Pluripotent Stem-Cell Derived Cardiomyocytes (Methods in Molecular Biology, 2320) 1071614835, 9781071614839
This volume provides methodologies for ES and iPS cell technology on the study of cardiovascular diseases. Chapters guid
135 86 10MB
English Pages 318 [295] Year 2021
Preface
Contents
Contributors
Part I: Overview
Chapter 1: Making Cardiomyocytes from Pluripotent Stem Cells
1 Introduction
2 Generating Cardiomyocytes at Large Scale
3 Confirming Function
4 Disease Modeling
5 Conclusions
References
Part II: Generation of Pluripotent Stem Cell-Derived Cardiomyocytes and Cardiac Tissues
Chapter 2: A Method for Cardiac Differentiation, Purification, and Cardiac Spheroid Production of Human Induced Pluripotent St...
1 Introduction
2 Materials
2.1 Cardiac Differentiation
2.2 Reseeding and Purification of Cardiomyocytes
2.3 Production of Cardiac Spheroids
2.4 Immunostaining of hiPSC-Derived Cardiomyocytes
3 Methods
3.1 Passaging hiPSCs and Cardiac Differentiation (Figs. 1 and 2)
3.2 Reseeding and Purification of Cardiomyocytes (Fig. 2)
3.3 Production of Cardiac Spheroids (Fig. 3)
3.4 Immunostaining of hiPSC-Derived Cardiomyocytes (Fig. 4)
4 Notes
References
Chapter 3: Large-Scale Differentiation of Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes by Stirring-Type Suspensi...
1 Introduction
2 Materials
3 Methods
4 Notes
References
Chapter 4: Efficient Method to Dissociate Induced Pluripotent Stem Cell-Derived Cardiomyocyte Aggregates into Single Cells
1 Introduction
2 Materials
2.1 Reagents
2.2 Equipment
3 Methods
4 Notes
References
Chapter 5: Isolation of Cardiomyocytes Derived from Human Pluripotent Stem Cells Using miRNA Switches
1 Introduction
2 Materials
2.1 Synthesis of miRNA Switch
2.2 Preparation of hPSCs and Cardiac Differentiation
2.3 Purification of hPSC-Derived Cardiomyocytes Using miRNA Switch
2.4 Assessment of hPSC-Derived Cardiomyocytes by Immunostaining
3 Methods
3.1 miRNA Switch mRNA Preparation
3.1.1 Preparation of Fragments (ORF, 5′UTR and 3′UTR)
3.1.2 Generating DNA Templates for IVT
3.1.3 IVT
3.2 Preparation of hPSCs and Cardiac Differentiation
3.3 miRNA Switch Transfection (Cell Sorting System)
3.4 miRNA Switch Transfection (Non-sorting System)
3.5 Assessment of hPSC-Derived Cardiomyocytes by Immunostaining
4 Notes
References
Chapter 6: Fabrication of Cardiac Constructs Using Bio-3D Printer
1 Introduction
2 Materials
2.1 Cardiac Spheroid Formation
2.2 Bio-3D Printer
2.3 Fabrication of Cardiac Constructs Using Bio-3D Printer
2.4 Contraction Analysis and Electrical Stimulation
3 Methods
3.1 Cardiac Spheroid Formation
3.1.1 Maintenance of Endothelial Cells and Fibroblasts
3.1.2 Thawing of Human iPSC-CMs
3.1.3 Cardiac Spheroid Formation
3.2 Cardiac Construct Fabrication
3.2.1 Preparation of the Bio-3D Printer
3.2.2 Fabrication of Cardiac Constructs by Bio-3D Printing
3.2.3 Maturation of Cardiac Constructs Using a Bioreactor
3.3 Evaluation of Cardiac Constructs
3.3.1 Contraction Analysis
3.3.2 Electrical Stimulation
4 Notes
References
Chapter 7: Fabrication of Thick and Anisotropic Cardiac Tissue on Nanofibrous Substrate for Repairing Infarcted Myocardium
1 Introduction
2 Materials
2.1 Nanofiber Spinning Materials
2.2 Cell Differentiation and Culture
2.3 Transplantation
3 Methods
3.1 Electrospinning of Nanofiber
3.2 Cardiomyocyte Differentiation
3.3 Cardiomyocyte Culture
3.4 Transplantation
4 Notes
References
Chapter 8: Construction of Three-Dimensional Cardiac Tissues Using Layer-by-Layer Method
1 Introduction
2 Materials
2.1 Reagents
2.2 Equipment
3 Methods
3.1 Cell Preparation
3.2 ECM Coating on the Cell Surface
3.3 Construction of 3D Cardiac Tissues
4 Notes
References
Chapter 9: Generation of Cylindrical Engineered Cardiac Tissues from Human iPS Cell-Derived Cardiovascular Cell Lineages
1 Introduction
2 Materials
2.1 Maintenance and Cardiovascular Differentiation of hiPSCs
2.2 Cell Harvest and Lineage Analysis
2.3 ECT Construction
3 Methods
3.1 Maintenance and Cardiovascular Differentiation of hiPSCs
3.2 Cell Harvest and Lineage Analysis
3.3 ECT Construction
4 Notes
References
Part III: Physiological Measurements Using Pluripotent Stem Cell-Derived Cardiomyocytes
Chapter 10: Protocol for Morphological and Functional Phenotype Analysis of hiPS-Derived Cardiomyocytes
1 Introduction
2 Materials
2.1 Reagents
2.2 Equipment and Software
3 Methods
3.1 Cell Preparation
3.2 Calcium Transients
3.2.1 Indo-1 Loading
3.2.2 Calcium Transient Measurement
3.2.3 Data Analysis
3.3 Field Membrane Potential
3.3.1 FluoVolt Loading
3.3.2 Field Membrane Potential Recording
3.3.3 Data Analysis
3.4 Optical Mapping
3.4.1 Dye Loading Procedure
3.4.2 Optical Membrane Potential Imaging
3.4.3 Data Processing
3.5 Morphological Analysis
3.5.1 Immunofluorescent Staining
3.5.2 Image Processing
4 Notes
References
Chapter 11: Application of FluoVolt Membrane Potential Dye for Induced Pluripotent Stem Cell-Derived Cardiac Single Cells and ...
1 Introduction
2 Materials
2.1 Embryoid Body (EB) Dissociation
2.2 Single-Cell Seeding onto a Fibronectin-Coated Plate
2.3 Fluorescence-Activated Cell Sorting (FACS) and Cryopreservation
2.4 Seeding iPSC-CM Single Cells on a Glass-Bottom Dish
2.5 Seeding High-Density iPSC-CM Monolayers on Glass-Bottom Dishes
2.6 Loading of FluoVolt (FV)
2.7 Recording of APs from Single Cells and Monolayers of Cardiomyocytes
3 Methods
3.1 Embryoid Body (EB) Dissociation on Day 29 (see Note 2)
3.2 Single-Cell Seeding onto a Fibronectin-Coated Plate
3.3 Fluorescence-Activated Cell Sorting (FACS) and Cryopreservation
3.4 Seeding Single iPSC-CMs on a Glass-Bottom Dish
3.5 Seeding High-Density iPSC-CM Monolayer on a Glass-Bottom Dish
3.6 Loading of FluoVolt (FV)
3.7 Recording APs from Single iPSC-CMs and iPSC-CM Monolayers
4 Notes
References
Chapter 12: Multielectrode Array Assays Using Human-Induced Pluripotent Stem Cell-Derived Cardiomyocytes
1 Introduction
2 Materials
2.1 MEA Measurement
2.2 Cell Preparation
3 Methods
3.1 Sterilization of MEA Probes When They Are Used for the First Time
3.2 Coating of MEA Probe
3.3 Plating of iPSC-CMs
3.3.1 Embryoid Body (EB) (See Note 5)
3.3.2 Sheet
3.4 Recording (See Note 7)
3.5 Measurement
3.6 Maintenance of MEA Probes for Reuse
4 Notes
References
Chapter 13: Electrophysiological Analysis of hiPSC-Derived Cardiomyocytes Using a Patch-Clamp Technique
1 Introduction
2 Materials
2.1 Equipment (Figs. 1 and 2)
2.2 Reagents
3 Methods
3.1 Cell Preparation
3.2 Pipette Preparation
3.3 Patch-Clamp Experiment
3.3.1 Current Clamp for Action Potential Recording
3.3.2 Voltage Clamp for Ion Current Recording
3.3.3 Analysis of Each Ion Channel Current
4 Notes
References
Chapter 14: Characterization of Ventricular and Atrial Cardiomyocyte Subtypes from Human-Induced Pluripotent Stem Cells
1 Introduction
2 Materials
2.1 hiPSC Differentiation
2.2 Flow Cytometry
2.3 Quantitative Real-Time PCR
2.4 Whole-Cell Patch-Clamp
3 Methods
3.1 Differentiation of Ventricular- and Atrial-Like Cardiomyocytes
3.2 Flow Cytometry and Cell Sorting
3.3 Quantitative Real-Time PCR
3.4 Whole-Cell Patch-Clamp
4 Notes
References
Chapter 15: Assessment of Contractility in Human iPS Cell-Derived Cardiomyocytes Using Motion Vector Analysis
1 Introduction
2 Materials
3 Methods
3.1 Preparation of iCell Cardiomyocytes
3.2 Application of Compounds
3.3 Video Microscopy for Motion Vector Analysis
3.4 Data Analysis
3.5 Video Microscopy for Motion-Based Propagation Analysis
3.6 Simultaneous Measurement of Motion Vectors and Field Potential
4 Notes
References
Chapter 16: Contractile Force Measurement of Engineered Cardiac Tissues Derived from Human iPS Cells
1 Introduction
2 Materials
2.1 Human iPSC-Derived Cardiomyocyte Culture
2.2 Fibrin Gel Sheet Preparation
2.3 Cardiac Tissue Preparation
2.4 Force Measurement
3 Methods
3.1 Human iPSC-Derived Cardiomyocyte Culture
3.2 Fibrin Gel Sheet Preparation
3.3 Cardiac Tissue Preparation
3.4 Contractile Force Measurement
4 Notes
References
Chapter 17: A Method for Contraction Force Measurement of hiPSC-Derived Engineered Cardiac Tissues
1 Introduction
2 Materials
2.1 ECT Formation
2.1.1 Dissociation of Human iPSC-Derived Cardiomyocytes (iPS-CMs)
2.2 Preparation of Normal Human Dermal Fibroblasts (NHDF)
2.3 ECT Construction
2.4 Contraction Force Measurement
2.5 Analysis of the Acquired Data
3 Methods
3.1 ECT Formation
3.1.1 Dissociation of iPS-CMs
3.1.2 Preparation of NHDF
3.1.3 ECT Construction
3.2 Contraction Force Measurement
3.3 Analysis of the Acquired Data
4 Notes
References
Part IV: Transcriptome and Bioinformatics Analysis
Chapter 18: Single-Cardiomyocyte RNA Sequencing to Dissect the Molecular Pathophysiology of the Heart
1 Introduction
2 Materials
2.1 The Langendorff Perfusion System (Buffers and Reagents)
2.2 The Langendorff Perfusion System (Instruments)
2.3 Single-Cell RNA-Seq
2.4 Single-Cell RNA-Seq Analysis
3 Methods
3.1 Isolation of Adult Mouse Cardiomyocytes Using the Langendorff Perfusion System
3.2 Single-Cell RNA-Seq
3.3 Single-Cell RNA-Seq Analysis (See Note 10)
4 Notes
References
Chapter 19: RNA-Sequencing Analysis of Differentially Expressed Genes in Human iPSC-Derived Cardiomyocytes
1 Introduction
2 Materials
2.1 Induced Cardiomyocytes
2.2 RNA Extraction
2.3 Library Construction
2.4 Sequencing
2.5 Analysis Computer/Server
2.6 Analysis Tools
2.7 Human Genome/Transcriptome Reference Files
3 Methods
3.1 RNA Extraction
3.2 Library Construction
3.3 Sequencing
3.4 Demultiplexing
3.5 Trimming of Adapter Sequences and Removal of Too Short Reads
3.6 Confirmation and Removal of Reads of rRNA Sequences
3.7 Mapping to the Human Genome
3.8 Quality Check by RSeQC
3.9 Counting Reads for Each Transcript
3.10 Normalization
3.11 Further Analysis
4 Notes
References
Chapter 20: Analysis of Transcriptional Profiling of Chamber-Specific Human Cardiac Myocytes Derived from Pluripotent Stem Cel...
1 Introduction
1.1 From Human Induced Pluripotent Stem Cells to Cardiomyocytes
1.2 Sample Preparation and Acquisition of Sequencing Raw Data as a Prerequisite Step in Setting Human iPSC-Derived Cardiomyocy...
1.3 Bioinformatics Workflow Vignettes for Comprehensive Analysis of RNA-Seq Data
1.4 Gene Enrichment Profiling for Cardiac Lineage Specificity
2 Materials
2.1 Computer
2.2 Software
2.3 Data Matrix Files
3 Methods
3.1 Loading the Expression Dataset to Run a Principal Component Analysis (PCA)
4 Notes
References
Part V: Gene Editing and CRISPR Technology for Pluripotent Stem Cells
Chapter 21: Genome Editing in Human Induced Pluripotent Stem Cells (hiPSCs)
1 Introduction
2 Materials
2.1 Plasmid Construction
2.2 Electroporation
2.3 hiPSC Culture and Antibiotics Selection
2.4 Genotype
3 Methods
3.1 Design of gRNA and the HDR Template
3.2 Electroporation into hiPSCs
3.2.1 Electroporation (Day 0)
3.2.2 Puromycin Selection (Day 1~)
3.2.3 Initial Evaluation by Direct Sanger Sequencing (Day 6-7)
3.2.4 Colony Pickup (Day 12-14)
3.2.5 Genotype the Isolated hiPSC Colonies (Day 21~)
4 Notes
References
Chapter 22: Generation of Efficient Knock-in Mouse and Human Pluripotent Stem Cells Using CRISPR-Cas9
1 Introduction
2 Materials
2.1 Mouse Embryonic Stem Cell Culture
2.2 Human Induced Pluripotent Stem Cell Culture
2.3 Electroporation
2.4 Cas9/gRNA-Expressing Plasmid
2.5 Knock-in Construct and Recombinase-Expressing Plasmid
2.6 Antibiotics for Drug Selection
2.7 Software
3 Methods
3.1 Construction of Cas9/gRNA-Expressing Plasmid
3.2 Construction of Knock-In Vectors
3.3 Design of Genotyping Primers
3.4 Knock-in to Mouse ESCs
3.4.1 Gene Targeting with CRISPR-Cas9
3.4.2 Removal of Drug-Resistant Gene
3.4.3 Cloning
3.4.4 Making Stock and Genotyping
3.5 Knock-in to Human iPSCs
3.5.1 Standard Human iPSC Culture
3.5.2 Lipofection
3.5.3 Removal of Drug-Resistant Gene
3.5.4 Cloning
4 Notes
References
Chapter 23: CRISPRi/a Screening with Human iPSCs
1 Introduction
2 Materials
2.1 CRISPRi/a Library
2.2 CRISPRi/a Plasmids
2.3 Lentivirus Production
2.4 iPSC Culture
2.5 Genomic DNA Extraction
2.6 NGS Library Prep and Sequencing
2.7 Data Analysis
3 Methods
3.1 Prepare a CRISPRi iPSC Line
3.1.1 Prepare a CRISPRi iPSC Line and Optimize Culture Condition
3.1.2 Decide the Concentration of Puromycin for Drug Selection
3.2 Test Small-Scale Transduction and Lentiviral Infection
3.2.1 Small-Scale Lentiviral Production
3.2.2 Small-Scale Lentiviral Infection of iPSCs
3.2.3 Evaluate the Knockdown Efficiency in Your Desired Cell Type
3.3 Large-Scale Lentiviral Production for the Pooled gRNA Library
3.3.1 Prepare Lentiviral sgRNA Library
3.3.2 Large-Scale Lentivirus Production
3.3.3 Functional Titration of Lentiviral Library
3.4 Large-Scale Lentiviral Infection of iPSCs
3.4.1 Prepare the Large-Scale Experiments
3.4.2 Lentiviral Library Infection of iPSCs
3.5 Large-Scale CRISPR Screening
3.6 Genomic DNA Extraction
3.7 Next-Generation Sequencing (NGS) to Determine sgRNA Distribution
3.7.1 PCR of Genomic DNA
3.7.2 Purification of Amplified NGS Library by QIAquick Column
3.7.3 Size Selection of NGS Library by AMPure XP Beads
3.7.4 Library QC and Submission for Sequencing
3.7.5 Analyze the Sequencing Results
3.7.6 After Screening
4 Notes
References
Part VI: Transplantation of Pluripotent Stem Cell-Derived Cardiomyocytes
Chapter 24: Transplantation of Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes in a Mouse Myocardial Infarction Mod...
1 Introduction
2 Materials
3 Methods
3.1 Intratracheal Intubation
3.2 Maintenance of Mechanical Ventilation
3.3 Thoracotomy and Myocardial Infarction Surgery
3.4 Transplantation of iPSC-CMs
3.5 Closure of Incision
4 Notes
References
Chapter 25: Transplantation of Pluripotent Stem Cell-Derived Cardiomyocytes into a Myocardial Infarction Model of Cynomolgus M...
1 Introduction
2 Materials
2.1 Medicines
2.2 Surgery-Related Equipment
3 Methods
3.1 Tubes for Blocking the Coronary Artery
3.2 Preoperative Preparation
3.3 Creation of Myocardial Infarction
3.4 Transplantation of Stem Cells-Derived Cardiomyocytes (On Day 14 After Myocardial Infarction)
4 Notes
References
Index
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Methods in Molecular Biology 2320
Yoshinori Yoshida Editor
Pluripotent Stem-Cell Derived Cardiomyocytes
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.
Pluripotent Stem-Cell Derived Cardiomyocytes Edited by
Yoshinori Yoshida Center for iPS Cell Research and Application, Kyoto University, Kyoto, Japan
Editor Yoshinori Yoshida Center for iPS Cell Research and Application Kyoto University Kyoto, Japan
ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1483-9 ISBN 978-1-0716-1484-6 (eBook) https://doi.org/10.1007/978-1-0716-1484-6 © Springer Science+Business Media, LLC, part of Springer Nature 2021 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, 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. 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 Cardiovascular diseases are one of the most common causes of death in the world, and the number of patients is increasing. Currently, strenuous efforts to overcome cardiovascular diseases are being made, and new treatments are being developed. Embryonic stem cells (ES cells) have the capacity of self-renewal and differentiation into many types of cells, including cardiovascular cells, which provide opportunities for researchers to use human cardiovascular cells for disease studies. In addition, induced pluripotent stem cell (iPS cell) technology, which was discovered by Takahashi and Yamanaka in 2006, can generate pluripotent stem cells from the somatic cells of patients by introducing reprogramming factors, thus enabling the study of cardiovascular cells that have the same genetic features as the patients for the investigation of disease mechanisms and development of new therapies for cardiovascular diseases. This book provides information on methodologies for ES and iPS cell technology toward the study of cardiovascular diseases. It includes protocols related to cardiomyocyte generation from pluripotent stem cells, physiological measurements, bioinformatic analysis, gene editing technology, and cell transplantation studies. The detailed protocols in this book will help researchers set up experiments using pluripotent stem cell-derived cardiac cells. I appreciate all the authors of this book for writing their step-by-step protocols and believe the reader will find Pluripotent Stem-Cell Derived Cardiomyocytes helpful for their research. Kyoto, Japan
Yoshinori Yoshida
v
Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
PART I
OVERVIEW
1 Making Cardiomyocytes from Pluripotent Stem Cells . . . . . . . . . . . . . . . . . . . . . . . Peter Karagiannis and Yoshinori Yoshida
PART II
v xi
3
GENERATION OF PLURIPOTENT STEM CELL-DERIVED CARDIOMYOCYTES AND CARDIAC TISSUES
2 A Method for Cardiac Differentiation, Purification, and Cardiac Spheroid Production of Human Induced Pluripotent Stem Cells . . . . . . . . . . . . . . . . . . . . . . Yuika Morita, Shugo Tohyama, Jun Fujita, and Keiichi Fukuda 3 Large-Scale Differentiation of Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes by Stirring-Type Suspension Culture . . . . . . . . . . . Nagako Sougawa, Shigeru Miyagawa, and Yoshiki Sawa 4 Efficient Method to Dissociate Induced Pluripotent Stem Cell-Derived Cardiomyocyte Aggregates into Single Cells. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Emiko Ito, Shigeru Miyagawa, Yoshinori Yoshida, and Yoshiki Sawa 5 Isolation of Cardiomyocytes Derived from Human Pluripotent Stem Cells Using miRNA Switches . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Kenji Miki, Hirohide Saito, and Yoshinori Yoshida 6 Fabrication of Cardiac Constructs Using Bio-3D Printer. . . . . . . . . . . . . . . . . . . . . Kenichi Arai, Daiki Murata, Shoko Takao, and Koichi Nakayama 7 Fabrication of Thick and Anisotropic Cardiac Tissue on Nanofibrous Substrate for Repairing Infarcted Myocardium. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Junjun Li, Li Liu, Itsunari Minami, Shigeru Miyagawa, and Yoshiki Sawa 8 Construction of Three-Dimensional Cardiac Tissues Using Layer-by-Layer Method. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Maki Takeda, Shigeru Miyagawa, Mitsuru Akashi, and Yoshiki Sawa 9 Generation of Cylindrical Engineered Cardiac Tissues from Human iPS Cell-Derived Cardiovascular Cell Lineages . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Hidetoshi Masumoto
vii
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29
35 53
65
75
81
viii
Contents
PART III 10
11
12
13
14
15
16
17
Protocol for Morphological and Functional Phenotype Analysis of hiPS-Derived Cardiomyocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jun Li and Jong-Kook Lee Application of FluoVolt Membrane Potential Dye for Induced Pluripotent Stem Cell-Derived Cardiac Single Cells and Monolayers Differentiated via Embryoid Bodies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tadashi Takaki and Yoshinori Yoshida Multielectrode Array Assays Using Human-Induced Pluripotent Stem Cell-Derived Cardiomyocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Daisuke Yoshinaga, Yimin Wuriyanghai, and Takeru Makiyama Electrophysiological Analysis of hiPSC-Derived Cardiomyocytes Using a Patch-Clamp Technique. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yuta Yamamoto, Sayako Hirose, Yimin Wuriyanghai, Daisuke Yoshinaga, and Takeru Makiyama Characterization of Ventricular and Atrial Cardiomyocyte Subtypes from Human-Induced Pluripotent Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Misato Nakanishi-Koakutsu, Tadashi Takaki, Kenji Miki, and Yoshinori Yoshida Assessment of Contractility in Human iPS Cell-Derived Cardiomyocytes Using Motion Vector Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yasunari Kanda, Ayano Satsuka, Sayo Hayashi, Mihoko Hagiwara-Nagasawa, and Atsushi Sugiyama Contractile Force Measurement of Engineered Cardiac Tissues Derived from Human iPS Cells. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Daisuke Sasaki, Katsuhisa Matsuura, and Tatsuya Shimizu A Method for Contraction Force Measurement of hiPSC-Derived Engineered Cardiac Tissues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yuya Fujiwara, Kohei Deguchi, Kenji Miki, Tomoyuki Nishimoto, and Yoshinori Yoshida
PART IV 18
19
20
PHYSIOLOGICAL MEASUREMENTS USING PLURIPOTENT STEM CELL-DERIVED CARDIOMYOCYTES 91
101
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171
TRANSCRIPTOME AND BIOINFORMATICS ANALYSIS
Single-Cardiomyocyte RNA Sequencing to Dissect the Molecular Pathophysiology of the Heart . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 183 Manami Katoh, Seitaro Nomura, Shintaro Yamada, Hiroyuki Aburatani, and Issei Komuro RNA-Sequencing Analysis of Differentially Expressed Genes in Human iPSC-Derived Cardiomyocytes. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 193 Chikako Okubo, Megumi Narita, Takuya Yamamoto, and Yoshinori Yoshida Analysis of Transcriptional Profiling of Chamber-Specific Human Cardiac Myocytes Derived from Pluripotent Stem Cells . . . . . . . . . . . . . . . . . . . . . 219 Antonio Lucena-Cacace and Yoshinori Yoshida
Contents
PART V 21 22
23
25
GENE EDITING AND CRISPR TECHNOLOGY FOR PLURIPOTENT STEM CELLS
Genome Editing in Human Induced Pluripotent Stem Cells (hiPSCs). . . . . . . . . 235 Shuichiro Higo, Shungo Hikoso, Shigeru Miyagawa, and Yasushi Sakata Generation of Efficient Knock-in Mouse and Human Pluripotent Stem Cells Using CRISPR-Cas9 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 247 Tatsuya Anzai, Hiromasa Hara, Nawin Chanthra, Taketaro Sadahiro, Masaki Ieda, Yutaka Hanazono, and Hideki Uosaki CRISPRi/a Screening with Human iPSCs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 261 Masataka Nishiga, Lei S. Qi, and Joseph C. Wu
PART VI 24
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TRANSPLANTATION OF PLURIPOTENT STEM CELL-DERIVED CARDIOMYOCYTES
Transplantation of Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes in a Mouse Myocardial Infarction Model . . . . . . . . . . . . . . . . . . . 285 Takeshi Hatani and Yoshinori Yoshida Transplantation of Pluripotent Stem Cell-Derived Cardiomyocytes into a Myocardial Infarction Model of Cynomolgus Monkey . . . . . . . . . . . . . . . . . 295 Hideki Kobayashi, Hajime Ichimura, Noburou Ohashi, and Yuji Shiba
Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
303
Contributors HIROYUKI ABURATANI • Genome Science Division, Research Center for Advance Science and Technology, The University of Tokyo, Tokyo, Japan MITSURU AKASHI • Department of Building Block Science, Osaka University Graduate School of Frontier Biosciences, Osaka, Japan TATSUYA ANZAI • Division of Regenerative Medicine, Center for Molecular Medicine, Jichi Medical University, Tochigi, Japan; Department of Pediatrics, Jichi Medical University, Tochigi, Japan KENICHI ARAI • Center for Regenerative Medicine Research, Faculty of Medicine, Saga University, Saga, Japan NAWIN CHANTHRA • Division of Regenerative Medicine, Center for Molecular Medicine, Jichi Medical University, Tochigi, Japan KOHEI DEGUCHI • Takeda-CiRA Joint Program (T-CiRA), Fujisawa, Kanagawa, Japan; T-CiRA discovery, Takeda Pharmaceutical Company Limited, Fujisawa, Kanagawa, Japan JUN FUJITA • Department of Cardiology, Keio University School of Medicine, Tokyo, Japan YUYA FUJIWARA • Department of Cell Growth and Differentiation, Center for iPS Cell Research and Application, Kyoto University, Kyoto, Japan; Takeda-CiRA Joint Program (T-CiRA), Fujisawa, Kanagawa, Japan KEIICHI FUKUDA • Department of Cardiology, Keio University School of Medicine, Tokyo, Japan MIHOKO HAGIWARA-NAGASAWA • Department of Pharmacology, Faculty of Medicine, Toho University, Tokyo, Japan YUTAKA HANAZONO • Division of Regenerative Medicine, Center for Molecular Medicine, Jichi Medical University, Tochigi, Japan; Translational Research Laboratory, Center for Development of Advanced Medical Technology, Jichi Medical University, Tochigi, Japan HIROMASA HARA • Division of Regenerative Medicine, Center for Molecular Medicine, Jichi Medical University, Tochigi, Japan; Animal Resource Laboratory, Center for Development of Advanced Medical Technology, Jichi Medical University, Tochigi, Japan TAKESHI HATANI • Department of Cell Growth and Differentiation, Center for iPS Cell Research and Application, Kyoto University, Kyoto, Japan SAYO HAYASHI • Division of Pharmacology, National Institute of Health Sciences (NIHS), Kawasaki, Japan SHUICHIRO HIGO • Department of Medical Therapeutics for Heart Failure, Osaka University Graduate School of Medicine, Osaka, Japan; Department of Cardiovascular Medicine, Osaka University Graduate School of Medicine, Osaka, Japan SHUNGO HIKOSO • Department of Cardiovascular Medicine, Osaka University Graduate School of Medicine, Osaka, Japan SAYAKO HIROSE • Department of Cardiovascular Medicine, Kyoto University Graduate School of Medicine, Kyoto, Japan HAJIME ICHIMURA • Division of Cardiovascular Surgery, Department of Surgery, Shinshu University School of Medicine, Nagano, Japan MASAKI IEDA • Department of Cardiology, Faculty of Medicine, University of Tsukuba, Tsukuba, Japan
xi
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Contributors
EMIKO ITO • Department of Cardiovascular Surgery, Osaka University Graduate School of Medicine, Osaka, Japan; Cuorips Incorporated, Tokyo, Japan YASUNARI KANDA • Division of Pharmacology, National Institute of Health Sciences (NIHS), Kawasaki, Japan PETER KARAGIANNIS • Center for iPS Cell Research and Application, Kyoto University, Kyoto, Japan MANAMI KATOH • Genome Science Division, Research Center for Advance Science and Technology, The University of Tokyo, Tokyo, Japan HIDEKI KOBAYASHI • Department of Cardiovascular Medicine, Shinshu University School of Medicine, Nagano, Japan ISSEI KOMURO • Department of Cardiovascular Medicine, Graduate School of Medicine, The University of Tokyo, Tokyo, Japan JONG-KOOK LEE • Department of Cardiovascular Regenerative Medicine, Osaka University Graduate School of Medicine, Suita, Japan JUN LI • Department of Cardiovascular Medicine, Osaka University Graduate School of Medicine, Suita, Japan JUNJUN LI • Department of Cardiovascular Surgery, Osaka University Graduate School of Medicine, Osaka, Japan; Department of Design for Tissue Regeneration, Osaka University Graduate School of Medicine, Osaka, Japan LI LIU • Department of Cardiovascular Surgery, Osaka University Graduate School of Medicine, Osaka, Japan; Department of Design for Tissue Regeneration, Osaka University Graduate School of Medicine, Osaka, Japan ANTONIO LUCENA-CACACE • Department of Cell Growth and Differentiation, Center for iPS Cell Research and Application, Kyoto University, Kyoto, Japan TAKERU MAKIYAMA • Department of Cardiovascular Medicine, Kyoto University Graduate School of Medicine, Kyoto, Japan HIDETOSHI MASUMOTO • Clinical Translational Research Program, RIKEN Center for Biosystems Dynamics Research (BDR), Kobe, Japan; Department of Cardiovascular Surgery, Graduate School of Medicine, Kyoto University, Kyoto, Japan KATSUHISA MATSUURA • Institute of Advanced Biomedical Engineering and Science, Tokyo Women’s Medical University, Tokyo, Japan; Department of Cardiology, Tokyo Women’s Medical University, Tokyo, Japan KENJI MIKI • Department of Cell Growth and Differentiation, Center for iPS Cell Research and Application, Kyoto University, Kyoto, Japan; Takeda-CiRA Joint Program (T-CiRA), Fujisawa, Kanagawa, Japan ITSUNARI MINAMI • Department of Cardiovascular Surgery, Osaka University Graduate School of Medicine, Osaka, Japan; Department of Cell Design for Tissue Construction Faculty of Medicine, Osaka University, Osaka, Japan SHIGERU MIYAGAWA • Department of Cardiovascular Surgery, Osaka University Graduate School of Medicine, Osaka, Japan; Department of Frontier Regenerative Medicine, Osaka University Graduate School of Medicine, Osaka, Japan YUIKA MORITA • Department of Cardiology, Keio University School of Medicine, Tokyo, Japan DAIKI MURATA • Center for Regenerative Medicine Research, Faculty of Medicine, Saga University, Saga, Japan MISATO NAKANISHI-KOAKUTSU • Department of Cell Growth and Differentiation, Center for iPS Cell Research and Application, Kyoto University, Kyoto, Japan; Takeda-CiRA Joint Program (T-CiRA), Fujisawa, Kanagawa, Japan KOICHI NAKAYAMA • Center for Regenerative Medicine Research, Faculty of Medicine, Saga University, Saga, Japan
Contributors
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MEGUMI NARITA • Department of Cell Growth and Differentiation, Center for iPS Cell Research and Application (CiRA), Kyoto University, Kyoto, Japan MASATAKA NISHIGA • Stanford Cardiovascular Institute, Stanford University School of Medicine, Stanford, CA, USA TOMOYUKI NISHIMOTO • Takeda-CiRA Joint Program (T-CiRA), Fujisawa, Kanagawa, Japan; T-CiRA discovery, Takeda Pharmaceutical Company Limited, Fujisawa, Japan SEITARO NOMURA • Genome Science Division, Research Center for Advance Science and Technology, The University of Tokyo, Tokyo, Japan; Department of Cardiovascular Medicine, Graduate School of Medicine, The University of Tokyo, Tokyo, Japan NOBUROU OHASHI • Division of Cardiovascular Surgery, Department of Surgery, Shinshu University School of Medicine, Nagano, Japan CHIKAKO OKUBO • Department of Cell Growth and Differentiation, Center for iPS Cell Research and Application (CiRA), Kyoto University, Kyoto, Japan LEI S. QI • Department of Bioengineering, Stanford University School of Medicine, Stanford, CA, USA; Department of Chemical & Systems Biology, Stanford University School of Medicine, Stanford, CA, USA; ChEM-H, Stanford University School of Medicine, Stanford, CA, USA TAKETARO SADAHIRO • Department of Cardiology, Faculty of Medicine, University of Tsukuba, Tsukuba, Japan HIROHIDE SAITO • Department of Life Science Frontiers, Center for iPS Cell Research and Application, Kyoto University, Kyoto, Japan YASUSHI SAKATA • Department of Cardiovascular Medicine, Osaka University Graduate School of Medicine, Osaka, Japan DAISUKE SASAKI • Institute of Advanced Biomedical Engineering and Science, Tokyo Women’s Medical University, Tokyo, Japan AYANO SATSUKA • Division of Pharmacology, National Institute of Health Sciences (NIHS), Kawasaki, Japan YOSHIKI SAWA • Department of Cardiovascular Surgery, Osaka University Graduate School of Medicine, Osaka, Japan YUJI SHIBA • Department of Regenerative Science and Medicine, Institute for Biomedical Sciences, Shinshu University, Nagano, Japan TATSUYA SHIMIZU • Institute of Advanced Biomedical Engineering and Science, Tokyo Women’s Medical University, Tokyo, Japan NAGAKO SOUGAWA • Department of Cardiovascular Surgery, Osaka University Graduate School of Medicine, Osaka, Japan ATSUSHI SUGIYAMA • Department of Pharmacology, Faculty of Medicine, Toho University, Tokyo, Japan TADASHI TAKAKI • Department of Cell Growth and Differentiation, Center for iPS Cell Research and Application (CIRA), Kyoto University, Kyoto, Japan; Takeda-CiRA Joint Program (T-CiRA), Fujisawa, Kanagawa, Japan; Department of Pancreatic Islet Cell Transplantation, National Center for Global Health and Medicine, Tokyo, Japan SHOKO TAKAO • Center for Regenerative Medicine Research, Faculty of Medicine, Saga University, Saga, Japan MAKI TAKEDA • Department of Cardiovascular Surgery, Osaka University Graduate School of Medicine, Osaka, Japan SHUGO TOHYAMA • Department of Cardiology, Keio University School of Medicine, Tokyo, Japan; Department of Organ Fabrication, Keio University School of Medicine, Tokyo, Japan
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Contributors
HIDEKI UOSAKI • Division of Regenerative Medicine, Center for Molecular Medicine, Jichi Medical University, Tochigi, Japan; Translational Research Laboratory, Center for Development of Advanced Medical Technology, Jichi Medical University, Tochigi, Japan JOSEPH C. WU • Stanford Cardiovascular Institute, Stanford University School of Medicine, Stanford, CA, USA; Division of Cardiovascular Medicine, Department of Medicine, Stanford University School of Medicine, Stanford, CA, USA; Department of Radiology, Stanford University School of Medicine, Stanford, CA, USA YIMIN WURIYANGHAI • Department of Cardiovascular Medicine, Kyoto University Graduate School of Medicine, Kyoto, Japan SHINTARO YAMADA • Genome Science Division, Research Center for Advance Science and Technology, The University of Tokyo, Tokyo, Japan; Department of Cardiovascular Medicine, Graduate School of Medicine, The University of Tokyo, Tokyo, Japan TAKUYA YAMAMOTO • Department of Life Science Frontiers, Center for iPS Cell Research and Application (CiRA), Kyoto University, Kyoto, Japan; Institute for the Advanced Study of Human Biology (WPI-ASHBi), Kyoto University, Kyoto, Japan; Medical-Risk Avoidance Based on iPS Cells Team, RIKEN Center for Advanced Intelligence Project (AIP), Kyoto, Japan; AMED-CREST, Tokyo, Japan YUTA YAMAMOTO • Department of Cardiovascular Medicine, Kyoto University Graduate School of Medicine, Kyoto, Japan; Department of Bioscience and Genetics, National Cerebral and Cardiovascular Center, Suita, Japan YOSHINORI YOSHIDA • Center for iPS Cell Research and Application, Kyoto University, Kyoto, Japan DAISUKE YOSHINAGA • Department of Pediatrics, Kyoto University Graduate School of Medicine, Kyoto, Japan
Part I Overview
Chapter 1 Making Cardiomyocytes from Pluripotent Stem Cells Peter Karagiannis and Yoshinori Yoshida Abstract The ability to differentiate pluripotent stem cells to cardiomyocyte lineages (PSC-CMs) has opened the door to new disease models and innovative drug and cell therapies for the heart. Nevertheless, further advances in the differentiation protocols are needed to fulfill the promise of PSC-CMs. Obstacles that remain include deriving PSC-CMs with proper electromechanical properties, coalescing them into functional tissue structures, and manipulating the genome to test the impact mutations have on arrhythmias and other heart disorders. This chapter gives a brief consideration of these challenges and outlines current methodologies that offer partial solutions. Key words Pluripotent stem cells, Cardiomyocytes, Assays, Gene editing, Animal models, RNA sequencing
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Introduction Cardiovascular disease remains one of the most common causes of death. More than 15 million people died of this disease in 2010 alone, and that number is estimated to reach more than 20 million in 2030 [1]. This increase will occur despite improvements in prevention, detection, and treatment. For the most severe cases, heart transplantation is the only curative option, but the number of donors significantly lags behind the number of patients. Donor scarcity is also an obstacle for drug development, as it limits the number of screenings and toxicity assays that can be done. Experimental cell regenerative therapies are hampered for similar reasons. Because of their pluripotency and self-proliferation capacity, human pluripotent stem cells (PSCs), including embryonic stem cells (ESCs) and induced pluripotent stem cells (iPSCs), have promised a bountiful supply of cardiomyocytes (CMs) for research and clinical applications. However, while the number of PSC-CMs from these sources exceeds the number of available primary CMs, the quality is inferior and the production remains costly and inefficient.
Yoshinori Yoshida (ed.), Pluripotent Stem-Cell Derived Cardiomyocytes, Methods in Molecular Biology, vol. 2320, https://doi.org/10.1007/978-1-0716-1484-6_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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The purpose of this book is to provide solutions to these problems by providing methods for PSC-CM differentiation and evaluation by leaders in the field. Other described methods complement these approaches by incorporating technologies that advance disease modeling and cell transplantation therapies.
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Generating Cardiomyocytes at Large Scale In principle, PSCs can be expanded indefinitely, providing a potentially unlimited source. In the case of CMs, large-scale production is especially necessary when considering that at least several hundred million CMs are required for a single cell transplantation therapy to treat the heart. To reach this massive number, Chapter 1 incorporates a metabolic shift into a large-scale culture system to produce PSC-CMs from cardiac spheroids [2]. As another option for the large-scale production of PSC-CMs, Chapter 2 describes a system in which shear stress in a suspension culture is used [3]. While these systems may achieve large scale, single cells must be dissociated from the spheroids before further testing for drug toxicity or cell transplantation. Chapter 3 provides a dissociation method that minimizes damage to the cells. However, the spheroids are a heterogeneous population that includes not only PSC-CMs, but endothelial cells, pericytes and other contaminating cell types; therefore, purification is needed. Transgenic enrichment is one option but disqualifies the cells from various clinical applications because of safety concerns. Traditionally, cell sorting by antibody selection is used, but there is no known surface antigen signature that is exclusive to CMs. Scientists have therefore considered intracellular markers, namely microRNA (miRNA) [4]. Chapter 4 explores this option by explaining how to apply the miRNA switch, a synthetic mRNA that purifies PSC-CM populations at higher efficiency than antibody-based sorting [5]. Of all organs, perhaps none depends more on the coupling of its individual components like the heart. A local perturbation in function can quickly propagate to cause severe heart failure. This feature could explain why the transplantation of individual cells to the heart results in relatively poor electromechanical coupling and limited therapeutic benefit [6, 7]. Instead, 3D structures constituted of organized PSC-CMs are preferred. Protocols that construct these structures are mostly divided into scaffold and non-scaffold. Chapter 5 describes a non-scaffold system that relies on 3D printing and the Kenzan method to create 3D structures from cardiac spheroids [8]. On the other hand, as explained in Chapter 6, scaffolds made of biodegradable synthetic material and coated with ECM components can assure that the PSC-CMs take the proper anisotropic organization to enhance function post-
Making Cardiomyocytes from Pluripotent Stem Cells
5
transplantation. As Chapter 7 shows, mixing mural cells and endothelial cells with PSC-CMs can improve the electromechanical properties of the construct [9]. Furthermore, by adding ECM components between PSC-CM sheets, researchers can prepare vascularized 3D tissues to maintain the PSC-CMs with regular oxygenation and nourishment, as seen in Chapter 8, permitting more complex assays [10].
3
Confirming Function The definition of PSC-CMs used by the above protocols is based primarily on the expression of biomolecules, be it surface antigens, intracellular miRNAs, or others. Once the PSC-CMs are validated molecularly, their function must be confirmed, since heart failure and drug toxicity are based not on marker expression but cardiac performance. Chapter 9 provides a protocol for basic assays on the membrane potential, conductance and other physiological properties of PSC-CMs. One way to directly observe these phenomena is through voltage-sensitive dyes. Chapter 10 describes FluoVolt, a membrane potential dye that responds to voltage fluctuations with microsecond resolution, making it ideal to study the action potential phenotypes of single cells [11]. Complementing these imaging assays are multi-electrode arrays, which are used to evaluate electrical transmissions across cells that could lead to arrhythmic events, as explained in Chapter 11. However, it is the patch clamp that has remained the gold standard for decades. Chapter 12 gives details on the setup of a manual patch clamp. The patch clamp can also be used to distinguish different subpopulations that emerge from the PSC-CM differentiation protocol, including ventricular CMs, atrial CMs and pacemaker cells, which each have their own distinctive electromechanical phenotypes and gene expressions (Chapter 13). Parallel to these electrophysiological assays are mechanical assays that assess the contractility of the PSC-CMs. Motion vector analysis can reveal the electromechanical relationship of PSC-CMs at high resolution (Chapter 14) [12], and there also exist assays to measure the contractile force of whole tissues made of PSC-CMs (Chapters 15 and 16) [13, 14].
4
Disease Modeling One revelation that has come from many functional assays is that PSC-CMs tend to show a fetal phenotype [15]. RNA-sequencing (RNA-seq) has revealed environmental conditions that promote maturation [16]. Moreover, there is significant heterogeneity in
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cell populations even cultured in the same dish. To precisely analyze the quality of PSC-CMs, Chapters 17–19 explore bulk and single cell RNA-seq. These techniques have also assisted in disease modeling by divulging perturbations in gene programs that define morphological and functional checkpoints for proper development [17]. The effects of these gene targets can now be investigated relatively easily by applying gene editing. Progress in this technology has allowed for efficient edits as small as a single base pair [18]. The large number of cells one must screen to identify the correctly edited line demands cells that are easy to expand, i.e., PSCs. CRISPR/Cas9 technology has simplified the production of isogenic PSC lines (Chapter 20), which have advanced our understanding of various cardiomyopathies. Most gene editing has knocked out a target gene, but using CRISPR/Cas9 technology can achieve high efficiency for knocking in a gene as well (Chapter 21) [19]. Finally, CRISPR/Cas9 technology can be used to activate or inhibit candidate causal genes without any edits, as explained in Chapter 22.
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Conclusions The discovery of human ESCs and iPSCs has given expectation of endless amounts of heart cells and tissues for disease modeling, drug discovery, and regenerative therapies. While PSC technology has expanded our resources for cardiac therapies, the scientific community has acknowledged the need for improving the cell product from the differentiation protocols. When developing clinical therapies, it is unlikely necessary to recapture the heart in its truest form. Nevertheless, for PSC-based research to fulfill its promise in heart research, extensive morphological and functional assays of high-quality CMs produced at large scale and low cost are necessary. Following acceptable results from these assays and subsequent preclinical transplantation studies in appropriate animal models (Chapters 23 and 24) [20], there is good reason to believe that patients will soon benefit from drug and cell therapies that are based on PSC-CM research.
References 1. Bansilal S, Castellano JM, Fuster V (2015) Global burden of CVD: focus on secondary prevention of cardiovascular disease. Int J Cardiol 201(Suppl 1):S1–S7 2. Tohyama S et al (2016) Glutamine oxidation is indispensable for survival of human pluripotent stem cells. Cell Metab 23:663–674
3. Chen VC et al (2015) Development of a scalable suspension culture for cardiac differentiation from human pluripotent stem cells. Stem Cell Res 15:365–375 4. Pang JKS, Phua QH, Soh BS (2019) Applications of miRNAs in cardiac development, disease progression and regeneration. Stem Cell Res Ther 10:336
Making Cardiomyocytes from Pluripotent Stem Cells 5. Miki K et al (2015) Efficient detection and purification of cell populations using synthetic microRNA switches. Cell Stem Cell 16:699–711 6. Shiba Y et al (2014) Electrical integration of human embryonic stem cell-derived cardiomyocytes in a guinea pig chronic infarct model. J Cardiovasc Pharmacol Ther 19:368–381 7. Park M, Yoon YS (2018) Cardiac regeneration with human pluripotent stem cell-derived cardiomyocytes. Korean Circ J 48:974–988 8. Moldovan NI, Hibino N, Nakayama K (2017) Principles of the Kenzan method for robotic cell spheroid-based three-dimensional bioprinting. Tissue Eng Part B Rev 23:237–244 9. Masumoto H et al (2016) The myocardial regenerative potential of three-dimensional engineered cardiac tissues composed of multiple human iPS cell-derived cardiovascular cell lineages. Sci Rep 6:29933 10. Amano Y et al (2016) Development of vascularized iPSC derived 3D-cardiomyocyte tissues by filtration layer-by-layer technique and their application for pharmaceutical assays. Acta Biomater 33:110–121 11. Takaki T et al (2019) Optical recording of action potentials in human induced pluripotent stem cell-derived cardiac single cells and monolayers generated from long QT syndrome type 1 patients. Stem Cells Int 2019:7532657 12. Sugiyama A et al (2019) Analysis of electromechanical relationship in human iPS cellderived cardiomyocytes sheets under proarrhythmic condition assessed by simultaneous
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field potential and motion vector recordings. J Pharmacol Sci 140:317–320 13. Ruan JL et al (2016) Mechanical stress conditioning and electrical stimulation promote contractility and force maturation of induced pluripotent stem cell-derived human cardiac tissue. Circulation 134:1557–1567 14. Sasaki D et al (2018) Contractile force measurement of human induced pluripotent stem cell-derived cardiac cell sheet-tissue. PLoS One 13:e0198026 15. Machiraju P, Greenway SC (2019) Current methods for the maturation of induced pluripotent stem cell-derived cardiomyocytes. World J Stem Cells 11:33–43 16. Branco MA et al (2019) Transcriptomic analysis of 3D cardiac differentiation of human induced pluripotent stem cells reveals faster cardiomyocyte maturation compared to 2D culture. Sci Rep 9:9229 17. Nomura S et al (2018) Cardiomyocyte gene programs encoding morphological and functional signatures in cardiac hypertrophy and failure. Nat Commun 9:4435 18. Kim SI et al (2018) Microhomology-assisted scarless genome editing in human iPSCs. Nat Commun 9:939 19. Zhang JP et al (2017) Efficient precise knockin with a double cut HDR donor after CRISPR/ Cas9-mediated double-stranded DNA cleavage. Genome Biol 18:35 20. Shiba Y et al (2016) Allogeneic transplantation of iPS cell-derived cardiomyocytes regenerates primate hearts. Nature 538:388–391
Part II Generation of Pluripotent Stem Cell-Derived Cardiomyocytes and Cardiac Tissues
Chapter 2 A Method for Cardiac Differentiation, Purification, and Cardiac Spheroid Production of Human Induced Pluripotent Stem Cells Yuika Morita, Shugo Tohyama, Jun Fujita, and Keiichi Fukuda Abstract Human induced pluripotent stem cells (hiPSCs) are one of the most promising cell sources for regenerative medicine. To realize the promise of hiPSCs for cardiac regenerative therapy, three major obstacles must be overcome: the first is the achievement of large-scale production of cardiomyocytes, the second is the successful elimination of non-cardiac cells containing residual pluripotent stem cells (PSCs) to prevent tumor formation, and the third is the achievement of high engraftment efficiency of transplanted cardiomyocytes. In this chapter, we introduce our protocols for cardiac differentiation, purification, and preparation of cardiac spheroids for safe and effective regenerative medicine. Key words Induced pluripotent stem cells (iPSCs), Cardiomyocytes, Differentiation, Purification, Spheroid, Transplantation
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Introduction Death due to heart failure is one of the leading causes of deaths worldwide [1]. Heart transplantation remains the treatment of choice for patients with severe heart failure, since an alternative fundamental therapeutic approach has not been established yet. However, heart transplantation has therapeutic limitations due to a shortage of donors. A solution to this problem may be provided by cardiac regenerative therapy with human induced pluripotent stem cells (hiPSCs), a potentially promising alternative to heart transplantation [2–6]. HiPSCs can differentiate into multiple cell types including cardiomyocytes, but not all hiPSCs follow cardiomyocyte lineage specification. As contamination of hiPSC-derivatives with residual pluripotent stem cells (PSCs) is known to be a risk factor for tumorigenesis [7, 8], it is necessary to establish a method to eliminate PSCs and purify only cardiomyocytes. We previously
Yoshinori Yoshida (ed.), Pluripotent Stem-Cell Derived Cardiomyocytes, Methods in Molecular Biology, vol. 2320, https://doi.org/10.1007/978-1-0716-1484-6_2, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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successfully developed a nongenetic fluorescence-activated cell sorting (FACS)-based method for purifying cardiomyocytes [9]. However, that method was not practical because several hundreds of millions of cells are required for the treatment of one heart failure patient. Then, we successfully eliminated residual PSCs and purified only cardiomyocytes with high efficiency and low cost, based on metabolic differences between hiPSCs and differentiated cardiomyocytes [10–12]. Next, to stably produce large numbers of pure cardiomyocytes, we developed a 2D large-scale culture system to obtain several hundreds of millions of cells in one batch [12]. Achieving a high engraftment rate after cell transplantation has been a big challenge for regenerative medicine. Many methodologies (e.g., pro-survival cocktails, gelatin hydrogels) have been developed [13]. We have also demonstrated a high engraftment rate of transplanted cardiac spheroids derived from PSCs [9]. This method has several advantages. For example, it does not require the use of recombinant proteins and scaffolds. However, there is still a need for an efficient cardiac spheroid production method that can be used as cardiac regenerative therapy for heart failure patients. We developed an efficient production system for cardiac spheroids using microwell culture plates on a clinically relevant scale. In this chapter, we would like to introduce these protocols, including protocols for medium-scale production of pure cardiomyocytes and cardiac spheroids.
2
Materials
2.1 Cardiac Differentiation
1. hiPSC line 253G4. 2. Corning® Matrigel® matrix. Store at
30 C.
3. Dulbecco’s phosphate buffered saline D-PBS( ). Store at room temperature. 4. TrypLE Select Enzyme (1), no phenol red. Store at 4 C. 5. Dulbecco’s Modified Eagle Medium (Gibco™ DMEM)/ F12 + Glutamax. Store at 4 C. 6. Stem Fit AS103C Medium. Store at 4 C (see Note 1). 7. Cardiac differentiation medium: Add 10 ml of B-27 without insulin to 500 ml of Roswell Park Memorial Institute (RPMI) 1640 medium with L-glutamine and phenol red. 8. Y-27632 (10 mM stock solution): Directly add 7.39 ml of sterile D-PBS( ) containing 0.1% BSA to 25 mg of Y-27632 to make a 10 mM stock solution. Store the solution at 30 C for up to several months. Use 0.5μl/ml for a final concentration of 5μM.
Production of Highly Purified Cardiomyocytes and Spheroids
13
Table 1 Materials for cardiac differentiation from hiPSCs Material
Company
Cat No.
AS103C
Ajinomoto
custom order
RPMI1640
FUJIFILM Wako Pure Chemical
189-02025
MEM alpha
Thermo Fisher Scientific
12571-048
AS501
Ajinomoto
custom order
DMEM/F12
FUJIFILM Wako Pure Chemical
048-29785
B27 without insulin
Thermo Fisher Scientific
A1895602
Sodium Pyruvate
Sigma
S8636-100ML
FBS
Biowest
S1560-500
CHIR99021
FUJIFILM Wako Pure Chemical
034-23103
BMP4
R&D systems
314-BP-010
IWR1
Sigma
10161-25MG
TrypLE Select
Thermo Fisher Scientific
12563-011
Y-27632
FUJIFILM Wako Pure Chemical
036-24023
Matrigel
Corning
354277
DMSO
Sigma
D2650
9. CHIR99021 (12 mM stock solution): Directly add 900 ml of DMSO to 5 mg of CHIR99021 to make a 12 mM stock solution. Store at 30 C for up to several months. Use 0.5μl/ml for a final concentration of 6μM. 10. BMP-4 (10μg/ml stock solution): Reconstitute 50μg of recombinant BMP-4 in 5 ml D-PBS( ) containing 0.1% BSA. Store at 30 C for up to several months. 11. IWR-1 (10 mM stock solution): Add 6.1 ml DMSO to a vial containing 25 mg of IWR1. Store at 30 C for up to several months. Use 0.5μl/ml for a final concentration of 5μM. 12. Day 0 cardiac differentiation medium: Add 15μl of CHIR99021 and 3μl of BMP-4 to 30 ml of cardiac differentiation medium. Store at 4 C. 13. Day 3 cardiac differentiation medium: Add 15μl of IWR-1 to 30 ml of cardiac differentiation medium. Store at 4 C. 14. Cardiac maintenance medium: Add 5% FBS and 20 ml sodium pyruvate to 1000 ml of α-MEM. Store at 4 C. 15. Falcon® 150 mm cell culture dish with 20 mm molded grid (Table 1).
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2.2 Reseeding and Purification of Cardiomyocytes
1. Fibronectin solution: 50μg/mL fibronectin in D-PBS. Store at 4 C. 2. Trypsin/EDTA solution: 2.5 g/L L-trypsin and 1 mmol/LEDTA in distilled water. Store at 4 C. 3. D-PBS( ). Store at room temperature. 4. Cardiac maintenance medium. Store at 4 C. 5. Stem Fit AS501 medium with lactate, without glucose and glutamine. Store at 4 C (see Note 2). 6. Falcon® 150-mm cell culture dish with 20 mm molded grid.
2.3 Production of Cardiac Spheroids
1. Trypsin/EDTA solution: 2.5 g/L L-trypsin and 1 mmol/LEDTA in distilled water. Store at 4 C. 2. D-PBS( ). Store at room temperature. 3. Cardiac maintenance medium. Store at 4 C. 4. Corning Elplasia 6-well plate.
2.4 Immunostaining of hiPSC-Derived Cardiomyocytes
1. D-PBS ( ). 2. 0.1% Tween20-PBST: 0.1% Tween20 in PBS. 3. 0.1% Triton-PBST: 0.1% TritonX-100 in PBS. 4. 4% Paraformaldehyde (PFA) solution: 4% PFA in PBS. 5. Immunoblock buffer (KAC Co., Ltd). 6. Blocking solution: Dissolve Immunoblock buffer in PBST at a ratio of 1 to 1. 7. Monoclonal anti-sarcomeric α-actinin antibody. 8. Rabbit polyclonal anti-troponin I antibody. 9. Alexa 488 donkey anti-mouse IgG (H + L) highly crossabsorbed secondary antibody. 10. Alexa 546 donkey anti-rabbit IgG (H + L) highly crossabsorbed secondary antibody. 11. DAPI solution: 1μg/ml DAPI in blocking solution.
3
Methods
3.1 Passaging hiPSCs and Cardiac Differentiation (Figs. 1 and 2)
Assume that hiPSCs are maintained in a 100-mm dish. 1. Mix well 1 ml of Matrigel and 150 ml of DMEM/F12, and add 30 ml to each 150-mm cell culture dish. Incubate the coated dishes for at least 1 h at room temperature (see Note 3). 2. After coating the dishes, aspirate the old medium from the hiPSC culture dishes, and wash once with 10 ml of D-PBS( ). 3. Add 2 ml of TrypLE select enzyme and incubate for 1 min at 37 C.
Production of Highly Purified Cardiomyocytes and Spheroids
D- 4 Small molecule Medium
D- 3
D- 1
Y27632
D0
matrigel AS103C
D1
CHIR99021 BMP4
D3
D6
D7
D13
15
D17~20
IWR1
RPMI +B27 insulin ( - )
MEMa
Glc (- ) Gln (- ) Lac (+)
Fig. 1 Schematic representation of the cardiac differentiation protocol
Fig. 2 Time course images of cardiac differentiation of human induced pluripotent stem cells (hiPSCs). Bright field images of the typical morphology at day 0, 1, 3, 6, 7, and 18 are shown. Scale bars, 200m
4. Discard the TrypLE select enzyme and add AS103C with Y27632. 5. Scrape the cells off the bottom of the dish until all the cells detach and pool the harvested cell suspension into a sterile conical tube. Mix gently five to ten times and count the number of cells. 6. Seed 3.0–9.0 106 cells into each Matrigel-coated 150-mm cell culture dish (Day-4). 7. Incubate for 2 days at 37 C, 5% CO2 without changing the medium. 8. After 2 days (Day-2), aspirate the medium and add 30 ml of fresh medium. 9. On the next day (Day-1), aspirate the medium and add 30 ml of fresh medium with 100μl of Matrigel (see Note 4).
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10. On the next day (Day 0), check if the cells are 80–90% confluent, aspirate the old medium and add 30 ml of Day 0 cardiac differentiation medium. Incubate at 37 C, in 5% CO2 incubator overnight (see Note 5). 11. After 24 h (Day 1), aspirate the old medium and wash once with 30 ml of D-PBS( ) (see Note 6). Then, replace the old medium with 30 ml of warm (37 C) RPMI/B-27 medium without insulin. Place the dishes in a 37 C, 5% CO2 incubator. 12. On day 3 of differentiation (Day 3), aspirate the old medium and add Day 3 cardiac differentiation medium. Place the dishes in a 37 C, 5% CO2 incubator. 13. On day 6 of differentiation (Day 6), aspirate the old medium and add 30 ml of RPMI/B27 medium without insulin. If cardiac differentiation was successful, you will see beating cardiomyocytes. Place the dishes in a 37 C, 5% CO2 incubator. 14. On day 7 of differentiation (Day 7), aspirate the old medium and add 30 ml of cardiac maintenance medium. Place the dishes in a 37 C, 5% CO2 incubator. Change the medium every other day until day 11 of differentiation. 3.2 Reseeding and Purification of Cardiomyocytes (Fig. 2)
1. Coat the 150-mm dish with fibronectin solution and incubate for at least 1 h at room temperature. 2. On day 11 of differentiation (Day 11), aspirate the old medium and wash with D-PBS. 3. Aspirate the D-PBS( ) and add 5 ml of 0.25% Trypsin/EDTA solution. Incubate for 5 min in 5% CO2 incubator at 37 C. 4. Check that all the cells are detached and if necessary, tap the dish to help detach all the cells. Pool the cells into a 50-ml conical tube containing 30 ml of cardiac maintenance medium. 5. Centrifuge the cells at 300 g for 3 min at room temperature. Aspirate and discard the supernatant with a sterilized Pasteur pipette. 6. Resuspend the cell pellet in 30 ml of cardiac maintenance medium and count the number of cells (see Note 7). 7. Reseed 2.0–6.0 107 cells into each 150-mm dish with 30 ml of cardiac maintenance medium. Move the dish back and forth and from side to side to disperse the cells on the surface of dish. Incubate for 2 days at 37 C, 5% CO2, without changing the medium. 8. On day 13 of differentiation, add 20 ml of D-PBS( ) (see Note 8). 9. Aspirate D-PBS( ), and add 30 ml of warm Stem Fit AS501 medium with lactate, without glucose and glutamine. Place the dish in a 37 C, 5% CO2 incubator. Change the AS501 medium
Production of Highly Purified Cardiomyocytes and Spheroids
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Fig. 3 Cardiac spheroids production from hiPSCs. Bright field images of the typical morphology of cardiac spheroids are shown. Scale bars, 200m
every day until day 17 of differentiation. You will see non-beating cells. These cells died during lactate selection. 3.3 Production of Cardiac Spheroids (Fig. 3)
1. Prepare the Corning Elplasia plate (6-well plate) and add 2 ml of cardiac maintenance medium and centrifuge at 300 g for 5 min at room temperature. Incubate the plate at 37 C, 5% CO2 (see Note 9). 2. On day 17 of differentiation, aspirate the old medium from the cell culture plate and wash with D-PBS( ). 3. Aspirate the D-PBS, add 5 ml of 0.25% Trypsin/EDTA solution and incubate for 5 min at 37 C, 5% CO2. 4. Check that all the cells are detached and if necessary, tap the dish to help detach all the cells. Pool the harvested cells into a 50-ml conical tube with 1 ml of cardiac maintenance medium and count the number of cells. 5. Reseed 3.0–9.0 106 cells into each well of Corning Elplasia 6-well plate. Move the dish back and forth and from side to side to disperse the cells into each micro space on the surface of the dish. Place the dish in a 37 C, 5% CO2 incubator. 6. On differentiation day 19, you will see cardiac spheroids in each micro space of the plate. Change half of the cardiac maintenance medium. Change the medium every 2 days (see Note 10).
3.4 Immunostaining of hiPSC-Derived Cardiomyocytes (Fig. 4)
1. Aspirate the old medium and wash twice with the D-PBS( ). 2. Fix the cultures for 15 min at room temperature with 4% PFA solution. 3. Discard the 4% PFA solution and wash twice with D-PBS( ). 4. Block the cultures with 0.1% Triton-PBST for 5 min at room temperature.
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Fig. 4 Immunostaining for α-actinin (green), troponin I (red), and DAPI (blue) on cardiac monolayer cultures on day 25. Scale bar, 100m
5. Wash twice with PBST at room temperature. 6. Block the cultures with the Immunoblock buffer for 30 min at room temperature. 7. Discard the Immunoblock buffer and incubate the cells with the anti-sarcomeric α-actinin antibody (1:500) and troponin I antibody (1:100) in blocking solution (Immunoblock buffer and PBST at 1:1 ratio) at 4 C overnight. 8. Collect the primary antibody solution for re-use and keep it at 4 C. 9. Wash twice with PBST. 10. Block the cells with Immunoblock buffer for 10 min at room temperature. 11. Stain the cells with the Alexa Fluor 488 anti-mouse IgG (H + L) antibody (1:200) and Alexa 546 anti-rabbit IgG (H + L) antibody (1:200) in blocking solution (Immunoblock buffer and PBST at 1:1 ratio) overnight at 4 C. Protect the cells from light since a fluorescent dye-conjugated secondary antibody was used. 12. Discard the secondary antibody solution and wash twice with PBST. 13. Stain with DAPI solution for 10 min at room temperature. 14. Wash twice with PBST. 15. Cultures are kept in Immunoblock buffer at 4 C.
Production of Highly Purified Cardiomyocytes and Spheroids
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Notes 1. Although AS103C medium is not commercially available, common medium for hiPSC maintenance culture will also work. 2. Although AS501 medium is not commercially available, glucose- and glutamine-depleted DMEM or RPMI (no glucose, no glutamine, no pyruvate) supplemented with 4 mM lactate will also work. 3. Matrigel solidifies irreversibly at room temperature. Therefore, it is advisable to dissolve the Matrigel in cold medium and coat the dishes immediately. 4. As mentioned in see Note 3, Matrigel should be dissolved in cold medium immediately. Do not add the Matrigel into warm AS103C medium. We found that the efficiency of cardiac differentiation is increased when Matrigel coating is performed the day before cardiac induction. 5. The step of changing the medium after 24 h from Day 0 has to be performed with great precision, so recording the time of medium change (from AS103C medium to Day 0 cardiac differentiation medium) is important. 6. As mentioned in see Note 5, you need to change the Day 1 cardiac induction medium 24 h later. 7. Cardiomyocytes tend to stick with each other when the volume of the culture medium is small. Therefore, it is advisable to use a larger volume of medium. 8. The purification efficacy of the lactate medium decreases if there are remnants of glucose or glutamine from the α-MEM medium. Therefore, it is advisable to wash off the α-MEM medium completely with D-PBS(-). 9. Simply adding the medium to this plate will not allow bubbles to escape into the micro space, and the medium will not enter the micro space. 10. It is advisable not to shake the plate because the cardiac spheroids might come out of the micro space. When changing the medium, use a PIPETMAN to slowly discard the old medium and slowly add the new medium.
Acknowledgments The present work was mainly supported by Projects for Technological Development, Research Center Network for Realization of Regenerative Medicine by Japan, the Japan Agency for Medical Research and Development grant (19bm0404023h0002 to S. T.), JSPS KAKENHI grant (20J01097 to Y.M.) and JSPS KAKENHI grant (19K22626 to S. T.).
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References 1. Benjamin EJ, Blaha MJ, Chiuve SE, Cushman M, Das SR, Deo R, de Ferranti SD, Floyd J, Fornage M, Gillespie C, Isasi CR, Jimenez MC, Jordan LC, Judd SE, Lackland D, Lichtman JH, Lisabeth L, Liu S, Longenecker CT, Mackey RH, Matsushita K, Mozaffarian D, Mussolino ME, Nasir K, Neumar RW, Palaniappan L, Pandey DK, Thiagarajan RR, Reeves MJ, Ritchey M, Rodriguez CJ, Roth GA, Rosamond WD, Sasson C, Towfighi A, Tsao CW, Turner MB, Virani SS, Voeks JH, Willey JZ, Wilkins JT, Wu JH, Alger HM, Wong SS, Muntner P, American Heart Association Statistics C, Stroke Statistics S (2017) Heart disease and stroke statistics2017 update: a report from the American Heart Association. Circulation 135(10):e146– e603. https://doi.org/10.1161/CIR. 0000000000000485 2. Shiba Y, Gomibuchi T, Seto T, Wada Y, Ichimura H, Tanaka Y, Ogasawara T, Okada K, Shiba N, Sakamoto K, Ido D, Shiina T, Ohkura M, Nakai J, Uno N, Kazuki Y, Oshimura M, Minami I, Ikeda U (2016) Allogeneic transplantation of iPS cellderived cardiomyocytes regenerates primate hearts. Nature 538(7625):388–391. https:// doi.org/10.1038/nature19815 3. Tabei R, Kawaguchi S, Kanazawa H, Tohyama S, Hirano A, Handa N, Hishikawa S, Teratani T, Kunita S, Fukuda J, Mugishima Y, Suzuki T, Nakajima K, Seki T, Kishino Y, Okada M, Yamazaki M, Okamoto K, Shimizu H, Kobayashi E, Tabata Y, Fujita J, Fukuda K (2019) Development of a transplant injection device for optimal distribution and retention of human induced pluripotent stem cellderived cardiomyocytes. J Heart Lung Transplant 38 (2):203–214. https://doi.org/10.1016/j. healun.2018.11.002 4. Chong JJ, Yang X, Don CW, Minami E, Liu YW, Weyers JJ, Mahoney WM, Van Biber B, Cook SM, Palpant NJ, Gantz JA, Fugate JA, Muskheli V, Gough GM, Vogel KW, Astley CA, Hotchkiss CE, Baldessari A, Pabon L, Reinecke H, Gill EA, Nelson V, Kiem HP, Laflamme MA, Murry CE (2014) Human embryonic-stem-cell-derived cardiomyocytes regenerate non-human primate hearts. Nature 510(7504):273–277. https://doi.org/10. 1038/nature13233 5. Kawamura M, Miyagawa S, Fukushima S, Saito A, Miki K, Funakoshi S, Yoshida Y, Yamanaka S, Shimizu T, Okano T, Daimon T, Toda K, Sawa Y (2017) Enhanced therapeutic
effects of human iPS cell derivedcardiomyocyte by combined cell-sheets with omental flap technique in porcine ischemic cardiomyopathy model. Sci Rep 7(1):8824. https://doi.org/10.1038/s41598-01708869-z 6. Liu YW, Chen B, Yang X, Fugate JA, Kalucki FA, Futakuchi-Tsuchida A, Couture L, Vogel KW, Astley CA, Baldessari A, Ogle J, Don CW, Steinberg ZL, Seslar SP, Tuck SA, Tsuchida H, Naumova AV, Dupras SK, Lyu MS, Lee J, Hailey DW, Reinecke H, Pabon L, Fryer BH, MacLellan WR, Thies RS, Murry CE (2018) Human embryonic stem cell-derived cardiomyocytes restore function in infarcted hearts of non-human primates. Nat Biotechnol 36 (7):597–605. https://doi.org/10.1038/nbt. 4162 7. Hentze H, Soong PL, Wang ST, Phillips BW, Putti TC, Dunn NR (2009) Teratoma formation by human embryonic stem cells: evaluation of essential parameters for future safety studies. Stem Cell Res 2(3):198–210. https:// doi.org/10.1016/j.scr.2009.02.002 8. Zhang Y, Wang D, Chen M, Yang B, Zhang F, Cao K (2011) Intramyocardial transplantation of undifferentiated rat induced pluripotent stem cells causes tumorigenesis in the heart. PLoS One 6(4):e19012. https://doi.org/10. 1371/journal.pone.0019012 9. Hattori F, Chen H, Yamashita H, Tohyama S, Satoh YS, Yuasa S, Li W, Yamakawa H, Tanaka T, Onitsuka T, Shimoji K, Ohno Y, Egashira T, Kaneda R, Murata M, Hidaka K, Morisaki T, Sasaki E, Suzuki T, Sano M, Makino S, Oikawa S, Fukuda K (2010) Nongenetic method for purifying stem cell-derived cardiomyocytes. Nat Methods 7(1):61–66. https://doi.org/10.1038/nmeth.1403 10. 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. https://doi.org/10.1016/j. stem.2012.09.013 11. Tohyama S, Fujita J, Hishiki T, Matsuura T, Hattori F, Ohno R, Kanazawa H, Seki T, Nakajima K, Kishino Y, Okada M, Hirano A, Kuroda T, Yasuda S, Sato Y, Yuasa S, Sano M, Suematsu M, Fukuda K (2016) Glutamine
Production of Highly Purified Cardiomyocytes and Spheroids oxidation is indispensable for survival of human pluripotent stem cells. Cell Metab 23 (4):663–674. https://doi.org/10.1016/j. cmet.2016.03.001 12. Tohyama S, Fujita J, Fujita C, Yamaguchi M, Kanaami S, Ohno R, Sakamoto K, Kodama M, Kurokawa J, Kanazawa H, Seki T, Kishino Y, Okada M, Nakajima K, Tanosaki S, Someya S, Hirano A, Kawaguchi S, Kobayashi E, Fukuda K (2017) Efficient large-scale 2D culture system for human induced pluripotent stem cells and differentiated cardiomyocytes. Stem Cell
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Rep 9(5):1406–1414. https://doi.org/10. 1016/j.stemcr.2017.08.025 13. Laflamme MA, Chen KY, Naumova AV, Muskheli V, Fugate JA, Dupras SK, Reinecke H, Xu C, Hassanipour M, Police S, O’Sullivan C, Collins L, Chen Y, Minami E, Gill EA, Ueno S, Yuan C, Gold J, Murry CE (2007) Cardiomyocytes derived from human embryonic stem cells in pro-survival factors enhance function of infarcted rat hearts. Nat Biotechnol 25(9):1015–1024. https://doi. org/10.1038/nbt1327
Chapter 3 Large-Scale Differentiation of Human Induced Pluripotent Stem Cell-Derived Cardiomyocytes by Stirring-Type Suspension Culture Nagako Sougawa, Shigeru Miyagawa, and Yoshiki Sawa Abstract Regenerative medicine using human-induced pluripotent stem cells (hiPSCs) is a promising approach to treat heart failure. However, a large number of cells are required to achieve the desired therapeutic effect. The stirring-type suspension culture method allows a large-scale production of hiPSC-derived cardiomyocytes (more than 1 108 cells/100 mL), leading to a stable cell supply. Here, we describe a method to scale-up hiPSC-derived cardiomyocyte production with a high differentiation efficiency. Key words hiPSCs, Cardiomyocytes, Large-scale, Differentiation, Stirring-type suspension culture
1
Introduction Regenerative medicine is considered a promising field for treatment of heart failure. Previous studies have shown that myoblast sheet transplantation improves cardiac function in various types of heart failure [1], and several lines of evidence have suggested that improvement of cardiac function mainly contributes to the paracrine mechanism [2, 3]. However, in severe heart failure with a massive loss of cardiomyocytes, it is difficult to regenerate the myocardium with a paracrine effect alone. Therefore, we aimed at a myocardial regeneration therapy using hiPSC-derived cardiomyocytes for clinical applications. A significant number of cells are required to attain therapeutic efficacy. Moreover, providing stable and inexpensive cell supplies is a must for clinical applications. hiPSCs differentiate into cardiomyocytes through formation of an embryoid body (EB) and monolayer culture [4–6]. The monolayer-based method is expensive and has a limitation in the culture area. In contrast, the EB formation method, especially the stirring-type suspension culture method, is able to expand the culture volume, leading to an easy scale-up.
Yoshinori Yoshida (ed.), Pluripotent Stem-Cell Derived Cardiomyocytes, Methods in Molecular Biology, vol. 2320, https://doi.org/10.1007/978-1-0716-1484-6_3, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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Here, we have described a method to obtain numerous cardiomyocytes derived from hiPSCs with a high efficiency of differentiation.
2
Materials 1. Primate ES Cell Medium (ReproCELL). 2. Mitomycin C-treated mouse embryonic fibroblasts (MEFs). 3. Dissociation solution for human ES/iPS cells (ReproCELL). 4. Accumax. 5. Y27632. 6. Differentiation medium: StemPro34 (gibco) medium containing 50 μg/mL ascorbic acid, 2 mM L-glutamine, and 400 μM 1-thioglycerol. 7. Ascorbic acid. 8. L-glutamine 200 mM, 100. 9. 1-thioglycerol. 10. Bone morphogenetic protein-4 (BMP4). 11. Activin A. 12. Basic fibronectin growth factor (bFGF). 13. IWR-1. 14. IWP-2. 15. Vascular endothelial growth factor (VEGF). 16. A single-use bioreactor (100 mL or 30 mL) (ABLE Co.). 17. Magnetic stirrer. 18. 70-μm nylon cell strainer. 19. Phosphate buffered saline (PBS).
3
Methods The method of cardiac differentiation described here is based on a previous report [7] with some modifications. Bring the medium or PBS to room temperature before use. 1. Human iPSCs are maintained in the Primate ES Cell Medium supplemented with 5 ng/mL bFGF on MEFs. Cells are passaged every 3–4 days using the Dissociation solution. 2. Following a wash with PBS, collect hiPSCs on the MEF in culture dishes (100 mm dish) using Accumax (see Note 1). 3. Centrifuge at 190 g for 3 min at room temperature.
Large-Scale Differentiation of Human Induced Pluripotent Stem Cell-Derived. . .
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Fig. 1 Scheme of the cardiac differentiation from hiPSCs. Cells are cultured with differentiation medium containing suitable additives by stirring-system. Cytokines are added at day 1. Medium is changed every 2 days after day 4
Fig. 2 Representative phase-contrast images at day 4, 6, 8, 11, and 13. Scale bar, 100 m. Some spontaneous beating EBs are observed from day 11
4. Discard the supernatant, resuspend the cells in the differentiation medium containing 5 μM Y27632, pass through a 70-μm nylon cell strainer, and count the cell number (see Note 2). 5. Suspend 1.2 107 to 1.8 107 cells in 100 mL of the differentiation medium containing 5 μM Y27632 and 5 ng/ mL BMP4, and seed them into a bioreactor. 6. Stir the floating culture in a 37 C incubator supplied with 20% O2. Set the agitation rate at 40 rpm (day 0). Thereafter, keep stirring the floating culture until the end of induction of differentiation (see Note 3). 7. After 24 h (day 1), add BMP4, bFGF, and Activin A (see Note 4). 8. At day 4, replace the medium with a fresh differentiation medium containing IWR-1 and IWP-2. 9. After day 6, replace the medium with a fresh differentiation medium containing VEGF and bFGF every 2 days (see Note 5). 10. At day 15, collect cells from the bioreactor and use them for the following experiments (see Note 6) (Figs. 1 and 2).
4
Notes 1. Recover the hiPSCs, including the feeder cells. If the feeder cells are removed in advance, EB formation does not proceed well.
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2. When the number of hiPSCs is less than 1 107 cells/100 mm dish, the differentiation efficiency tends to be low. 3. During stirring, ensure that the temperature in the bioreactor is maintained. 4. The optimal concentration of additives depends on the manufacturers of the additives or the hiPSCs you use; therefore, optimization of the concentration is required in each experiment. The concentrations of the additives used generally are as follows: Additive
Concentration
Day 1
BMP4 Activin A bFGF
8–10 6–14 5–10
ng/mL ng/mL ng/mL
Day 4
IWR-1 IWP-2
4–8 4–8
μM μM
Day 6~
bFGF VEGF
5–10 2.5–10
ng/mL ng/mL
5. At day 13 or day 14, the differentiation efficacy is checked. If the differentiation efficiency is 60% or higher, addition of bFGF is not necessary. 6. In the case of continuing culture until day 17, replace the utilized medium with a fresh differentiation medium containing additives on day 15 or day 16. Culturing for more than 17 days makes the EB hard and makes them difficult to dissociate.
Acknowledgment This was supported by Japan Agency for Medical Research and Development (AMED) under Grant Number JP20bm0204003. References 1. Sawa Y, Yoshikawa Y, Toda K et al (2015) Safety and efficacy of autologous skeletal myoblast sheets (TCD-51073) for the treatment of severe chronic heart failure due to ischemic heart disease. Circ J 79:991–999 2. Matsuura K, Honda A, Nagai T et al (2009) Transplantation of cardiac progenitor cells ameliorates cardiac dysfunction after myocardial infarction in mice. J Clin Invest 119:2204–2217
3. Masumoto H, Matsuo T, Yamamizu K et al (2012) Pluripotent stem cell-engineered cell sheets reassembled with defined cardiovascular populations ameliorate reduction in infarct heart function through cardiomyocyte-mediated neovascularization. Stem Cells 30:1196–1205 4. Yang L, Soonpaa MH, Adler ED et al (2008) Human cardiovascular progenitor cells develop
Large-Scale Differentiation of Human Induced Pluripotent Stem Cell-Derived. . . from a KDR+ embryonic-stem-cell-derived population. Nature 453:524–528 5. Willems E, Spiering S, Davidovics H et al (2011) Small-molecule inhibitors of the Wnt pathway potently promote cardiomyocytes from human embryonic stem cell-derived mesoderm. Circ Res 109:360–364 6. Uosaki H, Fukushima H, Takeuchi A et al (2011) Efficient and scalable purification of
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cardiomyocytes from human embryonic and induced pluripotent stem cells by VCAM1 surface expression. PLoS One 6:e23657 7. Matsuura K, Wada M, Shimizu T et al (2012) Creation of human cardiac cell sheets using pluripotent stem cells. Biochem Biophys Res Commun 425:321–327
Chapter 4 Efficient Method to Dissociate Induced Pluripotent Stem Cell-Derived Cardiomyocyte Aggregates into Single Cells Emiko Ito, Shigeru Miyagawa, Yoshinori Yoshida, and Yoshiki Sawa Abstract The human adult heart consists of approximately four billion cardiomyocytes, which do not possess selfrenewal abilities. Severe myocardial infarction and dilated cardiomyopathy result in the loss of more than a billion cardiomyocytes. Induced pluripotent stem cells (iPSCs) can differentiate into various types of cells. Due to this ability, these cells could potentially serve as a new resource for cell therapy. Many studies have utilized cardiomyocytes derived from iPSCs for myocardial regeneration therapy. To obtain large number of cardiomyocytes for transplantation, we need to develop effective methods that would allow us to dissociate multiple cardiomyocyte aggregates simultaneously. Here, we describe a method to efficiently dissociate large number of iPSC-derived cardiomyocyte aggregates. Key words Large scale culture, Induced pluripotent stem cells, Cardiomyocyte, Aggregate, Dissociate
1
Introduction Replacement therapies such as ventricular assist device and heart transplantation are useful for severe heart failure, especially in the cases where medical treatment alone is insufficient for complete recovery. However, these replacement therapies for severe heart failure pose many problems, such as chronic donor shortage and immunosuppression [1–4]. There are no universally applicable treatments that can be used successfully for all patients with severe heart failure. In recent years, studies have shown that cell transplantation is useful for recovering cardiac function. The clinical application of somatic stem cells for treatment of severe heart failure has already started in several countries in Europe, the United States, and Japan [5, 6]. Embryonic stem cells and induced pluripotent stem cells (iPSCs) have self-renewal abilities and differentiate into all cell types of the human body [7, 8]. This property makes them potentially useful resources in the development of regenerative therapy.
Yoshinori Yoshida (ed.), Pluripotent Stem-Cell Derived Cardiomyocytes, Methods in Molecular Biology, vol. 2320, https://doi.org/10.1007/978-1-0716-1484-6_4, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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The heart requires a large number of highly safe cardiomyocytes [9]. Therefore, it is necessary to develop a new method that efficiently produces numerous cardiomyocytes from iPSCs. Some of the methods used for cardiac differentiation include two-dimensional culture, embryoid body formation, and threedimensional agitation culture in the bioreactor [10–13]. The three-dimensional agitation culture in the bioreactor system is able to achieve stable and efficient production of cardiomyocytes on a large scale because of its ability to culture cells at high density and to control the pH and oxygen levels in the medium during differentiation [14]. To achieve a wider application range, it is necessary to efficiently dissociate iPSC-derived cardiomyocyte aggregates. In cardiac regenerative treatments for transplantation, these cells reconstruct a three-dimensional myocardial tissue such as a cell sheet. It is also possible to construct three-dimensional cardiac tissues with rich vascular networks by co-culturing cardiomyocytes and vascular endothelial cells [15–17]. In this section, we describe an efficient method to dissociate large quantities of iPSC-derived cardiomyocyte aggregates into single cells with minimal damage.
2 2.1
Materials Reagents
1. Collagenase solution (2 mg/mL): Dissolve 500 mg of collagenase type I in 197.5 mL of D-PBS ( ). Add 50 mL of fetal bovine serum and 2.5 mL of PBS containing 10 g/L of calcium chloride and 10 g/L of magnesium chloride 6H2O. After filter sterilization by using 0.22 μm filter, store the solution at 30 C. 2. EDTA solution (1 mmol/L): Prepare 1 mmol/L EDTA solution by mixing 0.5 mol/L EDTA solution and D-PBS ( ) and store at room temperature. 3. TrypLE select solution (3): Prepare 3 TrypLE select solution by mixing 6 mL of TrypLE™ Select (10), 14 mL of 1 mmol/L EDTA solution, 2 μL of 2 U/μL DNase I, and 20 μL of 10 mM Y-27632 before use. 4. DMEM buffer: Dulbecco’s Modified Eagle Medium (DMEM) containing 10% fetal bovine serum. Store at 4 C.
2.2
Equipment
1. Nalgene™ Rapid-Flow™ Sterile Disposable Bottle Top Filters with PES Membrane 0.2 μm (Thermo Fisher). 2. 50-ml tube. 3. Beaker. 4. Stirring bar. 5. Magnetic stirrer.
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6. CO2 incubator (5%). 7. 100-μm Cell Strainer. 8. Cell counter.
3
Methods 1. Collect the iPSC-derived cardiomyocyte aggregates in a (Fig. 1a) 50 mL tube and centrifuge at 240 g for 1 min (see Note 1). 2. Aspirate the supernatant and add 20 mL of HBSS. Centrifuge at 240 g for 1 min. 3. Repeat step 2 once. 4. Aspirate the HBSS and add collagenase solution. Use 20 mL of collagenase solution for 1 108 cardiomyocytes after dissociation. 5. Incubate at 37 C for 1 h (see Note 2). 6. After treatment with the collagenase solution, centrifuge at 240 g for 1 min. 7. Aspirate the collagenase solution and add 20 mL of HBSS. Centrifuge at 240 g for 5 min (Fig. 1b). 8. Aspirate the HBSS and add 3 TrypLE select solution. Use 20 mL of 3 TrypLE select solution (see Note 3). 9. Incubate at 37 C for 10 min (see Note 2). 10. After treatment with 3 TrypLE select solution, suspend the iPSC-derived cardiomyocyte aggregates (see Note 4). 11. Add DMEM containing 10% FBS and an equal amount of 3 TrypLE select solution (see Note 4). 12. Centrifuge at 240 g for 5 min, at 4 C. Next, aspirate the supernatant. 13. Add the medium according to the subsequent assay (Fig. 1c). Count the cardiomyocytes and use for each assay.
Fig. 1 Representative image of dissociative iPS-derived cardiomyocyte. (a) After treatment with the collagenase solution. (b) After treatment with 3 TrypLE select solution. (c) After filter by using 100-m cell strainer
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Notes 1. In steps 1–6, the deceleration setting of the centrifuge could be used to perform centrifugation at a slow speed. 2. Performing the collagenase and 3 TrypLE select solution treatments with stirring yields a large number of cardiomyocytes. 3. Enzyme treatment results in higher instances of cell death in cardiomyocyte clumps. To avoid this, add DNase I and Y-27632 during the 3 TrypLE select solution treatment. For dissociation of 1 108 cardiomyocytes, add 2 μL of 2 U/μL DNase I and 20 μL of 10 mM Y-27632 to 20 mL of 3 TrypLE select solution. Addition of Y-27632 to the medium used for suspending the cardiomyocytes is also recommended. 4. The cardiomyocytes are resuspended by tapping the tube gently. The cardiomyocytes should be pipetted gently. Avoid pipetting with the pipette tip.
References 1. Lund LH, Edwards LB, Kucheryavaya AY et al (2014) The registry of the International Society for Heart and Lung Transplantation: thirtyfirst official adult heart transplant report. J Heart Lung Transplant 33:996–1008 2. El-Banayosy A, Ko¨rfer R, Arusoglu L et al (2001) Device and patient management in a bridge-to-transplant setting. Ann Thorac Surg 71:S98–S102 3. Piccione W Jr (2001) Bridge to transplant with the HeartMate device. J Card Surg 16:272–279 4. Miyagawa S, Domae K, Kainuma S et al (2018) Long-term outcome of a dilated cardiomyopathy patient after mitral valve surgery combined with tissue-engineered myoblast sheets-report of a case. Surg Case Rep 4:142 5. Vrtovec B, Sever M, Jensterle M et al (2016) Efficacy of CD34+ stem cell therapy in nonischemic dilated cardiomyopathy is absent in patients with diabetes but preserved in patients with insulin resistance. Stem Cells Transl Med 5:632–638 6. Miyagawa S, Domae K, Yoshikawa Y et al (2018) Phase I clinical trial of autologous stem cell-sheet transplantation therapy for treating cardiomyopathy. J Am Heart Assoc 6: e003918 7. Efthymiou AG, Chen G, Rao M et al (2019) Self-renewal and cell lineage differentiation
strategies in human embryonic stem cells and induced pluripotent stem cells. Expert Opin Biol Ther 14:1333–1344 8. Kempf H, Olmer R, Kropp C et al (2014) Controlling expansion and cardiomyogenic differentiation of human pluripotent stem cells in scalable suspension culture. Stem Cell Rep 3:1132–1146 9. Burridge PW, Holmstro¨m A, Wu JC (2015) Chemically defined culture and cardiomyocyte differentiation of human pluripotent stem cells. Curr Protoc Hum Genet 87:21.3.1–21.3.15 10. Lian X, Zhang J, Azarin SM et al (2013) Directed cardiomyocyte differentiation from human pluripotent stem cells by modulating Wnt/β-catenin signaling under fully defined conditions. Nat Protoc 8:162–175 11. Hemmi N, Tohyama S, Nakajima K et al (2014) A massive suspension culture system with metabolic purification for human pluripotent stem cell-derived cardiomyocytes. Stem Cells Transl Med 3:1473–1483 12. Matsuura K, Wada M, Shimizu T et al (2012) Creation of human cardiac cell sheets using pluripotent stem cells. Biochem Biophys Res Commun 425:321–327 13. Matsuura K, Wada M, Konishi K et al (2012) Fabrication of mouse embryonic stem cellderived layered cardiac cell sheets using a bioreactor culture system. PLoS One 7:e52176
Dissociate iPS-Derived Cardiomyocyte Aggregates 14. Kawamura M, Miyagawa S, Miki K et al (2012) Feasibility, safety, and therapeutic efficacy of human induced pluripotent stem cell-derived cardiomyocyte sheets in a porcine ischemic cardiomyopathy model. Circulation 126:S29–S37 15. Rogozhnikov D, O’Brien PJ, Elahipanah S et al (2016) Scaffold free bio-orthogonal assembly of 3-dimensional cardiac tissue via cell surface engineering. Sci Rep 6:39806 16. Takeda M, Miyagawa S, Fukushima S et al (2018) Development of in vitro drug-induced
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cardiotoxicity assay by using three-dimensional cardiac tissues derived from human induced pluripotent stem cells. Tissue Eng Part C Methods 24:56–67 17. Beauchamp P, Jackson CB, Ozhathil LC et al (2020) 3D co-culture of hiPSC-derived cardiomyocytes with cardiac fibroblasts improves tissue-like features of cardiac spheroids. Front Mol Biosci 7:14
Chapter 5 Isolation of Cardiomyocytes Derived from Human Pluripotent Stem Cells Using miRNA Switches Kenji Miki, Hirohide Saito, and Yoshinori Yoshida Abstract The most common method for isolating cells of interest is an antibody method that recognizes cell surface antigens. However, specific surface antigens for many cell types have not been identified. We have developed the microRNA (miRNA)-responsive synthetic mRNA systems (miRNA switches), which isolate target cells based on endogenous miRNA activity. In this chapter, we describe protocols for isolating human pluripotent stem cell (hPSC)-derived cardiomyocytes using miRNA switches with or without cell sorting. Key words miRNA switch, Synthetic modified mRNA, Human pluripotent stem cells, Cardiomyocytes, Isolation
1
Introduction Human embryonic and induced pluripotent stem cell (hPSC)derived cardiomyocytes provide opportunities to develop new drugs and cell therapies for cardiovascular diseases [1–3]. There are many publications on cardiac differentiation methods using hPSCs [4–8], but no matter which method is used, hPSC-derived differentiated cells include not only cardiomyocytes but also nontarget cells such as fibroblasts, endothelial cells, pericytes, and/or undetermined cells. Several approaches for the isolation of hPSCderived cardiomyocytes have been devised. For example, genetic manipulation methods using a reporter gene under the control of cardiac promotors can isolate hPSC-derived cardiomyocytes with high efficiency [9–12]. However, this strategy has safety concerns for cell therapy and is unsuitable for studies using cells derived from dozens of patients. On the other hand, antibody methods are the most common methods for isolating target cells. For example, some surface antigens such as signal-regulatory protein alpha (SIRPA) and vascular cell adhesion molecule 1 (VCAM1) have been shown to isolate cardiomyocytes efficiently [13, 14]. However, these surface antigens are also expressed on other cell types
Yoshinori Yoshida (ed.), Pluripotent Stem-Cell Derived Cardiomyocytes, Methods in Molecular Biology, vol. 2320, https://doi.org/10.1007/978-1-0716-1484-6_5, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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miRNA switch target miRNA reporter protein complementary sequence sequence
synthetic modifed mRNA ・Cells do not express target miRNA. miRNAs
translation
Reporter proteins are expressed.
nucleous
・Cells express target miRNA. miRNAs RISC
nucleous
RISC
translational repression and mRNA cleavage
Reporter proteins are repressed.
Fig. 1 Purification outline using the miRNA switch. Top: Construction of the miRNA switch. Bottom: Mechanism of the miRNA switch
[13, 15], and no cardiomyocyte-specific surface antigen has been identified. Previously, we developed the miRNA switch system to isolate hPSC-derived cardiomyocytes with or without cell sorting [16]. The miRNA switch is a synthetic modified mRNA that contains the sequence complementary to a target miRNA at the 50 -UTR and a sequence encoding genes of interest, such as a fluorescent protein or apoptosis-related gene (Fig. 1 top). The protein expression from the mRNA depends on the activity of the target miRNAs in the cell (Fig. 1 bottom). This system has great potential for preparing safe cardiomyocytes because this system uses RNA transfection, and most of the transfected mRNAs are degraded within 48 h after the transfection. In this chapter, we outline protocols to synthesize mRNA switches and isolate hPSCderived cardiomyocytes.
2
Materials
2.1 Synthesis of miRNA Switch
1. Primers (see Table 1). 2. KOD-Plus Neo. 3. DpnI. 4. MegaScript T7 Kit. 5. RNeasy MiniElute Cleanup Kit. 6. Antarctic Phosphatase. 7. Pseudouridine-50 -triphosphate.
Rev
CAGTGAATTGTAATACGACTCACTATAGGGC
TAP_T7_G3C fwd primer
5UTRtemp_T208a-3p CGACTCACTATAGGTTCCGCGATCGCGGATCCACAAGCTTTTTGCTC GTCTTATAGATCACACCGGTCGCCACCATG
3UTR120A rev primer TTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTT TTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTTT TTTTTTTTTTCCTACTCAGGCTTTATTCA
GCTAATACGACTCACTATAGGTTCCTTAATCGCGGATCC
GCCCCGCAGAAGGTCTAGATCAGGCACCGGGCTTGC
CACCGGTCGCCACCATGACCGAGTACAAGCCCACG
GCCCCGCAGAAGGTCTAGAATCAATGCATTCTCCACACCAG
CACCGGTCGCCACCATG
GCCCCGCAGAAGGTCTAGACTATCACTCGAGATGCATATGAGATC
CACCGGTCGCCACCATGGGATCCGTGAGCAAGGGC
GCCCCGCAGAAGGTCTAGACTATCACTCGAGATGCATATGAGATC
CACCGGTCGCCACCATGGGATCCAGCGAG
TCTAGACCTTCTGCGGGGC TTTTTTTTTTTTTTTTTTTTCCTACTCAGGCTTTATTCAAAGACCAAG TCTAGACCTTCTGCGGGGCTTGCCTTCTGGCCATGCCCTTCTTCTCTCCCT TGCACCTGTACCTCTTGGTCTTTGAATAAAGCCTGAGTAGG
CATGGTGGCGACCGGTGTCTTATATTTCTTCTTACTC CAGTGAATTGTAATACGACTCACTATAGGGCGAATTAAGAGAGA AAAGAAGAGTAAGAAGAAATATAAGACACCGGTCGCCACCATG
CAGTGAATTGTAATACGACTCACTATAGGGC
Sequence (50 to 30 )
GCT7pro_5UTR2
Puror_IVTrev
Rev
Fwd(miRNA switch) Fwd(control)
Puror_IVTfwd
BimEL_IVTrev
Rev
Fwd
BimEL_IVTfwd
TAP_IVTrev
Rev
Fwd
TAPEGFP_IVTfwd
TAP_IVTrev
Rev
Fwd
tagBFP fwd
Fwd3UTR primer Rev3UTR2T20 IVT_3prime_UTR
TAP_T7_G3C fwd primer Rev5UTR primer IVT_5prime_UTR
Primer name
Fwd
Fwd Rev Template
Rev Template
Fwd
50 oligo miR-208a-3p switch
DNA template for IVT
Puror ORF
BimEL ORF
EGFP ORF
tagBFP ORF
30 UTR
50 UTR
Table 1 Primer list
A method for Isolation of Cardiomyocytes Using miRNA Switch 37
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8. 5-methylcytidine-50 -triphosphate. 9. Anti-Reverse Cap Analog. 10. MiniElute PCR purification Kit. 11. Rneasy Minielute Cleanup Kit. 12. Total RNA Extraction Column (FAVORGEN, FARBC-C50). 13. NanoDrop 2000c. 2.2 Preparation of hPSCs and Cardiac Differentiation
1. Primate ES cell medium. 2. D-PBS () without Ca and Mg. 3. Penicillin/streptomycin. 4. Transferrin. 5. Collagenase type IV. 6. Knockout Serum Replacement (KSR). 7. 10 Gelatin stock solution: 1 g of gelatin powder in 100 ml of distilled water. Autoclave and store at 4 C for up to 2 months. 8. Gelatin working solution: Warm the 10 gelatin stock solution to 37 C, and mix 50 ml of 10 gelatin stock solution with 450 ml of distilled water. Filter the solution using a bottle-top filter (0.22 μm) and store at 4 C for up to 2 weeks. 9. Gelatin-coated culture dishes: Add a sufficient volume of gelatin working solution so that a culture dish is totally covered. For example, 3 ml of gelatin working solution is used for a 60-mm dish. Incubate the dish for at least half an hour and no more than 4 h at 37 C. Aspirate excess gelatin working solution before use. 10. Trypsin-EDTA (0.25%). 11. Human recombinant basic fibroblast growth factor (bFGF) for hPSC medium: Dissolve 100 μg of human recombinant bFGF powder in 10 ml of 0.1% (w/v) BSA/PBS to a concentration of 10μg/ml and store the aliquots at 30 C. 12. hPSC medium: Add 200 μl of 10 μg/ml bFGF and 2.5 ml of 10,000 U and 10,000 mg/ml penicillin and streptomycin in Primate ES medium. 13. CTK solution: Mix 50 ml of 2.5% Trypsin, 100 ml of KSR, 50 ml of 1 mg/ml collagenase type IV, 5 ml of 0.1 M CaCl2 solution and 295 ml of sterilized water, filter the solution with a bottle-top filter (0.22 μm) and store the aliquots at 30 C. 14. StemPro™-34 SFM (1). 15. Matrigel Matrix Growth Factor Reduced: Mix 10 ml of Matrigel and 10 ml of IMDM and store the aliquots at 30 C. 16. Accumax. 17. L-GIutamin 200 mM.
A method for Isolation of Cardiomyocytes Using miRNA Switch
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18. Ascorbic acid solution: Dissolve 250 mg of ascorbic acid in 50 ml of water to a concentration of 5 mg/ml, filter the solution with a bottle-top filter and store the aliquots at 30 C. 19. MTG (1-Thyoglycerol). 20. Collagenase type I: Dissolve 500 mg of Collagenase type I powder in 50 ml FBS, 200 ml PBS and 2 ml Ca2+Mg2+ solution, filter the solution with a bottle-top filter (0.22 μm) and store the aliquots at 30 C. 21. Iscove’s modified Dulbecco’s media (IMDM). 22. ROCK inhibitor: Dissolve 5 mg of ROCK inhibitor in 1.48 ml of cell culture–grade water to a concentration of 10 mM stock solution and store the aliquots at 30 C. 23. BMP4 recombinant human: Dissolve 50 μg of human recombinant BMP4 powder in 5 ml of 4 mM HCl and 0.1% (w/v) BSA/PBS to a concentration of 10 μg/ml and store the aliquots at 80 C. 24. Activin A recombinant human: Dissolve 50 μg of human recombinant Activin A powder in 5 ml of 0.1% (w/v) BSA/PBS to a concentration of 10 μg/ml and store the aliquots at 80 C. 25. bFGF for cardiac differentiation: Dissolve 25 μg of human recombinant bFGF powder (R&D) in 2.5 ml of 1 mM DTT and 0.1% (w/v) BSA/PBS to a concentration of 10 μg/ml and store the aliquots at 80 C. 26. VEGF recombinant human: Dissolve 50 μg of human recombinant VEGF powder in 5 ml of 0.1% (w/v) BSA/PBS to a concentration of 10 μg/ml and store the aliquots at 80 C. 27. IWP-3: Dissolve 2 mg of IWP-3 in 413 μl of DMSO to a concentration of 10 mM and store the aliquots at 30 C. 28. Aggregation medium: StemPro™-34 media containing 2 mM L-glutamine, 4 10–4 M MTG, 50 μg/ml ascorbic acid, 150 μg/ml transferrin, 0.5% penicillin/streptomycin, 10 μM ROCK inhibitor and 2 ng/ml BMP4. 29. Induction medium (I1): StemPro™-34 media containing 2 mM L-glutamine, 4 10–4 M MTG, 50 μg/ml ascorbic acid, 150 μg/ml transferrin, 0.5% penicillin/streptomycin, bFGF (10 ng/ml, 5 ng/ml final), activin A (12 ng/ml, 6 ng/ml final) and BMP4 (18 ng/ml, 10 ng/ml final). 30. Induction medium (I2): StemPro™-34 media containing 2 mM L-glutamine, 4 10–4 M MTG, 50 μg/ml ascorbic acid, 150 μg/ml transferrin, 0.5% penicillin/streptomycin, 10 ng/ml VEGF and 1 μM IWP-3.
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31. Induction medium (I3): StemPro™-34 media containing 2 mM L-glutamine, 4 10–4 M MTG, 50 μg/ml ascorbic acid, 150 μg/ml transferrin, 0.5% penicillin/streptomycin and 5 ng/ml VEGF. 32. STOP medium: Mix 50 ml of FBS into 50 ml of IMDM. 33. SNL feeder cells [1]. 34. 60-mm cell culture dishes. 35. 15-ml conical tubes. 2.3 Purification of hPSC-Derived Cardiomyocytes Using miRNA Switch
1. Fibronectin stock solution: Dissolve 5 mg of fibronectin powder in 5 ml sterilized water to a concentration of 1 mg/ml and store the aliquots at 80 C. 2. Fibronectin working solution: Dilute the fibronectin stock solution with PBS to a concentration of 50 μg/ml just before use. 3. Fibronectin-coated plates: Add a sufficient volume of fibronectin working solution to fully cover a culture plate. For example, 500 μl or 1.5 ml of fibronectin working solution is required for one well of a 24-well plate or 6 well-plate, respectively. Incubate the plate for at least 2 h at 37 C, and aspirate excess fibronectin working solution before use. 4. Opti-MEM medium. 5. Lipofectamine™ MessengerMAX™ Transfection Reagent. 6. FACS buffer: Mix 2 ml of FBS into 98 ml of PBS. 7. BD FACS Aria Flow cytometer. 8. Puromycin: Dissolve 25 mg of puromycin powder in 2.5 ml of PBS and store the aliquots at 30 C.
2.4 Assessment of hPSC-Derived Cardiomyocytes by Immunostaining
1. 4% Paraformaldehyde (PFA). 2. Normal goat serum. 3. Bovine serum albumin (BSA). 4. Triton X-100. 5. 0.1% (v/v) Triton X-100 in D-PBS. 6. Blocking buffer: 5% (v/v) normal goat serum and 0.1% (v/v) Triton X-100 in D-PBS. 7. Cardiac Troponin T Monoclonal Antibody (ThermoFisher, MA5-12960, 1:200). 8. Goat anti-mouse IgG-Alexa Fluor 488 (Life Technologies, 1:500). 9. Hoechst 33342 (Life Technologies; 1:10,000).
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Methods
3.1 miRNA Switch mRNA Preparation
We first prepared fragments of the open reading frame (ORF) of a reporter protein, the 50 UTR and the 30 UTR. We fused these fragments to generate DNA templates for in vitro transcription (IVT). miRNA-responsible mRNAs were synthesized using a MegaScript T7 kit via IVT.
3.1.1 Preparation of Fragments (ORF, 50 UTR and 30 UTR)
1. Prepare the PCR reaction mixtures (see Tables 2, 3 and 4). 2. Run the PCR (see Tables 5 and 6).
Table 2 PCR reaction components for the ORF fragment Component
Volume (μl)
Final concentration
10 KOD-Plus-Neo buffer
5.0
1
2 mM dNTPs
5.0
200 μM
25 mM MgSO4
3.0
1.5 mM
10 μM Fwd 30 UTR
1.5
0.3 μM
10 μM Rev 3 UTR
1.5
0.3 μM
50 ng/ml plasmid
5.0
10 nM
KOD-Plus-Neo (1 U/μl)
1.0
0.02 U/μl
D2W
28.0
up to 50 μl
Total
50.0
0
Table 3 PCR reaction components for the 50 UTR Component
Volume (μl)
Final concentration
10 KOD-Plus-Neo buffer
5.0
1
2 mM dNTPs
5.0
200 μM
25 mM MgSO4
3.0
1.5 mM
10 μM Fwd 50 UTR
1.5
0.3 μM
1.5
0.3 μM
100 nM temp 5 UTR
5.0
10 nM
KOD-Plus-Neo (1 U/μl)
1.0
0.02 U/μl
D2W
28.0
up to 50 μl
Total
50.0
0
10 μM Rev 5 UTR 0
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Table 4 PCR reaction components for the 30 UTR Component
Volume (μl)
Final concentration
10 KOD-Plus-Neo buffer
5.0
1
2 mM dNTPs
5.0
200 μM
25 mM MgSO4
3.0
1.5 mM
1.5
0.3 μM
10 μM Rev 3 UTR
1.5
0.3 μM
100 nM temp 30 UTR
5.0
10 nM
KOD-Plus-Neo (1 U/μl)
1.0
0.02 U/μl
D2W
28.0
up to 50 μl
Total
50.0
0
10 μM Fwd 3 UTR 0
Table 5 PCR cycle condition for the ORF fragment 94 C
2 min
10 s
68 C
30 s
15 C
1
98 C
20 cycles
Table 6 PCR cycle condition for the 50 and 30 UTR fragments 94 C
2 min
98 C
10 s
68 C
10 s
15 C
13 cycles
1
3. Regarding the ORF fragments, add 1 μl of DpnI to the reactions and incubate at 37 C for 30 min. 4. Prepare 3 new 1.5 ml tubes and add 250 μl of PB solution (MiniElute PCR purification kit) to each. 5. Add 5 μl of 3 M sodium acetate (pH 5.2) to each PCR product (50 UTR, 30 UTR and DpnI-treated ORF). 6. Add each mixture in step 5 to a tube in step 4 and mix well. 7. Transfer each mixture to a MiniElute column.
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Table 7 PCR reaction components for the DNA template Component
Volume (μl)
Final concentration
10 KOD-Plus-Neo buffer
5.0
1
2 mM dNTPs
5.0
200 μM
25 mM MgSO4 aq
3.0
1.5 mM
10 μM Fwd primer
1.5
0.3 μM
10 μM Rev primer
1.5
0.3 μM
10 ng/μl ORF PCR product
1.0
0.2 ng/μl
100 nM 50 UTR
5.0
10 nM
100 nM 3 UTR PCR product
5.0
10 nM
KOD-Plus-Neo (1 U/μl)
1.0
0.02 U/μl
D2W
22.0
up to 50 μl
Total
50.0
0
Table 8 PCR cycle condition for the DNA template 94 C
2 min
98 C
10 s
30 s
45 s
1
60 C 68 C 15 C
20 cycles
8. Centrifuge at 10,000 g, 4 C for 1 min and discard the flowthrough. 9. Add 650 μl of Buffer PE to the columns. 10. Centrifuge at 10,000 g, 4 C for 1 min at and discard the flow-through. 11. Transfer each column to a new 1.5 ml tube, add 15 μl of DNase-free water and centrifuge at 10,000 g, 4 C for 1 min. 12. Measure the concentration of the samples with NanoDrop 2000c. 3.1.2 Generating DNA Templates for IVT
1. Prepare the PCR reaction mixtures using the fragments in step 12 of Subheading 3.1.1 (see Table 7 and see Note 1). 2. Run the PCR (see Table 8). 3. Purify the PCR products using steps 4–12 in Subheading 3.1.1.
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Table 9 Components for IVT Component
Volume (μl)
Final concentration
PCR product (100 ng/μl)
1.0
10 ng/μl
10 buffer
1.0
1
75 mM CAP analog
0.8
6 mM
75 mM GTP
0.2
1.5 mM
75 mM ATP
1.0
7.5 mM
75 mM 5-me-CTP
1.0
7.5 mM
75 mM pseud-UTP
1.0
7.5 mM
T7 enzyme
1.0
10 nM
D2W
3.0
10 μl
Total
10.0
3.1.3 IVT
1. Prepare the reaction mixtures using the MEGAscript T7 Transcription Kit (see Table 9). 2. Incubate the mixtures at 37 C for 4 h. 3. Add 1 μl of TURBO DNase to the mixtures and incubate at 37 C for 30 min. 4. Add 90 μl of RNase-free water to the mixtures, mix them, and add 350 μl of RLT buffer (RNeasy MiniElute Cleanup Kit). 5. Add 250 μl of EtOH to the mixtures, mix them, and transfer them to a column. 6. Centrifuge at 10,000 g, room temperature for 15 s and discard the flow-through. 7. Add 500 μl of buffer RPE (RNeasy MiniElute Cleanup Kit), centrifuge at 10,000 g, room temperature for 15 s and discard the flow-through. 8. Add 500 μl of 80% EtOH, centrifuge at 10,000 g for 15 s at room temperature and discard the flow-through. 9. Centrifuge at 20,400 g, room temperature for 5 min. 10. Prepare a new 1.5 ml tube and put the column in it. 11. Add 40 μl of RNase-free water to the column and centrifuge at 20,400 g, room temperature for 1 min. 12. Add the full dosage of the eluate to the column and centrifuge at 20,400 g, room temperature for 1 min. 13. Add 4 μl of 10 Antarctic Phosphatase reaction buffer and 1μl of Antarctic Phosphatase to the eluted samples and incubate them at 37 C for 30 min.
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14. Repeat steps 4–12 in Subheading 3.1.3. 15. Measure the concentration of the samples with NanoDrop 2000c. 16. Store at 30 C. 3.2 Preparation of hPSCs and Cardiac Differentiation
1. Maintain hPSCs on SNL feeder cells (60-mm dish) with hPSC medium until the cells reach about 80–90% confluence. 2. Aspirate the medium from an hPSC dish and wash the cells with 2 ml of PBS twice. 3. Remove the PBS completely, add 1 ml of CTK solution to remove the SNL feeder cells and incubate at 37 C for 2–3 min. 4. Remove the CTK solution completely and wash the cells with 2 ml of PBS. 5. Remove the PBS completely, add 1 ml of Accumax and incubate at 37 C for 3–5 min until the cells detach from the dish with a tap. 6. Pipette up and down to break up the colony to single cells (pipette a maximum of five times) (see Note 2). 7. Transfer the cells into a 15 ml conical centrifuge tube with 5 ml of hPSC medium and centrifuge at 300 g, room temperature for 5 min. 8. Remove the supernatant and suspend the pellet in 3 ml of aggregation medium. 9. Count the cell number and adjust the concentration to 1 106 cells per ml with aggregation medium. 10. Seed the hPSCs at 5 105 cells (500 μl) per well into a 6-well ultra-low attachment plate with 1.0 ml aggregation medium/ well (final 1.5 ml/well) and incubate at 37 C in a 5% CO2, 5% O2, 90% N2 environment for 1 day. During this time, the hPSCs make embryoid bodies (EBs) (see Note 3). 11. On day 1, add 1.5 ml of I1 medium to well and incubate at 37 C in a 5% CO2, 5% O2, 90% N2 environment for 2 days. At this point, the total amount of medium is 3 ml, and the concentrations of Activin A, BMP4, and bFGF are 6 ng/ml, 10 ng/ml and 5 ng/ml, respectively. 12. On day 3, collect the EBs into a 15 ml conical centrifuge tube and stand for 2–3 min at room temperature until the EBs sink down. 13. Remove the supernatant carefully, add 5 ml of IMDM and stand for 2–3 min at room temperature until the EBs sink down.
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14. Remove IMDM carefully, transfer the EBs into a 6-well ultralow attachment plate with 2 ml of I2 medium and incubate at 37 C in a 5% CO2, 5% O2, 90% N2 environment for 3 days. 15. On day 6, collect the EBs into a 15 ml conical centrifuge tube and stand for 2–3 min at room temperature until the EBs sink down. 16. Remove the supernatant carefully, add 5 ml of IMDM and stand for 2–3 min at room temperature until the EBs sink down. 17. Remove IMDM carefully, transfer the EBs into a 6-well ultralow attachment plate with 2 ml of I3 medium and incubate at 37 C in a 5% CO2, 5% O2, 90% N2 environment for 4 days. Refresh I3 medium every 2 days. 18. On day 10, transfer the plate to a 5% CO2 incubator at 37 C (normoxia environment). Refresh I3 medium every 2 days. 19. In this protocol, differentiation efficiencies are 20–60% depending on the cell line. 3.3 miRNA Switch Transfection (Cell Sorting System)
This protocol is for a 6-well plate. 1. On day 17, remove the medium from the EBs, add 2 ml of collagenase type I solution and incubate at 37 C for 1–2 h. 2. Remove the collagenase type I solution from the EBs, add 1 ml of 0.25% (w/v) trypsin/1 mM EDTA and incubate at 37 C for 5 min. 3. Add 1 ml of Stop medium and pipette up and down to break up the EBs to single cells (pipette a maximum of ten times) (see Note 4). 4. Transfer the cells into a 15 ml conical centrifuge tube with 5 ml of IMDM and centrifuge at 300 g for 5 min at room temperature. 5. Remove the supernatant and suspend the pellet in 3 ml of I3 medium. 6. Count the cell number and seed the cells at 1.0–1.5 106 cells/well in 6-well fibronectin-coated plates (see Note 5). 7. Incubate at 37 C and 5% CO2 for 2 days. 8. Remove the medium, wash with IMDM, and add 2 ml of fresh I3 medium without antibiotics to the well before the transfection. 9. Prepare two sterilized 1.5 ml tubes and add 125 μl/tube of Opti-MEM medium into both tubes. 10. In the first tube, add 3.75 μl of Lipofectamine™ MessengerMAX™ Transfection Reagent and incubate for 10 min.
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11. In the second tube, add 4 μl of 100 ng/μl miRNA-208a-BFP switch and 4 μl of 100 ng/ml control EGFP mRNA and mix by pipetting (see Note 6). 12. Add the mixture in step 10 to the mixture in step 11, mix by pipetting, and incubate for 5 min at room temperature. 13. Add the mRNA complex into the well and incubate at 37 C and 5% CO2 for 4 h (see Note 7). 14. Remove the medium containing the complex and wash with IMDM, add 2 ml of fresh I3 medium to the well and incubate at 37 C and 5% CO2 (see Note 8). We always use the sample the next day for cell sorting. 15. Aspirate the medium from the plate and wash the cells with 2 ml of PBS. 16. Add 1 ml of 0.25% (w/v) trypsin/1 mM EDTA and incubate at 37 C for 5 min. 17. Add 1 ml of Stop medium and pipette to detach the cells (pipette a maximum of five times). 18. Transfer the cells into a 15 ml conical centrifuge tube with 5 ml of IMDM and centrifuge at 300 g for 5 min at room temperature. 19. Discard the supernatant and suspend the pellet in 1–2 ml of FACS buffer. 20. Sort the EGFP+BFP population using a BD FACS Aria flow cytometer (Fig. 2a, b).
a
b
miRNA-208a-BFP switch
miRNA switch
BFP
Control mRNA
EGFP
Non-target cells
BFP
target miRNA site
Target cells
EGFP
BFP
BFP
c
EGFP
cTNT/Hoechst
Repress BFP signals EGFP
Fig. 2 miRNA switch-purification of target cells using cell sorting. (a) Difference of expression patterns of two fluorescent proteins in nontarget cells and target cells. BFP translation is repressed in target cells because the miRNA switch interacts with the endogenous target miRNA. (b) Practical example of miRNA208a switch using hPSC-derived differentiated cells. (c) Immunostaining of cTNT in miR-208a-BFP switch sorted cells. Scale bar, 100 μm
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21. Centrifuge at 300 g for 5 min at room temperature. 22. Resuspend the cells in 1 ml of I3 medium, count the cell number and seed the cells to a fibronectin-coated plate at 1–1.5 105 cells/cm2 for immunostaining. 3.4 miRNA Switch Transfection (Non-sorting System)
This protocol is for 6-well plates. 1. Conduct steps 1–10 in Subheading 3.3. 2. In the second tube, add 1.5 or 2 μl of 100 ng/μl miRNA-208aBim switch and 2 μl of 100 ng/μl puromycin resistance mRNA and mix by pipetting (Fig. 3a). 3. Add the first mixture to the second mixture, mix by pipetting, and incubate for 5 min at room temperature. 4. Add the mRNA complex into the well and incubate at 37 C and 5% CO2 for 4 h. 5. Remove the medium containing the complex, wash with IMDM and add 2 ml of fresh I3 medium containing 2 μg/ml puromycin to the well and incubate at 37 C and 5% CO2 for 2–3 days (see Note 9).
3.5 Assessment of hPSC-Derived Cardiomyocytes by Immunostaining
hPSC-cardiomyocytes are evaluated by immunostaining using cardiac troponin T antibody. This protocol is for 24-well plates. 1. Aspirate the medium from a plate and wash the cells with 500 μl of D-PBS. 2. Add 300 μl of 4% PFA and incubate at room temperature for 15–20 min. 3. Discard 4% PFA solution and wash the cells with 500 μl of PBS twice. 4. Add 300 μl of 0.1% (v/v) Triton X-100 in D-PBS and incubate at room temperature for 15 min. 5. Aspirate the 0.1% (v/v) Triton X-100 in D-PBS solution, add 300 μl of blocking buffer and incubate at room temperature for 1 h. 6. Aspirate the blocking buffer, add 300 μl of cardiac troponin T monoclonal antibody solution diluted by blocking buffer and incubate overnight at 4 C. 7. Aspirate the solution and wash the cells with 500 μl of D-PBS twice. 8. Add 300 μl of goat anti-mouse IgG-Alexa Fluor 488 solution diluted by D-PBS and incubate at room temperature for 3 h in the dark. 9. Aspirate the solution and wash the cells with 500 μl of D-PBS twice.
A method for Isolation of Cardiomyocytes Using miRNA Switch
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transfection
miRNA-Bim switch Bim
Puromycinr mRNA Puror
hPSC-derived differentiated cells Puromycin selection
Death
Untransfected cells
Bim induction
Apoptosis
Non-cardiomyocytes Bim repression Puromycin resistance
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cTNT/Hoechst w/o selection miR-208a-Bim selection
Fig. 3 Isolation of hPSC-derived cardiomyocytes using miRNA-Bim switch without cell sorting. (a) Strategy of the non-sorting system using miRNA-Bim switch and puromycin resistant mRNA. Puromycin in the medium causes the death of untransfected cells, and the transfection of miRNA-Bim switch induces apoptosis in non-cardiomyocytes. On the other hand, transfected cardiomyocytes have resistance to puromycin and repress the translation of Bim. Consequently, only cardiomyocytes survive. (b) Immunostaining of cTNT without selection and with miR-208a-Bim switch-selected cells. Scale bars, 100 μm
10. Add 300 μl of Hoechst solution diluted by D-PBS and incubate at room temperature for 1 min. 11. Aspirate the solution and wash the cells with 500 μl of D-PBS twice. 12. Observe the cells using fluorescence microscopy (Figs. 2c and 3b).
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Notes 1. The ORF of BFP and BimEL are used in the cell sorting system and non-sorting system, respectively. The ORF of EGFP is used as a control in the cell sorting system, and the ORF of Puror is used for selecting transfected cells in the non-sorting system. 2. Too much pipetting will trigger cell death and yield poor differentiation efficiency. 3. Cell numbers are very important and need to be optimized according to the hPSC line. 4. Pipetting should be performed gently. 5. Confluency is very important for the transfection efficiency. Seeding cell numbers depend on the differentiation efficiency because non-myocytes will proliferate during the 2 days prior the transfection. 6. The total amount of mRNAs is important because high-dose mRNAs might trigger cell death. 7. After adding the mRNA complex into the well, immediately shake the plate. 8. The medium changes should be performed gently. 9. The sensitivity of Bim and puromycin depend on the cell lines. Their optimal concentrations must be confirmed.
Acknowledgments We greatly thank Yoko Uematsu and Kaoru Shimizu for their administrative support and Peter Karagiannis for proofreading the manuscript. This work was supported by a grant from Leducq foundation (18CVD05), JSPS KAKENHI Grants (18K15120 and 17H04176), grants from the Research Center Network for Realization of Regenerative Medicine (JP19bm0104001, JP19bm0204003, and JP19bm0804008), Research on Regulatory Science of Pharmaceuticals and Medical Devices (JP19mk0104117), and Research Project for Practical Applications of Regenerative Medicine (JP19bk0104095) provided by the Japan Agency for Medical Research and Development, and iPS research fund. 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. Yu J, Vodyanik MA, Smuga-Otto K, Antosiewicz-Bourget J, Frane JL, Tian S, Nie J, Jonsdottir GA, Ruotti V, Stewart R, Slukvin II, Thomson JA (2007) Induced
A method for Isolation of Cardiomyocytes Using miRNA Switch pluripotent stem cell lines derived from human somatic cells. Science 318(5858):1917–1920 3. 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 4. Kattman SJ, Witty AD, Gagliardi M, Dubois NC, Niapour M, Hotta A, Ellis J, Keller G (2011) Stage-specific optimization of activin/ nodal and BMP signaling promotes cardiac differentiation of mouse and human pluripotent stem cell lines. Cell Stem Cell 8 (2):228–240 5. 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 6. Lee JH, Protze SI, Laksman Z, Backx PH, Keller GM (2017) Human pluripotent stem cell-derived atrial and ventricular cardiomyocytes develop from distinct mesoderm populations. Cell Stem Cell 21(2):179–194.e174 7. Cyganek L, Tiburcy M, Sekeres K, Gerstenberg K, Bohnenberger H, Lenz C, Henze S, Stauske M, Salinas G, Zimmermann WH, Hasenfuss G, Guan K (2018) Deep phenotyping of human induced pluripotent stem cell-derived atrial and ventricular cardiomyocytes. JCI Insight 3(12):e99941 8. Hatani T, Miki K, Yoshida Y (2018) Induction of human induced pluripotent stem cells to cardiomyocytes using Embryoid bodies. Methods Mol Biol 1816:79–92 9. Anderson D, Self T, Mellor IR, Goh G, Hill SJ, Denning C (2007) Transgenic enrichment of cardiomyocytes from human embryonic stem cells. Mol Ther 15(11):2027–2036 10. Bizy A, Guerrero-Serna G, Hu B, PonceBalbuena D, Willis BC, Zarzoso M, Ramirez RJ, Sener MF, Mundada LV, Klos M, Devaney EJ, Vikstrom KL, Herron TJ, Jalife J (2013) Myosin light chain 2-based selection of human iPSC-derived early ventricular cardiac myocytes. Stem Cell Res 11(3):1335–1347
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11. Elliott DA, Braam SR, Koutsis K, Ng ES, Jenny R, Lagerqvist EL, Biben C, Hatzistavrou T, Hirst CE, Yu QC, Skelton RJ, Ward-van Oostwaard D, Lim SM, Khammy O, Li X, Hawes SM, Davis RP, Goulburn AL, Passier R, Prall OW, Haynes JM, Pouton CW, Kaye DM, Mummery CL, Elefanty AG, Stanley EG (2011) NKX2-5 (eGFP/w) hESCs for isolation of human cardiac progenitors and cardiomyocytes. Nat Methods 8(12):1037–1040 12. Chen Z, Xian W, Bellin M, Dorn T, Tian Q, Goedel A, Dreizehnter L, Schneider CM, Ward-van Oostwaard D, Ng JK, Hinkel R, Pane LS, Mummery CL, Lipp P, Moretti A, Laugwitz KL, Sinnecker D (2017) Subtypespecific promoter-driven action potential imaging for precise disease modelling and drug testing in hiPSC-derived cardiomyocytes. Eur Heart J 38(4):292–301 13. Dubois NC, Craft AM, Sharma P, Elliott DA, Stanley EG, Elefanty AG, Gramolini A, Keller G (2011) SIRPA is a specific cell-surface marker for isolating cardiomyocytes derived from human pluripotent stem cells. Nat Biotechnol 29(11):1011–1018 14. Uosaki H, Fukushima H, Takeuchi A, Matsuoka S, Nakatsuji N, Yamanaka S, Yamashita JK (2011) Efficient and scalable purification of cardiomyocytes from human embryonic and induced pluripotent stem cells by VCAM1 surface expression. PLoS One 6(8):e23657 15. Osborn L, Hession C, Tizard R, Vassallo C, Luhowskyj S, Chi-Rosso G, Lobb R (1989) Direct expression cloning of vascular cell adhesion molecule 1, a cytokine-induced endothelial protein that binds to lymphocytes. Cell 59 (6):1203–1211 16. Miki K, Endo K, Takahashi S, Funakoshi S, Takei I, Katayama S, Toyoda T, Kotaka M, Takaki T, Umeda M, Okubo C, Nishikawa M, Oishi A, Narita M, Miyashita I, Asano K, Hayashi K, Osafune K, Yamanaka S, Saito H, Yoshida Y (2015) Efficient detection and purification of cell populations using synthetic microRNA switches. Cell Stem Cell 16 (6):699–711
Chapter 6 Fabrication of Cardiac Constructs Using Bio-3D Printer Kenichi Arai, Daiki Murata, Shoko Takao, and Koichi Nakayama Abstract The fabrication of three-dimensional (3D) cardiac tissue using human induced pluripotent stem cellderived cardiomyocytes (iPSC-CMs) is useful not only for regenerative medicine, but also for drug discovery. Here, we report a bio-3D printer that can fabricate tubular cardiac constructs using only human iPSC-CMs. Protocols to evaluate the contractile force and response to electrical stimulation in the cardiac constructs are described. We confirmed that the constructs can be applied for transplantation or drug response testing. In the near future, we expect that the constructs will be used as alternatives for heart transplantation and in animal experiments for new drug development. Key words Bio-3D printer, Scaffold-free, Biofabrication, Cardiac construct, Human induced pluripotent stem cell-derived cardiomyocytes
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Introduction In 2010, approximately 16.7 million deaths worldwide were attributed to heart disease according to statistics from the World Health Organization (WHO), with heart failure as one of the leading causes of death [1]. Heart failure is the most common heart disease, and although heart transplantation is effective for treatment of end-stage intractable heart failure, many problems such as donor shortage and the need for patients to be placed under an immunosuppressive regimen in remain. In order to solve these problems, intramyocardial injection (cell transplantation method) of induced pluripotent stem cell-derived cardiomyocytes (iPSC-CMs) into mouse heart tissue has been attempted. It was reported that the cardiac contractility of a myocardial infarction animal model was improved by transplantation of iPSC-CMs [2]. However, the method is limited by the low survival and retention rate of transplanted cardiomyocytes [3]. Tissue engineering has been employed to artificially fabricate heart-like tissue using iPSC-CMs in order to overcome the aforementioned challenges of cell transplantation. Tissue engineering of
Yoshinori Yoshida (ed.), Pluripotent Stem-Cell Derived Cardiomyocytes, Methods in Molecular Biology, vol. 2320, https://doi.org/10.1007/978-1-0716-1484-6_6, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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Fig. 1 Schematic illustration of bio-3D printing technology. Spheroids, needle array, and 3D design are first prepared. The spheroids are loaded onto the needle array by the bio-3D printer
3D constructs using cardiomyocytes may be scaffold-based or scaffold-free (e.g., cell sheet engineering and the Kenzan method). Scaffold-based tissue engineering requires an extracellular matrix hydrogel or artificial polymer to support cell attachment and subsequently form cardiac tissue [4–6]. On the other hand, cell sheet engineering, a scaffold-free method, allows fabrication of cell sheets by culturing a monolayer of cells in a thermo-responsive polymercoated dish. Multi-layered cardiac constructs are fabricated by stacking prepared cell sheets [7, 8]. In addition, we developed the Kenzan method in which bio-3D printing technology automatically loads spheroids onto a needle array (Kenzan) according to a desired 3D design (Fig. 1). Because cardiac constructs are fabricated using a needle array instead of a scaffold, the fusion of spheroids in a construct on the needle array can be observed by culturing for several days. The constructs can be removed from the needle array and matured in a bioreactor; thus, the fabricated construct does not require a scaffold. This bio-3D printer has been used to fabricate 3D constructs, including those of heart, liver, blood vessels, and trachea [9–14]. In this chapter, we report the fabrication protocol for cardiac constructs using a bio-3D printer (spheroid formation, construct fabrication, maturation, and evaluation), which was improved from a previously reported protocol [9, 10].
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Materials
2.1 Cardiac Spheroid Formation
1. iPSC-CMs (Cellular Dynamics International, CDI, Madison, WI, USA). 2. Human umbilical vein endothelial cells (HUVECs) (Lonza, Inc. Walkersville, MD, USA). 3. Normal human dermal fibroblasts (NHDFs) (Lonza, Inc.) 4. 0.05% Trypsin/EDTA solution. 5. Dulbecco’s phosphate-buffered saline without Ca or Mg (DPBS()). 6. iCell plating medium (CDI). 7. iCell maintenance medium (CDI). 8. EGM™-2 medium: EBM™-2 basal medium with CC-4176 Single Quots™ Kit. 9. FGM-2 medium: FBM™-2 basal medium with CC4126 Single Quots™ Kit. 10. Gelatin solution: prepare a stock solution (1% w/v) dissolved in dH2O and store at 4 C. 11. 150-mm cell culture dish (or flask). 12. Ultra-low attachment 96U-well plates (see Note 1). 13. Reagent Reservoirs. 14. 50 ml centrifuge tube. 15. Multichannel pipette.
2.2
Bio-3D Printer
1. Bio-3D printer (Cyfuse Biomedical K.K, Tokyo, Japan) (Fig. 2a) (see Note 2). 2. 9 9 tubular needle array (Cyfuse Biomedical K.K.) 3. 26 G nozzle (Cyfuse Biomedical K.K.) 4. Head holder (Cyfuse Biomedical K.K.) 5. Cleaning container (Cyfuse Biomedical K.K.) 6. Kenzan holder (Cyfuse Biomedical K.K.)
2.3 Fabrication of Cardiac Constructs Using Bio-3D Printer 2.4 Contraction Analysis and Electrical Stimulation
1. Polydimethylsiloxane (PDMS) tube (see Note 3). 2. Bioreactor (Fig. 3c). 3. Peristaltic pump. 1. Stereomicroscope. 2. Digital camera. 3. Contraction analysis software (developed in-house). 4. Platinum rods (diameter: 1 mm, length: 50 mm).
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Fig. 2 Photographic and schematic views of bio-3D printer (Regenova). (a) Diagram of bio-3D printer. (b) Nozzle and head unit. The nozzle-connected head aspirates a spheroid from the 96-well plate and prints a spheroid onto the needle array. This unit is equipped with a camera at the top and bottom. The top camera recognizes the tip of the needle array, whereas the bottom camera recognizes the tip of the nozzle and measures the spheroid size and roundness. (c) Kenzan holder and 9 9 needle array. (d) Plate storage and transport unit. The 96-well plate is set up in the magazine rack and transported to the printing area
5. Electrical stimulation device (see Note 4). 6. Electric power supply.
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3.1 Cardiac Spheroid Formation 3.1.1 Maintenance of Endothelial Cells and Fibroblasts
1. Add 20 ml of growth medium to a sterile 50 ml centrifuge tube in advance. 2. Thaw the cryovial containing the cryopreserved cells (HUVECs or NHDFs) in a 37 C water bath. 3. Transfer the cell suspension in the cryovial to a sterile 50 ml centrifuge tube using a 1-ml pipettor and stir with growth medium. 4. Centrifuge HUVECs at 190 g for 3 min, and NHDFs at 220 g for 5 min. 5. After centrifugation, aspirate the supernatant fluid and resuspend with fresh growth medium.
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Fig. 3 Schematic procedure and time course for the fabrication of cardiac constructs using bio-3D printer. (a) Time-lapse imaging and time course of cardiac spheroid formation. (b) Culture of cardiac constructs. The spheroids in the cardiac construct shown fused at day 12 after printing. (c) Illustration (left) and photograph (right) of bioreactor for cardiac constructs
6. Count cells and seed each cell in a dish or flask (recommended to seed HUVECs at 2500 cells/cm2 and NHDF at 3500 cells/ cm2 in a 15 cm dish). 7. Culture each cell type until 70–80% confluence. 8. Aspirate the growth medium from the 15 cm dish (or flask) and rinse immediately with DPBS(). 9. Add 0.08 ml/cm2 of 0.05% trypsin/EDTA solution, swirl to cover the entire surface of the 15 cm dish (or flask), and incubate at 37 C for 5 min.
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10. Add 0.2 ml/cm2 of the growth medium to inactivate trypsin/ EDTA. 11. Transfer the cell suspension to a sterile 50 ml centrifuge tube. 12. Centrifuge HUVECs at 190 g for 3 min, and NHDFs at 220 g for 5 min. 13. Remove the supernatant fluid and resuspend with fresh growth medium. 14. Count cells and adjust the cell concentration according to the subsequent application. 3.1.2 Thawing of Human iPSC-CMs
1. Retrieve the cryovial containing cryopreserved human iPSCCMs from the liquid nitrogen storage tank. 2. Immerse the cryovial containing human iPSC-CMs in a 37 C water bath for 3 min (see Note 5). 3. Gently transfer the thawed human iPSC-CM suspension to a sterile 50 ml centrifuge tube using a 1-ml pipettor. 4. Rinse the empty cryovial with 1 ml of iCell plating medium to retrieve the remaining cells from the cryovial. 5. Transfer and mix the 1 ml suspension of remaining cells dropwise to the 50 ml centrifuge tube containing the cell suspension over 90 s (see Note 6). 6. Slowly add and mix 3 ml of iCell plating medium dropwise to the 50 ml centrifuge tube over 180 s. 7. Gently mix the contents of the 50-ml centrifuge tube using a 25-ml pipettor (see Note 7). 8. Count the cells and adjust the concentration according to the subsequent application.
3.1.3 Cardiac Spheroid Formation
1. Dilute the human iPSC-CM suspension with iCell plating medium to a density of 4.5 105 cells/ml. 2. Dilute the HUVEC suspension with EGM™-2 medium to a density of 2.25 105 cells/ml. 3. Dilute the NHDF suspension with FGM-2 medium to a density of 2.25 105 cells/ml. 4. Mix equal amounts of these three cell suspensions (see Note 8). 5. Seed 0.1 ml cell suspension containing the three cell-types into ultra-low attachment 96U-well plates using a multichannel pipette (see Note 9). 6. After 2 days, exchange half the medium twice with fresh medium composed of equal amounts of iCell maintenance medium, EGM™-2 medium, and FGM-2 medium. 7. Every 2 days, exchange half the medium with the medium described in step 6 (see Note 10).
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3.2 Cardiac Construct Fabrication 3.2.1 Preparation of the Bio-3D Printer
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1. Autoclave the parts (9 9 tubular needle array, Kenzan holder, cleaning container, 26 G nozzle, and head) of the bio-3D printer in advance (Fig. 2a). 2. Attach the 26 G nozzle-connected head to the mobile nozzle arm in the bio-3D printer (Fig. 2b). 3. Set up the cleaning container in the bio-3D printer. 4. Set up the 9 9 tubular needle array in the Kenzan holder filled with DPBS() solution in the bio-3D printer (Fig. 2c).
3.2.2 Fabrication of Cardiac Constructs by Bio-3D Printing
1. Set up the 96-well plate in the storage magazine (Fig. 2d). 2. Measure the size and roundness of the cardiac spheroids using the bio-3D printer software (Fig. 3a) (see Note 11). 3. Set the pitch in the Z-direction as 0.8 times the spheroid size (see Note 12). 4. Create a 3D design for the cardiac construct (see Note 13). 5. Ensure that the camera on top of the mobile nozzle arm properly recognizes the tip of the needle array. 6. Begin cardiac construct printing. The camera on the bottom of the mobile nozzle arm is programmed to initiate the fabrication of the constructs after the tip of the 26 G nozzle is properly recognized. 7. The 96-well plate is automatically retrieved from the storage magazine and transported to the printing area. 8. The spheroids in the 96-well plate are aspirated by the tip of the nozzle connected to the head holder (Fig. 2b). 9. The aspirated spheroids are loaded on the needle array in the Kenzan holder according to the desired 3D design. 10. By repeating this procedure, the cellular construct can be automatically fabricated according to the 3D design.
3.2.3 Maturation of Cardiac Constructs Using a Bioreactor
1. Add medium to the bioreactor. 2. Set up the printed cardiac spheroids on the needle array in the bioreactor (Fig. 3b, c). 3. Connect the bioreactor to the peristaltic pump (Fig. 3c). 4. Adjust the flow rate of the peristaltic pump to 7.3 ml/min. 5. Culture the printed cardiac spheroids on the needle array in the bioreactor for 7 days (see Note 14). 6. After confirming spheroid fusion on the needle array, remove the cardiac construct from the needle array and transfer to the PDMS tube (Fig. 3b). 7. Culture the cardiac construct in the bioreactor for 7 days (Fig. 3).
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Fig. 4 Motion analysis of the contractile area in cardiac constructs. (a) A movie of the contraction of the cardiac construct is recorded. (b) An individual frame of the movie is selected and used to measure the cardiac construct area by analysis software. (c) The fractional area change in the spheroids and constructs is calculated and graphed 3.3 Evaluation of Cardiac Constructs 3.3.1 Contraction Analysis
1. Record a movie of the cardiac construct using a digital camera to analyze contraction of the construct (Fig. 4a) (see Note 15). 2. Import the recorded movie to an in-house analysis software. 3. Analyze the beating rate and fractional area change of the construct using an in-house analysis software (Fig. 4b). 4. Calculate the fractional area change as the contracted area/ minimum area (Fig. 4c).
3.3.2 Electrical Stimulation
1. Connect the two platinum electrodes and electrical stimulation device and place the cardiac construct between the two platinum electrodes (Fig. 5a). 2. Connect the electrical stimulation device to the electric power supply. 3. Set the voltage (20 V), frequency (2 Hz), and pulse (10 ms) for electrical stimulation (Fig. 5b). 4. Apply electrical stimulation to the cardiac construct. 5. Record the contraction of the cardiac construct using a microscope and digital camera. 6. Analyze the beat rate and fractional area change of the construct using an in-house analysis software (Fig. 5d).
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Fig. 5 Electrical stimulation of cardiac constructs. (a) Basic diagram of the electrical stimulation system. (b) Electrical stimulation conditions (voltage and frequency). (c) Mesh-type tubular cardiac constructs. (d) Changes in beat rate in response to electrical stimulation of the tubular cardiac constructs
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Notes 1. The PrimeSurface 96U Plate (MS-9096U; Sumitomo Bakelite) is recommended for spheroid generation. 2. Figure 2a shows a schema of the bio-3D printer, Regenova. The machine was designed for use on a clean bench. 3. The PDMS tube should have an external diameter of 2 mm and internal diameter of 1.73 mm. 4. The electrical stimulation device shown here was developed in-house. Electrical stimulators can be purchased commercially [15] (e.g., Grass Astro-Med stimulators model S88X Dual Output Square Pulse Stimulator). 5. A floating microtube rack is recommended for thawing the cryovial containing human iPSC-CMs. 6. Gently swirl the tube while adding the medium to mix completely and minimize osmotic shock of the thawed cells. 7. Avoid repeated strong pipetting of the cell suspensions.
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8. The cell suspension should be adjusted so that the cell ratio of human iPSC-CMs, HUVECs, and NHDFs is 2:1:1. 9. The cell suspension in the reservoir should be mixed regularly by pipetting. 10. For fabrication of cardiac constructs, 5-day-old cardiac spheroids are recommended. 11. The optimal construct size should be 500–600 μm and roundness >60%. Roundness (%) ¼ 100—(R – r)/R 100. “R” is the radius of the minimum circumscribed circle, and “r” is the radius of the inscribed circle. 12. The pitch in the Z-direction should be set as 0.8 times the spheroid size to allow fusion with other spheroids. If the cardiac spheroids do not fuse, the researcher should optimize the pitch in the Z-direction. 13. The tubular cardiac constructs should be fabricated to allow delivery of sufficient medium to the inner part of the construct. At an early stage of development, the primitive heart tube first forms by cardiac looping and is then organized as a tubular assembly [16]. 14. After 2 days of culture, exchange half the medium with fresh medium containing equal amounts of iCell maintenance medium, EGM™-2 medium, and FGM-2 medium. 15. The background of the movie should be dark to distinguish the cardiac construct and background.
Acknowledgments This work was supported by grants from JSPS KAKENHI Grant Number 16K19968, 16K15633 and 18K08763 and the Nakatani Foundation. And this work has received funding by FUJIFILM Corporation in Japan. Conflicts of Interest: Nakayama is a co-founder and shareholder of Cyfuse Biomedical KK and an inventor/developer designated on patents for the bio-3D printer (Patent title: Method for production of three-dimensional structure of cell; patent number: JP4517125; Patent title: Cell structure production device; patent number; JP5896104). All other authors have declared that no competing interests exist.
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References 1. Bansilal S, Castellano JM, Fuster V (2015) Global burden of CVD: focus on secondary prevention of cardiovascular disease. Int J Cardiol 201(Suppl 1):S1–S7 2. van Laake LW, Passier R, Monshouwer-Kloots J et al (2007) Human embryonic stem cellderived cardiomyocytes survive and mature in the mouse heart and transiently improve function after myocardial infarction. Stem Cell Res 1(1):9–24 3. Park M, Yoon YS (2018) Cardiac regeneration with human pluripotent stem cell-derived cardiomyocytes. Korean Circ J 48(11):974–988 4. Wang F, Guan J (2010) Cellular cardiomyoplasty and cardiac tissue engineering for myocardial therapy. Adv Drug Deliv Rev 62 (7–8):784–797 5. Godier-Furne´mont AF, Tiburcy M, Wagner E et al (2015) Physiologic force-frequency response in engineered heart muscle by electromechanical stimulation. Biomaterials 60:82–91 6. Tandon N, Taubman A, Cimetta E et al (2013) Portable bioreactor for perfusion and electrical stimulation of engineered cardiac tissue. Conf Proc IEEE Eng Med Biol Soc 2013:6219–6223 7. Sakaguchi K, Shimizu T, Okano T (2015) Construction of three-dimensional vascularized cardiac tissue with cell sheet engineering. J Control Release 205:83–88 8. Seta H, Matsuura K, Sekine H et al (2017) Tubular cardiac tissues derived from human induced pluripotent stem cells generate pulse pressure in vivo. Sci Rep 7:45499
9. Arai K, Murata D, Verissimo AR et al (2018) Fabrication of scaffold-free tubular cardiac constructs using a bio-3D printer. PLoS One 13(12):e0209162 10. Arai K, Murata D, Takao S et al (2020) Drug response analysis for scaffold-free cardiac constructs fabricated using bio-3D printer. Sci Rep 10:8972 11. Yanagi Y, Nakayama K, Taguchi T et al (2017) In vivo and ex vivo methods of growing a liver bud through tissue connection. Sci Rep 7:14085 12. Itoh M, Nakayama K, Noguchi R et al (2015) Scaffold-free tubular tissues created by a bio-3D printer undergo remodeling and endothelialization when implanted in rat aortae. PLoS One 10(9):e0136681 13. Taniguchi D, Matsumoto K, Tsuchiya T et al (2018) Scaffold-free trachea regeneration by tissue engineering with bio-3D printing. Interact Cardiovasc Thorac Surg 26(5):745–752 14. Zhang XY, Yanagi Y, Sheng Z et al (2018) Regeneration of diaphragm with bio-3D cellular patch. Biomaterials 167:1–14 15. Tandon N, Cannizzaro C, Chao PH et al (2009) Electrical stimulation systems for cardiac tissue engineering. Nat Protoc 4 (2):155–173 16. Sedmera D (2011) Function and form in the developing cardiovascular system. Cardiovasc Res 91(2):252–259
Chapter 7 Fabrication of Thick and Anisotropic Cardiac Tissue on Nanofibrous Substrate for Repairing Infarcted Myocardium Junjun Li, Li Liu, Itsunari Minami, Shigeru Miyagawa, and Yoshiki Sawa Abstract In this chapter, we introduce the method for fabricating thick and anisotropic cardiac tissue for heart regeneration. Aligned and biodegradable nanofiber can be prepared by electrospinning Food and Drug Administration-approved poly (lactic-co-glycolic acid) on a rotating drum. After the nanofibers are transferred on to a polydimethylsiloxane frame, the cardiomyocytes could be plated on the nanofiber to form thick and anisotropic cardiac tissue rapidly. Cardiac tissue-like construct could be easily created by one-step method, and transplanted onto the hearts of myocardium infarction models and lead to their functional recovery. Key words Cardiac tissue engineering, Pluripotent stem cells, Cardiomyocytes, Electrospun nanofiber, Three-dimensional, Extracellular matrix, Transplantation
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Introduction Although human pluripotent stem cells (hPSCs) have been differentiated into cardiomyocytes (CMs) in high purity [1–3], however, hPSC-CMs resemble more immature CMs rather than adult CMs in both morphologies and functional characteristics [4–6]. In addition, the state-of-art myocardial regeneration technologies such as cell injection [7, 8], cell sheet [9, 10], or patch [11] are still using randomly organized hPSC-CMs, that do not reproduce those in in vivo conditions [12, 13]. We electrospun poly(lactic-co-glycolic acid) (PLGA), a biodegradable polymer approved by Food and Drug Administration (FDA), into aligned nanofibers [14] with the thickness 10–40fold lower than previous reports [15–18]. Despite the low thickness, the nanofibers demonstrated excellent operability as they were fixed to a silicone frame, which also allowed floating cultures in the medium. Thick and anisotropic cardiac tissue, termed cardiac
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Immature cardiomyocytes
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Drug assessment Side view
Differentiation Transplantation
Electrospinning
Aligned nanofibers (ANFs)
Anisotropy + 3D
Fig. 1 Cardiac tissue-like constructs formed on aligned nanofibers for drug assessment and repairing infarcted myocardium. (Figures reproduced with permission [14])
tissue-like construct (CTLC), was then created by one-step plating of CMs derived from hPSCs [2] on the aligned nanofibrous scaffold (Fig. 1). During the following culture, the CMs infiltrated and enveloped the nanofibers, demonstrated anisotropic organization as well as the upregulated expression of cardiac biomarkers. In addition, in vivo transplantation of CTLCs in myocardial infarcted rat heart showed excellent survival of CTLC as well as improved cardiac function 4 weeks post transplantation. In this chapter, we introduce the methods to fabricate thick and anisotropic cardiac tissue and transplant these tissues onto the animal heart for repairing the infarcted heart.
2
Materials
2.1 Nanofiber Spinning Materials
1. High voltage supply (Tech Dempaz, Japan). 2. Rotating drum (Tongliweina Nanotechnology Co., Ltd., China). 3. Syringes: 1 mL. 4. Syringe needles: 23G 1 ¼00 . 5. Fume hood. 6. Electrospinning solution: 20% (w/v) PLGA in tetrahydrofuran (THF). In order to reduce the beads within the produced nanofiber, ionic surfactant sodium dodecyl sulfate (SDS) in de-ionized water was added to a final concentration of 0.92 g/L. 7. Tube rotator. 8. Vortex mixer. 9. Polydimethylsiloxane (PDMS): 1:10 ratio for cross-linker to base. 10. Bistoury and cutting mat for making PDMS frame. 11. Thermal press machine (AS ONE, Japan).
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2.2 Cell Differentiation and Culture
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1. ESC Culture Medium (ReproCELL, Japan). 2. Protease solution: 0.1% collagenase type I, 0.25% trypsin, 1 U/ mL DNase I, 116 mM NaCl, 20 mM HEPES, 12.5 mM NaH2PO4, 5.6 mM glucose, 5.4 mM KCl, and 0.8 mM MgSO4 (pH 7.35). 3. Cardiac differentiation medium: IMDM containing 1% MEM nonessential amino acid solution, 1% penicillin-streptomycin, 2 mM L-glutamine, 0.5 mM L-carnitine, 0.001% 2-mercaptoethanol, 0.4% HSA. 4. Cardiac differentiation chemicals: KY02111 (Wako, Japan), XAV939.
CHIR99021,
BIO,
5. Ultralow attachment culture dishes. 6. 40-μm cell strainer. 7. Multielectrode array (MEA), and MEA data acquisition system (USB-ME64-System, Multi Channel Systems, Germany). 2.3
Transplantation
1. Cellnest recombinant peptide (Fujifilm, Japan). 2. Beriplast P (CSL Behring, USA). 3. Autoclaved surgical tools: scalpel, scissors, forceps. 4. Ultrasound system (SONOS 5500, Agilent Technologies). 5. Suture line. 6. Culture bag (CultiLife™, Takara, Japan). 7. 8-week-old male nude rats (CLEA Japan Inc., Japan).
3
Methods
3.1 Electrospinning of Nanofiber
1. All components are mixed in a fume hood, then the electrospinning solution was vortexed for 30 min and rotated overnight (see Note 1). 2. The solution was loaded in a 1-mL syringe with a 23G 1 ¼00 needle. The PLGA nanofibers were electrospun at a voltage of 10 kV provided by a DC high-voltage generator. The positive electrode of the generator was connected to the needle. A grounded rotating drum was set to rotate at a speed of 11.4 m/s to generate aligned nanofibers (ANFs). A layer of aluminum foil was used to cover the drum for collecting the nanofibers. The distance between the needle tip and the drum is 8 cm (see Note 2). 3. The nanofiber sheets with the different thickness could be obtained by varying spin time (Fig. 2a): high-density ANFs (H-ANFs, spinning time: 10 min, thickness: 11.3 1.2 μm) and low-density ANFs (L-ANFs, spinning time: thickness: 1.5 0.1 μm) (see Notes 3 and 4).
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Fig. 2 Morphology of ANFs. (a) Scanning electron microscopy (SEM) image of low-density and high-density aligned nanofibers (L-ANFs and H-ANFs). (b) H-ANFs transferred to a PDMS frame. (c) Cross-section of PDMS frame with H-ANFs after being cut in the middle. (Figures reproduced with permission [14])
4. PDMS was poured into a rectangular dish to get 2-mm PDMS sheet. Place the dish into an oven at 70 C for 2 h. The sheet was cut into PDMS frame. 5. After spinning, the nanofibers were pressed onto the substrate by a thermal press machine or transferred to a frame made of PDMS (Fig. 2b, c) (see Note 5). 3.2 Cardiomyocyte Differentiation
The direct cardiomyocyte differentiation was carried out as previously reported [2]: 1. Confluent hiPSCs were enzymatically detached and transferred into Petri dishes with ESC Culture Medium. 2. To form an aggregate with 0.3–1 mm diameter, cells were held in suspension culture for 8–24 h. 3. Cell aggregates were transferred to ultralow attachment culture dishes (3–6 105 cells/cm2) in cardiac differentiation medium. 4. The medium was changed to cardiac differentiation medium with 2 μM CHIR99021 and 0.5 μM BIO until day 3.
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5. On day 3–9, 10 μM KY02111 and 2 μM XAV939 were added to the cell cultures, and the medium was changed every 2 days. 6. After day 9, the floating cardiac colonies were maintained until the protease treatment and the media were changed every 5 days. 3.3 Cardiomyocyte Culture
1. Floating colonies of day 28–35 CMs were collected and dispersed in 10 mL protease solution. The mixture was stirred for up to 1 h. 2. 40 mL 10% serum-supplemented cardiac differentiation medium was added to stop the reaction. 3. The cells were then filtered by using a 40 μm cell strainer, counted and spun at 300 g for 5 min. 4. Aspirate supernatant, resuspend the cells in serumsupplemented cardiac differentiation medium. The volume of medium should be decided by cell seeding area (e.g., 200 μL medium for 1 cm2 area). 5. The cell suspension was then added on an ANF sheet in a 6-well plate. CTLCs with different thicknesses could be obtained by varying the density of seeded CMs (Fig. 3a–c) (see Note 6). 6. The cells suspension will be kept on the nanofiber sheet for more than 3 h to allow the attachment of CMs. 3 mL medium is then added gently to the dish (see Note 7).
a
a-actinin / DAPI
b
c Thickness (μm)
4㽢106 cells cm-2
50 µm
⃰⃰⃰ 160 120 ⃰
80 40
50 µm 0
d
0.3 1 4 (㽢106 cells cm-2) L-ANFs + CMs
MEA system
GND #74
Fig. 3 Characterization of CTLC. (a) Immunostaining images of α-actinin. Cardiomyocytes were cultured on ANFs for 14 days. (b) Histology of CTLCs. (c) The thickness of CTLCs with different cell density. (d) Images of the MEA system and MEA chip with CTLC. The enlarged images indicate the homogeneous electrical signals recorded by electrodes. (Figures reproduced with permission [14])
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a
Culture
On-line function assessment
CTLC
CTLC
Petri-dish
MEA
Transportation
b
CTLC
CTLC
MEA Gasket
Culture bag
Rat heart
Culture bag
Fig. 4 Transplantation of the CTLC to a rat heart. (a) After cell seeding, the CTLC can be functionally evaluated before transportation. A gasket could be used to fix the CTLC on the MEA for signal recording. (b) Transplantation of CTLC on a rat heart. (Figures reproduced with permission [14])
7. The medium was changed to serum-free medium (cardiac differentiation medium without FBS) since day 2 and to fresh medium every 4 days. 8. During the culture, the electrical signal could be recorded by placing the CTLC in an MEA data acquisition system. (Fig. 3d, see Note 8). 3.4
Transplantation
1. 8-week-old male transplantation.
nude
rats
were
used
for
CTLC
2. Thoracotomy was performed between the fourth and fifth intercostal spaces, and myocardial infarction (MI) was created by ligation of the left coronary artery in rat hearts, as described previously [19]. 3. hiPSC-CMs (5–8 106 cells) were seeded on the H-ANFs with a 1 1 cm2 PDMS frame for 1 day. The CTLC can be also optionally recorded with a MEA (see Note 8) to check the electrophysiology or sealed into a culture bag for transportation (Fig. 4a). 4. Before transplantation, CTLC were immersed in recombinant peptide solution with a concentration of 5 mg cm2 at room temperature for 5 min. 5. The PDMS frame was cut and the CTLCs were placed on the epicardium (Fig. 4b) and covered with Beriplast P. (see Notes 9–11). 6. 4 week after transplantation, echocardiography was performed by using an ultrasound machine with a 12 MHz transducer [19]. Ejection fraction (EF) was calculated based on the formula: EF (%) ¼ (LVEDD3 LVESD3)/LVEDD3 100 (%).
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7. Rats with MI were sacrificed 4 weeks post transplantation. The hearts were harvested and cryosliced into 7-μm-thick sections for following histology and immunostaining.
4
Notes 1. When dissolving PLGA in the THF solvent, the tube should be vortexed until the majority of PLGA is dissolved. A large chunk of PLGA remained in the bottom of the tube could keep undissolved even after overnight rotating. 2. During the nanofiber fabrication, the humidity and the temperature could affect the nanofiber morphology. The ideal condition should be 25 C and 30%. 3. If there are beads within the nanofibers, the SDS could be slightly increased. 4. During the spinning the needle of the syringe should be cleaned from time to time. Otherwise, the accumulated debris on the syringe tip could significantly affect the size and the morphology of nanofibers. 5. The Nanofiber could be fixed on the frame of PDMS by using the liquid PDMS. The quantity of liquid PDMS should be limited; otherwise, the PDMS could be leaked into the inner area (within the PDMS frame) on the nanofiber sheet. In this case, the cells could attach on the PDMS layer rather than on the PLGA fiber sheet. 6. Before cell seeding, the PLGA nanofiber sheet could be sterilized by ultraviolet light. The treating time must be limited to 30 min as long-time UV treatment (e.g., over 1 h) could damage the PLGA nanofiber, which would easily break after coculture with cells for several days. 7. After medium addition, the nanofiber sheet is recommended to be immersed in the medium. The immersion will allow the CMs to receive the nutrient from both sides and improve the CMs viability. 8. During the MEA recording, the distance between CTLC and the electrode should be kept as short as possible to enable the recording of nice field potential from the CMs. Reducing the medium volume (e.g., to 0.5 mL) could help reduce the distance between CTLC and electrode. Moreover, one gasket (or Harp slice grid, Multichannel System, Germany) could be used to press the CTLC on the MEA as shown in the Fig. 4a. Heavier gasket (or Harp slice grid) could be used for better recording. After recording, the gasket should be removed to avoid permanent damage to the CMs underneath.
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9. During the CTLC transplantation, the PDMS frame should be cut after the fiber sheet attached to the surface of the heart. The CMs side of CTLC should be placed close to the heart. 10. During the operation, if the CTLC twist and crimp, simply reimmerse it gently into the medium and the sheet will quickly unfold. 11. After transplantation, the epicardium should be sutured to cover and protect the CTLC.
Acknowledgment This work was supported by the Japan Society for the Promotion of Science (JSPS): Grants-in-Aid for Scientific Research (B) (15H03948, 26289065, 23310087, 26286027), Grants-inAid for Scientific Research (C) (15K08270), and Grants-in-Aid for JSPS Fellows (26.04046). The research was (partially) supported by Japan Agency for Medical Research and development (AMED) program. References 1. Burridge PW, Matsa E, Shukla P, Lin ZC, Churko JM, Ebert AD, Lan F, Diecke S, Huber B, Mordwinkin NM (2014) Chemically defined generation of human cardiomyocytes. Nat Methods 11(8):855–860 2. Minami I, Yamada K, Otsuji TG, Yamamoto T, Shen Y, Otsuka S, Kadota S, Morone N, Barve M, Asai Y (2012) A small molecule that promotes cardiac differentiation of human pluripotent stem cells under defined, cytokine-and xeno-free conditions. Cell Rep 2 (5):1448–1460 3. Lian X, Zhang J, Azarin SM, Zhu K, Hazeltine LB, Bao X, Hsiao C, Kamp TJ, Palecek SP (2013) Directed cardiomyocyte differentiation from human pluripotent stem cells by modulating Wnt/β-catenin signaling under fully defined conditions. Nat Protoc 8(1):162–175 4. Matsa E, Burridge PW, Wu JC (2014) Human stem cells for modeling heart disease and for drug discovery. Sci Transl Med 6 (239):239ps236 5. Snir M, Kehat I, Gepstein A, Coleman R, Itskovitz-Eldor J, Livne E, Gepstein L (2003) Assessment of the ultrastructural and proliferative properties of human embryonic stem cellderived cardiomyocytes. Am J Physiol Heart Circ Physiol 285(6):H2355–H2363 6. Robertson C, Tran DD, George SC (2013) Concise review: maturation phases of human
pluripotent stem cell-derived cardiomyocytes. Stem Cells 31(5):829–837 7. Shiba Y, Fernandes S, Zhu W-Z, Filice D, Muskheli V, Kim J, Palpant NJ, Gantz J, Moyes KW, Reinecke H (2012) Human EScell-derived cardiomyocytes electrically couple and suppress arrhythmias in injured hearts. Nature 489(7415):322–325 8. Shiba Y, Gomibuchi T, Seto T, Wada Y, Ichimura H, Tanaka Y, Ogasawara T, Okada K, Shiba N, Sakamoto K, Ido D, Shiina T, Ohkura M, Nakai J, Uno N, Kazuki Y, Oshimura M, Minami I, Ikeda U (2016) Allogeneic transplantation of iPS cellderived cardiomyocytes regenerates primate hearts. Nature 538(7625):388–391 9. Kawamura M, Miyagawa S, Fukushima S, Saito A, Miki K, Ito E, Sougawa N, Kawamura T, Daimon T, Shimizu T (2013) Enhanced survival of transplanted human induced pluripotent stem cell–derived cardiomyocytes by the combination of cell sheets with the pedicled omental flap technique in a porcine heart. Circulation 128(11 suppl 1): S87–S94 10. Masumoto H, Matsuo T, Yamamizu K, Uosaki H, Narazaki G, Katayama S, Marui A, Shimizu T, Ikeda T, Okano T (2012) Pluripotent stem cell-engineered cell sheets reassembled with defined cardiovascular
Cardiac Tissue-Like Construct for Healing Myocardium populations ameliorate reduction in infarct heart function through cardiomyocytemediated neovascularization. Stem Cells 30 (6):1196–1205 11. Menasche´ P, Vanneaux V, Fabreguettes J-R, Bel A, Tosca L, Garcia S, Bellamy V, Farouz Y, Pouly J, Damour O, Pe´rier M-C, Desnos M, Hage`ge A, Agbulut O, Bruneval P, Tachdjian G, Trouvin J-H, Larghero J (2015) Towards a clinical use of human embryonic stem cell-derived cardiac progenitors: a translational experience. Eur Heart J Cardiovasc Imaging 36(12):743–750 12. Mathur A, Ma Z, Loskill P, Jeeawoody S, Healy KE (2016) In vitro cardiac tissue models: current status and future prospects. Adv Drug Deliv Rev 96:203–213 13. Shao Y, Sang J, Fu J (2015) On human pluripotent stem cell control: the rise of 3D bioengineering and mechanobiology. Biomaterials 52:26–43 14. Li J, Minami I, Shiozaki M, Yu L, Yajima S, Miyagawa S, Shiba Y, Morone N, Fukushima S, Yoshioka M, Li S, Qiao J, Li X, Wang L, Kotera H, Nakatsuji N, Sawa Y, Chen Y, Liu L (2017) Human pluripotent stem cell-derived cardiac tissue-like constructs for repairing the infarcted myocardium. Stem Cell Rep 9 (5):1546–1559 15. Joanne P, Kitsara M, Boitard S-E, Naemetalla H, Vanneaux V, Pernot M,
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Larghero J, Forest P, Chen Y, Menasche´ P, Agbulut O (2016) Nanofibrous clinical-grade collagen scaffolds seeded with human cardiomyocytes induces cardiac remodeling in dilated cardiomyopathy. Biomaterials 80:157–168 16. Kharaziha M, Shin SR, Nikkhah M, Topkaya SN, Masoumi N, Annabi N, Dokmeci MR, Khademhosseini A (2014) Tough and flexible CNT–polymeric hybrid scaffolds for engineering cardiac constructs. Biomaterials 35 (26):7346–7354 17. Masoumi N, Larson BL, Annabi N, Kharaziha M, Zamanian B, Shapero KS, Cubberley AT, Camci Unal G, Manning K, Mayer JE (2014) Electrospun PGS: PCL microfibers align human valvular interstitial cells and provide tunable scaffold anisotropy. Adv Healthc Mater 3(6):929–939 18. Han J, Wu Q, Xia Y, Wagner MB, Xu C (2016) Cell alignment induced by anisotropic electrospun fibrous scaffolds alone has limited effect on cardiomyocyte maturation. Stem Cell Res 16(3):740–750 19. Memon IA, Sawa Y, Fukushima N, Matsumiya G, Miyagawa S, Taketani S, Sakakida SK, Kondoh H, Aleshin AN, Shimizu T (2005) Repair of impaired myocardium by means of implantation of engineered autologous myoblast sheets. J Thorac Cardiovasc Surg 130(5):1333–1341
Chapter 8 Construction of Three-Dimensional Cardiac Tissues Using Layer-by-Layer Method Maki Takeda, Shigeru Miyagawa, Mitsuru Akashi, and Yoshiki Sawa Abstract Myocardial tissues in vivo are complex three-dimensional structures. Significant efforts are currently focused on developing functionally and structurally similar tissues in vitro to transplant them for regenerative therapy and to evaluate pharmacological agents. We describe a method for constructing threedimensional multilayered cardiac tissues by coating cells with extracellular matrix components (ECM). Key words Cardiac tissue, Three-dimensional, Layer by layer, Regenerative therapy, Drug discovery, Stem cells
1
Introduction Stem cells are expected to be used in regenerative therapy and drug discovery in the field of cardiology. Although the number of patients with heart failure is increasing year by year, many patients are waiting for transplantation owing to chronic shortage of donors. The development of myocardial regeneration therapy as a new treatment for severe heart failure, is being promoted [1]. Owing to some serious adverse effects, including cardiotoxicity, the development of new drugs has been discontinued [2]. There has been a demand for the development of a new assessment method to accurately predict drug safety using human stem cells. Furthermore, the use of patient-specific induced pluripotent stem cells (iPSCs) can be used to elucidate the pathological mechanism and to develop new therapeutic agents. Myocardial tissue in vivo is composed of multiple types of cells and extracellular matrix components (ECM) around the cells [3]. It is a three-dimensional structure that is precisely positioned and controlled. For use in therapeutic areas, development of an artificial myocardial tissue that mimics the microenvironment of the myocardial tissue in vivo is required. Various methods for producing a
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three-dimensional tissue have been developed, such as forming a spheroid, culturing with scaffolds, polymers, or a gel, constructing a cell sheet [4]. In this section, we describe a method for producing a threedimensional cardiac tissue using the layer-by-layer (LbL) method, which is a scaffold-free method and can produce a three-dimensional tissue using multiple cell types (Fig. 1) [5–7].
2 2.1
Materials Reagents
1. Fibronectin buffer: 0.2 mg/mL fibronectin in Dulbecco’s Phosphate Buffered Saline without Ca and Mg (D-PBS( )). 2. Gelatin buffer: 0.2 mg/mL gelatin in D-PBS( ). Solubilize and sterilize by autoclaving for 20 min at 121 C. Store at 4 C. 3. DMEM buffer: Dulbecco’s Modified Eagle Medium (DMEM) containing 10% Fetal Bovine Serum. Store at 4 C. 4. Coating buffer: 0.1% gelatin in distilled water (DW). Store at room temperature.
2.2
Equipment
1. Shaking incubator (SI-300; AS ONE). 2. 6 well insert: 24-mm transwell with 3.0 μm pore size (Corning). 3. 24 well insert: 6.5-mm transwell with 0.4 μm pore size (Corning).
3 3.1
Methods Cell Preparation
1. Wash dissociated human iPSCs derived cardiomyocytes with D-PBS( ). 2. Aspirate the supernatant and resuspend the pellet in D-PBS ( ). 3. Count the cell number.
3.2 ECM Coating on the Cell Surface
1. Set a 6-well plate on a shaking incubator. 2. Add 2.5 mL of fibronectin buffer to the upper left well (A1). 3. Add 2.5 mL of D-PBS( ) to the upper middle well (A2). 4. Add 2.5 mL of gelatin buffer to the upper right well (A3). 5. Set a 6-well insert in an empty well (see Note 1) and add the cell suspension into the 6-well insert (see Note 2). 6. Shake at 1000 rpm and drop the liquid under the insert. 7. Transfer the 6-well insert to A1 and add 550 μL of fibronectin buffer inside the 6-well insert.
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Fig. 1 Schematic image of LbL method
8. Incubate at 500 rpm for 1 min, wave mode. 9. Transfer the 6-well insert in an empty well. Shake at 1000 rpm and drop the liquid under the insert. 10. Transfer the 6-well insert to A2 and add 550 μL of D-PBS( ) inside the 6 well insert. 11. Incubate at 500 rpm for 1 min, wave mode. 12. Transfer the 6-well insert in an empty well. Shake at 1000 rpm and drop the liquid under the insert. 13. Transfer the 6-well insert to A3 and add gelatin buffer inside the 6-well insert. 14. Incubate at 500 rpm for 1 min, wave mode. 15. Transfer the 6-well insert in an empty well. Shake at 1000 rpm and drop the liquid under the insert. 16. Transfer the 6-well insert to A2 and add D-PBS( ) inside the 6-well insert. 17. Incubate at 500 rpm for 1 min, wave mode. 18. Transfer the 6-well insert in an empty well. Shake at 1000 rpm and drop the liquid under the insert. 19. Repeat Subheading 3.2, steps 7–19, three times. 20. Repeat Subheading 3.2, steps 7–9. 21. Add 3 mL of DMEM buffer to the empty well. 22. Transfer the 6-well insert to the well containing DMEM buffer. 23. Pipette the cells on the 6-well insert using P1000 Pipetman and collect cells in a tube. 24. Count the cell number.
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Fig. 2 Representative hematoxylin and eosin staining image of 3D cardiac tissue 3.3 Construction of 3D Cardiac Tissues
1. Set a 24 well insert to a 24-well plate. 2. Add coating buffer to the 24-well insert and incubate for at least 30 min in a CO2 incubator. 3. Aspirate coating buffer before cell seeding. 4. Seed the required amount of ECM coating cells into the 24-well insert (see Note 3). 5. At least 1 h after step 4, add DMEM buffer outside the 24-well insert (see Note 4). 6. Culture the plates at 37 C, CO2 5%, O2 20% (Fig. 2).
4
Notes 1. Before adding cells to the 6-well insert, slightly moisten the outer surface of the insert using D-PBS( ). The liquid will drain well. 2. If the cell concentration is too high, it tends to clump, and the liquid does not drop under the insert, resulting in reduced coating efficiency. It is better to prepare the cell concentration of less than 1 107 cells/mL. 3. To construct multilayer cardiac tissue, ECM coating cells are seeded at around 1 105 cells/a layer into a 24-well insert. 4. Once the cells have adhered, add DMEM buffer to connect media inside and outside the insert.
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References 1. Miyagawa S, Sawa Y (2018) Building a new strategy for treating heart failure using induced pluripotent stem cells. J Cardiol 72(6):445–448 2. Stevens JL, Baker TK (2009) The future of drug safety testing: expanding the view and narrowing the focus. Drug Discov Today 14:162 3. Howard CM, Baudino TA (2014) Dynamic cellcell and cell-ECM interactions in the heart. J Mol Cell Cardiol 70:19–26 4. Tomov ML, Gil CJ, Cetnar A, Theus AS, Lima BJ, Nish JE, Bauser-Heaton HD, Serpooshan V (2019) Engineering functional cardiac tissues for regenerative medicine applications. Curr Cardiol Rep 21(9):105 5. Matsusaki M, Kadowaki K, Nakahara Y, Akashi M (2007) Fabrication of cellular multilayers
with nanometer-sized extracellular matrix films. Angew Chem Int Ed Engl 46(25):4689–4692 6. Nishiguchi A, Yoshida H, Matsusaki M, Akashi M (2011) Rapid construction of threedimensional multilayered tissues with endothelial tube networks by the cell-accumulation technique. Adv Mater 23(31):3506–3510 7. Amano Y, Nishiguchi A, Matsusaki M, Iseoka H, Miyagawa S, Sawa Y, Seo M, Yamaguchi T, Akashi M (2016) Development of vascularized iPSC derived 3D-cardiomyocyte tissues by filtration layer-by-layer technique and their application for pharmaceutical assays. Acta Biomater 33:110–121
Chapter 9 Generation of Cylindrical Engineered Cardiac Tissues from Human iPS Cell-Derived Cardiovascular Cell Lineages Hidetoshi Masumoto Abstract The present protocol describes a method to generate cylindrical engineered cardiac tissues (ECTs) composed of cardiovascular cell lineages induced from human induced pluripotent stem cells (hiPSCs). Cardiomyocytes, endothelial cells, and vascular mural cells induced from hiPSCs are mixed with gel matrix and poured into a tissue mold with posts. By culture day 14, the mixed culture matures into a cylindrical ECT which beats spontaneously and synchronously. Cardiomyocytes align to the long axis of the ECT. The ECTs generated by the present method may be regarded as a surrogate of human myocardium and be served as researches in cardiac regenerative medicine, disease modeling, drug discovery, and cardiac toxicity tests. Key words Engineered cardiac tissue, Tissue engineering, Induced pluripotent stem cell, Cardiac differentiation, Cell culture
1
Introduction Numerous preclinical and clinical studies have revealed the efficiency of cell-based cardiac regenerative therapies for heart failure [1, 2]. Among various cell types, human induced pluripotent stem cells (hiPSCs) are recognized to be a promising cell source by virtue of their theoretically unlimited proliferative potential and the ability to generate various somatic cell lineages including cardiovascular cells [3]. HiPSC-derived cardiovascular cells are also potentially be applied for researches in disease modeling, drug discovery, and cardiac toxicity tests in vitro through providing a novel human cell-based platform [4, 5]. To promote the abovementioned possibilities of hiPSCs for medical research, establishment of three-dimensional (3-D) construction based on tissue engineering technologies might be a promising strategy to generate human myocardial tissue surrogates [6–10]. We have previously reported a 3-D cylindrical engineered cardiac tissues (ECTs) from hiPSC-derived cardiovascular lineages
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using a commercially available system [7]. In the report, we found that the coexistence of vascular lineage cells with cardiomyocytes within the ECT promoted structural and electrophysiological tissue function. Furthermore, we confirmed the therapeutic potential of the transplantation of the cylindrical ECTs in a rat myocardial infarction model to ameliorate cardiac dysfunction accompanied by restoration of human myocardium in the rat heart. In this protocol chapter, we report a method to generate hiPSC-derived ECTs utilizing tissue molds to generate porous engineered tissues. We coated tissue molds with a surfactant Pluronic F127 to prevent cell/gel matrix adhesion to the tissue mold which secured successful detachment of ECTs from the internal loading posts and the mold bottom during in vitro maturation facilitating gel compaction and ECT removal from the mold. Matured ECTs started intrinsic spontaneous beating and continued beating throughout the culture period. The hiPSC-derived ECTs would be utilized for medical researches of regenerative medicine, disease modeling, drug discovery and cardiac toxicity tests.
2
Materials
2.1 Maintenance and Cardiovascular Differentiation of hiPSCs
1. Cell culture dishes 100 20 mm style for hiPSC maintenance culture. 2. Multiwell plates for cell culture 6-well for differentiation culture. 3. Phosphate-buffered saline (PBS) 1 for cell wash before cell dissociation. 4. RPMI 1640 + B27 medium: B27 supplement minus insulin (50) and L-Glutamine (100) in RPMI1640 medium. 5. Growth factor-reduced Matrigel [1:60 diluted in Knock-out Dulbecco’s Modified Eagle Medium (DMEM)] for thincoating of culture dishes/plates (see Subheading 3.1, steps 1 and 3) and covering of the cells (see Subheading 3.1, step 4) 6. Recombinant human basic fibroblast growth factor (hbFGF). 7. Versene solution for cell dissociation. 8. Recombinant Human/Mouse/Rat ActivinA. 9. Recombinant Human Wnt3A (optional). 10. Recombinant bone morphogenetic protein 4 (BMP4). 11. Human vascular endothelial cell growth factor (VEGF)165 IS premium grade.
2.2 Cell Harvest and Lineage Analysis
1. ECT culture medium: 10% fetal bovine serum (FBS), 5 10 5 M 2-mercaptoethanol and 100 U/ml PenicillinStreptomycin in alpha minimum essential medium (αMEM).
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2. Accumax for differentiated cell dissociation (1). 3. FACS staining buffer: 5% FBS in PBS. 4. 0.75% Saponin in FACS staining buffer. 5. LIVE/DEAD® Fixable Aqua Dead Cell Stain Kit for 405 nm excitation (1:1000). 6. Anti-TRA-1-60-fluorescein isothiocyanate (FITC) antibody (1:20). 7. Anti-vascular (1:100).
endothelial
(VE)-cadherin-FITC
antibody
8. Anti-CD140 (PDGFRβ)-PE antibody (1:100). 9. Anti-cardiac isoform of troponin-T antibody. 10. Zenon® Alexa Fluor® 488 Mouse IgG1 Labeling Kit (1:50 as mixture of 9 and 10). 11. FACS tubes with cell strainer cap. 2.3
ECT Construction
1. Acrylic resin block for tissue molds. 2. 1% Pluronic® F-127 0.2 μm filtered (10% Solution in Water) (1:10 diluted in PBS) for coating of tissue mold. 3. Cell suspension medium: 20% FBS, 1% Penicillin-Streptomycin in high-glucose DMEM. 4. Collagen Type I solution from rat tail (2 mg/ml, pH 3). 5. Alkali buffer: 0.2 M NaHCO3, 0.2 M HEPES and 0.1 M NaOH in PBS. 6. Matrigel Basement Membrane Matrix (1).
3
Methods
3.1 Maintenance and Cardiovascular Differentiation of hiPSCs
1. Maintain undifferentiated hiPSCs on thin-coat Matrigel (growth factor-reduced, 1:60 dilution) in conditioned medium collected from culture of mouse embryonic fibroblasts (MEF-CM) added with hbFGF (see Notes 1–3). 2. Use Versene [0.48 mM ethylenediaminetetraacetic acid (EDTA) solution] to detach and dissociate cells when cell confluency becomes 90–100% (see Note 4). 3. Seed cells at a density of 10,000/mm2 on Matrigel-coated cell culture plates in MEF-CM added with hbFGF. 4. Cover the cells with medium containing Matrigel (1:60 dilution with MEF-CM) for 1 day when the culture reaches 100% confluent. 5. Replace the MEF-CM with RPMI 1640 + B27 medium. Add 100 ng/ml of Activin A to the medium for 1 day (this is day 0 of differentiation) (see Note 5).
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Fig. 1 Human iPS cell differentiation protocol toward cardiovascular cell lineages. MEF-CM mouse embryonic fibroblast conditioned medium, bFGF basic fibroblast growth factor, ActA Activin A, BMP4 bone morphogenetic protein 4, VEGF vascular endothelial cell growth factor. (Referred from ref. 6 under a Creative Commons Attribution-NonCommercialNoDerivs 4.0 International License)
6. On day 1 of differentiation, change the medium to RPMI 1640 + B27 with 10 ng/ml of BMP4 and 10 ng/ml of hbFGF. 7. On day 5 of differentiation, replace the medium with RPMI 1640 + B27 supplemented with 50 ng/ml of VEGF165. Change the culture medium every 48 h until day 13–15 of differentiation (Fig. 1). 3.2 Cell Harvest and Lineage Analysis
1. Wash the cells with Ca2+/Mg2+ free PBS. 2. Add Accumax (cell dissociation solution including proteases, collagenases and DNAses) cover the cultured cell. Incubate the plate for 15 min at 37 C. 3. Collect and dissociate the cells with culture medium and pipette them to have single cell suspension. 4. Allocate 1 106 cells for lineage analysis by flow cytometry. To eliminate dead cells, stain the cells with LIVE/DEAD fixable Aqua dead cell staining kit according to manual instructions. 5. Stain the cells with membrane surface markers in FACS staining buffer (PBS with 5% FBS). Use the following dilutions of antibody in FACS staining buffer: anti-PDGFRβ-PE (1:100), anti-VE cadherin-FITC (1:100), anti-TRA-1-60-FITC (1:20). 6. For the labeling of intracellular proteins such as cTnT, resuspend and fix the cells with 4% paraformaldehyde (PFA) in PBS. Stain the cells with anti-cTnT in PBS with 5% FBS and 0.75% Saponin. Zenon® Alexa Fluor® 488 Mouse IgG1 Labeling Kit can be used to fluorescently label the cTnT antibody (dilution 1:50) (see Note 6). 7. Resuspend the stained cells in PBS with 5% FBS and put them in FACS tubes through cell strainer. 8. Analyze the cell composition of the stained cells by flow cytometry to check cellular composition (see Note 7). Dead cells are
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detected with UV filter. CTnT-positive cardiomyocytes, VEcadherin-positive vascular endothelial cells and TRA-1-60-positive undifferentiated hiPSCs are detected by FITC filter. PDGFRβ-positive vascular mural cells are detected by PE filter. 3.3
ECT Construction
1. Prepare tissue mold from acrylic resin by cutting resin block with any methods such as laser or ultrasonic cutters (see Note 8) (Fig. 2). Sterile the tissue mold by immersion in 70% ethanol for 1–2 h. 2. Coat the tray with 1% Pluronic F127 in PBS for 1 h. Rinse the mold with PBS sufficiently (at least twice) prior to use. 3. Suspend induced cardiovascular cells in ECT culture medium (αMEM alpha minimum essential medium supplemented with 10% FBS, 5 10 5 M 2-mercaptoethanol and 100 U/ml Penicillin-Streptomycin) (see Notes 9 and 10). 4. Centrifuge the prepared cell suspension and resuspend the cells with 1.4 μl/mm2 of cell suspension medium (20% FBS and 1% Penicillin-Streptomycin in high-glucose DMEM). 5. Prepare matrix solution: Mix 1.1 μl/mm2 of acid-soluble rat tail collagen type I solution (2 mg/ml, pH 3) with 0.14 μl/ mm2 of 10 MEM. Then mix the solution with 0.14 μl/mm2 of alkali buffer to neutralize (0.2 M NaHCO3, 0.2 M HEPES, and 0.1 M NaOH) (see Notes 11–13). 6. Add 0.56 μl/mm2 of Matrigel to the neutralized matrix solution (see Note 14). 7. Mix the cell suspension and the matrix solution. The total volume of cell/matrix mixture for one construct is 3.3 μl/ mm2 (100 μl when the area of tissue mold is 30 mm2) (see Note 15). 8. Pour the cell/matrix mixture evenly into a Pluronic F127coated tissue mold (see Note 16). 9. Incubate the cell/matrix mixture in a standard CO2 incubator (37 C, 5% CO2) for 60 min. 10. After the tissue is formed, soak the tissue mold with ECT culture medium (see Note 17). 11. Culture the tissue for 14 days with medium change every day. 12. Unload the ECTs from the tissue mold for research use (see Notes 18 and 19) (Fig. 3).
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Notes 1. Adjust hbFGF at the appropriate concentration for each iPS cell line.
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Fig. 2 An example of acrylic resin tissue mold
Fig. 3 Human iPS cell-derived engineered cardiac tissues before (a) and after (b) removal from tissue molds
2. Laminin-511 E8 fragment can also be used for the coating of culture dish instead of Matrigel. 3. Commercially available medium designed for feeder-free culture of hiPSCs might be used as a substitute for MEF-CM. 4. Other commercially available reagents for cell dissociation can be used as well. 5. To promote differentiation efficiency of cardiomyocytes and vascular endothelial cells, Wnt3A or GSK3β inhibitors can be used on differentiation day 0 to upregulate canonical Wnt signaling. 6. Cell surface markers and intracellular markers can be simultaneously stained. Keep order: cell surface markers first, and then intracellular markers after fixation. 7. While performing this procedure, the remaining cell suspension for ECTs can be preserved in 4 C.
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8. For tissue molds, other biocompatible materials such as polydimethylsiloxane can be used as well [8]. 9. For higher tissue function, vascular mural cells should be ideally 10–20% of total cells [7]. 10. Ideal cell number for one ECT is 50,000 cells/mm2 (area of tissue mold) (1,500,000 cells when the area of tissue mold is 30 mm2). 11. Check buffer color. If the medium does not become pinkish when all solutions are mixed, add additional alkali buffer. 12. The mixing step must be collagen I + 10 MEM, and then + alkali buffer. DO NOT change this order. 13. Do NOT generate bubbles in the mixture of matrix solution. 14. The mixed solution must be kept on ice (0–4 C). 15. The cell/matrix mixture should be pinkish at this step. Keep the mixture on ice until being poured into tissue mold not to make it solidified. 16. Pour the mixture carefully not to contaminate bubbles inside of the poured gel which leads to defects in the ECT. 17. Add medium gently not to damage the gel (the gel is still too fragile). 18. Unloaded ECT exhibits intrinsic spontaneous beating in warm culture medium. The size of ECT after removal from the tissue molds is less than the original mold. Although unloaded ECT initially maintains a cylindrical structure, it shrinks over time without anchorages at both ends. 19. It is feasible to gently hold the ECTs with fine forceps and pierce them with surgical sutures such as 7-0 silk/polypropylene sutures.
Acknowledgment This work was supported by the Organoid Project at the RIKEN Center for Biosystems Dynamics Research. HiPSCs used in our published protocols were provided by the Center for iPS Cell Research and Application (CiRA), Kyoto University, Kyoto, Japan. I thank Dr. Jun K. Yamashita (CiRA, Kyoto University) for instructions on human iPS cell differentiation methods toward cardiovascular cells. I thank Dr. Bradley B. Keller (University of Louisville, Louisville, KY, USA) for instructions on the preparation of ECTs. The author has nothing to disclose regarding the conflict of interest.
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References 1. Sanganalmath SK, Bolli R (2013) Cell therapy for heart failure: a comprehensive overview of experimental and clinical studies, current challenges, and future directions. Circ Res 113 (6):810–834. https://doi.org/10.1161/ CIRCRESAHA.113.300219 2. Menasche P (2020) Cardiac cell therapy: current status, challenges and perspectives. Arch Cardiovasc Dis 113(4):285–292. https://doi. org/10.1016/j.acvd.2020.01.002 3. 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. https://doi.org/10.1016/j.cell. 2007.11.019 4. Kitaguchi T, Moriyama Y, Taniguchi T, Ojima A, Ando H, Uda T, Otabe K, Oguchi M, Shimizu S, Saito H, Morita M, Toratani A, Asayama M, Yamamoto W, Matsumoto E, Saji D, Ohnaka H, Tanaka K, Washio I, Miyamoto N (2016) CSAHi study: evaluation of multi-electrode array in combination with human iPS cell-derived cardiomyocytes to predict drug-induced QT prolongation and arrhythmia—effects of 7 reference compounds at 10 facilities. J Pharmacol Toxicol Methods 78:93–102. https://doi. org/10.1016/j.vascn.2015.12.002 5. Kanda Y, Yamazaki D, Osada T, Yoshinaga T, Sawada K (2018) Development of torsadogenic risk assessment using human induced pluripotent stem cell-derived cardiomyocytes: Japan iPS Cardiac Safety Assessment (JiCSA) update. J Pharmacol Sci 138(4):233–239. https://doi.org/10.1016/j.jphs.2018.10.010
6. Masumoto H, Ikuno T, Takeda M, Fukushima H, Marui A, Katayama S, Shimizu T, Ikeda T, Okano T, Sakata R, Yamashita JK (2014) Human iPS cell-engineered cardiac tissue sheets with cardiomyocytes and vascular cells for cardiac regeneration. Sci Rep 4:6716. https://doi.org/10.1038/srep06716 7. Masumoto H, Nakane T, Tinney JP, Yuan F, Ye F, Kowalski WJ, Minakata K, Sakata R, Yamashita JK, Keller BB (2016) The myocardial regenerative potential of threedimensional engineered cardiac tissues composed of multiple human iPS cell-derived cardiovascular cell lineages. Sci Rep 6:29933. https://doi.org/10.1038/srep29933 8. Nakane T, Masumoto H, Tinney JP, Yuan F, Kowalski WJ, Ye F, LeBlanc AJ, Sakata R, Yamashita JK, Keller BB (2017) Impact of cell composition and geometry on human induced pluripotent stem cells-derived engineered cardiac tissue. Sci Rep 7:45641. https://doi.org/ 10.1038/srep45641 9. Kawatou M, Masumoto H, Fukushima H, Morinaga G, Sakata R, Ashihara T, Yamashita JK (2017) Modelling Torsade de Pointes arrhythmias in vitro in 3D human iPS cellengineered heart tissue. Nat Commun 8 (1):1078. https://doi.org/10.1038/s41467017-01125-y 10. Masumoto H, Yamashita JK (2013) Strategies in cell therapy for cardiac regeneration. Inflamm Regen 33(2):114–120. https://doi. org/10.1152/ajpheart.00291.2008.-The
Part III Physiological Measurements Using Pluripotent Stem CellDerived Cardiomyocytes
Chapter 10 Protocol for Morphological and Functional Phenotype Analysis of hiPS-Derived Cardiomyocytes Jun Li and Jong-Kook Lee Abstract Induced pluripotent stem cells (iPSCs) have been utilized to study physiological development and also the pathogenesis of heart diseases. iPS-derived cardiomyocytes and engineered cardiac tissues provide a promising capacity for investigating cardiac development and disease modeling. In addition to protocols for cardiac differentiation and 3D cardiac tissue construction, the establishment of protocols for the comprehensive evaluation of the physiological and/or pathophysiological properties for the iPS-derived cells/tissues are indispensable. Key words iPS-derived cardiomyocytes, Morphology, Contraction, Calcium transient, Membrane potential, Cardiac conduction
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Introduction Induced pluripotent stem cells (iPSCs) have been considered as a promising platform for the investigation of cardiac differentiation/ development [1, 2] and disease onset/progress [3, 4] due to their potentials of indefinite proliferation and cardiac differentiation [1, 5, 6]. However, the immature nature of hiPSC-CMs has limited further steps for clinic application [7]. Although many efforts have been made to facilitate the maturation of hiPS-CMs [8], the appropriate and efficient evaluation methods are particularly important [3, 5]. Here, we show a simple and comprehensive method for morphological and functional evaluation (calcium transient, membrane potential, conductivity) of human iPS-derived cardiomyocytes (hiPS-CMs) in vitro.
Yoshinori Yoshida (ed.), Pluripotent Stem-Cell Derived Cardiomyocytes, Methods in Molecular Biology, vol. 2320, https://doi.org/10.1007/978-1-0716-1484-6_10, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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Materials Reagents
1. Geltrex® LDEV-Free hESC-qualified Reduced Growth Factor Basement Membrane Matrix (Gibco, A1048002), stored at 30 C. 2. 0.25% Trypsin EDTA, stored at 4 C. 3. FluoroBrite™ DMEM (Gibco, A18967-01), stored at 4 C. 4. 1 M HEPES sodium salt solution, stored at room temperature. 5. GlutaMAX™ supplement (Gibco, 35050061), stored at room temperature. 6. Pluronic F-127™ (Invitrogen, P3000MP), stored at room temperature. 7. Indo-1 solution: 1 mmol/L Indo-1 AM (Dojindo, 1006) in DMSO. 8. Indo-1 loading solution: dilute 1 mmol/L Indo-1 solution 1:200 with FluoroBrite™ DMEM contained 10% FBS, 10 mM HEPES and 1 GlutaMAX, supplemented with 0.1% Pluronic F-127. Pipetting well or vortex to mix. The procedure application should avoid direct exposure to light. 9. Voltage-sensitive fluorescent dye: FluoVolt™ Membrane Potential Kit (Invitrogen, F10488), stored at 4 C. 10. FluoVolt loading solution: dilute 100 PowerLoad™ 1:100 and 1000 FluoVolt™ dye 1:1000 with pre-warmed FluoroBrite™ DMEM. Pipet well. Direct exposure to light should be avoided during the procedure. 11. Blebbistatin (Wako, 027-17043) is diluted to 10 mM stock solution with DMSO. (Working concentration: 10 μM). 12. 4%Paraformaldehyde (PFA), stored at 4 C. 13. 0.1%Triton X in PBS, stored at 4 C. 14. 2%Bovine serum albumin (BSA) in PBS, stored at 4 C. 15. Primary antibody: cardiac troponin-T (cTnT, Lab vision, MS-295-P1) diluted with 1:200 in 2%BSA. 16. Secondary antibody: Alexa Fluor® 488 (Invitrogen, A-21202) was diluted with 1:500 in 2% BSA. 17. Hoechst® 33342 solution: Hoechst® 33342 (Dojindo, H342) was diluted to 1 μg/mL with PBS ( ).
2.2 Equipment and Software
1. FDSS/μCELL equipped with EFS 96 channel multi-electrode array (Hamamatsu Photonics, Hamamatsu). 2. FDSS software U8524-12 (Hamamatsu Photonics): data analysis for FDSS/μCELL. 3. MiCAM02 imaging system (BrainVision) combined with MyoPacer EP (IonOptix, Westwood, MA).
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4. BV_Ana software (BrainVision): data analysis for MiCAM02 imaging system. 5. High-content imaging system (Operetta™, PerkinElmer). 6. Harmony® high-content analysis software (PerkinElmer): data analysis for Operetta™.
3 3.1
Methods Cell Preparation
On the day 25–30 after the initiation of cardiac differentiation, hiPS-derived cardiomyocytes monolayer are enzymatically dissociated into single cells by 0.25% Trypsin EDTA. Re-plating hiPSCMs onto Geltrex® Matrix pre-coated 96 well plates (see Note 1), further cultured for 5–7 days until cells show synchronized beating in throughout the well. The density of cell seeding depends on the purpose of the experiment. For functional analysis, higher cell density (1 105 ~ 2 105cells/well) is recommended. On the other hand, for morphological evaluation, the lower cell density (5 103 ~ 1.5 104cells/well) will be more suitable (see Note 2).
3.2 Calcium Transients
1. Remove culture medium and rinse cells twice in pre-warmed PBS ( ) (see Note 3).
3.2.1 Indo-1 Loading
2. Add 100 μl/well Indo-1 loading solution to cells, and incubate at 37 C for 60 min. 3. Remove Indo-1 loading solution, and wash cells twice in pre-warmed PBS ( ) (see Note 4). 4. Replace with 100 μl/well FluoroBrite™ DMEM containing 10% FBS, incubate at 37 C for 30 min.
3.2.2 Calcium Transient Measurement
1. Setting culture plate into FDSS/μCELL, stabilize for 3–5 min before the measurement. 2. Fluorescence signals are obtained throughout 450–490 nm excitation bandpass and 540 nm emission filter set (see Note 5). 3. Calcium transients are recorded for ~100 s, under spontaneous beating condition for the first 30s, and then under paced conditions at 1 Hz and 1.5 Hz stepwise for 30s respectively (see Note 6).
3.2.3 Data Analysis
1. Parameters are automatically detected and adjusted as appropriate (Fig. 1). Waveforms are obtained under unfiltered condition. Adjust the peak threshold so that the real peak identified correctly. 2. Table 1 shows the main parameters including beating rate, wave amplitude, rising/falling slope, and calcium transient duration at 90% relaxation.
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Fig. 1 Calcium transient recording. (a) Representative traces of calcium transients of hiPS-CMs under the conditions of (a) spontaneous beating, (b) pacing at 1 Hz, and (c) at 1.5 Hz. Scale bars indicate 1 s. (b) The waveforms of calcium transient processed by FDSS software. The parameters marked with relevant symbols. Both baseline and bottom were recognized automatically, the red triangles represent peaks that were identified above the P threshold line Table 1 Major parameters of calcium transients P rate (/min) (BPM)
Amplitude (AMP)
Rising slope
Falling slope
PWD90 (ms)
23.46
0.31
0.00084
0.00024
1708.52
3.3 Field Membrane Potential
1. Remove culture medium and wash cells twice in pre-warmed PBS ( ) (see Note 7).
3.3.1 FluoVolt™ Loading
2. Add 100 μl/well FluoVolt loading solution, and incubate at 37 C or room temperature for 30 min (see Note 8). 3. Remove the loading solution, and wash cells twice in pre-warmed PBS ( ) (see Note 9). 4. Add 100 μl/well FluoroBrite™ DMEM containing 10% FBS, incubate at 37 C for 30 min.
3.3.2 Field Membrane Potential Recording
1. Set culture plates into FDSS, stabilize for 3–5 min before measurement. 2. Fluorescence signals are recorded using 480 nm excitation and 540 nm emission filter set. 3. Field membrane potentials are recorded for ~100 s, under spontaneous beating condition for the first 30s, and then
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under paced conditions at 1 Hz and 1.5 Hz stepwise for 30s respectively. 3.3.3 Data Analysis
1. Parameters are automatically detected and can be adjusted as appropriate (Fig. 2). 2. Table 2 shows the main parameters including beating rate, wave amplitude, rising/falling slope, and membrane potential duration at 90% relaxation.
3.4
Optical Mapping
Same as Subheading 3.3.1, “FluoVolt™ loading.”
3.4.1 Dye Loading Procedure 3.4.2 Optical Membrane Potential Imaging
1. Before recording, 10 μM blebbistatin is applied to avoid the effects of motion artifact. 2. Optical mapping was taken by a high-speed and low-noise CMOS camera, throughout the field of view (5.76 mm 4.8 mm), setting with 520/35–25 nm excitation and 580 nm emission filter. 3. Membrane potential signals are acquired at 5 ms/frame of sampling rate by MiCAM02 imaging system. 4. Electrical stimulation is performed with a bipolar electrode at the sites of the culture, with the voltage at 5 V. Pacing is started at 0.5 Hz and increased stepwise (see Note 10).
3.4.3 Data Processing
1. The fluorescent signal was normalized and underwent spatial filtering of 3 3 uniform binning and fast Fourier transform with a cutoff value at 20% of the sampling rate frequency (Fig. 3). 2. Activation map is shown as a color isochronous curve, the activation time is defined as the time for the max intensity of optical potential signal to reach each time point (Fig. 4).
3.5 Morphological Analysis
1. Remove culture medium, wash cells twice with PBS ( ), fixed with cold 4% PFA for 15 min at room temperature.
3.5.1 Immunofluorescent Staining
2. Permeabilized with 0.1% Triton X-100 for 10 min at room temperature. 3. Then blocking nonspecific antibody binding site with 2% BSA for 60 min at room temperature. 4. Incubate with primary antibody overnight at 4 C. 5. Rinse cells with PBS ( ) three times 5 min, then followed by secondary antibody application in the dark for 60 min at room temperature.
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Fig. 2 Membrane potential recording. (a) Representative traces of membrane potentials of hiPS-CMs under the conditions of (a) spontaneous beating, (b) pacing at 1 Hz, and (c) at 1.5 Hz. Scale bars indicate 1 s. (b) The waveforms of membrane potential processed by FDSS software. The parameters marked with relevant symbols. The baseline and bottom were recognized automatically, the red triangles represent the peaks that were identified above the P threshold line Table 2 Major parameters of membrane potentials P rate (/min) (BPM)
Amplitude (AMP)
Rising slope
Falling slope
PWD90 (ms)
36.15
4091.17
28.92
7.66
667.10
6. Wash cells with PBS ( ) three times 5 min. Afterward stain the nuclei with Hoechst® 33,342 solution (1 μg/mL) for 10 min. 3.5.2 Image Processing
1. Immunostaining imaging is obtained by Harmony highcontent imaging system and follows on to processed by the analysis software. 2. Cardiac troponin-T (cTnT) positive cells are quantified by calculating the intensity of relevant fluorescent dye (Fig. 5a). The cell shape characteristic is quantified by evaluating the cell roundness and the ratio of width to length. When cells show rounder shape, both parameters are closer to 1 (Fig. 5b).
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Fig. 3 Optical mapping of membrane potentials. The membrane potential of hiPS-CMs under the conditions of (a) spontaneous beating, (b) pacing at 1 Hz, and (c) at 1.5 Hz. The three lines of tracings with different colors represent membrane potential signals detected from ROI at three different sites
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Notes 1. (a) Undiluted Geltrex®Matrix solution is gelatinous, the stock solution remove from 30 C should be thawing at 4 C. (b) Before coating plate, dilute 1:100 in cold PBS ( ), pipetting evenly on ice and distributed to 96-well plate quickly, then incubate for 60 min at 37 C. (c) The various dilution ratio from 1:100 to 1:200 will be fine to hiPS-CMs. 2. For functional analysis, the higher cell density will reduce the time duration from re-seeding to the hiPS-CMs get a synchronized beating, as well as have lower noise. However, for morphological study, fewer cells are needed so that the single cell identity will be more accurate.
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Fig. 4 Activation map. Optical mapping (left: image, right: movie) of hiPS-CMs under the conditions of (a) spontaneous beating, (b) pacing at 1 Hz, and (c) at 1.5 Hz. Color bars at the bottom of each activation map showed activation time (ms). The representative movies play at 60 frames per second
3. Rinsing of cells with PBS ( ) may cause cell detachment occasionally. In such cases, FluoroBrite™ DMEM can be substituted for rinsing. 4. See Note 3. 5. Filter selection depends on which fluorescent dye for calcium imaging you choose. Usually like fura-2, Cal520, fura-3, fura-4, and Indo-1. Filter wavelength should cover the spectrum of fluorescent dye. 6. (a) The time course for spontaneous beating, pacing duration, and interval time are artificial setting. (b) Try various electrical stimulation voltages and pacing rates according to the spontaneous beating rate, as well as cell condition (stimulus threshold). 7. See Note 3.
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Fig. 5 Morphological analysis. (a) Immunostain images of hiPS-CMs. The cTnT positive cells are about 60%. cTnT, green; nuclei, blue. Scale bar, 100 m. (b) The examples of different cell shapes, the rounder cell (a) and rod-like cell (b). The parameters “Roundness” and “Ratio of width to length” were calculated by Harmony analysis system. The values are shown in the table below
8. Although the voltage-sensitive dye can be load either at 37 C or at room temperature, the procedure at 37 C is recommended. 9. See Note 3. 10. See Note 6(b).
Acknowledgments The authors are grateful to Mayuko Matsushima for technical assistance, and Keiko Miwa, Yuhki Kuramoto, Taku Sakai, Hiroyuki Nakanishi, Akira Yoshida, Teruki Yokoyama, Satoki Tomoyama, Hideki Yasutake and Kiyoshi Masuyama for the share of invaluable experimental experiences as well as for useful advice. The present studies were supported in part by AMED under Grant Number JP17bm0804008h0001, JSPS KAKENHI Grant Number JP18H03517 and Co-Create Knowledge for Pharma Innovation with Takeda (COCKPI-T®) Funding. References 1. Germanguz I, Sedan O, Zeevi-Levin N et al (2011) Molecular characterization and functional properties of cardiomyocytes derived from human inducible pluripotent stem cells. J Cell Mol Med 15:38–51
2. Kamp TJ, Lyons GE (2009) On the road to iPS cell cardiovascular applications. Circ Res 105 (7):617–619 3. Yoshida Y, Yamanaka S (2017) Induced pluripotent stem cells 10 years later: for cardiac applications. Circ Res 120:1958–1968
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4. Kawatou M, Masumoto H, Fukushima H et al (2017) Modelling Torsade de Pointes arrhythmias in vitro in 3D human iPS cell-engineered heart tissue. Nat Commun 8(1):1078 5. Denning C, Borgdorff V, Crutchley J et al (2016) Cardiomyocytes from human pluripotent stem cells: from laboratory curiosity to industrial biomedical platform. Biochim Biophys Acta 1863:1728–1748 6. Haraguchi Y, Matsuura K, Shimizu T et al (2015) Simple suspension culture system of
human iPS cells maintaining their pluripotency for cardiac cell sheet engineering. J Tissue Eng Regen Med 9:1363–1375 7. Kolanowski TJ, Antos CL, Guan K (2017) Making human cardiomyocytes up to date: derivation, maturation state and perspectives. Int J Cardiol 241:379–386 8. Scuderi GJ, Butcher J (2017) Naturally engineered maturation of cardiomyocytes. Front Cell Dev Biol 5:50
Chapter 11 Application of FluoVolt Membrane Potential Dye for Induced Pluripotent Stem Cell-Derived Cardiac Single Cells and Monolayers Differentiated via Embryoid Bodies Tadashi Takaki and Yoshinori Yoshida Abstract FluoVolt, a membrane potential dye, has been used to depict the action potentials of cardiomyocytes derived from human-induced pluripotent stem cells (hiPSC-CMs) in order to classify the cardiac cell subtype, evaluate long QT syndrome, and conduct cardiotoxic drug-responsive tests. To apply FluoVolt, users must prepare the hiPSC-CMs, assess the dye loadings, and apply the excitation light. Here we describe the steps to measure action potentials from single hiPSC-CMs and hiPSC-CM monolayers using this dye. Key words FluoVolt, Membrane potential dye, Action potential, Induced pluripotent stem cell, Cardiac single cells, Cardiac cell subtype
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Introduction Although the patch clamp technique is considered the gold standard to analyze the electrophysiological properties of cardiomyocytes, it requires much time to obtain data with advanced techniques [1]. FluoVolt (FV), a membrane potential dye, responds faster than Fo¨rster resonance energy transfer (FRET)-based voltage sensitive dyes (VSDs) and is more sensitive (about 20% ΔF/F per 100 mV greater) than classical electrochromic dyes because it modulates the photo-induced electron transfer (PeT) from an electron donor through a synthetic molecular wire to a fluorophore [2]. FV has been used to analyze the subtypes of single cardiomyocytes derived from human-induced pluripotent stem cells (hiPSC-CMs) [3, 4] and action potentials (APs) in hiPSC-CM monolayers [4– 6]. To apply FV, a series of steps including preparation of the hiPSC-CMs, loading of the dye, and the application of excitation light are needed. Here we describe the methods for measuring APs from single hiPSC-CMs and hiPSC-CM monolayers using FV.
Yoshinori Yoshida (ed.), Pluripotent Stem-Cell Derived Cardiomyocytes, Methods in Molecular Biology, vol. 2320, https://doi.org/10.1007/978-1-0716-1484-6_11, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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2.1 Embryoid Body (EB) Dissociation
1. StemPro-34 SFM added with StemPro-Nutrient Supplement and stored at 4 C protected from light. 2. L-glutamine: Make a 200 mM stock solution and store at 30 C. 3. L-ascorbic acid: Make a 5 mg/mL stock solution dissolved in dH2O and store at 30 C. 4. Transferrin: Make a 30 mg/mL stock solution and store at 4 C. 5. Monothioglycerol (MTG): Make a 0.135 M working solution dissolved in StemPro-34 SFM and store at 4 C. 6. rhVEGF: Make a 100 μg/mL stock solution dissolved in 0.1% BSA–DPBS ( ) and store at 80 C, and make a 10 μg/mL working solution dissolved in 0.1% BSA–DPBS ( ) and store at 4 C. 7. 10,000 U/mL of penicillin and 10,000 μg/mL of streptomycin. 8. Maintenance medium 1: StemPro-34 SFM supplemented with 2 mM L-glutamine, 50 μg/mL ascorbic acid, 150 μg/mL transferrin, 4 10 4 M monothioglycerol (MTG), 10 ng/ mL rhVEGF, 50 U/mL penicillin, and 50 μg/mL streptomycin. Add 1 mL of 200 mM L-glutamine, 1 mL of 5 mg/mL Lascorbic acid, 500 μL of 30 mg/mL transferrin, 300 μL of 0.135 M MTG, 100 μL of 10 μg/mL rhVEGF, and 500 μL of penicillin–streptomycin liquid to 100 mL StemPro-34 SFM. 9. Collagenase type I: Make a 2 mg/mL stock solution dissolved in 20% (v/v) FBS-1 DPBS (+) and store at 30 C. 10. Accumax. 11. Low-retention chip for 1 mL micropipettes.
2.2 Single-Cell Seeding onto a Fibronectin-Coated Plate
1. 1 mg/mL fibronectin stock solution: 5 mg fibronectin dissolved in 5 mL dH2O and stored at 80 C. 2. 50 μg/mL fibronectin working solution: 1 mg/mL fibronectin stock solution dissolved in DPBS ( ) and stored at 4 C. 3. 60-mm cell culture dishes.
2.3 FluorescenceActivated Cell Sorting (FACS) and Cryopreservation
1. Flow cytometry buffer: 2% (v/v) FBS dissolved in DPBS ( ) and stored at 4 C. 2. PE mouse anti-human CD140b (BD Pharmingen; 100 μL/ mL flow cytometry buffer). 3. PE mouse anti-human CD49a (BD Pharmingen; 60 μL/mL flow cytometry buffer).
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4. PE/Cy7 mouse anti-human SIRPa (Biolegend; 50 μL/mL flow cytometry buffer). 5. PE mouse anti-human CD31 (BD Pharmingen; 40 μL/mL flow cytometry buffer). 6. Alexa Fluor 488 mouse anti-human TRA-1-60 antigen (BD Pharmingen; 25 μL/mL flow cytometry buffer). 7. APC mouse anti-human CD90 (BD Pharmingen; 20 μL/mL flow cytometry buffer). 8. 5 mg/mL DAPI stock solution: Dissolved in DPBS ( ) and stored at 30 C. 9. 50 μg/mL DAPI working solution: Dissolved in DPBS ( ) and stored at 4 C. 10. STEM-CELLBANKER. 11. Ice. 12. Accumax. 13. 1.5 mL tubes. 14. 5 mL round-bottom tubes with 40 μm cell strainer caps. 15. 15 mL or 50 mL conical tubes (for collecting sorted cells). 16. 2 mL cryotube. 2.4 Seeding iPSC-CM Single Cells on a Glass-Bottom Dish
1. 50 μg/mL fibronectin working solution. 2. Maintenance medium 1 (item 8 in Subheading 2.1). 3. 35-mm glass-bottom dishes. 4. A cryotube containing 1–2 105 cells. 5. 37 C water bath.
2.5 Seeding High-Density iPSC-CM Monolayers on Glass-Bottom Dishes
1. 50 μg/mL fibronectin working solution. 2. rhbFGF: Make a 100 μg/mL stock solution dissolved in 1 mM DTT-0.1% BSA–DPBS ( ) and store at 80 C, and make a 10 μg/mL working solution dissolved in 1 mM DTT-0.1% BSA–DPBS ( ) and store at 4 C. 3. Maintenance medium 2: Maintenance medium 1 plus 5 ng/ mL rhbFGF; add 50 μL of 10 μg/mL rhbFGF working solution to 100 mL maintenance medium 1. 4. 35-mm glass-bottom dishes. 5. A cryotube containing 2.5–10 105 cells. 6. 37 C water bath.
2.6 Loading of FluoVolt (FV)
1. FluoVolt dye (see Note 1). 2. Tyrode’s solution: 120 mM NaCl, 5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 0.84 mM Na2HPO4, 0.28 mM MgSO4, 0.22 mM KH2PO4, 27 mM NaHCO3, and 5.5 mM glucose, pH 7.4.
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2.7 Recording of APs from Single Cells and Monolayers of Cardiomyocytes
1. Optical microscope setting including a cell culture environment and blue light irradiation apparatus. 2. Optical movie recording software with the function depicting intensity over time. 3. A thermometer connected with the medium in a 35-mm glassbottom dish.
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3.1 Embryoid Body (EB) Dissociation on Day 29 (see Note 2)
1. Thaw the collagenase type I stock solution in a 37 C water bath or at room temperature. 2. Tilt the low-bind plate culturing spontaneous beating EBs (see Fig. 1) and send the EBs to the bottom. 3. Exchange the culture medium (maintenance medium 1 or 2) for the collagenase type I stock solution of the same quantity and incubate at 37 C, 5% CO2 for 5–7 h. During this time, prepare a fibronectin-coated 60-mm dish (see step 1 in Subheading 3.2). 4. 5–7 h later, tilt the low-bind plate in order to send the EBs to the bottom. 5. Discard the collagenase type I solution, add 1 mL Accumax and transfer the EBs to a 1.5 mL tube using a 1 mL low-retention chip. 6. Incubate at 37 C, 5% CO2 sideways for 35 min. 7. Pipette three times with a 1 mL low-retention chip to get single cells (see Note 3). 8. Centrifuge at 300 g for 5 min. 9. Discard the supernatant and resuspend with the maintenance medium 1 (see Note 4).
Fig. 1 Spontaneous beating iPSC-CM aggregates 29 days after the start of the iPSC differentiation
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1. Coat a 60-mm dish with 50 μg/mL fibronectin working solution and leave in an incubator or on a clean bench. 2. Remove the fibronectin solution from the 60-mm dish and seed 3–4 106 cells in 4 mL maintenance medium 1. 3. Incubate at 37 C, 5% CO2 until the next day.
3.3 FluorescenceActivated Cell Sorting (FACS) and Cryopreservation
1. On the following day, prepare ice and the solution of conjugated antibodies in flow cytometry buffer before collecting the cells. For a 60-mm dish, add 50 μL PE mouse anti-human CD140b, 60 μL PE mouse anti-human CD49a, 50 μL PE/Cy7 mouse anti-human SIRPa, 40 μL PE mouse antihuman CD31, 25 μL Alexa Fluor 488 mouse anti-human TRA-1-60 antigen, and 20 μL APC mouse anti-human CD90 in 500 μL flow cytometry buffer. 2. Remove the maintenance medium, add 1 mL Accumax, and incubate for 7–10 min in 37 C, 5% CO2. 3. Collect the cells into a 1.5 mL tube. 4. Centrifuge at 300 g for 5 min. 5. Remove the supernatant and add the antibodies solution. 6. Lay the tube flat on ice protected from the light for 15 min. 7. Centrifuge at 300 g, 4 C for 5 min. 8. Remove the supernatant and suspend with 1 mL flow cytometry buffer. 9. Centrifuge at 300 g, 4 C for 5 min. 10. Remove the supernatant and suspend with 1 mL flow cytometry buffer. 11. Centrifuge at 300 g, 4 C for 5 min. 12. Remove the supernatant and suspend with 1 mL flow cytometry buffer. 13. Add 10 μL DAPI working solution for a final concentration of 0.5 μg/mL DAPI. 14. Filter the mixture through a 40-μm nylon mesh cell strainer and assay fluorescence-activated cell sorting. 15. Collect DAPI-negative, lineage (CD31, CD49a, CD140b, CD90, or TRA-1-60)-negative, and SIRPa-positive cells (see Fig. 2) in maintenance medium 1 cooled at 4 C (see Note 5). 16. Centrifuge at 300 g, 4 C for 10 min. 17. Remove the supernatant and CELLBANKER (see Note 6).
add
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19. A few days later, transfer the cryotube to a liquid nitrogen storage tank.
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Fig. 2 Flow cytometry contour plots on day 30. (a) PE-CD140b, CD49a, or CD31-positive cells were excluded. FITC-Tra-1-60-positive cells were not detected. (b) PE-Cy7-SIRPa-positive and APC-CD90-negative cells were sorted. Most CD90-positive cells were CD49a-positive and therefore excluded in (a) 3.4 Seeding Single iPSC-CMs on a GlassBottom Dish
1. Place 200 μL of 50 μg/mL fibronectin working solution on the center of a 35-mm diameter glass-bottom dish (see Fig. 3a). Distribute maintenance medium 2 on the surrounding so as not to dry the cells. Wait over 60 min at room temperature or 37 C (see Note 7). 2. Thaw a cryotube in a 37 C water bath, centrifuge at 300 g for 5 min, and resuspend in 100 μL maintenance medium 1. Count the number of cells and make a solution which includes 2 104 cells in 200 μL maintenance medium 1. 3. Aspirate the fibronectin solution in step 1, seed 2 104 cells, and incubate at 37 C, 5% CO2 until the following day. 4. Add 1.6–1.8 mL maintenance medium 1 the next day (see Fig. 4a), so that the final volume is around 2 mL. Exchange the medium every 2–3 days. The recommended incubation time is 5–10 days.
3.5 Seeding High-Density iPSC-CM Monolayer on a Glass-Bottom Dish
1. Place 5 μL of 50 μg/mL fibronectin working solution on the center of a 35-mm diameter glass-bottom dish (see Fig. 3b). Distribute maintenance medium 2 on the surrounding so as not to dry the cells. Wait over 60 min at room temperature or 37 C (see Note 7). 2. Thaw a cryotube in a 37 C water bath, centrifuge at 300 g for 5 min, and resuspend in 10 μL maintenance medium 2. Prepare a solution which includes 5 104 cells in 5 μL maintenance medium 1. To count cells, prepare 100 times diluted solution by mixing 1 μL with 99 μL medium.
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Fig. 3 Coated by fibronectin on a 35-mm glass-bottom dish. (a) Coating with 200 μL of 50 μg/mL fibronectin working solution on the center of a 35 mm diameter glass-bottom dish. (b) 5 μL fibronectin working solution placed on the center of a 35-mm glass-bottom dish. The media is distributed so as not to dry the center region
Fig. 4 Seeded iPSC-CMs on the following day. (a) Single iPSC-CMs. (b) High-density iPSC-CM monolayer
3. Aspirate the fibronectin solution in step 1 (see Note 8), seed 5 104 cells, and incubate at 37 C, 5% CO2 for 60 min. 4. Add about 1.8 mL maintenance medium 2 so that the final volume is around 2 mL (see Fig. 4b). Exchange the medium every 2–3 days. The recommended incubation time is 10–15 days. 3.6 Loading of FluoVolt (FV)
1. Make 0.1% volume FV solution by, for example, adding 1 μL FV to 1 mL Tyrode’s solution (see Note 9). 2. Aspirate the maintenance medium in a 35-mm glass-bottom dish and load 0.1% volume FV solution (see Note 9). 3. Twenty minutes after loading, aspirate the FV solution and add 1 mL Tyrode’s solution (see Note 10).
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3.7 Recording APs from Single iPSC-CMs and iPSC-CM Monolayers
1. Prepare the cell culture environment in an optical microscope setting. Set the 35-mm dish loaded with FV on a humid environment fed with 5% CO2 gas and maintain the temperature of the incubation apparatus to warm the Tyrode’s solution at 36–37 C (see Note 11). 2. Incubate for 1 h (see Note 12). 3. Apply 490 nm light from a Xe lamp or LED lamp to the cells and record a video of the flashing cells (see Note 13). 4. APs are calculated by plotting the light intensity in a region of interest (ROI) over time (see Fig. 5) (see Note 14).
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Notes 1. You do not need to use the PowerLoad™ Concentrate and Neuro Background Suppressor attachments of the FV kit. 2. The differentiation protocol is based on previous reports [4, 7, 8]. If you dissociate EBs on another day, you should consider the conditions for collagenase I and Accumax. 3. You do not need to disperse completely. 4. Do not use maintenance medium 2. Basic FGF-included medium will increase the ratio of non-cardiomyocytes. 5. Do not exceed 1.5 times the initial volume in the collected tube. For example, if the initial volume of the maintenance medium 1 was 10 mL, stop sorting when the liquid level reaches 15 mL. 6. We recommend 1–2 105 cells for single-cell seeding and 2.5–10 105 cells for monolayer seeding so as not to waste the cells. 7. It is acceptable to leave for 30 min or incubate for several days. 8. Do not aspirate fibronectin completely. Leave a little. 9. To save FV dye, you can make 0.1% volume FV solution by adding 0.2 μL FV into 0.2 mL Tyrode’s solution and load 200 μL in the center of the 35-mm glass-bottom dish. 10. The final volume should be 0.8–2 mL. 11. We recommend measuring the actual temperature of the Tyrode’s solution in a 35-mm dish. 12. If you start an experiment with less incubation time, the beating rate may not be stable. 13. We recommend using weaker light in order not to harm the cells. Therefore, ND (neutral density) filters are sometimes used. However, if you weaken the light too much, the intensity signal of the cells may be lost. Adjust the conditions accordingly.
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Fig. 5 Action potentials (APs) measured by using FluoVolt (FV) dye. (a, b) Single iPSC-CMs 6 days after the seeding. Scale bar, 50 μm. (b) APs of all 10 cells measured for 30 s on day 30 + 6 showed ventricular-like AP-morphology. (c, d) An iPSC-CM monolayer on day 11 days after the seeding. (c) The region of interest (ROI) was set to include almost whole cells. Scale bar, 100 μm. (d) AP of an iPSC-CM monolayer measured for 15 s on day 30 + 11 showed ventricular-like AP-morphology
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14. If you want to shorten the exposure time and shutter interval to get data in milliseconds from a beating monolayer, you may need to set the subarray recording mode or narrow the ROI.
Acknowledgments We thank Yoko Uematsu and Kaoru Shimizu for their administrative support, and Peter Karagiannis for proofreading the manuscript. This work was supported by a grant from Takeda Pharmaceutical Company Limited, a grant from Leducq foundation (18CVD05), JSPS KAKENHI Grant (17H04176), grants from the Research Center Network for Realization of Regenerative Medicine (JP19bm0104001, JP19bm0204003, and JP19bm0804008), Research on Regulatory Science of Pharmaceuticals and Medical Devices (JP19mk0104117), and Research Project for Practical Applications of Regenerative Medicine (JP19bk0104095) provided by the Japan Agency for Medical Research and Development, and iPS research fund. References 1. Yajuan X, Xin L, Zhiyuan L (2012) A comparison of the performance and application differences between manual and automated patchclamp techniques. Curr Chem Genomics 6:87–92. https://doi.org/10.2174/ 1875397301206010087 2. Miller EW, Lin JY, Frady EP, Steinbach PA, Kristan WB Jr, Tsien RY (2012) Optically monitoring voltage in neurons by photo-induced electron transfer through molecular wires. Proc Natl Acad Sci U S A 109(6):2114–2119. https://doi.org/10.1073/pnas.1120694109 3. Yechikov S, Copaciu R, Gluck JM, Deng W, Chiamvimonvat N, Chan JW et al (2016) Same-single-cell analysis of pacemaker-specific markers in human induced pluripotent stem cell-derived cardiomyocyte subtypes classified by electrophysiology. Stem Cells 34 (11):2670–2680. https://doi.org/10.1002/ stem.2466 4. Takaki T, Inagaki A, Chonabayashi K, Inoue K, Miki K, Ohno S et al (2019) Optical recording of action potentials in human induced pluripotent stem cell-derived cardiac single cells and monolayers generated from long QT syndrome type 1 patients. Stem Cells Int 2019:7532657. https://doi.org/10.1155/2019/7532657
5. Bedut S, Seminatore-Nole C, Lamamy V, Caignard S, Boutin JA, Nosjean O et al (2016) High-throughput drug profiling with voltageand calcium-sensitive fluorescent probes in human iPSC-derived cardiomyocytes. Am J Physiol Heart Circ Physiol 311(1):H44–H53. https://doi.org/10.1152/ajpheart.00793. 2015 6. McKeithan WL, Savchenko A, Yu MS, Cerignoli F, Bruyneel AAN, Price JH et al (2017) An automated platform for assessment of congenital and drug-induced arrhythmia with hiPSC-derived cardiomyocytes. Front Physiol 8:766. https://doi.org/10.3389/fphys.2017. 00766 7. Dubois NC, Craft AM, Sharma P, Elliott DA, Stanley EG, Elefanty AG et al (2011) SIRPA is a specific cell-surface marker for isolating cardiomyocytes derived from human pluripotent stem cells. Nat Biotechnol 29(11):1011–1018. https://doi.org/10.1038/nbt.2005 8. Miki K, Endo K, Takahashi S, Funakoshi S, Takei I, Katayama S et al (2015) Efficient detection and purification of cell populations using synthetic microRNA switches. Cell Stem Cell 16(6):699–711. https://doi.org/10.1016/j. stem.2015.04.005
Chapter 12 Multielectrode Array Assays Using Human-Induced Pluripotent Stem Cell-Derived Cardiomyocytes Daisuke Yoshinaga, Yimin Wuriyanghai, and Takeru Makiyama Abstract Induced pluripotent stem cell (iPSC)-derived cardiomyocytes (iPSC-CMs) have been shown to have great potential to play a key role in investigating cardiac diseases in vitro. Multielectrode array (MEA) system is sometimes preferable to patch-clamp in electrophysiological experiments in terms of several advantages. Here we show our protocol of electrophysiological examinations using MEA. Key words Multielectrode array, Human-induced pluripotent stem cell-derived cardiomyocyte, Electrophysiology, Field potential duration, Extracellular recording
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Introduction Human cardiomyocytes differentiated from patient-specific– induced pluripotent stem cells (iPSCs) have been successfully used to model and study the cellular, molecular, and electrical features of several cardiac arrhythmias, including long QT syndrome (LQTS) [1–7], Brugada syndrome [8], and catecholaminergic polymorphic ventricular tachycardia [9, 10]. Furthermore, multiple drugs have been added to patient-specific human iPSCderived cardiomyocytes (hiPSC-CMs) to recapitulate the rescue of the pathological phenotypes [2, 9, 11–15]. More recently, screening platforms based on hiPSC-CMs have been developed, in response to the drug discovery [16–18]. Major technologies for the recording of the electrical activity of CMs include intracellular recordings using patch-clamp and extracellular recordings using multielectrode arrays (MEAs). Intracellular recording of action potentials (APs) is currently the standard method. However, the patch-clamp method is unsuitable for the drug development which requires qualified and high-throughput
Daisuke Yoshinaga and Yimin Wuriyanghai contributed equally to this work. Yoshinori Yoshida (ed.), Pluripotent Stem-Cell Derived Cardiomyocytes, Methods in Molecular Biology, vol. 2320, https://doi.org/10.1007/978-1-0716-1484-6_12, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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Fig. 1 The FPD has been shown to correlate with APD measurements. FPD field potential duration, APD action potential duration
technique. In contrast, extracellular recording of field potentials (FPs) by MEA provides a rapid and efficient approach to analysis of CM electrophysiology and reduced operator skill requirements [19, 20]. It also allows long-term chronic drug exposure, use of mixed cell populations, and analysis of drug impact on cell-to-cell transmission of electrical signaling. The cell culture dish for MEA has centralized electrodes embedded on the surface that can sense the electrical activity of the cells. FP duration (FPD) is defined as the time interval between the peak of the first spike in the waveform (Na+ influx during membrane depolarization) to that of the second deviation (K+ efflux during membrane repolarization) [19]. FPD is correlated with action potential duration (APD) in the intracellular recording, reflecting the QT interval in the electrocardiogram (Fig. 1).
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2.1 MEA Measurement
1. MED64-Basic, Alpha MED scientific, Osaka, Japan. 2. MED64 Main Amplifier (MED-A64MD1).
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3. MED64 Head Amplifier (MED-A64HE1S). 4. MED connector: MED-C03. 5. MEA probe: MED-P515A. 6. Mo¨bius QT software. 7. Mini-incubator. 2.2
Cell Preparation
1. iPSC-CMs (see Notes 1 and 2). 2. Matrigel/KnockOut DMEM: 2% Matrigel (Corning Life Sciences) in KnockOut DMEM (Thermo Fisher Scientific). 3. 2% FBS/DMEM F12: 2% FBS, 1% GlutaMAX (Thermo Fisher Scientific), 1% MEM Non-Essential Amino Acids Solution (Thermo Fisher Scientific), 0.1 mM 2-mercaptoethanol, and 0.5% penicillin/streptomycin in DMEM F12. 4. B-27 supplemented RPMI: 2% B-27 Supplement (Thermo Fisher Scientific) in RPMI. 5. Collagenase B: 1.0 mg/mL collagenase B (Merck) in DMEM F12. 6. ACCUMAX (Merck). 7. PBS () 1. 8. Sterile distilled water. 9. 10 cm cell culture dish or petri dish.
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3.1 Sterilization of MEA Probes When They Are Used for the First Time
1. Place an MEA probe in a 10 cm sterile dish.
3.2 Coating of MEA Probe
1. Add 20 mL sterile distilled water into the 10 cm sterile dish to prevent electrodes on the MEA probe from drying.
2. Add 1 mL 70% ethanol into the probe dish and leave the probe in a clean bench under UV radiation for at least 30 min. 3. Wash the probe three times with 1 mL water (see Note 3).
2. Put a drop of 2μL Matrigel/KnockOut DMEM solution onto the electrodes (see Note 4). Close the lid of the 10 cm dish and incubate at 37 C. 3. Move on to the Subheading 3.3 step. 3.3 Plating of iPSC-CMs 3.3.1 Embryoid Body (EB) (See Note 5)
1. Aspirate the Matrigel to the extent that a little moisture remains. 2. Transfer 1–4 beating EBs with 20μL culture medium onto the coating area. 3. Move to incubator slowly and incubate overnight. 4. Add additional 2 mL medium on the third day.
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3.3.2 Sheet
1. After removing cell culture medium, add 0.3 mL of collagenase B (1 mg/mL) per dish (if iPSC-CMs are cultivated in a 24 wellplate). 2. Incubate for 15 min at 37 C. 3. After removing collagenase B, then add 0.3 mL of ACCUMAX per dish. 4. Incubate for 5–10 min at 37 C. 5. Dissociate aggregates of iPSC-CMs gently, pipetting several times with a P1000 pipette. 6. Add 1.5–2.0 mL of 2% FBS medium and pipette several times with a pipette controller. 7. Collect cells in a round-bottom tube and centrifuge at 20 G for 5 min. 8. Aspirate the supernatant. 9. Add 1 mL of 2% FBS medium and transfer the cell suspension into a microtube. 10. Centrifuge at 20 G for 5 min. During this centrifugation, count the cells. 11. Aspirate the supernatant as much as possible and add 2% FBS to 1.5 104 cells/μL concentration. 12. Suction the Matrigel to the extent that some moisture remains. 13. Tap the microtube to homogenize the cell suspension and place 2μL (3.0 104 cells) on the electrode. 14. Check with microscope. 15. Incubate the MEA probe with overnight. 16. Add 1 mL of B27 supplemented RPMI. 17. Measure field potential of the iPSC-CMs 6–19 days after the plating (see Note 6).
3.4 Recording (See Note 7)
1. Filter: Sample FP signals digitally at 20 kHz through 0.1 Hz high-pass and 10 kHz low-pass filters. 2. Atmosphere condition: Incubate the recording probe in a mini-incubator, where the iPSC-CMs should be maintained at 37 C and under 5% CO2. Make sure to keep humidity. 3. Recording period: As long as cells are incubated in the environment described above, long time recording can be performed. For measurement of acute response, recording should be done 5–15 min after drug administration. For measurement under stable conditions, FPD should be measured 30–60 min after drug administration.
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4. Drug administration: Drugs should be gently administered by taking half of the medium out, diluting it, and then returning it to the solution (see Note 8). 3.5
Measurement
1. Definition of FPD and EAD: FPD was defined as the interval between a positive or negative spike and a subsequent positive deflection, and interspike interval (ISI) was defined as the interval between adjacent spikes (Fig. 2). These parameters were automatically measured and analyzed using Mo¨bius QT. More than 30 beats were recorded, and the FPDs and ISIs of the final 30 beats were averaged, as described previously [21]. EAD was defined as relatively slow negative spikes during the repolarizing phase (Fig. 3). EAD-positive samples were defined as showing more than 5 EADs among 30 beats. All the data should be acquired from at least three independent experiments. 2. Correction of FPD: The FPD measurements are generally normalized (corrected FPD [cFPD]) to the beating rate using the following
Fig. 2 Representative field potential waveforms. (a) Every field potential waveform acquired from 64 electrodes in an MEA probe. (b) Representative field potential waveform adequate for measurement of FPD. FPD field potential duration, ISI inter-spike interval
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Fig. 3 Representative waveforms of EADs. The arrowheads represent EADs. EAD early-after depolarization
Fig. 4 Correction of FPD using Fridericia’s formula. Uncorrected relationship between FPD and ISI (red) and corrected relationship between cFPD and ISI (blue) are shown. Among each cell line, dispersion of FPD depending on ISI was reduced by using the correction formula. FPD field potential duration, cFPD corrected field potential duration, ISI inter-spike interval
Fridericia’s correction formula: cFPD ¼ FPD/(RR interval)1/3 in order to minimize influence of a wide range of ISIs on FPDs (see Note 9) (Fig. 4).
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1. After measurement, add sterile distilled water into the probe dish. 2. Exchange the water with 300μL of sodium hypochlorite solution and leave the probe in the solution for 1–3 min. 3. Wash the probe three times with 1 mL water. 4. Change the 10 cm dish that contains the MEA probe. 5. Add 1 mL 70% ethanol into the probe dish and leave the probe in a clean bench under UV radiation for at least 30 min (see Note 10). 6. Wash the probe three times with 1 mL water. 7. Leave the probe until it is used for the next experiment (see Notes 11 and 12).
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Notes 1. Purity of iPSC-CMs should be monitored if possible because arrhythmogenicity could be influenced by the purity [22]. 2. The age of iPSC-CMs evaluated in experiments should be determined according to what is focused on. 3. MEA probes are not shipped in sterilized conditions. 4. Use a 1μL tip and be careful not to touch the electrodes to avoid damaging the electrodes. 5. Approximately 14 days after the cardiac differentiation starts, the EBs are once attached on a fibronectin coating dish. Isolate from dish with 27 G needle under the microscope after 6–7 days incubation and transfer to MEA dish. Manage the direction of EBs, and keep the flat side which was attached to the previous dish downward. 6. In many cases, at least 6 days are required to observe iPSC-CMs beating stably. 7. Before recording, make sure to manipulate the cells gently. The duration when the cells are outside incubators should be minimized. 8. It is recommended to keep drugs and P1000 pipettes at 37 C since beating rate and FPD are sensitive to temperature. 9. Several formulas other than Fridericia’s formula are published to correct FPDs [23]. However, definitive formula seems to remain undetermined. 10. UV radiation is indispensable in this step. 11. Be careful to keep the probes in a wet condition. 12. MEA probes can be repeatedly used for several times.
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Acknowledgment This work was supported by JSPS KAKENHI (Grant Number JP20K17147 to D.Y., JP19K17595 to Y.W., and JP16K09499, JP19K08538 to T.M.). References 1. Bellin M, Casini S, Davis RP et al (2013) Isogenic human pluripotent stem cell pairs reveal the role of a KCNH2 mutation in long-QT syndrome. EMBO J 32:3161–3175 2. Moretti A, Bellin M, Welling A, Jung CB, Lam JT, Bott-Flu¨gel L, Dorn T, Goedel A, Ho¨hnke C, Hofmann F, Seyfarth M (2010) Patient-specific induced pluripotent stem-cell models for long-QT syndrome. N Engl J Med 363(15):1397–1409 3. Yazawa M, Hsueh B, Jia X et al (2011) Using induced pluripotent stem cells to investigate cardiac phenotypes in Timothy syndrome. Nature 471:230–236 4. Limpitikul WB, Dick IE, Tester DJ et al (2017) A precision medicine approach to the Rescue of Function on malignant Calmodulinopathic long-QT syndrome. Circ Res 120:39–48 5. Gibson JK, Yue Y, Bronson J et al (2014) Human stem cell-derived cardiomyocytes detect drug-mediated changes in action potentials and ion currents. J Pharmacol Toxicol Methods 70:255–267 6. Peng S, Lacerda AE, Kirsch GE et al (2010) The action potential and comparative pharmacology of stem cell-derived human cardiomyocytes. J Pharmacol Toxicol Methods 61:277–286 7. Yamazaki K, Hihara T, Taniguchi T et al (2012) A novel method of selecting human embryonic stem cell-derived cardiomyocyte clusters for assessment of potential to influence QT interval. Toxicol In Vitro 26:335–342 8. Liang P, Sallam K, Wu H et al (2016) Patientspecific and genome-edited induced pluripotent stem cell–derived cardiomyocytes elucidate single-cell phenotype of Brugada syndrome. J Am Coll Cardiol 68:2086–2096 9. Jung CB, Moretti A, Mederos y Schnitzler M et al (2012) Dantrolene rescues arrhythmogenic RYR2 defect in a patient-specific stem cell model of catecholaminergic polymorphic ventricular tachycardia. EMBO Mol Med 4:180–191 10. Novak A, Barad L, Zeevi-Levin N et al (2012) Cardiomyocytes generated from CPVT D307H patients are arrhythmogenic in
response to β-adrenergic stimulation. J Cell Mol Med 16:468–482 11. Bellin M, Marchetto MC, Gage FH et al (2012) Induced pluripotent stem cells: the new patient? Nat Rev Mol Cell Biol 13:713–726 12. Sinnecker D, Laugwitz KL, Moretti A (2014) Induced pluripotent stem cell-derived cardiomyocytes for drug development and toxicity testing. Pharmacol Ther 143:246–252 13. Terrenoire C, Wang K, Chan Tung KW et al (2013) Induced pluripotent stem cells used to reveal drug actions in a long QT syndrome family with complex genetics. J Gen Physiol 141:61–72 14. Yoshinaga D, Baba S, Makiyama T et al (2019) Phenotype-based high-throughput classification of long QT syndrome subtypes using human induced pluripotent stem cells. Stem Cell Rep 13:394–404 15. Wuriyanghai Y, Makiyama T, Sasaki K et al (2018) Complex aberrant splicing in the induced pluripotent stem cell–derived cardiomyocytes from a patient with long QT syndrome carrying KCNQ1-A344Aspl mutation. Hear Rhythm 15:1566–1574 16. Abi-Gerges N, Pointon A, Oldman KL et al (2017) Assessment of extracellular field potential and Ca2+ transient signals for early QT/ pro-arrhythmia detection using human induced pluripotent stem cell-derived cardiomyocytes. J Pharmacol Toxicol Methods 83:1–15 ´ lamo JC, Lemons D, Serrano R et al 17. del A (2016) High throughput physiological screening of iPSC-derived cardiomyocytes for drug development. Biochim Biophys Acta, Mol Cell Res 1863:1717–1727 18. Blinova K, Stohlman J, Vicente J et al (2017) Comprehensive translational assessment of human-induced pluripotent stem cell derived cardiomyocytes for evaluating drug-induced arrhythmias. Toxicol Sci 155:234–247 19. Meyer T, Boven KH, Gu¨nther E et al (2004) Micro-electrode arrays in cardiac safety pharmacology: a novel tool to study QT interval prolongation. Drug Saf 27(11):763–772
MEA Assays Using hiPSC-CMs 20. Halbach MD, Egert U, Hescheler J et al (2003) Estimation of action potential changes from field potential recordings in multicellular mouse cardiac myocyte cultures. Cell Physiol Biochem 13:271–284 21. Asakura K, Hayashi S, Ojima A et al (2015) Improvement of acquisition and analysis methods in multi-electrode array experiments with {iPS} cell-derived cardiomyocytes. J Pharmacol Toxicol 75:17–26
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22. Kawatou M, Masumoto H, Fukushima H et al (2017) Modelling torsade de pointes arrhythmias in vitro in 3D human iPS cell-engineered heart tissue. Nat Commun 8:1–11 23. Izumi-Nakaseko H, Kanda Y, Nakamura Y et al (2017) Development of correction formula for field potential duration of human induced pluripotent stem cell-derived cardiomyocytes sheets. J Pharmacol Sci 135:44–50
Chapter 13 Electrophysiological Analysis of hiPSC-Derived Cardiomyocytes Using a Patch-Clamp Technique Yuta Yamamoto, Sayako Hirose, Yimin Wuriyanghai, Daisuke Yoshinaga, and Takeru Makiyama Abstract Electrophysiological analysis of human-induced pluripotent stem cell-derived cardiomyocytes (hiPSCCMs) using a patch-clamp technique enables the most precise evaluation of electrophysiological properties in single cells. Compared to multielectrode array (MEA) and membrane voltage imaging, patch-clamp recordings offer quantitative measurements of action potentials, and the relevant ionic currents which are essential for the research of disease modeling of inherited arrhythmias, safety pharmacology, and drug discovery using hiPSC-CMs. In this chapter, we describe the detail flow of patch-clamp recordings in hiPSC-CMs. Key words Human-induced pluripotent stem cell-derived cardiomyocyte, Patch-clamp technique, Electrophysiology, Action potential recording, Voltage clamp
1
Introduction The patch-clamp technique is a gold standard technique to record electrophysiological properties such as action potentials and ion currents in excitable cells including cardiomyocytes. This technique was developed by Erwin Neher and Bert Sakmann in 1970s who received the Nobel Prize in Physiology or Medicine in 1991 for this work [1]. This technology enables us to measure the membrane voltage or ion currents using glass pipette forming tight electrical gigaohm seals. To date, several methods related to patch-clamp technique have been developed. In this chapter, a whole-cell patch-clamp, the most popular method used in the analysis of hiPSC-CMs, is described below. hiPSC-CMs have currently been widely used for the research of heart diseases, safety pharmacology, and drug discovery because they have similar characteristics to human adult cardiomyocytes [2–8]. In spite of the great potentials of hiPSC-CMs, there are several concerns, especially, their immaturity of electrophysiological
Yoshinori Yoshida (ed.), Pluripotent Stem-Cell Derived Cardiomyocytes, Methods in Molecular Biology, vol. 2320, https://doi.org/10.1007/978-1-0716-1484-6_13, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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properties (e.g., spontaneous electrical activity and lack of the inward rectifier current, IK1). To overcome this issue, biochemical methods to mature hiPSC-CMs using chemicals, hormones, electrical, and/or mechanical stimulation, cocultures, long-term culture, etc., have been studied, and also, real-time electrical injection of calculated IK1 by “dynamic clamp system” was developed [9– 11]. In addition, various endogenous ion channels are expressed in hiPSC-CMs; therefore, the setting of voltage-clamp protocol and the use of ion channel blockers is important for recording the target ion currents in hiPSC-CMs. Here, we describe the protocol of the manual patch-clamp technique to record action potentials and major cardiac ion channel currents in hiPSC-CMs.
2
Materials
2.1 Equipment (Figs. 1 and 2)
1. Air table (NIKON INSTECH, NIT-86LA(TP)). 2. Faraday cage formed by a copper wire mesh. 3. Microscope (NIKON INSTECH, Ti2-U). 4. In-line heating system (CL-100 Single Channel Bipolar Temperature Controller (WARNER INSTRUMENT), Dual In-line Heater (Harvard Apparatus)). 5. Patch-clamp data acquisition system (Digidata 1440 (Molecular Devices)). 6. Patch-clamp Devices)).
amplifier
(MultiClamp
700B
(Molecular
7. Patch-clamp software (pCLAMP10 (Molecular Devices)). 8. Micromanipulator (uMp-Micromanipulator (Sensapex)). 9. Micropipette puller (P-1000 Micropipette Puller (Sutter Instruments)). 10. Borosilicate glass capillary. 11. 0.22μm pore size filter. 2.2
Reagents
1. 2% FBS/DMEM F12 culture medium: DMEM/Ham’s F-12, 500 mL, fetal bovine serum (FBS) 10 mL, GlutaMAX (Thermo Fisher Scientific) 5 mL, MEM Non-Essential Amino Acids Solution (10 mM) 5 mL, 2-mercaptoethanol (55 mM) 1 mL, penicillin/streptomycin solution (100) 2.5 mL. 2. 0.1% gelatin with PBS. 3. Matrigel Matrix Basement Membrane (CORNING). 4. Collagenase B (Roche). 5. Extracellular and intracellular solution (Tables 1 and 2).
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Rf
+
Output signal (V)
Vc Amplifier
Electrode
Rs R
C
Cardiomyocytes
Fig. 1 Schematic diagram of whole-cell patch-clamp system. C condenser, R seal resistance, Rs series resistance, Rf feedback resistance, Vc command voltage
Fig. 2 Basic setup for the patch-clamp experiment
3 3.1
Methods Cell Preparation
1. Before cell dissociation, glass coverslips are coated by 0.1% gelatin or 2% Matrigel in IMDM in each well of a six-well plate over 3 h at 37 C (see Note 1).
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Table 1 Intracellular pipette solution mM
AP
KCl
150
INa
TEA-Cl
Iks
IKr
20
150
20 70
CsCl2 NaCl
ICaL
5
CsAsp
120
5
5
40
CaCl2
2
2
2
EGTA
5
10
10
10
5
HEPES
10
10
5
5
10
MgATP
5
5
5
5
5
K-aspartate
125 3
MgCl2
1
Na2-phosphocreatine
2
Na2-GTP
2
Amphotericin B (μg/mL)
200–400
Adjusted to pH 7.2 with
KOH
CsOH
CsOH
KOH
KOH
2. hiPSC dissociation using collagenase B and trypsin–EDTA reagents. Aspirate the culture medium, and incubate the cardiomyocytes in 1 mL of 1 mg/mL collagenase B for 30 min at 37 C (1/6 well plate scale) (see Note 2). 3. Retrieve the cardiomyocytes with collagenase B into the tube and centrifuge at 120 g for 5 min. Remove the supernatant, and the pellet is resuspended in 1 mL trypsin–EDTA and incubated at 37 C (around 4 min). 4. After incubation, disperse into single cells by pipetting using P1000 around five times, and add the 10% FBS medium (see Note 3). 5. Centrifuge the tube, aspirate the supernatant, and resuspend the cells with 10% FBS medium. 6. The dissociated cardiomyocytes are placed into culture dishes containing gelatin-coated glass coverslips. Next day, change the medium to the 2% FBS culture medium. 7. Patch-clamp recordings are performed 3–5 days after dissociation.
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Table 2 Extracellular solution mM
AP
INa
NaCl
150
37
TEA-Cl
100
ICaL
Iks
IKr
150
150
5.4
5.4
140
KCl
5.4
CaCl2
1.8
1.8
1.8
1.8
1.8
MgCl2
1
2
1.2
1
1
5.4
CsCl2 Glucose
15
10
10
15
15
HEPES
15
10
5
15
15
Na-pyruvate
1
1
1
0.001
0.002
NiCl2
0.2
Nifedipine
0.005
E-4031
0.0005
Tetrodotoxin
0.01
4-Aminopyridine
1
Adjusted to pH 7.4 with
3.2 Pipette Preparation
3.3 Patch-Clamp Experiment 3.3.1 Current Clamp for Action Potential Recording
NaOH
CsOH
CsOH
NaOH
NaOH
Prepare glass pipette having a tapered shape and a resistance of 1.0–7.0 MΩ (depends on your purpose. See in each section) using pipette puller. After pulling, the tip of the pipette is polished by heating to smooth the rim of the tip (see Note 4). 1. Pull recording pipettes having resistance of 3.0–7.0 MΩ. 2. Load extracellular solution into the recording chamber using perfusion system at a rate of 1–2 mL/min and heat to 36 C 1 C using a heating system. 3. Set the coverslips on which cardiomyocytes are attached into the recording chamber. 4. To perform action potential recordings using perforated patchclamp technique, 1 mg amphotericin B is dissolved in 10μL DMSO and 2–4μL solution is added to 1 mL intracellular solution (see Note 5). 5. Dip the tip of the electrode into amphotericin B-free intracellular solution for several seconds. Backfill the recording electrode with intracellular solution containing amphotericin B and remove micro air bubbles in the tips by tapping gently. Set the microelectrode into the electrode holder.
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Fig. 3 Action potentials recorded from spontaneously beating hiPSC-CMs
6. Find a single cardiomyocyte on a coverslip. 7. Immerse the tip of electrode into extracellular solution in the recording chamber, and check pipette resistance using the membrane test protocol. Compensate for electrode offset. 8. Advance the tip of electrode towards the targeted cardiomyocyte carefully using micromanipulator. Before touching the cell, check the pipette resistance again and zero the pipette using pipette offset. The resistance will increase suddenly when the pipette touches the cell. 9. Using syringe suction, gently apply negative pressure through the electrode to the cell membrane to seal the cell while continuously monitoring the pipette resistance until the resistance approaches to ≧GΩ. Do not rupture the cell membrane by too much negative pressure. 10. Switch the amplifier setting from a voltage-clamp to a currentclamp mode with null current. Wait 5–10 min for perforation with monitoring the baseline (usually 60 to 70 mV). 11. An action potential waveform will become visible as perforation of the cell membrane (Fig. 3). 12. Switch the amplifier setting to a voltage-clamp mode and check the access resistance and cell capacitance using the membrane test protocol. The access resistance should be ≦30 MΩ. 13. If you want to use pacing protocol, use episodic stimulation in the acquisition mode of pCLAMP to apply 50–200 pA, 3 ms current pulses. Modify the duration or magnitude of stimulus to establish the stable pacing. 14. When you want to apply the extracellular solution containing a compound, set a constant flow of solution 3–5 min before you measure the action potential (see Note 6). 3.3.2 Voltage Clamp for Ion Current Recording
1. Prepare appropriate solution, glass pipette, and protocol for recording each ion current [5, 6, 8, 12]. 2. Follow the protocol stated above (Subheading 3.3.1, steps 1– 9). Ruptured patch-clamps are performed without amphotericin B (see Note 7).
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3. After formation of the “giga-seal” condition, switch to the membrane test protocol and set the membrane potential to 60 mV. Rupture the cell membrane by applying a negative pressure using gentle syringe suction. When the cell membrane is ruptured, you will identify the capacitive transient current that decays mono-exponentially. Apply a small negative pressure to maintain stable whole-cell condition (see Note 8). 4. After the rupture of the cell membrane, perform the compensation of whole-cell capacitance and series resistance. Turn on the whole-cell parameter. 5. Record the cell capacitance to calculate the current density in the Membrane Test Protocol. Also check the access resistance and membrane resistance which reflect the condition of sealing. 6. Set the holding potential appropriately. 7. Run the protocol for recording each ion current. The recording is started in approximately 5 min after the rupture of the cell membrane. 3.3.3 Analysis of Each Ion Channel Current
1. INa recording (Fig. 4): To record INa, calcium currents are blocked by nifedipine (L-type Ca2+ channel) and Ni2+ (T-type Ca2+ channel) included in the extracellular solution. A representative INa trace is shown in Fig. 4a. l
Pipette resistance: 1.0–2.0 MΩ.
l
Access resistance: 0.6). (c) Percentage of Q30, quality of base calling (see Note 10). (d) Density: lower than 280 103 clusters/mm2 (though Illumina recommends 170–250 103 clusters/mm2). (e) Total number of reads: approximately 550 million. 5. After sequencing, make the sample sheet (SampleSheet.csv) by using the Illumina experiment manager. 3.4
Demultiplexing
1. Sequencers generate a multiplex per-cycle base call file (bcl file) using RTA (illumine) software on the instruments as the primary data. “bcl2fastq” command (Illumina) converts the bcl file to a fastq file with demultiplexing.
bcl2fastq --runfolder-dir path_to_rawdata_directory #which includes bcl file --sample-sheet path/SampleSheet.csv --no-lane-splitting --output-dir path_to_output_directory
2. After execution, confirm the output files of each fastq file (. fastq.gz), Reports and Stats, in path_to_output_directory (see box above). 3. Open laneBarcode.html in path_to_output_directory/ Reports/html/(Flowcell_ID)/all/all/all/and check whether
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the used indexes (barcode sequence) show proper percentages of the lane, the perfect barcode, one mismatch barcode, and PF Clusters in Lane Summary. Also, check whether unexpected index sequences show many counts in Top Unknown Barcode in the html file. 3.5 Trimming of Adapter Sequences and Removal of Too Short Reads
1. To trim the adapter sequences from each read, we use cutadapt [1]. The “-m 20” option can discard multiple processed reads that are too short ( sample_name.cutadapt.stat
2. After execution, confirm the output files, sample_name.fastq. gz in path_to_output_directory (see box above) and sample_name.cutadapt.stat. To verify the samples, check the percentage of the too short reads and percentage of reads with adapter sequences by using the stat files. 3.6 Confirmation and Removal of Reads of rRNA Sequences
Even though TruSeq Stranded Total RNA Library Prep Gold involves rRNA depletion, confirm the percentage of reads that have been aligned to rRNA sequences. Remove these aligned reads for further analysis. 1. All reads in the fastq file are mapped to rRNA sequences by bowtie2 with index files (Subheading 2.7, item 2) [2] (see Note 11). 2. Convert the sam file into a bam file. 3. Perform samtools with the “flagstat” option to get the counts of each FLAG type of mapped reads [3] (see Note 12). 4. Reads that have not been mapped to an rRNA sequence are extracted by samtools with the “view -b -f” option. 5. Convert the unmapped bam file to a fastq file by the “bam2fastq” command.
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#1 bowtie2 --very-sensitive -x path/human_rRNAtRNA -U \ path/sample_name.fastq.gz -S sample_name.sam #2 sam -> bam samtools view -b sample_name.sam > sample_name.bam #3 flagstat all samtools flagstat sample_name.bam > sample_name. rRNA.stat #4 unmapped samtools view -b -f 4 sample_name.bam > sample_name. unmapped.bam #5 bam2fastq --no-aligned --unaligned --overwrite -o sample_name.fastq sample_name.unmapped.bam
6. Confirm the output files, sample_name.bam, sample_name. unmapped.bam, sample_name.rRNA.stat, and sample_name. fastq, and verify the percentage of reads which have been mapped to rRNA by a calculation using sample_name.rRNA. stat (see Note 13). 3.7 Mapping to the Human Genome
1. Using the unmapped reads of Subheading 3.6, use STAR to align the reads to the human genome rapidly with the index files (Subheading 2.7, item 5) [4]. This command uses default settings with basic options. In the ENCODE project, a few other options are used (see STAR manual (https://github. com/alexdobin/STAR/blob/master/doc/STARmanual. pdf)). 2. Perform samtools with the “flagstat” option to get the counts of FLAG types of all processed reads (Aligned.sortedByCoord. out.bam in path_to_output_directory (see box below)) (see Note 12). 3. Extract the unmapped reads with the samtools “view -b -f 4” option to check reads that have not been aligned to the human genome. 4. Perform samtools with the “flagstat” option to obtain the number of unmapped reads. 5. Extract the reads that have been mapped to the human genome uniquely (mapping quality MAPQ ¼ 255) as sample_name. q255.bam by samtools with the “view -b -q 255” option (see Note 14). These reads will be used for further analysis.
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6. Perform samtools with the “flagstat” option to obtain the number of uniquely mapped reads. 7. In order to see the mapping results with Integrated Genomics Viewer (IGV), perform samtools with the “index” option to generate bam index files (sample_name.q255.bam.bai) (see Note 15) [12–14].
#1 STAR --genomeDir path_to_STAR_index_directory \ --readFilesIn sample_name.fastq \ --outSAMtype BAM SortedByCoordinate \ --outSAMunmapped Within \ --runThreadN 12 \ --outFileNamePrefix path_to_output_directory/ INPUT= path_to_output_directory/Aligned.sortedByCoord.out.bam #2 flagstat all samtools flagstat ${INPUT} > sample_name.all.flagstat #3 Extracting unmapped reads samtools view -b -f 4 ${INPUT} > sample_name. unmapped.bam #4 flagstat unmapped samtools flagstat sample_nameunmapped.bam \ > sample_name.unmapped.flagstat #5 Extracting uniquely mapped reads samtools view -b -q 255 ${INPUT} > sample_name.q255. bam #6 flagstat uniquely mapped reads samtools flagstat sample_name.q255.bam > sample_name.q255.flagstat #7 Generation of index file for IGV samtools index sample_name.q255.bam
8. Verify the percentage of unmapped reads and percentage of uniquely mapped reads that can be calculated with flagstat files (see Note 16). 3.8 Quality Check by RSeQC
Using the bam files produced by the alignment tool, use the modules in RSeQC to evaluate the quality of the RNA-seq data with the reference files (Subheading 2.7, item 6). We introduce modules that we routinely use. For more details or other modules, please refer to the original manual [5].
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1. infer_experiment: The TruSeq stranded library kit is strandspecific. This module calculates strand specificity. The “-s” option can assign a number of reads sampled from the bam file. Set the “-s” option with a number to include all reads of the bam file (e.g., 60,000,000 in the box below). As a result, values should be high in “+,+,” which means that a read mapped to the “+” strand indicates the parental gene on the “” and vice versa. As a result, we usually observe that “+ +,” is about 0.01–0.02 and “+ . +” is over 0.88. If genomic DNA is not removed, this bias is small. 2. geneBody_coverage: This module shows read coverage over the whole gene body. If the input RNAs were degraded, coverage at the 50 end is lower than at the 30 end (e.g. Fig. 3d). 3. geneBody_coverage.house: This is the same as geneBody_coverage, except that this module only uses housekeeping genes for the calculation. 4. read_distribution: This module shows the distribution of reads of the genomic features (CDS_Exons, 50 UTR_Exons, 30 UTR_Exons, Introns, etc.). Usually “Intron” shows less than 0.27 tags/Kb/million for iPSC-CMs. If genomic DNA is not removed, this value is high. 5. read_duplication: To check whether the reads were duplicated in the PCR step of the library construction, compare the occurrence of the sequence-based and mapping-based reads. 6. read_GC: This module shows the distribution of reads regarding GC content. The median is approximately 50% for iPSCCMs. 7. read_NVC: This module shows the nucleotide composition bias. An approximately uniform value of 0.25 is expected for each nucleotide (in reality, C and G values are usually lower than A and T values), although this is not true near the 50 end of the reads. 8. read_quality: This module shows the distribution of the quality (Phred quality score, see Note 10) across reads. INPUT=sample_name.q255.bam SAMPLE_NAME=sample_name RSEQC_REF=path/hg38_Gencode_V28.bed.gz RSEQC_REF_HOUSE=path/hg38.HouseKeepingGenes.bed.gz OUTDIR=path_to_output_directory mkdir ${OUTDIR}
Fig. 3 RNA-seq using a small amount of RNA. Total RNA is extracted from the purified iPSC-derived cardiomyocytes on day 20 of the differentiation. No technical replicates. (a) DNA concentration of each library by Bioanalyzer analysis. (b) Percentage of reads whose insert lengths are short (< 20 bases) as calculated by Cutadapt analysis. (c) Percentage of reads mapped to rRNA by Bowtie2 analysis. (d) Percentage of uniquely mapped reads, unmapped reads, and other reads by STAR analysis. (e) Number of detected (count >0) genes by HTSeq analysis. Each values are calculated with the results of the workflow using randomly sampled 15 million reads after bcl2fastq. (f) Gene body coverage plot by RSeQC analysis
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mkdir ${OUTDIR}/infer_experiment OUTPUT= ${OUTDIR}/infer_experiment/${SAMPLE_NAME}. txt infer_experiment.py -r ${RSEQC_REF} -s 60000000 -i ${INPUT} > ${OUTPUT} mkdir ${OUTDIR}/geneBody_coverage OUTPUT= ${OUTDIR}/geneBody_coverage/${SAMPLE_NAME} geneBody_coverage.py -r ${RSEQC_REF} -i ${INPUT} -o ${OUTPUT} mkdir ${OUTDIR}/geneBody_coverage.house OUTPUT=${OUTDIR}/geneBody_coverage.house/${SAMPLE_NAME} geneBody_coverage.py -r ${RSEQC_REF_HOUSE} -i ${INPUT} -o ${OUTPUT} mkdir ${OUTDIR}/read_distribution OUTPUT= ${OUTDIR}/read_distribution/${SAMPLE_NAME}. txt read_distribution.py -r ${RSEQC_REF} -i ${INPUT} > ${OUTPUT} mkdir ${OUTDIR}/read_duplication OUTPUT=${OUTDIR}/read_duplication/${SAMPLE_NAME} read_duplication.py -i ${INPUT} -o ${OUTPUT} mkdir ${OUTDIR}/read_GC OUTPUT=${OUTDIR}/read_GC/${SAMPLE_NAME} read_GC.py -i ${INPUT} -o ${OUTPUT} mkdir ${OUTDIR}/read_NVC OUTPUT=${OUTDIR}/read_NVC/${SAMPLE_NAME} read_NVC.py -i ${INPUT} -o ${OUTPUT} mkdir ${OUTDIR}/read_quality OUTPUT=${OUTDIR}/read_quality/${SAMPLE_NAME} read_quality.py -i ${INPUT} -o ${OUTPUT}
3.9 Counting Reads for Each Transcript
1. To quantify gene expression levels from the bam file, use HTSeq to count the number of reads corresponding to each gene with the GTF file (Subheading 2.7, item 4) [6].
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htseq-count -s reverse -i gene_name -f bam sample_name.q255.bam \ path/gencode.v33.annotation.gtf > sample_name.txt
2. Confirm the output file with a matrix that includes columns of gene names and gene levels. 3.10
Normalization
Using a negative binomial generalized linear model, use DESeq2, which is a package of R, to normalize with “size factor” among all compared samples, estimate the dispersions of each gene, and conduct the Wald test and Likelihood ratio test (LRT) [7] (see Note 17). The Wald test can compare two groups, and LRT can be used for two groups or more. 1. For the DEG analysis, the sample information and the model design are needed (e.g., SampleFormula.csv). SampleFormula.csv. SampleName
FileName
Condition1
A1
File_name_A1.txt
A
A2
File_name_A2.txt
A
B1
File_name_B1.txt
B
B2
File_name_B2.txt
B
C1
File_name_C1.txt
C
C2
File_name_C2.txt
C
2. Import all HTSeq txt files and the SampleFormula.csv with the “DESeqDataSetFromHTSeqCount” commands. 3. Remove genes with no expression. 4. Perform the “DESeq” command to normalize and Wald tests on all combinations of two groups. For example, export the results of the test between conditions A and B as “DESeq2_res_wald_AvsB.txt.” This result file includes the gene name, mean value, log2 fold change, p-value, and adjusted p-value (FDR) (see Note 18). 5. For an example of multiple comparisons, compare conditions A, B, and C with the LRT, which uses two models: full model and reduced model. When there is only one condition, the reduced model is “1,” and the result file is the same as that from the Wald test.
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6. To export the data matrix of the normalized count of genes and samples as a text file, use the “counts” command. **The following commands are performed in R. library(DESeq2) library(tidyverse) #2 import data directory plot(hc5, cex = 0.6) # Cluster dendogram >
fviz_cluster(list(data
=
cormat,
cluster
=
sub_grp)) > gap_stat BiocManager::install("NOISeq") > library(NOISeq) BiocManager::install("org.Hs.eg.db") > library("org.Hs.eg.db") > library("AnnotationDbi") > ReadCountMatrix1 ReadCountMatrix2 nams = ReadCountMatrix2[,1] > rownames(ReadCountMatrix1) = make.names(nams, unique=TRUE) > mycounts = ReadCountMatrix1[,1:7] > myfactors = data.frame(hipscms = c("A1","A2", "A3", “B1”, "B2", "B3", "C1")) > mydata exprs_data myTMM = tmm(exprs_data) > head(assayData(mydata)$exprs) > meanexpression = rowMeans(myTMM) > threshold = quantile(meanexpression, 0.33) > dim(myTMM) > mydata mynoiseq.uqua = noiseq(mydata, k = 0.5, norm = "uqua", replicates = "technical", factor="hipscms", conditions = c("C","A"), pnr = 0.2, nss = 5, v = 0.02, lc = 1) > mynoiseq.deg = degenes(mynoiseq.uqua, q = 0.7, M = NULL) > mynoiseq.deg_up = degenes(mynoiseq.uqua, q = 0.7, M = "up") > mynoiseq.deg_down = degenes(mynoiseq.uqua, q = 0.7, M = "down")
9. Generation of expression plot: The following command will generate a plot to depict values of mean expression for each condition (Fig. 2e)
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highlighting those genes declared as differentially expressed (threshold of 70% referred as q ¼ 0.7). > DE.plot(mynoiseq.uqua, q = 0.7, graphic = "expr", log.scale = TRUE)
10. Heatmap generation to visualize differentially expressed genes (DEGs): We will generate a file named as gr containing a matrix reads of mynoiseq.deg. object from NOISeq functions, and we will generate a heatmap to visualize DEGs. With colorRampPalette function, we can customize displayed colors of the resulted heatmap.
> gr quantile.range palette.breaks
color.palette
heatmap(gr, col = color.palette)
11. Heatmap generation to visualize downregulated elements: By using only downregulated genes generated in the function mynoiseq.deg_down, we will generate a file named as gr2 containing a matrix reads in order to generate a heatmap to visualize downregulated elements in the defined condition (Fig. 2f).
> gr2 quantile.range heatmap(gr2, col = color.palette) > write.table(mynoiseq.deg_down,file=’C_vs_A_down. txt’,sep=’\t’,quote=FALSE, row.names=TRUE,col.names=NA)
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12. Heatmap generation to visualize upregulated elements: By using only upregulated genes generated in the function mynoiseq.deg_down, we will generate a file named as gr3 containing a matrix reads in order to generate a heatmap to visualize upregulated elements in the defined condition (Fig. 2g).
> gr3 quantile.range palette.breaks heatmap(gr3, col = color.palette) > write.table(mynoiseq.deg_up,file=’C_vs_A_up.txt’, sep=’\t’,quote=FALSE, row.names=TRUE,col.names=NA)
13. High-level analysis and GO enrichment: To review gene lists descriptions used here as general examples, please see Note 4. 14. Loading gene lists as R-derived readable matrix:
> WER WER2 gene.df ego2 dotplot(ego2, showCategory=30) > head(summary(ego2)) > geneList = sort(gene.df$ENTREZID, decreasing = TRUE)
We will create a new object called “ego3,” with the same command lines as before, but running a command to define enrichment for biological process (BP) category. Dotplot displaying results of these commands is shown in Fig. 3b. > ego3 dotplot(ego3, showCategory=30)
A final object called “ego4,” will run a command line to define enrichment for cellular process (CC) category. Dotplot displaying results of these commands is shown in Fig. 3c. > ego4 dotplot(ego4, showCategory=30)
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Fig. 3 Visualization of functional enrichment results (a) Visualization of molecular functions (MF) GO terms. Dot size remarks gene count and enrichment score ( p-value) is depicted by dot colors. (b) Visualization of biological processes (BP) GO terms. Dot size remarks gene count and enrichment score ( p-value) is depicted by dot colors. (c) Visualization of cellular processes (CC) GO terms. Dot size remarks gene count and enrichment score ( p-value) is depicted by dot colors
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Notes 1. M, D are statistics parameters of differential expression. The sign of M represents the direction of expression (overexpression or inhibition). prob remarks the probability of differential expression (not equivalent to p-value), hence the higher the prob value, the more likely the difference in expression is due to the change in the experimental condition and not occurred randomly. Ranking, statistic parameter qffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi ffi that synthesizes the values M and D to M M 2 þ D2 . 2. x, y, and z values must be adjusted according to the read count matrix of the user. 3. “ReadCountMatrix1.txt” contains 12 samples: "A1","A2", “B1”, "B2", "C1", "C2", “D1, “D2”, “E1”, “E2”, “F1” and "F2". Numbers next to greek letters indicate biological replicates. All samples allocate to human iPSC- derived cardiomyocytes. “ReadCountMatrix2.txt” contains gene name
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(symbol) in column 1. This pipeline extracts gene names column from “ReadCountMatrix2.txt” to create a file read by R Studio workflow shown here. 4. “WER.txt” is a plane text file containing gene symbol (names) of the target genes we want to enrich for gene ontology categories.
Acknowledgments This work was supported by a grant from Leducq foundation (18CVD05), JSPS KAKENHI Grants (number 20K17078 and 17H04176), grants from the Research Center Network for Realization of Regenerative Medicine (JP19bm0104001, JP19bm0204003, JP19bm0804008, and JP19bm0804004), Research on Regulatory Science of Pharmaceuticals and Medical Devices (JP19mk0104117), and Research Project for Practical Applications of Regenerative Medicine (JP19bk0104095) provided by the Japan Agency for Medical Research and Development, and iPS research fund. We greatly appreciate colleagues at our laboratory and we would like to express our heartfelt gratitude to Yoko Uematsu and Kaoru Shimizu for their administrative support. Declaration of Interests: The authors declare no potential conflicts of interest. Conflicting Interest: The authors declare no potential conflicts of interest. References 1. Burridge PW et al (2012) Production of de novo cardiomyocytes: human pluripotent stem cell differentiation and direct reprogramming. Cell Stem Cell 10(1):16–28 2. Mummery CL et al (2012) Differentiation of human embryonic stem cells and induced pluripotent stem cells to cardiomyocytes: a methods overview. Circ Res 111(3):344–358 3. Hatani T, Miki K, Yoshida Y (2018) Induction of human induced pluripotent stem cells to cardiomyocytes using embryoid bodies. Methods Mol Biol 1816:79–92 4. Tarazona S et al (2015) Data quality aware analysis of differential expression in RNA-seq with NOISeq R/Bioc package. Nucleic Acids Res 43(21):e140 5. Yu G et al (2012) clusterProfiler: an R package for comparing biological themes among gene clusters. OMICS 16(5):284–287
6. Mortazavi A et al (2008) Mapping and quantifying mammalian transcriptomes by RNA-Seq. Nat Methods 5(7):621–628 7. Muraoka N et al (2019) Role of cyclooxygenase-2-mediated prostaglandin E2-prostaglandin E receptor 4 signaling in cardiac reprogramming. Nat Commun 10(1):674 8. Ronaldson-Bouchard K et al (2018) Advanced maturation of human cardiac tissue grown from pluripotent stem cells. Nature 556 (7700):239–243 9. Ng SY, Wong CK, Tsang SY (2010) Differential gene expressions in atrial and ventricular myocytes: insights into the road of applying embryonic stem cell-derived cardiomyocytes for future therapies. Am J Physiol Cell Physiol 299(6):C1234–C1249 10. Zhao Y et al (2019) A platform for generation of chamber-specific cardiac tissues and disease modeling. Cell 176(4):913–927. e18
Part V Gene Editing and CRISPR Technology for Pluripotent Stem Cells
Chapter 21 Genome Editing in Human Induced Pluripotent Stem Cells (hiPSCs) Shuichiro Higo, Shungo Hikoso, Shigeru Miyagawa, and Yasushi Sakata Abstract Cardiomyocytes differentiated from human induced pluripotent stem cells (hiPSCs) are powerful tools for elucidating the pathology behind inherited cardiomyopathies. Genome editing technologies enable targeted genome replacement and the generation of isogenic hiPSCs, allowing investigators to precisely determine the roles of identified mutations. Here, we describe a protocol to obtain isogenic hiPSCs with the corrected allele via homology-directed repair (HDR) using CRISPR/Cas9 genome editing under feeder-free conditions. Seeding hiPSCs in a 24-well plate and conducting the initial evaluation using direct genomic sequencing after 1 week is cost- and time-effective. Following optimization of the protocol, sequence confirmation of the corrected HDR clone is completed within 21 days. Key words Human induced pluripotent stem cell, CRISPR/Cas9 genome editing, Cardiomyopathy, Homology-directed repair, Electroporation
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Introduction Cardiomyopathies are defined by structural and functional abnormalities of the ventricular myocardium in the absence of coronary artery disease or related conditions. Recent advances in high-throughput sequencing technologies have led to the discovery of a high incidence of mutations in hereditary cardiomyopathies [1–3]. hiPSC-derived cardiomyocytes (hiPSC-CMs) are powerful tools for elucidating the pathological behind inherited cardiomyopathies and helping expand disease modeling and drug discovery. Although numerous studies using hiPSC-CMs have clarified the pathological significance of many genetic variants [4], using hiPSCCMs from healthy subjects as control cannot completely exclude the effects of different genetic backgrounds. Recent advances in CRIPSR/Cas9 genome editing technologies have enabled targeted genome replacement in hiPSCs and allowed for functional evaluation of the pathological mutations against an isogenic background. Because reagents and drugs necessary to maintain hiPSCs are
Yoshinori Yoshida (ed.), Pluripotent Stem-Cell Derived Cardiomyocytes, Methods in Molecular Biology, vol. 2320, https://doi.org/10.1007/978-1-0716-1484-6_21, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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expensive and the procedure to isolate correctly edited hiPSC clones requires substantial time and laboratory equipment, the establishment of a simpler, optimized protocol is highly desired.
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Materials
2.1 Plasmid Construction
1. pX459 vector [5] (Addgene, cat # 48139). 2. pCR-Blunt-II-TOPO vector (ZeroBlunt TOPO PCR Cloning Kit, Thermo Fisher Scientific, cat # 450245). 3. EndoFree Plasmid Maxi Kit (QIAGEN, cat # 12362).
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Electroporation
1. NEPA21 Electroporator (Nepa Gene). 2. Cuvette Chamber (Nepa Gene, cat # CU500). 3. Cuvette Stand Holder (Nepa Gene, cat # CU600). 4. NEPA Electroporation Cuvettes, 2 mm gap (a pipette is included with each cuvette; Nepa Gene, cat # EC-002S). 5. Opti-MEM® Medium (Thermo Fisher Scientific, cat # 31985062).
2.3 hiPSC Culture and Antibiotics Selection
1. StemFit® AK02N (AJINOMOTO). 2. 10 μM of Y-27632 (ROCK inhibitor). 3. 0.5 mg/mL of i-Matrix 511 (nippi). 4. Cell scraper. 5. 0.4% (w/v) trypan blue solution. 6. 10 mg/mL puromycin dihydrochloride. 7. TrypLE Select (Thermo Fisher Scientific).
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1. KAPA Express Extract (Nippon Genetics, cat # KK7102). 2. KOD FX Neo (TOYOBO, cat # KFX-201). 3. QIAquick PCR purification kit (QIAGEN). 4. Nanodrop (Thermo Fisher Scientific).
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Methods Our protocol to obtain the edited hiPSC clone via homologydirected repair (HDR) is a multistep procedure using a small scale using 24-well plate (Fig. 1). The initial evaluation by direct Sanger sequencing is performed at day 6–7. After optimization of the protocol, sequence confirmation of the corrected clone is completed within 21 days.
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Fig. 1 Protocol overview of the genome editing in hiPSCs for selecting corrected clones with HDR 3.1 Design of gRNA and the HDR Template
Figure 2a shows a schema of the electropherogram of the Sanger sequence analysis obtained from a patient with cardiomyopathy harboring a heterozygous frameshift mutation in PKD1 (c.11,143 delC) [6]), as an example of the targeted mutation by genome editing. The double signals in the electropherogram are caused by a heterozygous frameshift mutation due to the deletion of C (delC). To correct the mutation (delC), the gRNA sequence can be designed to specifically recognize the mutated sequence if the targeted sequences at the 30 end of the gRNA are different from the wild type (WT) and mutation (Fig. 2a). The designed oligo DNA must be cloned into the pX459 vector encoding Streptococcus pyogenes Cas9 (SpCas9), then the T2A self-cleaving peptide followed by puromycin N-acetyltransferase will allow for puromycin resistance (see Note 1). To replace the targeted genome, the sequence around the mutation must be precisely cloned from human DNA into the cloning vector (pCR-bluntII-TOPO vector) to generate a repair template vector. A point mutation to avoid Cas9-mediated re-cleavage should be inserted in the repair template DNA sequence (Fig. 2b) (see Note 2). The generated vectors must be purified to remove endotoxins using EndoFree Plasmid Maxi Kit.
3.2 Electroporation into hiPSCs
A 24-well plate format can reduce the need for excessive use of reagents and the typical number of hiPSCs required for electroporation using the NEPA21 Electroporator. Usually, a cell suspension at 100 μL containing 1 106 hiPSCs with 10 μg plasmid vectors is required for electroporation [7, 8]; however, the amount of
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A
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CCTTCCTGGCCATC CCTTCTGGCCATCA gRNA PAM
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Fig. 2 (a) Illustration of the representative electropherogram obtained from the patient with dilated cardiomyopathy harboring a 11,143 delC mutation. One allele shows a normal sequence, the other shows the deletion of a C at position 11,143. Direct Sanger sequencing shows double signals for both alleles. (b) Design of the HDR repair template. A resistant mutation for avoiding Cas9-mediated re-cleavage is introduced into the repair template. The arrows indicate the sequence primers for detecting HDR. The forward or reverse primer must be designed outside of the homology arms
hiPSCs, medium, and plasmid vectors can be reduced in equal proportion to a 50 or 20 μL scale, using the same cuvette. 3.2.1 Electroporation (Day 0)
1. Grow hiPSCs to 80% or 90% confluency in a P3.5 dish. This will yield approximately 5 105–1 106 hiPSCs. 2. Precoat a 24-well plate and P3.5 dish (for passaging) with laminin (i-Matrix 511) 1 h before electroporation. 3. Wash the dish with PBS(). 4. Add 300 μL of 0.5 TrypLE Select and incubate at 37 C for 4 min. 5. Gently scrape the hiPSCs with a cell scraper, transfer to a tube, and pipet the cell suspension to dissolve the cells so they do not remain aggregated. 6. Count the cell number and passage 13,000 hiPSCs onto a P3.5 culture dish. 7. Transfer the suspension of hiPSCs into a 15 mL conical tube and centrifuge at 120 g for 5 min at room temperature. 8. Aspirate the supernatant, add 1 mL of Opti-MEM and pipet the hiPSCs gently but thoroughly, ensuring no clumps remain. Add 9 mL of Opti-MEM to yield a total of 10 mL. 9. Centrifuge at 120 g for 5 min and aspirate the supernatant. 10. Prepare a 24-well plate with a warmed StemFit medium (ABC) that contains 10 μM Y-27632. 11. Gently dissolve the hiPSCs with the estimated amount of OptiMEM (see below), ensuring no clumps remain.
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12. Add the plasmid vectors (pX459 vector and repair template vector), mix well, and transfer to a cuvette. The control plasmid vector expressing fluorescent protein (e.g., EGFP) is transfected to evaluate the transfection efficiency. The hiPSCs electroporated with the control vector will be used as the negative control for puromycin selection. 13. Electroporate the hiPSCs with the following protocol. Each parameter should be optimized according to the hiPSC lines. 14. Aspirate the electroporated hiPSCs from the cuvette using the attached dropper and transfer into the 24-well plate with the warmed medium. 15. Twenty-four hour after electroporation, evaluate the transfection efficiency using a fluorescence microscope. The ratio of normally attached hiPSCs after electroporation and the transfection efficiency of the incorporated plasmid vectors are some of the key identifiers for successful genome editing (see Note 3).
Poring pulse
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3.2.2 Puromycin Selection (Day 1~)
Because the transient expression of Cas9-linked puromycin N-acetyltransferase protein encoded by the pX459 vector gradually decreases according to cell proliferation, the addition of puromycin should not be delayed beyond 48 h after electroporation. Because the range of puromycin concentration for successful selection of the transfected hiPSCs is very narrow, optimization of the
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Fig. 3 A bright-field image (left) and fluorescent image (right) of the hiPSCs 24 h after electroporation. The expression vector encoding EGFP-fused nuclear protein (HP1) is electroporated. Transfection efficiency is estimated from the expression of the fluorescent protein. Scale bar: 50 μm
concentration of puromycin and the duration of drug treatment are recommended (see Notes 4 and 5). 1. Twenty-four hour after electroporation, confirm normal growth of the attached hiPSCs and the expression of EGFP in the control well (Fig. 3). 2. Exchange the medium with fresh, warmed medium and 0.3 μg/mL (requires optimization) puromycin. 3. Incubate the cells until the control cells transfected with the fluorescent protein are all dead and detached from the well. The incubation time with puromycin depends on the cell conditions (e.g., survival rate after electroporation, cell confluency). Refresh the medium with fresh medium containing puromycin every 24 h. Termination of puromycin treatment when surviving hiPSCs remain in the control well may result in low efficiency of genome editing. Usually, a 24–36 h puromycin selection is sufficient to select the transfected cells. 3.2.3 Initial Evaluation by Direct Sanger Sequencing (Day 6–7)
Incubate the electroporated hiPSCs until they reach approximately 50%–70% confluency in 24-well plate. 1. Six to seven days after electroporation, passage the hiPSCs into two P3.5 dishes. Gently mix the cell suspension and transfer the hiPSCs into a P3.5 dish, plating 250–500 cells per dish for colony isolation and 13,000 cells per dish for passaging. 2. Centrifuge the remaining cells and remove the supernatant, then add the KAPA genotype mix. Incubate the cells according to the manufacturer’s protocol.
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Direct sequence
Intact
NHEJ
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CCTTCCTGGCCATC CCTTCTGGCCATCA
CCTTCCTGGCCATC CCTCTGGCCATCAC
CCTTCCTGGCCATC CCTTCCTGGCCATC
Fig. 4 Illustration of the expected electropherogram after electroporation in hiPSCs. At the initial evaluation using direct sequencing, small signals from around 3 bp upstream of the PAM sequence indicate the result of induced NHEJ. The expected electropherograms after colony selection (Intact, NHEJ, and HDR) are shown. In the NHEJ clone, double signals consisting of bi-allelic sequences with an introduced mutation are observed. In the successful HDR clone, a single DNA sequence corresponding to the normal WT sequence is observed
3. The supernatant after incubation can be used for the PCR when using the KOD Fx Neo PCR mix. Usually, a 30 μL scale of PCR is sufficient for the following steps. The forward or reverse PCR primer must be designed outside the homology arms, so as not to amplify the DNA sequence of the repair template vector (Fig. 2b). Creating a forward primer near the targeted mutation (typically 100 bp upstream) is useful for direct sequence analysis. 4. Electrophorese 2–4 μL of the PCR products and confirm the distinct PCR bands at the expected molecular weight. 5. Purify the PCR products using the QIAquick PCR purification kit. Elute the PCR bands with ddW and measure the absorbance using a Nanodrop for direct Sanger sequencing with the forward PCR primer. 6. When successful genome editing has occurred, the small electropherogram signals can be observed from the estimated genomic region (approximately 3 bp upstream of the protospacer adjacent motif (PAM) sequence) due to the result of nonhomologous end joining (NHEJ) at the targeted site (Fig. 4). If the Sanger sequencing results show no signals of NHEJ, terminate the protocol at the initial evaluation and prepare for the next session (see Note 6).
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Fig. 5 The grown colonies are marked with a permanent marker under the microscope. The bright-field images of the hiPSC colony before (left) and after (right) pickup are shown. Scale bar: 1 mm 3.2.4 Colony Pickup (Day 12–14)
1. Precoat two 96-well plates with laminin (i-Matrix 511). Usually, 24 or more wells per one plate are prepared. 2. Select the isolated hiPSC colonies with a round-shape morphology and no differentiated cells, and mark them from the bottom of the dish using a permanent marker under the microscope (Fig. 5). 3. Prepare plastic tubes (according to the number of obtained colonies) with 100 μL StemFit medium containing 10 μM Y-27632. 4. Treat the hiPSCs with 0.5 TrypLE Select as the ordinary protocol for passaging. 5. Add 1 mL of StemFit containing 10 μM Y-27632. 6. Set the calibration of the micropipette to 20 μL. Place the pipette over the marked colony and aspirate the cells with a 20 μL medium. Transfer the cells into prepared tubes containing 100 μL of StemFit medium. 7. Pipette the cell suspension thoroughly, then transfer equal amounts of cells into the two 96-well plates. One plate will be for genotyping, and one plate is for passage of the hiPSCs.
3.2.5 Genotype the Isolated hiPSC Colonies (Day 21~)
Genotyping of the split hiPSC colonies in one of the 96-well plates should be conducted before the hiPSCs in the corresponding plate reach high confluency. hiPSCs with 20–30% confluency in a 96-well plate is sufficient for genotyping using the KAPA genotyping kit. 1. Aspirate the medium in the 96-well plate for genotyping. 2. Directly add the KAPA genotyping mix into the wells. 3. Pipette the cells with the KAPA genotyping mix and transfer them into eight strip tubes.
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4. Perform direct Sanger sequencing according to the protocol described above. 5. Results of the genome editing are judged according to the electropherogram for direct Sanger sequencing. The results are classified as Intact, NHEJ (different double signal waveform indicates insertion or substitution mediated by NHEJ), or HDR (Fig. 4). 6. Typically, within a 3-week timespan, HDR and NHEJ clones are obtained at a rate of 4.2–22.5%, respectively. After genotype confirmation, the corrected clone will be passaged for clonal expansion (see Notes 7 and 8).
4
Notes 1. Specific gRNAs can be designed using any online available tools [9, 10]. Generally, mismatches of gRNA are tolerated at the 50 end of the 20-nt gRNA sequence, compared to at the 30 end [11]. Preevaluation of cleavage activity of the designed gRNA using a single-strand annealing assay [12] or a Cel-I nuclease assay [13] using HEK293T cells is recommended before the experiment. 2. Genomic sequence of the target gene can be obtained from the UCSC genome browser (https://genome.ucsc.edu/). Typically, an HDR repair template with 600–800 bp 50 - and 30 -homology arms is desirable for a correct HDR. If the targeted sequence resides within an exonic region, the mutation must be designed for synonymous substitution in order to avoid an amino acid change. 3. Because survival of hiPSCs after electroporation is one of the most critical factors for successful genome editing, the parameters of electroporation, defined in the use of NEPA21 (e.g., voltage), should be optimized according to the hiPSC lines in order to improve survival rate and transfection efficiency. 4. The optimal concentration of puromycin and the optimal duration of puromycin treatment for selecting hiPSCs that have successfully incorporated the transgene after electroporation should be optimized according to the hiPSC lines. Cellular conditions before the procedure (e.g., proliferation rate) and the density of the surviving cells after electroporation both affect the results. 5. The conventional method for isolating the hiPSC clone with HDR uses the selection cassette containing antibiotic resistance genes in the repair template DNA, which requires the subsequent excision step. However, transfection of the pX459 plasmid encoding gRNA and the puromycin resistance gene-
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fused Cas9 combined with the repair template DNA without the selection cassette into hiPSCs also enables acquisition of the corrected hiPSC clones. These findings are consistent with a recent report noting that hiPSC clones with a correct HDR can be selected after transient expression of the puromycinresistant gene whose expression is linked to that of Cas9 [14]. 6. NHEJ signals in the electropherogram of the direct Sanger sequencing analysis at the initial evaluation will be subtle. To avoid noise in the electropherogram, the experimental conditions of genomic PCR analysis must be optimized (e.g., annealing temperatures, designing PCR primers) to obtain a clear single band without nonspecific PCR bands. 7. The pluripotency of the genome-edited hiPSCs after clonal selection must be maintained for further functional analysis. However, the multistep and long-time procedures for selecting hiPSC clones after electroporation may affect cellular pluripotency and increase the proportion of differentiated cells. The presence of stem cell-like morphology and contamination should be checked constantly, and hiPSCs can be prepared from frozen stock should either of these issues arise. 8. Although plasmid-based preparation of the repair template DNA as double-stranded DNA allows HDR with high efficiency in hiPSCs and homology arms longer than 500 bp are recommended for increasing the rate of HDR [7], single-stranded DNA with homology arms less than 200 bp, combined with a Cas9-expressing plasmid, promotes highly efficient HDR in hiPSCs [14].
Acknowledgments This work was supported by JSPS KAKENHI Grant Numbers 18K08069, 18K19543, 19K08489, 19K16518, grants-in-aid from the Japanese Ministry of Health, Labor, and Welfare, the Japan agency for medical research and development (19bm0804008h0003). This work was also supported by the Cell Science Research Foundation and the Grant for Basic Research of the Japanese Circulation Society (2018). Department of Medical Therapeutics for Heart Failure is an endowment department, supported by Actelion Pharmaceuticals Japan. We thank M. Moriyasu for technical assistance. This study was supported by Center of Medical Innovation and Translational Research, and Center for Medical Research and Education in Graduate School of Medicine, Osaka University.
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References 1. Herman DS, Lam L, Taylor MR, Wang L, Teekakirikul P, Christodoulou D, Conner L, DePalma SR, McDonough B, Sparks E, Teodorescu DL, Cirino AL, Banner NR, Pennell DJ, Graw S, Merlo M, Di Lenarda A, Sinagra G, Bos JM, Ackerman MJ, Mitchell RN, Murry CE, Lakdawala NK, Ho CY, Barton PJ, Cook SA, Mestroni L, Seidman JG, Seidman CE (2012) Truncations of titin causing dilated cardiomyopathy. N Engl J Med 366 (7):619–628 2. Haas J, Frese KS, Peil B, Kloos W, Keller A, Nietsch R, Feng Z, Muller S, Kayvanpour E, Vogel B, Sedaghat-Hamedani F, Lim WK, Zhao X, Fradkin D, Kohler D, Fischer S, Franke J, Marquart S, Barb I, Li DT, Amr A, Ehlermann P, Mereles D, Weis T, Hassel S, Kremer A, King V, Wirsz E, Isnard R, Komajda M, Serio A, Grasso M, Syrris P, Wicks E, Plagnol V, Lopes L, Gadgaard T, Eiskjaer H, Jorgensen M, Garcia-Giustiniani D, Ortiz-Genga M, Crespo-Leiro MG, Deprez RH, Christiaans I, van Rijsingen IA, Wilde AA, Waldenstrom A, Bolognesi M, Bellazzi R, Morner S, Bermejo JL, Monserrat L, Villard E, Mogensen J, Pinto YM, Charron P, Elliott P, Arbustini E, Katus HA, Meder B (2015) Atlas of the clinical genetics of human dilated cardiomyopathy. Eur Heart J 36(18):1123–1135a 3. Ware JS, Seidman JG, Arany Z (2016) Shared genetic predisposition in Peripartum and dilated cardiomyopathies. N Engl J Med 374 (26):2601–2602 4. Sayed N, Liu C, Wu JC (2016) Translation of human-induced pluripotent stem cells: from clinical trial in a dish to precision medicine. J Am Coll Cardiol 67(18):2161–2176 5. Ran FA, Hsu PD, Wright J, Agarwala V, Scott DA, Zhang F (2013) Genome engineering using the CRISPR-Cas9 system. Nat Protoc 8 (11):2281–2308 6. Suwa Y, Higo S, Nakamoto K, Sera F, Kunimatsu S, Masumura Y, Kanzaki M, Mizote I, Mizuno H, Fujio Y, Hikoso S, Sakata Y (2019) Old-age onset progressive cardiac
contractile dysfunction in a patient with polycystic kidney disease harboring a PKD1 Frameshift mutation. Int Heart J 60(1):220–225 7. Sharma A, Toepfer CN, Ward T, Wasson L, Agarwal R, Conner DA, Hu JH, Seidman CE (2018) CRISPR/Cas9-mediated fluorescent tagging of endogenous proteins in human pluripotent stem cells. Curr Protoc Hum Genet 96:21.11.1–21.11.20 8. Li HL, Fujimoto N, Sasakawa N, Shirai S, Ohkame T, Sakuma T, Tanaka M, Amano N, Watanabe A, Sakurai H, Yamamoto T, Yamanaka S, Hotta A (2015) Precise correction of the dystrophin gene in duchenne muscular dystrophy patient induced pluripotent stem cells by TALEN and CRISPR-Cas9. Stem Cell Rep 4(1):143–154 9. Heigwer F, Kerr G, Boutros M (2014) E-CRISP: fast CRISPR target site identification. Nat Methods 11(2):122–123 10. Doench JG, Fusi N, Sullender M, Hegde M, Vaimberg EW, Donovan KF, Smith I, Tothova Z, Wilen C, Orchard R, Virgin HW, Listgarten J, Root DE (2016) Optimized sgRNA design to maximize activity and minimize off-target effects of CRISPR-Cas9. Nat Biotechnol 34(2):184–191 11. Sander JD, Joung JK (2014) CRISPR-Cas systems for editing, regulating and targeting genomes. Nat Biotechnol 32(4):347–355 12. Mashiko D, Fujihara Y, Satouh Y, Miyata H, Isotani A, Ikawa M (2013) Generation of mutant mice by pronuclear injection of circular plasmid expressing Cas9 and single guided RNA. Sci Rep 3:3355 13. Qiu P, Shandilya H, D’Alessio JM, O’Connor K, Durocher J, Gerard GF (2004) Mutation detection using surveyor nuclease. BioTechniques 36(4):702–707 14. Steyer B, Bu Q, Cory E, Jiang K, Duong S, Sinha D, Steltzer S, Gamm D, Chang Q, Saha K (2018) Scarless genome editing of human pluripotent stem cells via transient Puromycin selection. Stem Cell Rep 10(2):642–654
Chapter 22 Generation of Efficient Knock-in Mouse and Human Pluripotent Stem Cells Using CRISPR-Cas9 Tatsuya Anzai, Hiromasa Hara, Nawin Chanthra, Taketaro Sadahiro, Masaki Ieda, Yutaka Hanazono, and Hideki Uosaki Abstract A knock-in can generate fluorescent or Cre-reporter under the control of an endogenous promoter. It also generates knock-out or tagged-protein with fluorescent protein and short tags for tracking and purification. Recent advances in genome editing with clustered regularly interspaced short palindromic repeat (CRISPR) and CRISPR-associated protein 9 (Cas9) significantly increased the efficiencies of making knock-in cells. Here we describe the detailed protocols of generating knock-in mouse and human pluripotent stem cells (PSCs) by electroporation and lipofection, respectively. Key words Genome editing, CRISPR-Cas9, Pluripotent stem cells, Homology-directed repair, Knock-in
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Introduction Recent progresses in genome editing open a new era of modifying a genome in mammalian cells [1–3]. Previously, most of experiments used transgenic lines rather than knock-in because of low probability to obtain proper knock-in lines. Due to silencing and position effects of insertion sites [4], there are huge line-to-line variations and a limited number of cell-type–specific promoters that produce reproducible results in different labs. In terms of reliability and stability, knock-in is more preferable. However, knock-in lines were rarely used because a lot of effort and time were required and only 1% or less had successfully knocked-in alleles even with drug selection [5]. CRISPR-Cas9 and other nucleases such as zinc-finger nucleases (ZFNs) and transcription activator-like effector nucleases (TALENs) can cause a double-strand break (DSB) to a specific site
Tatsuya Anzai and Hiromasa Hara contributed equally to this work. Yoshinori Yoshida (ed.), Pluripotent Stem-Cell Derived Cardiomyocytes, Methods in Molecular Biology, vol. 2320, https://doi.org/10.1007/978-1-0716-1484-6_22, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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of choice. Because a DSB can be repaired by homologous recombination and an exogenous knock-in vector serves as a repair template, usage of CRISPR-Cas9 increases knock-in efficiencies dramatically [6]. However, based on our own experiences and reports from other labs, knock-in efficiencies to most of genes without drug selection seem to be around 1% or less [7, 8] (see Note 1). Because of high demands on making knock-in pluripotent stem cells (PSCs) for cell-type–specific gene expression, such as reporter system and gene-tagging, we describe here the details of our method for knock-in, achieving up to 90% efficiency (approximately 50% efficiency in average depending on the targets), to mouse embryonic stem cells (ESCs) and human induced pluripotent stem cells (iPSCs), which uses a drug selection cassette and subsequent removal of it before cloning.
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Materials
2.1 Mouse Embryonic Stem Cell Culture
1. Mouse ESCs: Sub-strains from E14tg2a [9] and mouse ESCs newly derived from C57BL/6 mice (from Dr. Hirofumi Nishizono, Max Planck Florida Institute for Neuroscience). 2. 2i Medium: 450 mL of Mix Glasgow’s Minimum Essential Medium (SIGMA G5154), 50 mL (final 10%) of fetal bovine serum (FBS; FBS requires lot tests prior to use to maintain mouse ESCs, heat inactivated), 5 mL (final 1 mM) of sodium pyruvate (100 mM), 5 mL of MEM-NEAA (GIBCO 11140050, 100), 5 mL of GlutaMax (GIBCO 35050-061, 100), 909 μL (final 0.1 mM) of 2-mercaptoethanol (GIBCO 219850-023, 55 mM); 50 μL (final 1000 U/mL) of ESGRO/mLIF (Millipore ESG1107—1 107 U/mL), or 500 μL of StemSure mouse LIF (Wako 195-16,053, 1 106 U/mL), 150 μL (final 3 μM) of CHIR99021 (stock 10 mM), and 100 μL (final 1 μM) of PD0325901 (stock 5 mM). Sterilize the mixture with a 0.22-μm filter. 3. 0.1% gelatin: EmbryoMax (Millipore), or prepare from gelatin powder (SIGMA G1890) in distilled water and autoclave the solution. 4. A freezing container: Mr. Frosty (Thermo Fisher), Bicell (Nihon Freezer), CoolCell (Wakebtech), or any alternates to slowly cooldown (1 C/min) the tubes in 80 C freezer.
2.2 Human Induced Pluripotent Stem Cell Culture
1. Human iPSCs: 201B6 [10], 610B1 [11], 648A1 [12], and BC1 [13]. 2. Maintenance Medium: AK02N (Ajinomoto). 3. iMatrix-511 (Matrixome).
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4. Rock Inhibitor, Y-27632 (Wako, STEMCELL Tech or other companies. Stock 10 mM in PBS). 5. TransIT-LT1 (Mirus). 2.3
Electroporation
1. Electroporator: NEPA21 (Nepa Gene). 2. Cuvette (NEPA Cuvette, 2 mm gap; EC-002S). 3. OPTI-MEM.
2.4 Cas9/ gRNA-Expressing Plasmid
1. px330 (Addgene, 422309), px458 (48138), or px459 (62988) (Fig. 1a). 2. BbsI-HF (New England BioLabs). 3. 10 H buffer (Takara) or equivalent solution consisting 500 mM Tris–HCl [pH 7.5], 100 mM MgCl2, 10 mM dithiothreitol, and 1000 mM NaCl. 4. Ligation high ver.2 (Toyobo). 5. DH5α chemically competent E. coli. 6. An oligo pair to an intended target: top, 50 -CACCG+20-nt guide sequence without PAM-30 and bottom, 50 -AAAC+20 nt reverse complemental guide sequence without PAM+C-30 .
2.5 Knock-in Construct and RecombinaseExpressing Plasmid
1. Circular plasmid vector with 500–1000 bp homology arm on both sides of a reporter gene such as green fluorescent protein (GFP) and red fluorescent protein (RFP, we recommend mCherry for combination with GFP and mScarlet for the strongest brightness; Fig. 1b). The reporter gene must be in-frame to the target gene and a polypeptide linker such as Gly-Gly-Gly-Gly-Ser between the target and the reporter gene is recommended. A drug selection cassette with excision recombination sites (FRT or loxP) is also recommended. As loxP sites are frequently used to knockout a gene conditionally, FRT (50 -GAAGTTCCTATTCtctagaaaGtATAGGAACTTC-30 ) is more preferable for this purpose. 2. pCAGGS-flpE-Puro (addgene #20733) or equivalent plasmidexpressing flippase (or Flp recombinase) to remove the drug selection cassette.
2.6 Antibiotics for Drug Selection
1. Blasticidin (10 mg/mL stock, working concentration is around 5–20 μg/mL). 2. Puromycin (1 mg/mL stock, working concentration is 1–2 μg/mL). 3. Hygromycin B (100 mg/mL stock, working concentration is 50–200 μg/mL).
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A: px330 and its family gRNA cloning site gRNA scaffold
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Fig. 1 Typical Cas9/sgRNA-expressing vector and knock-in construct design. (a) Structure of Cas9- and gRNA-expressing vector: px330 and its family encode spCas9 under the control of strong ubiquitous promoter CBh and sgRNA under the control of Pol III promoter U6. After BbsI (or BpiI) digestion, gRNA oligo can be cloned. (b) Design of a knock-in construct: A knock-in construct consists of 500–1000 bp homology arms on both sides, a fluorescent protein or gene of interest, and FRT-ubiquitously expressing drug-resistant gene-FRT. The first step is to deliver the knock-in construct and px330 vector encoding gRNA to PSCs. After selection with blasticidin, flippase is introduced to excise out the drug-resistant cassette 2.7
Software
1. Web-based services or commercial software to design specific gRNA are recommended to design gRNAs: e.g., Benchling (https://benchling.com/), CCTop (http://crispr.cos.uniheidelberg.de) [14], CRISPRdirect (http://crispr.dbcls.jp) [15], and Geneious (Biomatters Ltd).
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2. To design genotyping primer, primer-blast (https://www.ncbi. nlm.nih.gov/tools/primer-blast/) is one of the wellestablished web-based services.
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Methods
3.1 Construction of Cas9/ gRNA-Expressing Plasmid
To construct a Cas9/gRNA-expression plasmid, detailed method is described in the previous protocol paper by Dr. Feng Zhang’s group [16]. 1. To design gRNA, there are good web-based services and commercial software are available: e.g., Benchling (https:// benchling.com/), CCTop (http://crispr.cos.uniheidelberg.de) [14], CRISPRdirect (http://crispr.dbcls.jp) [15], and Geneious (Biomatters Ltd). To knockout a gene, gRNA is designed at a first few exons (preferably around the start codon or around a most important domain for the protein function). To make a reporter or a fusion protein, the last exon and right before the stop codon is more preferable. However, to design more specific gRNA is even more important (see Note 2). 2. Synthesize an oligo pair as follows: top, 50 -CACCG+20-nt guide sequence without PAM-30 and bottom, 50 -AAAC +20 nt reverse complemental guide sequence without PAM +C-30 and resolve at 100 μM with Tris–EDTA (pH 8.0) solution (Fig. 1a). 3. Mix 1 μL of each oligo, 16 μL of ultra-pure water, and 2 μL of 10 H buffer. 4. Anneal the mixture by denaturing at 95 C for 5 min and cooling down to 20 C with slow ramp (0.1 C/s). Then, dilute the annealed oligo with 180 μL of Tris–EDTA (pH 8.0) solution. 5. To prepare a plasmid vector backbone, digest px330 plasmid with BbsI for 1 h and purify by gel extraction following gel electrophoresis. 6. To ligate the oligo into plasmid backbone, mix 0.5 μL (10–50 ng) of digested px330, 0.5 μL of diluted-annealed oligo, 1 μL of ligation high ver.2; incubate at 16 C for 30 min, then transform DH5α chemically competent E. coli with the ligated plasmid. 7. Confirm sequence and prepare enough plasmid with midi-prep (we prepare at 1 μg/μL).
3.2 Construction of Knock-In Vectors
1. Obtain homology arms by PCR or synthetic DNA fragment (Integrated DNA Technologies). Homology arms are 500–1000 bp for both sides of the CRISPR-Cas9 target site.
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To note, removing CRISPR-Cas9 target site from the homology arms is important; otherwise, CRISPR-Cas9 would cut the knock-in construct or redigest the genome after knock-in occurs (see Note 3). For the efficient construction, add a multi-cloning site between the homology arms. 2. Clone the homology arms to cloning plasmid such as pBS and confirm the sequence. 3. Insert reporter genes such as GFP and mCherry to the multicloning site. Keep reading frame intact, so that reporter gene can be expressed properly as a fusion protein. 4. To improve knock-in efficiencies, a drug-resistant cassette can also be added. We typically use BamHI-linker-mChery-FRTSV40-Blast-pA-FRT-ClaI cassette (Fig. 1b). By our hands, adding drug selection cassette increased the insert size, but the knock-in efficiency itself remained similar (data note shown). 5. Prepare the plasmid at 1 μg/μL. 3.3 Design of Genotyping Primers
3.4 Knock-in to Mouse ESCs 3.4.1 Gene Targeting with CRISPR-Cas9
To test if a knock-in occurs, genotyping primer sets are critical. We recommend primer-blast (https://www.ncbi.nlm.nih.gov/tools/ primer-blast/) to design genotyping primer sets outside of the homology arms. We generally design two pairs of primer sets and test all four combinations to determine the most specific combination with good amplification. Amplicon with this primer pair can also be used to prepare homology arms. 1. Plasmid preparation: Mix 10 μL of OPTI-MEM, 2 μg (2 μL) of CRISPR-Cas9–expressing plasmid, and 8 μg (8 μL) of knock-in construct. Plasmid ratio 1:3 (2.5 μg of Cas9 plasmid and 7.5 μg of knock-in construct) also works, and in some cases, plasmid ratio 3:1 works better. 2. Plate preparation: Coat a well of a 6-well plate with 1 mL of 0.1% gelatin. 3. Mouse ESCs are cultured as previously described [17]. Harvest mouse ESCs and count cell numbers: Aspirate culture media from a plate and wash the cells with PBS. Add 1 mL of TrypLE for a 6-cm dish and incubate it for 3 min at 37 C. Resuspend the cells with 4 mL of 10% FBS-containing medium to neutralize trypsin. Count cell numbers. 4. Transfer 1.5–3.0 105 cells to a new tube and centrifuge the cells down at 300 g for 3 min. This knock-in method works with up to 1.0 106 cells but efficiency could be lower. 5. Resuspend the cells with 80 μL of OPTI-MEM and mix the cells with the plasmid prepared in Subheading 3.4.1, step 1 (20 μL). Then, transfer all of 100 μL to an electroporation
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cuvette. Make sure not to produce air bubbles that adversely affect electroporation. 6. Set parameters of NEPA21 as follows: Poring pulse 175 V, pulse length 2 ms, pulse interval 50 ms, pulse number four times, decay rate 10%, and polarity +; transfer pulse 20 V, pulse length 50 ms, pulse interval 50 ms, pulse number five times, decay rate 40%, and polarity +/. Place the cuvette in the electroporation chamber and check resistance of cuvettes (35–45 Ω, a large variation from a usual value implicates inaccuracy in the amounts of buffer, cells, or DNA) (see Note 4). 7. Apply the pulse to the electroporation cuvette. 8. Aspirate 0.1% gelatin from the plate and add 2 mL of 2i medium. 9. Use a pipette provided with the cuvette to transfer approximately 1 mL medium from the plate to the cuvette. Suspend the cells gently in the cuvette and transfer them back to the plate. 10. Culture the cells overnight after the electroporation and change medium. If px459 (Cas9-Puro) is used, a short puromycin selection can be performed between day 1 and 2 [18]. 11. Start drug selection from as early as day 3 after electroporation. Starting it later is better for specificity as nonintegrated plasmid will disappear from the cells. 12. Continue drug selectin at least 3 days for blasticidin or until all control cells are killed. 3.4.2 Removal of Drug-Resistant Gene
1. Plasmid preparation: Mix 10 μL of OPTI-MEM and 10 μg (10 μL) of pCAG-flpE-puro. 2. Plate preparation: Coat a well of a 6-well plate with 1 mL of 0.1% gelatin. 3. Harvest the drug-resistant cells, centrifuge, and resuspend the cells with 80 μL of OPTI-MEM. 4. Mix the cells and plasmid, and transfer the mixture to an electroporation cuvette. 5. Electroporate the cells with the same conditions as described in Subheading 3.4.1. 6. Replate the cells to two wells of a 6-well plate. 7. Culture the cells overnight. 8. Perform drug selection (puromycin) for 1 day. Treat only one well in case cells cannot tolerate to the treatment with puromycin. 9. Culture for 2–3 days until colonies grow.
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10. Replate 1000–10,000 cells to a 10-cm dish and change medium every other day until colonies grow. The remaining cells can be plated to a well of a 6-well plate, frozen and/or subjected to DNA purification. 3.4.3 Cloning
1. Once cells plated in Subheading 3.4.2, step 10, grow (typically takes 1 week or so), colonies are ready for cloning by colony picking. 2. To clone colonies, set up a microscope in a culture hood. Prepare 25–50 μL of TrypLE per a well in 24–96 wells of a 96-well plate depending on the number of colonies to be picked up. Coat 24-well plates with 0.1% gelatin. 3. Pick up a colony with a P10 pipette (set to 10 μL) by scraping the colony off from the dish and aspirating it into the pipette. Transfer the colony into a well of the 96-well plate with TrypLE. 4. After 6–24 colonies are picked up and transferred to the 96-well plate, place the 96-well plate in to a 37 C incubator for 1 min. 5. Replace 0.1% gelatin to culture medium (0.5 mL/well). 6. Triturate the colonies to make sure each colony is dissociated and then transfer all solution to a well of the 24-well plate with culture medium. 7. Culture the cells until each clone becomes sub-confluent (typically ~1 week). Refresh medium every other day.
3.4.4 Making Stock and Genotyping
1. Prepare freezing medium with 120 μL of DMSO, 120 μL of FBS, and 760 μL of IMDM per a clone and load the medium into each of 2-mL cryotubes. Label the cryotubes and new 1.5mL tube. For a clone, one cryotube with freezing medium and a 1.5-mL tube are needed. 2. Wash cells with PBS, then add 200 μL of TrypLE and incubate the culture plate at 37 C for 3 min. 3. Triturate cells to dissociate, transfer the cells in TrypLE in a well to one cryotube with the freezing medium, mix well, and transfer 200 μL to a new 1.5-mL tube. 4. Close the lids and freeze the cryotubes using a freezing container. 5. Centrifuge the 1.5-mL tubes to pellet down the cells for DNA purification. 6. DNA purification can be done with any method using purification columns (e.g., DNeasy) or simple lysis buffers (e.g., Phire Animal Tissue Direct PCR Kit).
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7. PCR and sequence to identify properly knocked-in clones. Knock-in efficiency varies from 16% to 83% depending on the targets or gRNA (50% in average for five different targets we have generated in a previous report [19, 20] and unpublished data by Anzai: all of the target genes are not expressed in mouse ESCs) (see Note 5). 3.5 Knock-in to Human iPSCs 3.5.1 Standard Human iPSC Culture
1. Human iPSCs (201B6, 610B1, 648A1, and BC1) are cultured in 6-well plates with AK02N according to the previously reported method with some modifications [21]. 2. Wash cells with PBS, add 500 μL of TrypLE to each well, and incubate the cells at 37 C for 3 min. Collect cells in 2 mL of DMEM with 10% FBS and count cell numbers. 3. Centrifuge 75,000–150,000 cells depending on cell lines and culture schedule (we plate 75,000 cells/w for 4-day culture before the next passage and 150,000 cells for 3 days) at 300 g, 3 min. 4. Resuspend cells in 2 mL of AK02N supplemented with 10 μM of Y-27632 and 2 μL of iMatrix-511 and plate the cells to a well of a 6-well plate. 5. Culture the cells overnight and refresh medium to AK02N on the next day. Medium needs to be changed every other day.
3.5.2 Lipofection
1. Prepare lipofection reagents as follows: (A) 10 μL of OPTIMEM and 0.3 μL of TransIT-LT1, and (B) 2 μL of px330gRNA (10 ng/μL) and 8 μL of knock-in vector (10 ng/μL). Mix A and B well by pipetting at least 20 times (see Note 4). 2. Harvest cells as explained in Subheading 3.5.1 and centrifuge 20,000 cells per a well of a 24-well plate. 3. Resuspend cells with culture medium (500 μL of AK02N supplemented with 10 μM of Y-27632 and 0.5 μL of iMatrix) and plate cells to a well of a 24-well plate. 4. Add the lipofection reagents prepared in step 1 to the cells. 5. Culture the cells overnight and change medium on the following day. 6. Passage and expand cells from a well of a 24-well plate to a well of a 6-well plate at around day 4 (see Note 6). 7. Between day 5 and 7 or 8, add blasticidin to the culture to select knock-in cells.
3.5.3 Removal of Drug-Resistant Gene
1. Once cells grow, transfect 100 ng of pCAG-flpE-puro to 20,000 cells as same as in Subheading 3.5.2, step 1 to exclude the drug-resistant gene. 2. Treat cells with puromycin from day 1 to 2.
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3. Passage cells from a well of 24-well to a 6-cm or 10-cm dish when cells grow. 3.5.4 Cloning
1. Prepare culture medium (for a clone, 500 μL of AK02N, 10 μM of Y27632, and 0.5 μL of iMatrix). Split 450 μL of the medium to a well of a 24-well plate and 50 μL to a well of 96-well plate. 2. Pick up a colony into a well of a 96-well plate using a P10 pipette, and triturate each colony well to breakdown it to small clumps using a P200 pipette. Then, transfer the clumps to a well of the 24-well plate and culture the cells overnight followed by refreshing medium to AK02N. 3. Following Subheading 3.2, step 4 to stock and genotype cells. We anticipate similar knock-in efficiencies as mouse ESCs based on our unpublished data (see Note 4).
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Notes 1. Since this is a rapidly evolving field, newer and more efficient methods can be introduced. For example, adeno-associated virus (AAV)-mediated methods would work very efficiently for PSCs as well as for zygotes and other cells such as hematopoietic stem cells [22, 23]. In our preliminary results, AAV could enhance knock-in efficiency up to 40% without drug selection. 2. When Cas9 and gRNA are expressed from plasmid vectors, they tend to be retained longer in the cells compared to Cas9 ribonucleoprotein (RNP). Thus, specificity of gRNA is very important. We use multiple software and compare the results to obtain the best gRNA as each software provide different results. 3. In case of leaving CRISPR-Cas9 target site to keep protein sequence intact (e.g., generation of fusion protein with knock-in to 30 side of the target site), the CRISPR-Cas9 target site needs to be removed by codon optimization. 4. To achieve the effective knock-in condition, it is important to optimize conditions for each laboratory, especially when different types of electroporators are used, by targeting some expressing genes in undifferentiated stage. Figure 2 depicts examples of knock-in to gene loci that are expressed in mouse ES (Actb) and human iPSCs (LMNB1). In these conditions, we used 1-kb homology arms and achieved 5–10% of knock-in efficiencies to these genes without drug selection. 5. For some targets, knock-in efficiency was lower. It could be due to the weaker activities of gRNA.
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Fig. 2 Representative results of knock-in to mouse ESCs and human iPSCs. (a) Knock-in of GFP to β-actin locus of mouse ESCs: After knock-in, cytoplasmic localization of GFP was observed, indicating GFP was fused with β-actin. Knock-in efficiencies were 5–10%. (b) Knock-in of GFP to LAMNB1 locus of human iPSCs: After knock-in, GFP exclusively localized to nuclear membrane of human iPSCs, indicating GFP was fused to LMNB1, a membrane protein. Knock-in efficiencies were 5–10%
6. We occasionally observed massive cell death a few days after transfection. In such cases, we performed transfection in a larger scale (e.g., a 6-well) or delayed blasticidin treatment.
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Acknowledgments We would like to thank all the lab members in the Division of Regenerative Medicine, Jichi Medical University for technical supports and helpful discussions. The methods described here were developed with the funding supports from The Program for Technological Innovation of Regenerative Medicine, Research Center Network for Realization of Regenerative Medicine from Japan Agency for Medical Research and Development (AMED, JP18bm0704012), Fund for the Promotion of Joint International Research (Fostering Joint International Research (B), JP19KK0219) from Japan Society for the Promotion of Science, and the Grant for Basic Research of the Japanese Circulation Society (to HU). This study was also supported in part by the Basic Science and Platform Technology Program for Innovative Biological Medicine from AMED (JP18am0301002 to Y.H. as co-PI). References 1. Mali P, Yang L, Esvelt KM et al (2013) RNA-guided human genome engineering via Cas9. Science 339:823–826. https://doi.org/ 10.1126/science.1232033 2. Cong L, Ran FA, Cox D et al (2013) Multiplex genome engineering using CRISPR/Cas systems. Science 339:819–823. https://doi.org/ 10.1126/science.1231143 3. Jinek M, East A, Cheng A et al (2013) RNA-programmed genome editing in human cells. elife 2:e00471. https://doi.org/10. 7554/eLife.00471 4. Strathdee D, Ibbotson H, Grant SGN (2006) Expression of transgenes targeted to the Gt (ROSA)26Sor locus is orientation dependent. PLoS One 1:e4. https://doi.org/10.1371/ journal.pone.0000004 5. Reid LH, Shesely EG, Kim HS, Smithies O (1991) Cotransformation and gene targeting in mouse embryonic stem cells. Mol Cell Biol 11:2769–2777. https://doi.org/10.1128/ mcb.11.5.2769 6. Zhang J-P, Li X-L, Li G-H et al (2017) Efficient precise knockin with a double cut HDR donor after CRISPR/Cas9-mediated doublestranded DNA cleavage. Genome Biol 18:35. https://doi.org/10.1186/s13059-017-11648 7. Roberts B, Haupt A, Tucker A et al (2017) Systematic gene tagging using CRISPR/Cas9 in human stem cells to illuminate cell organization. Mol Biol Cell 28:2854–2874. https:// doi.org/10.1091/mbc.E17-03-0209
8. Roberts B, Hendershott MC, Arakaki J et al (2019) Fluorescent gene tagging of transcriptionally silent genes in hiPSCs. Stem Cell Rep 12:1145–1158. https://doi.org/10.1016/j. stemcr.2019.03.001 9. Hooper M, Hardy K, Handyside A et al (1987) HPRT-deficient (Lesch-Nyhan) mouse embryos derived from germline colonization by cultured cells. Nature 326:292–295. https://doi.org/10.1038/326292a0 10. Takahashi K, Tanabe K, Ohnuki M et al (2007) Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell 131:861–872. https://doi.org/10.1016/j. cell.2007.11.019 11. Kawada J, Kaneda S, Kirihara T et al (2017) Generation of a motor nerve Organoid with human stem cell-derived neurons. Stem Cell Rep 9:1441–1449. https://doi.org/10. 1016/j.stemcr.2017.09.021 12. Okita K, Yamakawa T, Matsumura Y et al (2013) An efficient nonviral method to generate integration-free human-induced pluripotent stem cells from cord blood and peripheral blood cells. Stem Cells 31:458–466. https:// doi.org/10.1002/stem.1293 13. Chou B-K, Mali P, Huang X et al (2011) Efficient human iPS cell derivation by a non-integrating plasmid from blood cells with unique epigenetic and gene expression signatures. Cell Res 21:518–529. https://doi.org/ 10.1038/cr.2011.12
Generation of Efficient Knock-In Pluripotent Stem Cells 14. Stemmer M, Thumberger T, del Sol KM et al (2015) CCTop: an intuitive, flexible and reliable CRISPR/Cas9 target prediction tool. PLoS One 10:e0124633. https://doi.org/10. 1371/journal.pone.0124633 15. Naito Y, Hino K, Bono H, Ui-Tei K (2015) CRISPRdirect: software for designing CRISPR/Cas guide RNA with reduced off-target sites. Bioinformatics 31:1120–1123. https://doi.org/10.1093/bio informatics/btu743 16. Ran FA, Hsu PD, Wright J et al (2013) Genome engineering using the CRISPR-Cas9 system. Nat Protoc 8:2281–2308. https://doi. org/10.1038/nprot.2013.143 17. Uosaki H, Patrick C, Lee DI et al (2015) Transcriptional landscape of Cardiomyocyte maturation. Cell Rep 13:1705–1716. https://doi. org/10.1016/j.celrep.2015.10.032 18. Sluch VM, Chamling X, Wenger C et al (2018) Highly efficient scarless knock-in of reporter genes into human and mouse pluripotent stem cells via transient antibiotic selection. PLoS One 13:e0201683. https://doi.org/10. 1371/journal.pone.0201683 19. Sadahiro T, Isomi M, Muraoka N et al (2018) Tbx6 induces nascent mesoderm from
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pluripotent stem cells and temporally controls cardiac versus somite lineage diversification. Cell Stem Cell 23:382–395.e5. https://doi. org/10.1016/j.stem.2018.07.001 20. Chanthra N, ABE T, Miyamoto M et al (2020) A novel fluorescent reporter system identifies laminin-511/521 as potent regulators of Cardiomyocyte maturation. Sci Rep 10(1):4249. https://doi.org/10.1038/s41598-02061163-3 21. Miyazaki T, Isobe T, Nakatsuji N, Suemori H (2017) Efficient adhesion culture of human pluripotent stem cells using laminin fragments in an uncoated manner. Sci Rep 7:41165. https://doi.org/10.1038/srep41165 22. Mizuno N, Mizutani E, Sato H et al (2018) Intra-embryo gene cassette knockin by CRISPR/Cas9-mediated genome editing with adeno-associated viral vector. iScience 9:286–297. https://doi.org/10.1016/j.isci. 2018.10.030 23. Bak RO, Dever DP, Reinisch A et al (2017) Multiplexed genetic engineering of human hematopoietic stem and progenitor cells using CRISPR/Cas9 and AAV6. elife 6:18. https:// doi.org/10.7554/eLife.27873
Chapter 23 CRISPRi/a Screening with Human iPSCs Masataka Nishiga, Lei S. Qi, and Joseph C. Wu Abstract Identifying causative genes in a given phenotype or disease model is important for biological discovery and drug development. The recent development of the CRISPR/Cas9 system has enabled unbiased and largescale genetic perturbation screens to identify causative genes by knocking out many genes in parallel and selecting cells with desired phenotype of interest. However, compared to cancer cell lines, human somatic cells including cardiomyocytes (CMs), neuron cells, and endothelial cells are not easy targets of CRISPR screens because CRISPR screens require a large number of isogenic cells to be cultured and thus primary cells from patients are not ideal. The combination of CRISPR screens with induced pluripotent stem cell (iPSC) technology would be a powerful tool to identify causative genes and pathways because iPSCs can be expanded easily and differentiated to any cell type in principle. Here we describe a robust protocol for CRISPR screening using human iPSCs. Because each screening is different and needs to be customized depending on the cell types and phenotypes of interest, we show an example of CRISPR knockdown screening using CRISPRi system to identify essential genes to differentiate iPSCs to CMs. Key words Genome editing, CRISPR/Cas9, Induced pluripotent stem cells, Cardiomyocytes
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Introduction Although recent development in next-generation sequencing (NGS) has made it much faster and easier to study genome-wide gene expression changes and epigenetic changes even on a singlecell level, it is not straightforward to study causality of a biological phenotype from the large datasets because they are mostly consequences of upstream changes [1–3]. Thus, identifying causative genes and pathways for biological discovery and drug development is still a time-consuming process. To accelerate identification of causative genes and pathways in biological phenotypes of interest, systematic and high-throughput genetic perturbation approaches have been done using chemical DNA mutagens or shRNA libraries. However, these methods had still limitations of efficiency and accuracy [1–3]. Recent CRISPR/Cas9-based systems enabled more efficient and accurate gene knockout screening even in a genome-wide scale [3–7] (Fig. 1). Transcriptional inhibition or
Yoshinori Yoshida (ed.), Pluripotent Stem-Cell Derived Cardiomyocytes, Methods in Molecular Biology, vol. 2320, https://doi.org/10.1007/978-1-0716-1484-6_23, © Springer Science+Business Media, LLC, part of Springer Nature 2021
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Fig. 1 CRISPR screening in combination with iPSC technology. Identifying causative genes is important for biological discovery and drug development but has been time-consuming. On the other hand, forward genetic screening with pooled CRISPR libraries enables unbiased identification of causative genes even in genomewide scale by knocking out (or knocking down or activating) many genes at once and selecting cells with a phenotype of interest. Thus, CRISPR screening in combination with iPSC technology, which can make any cell type in principle, is a powerful approach to identify causative genes of diverse phenotypes in human somatic cells
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activation screening is also available by using nuclease-deactivated Cas9 (dCas9) [1, 2, 8, 9]. These CRISPR/Cas9-based screening methods have been used mainly in cancer cells to identify essential genes for proliferation or resistance to anticancer drugs [1–3, 7, 9] (Fig. 2a). Although forward genetic screening with loss-of-function or gain-of-function is a powerful approach to identify causative genes, it is not easy to apply CRISPR screening to human somatic cells because CRISPR screening requires a large number of cells to be cultured. Typically, the number of cells needs to be more than 1000-fold of the number of gRNAs in a systematic, high-coverage screen to avoid the uncovered bias (statistically, small sample size leads to a high variability) [1]. For example, for genome-wide libraries which contain 100,000 gRNAs, assuming that the transfection efficiency of the gRNA library is 30%, more than 300 million cells need to be cultured. Thus, human primary cells are not ideal for large-scale screening. On the other hand, human iPSCs are a good platform to perform CRISPR screening because iPSCs are expandable and can be differentiated to any cell type in principle [10, 11] (Fig. 1). Here we describe an example protocol for CRISPR screening with human iPSCs. It is difficult to generalize the protocol of CRISPR screening with iPSCs because each screening is unique and different depending on the purposes, cell types of interest, phenotypes of interest, and types of genome editors (Cas9, dCas9, base editors, etc) [1]. In this chapter, we show an example protocol of CRISPRi screening to identify essential genes during CM (cardiomyocyte) differentiation. This protocol is modifiable to fit many other applications (Fig. 2).
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Materials
2.1 CRISPRi/a Library
1. Human Genome-wide CRISPRi-v2 Libraries or your custom library (Addgene, #83969) or your custom sgRNA library: Follow the protocols in Addgene #83969. 2. Escherichia coli MegaX DH10B T1R Electrocomp Cells (Thermo Fisher Scientific, C640003). 3. Escherichia coli Stellar competent cells (Clontech, 636763). 4. Escherichia coli DH5a competent cells (e.g., Zymo Research, T3007). 5. Large plate for library amplification (e.g., Fisher Scientific, 12-565-224). 6. LB Agar. 7. LB medium. 8. Ampicillin. 9. Plasmid Midi prep kit. 10. Plasmid Maxi or Giga prep kit.
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Fig. 2 Unbiased genetic screening using CRISPR sgRNA libraries. (a) CRISPR screening can detect causative genes in a phenotype of interest by comparing the distributions of sgRNAs. After each cell is labeled by an sgRNA using pooled lentiviral library, the cells with a phenotype of interest are selected. Then, the distribution of sgRNAs in each population can be measured by next-generation sequencing. (b) We show an example schedule of CRISPRi screening with iPSC-CMs that identifies essential genes for CM differentiation. The CRISPRi iPSCs were infected with an sgRNA library and underwent puromycin selection. Then, iPSCs were differentiated to CMs. To screen essential genes for CM differentiation, CMs were stained by TNNT2 and sorted by flow cytometry. The volcano plot shows multiple hit genes and a positive control (TNNT2) as essential genes for CM differentiation 2.2 CRISPRi/a Plasmids
1. sgRNA backbone plasmid (pU6-sgRNA EF1Alpha-puro-T2ABFP) (Addgene, #60955). 2. Lentiviral CRISPRi plasmid KRAB) (Addgene, #85969).
(UCOE-SFFV-dCas9-BFP-
3. Lentiviral CRISPRa plasmid (pHRdSV40-dCas910xGCN4_v4-P2A-BFP) (Addgene, #60903). 4. Lentiviral CRISPRa plasmid (pHRdSV40-scFv-GCN4-sfGFPVP64-GB1-NLS) (Addgene #60904). 5. BxtXI (Thermo Fisher Scientific, FD1024). 6. BlpI (New England Biolabs, R0585S).
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2.3 Lentivirus Production
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1. psPAX2 plasmid (Addgene #12260). 2. pMD2.G plasmid (Addgene #12259). 3. HEK 293T cells (Clontech, Lenti-X™ 293T Cell Line, 632180). 4. PEI MAX (Polysciences, Linear Polyethylenimine Hydrochloride MW 40K, 24765-1). 5. Polybrene solution: 10 mg/mL polybrene in sterile water. 6. ViralBoost (ALSTEM, VB100). 7. DMEM high glucose. 8. Fetal bovine serum (FBS). 9. Penicillin–streptomycin (10,000 U/mL of penicillin and 10,000 μg/mL of streptomycin). 10. 0.45-μm PES syringe filter. 11. Vacuum filtration system (e.g., Millipore, S2GVU02RE). 12. 10-cm cell culture dish. 13. T225 cell culture flask.
2.4
iPSC Culture
1. CRISPRi iPSC line: If you want to make a new CRISPRi or CRISPRa line from iPSCs you have, follow the protocol provided on Addgene (Addgene, #83969). 2. DMEM/F-12. 3. Essential 8 medium (Thermo Fisher Scientific, A1517001). 4. RPMI 1640. 5. RPMI 1640 no glucose. 6. Matrigel Matrix Basement Membrane (Corning, 356231). 7. Y-27632 2HCl (ROCK inhibitor) (Selleck Chemicals, S1049). 8. CHIR-99021 (Selleck Chemicals, S2924). 9. IWR-1 (Selleck Chemicals, S7086). 10. Puromycin (Thermo Fisher Scientific, A1113803). 11. EDTA. 12. B-27 Supplement, minus insulin (Thermo Fisher Scientific, A1895601). 13. B-27 Supplement (50), serum free (Thermo Fisher Scientific, 17504044). 14. 15-cm cell culture dish.
2.5 Genomic DNA Extraction
1. NK lysis buffer (50 mM Tris, 50 mM EDTA, 1% SDS, pH 8). 2. Proteinase K (Qiagen, 19131). 3. RNaseA (Qiagen, 19101). 4. Ammonium acetate (Sigma-Aldrich, A1542).
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5. Isopropanol. 6. Ethanol, molecular biology grade. 7. 1 TE solution pH 8.0. 8. NanoDrop (Thermo Fisher Scientific). 2.6 NGS Library Prep and Sequencing
1. KAPA HiFi HotStart DNA Polymerase with 5 Fidelity Buffer (Roche Diagnostics, KK2502). 2. PCR Thermal Cyclers. 3. QIAquick PCR Purification Kit (Qiagen, 28106). 4. 3 M sodium acetate. 5. Agencourt AMPure XP (Beckman Coulter, A63881). 6. DNA LoBind tube (Fisher Scientific, 13-698-791). 7. Access to Bioanalyzer (High sensitivity DNA chip). 8. Access to Illumina NextSeq (or HiSeq).
2.7
Data Analysis
1. ScreenProcessing (GitHub, https://github.com/mhorlbeck/ ScreenProcessing). 2. Access to Python 3.7 (or 2.7).
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Methods Here we show an example protocol for CRISPRi screening with human iPSCs. It can be modified to other types of screening including activation screening (see Note 1). Make sure to start with optimization in a small-scale experiment of 2–3 individual sgRNAs instead of directly proceeding to a screening that requires a large number of cells (see Note 2).
3.1 Prepare a CRISPRi iPSC Line 3.1.1 Prepare a CRISPRi iPSC Line and Optimize Culture Condition
Prepare a CRISPRi iPSC line which stably expresses dCas9-KRAB. To generate a new CRISPRi iPSC line by lentivirus (transgenic) or genome editing (knock-in), follow the protocol provided by Weissman lab on Addgene [8, 9, 12]. Here we use an inducible dCas9KRAB knock-in iPSC line generated by TALEN-assisted gene-trap approach [8]. Once you have a CRISPRi iPSC line, optimize culture condition and differentiation protocol. We use Essential 8 (E8) medium on Matrigel-coated (1:200 dilution) 6-well plates to maintain iPSCs. For passaging of iPSCs, we detach iPSCs from the plates with 0.5 mM EDTA and culture them in E8 medium with 10 μM Y-27632 ROCK inhibitor (E8 + Y medium) for 24 h. To differentiate CMs from this iPSC line, we use 8 μM CHIR99021 in RPMI with B27 minus insulin supplement for 48 h and then 5 μM IWR-1.
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It is important to find an appropriate concentration of puromycin using your iPSC lines. For screening, too low concentration of puromycin may cause too many sgRNA-negative cells in your samples. Too high concentration may increase the percentage of cells that have two or more sgRNAs. If you plan to do drug selection after differentiation instead of doing selection at the stem cell stage, you need to test the concentration in your cell type. 1. Day 0: Plate iPSCs in a Matrigel-coated 24-well plate (40,000 cells/well) in E8 + Y medium. Culture cells overnight. 2. Day 1: Start puromycin. Try different concentration of puromycin (0.1–5 μM) in E8 medium (400 μL/well). 3. Continue puromycin selection for 3 days. Replace medium every day. 4. Day 4: Evaluate the confluency of cells. Find the lowest concentration that killed all the iPSCs.
3.2 Test Small-Scale Transduction and Lentiviral Infection
3.2.1 Small-Scale Lentiviral Production
Before starting a large-scale screening, confirm the knockdown efficiency in the cell type you will use in the screening afterwards because cell types affect chromatin accessibility of genome editors (e.g., dCas9-KRAB). Even if the knockdown efficiency is high in stem cell state, it does not guarantee a high efficiency in your cell type after differentiation. Small-scale experiments to evaluate the knockdown efficiency in your desired cell type targeting 2 or 3 genes are highly recommended. 1. Prepare lentiviral plasmids that expresses sgRNAs (backbone: Addgene#60955). You need a non-targeting sgRNA in addition to sgRNAs that target your gene of interest (e.g., control sgRNA, sgRNA#1, and sgRNA#2). To design sgRNAs, use CRISPRiaDesign on GitHub or pick up the sequences from the Human Genome-wide CRISPRi-v2 Library (Addgene#83969) [12]. 2. Culture HEK293T cells in DMEM 10% FBS (DMEM GlutaMax with 10% FBS and penicillin/streptomycin). On the day before transfection (Day 0), plate 800,000 cells per well in a 6-well plate (Table 1). 3. Day 1: For each sgRNA, prepare a lentiviral plasmid mixture and PEI MAX mixture (A and B below) and wait for 5 min. Mix A and B and incubate for 20 min in room temperature. Then gently drop the mixture to HEK293T cells in one well of a 6-well plate. Return the plate to the incubator and culture cells for overnight.
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Table 1 Small-scale lentivirus production (6-well plate) Day 0 Prepare HEK293T l
Plate HEK293T cells in a 6-well plate (800 K cells/well for each sgRNA)
Day 1 PEI transfection
Day 2 Check BFP
Day 4 Collect virus
Replace medium (1.5 mL, DMEM 10% FBS) l Prepare PEI mixture l Incubate 20 min l Gently drop into each well
l
Check BFP Replace medium (2 mL, E8 medium) l Incubate for 48 h
l
l
l
Collect supernatants l Filter supernatants l Make aliquots and freeze them in 80 C
Components
For each sgRNA
A
DMEM (serum free) psPAX2 pMD2G Lenti sgRNA plasmid
250 μL 1.8 μg 0.6 μg 1.8 μg
B
DMEM (serum free) PEI MAX
250 μL 8.4 μL
4. Day 2: Check BFP (blue fluorescent protein) on a fluorescent microscope to confirm PEI transfection is successful. Replace the medium with 2 mL of E8 medium per well. Culture cells for 48 h. 5. Day 4: Collect supernatant from each well. Filter the virus supernatant using 5-ml syringes with 0.45-μm (or 0.22-μm) syringe filters. Freeze the filtered supernatant in 80 C. (Optional) Using E8 medium on Day 2 as basal medium for virus production enables you to skip the virus concentration step and avoid exposing iPSCs to components in 293T medium (e.g., FBS). The functional titer of the supernatant virus is normally high enough without concentration. If you need higher titer and purity, you can concentrate virus with concentration reagents (e.g., Lenti-X Concentrator, Clontech). 3.2.2 Small-Scale Lentiviral Infection of iPSCs
1. Culture iPSCs on a Matrigel-coated 6-well plate until they reach 80% confluency and then proceed to lentiviral infection (Table 2). 2. Day 0 (iPSC plating and lentivirus infection): Detach iPSCs with 0.5 mM EDTA and suspend them in E8 + Y medium in the same way with regular iPSC passaging. Plate iPSCs to a new Matrigel-coated 6-well plate (300,000 cells/1.5 mL/ well ¼ 30,000/cm2, one well for each sgRNA). If you have three sgRNAs, prepare three wells (e.g., control sgRNA,
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Table 2 Small-scale lentiviral infection of iPSCs (6-well plate) Day 0 iPS plating and Lenti infection
Day 1
Plate iPSCs (300 K/well) Add polybrene (final 8 μg/mL) l Add lentivirus 500 μL l Incubate for 3–4 h l Replace medium (E8 + Y medium)
l
l
l
Continue selection (E8 + puromycin)
Day 3
Puromycin selection
l
Day 4
Day 2
Replace medium (E8)
Day 5 Passaging l
Passage to a new plate
Check BFP (40–50% positive) l Start puromycin (2 μg/mL) l
Day 6 l
If BFP < 80%, start puromycin again
l
Continue selection (E8 + puromycin)
Day 7 l
Make frozen stocks and start differentiation
sgRNA#1, and sgRNA#2). The cell density can be increased up to 600,000 cells/well depending on the growing speed of your iPSC line (see Note 3). 3. Add 1.6 μL of polybrene 10 mg/mL (final concentration 8 μg/mL). 4. Add 500 μL of lentivirus supernatant (total volume 2 mL). Return the 6-well plate to the incubator and wait for 3–4 h. 5. Remove the virus-containing medium and feed the cells with 2 mL of E8 + Y medium. Culture cells overnight. 6. Day 1: Replace medium to E8 medium (2 mL). 7. Day 2 (Start puromycin selection): Check BFP on a fluorescent microscope (expect 40–50% positive) to confirm the lentivirus infection is successful. Then start puromycin at the concentration determined in Subheading 3.1.2 (typically, around 2 μg/mL). 8. Day 3: Replace medium (E8 with puromycin). 9. Day 4: Replace medium (E8 with puromycin). 10. Day 5: Finish puromycin selection. Typically, the cells reach around 80% confluency at this time point. Passage iPSCs to a new Matrigel-coated 6-well plate. 11. Day 6: Check BFP and expect 90% positive. If the BFP-positive cells are still