACA

J22

TCGGGCCT

B23

CCAGAATC

J23

GTTATATG

B24

CCATCCTG

J24

ATAGAAGT

A19

AGATTCAT

I19

GAATGAGG

A20

TCTTGCTC

I20

TGCATACC

A21

ACGTCCTA

I21

AACGATAA

A22

TGACATAC

I22

CATCTTGG

A23

TGTAGTGG

I23

TGATAGCT

A24

AACAGTGT

I24

TACTCGCC

Table 2 (continued)

CGTATTTG

K24

GTTCAGCC

C24

CACCCTGA

K23

AAGCCGCA

C23

GTCACGTA

K22

CCGACGTG

C22

TGGAGATG

K21

GACATTCC

C21

GAAGGTGT

K20

GAACTGCG

C20

ATTGCCAC

K19

GCTCCTGG

C19

GCGCGCAA

L24

TGGGTAAA

D24

ACGGGCAC

L23

GTCTTCAT

D23

AGATAAAC

L22

AATGTCGT

D22

CTATCCGC

L21

CTACAAGG

D21

CCGCACTA

L20

CGGGATAT

D20

TCCCATTA

L19

TACGAGCA

D19

ATGCTGTC

M24

GCAATTAC

E24

ATCAACAA

M23

TTATCCTA

E23

GGAATGGC

M22

GGTCATTA

E22

CTTAGAAT

M21

GCTGGCGA

E21

AGCTCATA

M20

CGCGTCAG

E20

CACATCGC

M19

TAGAATCA

E19

TGTACAAA

N24

AAGCGACT

F24

GGTTGAGC

N23

GACAAAGG

F23

AATTCTTG

N22

TAAGGCAT

F22

TAATCGGG

N21

CTAACATT

F21

GCTATCGG

N20

GTACGACC

F20

TTGTATCT

N19

CTCCTAGT

F19

CACGGCCT

O24

TTCTCGTG

G24

CCACTTTG

O23

CCGCGTAC

G23

TCCGGCAA

O22

ATCTCGGC

G22

AGCCTCCA

O21

AGCTATCG

G21

TTGGGTAT

O20

AAGAAGGA

G20

AGTGGAAA

O19

AGTGGCTG

G19

GCATATGG

P24

CGTGACGA

H24

TAGGCGCT

P23

AGTGTGCT

H23

CTGCAACT

P22

CCGATACG

H22

GCGGATTC

P21

TAGCTGAC

H21

CAACAGCC

P20

TCTTCTTT

H20

GCACCGTG

P19

GCATCGAC

H19

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Table 3 Plate index N701

TCGCCTTA

N702

CTAGTACG

N703

TTCTGCCT

N704

GCTCAGGA

N705

AGGAGTCC

N706

CATGCCTA

N707

GTAGAGAG

N710

CAGCCTCG

N711

TGCCTCTT

N712

TCCTCTAC

N714

TCATGAGC

N715

CCTGAGAT

N716

TAGCGAGT

N718

GTAGCTCC

N719

TACTACGC

N720

AGGCTCCG

N721

GCAGCGTA

N722

CTGCGCAT

N723

GAGCGCTA

N724

CGCTCAGT

N726

GTCTTAGG

N727

ACTGATCG

N728

TAGCTGCA

N729

GACGTCGA

3

Methods

3.1 Thawing and Culturing mESCs in Feeder-Free Condition

1. Coat the flask with Plate Coating Solution: add just enough solution to cover the bottom of the plastic. Leave at room temperature for at least 5 min. Remove just before plating the cells. 2. Thaw frozen vials of mESCs in 37  C water batch. 3. Centrifuge the cells at 250  g for 3 min and remove supernatant. 4. Wash the cell pellet with prewarmed Stem Cell Culture Media, centrifuge 250  g for 3 min and remove supernatant. 5. Resuspend the cell pellet in prewarmed Stem Cell Culture Media, and plate cells into the coated flask. 6. Shake the flask back and forth gently to distribute cells evenly. 7. Incubate the cells in a 37  C, 5% CO2 incubator. Change the medium once every day until they reach 80% confluency.

3.2 Passaging Feeder-Free mESCs

1. Aspirate the medium, prewarmed DPBS.

wash

the

cells

twice

with

2. Add just enough prewarmed 0.05% trypsin to cover all cells. 3. Incubate the cells at room temperature for 3 min, and check under a microscope whether cells are detached from the bottom of the plate or dish. 4. Add equal volume of Stem Cell Culture Media to inactivate trypsin and pipette ups and downs to gently dissociate cells. 5. Transfer the cell suspension into a centrifuge tube, spin down at 250  g for 3 min, and remove the supernatant. 6. Resuspend the cell pellet with prewarmed Stem Cell Culture Media by gently pipetting ups and downs. 7. Passage the cells at 1:5 or 1:8 ratios in coated flasks.

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3.3 EB Differentiation

1. Prewarm the EB Differentiation Media in a 37  C water batch. 2. Follow steps 1–5 from Subheading 3.2 to obtain mESC pellet with EB Differentiation Media. 3. Resuspend the cell pellet with prewarmed EB Differentiation Media by gently pipetting ups and downs. 4. Count cells using a cell counter. Then seed 2  106 of them onto a 9-cm low-attachment dish containing EB Differentiation Media. 5. After 1 day (day 1), transfer EB suspension into a centrifuge tube, spin down at 250  g for 3 min, and remove the supernatant. 6. Resuspend the EBs gently with prewarmed EB Differentiation Media by gently pipetting ups and downs. 7. Transfer 1/10 EB suspension to a 9-cm low-attachment dish containing EB Differentiation Media. 8. Culture the EBs to the desired time points and renew the media every other day.

3.4 Prepare SingleCell Suspension

1. Transfer EB suspension at the desired time points to a centrifugation tube. Let EBs settle down to the bottom of the tube by gravity, and carefully remove the supernatant without disturbing EBs. 2. Add DPBS to wash EBs, let them settle for 1 min, and carefully remove the supernatant. 3. Add 0.05% trypsin to EBs and incubate at room temperature for 5 min. Carefully remove the supernatant. 4. Add EB Differentiation Media to neutralize the leftover trypsin. Let EBs settle for 1 min, and carefully remove the supernatant. 5. Add FACS buffer to EBs and dissociate the cells by gently pipetting ups and downs. 6. Pass the cell suspension through a 30 μm filter to get single-cell suspension, and count cells using a cell counter. 7. Adjust cell concentration to 1  106/mL using FACS buffer and transfer the cell suspension to an FACS tube. 8. Add DAPI at a final concentration of 1 μg/mL to the cell suspension, leave the tube on ice and proceed to FACS.

3.5 Single-Cell Sorting

1. Dilute the 100 μM Barcoded RT Primer stock into 2 μM concentration in a 384-well plate with ddH2O. Seal the plate and it can be stored in 80  C for up to 6 months (see Note 1). 2. Prepare the lysis plate: this can be done by preparing enough Lysis Buffer, and aliquot 2 μL into each well of a 384-well

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Fig. 2 A picture showing an example of adjusting the FACS machine. Note all the liquid drops, containing 30 single cells, are around the center of those wells (red rectangles)

plate. Then add 1 μL Barcoded RT Primer (2 μM) to each well to reach a final volume of 3 μL per well. This lysis plate can be stored in 80  C for up to 6 months (see Note 2). 3. On the day of the experiment, take the lysis plate out of the 80  C freezer, and thaw on ice. 4. Adjust the FACS machine setting and nozzle position: put an empty plate with seal in the FACS machine, and sort 20–30 single cells per well to the wells A1–4, A21–24, H1–4, H21–24, P1–4, P21–24. Check the drop on the seal, and adjust the nozzle position such that all drops are in the center of those wells (Fig. 2) (see Note 3). 5. After the nozzle adjustment, sort DAPI negative live single cells (Fig. 3) (see Note 3 for detailed comments) into all the wells (one cell per well), except the well P24 which is used as a negative control. 6. Seal the plate and immediately spin down the plate using a plate centrifuge at 1000  g for 1 min. The plate can be stored in 80  C for up to 3 months (see Note 4). 3.6 Single-Cell cDNA Amplification

1. Take the lysis plate with sorted cells out of 80  C freezer and thaw on ice. Briefly spin down to collect any evaporation. Incubate on a thermocycler at 72  C for 10 min followed by 4  C hold, with the lid temperature set to 105  C. 2. Prepare enough Reverse Transcription Mix (4), remove the seal and add 1 μL to each well. Seal the plate again, and briefly spin down the plate.

Tube: 20210310EB

(⫻ 1,000)

250 200 150

(⫻ 1,000)

250 200 150

100

100

50

50

50

FSC-A

#Events %Parent %Total 10,000 #### 100.0 7,309 73.1 73.1 6,677 91.4 66.8 6,440 96.5 64.4

FSC-W

SSC-A

(⫻ 1,000)

Population All Events P1 P2 P3

150 100 FSC-A

200 250 (⫻ 1,000)

50

100 150 FSC-A

200 250 (⫻ 1,000)

250

200

150

100

50

102

103

DAPI-A

104

105

300

Count

250 200 150 100 50 0 102

103

DAPI-A

104

105

Fig. 3 FACS results of single cells from day 4 mouse EBs. First, FSC-A vs SSC-A was used to gate out the cell debris. Only intact cells (P1) were retained. Within P1, FSC-A vs FSC-W was used to gate out the cell multiplets. Only singlets (P2) were retained. Within P2, DAPI stain was used to gate out dead cells. Only DAPI negative live cells (P3) were sorted into the wells

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3. Reverse transcription and template switching: put the plate on a thermocycler with the following program, with the lid temperature set to 105  C. Temperature ( C)

Duration

Cycle number

42

90 min

1

50 42

2 min 2 min

10

85

5 min

1

4. Prepare enough cDNA Amplification Mix, remove the seal and add 6 μL to each well. Seal the plate again, and briefly spin down the plate. 5. cDNA amplification: put the plate on a thermocycler with the following program, with the lid temperature set to 105  C. Temperature ( C)

Duration

Cycle number

98

5 min

1

98 65 72

20 s 30 s 4 min

16

72

5 min

1

10

Hold

N/A

6. The plate can be stored in 20  C for up to a week. 3.7 Single-Cell Pooling, cDNA Purification, and Quality Check

1. Take the plates out of the 20  C freezer, thaw on ice, and pool all reactions from a plate into one 50-ml tube. This can be done by using either a multichannel pipette (Fig. 4a) or a plate reservoir (e.g., Clickbio, cat. no. VBLOK200) (Fig. 4b). Each plate should be done separately, and do not mix wells from different plates at this stage (see Note 5). 2. Add 5 volumes of Buffer PB (Qiagen) to the pooled reaction, and mix by inverting the tube until the solution becomes homogeneous. A full 384-well plate normally yield a total of around 3.8 mL reaction. Therefore, 19 mL Buffer PB should be added. 3. Pass each plate pool through a single column from the QIAquick PCR Purification kit (Qiagen, cat. no. 28104) by putting a 20 mL Extender Tube (e.g., Angen Biotech, cat. no. D50071) or a 50 mL tube (with a puncture at the bottom) (Fig. 5a) on top of the column and connecting the column to a vacuum (Fig. 5b). 4. Pass 25 mL column wash buffer to wash the column.

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Fig. 4 Demonstration of the plate pooling procedure. (a) Plate pooling using a multichannel pipette. (b) Plate pooling using a plate reservoir

Fig. 5 Demonstration of the purification of large volume of cDNA using a single column. (a) Examples of the extender tube and home-made 50-mL tube with a puncture at the bottom. (b) An example showing how to connect the extender tube or 50-mL tube with the column and vacuum

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5. Take the column off the vacuum, put on to a 2 mL collection tube and centrifuge on a table top centrifuge at top speed for 1 min to remove trace of ethanol on the column. 6. Add 50 μL 10 mM Tris–HCl, pH 8.5 to the center of the column, leave at room temperature for 1 min. 7. Put the column on a 1.5 mL Eppendorf tube and centrifuge on a table top centrifuge at top speed for 1 min to elute the cDNA. This will yield about 45 μL purified cDNA and can be stored in 80  C indefinitely. 8. Use Exo I to remove the excess of primers by assembling the following reaction. Stock

Amount

Final concentration

10 NEB 3.1 Buffer

5 μL

1

Exonuclease I (NEB, cat. no. M0568)

2.5 μL

1 U/μL

Purified cDNA from above

42.5 μL

N/A

Total

50 μL

N/A

9. Incubate at 37  C for 30 min to digest the primer left-over, and then at 80  C for 2 min to inactivate the Exonuclease I (see Note 6). 10. Purify the full-length cDNA using SPRI beads (see Note 7) by adding 35 μL (0.7 volumes) to the Exo I digested reaction and mix well by pipetting ups and downs. 11. Leave at room temperature for 5 min, and put the tube on a magnetic stand. 12. Wait until the supernatant becomes clear, which usually takes a few minutes, and remove the supernatant. 13. While the tube is on the magnet, add 200 μL 80% ethanol to the beads and wait for 20 s. 14. Carefully remove the ethanol without disturbing the beads. 15. Repeat the ethanol wash for a total of three washes. 16. Let the beads air-dry until there is no shiny reflections of liquid on the surface of the beads. 17. Remove the tube from the magnet, and resuspend the beads in 20.5 μL 10 mM Tris–HCl, pH 8.5. 18. Leave at room temperature for 2 min, and then put the tube on a magnetic stand. 19. Wait until the supernatant becomes clear, and transfer 20 μL supernatant to a new 1.5 mL Eppendorf tube. 20. Measure the concentration of the cDNA using a Qubit dsDNA HS Assay Kits according to the manufacturer’s instructions. A

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Successful cDNA (low yield)

Successful cDNA

Failed cDNA (no amplification)

Failed cDNA (RNA degradation) 31.08 26.16

RFU

21.24

16.32

11.40

6.48 5k 3k 2k

1k 700 500 400

300

200

100

20

Length(bp)

Fig. 6 Examples of successful and failed cDNA profiles

typical concentration is 20–40 ng/μL from a full 384-well plate of cells of EBs (see Note 8). 21. If needed, dilute the cDNA to a concentration of 1–10 ng/μL. Run 1 μL of cDNA on a Agilent Bioanalyzer to check the integrity of the cDNA (Fig. 6). In Fig. 6, examples of both successful and failed cDNA profiles are shown. See Note 9 for detailed comments. 3.8 Sequencing Library Preparation and Quality Check

1. Perform tagmentation of full-length cDNA by assembling the following reaction in a 1.5 mL Eppendorf tube (see Note 10). Component

Amount

Final concentration

Purified full-length cDNA X μL (50 ng total cDNA) 10 ng/μL Tn5 transposase

1 μL

N/A

2 TD Buffer

25 μL

1

ddH2O

24–X μL

N/A

Total

50 μL

N/A

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2. Incubate the reaction at 55  C for 5 min. 3. Stop the tagmentation reaction by adding 12.5 μL 0.2% SDS, mix by pipetting ups and downs and incubate at 55  C for 5 min. 4. Purify the cDNA after tagmentation by adding 50 μL (0.8 volumes) SPRI beads to the tagmentation reaction and mix well by pipetting ups and downs. 5. Follow steps 11–19 from Subheading 3.7 to purify the tagmented cDNA. This yields 20 μL purified tagmented cDNA. 6. Prepare the sequencing library by PCR amplification by assembling the reaction as follows (see Note 11). Component

Amount

Final concentration

Purified tagmented cDNA

20 μL

N/A

Library Universal PCR Primer (10 μM)

2.5 μL

0.5 μM

Library Index PCR Primer (10 μM)

2.5 μL

0.5 μM

KAPA HiFi HotStart ReadyMix (2) (KAPA, cat. no. KK2601)

25 μL

1

Total

50 μL

N/A

7. Amplify using the following PCR program. Temperature ( C)

Duration

Cycle number

72

5 min

1

98

2 min

1

98 65 72

10 s 30 s 20 s

8

10

Hold

N/A

8. Purify the library after PCR by adding 40 μL (0.8 volumes) SPRI beads the reaction and mix well by pipetting ups and downs. 9. Follow steps 11–19 from Subheading 3.7 to purify the library. This yields 20 μL purified library per plate. 10. Run 1 μL on an Agilent Bioanalzyer to see the size distribution of the library (Fig. 7) (see Note 12). 11. Send the library for pair-end sequencing with the following setting (see Note 13):

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**

Fig. 7 An example of a successful sequencing library. Asterisks indicate primer leftover, which needs to be removed by another round of SPRI beads purification before sequencing

3.9 Data Processing and Quality Control

Read

Cycle number (read length)

Identity of the read

Read 1

At least 18 cycles (18 bp)

Well barcode and UMIs

Index 1

8 cycles (8 bp)

Plate barcode

Index 2

8 cycles (8 bp)

N/A

Read 2

At least 50 cycles (50 bp)

cDNA reads

1. Once the sequencing is done, two fastq files are returned per plate. In this example, single cells from EBs at day 4 of differentiation were profiled. For example, in this protocol, the file names associated with the experiments are “mEB_day4_r1.fq. gz” (Read 1 file) and “mEB_day4_r2.fq.gz” (Read 2 file). 2. STARsolo [34] can be used to process the data to get the gene expression matrix, containing the UMI count for each gene in each cell (see Note 14). 3. Build the genome index with STAR by running the following command (see Note 15): STAR --runThreadN 20 --runMode genomeGenerate --genomeDir --genomeFastaFiles --sjdbGTFfile

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4. Get the count matrix by running the following command (see Note 16): STAR --genomeDir --readFilesCommand zcat --readFilesIn mEB_day4_r2.fq.gz mEB_day4_r1.fq.gz --soloCBstart 1 --soloCBlen 8 --soloUMIstart 9 --soloUMIlen 10 --soloType CB_UMI_Simple --soloCBwhitelist whitelist.csv --runThreadN 20

--outSAMattributes

CB

UB

--outSAMtype

BAM

SortedBy

Coordinate

5. Once the program finishes, a BAM file called “Aligned.sortedByCoord.out.bam” and a directory named “Solo.out” should appear (see Note 17). The gene expression matrix is inside the “Solo.out” directory and can be easily analysed by other scRNA-seq analysis package, such as Seurat [35] and Scanpy [36]. 6. Plot the some quality control related metrics (Fig. 8a, b), perform dimensionality reduction using UMAP [37] (Fig. 8c), carry out cell clustering using the Leiden algorithm [38] (Fig. 8c) and identify marker genes of each cluster. Markers of three germ layers, such as Cdx2, Gata2 and Gata4, should start expressing (Fig. 8d). See Note 18 for a detailed comments.

4

Notes 1. We find slow evaporation still happens in 80  C, possibly due to the opening and closing of the freezer door. Make sure to centrifuge the plate to collect evaporation every time before use. When removing the seal, be very careful to avoid splash and cross-well contaminations. 2. We routinely use 384-well plates as units for experiments, but 96-well plates also work. The lysis plates can be prepared in bulk in advance and stored in 80  C. This saves time on the experimental day. For a pilot experiment to test the protocol, you do not need to perform a full plate. 12 wells should be more than enough to test if the experiments work or not. 3. It is very important to make sure the drops are all in the centre of those 24 wells. During the actual sorting, we use FSC-A vs SSC-A to gate out the cell debris, and use FSC-A vs FSC-W to remove doublets. Finally, we only sort DAPI negative live single cells (gate P3 in Fig. 3). In most places, FACS machines are operated by specialists. Check with your local FACS experts for the gating strategy and sorting accuracy adjustment. 4. At this stage, the cells are in lysis buffer. Although it is safe to store the plate in 80  C, we still recommend proceed to the

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d Row scaled expression

b

c

Fig. 8 Results of single cells from day 4 EBs from the sequencing data. (a) The number of UMIs (in log scale) in each well on a plate. Note there are very few UMIs in the empty well (P24). (b) Scatter plot showing the number of UMIs (log) and the number of detected genes per well. Note the empty wells and a few failed single cells have orders of magnitude difference in terms of UMIs comparing to the majority of single cells. (c) UMAP representation of each single cell. The cells are colored by the Leiden cluster. (d) Row scaled expression of top 10 marker genes in each cluster

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next stage as soon as possible. When you have multiple plates for sorting, put the plates with cells on dry ice while waiting for other plates to finish sorting. Put all plates together into 80  C when all of them are done sorting. 5. At this stage, the 30 end of the cDNA is tagged by the well barcodes, but the plate barcode has not been incorporated yet. Therefore, you can only pool wells from the same plate. Different plate pools should be kept separately. 6. This step is critical, as primer leftover results in the mispriming of oligos from different wells in the subsequent PCR reaction. Digestion of the excess of primers eliminate index swapping in the final library amplification. 7. There are quite a few choices of the beads, and we have successful experience with AmpureXP beads and VAHTS DNA clean beads. 8. With mESCs, we normally get hundreds of nanograms of cDNA, which is more than enough to make the final library. Cells with low RNA content might yield less cDNA. When the total cDNA is less than 10 ng, a few more PCR cycles (step 5 from Subheading 3.6) can be done using the cDNA PCR Fwd/Rev Primers. If the amount of cDNA is between 10 ng and 50 ng, the tagmentation reaction can be proportionally scaled down in the step 1 from Subheading 3.8. 9. In the profile from the Bioanalyzer, the main peak of cDNA should be above 1000 bp. The two successful examples shown in Fig. 6 represent typical cDNA size profiles. The exact amount of cDNA varies across cell types and experiments, but intact cDNA should always have a major peak larger than 1000 bp. There should be nearly flat or very few peaks with low intensity below 1000 bp. No visible peaks means failure of cDNA amplification. Many small peaks below 1000 bp indicates RNA degradation. Two examples of failed ones are shown in Fig. 6. 10. This is an important step where cDNA is “tagmented,” where the Tn5 transposases cut the cDNA and paste the partial sequencing adaptors to cDNA. The key component here is the Tn5 transposase. There are a few choices for the Tn5 Transposase. It can be purchased from Illumina (Illumina Tagment DNA TDE1 Enzyme and Buffer Kit, cat. no. 20034197). You can also get it from other vendors, such as Fapon (cat. no. NK001) or Vazyme (TD-501). One can also makes the Tn5 transposase in house following the procedure described by Picelli et al. [39]. 11. The Library Index PCR Primer is basically Illumina’s Nextera XT Index PCR Primer. This introduces plate barcode at the right-hand side of the library. Use different primers for

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different plates if you intend to pool multiple plates to sequence together. 12. A typical library profile should be a smooth bell shaped curve with the peak around 300–400 bp. Small sharp peak at 200 bp indicates over-tagmentation, and large peak above 700 bp suggest under-tagmentation. The most efficient way of solving the problem is to adjust the amount of Tn5 accordingly. In general, we found under-tagmentation is not a big problem and it still produces very successful sequencing results. 13. In most places, sequencing is done in a genomic core facility. Talk to your local sequencing specialist for the library requirement. The exact sequencing mode will depend on the machine and whether you are sequencing alone or together with libraries from others. In general, the first read (Read 1) contain 8 bp well barcode and 10 bp UMI at the beginning. Therefore, you need at least 18 cycles for Read 1. Index 1 is the plate index that has 8 bp in length. You need 8 cycles here. Index 2 is optional in the current protocol due to the absence of an index at the left-hand side. Read 2 is the actual mRNA read, at least 50 bp are needed, and we normally perform 75 cycles (75 bp) for Read 2. 14. To convert the fastq files to gene expression matrix, a flexible program that allows you to specify the position of barcodes and UMIs is very important. Both STARsolo [34] and kallistobustools [40] can do this. In this protocol, STARsolo is demonstrated as it provides the alignment files which might be useful for other purposes. 15. The command described here should be in one single line. The files and can be obtained from the UCSC genome browser [41] and GENCODE [42], respectively. The number of CPUs to run the job can be specified by the “--runThreadN” option. 16. The command described here should be in one single line. After the “--readFilesIn” flag, two fastq files should be provided. The first fastq file should contain the mRNA reads, and the second fastq file the barcode and UMIs. In our method, Read 1 contains the barcode and UMIs, and Read 2 the mRNA reads. Therefore, in the example, Read 2 file “mEB_day4_r2.fq.gz” appears before the Read 1 file “mEB_day4_r1.fq.gz”. The “whitelist.csv” file is a simple text file which basically contains the 8-bp well barcode in the Barcoded RT Primers, one barcode per line. See Table 1 for the sequence information. 17. In the “Solo.out” directory, there are a few text files that contain some basic quality control information. The files inside “Gene/raw” and “Gene/filtered” contain expression values of

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each gene in each cell. The files in the “Gene/raw” directory contain the gene expression of every barcode in the whitelist. The files in the “Gene/filtered” contain similar information with barcodes that have too few reads removed. 18. Simple plots of some basic metrics, such as number total UMIs and number detected genes, like shown in Fig. 8a, b are useful to check if the technique works or not. When plotting the total number of UMIs in a 384-well plate layout (Fig. 8a), the empty well at the bottom right (P24) should have extremely faint colour comparing to other wells. If pipetting or sorting error happens, clear patterns will be visible; for example, some rows or columns have very few UMIs. When draw a scatter plot like shown in Fig. 8b, the empty well should have orders of magnitude fewer reads comparing to the cells. Failed cells also have very few UMIs.

Acknowledgments We thank all members from the Chen lab and the Zhang lab for the helpful comments on the manuscript. The work was supported by Shenzhen Science And Technology Innovation Committee (ZDSYS20200811144002008) and National Natural Science Foundation of China (31970812). The computational work was supported by Center for Computational Science and Engineering at Southern University of Science and Technology. References 1. Smith AG (2001) Embryo-derived stem cells: of mice and men. Cell Dev Biol 17:435–462. https://doi.org/10.1146/annurev.cellbio.17. 1.435 2. Evans MJ, Kaufman MH (1981) Establishment in culture of pluripotential cells from mouse embryos. Nature 292:154–156. https://doi.org/10.1038/292154a0 3. Martin GR (1981) Isolation of a pluripotent cell line from early mouse embryos cultured in medium conditioned by teratocarcinoma stem cells. Proc Natl Acad Sci U S A 78:7634–7638. https://doi.org/10.1073/pnas.78.12.7634 4. Ying Q-L, Wray J, Nichols J et al (2008) The ground state of embryonic stem cell selfrenewal. Nature 453:519–523. https://doi. org/10.1038/nature06968 5. Doetschman TC, Eistetter H, Katz M et al (1985) The in vitro development of blastocyst-derived embryonic stem cell lines: formation of visceral yolk sac, blood islands and myocardium. J Embryol Exp Morph

87:27–45. https://doi.org/10.1242/dev.87. 1.27 6. Chen X, Teichmann SA, Meyer KB (2018) From tissues to cell types and back: single-cell gene expression analysis of tissue architecture. Annu Rev Biomed Data Sci 1:1–23. https:// doi.org/10.1146/annurev-biodatasci080917-013452 7. Clark SJ, Lee HJ, Smallwood SA et al (2016) Single-cell epigenomics: powerful new methods for understanding gene regulation and cell identity. Genome Biol 17:72. https://doi. org/10.1186/s13059-016-0944-x 8. Wang Y, Navin NE (2015) Advances and applications of single-cell sequencing technologies. Mol Cell 58:598–609. https://doi.org/10. 1016/j.molcel.2015.05.005 9. Tang F, Barbacioru C, Wang Y et al (2009) mRNA-seq whole-transcriptome analysis of a single cell. Nat Methods 6:377–382. https:// doi.org/10.1038/nmeth.1315

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Osteogenic Differentiation from Mouse Embryonic Stem Cells Zahra Alvandi and Michal Opas Abstract Embryonic stem cells (ESCs) are a unique model that allows the study of molecular pathways underlying commitment and differentiation. We have studied signaling pathways and their contributions to osteogenic differentiation. In addition to our previously published protocol where we recommended the addition of retinoic acid with later addition of dexamethasone to boost osteogenic lineage cells, here we describe an optimized protocol for osteogenic differentiation from R1 ESCs with suggestions for inhibition of Src activity. Key words Embryonic stem cells, Osteogenic differentiation, Embryoid bodies, c-Src kinase

1

Introduction ESCs can be differentiated to produce bone cell lineages through the combination of defined media and the addition of bioactive signaling molecules [1–3]. Differentiation of osteoblasts from mesenchymal osteo-progenitors progresses through a sequence of steps comprising formation of immature and mature osteoprogenitors, preosteoblasts, mature osteoblasts, and osteocytes and involves proliferation, maturation, extracellular matrix (ECM) development, and mineralization, which are linked to variable gene expression of osteogenic markers [4–7]. Src family kinases play crucial roles in regulating cell differentiation [8]. c-Src has been shown to decrease osteoblast differentiation in vitro [9–11]. Inhibition of c-Src activity in adult mice increases bone mass at least in part by stimulating osteoblast differentiation [12] . Using a hanging dropbased protocol to induce embryoid bodies (EBs), we recently showed the effect of Src kinase inhibition for eight different time periods during osteogenic differentiation [13]. EBs are 3-dimensional aggregates of differentiating stem cells providing a unique model in which to study early embryonic development. There are many different ways one can generate EBs overviewed by Kurosaw [14] and in our hands most efficacious was a hanging drop method [15]. Our results showed that inhibition of Src kinase

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activity starting from day 6 of osteogenic differentiation for 4 days significantly improved mineralization and the expression of osteocalcin at the end of the 21-day differentiation protocol. Therefore, the inhibition of Src activity for the indicated time has been incorporated in this protocol.

2

Materials 1. Maintaining media: Dulbecco’s modification eagle’s medium 1 (DMEM, Catalog No. 319-005-ES) supplemented with 10% FBS (FBS, Premium, Catalog No. 088150), 1% MEM nonessential amino acids (MEM NEAA 100, Gibco, Catalog No. 11140050), 0.1 mM 2-mercaptoethanol (Bioshop, Catalog No. MER 002), and 10 ng/ml leukemia inhibitory factor (LIF) (see Note 1). 2. L-ascorbic acid (Sigma, catalog No. A5960) (see Note 2). 3. β-glycerophosphate disodium salt hydrate (Sigma, catalog No. G9422) (see Note 3). 4. Retinoic acid (Sigma, Catalog No. R2625) (see Note 4). 5. PP2 (Calbiochem, Catalog No. 529573) (see Note 4). 6. Dexamethasone (Sigma, Catalog No. D4902) (see Note 4).

3

Methods 1. Day 0–3: Mouse ES cells R1 derived from J1 129/Sv mice can be maintained in their undifferentiated status on mitomycin C-treated mouse embryonic fibroblast feeder cells on gelatincoated plates in maintaining media. At passage 2, trypsinize and collect the cells for hanging drops. Drops containing 250 cells per 25 μL DMEM should be supplemented with 20% FBS and be placed on the lids of tissue culture dishes for 3 days to form EBs. 2. Day 3–5: After 3 days, transfer EBs into the floating cell culture dishes containing medium supplemented with 0.1 μM retinoic acid for 2 days. On day 5, collect EBs and plate them in tissue culture dishes coated with 0.1% gelatin. 3. Day 6–10: On day 6, the differentiation medium has to be supplemented with 50 μg/mL L-ascorbic acid and 10 mM β-glycerophosphate disodium salt hydrate to promote osteogenic differentiation. 10 μM Src inhibitor PP2 may be added to the culture to enhance osteogenic differentiation. 4. Day 10–21: Add 100 nM dexamethasone on day 10 to further enrich cells of the osteogenic lineage. The medium must be

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Fig. 1 Scheme represents the 21-day osteogenic differentiation protocol with the von Kossa staining at the day 21 that evaluates the mineralization level

replaced with freshly made media every 2 days for the entire 21 days of differentiation. Evaluate the mineralization of the osteonodules and expression level of osteogenic markers on day 21. A scheme of the suggested protocol is presented in Fig. 1.

4

Notes 1. Media are made completely antibiotic free. 2. L-ascorbic acid is heat and light sensitive. Prepare fresh stock before use. 3. β-glycerophosphate disodium salt hydrate can be dissolved in warm ddH2O. 4. Retinoic acid, dexamethasone, and PP2 may be stored as recommended by the manufacturers. We highly recommend preparing small aliquots for storage to avoid multiple freeze and thaw cycles.

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Acknowledgement This work was supported by Canadian Institutes of Health Research (CIHR) grants MOP-130551 and MOP-106461 to M.O. References 1. Pilquil C, Alvandi Z, Opas M (2020) Calreticulin regulates a switch between osteoblast and chondrocyte lineages derived from murine embryonic stem cells. J Biol Chem 295 (20):6861–6875 2. Yu Y, Al-Mansoori L, Opas M (2015) Optimized osteogenic differentiation protocol from R1 mouse embryonic stem cells in vitro. Differentiation 89(1–2):1–10 3. Yu Y, Pilquil C, Opas M (2016) Osteogenic differentiation from embryonic stem cells. Methods Mol Biol 1341:425–435 4. Aubin JE (1998) Advances in the osteoblast lineage. Biochem Cell Biol 76(6):899–910 5. Roeder E, Matthews BG, Kalajzic I (2016) Visual reporters for study of the osteoblast lineage. Bone 92:189–195 6. Jensen ED, Gopalakrishnan R, Westendorf JJ (2010) Regulation of gene expression in osteoblasts. Biofactors 36(1):25–32 7. Rutkovskiy A, Stensløkken KO, Vaage IJ (2016) Osteoblast differentiation at a glance. Med Sci Monit Basic Res 22:95–106 8. Parsons SJ, Parsons JT (2004) Src family kinases, key regulators of signal transduction. Oncogene 23(48):7906–7909

9. Marzia M et al (2000) Decreased c-Src expression enhances osteoblast differentiation and bone formation. J Cell Biol 151(2):311–320 10. Peruzzi B et al (2012) c-Src and IL-6 inhibit osteoblast differentiation and integrate IGFBP5 signalling. Nat Commun 3:630 11. Alvandi Z, Al-Mansoori LJR, Opas M (2020) Calreticulin regulates Src kinase in osteogenic differentiation from embryonic stem cells. Stem Cell Res 48:101972 12. Thouverey C, Ferrari S, Caverzasio J (2018) Selective inhibition of Src family kinases by SU6656 increases bone mass by uncoupling bone formation from resorption in mice. Bone 113:95–104 13. Alvandi Z, Opas M (2020) c-Src kinase inhibits osteogenic differentiation via enhancing STAT1 stability. PLoS One 15(11):e0241646 14. Kurosawa H (2007) Methods for inducing embryoid body formation: in vitro differentiation system of embryonic stem cells. J Biosci Bioeng 103(5):389–398 15. Kurosawa H et al (2003) A simple method for forming embryoid body from mouse embryonic stem cells. J Biosci Bioeng 96(4):409–411

Methods in Molecular Biology (2022) 2520: 265–273 DOI 10.1007/7651_2021_447 © Springer Science+Business Media, LLC 2021 Published online: 02 November 2021

Murine Embryonic Stem Cell Culture, Self-Renewal, and Differentiation Manar Elkenani and Belal A. Mohamed Abstract Embryonic stem cells (ESCs), derived from the inner cell mass of the blastocyst, can proliferate indefinitely in vitro (self-renewal) and can differentiate into cells of all three germ layers (pluripotency). These unique properties make them exceptionally valuable in basic science and clinical researches, including cell replacement therapies, drug discovery, and regenerative medicine. Mouse ESCs represent an important model system for studying gene function during development and disease. ESCs culture is time-consuming, laborious, and costly. Suboptimal ESCs culture conditions can alter their identity, pluripotency, and their compatibility with downstream differentiation protocols. In this chapter, we provide a general guideline for murine ESCs culture on murine fibroblast feeder layers. Moreover, we describe protocols for maintenance of ESCs pluripotency and induction of ESCs differentiation. Key words Murine embryonic stem cells, Cell culture, Self-renewal, Differentiation, Knockout serum replacement, Leukemia inhibitory factor

1

Introduction Embryonic stem cells (ESCs) are cells derived from the inner cell mass of the preimplantation blastocyst-stage embryo [1, 2]. The ESCs are characterized by their capacity for self-renewal and their pluripotency allowing them to differentiate into all derivatives of the primary germ layers, including ectoderm, endoderm, and mesoderm under certain conditions. Several key intrinsic factors such as Oct-4 and Nanog [3–5] and extrinsic factors like leukemia inhibitory factor (LIF), bone morphogenetic protein (BMP) and wingless (Wnt)/β-catenin for maintenance of ESCs pluripotency have been identified [6–9]. Mouse ESCs represent an important model system for studying gene function during development and disease. Several gene editing technologies in ESCs, including but not limited to, gene targeting through traditional homologous recombination, forced expression through insertion of transgenes, and recently discovered zinc finger nucleases, TALENs and CRISPR strategies [10, 11], has

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extensively facilitated research across all fields and offer enormous promise in treating difficult disorders. Murine ESCs require specialized, high-quality media and expert culture techniques for propagation in the laboratory. Suboptimal ESCs culture conditions can alter their purity, potency, functional capability and therefore their compatibility with downstream differentiation protocols. The purpose of this chapter is to provide detailed protocol information on the establishment of the mouse embryonic fibroblasts (MEF) feeder layers, the culture of murine ESCs on these feeder layers, and the induction of ESCs differentiation upon LIF withdrawal. We believe that these basic protocols will be helpful for the researches in the field of murine ESCs.

2 2.1

Materials Equipment

1. Biosafety cabinet (BSC). 2. Centrifuge. 3. Microscope. 4. 37
C–5% CO2 incubator. 5. 37
C water bath. 6. Cryogenic handling gloves and eye protection. 7. 15 cm petri dish. 8. Sterile fine scissors and scalpel blades. 9. 1.8 mL cryogenic vials. 10. Culture plates. 11. T-75 cell culture flasks. 12. 5 mL sterile serological pipettes and 10 mL sterile serological pipettes or equivalent. 13. 2 mL aspirating pipette. 14. 15 and 50 mL conical plastic tube. 15. 70% ethanol.

2.2 FeederDependent ESCs Culture

1. Gelatin solution: 0.1% gelatin in phosphate buffered saline (PBS) (see Note 1) stored at 4
C. 2. Mitomycin C-treated MEF feeder layers: Final concentration 10 μg/mL (see Note 2). 3. MEF culture medium: Dulbecco’s modified Eagle’s medium (DMEM) medium, 10% fetal bovine serum (FBS), 2 mM Lglutamine, 1% penicillin–streptomycin. 4. ESC culture medium: DMEM, 20% FBS, 1 mM nonessential amino acids (NEAA), 1 mM sodium pyruvate, 10 μM

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2-mercaptoethanol, 2 mM L-glutamine, 1000 U/mL LIF, 1 mM penicillin–streptomycin. 5. ESC knockout culture medium: DMEM, 20% knockout™ serum replacement, 1 mM NEAA, 1 mM sodium pyruvate, 10 μM 2-mercaptoethanol, 2 mM L-glutamine, 1 mM penicillin–streptomycin. 2.3

ESCs Passaging

1. PBS without CaCl2 or MgCl2. 2. Trypsin: 0.25% trypsin solution.

2.4

3

ESCs Freezing

Cryopreservation medium: 20% FBS, 10% DMSO, 65% MEF culture medium (see Note 3).

Methods Carry out all procedures under aseptic conditions except as otherwise provided. Before start, sterilize the BSC with 70% ethanol and use ultraviolet light for 15 min as a secondary decontaminant. Run the BSC for at least 10 min before working. Prewarm to 37
C any solutions/medium needed before use.

3.1 Isolation and Expansion of MEF

1. Before start coat T-75 culture flasks with 0.1% gelatin. Make sure surface area of plate is covered by gelatin solution and set them in the incubator for later use (step 11) (see Table 1 for the required volume). 2. Euthanize a pregnant female(s) at E13.5–14.5 day of gestation (i.e., 13–14 days after the appearance of the copulation plug) and dissect out the uterine tract containing embryos. 3. Place them in a petri dish containing PBS (see Note 4). 4. Isolate the embryos from the embryonic sac using sterile instruments. 5. Dissect the head, remove the internal organs, and wash with PBS. 6. Mince the embryos in small pieces (1–2 mm) with scalpel and place them in 0.25% trypsin/EDTA solution (use 2 mL of trypsin/EDTA pro embryo). 7. Pipet up and down several times with a 10 mL pipet. 8. Transfer the tissue into a 50 mL falcon tube and incubate for 15 min at 37
C in a tissue culture incubator. Each 5–7 min of incubation, pipet up and down thoroughly to dissociate the cells. 9. Inactivate the trypsin-EDTA by adding equal volume of MEF medium.

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Table 1 Recommended volumes of gelatin solution Cell culture plastic ware

Amount of 0.1% gelatin

T-75 flask

7–10 mL

24 well plate

0.5 mL each well

12 well plate

1 mL each well

6 well plate

2 mL each well

10. Pass the digested tissues through a 100-μm mesh to remove particulates and to obtain single cell suspension. 11. Transfer the cell suspension onto the gelatinized T-75 flasks after aspiration of the gelatin solution. 12. Place the flasks in the incubator and allow them grow to extreme confluency. 13. Change the medium on the next day, remove the old medium (with dead cells and debris), wash twice with prewarmed PBS and add fresh MEF medium. 14. Check the cells and change the medium daily, when the cells reach 80% confluency, split the cells. 15. Use 0.25% trypsin solution for splitting. Wash the cells twice with prewarmed PBS. Add 7 mL trypsin for each flask and incubate for 30 s (see Note 5). Pipet up and down into flask to break up clumps of cells. Add 7 mL of MEF medium and then transfer medium and cells to 15 mL conical tube. 16. Centrifuge gently to pellet the cells (1000 rpm  5 min). 17. Aspirate the medium carefully without disturbing the pellet, resuspend the pellet in 10 mL warm medium and transfer into new gelatinized T-75 flasks (1:3) (i.e., split each T-75 flask into 3 T75 flasks) and allow them to grow. This is passage 1. You can repeat the splitting procedure until passage 3 (see Note 6). 3.2 Mitotic Inactivation of MEF

1. Begin with a confluent monolayer of MEF cells. 2. Wash the cells with PBS and add 10 mL of fresh MEF medium containing 100 μL of Mitomycin C stock solution for each T-75 flask (final concentration: 10 μg/mL) and place the flasks in incubator for approximately 2 h. 3. After incubation, aspirate the inactivation medium and wash the cells three times with a minimum of 10 mL of PBS. 4. Collect the cells by trypsinization and centrifugation (1000 rpm  5 min). 5. Suspend the cells with cryopreservation medium (see Subheading 2.3).

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6. Aliquot the MEFs into cryovials (2  106cells in 0.5 mL/vial) and slowly freeze the cells at 70
C and then transfer to temperatures below 130
C. 3.3 Plates Preparation for ESCs Culture (See Note 7)

1. Pretreat 24 well plate with 0.1% gelatin for 60 min. 2. Quick thaw MEF cells in vial in 37
C water bath until barely thawed (see Note 8). 3. Immediately add 1 mL of MEF medium to the vial and pipet gently up and down (see Note 9). 4. Transfer the 1.5 mL in vial to sterile 15 mL conical tube and slowly add 3.5 mL medium then centrifuge at 1000 rpm  5 min. 5. Aspirate the medium carefully without disturbing the pellet, resuspend the pellet in warm MEF medium and count the cells. 6. Plate the feeder cells at a density of about 5  104 cells/cm2 (see Note 10). 7. Replace the media the next day and thereafter every other day for maximum 5 days.

3.4 Plating ESCs and Maintenance of Pluripotency

1. Aspirate the MEF medium and pipet enough sterile PBS into the flask to wash and to get rid of any dead cells. 2. Add prewarmed fresh ESC culture medium, containing LIF to the feeder layer. 3. Thaw the ESCs vial as described in Subheading 3.3, but use the ESC medium with LIF instead of the MEF medium. 4. Plate the cells over the feeder layer into ESC medium at a density of 2  105/cm2 (see Note 11). 5. Incubate overnight at 37
C, 5% CO2 and allow cells to attach. Change medium every day for optimal growth. 6. Check growth and the degree of confluency (see Note 12).

3.5

ESCs Passaging

1. Prepare new feeder layer on a gelatinized plates as previously described. Change the medium 4 h before passaging. 2. Wash the cells with PBS and add an adequate volume 0.25% trypsin. 3. Set the plates back in the incubator for approximately 2–3 min. 4. Check the cell under the microscope and ensure the complete detachment of the cells. 5. Stop the trypsin action by adding equal volume to ESC medium with LIF. 6. Pipet gently up and down to create a single cell suspension. 7. Add the cell suspension to a sterile conical tube and centrifuge for 5 min at 1000 rpm.

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8. Resuspend the cells with ESC medium and passage them at 1:6. 9. If freezing cells, then resuspend in cell freezing medium at an appropriate concentration 2  106 per 0.5 mL of freezing medium. 10. Slowly freeze the cells at 70
C and then transfer to temperatures below 130
C until later use (see Note 13). 3.6 ESCs Differentiation

1. Plate ESCs at a low density (5  103/cm2) as described in Subheading 3.4 (see Note 14). 2. Allow the cells to attach (6 h after plating). 3. Aspirate the ESC medium carefully without disturbing the cells. 4. Rinse the cells with warm PBS. 5. Remove PBS and add adequate volume of ESC knockout culture medium without LIF (see Note 15). This day is considered as differentiation day 0. 6. Change the medium daily and observe the cell morphology (see Note 16).

4

Notes 1. Prepare 1% gelatin stock in distilled water (5 g gelatin into 500 mL distilled water). Autoclave and store at 4
C. For the working solution (0.1%), take 50 mL of the stock solution and add 450 mL of autoclaved distilled water and store at 4
C. Coat the cell culture plastics with gelatin working solution 0.1% with the adequate volume (Table 1) then incubate for at least 60 min or overnight at 37
C in the incubator. Shortly before culturing the cells, aspirate the gelatin solution and then add the cells and media to the coated plates or flasks. 2. Mitomycin C inhibits proliferation of the MEF in long-term culture assays so that they can be used as nonreplicating viable supportive cells. Mitotically inactive mouse embryonic fibroblasts (MEFs) secrete necessary nutrients and factors that help the stem cells to maintain the pluripotency in the presence of LIF. Additionally, feeder cells can synthetize extracellular matrix proteins that act as a substrate for the attachment of cultured cells. For stock solution preparation, dissolve 2 mg of Mitomycin C powder into 2 mL PBS/ medium (1 mg/mL). 3. Aliquot the cryopreservation medium (0.5 mL/tube) and store at 20
C up to 6 months. 4. Steps 2 and 3 are performed under nonaseptic conditions.

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5. Trypsin is harmful to the cells. Pay special attention that cells are not in trypsin longer than 30 s. 6. To obtain healthy MEFs, you should carefully monitor their growth and morphology. Because they are primary cells, they have a limited life span in culture (1–3 passages). Overgrowth of the culture results in early senescence. 7. Plate the feeder at least 1 day in advance of thawing or splitting the ESCs. 8. DMSO in freezing medium will damage cells, so do these steps quickly. 9. Adding fresh medium to the thawed cells dilutes DMSO and protect the cells. 10. Plating density of the feeder layer is important as it can profoundly affect the growth of ESCs. Too sparse feeder cell density would result in ESCs differentiation. On the other hand, too dense feeder layer may result in rapid depletion of nutrients and oxygen as well as physically hinder the attachment and growth of ESCs colonies. 11. After plating the cells, limit opening and closing the incubator doors to prevent disturbing the even distribution of cells to the surface of the well and to allow the cells to attach. 12. If medium is yellow or tinged yellow-change medium immediately to bring pH back to neutral. You should split the ESCs as they reach 70–80% of confluency. Overconfluent cultures increases the chances for differentiation (presence of large and flat cells between the adjacent colonies), and induces terminal loss of pluripotency. 13. The passage number of ESCs should be monitored since late passages are associated with epigenetic and chromosomal instability. Karyotype analysis is highly recommended, particularly when the ESC lines are reaching a high passage number. 14. For differentiation, we recommend plating the ESCs on feeder layer to help ESCs attachment and in absence of LIF to induce monolayer differentiation. Please note that culturing ESCs on nonadherent dishes in a feeder-free condition and in absence of LIF will induce differentiation of ESCs but into embryoid bodies. 15. To initiate differentiation, ESCs is cultured in medium lacking LIF and any growth factors. Fetal bovine serum is replaced by knockout serum replacement in order to avoid the effect growth factors presenting in serum. 16. Culturing ESCs under differentiating conditions (knockout culture medium without LIF) leads to rapid morphological changes of ESC colonies into an epithelium-like shape

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Fig. 1 (a) Representative images showing the dome shape morphology of pluripotent ESC colony (left lane), whereas differentiated cells on the right lane exhibit an elongated-flattened morphology. Scale bar ¼ 100 μm. (b) Representative Western blotting for pluripotent markers (SOX2 and OCT4) and differentiation marker (DAB2) expression at day 4 and 8 of culture in undifferentiating (ESCs culture medium with LIF), and differentiating (ESCs Knockout culture medium without LIF) conditions. α-Tubulin was used as a loading control; KO Knockout

(Fig. 1a), which is accompanied by marked decrease in the selfrenewal markers, SOX2 and OCT4, and increase in differentiation marker, DAB2 (Fig. 1b). 17. We do not recommend passaging or freezing of the differentiated ESCs since monolayer differentiated ESCs loose their ability to attach and hence will not survive upon passaging.

Acknowledgments This work was supported by the German Research Foundation DFG grant MO 3373/1-1 to BAM. References 1. Kaufman MH, Evans MJ (1981) Establishment in culture of pluripotential cells from mouse embryos. Nature 292:154–156 2. Martin GR (1981) Isolation of a pluripotent cell line from early mouse embryos cultured in medium conditioned by teratocarcinoma stem cells. Proc Natl Acad Sci U S A 78:7634–7638. https://doi.org/10.1073/pnas.78.12.7634 3. Loh YH, Wu Q, Chew JL et al (2006) The Oct4 and Nanog transcription network regulates pluripotency in mouse embryonic stem cells. Nat Genet 38:431–440. https://doi. org/10.1038/ng1760 4. Mitsui K, Tokuzawa Y, Itoh H et al (2003) The homeoprotein nanog is required for maintenance of pluripotency in mouse epiblast and

ES cells. Cell 113:631–642. https://doi.org/ 10.1016/S0092-8674(03)00393-3 5. Niwa H, Miyazaki JI, Smith AG (2000) Quantitative expression of Oct-3/4 defines differentiation, dedifferentiation or self-renewal of ES cells. Nat Genet 24:372–376. https://doi. org/10.1038/74199 6. Smith AG, Heath JK, Donaldson DD et al (1988) Inhibition of pluripotential embryonic stem cell differentiation by. Nature 336:688–690 7. Niwa H, Burdon T, Chambers I, Smith A (1998) Self-renewal of pluripotent embryonic stem cells is mediated via activation of STAT3. Genes Dev 12:2048–2060. https://doi.org/ 10.1101/gad.12.13.2048

Murine Embryonic Stem Cells Culture 8. Nyamsuren G, Kata A, Xu X et al (2014) Pelota regulates the development of extraembryonic endoderm through activation of bone morphogenetic protein (BMP) signaling. Stem Cell Res 13:61–74. https://doi.org/10.1016/j. scr.2014.04.011 9. Elkenani M, Nyamsuren G, Toischer K et al (2021) Perturbed differentiation of murine embryonic stem cells upon Pelota deletion due to dysregulated FOXO1/β-catenin signaling. FEBS J 288:3317–3329. https://doi.org/ 10.1111/febs.15643

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10. Kim H, Kim JS (2014) A guide to genome engineering with programmable nucleases. Nat Rev Genet 15:321–334. https://doi.org/ 10.1038/nrg3686 11. Li Q, Qin Z, Wang Q et al (2019) Applications of genome editing technology in animal disease modeling and gene therapy. Comput Struct Biotechnol J 17:689–698. https://doi.org/ 10.1016/j.csbj.2019.05.006

Methods in Molecular Biology (2022) 2520: 275–294 DOI 10.1007/7651_2021_438 © Springer Science+Business Media, LLC 2021 Published online: 19 October 2021

Mouse Embryonic Stem Cell Culture in Serum-Containing or 2i Conditions Emre Balbasi, Gozde Guven, and Nihal Terzi Cizmecioglu Abstract With their unique capabilities of self-renewal and differentiation into three germ layers, mouse embryonic stem cells (mESCs) are widely used as an in vitro cellular model for early mammalian developmental studies. mESCs are traditionally cultured in high-serum and LIF-containing medium on a growth-deficient mouse embryonic fibroblast layer. A more recent culturing system with two inhibitors (for GSK3β (CHIR99021) and MEK1/2 (PD0325901)) and LIF enables the derivation of mESC lines from various mouse strains. Here we describe methods for the mESC growth and maintenance in each medium composition as well as their adaptation to either condition. Key words Mouse embryonic stem cells, mESCs, MEFs, Cell culture, 2i, 2i medium, LIF, CHIR, PD

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Introduction Mouse embryonic stem cells (mESCs) originate from the inner cell mass of blastocyst-stage embryos. Due to their self-renewal and pluripotency features, they are widely used as a model to study early mammalian development [1]. Standard mESC culturing technique relies on growing them on a layer of irradiated or mitomycintreated mouse embryonic fibroblasts (MEF) on gelatinized tissue culture dishes in high-serum and leukemia inhibitory factor (LIF) containing media. LIF sustains pluripotency by activating the JAK-STAT3 pathway [2]. A more recent mESC culturing technique uses a defined, serum-free base medium supplemented with GSK3β and MEK inhibitors (2i; CHIR99021 and PD0325901, respectively) along with LIF on gelatinized tissue culture dishes [3, 4]. The mESCs growing in 2i medium tend to stay in the naı¨ve pluripotent stage longer [5]. The components of serum are not well-defined and can have batch to batch variation, leading to spontaneous differentiation and heterogeneity of the mESCs [6, 7]. Fibroblast layer underneath mESCs is another source of variation in terms of conditioning of the medium [8]. In this chapter, we provide protocols for mESC culturing conditions for both standard high-serum containing medium on the MEF layer

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and low-serum or serum-free 2i medium, as well as adaptation of mESCs to either condition.

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Materials Gelatin from bovine skin (Cat. No.: G9391-100G, Sigma). Water, cell culture grade (Cat. No.: BI03-055-1A, Biological Industries). DMEM, high glucose, pyruvate (Cat. No.: 41966029, Gibco). Fetal Bovine Serum (FBS) (Cat. No.: 10270106, Gibco). GlutaMAX I (Cat. No.: 35050061, Gibco). Pen-Strep (Cat. No.: 15140-122, Gibco). MEM NEAA (Cat. No.: 11140-035, Gibco). β-Mercaptoethanol (Cat. No.: M-6250, Sigma). Leukemia Inhibitory Factor (LIF) (Cat. No.: ESG1107, Millipore). CHIR-99021 (Cat. No.: S2924, Selleckchem). PD0325901 (Cat. No.: S1036, Selleckchem). Adenosine (Cat. No.: A4036-5G, Sigma). Uridine (Cat. No.: U3003-5G, Sigma). Thymidine (Cat. No.: T1895-1G, Sigma). Cytidine (Cat. No.: C4654-1G, Sigma). Guanosine (Cat. No.: G5264-1G, Sigma). N-2 Supplement (100) (Cat. No.: 17502048, Gibco). B-27 Supplement (50) (Cat. No.: 17504044, Gibco). DMEM/F-12 (Cat. No.: 11320074, Gibco). Neurobasal Medium (Cat. No.: 21103049, Gibco). Bovine serum albumin (BSA) (Cat. No.: A3311-50G, Sigma). Trypsin EDTA Solution B (0.25%), EDTA (0.05%), with Phenol Red (Cat. No.: BI03-052-1B, Biological Industries). Mitomycin C lyophil. Research grade (Cat. No.: SE2980501, Serva). Nutrifreez™ D10 Cryopreservation Medium (Cat. No.: BI05713-1E, Biological Industries). Dimethyl sulfoxide for molecular biology (DMSO) (Cat. No.: D8418-50ML, Sigma). TrypLE Express Enzyme (1), phenol red (Cat. No.: 12605-010, Gibco). Accutase cell detachment solution (Cat. No.: SCR005, Millipore).

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Dulbecco’s phosphate buffered saline (DPBS) (Cat. No.: 02-0231A, Biological Industries). 1-Thioglycerol (MTG) (Cat. No.: M6145-25ML, Sigma). 2.1 Solution Preparations

0.1% Gelatin Solution: Dissolve 0.5 g gelatin in 500 mL cell culture grade water. Autoclave the solution. Shake the bottle after removing from the autoclave to ensure resuspension. Let it cool for a few hours. Store at 4  C. 10% BSA (w/v): Dissolve 5 g bovine serum albumin (BSA) in 50 mL Dulbecco’s phosphate buffered saline (DPBS). Filtersterilize and store at 4  C. Nucleoside Mix: Add 80 mg Adenosine, 85 mg guanosine, 73 mg uridine, 73 mg cytidine, and 24 mg thymidine into 100 mL distilled water and dissolve by warming to 45  C. Filter-sterilize and aliquot while warm. Store at 20  C. Mitomycin-C stock: Dissolve 2 mg Mitomycin-C powder in 10 mL DPBS (the final concentration is 200 μg/mL). Aliquot and store at 80  C.

2.2 Media Preparations

All media should be prepared in filter bottles and filtered. 2i4 Medium (Low-serum medium, 100 mL): 50 mL Neurobasal Medium, 50 mL DMEM/F-12, 500 μL N-2 Supplement (100), 1 mL B-27 Supplement (50) (see Note 1), 500 μL 10% BSA, 1 mL GlutaMAX I, 1 mL Pen/Strep, 1.3 μL MTG (1-Thioglycerol), 4 mL Fetal Bovine Serum (FBS). Supplement with final concentrations of 1000 units/mL Leukemia Inhibitory Factor (LIF), 3 μM CHIR-99021 and 1 μM PD0325901. 2i4 medium (without LIF, CHIR, and PD) can be prepared and stored at 4  C up to a month as “incomplete medium.” For long-term storage, the media can be aliquoted and stored at 20  C. LIF, and 2i (CHIR and PD) should be added freshly. Media older than a week should be resupplemented with LIF and 2i. The name 2i4 comes from the 2 inhibitors (CHIR-99021 and PD0325901) and 4% serum in it. This medium is also referred to as low-serum medium along with 2i2 and 2i1. Prepare 2i medium (2i4 medium without FBS) and dilute 2i4 medium with 2i medium to obtain 2i2 and 2i1 media. MEF Medium (100 mL): 88 mL DMEM, 10 mL fetal bovine serum (FBS), 1 mL GlutaMAX I, 1 mL Pen/Strep. MEF medium can be used up to a month when stored at 4  C. The media older than a month (when stored at 4  C) must be resupplemented with GlutaMAX I. For long term storage, the media can be aliquoted and stored at 20  C.

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Mitomycin-C Treatment Medium (100 mL): 95 mL MEF Medium, 5 mL Mitomycin-C stock solution. Final concentration of Mitomycin-C in the medium must be 10 μg/mL. ESC Medium (High-serum medium, 100 mL): 80 mL DMEM, 15.04 mL Fetal Bovine Serum (FBS), 2 mL Pen/Strep, 1 mL Nucleoside Mix, 1 mL GlutaMAX I, 1 mL MEM NEAA, 0.704 μL β-Mercaptoethanol, 10 μL Leukemia Inhibitory Factor (LIF, 107 units/mL). ESC medium should be prepared, and filter sterilized without LIF. LIF should be added freshly, just before use. Store at 4  C up to a month. For long term storage, the media can be aliquoted and stored at 20  C. After the addition of LIF, the medium should be used within a week. Media older than a week should be resupplemented with fresh LIF. 2 Freezing Medium: 40 mL Fetal Bovine Serum (FBS), 10 mL DMSO. Store 2 freezing medium at 4  C, do not freeze.

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Methods For suggested appropriate amounts of media, solutions, and cells, please refer to Table 1. All steps must be aseptically performed in a laminar flow hood.

Table 1 Required solution volumes and cell numbers for different culture dishes 1 well of 6-well plate

10 cm plate

Culture medium volume

3 mL

8 mL

Wash volume

2 mL

5 mL

Trypsin

300 μL

1.5 mL

TrypLE

150–200 μL

1–1.5 mL

Accutase

300 μL

1.5 mL

Quenching volume

3 mL

5 mL

Mitomycin-C treatment medium

2 mL

4–5 mL

De-MEFfing volume

2 mL

4–5 mL

Gelatin volume

2 mL

5 mL

mitoMEF number

5

2.5  10 cells

1.5  106 cells

mESC number

2.5–3.5  105 cells

1.5  106 cells

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3.1 mESC Growth in Low-Serum 2i4 Medium 3.1.1 Gelatin Coating of Culture Dishes

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1. Fill the culture dish with 0.1% gelatin solution. 2 mL for 1 well of a 6-well plate, 5 mL for a 10 cm plate is sufficient. 2. Incubate it in a CO2 incubator, at 37  C, 5% CO2 for at least 10 min. 3. Aspirate the gelatin solution. 4. Leave the plates in an inclined position in the laminar flow hood for residual gelatin to dry for 30 min. Aspirate again if necessary. 5. After the gelatin is completely dried out, the culture dishes can be used immediately or can be stored (stacked and covered with cling film (Saran wrap)) at room temperature for further use.

3.1.2 Thawing mESCs

1. Add 9 mL DMEM into a 15 mL falcon tube. 2. Quickly thaw the cryovial in a water bath at 37  C until a small piece of ice remains in the cryovial. Thawing mESCs slowly reduces their viability. Make sure the cap of the cryovial stays afloat to minimize risk of contamination. 3. Thoroughly spray the outside of the cryovial with 70% Ethanol (EtOH) before bringing it into the laminar flow hood. 4. With a 1000 μL micropipette, slowly add the content of the cryovial into 9 mL DMEM prepared in step 1. 5. Centrifuge the falcon tube at 300 rcf, 4  C for 5 min (see Notes 2 and 3). 6. Aspirate the supernatant (see Note 4). 7. Gently resuspend the pellet in 1 mL complete 2i4 medium. 8. Seed cells in 2i4 medium. 3 mL for 1 well of a 6-well plate and 8 mL for a 10 cm plate is sufficient. Generally, 1–2  106 cells can be thawed into one well of a 6-well plate in 3 mL 2i4 medium. Roughly, a cryovial frozen from a confluent one well of a 6-well plate can be thawed into 1 or 2 wells of a 6-well plate. Make sure you consider the duration of time cells spent frozen as well as the viability after thaw. 9. Shake the culture dish, back and forth and, left and right, virtually creating a plus (+) sign while shaking (see Note 5). 10. Incubate the cells at 37  C, 5% CO2. mESCs require passaging every 2–3 days. Morphology and the density of the colonies are the key factors to consider passaging. Pluripotent mESCs grow as round, dome shaped colonies that are uniform in size. mESC colonies look bright under the light microscope and have clear, sharp edges (Fig. 1a). mESCs should not be seeded too sparse or dense, as they start to differentiate in either case (Fig. 1b). Some differentiation is expected on the edges of the well, especially after a few days from passaging (Fig. 1c). You need

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Fig. 1 mESCs grown in low-serum 2i4 medium. (a) Healthy mESC colonies in 2i4 medium, 40 and 100 magnification. (b) Crowded mESC colonies in 2i4 medium, 40 and 100 magnification. (c) Differentiation at the edge of the well, 2i4 medium, 40 magnification. (d) Merged mESC colonies with cell death, 2i4 medium, 40 magnification

to monitor average colony size closely as larger colonies tend to have cell death in the middle or they can merge and differentiate from the edges (Fig. 1d). 3.1.3 Passaging mESCs

1. Aspirate the medium holding the plate in a slanted position. 2. Add DMEM to wash the culture dish. 2 mL for 1-well of a 6-well plate or 5 mL for 10 cm plate would suffice. Be gentle, as the mESC colonies grown in low-serum or serum-free conditions may detach due to shear force (see Note 6). 3. Aspirate DMEM. 4. Add TrypLE (150–200 μL for 1 well of a 6-well plate or 1–1.5 mL for 10 cm plate) onto the cells and incubate at 37  C for 1 min and 15 s (see Note 7). 5. After removing the plate from the incubator, slant it back and forth for a couple of times to see the detachment of the colonies. Cell clumps should be visible to the naked eye. When treated with TrypLE, they would slide down from the bottom of the culture dish. When this is observed, proceed to the next step.

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6. Add DMEM to dilute the TrypLE and quench the reaction. 3 mL for 1 well of a 6-well plate or 5 mL for a 10 cm plate is sufficient. 7. Transfer the cell suspension into a 15 mL falcon tube using a serological pipette (see Note 8). 8. Wash plate with DMEM to collect the remaining cells. 2 mL for 1 well of a 6-well plate or 5 mL for 10 cm plate would suffice. 9. Centrifuge at 300 rcf, 4  C for 5 min (see Notes 2 and 3). 10. Aspirate the supernatant (see Note 4). 11. Resuspend the pellet in 1 mL 2i4 medium. Pipette up and down slowly 10–15 times using 1000 μL micropipette. Do not be harsh with the pellet, as mESCs are fragile after dissociation. 12. Count the cells using Trypan Blue stain. Dilute the cells if necessary. Keep the cells on ice while counting as ambient temperatures may reduce the viability. 13. Seed cells onto gelatin coated plates in 2i4 medium. We found 3.5  105 live cells per well of a 6-well plate in 3 mL 2i4 medium is optimal for most mESC lines we use. For a 10 cm plate, we seed 1.5  106 cells in 8 mL 2i4 medium (see Note 9). 14. Shake the culture dish, back and forth and, left and right, virtually creating a plus (+) sign while shaking (see Note 5). 15. Incubate the cells at 37  C, 5% CO2. 16. It takes up to a full day for mESCs to settle and attach to the plate bottom if they are grown in low-serum or serum-free 2i medium. If the cells have not attached after 12–16 h, shake the plate in + shape a couple of times to disperse the cells gathered in the middle of the plate. The number of the cells seeded into each well should be optimized for each mESC line. mESCs grown in low-serum or serumfree media should be passaged infrequently. As the frequency of passaging increases, healthy mESC colony morphology is progressively lost and mESCs fail to attach properly. Since 2i media can sustain pluripotency well, it is advisable to let the mESC colonies grow large rather than split frequently. Obtaining single cell suspension during passaging is critical to ensure uniformly sized mESC colonies. 3.1.4 Freezing and Storage

1. Remove the plate from the CO2 incubator. 2. Aspirate the medium holding the plate in a slanted position.

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3. Add DMEM to wash. 2 mL for 1-well of a 6-well plate or 5 mL for 10 cm plate would suffice (see Note 6). 4. Aspirate DMEM. 5. Add TrypLE (150–200 μL for 1 well of a 6-well plate or 1–1.5 mL for a 10 cm plate) onto the cells and incubate at 37  C for 1 min and 15 s (see Note 7). 6. After removing the plate from the incubator, slant it back and forth for a couple of times to see the detachment of colonies. Cell clumps should be visible to the naked eye. When treated with TrypLE, they would slide down from the bottom of the culture dish. When this is observed, proceed to the next step. 7. Add DMEM to dilute the TrypLE and quench the reaction. 3 mL for 1 well of 6-well plate or 5 mL for a 10 cm plate is sufficient. 8. Collect the cell suspension using serological pipettes into a 15 mL falcon tube (see Note 8). 9. Wash plate with DMEM to collect the remaining cells. 2 mL for 1 well of a 6-well plate or 5 mL for a 10 cm plate is sufficient. 10. Centrifuge at 300 rcf, 4  C for 5 min (see Notes 2 and 3). 11. Aspirate the supernatant (see Note 4). 12. Resuspend the pellet in 1 mL 2i4 medium. Gently pipette 10–15 times using 1000 μL micropipette. Do not be harsh with the pellet, as mESCs are fragile and already stressed at this stage. 13. Count the cells using Trypan Blue stain. Dilute the cells if necessary. Keep the cells on ice while counting as ambient temperatures may reduce the viability. 14. Centrifuge at 300 rcf, 4  C for 5 min (see Notes 2 and 3). 15. Resuspend around 2  106 cells in 800 μL Nutrifreez™ D10 Cryopreservation Medium using a 1000 μL micropipette. 16. Alternatively, 1 well of a 6-well plate can be directly frozen without counting. In that case, skip the steps 12–14, and gently resuspend the pellet in 800 μL Nutrifreez™ D10 Cryopreservation Medium using a 1000 μL micropipette. 17. Transfer 800 μL cell suspension into a cryovial. Immediately put the cryovial on ice. 18. Store at 80  C in a Styrofoam for up-to a week but at least overnight. Transfer the cryovial into a liquid nitrogen container for long term storage (see Note 10).

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3.2 mESC Growth in High-Serum Media with MEFs 3.2.1 MEF Growth and Maintenance Thawing MEFs

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1. Add 9 mL DMEM into a 15 mL falcon tube. 2. Thaw the cryovial of primary MEFs (see Note 11) in a water bath until a small piece of ice remains. To minimize the risk of contamination, keep the cap of the cryovial above water. 3. Thoroughly spray the outside of the cryovial with 70% EtOH. 4. Immediately bring it into the laminar hood. Slowly add all the content of the cryovial into the falcon tube with 9 mL DMEM using a 1000 μL micropipette. 5. Centrifuge at 300 rcf, 4  C for 5 min. 6. Aspirate the supernatant without disturbing the cell pellet. Leave a small amount of medium (100 μL) in the tube if necessary. 7. Resuspend the pellet in 1 mL MEF medium using a 1000 μL micropipette. Pipet 10–15 times to completely resuspend the pellet. 8. Count the cells using Trypan Blue and seed in MEF medium. For a 6-well plate, 5–6  105 cells should suffice. To expand the MEFs, 10 cm tissue culture plates without gelatin coating can be used. If mESCs will be seeded on MEFs after plating, gelatin coated dishes are needed. 9. Depending on the initial density of MEFs, 2–3 days might be required for 70–80% confluency (Fig. 2a).

Passaging MEFs

1. Aspirate the MEF medium on cells and wash the bottom of plate with DMEM, 5 mL for 10 cm plate. 2. Trypsinize the cells using 0.25% Trypsin—0.5% EDTA solution for 3–5 min. 1.5 mL trypsin solution is sufficient for 10 cm plates. 3. Gently shake the plate to see the detachment of the cells. Detached cells will form clumps.

Fig. 2 Mouse embryonic fibroblasts (MEFs). (a) Expanded MEFs, ready for passaging, 40 magnification. (b) mitoMEFs, ready to support mESC growth, 40 magnification

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4. Immediately quench the trypsin using 5 mL DMEM. 5. Collect the cells using a 10 mL serological pipette and transfer into a falcon tube. 6. Wash the culture dish with 5 mL DMEM to collect possible remaining cells (see Note 12). 7. Centrifuge at 300 rcf, 4  C for 5 min. 8. Aspirate the supernatant without disturbing the cell pellet. 9. Resuspend the pellet in 1 mL MEF medium using a 1000 μL micropipette. Pipette 10–15 times to completely resuspend the pellet. 10. Seed MEFs in MEF medium, 1:6 passaging ratio is appropriate while expanding the MEFs. 8 mL MEF medium is sufficient for 10 cm plates. Mitomycin-C Treatment of MEFs (See Note 13)

1. When MEFs are 70% confluent (Fig. 2a), prepare the Mitomycin-C treatment medium. 2. Aspirate the MEF medium on cells and wash the culture dish with DMEM. 3. Add the treatment medium into the culture dish. 2 mL for 1 well of a 6-well plate or 5 mL for a 10 cm plate is sufficient. 4. Incubate for 2 h at 37  C, 5% CO2 (see Note 14). 5. Aspirate the Mitomycin-C treatment medium while holding the plate in a slanted position. Wash the cells thoroughly with DMEM twice, as the residual Mitomycin-C can negatively affect ESC growth. 6. Add 8 mL MEF medium in each plate. These MEFs are now called mitoMEFs. 7. mitoMEFs can be frozen as stock after Mitomycin-C treatment and replated at desired density when needed. 8. When replated, mitoMEFs completely settle and attach in 1–3 days and can support mESCs up to 10 days (Fig. 2b).

Freezing MEFs

1. Add 400 μL 2 freezing medium to each cryovial. Keep the cryovials on ice. 2. Aspirate the MEF medium on mitoMEFs and wash plate with 5 mL DMEM. 3. Trypsinize the cells using 1.5 mL 0.25% Trypsin—0.5% EDTA solution for 3–5 min at 37  C. 4. Gently shake the plate to see the detachment of the cells. 5. Immediately quench the trypsin using 5 mL DMEM. 6. Collect the cells using a 10 mL serological pipette and transfer into a falcon tube.

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7. Wash the plate with 5 mL DMEM and collect into the same falcon tube (see Note 12). 8. Centrifuge at 300 rcf, 4  C for 5 min. 9. Aspirate the supernatant without disturbing the cell pellet. 10. Resuspend the pellet in 400 μL DMEM per cryovial using 1000 μL micropipette. Pipette 10–15 times to completely resuspend the pellet. 11. Add all the cell suspension into the cryovial with 400 μL 2 freezing medium, the total volume will be 800 μL. 12. Put the cryovial immediately on ice. Store the cryovials at 80  C, in Styrofoam. After a few days, transfer the cryovials into a liquid nitrogen tank (see Note 10). 13. 5  106 live mitoMEFs per cryovial is ideal. 1 cryovial of mitoMEFs can be thawed directly into 3  10 cm plates or 3  6-well plates. 3.2.2 Thawing mESCs on mitoMEFs

1. Add 9 mL DMEM into a 15 mL falcon tube. 2. Thaw the cryovial of mESCs in a water bath at 37  C until a small piece of ice remains. To minimize the risk of contamination, keep the cap of the cryovial above water. 3. Thoroughly spray the outside of the cryovial with 70% EtOH before bringing it into the laminar flow hood. 4. With a 1000 μL micropipette, slowly add the content of the cryovial into 9 mL DMEM prepared in step 1. 5. Centrifuge the falcon tube at 300 rcf, 4  C for 5 min (see Notes 2 and 3). 6. Aspirate the supernatant without disturbing the cell pellet. Leave a small amount (100 μL) of supernatant to prevent cell loss. 7. Resuspend the pellet in 1 mL ESC medium using a 1000 μL micropipette. Gently pipette 10–15 times to ensure single mESCs. 8. Seed cells on mitoMEFs in ESC medium. 3 mL for 1 well of a 6-well plate or 8 mL for a 10 cm plate is sufficient. If the cryovial contains a pure population of mESCs they can be counted before seeding. 2.5  105 live cells for gelatin coated 1 well of a 6-well plate or 1.5  106 live cells for a gelatin coated 10 cm plate is optimal. If mESCs were directly frozen with mitoMEFs, then optimization of seeded cell density may be required. We generally freeze mESCs as a confluent 1 well of a 6-well plate in 1 cryovial. This cryovial can be thawed into 2 or 3 wells of a 6-well plate (see Note 15). 9. Shake the plate, back and forth and, left and right, virtually creating a plus (+) sign while shaking (see Note 5).

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Fig. 3 mESCs grown in high-serum ESC medium on mitoMEFs. (a) Healthy mESC colonies on mitomycin C-treated MEFs, 40 and 100 magnification. (b) Differentiated mESC colonies on mitomycin C-treated MEFs. Cobblestone morphology, 40 and 100 magnification. (c) mESC colonies grown on mitomycin C-treated BALBc MEFs (left) or SNLP mouse fibroblast cell line (right), 40 magnification

10. Incubate the cells at 37  C, 5% CO2. As mESCs can attach better in high-serum media, they should be attached to the bottom and start forming colonies the next day. Similar to mESCs grown in 2i4 medium, proper colony morphology and density are key factors to determine passage time. When properly grown, mESC colonies are bright on the edges and show dome-shaped morphology (Fig. 3a). When grown improperly they start to flatten and eventually show cobblestonelike morphology (Fig. 3b). Generally, sparsely seeded colonies flatten and show cobble-stone morphology much faster than densely seeded colonies. Dense ones tend to merge and then flatten.

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Flattened colonies are an indication of differentiation, yet they can still be pluripotent and can be rescued after passaging. However, cobblestone-like cells are irreversibly differentiated ones and cannot be rescued by passaging. 3.2.3 Passaging mESCs on mitoMEFs

1. Remove the plate from the CO2 incubator. 2. Aspirate the medium while holding the plate in a slanted position. 3. Add DMEM to wash. 2 mL for 1 well of a 6-well plate or 5 mL for a 10 cm plate is sufficient. 4. Aspirate DMEM. 5. Add 0.25% Trypsin—0.5% EDTA solution onto the cells and incubate at 37  C for 2–3 min (300 μL for 1 well of a 6-well plate or 1.5 mL for 10 cm plate) (see Note 16). 6. After removing the culture dish from the incubator, slant it back and forth for a couple of times to see the detachment of the colonies as clumps. 7. Add DMEM to dilute the 0.25% Trypsin – 0.5% EDTA solution and quench the reaction (3 mL for 1-well of a 6-well plate or 5 mL for a 10 cm plate). 8. Collect the cell suspension using a serological pipette and transfer in a falcon tube. 9. Wash the bottom of the plate with DMEM (2 mL for 1-well of a 6-well plate or 5 mL for 10 cm plate) to collect residual cells. 10. Centrifuge at 300 rcf, 4  C for 5 min (see Note 3). 11. Aspirate the supernatant. If needed, no more than 100 μL supernatant can be left in the tube (see Note 17). 12. Resuspend the cell pellet in 1 mL ESC medium using 1000 μL micropipette. Gently pipet up-down for 10–15 times to ensure single mESCs. Do not be harsh with the pellet, as mESCs are fragile and it may affect their morphology during culturing. 13. Generally, 1:8 to 1:10 passaging ratio is appropriate. Alternatively, mESCs can be counted following mitoMEFs removal (see Subheading 3.2.4). In that case, plate proper amounts of cells in ESC medium. 2.5  105 cells for gelatin coated 1 well of a 6-well plate or 1.5  106 cells for a gelatin coated 10 cm plate is optimal. 14. Shake the culture dish, back and forth and, left and right, virtually creating a plus (+) sign while shaking (see Note 5). 15. Incubate the cells at 37  C, 5% CO2.

3.2.4 Removal of mitoMEFs from mESCs (De-MEFfing)

For downstream experiments that require pure mESC populations such as mESC differentiation, protein purification or RNA isolation from mESCs, mitoMEFs need to be removed.

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1. Remove the plate from the CO2 incubator. 2. Aspirate the medium while holding the plate in a slanted position. 3. Add DMEM to wash, 2 mL for 1 well of a 6-well plate or 5 mL for a 10 cm plate is sufficient. 4. Aspirate DMEM. 5. Add 0.25% Trypsin—0.5% EDTA solution onto the cells and incubate at 37  C for 2–3 min (300 μL for 1 well of a 6-well plate or 1.5 mL for a 10 cm plate) (see Note 16). 6. After removing the culture dish from the incubator, slant it back and forth for a couple of times to see the detachment of the colonies as clumps. 7. Add DMEM to dilute the 0.25% Trypsin—0.5% EDTA solution and quench the reaction (2 mL for 1-well of a 6-well plate or 5 mL for a 10 cm plate). 8. Collect the cell suspension using a serological pipette and transfer into a falcon tube. 9. Wash the bottom of the culture dish with DMEM (2 mL for 1-well of a 6-well plate or 5 mL for a 10 cm plate) to collect residual cells. 10. Centrifuge at 300 rcf, 4  C for 5 min (see Note 3). 11. Aspirate the supernatant. If needed, no more than 100 μL supernatant can be left in the tube (see Note 17). 12. Resuspend the pellet in 1 mL ESC medium using 1000 μL micropipette. Gently pipet 10–15 times. Do not be harsh with the pellet, as mESCs are fragile and it may affect their morphology during culturing. 13. Replate the collected cells into the same size of culture dish in ESC medium (i.e., if the cells were collected from one well of a 6-well plate, replate the cells in a gelatin coated one well of a 6-well plate). Using a lower amount of ESC medium is important as it helps cells settle faster, saving time. 2 mL ESC medium for 1 well of a 6-well plate or 5 mL ESC medium for a 10 cm plate would suffice. 14. Incubate the cells at 37  C, 5% CO2 for 45 min. 15. At this stage, mitoMEFs should be attached to the bottom, whereas the mESCs are in suspension. Slant the plate and gently collect the medium on cells into a falcon tube using a 1000 mL micropipette. Be careful not to disturb the bottom of the dish as MEFs are not strongly attached and can come off due to shear force. Gently wash the plate by very slowly pouring the medium dropwise from the wall of the plate. mitoMEFs can be seen attached to the bottom of the dish under light microscope, while most mESCs remain unattached.

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16. Centrifuge at 300 rcf, 4  C for 5 min (see Note 3). 17. Aspirate the supernatant and resuspend the pellet in required medium depending on the purpose. 18. mESCs can be counted using Trypan Blue in this stage. Some mitoMEFs are to be expected. They appear much bigger than the mESCs. If there is extensive mitoMEF contamination, repeat steps 12–19. 19. Obtained mESCs can be replated, frozen, and used for further analysis. 3.2.5 Freezing and Storage

1. Add 400 μL 2 freezing medium to each cryovial. Keep the cryovials on ice. 2. Remove the plate from the CO2 incubator. 3. Aspirate the medium while holding the plate in a slanted position. 4. Add DMEM to wash. 2 mL for 1 well of a 6-well plate or 5 mL for a 10 cm plate is sufficient. 5. Aspirate DMEM. 6. Add 0.25% Trypsin—0.5% EDTA solution onto the cells and incubate at 37  C for 2–3 min (300 μL for 1 well of a 6-well plate or 1.5 mL for a 10 cm plate) (see Note 16). 7. After removing the culture dish from the incubator, slant it back and forth for a couple of times to see the detachment of the colonies as clumps. 8. Add DMEM to dilute the 0.25% Trypsin—0.5% EDTA solution and quench the reaction (2 mL for 1-well of a 6-well plate or 5 mL for a 10 cm plate). 9. Collect the cell suspension using a serological pipette and transfer into a falcon tube. 10. Wash the bottom of the culture dish with DMEM (2 mL for 1-well of a 6-well plate or 5 mL for a 10 cm plate) to collect residual cells. 11. Centrifuge at 300 rcf, 4  C for 5 min (see Note 3). 12. Aspirate the supernatant. If needed, no more than 100 μL supernatant can be left in the tube (see Note 17). 13. Resuspend the pellet in 400 μL DMEM per cryovial. 2–3  106 mESCs per cryovial is ideal. To count the mESCs before freezing, they should be separated from mitoMEFs (see Subheading 3.2.4). Alternatively, cells in 1 well of a 6-well plate can be directly frozen in 1 cryovial, similarly, mESCs from a 10 cm plate can be frozen in 6 cryovials. 14. Add 400 μL of cell suspension into the cryovial with 400 μL 2 freezing medium using 1000 μL micropipette. Gently pipet

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4–5 times to mix cell suspension and 2 freezing medium in the cryovial. Total volume should be 800 μL. 15. Put the cryovial immediately on ice. Store the cryovials at 80  C, in Styrofoam. After a few days, transfer the cryovials into a liquid nitrogen tank (see Note 10). 3.2.6 Adaptation of mESCs to Low-Serum and Serum-Free 2i Media

mESCs adapted to high-serum containing ESC medium might have attachment problems when thawed directly in serum-free 2i medium. We found gradual decrease of serum in the growth medium is useful to circumvent attachment problems. Serum-free 2i medium formulation can be supplemented with a low percentage of serum. With each passage, the serum percentage can be halved to reach desired serum percentage. We found keeping 4% serum in 2i condition was helpful for attachment and morphology of mESCs. However, serum-free 2i medium formulation should be used if mESCs will be differentiated toward neuroectodermal lineage (see Directed Differentiation of Mouse Embryonic Stem Cells to Mesoderm, Endoderm, and Neuroectoderm Lineages chapter, Subheading 3.2). 1. Thaw a cryovial of confluent mESCs frozen from one well of a 6-well plate into 2–3 wells of a 6-well plate coated with gelatin in 3 mL 2i4 medium as previously described (see Subheading 3.1.2) (see Note 18). 2. The cells will take 2–4 days before passaging. If the cell death is high but the remaining colonies are attached properly, you can wash the well with 2 mL DMEM, twice, to remove the dead mESCs as they can be toxic to the healthy colonies and negatively affect their adaptation. 3. If the thawed cryovial contains mitoMEFs, removal of the mitoMEFs is preferred (see Subheading 3.2.4). 4. Passage the mESCs when they are 70–80% confluent as previously described (see Subheading 3.1.3). Due to flattened morphology in earlier passages, cells can reach confluency faster than expected. Since the mESCs are now in 2i4 medium, use TrypLE instead of Trypsin for passaging (see Note 19). 5. Seed 2.5–3.5  105 cells into gelatin coated 1 well of a 6-well plate in 3 mL 2i4 medium. 6. Repeat the steps 2–5 until the colonies show healthy mESC morphology (Fig. 1a). mESCs can be grown and maintained in low-serum 2i4 medium after the adaptation is completed. Alternatively, mESCs can be further adapted to serum-free 2i medium. 7. To adapt mESCs to 2i medium, repeat steps 2–5. For each passage, reduce the serum concentration to half. That is, as the mESCs are in 2i4 medium with 4% serum, seed the mESCs in

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2% serum containing 2i medium (we refer to it as 2i2 medium) when they are passaged. 8. In 2i2 media, mESC attachment might be impaired. Therefore, some colonies will first form in suspension the next day and attach to the bottom of the dish a day later. This might lead to uneven colony size since they might merge when they settle (see Note 20). 9. During the following passages, use Accutase instead of TrypLE solution to detach the colonies from the well bottom. 300 μL Accutase for 1-well of a 6-well plate is sufficient. Incubate the cells in Accutase for 3–4 min at 37  C, 5% CO2. All other steps are the same as previously described (see Subheading 3.1.3) 10. Seed 2.5–3.5  105 mESCs into gelatin coated 1 well of a 6-well plate in 3 mL 2i1 medium (2i medium with 1% serum). 11. mESCs in 2i2 and 2i1 media show much less attachment capabilities. They form colonies in suspension and attach to the bottom of the well in 1 or 2 days. Generally, they become confluent in 3 days. Therefore, observe the well under a light microscope every day and shake the plate to disperse the colonies until they attach. 12. Final passage will be conducted as the previous one, using Accutase and the mESCs will be seeded in serum-free 2i medium. 13. After full adaptation to serum-free 2i medium, mESC attachment will cease to be a problem. However, it is still recommended to avoid serial passaging for an extended time in 2i medium (see Note 21). 14. Use Accutase for passaging, and all other steps are the same as described for mESCs in 2i4 medium. 3.2.7 Adaptation of mESCs to High-Serum Medium with MEFs

Adaptation of mESCs to high-serum medium is much easier. mESCs can strongly attach in ESC medium, their viability after thaw and passages are much higher when compared to mESCs grown in low-serum or serum-free 2i media. 1. Prepare mitoMEFs as previously described (see Subheading 3.2.1). 2. A cryovial of 5  106 live mitoMEFs can be thawed into gelatin coated 3  10 cm plates or 3  6-well plates. They settle within 1–3 days after thawing and become ready to support mESC growth. 3. Thaw a cryovial of confluent mESCs frozen from 1-well of a 6-well plate into 2–3 wells of a 6-well plate coated with gelatin and mitoMEFs in 3 mL ESC medium as previously described (see Subheading 3.2.2).

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4. Two or three passages are required for the full adaptation into ESC medium and mitoMEFs. Trypsin is preferred for passaging mESCs in high-serum medium. 5. Generally, most colonies show healthy mESC morphology starting from the beginning (Fig. 3a). Some colonies may differentiate, flatten and even some of them show cobblestone-like morphology (Fig. 3b). With every passage, those unhealthy colonies will diminish in number (no more than 5% of colonies).

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Notes 1. B-27 Supplement has two different formulations. During mESC growth and maintenance, we use the one with retinoic acid (or vitamin A). During the differentiation into mesendoderm, we use the B-27 variant with no retinoic acid, as the retinoic acid promotes neuroectoderm differentiation. 2. Use a swing-out rotor as it helps better attachment of mESCs to the tube bottom. mESCs grown in low-serum or serum-free conditions tend to not pellet well. 3. Do not forget to cool the centrifuge beforehand, as centrifuging at higher temperatures may lead to low cell viability. 4. The pellet may be very small, depending on the cell number. To prevent aspirating the pellet, leave a small amount of the supernatant (not more than 100 μL). 5. This is especially important when seeding the cells into a small well. If it is not properly done, mESCs can gather in the middle of the well, and merge as they grow. This would lead to differentiation of the merged colonies. 6. mESCs in low-serum or serum-free 2i media have difficulties in attachment to the surface when compared to the mESCs grown in high-serum medium. 7. Be careful here, as overtreatment with TrypLE may lead to low colony attachment after passage. On the other hand, undertreatment would result in uneven colony size after passaging, and cause problems while counting the cells. When mESCs are grown in low-serum or serum-free 2i media, a gentle trypsinization method should be used, as they face difficulties in attachment. As an alternative to TrypLE, Accutase can be used. 8. Use a 15 mL falcon tube rather than a 50 mL one, as mESCs grown in low-serum or serum-free 2i media form a loose pellet when compared to mESCs grown in high-serum media. The narrower bottom of the 15 mL falcon tube helps reduce cell loss during wash steps.

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9. The cell numbers for seeding may require optimization depending on the background of the mESCs. For example, some knock-out mESC lines may grow slowly, or form smaller colonies than wild type ones. 10. Cells may remain at 80  C for a month if frequently used. However, the long term storage at 80  C may lead to low viability upon thawing. 11. Murine embryonic fibroblast (MEFs) cells are widely used as a feeder layer for the growth and maintenance of both mouse and human ESCs. There are a variety of MEFs which can be used for this purpose. Initially, we used a commercial line of LIF-secreting mouse fibroblast cell line, SNLP 76/7-4 (ATCC® SCRC-1050™). However, we were not able to obtain healthy mESC morphology with this line (Fig. 3c). Primary MEFs from BALB-c mice provided us with healthy mESC morphology. 12. MEFs attach to the plate very strongly. Wash the plate as thoroughly as possible to yield maximum MEF numbers. 13. Primary MEFs stop dividing and become senescent after 3–4 passages. Therefore, once a vial of primary MEFs is thawed, try to expand it for 3–4 passages in 15 cm plates and treat with Mitomycin-C afterward. This way, stocks of mitoMEFs can be prepared in large quantities and the amount of Mitomycin-C required is minimized. 14. Longer periods of Mitomycin-C treatment may reduce the viability of the MEFs. Generally, 1–3 h of treatment is necessary depending on the MEF line. Optimization may be required. 15. Generally, 2–4 days are required before passaging if the thawed mESC cryovial was frozen recently. mESCs remained frozen for 4–5 years should be thawed into 1 well of a 6-well plate even if they were frozen directly from 1 well of a 6-well plate, as they show poor viability. 16. Be careful here, as overtreatment with 0.25% Trypsin—0.5% EDTA solution may lead to low colony attachment after passage. On the other hand, undertreatment would result in uneven colony size after passaging as clumps remained. 17. mESCs grown in high-serum containing medium form a firmer pellet than mESC grown in low-serum or serum-free media. 18. As they need time to adapt to their new medium, the colonies will look unhealthy for a couple of passages. That is, most colonies will be flattened, losing their dome-shaped morphology. They will not be as bright as before, and they may have difficulties in attachment to the bottom of the well.

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19. Trypsin is harsher on the mESCs and can negatively affect attachment or colony morphology. 20. During the adaptation process, mESCs in low-serum or serumfree 2i media may form colonies in suspension while some colonies grow attached to the bottom. In such a case, colonies in suspension can be collected, and single cells can be obtained using Accutase. When reseeded, these mESCs have been observed to attach to the bottom. 21. While thawing the mESCs grown in 2i medium, using preheated media (37  C) can alleviate possible attachment problems and reduce cell death. References 1. Huang G, Ye S, Zhou X et al (2015) Molecular basis of embryonic stem cell self-renewal: from signaling pathways to pluripotency network. Cell Mol Life Sci 72:1741–1757 2. Niwa H (2014) The pluripotency transcription factor network at work in reprogramming. Curr Opin Genet Dev 28:25–31 3. Ying QL, Smith A (2017) The art of capturing pluripotency: creating the right culture. Stem Cell Rep 8:1457–1464 4. Wray J, Kalkan T, Smith AG (2010) The ground state of pluripotency. Biochem Soc Trans 38:1027–1,032

5. Nichols J, Smith A (2009) Naive and primed pluripotent states. Cell Stem Cell 4:487–492 6. Tamm C, Galito´ SP, Annere´n C (2013) A comparative study of protocols for mouse embryonic stem cell culturing. PLoS One 8:1–17 7. Chaudhry MA, Vitalis TZ, Bowen BD et al (2008) Basal medium composition and serum or serum replacement concentration influences on the maintenance of murine embryonic stem cells. Cytotechnology 58:173–179 8. Marks H, Kalkan T, Menafra R et al (2012) The transcriptional and epigenomic foundations of ground state pluripotency. Cell 149:590–604

Methods in Molecular Biology (2022) 2520: 295–307 DOI 10.1007/7651_2021_439 © Springer Science+Business Media, LLC 2021 Published online: 06 October 2021

Directed Differentiation of Mouse Embryonic Stem Cells to Mesoderm, Endoderm, and Neuroectoderm Lineages Emre Balbasi, Dersu Sezginmert, Ceren Alganatay, and Nihal Terzi Cizmecioglu Abstract The self-renewal and pluripotency features of mouse embryonic stem cells (mESCs) make them a great tool to study early mammalian development. Various signaling pathways that shape early mammalian development can be mimicked for in vitro mESC differentiation toward primitive lineages first and more specialized cell types later. Since the precise nature of the molecular mechanisms and the crosstalk between these signaling pathways is yet to be fully understood, there is a high level of variability in the efficiency and synchronicity among available differentiation protocols. Commitment of mESCs toward mesoderm, endoderm, or neuroectoderm lineages happens over only a few days and is highly efficient. Here, we provide protocols for the directed differentiation of mESCs toward these lineages in vitro. Key words Mouse embryonic stem cells, mESCs, mESC differentiation, Primary germ layers, Endoderm, Mesoderm, Ectoderm

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Introduction Embryonic stem cells (ESCs) have the capability of self-renewal and differentiation into three germ layers called mesoderm, endoderm, and ectoderm. Due to their potential to form all primary germ layers, mouse embryonic stem cells (mESCs) are a great tool and model to study early mammalian development in vitro [1]. mESCs are derived from the inner cell mass (ICM) of the preimplantation blastocyst stage of the developing mouse embryo. After implantation, the ICM forms the pluripotent cells called epiblast stem cells (EpiSCs). Through spatiotemporal control of Wnt, Nodal, and BMP signaling pathways, EpiSCs form the primitive streak on the posterior side of the developing embryo, which gives rise to mesoderm and later definitive endoderm layers [2, 3]. Graded signaling of Wnt, Nodal and BMP throughout the primitive streak defines the differentiation into either layer. While canonical Wnt, Nodal and BMP signaling drive the differentiation into mesodermal lineages [4–8], a robust Nodal signaling is required for the definitive endoderm formation during gastrulation

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[7, 9]. On the other hand, inhibition of Nodal and Wnt signaling on the anterior side of the epiblast leads to the formation of ectoderm layer [2]. Using the knowledge gained from in vivo studies of early mouse development, efficient and successful in vitro differentiation protocols of mESCs for all primary germ layers have emerged [10–14]. The use of various cytokines at the right concentration enables the formation of desired primary germ layer cells from mESCs in vitro. Here we present the methods which were successfully adapted and used in our laboratory for the directed differentiation of mESCs into mesoderm, endoderm and neuroectoderm lineages in vitro.

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Materials IMDM (Cat. No.: 21980032, Gibco). Ham’s F-12 Nutrient Mix, GlutaMAX™ Supplement (Cat. No.: 31765027, Gibco). DMEM/F-12 (Cat. No.: 11320074, Gibco). Neurobasal Medium (Cat No.: 21103049, Gibco). B-27™ Supplement (50), minus vitamin A (Cat. No.: 12587010, Gibco). B-27™ Supplement (50), serum free (Cat. No.: 17504044, Gibco). N-2 Supplement (100) (Cat. No.: 17502048, Gibco). N-2 MAX Media Supplement (100) (Cat No.: AR009, R&D Systems). GlutaMAX I (Cat. No.: 35050061, Gibco). L-Ascorbic

acid (Cat. No.: A4544-25G, Sigma).

1-Thioglycerol (MTG) (Cat. No.: M6145-25ML, Sigma). Recombinant Human/Murine/Rat Activin A (Cat. No.: 120-14P, Peprotech). Recombinant Mouse Wnt3a Protein (Cat. No.: ab81484, Abcam). Recombinant Human BMP-4 Protein (Cat. No.: 314-BP, R&D Systems). Accutase cell detachment solution (Cat. No.: SCR005, Millipore). Bovine serum albumin (Cat. No.: A3311-50G, Sigma). Water, cell culture grade (Cat. No.: BI03–055-1A, Biological Industries). Dulbecco’s phosphate buffered saline (DPBS) (Cat. No.: 02–0231A, Biological Industries). 6-cm petri dishes. 10-cm petri dishes. 6-well cell culture plates.

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L-Ascorbic Acid Solution: Dissolve 5 mg L-Ascorbic acid in 10 mL water by heating the solution to 37  C. Use cell culture grade water. Prepare fresh before each use; do not store.

MTG (1-Thioglycerol) Solution: Dilute 13 μL MTG in 1 mL IMDM. Mix well. Prepare fresh before each use; do not store. 10% BSA (w/v): Dissolve 5 g bovine serum albumin in 50 mL Dulbecco’s Phosphate Buffered Saline. Filter-sterilize and store it at 4  C. 2.2 Media Preparations

Serum-free Differentiation Base Medium (100 mL): 75 mL IMDM, 25 mL Ham’s F-12 Nutrient Mix with GlutaMAX™ Supplement, 5 mL 10% BSA, 1 mL B-27 Supplement without Vitamin A (50) (see Note 1), 500 μL N-2 Supplement (100), 1 mL GlutaMAX I, 1 mL L-ascorbic acid solution, 300 μL MTG solution. Prepare the Serum-free Differentiation Base Medium in a filter bottle and filter-sterilize before use. The medium should be freshly prepared just before setting up differentiation experiments. Use within 10 days. The same medium can be used throughout the experiment. Store at 4  C. For endoderm commitment, Serum-free Differentiation Base Medium is supplemented with 75 ng/mL Activin A after the second day of differentiation. For mesoderm commitment, Serum-free Differentiation Base Medium is supplemented with 1 ng/mL Activin A, 3 ng/mL Wnt3a, and 1 ng/mL BMP4 after the second day of differentiation. Neuroectoderm Differentiation Medium (N2B27) (100 mL): 50 mL DMEM/F-12, 50 mL Neurobasal Medium, 2 mL B-27 Supplement (50), 1 mL N-2 MAX Media Supplement (100), 50 μL 10% BSA, 300 μL MTG Solution. Prepare the N2B27 medium in a filter bottle and filter-sterilize before use. The medium should be freshly prepared just before setting up differentiation experiments. Use within 10 days. Same medium can be used throughout the experiment. Store at 4  C.

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Methods

3.1 Endoderm and Mesoderm Differentiation of mESCs

Day 0 and Day 1 of differentiation are common for endoderm and mesoderm. On Day 2, use the specific protocol for endoderm or mesoderm differentiation as explained below. Day 0

1. Collect mESCs as previously described (see Mouse Embryonic Stem Cell Culture in Serum-Containing or 2i Conditions

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Fig. 1 Endoderm differentiation of mESCs. Endoderm differentiation time course. EBs on Day 2 (a), Day 3 (b), Day 4 (c), and Day 5 (d) of endoderm differentiation. 40 magnification (left), 100 magnification (right)

chapter, Subheading 3.1.3 or Subheading 3.2.4). Resuspend the pellet in 1 mL IMDM after centrifugation using 1000 μL micropipette. Gently pipet up and down for 10–15 times. Make sure to resuspend cell clumps completely as a single-cell suspension. 2. Count the cells using Trypan Blue stain. Keep the cells on ice while counting as ambient temperatures may reduce the viability. 3. Seed 7.5  105 live mESCs in a 10-cm petri dish (see Note 2) in 8 mL Serum-free Differentiation Base medium. 4. Shake the petri dish to disperse the cells homogeneously in the medium (see Note 3). 5. Incubate the dish at 37  C, 5% CO2 for 48 h (see Note 4). Day 2: Endoderm Differentiation

6. After 48 h incubation, collect the embryoid bodies using a 10 mL serological pipette into a 50 mL falcon tube. In this stage, the embryoid bodies should be visible to naked eye as small dots. They are spherical in shape, transparent and bright (Fig. 1a). 7. Centrifuge at 180 rcf (1000 rpm), 4  C for 5 min, using a swing-out rotor. 8. Aspirate the supernatant (see Note 5).

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9. Resuspend the pellet in 0.8–1 mL Accutase using a 1000 μL micropipette. Gently pipet up and down for 4–5 times. The Accutase solution should look clear with visible embryoid bodies in it. 10. Immediately bring the tube into a 37  C water bath. Incubate the embryoid bodies in Accutase for 2–4 min while gently swirling the tube in the water bath. After 2 min of incubation, remove the tube from the water bath, gently shake and check if the embryoid bodies dissociated. As the single cells form, the Accutase solution gets cloudy and embryoid bodies get smaller. If this is not observed, incubate for 30 more seconds in the water bath, and check again (see Note 6). 11. Once the embryoid bodies dissociate, bring the tube into the laminar flow hood. 12. Quench the Accutase reaction with 5 mL IMDM. 13. Centrifuge at 400 rcf (1500 rpm), 4  C for 5 min, using a swing-out rotor. 14. Resuspend the pellet in 1 mL IMDM using a 1000 μL micropipette. Gently pipet up and down for 10–15 times. 15. Count the cells using Trypan Blue stain. Keep the cells on ice while counting as ambient temperatures may reduce the viability. 16. Seed 5  105 live cells in a 6-cm petri dish in 3 mL Serum-free Differentiation Base Medium supplemented with 75 ng/mL Activin A (Serum-free Endoderm Differentiation Medium). Prepare 2 or 3 6-cm petri dishes for each day of interest depending on how many cells are needed for downstream applications (see Note 7). 17. Shake the petri dishes to homogeneously disperse the cells (see Note 3). 18. Incubate the dishes at 37  C, 5% CO2. 19. We generally collect samples every day starting with the third day of differentiation. Sample collection period may vary based on the downstream application. Day 3–6 of Endoderm Differentiation

20. Collect the embryoid bodies using a 10 mL serological pipette into a 15 mL falcon tube. 21. Centrifuge at 180 rcf (1000 rpm), 4  C for 5 min, using a swing-out rotor. 22. Aspirate the supernatant (see Note 5). 23. Resuspend the pellet in 200–300 μL Accutase (for 2 or 3 6-cm dishes) using a 1000 μL micropipette. Pipet up and down for 4–5 times. At this stage, the embryoid bodies should be visible

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to naked eye as small dots. The Accutase solution should look clear with visible embryoid bodies in it. 24. Immediately bring the tube into the water bath at 37  C. Incubate the embryoid bodies in Accutase for 2–3 min while gently swirling the tube in the water bath. After 2 min of incubation, remove the tube from the water bath and check if the embryoid bodies dissociate. As the single cells form, the Accutase solution gets cloudy and the embryoid bodies get smaller. If this is not observed, incubate for 30 more seconds in the water bath, and check again (see Note 6). 25. Once the embryoid bodies dissociate, bring the tube into the laminar flow hood. 26. Quench the Accutase reaction with 5 mL IMDM. 27. Centrifuge at 400 rcf (1500 rpm), 4  C for 5 min, using a swing-out rotor. 28. Resuspend the pellet in 1 mL IMDM using a 1000 μL micropipette. Gently pipet up and down for 10–15 times. 29. Obtained cells are ready to be used for a variety of experiments such as RNA isolation followed by qPCR analysis or protein extraction for western blot analysis (see Note 8). We generally extend the differentiation to 6 days. This medium does not support embryoid body viability post day 6. To further extend the differentiation, the medium can be refreshed. If the desired markers are not expressed within 6 days (see Note 9), embryoid bodies can be further differentiated into desired lineages with appropriate cytokines. Day 2: Mesoderm Differentiation

6. After 48 h incubation, collect the embryoid bodies using a 10 mL serological pipette into a 50 mL falcon tube. In this stage, the embryoid bodies should be visible to naked eye as small dots. They are spherical in shape, transparent and bright (Fig. 1a). 7. Centrifuge at 180 rcf (1000 rpm), 4  C for 5 min, using a swing-out rotor. 8. Aspirate the supernatant (see Note 5). 9. Resuspend the pellet in 0.8–1 mL Accutase using a 1000 μL micropipette. Gently pipet up and down for 4–5 times. The Accutase solution should look clear with visible embryoid bodies in it. 10. Immediately bring the tube into the water bath at 37  C. Incubate the embryoid bodies in Accutase for 2–4 min while gently swirling the tube in the water bath. After 2 min of incubation, remove the tube from the water bath, gently

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shake and check if the embryoid bodies dissociated. As the single cells form, the Accutase solution gets cloudy and the embryoid bodies get smaller. If this is not observed, incubate for 30 more seconds in the water bath, and check again (see Note 6). 11. Once the embryoid bodies dissociate, bring the tube into the laminar flow hood. 12. Quench the Accutase reaction with 5 mL IMDM. 13. Centrifuge at 400 rcf (1500 rpm), 4  C for 5 min, using a swing-out rotor. 14. Resuspend the pellet in 1 mL IMDM using a 1000 μL micropipette. Gently pipet up and down for 10–15 times. 15. Count the cells using Trypan Blue stain. Keep the cells on ice while counting as ambient temperatures may reduce the viability. 16. Seed 7.5  105 live cells in a 6-cm petri dish in 3 mL Serumfree Differentiation Base Medium supplemented with 1 ng/mL Activin A, 3 ng/mL Wnt3a, and 1 ng/mL BMP4 (Serum-free Mesoderm Differentiation Medium). Prepare 2 or 3 6-cm petri dishes for each day of interest depending on how many cells are needed for downstream applications (see Note 7). 17. Shake the petri dishes to homogeneously disperse the cells (see Note 3). 18. Incubate the dishes at 37  C, 5% CO2. 19. In our lab, we generally collect samples every day starting with the third day of differentiation. Depending on the experiments, sample collection periods may vary. Day 3–6 of Mesoderm Differentiation

20. Collect the embryoid bodies using 10 mL serological pipettes into a 15 mL falcon tube. 21. Centrifuge at 180 rcf (1000 rpm), 4  C for 5 min, using a swing-out rotor. 22. Aspirate the supernatant (see Note 5). 23. Resuspend the pellet in 200–300 μL Accutase (for 2 or 3 6-cm dishes) using a 1000 μL micropipette. Pipet up and down for 4–5 times. In this stage, the embryoid bodies should be visible to naked eye as small dots. The Accutase solution should look clear with visible embryoid bodies in it. 24. Immediately bring the tube into the water bath at 37  C. Incubate the embryoid bodies in Accutase for 2–3 min while gently swirling the tube in the water bath. After 2 min of incubation, remove the tube from the water bath and check if the embryoid bodies dissociate. As the single cells form, the

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Accutase solution gets cloudy and the embryoid bodies get smaller. If this is not observed, incubate for 30 more seconds in the water bath, and check again (see Note 6). 25. Once the embryoid bodies dissociate, bring the tube into the laminar flow hood. 26. Quench the Accutase reaction with 5 mL IMDM. 27. Centrifuge at 400 rcf (1500 rpm), 4  C for 5 min, using a swing-out rotor. 28. Resuspend the pellet in 1 mL IMDM using a 1000 μL micropipette. Gently pipet up and down for 10–15 times. 29. Obtained cells are ready to be used for a variety of experiments such as RNA isolation followed by qPCR analysis or protein extraction for western blot analysis (see Note 8). 3.2 Neuroectoderm Differentiation of mESCs

mESCs are grown in serum-free 2i medium to 70–80% confluency prior to experiment (Fig. 2a). Since the Neuroectoderm Differentiation Medium (N2B27) is a serum-free medium, it is important to adapt the cells to serum-free medium (2i) prior to the experiment as described (see Mouse Embryonic Stem Cell Culture in SerumContaining or 2i Conditions chapter, Subheading 3.2.6). Culturing mESCs in low-serum medium (2i4) results in decreased cell attachment and increased cell death during neuroectoderm differentiation process. Neuroectoderm differentiation process is accompanied by high numbers of cell death, especially after the third day of differentiation. It is advised that the number of wells is scaled up accordingly if the final experiment requires substantial cell numbers (see Note 10). The following protocol utilizes a monolayer adherent differentiation method. 1. Collect mESCs using Accutase. Accutase treatment should be adjusted in amount and duration according to the colony size and confluency. 300 μL Accutase for 1-well of a 6-well plate is sufficient. Incubate the cells in Accutase for 2.5–4 min at 37  C, 5% CO2. All other steps are the same as previously described (see Mouse Embryonic Stem Cell Culture in Serum-Containing or 2i Conditions chapter, Subheading 3.1.3). 2. Centrifuge at 300 rcf, 4  C for 5 min, using a swing-out rotor. 3. Aspirate the supernatant. 4. Resuspend the pellet in 1 mL DMEM using 1000 μL micropipette. Gently pipet up and down for 10–15 times to obtain a single cell suspension. 5. Count cells using Trypan Blue stain. Keep the cells on ice while counting as ambient temperatures may reduce the viability.

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Fig. 2 Neuroectoderm differentiation of mESCs. Neuroectoderm differentiation time course. mESCs (a), Day 2 (b), Day 3 (c), Day 4 (d), and Day 5 (e) of neuroectoderm differentiation. (a–c) 100 magnification. (d, e) 40 magnification (left), 100 magnification (middle), 200 magnification (right)

6. Seed 1.1  105 live mESCs for each well of gelatinized 6-well plate in 1.5 mL N2B27 medium (see Note 11). 7. Shake the plate to disperse the cells homogeneously. 8. Incubate the plate at 37  C, 5% CO2. 9. Change the medium every other day (Day 2, Day 4, etc.) as follows (see Notes 4 and 12):

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(a) Aspirate the old medium while keeping the plate tilted. (b) Keep the plate tilted all the time and add 2 mL DMEM slowly by leaning the tip of the serological pipette against the upper wall of the well (see Note 13). (c) Keep the plate tilted and aspirate the medium. (d) Add 3 mL fresh N2B27 medium dropwise by leaning the tip of the serological pipette against the wall of the well. Addition of fresh medium should be done as slowly as possible to avoid detachment of cells. 10. Collect cells on desired days using Accutase as follows: (a) Aspirate the old medium while keeping the plate tilted. (b) Keep the plate tilted and add 2 mL DMEM slowly by leaning the tip of the serological pipette against the upper wall of the well. (c) Keep the plate tilted and aspirate the medium. (d) Add 200–300 μL Accutase to each well and incubate the plate at 37  C for 2 min. (e) Remove the plate from the incubator and slant it back and forth for a couple of times to see the detachment of cells. If sliding of the cells is not observed, the plate can be incubated for an extended period by checking every 20–30 s. (f) Add 3 mL DMEM to dilute the Accutase and quench the reaction. (g) Collect the cell suspension using a serological pipette into a 15 mL or 50 mL falcon tube. (h) Wash the wells with 2 mL DMEM to collect the remaining cells. (i) Centrifuge at 300 rcf, 4  C for 5 min, using a swing-out rotor. (j) Aspirate the supernatant. (k) Resuspend the pellet in 1 mL DMEM using 1000 μL micropipette. Gently pipet up and down for 10–15 times to obtain a single cell suspension. (l) Obtained cells are ready to be used for a variety of experiments such as RNA isolation followed by qPCR analysis or protein extraction for western blot analysis. We usually collect samples starting from Day 3 of neuroectoderm differentiation. Depending on the desired final experiment, sample collection periods may vary (see Notes 10 and 11).

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Notes 1. B-27 supplement has two variations, with or without vitamin A (retinoic acid). Vitamin A suppresses the mesendoderm differentiation while promoting neuroectoderm differentiation. Therefore, it should not be included in the media for endoderm or mesoderm differentiation. Make sure to use the B-27 supplement without vitamin A in the media composition. 2. Embryoid body formation can be promoted using dishes that prevent ESCs from settling down and attaching. We routinely use petri dishes during EB based differentiation. There are also commercially available low attachment dishes with modified wells to allow formation of homogeneously sized EBs. If your experiments need only very few EBs and that you do not need to scale up, you can also use the hanging drop method explained elsewhere. 3. During endoderm and mesoderm differentiation, mESCs will form embryoid bodies in suspension. Dispersing the mESCs homogeneously will result in more uniform embryoid body sizes. 4. After starting differentiation on Day 0, the timing of the following days need to be consistent. That is, if the cells were placed into the CO2 incubator at 13.00 PM on Day 0, collect the cells at 13.00 PM on Day 2 and on consecutive days as well. 5. The pellet will be loose. Slant the tube and aspirate the supernatant without getting the tip near the pellet. The pellet may slide down, try to be quick after centrifugation. 6. Incubation time is dependent on embryoid body size. The larger ones will dissociate later than the small ones. If the embryoid body sizes are not uniform, the small ones may die due to overtreatment with Accutase until the large embryoid bodies dissociate. The embryoid bodies collected on Day 3 (Fig. 1b) will be much smaller than the ones collected on further days (Fig. 1c, d). Therefore, they should be treated less. 7. We use 5  105 and 7.5  105 live cells per 6-cm petri dish for endoderm and mesoderm differentiation respectively. However, the cell numbers might need to be optimized for different mESC lines. Also, we find 6-cm petri dishes suitable for the differentiation as one to two million live cells per day are sufficient for the analyses routinely done in our laboratory. If the following experiments require high numbers of cells, use a larger petri dish, and optimize the cell numbers to seed based on the volume of the medium you need to use. 8. Using these differentiation methods, we observe >70% Brachyury (Bry, T) positive cells using reporter mESC lines. Bry

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expression occurs first at endoderm or mesoderm differentiation day 3–5. To validate the definitive endoderm differentiation success, we perform qPCR analysis for Foxa2 and Sox17 whose expression follows Bry expression, usually with a delay of 12–24 h. 9. Culturing conditions of the mESCs affect when lineage marker expressions peak. mESCs cultured in 2i containing medium tend to express lineage markers a day or two later than the mESCs cultured in serum-containing medium. 10. When we initiate the neuroectoderm differentiation with 1.1  105 serum-free adapted, live mESCs, we obtain about 8.0  105 live cells per well on Day 3, 6.5  105 live cells per well on Day 4, and 3.0  105 live cells on Day 5. 11. The cell numbers should be optimized for each cell line; seeding too many cells results in poor differentiation while seeding too few cells may result in higher cell death. Especially during cell number optimization, qPCR validation of differentiation process should be done with key neuroectoderm markers (such as Sox1, Pax6 and N-cadherin) and key pluripotency markers (such as Nanog and Oct4) since changing cell numbers might result in poor differentiation. Sox1 expression reaches a peak at Day 4 while the peak for Pax6 expression is observed at Day 5. Nanog expression decreases sharply on Day 3 and remains low on the following days of the neuroectoderm differentiation. 12. Formation of attached nonspherical, monolayer colonies should be visible starting from Day 2 (Fig. 2b–e). It is expected to observe some detached cells in aggregates after medium change on Days 2 and 4 (Fig. 2b, d). It should be noted that differentiation occurs in patches and the whole surface of the well may not be covered. 13. This step aims to wash off dead cells and the old medium. However, the attached cells easily detach during neuroectoderm differentiation protocol, therefore swirling or shaking the plate is discouraged. References 1. Huang G, Ye S, Zhou X et al (2015) Molecular basis of embryonic stem cell self-renewal: from signaling pathways to pluripotency network. Cell Mol Life Sci 72:1741–1757 2. Arnold SJ, Robertson EJ (2009) Making a commitment: cell lineage allocation and axis patterning in the early mouse embryo. Nat Rev Mol Cell Biol 10:91–103 3. Tam PPL, Loebel DAF (2007) Gene function in mouse embryogenesis: get set for gastrulation. Nat Rev Genet 8:368–381

4. Mishina Y, Suzuki A, Ueno N et al (1995) Bmpr encodes a type I bone morphogenetic protein receptor that is essential for gastrulation during mouse embryogenesis. Genes Dev 9:3027–3037 5. Zeng L, Fagotto F, Zhang T et al (1997) The mouse fused locus encodes axin, an inhibitor of the Wnt signaling pathway that regulates embryonic axis formation. Cell 90:181–192

Directed mESC Differentiation 6. Liu P, Wakamiya M, Shea MJ et al (1999) Requirement for Wnt3 in vertebrate axis formation. Nat Genet 22:361–365 7. Vincent SD, Dunn NR, Hayashi S et al (2003) Cell fate decisions within the mouse organizer are governed by graded nodal signals. Genes Dev 17:1646–1662 8. Dunn NR, Vincent SD, Oxburgh L et al (2004) Combinatorial activities of Smad2 and Smad3 regulate mesoderm formation and patterning in the mouse embryo. Development 131:1717–1728 9. Chu GC, Dunn NR, Anderson DC et al (2004) Differential requirements for Smad4 in TGFβ-dependent patterning of the early mouse embryo. Development 131:3501–3512 10. Kubo A, Shinozaki K, Shannon JM et al (2004) Development of definitive endoderm from

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embryonic stem cells in culture. Development 131:1651–1662 11. Irion S, Nostro MC, Kattman SJ et al (2008) Directed differentiation of pluripotent stem cells: from developmental biology to therapeutic applications. Cold Spring Harb Symp Quant Biol 73:101–110 12. Gadue P, Huber TL, Paddison PJ et al (2006) Wnt and TGF-β signaling are required for the induction of an in vitro model of primitive streak formation using embryonic stem cells. Proc Natl Acad Sci U S A 103:16806–16811 13. Ying QL, Smith AG (2003) Defined conditions for neural commitment and differentiation. Methods Enzymol 365:327–341 14. Ying QL, Stavridis M, Griffiths D et al (2003) Conversion of embryonic stem cells into neuroectodermal precursors in adherent monoculture. Nat Biotechnol 21:183–186

Methods in Molecular Biology (2022) 2520: 309–319 DOI 10.1007/7651_2021_437 © Springer Science+Business Media, LLC 2021 Published online: 06 October 2021

Accessing the Human Pluripotent Stem Cell Translatome by Polysome Profiling Rubens Gomes-Ju´nior, Patrı´cia Shigunov, Bruno Dallagiovanna, and Isabela Tiemy Pereira Abstract Polysome profiling is a technique that uses sucrose density gradient ultracentrifugation to separate complexes of mRNAs associated with one or more ribosomes. Here we describe polysome profiling analysis in human pluripotent stem cells (hPSCs) using a continuous ultraviolet spectrophotometer and a gradient fractionator. We provide protocols for processing sucrose gradient fractions for isolation of RNA for RT-qPCR or large-scale sequencing analysis, used to establish the translational status of specific mRNAs and identify the role of noncoding RNA in translation. Key words Polysome profiling analysis, Pluripotent stem cells, Sucrose density gradient, Ribosome, RNA isolation

1

Introduction Post-transcriptional regulation is a mechanism for controlling gene expression, for example by potentiating or inhibiting the translation of specific mRNAs. The mRNAs have different translation efficiencies and undergo modulations according to the activation of cellular processes such as proliferation, pluripotency maintenance, differentiation, and cell death, which are triggered by internal and environmental factors. The mRNAs can be actively translated when associated with multiple ribosomes, in structures called polysomes, polyribosomes, or translating ribosomes [1]. Most translational responses alter the density of ribosomes at a given mRNA [2], and the number of ribosomes, among other features, allows inferring its translation status. The abundance of proteins in mammalian cells is highly controlled through translation and beyond transcript concentration [3–5]. Stem cell self-renewal and differentiation are the result of dynamic cellular environments that must tightly adjust protein levels, with translational regulation being a crucial step in the control of gene expression [6–9]. For instance, the translational

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output largely contributes to the pluripotent chromatin state of embryonic stem cells (ESC) [10]. Polysome fractionation by sucrose density gradient centrifugation followed by the analysis of RNA and protein constitutes a methodology that allows understanding the changes in the translation of individual mRNAs as well as genome-wide effects on the translatome [11]. The detachment of ribosomes from the aggregates is strongly inhibited by cycloheximide [12]. A high concentration of cycloheximide is used to freeze ribosomes on the translating mRNAs, followed by fractionation in a sucrose density gradient centrifugation to separate populations of mRNAs with various numbers of ribosomes loaded [13]. The mass of mRNP complexes increases as a function of the number of ribosomes attached. Thus, more actively translated mRNAs, which means heavier mRNPs, sediment further in the gradient [14]. Human pluripotent stem cells (hPSCs), embryonic and induced pluripotent stem cells, are delicate to handle and the relative intensity of the polysome peaks are lower than that of other human cells. Nevertheless, they still provide data allowing to establish the translational status of specific mRNAs as well as identify the role of noncoding RNA in translation, among others. Here, we provide protocols for processing hPSCs and sucrose gradient fractions for isolation of RNA and RT-qPCR analysis (Fig. 1). Using cycloheximide and puromycin treatment, we show both the usual polysome profile as well as the profile with disassembled polysomes as a control. RT-qPCR analysis of POU5F1 was performed as an example of an actively translated gene in the pluripotent condition (Fig. 2).

2

Materials All solutions described below must be prepared using ultrapure RNase-free water and all instruments should be clean. The solutions can all be prepared in advance of time and stored accordingly.

2.1 Sucrose Gradients

1. Polysome buffer: 15 mM Tris–HCl, 15 mM MgCl2, 0.3 M NaCl, pH 7.4. Store at 4
C. 2. Sucrose 50%: 50 g of sucrose in polysome buffer up to 100 mL. Store at 4
C. 3. Sucrose 10%: 10 g of sucrose in polysome buffer up to 100 mL. Store at 4
C. 4. Cycloheximide: stock solution at 10 mg/mL in water. Store at 20
C.

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Fig. 1 Polysome profiling analysis of hPSC. (a) Representative image of hPSC in culture. (b) Schematic illustration of sucrose gradient showing distribution of mRNP complex sedimentation by ultracentrifugation. (c) Polysome profiles of hPSCs treated with cycloheximide and puromycin, recorded by an ultraviolet spectrophotometer at 254 nm

Fig. 2 Relative polysome engagement of POU5F1. Percentage of fraction distribution of POU5F1 transcripts throughout the sucrose gradient. Polysome profile fractions were analyzed by RT-qPCR. POU5F1 is actively translated in hPSC, enriched in light-polysome fractions in cycloheximide treated cells. Once polysomes are disassembled with puromycin treatment, POU5F1 transcripts shifted leftward to the lighter gradient fractions

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5. Gradient tubes: (14  95 mm).

model

SW40

or

with

similar

size

6. Gradient system: Gradient Master (model 108)—Biocomp. 2.2

Cell Procedure

1. Phosphate-buffered saline (PBS) 1: 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4 and 1.8 mM KH2PO4. Store at 4
C. 2. Accutase®—Innovative Cell Technologies (see Note 1). 3. Puromycin: stock solution at 10 mg/mL in water. Store at 20
C 4. Lysis buffer: Polysome buffer containing 1% Triton X-100 (see Note 2). For each sample, prepare 400 μL of lysis buffer supplemented with 1 μL of RNaseOUT™ (40 U/μL), 1 μL of deoxyribonuclease I (50 U/μL) (see Note 3), and 4 μL of cycloheximide (10 mg/mL) or 40 μL of puromycin (10 mg/ mL). Keep on ice (see Note 4). 5. Ultracentrifuge.

2.3

Fraction Isolation

1. Density Gradient Fractionation System—Teledyne ISCO. 2. Dyed-sucrose 65%: 65 g of sucrose in polysome buffer up to 100 mL. Add bromophenol blue enough to make the solution turn blue (see Note 5). Store at 4
C.

2.4 RNA Extraction and cDNA Synthesis

1. Direct-zol™ RNA Miniprep kit—Zymo Research (see Note 6). 2. TRI reagent®—Merck or similar acid-guanidinium-phenol reagent. 3. Ethanol (95–100%). 4. ImProm-II™ Reverse Transcription System (Promega) or similar reverse transcription system. 5. Synthetic RNA spike.

2.5 RT-qPCR Analysis

1. GoTaq® qPCR Master Mix (Promega) or your preferred chemistry. 2. LightCycler® instrument.

3

96

System

(Roche)

or

any

compatible

Methods Before starting the protocol, clean all workspaces and equipment to maintain an RNase-free environment.

3.1 Preparation of Sucrose Gradients

1. Bring 50 and 10% sucrose solutions to room temperature (see Note 7) and thaw cycloheximide stock on ice.

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2. Freshly prepare 50 and 10% sucrose solutions supplemented with 0.1 mg/mL cycloheximide. To make two gradients, add 130 μL of cycloheximide stock in 13 mL of 50% sucrose, and 150 μL of cycloheximide stock in 15 mL of 10% sucrose, mix thoroughly. For puromycin control, the gradients are made without adding cycloheximide to the sucrose solutions, nor puromycin. 3. Set up the gradient tubes first adding 5.9 mL of 50% sucrose solution to the bottom. Very carefully add 10% sucrose solution on top using a syringe with needle or 1 mL automatic pipette until the gradient tube is full (see Note 8). 4. Cap the tube using the short cap with the air hole facing up, allowing the bubble to escape (see Note 9). A line between both sucrose solutions should be visible. 5. Level the Gradient Master tube holder and balance the tubes. Set the program to: SHORT SUCR 10–50% WW 1st (Time 1:55; Angle 81.5
; Speed 25 rpm; 1 cycle) and run (see Note 10). 6. Once the gradient is done, the line between the solutions is not visible because they have been mixed. Store the gradients in a tube holder at 4
C until lysates are ready to be loaded. 3.2 CycloheximideTreated Cells and Lysis (See Note 11)

1. Freshly prepare the cycloheximide lysis buffer and keep it on ice. 2. When the cell culture is ready for use (Fig. 1a), add 0.1 mg/mL cycloheximide to the cell medium. 3. Incubate for 10 min at 37
C. 4. Aspirate and discard the medium, wash the cells with 1 PBS and dissociate them using Accutase. 5. Wash the cell pellet twice using 1 PBS supplemented with 0.1 mg/mL cycloheximide. 6. Add the lysis buffer and gently pipet up and down to disrupt the cells. 7. Incubate the cell lysate on ice for 10 min. 8. Centrifuge the cell lysate to 12,000  g for 10 min, at 4
C. 9. Save the supernatant to load on the gradient (see Subheading 3.4).

3.3 PuromycinTreated Cells and Lysis (See Note 11)

1. Freshly prepare the puromycin lysis buffer and keep it on ice. 2. When the cell culture reaches 80% confluence, add 1 mg/mL puromycin to the cell medium. 3. Incubate for 1 h at 37
C.

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4. Aspirate and discard the medium, wash the cells with 1 PBS and dissociate them using Accutase. 5. Wash the cell pellet twice using 1 PBS. 6. Add the lysis buffer and gently pipet up and down to disrupt the cells. 7. Incubate the cell lysate on ice for 10 min and then for 20 min at 37
C. 8. Centrifuge the cell lysate at 12,000  g for 10 min, at 4
C. 9. Save the supernatant to load on the gradient (see Subheading 3.4). 3.4 Setting Up the Gradient

1. To load the cell lysate supernatant on the gradient, first open the cap and carefully remove 400 μL of sucrose solution from the very top of the gradient. 2. Load slowly the cell lysate supernatant over the gradient. 3. Balance the tubes by weighing them (see Note 12). Add polysome buffer if necessary. 4. Centrifuge at 39,000 rpm (270,000  g) for 2 h, at 4
C (see Note 13).

3.5 Setting Up the Fractionation System

1. Twenty minutes before the end of the centrifugation, connect the fractionation system (detector, pump, and chart recorder) and turn on the UV lamp to start warming it up (see Note 14). 2. Flush the entire system, letting the RNase-free water pass through to the final sample collector. 3. Turn on the peristaltic pump to flow 65% dyed-sucrose through the system to the needle of the gradient tube holder. 4. Identify twenty-two 1.75 mL microcentrifuge tubes and prepare an ice bucket. 5. Set the baseline (see Note 15).

3.6 Polysome Fractionation and Sample Collection (See Note 16)

1. Carefully remove the gradients from the centrifuge and keep them on ice. 2. Load one gradient tube on the fractionation system by securing it into the top holder and raising the bottom holder (see Note 17). 3. Place the labeled 1.75-mL microcentrifuge tubes on the sample collector rack (see Note 18). 4. Turn on the fractionation system (see Note 19). The sample will be pushed through the detector and fractionated. 5. Collect the fractions in the microcentrifuge tubes as they exit and place them on ice. The polysome profile will be recorded in the chart recorder.

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Downstream Analysis of Polysome Profile and Fractions Many downstream analyzes on the association of specific proteins, RNA and ribosomes can be performed using the polysome profile and sample fractions. Among others, semiquantitative observations of the overall level of translational activity in the cell might be carried out (see Note 20). Here, we will focus on the relative engagement on translation of a particular RNA, specifically describing RNA isolation and RT-qPCR protocols (see Note 21).

4.1 Extraction of RNA from Sucrose Gradient Fractions (See Note 22)

1. Add an equal volume of TRI reagent® or similar acidguanidinium-phenol reagent to each fraction or pooled fractions (see Note 21). 2. Add an equal volume of ethanol (95–100%) to each sample (fraction(s) + TRI reagent®) and mix thoroughly. For instance, if you are working with pooled heavy-polysome fractions to a final volume of 5 mL, add 5 mL of TRIzol and then 10 mL of ethanol. 3. Transfer the mixture into the kit spin column in a collection tube and centrifuge at 10,000–16,000  g for 30 s (see Note 23). Discard the flow-through. 4. Add 400 μL PreWash Buffer to the column and centrifuge at 10,000–16,000  g for 30 s. Discard the flow-through and repeat this step. 5. Add 700 μL Wash Buffer to the column and centrifuge at 10,000–16,000  g for 2 min to ensure complete removal of the wash buffer. 6. Transfer the column carefully into an RNase-free microcentrifuge tube and elute the RNA adding 25–50 μL of DNase/ RNase-Free water directly to the column matrix and centrifuge (see Note 24). 7. Use the RNA immediately or store appropriately at 80
C.

4.2

cDNA Synthesis

1. Use an equal volume of each RNA fraction/sample to perform the cDNA synthesis (see Note 25). 2. It is recommended to add an RNA spike to use as a control for amplification (see Note 26), since regular endogenous controls such as GAPDH are not applicable in this analysis (see Note 27).

4.3 RT-qPCR Analysis

1. Perform qPCR reaction to amplify specific transcripts using gene-specific forward and reverse primer mix and your desired chemistry. Use primer sets for spike control amplification as well.

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2. The data analysis can be performed by subtracting the Ct value of the target gene from the spike control of each sample (ΔCt). Then, calculate the “fold to spike” using the formula ½^ΔCt to determine a value of relative expression (Vx) of the target to the spike control in each fraction or pool (x). Sum the Vx for all fractions to calculate the total value of relative expression (Vt) in one condition (e.g., cycloheximide) and, finally, calculate the proportion of the target gene found in a fraction or pool by P ¼ 100  Vx/Vt. 3. The proportion of a transcript found in each fraction or pool is represented as a percentage. Actively translated RNAs are expected to be enriched in light and heavy-polysome fractions, in cycloheximide treated cells. When comparing cycloheximide and puromycin treated cells, polysome-associated RNAs would display a leftward shift on the gradient fractions in puromycin conditions, due to polysomes disassemble (Fig. 2).

5

Notes 1. We recommend using Accutase, but you can use a similar detachment solution. 2. The polysome buffer might be premixed with Triton X-100 and store at 4
C. 3. We recommend adding DNase in the lysis buffer because the nucleus of hPSC can be disrupted in this process, and the releasing of the DNA might interfere with the fractionation. 4. RNaseOUT, DNase I and cycloheximide or puromycin must be freshly added to the lysis buffer before use. 5. To facilitate the solubilization, the solution can be warmed at 37
C while mixing. Dying this solution will help to track the sample movement throughout the fractionation system. 6. We recommend this RNA extraction kit, which has the best efficiency for sucrose-based solution. 7. If the sucrose solutions are cold, they form bubbles on the wall of the gradient tube and change the viscosity of the solution, which impairs the gradient formation. Always use them at room temperature. 8. Touch the gradient tube wall with the needle or pipette tip close to the liquid surface and dispense the 10% sucrose solution very slowly and carefully to not disturb the 50% sucrose solution. Ideally, both sucrose solutions should not mix up at this point.

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9. Always check the sucrose solutions for bubbles as they impair the gradient formation. If there are any bubbles, remove the cap to safely eliminate them. 10. Handle the tubes carefully during the preceding and following steps in the protocol to not disrupt the gradient. 11. At least 3  106 cells (an area of 9 cm2 of hPSC monolayer culture 80% confluent) are needed for each gradient. 12. Balance the tubes by weight before any ultracentrifugation. Please refer to the best practices of your ultracentrifuge. 13. Set up acceleration and brake to 9. Prechill the rotor and buckets at 4
C before use. 14. Turn on the UV lamp at least 15–30 min before use to warm it up. The indicator light usually starts red and turns green when ready. 15. The baseline should be around the middle of the chart to allow it record ups and downs. Adjust it using the Recorder Offset or Baseline adjust. Set up the Sensitivity to 1.0. 16. The polysomal complexes can be visualized using a UV spectrophotometer. The sucrose density gradient under ultracentrifugation allows the separation of cytoplasmic RNA mainly based on the number of ribosomes bound. Peaks corresponding to the 80S monosome and increasing numbers of polysomes should be distinguishable. For human pluripotent stem cells, the relative intensity of these peaks will be lower than from other human cells. 17. Make sure the tube is tightly capped/connected, otherwise the sample can leak out. 18. Typically, samples start to drop around the tube at the sixth position. Place just one or two microcentrifuge tubes and wait for the samples start dropping to place the other tubes appropriately. 19. Configure the pump velocity to 50, chart speed of chart recorder to 60, start at the touchscreen. 20. Since the amount of polysomes reflects the overall level of translational activity in the cell, the relative area under the peaks of the polysome profile chart can be determined. The ratios of these areas over the total area under the peaks can be used as a comparison between distinct conditions to make semiquantitative observations. For instance, the polysome– monosome area ratio can be quantified by calculating the area beneath the polysome and monosome peaks using ImageJ software and be used to infer the global translational state in distinct cell conditions. It is not recommended to compare absolute absorbance intensities between samples because

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there can be a lot of variability, even when loading the same relative amount of sample per gradient. 21. Polysome profiling comprises the fractionation of samples according to the association with no ribosomes or varying numbers of ribosomes, including fractions without ribosomal material (ribosome-free), fractions with 80S (monosomes) and light- and heavy-polysome fractions, which can be identified on the polysome profile chart (Fig. 1b, c). For the analysis, it is possible to process each fraction individually or, alternatively, pool them into main subclasses (e.g., ribosome-free, monosome, light polysomes, and heavy polysomes). The relative distribution of a specific RNA along the gradient shows, for example, whether it is more abundant in the actively translating fractions (light/heavy polysomes) or not. The isolated RNA is also applicable to large-scale sequencing analysis. 22. The RNA extraction protocol was adapted due to the high volume of sucrose in the solution. 23. If you end up working with high volumes of samples, one spin column of the kit might be loaded multiple times with the same sample before going to the next step. After loading the whole sample volume into the spin column, proceed to the next steps according to the kit datasheet. 24. To ensure a suitable final concentration, elute the RNA with 25 μL of DNase/RNase-Free water. The final volume can be reduced even more when working with individual fractions. Once the ideal final volume is defined, it is mandatory to keep the same volume for all fractions/samples. 25. We recommend using 5 pg of RNA spike per cDNA synthesis reaction (total volume: 20 μL). 26. Any cDNA synthesis enzyme or kit might be used. To analyze a particular class of RNA, such as miRNA or non-poly-A RNAs, proceed with a specific appropriate kit. It might be necessary to use the whole final volume of RNA when working with individual fractions, to ensure suitable synthesis for further amplifications. 27. Frequent endogenous controls, such as GAPDH or POLR2A transcripts, might be differentially distributed between polysomal fractions and cannot be used for normalization in RT-qPCR analysis. The RNA spike chosen must not be present in any fraction or condition. Appropriate controls are, for instance, green fluorescent protein (GFP) or firefly luciferase (FLuc) RNAs synthesized in vitro. Alternatively, the RNA spike can be added directly to the sucrose fractions or their pool prior RNA extraction, as a control for RNA recovery.

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References 1. Warner JR, Knopf PM, Rich A (1963) A multiple ribosomal structure in protein synthesis. Proc Natl Acad Sci U S A 49:122–129. https://doi.org/10.1073/pnas.49.1.122 2. King HA, Gerber AP (2016) Translatome profiling: methods for genome-scale analysis of mRNA translation. Brief Funct Genomics 15(1):22–31. https://doi.org/10.1093/ bfgp/elu045 3. Schwanh€ausser B, Busse D, Li N, Dittmar G, Schuchhardt J et al (2011) Global quantification of mammalian gene expression control. Nature 473:337–342. https://doi.org/10. 1038/nature10098 4. Tahmasebi S, Amiri M, Sonenberg N (2019) Translational control in stem cells. Front Genet 10:1–9. https://doi.org/10.3389/fgene. 2018.00709 5. Gabut M, Bourdelais F, Durand S (2020) Ribosome and translational control in stem cells. Cells 9(2):497–527. https://doi.org/ 10.3390/cells9020497 6. Blair JD, Hockemeyer D, Doudna JA et al (2017) Widespread translational remodeling during human neuronal differentiation. Cell Rep 21(7):2005–2016. https://doi.org/10. 1016/j.celrep.2017.10.095 7. Fujii K, Shi Z, Zhulyn O et al (2017) Pervasive translational regulation of the cell signalling circuitry underlies mammalian development. Nat Commun 8:1–13. https://doi.org/10. 1038/ncomms14443 8. Kristensen AR, Gsponer J, Foster LJ (2013) Protein synthesis rate is the predominant

regulator of protein expression during differentiation. Mol Syst Biol 9:689–700. https:// doi.org/10.1038/msb.2013.47 9. Lu R, Markowetz F, Unwin RD et al (2009) Systems-level dynamic analyses of fate change in murine embryonic stem cells. Nature 462 (7271):358–362. https://doi.org/10.1038/ nature08575 10. Bulut-Karslioglu A, Macrae TA, Oses-Prieto JA et al (2018) The transcriptionally permissive chromatin state of embryonic stem cells is acutely tuned to translational output. Cell Stem Cell 22(3):369–383.e8. https://doi. org/10.1016/j.stem.2018.02.004 11. Zuccotti P, Modelska A (2016) Studying the translatome with polysome profiling. Methods Mol Biol 1358:59–69. https://doi.org/10. 1007/978-1-4939-3067-8_4 12. Godchaux W 3rd, Adamson SD, Herbert E (1967) Effects of cycloheximide on polyribosome function in reticulocytes. J Mol Biol 27 (1):57–72. https://doi.org/10.1016/00222836(67)90351-8 13. He SL, Green R (2013) Polysome analysis of mammalian cells. Methods Enzymol 530:183–192. https://doi.org/10.1016/ B978-0-12-420037-1.00010-5 14. Pringle ES, McCormick C, Cheng Z (2019) Polysome profiling analysis of mRNA and associated proteins engaged in translation. Curr Protoc Mol Biol 125(79):1–13. https://doi. org/10.1002/cpmb.79

Methods in Molecular Biology (2022) 2520: 321–333 DOI 10.1007/7651_2022_461 © Springer Science+Business Media, LLC 2022 Published online: 18 May 2021

Genome Engineering Human ESCs or iPSCs with Cytosine and Adenine Base Editors Giulia Pavani, Joshua G. Klein, Deborah L. French, and Paul Gadue Abstract The ability to engineer specific mutations in human embryonic stem cells (ECSs) or induced pluripotent stem cells (iPSCs) is extremely important in the modeling of human diseases and the study of biological processes. While CRISPR/Cas9 can robustly generate gene knockouts (KOs) and gene loci modifications in coding sequences of iPSCs, it remains difficult to produce monoallelic mutations or modify specific nucleotides in noncoding sequences due to technical constraints. Here, we describe how to leverage cytosine (BE4max) and adenine (ABEmax) base editors to introduce precise mutations in iPSCs without inducing DNA double-stranded breaks. This chapter illustrates how to design and clone gRNAs, evaluate editing efficiency, and detect genomic edits at specific sites in iPSCs through the utilization of base editing technology. Key words Genome editing, Induced pluripotent stem cells (iPSC), Disease modeling, Base editing, Gene correction

1

Introduction This protocol describes how to generate specific genome edits in iPSCs using cytosine (BE4max) and adenine (ABEmax) base editors developed by Koblan et al. [1]. These base editors (BEs) are generated by fusing a spCas9 nickase to a cytidine (BE4max) or to an adenosine (ABEmax) deaminase domain to promote C > T or A > G changes, respectively, in the genome. Since gRNAs rely on spCas9 for targeting, they can be designed to edit a specific genomic location. BE4max and ABEmax preferentially target cytosines and adenines present between the fourth and the eighth bases of the gRNA protospacer, as highlighted in Fig. 1. Adjacent bases can also be targeted, but with lower efficiency. Over the years, several new BEs with various editing capabilities, protospacer adjacent motif (PAM) requirements, and accuracies have been developed [2]. Each BE will have different editing windows and efficiencies, so it is important to choose a BE best suited for the desired application and design the targeting gRNA accordingly. This protocol can also be used with alternative editors if the BE is spCas9based.

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Fig. 1 Schematic representation of BE4max (left) and ABEmax (right) and their respective editing windows. BE activity window is shown as a gray box (dark and light gray indicate higher and lower editing efficiency, respectively). The location of a DNA nick in the target sequence is indicated by an arrow. Base positions are numbered relative to the PAM end (NGG) of the guide RNA. APOBEC cytidine deaminase domain, UGI uracil DNA glycosylase inhibitor, D10A Cas9 nickase Cas9, TadA adenosine deaminase domain, sgRNA single guide RNA

Editing with BEs has multiple advantages. In addition to modeling single nucleotide variants, cytosine BEs can also generate gene knockouts (specifically, by changing glutamine codons to stop codons). One of the main benefits of using BEs over classic Cas9 editing, beside the absence of DNA double-stranded breaks, resides in their capability to produce both heterozygous and homozygous edits. If a sufficient number of clones are screened, it is feasible to isolate individual iPSC lines from the same editing experiment containing single and biallelic modifications. This results in a faster turnaround time for the generation of specific isogenic iPSC lines and controls, without the risk of inducing large genomic deletions and/or translocations.

2

Materials

2.1 gRNA Cloning and Base Editor Plasmids

pCMV ABEmax-P2A-GFP plasmid (Addgene, 112101). pCMV BE4max-P2A-GFP plasmid (Addgene, 112099). phU6-gRNA plasmid (Addgene, 53188). ATP (NEB, P0756S). T4 Polynucleotide Kinase (NEB, M0201S).

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T4 DNA ligase (M0202S). FastAP (Thermo Scientific, EF0654). FastDigest BpiI (BbsI, Thermo Scientific, FD1014). Agarose (e.g., VWR N605). Gel extraction kit (e.g., Macherey-Nagel 740609). Kanamycin (50 μg/ml) LB-Agar plates. LB broth (Miller) (Thermo Scientific, L3522). Kanamycin Sulfate (Thermo Scientific, 11815024). Competent cells: for example, Stellar™ Escherichia coli cells (Takara 636763). Miniprep plasmid isolation kit: for example, Purelink™ Quick Plasmid Miniprep Kit (Invitrogen K210011). PCR strip tubes. M13F primer (GTAAAACGACGGCCAG). Additional reagents and equipment for agarose gel electrophoresis. 2.2 Efficiency Testing and Clonal Screening

Corning® Matrigel® Growth Factor Reduced (GFR) Basement Membrane Matrix (354230). Induced pluripotent stem cells (iPSCs) or embryonic stem cells (ESC), human murine embryonic fibroblasts, irradiated (MEF). Base editor-GFP plasmid, filtered 0.22 μm (recommended concentration > 500 μg/ml). Guide RNA plasmid, filtered 0.22 μm (recommended concentration > 500 μg/ml). Corning Dulbecco’s Modified Eagle’s Medium/Ham’s F-12 50:50 mix with L-glutamine and 15 mM HEPES (DMEM/F12) (Ref 10-092-CV). ThermoFisher® Lipofectamine Stem Reagent. Human Embryonic Stem Cell (hESC-10) media containing 10 ng/ ml basic fibrinogen growth factor (bFGF), sterile. Y-27632 dihydrochloride/ROCK inhibitor (ROCKi). Gibco StemPro® Accutase® (REF A11105-01). Sterile 1.5 ml microcentrifuge tubes. Sterile 5 ml round-bottom tubes with cell strainer caps. 6-well multiwell sterile plate (e.g., Falcon 353046). Sterile 10 cm dishes and 12-well plates. Fluorescence-activated cell sorter (FACS) apparatus. Sterile 15 ml conical centrifuge tubes.

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Sterile 12-well and 6-well plates. Gibco TrypLE™ Express (REF 12605010). Corning® Iscove’s modification of Dulbecco’s medium (IMDM). Lucigen QuickExtract™ (Cat. No. QE0905T). KAPA2G Fast ReadyMix (Roche) + PCR Primers. Invitrogen™ TOPO™ TA Cloning™ Kit for Sequencing (Cat. No. 450030). SOC Medium (Takara). Competent cells: for example, Stellar™ Escherichia coli cells (Takara, 636763). Kanamycin (50 μg/ml) LB-Agar plates. Sterile PCR tubes. Agarose (e.g., VWR N605). Additional reagents and equipment for agarose gel electrophoresis.

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Methods

3.1 gRNA Design and Cloning

There are multiple available programs for designing gRNA and predicting BE outcomes in different genomes (https://www. crisprbehive.design [3]; http://www.rgenome.net/be-designer/ [4]; https://fgcz-shiny.uzh.ch/PnBDesigner/). An example of anticipated BE outcomes generated by the BeHive algorithm is shown in Fig. 2. It is important to note that predictions consider allele edits as independent events; however, the combination of edited alleles within a cell can be affected by several factors such as gRNA efficiency, delivery method, and cell state. When possible, choose gRNAs with high efficiency and the lowest number of targetable bases (C or A) in the editing window.

Fig. 2 Representative outcomes of BeHive prediction algorithm for cytosine (left) or adenine (right) base editors. Highlighted bases are potential base editor targets, and the darker color indicates a higher probability of editing. Predicted outcomes are scored and ranked according to efficiency scores. Base positions are numbered relative to the PAM end (NGG) of the guide RNA

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A dedicated prediction algorithm is also available when targeting a splice site for gene modulation or knockout (https:// moriaritylab.shinyapps.io/splicer/ [5]). This cloning protocol uses a gRNA expression plasmid with a hU6 promoter (Addgene #53188) from Kabadi et al. [6] (see Note 1). 1. To clone the gRNA target sequence under the control of the hU6 promoter, overhangs should be added to DNA oligos according to the scheme below. sense

5' CACCG(N)20

antisense 3'

5'

C(N)20 complement CAAA 5'

where (N)20 is your guide RNA target sequence. 2. To anneal sense and antisense oligos, add to a PCR tube. 8 μl sense oligo (10 μM) 8 μl antisense oligo (10 μM) 2 μl 10 NEB ligase buffer After mixing, proceed with the annealing step in a thermocycler. 3. Heat the oligos at 96  C for 5 min, then cool to 25  C using a 0.3  C/s ramp (see Note 2). 4. Phosphorylate overhangs by adding 1 μl 25 mM ATP and 1 μl T4 NEB Polynucleotide Kinase to the same tube. 5. Incubate at 37  C for 60 min, then heat inactivate at 65  C for 20 min. The oligos are now ready to be ligated in the gRNAexpressing plasmid. 6. Before proceeding with ligation, hU6 plasmid should be digested and linearized with BpiI (BbsI) enzyme. Set up the digestion in a PCR tube according to this Scheme. 2.5 μg of hU6 plasmid 2 μl BpiI (BbsI) 3 μl of Enzyme Buffer Up to 30 μl of H2O 7. Incubate at 37  C for 1 h. 8. Add 1 μl of FastAP to dephosphorylate the linearized backbone. 9. Incubate for 1 h at 37  C, then 20 min at 65  C to deactivate the phosphatase. 10. Run the digestion in a 1% agarose gel and cut the ~3500 bp band corresponding to the linearized hU6 plasmid.

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11. Purify the linearized plasmid according to manufacturer’s instructions, and elute in 15–20 μl of H2O. 12. Quantify the linearized backbone before proceeding to ligation. 13. Set up the ligation, including a ligation control using the same linearized plasmid without adding the annealed oligos. 1 μl T4 DNA buffer 1 μl T4 DNA Ligase 1 μl annealed oligos (or H2O for control) 25–50 ng of digested hU6 plasmid Up to 10 μl of H2O 14. Incubate for 15 min at RT, then 16  C for at least 60 min. 15. After ligation, transform 50 μl of competent cells using 5 μl of the ligation mixture according to manufacturer’s instructions. 16. Plate transformed bacteria on Kanamycin LB-Agar plates. 17. Incubate at 37  C overnight. 18. Check ligation efficiency by counting colonies in your ligation plates vs control plates. You should have at least twofold more colonies on ligation plates. 19. Pick 2–4 colonies per plate and grow them in individual conical tubes containing 5 ml of LB-Kanamycin on a shaker at 37  C, 200 rpm. 20. Purify plasmid from the miniprep according to manufacturer’s instructions. 21. Confirm the presence of the gRNA protospacer by Sanger sequencing the plasmid with the M13F primer. 3.2 iPSC Lipofection and Sorting

1. Feed iPSC with hESC-10 media at least 2 h before lipofection. 2. For each well of cells to be transfected, reserve two microcentrifuge tubes and add 25 μl room temperature DMEM-F12 media to each. Media must be serum and antibiotic-free, and Opti-MEM can also be used instead of DMEM-F12. 3. To Tube 1, add 0.5–1 μg of the guide RNA plasmid and 1–2 μg of the base editor-GFP plasmid (see Note 3). To Tube 2, add 5 μl of Lipofectamine Stem. 4. Gently transfer the contents of Tube 1 into Tube 2 by pipetting (see Note 4). 5. Let the mixture rest for 10 min at room temperature. 6. Add the 50 μl reaction mixture, dropwise, to a single well of iPSCs in hESC-10 media. 7. Culture the cells for 48 h at 37  C and 5% oxygen, changing hESC-10 media daily.

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8. Prepare a 10 cm dish plate with MEF feeder cells the day before sorting. If planning on conducting bulk sequencing, also prepare a 12-well plate with MEF feeder cells. 9. 48 h after lipofection, aspirate hESC-10 off of the lipofected wells. 10. Wash cells once with 2 ml room-temperature DMEM-F12 per well. Aspirate DMEM-F12. 11. Add 1 ml room temperature Accutase to each well. Other dissociation reagents, such as TrypLE, can be used; we found that Accutase was the most efficient in generating single cells and maintaining high cell viability. 12. Incubate at 37  C until the cells have detached from the well (do not leave cells in Accutase for longer than 15 min). 13. Transfer the cells to a 15 ml conical centrifuge tube containing 10 ml of room-temperature DMEM-F12. 14. Pellet the cells via centrifugation for 3 min at 330  g. 15. Aspirate supernatant from the cell pellet, and resuspend the cells in 0.5 ml hESC-10 media +10 μM ROCKi. If multiple iPSC wells were lipofected with identical plasmids, the resuspended cells can now be combined. 16. Filter the resuspended cells through a cell strainer cap into a 5 ml round-bottom tube. Keep resuspended cells on ice until ready to sort. 17. Prepare a collection 15 ml centrifuge tube containing 1.5 ml hESC-10 media + ROCKi. 18. Sort GFP-positive cells using a fluorescence-activated cell sorter (see Note 4). A representative flow plot of the GFP-positive cell sort is shown in Fig. 3.

Fig. 3 Representative flow plot with sorting gate. Cells expressing a high level of GFP are collected to maximize editing efficiency in sorted cells

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3.3 Measuring Editing Efficiency

1. After sorting, plate 10,000 cells in 10 ml hESC-10 + ROCKi on a MEF-coated 10 cm dish (see Note 5). If desired, plate 1000–5000 cells in 1 ml hESC-10 + ROCKi in a single well of a MEF-coated 12-well plate to measure editing efficiency before picking colonies (optional: see steps 3–12). 2. Culture cells with hESC-10 media until visible colonies form. Depending on the number of cells plated, this should happen around day 10–14. 3. Optional (steps 3–12): If plating on a 12-well, culture cells for approximately 5–7 days until small colonies can be seen. 4. Aspirate hESC-10 media from iPSCs, and wash once with 1 ml room-temperature IMDM. 5. Aspirate IMDM, then add 0.5 ml room temperature TrypLE to the well. Let stand for 3 min at room temperature. 6. Aspirate TrypLE, then gently wash the well twice with 1 ml IMDM. 7. Using a cell scraper, scrape cells into 1 ml IMDM. Transfer the cell suspension to a 1.5 ml microcentrifuge tube. 8. Pellet the cells using a tabletop centrifuge, then resuspend the cell pellet in 25 μl QuickExtract™. 9. Incubate at 65  C for 15 min, then 95  C for 3 min. 10. Dilute extracted DNA solution with 50 μl of PCR grade water. 11. Use 5 μl of diluted DNA mixture to perform PCR, and analyze via Sanger sequencing. Include an unedited sample as a control when performing PCR and sequencing (see Notes 6 and 7). 12. Analyze chromatogram to measure editing efficiency via EditR [7]. EditR is a free online software and R-based package, designed to quantitively calculate base editing efficiency from Sanger sequencing chromatogram data (https://moriaritylab. shinyapps.io/editr_v10/) (see Note 8).

3.4 Subcloning and Genotyping

1. At least 12 h before picking colonies from the sorted cells, prepare MEF-coated 12 well plates. 2. Wash MEFs and replace media with 1 ml hESC-10 media containing 10 μM ROCKi right before picking colonies. 3. Using a microscope under a laminar flow hood, pick sizeable colonies into individual sterile PCR tubes in 80 μl hESC10 + ROCKi. Gently pipette up and down 2–3 times to thoroughly resuspend the colony. To maintain pluripotency, pick colonies before the colony shows a phase-bright center. 4. Transfer 50 μl of the cell suspension onto a MEF-coated well.

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5. Pellet the remaining 30 μl of the cell suspension using a tabletop centrifuge and aspirate the supernatant. 6. Resuspend pellets in 15 μl QuickExtract. 7. Incubate at 65  C for 15 min, then 95  C for 3 min. 8. Dilute DNA with 30 μl of PCR grade water. The DNA solution can be stored long term at 20  C. 9. Use 5 μl of diluted DNA mixture to perform PCR, and analyze via Sanger sequencing. Include an unedited sample as a control when performing PCR and sequencing (see Notes 6 and 7). 10. Analyze individual clones via Sanger sequencing and confirm edits using an alignment software such as SnapGene or Benchling (see Note 9). For a rapid qualitative analysis of base editing efficiency, the chromatogram can be directly analyzed within the editing window; however, free software such as CRISP-ID (http://crispid.gbiomed.kuleuven.be) [8] may also be utilized. Table 1 Table displays results from three genes edited with the pCMV-BE4max-P2A-GFP plasmid. While the overall editing efficiency varied depending on the target gene, base editing was observed in all three genes after sequencing. Both homozygous and heterozygous edited clones were obtained in all three editing experiments. Notably, one editing experiment also yielded clones containing indels and/or transversions (see Note 10) BE4max editing efficiencies # Clones edited # Clones edited # Clones edited # Clones Guide RNA # Clones picked (homozygous) (heterozygous) (indels or transversions) no editing gRNA 1

42

19 (45%)

2 (5%)

0 (0%)

21 (50%)

gRNA 2

19

3 (16%)

4 (21%)

0 (0%)

12 (63%)

gRNA 3

37

1 (3%)

5 (13.5%)

5 (13.5%)

26 (70%)

Table 2 Table displays results from three genes edited with the pCMV-ABEmax-P2A-GFP plasmid. While the overall editing efficiency varied depending on the target gene, base editing was observed in all three genes after sequencing. Heterozygous edited clones were obtained in all three editing experiments. However, homozygous edited clones were only observed in two experiments ABEmax editing efficiencies # Clones edited # Clones edited # Clones edited # Clones Guide RNA # Clones picked (homozygous) (heterozygous) (indels or transversions) no editing gRNA 4

46

30 (65%)

6 (13%)

0 (0%)

10 (22%)

gRNA 5

37

18 (49%)

15 (40%)

0 (0%)

4 (11%)

gRNA 6

15

0 (0%)

5 (33%)

0 (0%)

10 (67%)

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Table 3 Table displays results from two genes edited with the pCas9-GFP plasmid. Editing was observed in both genes after sequencing. However, only homozygous edited clones were observed in each experiment. These indels were of different character and were present on both alleles of each edited clone Cas9 editing efficiencies

Guide RNA

# Clones picked

# Clones edited (homozygous)

# Clones edited (heterozygous)

# Clones no editing

gRNA 4

27

10 (37%)

0 (0%)

17 (63%)

gRNA 7

10

5 (50%)

0 (0%)

5 (50%)

Overviews of editing results and efficiencies using BE4max, ABEmax, and Cas9 are shown in Tables 1, 2 and 3. 11. Further subclone the clones with desirable genotypes as per steps 3–11, then expand the subclones for further applications and freezing. We recommend subcloning to reduce the risk of heterogenous populations within individual clones. If a genotype remains identical upon subcloning, the genetic identity of the clone can be confirmed. 3.5 Optional: Single Allele Genotyping

If an iPSC clone carries multiple heterozygous edits, TOPO-TA cloning may be necessary to determine the mutations carried by each allele. 1. Combine the following reagents in a clean PCR tube. 1 μl fresh PCR product 1 μl Salt Solution 3 μl cell culture grade water 1 μl PCR® 4-TOPO® vector 2. Allow the reaction to incubate for 20 min at room temperature. 3. Add 5 μl of the reaction to competent cells on ice and transform according to manufacturer’s instructions. 4. Plate on a kanamycin (50 μg/ml) LB-agar plate and incubate overnight at 37  C. 5. Pick individual colonies using 10 μl pipette tips, and dissolve individual colonies into 10 μl of water in PCR tubes. Optional: Keep a “duplicate” of each colony by tapping the colony into a new Kanamycin LB-agar plate prior to dissolving in water. 6. Heat the dissolved colonies for 7 min at 95  C.

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Fig. 4 Determining genotype from a BE4max-edited subclone. (a) PCR of TOPOTA colonies run on a 1% agarose gel. Bands representing successful amplification of individual alleles can be observed in lanes 1, 3, and 4. No band is present in lane 2, suggesting unsuccessful ligation of fresh PCR product to the TOPO-TA vector prior to transformation. (b) Chromatogram of the original subclone alongside two chromatograms of its corresponding TOPO-TA products. Analysis of the TOPO-TA product chromatograms confirms the identities of both alleles

7. Use 5 μl of the mixture to perform PCR on the extracted DNA. We recommend using the M13F and M13R primers that come with the kit. 8. Perform agarose gel electrophoresis on the PCR products to identify colonies that successfully incorporated the PCR product. Figure 4a shows an example agarose gel used to identify clones that have incorporated the PCR product. 9. Analyze PCR samples (or miniprep of desired colonies) via Sanger sequencing (see Note 11). We recommend using T3 as sequencing primer, and sequencing at least five colonies to increase the odds of reading both alleles. Figure 4b displays how sequencing TOPO-TA colonies can determine the allele identities of an original subclone.

4

Notes 1. Different plasmids can be used to express gRNAs as long as the scaffold is compatible with spCas9, but the cloning protocols will change. If your target sequence starts with G, you can

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remove it from the protospacer sequence. The gRNA will be transcribed by the RNA pol III, as the G present in the overhang is used as the preferred initiation nucleotide. 2. The annealing procedure can also be performed by heating the oligos to 96  C for 5 min and allowing to cool at room temperature (15–20 min). 3. Do not exceed 3 μg total plasmid quantity, as excess DNA may be toxic to iPSCs. The optimal molar ratio of base editor-GFP plasmid to guide RNA plasmid should be approximately 1:3–1: 2. The molecular weights of the ABEmax-GFP and BE4maxGFP plasmids are 9.6 kb and 9.8 kb, respectively. The molecular weight of the phU6-gRNA plasmid is 3.5 kb. Our standard transfection protocol requires 1.5 μg of BE and 1 μg of gRNA; however, plasmid concentrations can be varied to obtain specific edits (low frequency outcomes or heterozygous mutations). 4. If a low percentage of cells are GFP+ upon sorting, there may be low transfection efficiency during lipofection. Ensure that both plasmids are within the concentration range specified and are high quality, endotoxin-free, and sterile. In addition, take care to pipette Tube 1 into Tube 2 gently. Do not further mix the two solutions after the initial transfer of Tube 1 into Tube 2, as pipetting up and down may compromise micelle integrity and therefore lower transfection efficiency. 5. Plating density for sorted cells can range between 5000 and 20,000 cells in a 10 cm MEF-coated plate. Lower density (20,000) will increase the risk of picking merged/mixed colonies. If the yield of GFP+ cells after sorting is lower than desired, add lipofectamine and DNA to additional wells under identical conditions and combine before sorting. If no colonies are observed following a successful sort, or if the only surviving colonies are wild type or heterozygous for the mutation of interest, the targeted wild type sequence may be essential for cell survival. 6. To analyze editing outcomes via Sanger sequencing, primers should be designed to obtain 200–500 bp amplicons. The sequencing primer should be located at least 100 bp away from the expected base editing window. 7. If no band is observed following PCR and agarose gel electrophoresis of a colony, ensure that the colony pellet was not aspirated along with the supernatant during the DNA extraction protocol. In addition, insufficiently sized colonies may not yield enough cells for adequate DNA extraction. Lastly, ensure that extracted DNA is properly stored, as the QuickExtract™

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product is unstable at room temperature for extended periods of time. 8. If post-sorting the bulk population shows a low editing efficiency within the predicted window, plasmid concentrations of gRNA and BE can be increased. Alternatively, additional prediction software can be used to explore different guide RNAs and base editors that may have increased efficiency in modifying the nucleotides of interest. 9. If a mixed chromatogram is observed upon Sanger sequencing of individual clones, it is likely that the picked colony was comprised of multiple cell genotypes. Further subclone the population until the chromatogram appears clean. 10. On occasion, unexpected edits, off-target edits, or indels can be observed. We have noticed these occurrences while using BE4max, especially when targeting sequences in GC-rich regions. If these unexpected edits are a hinderance toward obtaining the mutation of interest, other base editors with more limited editing windows and less off-target effects can be used. In addition, the guide RNA can be modified to target a different site. 11. If only one genotype is observed upon sequencing individual TOPO-TA colonies from a heterozygous clone, sequence additional colonies until the second allele is observed.

Acknowledgments This research was supported by NIH grants U01HL134696, R01DK118155, R01DK123162, and UG3DK122644. References 1. Koblan LW et al (2018) Improving cytidine and adenine base editors by expression optimization and ancestral reconstruction. Nat Biotechnol 36(9):843–846 2. Rees HA, Liu DR (2018) Base editing: precision chemistry on the genome and transcriptome of living cells. Nat Rev Genet 19(12):770–788 3. Arbab M et al (2020) Determinants of base editing outcomes from target library analysis and machine learning. Cell 182(2): 463–480.e30 4. Hwang GH et al (2018) Web-based design and analysis tools for CRISPR base editing. BMC Bioinformatics 19(1):542

5. Kluesner MG et al (2021) CRISPR-Cas9 cytidine and adenosine base editing of splice-sites mediates highly-efficient disruption of proteins in primary and immortalized cells. Nat Commun 12(1):2437 6. Kabadi AM et al (2014) Multiplex CRISPR/ Cas9-based genome engineering from a single lentiviral vector. Nucleic Acids Res 42(19):e147 7. Kluesner MG et al (2018) EditR: a method to quantify base editing from sanger sequencing. CRISPR J 1:239–250 8. Dehairs J et al (2016) CRISP-ID: decoding CRISPR mediated indels by Sanger sequencing. Sci Rep 6:28973

Methods in Molecular Biology (2022) 2520: 335–360 DOI 10.1007/7651_2022_462 © Springer Science+Business Media, LLC 2022 Published online: 18 May 2021

Proteomic Analysis of Human Neural Stem Cell Differentiation by SWATH-MS Jirina Tyleckova, Jakub Cervenka, Ievgeniia Poliakh, Jaromir Novak, Katerina Vodickova Kepkova, Helena Kupcova Skalnikova, and Petr Vodicka Abstract The unique properties of stem cells to self-renew and differentiate hold great promise in disease modelling and regenerative medicine. However, more information about basic stem cell biology and thorough characterization of available stem cell lines is needed. This is especially essential to ensure safety before any possible clinical use of stem cells or partially committed cell lines. As proteins are the key effector molecules in the cell, the proteomic characterization of cell lines, cell compartments or cell secretome and microenvironment is highly beneficial to answer above mentioned questions. Nowadays, method of choice for large-scale discovery-based proteomic analysis is mass spectrometry (MS) with data-independent acquisition (DIA). DIA is a robust, highly reproducible, high-throughput quantitative MS approach that enables relative quantification of thousands of proteins in one sample. In the current protocol, we describe a specific variant of DIA known as SWATH-MS for characterization of neural stem cell differentiation. The protocol covers the whole process from cell culture, sample preparation for MS analysis, the SWATH-MS data acquisition on TTOF 5600, the complete SWATH-MS data processing and quality control using Skyline software and the basic statistical analysis in R and MSstats package. The protocol for SWATH-MS data acquisition and analysis can be easily adapted to other samples amenable to MS-based proteomics. Key words Proteomics, Data independent acquisition, SWATH-MS, Neural stem cell, Neural differentiation, Spectral library, Skyline, Mass spectrometry

1

Introduction The unique ability of embryonic stem cells (ESCs) to differentiate into different cell types of the adult body makes them an attractive model system for various human diseases. They are useful for hypothesis testing, understanding underlining biological processes and developmental mechanisms of these diseases, testing of new pharmaceuticals or cell transplantation therapies [1]. However, isolation of ESCs from human embryos is burdened with ethical issues [2]. These obstacles can be overcome by using induced pluripotent

Supplementary Information The online version contains supplementary material available at [https://doi.org/ 10.1007/7651_2022_462].

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stem cells (iPSCs) derived by reprograming of adult fibroblasts or other terminally differentiated cells [3]. ESC-/iPSC-derived neural stem cells (NSCs) that can differentiate into all neural subtypes are in addition a great hope for treatment of nervous system injuries, neurodegenerations and autoimmune diseases affecting nervous system [4, 5]. For example, NSCs could compensate deficits of cholinergic neurons in the cerebral cortex in Alzheimer disease [6], medium spiny neurons in striatum in Huntington disease [7] or dopaminergic neurons in the substantia nigra in Parkinson disease [8]. However, the translation of NSCs into cell-replacementbased application has been limited so far, with many concerns regarding ethical and safety issues (e.g., potential for tumorigenesis). Moreover, availability of numerous cell lines with unclear origin and high variability in both ESCs and NSCs differentiation protocols make proper standardization and characterization required for clinical use difficult. Presence of mixed cell populations of various differentiation stages common to many protocols used in research settings is unacceptable in clinic and reliable method to test for presence of such undesired populations is needed. Transcriptomic approaches are often used to identify changes in gene expression connected to proliferation and differentiation of NSCs and indirectly gain insight into regulation of these processes. However, changes in protein abundance and post-translational modifications are ultimately responsible for changes in cellular phenotypes and functions. Therefore, proteomic approaches that enable the detection and quantification of a large number of proteins in multiple conditions without any prior knowledge by largescale experiments employing mass spectrometry (MS) represent a method of choice [9]. Traditional shot-gun MS proteomic data-dependent acquisition (DDA) strategy is due to the partially stochastic sampling of MS2 spectra not suitable for fully reliable quantification. Other approach is targeted analysis using Selected Reaction Monitoring (SRM) method, which is precise and sensitive, but limited to approximately one hundred of preselected protein targets in one analysis. Recently, we developed SRM-based assay for specific and sensitive monitoring of cell type-specific (human ESCs, NSCs, glial cells, and neurons) protein markers expression, which allows characterization of cell populations in culture and further optimization of differentiation protocols [10]. Nowadays, another approach called data-independent acquisition (DIA) is gaining more popularity, mainly due to the reproducibility and accuracy of label-free protein quantification. In DIA, MS1 scan is divided into windows with specific m/z values and all precursor ions in every particular window are fragmented, which should theoretically lead to identification and quantification of all detectable spectra. The same precursor isolation windows are measured in repeated cycles across the whole chromatographic separation [11]. This process results in

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very complex fragmentation spectra from different precursor ions within the same isolation window, making classical peptide identification by search against protein sequence databases impossible. This can be resolved by generation of sample specific spectral library from parallel measurement of DDA, ideally on the same sample and instrument [12], or computationally by generation of spectral libraries in silico [13]. Currently, number of different DIA protocols for distinct MS instruments have been developed [14]. One of specific DIA implementations represents Sequential Window Acquisition of all Theoretical Mass Spectra (SWATH-MS) data acquisition on TripleTOF 5600 (AB Sciex). SWATH-MS combines advantages of sensitivity of SRM-like data extraction with in depth proteome coverage. Thousands of proteins may be measured and quantified in a single sample. This chapter presents complete protocol for SWATH-MS characterization of NSC differentiation starting from cell culture, followed by sample preparation protocol for MS analysis, the MS analysis on TTOF 5600+ itself, then SWATH-MS data processing and quality control in Skyline [15] and finally the basic statistical analysis using MSstats package in R [16]. The aim of the protocol is to provide an overview of one of the numerous possibilities of DIA analyses of human cell lines using procedures that are easily to repeat in a general wet lab with the main software tools freely available (Skyline, R). This protocol is focused on the analysis of a complete cell lysate. We performed a comprehensive characterization of cellular proteome during spontaneous differentiation of human H9 ESC-derived NSCs and quantified changes in expression of more than 2500 proteins by employing this protocol [17]. Further modifications of sample preparation allow for analysis of a specific cell compartment or cell conditioned media/secretome as well.

2

Materials

2.1 Cell Culture and Differentiation

All material for cell culture should be prepared in sterile conditions according to basic laboratory practice (see Note 1). 1. Sterile deionized water. 2. Sterile Phosphate buffered saline 1 (PBS), pH 7.4 at 25  C. 3. Poly-L-ornithine working solution 20 μg/mL in sterile deionized water. 4. Laminin mouse protein working solution 10 μg/mL in sterile PBS. 5. Human Neural Stem Cells (H9 ESC-derived) (Thermo Fisher Scientific, N7800100) (see Note 2).

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6. Stem Pro™ Neural Stem Cell Serum Free Medium (SFM) (Thermo Fisher Scientific) containing KnockOut™ DMEM/ F-12 with 2 mM GlutaMAX™, 2% StemPro™ Neural Supplement (v/v) without the growth factors, 20 ng/mL recombinant human EGF, 20 ng/mL recombinant human FGF basic and combination of antibiotics 1% penicillin–streptomycin (see Note 3). 7. Stem Pro™ Neural Stem Cell differentiation medium: KnockOut™ D-MEM/F-12 including 2 mM GlutaMAX™ with 2% StemPro™ Neural Supplement without the growth factors and with combination of antibiotics 1% penicillin–streptomycin. 8. Cell culture plastics. 2.2 Consumables for Cell Lysis and Sample Preparation

1. Sterile polypropylene 15 mL and 50 mL tubes. 2. Sterile polypropylene 2 mL microtubes that withstand centrifugation forces of 20,000  g. 3. Precision pipettes with tips up to 1000 μL. 4. 96-well flat bottom plates or cuvettes for protein concentration determination assay. 5. Cell scrapers. 6. Hamilton glass syringe. 7. pH indicator strips for pH 1–7. 8. Ready-to-use spin columns of porous C18 reverse-phase resin (see Note 4).

2.3 Chemicals for Cell Lysis and Sample Preparation

All the solutions for peptide cleanup, peptide sample dilution and mobile phases for liquid chromatography (LC) have to be in LC-MS grade quality. 1. Ammonium Bicarbonate, 50 mM in LC-MS water. 2. Denaturation buffer containing 8 M urea, 5 mM EDTA, and 50 mM ammonium bicarbonate. 3. Assay kit for determination of protein concentration (see Note 5). 4. ProteaseMAX™ Surfactant, Trypsin Enhancer (Promega), prepare 1% solution by adding 100 μL of 50 mM ammonium bicarbonate to 1 mg vial of lyophilized ProteaseMAX™. 5. Tris(2-carboxyethyl)phosphine hydrochloride (TCEP), 100 mM stock solution in LC-MS water (see Note 6). 6. Iodoacetamide (IAA), 400 mM stock solution in LC-MS water. 7. Proteases Endoproteinase Lys-C MS grade and Trypsin MS grade, 1 μg/μL LysC/Trypsin in 50 mM acetic acid. 8. Formic acid (FA).

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9. Acetonitrile (ACN). 10. Water. 2.4 Hardware and Software for Sample Preparation

1. Spectrophotometer for determination of protein and peptide concentration with appropriate software. 2. Ultrasonic water bath. 3. Vortex. 4. Refrigerated centrifuge with fixed angle rotor for microtubes, up to 20,000  g at 4  C. 5. Thermo Shaker incubator for microtubes. 6. Acid-Resistant Centrifugal Vacuum Concentrator.

2.5 Hardware and Software for Data Acquisition and Analysis of SWATH-MS Measurement in TrapElute Mode

Workflow described in this protocol is designed and optimized on following hardware and software setup. 1. 5600+ TripleTOF mass spectrometry system (AB Sciex). 2. Eksigent 425 Nano-LC system (AB Sciex). 3. Trap column Acclaim™ PepMap™ 100 C18 (100 Å 5 μm, 0.1  20 mm; Thermo Fisher Scientific). 4. 25-cm fused-silica analytical column (75-μm inner diameter) packed in-house with ProntoSIL 120 Å 3 μm C18 AQ beads (Bischoff Analysentechnik GmbH). 5. Analyst TF software 1.7.1 Components for NanoCell (AB Sciex). 6. Eksigent Control Software 4.2 (AB Sciex). 7. PeakView 2.2 (AB Sciex). 8. SWATH Acquisition Variable Window Calculator Excel Sheet (AB Sciex). 9. Mascot Distiller 2.7.1 (Matrix Science Ltd.). 10. Mascot Server 2.6 (Matrix Science Ltd.). 11. Skyline-daily (version Software) [15].

20.1

or

higher,

MacCoss

Lab

12. R (R Core Team) interactive statistical environment [18] (see Note 7). 2.6 Standards for Nano-LC/MS Analysis

1. Mix of indexed retention time (iRT) standard peptides (see Note 8). 2. 25 fmol Beta-galactosidase (Bgal) digest (AB Sciex) with 67 fmol Glu1-Fibrinopeptide B peptide (AB Sciex) in aqueous solution of 10% acetonitrile and 0.1% formic acid for calibration mass spectra of precursor and product ions. 3. A complex digested lysate as quality control of mass spectrometer performance (see Note 9).

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Methods

3.1 Cell Culture and Differentiation

1. Culture the NSCs under sterile condition according to guidelines for culturing GIBCO human H9 ESC-derived NSCs (GIBCO H9 Protocol). In summary, culture the NSCs on poly-L-ornithine/laminin coated tissue-culture dishes in StemPro NSCs SFM complete medium at 37  C in a humidified atmosphere of 5% CO2 in the air (see Note 10). Replace the cell culture media with fresh media every other day (see Note 11). 2. When cells reach approx. 70% confluence, the differentiation may be started. Perform the spontaneous differentiation by growth factors withdrawal from the cell culture media (Stem Pro NSCs differentiation medium). Change half of the media (see Note 12) every other day for 7 and 21 days after growth factors withdrawal (Fig. 1). Prepare at least four biological replicates per condition.

Fig. 1 Morphology and immunocytochemistry characterization of spontaneously differentiated NSCs. Human H9 ESC-derived NSCs were spontaneously differentiated by withdrawal of growth factors for 21 days. Cellular morphology is significantly changed from still NSC-like at day 7 (a) to neuronal at day 21 (b). Sporadic signals of neuronal marker Tubulin beta-3 chain (green color) and astrocyte marker protein S100-B (red color) were detectable at day 7 (c), but expression of both markers was highly increased at day 21 (d). Nuclei were stained by DAPI (blue color)

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1. Remove medium from the plate with differentiated cells. 2. Carefully wash the plate with prewarmed PBS solution (37  C). 3. Add cold (4  C) PBS to the plate. 4. Place the plate on box with ice. 5. Using a cell scraper, scrape cells from the bottom and collect them in the corner of the dish (see Notes 13 and 14). 6. Transfer the cells in PBS solution from plate to polypropylene tube with polypropylene pipette (see Note 15). 7. Add more PBS (4  C) to the plate to wash it and transfer PBS with remaining cells to the same tube from step 6. 8. Pellet cells by centrifugation at 4  C, 300  g for 5 min. 9. Remove PBS with fluid aspirator. 10. Resuspend cells in PBS (4  C). 11. Repeat steps 8–10 two more times. 12. Centrifuge at 4  C, 700  g for 5 min. 13. Remove supernatant (see Note 16). 14. Add denaturation buffer to the pellet (minimally 50 μL of denaturation buffer/1  106 of cells). 15. Disrupt the cell pellet by vortexing for 10 s and centrifuge at room temperature, 1000  g for 2 min. Transfer the cell samples to 2 mL microtubes. 16. Add ProteaseMAX™ to the protein lysate to final concentration of 0.1% (v/v) (see Note 17). 17. Incubate on ice for approx. 12 min with occasional vortexing and spinning (see Note 18). 18. Sonicate lysate on ice in an ultrasonic water bath with frequency approximately 40 kHz for 15 min (see Note 19). 19. Repeatedly pipette the sample through a thin pipet tip (see Note 20). 20. Centrifuge samples at 4  C, 20,000  g for 15 min, keep the supernatant (see Note 21). 21. Determine protein concentration to ensure that initial amount of proteins is the same in all samples (optimal is approx. 50–100 μg of proteins per sample) (see Note 22).

3.3 In-Solution Cell Lysate Digestion and Peptide Desalting

There are numerous methods for sample preparation for MS proteomic analysis. Here we describe a universal technique for peptide isolation, which is in-solution protein digestion. It is one of the most common, reproducible and error-proof protocol. Alternatively, Filter Aided Sample Preparation (FASP) [19] protocol is often used for removing detergents from lysis buffer, which interfere with digestion and MS measurement. For deeper proteome

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coverage abundant protein depletion, enrichment and fractionation methods are highly recommended. 1. Add reducing agent TCEP to the protein lysate to a final concentration of 10 mM. 2. Incubate for 30 min at 32  C under agitation. 3. Add alkylating agent IAA to the protein lysate to a final concentration of 40 mM. 4. Incubate for 45 min at room temperature in the dark under agitation. 5. Dilute the sample with freshly prepared 50 mM Ammonium Bicarbonate and 1% ProteaseMAX™ to final concentration of 1 M urea and about 0.02% ProteaseMAX™. 6. Add LysC in 1:100 enzyme–substrate ratio. 7. Incubate for 2–4 h at 37  C on a shaker under agitation. 8. Add Trypsin in 1:100 enzyme–substrate ratio. 9. Incubate for 14–16 h (overnight) at 37  C in the dark under agitation. 10. Stop the digestion by adding FA to a final concentration of 2% (see Notes 23 and 24). 11. Centrifuge at 20,000  g for 15 min (4  C) and keep the supernatant. 12. Use pH indicator strip to check that the pH is lower than 3 for the peptide cleanup. 13. Desalt samples, for example, on C18 spin columns according to manufacturer’s directions. Select the C18 spin column size according to sample load (see Note 25). 14. Vacuum centrifuge the eluted peptides after desalting step to dryness at 45  C. 15. Resuspend dry peptides in aqueous solution with 2% ACN and 0.5% FA and sonicate in water with ice bath for 5 min. 16. Centrifuge at 20,000  g for 15 min (4  C) and collect pure supernatants. 17. Determine the peptide concentration by measuring absorbance at 280 nm with Nanodrop or equivalent spectrophotometer with low sample volume requirement. Prepare peptide samples of defined concentration (e.g., 1 μg/μL) with 1:30 of spike-in iRT peptides in aqueous solution with 2% ACN and 0.5% FA into MS vials (see Note 26). 3.4 LC-MS/MS Methods and Measurements

Separate the peptide mixture by reversed phase C18 in a trap-elute mode using the nano-LC system coupled online to MS.

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Fig. 2 Example of LC method settings. (a) Graphical summary of LC method of 120-min linear gradient of 5–35% ACN followed by column wash with 95% ACN and column equilibration with 5% ACN with flowrate 200 nL/min and total run time 168 min. (b) Settings of LC method. For trap-and-elute configuration choose standard type for sample injection

1. Set the autosampler method injecting 1 μL of sample. 2. Set the LC loading method using a loading pump flow rate 2 μL/min of 2% ACN and 0.5% FA for 10 min and the analytical gradient method using a flow rate 200 nL/min on a linear gradient of 5–35% (v/v) ACN with 0.1% FA (from 0 to 120 min), 35–95% ACN with 0.1% FA (from 125 to 130 min), 95% ACN with 0.1% FA (from 130 to 135 min), 95–5% ACN with 0.1% FA (from 135 to 140 min) and column equilibration in 5% ACN with 0.1% FA (from 140 to 168 min) (Fig. 2). 3. In Analyst 1.7.1 create a new IDA acquisition method (see Note 27). Set Experiment 1 to TOF MS with m/z 400–1250 (see Note 28), the accumulation time 300 ms and positive polarity (Fig. 3a). Click on Edit parameters button and fill the Source/Gas and Compound Parameters settings according to Table 1, and Fig. 3a (see Note 29). Set Experiment 2 to Product Ion, select IDA experiment. Set accumulation time to 150 ms and m/z to 170–2000. Monitor 20 candidate ions (see Note 30) per one survey scan with charge state 2–5, which exceeds 150 cps with mass tolerance 50 mDa and exclude former target ions for 13 s after one MS/MS spectra is collected (Fig. 3b) (see Note 31). Check Rolling collision energy (RCE) (see Note 32) box in IDA advanced settings. Set the duration of MS method to 140 min. Save the LC-MS acquisition method with LC sync. 4. Ahead of creating SWATH-MS method prepare the Variable Window text file by computing from previously acquired IDA

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Fig. 3 (a) Example of typical IDA acquisition method settings for TOF MS. Typical Source/Gas Parameter Settings are 2300–2500 V for ISVF; 10–15 for GS1; 0 for GS2; 25–35 for the Curtain Gas and 100–150 for IHT. (b) Example of Switch Criteria settings and the IDA Collision Energy Parameters that need to be actualized time-to-time

data of your specific sample. Use the Variable windows calculator that can be downloaded from AB Sciex webpage after registration (SWATH Variable Window Calculator). Follow the steps in the Instructions. Select the number of variable windows (see Note 30), set the mass range (typically 400–1250 for TripleTOF 5600) and the window overlap (usually 1 Da). Set the CES to 15 and minimum window width

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Table 1 Typical Source/Gas and Compound settings that must be optimized for each TTOF source Parameter

Typical settings

Ion source gas 1 (GS1)

10–15

Ion source gas 2 (GS2)

0

Curtain gas (CUR)

25–35

Interface heater temperature (IHT)

100–150

IonSpray voltage floating (ISVF)

2300–2500 V

Declustering potential (DP)

80–100 V

Collision energy spread (CES)

10–15 V

Note

see Note 29

(no lower than 4). After calculating the windows, copy the Q1 start, Q1 stop and CES columns in the OUTPUT for Analyst tab and save it as text file (Fig. 4a, b). 5. Create a new SWATH-MS method. Open the IDA method and in the acquisition method tab click on Create SWATH Exp button, choose Manual button. Under the fragmentation conditions check RCE. SWATH Detection Parameters set to 170–2000 with accumulation time 100 ms (see Note 33). Tick the Read SWATH Windows from Text File and upload the variable windows text file (Fig. 4c). Save the SWATH-MS acquisition method and check the TOF MS settings. 6. Set the queue with random NSC differentiation samples positions and both IDA and SWATH-MS methods for each sample and with quality control and calibration Bgal samples among them and run the queue. 3.5

Data Analysis

For processing SWATH-MS data (or DIA data in general), several workflows using different tools have been introduced. There are two main strategies of processing DIA data. The first one and the most prevalent is a peptide-centric approach that uses spectral libraries established from previous DDA measurements for targeted extraction of the SWATH-MS (DIA) data. The most common tools used for this type of SWATH-MS data analyses are OpenSWATH (ETH Zu¨rich) [20, 21] and PeakView (AB Sciex) [22]. The second one is a spectral-centric approach that uses data from DIA measurement for the library building. This kind of analysis still has lower number of identified precursor ions when compared to the peptidecentric approach, but software tools are regularly improving. This approach is supported by a DIA-Umpire [13] and Group-DIA [23]. MaxDIA (MaxQuant software from Cox laboratory) [24], Skyline (MacCoss Lab Software) [25], and Spectronaut (Biognosys) [26, 27] allow both types of analyses.

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Fig. 4 (a) An example of using SWATH-MS Variable windows calculator for constructing variable windows pattern (red line) from m/z density histogram (blue line) from the NSCs sample TOF MS IDA data. In high m/z dense regions with high number of precursors smaller Q1 windows are used (b). A text file characterizing the Q1 isolation window strategy that can be loaded into the SWATH-MS method editor for SWATH-MS method building (c)

In the next section, we describe the use of Skyline for SWATHMS data analysis of spontaneous differentiation of H9 ESC-derived NSCs and MSstats package in R for statistical analysis of the data. We choose Skyline because it is a free software with incorporated options for spectral library building (see Note 34) and mProphet peak picking for SWATH-MS data. Unlike OpenSWATH and DIA-Umpire, Skyline has a user-friendly graphical interface with unique options of data visualization. Skyline is optimal for small laboratories and beginners, who need to quickly learn how to process data from MS measurements. Skyline creators provide their users with free webinars, tutorials and quick support feedback. Skyline may be also preferred by users familiar with SRM data analysis. For SWATH-MS data analysis in Skyline you will need to build a spectral library and create a retention time calculator. Before this, you will specify peptide and transition settings in Skyline document,

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accordingly to parameters from your measurements. For peak picking and finding a match in your assay library Skyline has incorporated the mProphet model. Finally, Skyline allows basic statistical analysis using MSstats plug-in or export of data for further advanced analysis in other statistical programs such as R. Complete published dataset [17] comprising spectral library and SWATH-MS measurements analyzed in Skyline is available in the Panorama Public repository (https://panoramaweb.org/ NSCsdifferentiation.url). For simplicity, selected samples of this dataset were processed separately and are provided as Electronic Supplementary Material (Data 1 and 2) for this protocol in the form of the example input data for further MSstats analysis. 3.5.1 Skyline Document Transition Settings

1. Start with resetting Skyline parameters by clicking on Settings/ Default (see Note 35). 2. In Settings/Transition Settings/Full-Scan (Fig. 5) both for MS1 and MS/MS filtering choose your Product mass analyzer and set Resolving power from your measurements (see Note 36).

Fig. 5 Example of full-scan parameters in transition settings

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(a) In MS1 filtering choose Isotope peaks included/Count (see Note 37). (b) In MS/MS filtering choose Acquisition method/DIA. (c) In MS/MS filtering create isolation scheme through Isolation scheme/Add/Prespecified isolation windows/ Import—choose one of the raw SWATH-MS data .wiff files (see Note 38). (d) Tick Use high-selectivity extraction in case you measured the data in high-selectivity mode. (e) Set window width in minutes in Retention time filtering/Use only scans within N minutes of predicted RT (see Note 39). 3. In Settings/Transition Settings/Filter set Peptides/Precursor charges: 2, 3, 4; Ion charges: 1, 2, 3 and Ion types: y, b. In Product ion selection select From: ion 3 and To: last ion. Tick Use DIA precursor window for exclusion and Auto-select all matching transitions (see Note 40). 4. For future assay library settings choose in Settings/Transition Settings/Library/Ion match tolerance: 0.05 m/z; tick If a library spectrum is available, pick its most intense ions; Pick: 6 product ions and 3 minimum product ions and select From filtered product ions. 5. In Settings/Transition Settings/Instrument copy parameters from your SWATH-MS method settings. For example, on TTOF 5600+ we measure with Min m/z: 170 m/z and Max m/z: 2000 m/z with Method match tolerance m/z: 0.05 m/z. 6. In Settings/Transition Settings/Prediction and in Settings/ Transition Settings/Ion Mobility leave everything default. 3.5.2 Spectral Library Building from IDA Measurements in Skyline and Peptide Settings

1. Run protein sequence database search of your IDA measurements against annotated database of your sample species (see Notes 41 and 42). For example, we use Mascot Distiller with Mascot Server (see Note 43) and a human Swiss-Prot FASTA file extended by a list of common protein contaminants for MS. 2. Export sequence database search from your search engine in format acceptable in Skyline like .dat file. 3. Specify in Settings/Peptide settings/Digestion, which enzyme/s you used (see Note 44) and how many missed cleavages you allow (see Note 45). If you used combination of enzymes, build background proteome in Background proteome/Add using the same FASTA file as set in your search engine and Enforce peptide uniqueness by: None (Fig. 6a).

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Fig. 6 Example of library building in Skyline with steps (a) to (c)

4. In Settings/Peptide settings/Filter choose Min length: 7 and Max length: 36, set 0 to Exclude N-terminal AAs and tick Auto-select all matching peptides. 5. In Settings/Peptide settings/Modifications set your peptide modifications, most common are carbamidomethylation of cysteine, oxidation of methionine and acetylation of N-term, Max variable mods: 3.

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6. Build your spectral library and iRT calculator predictor in one window through Settings/Peptide settings/Library/Build, name your library and choose output path. Choose your type of spiked-in standard peptides for building retention time predictor (Fig. 6b). In our case, we used 11 iRT peptides from Biognosys and set cut-off score: 0.99 (see Note 46). Click on Next and Add files from your search engine (e.g., .dat files) and click on Finish. After finishing the library building, click on OK for Add iRT Peptides. Then click on Yes for Do you want to recalibrate the iRT standard values relative to the peptides being added? (see Note 47). Skyline will ask you for Add Retention Time Predictor—name it and set Time window: 10 min. When library is prepared, in Settings/Peptide settings/Library set Pick peptides matching: Library and Filter. 7. In Settings/Peptide settings/Prediction and in Settings/Peptide settings/Quantification leave everything default. 8. After confirming all Settings/Peptide settings Skyline may ask you for Add Standard Peptides—set Maximum transitions per peptide: 3 and click on Yes. This step will add Biognosys standard peptides to your protein targets. 9. To add protein targets to your Skyline go to View/Spectral Libraries, in Library: choose your library, tick Associate proteins and click on Add all (Fig. 6c). Click on Do not add for peptides matching multiple proteins as well as for peptides not matching the current filter settings. 3.5.3 Importing the SWATH-MS Data into Skyline and Peak Picking with mProphet

1. Adjust settings for your data analysis in Refine/Advanced—set Remove repeated peptides and Remove duplicate peptides (see Note 48). In the same window set Min peptides per protein: 2 and Min transitions per precursor: 3. In Refine/Remove Empty Proteins. In case your FASTA file for background proteome contains iRT peptides, delete them from the Targets prior this step. Otherwise, both your iRT peptides and Biognosys standards added in Subheading 3.5.2, step 8 will be removed from your Targets. 2. In Refine/Add Decoys choose Decoy generation method: Shuffle Sequence (see Note 49). Number of decoy peptides should correspond to the number of all peptides in protein targets without iRT peptides. 3. File/Import/Results—choose Add single-injection replicates in files with Optimizing: None (see Note 50). 4. In Refine/Reintegrate tick Integrate all peaks and in Peak scoring model choose Add. Name your model, Choose model: mProphet, tick Use decoys and click on Train Model. Perform a quality control of mProphet model. View graph of decoys and targets discrimination (Fig. 7).

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Fig. 7 Example of mProphet quality control. The generated Decoys (red histogram) should match the Decoy normal distribution. When using sample specific library, the Targets distribution (blue histogram) should be clearly distinct from the Decoys distribution

5. Perform a manual quality control check of your extraction parameters (see Note 51, Fig. 8). To see whether mass error was set correctly, click on View/Mass Errors/Histogram (Fig. 8e) and View/Mass Errors/Replicate Comparison. To see the linear regression used to predict the target retention time, click on View/Retention Time/Regression/Score To Run. 3.5.4 Sample Annotation, Group Comparison and Data Export

1. Annotate your data for downstream statistical analysis in Settings/Document Settings/Annotations—click on Add and define annotations for Condition (set Type: Value List, Applies to: Replicates) and BioReplicate (set Type: Number, Applies to: Replicates). Then go to View/Document Grid and annotate the samples (see Note 52). 2. Basic group comparison can be performed directly in Skyline, using MSstats as an external tool. If not already installed, add MSstats tool to your Skyline in Tools/External Tools—click on Add, choose From Tool Store and select MSstats (see Note 53). Click on Tools/MSstats/Group comparison and select groups you want to compare, control group and type of normalization. The results from group comparison will be by default automatically saved to your Skyline folder.

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Fig. 8 Example of quality control check of extraction parameters on Neuropilin-1 peptide TGPIQDHTGDGN FIYSQADENQK on day 7 (D7) and day 21 (D21) of differentiation. (a) Check the correct peak picking (identification) together with checking the peak boundaries and matching in relative transition intensities to the library spectrum (b). (c) Check the reproducibility of relative transition peak intensities among samples (Peak Areas) and look at Retention Time Replicate Comparison where the box plots should appear approx. at the same retention time and with similar distributions (d). (e) Look at histogram of mass errors and adjust parameters in Settings/Transition Settings/Full-Scan if needed

3. Customize your data export for external MSstats analysis in File/Export/Report click on Edit list—choose MSstats Input and click on Edit to add desired information to your export. In case that MSstats Input is not in your report options, you can create it in File/Export/Report click on Edit list, click on Add and then name your export report and configure it. 4. Remove Biognosys standards (or iRT peptides), contaminants and Decoys from the Targets before exporting the results as MSstats Input, because these are not needed for quantification (see Note 54). 5. Export data for further analysis in File/Export/Report/ MSstats Input—click on Preview (Fig. 9), control data and then click on Export. 3.6 Statistical Analysis: Relative Quantification Using R

Commented script replicating above described statistical analysis with up to date R [18] and MSstats version (R version 4.1.2, released 2021-11-01, MSstats_4.0.1) is available as Electronic Supplementary Files (Data 1 and 2) with this chapter. In our example, we use data preprocessed in Skyline and convert the Skyline output into correctly formatted MSstats input using SkylinetoMSstatsFormat function. However, MSstats provides several convenient

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Fig. 9 The MSstats required parameters for MSstats Input are Protein Name, Peptide Modified Sequence, Precursor Charge, Fragment Ion, Product Charge, Isotope Label Type, Condition, BioReplicate, File Name, Area, Standard Type, Truncated and Detection Q Value

conversion functions for preparing output of other SWATH-MS data processing pipelines for statistical analysis (e.g., OpenSWATHtoMSstatsFormat). The main function for transforming and summarizing original raw data into quantitative data for model fitting and comparison of groups of interest is “dataProcess.” We refer readers to MSstats documentation for detailed explanation of all available parameters of this function. One important parameter of note is “censoredInt,” which defaults to “NA,” but should be set to “0” for Skyline input. Function “quantification” provides log-transformed protein abundances summarized either on sample or biological group level. Sample level output is useful if more complex statistical analysis beyond simple between group (e.g., Control vs Treatment) or repeated measure (Pretreatment vs Posttreatment on the same subject) comparison is required. Other R packages, such as limma [28, 29], lme4 [30, 31] and others can be used to fit appropriate models to exported sample level data.

4

Notes 1. The following products are recommended to prepare solutions for cell culture: Phosphate buffered saline (Sigma-Aldrich), poly-L-ornithine (Sigma-Aldrich), Laminin mouse protein (Thermo Fisher Scientific), EGF (Thermo Fisher Scientific), FGF basic (Thermo Fisher Scientific), penicillin–streptomycin (Thermo Fisher Scientific). 2. Discontinued product. 3. Each SFM component can be purchased independently from Thermo Fisher Scientific; KnockOut™ DMEM/F-12, StemPro™ Neural Supplement without the growth factors, EGF, FGF basic and GlutaMAX™. GlutaMAX™ is the L-alanine-Lglutamine dipeptide, which is more stable source of L-

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glutamine in cell culture media as L-glutamine spontaneously degrades into toxic ammonia over time. 4. Choose the columns accordingly to the amount of peptide in the sample. There are several manufacturers (e.g., The Nest Group, Thermo Fisher Scientific, Oasis, Merck), and all of them have been proved to be good. 5. Pierce™ 660 nm Protein Assay Kit is recommended and compatible with denaturation buffer. 6. Aliquots of stock solutions of denaturation buffer, 100 mM TCEP in LC-MS water, 400 mM IAA in LC-MS water, 1% ProteaseMAX™ in 50 mM ammonium bicarbonate, 1 μg/μL LysC in 50 mM acetic acid, and 1 μg/μL trypsin in 50 mM acetic acid can be stored at 20  C. Ammonium bicarbonate: prepare always fresh. 7. Optionally RStudio [32] (R-Tools Technology, Inc) integrated development environment may be used. 8. In our workflow we use iRT peptides from Biognosys. These iRT standards are highly stable non–naturally occurring peptides that can also be used for calibration of LC system. Other alternatives are offered by, for example, Thermo Fisher Scientific, Polyquant (RePLiCal), AB Sciex (PepCalMix), SigmaAldrich. Alternatively, you can design and use your own iRTs. As far as we know, iRT standards from Biognosys are incorporated not only in Skyline, but also in other software tools for SWATH-MS (DIA) data processing. 9. We use in-house prepared peptide mixture of cell lysate of uniform NSCs for quality control of MS performance and sample preparation procedure. This should be prepared in bulk to be used for approx. 1 year. Alternatively, commercial HeLa Protein Digest Standard (Thermo Fisher Scientific, Cat. No. 88329) or K562 Protein Extract Digest (part of SWATH Acquisition Performance Kit, AB Sciex, Cat. No. 5045757) can be used. The advantage of the commercial protein digests is their possible usage in interlaboratory comparison including some automated quality control systems, for example, QCloud [33]. 10. The NSCs may benefit from culture at lower oxygen partial pressure (10% of oxygen in the air) in order to decrease the oxidative stress. 11. Six-well culture plates can be used for cell growth and differentiation to obtain sufficient material for SWATH-MS analysis as minimal requirements are approx. 3  105 cells. Cells from one well (surface about 9 cm2) represent one sample replicate. At least four replicates are recommended for each condition. Use

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3 mL of culture medium/PBS per well or adjust the volume as needed for different cell culture vessels. 12. The medium should be never completely aspirated from differentiating cells. Medium should be changed carefully by very slow pipetting against the wall of the culture dish, as cells tend to detach in sheets after prolonged differentiation (high risk of detachment from day 7 on). 13. Check under microscope that the cells are detached from the bottom to the solution. If needed, scrape remaining cells from the plate again. 14. Alternatively, detach the cells using StemPro™ Accutase™ Cell Dissociation Reagent (Thermo Fisher Scientific) or 0.05% solution of Trypsin. Check under microscope for cell detachment. When using Trypsin, do not extend incubation over 3 min and stop the enzyme action with soybean Trypsin inhibitor. Accutase preserves higher cell viability in comparison to Trypsin and should be more appropriate for NSCs. 15. Prewet the pipette in PBS before use, to prevent cells from sticking to pipette walls, resulting in loss of material. 16. You can freeze the cell pellet at keep until further use.

80  C at this moment and

17. ProteaseMAX™ is surfactant that solubilizes proteins and thus enhances protein digestion. 18. Combispin (Biovendor) or similar combined centrifuge/vortex device installed in a refrigerator or cold room can be used for this purpose. Recommended settings are 10 cycles, 1 min at 6000 RPM, 20 s hard shaking. 19. You may optionally repeat steps 17 and 18 to enhance the protein yield. 20. This step helps to fragment DNA when the sample tends to become highly viscous. You may add more denaturation buffer to reduce viscosity. 21. You can freeze the samples at until further use.

80  C at this moment and keep

22. Optional: You can spike-in proteins from another nonhomologous protein species to your protein sample at this moment as quality control of your sample preparation protocol and your protein quantification workflow. For human samples we use chicken ovalbumin protein and alcohol dehydrogenase protein from S. cerevisiae. The spike-in proteins spiked in equal amounts per sample could be used only as simple quality control, for example, significantly lower abundance of these proteins in a specific sample would mean problem with your sample preparation. When proteins are spiked into samples in

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different ratios one can inspect the quantification workflow correctness. For less complex samples or for samples where more than 50% of protein content will change in abundance the spike-in proteins could be used as normalization standards as well. 23. When handling FA, work under the hood and use a glass syringe. 24. Alternatively, you can stop the digestion by heating the sample to 80  C for 20 min. 25. The appropriate cleanup column for this type of sample should be, for example, MacroSpin™ (NestGroup) or BioPureSPN MACRO™ SPE column with sample capacity 30–350 μg of peptide/protein. This depends on the initial amount of proteins in a sample lysate, which in our case was approx. 50–100 μg of protein lysate from NSCs differentiated for 7 and 21 days and harvested from one well of about 9 cm2 per sample. 26. The peptide sample load should be optimized for each sample type and instrument. Too high sample load may lead to saturation of the detector and to decrease in LC-MS/MS performance over time due to contamination of MS front end. Our experience is that sample load of 0.5 μg of peptides per sample run is usually enough for 5600+ TripleTOF, the sample load may be increased to 1 μg if necessary for a higher number of spectra and peptide/protein identifications. 27. IDA is an abbreviation for Information Dependent Acquisition in AB Sciex terminology used in AB Sciex software with the same meaning as DDA (Data-Dependent Acquisition). 28. TripleTOF 6600 can go up to m/z 2250. 29. IHT depends mainly on the ESI source. In our settings with black NanoSpray Source III that maintains higher inner temperature IHT 100 may be enough. Low IHT may lead to increased front-end contamination, too high IHT may lead to column tip damage. 30. Accumulation time and ions to monitor are parameters that we recommend to optimize. Depending on chromatographic peak width you should have 6–8 points across the chromatographic peak at half height, that is, for a peak width of 20 s at half height your cycle time should be around 3 s. If you set shorter accumulation time, you can monitor more candidate ions and the other way around. The same principle applies to the number of variable windows that can be used in SWATH-MS mode. 31. These parameters should be again optimized for individual system and sample type. The exclusion time should be set that the peptide is triggered around the highest intensity of

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chromatographic peak. Mass tolerance in mDa is fixed across the m/z, whereas the absolute mass tolerance in ppm will change across the m/z. 32. RCE is calculated based on the charge state of the ion. RCE slope and intercept values needed for determining the unknown charge state should be updated according to AB Sciex instructions over time in the IDA CE Parameters Table (Fig. 3b). 33. We prefer to use longer accumulation time and lower number of windows. Probably more popular is to keep shorter accumulation time and higher number of windows, that is, 30 ms accumulation time would allow 100 variable windows, which is the maximum for TripleTOF 5600. However, with higher number of windows the intensity of detected fragment ions will be lower. 34. Skyline uses term “assay library” for spectral library including retention time calculator. 35. If you already have an optimized profile, for example for SRM analyses, do not forget to save it first—Settings/Save Current. 36. Mass accuracy in ppm is used for centroid data, whereas resolving power is used for raw (profile) data. Approximately, resolving power of 30,000 equals to 15 ppm, 30 ppm equals to resolving power of 15,000 etc. According to Skyline creators, profile data should perform better than centroid data for TTOF 5600+, but the trade-off is a larger data file size and slower data import and processing. Skyline and OpenSWATH creators recommend msConvert (ProteoWizard) for data conversion to centroid data format. 37. Choosing MS1 filtering is optional, but it is assumed that it will increase quality of spectral library. 38. This is an important step. All of your data have to be measured with the same DIA windows parameters/method. 39. The retention time window depends on stability of your LC separation. In our experience 3–10 min is enough. If setting 5 min, the time window will be 10 min (5 min from predicted time). 40. Setting Ion types: p for precursors can be used to check how MS1 spectra match with MS/MS spectra; however, it is not recommended for our current protocol. For more information about using precursor ions please refer to Skyline tutorials. 41. For organisms other than human and mouse, UniProtKB reference proteomes with one protein sequence per gene provide better results than Swiss-Prot as the number of manually reviewed proteins is usually low.

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42. It is possible to use sample nonspecific libraries, for example, pan-Human Library [34]. The currently available SWATH-MS libraries can be downloaded from swathatlas.org. When using sample nonspecific library, it is necessary to use strict cut-off score (0.95 or higher), otherwise the number of the false positive protein identifications may be high. 43. Skyline allows creation of spectral libraries from several search engines as well as employment of Peptide Search tool integrated in Skyline (both for profile and centroid data). 44. The most common enzyme is Trypsin, often used in combination with Lys-C, as in our case. Unfortunately, Skyline does not support combination of Lys-C and Trypsin directly, but there are two different ways to use Skyline with this combination of enzymes: (1) Select trypsin and continue according to this protocol. This way will result in underreporting of missed cleavages after lysine followed by proline. (2) Select Trypsin/ P and import your protein list (File/Import/FASTA). This way will result in overreporting of missed cleavages after arginine, thus increase of number of allowed missed cleavages in Skyline may be required. 45. Set the same number of missed cleavages as in search engine, as Skyline fully relies on missed cleavage results from the search engine. Typically allowing one to two missed cleavages is optimal to see all possible peptides. However, only peptides without missed cleavages should be used for quantification. 46. At this step iRT calculator will be created automatically. The cut-off score 0.99 should approximately correspond to 1% false discovery rate (FDR) for Mascot protein identifications. 47. If you created a sample-specific library, the recalibration of the iRT values relative to the peptides being added can improve retention time alignment under stable chromatographic conditions. 48. Only unique (proteotypic) peptides should be used for statistical analysis. It is a common practice to use at least two peptides per protein and three transitions per peptide for quantification. 49. Decoy transitions are necessary for implemented mProphet algorithm to work as they serve as negative controls [35]. Setting decoy generation method is optional, however, Skyline creators suggest using “shuffle sequence” setting. 50. This part may take some time depending on the size of your results file. Do not import results before adding decoys, otherwise you will have to reimport the data. You may choose to import many files simultaneously. 51. You can manually inspect your data by browsing the Skyline document or by searching for specific proteins/peptides of

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interest. Pay attention to peak picking, to chromatographic pattern and/or to the retention time shifts. 52. Here you can add classification of your samples into biologically meaningful groups (Condition), that is, day 7 (D7) and day 21 (D21) in our example of NSC differentiation. The BioReplicate should be numerical value, that is, with eight samples in our analysis the BioReplicate values are 1–8. 53. The MSstats version available in Skyline external tool store is currently outdated and relies on R version 3.5.0. R and several required packages will be installed when you select this option. Presence of other R versions on the same system or customization of R library location will interfere with correct MSstats installation and function and should be avoided. Installation of MSstats from Skyline store can also overwrite your already installed R packages, so care should be taken if any other analysis depends on specific package versions. 54. You can remove iRTs, decoys and contaminants before exporting the results or you can remove it later on in R data preprocessing, because MSstats does not recognize them. Depending on Skyline version, decoys may be removed automatically by Skyline during MSstats Input export generation.

Acknowledgments This work was supported by the Czech Ministry of Education, Youth and Sports project InterCOST (LTC18079) under CellFit COST Action (CA16119), the Charles University, projects GA UK No. 1767518 and No. 1460217, and the Czech Science Foundation (19-01747S). References 1. Thomson JA, Itskovitz-Eldor J, Shapiro SS et al (1998) Embryonic stem cell lines derived from human blastocysts. Science 282: 1145–1147 2. Barker RA, de Beaufort I (2013) Scientific and ethical issues related to stem cell research and interventions in neurodegenerative disorders of the brain. Prog Neurobiol 110:63–73 3. Takahashi K, Yamanaka S (2006) Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell 126:663–676 4. Goldman SA (2016) Stem and progenitor cellbased therapy of the central nervous system: hopes, hype, and wishful thinking. Cell Stem Cell 18:174–188

5. Zizkova M, Sucha R, Tyleckova J et al (2015) Proteome-wide analysis of neural stem cell differentiation to facilitate transition to cell replacement therapies. Expert Rev Proteomics 12:83–95 6. Boese AC, Hamblin MH, Lee J-P (2020) Neural stem cell therapy for neurovascular injury in Alzheimer’s disease. Exp Neurol 324:113112 7. Choi K-A, Hong S (2017) Induced neural stem cells as a means of treatment in Huntington’s disease. Expert Opin Biol Ther 17: 1333–1343 8. Fan Y, Winanto, Ng S-Y (2020) Replacing what’s lost: a new era of stem cell therapy for Parkinson’s disease. Transl Neurodegener 9:2

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9. Aebersold R, Mann M (2003) Mass spectrometry-based proteomics. Nature 422: 198–207 10. Sucha R, Kubickova M, Cervenka J et al (2021) Targeted mass spectrometry for monitoring of neural differentiation. Biol Open 10: bio058727 11. Gillet LC, Navarro P, Tate S et al (2012) Targeted data extraction of the MS/MS spectra generated by data independent acquisition: a new concept for consistent and accurate proteome analysis. Mol Cell Proteomics 11(6): O111.016717 12. Ludwig C, Gillet L, Rosenberger G et al (2018) Data-independent acquisition-based SWATH-MS for quantitative proteomics: a tutorial. Mol Syst Biol 14:e8126 13. Tsou C-C, Avtonomov D, Larsen B et al (2015) DIA-umpire: comprehensive computational framework for data-independent acquisition proteomics. Nat Methods 12:258–264, 7 p following 264 14. Meyer JG, Schilling B (2017) Clinical applications of quantitative proteomics using targeted and untargeted data-independent acquisition techniques. Expert Rev Proteomics 14: 419–429 15. MacLean B, Tomazela DM, Shulman N et al (2010) Skyline: an open source document editor for creating and analyzing targeted proteomics experiments. Bioinformatics 26:966–968 16. Choi M, Chang C-Y, Clough T et al (2014) MSstats: an R package for statistical analysis of quantitative mass spectrometry-based proteomic experiments. Bioinformatics 30: 2524–2526 ˇ ervenka J, Tylecˇkova´ J, Kupcova´ Skalnı´kova´ 17. C H et al (2021) Proteomic characterization of human neural stem cells and their secretome during in vitro differentiation. Front Cell Neurosci 14:612560 18. R Core Team (2021) R: a language and environment for statistical computing. R Foundation for Statistical Computing, Vienna 19. Wis´niewski JR (2016) Quantitative evaluation of filter aided sample preparation (FASP) and multienzyme digestion FASP protocols. Anal Chem 88:5438–5443 20. Ro¨st HL, Rosenberger G, Navarro P et al (2014) OpenSWATH enables automated, targeted analysis of data-independent acquisition MS data. Nat Biotechnol 32:219–223 21. Ro¨st HL, Aebersold R, Schubert OT (2017) Automated SWATH data analysis using targeted extraction of ion chromatograms. In:

Comai L, Katz JE, Mallick P (eds) Proteomics. Springer, New York, pp 289–307 22. Holewinski RJ, Parker SJ, Matlock AD et al (2016) Methods for SWATH™: data independent acquisition on TripleTOF mass spectrometers. Methods Mol Biol 1410:265–279 23. Li Y, Zhong C-Q, Xu X et al (2015) GroupDIA: analyzing multiple data-independent acquisition mass spectrometry data files. Nat Methods 12:1105–1106 24. Sinitcyn P, Hamzeiy H, Salinas Soto F et al (2021) MaxDIA enables library-based and library-free data-independent acquisition proteomics. Nat Biotechnol 39:1–11 25. Egertson JD, MacLean B, Johnson R et al (2015) Multiplexed peptide analysis using data independent acquisition and skyline. Nat Protoc 10:887–903 26. Kelstrup CD, Bekker-Jensen DB, Arrey TN et al (2018) Performance evaluation of the Q exactive HF-X for shotgun proteomics. J Proteome Res 17:727–738 27. Koopmans F, Ho JTC, Smit AB et al (2018) Comparative analyses of data independent acquisition mass spectrometric approaches: DIA, WiSIM-DIA, and untargeted DIA. Proteomics 18:1700304 28. Ritchie ME, Phipson B, Wu D et al (2015) limma powers differential expression analyses for RNA-sequencing and microarray studies. Nucleic Acids Res 43:e47 29. Smyth G, Hu Y, Ritchie M et al (2020) limma: linear models for microarray data, bioconductor version: release (3.10) 30. Bates D, M€achler M, Bolker B et al (2015) Fitting linear mixed-effects models using lme4. J Stat Softw 67:1–48 31. Bates D, Maechler M, Bolker B et al (2018) lme4: linear mixed-effects models using “Eigen” and S4 32. RStudio Team (2021) RStudio: Integrated Development Environment for R. RStudio, Inc., Boston, MA 33. Chiva C, Olivella R, Borra`s E et al (2018) QCloud: a cloud-based quality control system for mass spectrometry-based proteomics laboratories. PLoS One 13:e0189209 34. Rosenberger G, Koh CC, Guo T et al (2014) A repository of assays to quantify 10,000 human proteins by SWATH-MS. Sci Data 1:140031 35. Reiter L, Rinner O, Picotti P et al (2011) mProphet: automated data processing and statistical validation for large-scale SRM experiments. Nat Methods 8:430–435

INDEX B BAP-EB ................................................................ 182–186 Base editing ................................................. 328, 329, 332

C Cardiac differentiation ............... 38–40, 42, 44–49, 162, 165, 167, 168 Cardiac myocytes ......................................................37–49 Cardiomyocytes..................38–40, 46–48, 161–169, 197 Cell culture .................. 5, 25–35, 37–49, 54, 55, 63–65, 67, 82, 84–86, 94, 101–113, 118–120, 142, 153–157, 162, 169, 183, 190–191, 193, 195, 203, 208, 209, 215, 217, 219, 220, 223–227, 229, 235, 243, 262, 265–272, 275–294, 296, 297, 302, 313, 330, 337–338, 340, 353–355 Cell culture and differentiation ..........108, 337–338, 340 Cell dissociation .............................................65, 183–185 Cell therapy ...................................................... 81–99, 189 CHIR .................... 54, 55, 140, 145–148, 168, 276, 277 Chromatin immunoprecipitation (ChIP) ..................118, 120–129, 131, 132 CRISPR screen............................................................ 1–23

D Data independent acquisition (DIA) ................ 336, 337, 345, 346, 348, 354, 357 Differentiation....................... 4, 25, 38, 53, 59, 82, 102, 118, 136, 152, 161, 173, 181, 190, 199, 216, 234, 261, 266, 275, 295, 309, 336 Disease modeling ................................................... 38, 189

E Ectoderm ........................... 37, 102, 118, 135, 138, 148, 181, 190, 195, 196, 199, 233, 265, 295, 296 Embryoid body (EBs).................. 27, 30–31, 34, 35, 39, 40, 43, 44, 48, 59–78, 82, 83, 88–90, 97, 102, 103, 108–110, 113, 119, 120, 122, 123, 130, 131, 182, 185, 186, 189–197, 215–230, 233–257, 261, 262, 298–302, 305 Embryonic development ....................135, 199, 216, 261 Embryonic stem cell (ESCs) ......................... v, 5, 38, 61, 64–66, 77, 78, 101, 102, 106, 108, 117, 124,

137, 151, 152, 154, 182, 199–212, 215, 216, 261, 265–267, 269–272, 278, 284–288, 290–293, 295, 305, 310, 321–333, 335–337, 340, 346 Endoderm.........................................................37, 69, 71, 102, 118, 181, 190, 195, 196, 199, 233, 265, 290, 295–306 Epiblast-like cells .......................................................53–58 Erythrocytes ........................................172–174, 176, 177

F Feeder free ........................................25–35, 55, 101, 103, 108, 109, 122, 143, 165, 189–197, 218, 219, 223, 224, 226–227, 230, 243, 271 Feeder-free culture ............................26, 64–65, 101, 223 Flow-FISH............................................... 3–5, 7, 8, 15, 21

G Gene silencing ...................................................... 200, 207 Genome editing ............................................................ 171 Genomics ..................................5–6, 8, 12, 17, 129, 164, 166, 168, 234, 256, 322 Glycolysis .............................................................. 152–157 Good manufacturing practice (GMP) ............82, 83, 85, 88, 96, 190

H Hanging drop method...............................39–41, 43–44, 65–67, 113, 119, 130, 305 Hematopoietic stem cell (HSC)................................... 172 Hemogenic endothelial cell (HEC) .................... 172–175 High content image analysis (HCIA) ......................59–78 Human........................................12, 25, 54, 59, 81, 117, 135, 161, 172, 181, 189, 201, 215, 293, 296, 310, 323, 335 Human embryonic stem cells (hESC) ...................25–35, 81–99, 122–123, 129, 130, 132, 137, 148, 152, 163, 181–186, 189–197, 216, 218, 219, 223, 230, 323, 326–328 Human induced pluripotent stem cells (hiPSC) ...........................31, 122, 129, 130, 137, 161–169, 189, 190, 193, 216, 218, 223, 230

Kursad Turksen (ed.), Embryonic Stem Cell Protocols, Methods in Molecular Biology, vol. 2520, https://doi.org/10.1007/978-1-0716-2437-1, © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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P

2i ....................................54, 55, 57, 109, 154, 275–294, 297, 302, 306 2i medium ..........................................153, 275–277, 281, 290–292, 294, 302 Induced pluripotent stem cells (iPSC).................. 25, 31, 37–49, 59, 165, 167, 168, 189, 201, 215, 310, 321–333, 336 Inner ear ............................................................... 135–149 Intracellular antibody staining...................................... 4, 8

PD .....................................................................54, 55, 277 Pluripotency ................................. 25, 26, 38, 53, 56, 57, 101, 102, 117, 130, 131, 151, 152, 181, 189, 190, 196, 199, 209, 211, 216, 233, 265, 269–271, 275, 281, 306, 309, 328 Pluripotent stem cells (PSCs)................... 25, 53, 56, 69, 71, 81, 109, 119, 130, 136, 215, 216, 218, 224 Polycomb group proteins (PcG) ......................... 117–132 Polycomb repressive complex 1 (PRC1) ..................... 118 Polysome profiling analysis........................................... 311 Primary germ layers ..............................38, 265, 295, 296 Proteomics............................................................ 335–359

K Knockout serum replacement (KSR) ............54, 61, 192, 221, 223, 271

L Leukemia inhibitory factor (LIF)............... 5, 40, 54, 55, 57, 61, 62, 64, 65, 101–103, 106–109, 113, 154, 202, 221, 224, 235, 262, 265–267, 269–272, 275–278, 293 Long-term neuroepithelial-like stem cells (Lt-NES).................................... 81–83, 85, 90–99

M MAPK pathway ................................................................. 4 Mass spectrometry (MS)........................... 326, 336–339, 341–346, 354, 356 mESC differentiation ..................... 55, 57, 287, 295–306 Mesoderm...................................37, 102, 118, 172, 181, 190, 195–197, 233, 265, 290, 295–306 Metabolism........................................................... 152, 168 Mouse embryonic fibroblasts (MEFs) .................. 25, 26, 55, 101–113, 183, 184, 186, 189, 218, 220, 222–224, 226, 262, 266–271, 275, 277, 278, 283–286, 288, 291, 293, 323, 327, 328, 332 Mouse embryonic stem cells (mESCs) ............ 53–58, 61, 64, 65, 101–113, 151–158, 200–203, 207–209, 211, 216, 217, 219–226, 228, 230, 233, 235, 243, 244, 255, 261–264, 275–306 Murine embryonic stem cells .......................... 4, 265–272

N Neural differentiation .........................82, 118, 120, 128, 129, 222, 226 Neural progenitors .................................. 81–99, 128, 129 Neural stem cell.................................................... 335–359 Neuronal differentiation .....................117–132, 216, 217 Neutrophil ...........................................173, 174, 176–178

O Organoid ............................................... 59, 135–149, 162 Osteogenic differentiation ................................... 261–264

Q qPCR ........................ 41, 118, 126, 128, 131, 132, 300, 302, 304, 306, 312, 315

R Regenerative medicine ................... 53, 99, 189, 199, 215 Replating cardiomyocytes .................................... 161–169 Ribosome....................................309, 310, 315, 317, 318 RING1B (Really interesting new gene 1B/RING-type E3 ubiquitin transferase RING1)................ 118, 121, 127, 129 RNA ................................... 1–5, 7, 8, 15, 21, 41, 45, 46, 128, 162, 255, 287, 300, 302, 304, 310, 312, 315–318, 322–326, 329, 330, 332, 333 RNA isolation.............................287, 300, 302, 304, 315

S Seahorse ................................................................ 152–157 Self-renewal .................................... 38, 81, 82, 117, 199, 211, 233, 265–272, 275, 295, 309 Sequential Window Acquisition of all Theoretical Mass Spectra (SWATH-MS) ............................. 335–359 sgRNA library.................................................1, 2, 4, 7–20 shRNA .................................................................. 199–212 shRNA screening........................................................... 201 Signaling ............................ 4, 54, 55, 57, 135, 136, 138, 162, 169, 172, 216, 261, 295, 296 Single-cell RNA-seq (scRNA-seq) ........................ 2, 162, 235–243, 253 Skyline................................ 337, 339, 345–354, 357–359 Spectral library............................ 337, 346–350, 357, 358 Stem cells .................... 4, 25, 37, 59, 81, 103, 117, 138, 154, 162, 191, 215, 234, 261, 270, 290, 297, 309, 337 Sucrose density gradient ...................................... 310, 317 Supporting cells.................................................... 135–139

EMBRYONIC STEM CELL PROTOCOLS Index 363 T

Z

Trophectoderm spheroids ................................... 181–186 3D cell models ..........................................................60, 77 3-Dimensional culture .....................................38, 39, 190

Zebrafish ............................................................... 171–179