Cardiac Regeneration: Methods and Protocols (Methods in Molecular Biology, 2158) 1071606670, 9781071606674

Table of contents :
Preface
Contents
Contributors
Part I: Cardiac Injury Models
Chapter 1: Myocardial Infarction Techniques in Adult Mice
1 Introduction
1.1 Cardiac Injuries
1.2 Assessing Cardiac Function in a High Variation Background
1.3 Cell Cycle Markers as Proxies for Proliferation
1.4 Discrepancies Between Different Attributes
2 Materials
2.1 Left Anterior Descending Artery Occlusion
2.2 Cardiac Function Assessment by Echocardiography
2.3 Histological Scar Quantification
2.4 Immunofluorescence Analysis of Cardiomyocyte Proliferation
3 Methods
3.1 Left Anterior Descending Artery Occlusion, See Also
3.1.1 One Day Prior to Surgery
3.1.2 Preparation for Surgery
3.1.3 Anesthesia and Intubation
3.1.4 Surgery
3.2 Cardiac Function Assessment by Echocardiography, See Also
3.2.1 Preparation for Echocardiography
3.2.2 Parasternal Long-Axis Acquisition
3.2.3 Short-Axis Acquisition
3.2.4 Analysis of the Long-Axis Echocardiographic Measurements B-Mode
3.2.5 Analysis of the Short-Axis Echocardiographic Measurements M-Mode
3.3 Histological Scar Quantification
3.3.1 Specimen Preparation and Image Acquisition
3.3.2 Maximum Damage Area Analysis
3.3.3 Scar Volume Analysis
3.3.4 Scar Degree Analysis
3.4 Immunofluorescence Analysis of Cardiomyocyte Proliferation
3.4.1 Staining
3.4.2 Imaging and Analysis
4 Notes
References
Chapter 2: Apical Resection and Cryoinjury of Neonatal Mouse Heart
1 Introduction
2 Materials
2.1 Equipment
2.2 Mice
2.3 Reagents
2.3.1 Picro-Sirius Red Staining
2.3.2 AFOG Staining
2.3.3 Cardiomyocyte Cell Cycle Assay by Immunostaining
3 Methods (Procedure Outlined in Fig. 2, See Note 1)
3.1 Prepare the Pups for Surgery and Anesthesia by Hypothermia
3.2 Ventricular Resection
3.3 Cryoinjury
3.4 Suture and Post-operative Care
3.5 Histological and Proliferation Analysis
3.5.1 Histological Analysis
3.5.2 Proliferation Analysis
4 Notes
References
Chapter 3: Left Ventricular Pressure Volume Loop Measurements Using Conductance Catheters to Assess Myocardial Function in Mice
1 Introduction
2 Materials
3 Methods
4 Notes
References
Chapter 4: Myocardial Infarction in Pigs
1 Introduction
2 Materials
2.1 Percutaneous Transluminal Coronary Angioplasty to Induce Coronary Ischemia
2.2 Surgical Equipment
2.3 Medications
3 Methods
3.1 Anesthetizing the Pig
3.2 Obtaining Baseline Functional Measurements
3.3 Inducing Myocardial Ischemia
3.4 Removing the Arterial and Venous Sheaths
3.5 Options for Administration of Compounds During the Procedure
3.6 Post-procedure (24 h-2 Months of Reperfusion, or Longer)
References
Chapter 5: Ventricular Cryoinjury as a Model to Study Heart Regeneration in Zebrafish
1 Introduction
2 Materials
2.1 Reagents and Equipment
2.2 Solutions
2.3 Material Preparation
3 Methods
3.1 Surgical Procedure
3.2 Heart Dissection and Fixation
4 Notes
References
Chapter 6: Cardiac Resection Injury in Zebrafish
1 Introduction
2 Materials
2.1 Zebrafish Handling
2.2 Anesthesia
2.3 Resection Injury
3 Methods
3.1 Preparation for Resection Injury
3.2 Anesthesia
3.3 Resection Injury
3.4 Post-injury Procedure
4 Notes
References
Chapter 7: A Genetic Cardiomyocyte Ablation Model for the Study of Heart Regeneration in Zebrafish
1 Introduction
2 Materials
3 Methods
3.1 Zebrafish Cardiomyocyte Ablation
3.2 Assay for Cardiomyocyte Proliferation During Regeneration
3.3 Histological Analysis of Cardiomyocyte Ablation and Regeneration
4 Notes
References
Chapter 8: Cardiac MRI Assessment of Mouse Myocardial Infarction and Regeneration
1 Introduction
2 Materials
2.1 T1-Weighted Gd-Based Contrast Agents (See Note 1)
2.2 T2-Weighted Iron Oxide Nanoparticles
3 Methods
3.1 Animal Preparation for In Vivo CMR
3.2 CMR Preparation
3.3 Cine CMR
3.4 Tagging MRI
3.5 Dynamic Contrast Enhancement (DCE)
3.6 Late Gadolinium Enhancement (LGE)
3.7 Extracellular Volume (ECV)
3.8 Cellular MRI for Intramyocardial Inflammation
4 Notes
References
Part II: Ex Vivo and In Vivo Approaches
Chapter 9: Isolation, Culture, and Live-Cell Imaging of Primary Rat Cardiomyocytes
1 Introduction
2 Materials
2.1 SADO Mix
2.2 Pre-plating and Plating Neonatal and Embryonic Cardiomyocytes
2.3 MACS Miltenyi Biotec Protocol
2.4 Adult Cardiomyocyte Isolation
3 Methods
3.1 SADO Mix Protocol
3.1.1 Neonatal Cardiomyocyte Isolation
3.1.2 Embryonic Cardiomyocyte Isolation
3.2 MACS Miltenyi Biotec Protocol
3.3 Adult Cardiomyocyte Isolation
3.4 Live-Cell Imaging of Cardiomyocytes to Detect and Analyze Mitosis
4 Notes
References
Chapter 10: Generation of Human Induced Pluripotent Stem Cells and Differentiation into Cardiomyocytes
1 Introduction
2 Materials
2.1 Fibroblast Transduction
2.2 Blood Cell Transduction
2.3 Checking Transgene by RT-PCR
2.3.1 Maintenance of Human iPSCs in a Feeder-Free System
2.3.2 Cardiomyocyte Differentiation Media and Reagents
3 Methods
3.1 Fibroblast Transduction
3.1.1 Fibroblast Preparation
3.1.2 Transduction
3.1.3 Clone Expansion
3.1.4 Confirmation of Vector-Free iPS Clones
3.2 Blood Cell Transduction
3.2.1 PBMC Isolation and Expansion
3.2.2 PBMC Expansion
3.2.3 Viral Transduction, Clone Expansion, and Confirmation of Vector-Free iPS Clones
3.3 Maintenance of Human iPSCs
3.3.1 Preparing Matrigel Plates
3.3.2 Passaging of iPSCs (See Note 9)
3.4 Human Cardiomyocyte Differentiation
3.4.1 iPS Passaging
3.4.2 Differentiation
3.4.3 CM Dissociation
4 Notes
References
Chapter 11: Differentiation of Human Induced Pluripotent Stem Cells into Epicardial-Like Cells
1 Introduction
2 Materials
2.1 Differentiation of iPSCs into iECs
2.2 Enrichment of CDH1+ iECs
2.3 Long-Term Maintenance of iPSC-Derived iECs
2.4 Differentiation of iECs into Cardiac Fibroblasts and Smooth Muscle Cells
2.5 Immunostaining of iECs and iEC Derivatives
2.6 Titration of CHIR99021 to Increase Efficiency of Mesoderm Differentiation
3 Methods
3.1 Epicardial Differentiation of iPSCs
3.2 Enrichment of CDH1+ iECs
3.3 Long-Term Maintenance of iECs
3.4 Differentiation of iECs into Cardiac Fibroblasts and Smooth Muscle Cells
3.5 Immunostaining Analysis
3.6 Titration of CHIR99021 to Increase Efficiency of Mesoderm Differentiation
4 Notes
References
Chapter 12: In Vitro Conversion of Murine Fibroblasts into Cardiomyocyte-Like Cells
1 Introduction
2 Materials
2.1 Generation of Retroviruses
2.2 Generation of Mouse Embryonic Fibroblasts (MEFs)
2.3 Generation of Neonatal Mouse Fresh Cardiac Fibroblasts
2.4 Cardiac Reprogramming In Vitro and Evaluation of Reprogramming Efficiency
3 Methods
3.1 Generation of Retroviruses (See Note 1)
3.2 Generation of Mouse Embryonic Fibroblasts (MEFs)
3.3 Generation of Neonatal Mouse Fresh Cardiac Fibroblasts (See Note 6)
3.3.1 Generate Cardiac Fibroblasts from Explant Culture Method
3.3.2 Generate Cardiac Fibroblasts from Enzyme Digestion Method
3.3.3 Isolation of Thy1.2+ Fibroblasts by Magnetic Activated Cell Sorting (MACS)
3.4 Cardiac Reprogramming In Vitro
3.5 Evaluation of Reprogramming Efficiency
3.5.1 FACS Analysis of Reprogramming Efficiency
3.5.2 RT-qPCR Analysis to Test Gene Expression
3.5.3 Immunocytochemical (ICC) Analysis of Reprogramming Efficiency
3.5.4 Evaluation of Calcium Activity of iCMs at Week 4
4 Notes
References
Chapter 13: Frame-Hydrogel Methodology for Engineering Highly Functional Cardiac Tissue Constructs
1 Introduction
2 Materials
2.1 Manufacturing of Tissue Molds
2.2 3D Engineered Tissue Culture Media and Cell Sources
2.3 3D Tissue Generation
3 Methods
3.1 Manufacturing of PDMS Tissue Molds
3.2 Frame Manufacturing
3.3 Initial Mold Preparation for Tissue Culture
3.4 Cell Preparation
3.5 Final Mold Preparation
3.6 Hydrogel Preparation
3.7 Casting of Engineered Tissues in PDMS Molds
3.8 Removing Engineered Tissues from Molds
3.9 Tissue Culture
4 Notes
References
Chapter 14: Efficient Protocols for Fabricating a Large Human Cardiac Muscle Patch from Human Induced Pluripotent Stem Cells
1 Introduction
2 Materials
2.1 Maintaining hiPSCs
2.2 Supplies/Reagents for Differentiating hiPSCs into CMs
2.3 Supplies/Reagents for Differentiating hiPSCs into ECs
2.4 Supplies/Reagents for Differentiating hiPSCs into SMCs
2.5 Supplies and Reagents for hCMP Manufacture
2.6 Supplies and Reagents Needed for Patch Transplantation in Swine MI Model
2.7 Equipment
3 Methods
3.1 HiPSC Culture and Maintenance
3.2 Differentiation of hiPSCs into CMs
3.3 Differentiation of hiPSCs into ECs
3.4 Differentiation of hiPSCs into SMCs
3.5 Construction of a Large hCMP
3.6 hCMP Transplantation
3.7 Analysis of hCMP Engraftment
4 Notes
References
Chapter 15: Isolation and Characterization of Intact Cardiomyocytes from Frozen and Fresh Human Myocardium and Mouse Hearts
1 Introduction
2 Materials
2.1 Tools and Instruments
2.2 Reagents
3 Methods
3.1 Sample Fixation and Enzyme Digestion
3.2 Count the Total Number of Cardiomyocytes in the Mouse Heart
3.3 Immunofluorescence Microscopy
3.4 Image Acquisition and Cardiomyocyte Characterization
3.5 Other Species and Organs
4 Notes
References
Chapter 16: Ex Vivo Techniques to Study Heart Regeneration in Zebrafish
1 Introduction
2 Materials
2.1 Zebrafish
2.2 Reagents and Key Equipment
3 Methods
3.1 Preparation
3.2 Heart Extraction and Cleaning
3.3 Standard Explant Culture
3.4 Epicardial Cell Ablation
3.5 Proliferation Assay
3.6 Partial Ventricular Explant Culture
3.7 Chemical Treatments and Screens
4 Notes
References
Chapter 17: Purification of Pluripotent Stem Cell-Derived Cardiomyocytes Using CRISPR/Cas9-Mediated Integration of Fluorescent...
1 Introduction
2 Materials
2.1 iPSC Culture Maintenance, Passaging, and Freezing
2.2 SgRNA Cloning into PX458 Plasmid
2.3 Integration of Reporter Constructs into iPSCs
2.4 iPSC Cardiac Differentiation
2.5 Fluorescence-Activated Cell Sorting of Fluorescently Labeled Cardiomyocytes
3 Methods
3.1 Maintenance of Undifferentiated iPSCs
3.2 Passaging Undifferentiated iPSCs
3.3 Freezing Undifferentiated iPSCs
3.4 SgRNA Design
3.5 Cloning and Validation of sgRNAs
3.6 Design of Reporter Constructs for Isolation of PSC-Derived Cardiomyocytes
3.7 Integration of Reporter Constructs into iPSCs
3.8 Colony Picking and Clone Validation
3.9 hiPSC Cardiac-Directed Differentiation
3.10 Fluorescence-Activated Cell Sorting of Fluorescently Labeled Cardiomyocytes
4 Notes
References
Part III: Visualizing and Manipulating Heart Regeneration
Chapter 18: In Vivo Clonal Analysis of Cardiomyocytes
1 Introduction
2 Materials
2.1 Mouse Models
2.2 Genotyping Primers
2.3 Hydroxytamoxifen Preparation and Administration
2.4 2-Dimensional Quantification of CM Clone Number and Size
2.5 3-Dimensional Quantification of CM Clone Volume
2.5.1 Clarity
2.5.2 Light-Sheet Imaging
2.5.3 Imaris Processing
3 Methods
3.1 Generation of aMHCCreER;Rainbow
3.2 Hydroxytamoxifen Preparation and Administration
3.3 2-Dimensional Quantification of Cardiomyocyte Clones
3.3.1 Tissue Harvest and Processing for Histological Analysis
3.3.2 Fluorescent Imaging of Tissue Sections
3.3.3 Quantification of Clone Number and Size
3.4 3-Dimensional Quantification of Clone Volumes
3.4.1 Clearing of Postnatal Day 2 (P2) Mouse Hearts Using CLARITY
3.4.2 Light-Sheet Calibration
3.4.3 Light-Sheet Imaging
3.4.4 Image Reconstruction and 3D Clonal Volume Measurement Using Imaris
4 Notes
References
Chapter 19: High-Fidelity Quantification of Cell Cycle Activity with Multi-Isotope Imaging Mass Spectrometry
1 Introduction
2 Materials
2.1 In Vivo Labeling
2.2 Tissue Fixation
2.3 Tissue Embedding
2.4 Preparing Silicon Wafers
2.5 Sectioning and Preparing the Mounted Sample
2.6 NanoSIMS Analysis
2.7 Image Visualization and Data Analysis
3 Methods
3.1 In Vivo Labeling
3.2 Tissue Fixation
3.3 Tissue Embedding
3.4 Preparing Silicon Wafers
3.5 Sectioning and Preparing of the Mounted Sample
3.6 NanoSIMS Analysis
3.7 Image Visualization and Data Analysis
4 Notes
References
Chapter 20: AAV Gene Transfer to the Heart
1 Introduction
2 Materials
3 Methods
3.1 Preparation of DNA for AAV Production
3.2 Transfection of HEK293 Cells
3.3 Harvesting of Crude AAV Prep
3.4 Purification of AAV
3.5 QPCR-Based Quantification of AAV Titer
3.6 Delivery of AAV to Mice
4 Notes
References
Chapter 21: In Vitro Synthesis of Modified RNA for Cardiac Gene Therapy
1 Introduction
2 Materials
2.1 Equipment
2.2 Supplies
2.3 In Vitro Synthesis of modRNA
2.3.1 DNA Plasmid Order
2.3.2 Plasmid Transformation
2.3.3 Plasmid Miniprep
2.3.4 Synthesizing Tailed DNA Template for IVT Reaction
2.3.5 Purifying PCR Product
2.3.6 Checking Quality of Tailed DNA
2.3.7 In Vitro Transcription Reaction
2.3.8 Purify modRNA Using MegaClear
2.3.9 RNA Phosphatase Treatment
2.3.10 Purify modRNA Using MegaClear
2.3.11 Concentrate modRNA for In Vivo Use by Amicon Ultra-4 Centrifugal Filters
2.3.12 Measure the Quality of modRNA by TapeStation
3 Methods
3.1 In Vitro Synthesis of modRNA
3.1.1 Plasmid Order
3.1.2 Plasmid Transformation
3.1.3 Plasmid Miniprep
3.1.4 Synthesizing Tailed DNA Template for IVT Reaction
3.1.5 Purify PCR Product Using the QIAquick PCR Purification Kit
3.1.6 Checking Quality of Tailed DNA
3.1.7 In Vitro Transcription (IVT) Reaction
3.1.8 Purify modRNA Using MEGAclear
3.1.9 RNA Phosphatase Treatment
3.1.10 Purify modRNA Using MEGAclear
3.1.11 Concentrate modRNA for In Vivo Use by Amicon Ultra-4 Centrifugal Filters
3.1.12 Measure modRNA Quality by Agilent High-Sensitivity RNA Assay
3.2 modRNA Preparation for In Vitro or In Vivo Delivery, Detection, and Analysis of modRNA Translation
3.2.1 In Vitro Transfection
3.2.2 In Vivo Delivery
3.2.3 Detecting Luciferase Expression Using the IVIS System
4 Notes
References
Chapter 22: Generation and Manipulation of Exosomes
1 Introduction
2 Materials
2.1 Exosome Generation
2.2 Exosome Manipulation: Cloaking Ischemic Myocardium Homing Peptide
2.3 Exosome Manipulation: Cloaking Platelet Membrane
3 Methods
3.1 Exosome Generation
3.2 Exosome Manipulation: Cloaking Ischemic Myocardium Homing Peptide
3.3 Exosome Manipulation: Cloaking Platelet Membrane
4 Notes
References
Chapter 23: Epigenetic Assays in Purified Cardiomyocyte Nuclei
1 Introduction
2 Materials
2.1 Cardiomyocyte Nuclei Isolation
2.2 ATAC-Seq
2.3 CUT&RUN and High-Throughput Sequencing
3 Methods
3.1 Density Gradient Centrifugation
3.2 Magnetic Immunoprecipitation of Cardiomyocyte Nuclei
3.3 CM-Specific ATAC-Seq
3.4 ATAC-Seq Library Preparation and Quality Control
3.5 CUT&RUN
3.6 CUT&RUN DNA Extraction
3.7 CUT&RUN Library Preparation
4 Notes
References
Chapter 24: Genetic Lineage Tracing of Non-cardiomyocytes in Mice
1 Introduction
2 Materials
3 Methods
3.1 Generation of c-Kit-Based Lineage-Tracing Mice
3.2 Tamoxifen Treatment
3.3 Harvest Tissues for Frozen Sections
3.4 Immunofluorescence to Identify Cells Derived from c-Kit+ Cells
3.5 Quantification of c-Kit-Derived Lineages, Such as Cardiomyocytes
4 Notes
References
Chapter 25: Experimental Hypoxia as a Model for Cardiac Regeneration in Mice
1 Introduction
2 Materials
2.1 Hypoxia Chamber (Coy Laboratory Products) (See Fig. 1)
2.1.1 Principle
2.1.2 Standard Equipment
2.2 Live-Animal Waste Filtration System
2.2.1 Standard Equipment
2.2.2 Principle
3 Methods
3.1 Mild Hypoxia in Neonatal Mice (See Note 2)
3.2 Severe Hypoxia in Adult Mice (See Schematic)
3.3 Controlling the Atmosphere by Nitrogen Gas
3.3.1 Principle
3.3.2 Calibration Setup
3.4 O2 Calibration Procedure
3.4.1 Zero Calibration
3.4.2 Span Calibration
3.5 Daily Maintenance of the Hypoxia Chamber
4 Notes
References
Correction to: Ventricular Cryoinjury as a Model to Study Heart Regeneration in Zebrafish
Index

Citation preview

Methods in Molecular Biology 2158

Kenneth D. Poss Bernhard Kühn Editors

Cardiac Regeneration Methods and Protocols

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.

Cardiac Regeneration Methods and Protocols

Edited by

Kenneth D. Poss Department of Cell Biology, Duke University Medical Center, Durham, NC, USA

Bernhard Kühn Division of Cardiology, UPMC Children’s Hospital of Pittsburgh, Pittsburgh, PA, USA

Editors Kenneth D. Poss Department of Cell Biology Duke University Medical Center Durham, NC, USA

Bernhard Ku¨hn Division of Cardiology UPMC Children’s Hospital of Pittsburgh Pittsburgh, PA, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-0667-4 ISBN 978-1-0716-0668-1 (eBook) https://doi.org/10.1007/978-1-0716-0668-1 © Springer Science+Business Media, LLC, part of Springer Nature 2021, Corrected Publication 2021 Chapter 5 is licensed under the terms of the Creative Commons Attribution 4.0 International License (http:// creativecommons.org/licenses/by/4.0/). For further details see license information in the chapter. 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. Cover Illustration Caption: Cover image courtesy of Fei Sun (Duke University) and Honghai Liu (University of Pittsburgh Medical Center). 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 Myocardial infarction (MI) is a life-threatening injury causing permanent loss of cardiomyocytes. MI events are experienced by over 750,000 people each year in the United States, with three out of four surviving. Scars form in MI survivors, increasing susceptibility to compensatory pathology, aneurysm, additional MI events, heart failure, and sudden death. Therefore, a major goal of regenerative medicine and of the cardiovascular research community is to replenish lost cardiomyocytes, avoid scar-associated pathology, and improve MI outcomes. The path to cardiac regenerative medicine requires the use of relevant animal models with different capacities for innate heart regeneration. It also requires technologies to generate large numbers of cardiomyocytes from stem cells in vitro and transplant them into injured hearts, or to reprogram cells to switch to a cardiomyocyte cell fate. The field of heart regeneration has matured over the past 20 years, enduring many breakthroughs and some controversies. One of these controversies may, thankfully, be resolved and behind us because recent findings argue strongly against the notion that the adult mammalian heart has a robust population of cardiac stem cells with substantial cardiomyogenic potential in vitro or in vivo (see Chap. 23). Other topics in the field are in the process of finding consensus; for instance, that the extent to which the mouse heart regenerates after an extreme injury at early postnatal days can depend on surgical technique, strain background, or type of injury (see Chap. 2). Some controversies, and a more general challenge, are that many of the methods in the field of heart regeneration are complex or require great skill and experience. For instance, a common procedure like surgical myocardial infarction in adult mice takes months to master and results in variable injury size. The field requires detailed protocols that are reproducible in different laboratories. Part 1 is a section on “Cardiac Injury Models.” Chaps. 1–8 describe protocols on different methods of heart injury to key models of repair and regeneration: zebrafish, neonatal and adult mice, and pigs. These include surgical removal of myocardial tissue, experimental freezing of tissue, genetic methods to inducibly and specially ablate cardiomyocytes, as well as classic myocardial infarction injuries. It is now clear that cardiac tissues respond in tailored ways to different injury types, for instance, those that kill tissue in place versus those that extract tissue from the animal. The ability to perform injuries in species with high regenerative capacity (e.g., zebrafish, neonatal mammals) and poor capacity (adult mammals) provides single or collaborating groups a powerful ability to test whether specific factors/manipulations are necessary and sufficient for heart regeneration. In Part II, we describe techniques for culturing cardiomyocytes from different species, either directly isolated from the heart or generated by differentiation protocols. An exciting route to therapy is the potential for autologous transplantation of patient-derived cardiomyocytes to an infarct injury. Yet, major challenges exist in poor survival, inefficient coupling, and ineffective maturation (if derived from stem cells) of transplanted cardiomyocytes. Chaps. 8–15 describe methods for isolating and culturing cardiomyocytes, for generating a large, pure population of cardiomyocytes (or other cardiac cell types) from pluripotent stem cells or by direct reprogramming from fibroblasts, and for creating tissue patches comprising muscle and other cardiac cell types. The last of these can be transplanted as a mass of functional muscle, trained in culture to rhythmically contract. Also included is a chapter

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on methodology to study regenerative phenomena in intact hearts of adult zebrafish outside of the animal and able to beat in culture for several weeks. Part III is composed of methods for labeling or manipulation of cardiac tissue for the purpose of answering questions in regeneration, or for potentially changing the regenerative response. Analysis methods include chromatin signature profiling from cardiomyocyte nuclei, methods for genetic fate-mapping or multicolor clonal analysis, and use of isotopes to assess cardiac proteomes by mass spectroscopy. We include methods to generate and infect the heart with adeno-associated viral vectors, a key mode of contemporary gene therapy, and to treat the injured heart with either purified exosomes or modified mRNA molecules with enhanced stability. Finally, we include a chapter that describes equipment and methods for inducing hypoxia in mice, which has been reported to enhance the proliferative capacity of cardiomyocytes in mice under certain experimental conditions. These chapters describe the latest models and methods used in the field of heart regeneration, designed for researchers interested in establishing these assays in their laboratories to reproduce or extend findings, and for familiarizing themselves with the field if it is new to them. We are indebted to all of the contributors to this volume. In addition, we effusively thank Lisa M. Summe for shepherding all communications with authors and extensive help with editing. We wish the best to all new and established researchers in the field in their efforts toward understanding and modulating the heart’s regenerative capacity! Durham, NC, USA Pittsburgh, PA, USA

Kenneth D. Poss ¨ hn Bernhard Ku

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

PART I

CARDIAC INJURY MODELS

1 Myocardial Infarction Techniques in Adult Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . Elad Bassat, Dahlia E. Perez, and Eldad Tzahor 2 Apical Resection and Cryoinjury of Neonatal Mouse Heart . . . . . . . . . . . . . . . . . . Hua Shen, Ali Darehzereshki, Henry M. Sucov, and Ching-Ling Lien 3 Left Ventricular Pressure Volume Loop Measurements Using Conductance Catheters to Assess Myocardial Function in Mice . . . . . . . . . . . . . . . Tilman Ziegler, Karl-Ludwig Laugwitz, and Christian Kupatt 4 Myocardial Infarction in Pigs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Andrea B€ a hr, Nadja Hornaschewitz, and Christian Kupatt 5 Ventricular Cryoinjury as a Model to Study Heart Regeneration in Zebrafish . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ines J. Marques, Andre´s Sanz-Morejo n, and Nadia Mercader 6 Cardiac Resection Injury in Zebrafish . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Delicia Z. Sheng, Dawei Zheng, and Kazu Kikuchi 7 A Genetic Cardiomyocyte Ablation Model for the Study of Heart Regeneration in Zebrafish . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fei Sun, Adam R. Shoffner, and Kenneth D. Poss 8 Cardiac MRI Assessment of Mouse Myocardial Infarction and Regeneration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Yijen L. Wu

PART II

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EX VIVO AND IN VIVO APPROACHES

9 Isolation, Culture, and Live-Cell Imaging of Primary Rat Cardiomyocytes . . . . . Marina Leone and Felix B. Engel 10 Generation of Human Induced Pluripotent Stem Cells and Differentiation into Cardiomyocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ¨ hn Lu Han, Jocelyn Mich-Basso, and Bernhard Ku 11 Differentiation of Human Induced Pluripotent Stem Cells into Epicardial-Like Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Navid A. Nafissi, Paige DeBenedittis, Michael C. Thomas, and Ravi Karra 12 In Vitro Conversion of Murine Fibroblasts into Cardiomyocyte-Like Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jun Xu, Li Wang, Jiandong Liu, and Li Qian 13 Frame-Hydrogel Methodology for Engineering Highly Functional Cardiac Tissue Constructs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Abbigail Helfer and Nenad Bursac

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Contents

Efficient Protocols for Fabricating a Large Human Cardiac Muscle Patch from Human Induced Pluripotent Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ling Gao and Jianyi (Jay) Zhang Isolation and Characterization of Intact Cardiomyocytes from Frozen and Fresh Human Myocardium and Mouse Hearts. . . . . . . . . . . . . . ¨ hn Honghai Liu, Kevin Bersell, and Bernhard Ku Ex Vivo Techniques to Study Heart Regeneration in Zebrafish . . . . . . . . . . . . . . . Sierra Duca and Jingli Cao Purification of Pluripotent Stem Cell-Derived Cardiomyocytes Using CRISPR/Cas9-Mediated Integration of Fluorescent Reporters. . . . . . . . . Francisco X. Galdos, Adrija K. Darsha, Sharon L. Paige, and Sean M. Wu

PART III

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VISUALIZING AND MANIPULATING HEART REGENERATION

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In Vivo Clonal Analysis of Cardiomyocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Ngoc B. Nguyen, G. Esteban Fernandez, Yichen Ding, Tzung Hsiai, and Reza Ardehali 19 High-Fidelity Quantification of Cell Cycle Activity with Multi-Isotope Imaging Mass Spectrometry. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Frank Gyngard, Louise Trakimas, and Matthew L. Steinhauser 20 AAV Gene Transfer to the Heart. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Suya Wang, Yuxuan Guo, and William T. Pu 21 In Vitro Synthesis of Modified RNA for Cardiac Gene Therapy. . . . . . . . . . . . . . . Nishat Sultana, Mohammad Tofael Kabir Sharkar, Yoav Hadas, Elena Chepurko, and Lior Zangi 22 Generation and Manipulation of Exosomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Shiqi Hu, Li Qiao, and Ke Cheng 23 Epigenetic Assays in Purified Cardiomyocyte Nuclei. . . . . . . . . . . . . . . . . . . . . . . . . Matthew C. Hill and James F. Martin 24 Genetic Lineage Tracing of Non-cardiomyocytes in Mice . . . . . . . . . . . . . . . . . . . . Zhongming Chen and Jop H. van Berlo 25 Experimental Hypoxia as a Model for Cardiac Regeneration in Mice . . . . . . . . . . Yuji Nakada and Hesham A. Sadek Correction to: Ventricular Cryoinjury as a Model to Study Heart Regeneration in Zebrafish. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors REZA ARDEHALI • Division of Cardiology, Department of Internal Medicine, David Geffen School of Medicine, University of California, Los Angeles, Los Angeles, CA, USA; Eli and Edythe Broad Center for Regenerative Medicine and Stem Cell Research, University of California, Los Angeles, Los Angeles, CA, USA; Molecular, Cellular and Integrative Physiology Graduate Program, University of California, Los Angeles, Los Angeles, CA, USA; Molecular Biology Institute, University of California, Los Angeles, Los Angeles, CA, USA ANDREA BA€ HR • Internal Medicine I, Klinikum rechts der Isar der TU Mu¨nchen, Munich, Germany; DZHK (German Center for Cardiovascular Research), Partner Site Munich Heart Alliance, Munich, Germany ELAD BASSAT • Department of Molecular Cell Biology, Weizmann Institute of Science, Rehovot, Israel KEVIN BERSELL • Department of Medicine, Brigham and Women’s Hospital, Harvard Medical School, Boston, MA, USA NENAD BURSAC • Department of Biomedical Engineering, Duke University, Durham, NC, USA JINGLI CAO • Department of Cell and Developmental Biology, Cardiovascular Research Institute, Weill Cornell Medical College, Cornell University, New York, NY, USA KE CHENG • Department of Molecular Biomedical Sciences and Comparative Medicine Institute, North Carolina State University, Raleigh, NC, USA; Joint Department of Biomedical Engineering, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA ZHONGMING CHEN • Department of Medicine, Cardiovascular Division, Lillehei Heart Institute, University of Minnesota, Minneapolis, MN, USA ELENA CHEPURKO • Department of Genetics and Genomic Sciences, Icahn School of Medicine at Mount Sinai, New York, NY, USA; Cardiovascular Research Center, Icahn School of Medicine at Mount Sinai, New York, NY, USA; Black Family Stem Cell Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA ALI DAREHZERESHKI • Saban Research Institute and Heart Institute, Children’s Hospital Los Angeles, Los Angeles, CA, USA; Department of Surgery, MedStar Health Baltimore, Franklin Square Medical Center, Baltimore, MD, USA ADRIJA K. DARSHA • Cardiovascular Institute, Stanford University, Stanford, CA, USA; Institute of Stem Cell and Regenerative Biology, School of Medicine, Stanford University, Stanford, CA, USA PAIGE DEBENEDITTIS • Department of Medicine, Duke University Medical Center, Durham, NC, USA YICHEN DING • Division of Cardiology, Department of Internal Medicine, David Geffen School of Medicine, University of California, Los Angeles, CA, USA SIERRA DUCA • Department of Cell and Developmental Biology, Cardiovascular Research Institute, Weill Cornell Medical College, Cornell University, New York, NY, USA

ix

x

Contributors

FELIX B. ENGEL • Experimental Renal and Cardiovascular Research, Department of Nephropathology, Institute of Pathology, Friedrich-Alexander-Universit€ at ErlangenNu¨rnberg (FAU), Erlangen, Germany; Muscle Research Center Erlangen (MURCE), Erlangen, Germany G. ESTEBAN FERNANDEZ • Children’s Hospital Los Angeles, The Saban Research Institute, Los Angeles, CA, USA FRANCISCO X. GALDOS • Cardiovascular Institute, School of Medicine, Stanford University, Stanford, CA, USA; Institute of Stem Cell and Regenerative Biology, School of Medicine, Stanford University, Stanford, CA, USA LING GAO • Department of Biomedical Engineering, School of Medicine and School of Engineering, University of Alabama at Birmingham, Birmingham, AL, USA; Translational Medical Center for Stem Cell Therapy & Institute for Regenerative Medicine, Shanghai East Hospital, School of Life Sciences and Technology, Tongji University, Shanghai, China YUXUAN GUO • Department of Cardiology, Boston Children’s Hospital, Boston, MA, USA FRANK GYNGARD • Center for NanoImaging, Division of Genetics, Brigham and Women’s Hospital, Boston, MA, USA; Harvard Medical School, Boston, MA, USA YOAV HADAS • Cardiovascular Research Center, Icahn School of Medicine at Mount Sinai, New York, NY, USA; Department of Genetics and Genomic Sciences, Icahn School of Medicine at Mount Sinai, New York, NY, USA; Black Family Stem Cell Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA LU HAN • Division of Cardiology, Pediatric Institute for Heart Regeneration and Therapeutics (I-HRT), UPMC Children’s Hospital of Pittsburgh, Pittsburgh, PA, USA; Department of Pediatrics, University of Pittsburgh, Pittsburgh, PA, USA; Richard King Mellon Foundation Institute for Pediatric Research, Pittsburgh, PA, USA ABBIGAIL HELFER • Department of Biomedical Engineering, Duke University, Durham, NC, USA MATTHEW C. HILL • Program in Developmental Biology, Baylor College of Medicine, Houston, TX, USA NADJA HORNASCHEWITZ • Internal Medicine I, Klinikum rechts der Isar der TU Mu¨nchen, Munich, Germany; DZHK (German Center for Cardiovascular Research), Partner Site Munich Heart Alliance, Munich, Germany TZUNG HSIAI • Division of Cardiology, Department of Internal Medicine, David Geffen School of Medicine, University of California, Los Angeles, CA, USA SHIQI HU • Department of Molecular Biomedical Sciences and Comparative Medicine Institute, North Carolina State University, Raleigh, NC, USA; Joint Department of Biomedical Engineering, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA RAVI KARRA • Department of Medicine, Duke University Medical Center, Durham, NC, USA; Regeneration Next, Duke University, Durham, NC, USA KAZU KIKUCHI • Developmental and Stem Cell Biology Division, Victor Chang Cardiac Research Institute, Darlinghurst, NSW, Australia; St. Vincent’s Clinical School, University of New South Wales, Kensington, NSW, Australia BERNHARD KU¨HN • Division of Cardiology, Pediatric Institute for Heart Regeneration and Therapeutics (I-HRT), UPMC Children’s Hospital of Pittsburgh, Pittsburgh, PA, USA; Department of Pediatrics, University of Pittsburgh, Pittsburgh, PA, USA; McGowan Institute of Regenerative Medicine, Pittsburgh, PA, USA; Richard King Mellon Foundation Institute for Pediatric Research, Pittsburgh, PA, USA

Contributors

xi

CHRISTIAN KUPATT • Internal Medicine I, Klinikum rechts der Isar der TU Mu¨nchen, Munich, Germany; DZHK (German Center for Cardiovascular Research), Partner Site Munich Heart Alliance, Munich, Germany KARL-LUDWIG LAUGWITZ • Internal Medicine I, Klinikum rechts der Isar der TU Mu¨nchen, Munich, Germany; DZHK (German Center for Cardiovascular Research), Partner Site Munich Heart Alliance, Munich, Germany MARINA LEONE • Experimental Renal and Cardiovascular Research, Department of Nephropathology, Institute of Pathology, Friedrich-Alexander-Universit€ at ErlangenNu¨rnberg (FAU), Erlangen, Germany CHING-LING LIEN • Saban Research Institute and Heart Institute, Children’s Hospital Los Angeles, Los Angeles, CA, USA; Department of Surgery, Biochemistry and Molecular Medicine, University of Southern California, Los Angeles, CA, USA HONGHAI LIU • Richard King Mellon Foundation Institute for Pediatric Research, Pittsburgh, PA, USA; Division of Cardiology, UPMC Children’s Hospital of Pittsburgh, Pittsburgh, PA, USA; Department of Pediatrics, University of Pittsburgh, Pittsburgh, PA, USA; Pediatric Institute for Heart Regeneration and Therapeutics (I-HRT), Pittsburgh, PA, USA JIANDONG LIU • Department of Pathology and Laboratory Medicine, McAllister Heart Institute, University of North Carolina, Chapel Hill, NC, USA INES J. MARQUES • Institute of Anatomy, University of Bern, Bern, Switzerland JAMES F. MARTIN • Program in Developmental Biology, Baylor College of Medicine, Houston, TX, USA; Department of Molecular Physiology and Biophysics, Baylor College of Medicine, Houston, TX, USA; Texas Heart Institute, Houston, TX, USA; Cardiovascular Research Institute, Baylor College of Medicine, Houston, TX, USA NADIA MERCADER • Institute of Anatomy, University of Bern, Bern, Switzerland; Centro Nacional de Investigaciones Cardiovasculares (CNIC), Madrid, Spain JOCELYN MICH-BASSO • Division of Cardiology, Pediatric Institute for Heart Regeneration and Therapeutics (I-HRT), UPMC Children’s Hospital of Pittsburgh, Pittsburgh, PA, USA; Department of Pediatrics, University of Pittsburgh, Pittsburgh, PA, USA; Richard King Mellon Foundation Institute for Pediatric Research, Pittsburgh, PA, USA NAVID A. NAFISSI • Department of Medicine, Duke University Medical Center, Durham, NC, USA YUJI NAKADA • Department of Internal Medicine, University of Texas Southwestern Medical Center, Dallas, TX, USA NGOC B. NGUYEN • Division of Cardiology, Department of Internal Medicine, David Geffen School of Medicine, University of California, Los Angeles, CA, USA; Eli and Edythe Broad Center for Regenerative Medicine and Stem Cell Research, University of California, Los Angeles, CA, USA; Molecular, Cellular and Integrative Physiology Graduate Program, University of California, Los Angeles, CA, USA SHARON L. PAIGE • Cardiovascular Institute, School of Medicine, Stanford University, Stanford, CA, USA; Institute of Stem Cell and Regenerative Biology, School of Medicine, Stanford University, Stanford, CA, USA; Division of Cardiovascular Medicine, Department of Medicine, School of Medicine, Stanford University, Stanford, CA, USA DAHLIA E. PEREZ • Department of Molecular Cell Biology, Weizmann Institute of Science, Rehovot, Israel KENNETH D. POSS • Regeneration Next and Department of Cell Biology, Duke University School of Medicine, Durham, NC, USA WILLIAM T. PU • Department of Cardiology, Boston Children’s Hospital, Boston, MA, USA

xii

Contributors

LI QIAN • Department of Pathology and Laboratory Medicine, McAllister Heart Institute, University of North Carolina, Chapel Hill, NC, USA LI QIAO • Department of Molecular Biomedical Sciences and Comparative Medicine Institute, North Carolina State University, Raleigh, NC, USA; Joint Department of Biomedical Engineering, University of North Carolina at Chapel Hill, Chapel Hill, NC, USA HESHAM A. SADEK • Department of Internal Medicine, University of Texas Southwestern Medical Center, Dallas, TX, USA; Division of Cardiology, Department of Internal Molecular Biology, University of Texas Southwestern Medical Center, Dallas, TX, USA ANDRE´S SANZ-MOREJO´N • Institute of Anatomy, University of Bern, Bern, Switzerland; Centro Nacional de Investigaciones Cardiovasculares (CNIC), Madrid, Spain MOHAMMAD TOFAEL KABIR SHARKAR • Department of Genetics and Genomic Sciences, Icahn School of Medicine at Mount Sinai, New York, NY, USA; Cardiovascular Research Center, Icahn School of Medicine at Mount Sinai, New York, NY, USA; Black Family Stem Cell Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA DELICIA Z. SHENG • Developmental and Stem Cell Biology Division, Victor Chang Cardiac Research Institute, Darlinghurst, NSW, Australia HUA SHEN • Department of Stem Cell Biology and Regenerative Medicine, Broad CIRM Center, Keck School of Medicine, University of Southern California, Los Angeles, CA, USA ADAM R. SHOFFNER • Regeneration Next and Department of Cell Biology, Duke University School of Medicine, Durham, NC, USA MATTHEW L. STEINHAUSER • Center for NanoImaging, Division of Genetics, Brigham and Women’s Hospital, Boston, MA, USA; Harvard Medical School, Boston, MA, USA; Division of Cardiovascular Medicine, Brigham and Women’s Hospital, Boston, MA, USA; Aging Institute, University of Pittsburgh School of Medicine, Pittsburgh, PA, USA HENRY M. SUCOV • Department of Stem Cell Biology and Regenerative Medicine, Broad CIRM Center, Keck School of Medicine, University of Southern California, Los Angeles, CA, USA; Department of Regenerative Medicine and Cell Biology, Medical University of South Carolina, Charleston, SC, USA; Division of Cardiology, Department of Medicine, Medical University of South Carolina, Charleston, SC, USA NISHAT SULTANA • Department of Genetics and Genomic Sciences, Icahn School of Medicine at Mount Sinai, New York, NY, USA; Cardiovascular Research Center, Icahn School of Medicine at Mount Sinai, New York, NY, USA; Black Family Stem Cell Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA FEI SUN • Regeneration Next and Department of Cell Biology, Duke University School of Medicine, Durham, NC, USA MICHAEL C. THOMAS • Department of Medicine, Duke University Medical Center, Durham, NC, USA LOUISE TRAKIMAS • Harvard Medical School, Boston, MA, USA ELDAD TZAHOR • Department of Molecular Cell Biology, Weizmann Institute of Science, Rehovot, Israel JOP H. VAN BERLO • Department of Medicine, Cardiovascular Division, Lillehei Heart Institute, University of Minnesota, Minneapolis, MN, USA; Stem Cell Institute and Department of Integrative Biology and Physiology, University of Minnesota, Minneapolis, MN, USA LI WANG • Department of Pathology and Laboratory Medicine, McAllister Heart Institute, University of North Carolina, Chapel Hill, NC, USA SUYA WANG • Department of Cardiology, Boston Children’s Hospital, Boston, MA, USA

Contributors

xiii

SEAN M. WU • Cardiovascular Institute, School of Medicine, Stanford University, Stanford, CA, USA; Institute of Stem Cell and Regenerative Biology, School of Medicine, Stanford University, Stanford, CA, USA; Departments of Pediatrics and Medicine, Division of Cardiovascular Medicine, Stanford University, Stanford, CA, USA YIJEN L. WU • Department of Developmental Biology, Rangos Research Center Animal Imaging Core, School of Medicine, University of Pittsburgh, Pittsburgh, PA, USA JUN XU • Department of Pathology and Laboratory Medicine, McAllister Heart Institute, University of North Carolina, Chapel Hill, NC, USA LIOR ZANGI • Cardiovascular Research Center, Icahn School of Medicine at Mount Sinai, New York, NY, USA; Department of Genetics and Genomic Sciences, Icahn School of Medicine at Mount Sinai, New York, NY, USA; Black Family Stem Cell Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA JIANYI (JAY) ZHANG • Department of Biomedical Engineering, School of Medicine and School of Engineering, University of Alabama at Birmingham, Birmingham, AL, USA DAWEI ZHENG • Developmental and Stem Cell Biology Division, Victor Chang Cardiac Research Institute, Darlinghurst, NSW, Australia TILMAN ZIEGLER • Internal Medicine I, Klinikum rechts der Isar der TU Mu¨nchen, Munich, Germany; DZHK (German Center for Cardiovascular Research), Partner Site Munich Heart Alliance, Munich, Germany

Part I Cardiac Injury Models

Chapter 1 Myocardial Infarction Techniques in Adult Mice Elad Bassat, Dahlia E. Perez, and Eldad Tzahor Abstract The discovery of endogenous regenerative potential of the heart in zebrafish and neonatal mice has shifted the cardiac regenerative field towards the utilization of intrinsic regenerative mechanisms in the mammalian heart. The goal of these studies is to understand, and eventually apply, the neonatal regenerative mechanisms into adulthood. To facilitate these studies, the last two decades have seen advancements in the development of injury models in adult mice representative of the diversity of cardiac diseases. Here, we provide an overview for a selection of the most common cardiac ischemic injury models and describe a set of methods used to accurately analyze and quantify cardiac outcomes. Importantly, a comprehensive understanding of cardiac regeneration and repair requires a combination of multiple functional, histological, and molecular analyses. Key words Cardiac regeneration, Cardiac function, Echocardiography, Cardiomyocyte proliferation, Scar quantification

1

Introduction Cardiovascular diseases are the leading cause of mortality worldwide. The magnitude and urgency of this unmet clinical need have engendered many scientific approaches that are fundamentally diverse, but unified in the goal of replenishing the damaged myocardium. These approaches may include stem cell-based procedures, trans-differentiation of cardiac fibroblasts, or dedifferentiation and proliferation of existing cardiomyocytes [1– 4]. Each approach between and within these categories varies in its success in promoting cardiac regeneration. However, despite the obvious importance of a clear standard for comparison between experimental methods, to date different injury models and quantification methods for heart functionality confound the direct comparison of the efficacy between different therapeutic approaches. Thus, the standardization of methods for injury and the assessment of the cardiac outcome are paramount for the communication between different fields of experimentation and inquiry underneath the umbrella of cardiac regeneration. Here, we take advantage of

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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our experience in cardiac regenerative biology to introduce the common injury models in mice and debate some of the most commonly used methods to accurately assess their outcomes. Although we attempted to include many cardiac regenerative readouts in this chapter, some were omitted, such as electrophysiological coupling, angiogenesis, and cell death. We also discuss the variability potentially introduced between studies of various mouse strains, sex, and a variety of parameters of cardiac functionality. 1.1

Cardiac Injuries

The different cardiac injury models for cardiomyocyte loss can be broadly divided into groups: nonischemic injuries, such as cryogenic [5, 6] genetic ablation [7, 8], and resection injuries [9–11], or ischemic injury [12–14]. Though these injury categories result in the loss of cardiac muscle and contribute to the study of cardiomyocyte injury responses, ischemic injury better mimics the human pathology [12]. Another category of injury, pressure overload, is also utilized extensively for studying cardiac hypertrophy, which includes pulmonary artery banding (PAB) [15, 16] and transverse aortic constriction (TAC) [16, 17]. The canonical TAC and PAB methods promote remodeling by increasing the pressure of the ventricles; over time this pressure induces adverse ventricular remodeling and emergence of pathologies, such as dilated cardiomyopathy and, eventually, heart failure [16, 17]. The prevalence of myocardial infarction-related mortality has precipitated a shift in high-impact publications towards the study of ischemic injury models, such as left anterior descending artery ligation (LAD ligation). Therefore, we also focus on the modeling of ischemic injury. To this end, we further separate ischemic injury models into permanent ligation and ischemia-reperfusion (I/R) models, though both may be achieved by the LAD ligation technique (described below). Both injuries share the primary outcome of the occlusion, i.e., the necrotic death of the myocardium and other cell types affected by the loss of blood supply, and the subsequent remodeling of the infarcted area [13, 14, 18]. The main differences between the injury models involve the secondary damages sustained by the release of the occlusion, the release permits the massive influx of detrimental factors causing reperfusion injury, which can expand the area of damage [18]. The full comprehension of these considerations underpins the planning of any in vivo cardiac experiment. Notably, nonischemic models may be efficiently and reproducibly employed for the study of post-injury cardiomyocyte proliferation, independent of other cell types. However, questions that require a systemic understanding of remodeling in the heart benefit from the utilization of ischemic models. Finally, the research of secondary injuries requires the I/R method; though this technique represents the most clinically relevant model, it also has the highest morbidity of any model heretofore addressed.

MI in Adult Mice

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Fig. 1 Differences in cardiac baseline function in adult mice. Serial echocardiographic measurements of ejection fraction (EF; panels a, d), fractional shortening (FS; panels b, e), and cardiac output (CO; panels c, f) of uninjured 12-week-old C57BL/6 (BL6) compared to ICR(CD1) and C57BL/6 (BL6) male-to-female mice, respectively 1.2 Assessing Cardiac Function in a High Variation Background

A crucial motivation for studying cardiac regeneration in adult mice is the ability to accurately and reliably measure outcomes of cardiac functionality. Recent publications have shed light on the variability between mouse strains and their regenerative potential, with emphasis on the endogenous proliferative and polyploid properties of cardiomyocytes [19, 20]. However, it is important to note that high variability of cardiac function may exist even within the same mouse line. The comparison of baseline cardiac function in two different mouse strains (C57BL/6 and ICR/CD1) yielded differences in ejection fraction, fractional shortening, and cardiac output (Fig. 1a–c), which demonstrates the variability, even in the absence of injury. In addition to the strain, mouse sex also contributes to variability in baseline cardiac function. For reasons yet unknown, cardiac baseline and response to ischemic injury differ in amplitude of variation between the male and female mice (Fig. 1d–f). In consideration of these issues, we adopted a methodology in which we assess cardiac function for mice prior to surgery to exclude outliers in baseline cardiac function that could skew the data postinjury.

1.3 Cell Cycle Markers as Proxies for Proliferation

The primary focus of the cardiac regeneration field has been the preferential replenishment of cardiomyocytes over scar tissue in the damaged myocardium [1, 13, 14]. The continued study of the latter hinges on the ability to accurately detect and analyze cardiomyocyte proliferation. To date, cardiomyocyte proliferation may be visualized in vitro by time-lapse microscopy of lineage-

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labeled cardiomyocytes [13]; however, the straightforward in vivo assessment of cardiomyocyte proliferation remains unresolved. A known attribute of the cardiomyocytes that hinders detection of proliferation is their ability to undergo “incomplete” cell division. Cardiomyocytes can undergo karyokinesis, duplication of the nucleus, without the subsequent cytokinesis that typically occurs during normal mitotic division [21]. Cardiomyocytes, therefore, are frequently multinucleated; thus, the cell cycle markers that measure nuclear duplication are not competent to discern between cell division and multinucleation [21]. Recent efforts in advancing methods to directly analyze cardiomyocyte proliferation in vivo resulted in the development of several genetic tools. The fluorescent ubiquitinated cell cycle indicator (FUCCI) system fuses specific cell cycle proteins to fluorescent proteins, thereby allowing direct visualization of cell cycle activity state. By implementing the FUCCI model, researchers demonstrated a proliferative peak of cardiomyocytes at postnatal day 2, whereas the vast majority of cardiomyocytes in adult mice are cell cycle arrested [22]. Recently, researchers implemented the single-cell fate mapping technique, mosaic analysis with double markers (MADM) in vivo, and verified low turnover of cardiomyocytes following adult myocardial infarction [23]. Other techniques involve lineage tracing of existing cardiomyocytes with the confetti system, and reported visualization of clonal expansion in vitro [24], as well as at regions bordering the infarct area (border zones) in vivo, following myocardial infarction [25]. In the absence of alternative markers, and due to the cumbersome, albeit elegant, nature of genetic approaches, we recommend the combination of cell cycle markers from various stages in the mitotic process. Examples of informative combinations may include Ki67 for cell cycle activity, phospho-histone H3 for mitosis, Aurora B kinase for cytokinesis [21], and cumulative markers, such as BrdU and nuclei counting. Taken together, these markers provide a more complete picture of cardiomyocyte cell cycle status. 1.4 Discrepancies Between Different Attributes

Our cumulative experience attests that a single parameter of cardiac function or quantification of injury often cannot accurately depict the regenerative response. The comprehensive understanding of cardiac injuries requires a combination of multiple functional and histological analyses (Fig. 2). Paradoxically, cardiac function and scar size do not necessarily correlate with each other. Though these parameters do correlate the majority of the time, we have, with some frequency, observed hearts with measurably dramatic scars not reflected by functional cardiac analyses (Fig. 3, mouse 5). Interestingly, this phenomenon is also common in human patients classed as “preserved ejection fraction heart failure” patients [26]. This highlights the need to employ multiple measurement

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Fig. 2 Methods to induce and accurately quantify cardiac injury and regeneration. Schematic representation of various parameters discussed in this chapter. Functional analysis, performed by echocardiography, measures different parameters of the heart, including ejection fraction, fractional shortening, stroke volume, and cardiac output, by calculating left ventricle diastole and systole volumes; histological and molecular analysis refers to CM proliferation and molecular markers of other cell types, as well as scar analysis at the morphological level; LV left ventricle, IR infarct region

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techniques to capture the most comprehensive snapshot of the heart following injury and treatment (Fig. 2).

2

Materials

2.1 Left Anterior Descending Artery Occlusion

1. Hair removal cream (see Note 1). 2. 12-week-old ICR(CD1) outbred female mice (see Note 2) of similar weight (ideally ~35 g). 3. Isoflurane induction chamber.

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4. Isoflurane intubation equipment, 20-gauge intravenous catheter with blunt-tip needle guide. 5. Styrofoam board. 6. Magnifying binocular with high illumination source. 7. Blunt-edge forceps. 8. Rodent ventilator. 9. Disinfection iodine solution. 10. Gauze pads. 11. Fine-tip forceps, 2
. 12. Surgical scissors, 2
. 13. Sternal retractor. 14. 8-0 Polypropylene suture (3/8 circle needle). 15. 5-0 Polypropylene suture (3/8 circle needle). 16. Needle holder, 2
. 17. 4% PFA. 2.2 Cardiac Function Assessment by Echocardiography

1. Fujifilm VisualSonics Vevo 3100 ultrasound machine and software. 2. Heating pad with embedded ECG leads. 3. Isoflurane induction chamber. 4. Isoflurane rodent face mask. 5. Ultrasound gel. 6. Electrode gel.

2.3 Histological Scar Quantification

1. Masson Trichrome-stained slides. 2. Slide scanner (or microscope containing tiling software). 3. ImageJ/FIJI software.

2.4 Immunofluorescence Analysis of Cardiomyocyte Proliferation

1. Slides of paraffin-embedded transverse heart sections. 2. Slide chambers for deparaffinization. 3. Xylene. 4. Absolute EtOH. 5. PBS. 6. EDTA antigen retrieval buffer. 7. Horse or goat serum. 8. Bovine serum albumin. 9. Triton X-100. 10. Antibodies: Anti-cTnT (1:200, Abcam, ab33589), anti-cTni (1:200, Abcam, ab47003), anti-Ki67 antibody (1:200, 275R, Cell Marque), anti-phosphorylated-histone 3 (pH 3) (1:200,

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SC-8656-R, Santa Cruz Biotechnology), and anti-Aurora B (Aim1, 1:100, 611,082, BD Transduction Laboratories). 11. 40 ,6-Diamidino-2-phenylindole (DAPI) solution. 12. Immu-mount mounting solution. 13. Confocal microscope for staining visualization. 14. ImageJ/FIJI software.

3

Methods

3.1 Left Anterior Descending Artery Occlusion, See Also [27] 3.1.1 One Day Prior to Surgery

1. Place the mice in the isoflurane induction chamber with 2–2.5% in oxygen to induce sleep. 2. Remove the mice from the chamber and apply the hair removal cream to the chest and neck area. Return the mice to the isoflurane induction chamber. 3. After 2–3 min, remove the hair by wiping the hair removal cream off with a damp gauze pad. Repeat these steps, as needed, until all hair is removed from the relevant areas. 4. Place the mice under the heat lamp until fully recovered, and then return to housing.

3.1.2 Preparation for Surgery

1. Ensure that all surgical equipment has been properly sterilized. 2. Prepare 100 μL per mouse of 2 mg/kg buprenorphine analgesia solution by diluting it with sterile PBS. 3. Ensure that the isoflurane vaporizer contains sufficient isoflurane for the procedure. 4. Tape a small 5-0 suture loop to the edge of the surgical styrofoam board, to be used for the intubation of the mouse.

3.1.3 Anesthesia and Intubation

1. Place the mouse in the induction chamber and close the lid tightly, ensuring that the isoflurane will affect the animal, but not disperse into the operating room. Turn on the isoflurane to 3% in oxygen. 2. Once the mouse has lost its righting reflex and enters deep sleep, transfer the mouse to the surgical styrofoam board in a supine position. With its head facing the surgeon, place the incisor teeth in the suture loop and move the tongue to the side. 3. Pull the tail of the mouse and position yourself to see the trachea prior to intubation. 4. Gently insert a 20-gauge IV blunt-tip catheter with lead into the trachea, remove the lead, and connect the catheter to the isoflurane line via the mouse ventilator (Harvard Apparatus) at 250 μL/stroke and 150–200 stroke/min (see Note 3).

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5. Rotate the work surface 180 and place tape on the catheter edge and on the limbs. 3.1.4 Surgery

1. Clean the chest and neck with a gauze pad soaked with iodine. 2. Cut the skin with scissors to visualize the area 2 mm left of the sternum at the third intercostal space. 3. With a second pair of scissors, make a small incision at that location and insert the rib retractor. Expand the retractor sufficiently to visualize the heart and fix it in place. 4. Under dissecting microscope magnification, use the bluntedge curved forceps to grab the edge of the pericardium, and pull on it gently to expose the LAD artery (Fig. 4). 5. Under dissecting microscope magnification, pass an 8-0 polypropylene suture with the needle holder through the muscle in close proximity to the LAD in the area between the first and second diagonal branching (Fig. 4). 6. Pull the suture through the muscle from the other side, leaving a small tail. Loop the suture wire three times around the forceps and grab the short tail. Pull until the knot surrounding the LAD is tight; after a few seconds, the area below the suture should appear pale. 7. If intramyocardial treatment administration is required, use the needle holder to create a 90 bend at the edge of the needle of a 30-gauge insulin syringe that contains up to 50 μL of the desired treatment. Insert the needle shallowly into the ventricular wall and inject treatment slowly. Proper administration can be visualized as a transient edema forming on the wall.

D1 Suture D2

Left anterior descending artery

Fig. 4 Area of occlusion in the left anterior descending artery. Schematic representation of the optimal suture placement on LAD (highlighted) between the first and second diagonal branches (D1 and D2, respectively)

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8. Remove the rib spreaders from the incision. 9. Pass the 5-0 polypropylene suture under the skin, through the second and the fourth intercostal spaces, and perform the triple knot (described in step 6). 10. Administer lidocaine gel topically to the incision sight. 11. Inject 100 μL (2 mg/kg) of buprenorphine subcutaneously for analgesia. 12. For permanent LAD ligation, please continue at step 21. 13. Stop administration of the isoflurane while maintaining the intubation with normal air; this should allow the mouse to regain spontaneous breathing. Gently press the toes with forceps to test for recovery of the pain response. 14. Carefully remove the intubation. 15. Place the mouse under heat lamp for 45–60 min for proper ischemic injury. 16. Repeat anesthesia and intubation steps (described above). 17. Clean the chest and neck with a gauze pad soaked in iodine. 18. Open the chest suture and insert the rib spreaders to expose the sutured LAD. 19. Under dissecting microscope magnification, use spring scissors to carefully cut the suture and remove it using blunt-edge forceps. 20. If intramyocardial treatment administration is required, see steps 7–9. 21. Place surgical glue on the closed incision area and place the skin above it with forceps. 22. Stop administration of isoflurane while maintaining the intubation with normal air; the mouse should regain bilateral breathing. Gently press the toes with forceps to see that the mouse regains the pain response. 23. Carefully remove the intubation. 24. Place the mouse under the heat lamp until full recovery from anesthesia. 25. Pain-relief medication, 100 μL (2 mg/kg) of buprenorphine, should be administered subcutaneously twice daily for 3 days. 26. LAD ligation, as described, may be utilized to induce heart failure for the purposes of chronic heart failure (CHF) modeling and experimentation. To generate reproducibly low-functioning hearts, perform the LAD ligation 4 weeks prior to the desired experiment or treatment start point.

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3.2 Cardiac Function Assessment by Echocardiography, See Also [28] 3.2.1 Preparation for Echocardiography

1. Remove the hair from the chest and neck area of the mice (as described in 3.1.1) the day before (see Note 4). 2. Confirm that the equipment is in working order and that all necessary materials are available before retrieving the mice to mitigate any avoidable stress for the animals. This includes assuring that the stage is preheated to 37  C, there is sufficient isoflurane in the induction system, adhesive tape is conveniently located and a destination cage is under the heat lamp. Once the procedure has begun, efficient workflow is paramount in achieving high reproducibility. 3. Bring the animals into the echocardiography room. 4. Place the mouse in the induction chamber and close the lid tightly to ensure that the isoflurane will affect the animal but will not disperse into the room. Turn on the isoflurane to 2–2.5% in oxygen. 5. Place electrode gel on the ECG electrodes. 6. Once the mouse falls asleep, immediately transfer it to the preheated stage with the nose inside the mask. Place adhesive tape over the mask to ensure that the mouse receives sufficient gas and change the stopcock position to the mask from the induction chamber. 7. With the mouse in a supine position, securely tape the legs and arms on the ECG leads and make sure that the software reads the pulse, ECG, and respiratory rate properly. 8. For correct and reproducible acquisition, maintain pulse between 400 and 500 BPM and isoflurane levels between 1.5 and 2%. 9. Apply warmed ultrasound gel to the depilated chest of the mouse, avoiding bubbles, which could affect the ultrasound measurement.

3.2.2 Parasternal Long-Axis Acquisition

1. Place the MX550D transducer at a 30 –45 angle, relative to midsagittal plane, and 30 to the transverse plane (Fig. 5), and lower it into close proximity to the chest while touching the gel. 2. While scanning, use the rail system to move the transducer until the tip of the apex and the aorta are horizontal (Fig. 6) (see Note 5). 3. Move the transducer slightly to either side to find the visual field of the widest part of the ventricle with the apex tip and the aorta still in view. 4. Collect multiple B-mode cine loops for analysis of functional parameters.

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Transducer

Isoflurane

Face mask Long axis

900 turn Short axis

ECG leads

Heat pad

Fig. 5 Mouse placement for echocardiography. Schematic representation of mouse positioning for echocardiography. The placements on the ECG leads are depicted in gray. The transducer is positioned at 30 relative to both transversal and midsagittal planes Parasternal long axis

Systole

Diastole

Ap

Ao

LV

Ap

LV

Ao

Parasternal short axis

M-mode Anterior wall outer edge Anterior wall inner edge

Anterior wall systole Left ventricle diastole

Posterior wall inner edge Posterior wall outer edge

Posterior wall systole

Anterior wall diastole Left ventricle systole Posterior wall diastole

Fig. 6 Long- and short-axis echocardiographic analysis. Example of the LV (left ventricle) tracing for long-axis analysis, demonstrating proper positioning of the heart in systole and diastole. The aorta (Ao) and apex (Ap) appear horizontal from each other (top panels). Depiction of a representative short-axis tracing in the M-mode (bottom panel)

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3.2.3 Short-Axis Acquisition

1. To move from long axis to short axis, rotate the transducer 90 clockwise. While scanning, adjust the orientation of the heart such that visual field contains the widest part of the ventricle that maintains a “closed circle” of the posterior and anterior walls and visualization of the papillary muscle (Fig. 6). Collect multiple B-mode cine loops as references for M-mode measurement. 2. Switch to M-mode and define a scanning area spanning across the entire middle of the ventricle, including the muscle walls. Collect multiple cine loops for size measurements (Fig. 6).

3.2.4 Analysis of the Long-Axis Echocardiographic Measurements B-Mode

1. Import the cine loops to the Vevo lab software. 2. Observe the cine loop multiple times before analysis. Choose at least three consecutive systole-diastole cycles that are representative of the heart and in which the diaphragm of the mouse is immobile. 3. Select either AutoLV for automated left ventricle tracing analysis or select the manual trace LV button. Move the video frame to peak systole and trace the left ventricle inner wall from aorta to apex, move frame by frame to the consecutive peak diastole, and perform the trace again (Fig. 6). We recommend exclusion of the papillary muscles, which might be visualized as part of the LV wall. 4. Repeat step 3 for three full consecutive cycles, and adjust the tracing for each frame. 5. Measurements for stroke volume, ejection fraction, fractional shortening, cardiac output, and volumes in systole and diastole should appear.

3.2.5 Analysis of the Short-Axis Echocardiographic Measurements M-Mode

1. Import the cine loops to the Vevo lab software. 2. Play the short-axis B-mode cine loop multiple times to verify the location of the ventricular walls and the lumen. 3. Open the M-mode cine loop and trace the outermost part of posterior ventricle wall for at least three consecutive cycles. Repeat the trace for the inner posterior ventricle wall, inner anterior wall, and outer anterior wall (Fig. 6). Replay the B-mode cine loop as needed to more easily visualize wall boundaries. 4. Calculate the measurements for left ventricular posterior and anterior wall diameter, as well as systole and diastole diameter. Additional parameters measured include stroke volume, ejection fraction, fractional shortening, and cardiac output; however, these parameters should be treated with caution if left ventricular asymmetry following injury occurs or if area of occlusion is below the imaged plane [29].

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3.3 Histological Scar Quantification

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Scar content may be evaluated directly by MRI with contrasting agents [30]; however, histological analysis is more common and requires no special equipment. Our lab has published three different techniques to quantify the scar content based on Masson’s trichrome staining, to be utilized at the discretion of the scientist based on the morphology of the heart and scar. These approaches include measurement of scar area, volume, and angular degree. For the quantitation of maximum area of damage, the single section with the highest fibrosis represents the maximal scar area. This method is useful in cases of incomplete or highly variable injuries in which the fibrotic area does not span to the apex, but rather is limited to area surrounding occlusion. The second technique measures the scar volume by calculating the area in all heart sections while also taking into consideration the area in between tissue sections (skips). Finally, in cases of complete occlusion of the LAD, after sufficient time, the scar may span from the point of occlusion down to the apex. Typically, this scenario results in a thin ventricular wall composed entirely of scar, and an enlarged ventricle lumen. Quantification of the scarred area in these cases may be biased, as the thinning of the ventricle walls results in a lower scar to left ventricle size ratio compared to hearts in which the scar occurs in the absence of thinning. As this bias could lead to a misrepresentation of the scar severity, we recommend measurement of scar angular degree rather than scar area in these cases (methods described below, Fig. 7).

Mask

Maximal scar area

0.4

0.4

0.4

i=1.6mm

Scar volume 0.4

Angular scar size

3600

Apex

3600

1830

1520

1150

Base

Fig. 7 Scar quantification methods. Three methods of quantification for scarring in the histological sections of an infarcted heart stained with Masson’s trichrome: maximal/average scar area, scar volume, and angular scar size

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3.3.1 Specimen Preparation and Image Acquisition

1. Fix the injured hearts in 4% PFA for 3 days on a shaker at 4  C. 2. Embed the hearts in paraffin to facilitate the cutting of transverse sections (step 3), according to standard protocol (see Note 6). 3. Acquire 4–7 μm of serial sections of the heart with a microtome, with skips of 300–400 μm between each series to achieve full coverage of the heart within 1 slide. 4. Stain the slides for Masson’s trichrome according to standard protocol. 5. Acquire images of all stained sections of the heart from the area of damage to the apex with a slide scanner (Panoramic SCAN, 3dHisTech) or microscope with tiling software.

3.3.2 Maximum Damage Area Analysis

1. Identify the section in which the damage is greatest; this is visualized as the largest area containing fibrotic blue stain. 2. Using ImageJ/FIJI software, trace the blue area indicative of the injury by hand or with one of the automated tracing plugins available online. Measure the pixel area. 3. Trace the LV outer wall comprised of the anterior and posterior walls, and measure the pixel area. 4. Trace the lumen of the LV and disregard the papillary muscles. 5. Calculate maximum damage area percent by Scar area
100 ¼ Scar area outer LV area  inner LV area

3.3.3 Scar Volume Analysis

1. Define the proper scale units (from pixels to μm) in ImageJ/ FIJI. 2. Using the ImageJ/FIJI software, either trace the injured blue area by hand or use one of the automated tracing plug-ins available online and measure the area (as in step 2, above). Repeat for all sections containing damaged area. 3. Calculate scar volume according to the truncated pyramid formula, where a1 and a2 are scar areas (as calculated above) of two consecutive serial sections and i is the section interval (0.3–0.4 mm) between them: Scar volume ¼

3.3.4 Scar Degree Analysis

pffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi i
ða 1 þ a 2 þ a 1
a 2 Þ 3

1. Using the ImageJ/FIJI software, mark two lines from the middle of the LV lumen to the edges of the scar and measure the angle between them. 2. Repeat the measurement for every section-containing scar.

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3. Average the measurements between sections to determine the angular scar size. 3.4 Immunofluorescence Analysis of Cardiomyocyte Proliferation 3.4.1 Staining

1. Fix the injured hearts in 4% PFA for 3 days on a shaker at 4  C. 2. Embed the hearts in paraffin to facilitate the cutting of transverse sections (step 3), according to standard protocol (see Note 6). 3. Acquire 4–7 μm of serial sections of the heart with a microtome, with skips of 300–400 μm between each series to achieve full coverage of the heart within one slide. 4. Place the slides in a slide rack that is solvent resistant and microwave compatible. 5. Deparaffinize and rehydrate the slides at room temperature by immersing them in a staining jar containing the following: 100% xylene—10 min; 100% xylene—10 min; 100% EtOH— 10 min; 90% EtOH in dH2O—5 min; 70% EtOH in dH2O— 5 min; 50% EtOH in dH2O—5 min; 30% EtOH in dH2O— 5 min; dH2O—5 min. 6. Place the slides in the staining jar containing a sodium citrate buffer (10 mM sodium citrate acid, 0.05% Tween 20, pH 6.0) filled to the top. 7. Place the staining jar in the microwave, run at maximum power until boiling begins, then reduce the power, and continue microwaving for 15 min. The volume of buffer must not evaporate below the level of the tissue. 8. Cool the slides to room temperature for ~1 h. With the exception of the primary antibody incubation, from this point forward the slides and buffers can remain at room temperature. 9. Wash the slides with one staining jar volume of PBS. 10. Place the slides in a jar containing a permeabilization buffer (0.5% Triton X-100 in PBS) for 15 min. 11. Move the slides into blocking solution (0.1% Triton X-100, 3% BSA, 1% fetal bovine serum, or 10% horse serum) for 1 h. 12. Remove the slide from blocking buffer and carefully remove excess liquid with a KimWipe, though the tissue should not dry completely. 13. Draw a barrier surrounding the desired tissue area with a hydrophobic pen. 14. Add a sufficient amount of primary antibody to cover the selected tissue area. Typically 250 μL of primary antibodies (diluted 1:200 in blocking solution) is sufficient to cover four sections. 15. Create a humidity chamber to avoid evaporation of the primary antibody by lining the bottom of a slide box with water-soaked

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paper towels. Place the slides horizontally in the box such that they are elevated over the paper towels. Cover the slides with parafilm to prevent evaporation and disperse the solution equally. Incubate the slides in the primary antibody overnight at 4  C. 16. Equilibrate the humid chamber to room temperature for 1 h. 17. Wash the slides in PBS to remove the parafilm cover. If the parafilm resists removal, separate the parafilm from the slide by shaking the slide gently or using forceps. 18. Place the slides back a in slide rack, and wash with PBS three times for 5 min each. 19. Remove the slides from the washes and carefully remove excess liquid around the tissue with a KimWipe (as in step 12). 20. Reinforce the border of the hydrophobic pen, if needed, and add sufficient solution (250 μL) of fluorophore-conjugated secondary antibodies diluted in PBS (1:200). Place the slides in the humidity chamber, cover them with parafilm, and incubate for 1–2 h at room temperature in the dark to protect the fluorophores (see Note 7). 21. Wash the slides in PBS to remove the parafilm cover (as in step 17). 22. Place the slides back in a slide rack and wash with PBS three times for 5 min each. 23. Remove the slides from PBS, place on paper to dry, and carefully remove excess liquid with a KimWipe. 24. Mount the slides with mounting media containing DAPI and place a coverslip over the tissue. 25. Place the slides in dry, dark slide box for at least 1 day prior to imaging to dry the mounting media. Slides may be stored at 4  C for several days. 3.4.2 Imaging and Analysis

1. Visualize the stained slide by confocal microscope and identify the section with the largest infarct region. 2. Use tiling software to acquire images of the entire section in all relevant channels. 3. The area negative for all cardiomyocyte markers is the infarct region. Measure 400 μm around the infarct region and define this area as the “border zone” (Fig. 8). 4. Count the number of total cardiomyocytes (CMs) in the border zone. The percentage of proliferation is the number of CMs positive for proliferation markers over the total. 5. If relevant, repeat the counting for the total number of CMs and the number of CMs positive for proliferation markers in the remote region, and calculate the percent of proliferation.

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Border zone (BZ)

Infarct Region (IR)

IR Border zone (BZ)

19

BZ

BZ LV

Left ventricle

Remote region

cTnT

Fig. 8 Segmentation of left ventricle preceding quantification of cardiomyocyte proliferation. Schematic segmentation of left ventricle infarct region, border zone, and remote region (left panel). Segmentation of a 12-week-old mouse heart, 4 days following injury (right panel). The loss or reduction of cTnT immunofluorescence staining signal informs the designation of the infarct region

4

Notes 1. We used a local brand of hair removal cream containing sodium thioglycolate. Different hair removal products may be tested for efficacy. 2. After analysis of several mouse lines, we determined that ICR (CD1) females showed superior survivability following injury and most reproducible baseline parameters. Thus, we preferentially utilize this line and sex when compatible with our work. 3. Proper intubation is necessary for the surgeries described. If a normal breathing pattern of chest rising and falling cannot be observed following insertion of the catheter into the trachea, remove the catheter and attempt intubation again. 4. Complete hair removal is crucial prior to echocardiography because obstructions in the sonogram path obscure readings. However, as prolonged exposure to isoflurane affects measurement of cardiac function, we recommend removing the hair the day before acquisition of cardiac functionality to minimize exposure to isoflurane in any given day. 5. Though consistent positioning and acquisition are crucial for reproducible results, the ligation injury may result in changes in morphology, adherence of external tissues, and tilting that cause shadows. Several attempts may be required to obtain good images.

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6. Cutting the heart transversely in half and embedding both parts together allows for maximal coverage while reducing the number of sections on the slide. 7. Cardiomyocytes are autofluorescent in green wavelengths; this should be considered when selecting fluorophores. References 1. Tzahor E, Poss KD (2017) Cardiac regeneration strategies: staying young at heart. Science 356(6342):1035–1039. https://doi.org/10. 1126/science.aam5894 2. Pomeroy JE, Helfer A, Bursac N (2019) Biomaterializing the promise of cardiac tissue engineering. Biotechnol Adv. https://doi. org/10.1016/j.biotechadv.2019.02.009 3. Heallen Todd R, Kadow Zachary A, Kim Jong H, Wang J, Martin James F (2019) Stimulating cardiogenesis as a treatment for heart failure. Circ Res 124(11):1647–1657. https:// doi.org/10.1161/CIRCRESAHA.118. 313573 4. Hashimoto H, Olson EN, Bassel-Duby R (2018) Therapeutic approaches for cardiac regeneration and repair. Nat Rev Cardiol 15 (10):585–600. https://doi.org/10.1038/ s41569-018-0036-6 5. Gonza´lez-Rosa JM, Martı´n V, Peralta M, Torres M, Mercader N (2011) Extensive scar formation and regression during heart regeneration after cryoinjury in zebrafish. Development 138(9):1663–1674. https://doi.org/ 10.1242/dev.060897 6. van den Bos EJ, Mees BME, de Waard MC, de Crom R, Duncker DJ (2005) A novel model of cryoinjury-induced myocardial infarction in the mouse: a comparison with coronary artery ligation. Am J Phys Heart Circ Phys 289(3): H1291–H1300. https://doi.org/10.1152/ ajpheart.00111.2005 7. Wang J, Pana´kova´ D, Kikuchi K, Holdway JE, Gemberling M, Burris JS, Singh SP, Dickson AL, Lin Y-F, Sabeh MK, Werdich AA, Yelon D, MacRae CA, Poss KD (2011) The regenerative capacity of zebrafish reverses cardiac failure caused by genetic cardiomyocyte depletion. Development 138(16):3421–3430. https:// doi.org/10.1242/dev.068601 8. Akazawa H, Komazaki S, Shimomura H, Terasaki F, Zou Y, Takano H, Nagai T, Komuro I (2004) Diphtheria toxin-induced autophagic cardiomyocyte death plays a pathogenic role in mouse model of heart failure. J Biol Chem 279(39):41095–41103. https:// doi.org/10.1074/jbc.M313084200

9. Poss KD, Wilson LG, Keating MT (2002) Heart regeneration in zebrafish. Science 298 (5601):2188–2190. https://doi.org/10. 1126/science.1077857 10. Porrello ER, Mahmoud AI, Simpson E, Hill JA, Richardson JA, Olson EN, Sadek HA (2011) Transient regenerative potential of the neonatal mouse heart. Science 331 (6020):1078–1080. https://doi.org/10. 1126/science.1200708 11. Sarig R, Rimmer R, Bassat E, Zhang L, Umansky KB, Lendengolts D, Perlmoter G, Yaniv K, Tzahor E (2019) Transient p53-mediated regenerative senescence in the injured heart. Circulation 139 (21):2491–2494. https://doi.org/10.1161/ CIRCULATIONAHA.119.040125 12. Porrello ER, Mahmoud AI, Simpson E, Johnson BA, Grinsfelder D, Canseco D, Mammen PP, Rothermel BA, Olson EN, Sadek HA (2013) Regulation of neonatal and adult mammalian heart regeneration by the miR-15 family. Proc Natl Acad Sci U S A 110(1):187–192. https://doi.org/10.1073/pnas.1208863110 13. D’Uva G, Aharonov A, Lauriola M, Kain D, Yahalom-Ronen Y, Carvalho S, Weisinger K, Bassat E, Rajchman D, Yifa O, Lysenko M, Konfino T, Hegesh J, Brenner O, Neeman M, Yarden Y, Leor J, Sarig R, Harvey RP, Tzahor E (2015) ERBB2 triggers mammalian heart regeneration by promoting cardiomyocyte dedifferentiation and proliferation. Nat Cell Biol 17(5):627–638. https://doi.org/10. 1038/ncb3149. http://www.nature.com/ ncb/journal/v17/n5/abs/ncb3149. html#supplementary-information 14. Bassat E, Mutlak YE, Genzelinakh A, Shadrin IY, Baruch Umansky K, Yifa O, Kain D, Rajchman D, Leach J, Riabov Bassat D, Udi Y, Sarig R, Sagi I, Martin JF, Bursac N, Cohen S, Tzahor E (2017) The extracellular matrix protein agrin promotes heart regeneration in mice. Nature 547(7662):179–184 15. Urashima T, Zhao M, Wagner R, Fajardo G, Farahani S, Quertermous T, Bernstein D (2008) Molecular and physiological characterization of RV remodeling in a murine model of pulmonary stenosis. Am J Physiol Heart Circ

MI in Adult Mice Physiol 295(3):H1351–H1368. https://doi. org/10.1152/ajpheart.91526.2007 16. Tarnavski O, McMullen JR, Schinke M, Nie Q, Kong S, Izumo S (2004) Mouse cardiac surgery: comprehensive techniques for the generation of mouse models of human diseases and their application for genomic studies. Physiol Genomics 16(3):349–360. https://doi.org/ 10.1152/physiolgenomics.00041.2003 17. deAlmeida AC, van Oort RJ, Wehrens XHT (2010) Transverse aortic constriction in mice. J Vis Exp 38:1729. https://doi.org/10.3791/ 1729 18. Lindsey ML, Bolli R, Canty JM Jr, Du X-J, Frangogiannis NG, Frantz S, Gourdie RG, Holmes JW, Jones SP, Kloner RA, Lefer DJ, Liao R, Murphy E, Ping P, Przyklenk K, Recchia FA, Longacre LS, Ripplinger CM, Eyk JEV, Heusch G (2018) Guidelines for experimental models of myocardial ischemia and infarction. Am J Phys Heart Circ Phys 314(4):H812–H838. https://doi.org/10. 1152/ajpheart.00335.2017 19. Patterson M, Barske L, Van Handel B, Rau CD, Gan P, Sharma A, Parikh S, Denholtz M, Huang Y, Yamaguchi Y, Shen H, Allayee H, Crump JG, Force TI, Lien C-L, Makita T, Lusis AJ, Kumar SR, Sucov HM (2017) Frequency of mononuclear diploid cardiomyocytes underlies natural variation in heart regeneration. Nat Genet 49:1346. https:// doi.org/10.1038/ng.3929. https://www. nature.com/articles/ng.3929#supplemen tary-information 20. Gonza´lez-Rosa JM, Sharpe M, Field D, Soonpaa MH, Field LJ, Burns CE, Burns CG (2018) Myocardial polyploidization creates a barrier to heart regeneration in zebrafish. Developmental Cell 44(4):433–446.e437. https://doi.org/ 10.1016/j.devcel.2018.01.021 21. Hesse M, Doengi M, Becker A, Kimura K, Voeltz N, Stein V, Fleischmann Bernd K (2018) Midbody positioning and distance between daughter nuclei enable unequivocal identification of cardiomyocyte cell division in mice. Circ Res 123(9):1039–1052. https:// doi.org/10.1161/CIRCRESAHA.118. 312792 22. Alvarez R, Wang BJ, Quijada PJ, Avitabile D, Ho T, Shaitrit M, Chavarria M, Firouzi F, Ebeid D, Monsanto MM, Navarrete N, Moshref M, Siddiqi S, Broughton KM, Bailey BA, Gude NA, Sussman MA (2019) Cardiomyocyte cell cycle dynamics and proliferation revealed through cardiac-specific transgenesis of fluorescent ubiquitinated cell cycle indicator (FUCCI). J Mol Cell Cardiol 127:154–164. https://doi.org/10.1016/j.yjmcc.2018.12. 007

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Chapter 2 Apical Resection and Cryoinjury of Neonatal Mouse Heart Hua Shen, Ali Darehzereshki, Henry M. Sucov, and Ching-Ling Lien Abstract Neonatal mouse hearts have a regenerative capacity similar to adult zebrafish. Different cardiac injury models have been established to investigate the regenerative capacity of neonatal mouse hearts, including ventricular amputation, cryoinjury, and ligation of a major coronary artery. While the ventricular resection model can be utilized to study how tissue forms and regenerates de novo, cryoinjury and coronary artery ligation are methods that might better mimic myocardial infarction by creating tissue damage and necrosis as opposed to the removal of healthy tissue in the ventricular amputation model. Here we describe methods of creating ventricular resection and cardiac cryoinjury in newborn mice. Key words Ventricular apical resection, Cryoinjury, Neonatal mouse, Cardiomyocyte proliferation, Scar formation

1

Introduction Adult zebrafish can fully regenerate their hearts after cardiac injuries created by amputating 20% of the ventricle [1]. In 2011, Porrello and colleagues described an apical amputation model in neonatal mouse hearts that is analogous to the procedure in adult zebrafish [2]. The results of this study indicated that not only neonatal mice survive the cardiac amputation, but also myocardial regeneration without scar formation was observed when such injury happened within 24 h after birth. This outcome has been observed by different independent groups [3–8], although it should be noted that scar formation was also reported in neonatal mouse hearts after amputation [9, 10]. The degree of scar formation may be a function of the specific technique that was applied and the amount of myocardium resected [11]. Regardless, there is consensus that this capacity is lost by 7 days after birth. Apical resection creates myocardial damage by removing approximately 10–20% of healthy ventricular tissue. The missing apex will be reconstructed and restored to the normal apex morphology after the regeneration process of 21 days. Subsequent

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_2, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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histological studies revealed that cardiomyocyte proliferation and revascularization occur during neonatal mouse heart regeneration [2]. Furthermore, acute inflammation and macrophages were shown to stimulate the regenerative response after ventricular resection in neonatal hearts [4, 12]. From a pathophysiology standpoint, an injury created after ventricular resection is different from an injury induced by myocardial infarction. Other injury models that are more similar to myocardial infarction (e.g., coronary artery ligation) have been used in adult mice or larger mammals, but this is less feasible in zebrafish or neonatal mice due to small size and less obvious coronary vasculature [13, 14]. A cryoinjury model was established for adult zebrafish to create heart injuries to mimic the pathogenesis of myocardial infarction [15–17]. To compare the molecular mechanisms of cardiac regeneration in adult zebrafish and neonatal mice, we also developed a cryoinjury model using a cold probe to injure the left ventricle over the area that is commonly perfused by the left anterior descending (LAD) coronary artery. Newborn neonatal mice tolerate this injury with a 90–100% survival rate, depending on the severity of the injury. Subsequent histological studies and echocardiographic imaging can confirm the degree of injury as well as regeneration vs. repair mechanisms; large transmural cryoinjury induced scar formation [18]. Here, we describe the surgical procedure of ventricular resection and cardiac cryoinjury in neonatal mice since we have utilized both in our studies.

2 2.1

Materials Equipment

1. Ice pack. 2. Ice bed. 3. 70% Ethanol. 4. Sterile gloves. 5. Micro-fine-tip tweezers 2 (Fine Science Tools, Dumont #5). 6. Iridectomy scissors (straight or curved tips can be used; curved tips recommended for apical resection) (straight: Biomedical Research Instruments, 11-1020; curved: Natume, MB-54-3). 7. Fine needle holder (Sansyo 701-16 cm). 8. 7-0 Prolene suture (Ethicon, USA, Cat# 8701H). 9. Liquid nitrogen. 10. Metal probe (1.2 mm in diameter with long handle) (FST, Cat#10088-15). 11. Topical tissue adhesive (surgical glue) (VetBond™ Tissue Adhesive, Santa Cruz, SC-361931).

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Fig. 1 Surgical tools used in this procedure. (Left to right): Fine needle holder, iridectomy scissors (straight tips), iridectomy scissors (curved tips), micro-finetip tweezers 2

12. Surgical tape. 13. Sterile gauze. 14. Light source. 15. Warm pad. 16. Warm lamp. 17. Microtome (Leica). 18. Cryostat (Leica). 19. Light microscope with imaging system. 20. Fluorescent microscope with imaging system. Note: Instrument Preparation Before and Between Procedures 1. Prior to surgery, autoclave the wrapped or packaged instruments (Fig. 1) to sterilize. 2. During the surgery, while switching between neonates, place the tips of the instruments into the hot-bead sterilizer for ~5–10 s before moving onto the next neonates. 2.2

Mice

1. This injury model can be used for neonatal mice younger than 1 week. The day of birth is considered as postnatal day (P) 0. Apical resection and cryoinjury are mainly performed on P1 pups. 2. Before delivery, separate the pregnant female from the stud mouse to avoid cannibalization by the stud male after injury.

2.3

Reagents

Trichrome stain kit (Millipore Sigma, HT15).

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2.3.1 Picro-Sirius Red Staining

1. Sirius Red (Direct Red 80, Sigma-Aldrich Cat#365548). 2. Saturated aqueous solution of picric acid (Sigma, Cat# P67441GA). 3. Acetic acid (glacial) for acidified water (Sigma-Aldrich A6283).

2.3.2 AFOG Staining

1. Acid Fuchsin (Sigma, Cat# F8129). 2. Orange G (Sigma, Cat# 861286). 3. Aniline Blue (Sigma, Cat#415049). 3 g of Acid Fuchsin, 2 g of Orange G, 1 g of Aniline Blue dissolved in 200 mL of acidified distilled water, pH 1.09.

2.3.3 Cardiomyocyte Cell Cycle Assay by Immunostaining

1. Anti-NKX2.5 antibody (Santa Cruz SC-8697). 2. Anti-Phospho-Histone H3 antibody (pH 3) (Upstate 06-570). 3. Alexa Fluor donkey anti-goat 647 (Thermo Fisher A-21447). 4. Alexa Fluor donkey anti-rabbit 546 (Thermo Fisher A10040). 5. ProLong Antifade mounting medium with DAPI (Thermo Fisher P36931).

3

Methods (Procedure Outlined in Fig. 2, See Note 1)

3.1 Prepare the Pups for Surgery and Anesthesia by Hypothermia

1. Before the procedure, disinfect the procedure area with 70% ethanol. Sterilized instruments should be used for the entire procedure. 2. Set up the ice bed, light source, warm lamp, and sterilized surgical tools in the procedure area. 3. Right before surgery, remove all pups from the nursing dam and transfer them to a new cage. Keep all pups on a warm pad. Keep the nursing dam in the original cage. 4. Place one pup on the ice pack; use a Petri dish and other proper material (e.g., surgical gloves) to keep the mouse skin from contacting the ice. Hypothermia induces anesthesia in mouse pups within 5 min. 5. Transfer the anesthetized pup onto a flat ice bed. Place the pup in the supine position with the tail on the surgeon’s side. Use surgical tape to restrain the fore- and hind- limbs and the tail onto a Petri dish. Forelimbs need to be wide open for optimal chest exposure. 6. Wear sterile gloves. 7. Clean and prep the chest skin with 70% ethanol by gently scrubbing several times. 8. Using iridectomy scissors to make an ~1 cm transverse incision over the left side of chest at the armpit level. Hold the skin with

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3.4 Suture and postoperative care Fig. 2 Surgical procedures outlined for ventricular resection and cryoinjury of neonatal mouse hearts. 3.2 and 3.3 in flowchart were illustrated by Xiyu Erika Wang

tweezers to elevate it from the underlying muscle to facilitate the skin incision. 3.2 Ventricular Resection

1. Use tweezers to bluntly separate the intercostal muscles at fifth intercostal space to expose the chest cavity. 2. Use your fingers to gently press the abdomen to force the heart out of the chest cavity. The heart normally stands with the apex up. 3. Resect the apex with angled iridectomy scissors until the chamber of the left ventricle is exposed. Confirm that the left ventricle is exposed by initial bleeding. Use gauze to completely remove any blood. This will improve the visualization to reassure adequate resection. Blood clot usually takes some time to form; once it is formed, do not remove it. Sham-operated mice have their hearts exposed without ventricular resection.

3.3

Cryoinjury

1. Use tweezers to bluntly dissect the intercostal muscles at fifth intercostal space until the chest cavity can be accessed. 2. Use your fingers to gently press pup’s torso to force the heart out of the thoracic cavity. The heart normally stands with the apex up.

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3. Have the surgical assistant place a 2 mm metal probe in liquid nitrogen to chill for 30 s, while the surgeon is exposing the heart. 4. Once heart is exposed outside of the thoracic cavity, apply the pre-chilled probe to the left ventricular anterolateral wall to create an injury. Sham-operated hearts are only exposed without probe application (see Note 2). A drop of cold sterile saline can be used on the freezing probe to ease the separation of the probe from the frozen myocardium and avoid tissue rupture. Avoid probe sticking to the heart by not pre-chilling the probe in the liquid nitrogen for too long. 3.4 Suture and Postoperative Care

1. Close the chest by suturing using 7-0 Prolene, including the upper and lower ribs. Check for bleeding to reassure hemostasis and wipe any remaining blood from the skin with sterile gauze. Bleeding can be stopped by applying gentle pressure for few seconds. This is rarely needed. Reapproximate the skin edges and seal the wound by surgical glue. 2. Once the glue is dry, remove the tape and transfer the pup to a warm pad for recovery for 5–10 min. Spontaneous movements, urination, and change in skin color from pale to pink indicate appropriate recovery. 3. Transfer the recovered pups to the littermate cage after confirming their normal body temperature by touching their skin. Keep the pup with their littermates under a warm lamp until surgery is completed for the entire litter. 4. Return the entire litter of pups with bedding to the mother’s cage after the procedure is completed (see Note 3). Monitor the cage for maternal acceptance (see Note 4). Carrying and transferring the pups could be a sign of maternal acceptance (see Notes 5 and 6).

3.5 Histological and Proliferation Analysis 3.5.1 Histological Analysis

1. At different time points (e.g., 2, 3, 7, 14, and 21 days postinjury), sacrifice the pups to collect the hearts for histological analysis (Fig. 3). Fix heart in 4% PFA overnight, dehydrate by gradient ethanol, and then bring to paraffin. 2. Embed the neonatal hearts in paraffin or OCT (for proliferation assay) and perform either frontal or transverse cryo or paffin-sectioning at 8 mm thickness. 3. Stain the heart sections with H&E to examine the morphology and injury severities. Trichrome [2], Picro-Sirius Red, or AFOG staining [18] can be utilized to identify collagen scar tissues (Fig. 4) (see Notes 7 and 8).

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Fig. 3 Histological analysis of neonatal mouse hearts after injuries. (a) Example of heats at 2 days postventricular resection (2dpa). Trichrome staining of paraffin-embedded heart section. Ventricular resection is performed on P1 neonatal mice heart, and heart sections are stained on day 2 after ventricular resection. P3 shows normal heart morphology. 2dpa shows the morphology after resection. (b) Example of hearts 3 days post-cryoinjury and H&E staining of paraffin-embedded heart section

Fig. 4 Examining heart regeneration and scar formation by trichrome, Picro-Sirius Red and AFOG staining. Example of trichrome, Picro-Sirius Red, and AFOG staining of paraffin-embedded heart section. Shamoperated control is stained with trichrome. Ventricular resection is performed on wild-type (WT) and mutant P1 neonatal mice hearts, and heart sections are stained on 21 dpa with Picro-Sirius Red staining. Collagen is in red. Red color at apex indicates scar formation. Mild and severe cryoinjuries [18] were performed on P1 neonatal pups and analyzed at 21 days after mild (dpmc) and severe (dpsc) cryoinjuries with AFOG staining. For trichrome and Picro-Sirius Red staining, nuclei are stained with Weigert’s hematoxylin 3.5.2 Proliferation Analysis

Co-staining of phospho-histone 3 (pH3) and a cardiomyocyte nuclear marker can be used for detecting cardiomyocyte proliferation. Here, we show the representative images of pH 3 immunofluorescence staining of OCT-embedded cryosections of hearts within first week after apical resection (Fig. 5) [19].

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Fig. 5 Quantification of cardiomyocyte M-phase activity. Example of proliferation analysis with immunofluorescence at day 7 after ventricular resection on OCT-embedded wild-type heart section. Ventricular resection was performed on P1 neonatal mice heart. Phospho-histone 3 is in white color. Nkx2.5 in red color is used to label cardiomyocyte nuclei. Cardiomyocytes are in green and DAPI is in blue. White arrows indicate co-staining of pH 3 and NKX2.5. Gray two-way arrow indicates border zone, which is a fainter green. CM indicates cardiomyocytes. pH 3 and Nkx2.5 co-staining indicates cardiac cells in M-phase. In sham heart at postnatal day 8 (P8), pH 3/Nkx2.5 double-positive cardiomyocytes are barely detectable, and more pH 3/ Nkx2.5 double-positive cardiomyocytes are detectable in 7dpa heart

4

Notes 1. The whole procedure for one pup usually takes 10–15 min for experienced researchers. 2. The severity of cryoinjury can be controlled by placing the cryoprobe on the left ventricular anterolateral wall for different lengths of time, 5 s in order to create a transmural cryoinjury and 1 s to create a non-transmural cryoinjury. 3. Minimize the time that the entire litter of pups is away from nursing mother. 4. A small amount of vanilla extract can be introduced to the nursing mother’s cage the night before surgery and onto recovered pups before returning them to the mother’s cage. This will often increase the acceptance by the nursing mother, presumably by the vanilla masking the blood smell to allow the pups to be more easily reacquainted with the mother. 5. The pups can be transferred to a surrogate mother if the nursing mother does not accept them after the operation.

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6. The presence of milk in the stomach (visible through the skin) on postoperative day 1 confirms acceptance by the nursing mother. 7. Due to the autofluorescence of cardiomyocytes in the adult heart, the section can be previewed under the green channel before staining. Normal cardiomyocytes are visible under the green channel, and the scar area is fainter or invisible. 8. Differential scar formation and heart regeneration can be observed after mild vs. severe cryoinjury [18]. References 1. Poss KD, Wilson LG, Keating MT (2002) Heart regeneration in zebrafish. Science 298 (5601):2188–2190 2. Porrello ER, Mahmoud AI, Simpson E, Hill JA, Richardson JA, Olson EN, Sadek HA (2011) Transient regenerative potential of the neonatal mouse heart. Science 331 (6020):1078–1080. https://doi.org/10. 1126/science.1200708. 331/6020/1078 [pii] 3. Bassat E, Mutlak YE, Genzelinakh A, Shadrin IY, Baruch Umansky K, Yifa O, Kain D, Rajchman D, Leach J, Riabov Bassat D, Udi Y, Sarig R, Sagi I, Martin JF, Bursac N, Cohen S, Tzahor E (2017) The extracellular matrix protein agrin promotes heart regeneration in mice. Nature 547(7662):179–184. https://doi.org/10.1038/nature22978 4. Han C, Nie Y, Lian H, Liu R, He F, Huang H, Hu S (2015) Acute inflammation stimulates a regenerative response in the neonatal mouse heart. Cell Res 25(10):1137–1151. https:// doi.org/10.1038/cr.2015.110 5. Morikawa Y, Heallen T, Leach J, Xiao Y, Martin JF (2017) Dystrophin-glycoprotein complex sequesters Yap to inhibit cardiomyocyte proliferation. Nature 547(7662):227–231. https://doi.org/10.1038/nature22979 6. Morikawa Y, Zhang M, Heallen T, Leach J, Tao G, Xiao Y, Bai Y, Li W, Willerson JT, Martin JF (2015) Actin cytoskeletal remodeling with protrusion formation is essential for heart regeneration in Hippo-deficient mice. Sci Signal 8(375):ra41. https://doi.org/10. 1126/scisignal.2005781 7. Tao G, Kahr PC, Morikawa Y, Zhang M, Rahmani M, Heallen TR, Li L, Sun Z, Olson EN, Amendt BA, Martin JF (2016) Pitx2 promotes heart repair by activating the antioxidant response after cardiac injury. Nature 534

(7605):119–123. https://doi.org/10.1038/ nature17959 8. Yu W, Huang X, Tian X, Zhang H, He L, Wang Y, Nie Y, Hu S, Lin Z, Zhou B, Pu W, Lui KO, Zhou B (2016) GATA4 regulates Fgf16 to promote heart repair after injury. Development 143(6):936–949. https://doi. org/10.1242/dev.130971 9. Andersen DC, Ganesalingam S, Jensen CH, Sheikh SP (2014) Do neonatal mouse hearts regenerate following heart apex resection? Stem Cell Rep 2(4):406–413. https://doi. org/10.1016/j.stemcr.2014.02.008 10. Sampaio-Pinto V, Rodrigues SC, Laundos TL, Silva ED, Vasques-Novoa F, Silva AC, Cerqueira RJ, Resende TP, Pianca N, LeiteMoreira A, D’Uva G, Thorsteinsdottir S, Pinto-do OP, Nascimento DS (2018) Neonatal apex resection triggers cardiomyocyte proliferation, neovascularization and functional recovery despite local fibrosis. Stem Cell Rep 10 (3):860–874. https://doi.org/10.1016/j. stemcr.2018.01.042 11. Bryant DM, O’Meara CC, Ho NN, Gannon J, Cai L, Lee RT (2015) A systematic analysis of neonatal mouse heart regeneration after apical resection. J Mol Cell Cardiol 79:315–318. https://doi.org/10.1016/j.yjmcc.2014.12. 011 12. Aurora AB, Porrello ER, Tan W, Mahmoud AI, Hill JA, Bassel-Duby R, Sadek HA, Olson EN (2014) Macrophages are required for neonatal heart regeneration. J Clin Invest 124 (3):1382–1392. https://doi.org/10.1172/ JCI72181 13. Gamba L, Harrison M, Lien CL (2014) Cardiac regeneration in model organisms. Curr Treat Options Cardiovasc Med 16(3):288. https:// doi.org/10.1007/s11936-013-0288-8

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14. Rubin N, Harrison MR, Krainock M, Kim R, Lien CL (2016) Recent advancements in understanding endogenous heart regeneration-insights from adult zebrafish and neonatal mice. Semin Cell Dev Biol 58:34–40. https://doi.org/10.1016/j.semcdb.2016.04. 011 15. Chablais F, Jazwinska A (2010) IGF signaling between blastema and wound epidermis is required for fin regeneration. Development 137(6):871–879. https://doi.org/10.1242/ dev.043885. 137/6/871 [pii] 16. Gonzalez-Rosa JM, Martin V, Peralta M, Torres M, Mercader N (2011) Extensive scar formation and regression during heart regeneration after cryoinjury in zebrafish. Development 138(9):1663–1674. https://doi.org/ 10.1242/dev.060897. dev.060897 [pii] 17. Schnabel K, Wu CC, Kurth T, Weidinger G (2011) Regeneration of cryoinjury induced

necrotic heart lesions in zebrafish is associated with epicardial activation and cardiomyocyte proliferation. PLoS One 6(4):e18503. https://doi.org/10.1371/journal.pone. 0018503 18. Darehzereshki A, Rubin N, Gamba L, Kim J, Fraser J, Huang Y, Billings J, Mohammadzadeh R, Wood J, Warburton D, Kaartinen V, Lien CL (2015) Differential regenerative capacity of neonatal mouse hearts after cryoinjury. Dev Biol 399(1):91–99. https://doi.org/10.1016/j.ydbio.2014.12. 018 19. Shen H, Gan P, Wang K, Darehzereshki A, Wang K, Kumar SR, Lien CL, Patterson M, Tao G, Sucov HM (2020) Mononuclear diploid cardiomyocytes support neonatal mouse heart regeneration in response to paracrine IGF2 signaling. Elife. 13(9):e53071. https:// doi.org/10.7554/eLife.53071 [pii]

Chapter 3 Left Ventricular Pressure Volume Loop Measurements Using Conductance Catheters to Assess Myocardial Function in Mice Tilman Ziegler, Karl-Ludwig Laugwitz, and Christian Kupatt Abstract Left ventricular catheterization in mice allows for in-depth assessment of myocardial function in healthy and diseased animals with the advent of pressure volume loop recordings greatly enhancing the technique. While a powerful tool, proper execution of the procedure is paramount to ensure reproducibility and reliability of the results obtained. Here, we describe the technique of left ventricular catheterization using the Scisense conductance catheter system by Transonic; however, the basic method applies to all murine catheter systems. We furthermore indicate possible pitfalls during the procedure and how to avoid them. Key words Cardiac function, Pressure-volume loop, Left ventricular catheterization, Conductance catheter

1

Introduction While researching heart diseases in mouse models, it is important not only to assess morphological changes to the heart, but also to measure cardiac function in order to verify the functional impact of genetic alterations or treatments in disease models. While small rodent ultrasound imaging systems have recently made large strides [1], left ventricular catheterization represents the gold standard to assess heart function in rodents. Initially, catheters such as the SPR-1000 Mikro-Tip catheter by Millar were only able to record the left ventricular pressures in systole and diastole [2]; advances in catheter design, however, enable researchers to also assess left ventricular volumes during the whole heart cycle. Early catheter systems recorded blood flow via thermodilution of chilled saline, a procedure requiring accurate calibration using standard curves. Recently, conductance catheters have been developed that do not require the error-prone calibration needed for earlier catheter systems.

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_3, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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The conductance system of the Scisense pressure volume catheters functions by introducing a current in the heart that generates an electric field between two electrodes placed at each end of the catheter tip. Changes in the blood volume of the heart will lead to a proportional change in the conductivity between the two electrodes [3]. The voltage, which corresponds to the blood volume, is then converted into a volume signal via Baan’s equation [4]. The conductance catheter, which is equipped with a pressure transducer, allows for the measurement of pressure parameters (Pmax, Pmin, Pmean, Pdev, end-systolic pressure, end-diastolic pressure, and dP/dTmax, dP/dTmin), as well as volumetric parameters (stroke volume, ejection fraction, and cardiac output). Here, we describe a technique for introducing the catheter into the left ventricle of a mouse via a small incision at the neck that allows for accurate measurement of multiple cardiac parameters and a thorough assessment of the cardiac function. We describe proper handling of Scisense pressure volume catheters, but the application of other catheter systems does not differ in terms of the operation itself. Furthermore, we discuss common mistakes and pitfalls that occur while performing the procedure and highlight possible solutions.

2

Materials To conduct left ventricular pressure volume loop recordings utilizing Transonic Scisense PV Catheters, an operating station containing a stereoscopic microscope and a heating platform with a rectal probe is required. There is no need for mechanical ventilation since, during left ventricular catheterization, no thoracotomy is performed. For instrumentation, two tweezers (Fig. 1a), yarn, a clamp (Fig. 1b), surgical scissors, and microscissors (Figs. 1c, d) are needed. Furthermore, a venous catheter (22 gauge ¼ 0.9 mm diameter) is required for insertion into the jugular vein and a hypodermic needle bent at the tip to facilitate catheter insertion as well as a microliter syringe (Fig. 1e). The system to measure pressure-volume loops consists of the catheter, the control unit, a recording unit, and a computer running the appropriate software. For mice, a 1.2F catheter is used, the tip of which consists of proximal and distal conductance electrodes and a central pressure sensor (Transonic Scisense PV Catheters, Fig. 1f). The catheter is then connected to the control unit (Transonic ADVantage PV System ADV500), which is used to zero the pressure and calibrate the conductance. The control unit is connected to a recording unit (PowerLab 4/35, ADInstruments), which converts the analog output of the control unit into a digital signal suitable for the downstream software (LabChart 8, ADInstruments).

Left Ventricular Pressure Volume Loop Measurements Using Conductance. . .

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Fig. 1 Required instruments to perform left-ventricular catheterization: (a) tweezers, (b) clamp, (c) surgical scissors, (d) micro scissors, (e) microliter syringe and (f) conductance catheter

For the anesthesia of the mouse, the two sedatives medetomidine (1 mg/mL) and midazolam (1 mg/mL) are required as well as fentanyl (0.05 mg/mL) as an analgesic (MMF-anesthesia). Furthermore, norepinephrine is required to assess reserve capacity. You will also need pre-warmed saline (37  C), surgical sponges to drain excess liquid, syringes for intraperitoneal injection, and tape to fixate the animal.

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Methods 1. Bring the heating platform and saline solution to 37  C. 2. Prepare norepinephrine at concentrations of 1 μg/mL, 2.5 μg/ mL, 5 μg/mL, and 10 μg/mL. 3. Mix the MMF-anesthesia and inject it intraperitoneally according to body weight (see Table 1). After expiration of reflexes (pinching the animal between the toes does not result in twitching), place the animals on a warming platform with their extremities fixed with tape in supine position with the head facing the operator. 4. Tie a loop of yarn around the upper incisors, fixed above the head with tape, to extend the neck. 5. Insert the rectal probe of the warming platform. You are now ready to begin the operation. 6. Use the surgical scissors to make a longitudinal incision ~5 mm to the right of the midline of the neck. Pull apart both sides of the incision with the tweezers (see Fig. 2a). 7. The first structure now visible is the submandibular gland, which in mice is rather large. Remove the gland by pulling on the caudal end toward the head. This should expose the trachea in the middle, which is surrounded by a sheath of longitudinal muscle fibers. The carotid artery is to the right (slightly deeper inside the neck) and the jugular vein further lateral to the right (Fig. 2b). 8. Use the tweezers to separate the carotid artery from the vagus nerve. Place three pieces of yarn as purse strings around the carotid artery (Fig. 2c). Tighten the cranial yarn to occlude the carotid artery cranially. Place another piece of yarn around the most caudal section of the exposed carotid artery, but do not tie it. Infix both ends of the yarn with a clamp and pull it until the vessel stops perfusing. Place an additional tie in between without tightening the knot (Fig. 2c).

Table 1 Midazolam and fentanyl of the indicated concentrations are mixed with the volumes indicated Medetomidine

Midazolam

Fentanyl

mg/mL

1

1

0.05

mL

0.5

5

1

mg/kg BW

0.5

0.05

0.005

The lowest row indicates the appropriate required amount of each component per kg bodyweight

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Fig. 2 (a) Indicates the incision site about 5 mm lateral of the central line, about 5 mm. (b) After incision, three anatomical structures come into view: (1) trachea, (2) carotid artery and (3) jugular vein. (c) Carotid artery is ligated at the cranial end, blood flow is interrupted proximally via a yarn-loop, and a knot for fixation after catheter insertion is loosely placed around the carotid artery. (d) After advancing the catheter across the aortic valve, pressure and volume measurements can be performed. (e) Indicates the recorded pressures in the aorta (4) and the left ventricle (5) which allow to ensure proper catheter tip placement

9. Once this setting is completed, use the surgical microscissors to make an incision spanning one-third of the circumference of the carotid artery in the cranial third of the carotid artery. 10. Moisten the operating field with saline to ease the introduction of the catheter. 11. Introduce the catheter by lifting the incision with a bent hypodermic needle. Once the catheter tip is fully inserted into the carotid artery, tighten the loose knot prepared earlier. The catheter tip prevents blood from flowing past the catheter tip and out the incision. 12. Once the catheter is secured in the carotid artery, open the proximal yarn, held in place by the clamp, to allow the catheter

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to advance into the aorta and subsequently into the left ventricle. 13. Proper placement of the catheter tip can be monitored via the recorded pressure curves. Figure 2e shows typical aortic and left ventricular pressure measurements during insertion of the catheter. 14. Observe the pressure curves to determine proper catheter placement. The tip of the catheter occasionally rests against the left ventricular wall, leading to sharp end-systolic spikes in the pressure curve recordings. In this case, slightly retract the catheter until you achieve a smooth, dome-shaped pressure curve. 15. After proper catheter placement, calibrate the conductance and volume measurements on the ADV500 console. 16. Fix the catheter in place until cardiac function has stabilized. After this period of time, baseline measurements of cardiac function can be attained. 17. In order to assess the reserve capacity after norepinephrine stimulation, establish a central venous line. Prepare the jugular vein similarly to the carotid artery. 18. Tighten the knot at the cranial end of the jugular vein to occlude blood flow. Interrupt proximal blood flow with the yarn in the surgical clamps to exercise draw on the vessel. 19. After preparation of an additional loose loop of yarn around the jugular vein, incise the vein with the surgical microscissors and a peripheral venous catheter (G 22, 0.90 mm diameter) and insert it into the jugular vein after flushing it with saline. 20. After fixating the catheter by tightening the prepared yarn, advance the peripheral venous catheter into the jugular vein. 21. After allowing for the blood pressure measurements to stabilize, inject norepinephrine into the now established central line via the microliter syringe and record pressure-volume loops (Fig. 3). Make sure to allow for a return to baseline after simulation with norepinephrine before administering the next dose. 22. An increasing dosage scheme is recommended (10 ng, 25 ng, 50 ng, 100 ng). To this end, inject 10 μL of the prepared dilutions into the central line. 23. After recording the PV loops at rest and during stimulation with norepinephrine, remove the catheter from the animal and perform the steps used in the insertion in reverse. Soak the extracted catheter in distilled water or saline for 5 min. Next, soak in enzymatic cleaning solution for 30–120 min, depending on the level of soiling. Sacrifice the animal by injection of

Left Ventricular Pressure Volume Loop Measurements Using Conductance. . .

A

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Baseline

50ng

B

100ng

180 160 LV Pressure (mmHg)

140 120 100 80 60 40 20 0 10

20

30

40

50

60

LV Volume (ml)

Fig. 3 (a) Recording of left ventricular pressure during rest and after stimulation with increasing doses of norepinephrine. Note the increase in left-ventricular pressure corresponding to the increased doses of norepinephrine. (b) Example of a recorded pressure-volume loop at baseline conditions

potassium chloride into the central line or by cervical dislocation.

4

Notes 1. PV loop measurements in rodents have to be approved by the appropriate governing body, e.g., local animal welfare committee or comparable institutions. 2. While the anesthesia with MMF is known to have cardiodepressant properties, it is an anesthetic with high controllability, which renders it useful in these measurements. Other forms of sedation and analgesia might be used instead, such as intraperitoneal ketamine injections or a mix of inhalative sedation with isoflurane and a local anesthetic (xylocaine) at the site of

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operation. The latter mode of anesthesia is especially useful when attempting to measure animals that, due do certain interventions, are hemodynamically unstable. The mode of anesthesia has to be adjusted according to the experimental setup, followed by a consultation of the responsible governing body for approval [5]. 3. The whole procedure is time sensitive since prolonged anesthesia can influence the measured parameters. The operation should be practiced to achieve reproducibility. A trained operator can perform the procedure, including the norepinephrine stimulation, in 30–40 min. 4. While the carotid artery represents a robust vessel, which is easily separated from the surrounding connective tissue, the jugular vein is more fragile and encircled in thick layers of connective tissue. To expedite the operation and to avoid unnecessary bleeding, the jugular vein should not be completely separated from its surrounding connective tissue. 5. The incision of both the carotid artery and the jugular vein represents a crucial step in the procedure and accounts for a majority of measurement failures, if done incorrectly. One key mistake often seen in this step is the improper placement of the incision or an incision that is too large. This can lead to a total rupture of the vessel, after which the catheter cannot be advanced into the left ventricle. Furthermore, when the catheter is secured in the carotid artery, reopening the vessel can lead to blood flowing past the catheter. In this case, blood loss could influence the validity of the measurements. 6. Once the central line is established, negative pressure in the jugular vein can lead to aspiration of the saline solution in the venous catheter. If the catheter is completely drained, air will be sucked into the circulation, leading to air embolism, which can quickly kill the animal. Thus, the fluid amount in the venous catheter has to be closely monitored to avoid air embolism. 7. If desired, a tracheotomy can be performed to secure the airway passage. While this procedure adds time to the overall process, unstable animals can profit from the improved ventilation, reducing dropout rates. Perform the tracheotomy as a first step, after the lateral cervical incision. Separate the longitudinal muscle covering the trachea in the middle. Make an incision between the two most cervical tracheal cartilages. Insert a polyethylene tube into the incision and secure it with a knot. The length of the PE tubing should be chosen to reach the lower jaw of the mouse. Inserting the tubing prevents the aspiration of blood or saline from the operating field into the lung.

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References 1. Heinen A et al (2018) Echocardiographic analysis of cardiac function after infarction in mice: validation of single-plane long-axis view measurements and the bi-plane Simpson method. Ultrasound Med Biol 44(7):1544–1555 2. Lorenz JN, Robbins J (1997) Measurement of intraventricular pressure and cardiac performance in the intact closed-chest anesthetized mouse. Am J Phys 272(3 Pt 2):H1137–H1146 3. Kottam A et al (2011) Novel approach to admittance to volume conversion for ventricular

volume measurement. Conf Proc IEEE Eng Med Biol Soc 2011:2514–2517 4. Baan J et al (1981) Continuous stroke volume and cardiac output from intra-ventricular dimensions obtained with impedance catheter. Cardiovasc Res 15(6):328–334 5. Schmitz S et al (2016) Comparison of physiological parameters and anaesthesia specific observations during isoflurane, ketamine-xylazine or medetomidine-midazolam-fentanyl anaesthesia in male guinea pigs. PLoS One 11(9):e0161258

Chapter 4 Myocardial Infarction in Pigs Andrea B€ahr, Nadja Hornaschewitz, and Christian Kupatt Abstract Myocardial infarction is a major clinical challenge for interventional, pharmacological, and potential molecular treatment of the ischemic insult. A large animal model with clinic-derived instrumentation allows for detailed imitation of interventional catheterization routines and application routes, whereas similar anatomy and heart proportions raise the possibility to precisely evaluate the efficacy of application modes, e.g., antegrade or retrograde intracoronary application of locally acting pharmaceutical agents, viruses, and cells. Here, we describe the techniques of left ventricular catheterization and induction of ischemia and reperfusion, as well as hemodynamic monitoring and regional application of therapeutic agents in pigs. Key words Pig heart, Ischemia and reperfusion, Coronary catheterization, Regional treatment

1

Introduction Once the effort is committed to evaluate particular molecular interventions in the scenario of myocardial infarction in porcine hearts, it is important to maximize the outcome with regard to translatability toward the patient’s situation. Whereas basic findings are often discovered in cellular or small animal models, large animals offer the advantage of similarity of proportions, anatomy, and physiology, including heart rate and stroke volume. This predominant translational feature, however, requires careful analysis, as pigs used for this type of research are generally young and healthy, growing at rates of up to 5 kg/week. In order to mimic the cardiac risk factor profile of patient cohorts, one can now use also hyperglycemic [1] and hypercholesterinemic pigs [2]. The initiation of the myocardial ischemia is achieved by a balloon inserted into an infarct-related coronary artery of choice, either the left anterior descending (LAD) artery [3–7] or the ramus circumflexus (RCx) [8]. In the LAD, a percutaneous transluminal coronary artery (PTCA) balloon is inflated distal to the first or second diagonal branch, rendering a larger or smaller infarct size, respectively. Whereas >25% LV area can be infarcted by 60 min of

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_4, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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balloon inflation just distal to the first diagonal branch, the same time interval yields about 20% of the infarct size of the LV area if the balloon is inflated distally to the second diagonal branch. Doubling the time intervals increases the infarct size slightly, but may also increase side effects such as ventricular tachycardia. Permanent occlusion of the infarct-related artery may be percutaneously inflicted by placement of coils or vascular plugs, but by virtue of the model lacks the reperfusion component, which is currently treatment standard for myocardial infarction in Western and many developing countries. Thus, as a highly translational model, myocardial ischemia and reperfusion are to be preferred in the pig. The functional impact of myocardial infarction may be globally analyzed by left ventricular angiograms by analyzing contraction and relaxation velocities of a pressure tip placed in the left ventricle, and by regional assessment of contractile function via Piezo elements inserted into the subendocardial parenchyma, which signal the changes in distance to each other, called subendocardial segment shortening (SES), to a recorder. Whereas the former modalities (LV angiogram, pressure-tip catheter) can be conducted longitudinally, i.e., before and after ischemia induction as well as at the end of the experiment, SES requires an open chest and direct access to the heart, leaving it confined to the end of an experiment. Moreover, a Scisense® catheter (Transonic, Esloo, The Netherlands) may be used to generate pressure-volume loops by introducing a current in the heart that generates an electric field between two electrodes placed at each end of the catheter tip allowing for quantifying volume changes during the heart cycle (see Ziegler et al., in this volume). In this chapter, we describe the procedure of percutaneous induction of myocardial infarction as well the assessment of myocardial function (global and regional) in pigs. We also describe the possibilities of regional application of therapeutic agents into the infarcted area during or shortly after the termination of ischemia.

2

Materials To conduct myocardial infarction in pigs, you need a fully equipped operating room where you can perform sterile surgical interventions that has equipment to mechanically ventilate and monitor anesthesia comprehensively, including electrocardiogram and blood pressure as well as pulse oximetry monitoring. Additionally, a defibrillator is needed and a fluoroscopy system consisting of a c-arm and a control unit to visualize the intervention. A Scisense® catheter with control unit and appropriate software is necessary for taking pressure-volume loop measurements.

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2.1 Percutaneous Transluminal Coronary Angioplasty to Induce Coronary Ischemia

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1. Arterial sheath, 67F or larger (Terumo™ or Cordis™), for insertion into the right or left carotid artery. 2. Arterial sheath (Terumo™ or Cordis™) for insertion into the right or left jugular vein. 3. A 0.3500 standard wire is recommended for advancement into the ascending thoracic aorta. 4. A Cordis Judkins right 3.5 catheter (6F) for intubation in the left main coronary ostium. 5. Imeron 350™ (Bracco Imaging, Germany) as contrast agent to visualize the coronary tree and the LV. 6. A 0.1400 wire (e.g., Prowater, Asahi, Japan) to target the infarctrelated artery and serve as a guiding structure for a PTCA balloon (e.g., Trek 3.0 mm, 15 mm length, Abbott, USA). 7. Insufflator to inflate the balloon with a 50:50 mix of water and contrast agent. 8. External pacemaker.

2.2 Surgical Equipment

1. Cautery knife or scalpel. 2. Atraumatic forceps. 3. Surgical retractor. 4. Dissecting forceps. 5. Dissecting scissors. 6. Vascular scissor. 7. Mosquito clamps. 8. Suture material. 9. Needle holder. 10. Surgical sutures. 11. Swabs. 12. Saline.

2.3

Medications

1. Azaperone. 2. Atropine sulfate. 3. Ketamine. 4. Propofol. 5. Fentanyl. 6. Heparin. 7. Amiodarone. 8. Adrenaline. 9. Cefuroxime. 10. Meloxicam.

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Methods Prepare all surgical equipment, sheaths, and catheters. For prefilling the catheters, saline can be supplemented with 1000 I.E. heparin/L to prevent clotting. Prepare the pig for performing procedures.

3.1 Anesthetizing the Pig

1. For sedation, inject 2 mg/kg of azaperone, 0.02 mg/kg of atropine sulfate, and 20 mg/kg of ketamine into the lateral neck musculature of the pig. 2. After the animal has fallen asleep, insert a peripheral venous catheter (type Braun™) into an ear vein and fixate. 3. Insert an endotracheal tube into the trachea using a laryngoscope and guiding wire (for this it might be necessary to relax the animal further by intravenous application of approximately 1 mg/kg of propofol and 2.5 μg/kg of fentanyl, according to effect). 4. Under manual bag ventilation (Ambu™), transfer pig to the operating table and connect to the ventilator. Start continuous intravenous propofol infusion at a rate of approximately 10 mg/kg/h, according to effect. 5. For analgesia, apply 3 μg/kg/h of fentanyl intravenously. 6. Place electrocardiogram leads and a pulse oximeter to monitor vital signs. 7. Fixate the pig in supine position on the operating table while pulling the front legs toward its rear legs and tying the legs to the table using elastic tape. This is done to stretch and stabilize the neck. 8. Shave the whole throat area between the mandibles and the shoulder joint and clean and disinfect thoroughly. Cover the pig with sterile drapes, leaving the throat area open for access. You are now ready to begin the procedure. 9. For first-time intervention on an animal, use the vessels on the right side of the neck. Touch the right mandible with the little finger of your left hand, the right shoulder joint with the little finger of your right hand, and the sternum with the ring finger of your right hand. Now drop index fingers and thumbs of both hands to join in the middle. The area where they touch the throat of the pig is where you make an incision. You can now palpate the sternocleidomastoid muscle paramedian to the centerline. 10. Use a cautery knife or a scalpel to make a longitudinal incision ~3–4 cm right on top of the sternocleidomastoid muscle.

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11. Using a surgical retractor, pull apart the skin and carefully cut through the skin muscle and connective tissue below until you can visualize the sternocleidomastoid muscle. 12. You will find the carotid artery medial of the muscle and the jugular vein lateral. 13. First, carefully prepare the artery using dissecting forceps and scissors. The pulse of the artery can be palpated and this should be used as guidance. 14. Once you have reached the vessel, grab it with atraumatic forceps and gently pull upward. Remove as much of the surrounding connective tissue as possible on a stretch of around 1 cm. 15. Loosely tie a ligature around the cranial part of the exposed artery. 16. Now apply 10,000 IE of heparin intravenously and let circulate before tying the ligature. 17. To better expose the vessel, pull the tied ligature upwards and fixate both ends to the cranial corner of the skin incision using a mosquito clamp. 18. Twine another ligature to the caudal part of the exposed artery and pull upwards to stop blood flow. Do not tie at this stage! 19. Make a small incision into the vessel wall cranial of the untied ligature. 20. Insert the arterial sheath through this incision and gently push forward toward the heart while loosening the pull on the ligature. 21. Once the sheath is in place, fixate sheath in position by tying the ligature around the vessel and the sheath. 22. Repeat the procedure with the jugular vein lateral of the sternocleidomastoid muscle. 23. Remove all retractors and clamps and check that there is no bleeding. 24. Attach a pressure line to the arterial sheath and connect to a blood pressure-measuring device to take internal blood pressure measurements. 3.2 Obtaining Baseline Functional Measurements

1. Insert the external pacemaker into the right atrium through the venous sheath and test pacing threshold by setting the frequency to 100 beats per minute (bpm) and reducing the output from 20 to 1 mA. Note when the pacer stops capturing the preset frequency (output threshold) and set an output above this threshold. Reset the frequency to a rescue level of 50 or 60 bpm.

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2. Under fluoroscopy, advance the flexible 0.3500 wire in the 6F pigtail catheter through the arterial sheath into the ascending aorta. 3. Pass the aortic valve with the wire by gently rotating it while bending on the valve, and retracting, until it falls through the valve during systole. 4. Remove the wire. 5. Obtain a basic LV angiogram by either automated injection of contrast agent (20 mL at 10 mL/s) or manual injection through the pigtail catheter under fluoroscopy. 6. Obtain online pressure measurements of systolic and diastolic LV pressures as well as contraction and relaxation velocities dP/dtmax/min. 7. Obtain basic PV loop by advancing a Scisense® FFS-097-004 catheter over the aortic valve into the left ventricle. 8. Optional: Obtain pressure-volume loops at rest and at rapid atrial pacing (120 and 150 bpm under norepinephrine). 3.3 Inducing Myocardial Ischemia

1. After basic measurements, remove the LV catheter and insert the 0.3500 wire into the aortic root. 2. Advance the Judkins right 3.500 catheter near the ostium of the left main stem. Retract the wire and gently advance the catheter into the ostium. Inject the contrast agent to check position and visualize coronary anatomy. Select the site of the balloon inflation. 3. Now advance the 0.1400 wire through the Judkins catheter into the infarct-related artery to the distal part. 4. Mount the balloon on the wire and advance it through the guiding catheter into the coronary artery to the occlusion site. 5. Check the balloon location by contrast agent injection and recording of cine loops (saved on the fluoroscopy storage). 6. Inflate the balloon gently to a pressure of 4–6 bar (indicated on the manometer of the inflation pump). 7. Check correct location of the balloon and flow to diagonal branch(es) by a further contrast agent injection into the coronary artery. 8. Check for changes of blood pressure, electrocardiogram (ST elevations), and rhythm indicative of functional impairment. 9. Be prepared for cardiac resuscitation in case of ventricular tachycardia or fibrillation. Preventive amiodarone infusion (300 mg over 30 min) is recommended.

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10. In case of ventricular tachycardia or fibrillation, immediate closed chest cardiac resuscitation is necessary. Apply external defibrillation at 360 J and inject adrenaline. If unsuccessful, deflate and remove balloon, repeat resuscitation, and wait until cardiovascular stabilization is reached before continuing ischemia. 3.4 Removing the Arterial and Venous Sheaths

1. Visualize the vessels by pulling apart the tissue with a surgical retractor. 2. Place a ligature around the vessel caudal to the position where the vessel and the sheath were tied together. 3. Cut the ligature around the sheath and the vessel and pull out the sheath while simultaneously closing the new ligature. 4. Securely ligate the vessel and trim all ligature ends. 5. Repeat with the second vessel. 6. Close the muscle layer and skin by single sutures. 7. 30 min before the end of anesthesia, apply 0.4 mg/kg of meloxicam intravenously as postoperative analgesia. Analgesia is continued for the first 2 postoperative days by daily oral application of 0.4 mg/kg of meloxicam. 8. As a preventive measure for postoperative infections, a single injection of antibiotic treatment can be given during the procedure (e.g., 10 mg/kg of cefuroxime).

3.5 Options for Administration of Compounds During the Procedure

1. Intracoronary application can be performed when using an over-the-wire (OTW) balloon (e.g., 3  15 mm). The balloon carries a central lumen, through which therapeutics can be delivered directly into the infarcted area. The balloon is then deflated at the end of the ischemia, initiating reperfusion. 2. Retrograde intravenous application is conducted by advancing a Swan-Ganz catheter through the coronary sinus during fluoroscopy via the venous sheath. Advance a 0.1400 extra support wire (e.g., Grand Slam) selectively into the vein accompanying the infarct-related artery (e.g., the anterior interventricular vein AIV in case of the LAD). 3. Advance the Swan-Ganz catheter to the proximal third of the target vein and inflate the balloon at the tip of the catheter. 4. Inject contrast agent first, and if it is staining the target area, inject therapeutic agent by pressure-controlled reversal of the venous blood flow (do not increase pressure above 40 mmHg). 5. Retrograde injection benefits from antegrade occlusion of the infarct-related artery. Therefore, perform retrograde injection at the end of the ischemic period.

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6. Deflate the balloon of the Swan-Ganz catheter at the end of the injection and then deflate the balloon in the artery. 3.6 Post-procedure (24 h–2 Months of Reperfusion, or Longer)

1. Repeat steps for anesthesia and instrumentation, Subheading 3.1, steps 1–24. 2. Repeat functional measurements, Subheading 3.2, steps 1–8. 3. Open the chest with a suitable surgical scissors. 4. Remove pericardial fat and open the pericardial sac. 5. Perform a regional measurement of contraction by placing two ultrasonic crystals 1 and 4 cm distal to the occlusion site perpendicular to the LAD, and placing a second pair in the Cx perfusion region. 6. Ligate the occlusion site on the LAD with a 4.0 Prolene stitch. 7. Inject 20 mL of methylene blue into the left ventricular cavum and arrest the heart with 10 mL of KCl solution. 8. Quickly excise the heart under permanent suction. 9. Cannulate the LAD through the ligation (which may be gently opened and then tied down on the tip of the canula). 10. Gently inject tetrazolium-red (10%) under pressure control (2.5 cm), use a cryoprobe of 0.5 mm; for the smaller fish (2–2.5 cm), use a 0.3 mm cryoprobe. (B, C0 ) Extracted hearts from uninjured (B, B0 ) and hearts at 3 days post-injury (3dpi) (C, C0 ) Tg(ubb:mCherry) [16] zebrafish. In this line, mCherry is expressed ubiquitously under the ubb promoter. (B) Bright-field image of an uninjured heart. (B0 ) Fluorescent image of the heart depicted in B. Notice the ubiquitous RFP expression. (C) Bright-field image of injured heart at 3 dpi. The black dashed line indicates the injury area. (C0 ) Fluorescent image of the heart depicted in c. Notice the absence of mCherry signal in the injury area. (D, D00 ) Acid Fuchsin Orange G (AFOG) staining of paraffin sections of representative uninjured (D) and cryoinjured hearts at 47 dpi (D0 ) and 100 dpi (D00 ). At 47 dpi, large amounts of collagen are deposited in the injury area (D0 ), whereas at 100 dpi previously deposited collagen is strongly reduced (D00 ). Blue marks collagen and brown-orange stains myocardial tissue. ba bulbus arteriosus, v ventricle, at atrium. Scale bar: 1 cm (A), 200 μm (B and C)

4. Complete anesthesia is characterized by the lack of movement when the caudal fin is gently squeezed. 5. To remove the scales covering the skin on top of the heart, always lift them in the direction from the tail to the head. The heart region is clearly identifiable; you can see the heart beating under the skin (Fig. 2d and Supplementary Video 1).

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6. It is critical to perform the incision in the correct place. If you cut too much toward the caudal region, you risk exposing the liver and bursting the abdominal wall. Make sure that you pinch the skin on top of the heart in a position more toward the gills and that you cut immediately underneath the forceps (Supplementary Video 1). 7. In case the initial incision is too small, widen it. If you did not cut through the pericardium, gently tear it open with the forceps (Supplementary Video 1). 8. In some cases, the ventricle is deep inside the fish. If this is the case, do not squeeze the fish too hard. When experiencing problems to expose the ventricle, try to enlarge the incision and press the flanks of the fish close to the pectoral fins (Fig. 2e). 9. It is possible that the atrium is exposed before the ventricle. If this happens, gently push the atrium back into the pericardial cavity (using blunt forceps or a paper towel) and expose the ventricle. 10. It is important to thoroughly dry the apex of the ventricle prior to the cryoinjury. The presence of humidity covering the ventricle could potentially influence the cryoinjury efficiency and size. 11. Between surgeries, clean the cryoprobe thoroughly with a paper towel before inserting it back into the liquid nitrogen to avoid the formation of ice crystals. 12. Make sure that you only remove the cryoprobe after it is completely thawed; otherwise you risk tearing the heart from the fish (unless using procedure described in Note 15) (Fig. 2g). 13. Short freezing periods may lead to very small and superficial injuries. Longer freezing periods may lead to larger injuries that will take longer to regenerate. Adjust the freezing time according to the size of the fish (and the heart). Smaller fish, with smaller hearts, require shorter freezing times than larger fish with larger hearts. Adjusting the size of the cryoprobe and the duration of the cryoinjury for every experimental setup is recommended, but make sure to use the same setup within one experiment. For first-time users, remove some hearts 3 days post-injury to evaluate the injury extension (see Fig. 3b, c0 ). For smaller fish (~2 cm), use a cryoprobe of 0.3 mm (Fig. 3a). The following factors can influence the thawing speed of the probe: (1) the temperature of the surgery room, (2) unintended contact with other surfaces, (3) the presence of a light source very close to the probe (LED light sources may help prevent the heating of the probe), and (4) the time that the cryoprobe is immersed in the liquid nitrogen (very short immersion periods

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do not allow for the probe to cool down enough), among other factors. 14. If the probe is taking longer than expected to thaw, use a pipette to pour some fish water on top of it (once 10 s has passed, the probe must be removed). 15. We provide an alternative to the cryoinjury method described in the Surgical Procedure, point 12, which we recommend for more experienced personnel. After exposing the ventricular apex, touch the ventricle with the pre-cooled cryoprobe for 3–4 s. Use your index finger to gently detach the frozen cryoprobe from the heart. Make sure that you do not pull the frozen probe when in contact with the heart since this may result in tearing the fish heart. 16. Once the fish has been reanimated and starts to swim again, observe its behavior. Euthanize fish that, 15 min after waking up, are swimming in circles or that are unable to keep a homeostatic position in the water. Check the fish 2 h after surgery and remove dead animals. 17. Fish mortality upon ventricular cryoinjury depends on the age, health status and genetic background of the fish as well as of the, experience of the user, among other factors. The protocol described here should result in 80–90% average fish survival upon ventricular cryoinjury. 18. To identify the heart, look for the beating of the ventricle. Follow the ventricle until you find the bulbus arteriosus (BA), which resembles a conic-shaped, transparent structure that is attached to the base of the ventricle. To remove the heart, take hold the BA at its distal-most site between BA and aorta (not between BA and ventricle). 19. Alternatively, you can remove the heart by making three incisions: one between the gills and then another two underneath the left and right pectoral fins, parallel to the fish body. Using forceps, pull up the body wall to expose the heart. Alternative heart dissection protocols were published previously [14, 15]. 20. Upon collection, it is recommended to wash the hearts in the fish dissection buffer 1 to reduce the presence of circulating cells. Prior to fixation, the heart should no longer show contractility. 21. Long fixation times may affect further processing steps, such as sectioning or immunostainings. 22. For quantitative studies of cardiac regeneration, we recommend using at least 20 animals per group to increase the statistical power. Since the cardiac injury size will be

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proportional to the initial cryoinjured tissue, we suggest performing the surgeries in a blind manner and to switch between experimental and control groups every 3–5 fish. For qualitative studies, 5–10 animals per group are usually enough.

Acknowledgments This work was supported by The European Research council, grant ERC Consolidator grant 819717 – TransReg and SNF Project ForceInRegeneration 310030L_182575 from the Swiss National Science Foundation to N.M. References 1. Frangogiannis NG (2006) The mechanistic basis of infarct healing. Antioxid Redox Signal 8(11–12):1907–1939 2. Gonzalez-Rosa JM, Burns CE, Burns CG (2017) Zebrafish heart regeneration: 15 years of discoveries. Regeneration (Oxf) 4 (3):105–123 3. Poss KD, Wilson LG, Keating MT (2002) Heart regeneration in zebrafish. Science 298 (5601):2188 4. Raya A, Koth CM, Buscher D, Kawakami Y, Itoh T, Raya RM, Sternik G, Tsai HJ, Rodriguez-Esteban C, Izpisua-Belmonte JC (2003) Activation of Notch signaling pathway precedes heart regeneration in zebrafish. Proc Natl Acad Sci U S A 100(Suppl 1):11889–11895 5. Curado S, Anderson RM, Jungblut B, Mumm J, Schroeter E, Stainier DYR (2007) Conditional targeted cell ablation in zebrafish: a new tool for regeneration studies. Dev Dyn 236(4):1025–1035 6. Chablais F, Veit J, Rainer G, Jazwinska A (2011) The zebrafish heart regenerates after cryoinjury-induced myocardial infarction. BMC Dev Biol 11:21 7. Gonzalez-Rosa JM, Martin V, Peralta M, Torres M, Mercader N (2011) Extensive scar formation and regression during heart regeneration after cryoinjury in zebrafish. Development 138(9):1663–1674 8. Schnabel K, Wu CC, Kurth T, Weidinger G (2011) Regeneration of cryoinjury induced necrotic heart lesions in zebrafish is associated with epicardial activation and cardiomyocyte proliferation. PLoS One 6(4):e18503 9. Chablais F, Jazwinska A (2012) Induction of myocardial infarction in adult zebrafish using cryoinjury. J Vis Exp (62):3666 10. Gonzalez-Rosa JM, Mercader N (2012) Cryoinjury as a myocardial infarction model

for the study of cardiac regeneration in the zebrafish. Nat Protoc 7(4):782–788 11. Jopling C, Sleep E, Raya M, Martı´ M, Raya A, Belmonte JCI (2010) Zebrafish heart regeneration occurs by cardiomyocyte dedifferentiation and proliferation. Nature 464:606 12. Kikuchi K, Holdway JE, Werdich AA, Anderson RM, Fang Y, Egnaczyk GF, Evans T, MacRae CA, Stainier DYR, Poss KD (2010) Primary contribution to zebrafish heart regeneration by gata4+ cardiomyocytes. Nature 464:601 13. Gonzalez-Rosa JM, Guzman-Martinez G, Marques IJ, Sanchez-Iranzo H, JimenezBorreguero LJ, Mercader N (2014) Use of echocardiography reveals reestablishment of ventricular pumping efficiency and partial ventricular wall motion recovery upon ventricular cryoinjury in the zebrafish. PLoS One 9(12): e115604 14. Singleman C, Holtzman NG (2011) Heart dissection in larval, juvenile and adult zebrafish, Danio rerio. J Vis Exp (55):3165 15. Arnaout R, Reischauer S, Stainier DY (2014) Recovery of adult zebrafish hearts for highthroughput applications. J Vis Exp (94): e52248 16. Mosimann C, Kaufman CK, Li P, Pugach EK, Tamplin OJ, Zon LI (2011) Ubiquitous transgene expression and Cre-based recombination driven by the ubiquitin promoter in zebrafish. Development 138(1):169–177 17. Wang J, Pana´kova´ D, Kikuchi K, Holdway JE, Gemberling M, Burris JS, Singh SP, Dickson AL, Lin Y-F, Sabeh MK, Werdich AA, Yelon D, Macrae CA, Poss KD (2011) The regenerative capacity of zebrafish reverses cardiac failure caused by genetic cardiomyocyte depletion. Development 138(16):3421–3430

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Open Access This chapter is licensed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license and indicate if changes were made. The images or other third party material in this chapter are included in the chapter’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the chapter’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder.

Chapter 6 Cardiac Resection Injury in Zebrafish Delicia Z. Sheng, Dawei Zheng, and Kazu Kikuchi Abstract The zebrafish (Danio rerio) possesses a spectacular capacity for cardiac regeneration. Zebrafish have been used in cardiac regeneration research for nearly two decades, contributing to the identification of signals and cellular mechanisms as potential targets for human heart repair. Investigations into cardiac regeneration in zebrafish have been facilitated by multiple methods of inducing cardiac tissue damage. Among the established methods, cardiac resection injury is a relatively simple, yet robust approach traditionally used to induce cardiac tissue damage in a reproducible manner. Here, we describe a detailed protocol to perform a cardiac resection injury in adult zebrafish and discuss potential complications for researchers who are new to this technique. Key words Zebrafish, Heart regeneration, Cardiomyocytes, Injury, Myocardial infarction, Heart failure

1

Introduction Zebrafish can effectively regenerate a wider range of tissue structures than mammals, thus providing a model to learn how regeneration might be induced in damaged human tissues. Due to a robust capacity for cardiac regeneration and amenability to genetic approaches, the zebrafish has become a standard model system in cardiac regeneration research and has significantly contributed to the discovery of new signals and cellular mechanisms that regulate new muscle formation in injured hearts. The cardiac regenerative response was initially examined using a resection injury model, in which ~20% of the ventricle is removed at the apex using iridectomy scissors [1, 2]. Although severe bleeding occurs after injury, a clot mechanism can quickly seal the wound, allowing the circulation to be sustained and the animal to survive. Within 30 days, new myocardium replaces the fibrin clot and restores the contiguous wall of the ventricle [1]. In addition to the resection injury model, cryoinjury [3–5], hypoxia/reoxygenation injury [6], and a cardiomyocyte-specific

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_6, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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genetic ablation model [7] have been used in zebrafish heart regeneration research, thereby expanding our knowledge on the regenerative capacity of the zebrafish heart. Compared with these methods, cardiac resection injury is a relatively simple procedure that can be easily adopted and rapidly performed in a variety of laboratory settings. In this chapter, we present a detailed and reproducible protocol for performing a cardiac resection injury in adult zebrafish. Moreover, we will discuss the complications that may be experienced by those who are new to this technique.

2

Materials

2.1 Zebrafish Handling

1. Small fish net. 2. Plastic spoon. 3. Plastic transfer pipette, 7 mL.

2.2

Anesthesia

1. Flat-bottom dish (12  5 cm). 2. Tricaine stock solution (20): Dissolve 4 g of tricaine (ethyl 3-aminobenzoate methane sulfonate salt) powder in 900 mL of ultrapure water and adjust the pH to 7.4 with 1 M Tris–HCl (pH 9.5). Make up to 1 L with ultrapure water. The stock solution can be aliquoted into 50 mL conical tubes and maintained at 20  C. Protected from light, the dissolved stock solution can be stored at 4  C for up to 1 month.

2.3

Resection Injury

1. Stereomicroscope (Nikon SMZ745T). 2. McPherson-Vannas scissors, 8 cm, straight; 5 mm blades, 0.1 mm tip. 3. McPherson-Vannas scissors, 8 cm, curved; 5 mm blades, 0.1 mm tip. 4. ProSciTech pointed tweezers, style 5, 114 mm; anti-magnetic/ anti-acid (AM/AA). 5. Zebrafish holder: A cellulose sponge with a slot in the center to hold the fish securely (see Note 1). 6. Twisted pieces of KimWipes. 7. Breeding tank (1–2 L) without the inner tank and divider.

3

Methods

3.1 Preparation for Resection Injury

1. For a right-handed operator, place the straight scissors, curved scissors, and tweezers systematically from left to right on the right-hand side of the stereomicroscope (Fig. 1, instruments). Hereafter, all procedures are described for a right-handed

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Fig. 1 A typical setup for cardiac resection injury experiments

operator. A left-handed operator should reverse the instrument and placement order accordingly. 2. Pour the tricaine stock solution into a flat-bottom dish and dilute it with aquarium water to obtain a 1 tricaine solution (Fig. 1, tricaine for anesthesia). Place a plastic spoon in the dish to stir the tricaine solution and choose the anesthetized zebrafish for the injury procedure. 3. Moisten a cellulose sponge with aquarium water (Fig. 1, sponge) and place it on the stage of the stereomicroscope (Fig. 1, microscope). The sponge is necessary to prevent dehydration and provide stability for the zebrafish during the injury procedure. 4. Make spillage paper by twisting small pieces of KimWipes and placing them near the surgical instruments (Fig. 1, spillage paper). 5. Fill a breeding tank (1–2 L) with aquarium water to approximately 70% of its capacity (Fig. 1, recovery tank). Use this tank as a container for post-surgery zebrafish. 3.2

Anesthesia

1. Transfer the zebrafish to the tricaine solution using a fish net and gently stir using a plastic spoon (see Note 2). 2. Carefully monitor the fish while stirring and wait for the gill movements to slow down. This usually takes 60–90 s (see Note 3). 3. Transfer the anesthetized fish to the moist sponge using a plastic spoon. Place the fish into the slot of the sponge with the head to the left and the ventral side up (Fig. 2a). Quickly adjust the focus of the microscope onto the beating heart.

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Fig. 2 Procedure for cardiac resection injury. (a) Position of zebrafish on the holding sponge. Asterisks, pectoral fin muscle; dotted box, approximate area over the heart. (b) Penetration of the pericardial sac. Dotted line, penetration site; asterisk, the blade used for penetration. See text for arrows. (c) Resection of the ventricular apex. Dotted circle, the ventricular apex. See text for arrows 3.3

Resection Injury

Perform all procedures under the stereomicroscope unless otherwise mentioned. 1. Take the tweezers in your left hand and place the tips on the proximal part of the pectoral fin muscle (Fig. 2a, asterisks). Gently push the tweezers downward and slightly open it to widen the area between the pectoral fins (Fig. 2b, arrows). This procedure stretches and thins the connective tissue covering the heart (Fig. 2a, dotted box), making it easier to penetrate

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the tissue during the next step. Take care not to move the tweezers. 2. Keep the tweezers stationary and take hold of the straight scissors (see Note 4). Estimate the location of the skin covering the internal space between the heart and the abdominal organs (Fig. 2b, dotted line) (see Note 5) and penetrate the skin with one blade of the straight scissors (Fig. 2b, asterisk). Pull the scissors back slightly and carefully make a small incision of ~1 mm to the pericardial sac (see Note 6). Keep the incision open by maintaining the position of the tweezers. 3. Take the curved scissors (see Note 4) and position the convex side on the abdomen. Gently pressing the abdomen allows an apical portion of the ventricle to protrude from the incision (see Note 7). Press the abdomen until 30–50% of the ventricle is exposed, and then slow down the beating movement by constricting the ventricle using both the tweezers and the curved scissors (see Note 8). Once the movement of the ventricle slows, carefully slide the curved scissors toward the ventricle (Fig. 2c, arrow) and remove 15–20% of the ventricular apex (Fig. 2c, dotted circle) (see Note 9). 3.4 Post-injury Procedure

1. Place the curved scissors aside and use the twisted KimWipes to blot the bleeding (Figs. 1b and 3a). Release the holding tweezers to allow the heart to slip back into the pericardial sac. Gently close the incision with the tweezers and transfer the injured zebrafish to the recovery tank using a plastic spoon (see Note 10). 2. Gently and continuously squirt water into the gills using a plastic transfer pipette (Fig. 3b) until the fish is able to swim regularly on its own (see Note 11). 3. Leave the injured fish in the recovery tank for at least 5 min before transferring it into an aquarium tank in the zebrafish

Fig. 3 Post-injury procedures. (a) Blotting the bleeding with spillage paper. (b) Reviving post-injury zebrafish

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facility. The water flow must be set at a low speed for 24 h after injury.

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Notes 1. Make a slot in the center of the sponge of approximately 20 L  5 W  3D (mm) using a razor blade. To ensure that the fish is held securely, use the scissors to adjust the size of the slot by trimming the edges. 2. Add 4 mL of 20 tricaine stock (0.4% w/v) into 76 mL of fresh aquarium water in a glass dish of 8 cm diameter  5 cm height to achieve a final concentration of 0.02% (w/v). To avoid fish escaping during anesthesia, avoid filling the glass dish to more than half of its capacity. 3. Since deep anesthesia reduces the survival of injured zebrafish, it is critical to optimize the tricaine concentration and treatment time to the animal under investigation. The correct anesthetic condition will vary depending on the size and age of the fish; a practical approach to achieve the appropriate tricaine concentration is to start with a low tricaine concentration and then add stock solution in increments of 0.5–1 mL until a slowing of gill movement is observed within 1 min. 4. Avoid taking your eyes from the operation when reaching for the straight scissors. This helps to hold the position of the tweezers securely. 5. The silver-colored membrane between the heart and the abdominal organs is often discontinuous in adult zebrafish; this can help the targeting of the correct site. 6. Be careful not to damage the underlying atrium or ventricle. If these tissues are damaged, severe bleeding will occur and will cause death. The size of the incision should be kept small to minimize physical stress. Larger incisions often hamper the constriction procedure in the next step. If necessary, make the incision by tearing the pericardial sac and the connective tissue using a pair of tweezers. 7. Ensure that the ventricle, not the atrium, protrudes from the incision. A pink color indicates the ventricle, whereas the atrium presents as dark red. Access to the ventricle through the incision is usually straightforward, but the atrium can grow extremely large and block the access to the ventricle. In such a case, carefully push the atrium aside using a pair of tweezers to expose the ventricle. 8. The constriction can be achieved by slightly pushing the tweezers downward and pressing the abdomen toward the heart using the convex side of the curved scissors. These

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manipulations help to increase the pressure encircling the ventricle and slow down its movement. 9. Removal of more than 20% of the ventricle will significantly reduce survival after injury. A visual confirmation of the removed tissue (which usually remains on the blade of the scissors) is essential (Fig. 2c, dotted circle). The fish you have operated on should not be considered injured until the removed tissue is confirmed. 10. Suture is not necessary. 11. Reviving the injured fish usually takes 3–5 min. A rapid recovery from anesthesia is usually not a good sign; the fish may start swimming erroneously and eventually die. When fish are successfully injured, recovery times are longer and associated with characteristic behavioral features: the fish tend to elevate the head with the fin down, and slowly swim in circles just below the water surface. Fish showing this behavior usually remain conscious, so another fish can be anesthetized and ready for the next injury procedure.

Acknowledgments We thank M. Nakayama, C. Jenkin, and K. Brennan for zebrafish care. This work is supported by grants from NHMRC (APP1130247, APP1160466). References 1. Poss KD, Wilson LG, Keating MT (2002) Heart regeneration in zebrafish. Science 298 (5601):2188–2190 2. Raya A et al (2003) Activation of Notch signaling pathway precedes heart regeneration in zebrafish. Proc Natl Acad Sci U S A 100 (1):11889–11895 3. Chablais F, Veit J, Rainer G, Jaz´win´ska A (2011) The zebrafish heart regenerates after cryoinjuryinduced myocardial infarction. BMC Dev Biol 11:21 4. Gonza´lez-Rosa JM, Martı´n V, Peralta M, Torres M, Mercader N (2011) Extensive scar formation and regression during heart

regeneration after cryoinjury in zebrafish. Development 138(9):1663–1674 5. Schnabel K, Wu CC, Kurth T, Weidinger G (2011) Regeneration of cryoinjury induced necrotic heart lesions in zebrafish is associated with epicardial activation and cardiomyocyte proliferation. PLoS One 6(4):e18503 6. Parente V et al (2013) Hypoxia/reoxygenation cardiac injury and regeneration in zebrafish adult heart. PLoS One 8(1):e53748 7. Wang J et al (2011) The regenerative capacity of zebrafish reverses cardiac failure caused by genetic cardiomyocyte depletion. Development 138(16):3421–3430

Chapter 7 A Genetic Cardiomyocyte Ablation Model for the Study of Heart Regeneration in Zebrafish Fei Sun, Adam R. Shoffner, and Kenneth D. Poss Abstract Adult zebrafish possess an elevated cardiac regenerative capacity as compared with adult mammals. In the past two decades, zebrafish have provided a key model system for studying the cellular and molecular mechanisms of innate heart regeneration. The ease of genetic manipulation in zebrafish has enabled the establishment of a genetic ablation injury model in which over 60% of cardiomyocytes can be depleted, eliciting signs of heart failure. After this severe injury, adult zebrafish efficiently regenerate lost cardiomyocytes and reverse heart failure. In this chapter, we describe the methods for inducing genetic cardiomyocyte ablation in adult zebrafish, assessing cardiomyocyte proliferation, and histologically analyzing regeneration after injury. Key words Zebrafish, Heart, Cardiomyocyte, Regeneration, Ablation, Tamoxifen

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Introduction Due to the limited regenerative potential of cardiomyocytes in adult mammals, a myocardial infarction, or heart attack, often leads to the permanent loss of cardiomyocytes, which can proceed to heart failure [1–8]. By contrast, zebrafish robustly regenerate cardiomyocytes. After either surgically resecting [9] or cryoinjuring the cardiac ventricle [10–13], zebrafish can replace most or all of the lost muscle with new cardiomyocytes in 60 days. Genetic fate mapping has demonstrated that this new muscle is derived from preexisting cardiomyocytes [14–16]. Several years ago, a genetic ablation model was established that further challenged the cardiac regenerative capacity of zebrafish [17]. This system employs a cell type-specific depletion model to achieve inducible and targeted cardiomyocyte ablation via a dualtransgenic system, similar to those used in studies with mouse tissues [18–22]. In this model, termed zebrafish cardiomyocyte ablation transgenes (Z-CAT), expression of a tamoxifen-inducible Cre recombinase is directed in cardiomyocytes using the

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_7, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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cardiomyocyte-restricted cmlc2 (or myl7) promoter, enabling conditional expression of a diphtheria toxin A-chain (DTA) downstream of a beta-actin2 promoter and a loxP-flanked mCherry reporter cassette. Upon tamoxifen exposure, cmlc2:CreER; bactin2:loxp-mCherry-STOP-loxp-DTA (bact2:RSD) transgenes enable induced recombination at loxP sites and strong expression of cytotoxic DTA in a portion of cardiomyocytes [17]. Compared to cryoinjury models, the Z-CAT model allows for more extensive and reproducible cardiomyocyte-specific depletion. Over 60% of cardiomyocytes can be depleted from adult Z-CAT fish hearts without affecting animal viability [17]. Fish exhibit signs of heart failure, including lethargy, rapid gasping, and stress hypersensitivity, which has not been demonstrated in the other two injury models [17]. Despite the severe injury, Z-CAT fish respond to cardiomyocyte depletion by vigorous proliferation of the spared cardiomyocytes, as well as activation of injury responses in cardiac non-muscle cells, like endocardium and epicardium [17]. Fish can regenerate most or all destroyed cardiomyocytes in 30 days and reverse signs of heart failure [17]. This noninvasive injury is easy to execute on a large scale in unanesthetized adult animals, saving time versus laborious surgical procedures. Additionally, given that the ablation injury is distributed more or less evenly throughout the zebrafish cardiac chambers, it can facilitate identifying new regulators of cardiomyocyte proliferation and heart regeneration in zebrafish by genomic profiling [17, 23].

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Materials 1. Zebrafish: Maintain adult Tg(cmlc2:CreER) fish and Tg(bact2: RSD) fish at a density of 4–5 fish/L fish water at 27
C (see Note 1). 2. Mating tanks. 3. Fish water. 4. Tamoxifen: Aliquot tamoxifen powder into 1.7 mL Eppendorf tubes and store at 4
C in dark tubes (see Note 2). 5. 100% Ethanol. 6. Sponge: Cut a small groove in the middle of the sponge to fit an adult fish. 7. Stainless steel straight scissors. 8. Forceps. 9. Dissecting microscope. 10. Glass bowl. 11. 20 Tricaine solution: Dissolve 4 g of ethyl 3-aminobenzoate powder in 900 mL of water. Adjust the pH to 7.2–7.4 with

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Tris–HCL (pH ¼ 9.5) solution. Add the water to 1 L and filter-sterilize the solution. Ethyl 3-aminobenzoate powder should be stored at 20
C. 20 Tricaine solution can be aliquoted and stored at 20
C. Store thawed tricaine at 4
C for up to 1 week. 12. 4% PFA: Weigh 40 g of paraformaldehyde powder in chemical hood. Add 750 mL of PBS and heat the solution to 65–80
C on a heat plate. Once that temperature is reached, add sodium hydroxide to accelerate the process of dissolving. Cool the solution to room temperature after the powder is fully dissolved. Adjust the pH to 7.2–7.4 with HCl solution and add PBS to 1 L. Filter-sterilize and aliquot. Store at 20
C. Thaw 4% PFA fresh each time before tissue collection. 13. Formaldehyde solution: Store 36.5–38% in water at room temperature. Freshly dilute to working concentration before use. 14. PBST: PBS with 0.1% Tween 20. 15. PBS-Tx: PBS with 0.2% Triton-X. 16. 30% Sucrose solution: Dissolve 30 g of sucrose in 100 mL of ddH2O. The 30% sucrose solution can be stored for a long term at 20
C in small aliquots and kept at 4
C for shortterm storage. 17. Horse serum. 18. Heat-inactivated newborn calf serum. 19. Goat serum. 20. EdU stock solution: Prepare 25 mM of EdU stock solution in DMSO, aliquot, and freeze at 20
C for long-term storage. 21. EdU staining solution: Freshly prepare the EdU staining solution each time by mixing all the following reagents, with volumes listed for 300 μL (typically required for each slide): 150 μL 1 M Tris (pH 8.5) for 0.5 M final concentration, 1.5 μL 1 M CuSO4 for 5 mM final concentration, 1.5 μL 10 mM azide for a 50 μM final concentration, and 150 μL 0.5 M ascorbic acid for a 0.25 M final concentration. Freshly dissolve ascorbic acid in water each time before use and add last to the staining solution. 22. Anti-MEF2A + MEF2C antibody: This is a rabbit-derived monoclonal antibody from Abcam. Aliquot and store antibody at 20
C. 23. Troponin T antibody: This is a mouse-derived monoclonal antibody from Abcam. Aliquot and store antibody at 20
C. 24. Alexa Fluor 488 anti-rabbit antibody: Store antibody in the dark at 4
C.

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25. Alexa Fluor 488 anti-mouse antibody: Store antibody in the dark at 4
C. 26. DAPI solution: Dissolve DAPI in water to make a stock solution of 5 mg/mL. Store stock solution in the dark at 4
C. 27. Fluoromount G. 28. Hydrophobic pen. 29. Coplin jars. 30. Cryostat. 31. Coverslips. 32. Slide heater. 33. Fluorescence microscope. 34. 100 μl Hamilton syringe. 35. Tissue freezing medium.

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Methods

3.1 Zebrafish Cardiomyocyte Ablation

1. Mate adult Tg(cmlc2:CreER) fish and Tg(bact2:RSD) fish to acquire double-transgenic fish for experiments. Select embryos for mCherry-positive tissue using a fluorescent microscope at 3 days postfertilization (see Note 3). Keep fish at 4–5 fish/L in aquarium water to grow to adulthood after 4 weeks postfertilization. 2. Tg(cmlc2:creER) fish were established with a linked dsRed transgene driven by the alpha-crystallin promoter. Screen adult fish to separate CreER+ and CreER individuals based on the presence or absence of the eye marker, respectively, using a fluorescent dissecting microscope. Maintain CreER fish as uninjured controls in the following experiment. 3. Weigh tamoxifen powder, place it in a 1.7 mL Eppendorf tube, and dissolve it in 100% ethanol to make a 2 mM stock solution (see Note 4). Dilute the stock tamoxifen solution to 0.4–1 μM in fish water. Treat adult fish in mating tanks with 50–100 mL of diluted tamoxifen water per fish in the dark. Use tamoxifentreated CreER fish as uninjured controls in the experiment. 4. Remove fish from the tamoxifen after 16–24-h tamoxifen incubation (see Note 5). Rinse treated fish with fresh aquarium water in mating tanks before being returned to the circulating system (see Note 6).

3.2 Assay for Cardiomyocyte Proliferation During Regeneration

Zebrafish regenerate cardiac muscle through the proliferation of preexisting cardiomyocytes. To evaluate and quantify cardiomyocyte cell cycle entry in Z-CAT fish, EdU is injected interperitoneally to label cells in S phase. Mef2, a marker of cardiomyocyte nuclei, is

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Fig. 1 Ventricular cardiomyocyte cell cycle entry after genetic ablation. EdU was injected at 13 days postincubation (dpi) and hearts were collected 24 h post-EdU injection. (a) Z-CAT control heart shows rare Mef2+ and EdU+ cells. (b–d). The ablated ventricle shows more EdU-positive cells and evidence of more proliferating cardiomyocytes as indicated by arrows in (c) and (d). Scale bar: 100 μm

visualized with an antibody. Cells with both EdU and Mef2 are counted as proliferating cardiomyocytes (Fig. 1). 1. Prepare 10 mM of EdU solution by diluting the EdU stock solution with PBS. The EdU solution should be protected from light. 2. Anesthetize fish in 1 tricaine solution and interperitoneally inject them with 10 μL of 10 mM EdU solution using a 100 μL Hamilton syringe 24 h prior to heart collection. 3. Collect fish hearts for proliferation assays: Euthanize tamoxifen-treated fish in tricaine solution. After euthanization, expose their hearts with a single ventral incision using straight scissors. Use a pair of fine forceps to grasp the outflow tract and slowly pull to remove the heart. 4. Rinse hearts in cold PBS and then incubate in ice-cold 30% sucrose for at least 5 min. 5. Mount unfixed hearts in ice-cold tissue freezing medium, snap frozen in a dry ice/ethanol bath, and section at 10 μm thickness. 6. Dry slides on a slide warmer for 10 min. 7. Use a hydrophobic pen to draw around the tissue on the slide. 8. Dilute the formaldehyde solution to 3.7% with PBS and postfix tissue sections with freshly prepared 3.7% formaldehyde solution for 15 min at room temperature.

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9. Wash the slides with PBS for 5 min in a Coplin jar. Repeat for a total of three washes. 10. Stain the slides with fresh EdU staining solution for 30 min. 11. Wash the slides with PBS + 0.2% Triton X for 5 min in a Coplin jar. Repeat for a total of four washes. 12. Block the sections with 1% BSA + 5% goat serum in PBS-Tx for 1 h at room temperature. 13. Drain the blocking solution from the sections. Dilute the Mef2 primary antibody 1:100 in fresh blocking solution and incubate the slides in primary antibody overnight at 4
C. 14. Wash the slides with PBS-Tx for 5 min in a Coplin jar. Repeat for a total of four washes. 15. Dilute the Alexa Fluor 488 anti-rabbit antibody 1:250 in blocking solution. Incubate the slides with the secondary antibody for 2 h at RT. 16. Wash the slides for 5 min in PBS-Tx. Repeat for a total of four washes. DAPI can be added (1:5000) into the first PBS-Tx wash. 17. Mount the slides with Fluoromount G. Store them in the dark at 4
C before imaging with a fluorescence microscope. 18. Image the ventricular sections using a 20 objective on a compound fluorescence microscope. To calculate a cardiomyocyte proliferation index, sample 8–12 hearts in each group and at least three sections of each heart. 19. Count Mef2+ cell and Mef2+/EdU+ cells manually with the aid of ImageJ/FIJI software. 20. Calculate the cardiomyocyte proliferation index as the fraction of Mef2+/EdU+ cells versus total Mef2+ cells. Average the percentages from the three selected sections to determine the cardiomyocyte proliferation index for each animal. One may choose to calculate separate indices for trabecular and cortical muscle. 3.3 Histological Analysis of Cardiomyocyte Ablation and Regeneration

Evaluate cardiac muscle integrity with troponin-T staining after genetic ablation and at various stages of regeneration (Fig. 2). 1. Collect fish hearts as described above. 2. Place hearts into ice-cold 4% PFA solution and fix overnight at 4
C. 3. After overnight fixation, wash the hearts for 5 min in PBS + 0.1% Tween-20. Repeat for a total of four washes. 4. For cryopreservation, incubate the hearts in 30% sucrose overnight at 4
C until they sink to the bottom of the collection tube.

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Fig. 2 Cardiomyocyte regeneration after ablation of ventricular cardiomyocytes. (a–c). Troponin-T was visualized on ventricular sections by antibody staining. Approximately 60% of all ventricular cardiomyocytes were ablated as shown in (b). Most or all cardiomyocytes regenerate by 30 days following tamoxifen treatment (c). Scale bar: 100 μm

5. Mount the hearts in tissue freezing medium and freeze on dry ice for cryosection. 6. Section heart sections at 10 μm thickness and dry on the slide warmer for 30 min before staining. Store the slides for a long term at 20
C before immunostaining. 7. If the slides are taken from 20
C, dry the slides again on the slide warmer (37
C) for 30 min. 8. Use a hydrophobic pen to draw around the tissue on the slide. 9. Wash the slides for 5 min with PBST in a Coplin jar. Repeat for a total of four washes. 10. Block the tissue with 10% HI-NSC, 2% horse serum, and 1% DMSO in PBST for 1 h at 37
C. 11. Remove the blocking solution from slides. Dilute the mouse anti-Tnnt primary antibody 1:100 in PBST with 10% HI-NSC and 1% DMSO. Incubate the tissue with the primary antibody for 3 h at 37
C. 12. Wash the slides for 5 min with PBST in a Coplin jar. Repeat for a total of four washes. 13. Dilute the Alexa Fluor 488 anti-mouse antibody 1:250 in PBST with 10% HI-NSC and 1% DMSO. Incubate the slides with the secondary antibody at 37
C for 1 h. 14. Wash the slides for 5 min with PBST in a Coplin jar. Repeat for a total of four washes. DAPI can be added (1:5000) to the first PBST wash to stain the nuclei. 15. Mount the slides with Fluoromount G. Store them in the dark at 4
C before imaging with a fluorescence microscope.

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Notes 1. All fish should be maintained and handled under the guidelines of an institutionally approved animal use protocol. 2. Tamoxifen in solid form is stable for up to 1 year at 4
C, if stored carefully away from light. Tamoxifen solution should be freshly prepared each time before use and kept in the dark after dissolving and during treatment. 3. Due to the maternal expression from the beta-actin2 promoter, all embryos derived from female Tg(bact2:RSD) fish are red throughout their entire body as visualized by a fluorescence microscope until 3 dpf. Under these conditions, only embryos with strong red fluorescent hearts contain the bact2:RSD transgene. It is, therefore, preferable to mate male Tg(bact2:RSD) fish with female Tg(cmlc2:creER) fish to obtain Z-CAT doubletransgenic fish. 4. Tamoxifen is insoluble in aqueous solutions. It can be completely dissolved in 100% ethanol by vortexing briefly at room temperature. Propylene glycol can also be used as an organic solvent but requires vigorous shaking at 37
C to fully dissolve the tamoxifen. DMSO should not be used as the solvent since it can induce cardiomyocyte proliferation in adult zebrafish without injury. 5. Z-CAT fish are sensitive to small variations in tamoxifen dosage and treatment time. The ablation effect may vary between clutches. To achieve the desired level of ablation, pilot experiments should be performed with each clutch to determine the optimal tamoxifen dosage and incubation time. 6. Tamoxifen must be handled with gloves. Any surplus tamoxifen must be disposed of as a hazardous waste. Any spills should be cleaned with a 20% bleach solution. Any container that contacts tamoxifen solution should be flushed with hot water for 5 min and placed in a designated drug treatment area for cleaning. Wash hands with soap and water immediately after handling tamoxifen.

Acknowledgments K.D.P. acknowledges support from National Institutes of Health (R35 HL150713).

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References 1. Chiong M, Wang ZV, Pedrozo Z, Cao DJ, Troncoso R, Ibacache M, Criollo A, Nemchenko A, Hill JA, Lavandero S (2011) Cardiomyocyte death: mechanisms and translational implications. Cell Death Dis 2:e244. https://doi.org/10.1038/cddis.2011.130 2. Goldspink DF, Burniston JG, Tan LB (2003) Cardiomyocyte death and the ageing and failing heart. Exp Physiol 88(3):447–458 3. Cui B, Zheng Y, Sun L, Shi T, Shi Z, Wang L, Huang G, Sun N (2018) Heart regeneration in adult mammals after myocardial damage. Acta Cardiol Sin 34(2):115–123. https://doi.org/ 10.6515/ACS.201803_34(2).20171206A 4. Steinhauser ML, Lee RT (2011) Regeneration of the heart. EMBO Mol Med 3(12):701–712. https://doi.org/10.1002/emmm. 201100175 5. Dickstein K, Cohen-Solal A, Filippatos G, McMurray JJ, Ponikowski P, Poole-Wilson PA, Stromberg A, van Veldhuisen DJ, Atar D, Hoes AW, Keren A, Mebazaa A, Nieminen M, Priori SG, Swedberg K, Guidelines ESCCfP (2008) ESC guidelines for the diagnosis and treatment of acute and chronic heart failure 2008: the Task Force for the diagnosis and treatment of acute and chronic heart failure 2008 of the European Society of Cardiology. Developed in collaboration with the Heart Failure Association of the ESC (HFA) and endorsed by the European Society of Intensive Care Medicine (ESICM). Eur J Heart Fail 10 (10):933–989. https://doi.org/10.1016/j. ejheart.2008.08.005 6. Stewart S, MacIntyre K, Hole DJ, Capewell S, McMurray JJ (2001) More ‘malignant’ than cancer? Five-year survival following a first admission for heart failure. Eur J Heart Fail 3 (3):315–322. https://doi.org/10.1016/ s1388-9842(00)00141-0 7. Pfeffer MA, Braunwald E (1990) Ventricular remodeling after myocardial infarction. Experimental observations and clinical implications. Circulation 81(4):1161–1172 8. Kehat I, Molkentin JD (2010) Molecular pathways underlying cardiac remodeling during pathophysiological stimulation. Circulation 122(25):2727–2735. https://doi.org/10. 1161/CIRCULATIONAHA.110.942268 9. Poss KD, Wilson LG, Keating MT (2002) Heart regeneration in zebrafish. Science 298 (5601):2188–2190. https://doi.org/10. 1126/science.1077857 10. Chablais F, Veit J, Rainer G, Jazwinska A (2011) The zebrafish heart regenerates after

cryoinjury-induced myocardial infarction. BMC Dev Biol 11:21. https://doi.org/10. 1186/1471-213X-11-21 11. Gonzalez-Rosa JM, Martin V, Peralta M, Torres M, Mercader N (2011) Extensive scar formation and regression during heart regeneration after cryoinjury in zebrafish. Development 138(9):1663–1674. https://doi.org/ 10.1242/dev.060897 12. Gonzalez-Rosa JM, Mercader N (2012) Cryoinjury as a myocardial infarction model for the study of cardiac regeneration in the zebrafish. Nat Protoc 7(4):782–788. https:// doi.org/10.1038/nprot.2012.025 13. Schnabel K, Wu CC, Kurth T, Weidinger G (2011) Regeneration of cryoinjury induced necrotic heart lesions in zebrafish is associated with epicardial activation and cardiomyocyte proliferation. PLoS One 6(4):e18503. https://doi.org/10.1371/journal.pone. 0018503 14. Jopling C, Sleep E, Raya M, Marti M, Raya A, Izpisua Belmonte JC (2010) Zebrafish heart regeneration occurs by cardiomyocyte dedifferentiation and proliferation. Nature 464 (7288):606–609. https://doi.org/10.1038/ nature08899 15. Kikuchi K, Holdway JE, Werdich AA, Anderson RM, Fang Y, Egnaczyk GF, Evans T, Macrae CA, Stainier DY, Poss KD (2010) Primary contribution to zebrafish heart regeneration by gata4(+) cardiomyocytes. Nature 464 (7288):601–605. https://doi.org/10.1038/ nature08804 16. Foglia MJ, Poss KD (2016) Building and re-building the heart by cardiomyocyte proliferation. Development 143(5):729–740. https://doi.org/10.1242/dev.132910 17. Wang J, Panakova D, Kikuchi K, Holdway JE, Gemberling M, Burris JS, Singh SP, Dickson AL, Lin YF, Sabeh MK, Werdich AA, Yelon D, Macrae CA, Poss KD (2011) The regenerative capacity of zebrafish reverses cardiac failure caused by genetic cardiomyocyte depletion. Development 138(16):3421–3430. https:// doi.org/10.1242/dev.068601 18. Brockschnieder D, Lappe-Siefke C, Goebbels S, Boesl MR, Nave KA, Riethmacher D (2004) Cell depletion due to diphtheria toxin fragment A after Cre-mediated recombination. Mol Cell Biol 24(17):7636–7642. https://doi.org/10.1128/MCB.24.17.76367642.2004 19. Lee P, Morley G, Huang Q, Fischer A, Seiler S, Horner JW, Factor S, Vaidya D, Jalife J,

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pathogenic role in mouse model of heart failure. J Biol Chem 279(39):41095–41103. https://doi.org/10.1074/jbc.M313084200 22. Breitman ML, Clapoff S, Rossant J, Tsui LC, Glode LM, Maxwell IH, Bernstein A (1987) Genetic ablation: targeted expression of a toxin gene causes microphthalmia in transgenic mice. Science 238(4833):1563–1565 23. Choi WY, Gemberling M, Wang J, Holdway JE, Shen MC, Karlstrom RO, Poss KD (2013) In vivo monitoring of cardiomyocyte proliferation to identify chemical modifiers of heart regeneration. Development 140(3):660–666. https://doi.org/10.1242/dev.088526

Chapter 8 Cardiac MRI Assessment of Mouse Myocardial Infarction and Regeneration Yijen L. Wu Abstract Small animal models are indispensable for cardiac regeneration research. Studies in mouse and rat models have provided important insights into the etiology and mechanisms of cardiovascular diseases and accelerated the development of therapeutic strategies. It is vitally important to be able to evaluate the therapeutic efficacy and have reliable surrogate markers for therapeutic development for cardiac regeneration research. Magnetic resonance imaging (MRI), a versatile and noninvasive imaging modality with excellent penetration depth, tissue coverage, and soft-tissue contrast, is becoming a more important tool in both clinical settings and research arenas. Cardiac MRI (CMR) is versatile, noninvasive, and capable of measuring many different aspects of cardiac functions, and, thus, is ideally suited to evaluate therapeutic efficacy for cardiac regeneration. CMR applications include assessment of cardiac anatomy, regional wall motion, myocardial perfusion, myocardial viability, cardiac function assessment, assessment of myocardial infarction, and myocardial injury. Myocardial infarction models in mice are commonly used model systems for cardiac regeneration research. In this chapter, we discuss various CMR applications to evaluate cardiac functions and inflammation after myocardial infarction. Key words Cardiac MRI, Myocardial infarction, Mouse, Tagging, Strain, Fibrosis, Late-gadolinium enhancement, Dynamic contrast enhancement, Myocardial perfusion, Extracellular volume

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Introduction Small rodent models are an indispensable tool for cardiac regeneration research. Studies in mouse and rat models have provided important insights into the etiology and mechanisms of cardiovascular diseases and accelerated the development of therapeutic strategies. It is essential to be able to quantitively, accurately, and reliably evaluate the cardiac motility and functions to assess the degrees of injury and the therapeutic efficacy. Magnetic resonance imaging (MRI) is a versatile and noninvasive imaging modality with excellent penetration depth, tissue coverage, and soft-tissue contrast; hence it has increasing importance in both the clinical setting and research arena. Proton MRI is appropriate in animal and human imaging because the body is

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_8, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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composed of more than 70% of water. MRI is noninvasive, utilizing low-energy radio-frequency waves in magnetic fields to form images without ionized radiation, and thus MRI allows longitudinal imaging over time for temporal disease and therapeutic progression. MRI offers complete coverage of the whole body or organ volume without limitation in depth penetration as in other imaging modalities, such as optical imaging and ultrasound. MRI has excellent intrinsic soft-tissue contract because the relaxation characteristics of water protons are governed by the microenvironment of each tissue. The modern MRI platforms have made available a wide variety of pulse sequences to sensitize MRI acquisition to many different conditions, such as T1 weighting, T2 weighting, proton density weighting, diffusion weighting, flow encoding, displacement encoding, tissue perfusion, tissue oxygenation sensitivity, as well as rapid imaging for mechanical motions. In addition, various classes of MRI contrast agents are available for blood pool, infarction, cellular, and molecular imaging. Choosing a specific combination of relaxation and contrast mechanisms has broadened the diagnostic capabilities of MRI in both clinical and preclinical settings [1]. MRI is extremely versatile and has become in itself a multimodal imaging tool for structural and functional imaging. Cardiac MRI (CMR) is versatile and capable of measuring different aspects of systolic and diastolic cardiac functions [2], both global and regional, and is an emerging modality for quantitatively and accurately measuring cardiac regeneration outcome evaluation. Cine MRI can precisely image cardiac morphology and mass and can measure global systolic functions, such as stroke volume (SV) and ejection fraction (EF), for both left and right ventricles. The first-pass perfusion with gadolinium (Gd)-based contrast agents is capable of measuring myocardial perfusion and health of coronary vasculature in resting, stressed, and various pathological states. Late gadolinium enhancement (LGE) can delineate myocardial viability, necrosis, and infarction territories. Extracellular matrix expansion is a key element of ventricular remodeling and a potential therapeutic target. Contrast MRI can quantify extracellular volume (ECV) and fibrosis noninvasively in live animals. CMR is also capable of accessing valvular diseases, fat deposition, water contents, scarring, fibrosis, tumors, and blood flow, and can also perform MR angiography. In addition, diffusion tensor imaging (DTI) allows the tracking of myocardial fiber architecture and the quantification of myocardial fiber disarray after cardiac injury or diseases. With the advancement of high-field magnets and coils to increase the resolution and sensitivity, together with the increasing availability of various contrast agents, CMR is now becoming ideally suited for in vivo evaluation of cardiac functions in small animal models. This overview outlines the multi-parametric CMR methodology to access myocardial infarction in mouse models for cardiac injury and regeneration.

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Materials

2.1 T1-Weighted GdBased Contrast Agents (See Note 1)

1. The dosage for MultiHance is 0.1 mmol/kg bodyweight. 2. Dilute the clinical grade MultiHance (gadobenate dimeglumine injection, 529 mg/mL, Bracco, Inc.) to 1:50 concentration with sterile saline for mouse experiments. 3. Use 0.1 cm3/10 g of bodyweight of the 1/50 diluted MultiHance solution. For a 25 g adult mouse, use 0.25 cm3 of the 1/50 freshly diluted MultiHance solution. 4. MultiHance solution is light sensitive. Protect the diluted solution from light. 5. Use only freshly diluted solution. Dilute only small aliquots. Diluted solution is less effective after a few hours.

2.2 T2∗-Weighted Iron Oxide Nanoparticles

1. Iron oxide particles (see Note 2), such as micrometer-sized iron oxide particles (MPIO), need to be washed in sterile phosphate-buffered saline (PBS) prior to injection. 2. Draw 1 mL of MPIO stock solution into a 15 mL Falcon tube. Draw 10 mL of sterilized PBS solution into the tube. Invert and mix the solution gently ten times. 3. Put the tube near a magnet and let the MPIO particles precipitate. 4. Discard the supernatant by using a pipette. 5. Repeat washing twice. 6. Regular application dosage for iron oxide particles is 1.0–1.5 mg/mL. 7. MPIO particles tend to precipitate at the bottom of the tubes. Vortex well to form a homogenous suspension before injection.

3

Methods

3.1 Animal Preparation for In Vivo CMR

1. Give all mice general inhalation anesthesia with isoflurane (see Notes 3 and 4) for in vivo CMR. 2. Place a mouse into a clear plexiglass anesthesia induction box that allows unimpeded visual monitoring of the animals. Achieve induction by administering 2–3% isoflurane mixed with oxygen for a few minutes. Monitor the depth of anesthesia by toe reflex (extension of limbs, spine positioning) and respiration rate.

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3. Once the plane of anesthesia is established, maintain it with 1–2% isoflurane mixed with oxygen via a designated nose cone and transfer the mouse to the designated animal cradle for imaging. 4. Monitor the heart rate, respiration rate, and body temperature by continuous electrocardiogram (ECG) and respiration monitoring system throughout the imaging process. 5. Monitor the core body temperature by a rectal fiber-optic probe and maintain it at 37
C  0.5
C (see Note 5) by circulating warm water inside the animal bed or feedbackcontrolled MR-compatible air heater module for small animals (SA Instruments. Inc. Model 1025). 6. Place a pneumatic sensor pillow in between the cradle and one side of the mouse’s abdomen. Monitor respiration with the pneumatic sensors coupled to a pressure transducer. 7. Monitor the ECG with two subcutaneous MR-comparable Pt needle electrodes connected with MR-compatible ECG monitoring and gating system (MR-compatible Small Animal Monitoring & Gating System, SA Instruments. Inc. Model 1030). 8. If cardiac and/or respiratory gating is needed for the CMR, the same ECG and respiration signals can be used to synchronize the MR signal acquisition to the R-wave and the exhalation period of the respiration. 3.2

CMR Preparation

1. Carry out in vivo cardiac MRI (CMRI) on a Bruker Biospec 70/30 system spectrometer (Bruker Biospin MRI, Billerica, MA USA) operating at 7 T (see Note 3), equipped with an actively shielded gradient system and a quadrature radiofrequency (RF) volume coil with an inner diameter of 35 mm. 2. Acquire cine CMR with or without ECG and respiration gating. 3. Once the anesthetized mouse is in place with the proper physiological monitoring, center the mouse in the RF coil which is centered in the gradient and magnetic field. 4. Tune the coil to match the optimal circuitry impedance. 5. Shim the magnetic field and gradient to optimize the magnetic field homogeneity to reduce the background gradient inhomogeneity. 6. Acquire fast low-resolution scout pilot images to position the mouse heart to the center of the magnetic field and gradients. 7. Acquire a fast triplane localizer sequence to define the axial, sagittal, and coronal plane of the mouse. 8. Acquire a series of oblique imaging planes from the triplane views to define the short-axis, two-chamber, and four-chamber long-axis views of the mouse heart.

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Cine CMR

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1. Acquire a multi-planar short-axis cine MRI (see Note 6) to cover the whole heart volume. Typically, for an adult mouse with bodyweight around 25 g, field of view (FOV) is around 2.0–2.5 cm, slice thickness 0.8–1.2 mm, acquisition matrix ¼ 256  256, in-plane resolution ¼ 78–97 μm, typical flip angle (FA) ¼ 30
, echo time (TE) ¼ 2.2 ms, and repetition time (TR) ¼ 12 ms. 2. In addition to short-axis views, acquire two-chamber and fourchamber long-axis cine with similar parameters. 3. Export the MRI data to the universal Digital Imaging and Communications in Medicine (DICOM) format that can be viewed in any open-source DICOM viewers, such as Horos (https://horosproject.org/). 4. Derive various global systolic functional parameters (Table 1) from the multi-slice short-axis and long-axis cine MRI. 5. Define the epicardial and endocardial myocardium borders at the end-systole (ES) and the end-diastole (ED) for each imaging plane by an operator with common open-source imaging processing software, such as Image J (https://imagej.nih.gov/ij/) and Fiji (https://imagej.net/Fiji). 6. Use Table 1 to calculate left ventricular volume (LVV), stroke volume (SV), ejection fraction (EF), cardiac output (CO), Table 1 LV anatomical and functional parameters derived from in vivo cine MRI Parameter

Measurement/calculation

SAA

Area encompassed the epicardium in the SA view

LAA

Area encompassed the epicardium in the LA view

LVVED

Σ (SAA  SLTH) over all slices at ED

LVVES

Σ (SAA  SLTH) over all slices at ES

SV

LVVED  LVVES

EF

SV/LVVED  100%

CO

EF  HR

RS

ðDiameter out SA EDdiameter out SA ESÞ diameter out SA ED

LS

ðLA out EDLA out ESÞ LA out ED

RWT

ðDiameter out SAdiameter in SA Þ 2

LWT

ðLWTs ESLWTs EDÞ LWTs ED

 100%

LVW (wall volume)

½43ðSA outÞðLA outÞ

½43ðSA inÞðLA inÞ

2

LV mass

LVW  1.05



 100%

2

 100%

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radial shortening (RS), longitudinal shortening (LS), LV wall volume, longitudinal wall thickening (LWT), and radial wall thickening (RWT). 7. Calculate LV mass by applying myocardial density as 1.05 g/mL. 8. Directly apply similar parameters (RV) without geometrical assumption. 3.4

Tagging MRI

to

right

ventricle

1. Tagging MRI requires ECG and respiration gating. 2. Tagging MRI is commonly acquired with SPAMM pulse sequence (see Note 7). 3. For a typical adult mouse with 25 g body weight, acquire typical tagging MRI with the following parameters: FOV ¼ 2.5 cm, SLTH ¼ 1.1 mm, in-plane resolution ¼ 97 μm, tag distance ¼ 0.8 mm, tag thickness ¼ 0.15 mm, TE ¼ 2.514 ms, TR ¼ 14.612 ms, and FA ¼ 30
. 4. Export multi-planar short-axis tagging MRI to DICOM format to analyze it by various software, such as the Food and Drug Administration (FDA)-approved Harmonic phase (HARP) algorithm by Myocardial Solutions, Inc. (https:// www.myocardialsolutions.com/).

3.5 Dynamic Contrast Enhancement (DCE)

1. Fast T1-weighted dynamic CMR (see Note 8) with sub-minute temporal resolution is required for measuring myocardial perfusion with dynamic contrast enhancement (DCE). 2. Acquire an intravenous line (I.V.) for single bolus administration of (Gd)-based contrast agent MultiHance while the mouse is in the magnet. 3. Perform I.V. cannulation at the femoral vein or tail vein, using PE-10 tubing. 4. Measure the PE-10 tubing to cover the length from the 5-Gauss fringe field line to the mouse in the center of the magnet. Carefully measure the dead volume of the PE-10 line. Typical length is around 1.5–3 m, whereas the dead volume of the line is around 0.4–0.7 cm3. 5. (Optional) Coat the inner surface of the PE-10 line with clinical grade heparin to prevent blood clotting before cannulation. 6. Use Discofix three-way stopcock for convenient injection. 7. Before scanning, carefully fill the tubing with the desired volume of the diluted Gd solution. 8. To administer the Gd bolus, flush the Gd bolus by the same volume of saline. 9. Synchronize the fast dynamic MRI acquisition with the Gd bolus injection. 10. Quantify the fast wash-in and washout dynamic of the Gd contrast flowing through the myocardium on the T1-weighted time series.

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1. For LGE imaging (see Note 9), use the same I.V. and Gd bolus as in Subheading 3.5, but acquire T1-weighted images 15 min after the single Gd bolus injection to healthy myocardium when it has already returned to baseline intensity but the infarcted myocardium still retains high intensity of T1 contrast. 2. For multi-planar LGE imaging, a bolus subcutaneous administration of Gd contrast agent can prolong the time period when the infarcted myocardium still maintains bright contrast for longer imaging.

3.7 Extracellular Volume (ECV)

1. To calculate ECV (see Note 10), measure pre-contrast and post-Gd contrast myocardial T1 (or R1) in the blood and myocardium, as well as hematocrit. 2. To sample blood for hematocrit reading, first anesthetize the mouse. 3. Use a small prick with a lancet/sterile 27G needle to create a bead of blood from the tail, lateral saphenous, or facial vein. 4. Place a Hemato-tube on the blood bead to acquire 1 μL of blood. 5. Seal the Hemato-tube with Critoseal. 6. Apply gauze firmly to the mouse’s vein from which the blood was harvested for at least 1 min to stop the bleeding. 7. Place the Hemato-tube inside the micro-hematocrit centrifuge to be processed. 8. After spinning down, compare the length of the pallet in the Hemato-tube to the hematocrit reading chart to obtain the hematocrit values. 9. Recover the mouse and return to its cage. 10. To measure T1 (R1) of blood and myocardium, acquire a cine MRI with four flip angles (FA) to map the blood and myocardial T1 (or R1). 11. Administer single-bolus Gd. 12. Repeat T1 (R1) measurement after Gd contrast.

3.8 Cellular MRI for Intramyocardial Inflammation

1. Inject desired iron oxide nanoparticles (see Notes 2 and 11) 18–24 h prior to MRI sessions. 2. Acquire T2∗-weighted MRI with TE ¼ 4.5 ms or longer. The rest of the parameters can be similar to Subheading 3.3. 3. Data exporting and viewing processes are the same as in Subheading 3.3.

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Notes 1. Commonly used gadolinium (Gd)-based T1-weighted MRI contrast agents are available as clinical intravenous injection solutions, such as MultiHance (gadobenate dimeglumine injection, 529 mg/mL, Bracco, Inc.), and ProHance (gadoteridol injection, 279.3 mg/mL, Bracco, Inc.). The gadobenate ion is cleared from plasma by renal and biliary excretion [3]. Repetitive Gd-based contrast agent usage can lead to acute kidney injury, chronic renal dysfunction, and nephrogenic systemic fibrosis. In animals, 34 applications of MultiHance were found to cause toxicity [4]. Up to 20 applications are found to be safe without adverse side effects. 2. Paramagnetic [5–10] and super-paramagnetic iron oxide (FenOm) particles [11] are able to cause enormous 1H signal alterations, resulting in considerable hypo-intensity or negative contrast in T2∗-weighted MR images; thus, they have been utilized for noninvasive in vivo cell tracking. The dipole “blooming effect” [6, 7, 12] of the iron oxide particles causes significant background gradient perturbation to make even single cells detectable with in vivo T2∗-weighted MRI [13, 14], even though the size of the target cell is much smaller than an imaging voxel, thus being suitable for in vivo cell tracking [15–27]. Iron oxide particles (Fig. 5a) have different varieties suitable for different research goals. Depending on the types of coating, hydrodynamic diameters, and iron core sizes, various iron oxide particles have different zeta potentials, ranges of relaxivity, as well as different blood half-lives, biokinetics, and extrusion time span in the body. Superparamagnetic iron oxide (SPIO) and ultrasmall superparamagnetic iron oxide (USPIO) nanoparticles consist of compatible iron core sizes (4–8 nm), iron content (3–9  1019 g of Fe per particle), and similar surface coating materials, yet different hydrodynamic diameters (15–30 nm for USPIO and 120–400 nm for SPIO). SPIO and USPIO are usually coated with poly-sugars, such as dextran, so they are biocompatible and biodegradable [16, 17]. They are usually metabolized and absorbed into the body iron pool after about 1 month, depending on the specific structure and coating materials. On the other hand, the much larger (~1 to a few μm) micrometersized particles, MPIO [14, 17, 19, 28, 29], are usually composed of a magnetite core and an inert coat, such as polystyrene-divinyl-benzene, which can cause much larger signal perturbation and can stay in the body for a long time, making it suitable for longitudinal studies for an extended period of time.

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3. An average adult mouse (body weight 20–30 g) and rat (body weight 250–350 g) weigh only 0.03% and 0.5% of an average human (body weight 60 kg), respectively. In order to achieve a similar imaging quality to human MRI, small animal MRI scanners need to increase the signal-to-noise ratio (SNR) and detection sensitivity by 200–3000 to compensate for the small imaging voxels for high spatial resolution. The SNR is proportional to the power of 7/4 with the magnetic field (B0^ (7/4)), and affected by gradient strength, RF coil geometry, inductance, quality factor, and sample dissipation [30– 32]. Therefore, it is important to have high magnetic field strength to increase SNR and resolution. However, too high of a magnetic field strength with lengthened T1 and greater susceptibility artifact will decrease SNR. Therefore, most mouse and rat MRI studies are conducted at field strengths of 4.7 to 11.7-Tesla [33–35]. However, currently the 11.7-T animal MRI is only available as vertical bore, which requires placing animals in vertical stand-up position for imaging. It is reported [36] that mouse cerebral blood flow decreased by 40% when changing from horizontal position to vertical position, due to orthostatic response. How this positioning will affect the cardiovascular function and blood flow is not known. Therefore, the horizontal bore magnet with 7 and 9.4-T offers the best trade-off for mouse imaging. In addition, suitable RF coil size and geometry are important. For mouse cardiac MRI studies, optimal volume coils with inner diameter of 25–35 mm are typically used. Surface coils usually deliver better SNR, but the depth coverage can be limited depending on the coil size and geometry. All images presented in this overview were acquired with a 7-Tesla preclinical MRI with the standard 35 mm quadrature volume coil for mice and 72 mm bird-cage volume coil for rats. 4. Animals need to be anesthetized during MRI acquisition. Typically, the anesthesia level for mice is maintained with 1.0–1.5% isoflurane mixed with 100% oxygen or 20% oxygen/80% nitrogen. Anesthesia induction is typically first achieved by 2–3% isoflurane mixed with oxygen in an induction box for 1–3 min. The depth of anesthesia is monitored by toe reflex, extension of limbs, and spine positioning. Once the plane of anesthesia is established, the mouse is placed on a designated animal bed for imaging and the anesthesia is maintained by 1.0–1.5% isoflurane with 100% oxygen via a nose cone or ventilation. Some studies may choose 20% O2/80% N2 or 20% O2/80% N2O inhalation gas combination. The advantages of volatile inhalable anesthesia, such as sevoflurane and isoflurane, are fast acting and it is easy to manipulate the depth and timing of anesthesia and recovery. Other commonly used anesthesia for

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mouse imaging are injectable ketamine and barbiturates, such as pentobarbital, phenobarbital, and fentanyl. The types and doses of anesthesia depend on the animal strains and the studies. The types, depth, and dosages of anesthesia [37–39] are important because anesthesia can suppress cerebral and brain stem activities [37–39], mucociliary clearance in the airways [40, 41], and vascular constriction and hemodynamic functions [42–44], which can greatly affect cardiac functional measurements. Therefore, it is crucial to keep the general anesthesia conditions as light as possible and consistent from animal to animal to ensure accurate comparison among different animal groups. 5. It is important to maintain the physiological body temperature (37
C  0.5
C) throughout the CMR acquisition because body temperature greatly affects heart rates and hemodynamic functions. The body temperature is typically monitored by an optical rectal temperature probe which is used for feedback control of a warm air blower or warm water circulation padding around the animal to maintain the physiological body temperature. The respiration is continuously monitored by placing a small pneumatic pillow under the animal’s diaphragm which is connected to a magnet-comparable pressure transducer feeding to a physiological monitoring computer equipped with respiration waveform measuring software. The respiration waveform is automatically processed to detect inspiration, expiration, and respiration rate. To overcome cardiac and respiration motion artifact, CMR can be acquired with prospective or retrospective gating. Prospective gating synchronizes the CMR acquisition with the electrocardiogram (ECG) and respiration motion of the animals. K-space lines are only acquired at specific cardiac and respiration phases. This gating method is useful for most studies. However, it has the drawbacks of prolonged acquisition time and is unable to accommodate physiological fluctuation over time. Another approach is retrospective gating [45, 46] or self-gating strategies [47–49]. This approach uses deconvolution of cardiac and respiration navigator signals to reconstruct the CMR without motion artifact after the full k-space data have been acquired. This allows free-breathing cine acquisition with no ECG or respiration gating needed and is available for preclinical scanners for mice and rats [45, 49–51]. 6. Multi-planar free-breathing cine MRI [52, 53] covering the whole heart volume with retrospective gating [45, 49] is commonly used to evaluate systolic LV functions, volumes, and mass. Figure 1 shows a free-breathing gate-free cine MRI of a wild-type (WT) C57BL6 control mouse (Fig. 1a–d) and a

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Fig. 1 Mouse cine MRI. (a–d) Cine MRI of a wild-type C57BL6 control mouse (e–h) cine MRI of a mildly diseased mouse with toxin-induced hypertrophic cardiomyopathy (HCM) at 10 weeks of age acquired at 7-T with a 78 μm in-plane resolution. (a, b, e, f) Long-axis four-chamber views, (c, d, g, h) mid-level short-axis views, (a, e, c, g) end-diastole (ED), (b, f, d, h), end-systole (ES). LV—left ventricle, RV—right ventricle, Ao— aorta, L. Comm Ar.—left common carotid artery, R. Comm Ar.—right common carotid artery, ED—enddiastole, ES—end-systole, Cont—control, HCM—hypertrophic cardiomyopathy, L—left, R—right

mouse with toxin-induced hypertrophic cardiomyopathy (HCM, Fig. 1e–h), showing short-axis (Fig. 1c, d, g, h) and long-axis four-chamber (Fig. 1a, b, e, f) views acquired at 7-T. The bright-blood imaging condition yields very good blood myocardium contrast for quantification. The 78 μm in-plane resolution is sufficient to resolve anatomical structures in mice. Multi-planar cine MRI [52, 53] covering the whole heart is advantageous over 2D echocardiography in anatomical and functional evaluation in mice. The 2D echocardiography in mice usually measures ejection fraction (EF) in single shortaxis or long-axis imaging planes and then calculates the EF with 1D linear M-mode or 2D B-mode with mathematical models, such as modified Simpson’s method or an ellipsoid model using biplane data [54–57]. This standard vendor-provided function at the preclinical ultrasound system is quick and valid for anatomical and functional measurements in the left ventricle (LV). However, this model-based approach is difficult in right ventricle (RV) evaluation because the ellipsoidal model does not apply to RV. More importantly, in the ischemic infarction models (Fig. 3g, h), the myocardial infarct site atrophies over time, and LV shape changes over time due to LV remodeling. With the distorted LV morphology deviating from the ellipsoidal shape, this 1D or 2D model-based measurements become inaccurate. On the other hand, the multi-planar cine MRI with

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full coverage of the 3D heart volume allows more precise and direct measurements of the blood and myocardium volumes. The epicardial and endocardial boundaries of the myocardium can be traced at the end-systole (ES) and end-diastole (ED), typically on short-axis (SA) planes. The LV blood volume (LVV) at ED and ES can then be obtained by summation of the blood area (SAA) times the imaging slice thickness (SLTH) in the LV chamber over all the imaging planes. The stroke volume (SV) is defined as the difference of LVVED and LVVES. Ejection fraction (EF) is defined as SV/LVVED. Cardiac output (CO) is defined as the multiplication of EF and heart rate (HR). The same method can be applied to RV or distorted LV because no shape geometry needs to be assumed in the calculation. The typical values for 10-week-old C57BL6 mice are as follows: LVVED ~40 μL, LVVES ~15 μL, SV ~25 μL, HR ~500 beats per min, CO ~12.5 mL/min, and EF ~55–65% [33]. Systolic functional parameters include radial shortening (RS), longitudinal shortening (LS), radial wall thickening (RWT), and longitudinal wall thickening (LWT). RS and LS are defined as percent changes of short-axis (SA) and long-axis (LA) LV diameters, respectively. The LV wall volume can be calculated by differences between the epicardium and endocardium at ES and ED. RWT and LWT are defined as percent changes of SA and LA wall thickness at ES and ED. The literature value of myocardial specific density is 1.0526 g/mL [58]. Using the 1.05 g/mL value to calculate LV mass is shown to be fairly accurate in estimating mouse heart mass [33, 59, 60] in both normal mouse hearts and those with hypertrophy [59]. For mean mouse heart mass ~75 mg, the LV mass measured by the noninvasive in vivo CMR showed a mean error of only 0.9 mg [33, 60]. Table 1 summarizes the common cardiac parameters derived from multiplanar cine MRI. 7. With localized ischemic infarction, EF may or may not significantly decrease. Patients with heart failure with preserved ejection fraction (HFpEF) do not exhibit depressed EF. Diastolic dysfunction refers to abnormal diastolic filling properties of the left ventricle regardless of whether systolic function is normal, or the patient has symptoms [61, 62]. After myocardial infarction (MI), it is important to evaluate the region and extent of the myocardial damage. Regeneration therapy strategy often induces new cardiomyocyte proliferation. It is imperative to be able to quantitatively assess whether the newly born cardiomyocytes have functionally incorporated into the existing myocardium or these cardiomyocytes just from inner nonfunctional heart mass. The global systolic functional indexes are often not

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sensitive enough to detect the mentioned changes. More sensitive functional gauges are needed to accurately evaluate the extent of MI and the effectiveness and efficacy of cardiac regeneration therapy after MI. Tagging MRI [63–66] followed by strain analysis [67] is found to be the most sensitive tin evaluating diastolic dysfunction, regional wall motion, extent of post-MI myocardial damage, and regional function evaluation for cardiac regeneration. Tagging MRI [63–66] spatially modulates water spins magnetically to generate tags (Fig. 2a–l) on myocardial tissue to allow

Fig. 2 Mouse tagging MRI and strain analysis for regional wall motion and infarction evaluation acquired at 7-T. (a–l) Multi-planar tagging MRI of a wild-type C57BL6 mouse acquired at the (a–f) end-diastole (g–l) and the end-systole from apex (a, g) to base (f, l). (m) Diagram of a left ventricle (LV) showing the tensor direction of the circumferential strain (Ecc), radial strain (Err), and the longitudinal strain (Ell). (n–q) End-systolic Ecc strain maps overlaying tagging MRI of a mouse heart 28 days after anterolateral ischemic injury, showing four short-axis imaging planes. The red arrowheads point to the infarct site. (r) Peak Ecc and (s) percentage of transmural infarction of the infarcted mouse heart represented with the AHA 17-segment model [143]. Segments: 1-basal anterior, 2-basal anteroseptal, 3-basal inferoseptal, 4-basal inferior, 5-basal inferolateral, 6-basal anterolateral, 7-mid-anterior, 8-mid-anteroseptal, 9-mid-inferoseptal, 10-mid-inferior, 11-mid-inferolateral, 12-mid-anterolateral, 13-apical anterior, 14-apical septal, 15-apical inferior, 16-apical lateral, 17-apex

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evaluation of regional wall motion. Strains [67, 68] are values that quantify the extent of ventricular deformation throughout cardiac phases: stretching/elongation or compression/ shortening. Our previous study [19] in a heterotopic working heart transplantation model showed that diastolic dysfunction proceeded systolic dysfunction during allograft rejection after heart transplantation. The standard global systolic function parameters, such as EF and SV, did not change until the late rejection stage when the irreversible tissue damage has already occurred, and thus too late for intervention. On the other hand, strain derived from tagging MRI was most sensitive in detecting early regional myocardial changes [19]. MR tagging can detect regional differences in myocardial function post-MI in mice [69]. Longitudinal strain was found to be able to detect diastolic dysfunction and HFpEF in both patients [70–73] and mice [74]. Strains are categorized into two main classes in relation to the heart axes: normal strains are defined in relation to the short-axis planes, and principal strains are defined in relation to the direction of the myocardial fiber bundles. The most commonly used classification is the three orthogonal straintensor sets that define normal strains (Fig. 2m): circumferential strain (Ecc), radial strain (Err), and longitudinal strain (Ell), in which strain tensors are tangent to the epicardium surface, perpendicular to the epicardium surface toward the center of the LV, and perpendicular to the short-axis plane along the long axis of the LV, respectively. Several methods have been implemented for strain analysis from tagging MRI, such as harmonic phase (HARP) analysis [67, 68, 75–78], displacement-encoded imaging with stimulated echoes (DENSE) [79, 80] following spatial modulation of magnetization (SPAMM) [81, 82], or delayed alternating with mutations for tailored excitation (DANTE) [83, 84] tagging. The HARP method (Myocardial Solutions, Inc.) has been FDA approved for clinical applications. HARP is capable of calculating strains in all three heart layers (epimyocardium, mid-wall, and endomyocardium), and delineating regional strain with the AHA 17-segment convention to show regional wall motility to evaluate the diastolic function and regional wall motion for MI and regeneration evaluation. HARP has been shown in the MultiEthnic Study of Atherosclerosis (MESA) clinical study to yield good inter-observer and intra-observer consistency in strain analysis [68]. Strain rates reflect diastolic functions. In addition

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to short-axis planes, four-chamber long-axis slices can be used to derive a longitudinal strain (Ell). The Treatment of Preserved Cardiac Function Heart Failure with an Aldosterone Antagonist (TOPACT) clinical trial showed that Ell is most sensitive in showing ventricular abnormality with preserved EF [85–88]. Furthermore, the rotation and torsion of the heart can be analyzed at each level from the tagging MRI. Normal hearts exhibit systolic wringing motions with clockwise rotations at the base and counterclockwise rotations at the apex; the helical arrangement of myocardial fibers that run in opposite directions at the epicardium and endocardium generates systolic twist [89–95]. It has been found that abnormal systolic twisting and diastolic recoil are associated with pathological cardiac conditions [96], such as hypertension [93], aneurysm [97], type I diabetes [98], and heart failure [99, 100]. Torsion of the heart can be derived from the relative rotation between the apical and basal short-axis slices. It can potentially be a sensitive evaluation index for MI and cardiac regeneration. 8. Gadolinium (Gd)-based T1 contrast agents displayed differential tissue wash-in and washout kinetics (Fig. 3a) for healthy and infarcted myocardium, and thus can be used to evaluate myocardial perfusion, infarction, and fibrosis. Gd flows in myocardium with good perfusion quickly and clears from the heart quickly (Fig. 3a, blue) within 1–3 min after the single-bolus Gd injection. On the other hand, the infarcted myocardium with poor perfusion and damaged cardiomyocytes displays much slower wash-in and washout kinetics (Fig. 3a red). Figure 3c– f shows various time frames after the single intravenous Gd bolus injection in a rat heart with ischemic reperfusion injury (IRI) with 45-min transient occlusion of the left anterior descending (LAD). The infarct core showed lower intensity than the rest of the unaffected myocardium within the first 5 min. Time-to-peak contrast (TPC) expressed with the AHA 17-segment model (Fig. 3b) can clearly delineate the infarcted regions. 9. Late gadolinium enhancement (LGE), acquiring T1-weighted CMR 15–20 min after the intravenous Gd bolus when the contrast enhancement in the healthy myocardium has already subsided but the enhancement in MI or fibrosis persists, can be used to noninvasively detect MI or fibrosis in vivo. Myocardial atrophy was observed in the infarct site of a mouse 28 days after

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Fig. 3 Dynamic contrast enhancement (DCE) for myocardial perfusion evaluation and the late gadolinium enhancement (LGE) for infarction assessment acquired at 7-T. (a) Schematic presentation of gadolinium contrast enhancement in the heart for healthy myocardium (blue) and infarction (red). (b–f) Myocardial perfusion assessed by dynamic contrast enhancement (DCE) of a rat heart with ischemic-reperfusion injury after 45-min reversible occlusion of the left anterior descending (LAD). (b) Time-to-peak contrast (TPC) in seconds of the gadolinium contrast enhancement for a rat heart after 45-min LAD occlusion expressed with the AHA 17-segment model [143]. See Fig. 2 legend for the segment nomenclature. (c, d) End-diastolic DCE images of a rat heart after 45-min LAD occlusion acquired before (c) and 1 min 21 s (d), 2 min 3 s (e), and 5 min 54 s (f) after a single bolus of gadolinium contrast agent administration. The white arrows point to the infarct core. (g–i) CMR of a WT mouse acquired at 28 days after ischemic infarction. The yellow arrowheads point to the infarct site. (g, h) Cine MRI acquired at the end-diastole (g) and end-systole (h). (i) T1-weighted image of the same mouse acquired at 15 min after a single bolus of gadolinium contrast agent administration. The hyperintensity can be seen at the infarct site (yellow arrowhead) indicating fibrosis

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MI (Fig. 3g, h) with epicardial fibrotic tissue developed, which exhibited bright hyperintensity with LGE (Fig. 3i). 10. Myocardial fibrosis, characterized by expansion of the myocardial extracellular matrix and accumulation of interstitial collagen, is the hallmark of pathologic remodeling. However, LGE can only detect regional fibrosis foci, but cannot quantify diffuse fibrosis. It has been reported that the myocardial extravascular extracellular volume fraction [101–104] can quantify diffuse fibrosis not readily detectable by conventional late gadolinium enhancement. Extracellular matrix expansion is a key element of ventricular remodeling and a potential therapeutic target [105]. Myocardial extracellular volume (ECV) and fibrosis can be noninvasively quantified by CMR with quantitative longitudinal relaxation time constant T1 mapping [105–109]. ECV quantified by CMR has been found to be associated with diabetes [110], heart failure [111], and active idiopathic systemic capillary leak syndrome [112]. ECV evaluation was found to provide accurate prognosis for severity of the heart failure, hospital stay length, and death [111, 113]. ECV is quantified by pre- and post-Gd contrast T1 or R1 mapping in conjunction with hematocrit: ECV ¼ (1  hematocrit) (R1myopost  R1myopre)/(R1bloodpost  R1bloodpre) or in which longitudinal relaxation time (R1) is the inverse longitudinal relaxation time (T1). Hematocrit reading in mice can be easily obtained by a small droplet of venous blood collected from the lateral saphenous vein at the time of CMR. Myocardial T1 is commonly quantified by the modified Look-Locker inversion recovery sequence (MOLLI) [106, 107, 114–116] before and after Gd contrast administration. MOLLI in mouse imaging has a drawback of longer scanning time. We have stablished fast free-breathing gate-free T1/R1 mapping (Fig. 4) with varied flip angle (FA). It is capable of quantifying ECV in a rat IRI model (Fig. 4). 11. MI triggers robust local inflammatory responses [117– 122]. Inflammation plays a central role in acute MI, IRI [121, 123–126], post-MI cardiac repair, and subsequent LV remodeling [119–122, 127]. The amplitude of inflammation and the timely resolution affect the post-MI remodeling and the quality of cardiac repair. Early preclinical finding that antiinflammatory corticosteroid treatment could decrease infarct size [128] led to clinical trials using anti-inflammatory agents, such as methylprednisolone, to treat MI patents [129], but

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Fig. 4 Fibrosis quantification by extracellular volume (ECV) of a rat heart after ischemic reperfusion injury with 45-min transient occlusion of LAD. (a, c, e) Pre-contrast (b, d, f) post-gadolinium contrast. (a, b) CMR with varied flip angle (FA): 3
, 19
, 22
, and 28
. (c, d) T1 maps derived from the four FA. (e, f) R1 maps derived from the four FA. (g) ECV calculated from a–f and hematocrit measurements. Panel a: colored map; panel b and c: different mask options. White arrows point to the infarct site (adapted with permission with the dissertation [144] of Dr. Anthony G. Christodoulou)

resulted in catastrophic outcome. Growing evidence shows that monocytes/macrophages are imperative for infarct healing [130–132]. Both insufficient and excessive inflammation are detrimental to post-MI cardiac repair. Immune cells, particularly macrophages and monocytes, are emerging therapeutic targets [117, 127]; however, the noninvasive clinical tools for assessing their presence in the myocardium are lacking [117, 133, 134]. The systemic biomarkers in blood might not reflect the inflammation status in

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Fig. 5 Cellular MRI for imaging intramyocardial inflammation. (a) Examples of various iron oxide particles, including ultrasmall super-paramagnetic iron oxide (USPIO), super-paramagnetic iron oxide (SPIO), and micrometer-sized particles (MPIO). (b, c) T2∗-weighted MRI of a rat heart 2 days after ischemic reperfusion injury with 45-min transient occlusion of LAD, showing a short-axis view (b) and a long-axis view (c). The white arrows point to hypointensity after in situ MPIO labeling of macrophages. (d) 3D volume rendering of the macrophage infiltration (red) with in situ MPIO labeling

the myocardium. Diagnostic imaging tools that can assess myocardial inflammation will be very beneficial in this endeavor [117, 135–140]. We have previously established in vivo cellular MRI to detect intramyocardial inflammation in MI and rejection models [15–17, 19, 141, 142]. Post-MI inflammation can be detected in vivo as hypointensity in T2∗-weighted MRI (Fig. 5b–d) after in situ labeling of macrophages by intravenous administration of iron oxide particles. Cellular MRI provides noninvasive quantification for intramyocardial inflammation. Integrated multi-parametric CMR (Fig. 6) simultaneously evaluating intramyocardial inflammation (Fig. 6a–c), regional wall motion, and strain by tagging (Fig. 6d–f) and myocardial perfusion (Fig. 6g–i) provides comprehensive evaluation of MI and can quantify the effectiveness of cardiac regeneration.

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Fig. 6 Integrated multi-parametric CMR of three rat hearts with different degrees of ischemic reperfusion injury after 45-min transient LAD occlusion. (a–c) In vivo cellular MRI of MPIO-labeled macrophages (MΦ). (d–f) Systolic Ecc strain maps overlaid on tagging MRI. (g–i) Frist-pass myocardial perfusion map by DCE. Arrows point to the infarct sites

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Part II Ex Vivo and In Vivo Approaches

Chapter 9 Isolation, Culture, and Live-Cell Imaging of Primary Rat Cardiomyocytes Marina Leone and Felix B. Engel Abstract The heart is a complex organ consisting of a variety of different cardiomyocytes (ventricular vs. atrial, left vs. right ventricular, working vs. nodal) as well as other cell types, including endothelial cells and vascular smooth muscle cells. Pericytes, neurons, and immune cells are less abundant, yet still important. Whereas cardiomyocytes account for around 75% of the heart volume, 50–70% of the cells in the heart are non-myocytes. This complexity of the heart underlines the difficulties in interpreting data obtained in vivo. In the field of cardiac regeneration, it remains unclear whether it is possible to induce a significant number of cardiomyocytes to proliferate and whether the often-observed improvement in cardiac function after experimental therapies is due to the induction of cardiomyocyte proliferation. Therefore, the reductionist approach inherent to cultures of isolated cells continues to be of great importance, even though it is important to study heart disease in vivo due to interactions of the different cell types. Cultured cardiomyocytes allow for easy manipulation of cell behavior (e.g., cell division) and its analysis (e.g., live-cell imaging). In addition, isolated cells in culture are a valuable tool for pharmacological and toxicological studies. This chapter offers a practical guide to isolate and culture primary neonatal and adult rat cardiomyocytes and a detailed protocol for live-cell imaging of embryonic and neonatal cardiomyocytes. Key words Cardiomyocytes, Live-cell imaging, Neonatal, Adult, Rat, Langendorff-perfusion, MACS, SADO Mix

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Introduction In recent years, accumulated data have suggested that reentry of cardiomyocytes into the cell cycle is possible [1–3]. Yet, like in the stem cell field, there are controversies whether this approach will be efficient enough to reverse cardiomyocyte loss, for example, after myocardial infarction (MI) [1]. A single episode of MI may result in the loss of one billion cardiomyocytes or more (~25% of total cardiomyocytes) in humans [4]. Yet, there are several reasons why induction of cardiomyocyte proliferation is appealing: fetal mammalian cardiomyocytes can proliferate even though they already contract [5–7]; Newts, zebrafish, and neonatal mice can regenerate their hearts based on cardiomyocyte proliferation [5, 8, 9]; and

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_9, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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adult mammalian cardiomyocytes can be induced to enter the cell cycle and progress into cytokinesis [3, 10]. There are currently no established methods for cardiomyocyte proliferation in vivo [1]. Thus, it is important to verify in vitro that a certain therapy can induce robust neonatal and adult cardiomyocyte proliferation. A large number of markers have been previously used to determine whether a cardiomyocyte divides. Unfortunately, all of these markers are limited, as discussed previously in detail [2]. Thus, live-cell imaging appears to be the only current technology that proves that a certain manipulation induces cardiomyocyte proliferation [11]. In addition, it is not feasible to screen for new inducers of cardiomyocyte proliferation in vivo and, thus, isolated cardiomyocytes are often used as a screening tool before candidate compounds/factors are tested for their regenerative potential in vivo [12, 13]. As alternatives, many labs utilize cardiomyocytes derived from embryonic stem cells (ESCs) or induced pluripotent stem cells (iPSCs) [14–17]. However, ESC- and iPSC-derived cardiomyocytes are, under normal conditions, not well differentiated and represent a fetal cardiomyocyte with an inherent capacity to proliferate [18, 19]. Neonatal rat cardiomyocytes (1–3 days old, P1–P3) are commonly used since the isolation and culture are easier than those of adult rat cardiomyocytes. Yet, P1–P3 cardiomyocytes are also not yet fully differentiated and exhibit a markedly higher proliferative capacity than adult cardiomyocytes [12, 13]. Here, we provide detailed protocols for the isolation and culture of embryonic day 15 (E15), neonatal (P1–P5), and adult rat cardiomyocytes, as well as live-cell imaging of embryonic and neonatal cardiomyocytes that we routinely utilize in our laboratory.

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Materials Prepare all solutions using autoclaved ddH2O and use a 0.22 μm filter for sterile filtration. All procedures are performed at room temperature unless otherwise stated.

2.1

SADO Mix

1. SADO Mix: Add 50 mL of 200 mM HEPES-NaOH (pH 7.6), 50 mL of 1.3 M NaCl, 5 mL of 300 mM KCl, 5 mL of 100 mM NaH2PO4, and 1 mL of 2 M glucose to 390 mL of ddH2O. Sterile-filter the buffer and store at 4
C (see Note 1). 2. Neonatal digestion buffer: Dissolve 25.2 mg of collagenase type II (0.14 mg/mL) and 90 mg of pancreatin (0.50 mg/ mL) in 180 mL of SADO Mix and filter the solution (see Notes 2 and 3). Add 540 μL of 10 mg/mL DNase I solution. 3. Embryonic digestion buffer: Dissolve 12.6 mg of collagenase type II (0.14 mg/mL) and 45 mg of pancreatin (0.5 mg/mL) in 90 mL of SADO Mix and filter the solution. Add 270 μL of 10 mg/mL DNase I and 180 μL of 2 M glucose. Filter the solution (see Notes 2 and 3).

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4. For this protocol, a magnetic stirrer with heating (e.g., IKA C-MAG HS 7) and an electronic contact thermometer (e.g., IKA ETS-D5) are required (Fig. 1a).

A

equipment for SADO Mix protocol

B

equipment for adult cardiomyocyte isolation

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isolated neonatal cardiac cells

heart neonatal C neonatal heart D neonatal E minced ventricles heart ventricles

phase-contrast

actinin/DAPI

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day 3

day 1

G

Fig. 1 Cardiomyocyte isolation: (a) Magnetic stirrer with heating used for the SADO Mix protocol with an electronic contact thermometer and round-bottomed glass tube placed in the finger bowl. (b) An example for a possible setup required for adult rat cardiomyocyte isolation. (c) P3 rat heart (atria indicated by dotted yellow line). (d) P3 rat heart ventricles after removal of aorta and atria. (e) Minced P3 rat ventricles (left) and P3 rat heart ventricles (right) before being minced. (f) Image of a microscopic view (20) of a hemocytometer loaded with a cardiac cell suspension after isolation stained with trypan blue. Yellow arrows indicate examples of cardiomyocytes. (g) P3 rat cardiomyocyte culture isolated with MACS Miltenyi Biotec protocol at day 1 (before changing medium) and day 3 post-isolation on fibronectin-coated coverslips. Green: Cardiac-specific marker (actinin), blue: DNA (DAPI). Scale bars: 200 μm. (h) Schematic drawing of a system to isolate adult rat cardiomyocytes. LF: air trap, EV: valve to release air from the system, WB: water bath, WA: heating coil, K: cannula, T: optional water-jagged reservoir to collect digest solution, AB: waste reservoir, V: valves, P: pumps, GV: gas valves, GF: gas filter, Ca: carbogen, SV: optional valve to bubble collected digest solution

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2.2 Pre-plating and Plating Neonatal and Embryonic Cardiomyocytes

1. Cardiomyocyte pre-plating medium: Add 10% fetal bovine serum (FBS) and 5 mL of 10,000 U/μg/mL of penicillin/ streptomycin solution to 500 mL of DMEM/F-12, GlutaMAX™ Supplement Medium. 2. Cardiomyocyte medium: Add 15 mL of 100 mM Na-pyruvate, 0.5 mL of 100 mM ascorbic acid, 2.5 mL of ITS-A (insulintransferrin-selenium-sodium pyruvate) solution (Invitrogen Cat. Number 51300044), 2.9 mL of 35% bovine serum albumin (BSA) solution, and 5 mL of 10,000 U/μg/mL penicillin/streptomycin solution to 500 mL of DMEM/F-12, GlutaMAX™ Supplement Medium (see Note 4).

2.3 MACS Miltenyi Biotec Protocol

1. For this protocol, a gentleMACS™ Octo Dissociator (Miltenyi Biotec), the content of the “Neonatal Heart Dissociation Kit, mouse and rat” and gentleMACS C Tubes are required. 2. Prepare 5 mL of enzyme mix: Mix 125 μL of Enzyme P and 4.6 mL of Buffer X (Mix1). Then mix 25 μL of Enzyme A, 50 μL of Buffer Y, and 200 μL of Enzyme D (Mix2). Add Mix1 to Mix2. 3. PEB: Add 14.3 μL of 35% BSA solution to 985.7 μL of autoMACS Rinsing Solution. 4. RBC lysis solution: Add 1 mL of 10 red blood cell lysis solution to 9 mL of ddH2O. 5. RBC washing solution: Add 15 μL of Enzyme A to 10 mL of phosphate-buffered saline (PBS), without Ca2+ and Mg2+.

2.4 Adult Cardiomyocyte Isolation

1. Stock perfusion buffer (KHP): Add to 1 L of ddH2O, 7.42 g NaCl (127 mM), 0.34 g KCl (4.6 mM), 2.08 g NaHCO3 (24.8 mM), 0.13 g MgSO4  7 H2O (1.1 mM), 0.16 g KH2PO4 (1.2 mM), 1.5 g glucose (8.3 mM), 20 mL of a 100 mM Na-pyruvate solution (2.0 mM), 1.31 g creatine (10 mM), and 2.5 g taurine (20 mM). Filter sterilize and store at 4
C (up to 1 month). 2. Buffer A: Add 8.5 mL 100 mM sterile-filtered CaCl2  2 H2O to 500 mL of KHP. 3. Buffer B: Dissolve 0.5 g of fatty acid-free BSA in 500 mL of KHP. Sterilize the filter. 4. Digest buffer: Add 92 mg of fatty acid-free BSA and 16 mg of collagenase type II to 4 mL of KHP (see Note 5). 5. Washing buffer: Add 1 g of BSA to 100 mL of KHP and sterile filter. 6. Medium: Add 3.0 mL of 35% BSA solution (0.21%), 1.5 mL of insulin solution (4 mg/mL) (12 μg/mL), 0.327 g of creatine (5 mM), 0.313 g of taurine (5 mM), 0.2 g of L-carnitine (2 mM), and 5 mL of 10,000 U/μg/mL penicillin/streptomycin solution to 500 mL of DMEM-F12.

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7. Laminin solution: Dilute laminin solution (1 mg/2 mL Roche #1243217) 1:50 in PBS (10 μg/mL). 8. Perfusion system: The perfusion system consists of a water bath, a pump to circulate warm water, a pump to circulate perfusion solutions (e.g., Masterflex C/L Model 77120-62; Dual-Channel Variable-Speed Tubing Pump; 10–60 rpm), three glass containers to hold the three buffers (75–100 mL beakers), a heating coil with a de-gassing bubble trap (e.g., Radnoti #158830, 120 mm length, 35 mm outer diameter), valves (e.g., Cole-Parmer® Polycarbonate Luer Stopcocks, 4-way male lock), a gas filter, connectors (e.g., Cole-Parmer® Polypropylene fittings; male slip  1/1600 hose barb), and tubing for gas, as well as solutions (e.g., Ismatec SC0350, PharMed®BPT, inner diameter 2.06 mm, wall 0.85 mm). Assembly of our system is shown in Fig. 1b. Complete systems are also commercially available.

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Methods

3.1 SADO Mix Protocol 3.1.1 Neonatal Cardiomyocyte Isolation

This protocol is used to isolate cardiomyocytes from 20 to 30 E15 or 10 to 30 P0–P5 rat hearts. 1. Prior digestion: Fill a finger bowl (or any glass vessel) 2/3 with water (>5 cm water depth) and place it on a magnetic stirrer with heating. Submerge a contact thermometer probe and set to 37
C. Set the stirring at a speed just high enough that the tissue pieces slowly rotate in the digestion tube (see Notes 6–8) (Fig. 1a). 2. Prepare a total of 180 mL of neonatal digestion buffer. Place nine 50 mL tubes, each containing 20 mL of buffer, on ice along with four 50 mL tubes each containing 5 mL of horse serum (thawed in water bath). 3. Place two 10 cm Petri dishes each with 10 mL of PBS without Ca2+ and Mg2+ on ice and add 50 μL of 2 M glucose per dish. 4. Use the large curved scissors to decapitate neonates with one clean cut. To harvest a heart, use your thumb and middle finger to bend the forelimbs backwards and apply with the index finger pressure on the back. Use microdissection scissors to make a longitudinal incision from the neck along the sternum. Due to the pressure of the index finger and the contractions of the heart, the heart will emerge from the chest cavity and can be removed with a curved forceps by simple pulling. 5. Place harvested hearts in one of the Petri dishes prepared in step 3 (Fig. 1c).

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6. Once all of the hearts are harvested, use a scalpel blade to cut off the aorta and atria from the heart (upper third of the heart). Squeeze the remaining heart tissues (ventricles, Fig. 1d) gently with the forceps to remove blood from the heart ventricles and coronaries. Then transfer the tissue to the second Petri dish for a second wash. 7. Move the ventricles to a dry 10 cm Petri dish on ice and use the scalpel blade to mince them into pieces 1  1 mm (Fig. 1e) (see Note 9). 8. Transfer the minced neonatal heart tissue into roundbottomed glass tubes (Fig. 1a) with one micro stir bar (12.7  3 mm) for digestion. Add 20 mL of ice-cold neonatal digestion buffer. 9. Washing step 1: Place the glass tube in the finger bowl with water. Make sure that the water level is at or above the level of the digestion buffer in the tube. Stir the tissue for 3 min. Meanwhile, preheat another 20 mL of neonatal digestion buffer to 37
C (e.g., in the finger bowl). After 3 min, take the glass tube out of the finger bowl and let the tissue settle for 3 min under the laminar flow hood. Discard the supernatant (see Note 10). 10. Washing step 2: Add the pre-warmed neonatal digestion buffer to the tissue in the glass tube, place it in the finger bowl, and stir for 3 min. In the meantime, preheat another 20 mL of neonatal digestion buffer to 37
C. Remove the glass tube, let the tissue settle for 3 min, and discard the supernatant (see Note 11). 11. Digestion steps 1 and 2: Add the pre-warmed neonatal digestion buffer to the tissue in the glass tube, place it in the finger bowl, and stir it for 10 min. In the meantime, preheat another 20 mL of neonatal digestion buffer to 37
C. Remove the glass tube, let the tissue settle for 5 min, and remove the supernatant. Then, transfer the supernatant to one of the tubes on ice containing horse serum. Repeat this step. 12. Digestion steps 3–7: Repeat the digestion four times, but stir the tissue each time only for 8 min (see Note 12). 13. Pre-plating: Centrifuge the collected supernatants (330  g, 5 min, 4
C). Remove the supernatant and resuspend pellets in 20 mL of preheated (37
C) cardiomyocyte pre-plating medium per ten digested hearts. Seed the cells on cell culture plates (10 mL per round plate with 10 cm diameter). Incubate for 1.5 h at 37
C in a cell culture incubator (see Note 13). 14. Carefully rinse the plates with the supernatant medium. For this purpose, tilt the plate and let the medium run twice from top to bottom. This will markedly increase the yield of

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cardiomyocytes from the isolation. Subsequently, transfer the medium enriched in cardiomyocytes into 50 mL centrifuge tubes and centrifuge them (330  g, 5 min, 4
C). Discard the supernatant and resuspend the pellet in 1 mL of cardiomyocyte medium per each heart used in the isolation. 15. Mix an aliquot of cell suspension 1:1 with trypan blue and count the cells utilizing a hemocytometer (Fig. 1f) (see Notes 14 and 15). 16. Coating: Cardiomyocytes attach to plastic surface. However, in case of glass (e.g., glass coverslips), it is necessary to coat them. A cheap but suboptimal coating is 1% gelatin. Cover glass coverslips with 1% gelatin solution for 2 h at 37
C in a cell culture incubator. Replace the 1% gelatin solution with fresh 1% gelatin solution and incubate them for another 2 h. Allow the coverslips dry for 30 min under UV light in a laminar flow hood. Very fast adhering and efficient spreading are achieved by fibronectin coating (25 μg/mL). Add 100 μL of fibronectin solution on the middle of a round coverslip with a diameter of 12 mm. Glass coverslips will be incubated for 1 h at 37
C in a cell culture incubator. Then, remove the fibronectin and wash once in PBS (see Note 16). 17. Neonatal cardiomyocytes will attach and be ready for experimentation within 48 h of seeding. It is recommended to use 1% horse serum to ensure cardiomyocyte survival and the ability to maintain their differentiation status. The use of 10% FBS will greatly enhance cardiomyocyte attachment and beating behaviors. Yet, the cells will become hypertrophic and non-myocytes might overgrow the culture. 3.1.2 Embryonic Cardiomyocyte Isolation

1. Prior digestion: See Subheading 3.1.1, step 1. 2. Prepare a total of 90 mL of embryonic digestion buffer. Place six 50 mL tubes each containing 10 mL of buffer on ice along with two 50 mL tubes each containing 4 mL of horse serum (thawed in water bath). 3. Anesthetize a pregnant Wistar or Sprague-Dawley rat. First, subcutaneously inject 0.04 mg/kg of buprenorphine. After 30 min, anesthetize the rat with isoflurane (add 2 mL onto gaze in a 5 L beaker). Once the animals have reduced their breathing frequency and lost reflexes to stand, to keep their eyes open, and react to pain (squeezing paws), open the chest and excise the heart, which will result in death due to exsanguination (see also below Subheading 3.3, step 8). 4. Apply hysterectomy: Make a ventral midline incision into the abdomen, cut the fallopian tubes, and transfer the uterus to ice-cold, sterile PBS buffer. Free the individual embryos from their amniotic sac.

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5. Decapitate the embryos under a stereomicroscope/binocular with forceps and remove one arm to get access to the chest. Carefully open the chest with forceps/tweezers and collect the heart. 6. Follow the instructions under Subheading 3.1.1, steps 6 and 7. 7. Transfer the minced embryonic heart tissue into roundbottomed glass tubes (Fig. 1a) with one micro stir bar (12.7  3 mm) for digestion. Add 10 mL of ice-cold neonatal digestion buffer. 8. Washing step 1: Place the glass tube in the finger bowl with water, making sure that the water level is at or above the level of the digestion buffer in the tube. Stir the tissue for 3 min. In the meantime, preheat another 10 mL of embryonic digestion buffer to 37
C. After 3 min, take the glass tube out of the finger bowl and let the tissue settle for 3 min under the laminar flow hood. Discard the supernatant. 9. Digestion steps 1 and 2: Add the pre-warmed embryonic digestion buffer to the tissue in the glass tube, place it in the finger bowl, and stir it for 10 min. In the meantime, preheat another 10 mL of embryonic digestion buffer to 37
C. Remove the glass tube, let the tissue settle for 5 min, and remove the supernatant and transfer it to one of the tubes on ice containing horse serum. Repeat this step. 10. Digestion steps 3–5: Repeat the digestion three times but stir the tissue each time only for 8 min (see Note 12). 11. Centrifuge the collected supernatants (330  g, 5 min, 4
C). Remove the supernatant and resuspend pellets in 2 mL of preheated (37
C) cardiomyocyte medium (see Note 17). 12. Mix an aliquot of cell suspension 1:1 with trypan blue and count the cells utilizing a hemocytometer (see Notes 18 and 19). 13. Plate the cells as described under Subheading 3.1.1, steps 16 and 17. 3.2 MACS Miltenyi Biotec Protocol

An alternative to the traditional SADO Mix protocol is the MACS Miltenyi Biotec protocol. This procedure has several advantages, including standardization and saving time and workforce. However, it requires cost-intensive equipment and is, on average, more expensive per isolation. This protocol is used to isolate cardiomyocytes from 10 to 30 P0–P5 rat hearts and can also be applied to isolate P2–P7 mouse cardiomyocytes. 1. Harvest the hearts following the instructions under Subheading 3.1.1, steps 4–7.

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2. Transfer the minced hearts in a gentleMACS C Tube (maximum 20 hearts per tube) and add 5 mL of enzyme mix. 3. Place the gentleMACS C Tube into the gentleMACS™ Octo Dissociator and start the program “37C_mr_NHDK_1” (see Note 20). 4. Pipet 5 mL of cardiomyocyte pre-plating medium into the gentleMACS C Tube and transfer the content of the gentleMACS C Tube through a 100 μm cell strainer into a 50 mL tube. 5. Rinse the gentleMACS C Tube with 3 mL of cardiomyocyte pre-plating medium and repeat step 4. 6. Centrifuge the 50 mL tube at 300  g for 15 min at room temperature (RT) and discard the supernatant. 7. Resuspend the pellet in 1 mL of PEB. 8. Add 9 mL of RBC lysis solution to the resuspended pellet and wait for 2 min. 9. Repeat step 6. 10. Resuspend the pellet in 1 mL of RBC washing solution, add 9 mL of RBC washing solution, and repeat step 6. 11. Pre-plate, count, and plate the cells as described under Subheading 3.1.1, steps 13–17. Cardiomyocytes will be attached, spread, and ready for experimentation the morning after isolation (2 h (see Note 22). 3. Cleaning the perfusion system: Assemble (Fig. 1b, h) and clean the system before and after cardiomyocyte isolation by pumping ddH20 through the system for 10 min (see Note 23). 4. Set the water bath to 38–40
C and switch it on. At the same time, pump the water through the heat exchanger (here the temperature should be 37
C). 5. Set up for perfusion: Add 50 mL of buffer A, 50 mL of buffer B, and 36 mL of KHP into perfusion reservoirs in the water bath. 6. Start to introduce carbogen (5% CO2 and 95% oxygen) (>20 min before starting perfusion) in the solutions.

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7. Anesthesia: Use 12–14-week-old male Wistar or SpragueDawley rats (or 250–300 g). Subcutaneously inject 0.04 mg/ kg buprenorphine. After 30 min, anesthetize the rat with isoflurane (add 2 mL onto gaze in a 5 L beaker). Once animals have reduced their breathing frequency and have their lost reflexes to stand, to keep their eyes open, and react to pain (squeezing paws), open the chest and excise the heart. 8. Harvesting an adult heart: Use large scissors to snip off a longitudinal pinch of fur/skin extending from the belly up past the sternum. While holding a pinch of abdominal muscle up w/ forceps, cut transversely through the muscle just below the sternum. Then angle the scissors anterolaterally and cut through ribs on either side of the sternum. Pull up the sternum, and then cut through the diaphragm to view the heart. Grasp the top of the heart firmly with the thumb, index, and middle finger of your left hand, pull the heart up, and cut through the vessels with a single snip utilizing small scissors. 9. Transfer the heart rapidly to ice-cold PBS. 10. Turn on the perfusion pump with a flow rate of around 11 mL per min (see Note 24). 11. Take the aorta (big obvious whitish vessel atop the heart) and grasp each side with two sets of forceps. 12. Cannulate the heart while buffer A is pumped through the system by stretching the aorta around the cannula (e.g., plastic lock luer) with both sets of forceps (see Note 24). Pull the aorta up to the top of the cannula nozzle and hold it in this position with a set of forceps in one hand. Then, apply a small metal clamp (e.g., Roboz RS-5422: Micro Clip; Straight; 50–110 g pressure; 0.75 mm clip width; 6 mm jaw length) above the plastic ridge with the other hand and finally fix the aorta with a fine thread (e.g., Look® Braided Silk Nonabsorbable Uncoated, size 6/0, SP114) (see Note 25). 13. Once the heart is cannulated and beats regularly, and the coronaries have become clear (~1–3 min), switch to buffer B and perfuse until the heart stops beating because of lack of Ca2+ (~3–5 min). At the same time, add digest buffer to the reservoir with KHP (digest solution). 14. Switch perfusion to digest solution and perfuse for 20–30 min (see Note 26). 15. Once the perfusion is completed, use sharp scissors to cut the heart from the cannula. Place the heart in a 6 cm petri dish filled with 2 mL of the last perfusion buffer. Cut the atrium away and use two sets of forceps to carefully tear the heart into several pieces (see Note 27). 16. Collect the medium and add 10 mL of washing buffer.

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17. Place the tissue pieces in a beaker with 20 mL of the last perfusion buffer and bubble for 5 min with carbogen. If tissue is not completely disintegrated, take pieces out and tear again into pieces using forceps. 18. Add 10 mL of washing buffer, combine it with the cell suspension from step 14, divide it between two 50 mL tubes, and centrifuge it for 2 min (50  g) (see Note 28). 19. Resuspend the cells in 20 mL of washing buffer and centrifuge again. Repeat once. 20. Resuspend the cells in 20 mL of washing buffer and restore the calcium by adding 20, 30, 50, and 100 μL 100 mM CaCl2 and incubate the cells after each addition for 3 min. An alternative is to introduce calcium slowly during the end of the perfusion. 21. After centrifugation, resuspend the cells in 20 mL of adult cardiomyocyte medium. 22. Remove the laminin solution from the coverslips and add 500 μL of cell suspension per well (see Note 29). 23. After 2 h, cells should be attached (see Note 30). Wash two times carefully with medium to remove unattached adult cardiomyocytes. 24. Start experiments the next day (see Note 31). 3.4 Live-Cell Imaging of Cardiomyocytes to Detect and Analyze Mitosis

1. Coat one well of an 8-well gridded chamber slide (e.g., from Ibidi) with 200 μL of fibronectin solution (25 μg/mL) for 1 h at 37
C. Wash once with PBS before plating cells. 2. Seed between 20,000 and 30,000 cardiomyocytes per well to study proliferation of mononucleated cardiomyocytes. When studying binucleated cardiomyocytes, seed 40,000–50,000 cells per well. 3. Change the medium 24 h post-isolation (MACS: neonatal; SADO Mix: embryonic) and stimulate the cells with pro-proliferative (candidate) factors. 4. According to the expected timing of cell division, start video recording (see Note 32). 5. According to the need for temporal resolution (division yes/no vs. defects in chromosome segregation or cleavage furrow ingression), images should be taken every 10 min, and for high temporal resolution, around every 6 min. 6. In order to capture a significant number of mitotic cardiomyocytes, 10 or 15 positions should be recorded using a 20 objective for mononucleated and binucleated cardiomyocytes, respectively (see Note 33). 7. A major issue in live-cell imaging is stage drift, cell, and cell organelle movement. In order to compensate for such issues, a

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total of 5–7 Z-stacks can be obtained per position (see Note 34). 8. After the end of the recording (see Note 32), assign each movie to the position on the gridded chamber slides based on the provided coordinates. 9. Perform immunofluorescence analysis by using a cardiac marker, such as actinin or troponin I, a membrane marker, such as N-cadherin, and a nuclear cardiac-specific marker, such as Nkx2.5 (see Note 35). 10. Correlate the videos and immunofluorescence stainings by using the coordinates provided on the gridded chamber slide and analyze the outcome of cardiomyocyte mitosis.

4

Notes 1. 200 mM HEPES-NaOH (pH 7.6), 1.3 M NaCl, 300 mM KCl, and 100 mM NaH2PO4 can be stored after being autoclaved at room temperature. Sterile-filtered 2 M of glucose solution can be stored at 20
C in 2 mL aliquots. 2. Pancreatin is difficult to dissolve. You will have to stir for 5–10 min. 3. The concentration and/or activity of collagenase type II and pancreatin can vary significantly. Thus, it is important to test different samples to identify the best supplier. It might be necessary to optimize the concentrations. We use collagenase type II from Invitrogen (#17101-015) and pancreatin from Sigma (P3292). 4. Do not use BSA in powder form. 5. The digest buffer should always be prepared fresh. 6. The contact thermometer probe must not touch the bottom of finger bowl and 2.5 cm of the probe needs to be covered by water. 7. The water depth in the finger bowl must be sufficient to submerge the round-bottomed glass tubes in a way that the digestion solution in the tubes is surrounded by water. 8. In our system (see Subheading 2.1, step 4), we set the temperature of the magnetic stirrer with the heating at 65
C and the stirring at 2.5–3 units. 9. Mincing can harm cardiomyocytes. Thus, it is important to empirically test which grade of mincing gives the best results. 10. Remove as much of the buffer as possible without taking any tissue.

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11. If the total number of neonatal rat hearts is 70%). If there is something wrong (e.g., concentration of enzymes, too low temperature in the heat exchanger, too short digestion time) the heart will be tough and/or most of the released cells will be rounded up. Adding 2,3-butanedione 2-monoxime (final concentration: 20 mM), a well-known inhibitor of cardiac muscle contraction, to all buffers and medium will increase the survival but also affects cell behavior. 28. Adult cardiomyocytes are very sensitive to low temperatures; always handle cardiomyocytes at room temperature or warmer. 29. This procedure should generate enough adult cardiomyocytes from one heart to densely seed two 24-well plates. 30. Adult cardiomyocytes tend to cluster in the middle of the cell culture dish. Thus, the plate should be shaken every 30 min (side to side, no rotating movements). 31. The enzymatic digestion can cause damage to the isolated cardiomyocytes (e.g., loss of surface receptors, membrane damage). Experience demonstrates that experiments with cardiomyocytes 1 day after isolation result in significantly higher reproducibility. 32. In our previous studies, we have started video recording ~12 h, 36 h, and 48 h post-stimulation for embryonic, binucleated neonatal, and mononucleated cardiomyocytes, respectively. Recordings were stopped after ~24 h in case of embryonic and mononucleated neonatal cardiomyocytes or ~48 h in case of binucleated cardiomyocytes. 33. In case a higher magnification and the use of oil are required, apply the oil to the lens at least 30 min before starting the video to allow temperature equilibration. This will prevent problems with focus due to changes in the temperature-dependent viscosity of the oil. 34. In order to determine the optimum Z-stack number, the lowest and highest focus planes need to be determined among all positions to be recorded. While we have previously used 5–7 stacks, it should be noted that the number of stacks is limited

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by the number of positions and the recording frequency (1 image per 6–10 min). 35. It is fundamental to use a cardiac-specific nuclear marker because non-myocytes are more motile and grow on top of cardiomyocytes, which complicates the identification of cardiomyocyte nuclei.

Acknowledgments We thank all the past members of the Engel laboratory, Machteld van Amerongen, Tatyana Novoyatleva, Florian Diehl, Ingrid Schmalenberger, Ajit Magadum, and David C. Zebrowski, and the present members, Robert Becker, Jennifer Schmidt, and Jana Petzold, for establishing and optimizing the protocols published in this book chapter. We thank also Tilman Esser and Isabel Schoenauer for experimental support. This work was supported by the German Research Foundation (DFG, EN 453/12-1 to F.B.E.) and an ELAN Program Grant (ELAN-18-12-17-1-Leone to M.L.). References 1. Leone M, Engel FB (2019) Advances in heart regeneration based on cardiomyocyte proliferation and regenerative potential of binucleated cardiomyocytes and polyploidization. Clin Sci (Lond) 133(11):1229–1253 2. Leone M, Magadum A, Engel FB (2015) Cardiomyocyte proliferation in cardiac development and regeneration: a guide to methodologies and interpretations. Am J Physiol Heart Circ Physiol 309(8): H1237–H1250 3. Zebrowski DC, Becker R, Engel FB (2016) Towards regenerating the mammalian heart: challenges in evaluating experimentally induced adult mammalian cardiomyocyte proliferation. Am J Physiol Heart Circ Physiol 310 (9):H1045–H1054 4. Murry CE, Reinecke H, Pabon LM (2006) Regeneration gaps: observations on stem cells and cardiac repair. J Am Coll Cardiol 47 (9):1777–1785 5. van Amerongen MJ, Engel FB (2008) Features of cardiomyocyte proliferation and its potential for cardiac regeneration. J Cell Mol Med 12 (6A):2233–2244 6. Cluzeaut F, Maurer-Schultze B (1986) Proliferation of cardiomyocytes and interstitial cells in the cardiac muscle of the mouse during preand postnatal development. Cell Tissue Kinet 19(3):267–274

7. Manasek FJ (1968) Mitosis in developing cardiac muscle. J Cell Biol 37(1):191–196 8. Oberpriller JO, Oberpriller JC (1974) Response of the adult newt ventricle to injury. J Exp Zool 187(2):249–253 9. Poss KD, Wilson LG, Keating MT (2002) Heart regeneration in zebrafish. Science 298 (5601):2188–2190 10. Engel FB, Schebesta M, Duong MT, Lu G, Ren S, Madwed JB, Jiang H, Wang Y, Keating MT (2005) p38 MAP kinase inhibition enables proliferation of adult mammalian cardiomyocytes. Genes Dev 19(10):1175–1187 11. Leone M, Musa G, Engel FB (2018) Cardiomyocyte binucleation is associated with aberrant mitotic microtubule distribution, mislocalization of RhoA and IQGAP3, as well as defective actomyosin ring anchorage and cleavage furrow ingression. Cardiovasc Res 114(8):1115–1131 12. Magadum A, Ding Y, He L, Kim T, Vasudevarao MD, Long Q, Yang K, Wickramasinghe N, Renikunta HV, Dubois N, Weidinger G, Yang Q, Engel FB (2017) Live cell screening platform identifies PPARdelta as a regulator of cardiomyocyte proliferation and cardiac repair. Cell Res 27 (8):1002–1019 13. Eulalio A, Mano M, Dal Ferro M, Zentilin L, Sinagra G, Zacchigna S, Giacca M (2012)

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Functional screening identifies miRNAs inducing cardiac regeneration. Nature 492 (7429):376–381 14. Uosaki H, Magadum A, Seo K, Fukushima H, Takeuchi A, Nakagawa Y, Moyes KW, Narazaki G, Kuwahara K, Laflamme M, Matsuoka S, Nakatsuji N, Nakao K, Kwon C, Kass DA, Engel FB, Yamashita JK (2013) Identification of chemicals inducing cardiomyocyte proliferation in developmental stage-specific manner with pluripotent stem cells. Circ Cardiovasc Genet 6(6):624–633 15. Mills RJ, Titmarsh DM, Koenig X, Parker BL, Ryall JG, Quaife-Ryan GA, Voges HK, Hodson MP, Ferguson C, Drowley L, Plowright AT, Needham EJ, Wang QD, Gregorevic P, Xin M, Thomas WG, Parton RG, Nielsen LK, Launikonis BS, James DE, Elliott DA, Porrello ER, Hudson JE (2017) Functional screening in human cardiac organoids reveals a metabolic mechanism for cardiomyocyte cell cycle arrest. Proc Natl Acad Sci U S A 114(40): E8372–E8381 16. Titmarsh DM, Glass NR, Mills RJ, Hidalgo A, Wolvetang EJ, Porrello ER, Hudson JE,

Cooper-White JJ (2016) Induction of human iPSC-derived cardiomyocyte proliferation revealed by combinatorial screening in high density microbioreactor arrays. Sci Rep 6:24637 17. Chow M, Boheler KR, Li RA (2013) Human pluripotent stem cell-derived cardiomyocytes for heart regeneration, drug discovery and disease modeling: from the genetic, epigenetic, and tissue modeling perspectives. Stem Cell Res Ther 4(4):97 18. Lundy SD, Zhu WZ, Regnier M, Laflamme MA (2013) Structural and functional maturation of cardiomyocytes derived from human pluripotent stem cells. Stem Cells Dev 22 (14):1991–2002 19. Hirt MN, Boeddinghaus J, Mitchell A, Schaaf S, Bornchen C, Muller C, Schulz H, Hubner N, Stenzig J, Stoehr A, Neuber C, Eder A, Luther PK, Hansen A, Eschenhagen T (2014) Functional improvement and maturation of rat and human engineered heart tissue by chronic electrical stimulation. J Mol Cell Cardiol 74:151–161

Chapter 10 Generation of Human Induced Pluripotent Stem Cells and Differentiation into Cardiomyocytes Lu Han, Jocelyn Mich-Basso, and Bernhard Ku¨hn Abstract Failure to regenerate myocardium after injury is a major cause of mortality and morbidity in humans. Direct differentiation of human induced pluripotent stem cells (iPSCs) into cardiomyocytes provides an invaluable resource to pursue cardiac regeneration based on cellular transplantation. Beyond the potential for clinical therapies, iPSC technology also enables the generation of cardiomyocytes to recapitulate patient-specific phenotypes, thus presenting a powerful in vitro cell-based model to understand disease pathology and guide precision medicine. Here, we describe protocols for reprogramming of human dermal fibroblasts and blood cells into iPSCs using the non-integrative Sendai virus system and for the monolayer differentiation of iPSCs to cardiomyocytes using chemically defined media. Key words Heart regeneration, Induced pluripotent stem cells (iPSCs), Cardiomyocytes, Patientspecific, Reprogramming, Differentiation, Sendai virus, Chemically defined, Monolayer differentiation

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Introduction Induced pluripotent stem cells (iPSCs) provide an unprecedented opportunity for understanding cardiovascular development and regeneration, heart physiology and diseases, and drug discovery [1–6]. In 2006, Takahashi and Yamanaka et al. revolutionarily transformed mouse fibroblasts to embryonic stem cell-like pluripotent cells using four transcription factors: OCT4 (POU5F1), SOX2, KLF4, and MYC [7]. In 2007, two independent groups, Takahashi et al. and Yu et al., successfully generated human induced pluripotent cells from fibroblasts [8, 9]. These iPSCs resemble embryonic stem cells (ESCs) in that they possess the potential to undergo infinite self-renewal and have the ability to differentiate into any specialized cell lineage [10]. Most importantly, autologous iPSCs circumvent the ethical concerns and immunogenicity of human ESCs, thus becoming an attractive alternative for tissue repair and regeneration [1, 11]. Advances in Yamanaka’s original method have reduced the risk of inducing unwanted genome

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_10, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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instability and DNA aberration [12–17]. Delivery methods have evolved from integrative viral vectors (retrovirus and lentivirus) to non-integrative systems, including episomal DNA vectors, synthetic mRNA, protein, and Sendai virus (SeV) [15–19]. With progress in the iPSC field, generation of iPSCs could become safe and efficient for clinical applications. The principle for differentiating iPSCs into cardiomyocytes is to recapitulate key steps in embryonic cardiac development, leading from the formation of mesodermal cells to functional cardiomyocytes [20, 21]. These steps involve stage-specific activation and inhibition of specific signaling pathways with defined growth factors. Stimulation of activin/Nodal, bone morphogenetic protein (BMP), Wnt (wingless/INT protein), and fibroblast growth factor (FGF) signaling promotes cardiac mesoderm formation [22, 23]. Subsequent inhibition of Wnt, BMP, and TGF3β induces cardiac specification [23, 24]. In the most recent development of monolayer differentiation techniques, the combination of growth factors was replaced by small molecules that modulate only WNT signaling [25, 26]. The purity of cardiomyocytes from these protocols was enhanced by using glucose-deprived culture media containing abundant lactate, which favors the survival of cardiomyocytes over other cardiac cell lineages [27]. These chemically defined protocols have largely reduced cost and labor while increasing consistency and yield for large-scale de novo production of cardiomyocytes. In this chapter, we describe a detailed protocol to reprogram human iPS cells from dermal fibroblasts and blood cells by using a non-integrative Sendai (SeV) virus encoding the four Yamanaka factors (OCT4, SOX2, KLF4, and c-MYC). We also lay out a thorough procedure for differentiation of iPSCs into high-purity functional cardiomyocytes using a chemically defined monolayer method.

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Materials

2.1 Fibroblast Transduction

1. Sendai virus (encoding OSKM factors). 2. Inactivated MEF cells (feeder cells). 3. MEF media: 90% DMEM, 10% heat-inactivated Fetal Bovine Serum, 1% L-Glutamine, 1% Antibiotic-Antimycotic solution. 4. ES media: 80% Knockout DMEM (or Knockout DMEM/ F12), 20% KnockOut Serum Replacer, 1% L-Glutamine, 1% MEM Non-Essential Amino Acids, 1% AntibioticAntimycotic solution, 10 ng/mL bFGF.

2.2 Blood Cell Transduction

1. Sendai virus (encoding OSKM factors). 2. Inactivated MEF cells (feeder cells).

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3. Polybrene. 4. Expansion media: QBSF base medium, 1% AntibioticAntimycotic solution, 10 μg/mL Ascorbic Acid, 10 ng/mL IL3; 2 U/mL EPO, 40 ng/mL IGF1, 1 μM Dexamethasone, 50 ng/mL SCF. 5. MEF media: 8.7% IMDM base medium, 10% heat-inactivated Fetal Bovine Serum, 1% L-Glutamine, 1% AntibioticAntimycotic solution, 1% MEM Non-Essential Amino Acids, 1.8 μL/mL 55 mM 2-mercaptoethanol (2-ME). 6. MEF media with cytokines: 8.7% IMDM base medium, 10% heat-inactivated Fetal Bovine Serum, 1% L-Glutamine (Glutamax), 1% Antibiotic-Antimycotic Solution, 1% MEM Non-Essential Amino Acids, 1.8 μL/mL 55 mM 2-ME, 10 μg/mL Ascorbic Acid, 10 ng/mL IL3; 2 U/mL EPO, 40 ng/mL IGF1, 50 ng/mL SCF, 1 μM Dexamethasone, 10 ng/mL bFGF. 7. ES media: See Subheading 2.1. 2.3 Checking Transgene by RT-PCR

1. Primers (human):

SeV

Forward GGA TCA CTA GGT GAT ATC GAG C Reverse

SOX2

Forward ATG CAC CGC TAC GAC GTG AGC GC Reverse

KLF4

TCC ACA TAC AGT CCT GGA TGA TGA TG

Forward CCC GAA AGA GAA AGC GAA CCA G Reverse

2.3.1 Maintenance of Human iPSCs in a Feeder-Free System

AAT GTA TCG AAG GTG CTC AA

Forward TAA CTG ACT AGC AGG CTT GTC G Reverse

OCT3/ 4

AAT GTA TCG AAG GTG CTC AA

Forward TTC CTG CAT GCC AGA GGA GCC C Reverse

c-MYC

ACC AGA CAA GAG TTT AAG AGA TAT GTA TC

AAT GTA TCG AAG GTG CTC AA

1. mTESR 1 Kit (or other feeder-free hESC culture media). 2. Matrigel hESC-qualified Matrix. 3. ReLeSR. 4. DMEM/F-12. 5. PBS.

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2.3.2 Cardiomyocyte Differentiation Media and Reagents

1. Matrigel-growth factor reduced (GFR). 2. ROCK inhibitor (Y-27632). 3. PBS. 4. ReLeSR dissociation reagent. 5. STOP media (DMEM/FBS 1:1 mix). 6. TrypLE Express Enzyme (1
). 7. DNase. 8. RPMI1640 B-27 minus insulin: RPMI1640, 2% B-27 Supplement minus insulin, 1.5% Antibiotic-Antimycotic. 9. RPMI1640 B-27 with insulin: RPMI1640; 2% B-27 Supplement, 1.5% Antibiotic-Antimycotic. 10. CM enrichment media: RPMI 1640 without glucose; 2% B-27 Supplement, Sodium-D-lactate (final concentration 5 mM); 1.5% Antibiotic-Antimycotic.

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Methods

3.1 Fibroblast Transduction 3.1.1 Fibroblast Preparation

Workflow, see Fig. 1.

1. On day -5, coat a 10 cm culture dish with 1% gelatin and allow it to sit for 1 h at room temperature. 2. Thaw fibroblast cells: Remove 1 vial of MEF from liquid nitrogen and quick thaw in 37  C water bath. Transfer it to a 15 mL conical tube and add 2 mL of MEF media, dropwise. Pipet up and down to mix. Spin at 200
g for 5 min at room temperature. 3. Remove the supernatant and resuspend the pellet in 10 mL of fresh MEF media. 4. Plate the cells onto a gelatin-coated 10 cm dish and incubate at 37  C, 5% CO2, until confluent.

Fig. 1 Workflow of reprogramming human dermal fibroblast cells into iPSCs. D day

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Fig. 2 Reprogramming of human dermal fibroblast cells into iPSCs. (a) Freshly isolated human dermal fibroblast cells in vitro. (b) Representative image of human iPS clone derived from fibroblast cells by using Sendai virus encoding the four Yamanaka factors. (c) Characterization of iPS clone shows positive alkaline phosphatase (AP) staining. Scale bars, 100 μm

5. On day -2, remove the MEF media and rinse with 3 mL of PBS. Add 2 mL of TrypLE and incubate at 37  C, 5% CO2, for approximately 2–3 min until cells have lifted from the bottom of the dish. 6. Add 2 mL of MEF media to stop the reaction and transfer to a 15 cc conical tube. 7. Add 1 mL of MEF media to the plate to rinse the remaining cells from the dish and add to the conical tube. 8. Spin cells at 200
g for 5 min at room temperature. Decant the supernatant and add fresh MEF media. 9. Plate the cells onto 2 wells of a gelatin-coated 6-well plate at a density to achieve approximately 5
105 (80–90% confluency) per well after 48 h (Fig. 2a). 3.1.2 Transduction

1. On day 0, perform transduction by warming 2 mL of MEF media. 2. Add the Sendai virus at a MOI of 3 to 2 mL of prewarmed MEF media (see Note 1). 3. Thoroughly mix by gently pipetting up and down. 4. Within 5 min of mixing, aspirate the MEF media from the fibroblast cells. Add 1 mL of the virus mixture to each well. Incubate overnight at 37  C, 5% CO2. 5. On day 1, replace the medium with 2 mL of fresh MEF media 24 h after transduction (see Notes 2 and 3). 6. Continue to replace the media with fresh MEF media every 2 days until day 7.

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7. On day 6, prepare 3
10 cm of gelatin-coated MEF dishes for each cell line that has been transduced. Coat the dishes with 1% gelatin for 1 h. Plate 0.9-1
106 of inactivated MEFs per dish. Incubate until ready for use the next day. 8. On day 7, plate the transduced cells onto 10 cm feeder plates. 9. Remove the cells by adding 500 μL of TrypLE. 10. Incubate at room temperature for 1–3 min until the cells have rounded. 11. Add 1 mL of MEF media. Collect the cells and transfer to a 15 mL conical tube. 12. Rinse the wells with an additional 1 mL of MEF media. Spin the cells at 200
g for 5 min. 13. Count the cells and plate at a density of 5
104, 1
105, and 2
105 on the three feeder dishes containing inactivated MEFs. Add 8–10 mL of MEF media. Incubate the plates at 37  C, 5% CO2, overnight (see Note 4). 14. On days 8–28, change to ES media 24 h after plating onto the feeders. 15. Replace the media every other day. Start observing cells for colony formation on day 8. It may take up to 4 weeks before the clones will begin to appear. 16. Approximately 3–4 weeks post-transduction, colonies should be large enough to transfer (see Notes 5 and 6). 3.1.3 Clone Expansion

1. Once the clones have reached the appropriate size, seed a 12-well plate with inactivated MEF cells 1–2 days prior to picking the clones. 2. Identify several clones that are large enough to passage and mark them using either a cell marker or a felt-tip pen. 3. Cut 1 clone into several smaller pieces using a finely pulled glass pipet with a sealed tip. Transfer all pieces into 1 well of the 12-well feeder plate containing 1 mL of ES media. 4. Proceed to transfer the remaining marked colonies in the same manner, allowing for 1 clone per well. 5. Number each well of the 12-well plate to identify individual clones. 6. Incubate the plate for 48 h before changing media to allow the pieces to attach. Continue with media changes every day. 7. Continue to passage, expand, and maintain the newly named clones using standard culture procedures for ES cells (Fig. 2b).

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Fig. 3 Human iPSCs express pluripotency markers. Human iPSCs show positive staining for the nuclear marker Oct-4 (green, top) and the cytoplasmic marker TRA-1-60 (red, bottom) determined by immunofluorescent microscopy. Scale bars, 100 μm 3.1.4 Confirmation of Vector-Free iPS Clones

1. After ten passages, check iPS clones for removal of the Sendai reprogramming vectors by performing RT-PCR. Program: 94 for 5 min; 30–35 cycles of 95 for 30 s, 55 for 30 s, 72 for 30 s, followed by 72 for 10 min, 4  C forever. 2. If c-MYC is still present after ten passages, you can enhance removal by incubating the cultures at 38–39  C for 5 days. (c-MYC contains a temperature-sensitivity mutation, whereas the other genes do not.) Repeat RT-PCR. 3. Continue confirmation assays by performing AP staining (see Fig. 2c), qRT-PCR, and immunocytochemistry (see Fig. 3) with pluripotency markers.

3.2 Blood Cell Transduction

1. On day 14, mix blood 1:1 with warm PBS.

3.2.1 PBMC Isolation and Expansion

3. Spin at 300–400
g for 30 consecutive min.

2. Gently layer over 10 mL of warm Histopaque. 4. Use a sterile pipet to collect the buffy coat at the center of the interfaces. 5. Increase the volume by adding 5 mL of sterile PBS.

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6. Spin at 300
g for 10 min. 7. If necessary, perform RBC lysis by adding 5 mL of RBC lysis buffer and incubating at room temperature for 10 min. Fill with PBS and spin at 300
g for 10 min. 8. Remove the supernatant and resuspend it in 10 mL of sterile PBS. Spin at 300
g for 10 min. 9. Repeat the above wash step. Discard the supernatant, resuspend in 5 mL of PBS, and count the cells. 10. Transfer 1–2
106 cells to a conical tube and spin at 300
g for 10 min. 11. Resuspend in 2 mL of EM media and transfer to 1 well of a 12-well plate. Incubate for 3 days. 12. Freeze the remaining cells at ~2
106 using 90% FBS and 10% DMSO. 3.2.2 PBMC Expansion

1. On day -11, transfer the cells into a sterile 15 mL conical tube. 2. Wash the well once with 1 mL of QBSF base media to collect any adherent cells and add to the conical tube. 3. Spin the cells at 300
g for 10 min. 4. Resuspend in 2 mL of EM and plate the cell suspension in 1 well of a fresh 12-well plate. 5. On day -8 and day -4, repeat the procedure from day -11.

3.2.3 Viral Transduction, Clone Expansion, and Confirmation of Vector-Free iPS Clones

1. On day 0, count the cells, transfer 2.5–5.0
105 to a conical tube and spin at 300
g for 10 min. 2. Warm 1 mL of EM. 3. Add the Sendai virus at a MOI of 3 to 2 mL of prewarmed MEF media (see Note 1). 4. Add 4 μg/mL of polybrene. 5. Within 5 min, decant the supernatant from the spun cells, resuspend it in approximately 300 μL of virus mixture, and mix gently. 6. Plate each line onto 1 well of 24-well plate. Incubate overnight at 37  C, 5% CO2. 7. On day 1, 24 h after transduction, collect the cells in a 15 mL conical tube and spin at 300
g for 10 min. 8. Resuspend the cells in 500 μL of EM media and replate in a 24-well plate. 9. Incubate until day 3 (see Note 7). 10. On day 3, collect the cells and spin at 300
g for 10 min.

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11. Count the cells and plate 2
105 onto a 10 cm feeder plate using 10 mL of MEF media containing cytokines and 10 ng/ mL of bFGF. Incubate for 48 h. 12. On day 5, aspirate the media and replace it with 10 mL of MEF media containing 10 ng/mL bFGF (NO CYTOKINES). 13. Incubate for 48 h. 14. On day 7, aspirate the media and replace it with 10 mL of media made up of 50% MEF media containing bFGF and 50% ESC media. 15. Incubate for 48 h. 16. On day 9, replace the media with 10 mL of ESC media. 17. Continue to replace the media daily, watching for the development of clones. 18. On day 15+, clones should be reaching the appropriate sizes for transfer. To help avoid differentiation, try to transfer clones close to 3 weeks or later post-transduction. 19. Picking clones: See Subheading 3.1.3. 20. Confirmation of vector-free iPS clones: See Subheading 3.1.4. 3.3 Maintenance of Human iPSCs

1. Prepare the Matrigel (0.5 mg for 6 mL of DMEM/F-12) by thawing the Matrigel aliquot(s) on ice.

3.3.1 Preparing Matrigel Plates

2. Dilute the Matrigel into the appropriate amount of ice-cold DMEM/F-12, according to the protein concentration on the manufacturer’s protocol. 3. Distribute the diluted Matrigel into the appropriate number of wells for passaging iPS cells. 2 mL for each well of a 6-well plate. 4. Let the plates sit for a minimum of 1 h under the hood prior to use (see Note 8).

3.3.2 Passaging of iPSCs (See Note 9)

1. Aspirate the growth media from the currently growing undifferentiated iPSCs and rinse with PBS. 2. Add 1 mL of room-temperature ReLeSR to each well of a 6-well plate. 3. Aspirate off the ReLeSR and incubate at 37  C for 5 min. 4. After incubation, gently tap the plate and add 1 mL of the culture media. 5. Gently pipet up and down 5
with a P1000 pipet to break colonies into small clumps (see Note 10). 6. Transfer the cell suspension to a 1.5 mL microcentrifuge tube. 7. Pipet up and down gently 3–5
to ensure that the cells are in small clusters (5–10 cells/cluster).

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Fig. 4 Workflow of iPSC differentiation to cardiomyocytes using a chemically defined method

8. Obtain new Matrigel-coated plate(s). Aspirate off the Matrigel. Pipet the cell suspension up and down 1–2
gently, taking care not to break up any cell aggregates. Evenly divide the cell suspension into the new well(s) (see Note 11). 9. Add the culture media so that each well has a total volume of 2–3 mL (see Note 12). 10. Move the plate quickly back and forth and side to side to disperse the cells evenly across the well surface. Place in a 37  C, 5% CO2, incubator. 11. Change the media daily and check for contamination under an inverted phase-contrast microscope. 3.4 Human Cardiomyocyte Differentiation 3.4.1 iPS Passaging

Workflow, see Fig. 4.

1. Prepare the Matrigel as done in Subheading 3.3.1. 2. Refer to Subheading 3.3.2. for passaging of iPSCs. 3. Evenly distribute the cell suspension into Matrigel-coated wells/plates to achieve 80–90% confluency within 4 days (see Note 13). 4. Add the culture media containing 10 μM of the ROCK inhibitor to bring the final volume for each well of a 6-well plate to 2 mL (1 mL for one 12-well plate). 5. Move the plate quickly back and forth and front to back 5
to ensure even distribution of cells around the wells. 6. Incubate at 37  C, 5% CO2. 7. Replace the media with fresh culture media on the following day. 8. Continue with daily media changes until day 0, when cells should be ~70–80% confluent, approximately 4–5 days post passaging.

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1. On day 0 (CHIR99021), cells should be at least 70% confluent. Aspirate the culture media and replace with RPMI 1640 w/ B-27 () insulin media: 2 mL for each well of 12-well plate 4 mL for each well of 6-well plate 2. Add 12 μM of CHIR99021 (10 mM CHIR99021 stock). 3. Incubate for 24 h. 4. On day 1, aspirate off the media containing the CHIR99021 and replace it with fresh RPMI 1640 w/ B-27 (). 5. Incubate for 48 h. 6. On day 3, change the media on all wells to fresh RPMI1640 w/ B-27 (). 7. Add 5 μM of IWR-1 (10 mM IWR-1 stock). 8. Incubate for 48 h. 9. On day 5, remove the IWR-1 media from all wells and add fresh RPMI 1640 w/ B-27 (). 10. On days 7 and 10, remove the RPMI 1640 w/ B-27 () and add fresh RPMI 1640 w/ B-27 supplement media. You should be able to see cells starting to contract between days 8 and 9 (see Note 14). 11. On day 12, remove the RPMI 1640 w/ B-27 supplement and replace it with CM enrichment media. 12. Continue to culture with the CM enrichment media until the day of dissociation (see Note 14). 13. On day 15 perform iPSC-CM dissociation.

3.4.3 CM Dissociation

1. Prepare the Matrigel as done in Subheading 3.3.1 2. Distribute the diluted Matrigel into the number of wells needed for passaging cells. 3. Let the plate sit for a minimum of 1 h under the hood prior to use. 4. Wash the cells with PBS. 5. Add 1 mL of TrypLE to each well and incubate for 5–7 min at 37  C. 6. After 5–7 min, gently pipette up and down ~10
to dislodge and break up cell clumps. 7. Incubate the plate for an additional 5–7 min. 8. Add 1 mL of stop media containing 10 μg/mL of DNase and gently pipette up and down until all of the cells are dislodged from the bottom of the well and have dissolved into the cell suspension without visible clumps.

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Fig. 5 iPSC-derived cardiomyocytes at day 16 of differentiation. More than 90% of the cells show positive staining for the cardiomyocyte marker α-actinin. iPSC-derived cardiomyocytes exhibit sarcomere structure by immunofluorescent microscopy at high magnification (60
). Scale bars, 100 μm (brightfield), 20 μm

9. Pass the cells through a 70 μm cell strainer inserted into a sterile 15 or 50 mL conical tube. 10. Add an additional 1 mL of stop media to the well to rinse and filter. 11. Spin at 600
g for 8 min. 12. Aspirate off the supernatant and resuspend the pellet by gently tapping the bottom of the tube. 13. Add 1 mL of CM enrichment media. 14. Count the cells. 15. Plate the cells on the desired cell culture plates needed for experiments. Recommended densities: 24-well plate with 13 mm coverslips: 1–2
105/well 6-well plate: 0.8–1
106/well 16. Confirm the differentiation efficiency by flow cytometry or immunocytochemistry (see Fig. 5).

4

Notes 1. If using individual encoding Sendai viruses for OSKM, each one should have a MOI of 3. The volume of virus (μL) is calculated using the following formula: MOI
number of cells Titer of virus ðCIU=mLÞ
103 ðmL=μLÞ

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2. Cytotoxicity may be seen 24–48 h post-transduction, which can affect up to 50% of the cells. This indicates a high virus uptake. 3. High cell density before day 5 may be seen. Do not plate onto feeders prior to day 7. 4. This is to determine the correct plating density. Once it has been determined, three plates may not be necessary for future transductions. 5. During clone formation, if the feeder layer becomes overly spent and is thinning, add fresh feeders to the 10 cm plate. 6. After 3+ weeks of transduction, if the feeder layer appears thick, gently peel away the areas where possible and replace with fresh feeders. 7. You may see drastic cell death (>60%) 24–48 h posttransduction. Large aggregated cells may be seen. 8. Wrap plates in Parafilm and store at 4  C for up to 2 weeks. Bring to room temperature prior to use. 9. 1 well of a 6-well plate at an optimal density of 70–80%; cells should be split every 5–7 days. 10. Differentiated colonies will remain attached. 11. The typical split ratio is ~1:40 if passaging from a well that is approximately 70–80% confluent. 12. 10 μM of the ROCK inhibitor may be added to each well to ensure survival and reduce spontaneous differentiation. 13. The typical split ratio is ~1:30 if passaging from a well that is approximately 70–80% confluent. 14. Differentiation between days 7–10 and 12–15: Additional media changes with noted media may be required if the media turns yellow prior to the indicated media change days.

Acknowledgments This work was supported by the Richard King Mellon Foundation Institute for Pediatric Research (UPMC Children’s Hospital of Pittsburgh), by a Transatlantic Network of Excellence grant by Foundation Leducq (15CVD03), Children’s Cardiomyopathy Foundation, NIH grant R01HL106302 (to B.K.), and a Career Development Award from the AHA (18CDA34110053 to L.H.). We thank Dan Roden and Kevin Bersell (Vanderbilt) for generously sharing a human iPS cell line CiPS001-13 (Vanderbilt University).

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Chapter 11 Differentiation of Human Induced Pluripotent Stem Cells into Epicardial-Like Cells Navid A. Nafissi, Paige DeBenedittis, Michael C. Thomas, and Ravi Karra Abstract The epicardium is a multipotent cell layer that is vital to myocardial development and regeneration. Epicardial cells contribute to cardiac fibroblast and smooth muscle populations of the heart and secrete paracrine factors that promote cardiomyocyte proliferation and angiogenesis. Despite a central role in cardiac biology, the mechanisms by which epicardial cells influence cardiac growth are largely unknown, and robust models of the epicardium are needed. Here, we review our protocol for differentiating induced pluripotent stem cells (iPSCs) into epicardial-like cells through temporal modulation of canonical Wnt signaling. iPSC-derived epicardial cells (iECs) resemble in vivo epicardial cells morphologically and display markers characteristic of the developing epicardium. We also review our protocol for differentiating iECs into fibroblasts and smooth muscle cells through treatment with bFGF and TGF-β1, respectively. iECs provide a platform for studying fundamental epicardial biology and can inform strategies for therapeutic heart regeneration. Key words iPS, Epicardium, Smooth muscle cell, Fibroblast, Cell culture, Differentiation

1

Introduction The epicardium is critical to cardiac growth, serving as a reservoir of multipotent progenitor cells and as a source for paracrine growth factors. Epicardial cells comprise the outer mesothelial covering of the heart and are able to undergo epithelial to mesenchymal transition (EMT). Accordingly, genetic lineage tracing studies of the epicardium during heart development and regeneration have revealed major contributions to fibroblast and vascular smooth muscle cell (SMCs) populations of the heart [1, 2]. While epicardial fate transitions are important to the cellular makeup of the heart, epicardial derived factors are also essential to cardiac growth. Genetic ablation of the epicardium or interference with epicardial signaling pathways can lead to defects in cardiomyocyte proliferation and coronary angiogenesis, indicating that epicardial derived

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_11, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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factors influence the growth of surrounding tissues within the heart [3–7]. Because epicardial responses seemingly differ after injury in regenerative and non-regenerative organisms, a better understanding of epicardial signaling pathways may enhance attempts to regenerate the mammalian heart after injury [8]. As such, human iPSC-derived epicardial cells (iECs) are an important model system for studying fundamental human epicardial biology [9–14]. Approaches for generating epicardial cells from pluripotent cells leverage the developmental ontogeny of the epicardium during heart organogenesis. In vivo, the epicardium is derived from the proepicardium, an extracardiac structure that arises from the coelomic mesenchyme. Specification of the proepicardium from pre-cardiac mesoderm depends on Wnt, FGF, and WT1 pathways [15]. Human iPSCs can be differentiated to form self-renewing iECs through temporal modulation of canonical Wnt/β-catenin signaling and TGF-β inhibition in a fashion analogous to epicardial specification during development [9, 11, 14]. iECs express Wt1, Tbx18, and Adlh1a2, all markers of developing epicardium [14]. When treated with bFGF or TGF-β1, iECs undergo EMT in vitro to form fibroblasts and SMCs [9, 11, 14]. Transplanted iECs are able to invade the myocardium in vivo in a mouse model of myocardial infarction, demonstrating the potential for iEC-based regenerative therapies [10]. Here, we detail our protocol for differentiating iPSCs into iECs.

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Materials

2.1 Differentiation of iPSCs into iECs

1. Dulbecco’s modified Eagle medium/nutrient mixture F-12 (DMEM/F-12). 2. Geltrex LDEV-Free, hESC-Qualified Basement Membrane Mix (Gibco), 1:100 dilution in DMEM/F-12 (see Note 1). 3. 6-Well tissue culture plates. 4. mTeSR1 feeder-free maintenance medium for human ES and iPSCs (STEMCELL Technologies). 5. ROCK inhibitor Y-27632, 10 mM solution in DMSO. 6. CHIR99021, 10 mM solution in DMSO (see Note 2). 7. LaSR Media: 5 mL of 100
Glutamax (Gibco), 500 μL of 50 mg/mL gentamycin, and 500 μL of 100 mg/mL ascorbic acid in 500 mL of Advanced DMEM. 8. IWP-2, 5 mM in DMSO.

2.2 Enrichment of CDH1+ iECs

1. Accutase (Innovative Cell Technologies). 2. DMEM/F-12.

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3. EasySep™ Human Biotin Positive Selection Kit (STEMCELL Technologies). 4. EasySep™ Magnet (STEMCELL Technologies). 5. Biotinylated-CDH1 antibody (Miltenyi Biotec). 6. LaSR Media: 5 mL of 100
Glutamax (Gibco), 500 μL of 50 mg/mL gentamycin, and 500 μL of 100 mg/mL ascorbic acid in 500 mL of Advanced DMEM. 7. SB431542, 10 mg/mL in DMSO. 8. ROCK inhibitor Y-27632, 10 mM in DMSO. 9. Geltrex LDEV-Free, hESC-Qualified Basement Membrane Mix (Gibco), 1:100 dilution in DMEM/F-12. 10. 6-Well tissue culture plates. 11. Tabletop swinging bucket centrifuge. 2.3 Long-Term Maintenance of iPSC-Derived iECs

1. LaSR Media: 5 mL of 100
Glutamax (Gibco), 500 μL of 50 mg/mL gentamycin, and 500 μL of 100 mg/mL ascorbic acid in 500 mL of Advanced DMEM. 2. SB431542, 10 mg/mL in DMSO. 3. ROCK inhibitor Y-27632, 10 mM in DMSO. 4. 6-Well tissue culture plates. 5. Accutase (Innovative Cell Technologies). 6. DMEM/F-12. 7. Geltrex LDEV-Free, hESC-Qualified Basement Membrane Mix (Gibco), 1:100 dilution in DMEM/F-12.

2.4 Differentiation of iECs into Cardiac Fibroblasts and Smooth Muscle Cells

1. 8-Well chamber slides. 2. Geltrex LDEV-Free, hESC-Qualified Basement Membrane Mix (Gibco), 1:100 dilution in DMEM/F12. 3. LaSR Media: 5 mL of 100
Glutamax (Gibco), 500 μL of 50 mg/mL gentamycin, and 500 μL of 100 mg/mL ascorbic acid in 500 mL of Advanced DMEM. 4. ROCK inhibitor Y-27632, 10 mM in DMSO. 5. bFGF, 10 μg/mL in DMSO. 6. TGFβ1, 10 μg/mL in DMSO.

2.5 Immunostaining of iECs and iEC Derivatives

1. 1
Phosphate-buffered saline (PBS). 2. 4% Paraformaldehyde (PFA), prepared in PBS. 3. PBST: 0.1% Tween-20 in PBS. 4. Blocking solution: 10% Newborn calf serum, 2% horse serum, 1% DMSO in PBST.

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5. Antibody solution: 10% Newborn calf serum, 1% DMSO in PBST. 6. APC-Anti-Thy1 antibody 1:100 (BD Biosciences). 7. Anti-calponin antibody 1:200 (Abcam). 8. Anti-WT1 antibody 1:100 (Abcam). 9. 40 ,6-Diamidino-2-phenylindole (DAPI) 1:5000 dilution of 5 mM stock. 10. Alexa Fluor secondary antibodies, 1:200 (Thermo Fisher). 11. Mounting media. 12. 8-Well chamber slides 13. #1.5 thickness coverslips. 2.6 Titration of CHIR99021 to Increase Efficiency of Mesoderm Differentiation

1. 8-Well chamber slides. 2. mTeSR1 feeder-free maintenance medium for human ES and iPS cells (STEMCELL Technologies). 3. ROCK inhibitor Y-27632, 10 mM in DMSO. 4. CHIR99021, 10 mM in DMSO. 5. LaSR Media: 5 mL of 100
Glutamax (Gibco), 500 μL of 50 mg/mL gentamycin, and 500 μL of 100 mg/mL ascorbic acid in 500 mL of Advanced DMEM. 6. PBS. 7. 4% PFA, prepared in PBS. 8. PBST: 0.1% Tween 20 in PBS. 9. Blocking solution: 10% Newborn calf serum, 2% horse serum, 1% DMSO in PBST. 10. Antibody solution: 10% Newborn calf serum, 1% DMSO in PBST. 11. Anti-Brachyury Antibody, 1:100 (R&D Systems). 12. Alexa Fluor 594 chicken anti-Goat Antibody, 1:200 (Thermo Fisher). 13. DAPI, 1:5000 dilution of 5 mM stock solution. 14. Mounting media. 15. #1.5 thickness coverslips.

3

Methods

3.1 Epicardial Differentiation of iPSCs

A schedule of events is presented in Fig. 1. iPSCs should be free from spontaneous differentiation and over 95% confluent before starting the epicardial differentiation (Fig. 2a). Starting with this level of confluency is important to the final iEC yield because

Generation of iPSC-Derived Epicardial Cells

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Fig. 1 Overview of iEC differentiation protocol. iPSCs are plated to >98% confluency and then treated with the GSK3 inhibitor CHIR99021 to induce mesoderm progenitors. Cells are then transferred to LaSR medium and treated with the Wnt inhibitor IWP-2 to generate cardiac progenitor cells. After expansion, cardiac progenitors are again treated with the GSK3 inhibitor CHIR99021 to generate iECs. iECs can be further enriched by positive selection for CDH1

Fig. 2 Representative images of iECs. (a, b) Phase-contrast images of iPSCs and iECs show cell morphology differences. iECs are larger and have a cuboidal morphology (c, d). Immunostaining of iPSCs and iECs for the epicardial marker WT1 (green) and DAPI (blue). Epicardial cells express WT1 with approximately 80% purity. Scale bars indicate 50 μm

subsequent treatment with CHIR99021 results in moderate amounts of iPSC loss.

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1. In a Geltrex-coated 6-well tissue culture plate (see Note 3), add iPSCs in pre-warmed mTeSR1 medium (see Note 4) supplemented with 5 μM of ROCK inhibitor. Each well should have approximately 500,000 iPSCs in 2 mL of culture medium. This initial plating corresponds to day 3. 2. On days 2 and 1, feed the cells by aspirating the media and replacing it with 2 mL of prewarmed mTeSR1 into each well. 3. On day 0, feed the cells by replacing the medium in each well with 2 mL of LaSR medium containing 6 μM CHIR99021 (see Note 5). Record the time. 4. On day 1, exactly 24 h after the addition of CHIR99021, feed each well of cells with 2 mL of prewarmed LaSR basal medium (see Note 6). We typically observe a moderate amount of iPSC death after treatment with CHIR99021. Continue the differentiation with the surviving cells. 5. On day 3, 72 h after addition of CHIR99021, feed each well of cells with 2 mL of LaSR medium containing 5 μM IWP-2. 6. On days 4–5, monitor the cells for confluency. If the cells are becoming too confluent, cells can be split on day 5 (see Note 7). 7. On day 6, feed each well of cells with 2 mL of LaSR medium containing 3 μM CHIR99021. Repeat again on day 7. 8. On days 8–10, feed each well of cells daily with 2 mL of prewarmed LaSR basal medium. At this stage, iECs exhibit a cuboidal morphology and express the epicardial marker WT1 (Fig. 2). An immunostaining protocol for WT1 is provided in Subheading 3.4. 3.2 Enrichment of CDH1+ iECs

At this stage, we observe that greater than 80% of cells express WT1 (Fig. 2c). To further enrich the number of iECs, we perform a positive selection for iECs expressing the pan-epithelial marker, CDH1. 1. For each well of iECs, aspirate the medium and rinse with 1 mL of room-temperature DMEM/F-12 medium. Add 1 mL of room-temperature Accutase to each well. Put the plate in a 37  C, 5% CO2, incubator and wait for 5–7 min. 2. To each well, add 1 mL of DMEM/F-12 medium. Gently pipette up and down to singularize the cells. After pipetting, transfer the cells to a 15 mL conical tube (see Note 5). Centrifuge the cells at 300
g for 3 min using a swinging bucket centrifuge. 3. From the tube of pelleted cells, aspirate the supernatant. Resuspend the cells in 0.1–2.5 mL of EasySep Buffer from the EasySep™ Human Biotin Positive Selection Kit. Cells should be resuspended to a concentration of approximately 1
108 cells/

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mL. Transfer the cell suspension to a 5 mL polystyrene roundbottom tube. Add 100 μL/mL of FcR Blocker to each tube and gently mix by flicking the tube. Add the biotinylated antiCDH1 antibody at 3 μg/mL. Incubate at room temperature for 15 min. 4. To each tube, add 100 μL/mL of Selection Cocktail from the EasySep™ Human Biotin Positive Selection Kit. After mixing, incubate for 15 min at room temperature. 5. Vortex the RapidSpheres from the EasySep™ Human Biotin Positive Selection Kit for 30 s. Add 50 μL of RapidSpheres per mL of solution to each tube. Mix and incubate for 10 min at room temperature. Add 2 mL of EasySep Buffer to each tube and mix gently by pipetting up and down. Place each tube in the EasySep Magnet tube holder and incubate for 10 min at room temperature. Make sure to loosely cap each tube so that the pelleted beads will not be agitated when removing the cap. 6. Decant the supernatant by picking up the EasySep Magnet containing the tube of cells and pouring it into a waste container in one continuous motion. Hold the tube and magnet inverted for 2 s before returning upright. Remove the tube from the magnet and add 1 mL of EasySep Buffer, pipetting up and down to evenly wash the walls of the tube. Lightly tap the tube on each side and add another 1 mL of EasySep Buffer. Replace the tube in the magnet and repeat for a total of three washes. 7. After the last incubation, decant the supernatant, remove the tube from the magnet, and resuspend the cells in 2 mL of LaSR medium containing 2 μM of SB431542 and 5 μM of ROCK inhibitor. Plate the cells at 200,000 cells per well of a Geltrexcoated 6-well plate. 3.3 Long-Term Maintenance of iECs

1. For each well of cells, aspirate the medium and add 2 mL of prewarmed LaSR basal medium supplemented with 2 μM of SB431542. Return the plate to the 37  C, 5% CO2, incubator. Change the media daily. Maintain the iECs’ cuboidal morphology during maintenance. Loss of cuboidal morphology can indicate spontaneous transdifferentiation. 2. When the iECs are 80–90% confluent, the cells can be split. To split the iECs, aspirate the medium from each well of cells and rinse with 1 mL of room-temperature DMEM/F-12 medium. Next, add 1 mL of room-temperature Accutase to each well. Place the cells in a 37  C, 5% CO2, incubator and wait for 5 min. After incubation, add 1 mL of DMEM/F-12 medium to each well of cells and pipette up and down with a glass pipette to singularize the cells (see Note 5). Transfer the singularized cells to a 15 mL conical tube. Centrifuge the cells at

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bFGF bFGF bFGF bFGF bFGF bFGF OR SB

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Fig. 3 Timeline of growth factor treatment for differentiating iECs into cardiac fibroblasts or SMCs. iECs are removed from maintenance LASR medium supplemented with SB431542 and treated with either bFGF (10 ng/mL) or TGFβ1 (5 ng/mL) for 6 days to generate fibroblasts or SMCs, respectively. Medium and growth factor are changed daily

300
g for 3 min in a swinging bucket centrifuge. Aspirate the medium and resuspend the cell pellet in LaSR basal medium supplemented with 2 μM of SB431542 and 5 μM of ROCK inhibitor. Cells can be replated onto the Geltrex-coated plates using a split ratio of 1:3. 3.4 Differentiation of iECs into Cardiac Fibroblasts and Smooth Muscle Cells

A schedule for differentiating iECs into fibroblasts or SMCs is presented in Fig. 3. 1. On day 1, plate 30,000 iECs in 250 μL of LaSR medium supplemented with 5 μM ROCK inhibitor into each well of a Geltrex-coated 8-well chamber slide. 2. On day 0, aspirate the medium and replace it with LaSR medium supplemented with a differentiating growth factor. To generate fibroblasts, use bFGF at 10 ng/mL. To generate smooth muscle cells, add TGFβ1 at 5 ng/mL. Concentrations of growth factors are estimates based on our experience but can vary based on the manufacturer and the lot. 3. For days 1–5, change the media daily with LaSR supplemented with differentiating growth factor. 4. Efficiency of differentiation can be verified by immunostaining for Thy1 to detect fibroblasts or CNN1 to label SMCs as detailed below (Fig. 4).

3.5 Immunostaining Analysis

1. For each well of a chamber slide, aspirate the medium and add 1 mL of PBS to wash the cells. 2. Aspirate the PBS and add 300 μL of 4% PFA to each well. Incubate for 10 min at room temperature to fix the cells. Aspirate the PFA. Next, add 1 mL of PBST to each well and aspirate to rinse the cells. Repeat the PBST rinse step twice, waiting 5 min between rinses.

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Fig. 4 Differentiation of iECs in cardiac fibroblasts or SMCs. iECs were treated with either bFGF or TGFβ1 and immunostained for the fibroblast marker Thy1 (red), the SMC marker CNN1 (green), and DAPI (blue). iECs have very few Thy1+ or CNN1+ cells. However, a subset of iECs treated with bFGF express Thy1 and iECs treated with TGFβ1 express CNN1. Scale bars indicate 50 μm

3. Add 300 μL of blocking buffer to each well. Block for 1 h at room temperature. 4. Primary antibodies are added in antibody solution. To stain for fibroblasts, use anti-Thy1 at 1:100. To stain for smooth muscle cells, use anti-calponin at 1:200. To stain for epicardial cells, use anti-WT1 at 1:100. Incubate cells with the primary antibody solution at 4  C overnight. Alternatively, cells can be incubated with primary antibody staining solution for 3 h at 37  C. 5. Aspirate the primary antibody solution and wash each well of cells with 1 mL of PBST four times, waiting 5 min between washes. 6. Dilute the secondary antibody conjugated to a fluorophore of choice and DAPI in fresh antibody solution. Apply the secondary antibody solution to each well and incubate for 1 h at room temperature. Aspirate the secondary antibody solution from

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each well and rinse with 1 mL of PBST four times, waiting 5 min between washes. 7. Carefully remove the chamber portion of the chamber slide. Add the mounting medium and apply a coverslip. The slides can now be imaged. 3.6 Titration of CHIR99021 to Increase Efficiency of Mesoderm Differentiation

1. On day 3, add 30,000 iPSCs in 250 μL of mTeSR1 supplemented with 5 μM ROCK inhibitor to each well of a Geltrexcoated 8-well chamber slide. 2. On days 2 and 1, feed the cells by aspirating the medium and replacing it with 250 μL of prewarmed mTeSR1 into each well. 3. On day 0, feed the cells in each well with a different concentration of CHIR99021, ranging from 4 to 12 μM in LaSR. Note the time. 4. On day 1, exactly 24 h after the addition of CHIR99021, feed the cells with 250 μL of prewarmed LaSR basal medium. 5. On day 2, aspirate the medium and add 1 mL of PBS per well to wash the cells. Aspirate the PBS, add 300 μL of 4% PFA to each well, and incubate for 10 min at room temperature to fix the cells. Aspirate the PFA solution and then add 1 mL of PBST to each well. Aspirate the PBST after 5 min. Repeat the PBST wash two more times. 6. Add 300 μL of blocking buffer to each well. Block for 1 h at room temperature. 7. Add the anti-Brachyury antibody at 1:100 in 150 μL of antibody solution to each well. Incubate at 4  C overnight or at 37  C for 3 h. 8. Wash each well with 1 mL of PBST four times, waiting 5 min between washes. 9. Dilute the secondary antibody conjugated to a fluorophore of choice and DAPI in antibody solution. Apply 150 μL of secondary antibody solution to each well and incubate for 1 h at room temperature. Rinse with 1 mL of PBST four times, waiting 5 min between washes. 10. Remove the chamber portion of the chamber slide. Add mounting media and apply a coverslip. The slides are ready for imaging. 11. The concentration of CHIR99021 that yields the highest percentage of Brachyury+ cells with the least amount of cell death should be used for future experiments (Fig. 5).

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Fig. 5 Optimizing differentiation of iPSCs into mesodermal progenitors by modulating GSK3 inhibition. iPSCs were treated with increasing concentrations of CHIR99021 for 24 h and immunostained for the mesodermal marker Brachyury (red) and DAPI (blue). We typically observe that treatment with 10–12 μM CHIR99021 results in the largest percentage of Brachyury+ cells. Scale bar indicates 100 μm

4

Notes 1. An alternate basement membrane matrix, such as Matrigel (Corning), can also be used. 2. The effective concentration of CHIR99021 varies with the cell line, lot, and manufacturer. For each iPSC line and lot of CHIR99021, we optimize the CHIR99021 concentration as described in Subheading 3.6. 3. Coat the tissue culture vessels with Geltrex diluted 1:100 in DMEM/F-12 using the manufacturer’s guidelines. 4. Prewarm all cell culture reagents and vessels before use. Warm the media to 37  C and warm all other cell culture reagents and growth factors to room temperature. 5. When using Accutase to detach and singularize the cells, ensure that all cells are lifted and removed. If cells remain adherent to the plate, treat the wells with a second round of Accutase for 5 min at 37  C. Hold the singularized cells in DMEM/F-12 and combine before centrifugation. 6. For best practices, keep intervals between medium changes consistent and no longer than 24 h. 7. iPSCs proliferate well enough that they can become overconfluent too early in the epicardial differentiation protocol. To ensure maximal differentiation, the cells can be split to prevent

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overgrowth and clumping at day 5 before the next round of CHIR99021. Cells should be split heavily enough to ensure >75% confluency for day 6.

Acknowledgments This work was supported by R03 HL144812 (to R.K.), the Duke Strong Start Physician Scientist Program (R.K.), the Walker P. Inman Endowment (R.K.), and the Edna and Fred L. Mandel, Jr., Foundation (R.K.). References 1. Kikuchi K, Gupta V, Wang J, Holdway JE, Wills AA, Fang Y, Poss KD (2011) tcf21+ epicardial cells adopt non-myocardial fates during zebrafish heart development and regeneration. Development 138(14):2895–2902. https://doi.org/10.1242/dev.067041 2. Acharya A, Baek ST, Huang G, Eskiocak B, Goetsch S, Sung CY, Banfi S, Sauer MF, Olsen GS, Duffield JS, Olson EN, Tallquist MD (2012) The bHLH transcription factor Tcf21 is required for lineage-specific EMT of cardiac fibroblast progenitors. Development 139(12):2139–2149. https://doi.org/10. 1242/dev.079970 3. Wang J, Cao J, Dickson AL, Poss KD (2015) Epicardial regeneration is guided by cardiac outflow tract and Hedgehog signalling. Nature 522(7555):226–230. https://doi.org/10. 1038/nature14325 4. Lepilina A, Coon AN, Kikuchi K, Holdway JE, Roberts RW, Burns CG, Poss KD (2006) A dynamic epicardial injury response supports progenitor cell activity during zebrafish heart regeneration. Cell 127(3):607–619. https:// doi.org/10.1016/j.cell.2006.08.052 5. Shen H, Cavallero S, Estrada KD, Sandovici I, Kumar SR, Makita T, Lien CL, Constancia M, Sucov HM (2015) Extracardiac control of embryonic cardiomyocyte proliferation and ventricular wall expansion. Cardiovasc Res 105(3):271–278. https://doi.org/10.1093/ cvr/cvu269 6. Li P, Cavallero S, Gu Y, Chen TH, Hughes J, Hassan AB, Bruning JC, Pashmforoush M, Sucov HM (2011) IGF signaling directs ventricular cardiomyocyte proliferation during

embryonic heart development. Development 138(9):1795–1805. https://doi.org/10. 1242/dev.054338 7. Huang Y, Harrison MR, Osorio A, Kim J, Baugh A, Duan C, Sucov HM, Lien CL (2013) Igf signaling is required for cardiomyocyte proliferation during zebrafish heart development and regeneration. PLoS One 8(6): e67266. https://doi.org/10.1371/journal. pone.0067266 8. Kikuchi K, Holdway JE, Major RJ, Blum N, Dahn RD, Begemann G, Poss KD (2011) Retinoic acid production by endocardium and epicardium is an injury response essential for zebrafish heart regeneration. Dev Cell 20 (3):397–404. https://doi.org/10.1016/j. devcel.2011.01.010 9. Bao X, Lian X, Qian T, Bhute VJ, Han T, Palecek SP (2017) Directed differentiation and long-term maintenance of epicardial cells derived from human pluripotent stem cells under fully defined conditions. Nat Protoc 12 (9):1890–1900. https://doi.org/10.1038/ nprot.2017.080 10. Bao X, Lian X, Hacker TA, Schmuck EG, Qian T, Bhute VJ, Han T, Shi M, Drowley L, Plowright A, Wang QD, Goumans MJ, Palecek SP (2016) Long-term self-renewing human epicardial cells generated from pluripotent stem cells under defined xeno-free conditions. Nat Biomed Eng 1:0003. https://doi.org/10. 1038/s41551-016-0003 11. Iyer D, Gambardella L, Bernard WG, Serrano F, Mascetti VL, Pedersen RA, Talasila A, Sinha S (2015) Robust derivation of epicardium and its differentiated smooth

Generation of iPSC-Derived Epicardial Cells muscle cell progeny from human pluripotent stem cells. Development 142(8):1528–1541. https://doi.org/10.1242/dev.119271 12. Guadix JA, Orlova VV, Giacomelli E, Bellin M, Ribeiro MC, Mummery CL, Perez-Pomares JM, Passier R (2017) Human pluripotent stem cell differentiation into functional epicardial progenitor cells. Stem Cell Rep 9 (6):1754–1764. https://doi.org/10.1016/j. stemcr.2017.10.023 13. Zhao J, Cao H, Tian L, Huo W, Zhai K, Wang P, Ji G, Ma Y (2017) Efficient differentiation of TBX18(+)/WT1(+) epicardial-like cells from human pluripotent stem cells using

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small molecular compounds. Stem Cells Dev 26(7):528–540. https://doi.org/10.1089/ scd.2016.0208 14. Witty AD, Mihic A, Tam RY, Fisher SA, Mikryukov A, Shoichet MS, Li RK, Kattman SJ, Keller G (2014) Generation of the epicardial lineage from human pluripotent stem cells. Nat Biotechnol 32(10):1026–1035. https:// doi.org/10.1038/nbt.3002 15. Brade T, Pane LS, Moretti A, Chien KR, Laugwitz KL (2013) Embryonic heart progenitors and cardiogenesis. Cold Spring Harb Perspect Med 3(10):a013847. https://doi.org/10. 1101/cshperspect.a013847

Chapter 12 In Vitro Conversion of Murine Fibroblasts into Cardiomyocyte-Like Cells Jun Xu, Li Wang, Jiandong Liu, and Li Qian Abstract Direct reprogramming of fibroblasts into induced cardiomyocytes (iCMs) holds great promise as a potential treatment for cardiovascular disease, many of which are associated with tremendous loss of functional cardiomyocytes and simultaneous formation of scar tissue. Burgeoning studies have shown that the introduction of three minimal transcriptional factors, Gata4, Mef2c, and Tbx5 (G/M/T), could convert murine fibroblasts into iCMs that closely resemble endogenous CMs both in vitro and in vivo. Recent studies on iCM cell fate determination have demonstrated that the removal of genetic and epigenetic barriers could facilitate iCM reprogramming. However, varied reprogramming efficiency among research groups hinders its further study and potential applicability. Here, we provide a newly updated and detailed protocol for in vitro generation and evaluation of functional iCMs from mouse embryonic fibroblasts and neonatal cardiac fibroblasts using retroviral polycistronic construct encoding optimal expression of G/M/T factors. We hope that this optimized protocol will lay the foundation for future mechanistic studies of murine iCMs and further improvement of iCM generation. Key words Direct reprogramming, Induced cardiomyocyte, Fibroblast, MGT, Cardiac regeneration

1

Introduction Cardiovascular disease (CVD) produces immense health burdens worldwide. In 2016, approximately 17.6 million deaths were attributed to CVD globally, a 14.5% increase since 2006 [1]. It was reported that a myocardial infarction can destroy nearly 25% cells of the left ventricle within a few hours [2]. Due to the limited regenerative capacity of the human heart, cardiomyocyte (CM) loss postmyocardial infarction may result in a progressive decrease in the myocardial contractility with persistent severe pressure and volume load, eventually leading to heart failure [3]. Current medical treatment, including pharmacological and mechanical therapies, is largely focused on preventing cardiac remodeling toward heart failure but cannot overcome the challenge of permanent CM loss.

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_12, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Cardiac fibroblasts (CF) are one of the major cell populations in the heart and play a vital role in heart development and homeostasis under normal conditions[4]. In response to ischemia and other injurious stimuli, CFs migrate to the site of injury, proliferate, and finally form scar tissue that contributes to the dampening of heart function [5]. Even though the biological importance of CFs has long been recognized, their potential usage for cardiac regeneration has been explored only recently. The CF reservoir could serve as a resident source for cardiac regeneration if they could be directly converted into functional cardiomyocytes. Direct conversion between somatic cell types can be achieved using fibroblasts to generate various types of cells, including blood cells [6], neurons [7], hepatocytes [8, 9], and cardiomyocytes [10– 14], without passing through a pluripotent or progenitor stage. We and others have shown that retroviral delivery of three transcriptional factors, Mef2c, Gata4, and Tbx5 (M/G/T), is sufficient to reprogram cultured fibroblasts into induced cardiomyocyte-like cells (iCMs) that display similar cellular and physiological features of endogenous cardiomyocytes [10]. Such direct iCM conversion can be induced in vivo, resulting in reduced scar size and improved cardiac function in an infarcted heart [11]. These studies indicate that this iCM reprogramming approach holds great potential for healing an injured heart. However, several hurdles for iCM generation, such as inadequate reprogramming efficiency and unclear molecular mechanisms, hinder its further application. Recent studies have revealed that removing genetic and epigenetic barriers, adding microRNAs and small molecules, modulating cellular signaling pathways, and using a variety of delivery methods, could enhance reprogramming efficiency [15–24]. These studies differ in their sources of fibroblasts, reprogramming factor cocktails, methods to generate viruses for delivery, markers, and timelines for reprogramming evaluation. Therefore, it poses a difficulty in reaching a consensus protocol to generate iCM in dish. Our previous work has investigated the role of M/G/T stoichiometry on iCM reprogramming and found that optimal expression of M/G/T factors (high expression of M, and low expression of G and T) results in quantity- and quality-improved iCM generation both in vitro and in vivo [25]. Based on this research, we have developed a unique platform to efficiently and reproducibly convert CFs into iCMs using our MGT polycistronic construct. Here, we will describe in detail our updated step-by-step protocol for in vitro iCM reprogramming using this MGT system to efficiently generate iCMs from CFs and mouse embryonic fibroblasts in culture.

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Materials

2.1 Generation of Retroviruses

1. pMx-puro-MGT retroviral expression vector (Addgene ID number: 111809). 2. pMx-puro-dsRed retroviral expression vector [11, 25]. 3. Plat-E cell retroviral packaging cell line. 4. Dulbecco’s modification of Eagle’s medium (DMEM), liquid (high glucose), with L-glutamine (Gibco). 5. FBS (Hyclone). 6. Penicillin-streptomycin (Gibco). 7. Puromycin (ready-to-use solution, 10 mg/mL in H2O, 0.2 μm filtered; Sigma). 8. Blasticidin (Invitrogen). 9. Nonessential amino acid solution (NEAA; Invitrogen). 10. NanoFect transfection reagent (Alstem, NF101). 11. 0.45 μm Syringe filters. 12. Syringes, 30 and 10 mL (BD). 13. Retro-X qRT-PCR titration kit (Clontech). 14. PEG6000 (Sigma). 15. NIH3T3 cell line. 16. Centrifuge tubes, 15 mL (Corning). 17. Centrifuge tubes, 50 mL (Corning). 18. Cell culture dish, 100 mm
20 mm (Corning). 19. Cell strainer, 40 μm (BD Falcon). 20. Microcentrifuge tubes, 1.5 mL (Eppendorf). 21. Flow cytometer. 22. Plat-E culture medium: DMEM supplemented with 10% (vol/vol) fetal bovine serum (FBS), 1
nonessential amino acids, 1% (vol/vol) penicillin/streptomycin. While maintaining Plat-E cells, add 1 μg/mL puromycin and 10 μg/mL blasticidin. 23. Plat-E transfection medium: DMEM supplemented with 10% (vol/vol) fetal bovine serum, 1
nonessential amino acids.

2.2 Generation of Mouse Embryonic Fibroblasts (MEFs)

1. αMHC (α-myosin heavy chain)-GFP transgenic mouse embryos, 12.5–14.5 d.p.c. (day post-coitum). 2. PBS without Ca2+ and Mg2+ (Gibco). 3. Trypsin-EDTA, 0.05% (wt/vol) (Gibco, Life Technologies). 4. Cell culture dish, 100 mm
20 mm (Corning). 5. Delicate scissors (Fine Science Tools).

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6. Forceps (Fine Science Tools). 7. Sterile razor blade. 8. CO2 incubator (CO2 at 5%, humidified, 37  C). 9. Water bath maintained at 37  C. 10. MEF culture medium: DMEM supplemented with 10% (vol/vol) fetal bovine serum (FBS), 1% (vol/vol) penicillin/ streptomycin. 2.3 Generation of Neonatal Mouse Fresh Cardiac Fibroblasts

1. Neonatal αMHC (α-myosin heavy chain)-GFP transgenic mice. 2. M199 (BioWhittaker, cat. no. BE12-119F). 3. Iscove’s modified Dulbecco’s medium (IMDM; Gibco). 4. PBS without Ca2+ and Mg2+ (Gibco). 5. Trypsin-EDTA, 0.05% (wt/vol) (Gibco, Life Technologies). 6. Collagenase type 2 (Worthington Biochemical Corporation). 7. HBSS, 10
(Cellgro). 8. MACS kit (Miltenyi Biotec). 9. Anti-Biotin MicroBeads (Miltenyi Biotec). 10. Anti-mouse CD90.2 (Thy-1.2) antibody (BD Biosciences). 11. Delicate scissors (Fine Science Tools). 12. Forceps (Fine Science Tools). 13. 0.22 μm Syringe filters. 14. Water bath maintained at 37  C. 15. Tissue culture centrifuge. 16. Fibroblast (FB) medium: IMDM supplemented with 20% (vol/vol) FBS and 1% (vol/vol) penicillin-streptomycin.

2.4 Cardiac Reprogramming In Vitro and Evaluation of Reprogramming Efficiency

1. Monoclonal mouse anti-α-actinin (sarcomeric) clone EA-53 (Sigma). 2. Anti-GFP, rabbit IgG fraction (anti-GFP, IgG), 2 mg/mL (polyclonal; Invitrogen). 3. Mouse anti-troponin T, cardiac isoform (Thermo Scientific). 4. Alexa Fluor 488-conjugated donkey anti-rabbit IgG (Jackson ImmunoResearch Inc). 5. Alexa Fluor 647-conjugated donkey anti-mouse IgG (Jackson ImmunoResearch Inc). 6. Gelatin, 0.1% (wt/vol) solution (Sigma). 7. Polybrene (Chemicon). 8. BD Cytofix/Cytoperm Solution Kit (BD). 9. TRIzol reagent (Life Technologies).

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10. Chloroform. 11. Isopropyl alcohol. 12. Ethanol. 13. SuperScript IV VILO Master Mix (Invitrogen 00620085). 14. SYBR Green PCR Master Mix (Applied Biosystems, 1710515). 15. PFA, 32% (wt/vol), electron microscopy grade, 10
10 mL (Electron Microscopy Sciences). 16. Triton X-100 (Sigma). 17. Hoechst 33342 (Life Technologies, H3570). 18. Rhod-3 Calcium Imaging Kit (Thermo Fisher Scientific). 19. Cell culture plate, 24 wells (Corning). 20. CO2 incubator (CO2 at 5%, humidified, 37  C). 21. EVOS FL Auto imaging system. 22. Nanodrop spectrophotometer. 23. ABI Thermocycler. 24. QuantStudio 6 Flex (Thermo Fisher Applied Biosystems). 25. Cardiomyocyte medium: Supplement DMEM media with 18% (vol/vol) M199 media and 10% (vol/vol) FBS, 1% (vol/vol) NEAA, 1% (vol/vol) penicillin-streptomycin. 26. B27 medium: Supplement 500 mL of RPMI1640 media with 20 mL of B27 media. 27. Gelatin-coated 24-well plate: Add 0.5 mL of 0.1% (wt/vol) gelatin solution to a well of 24-well plate, incubate at 37  C for at least 10 min, and aspirate right before use. 28. FACS labeling buffer: Supplement 500 mL of DPBS with 10 mL of FBS and 2 mL of EDTA (0.5 M) to reach a final concentration of 2 mM. Pass through 0.22 μm filter and store at 4  C. 29. MACS sorting buffer: Supplement 500 mL of DPBS with 2.5 g of BSA and 2 mL of EDTA (0.5 M) to reach a final concentration of 2 mM. Pass through 0.22 μm filter and store at 4  C. 30. Immunocytochemistry (ICC) fixation buffer: Add 5 mL of paraformaldehyde (PFA, 32%) to 35 mL of DPBS to reach a final concentration of 4% (vol/vol). 31. ICC permeabilization buffer: Add 0.1 mL of Triton to 100 mL of DPBS to reach a final concentration of 0.1% (vol/vol). 32. ICC blocking buffer: Add 5 g of bovine serum albumin (BSA) to 100 mL of DPBS to reach a final concentration of 5% (wt/vol). Pass through 0.22 μm filter and store at 4  C.

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33. ICC staining buffer: Add 1 g of BSA to 100 mL of DPBS to reach a final concentration of 1% (wt/vol). Pass through 0.22 μm filter and store at 4  C.

3

Methods

3.1 Generation of Retroviruses (See Note 1)

1. A single polycistronic vector with an optimal ratio of Mef2c, Gata4, and Tbx5 significantly increased reprogramming efficiency both in vitro and in vivo [26, 25]. pMx-puro-dsRed serves as an indicator for the evaluation of transduction efficiency. 2. Maintain Plat-E cells in Plat-E media supplemented with 1 μg/ mL of puromycin and 10 μg/mL of blasticidin at 37  C with 5% CO2. 3. On day 2, withdraw puromycin and blasticidin from Plat-E cells. 4. On day 0, split Plat-E cells into 100 mm dishes at approximately 4–5
106 cells per dish. Cells should be ~80–90% confluent on the day of transfection. 5. On day 1, change the culture media to transfection media on cells at least 1 h prior to transfection. Transfect the Plat-E cells with NanoFect when the culture reaches ~80% confluency. Set up the following two mixtures: (1) dilute 20 μg of packaging vectors in 350 μL of DMEM medium (room temperature); (2) dilute 45 μL of NanoFect in 305 μL of DMEM medium. Slowly add the NanoFect mixture to the plasmid mixture and vortex at a slow speed (4–5/10) for 10 s. Incubate the DNA-NanoFect mixture at room temperature (RT) for 15 min. 6. Add the transfection mixture dropwise to the Plat-E cells. Mix it well by moving the plate in a back-and-forth, side-to-side motion. Incubate overnight in a 37  C incubator. 7. After ~16 h (day 2), replace with 8 mL of fresh prewarmed transfection culture media. Incubate for 24 h. 8. On day 3 and day 4, harvest the retrovirus by collecting culture supernatant from the Plat-E dishes with a 10 mL/30 mL sterile disposable syringe, filtering it through a 0.45 μm pore size cellulose acetate filter, and transferring into a 50 mL tube. Add 2 mL of 40% PEG6000 to every 8 mL of virus-containing supernatant. Gently mix it and incubate overnight at 4  C (see Note 2). 9. On the next day (day 4 and day 5, respectively), spin the viral mixture at 1500
g at 4  C for 30 min. The viral particles should appear in a white pellet at the bottom of the tube.

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Discard the supernatant. Spin down the residual solution by centrifugation at 1500
g for 5 min. Remove all traces of fluid by aspiration, taking great care not to disturb the precipitated retroviral particles in the pellet. 10. Resuspend the retroviral pellets with 100 μL of DMEM for every 8 mL of viral supernatant. Aliquot the virus for further usage (see Note 3). 11. Determine the total copy number of all packaged retroviruses using a Retro-X qRT-PCR titration kit. 12. Determine the infectious unit (IFU) of fluorescent retrovirus. Seed 3
104 NIH3T3 cells per well on a 24-well plate in 1 mL of medium per well and incubate the plate overnight. On the second day, count the cells in one well and transduce the cells with dsRed retrovirus with 5–10-fold serial dilution. Incubate the cells overnight. Digest the cells, analyze fluorescence by FACS, and read the percentage from linear values (usually 5–10%). The titer (IFU) is the percentage of the cells transduced by a given volume of the cell number counted above. 13. Calculate the percentage of infectious virus and fluorescent retrovirus by comparing the IFU with the total copy number of virus (total copy number divided by IFU). The IFU of nonfluorescent retroviruses can be calculated by multiplying the total copy number by the percentage of infectious fluorescent protein virus calculated above. For example, if the percentage of infectious virus in the dsRed retrovirus is 80% and the total copy number of the Gata4 retrovirus is 1
108 copies/μL, then the IFU of Gata4 is calculated as 80%
(1
108) ¼ 0.8
108 copies per μL (see Note 4). 3.2 Generation of Mouse Embryonic Fibroblasts (MEFs)

1. Euthanize a pregnant mouse at 12.5–14.5 d.p.c. 2. Dissect out the uterine horns, briefly rinse in 70% (vol/vol) ethanol, and place the embryos in a 100 mm sterile petri dish containing PBS without Ca2+ and Mg2+ (see Note 5). 3. Remove the limbs, head, and insides with forceps and scissors. 4. Place embryos in a 100 mm dish with a small volume of PBS. Finely mince the tissue with a sterile razor blade until it becomes possible to pipette. 5. Place the sludge into a 15 mL tube with 1 mL of 0.05% (wt/vol) trypsin-EDTA, including 100 Kunitz units of DNase I per embryo. Incubate at 37  C for 15 min. After each 5 min of incubation, dissociate the cells by pipetting up and down thoroughly. 6. Inactivate the trypsin by adding about one volume of freshly prepared MEF medium.

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7. Centrifuge the cells at the speed of 200
g for 5 min, then carefully remove the supernatant, and resuspend cell pellet in warm MEF medium. 8. Plate approximately one embryo’s worth of cells in each well of 6-well plate coated with 0.1% (wt/vol) gelatin. 9. Change media the next day. Ideally, cells are 80–90% confluent after 24 h. 10. Freeze the cells at a density of 5
106 for future usage. Keep in 80  C freezer for 1 week and then transfer to liquid nitrogen for long-term storage. 3.3 Generation of Neonatal Mouse Fresh Cardiac Fibroblasts (See Note 6) 3.3.1 Generate Cardiac Fibroblasts from Explant Culture Method

1. Clean a neonatal αMHC-GFP transgenic mouse, postnatal days 1 and 2 (P1 to P2) with 75% ethanol. Remove the head with sterile scissors. Then make a horizontal incision from under one armpit to the other. From here, the heart can be squeezed out. Use a sterile blunt and bent forceps to dissect out the heart and place it in one well of a 24-well plate containing ice-cold PBS buffer (see Note 7). 2. Check GFP expression in the heart by using a fluorescent microscope. Choose GFP-positive hearts and pool them into a 100 mm dish (see Note 8). 3. Cut the heart into small pieces less than 1 mm3 in size with a sterile blade or scissors. 4. Place 3–4 minced hearts into one 100 mm dish with 2 mL of mouse FB media. Let the tissues settle down for 3 h in a 37  C incubator. 5. Slowly add 8 mL of pre-warmed FB media to the dishes containing heart tissues, and continue to culture for 1 week. Do not disturb the tissues for 3 days. 6. Replace the media every 3 days. 7. On day 7, aspirate the culture medium and wash the cells with DPBS. Add 3 mL of 0.05% trypsin to each plate and digest at 37  C for 5 min. Add 5 mL of FB media to quench trypsin. Gently detach the cells with a cell scraper. Pipette the media up and down to further mechanically dissociate the tissue. 8. Collect cells and filter through 40 μm cell strainers to avoid contamination of heart tissue fragments, and then pellet the cells by spinning at 200
g for 5 min. 9. Wash cells once with the MACS buffer and cells are ready for sorting.

3.3.2 Generate Cardiac Fibroblasts from Enzyme Digestion Method

1. Harvest GFP-positive hearts as directed in Subheading 3.3.1. Transfer all of the hearts into a 100 mm dish containing 10 mL of ice-cold DPBS. Squeeze the ventricles with sterile forceps to remove blood and rinse once.

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2. Trim the hearts to remove other tissue and fat. Cut the hearts into small pieces, but do not disconnect them. 3. Incubate the heart tissues in a warm 0.05% trypsin-EDTA solution at 37  C for 15 min. If there are less than 20 hearts, transfer the hearts into a 15 mL conical tube with 8 mL of warm 0.05% trypsin-EDTA. If there are 20–30 hearts, transfer the hearts into a 50 mL conical tube with 15 mL of warm 0.05% trypsin-EDTA. 4. Gently aspirate the trypsin and add 5 mL (for less than 20 hearts) or 10 mL (for 20–30 hearts) of a warm type II collagenase (0.5 mg/mL) in HBSS. Vortex the tube for 1 min (if the liquid does not get up to the lid, then the speed is fine). Incubate the tube in a 37  C water bath for 7 min. 5. Vortex the tube again for 1 min and let the tissue settle down for 1 min. Collect the liquid into a new tube containing 10 mL of cold FB medium. 6. Repeat steps 4 and 5 four to five rounds until all tissue is digested. 7. Combine all the collections and filter through a 40 μm cell strainer to make a single-cell suspension. 8. Spin down at 200
g for 5 min, and then wash once with 10 mL of MACS buffer. 9. Resuspend cells in 10 mL of MACS buffer. 10. Optional: If there are a lot of blood cells, resuspend cells with 1 mL of an RBC lysis buffer (150 mM of NH4Cl, 10 mM of KHCO3, and 0.1 mM of EDTA), keep on ice for 1 min, then add 10 mL of MACS buffer, and centrifuge at 200
g for 5 min. Wash one more time with MACS buffer. 3.3.3 Isolation of Thy1.2+ Fibroblasts by Magnetic Activated Cell Sorting (MACS)

1. Count the viable cells using a trypan blue staining. Briefly, take 10 μL of cells out from 10 mL of the cell suspension described in Subheadings 3.3.1 and 3.3.2. Mix it with 10 μL of 0.4% trypan blue solution. Add the mixture to a hemacytometer. Allow it to stand for 3–5 min and then examine it immediately under a microscope. The dead cells are stained in blue and viable cells are unstained. Count all the viable cells in four 1 mm corner squares of the hemacytometer. The viable cell number will be determined by the average count per square
2 (dilution factor)
104
10 (total volume of cell suspension). 2. For less than 1
107 cells, resuspend them with 20 μL of Thy1.2 antibody in 80 μL of chilled MACS buffer. Add more antibody proportionally if there are more than 1
107 cells. Mix well and incubate in a refrigerator (2–8  C) for 30 min. 3. Add 10 mL of MACS buffer and spin samples at 200
g for 5 min.

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4. Add 20 μL of anti-biotin microbeads to 80 μL of MACS buffer and use the mixture to resuspend the cells. Incubate the cells with microbeads in a refrigerator (2–8  C) for 30 min. 5. Wash once with 10 mL of MACS buffer and centrifuge again. 6. Bring the volume to 2 mL in the MACS buffer. 7. Set up the MACS MultiStand and MidiMACS Separator in hood. Insert an LS column to the separator. Apply 3 mL of MACS buffer to the column to equilibrate it. 8. Pass the cell suspension through the equilibrated LS column. 9. Wash three times with 2 mL of MACS buffer each time. 10. Take the LS column off the separator, add 2 mL of MACS buffer to the column, and insert the plunge to flush out the bead-binding cells. Spin at 200
g for 5 min. 11. Resuspend the cells in 10 mL of FB media, count the cells, and seed into plates at proper density (see Note 9). 3.4 Cardiac Reprogramming In Vitro

1. Prepare 0.1% gelatin-coated 24-well plates. Seed the cells into plates at proper density. For MEF, the seeding cells’ density could be around 2–4
104 cells/well of a 24-well plate. For neonatal cardiac fibroblasts generated from the explant culture method, the seeding cell density could be around 2–3
104 cells/well of a 24-well plate. For fibroblasts generated from the enzyme digestion method, the cells’ density could be around 4–5
104 cells/well of a 24-well plate (see Note 10). Culture the cells overnight in the FB medium at 37  C. 2. On day 0, change the culture media to 0.5 mL of pre-warmed iCM media containing 4 μg/mL of polybrene. Add 10 μL of retrovirus to each well (see Note 11). Put the plate in a 37  C incubator for 24–48 h to perform the viral infection. 3. On day 2, change virus-containing media to 0.5 mL of regular iCM media. Change the media every 2–3 days. 4. On day 3, change the media to iCM media supplemented with 2 μg/mL of puromycin for positive selection of viral transduced cells. Keep it for 3 days. 5. On day 3, take the plate out of the incubator and place it under an inverted fluorescent microscope (20
) to observe dsRed expression. 6. On day 10, cells could be harvested for FACS analysis and ICC analysis to determine the reprogramming efficiency. 7. On day 14, replace the iCM media with B27 media. Change the media every 3 days. Spontaneous beating-cell loci may appear at 3 (cells from the enzyme digestion method) or 4 (cells from the explant culture method) weeks after viral transduction.

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1. Thoroughly wash the cells once with DPBS. Add 0.3 mL of 0.05% trypsin to each well and incubate at 37  C for 5 min. 2. Gently tap the plate to facilitate dissociating the cells. Check the cell detachment under the microscope. Add 1 mL of FACS buffer to each well once most cells are dissociated. 3. Transfer the cells to 2 mL of a 96-well plate with a V-shaped bottom. Spin at 200
g for 5 min to collect the cells. 4. Wash the cells once with the FACS buffer. 5. Resuspend the cells with 100 μL of fixation/permeabilization buffer and treat the cells for 20 min at 4  C. 6. Wash the cells twice with 500 μL of 1
perm/wash solution, pellet, and remove supernatant. 7. Prepare the primary antibody solution against GFP and cTnT in 1
perm/wash buffer. Thoroughly resuspend each sample with 50 μL of antibody solution and incubate at 4  C for 30 min. 8. Wash the cells once in 1
perm/wash buffer, pellet, and remove supernatant. 9. Add 50 μL of the secondary antibody solution containing Alexa Fluor 488-conjugated donkey anti-rabbit IgG and Alexa Fluor 647-conjugated donkey anti-mouse IgG to each sample. Incubate at 4  C for 30 min in the dark. 10. Wash the cells once in 1
perm/wash buffer, pellet, and remove supernatant. 11. Resuspend the cells in 300 μL of the fixation buffer (DPBS with 1% PFA). Transfer the cell into a Falcon tube with a cell strainer cap. Samples are now ready for FACS detection.

3.5.2 RT-qPCR Analysis to Test Gene Expression

1. Lyse the cells in 1 mL of TRIzol per well according to the manufacturer’s instructions, and transfer it into a 1.5 mL microcentrifuge tube (see Note 12). 2. Pipette the mixture up and down until no pellet is visible at the bottom of the tube (see Note 13). 3. Incubate the homogenized sample at RT for 5 min. 4. Add 0.2 mL of chloroform and cap the sample tubes securely. Shake the tubes vigorously by hand for 15 s and incubate at RT for 2–3 min. 5. Centrifuge the samples at no more than 12,000
g for 15 min at 4  C (now the mixture separates into three phases, the upper aqueous phase contains RNA). 6. Transfer the aqueous phase to a fresh tube and precipitate the RNA by mixing it with 0.5 mL of isopropyl alcohol.

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7. Incubate the samples at RT for 10 min and centrifuge at no more than 12,000
g for 10 min at 4  C. 8. Remove the supernatant, wash the RNA pellet with 1 mL of 75% ethanol, vortex the sample, and centrifuge at no more than 7500
g for 5 min at 4  C (keep it at 20  C until use). 9. Briefly dry the RNA pellet, dissolve the RNA in RNase-free water, and incubate for 10 min at 55–60  C. Quantify the RNA by Nanodrop spectrophotometer. 10. Prepare cDNA using a first-strand cDNA synthesis kit according to the manufacturer’s instructions. 11. Assess for gene expression using a SYBR Green PCR Master Mix together with designed primers. The genes involved in sarcomere structures (Actc1, TnnT2, Myh6, Myh7), ion channels (Pln, Slc8a1, Scn5a), and cell junctions (Gja1, Kcna5, Cacba1c) could be evaluated. Successful cardiac reprogramming shows significant upregulation of those genes and downregulation of genes represented as fibroblast markers including Col1a1 and Col3a1. GAPDH expression should be detected simultaneously as a housekeeping gene for normalization. 3.5.3 Immunocytochemical (ICC) Analysis of Reprogramming Efficiency

1. Rinse the cells with ice-cold PBS three times, and then remove the excess solution. 2. Add 0.5 mL of the ICC fixation buffer to each well of a 24-well plate. Fix the cells at RT for 15–20 min (see Note 14). 3. Rinse the cells with PBS three times. 4. Add 0.5 mL of the ICC permeabilization buffer to each well of a 24-well plate. Permeabilize the cells at RT for 15 min. 5. Rinse the cells with PBS three times. 6. Add 0.5 mL of the ICC blocking buffer to each well of a 24-well plate, and then incubate at RT for 1 h. 7. Dilute the primary antibodies with the ICC staining buffer. Add 200 μL of antibody solutions to each well and incubate overnight at 4  C. The antibody dilution factors are listed in Materials. 8. The next day, wash cells with PBS three times. 9. Add 200 μL of secondary antibody solutions to the cells and incubate for 1 h at RT in the dark. 10. Wash the cells with PBS three times. 11. Add 200 μL of Hoechst solution (1:5000) to each well for 40–60 s to stain the nuclei. 12. Aspirate the solution and wash the cells with PBS three times. 13. The samples are ready for imaging. The reprogrammed cells express cardiac marker cTnT and αActinin at day 14.

Murine iCM 3.5.4 Evaluation of Calcium Activity of iCMs at Week 4

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1. Calcium flux signals can be detected by using a Rhod-3 Calcium Imaging Kit according to the manufacturer’s instructions (see Note 15). 2. Reconstitute Rhod-3 in 100 μL of DMSO to yield a stock solution of 10 mM of Rhod-3. Store the stock solution at 20  C and protect from light in single-use aliquots. Reconstitute the probenecid in 1 mL of HBSS to prepare 250 mM of stock. 3. Wash the cells twice with HBSS buffer. 4. Prepare a fresh loading buffer containing Rhod-3 and probenecid. Add the loading buffer to the cells immediately and incubate the cells in the dark at RT for 30–60 min. 5. Wash the cells twice with HBSS. 6. Add 2 mL of the incubation buffer (2.5 mM of probenecid in HBSS) to the cells and incubate the cells in the dark at RT for 30–60 min. 7. Wash the cells once with HBSS. The cells are now ready for live-cell imaging.

4

Notes 1. Perform the following steps in a BSL2 Biological Safety Cabinet under sterile conditions. The proper disposal of transfected cells, pipette tips, and tubes is recommended to avoid the risk of environmental and health hazards. 2. Incubation should last at least 12 h. The retroviruses and precipitation solution mixture are stable for up to 2 weeks at 4  C. 3. Retroviruses should be used freshly. The titer of the virus should be the best 48 h after transfection of Plat-E cells. 4. A titration is recommended right before the virus infection because both excessive and insufficient viruses can reduce the reprogramming efficiency [27]. 5. The first two steps are performed under non-aseptic conditions. The following steps are carried out in a tissue culture hood under aseptic conditions and using sterile instruments. 6. The protocol outlined here uses neonatal mice. Animal care and experiments are performed in accordance with the guidelines established by the Division of Laboratory Animal Medicine (DLAM) at the University of North Carolina, Chapel Hill. 7. Keep the hearts in cold PBS and leave them on ice before fibroblast isolation. It is a critical step to keep the tissue cold

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to minimize cell death or senescence. All tissue should be used within 2 h. 8. Since the GFP expression is driven by the αMHC promoter, which is a cardiac specific promoter, the heart from a transgenic mouse should be in green, while the negative heart is quite dim. 9. Reprogramming efficiency relies on the freshness of cardiac fibroblasts. Fibroblasts can be used as early as 3 days after seeding as to the explant culture method, while fibroblasts from enzyme digestion method can be used as early as the next day of digestion. We do not recommend freezing or passaging the fibroblast for reprogramming, which will markedly decrease the reprogramming efficiency. 10. If shRNA viruses are used in the experiment, the number of seeding cells needs to increase to ensure the sufficient cells to be reprogrammed. 11. The amount of the virus should be proportionally increased based on the increased number of cells in different sizes of culture plates. 12. RNase-free reagents, consumables, and techniques are required for the RNA extraction. 13. Samples can be stored for up to 1 year at 80  C until ready for RNA extraction. 14. Fixed cells can be stored at 4  C for up to 1 month before use, but the fresh cells tend to stain better. 15. Rhod-3 staining has a high background, and the signal starts to decrease half an hour after the final washing step. Thus, we highly recommend imaging the cells within 1 h. References 1. Benjamin Emelia J, Muntner P, Alonso A, Bittencourt Marcio S, Callaway Clifton W, Carson April P, Chamberlain Alanna M, Chang Alexander R, Cheng S, Das Sandeep R, Delling Francesca N, Djousse L, Elkind Mitchell SV, Ferguson Jane F, Fornage M, Jordan Lori C, Khan Sadiya S, Kissela Brett M, Knutson Kristen L, Kwan Tak W, Lackland Daniel T, Lewis Tene´ T, Lichtman Judith H, Longenecker Chris T, Loop Matthew S, Lutsey Pamela L, Martin Seth S, Matsushita K, Moran Andrew E, Mussolino Michael E, O’Flaherty M, Pandey A, Perak Amanda M, Rosamond Wayne D, Roth Gregory A, Sampson Uchechukwu KA, Satou Gary M, Schroeder Emily B, Shah Svati H, Spartano Nicole L, Stokes A, Tirschwell David L, Tsao Connie W, Turakhia Mintu P, VanWagner LB, Wilkins

John T, Wong Sally S, Virani Salim S, American Heart Association Council on Epidemiology and Prevention Statistics Committee and Stroke Statistics Subcommittee (2019) Heart disease and stroke statistics—2019 update: a report from the American Heart Association. Circulation 139(10):e56–e528. https://doi. org/10.1161/CIR.0000000000000659 2. Laflamme MA, Murry CE (2011) Heart regeneration. Nature 473(7347):326–335. https:// doi.org/10.1038/nature10147 3. Quaini F, Urbanek K, Graiani G, Lagrasta C, Maestri R, Monica M, Boni A, Ferraro F, Delsignore R, Tasca G, Leri A, Kajstura J, Quaini E, Anversa P (2004) The regenerative potential of the human heart. Int J Cardiol 95 (Suppl 1):S26–S28

Murine iCM 4. Nag AC (1980) Study of non-muscle cells of the adult mammalian heart: a fine structural analysis and distribution. Cytobios 28 (109):41–61 5. Souders CA, Bowers SL, Baudino TA (2009) Cardiac fibroblast: the renaissance cell. Circ Res 105(12):1164–1176. https://doi.org/ 10.1161/circresaha.109.209809 6. Szabo E, Rampalli S, Risueno RM, Schnerch A, Mitchell R, Fiebig-Comyn A, LevadouxMartin M, Bhatia M (2010) Direct conversion of human fibroblasts to multilineage blood progenitors. Nature 468(7323):521–526. https://doi.org/10.1038/nature09591 7. Vierbuchen T, Ostermeier A, Pang ZP, Kokubu Y, Sudhof TC, Wernig M (2010) Direct conversion of fibroblasts to functional neurons by defined factors. Nature 463 (7284):1035–1041. https://doi.org/10. 1038/nature08797 8. Huang P, He Z, Ji S, Sun H, Xiang D, Liu C, Hu Y, Wang X, Hui L (2011) Induction of functional hepatocyte-like cells from mouse fibroblasts by defined factors. Nature 475 (7356):386–389. https://doi.org/10.1038/ nature10116 9. Sekiya S, Suzuki A (2011) Direct conversion of mouse fibroblasts to hepatocyte-like cells by defined factors. Nature 475(7356):390–393. https://doi.org/10.1038/nature10263 10. Ieda M, Fu JD, Delgado-Olguin P, Vedantham V, Hayashi Y, Bruneau BG, Srivastava D (2010) Direct reprogramming of fibroblasts into functional cardiomyocytes by defined factors. Cell 142(3):375–386. https://doi.org/10.1016/j.cell.2010.07.002 11. Qian L, Huang Y, Spencer CI, Foley A, Vedantham V, Liu L, Conway SJ, Fu JD, Srivastava D (2012) In vivo reprogramming of murine cardiac fibroblasts into induced cardiomyocytes. Nature 485(7400):593–598. https://doi.org/10.1038/nature11044 12. Jayawardena TM, Egemnazarov B, Finch EA, Zhang L, Payne JA, Pandya K, Zhang Z, Rosenberg P, Mirotsou M, Dzau VJ (2012) MicroRNA-mediated in vitro and in vivo direct reprogramming of cardiac fibroblasts to cardiomyocytes. Circ Res 110(11):1465–1473. https://doi.org/10.1161/circresaha.112. 269035 13. Protze S, Khattak S, Poulet C, Lindemann D, Tanaka EM, Ravens U (2012) A new approach to transcription factor screening for reprogramming of fibroblasts to cardiomyocyte-like cells. J Mol Cell Cardiol 53(3):323–332. https://doi.org/10.1016/j.yjmcc.2012.04. 010

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14. Song K, Nam YJ, Luo X, Qi X, Tan W, Huang GN, Acharya A, Smith CL, Tallquist MD, Neilson EG, Hill JA, Bassel-Duby R, Olson EN (2012) Heart repair by reprogramming non-myocytes with cardiac transcription factors. Nature 485(7400):599–604. https:// doi.org/10.1038/nature11139 15. Miyamoto K, Akiyama M, Tamura F, Isomi M, Yamakawa H, Sadahiro T, Muraoka N, Kojima H, Haginiwa S, Kurotsu S, Tani H, Wang L, Qian L, Inoue M, Ide Y, Kurokawa J, Yamamoto T, Seki T, Aeba R, Yamagishi H, Fukuda K, Ieda M (2018) Direct in vivo reprogramming with Sendai virus vectors improves cardiac function after myocardial infarction. Cell Stem Cell 22(1):91–103.e105. S1934-5909(17)30462-9 [pii]. https://doi. org/10.1016/j.stem.2017.11.010 16. Zhou H, Morales MG, Hashimoto H, Dickson ME, Song K, Ye W, Kim MS, Niederstrasser H, Wang Z, Chen B, Posner BA, Bassel-Duby R, Olson EN (2017) ZNF281 enhances cardiac reprogramming by modulating cardiac and inflammatory gene expression. Genes Dev 31 (17):1770–1783. https://doi.org/10.1101/ gad.305482.117. 31/17/1770 [pii] 17. Mohamed TM, Stone NR, Berry EC, Radzinsky E, Huang Y, Pratt K, Ang YS, Yu P, Wang H, Tang S, Magnitsky S, Ding S, Ivey KN, Srivastava D (2017) Chemical enhancement of in vitro and in vivo direct cardiac reprogramming. Circulation 135 (10):978–995. https://doi.org/10.1161/ CIRCULATIONAHA.116.024692 18. Liu Z, Wang L, Welch JD, Ma H, Zhou Y, Vaseghi HR, Yu S, Wall JB, Alimohamadi S, Zheng M, Yin C, Shen W, Prins JF, Liu J, Qian L (2017) Single-cell transcriptomics reconstructs fate conversion from fibroblast to cardiomyocyte. Nature 551(7678):100–104. https://doi.org/10.1038/nature24454 19. Abad M, Hashimoto H, Zhou H, Morales MG, Chen B, Bassel-Duby R, Olson EN (2017) Notch inhibition enhances cardiac reprogramming by increasing MEF2C transcriptional activity. Stem Cell Reports 8(3):548–560. https://doi.org/10.1016/j.stemcr.2017.01. 025 20. Zhou Y, Wang L, Vaseghi HR, Liu Z, Lu R, Alimohamadi S, Yin C, Fu JD, Wang GG, Liu J, Qian L (2016) Bmi1 is a key epigenetic barrier to direct cardiac reprogramming. Cell Stem Cell 18(3):382–395. https://doi.org/ 10.1016/j.stem.2016.02.003 21. Singh VP, Mathison M, Patel V, Sanagasetti D, Gibson BW, Yang J, Rosengart TK (2016) MiR-590 promotes transdifferentiation of

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porcine and human fibroblasts toward a cardiomyocyte-like fate by directly repressing specificity protein 1. J Am Heart Assoc 5(11): e003922. https://doi.org/10.1161/JAHA. 116.003922 22. Liu L, Lei I, Karatas H, Li Y, Wang L, Gnatovskiy L, Dou Y, Wang S, Qian L, Wang Z (2016) Targeting Mll1 H3K4 methyltransferase activity to guide cardiac lineage specific reprogramming of fibroblasts. Cell Discov 2:16036. https://doi.org/10.1038/celldisc. 2016.36 23. Zhou H, Dickson ME, Kim MS, BasselDuby R, Olson EN (2015) Akt1/protein kinase B enhances transcriptional reprogramming of fibroblasts to functional cardiomyocytes. Proc Natl Acad Sci U S A 112 (38):11864–11869. https://doi.org/10. 1073/pnas.1516237112 24. Zhao Y, Londono P, Cao Y, Sharpe EJ, Proenza C, O’Rourke R, Jones KL, Jeong MY, Walker LA, Buttrick PM, McKinsey TA,

Song K (2015) High-efficiency reprogramming of fibroblasts into cardiomyocytes requires suppression of pro-fibrotic signalling. Nat Commun 6:8243. https://doi.org/10. 1038/ncomms9243 25. Wang L, Liu Z, Yin C, Asfour H, Chen O, Li Y, Bursac N, Liu J, Qian L (2015) Stoichiometry of Gata4, Mef2c, and Tbx5 influences the efficiency and quality of induced cardiac myocyte reprogramming. Circ Res 116(2):237–244. https://doi.org/10.1161/circresaha.116. 305547 26. Ma H, Wang L, Yin C, Liu J, Qian L (2015) In vivo cardiac reprogramming using an optimal single polycistronic construct. Cardiovasc Res 108(2):217–219. https://doi.org/10.1093/ cvr/cvv223 27. Qian L, Berry EC, Fu JD, Ieda M, Srivastava D (2013) Reprogramming of mouse fibroblasts into cardiomyocyte-like cells in vitro. Nat Protoc 8(6):1204–1215. https://doi.org/10. 1038/nprot.2013.067

Chapter 13 Frame-Hydrogel Methodology for Engineering Highly Functional Cardiac Tissue Constructs Abbigail Helfer and Nenad Bursac Abstract Engineered cardiac tissues hold tremendous promise for in vitro drug discovery, studies of heart development and disease, and therapeutic applications. Here, we describe a versatile “frame-hydrogel” methodology to generate engineered cardiac tissues with highly mature functional properties. This methodology has been successfully utilized with a variety of cell sources (neonatal rat ventricular myocytes, human and mouse pluripotent stem cell-derived cardiomyocytes) to generate tissues with diverse 3D geometries (patch, bundle, network) and levels of structural and functional anisotropy. Maturation of such engineered cardiac tissues is rapidly achieved without the need for exogenous electrical or mechanical stimulation or use of complex bioreactors, with tissues routinely reaching conduction velocities and specific forces of 25 cm/ s and 20 mN/mm2, respectively, and forces per input cardiomyocyte of up to 12 nN. This method is reproducible and readily scalable to generate small tissues ideal for in vitro testing as well as tissues with large, clinically relevant dimensions. Key words Engineered cardiac tissues, Human pluripotent stem cells, Tissue engineering, Cardiac patch, Cardiac bundle, Cardiomyocytes, Hydrogel

1

Introduction Ischemic heart disease is a leading cause of death worldwide, resulting in approximately 17.7 million deaths per year [1]. In order to facilitate studies of cardiovascular disease, drug discovery, and regenerative therapies, extensive work has gone into mimicking myocardium in vitro [2–7]. Two-dimensional (2D) cultures of cardiomyocytes can provide valuable electrophysiological and structural information, as well as drug responses, but do not fully recapitulate the in vivo environment. Furthermore, cardiomyocytes cultured in 2D fail to exhibit mature structural characteristics and electromechanical function [8]. To combat these limitations, various methodologies for 3D cardiomyocyte culture have been developed. Early work relied on the use of neonatal rat ventricular myocytes (NRVMs) to generate 3D cardiac tissues due to their

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_13, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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wide availability and ease of culture [9–13]. More recent advances in cardiac differentiation protocols for human induced pluripotent and embryonic stem cells (iPSCs and ESCs) have enabled generation of functional human cardiac tissues [8, 14, 15]. Here, we describe a “frame-hydrogel” methodology, whereby cardiomyocytes are cast in a fibrin hydrogel and anchored to a flexible nylon frame to form highly functional 3D engineered cardiac tissues [2, 16–23]. Such formed cardiac tissues are subsequently cultured in a low-shear, dynamic culture environment to rapidly mature heart cells without the need for external electrical or mechanical stimulation or use of complex bioreactors [18]. After 2–3 weeks of culture, cardiac tissues generated using this methodology exhibit physiologically high cell density and electromechanical properties approximating those of adult myocardium. We have successfully utilized this method with both rodent (neonatal rat ventricular and mouse ESC-derived) and human (iPSC- and ESC-derived) cardiomyocytes (CMs), as well as mouse ESC- and iPSC-derived cardiovascular progenitors (CVPs) [2, 17–22]. The frame-hydrogel casting method also allows for the generation of a wide variety of tissue architectures and sizes while consistently yielding highly advanced levels of CM maturity. Specifically, we have applied this methodology to generate isotropic tissue patches with random CM orientations, network patches with staggered elliptical pores to guide local CM alignment and allow control of tissue anisotropy, epicardial-mimetic network patches with locally varying CM alignment replicating realistic fiber orientations of human myocardium determined with cardiac MRI, and cylindrical bundles with highly aligned CMs. The aforementioned tissues have dimensions that range from 7
7 to 36
36 mm in surface area.

2

Materials

2.1 Manufacturing of Tissue Molds

1. Polytetrafluoroethylene (PTFE). 2. Computer numerical control (CNC) router. 3. Laser cutter. 4. SYLGARD™ 184 Silicone (polydimethylsiloxane, PDMS).

Elastomer

Kit

5. Petri dishes, non-treated. 6. Oven set to 60  C. 7. Cerex® nylon material (Cerex® Advanced Fabrics Inc) (see Note 1). 8. 70% Ethanol solution. 9. Ultrasonic bath. 10. Razor blades.

Engineering Cardiac Tissue Constructs

2.2 3D Engineered Tissue Culture Media and Cell Sources 2.3 3D Tissue Generation

173

1. A list of cell types, sources, and 3D engineered tissue culture media is provided in Table 1.

All components should be prepared in a sterile fashion and filtered using a 0.22 μm filter before use. 1. Pluronic F-127: 0.2% (w/v) in deionized H2O. 2. Molecular biology-grade water. 3. Dulbecco’s phosphate-buffered saline (DPBS) without CaCl2 and MgCl2. 4. Bovine thrombin: 50 U/mL in 0.1% (w/v) BSA solution. 5. Human or bovine fibrinogen: 10 mg/mL in DPBS, dissolved at 37  C and stored on ice (see Note 2). 6. 2X Media: 2X concentrated low-glucose DMEM (reconstituted from powder at half recommended volume), 20% heatinactivated horse serum, 10 U/mL penicillin, 4 μg/mL B-12, 2 mg/mL aminocaproic acid. 7. Matrigel matrix. 8. Incubator set to 37  C, 5% CO2. 9. Water bath set to 37  C. 10. High-angle rocker set to 30 tilt and 0.4 Hz rocking frequency. 11. Hemocytometer. 12. Trypan blue 0.4%. 13. Standard forceps, curved. 14. Fine-point forceps, straight. 15. 10 cm culture dishes, non-treated. 16. 6 cm culture dishes, non-treated. 17. 12-well or 6-well culture plates, non-treated.

3

Methods

3.1 Manufacturing of PDMS Tissue Molds

1. Design a master negative using 3D design software (see Note 3). We have generated many different sizes and geometries for masters (Fig. 1a) that can be used to generate cardiac bundles and patches. 2. Machine polytetrafluoroethylene (PTFE, Teflon) into negative master shape using a CNC router (see Note 4). 3. Prepare the PDMS by thoroughly mixing SYLGARD™ 184 Silicone Elastomer base and SYLGARD™ 184 Silicone Elastomer curing solution in a 10:1 ratio (w:w).

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Table 1 Cardiogenic cell types, sources, and culture media used for maintenance of engineered cardiac tissues generated by the frame-hydrogel method Cell type

Cell source

Culture media

Differentiated from monolayers of Days 0–7 of culture: Human induced pluripotent human iPSCs using smallRPMI-1640, 2% B-27 stem cell-derived molecule modulation of the supplement, 2 mg/mL cardiomyocytes (hiPSC-CMs) Wnt pathway [24]. Purified aminocaproic acid, 50 ug/mL using metabolic selection [25]. ascorbic acid 2-phosphate, 1% penicillin-streptomycin, 1% nonessential amino acids, 1% sodium pyruvate, 0.45 μM 1-thioglycerol Days 7–21 of culture: Low-glucose DMEM, 5% fetal bovine serum, 2 mg/mL aminocaproic acid, 50 ug/mL ascorbic acid 2-phosphate, 1% penicillin-streptomycin, 1% nonessential amino acids, 0.45 μM 1-thioglycerol [2] Human embryonic stem cellderived cardiomyocytes (hESC-CMs)

DMEM, 5% fetal bovine serum, Differentiated from embryoid 1 mg/mL aminocaproic acid, bodies of human ESCs using 50 μg/mL ascorbic acid previously published protocols 2-phosphate, 1 mM sodium [26]. Purified using magnetic pyruvate, 2 mM glutamine, activated cell sorting for SIRPA0.1 mM nonessential amino positive cells [26]. acids, 0.45 mM 1-thioglycerol [20]

Neonatal rat ventricular myocytes (NRVMs)

Low-glucose DMEM, 10% horse Enzymatically digested from the serum, 1% chick embryo ventricles of 2-day-old Spragueextract, 1 mg/mL Dawley rats. Purified using two aminocaproic acid, 50 μg/mL pre-plating steps [11]. ascorbic acid 2-phosphate, 5 U/mL penicillin, 2 μg/mL vitamin B-12 [19]

Mouse induced pluripotent stem cell-derived cardiovascular progenitors (miPSC-CVPs)

1:1 ratio of DMEM/F12 and Differentiated from embryoid neurobasal base medium, 1
bodies of mouse iPSCs using N2 supplement, 1
B-27 published methods supplement, Glutamax-I, [22]. miPSCs stably transfected nonessential amino acids, with puromycin N-acetyl β-mercaptoethanol, 5 mg/mL transferase under the control of BSA, gentamicin, 1 mg/mL a Nkx2–5 cardiac-specific aminocaproic acid [22] enhancer element to allow for purification of progenitor cells by puromycin selection [22].

Mouse embryonic stem cellderived cardiomyocytes and cardiovascular progenitors

Differentiated from embryoid bodies of mouse ESCs using published protocols

1:1 ratio of DMEM/F12 and neurobasal base medium, 1% L-glutamine, 0.1% gentamicin (continued)

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Table 1 (continued) Cell type

Cell source

(mESC-CMs and mESCCVPs)

[21, 27]. mESCs stably transfected with puromycin aminotransferase under the control of a mouse Myh6 promotor or a Nkx2.-5 enhancer element to allow for purification by puromycin selection [21].

Culture media reagent solution, 1% B-27 supplement, 0.0125 mg/mL insulin, 0.05 mg/mL apo-transferrin, 2.5 ng/mL sodium selenite, 3 ng/mL progesterone, 8 ug/mL putrescine, 0.0334% BSA fraction V, 1 mg/mL aminocaproic acid [21]

4. Transfer the PDMS to a vacuum desiccator and degas for 30 min until all bubbles are removed. 5. Place the PTFE master(s) face up in a petri dish and pour the PDMS over the master until it is completely submerged. There should be ~2 mm of PDMS above the highest point of the master. 6. Transfer the dish containing the PDMS and master to a vacuum desiccator and degas for 30 min, ensuring that all bubbles are evacuated. Remaining bubbles may be removed with a needle or pipette tip. 7. Cure the PDMS in a 60  C oven for at least 4 h, ensuring that the dish is level. 8. After the curing is complete, remove the PDMS from the dish and carefully extract the PTFE master (see Note 5). 9. Remove the excess PDMS that surrounds the mold using a sharp razor blade to produce a final PDMS mold (Fig. 1c, d). 10. Submerge the PDMS molds in 70% ethanol and sonicate in an ultrasonic bath for 60 min at 60  C. 11. Remove the PDMS molds from 70% ethanol, allow them to dry at room temperature, and then steam autoclave to sterilize (see Note 6). 3.2 Frame Manufacturing

1. Design a frame template using 2D drawing software that is compatible with the laser cutter. The frame should surround the entire perimeter of the final tissue (Fig. 1b, e–g). Frames typically have edges that are, at minimum, 1 mm thick and dimensions slightly smaller than the mold interior in which they will be press-fitted. Frames are connected to the main sheet of the Cerex® nylon material by a thin bridge of uncut material (Fig. 1b).

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Fig. 1 (a) PTFE master negatives used to manufacture PDMS molds for generating a 7
2 mm bundle, a 7
7 mm patch, a 10
10 mm patch with border pores, and a 36
36 mm patch. Scale bar ¼ 10 mm. (b) Left: Laser cutter path to generate frames within a nylon sheet. Middle: Laser-cut frames attached to sheet with thin bridges (dashed lines). Right: Nylon frames cut from the main sheet at the bridges. Scale bar ¼ 5 mm. (c) Resulting PDMS molds generated

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2. Laser-cut frames out of nylon material, ensuring that the laser settings do not burn the Cerex® nylon material. 3. Carefully separate individual frames by cutting the thin bridges that are holding the frames to the larger sheet of nylon material (Fig. 1b). 4. Sterilize frames using UV or gas sterilization. 3.3 Initial Mold Preparation for Tissue Culture

All steps should be performed in a BSL-2 biosafety cabinet. 1. Using sterile curved forceps, place the autoclaved PDMS molds into a sterile 10 cm petri dish. Leave at least 3 mm of space between molds to facilitate eventual mold removal. A 10 cm petri dish can accommodate up to twelve 10
10 mm (length
width) molds, three 18
18 mm molds, or one 41
41 mm mold. 2. To decrease cell attachment, increase the hydrophilicity of the PDMS molds by filling each mold with Pluronic F-127 solution until the internal surface is fully covered, approximately 250 μL for a 10
10
1.5 mm mold (length
width
depth). Remove any air bubbles using a sterile pipette tip. 3. Allow Pluronic F-127-filled molds to sit undisturbed at room temperature for 1–2 h. Proceed to cell preparation after 1–2 h has passed. 1. Prepare the cardiac cells in suspension using previously published methods (Table 1) (see Note 7).

Cell Preparation

2. Resuspend the cells in recommended culture media (see Table 1) and count the number of live cells using a hemocytometer and trypan blue. 3. Centrifuge cells at 300
g for 5 min. 4. Determine the required live cell concentration using the following formula, where V is equal to the volume of cell/hydrogel mixture required for one engineered tissue (in mL): Cell concentration ¼

Cells per tissue cells=mL 0:48
V

For a 7
7 mm patch, V is equal to 120 μL and the number of cells per tissue is 500,000. Therefore, the required ä

3.4

Fig. 1 (continued) from the masters shown in (a) with nylon frames pressed in. Scale bar ¼ 10 mm. (d) PDMS molds for fabrication of a 7
7 mm and a 14
14 mm network patch and a 25
25 mm epicardial mimetic patch. These molds were generated using high-aspect-ratio soft lithography masters. Scale bar ¼ 10 mm. Appearance of 7
7 mm (e) bundle, (f) patch, and (g) networkengineered cardiac tissues after 2–3 weeks of culture. Scale bar ¼ 1 mm (figure contains elements reproduced from refs. 2, 16, and 18 with permission)

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cell concentration is 8.68
106 cells/mL. For a 7
2 mm bundle, V is equal to 37.5 μL and the number of cells per tissue is 375,000. Therefore, the required cell concentration is 20.8
106 cells/mL. 5. Resuspend the cells in culture media at the calculated concentration and store on ice. 6. Immediately proceed to final mold preparation and hydrogel preparation. Cells should be kept as a single-cell suspension on ice for a minimal duration of time. 1. Immediately after the cardiomyocytes are prepared, aspirate the Pluronic F-127 from the molds, taking care not to touch the molds with the aspirating tip.

3.5 Final Mold Preparation

2. Rinse the molds by filling each mold with molecular biologygrade water until the internal surface is covered. 3. Aspirate the water from the molds, drying the molds as much as possible while ensuring that the aspirating tip does not touch or scrape the mold surface. 4. Allow the molds to air-dry in the BSL-2 biosafety cabinet with the petri dish lids removed to improve airflow and evaporation. This step can occur while the hydrogel is prepared and aliquoted. Molds with more complex architecture should be allowed a longer period of time to dry. 5. Proceed immediately to hydrogel preparation. All solutions should be prepared and kept on ice in a BSL-2 biosafety cabinet.

3.6 Hydrogel Preparation

1. Place stock solutions of fibrinogen, Matrigel, thrombin, and 2X media on ice (see Note 8). Table 2 Hydrogel composition for generation of engineered cardiac tissues using the frame-hydrogel method. V is defined as the volume of cell/hydrogel mixture required for one engineered tissue (in mL) and Ntissues is defined as the number of tissues to make consecutively Starting concentration

Percent of final Final concentration hydrogel volume

Volume to prepare

Fibrinogen

10 mg/mL

2 mg/mL

20%

0.2
V
Ntissues

Matrigel

100%

10% (v/v)

10%

0.1
V
Ntissues

Thrombin

50 U/mL

1 U/mL

2%

0.02
V
Ntissues

20%

0.2
V
Ntissues

2X Media

100%

20%

Cells suspended in culture media

Cells per tissue 0:48
V

Cells per tissue V

cells=mL

cells=mL 48%

0.48
V
Ntissues

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2. Use Table 2 to calculate volumes required for each component (see Note 9). Define V as the volume of cell/hydrogel mixture required for one engineered tissue. For a patch with dimensions of 7
7 mm (length
width), V is equal to 120 μL. V can be scaled in proportion to the area of the tissue. Define Ntissues as the number of tissues to make consecutively. The number of tissues that can be cast consecutively (Ntissues) is limited by the rapid cross-linking of the fibrinogen by thrombin. Therefore, tissues must be generated in batches. We have found that the maximum Ntissues is 4. This value should be reduced for larger or more complex molds. 3. Using the volumes calculated in step 2, combine the fibrinogen, Matrigel, and cell solutions in an Eppendorf tube on ice, mixing thoroughly but carefully. 4. In a separate Eppendorf tube on ice, combine the 2X media and thrombin solutions using the volumes calculated in step 2. 5. If the number of needed engineered tissues is larger than Ntissues, repeat steps 3 and 4 until there are a sufficient number of Eppendorf tubes to generate the desired number of tissues. For example, if 12 total engineered cardiac tissues are to be generated, and Ntissues is 4, prepare three Eppendorf tubes containing the fibrinogen, Matrigel, and cell solutions and three Eppendorf tubes containing the 2X media and thrombin solutions. 6. Proceed to cast the engineered tissues in PDMS molds, limiting the amount of time between hydrogel preparation and tissue casting. 3.7 Casting of Engineered Tissues in PDMS Molds

1. Using sterile fine point forceps, place one sterile nylon frame into each PDMS mold, ensuring that the mold is entirely dry and that no water is wicking into the frame (see Note 10) (Fig. 1c). The frame should be in contact with the base of the PDMS mold along the entire perimeter of the frame. 2. To initiate polymerization and form the final hydrogel/cell mixture, add the contents of the 2X media/thrombin Eppendorf tube to the fibrinogen/Matrigel/cell Eppendorf tube and pipette up and down to mix, taking care not to introduce any bubbles. 3. Quickly and carefully pipette one volume’s worth (V) of the hydrogel/cell mixture into one PDMS mold containing a frame. Do not touch or scrape the PDMS mold with the pipette tip and avoid introducing bubbles into the hydrogel. The hydrogel should completely cover the bottom surface of the mold and wick into the nylon frame (see Note 11). 4. Repeat step 3 without changing pipette tips until hydrogel mixture is fully distributed into Ntissues molds (see Note 12).

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5. Repeat steps 2–4, making tissues in batches until the desired total number of molds are filled. 6. Place the 10 cm petri dish containing the hydrogel-filled molds into a 37  C 5% CO2 incubator for 30–45 min to allow for complete gelling of the hydrogel. 7. After 30–45 min, return the 10 cm petri dish containing the hydrogel-filled molds to the biosafety cabinet. Place each hydrogel-filled mold into a well of a tissue culture plate. 10
10 mm molds fit in the wells of a 12-well plate and 18
18 mm molds fit in the wells of a 6-well plate. To prevent the molds from floating, press the PDMS mold firmly to the bottom of dry plates. 8. Add culture media to each well, ensuring that the mold is fully submerged. 2 mL of media is sufficient to cover the 10 mm
10 mm molds in a well of a 12-well culture plate and 5 mL of media is sufficient to cover the 18 mm
18 mm molds in a well of a 6-well culture plate. 9. Place the tissue culture plate onto a high-angle rocker set to 30 tilt and 0.4 Hz rocking frequency in a 37  C 5% CO2 incubator for 24 h (see Note 13). The molds should not be free floating at this stage. 10. 24 h after the tissues were cast, proceed to removing the engineered tissues from the molds. 3.8 Removing Engineered Tissues from Molds

All steps should be performed in a BSL-2 biosafety cabinet. 1. Fill a 6 cm culture dish with 20 mL of DPBS or to a depth that will fully submerge the molds. 2. Remove the tissues from the high-angle rocker and transfer them to the biosafety cabinet. 3. Using a pair of sterile curved forceps, remove one hydrogelfilled mold from the culture plate and submerge it in the DPBS-filled dish. 4. Hold and stabilize the sides of the mold using a pair of curved forceps. Use another pair of fine-point forceps to remove the engineered tissue from the mold by gently sliding the finepoint forceps under the nylon frame and lifting the frame upward. Continue gently lifting the frame at several points along one edge of the mold. When the center of the hydrogel (the area not supported by the frame) has separated from the PDMS mold, repeat the lifting technique on each of the remaining three sides until entire tissue is separated from the PDMS mold and is free floating in the DPBS bath. Only touch the frame with the forceps—touching the hydrogel with forceps will damage the tissue (see Note 14).

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5. Return the engineered tissue to the cell culture plate, holding the frame with fine-point forceps. The tissue should sink to the bottom of the well. 6. Repeat steps 3–5 until all tissues are removed from molds and placed in culture plates (see Note 15). 7. Return the culture dish to a high-angle rocker in a 37  C 5% CO2 incubator. Tissues should be free floating in the culture media. 3.9

Tissue Culture

1. Replace the culture media (Table 1) every other day. (a) Remove 2/3 of the culture media with an aspirating pipette while keeping the culture plate flat. Take care not to place the aspirating pipette too close to the tissue. (b) Replace the removed cell culture media with new media, adding the new media away from the engineered tissue to avoid damaging it. 4. Culture the tissues on the high-angle rocker in a 37  C 5% CO2 incubator for 2–3 weeks. 5. Analyze the tissues for force generation, electrical propagation, and structure 2–3 weeks after initial tissue formation. Expected functional values for tissues generated from various cell sources and with different geometries are provided in Table 3 and Fig. 2.

4

Notes 1. We have also utilized standard Velcro® as a frame material. 2. Fibrinogen should be prepared less than 4 h before use and kept on ice. It is recommended to limit the amount of time the fibrinogen is in the 37  C water bath to 10 min or until the solids have just dissolved. Sterile filtration should occur when the solution is still warm, and then placed on ice. Ensure that the solution is fully cooled to 4  C before use. 3. It is recommended to first determine the desired final shape and size of tissues, subsequently design a tissue mold using 3D software, and then fabricate the master for tissue mold generation. 4. Some geometries, for example, small network patches, may have features that are smaller than can be achieved with a CNC router. In order to form these intricate molds, we recommend using high-aspect-ratio soft lithography, as has been previously described in detail [16]. Soft lithography will allow for the generation of a PDMS master that can be used in the same manner as the PFTE master described here.

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Table 3 Average functional outputs achieved by 2–3-week-old engineered cardiac tissues generated using the frame-hydrogel method. Results of functional measurements are presented for various cell types and tissue geometries achieved during the last decade

Cell type

Geometry

Max active force (mN)

Specific force (mN/mm2)

Force per input CM (nN)

Conduction velocity (cm/s)

Citation

hiPSCCMs

7
7 mm patch

5.2  0.2 22.4  0.9

11.9  0.5 28.5  1.0

Shadrin et al. (2017) [2]

hiPSCCMs

15
15 mm patch (megapatch)

9.4  1.0 19.4  2.1

4.7  0.5 27.2  1.1

Shadrin et al. (2017) [2]

hiPSCCMs

36
36 mm patch (gigapatch)

17.5  1.1 17.0  0.8

1.75  0.1 28.9  1.8

Shadrin et al. (2017) [2]

18.0  1.4 18.0  1.4

3.0  0.2 32.3  1.8

Jackman et al. (2018) [19]

NRVMs 7
2 mm bundle

2.10  0.10 59.7  4.3

5.6  0.27 52.5  0.9

Jackman et al. (2016) [18]

7
2 mm bundle

1.3  0.05 23.2  1.6

3.47  0.14 25.8  1.2

Jackman et al. (2016) [18]

4.35  0.45 26.8  0.8

Bian et al. (2014) [17]

NRVMs 10
10 mm patch with border pores

hiPSCCMs

NRVMs 7
7 mm network 2.39  0.25 8.9  1.1 patch, 1.2 mm pore length 3.0  1.1 11.8  4.5

5.7  1.1 25.1

hESCCMs

7
7 mm network patch, 1.2 mm pore length

mESCCMs

1.96  0.54 Not 7
7 mm network reported patch, 1.2 mm pore length

2.51  0.69 24.1  1.4

Liau et al. (2011) [21]

mESC- 7
7 mm network 1.28  0.11 Not CVPs patch, 1.2 mm pore reported length

1.64  0.14 19.2  0.4

Liau et al. (2011) [21]

Zhang et al. (2013) [20]

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Fig. 2 Functional properties of engineered cardiac tissues. Representative immunohistochemistry images of (a) a 7
7 mm hiPSC-CM patch, and (b) its cross section, (c) a 7
2 mm NRVM bundle and (d) its cross section, and (e) a 7
7 mm hESC-CM network patch. Representative isochrone activation maps from (f) a 7
7 mm hiPSC-CM patch, (g) a 7
2 mm NRVM bundle, and (h) a 7
7 mm mESC-CM network patch. (i) Representative force traces during stretch of a 1 Hz electrically stimulated 7
7 mm hESC-CM cardiac network patch. (j) Active and passive force-length relationships for 7
7 mm hESC-CM cardiac network patches (figure contains elements reproduced from refs. 2, 18, 21, and 20 with permission) (SAA sarcomeric alpha-actinin, Vim vimentin)

5. If the PDMS is difficult to remove from the petri dish, use a pair of cutting pliers to fracture the side of the petri dish. Carefully remove fragments of the petri dish until the PDMS can be extracted. 6. PDMS molds can be generated weeks to months in advance and stored in sterile autoclaving pouches until needed. 7. It is recommended to begin with a cell population that contains approximately 10% fibroblasts. If necessary, fibroblasts can be separately cultured and added to highly pure cardiomyocyte populations. 8. Thrombin, Matrigel, and 2X media stock solutions may be prepared in advance, aliquoted, and stored at 20  C. These components should be thawed on ice the day of tissue generation, ensuring that each solution is fully thawed before beginning cell collection. 9. There is some volume loss during the hydrogel generation procedure. Therefore, it is recommended to prepare a minimum of 105% of the total calculated volumes of each solution. 10. The plate lid will attract the frames through static electricity. Avoid placing the lid back on the plate once frames are pressfitted into the molds.

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11. Due to the surface tension of the hydrogel, it can be difficult to fully spread the small volume of hydrogel over the entire surface of the mold. We recommend pipetting from the walls of the mold toward the center, instead of ejecting the hydrogel into the middle of the mold. This will help the hydrogel wick up the side walls of the mold. 12. If the hydrogel begins to take on a viscous appearance while being injected into the molds, the hydrogel has begun to polymerize. The number of tissues made at one time (Ntissues) should be decreased. 13. If the media spills out of the culture plate or splashes onto the lid while the culture plate is on the high-angle rocker, reduce the media volume. If the mold cannot be completely submerged with a smaller volume of media, a shorter mold is recommended. If the mold exceeds the dimensions of a standard tissue culture dish (for example, the 41
41 mm patch molds), a custom-built chamber may be required [2]. 14. If the hydrogel is adhering to the PDMS mold, the Pluronic F-127 solution can be used at a higher concentration or applied to the molds for a longer period of time. Alternatively, tissues can be removed from the PDMS molds earlier than 24 h after formation. 15. PDMS molds can be reused after tissues are removed. Follow steps 9 and 10 in Subheading 3.1 to clean and prepare the molds for reuse.

Acknowledgments This work was supported by NIH grants U01HL134764, UG3TR002142, HL132389, and HL126524, and a grant from Foundation Leducq to NB and an NSF Graduate Research Fellowship (2017–2020) to AH. References 1. Thomas H, Diamond J, Vieco A, Chaudhuri S, Shinnar E, Cromer S, Perel P, Mensah GA, Narula J, Johnson CO, Roth GA, Moran AE (2018) Global atlas of cardiovascular disease 2000–2016: the path to prevention and control. Glob Heart 13(3):143–163. https://doi. org/10.1016/j.gheart.2018.09.511 2. Shadrin IY, Allen BW, Qian Y, Jackman CP, Carlson AL, Juhas ME, Bursac N (2017) Cardiopatch platform enables maturation and scale-up of human pluripotent stem cellderived engineered heart tissues. Nat Commun

8(1):1825. https://doi.org/10.1038/ s41467-017-01946-x 3. Lundy SD, Zhu WZ, Regnier M, Laflamme MA (2013) Structural and functional maturation of cardiomyocytes derived from human pluripotent stem cells. Stem Cells Dev 22 (14):1991–2002. https://doi.org/10.1089/ scd.2012.0490 4. Nunes SS, Miklas JW, Liu J, Aschar-Sobbi R, Xiao Y, Zhang B, Jiang J, Masse S, Gagliardi M, Hsieh A, Thavandiran N, Laflamme MA, Nanthakumar K, Gross GJ, Backx PH,

Engineering Cardiac Tissue Constructs Keller G, Radisic M (2013) Biowire: a platform for maturation of human pluripotent stem cellderived cardiomyocytes. Nat Methods 10 (8):781–787. https://doi.org/10.1038/ nmeth.2524 5. Mannhardt I, Breckwoldt K, Letuffe-BreniereD, Schaaf S, Schulz H, Neuber C, Benzin A, Werner T, Eder A, Schulze T, Klampe B, Christ T, Hirt MN, Huebner N, Moretti A, Eschenhagen T, Hansen A (2016) Human engineered heart tissue: analysis of contractile force. Stem Cell Reports 7(1):29–42. https:// doi.org/10.1016/j.stemcr.2016.04.011 6. Mills RJ, Titmarsh DM, Koenig X, Parker BL, Ryall JG, Quaife-Ryan GA, Voges HK, Hodson MP, Ferguson C, Drowley L, Plowright AT, Needham EJ, Wang QD, Gregorevic P, Xin M, Thomas WG, Parton RG, Nielsen LK, Launikonis BS, James DE, Elliott DA, Porrello ER, Hudson JE (2017) Functional screening in human cardiac organoids reveals a metabolic mechanism for cardiomyocyte cell cycle arrest. Proc Natl Acad Sci U S A. https://doi.org/10. 1073/pnas.1707316114 7. Ronaldson-Bouchard K, Ma SP, Yeager K, Chen T, Song L, Sirabella D, Morikawa K, Teles D, Yazawa M, Vunjak-Novakovic G (2018) Advanced maturation of human cardiac tissue grown from pluripotent stem cells. Nature 556(7700):239–243. https://doi. org/10.1038/s41586-018-0016-3 8. Pomeroy JE, Helfer A, Bursac N (2019) Biomaterializing the promise of cardiac tissue engineering. Biotechnol Adv. https://doi. org/10.1016/j.biotechadv.2019.02.009 9. Zimmermann WH, Fink C, Kralisch D, Remmers U, Weil J, Eschenhagen T (2000) Three-dimensional engineered heart tissue from neonatal rat cardiac myocytes. Biotechnol Bioeng 68(1):106–114 10. Zimmermann WH, Schneiderbanger K, Schubert P, Didie M, Munzel F, Heubach JF, Kostin S, Neuhuber WL, Eschenhagen T (2002) Tissue engineering of a differentiated cardiac muscle construct. Circ Res 90 (2):223–230 11. Bursac N, Papadaki M, Cohen RJ, Schoen FJ, Eisenberg SR, Carrier R, Vunjak-Novakovic G, Freed LE (1999) Cardiac muscle tissue engineering: toward an in vitro model for electrophysiological studies. Am J Phys 277(2): H433–H444. https://doi.org/10.1152/ ajpheart.1999.277.2.H433 12. Papadaki M, Bursac N, Langer R, Merok J, Vunjak-Novakovic G, Freed LE (2001) Tissue engineering of functional cardiac muscle: molecular, structural, and electrophysiological studies. Am J Phys Heart Circ Phys 280(1):

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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 26. 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. https://doi.org/ 10.1038/nbt.2005 27. Christoforou N, Miller RA, Hill CM, Jie CC, McCallion AS, Gearhart JD (2008) Mouse ES cell-derived cardiac precursor cells are multipotent and facilitate identification of novel cardiac genes. J Clin Invest 118(3):894–903. https://doi.org/10.1172/jci33942

Chapter 14 Efficient Protocols for Fabricating a Large Human Cardiac Muscle Patch from Human Induced Pluripotent Stem Cells Ling Gao and Jianyi (Jay) Zhang Abstract Human induced pluripotent stem cells (hiPSCs) are among the most promising tools for regenerative myocardial therapy and in vitro modeling of cardiac disease; however, their full potential cannot be met without robust methods for differentiating them into cardiac-lineage cells. Here, we present novel protocols for generating hiPSC-derived cardiomyocytes (CMs), endothelial cells (ECs), and smooth muscle cells (SMCs) and for assembling them into a patch of human cardiac muscle (hCMP). The differentiation protocols can be completed in just a few weeks and are substantially more efficient than conventional methods, while the hCMP fabrication procedure produces a patch of clinically relevant size and incorporates a simple method for maturing the engineered tissue via mechanical stimulation. We also describe how the patch can be evaluated in a large-animal (swine) model of myocardial injury. Key words Human iPS cells, Cardiac patch, Differentiation

1

Introduction Human induced pluripotent stem cells (hiPSCs) are among the most potent tools for advancing regenerative myocardial therapy and studying cardiac disease because they have an unlimited capacity for proliferation and can be differentiated into cells descended from any of the three primary germ layers, including all types of cells that compose the heart [1]. Ideally, hiPSC differentiation protocols should be simple, quick, and scalable, and they must reliably produce highly pure populations of functionally mature cells in quantities sufficient for studies in large-animal models and clinical trials [2–4]. Once differentiated, the cells can be delivered via direct intramyocardial injection; however, a patch-based approach [5] may be more effective because the patch can be engineered to maximize the survival and/or promote the proliferation of transplanted cells while simultaneously providing structural support to strengthen the injured myocardium and limit adverse remodeling [6, 7].

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_14, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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This chapter details a series of protocols that have been developed for differentiating hiPSCs into cardiomyocytes (CMs), smooth muscle cells (SMCs), and endothelial cells (ECs), and then assembling the differentiated cells into a human cardiac muscle patch (hCMP). The differentiation protocols can be completed in just a few weeks and produce exceptionally pure cell populations: 96.4% of the hiPSC-CMs expressed cardiac troponin T (cTnT), 94.5% of the hiPSC-SMCs expressed α-smooth muscle actin (αSMA), and >95% of the hiPSC-ECs expressed CD31 and/or vascular endothelial cadherin. Furthermore, because experiments in large-animal models [8] are crucial for defining the optimal cell type, dose, timing, and route of administration, for characterizing the mechanisms associated with any observed benefits [9], and for establishing the safety, efficacy, and reproducibility of treatment before clinical use, we also describe how the patch can be evaluated in a swine model of myocardial infarction (MI) [10].

2

Materials

2.1 Maintaining hiPSCs

1. Dulbecco’s phosphate-buffered saline (DPBS). 2. Dulbecco’s modified Eagle’s medium: Nutrient mixture F-12 (DMEM/F12). 3. Growth factor-reduced Matrigel. 4. mTeSR1 medium. 5. Versene. 6. Y-27632.

2.2 Supplies/ Reagents for Differentiating hiPSCs into CMs

1. DPBS. 2. Versene. 3. Y27632 (Rho-associated kinase inhibitor). 4. Growth factor-reduced Matrigel. 5. CHIR99021. 6. Roswell Park Memorial Institute 1640 (RPMI-1640) medium. 7. B27 supplement, minus. 8. B27 supplement, complete. 9. IWR-1. 10. 0.25% Trypsin-EDTA. 11. Fetal bovine serum (FBS). 12. Glucose-free RPMI-1640. 13. Pen-strep antibiotic.

Patch Fabrication and Transplantation

2.3 Supplies/ Reagents for Differentiating hiPSCs into ECs

189

1. DPBS. 2. B27 supplement, minus insulin. 3. B27 supplement, complete. 4. Accutase. 5. Y-27632. 6. DMEM. 7. Fibronectin. 8. Endothelial cell growth medium-2 MV (EGM2-MV). 9. Vascular endothelial growth factor (VEGF), human. 10. EasySep Magnet. 11. Human CD34 Positive Selection Kit.

2.4 Supplies/ Reagents for Differentiating hiPSCs into SMCs

1. Accutase. 2. mTeSR. 3. CHIR99021. 4. Y-27632. 5. B27 supplement, minus vitamin. 6. Activin-A. 7. N2 supplement. 8. Neurobasal medium. 9. Platelet-derived growth factor BB (PDGF-BB). 10. Heparin.

2.5 Supplies and Reagents for hCMP Manufacture

1. FBS. 2. Pen-strep antibiotic. 3. DMEM. 4. Rectangular nylon mold (dimensions: internal, 4 cm  2 cm; external, 5 cm x 3 cm; height, 1 cm) (tool die maker office of UAB, USA). 5. Fibrinogen. 6. Pluronic solution. 7. Hydroxyethyl piperazineethanesulfonic acid (HEPES). 8. Thrombin. 9. Agarose. 10. Calcium chloride (CaCl2). 11. Ɛ-Aminocaproic acid (εACA).

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2.6 Supplies and Reagents Needed for Patch Transplantation in Swine MI Model

1. Isoflurane.

2.7

1. Humidified tissue culture incubator (MCO-18ACL-PA; Panasonic, Japan).

Equipment

2. 4-0 Silk suture. 3. 2-0 or 3-0 monocryl suture. 4. Cyclosporine.

2. Flow cytometry Dickinson, USA).

FACS

Aria

II

instrument

(Becton

3. Inverted phase-contrast microscope (CKX53, Olympus, Japan). 4. Nunc™ 15 mL Graduated Centrifuge Tubes. 5. Flow round-bottom tube (5 mL). 6. 6-Well plate. 7. Serological pipettes (1, 5, 10, and 25 mL). 8. Stericup filtration system (250 mL). 9. Stericup filtration system (500 mL). 10. Compact digital waving rotator. 11. Centrifuge. 12. Eppendorf Realplex2 PCR system. 13. 150 cm3 Flask. 14. Bright-Line™ Hemacytometer.

3

Methods The hiPSCs used for the development of these protocols were generated from cardiac-lineage cells. Fibroblasts from the left atrium of adult male humans were transfected with Sendai viruses coding for OCT4, SOX2, KLF4, and C-MYC to generate the hiPSCs; then, the hiPSCs were engineered to constitutively express green fluorescent protein (GFP) and maintained with hiPSC growth medium in Matrigel-coated plates [11].

3.1 HiPSC Culture and Maintenance

1. Visually monitor hiPSCs and passage at 80% confluence (see Note 1); split cells in a ratio of 1:6–1:8. 2. Thaw one Matrigel aliquot (2 mg) on ice or in a 4  C refrigerator. 3. In a sterile hood, transfer the Matrigel solution to a 50 mL conical tube containing 23 mL cold (4  C) DMEM/F12, and then immediately add 1 mL of the Matrigel-DMEM/F12 mixture into each well of a 6-well plate; allow the Matrigel to set for 60 min at 37  C (see Note 2).

Patch Fabrication and Transplantation

191

4. Remove the 6-well plate of hiPSCs from the incubator and place in the hood; aspirate the media from the cells and wash the plate once with PBS. 5. Add 1 mL of room-temperature Versene to each well, and incubate the plate for 5 min at 37  C. 6. Aspirate the Versene with a Pasteur pipet and add 3 mL of mTeSR1 + 5 μM of Y-27632 to each well; disperse the medium over the surface of the plate until all cells are detached. 7. Remove the cells from the surface of the well and transfer into a sterile conical tube containing 9 mL of mTeSR1 + 5 μM of Y27632; gently mix 5–10 times with a 10 mL pipette. 8. Seed 2 mL of the cell suspension into each well of a Matrigelcoated 6-well plate and return the plate to the 37  C, 5% CO2, incubator; disperse the cells across the surface of the wells by moving the plate in three quick front-to-back and side-to-side motions. 9. On each subsequent day, aspirate the medium from each well and replace with 2 mL of fresh, room-temperature mTeSR1 medium. 3.2 Differentiation of hiPSCs into CMs

1. Day 0: When the hiPSCs reach 80% confluency, change the culture medium to 6 μM of CHIR99021 in RPMI-1640 + 2% B27 without insulin and culture for 48 h. 2. Day 2: Change the CHIR99021-containing medium to RPMI-1640 medium + 2% B27 without insulin and culture for 24 h. 3. Day 3: Change the medium to 5 μM of IWP-1 in RPMI1640 + 2% B27 without insulin and culture for 48 h. 4. Day 5: Change the medium to RPMI-1640 + 2% B27 without insulin and culture for 48 h. 5. Day 7: Change the medium to RPMI-1640 + 2% B27 complete and replace with fresh RPMI-1640 + 2% B27 complete every 3 days thereafter; spontaneously beating cells should begin to appear on days 8–10. 6. Day 12: Aspirate the medium and wash the cells once with PBS. 7. Dissociate the cells by culturing them in 1 mL of 0.25% trypsin for 5 min at 37  C and then using a 1 mL pipette to repeatedly draw up the dissociation solution and spray it over the cell layer. 8. Transfer the dissociated cells into a 15 mL conical tube containing 5 mL of 5% FBS in RPMI-1640 + 2% B27 complete and centrifuge for 5 min at 240  g; aspirate and discard the supernatant.

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9. Resuspend the cells in 3 mL of RPMI-1640 + 2% B27 complete + 10 μM of Y-27632 and plate onto a new Matrigelcoated 6-well plate (see Note 3). 10. Day 15: Prepare a low-glucose medium by combining 10 mL of B27 complete and 5 mL of pen-strep antibiotic with 500 mL of glucose-free RPMI-1640 medium. 11. Transfer the differentiated hiPSCs into 3 mL of the low-glucose medium and culture for 6 days with medium changes every 3 days; noncardiomyocytes will die during the glucose-deprivation period, producing a highly pure population of hiPSC-CMs. 12. Day 21: Change the medium to 3 mL of RPMI-1640 + 2% B27 complete. 3.3 Differentiation of hiPSCs into ECs

1. Day 3: Prepare the hiPSCs for differentiation by dissociating them with Accutase at 37  C for 5 min, and then seed the cells in mTeSR medium supplemented with 5 μM of Y-27632 onto a Matrigel-coated 6-well plate (50,000 cell/cm2) for 24 h; then culture cells in mTeSR medium and change medium daily. 2. Day 0: Aspirate the mTeSR from the plate and initiate differentiation into endothelial progenitor cells (EPCs) by adding a mixture of DMEM + 6 μM CHIR99021 + 100 μg/mL ascorbic acid (3 mL/well); culture the cells for 2 days. 3. Day 2: Aspirate the CHIR99021-containing medium and then culture the cells in DMEM + 100 μg/mL ascorbic acid (3 mL/ well) for 3 days. 4. Day 5: Dissociate the hiPSC-EPCs with Accutase for 8 min and purify via CD34 selection with an EasySep Magnet and a Human CD34 Positive Selection Kit as directed by the manufacturer’s instructions. 5. Resuspend the purified cells in EGM-2MV medium with 50 ng/mL of VEGF and transfer them into a fibronectincoated 150 cm2 flask (50,000 cells/cm2); replace the medium with fresh EGM-2MV/VEGF medium every 2 days (i.e., on days 8, 10, and 12). 6. Day 12: Purify the fully differentiated hiPSC-EC population by using a fluorescence-activated cell sorting (FACS) apparatus to collect cells that express both CD31 and vascular endothelial cadherin (VE-cadherin).

3.4 Differentiation of hiPSCs into SMCs

1. Day 0: Visually monitor the hiPSCs until they reach 80% confluency; then, dissociate the cells by incubating them with 1 mL warmed (37  C) Accutase for 5 min. 2. Add 2 mL of warmed mTeSR, transfer the cells to a 15 mL Falcon tube, and centrifuge the suspension at 200  g for

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5 min; aspirate the supernatant, and resuspend the cellcontaining pellet in 12 mL of warmed mTeSR + 5 μM Y-27632. Seed the cells in a 6-well plate (35,000–45,000 cells/cm2) and incubate overnight at 37  C and 5% CO2 (see Note 4). 3. Day 1: Prepare a N2B27 medium by combining 20 mL of B27 without vitamin A, 10 mL of N2 supplement, 1 mL of β-mercaptoethanol, 500 mL of neurobasal medium, and 500 mL of DMEM/F12; then, replace the mTeSR/Y-27632 medium with warmed N2B27 medium supplemented with 6 μM of CHIR99021 and culture the cells for 3 days without medium changes. 4. Day 4: Replace the N2B27/CHIR-99021 medium with the N2B27 medium supplemented with 10 ng/mL ofPDGF-BB and 2 ng/mL activin-A (3 mL/well); culture the cells for 2 days with daily medium changes. 5. Day 6: Dissociate the cells with warmed Accutase and them (30,000 cells/cm2) into a collagen-coated 175 flask; culture with N2B27 medium supplemented activin-A (2 ng/mL) and heparin (1:1000, 2 μg/mL concentration). 3.5 Construction of a Large hCMP

seed cm2 with final

1. In the hood, coat the bottom of a 10 cm petri dish with 5% pluronic solution for 2 h; then, remove the excess pluronic solution and allow the petri dish to dry. 2. Position a clean, autoclaved, rectangular nylon mold in the middle of the coated petri dish; seal the mold to the dish by pouring melted 2% agarose around the outside of the mold and allowing it to cool and solidify. 3. Suspend 4  106 hiPSC-CMs, 2  106 hiPSC-ECs, and 2  106 hiPSC-SMCs in a solution containing 0.12 mL of fibrinogen (25 mg/mL), 0.02 mL of Matrigel, and 0.56 mL of HEPES (20 mM, pH 7.4) (see Note 5). 4. Initiate fibrin polymerization by mixing the cell-containing fibrinogen solution with a solution containing 0.004 mL of thrombin (80 U/mL), 0.001 mL of CaCl2 (2 M), and 0.3 mL of DMEM (final volume: 1 mL) (see Note 6); the mixture typically solidifies in ~1 min to form the hCMP. 5. Add a high-serum medium composed of DMEM, 10% FBS (see Note 7), 2% B27 complete, 2 mg/mL of Ɛ-aminocaproic acid (see Note 8), 1  pen-strep, and 10 μM of Y-26732 to the dish, and culture at 37  C on a rocking (45 rpm) platform for 24 h. 6. Replace the high-serum medium with medium composed of DMEM, 2% FBS, 2% B27 complete, 2 mg/mL of Ɛ-aminocaproic acid, and 1  pen-strep, and culture the

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hCMP for another 6 days on the rocking platform (see Note 9) with medium changes every other day; synchronized beating of hiPSC-CMs across the entire hCMP will typically appear on the second day after hCMP manufacture. 3.6 hCMP Transplantation

1. Sedate the swine (~13 kg, 45 days of age) with intramuscular injections of Telazol (0.05–0.1 mL/kg) and maintain anesthesia with isoflurane (1.5–2.0%). 2. Intubate and ventilate the animals with a respirator and supplemental oxygen; monitor body temperature, electrocardiograms (ECG), blood pressure, and arterial oxygen saturation throughout the surgical procedure (see Note 10). 3. Perform a thoracotomy in the fourth intercostal space under sterile conditions to expose approximately 0.5 cm of the coronary artery, and then occlude the coronary artery with a ligature for 60 min; this procedure typically produces an infarction that occupies ~10% of the left ventricular mass. 4. Remove the ligature, and observe the animal for 15 min; then, position two hCMPs on the surface of the epicardium over the infarcted region and suture them to the heart with 4-0 silk sutures. 5. Close the muscle layers, subcutaneous tissue, and chest wall with 3-0 or 2-0 monocryl sutures, depending on the size of the animal. 6. Orally administer cyclosporine (15 mg/kg per day with food) and methylprednisolone (1.5 mg/kg per day with food) for immunosuppression and perform standard postoperative care, including analgesia, until the animal eats normally and becomes active. 7. Cardiac function, such as left ventricular ejection fractions and systolic thickening fractions, can be evaluated in living animals before the surgical procedure and during the recovery period via a cardiac magnetic resonance imaging (MRI).

3.7 Analysis of hCMP Engraftment

1. Stop the heart during diastole with injection of KCl (74 mg/kg); harvest the heart after beating ceases, and record the total heart weight. 2. Separate the right- and left-halves of the heart at the ventricular septal wall; include the ventricular septal wall in the left half, remove the left atrium, and record weight of the remaining tissue (i.e., the left ventricle). 3. Vertically cut the left ventricle into six short-axis rings (R1–R6), and cut each ring sequentially into eight samples (S1–S8); collect S1–S5 of R1–R4 (i.e., from the site of patch administration) for analysis (see Note 11).

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4. Calculate the engraftment rate: Cut about 0.5 g of myocardial tissue from the samples and digest with proteinase K overnight at 56  C; then, isolate the total DNA from the digested buffer with a QIAGEN DNA isolation kit. 5. Quantify the amount of Y-chromosomes present in the samples via quantitative real-time polymerase chain reaction (qPCR) on an Eppendorf Realplex2 PCR system with the SYBR Green kit and appropriate primers (sense: ATCAGCCTAGCCTGTCTTCAGCAA; antisense: TTCACGACCAACAGCACAGCAATG). 6. Determine the number of engrafted cells by comparing the number of cycles required for each tissue sample to a standard curve calculated from the DNA of known quantities of undifferentiated hiPSCs; then, multiply the masses of the samples by the number of engrafted cells per gram, and divide the total number of engrafted cells in all samples by the number of cells administered to calculate the engraftment rate for each animal. 7. Cut the stored tissues into 10 μm cryosections for additional immunohistological assessments; for example, engrafted hiPSC-CMs can be identified by the expression of the human isoform of cTnT (hcTnT), or by the co-expression of human nuclear antigen (HNA) and non-species-specific cardiac troponin T (cTnT); hiPSC-SMCs can be identified via the co-expression of α-smooth muscle actin (αSMA) and GFP, or the human-specific isoform of calponin 1 (hCalponin 1) and αSMA; hiPSC-ECs can be identified via the expression of the human-specific CD31 isoform (hCD31); and the sarcomeric structure can be visualized by staining sections for the co-expression of GFP and α-actinin.

4

Notes 1. hiPSCs will begin to differentiate if they reach confluency. 2. All the media used in this protocol should be stored at 4  C and used within 2 weeks of production. 3. Survival of the differentiated hiPSC-CMs can be improved by culturing the cells with 10 μM of Y-27632 for 24 h. 4. For maximum hiPSC-SMC yield, the optimal density of seeded hiPSCs can vary, depending on which hiPSC line is used, and should be adjusted accordingly. 5. If the fibrinogen begins to fall out of the solution (forming visible particulate matter), warm the solution to 37  C until it becomes clear and then chill on ice.

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6. Keep the cells, fibrinogen, and thrombin solutions on ice until mixing; warming will reduce cell viability and accelerate the fibrin polymerization reaction. 7. Culturing the hCMP in a high-serum medium (10%) for the first 24 h will attenuate the shock-induced cells by entrapment in the matrix and facilitate cell-matrix connections. 8. Ɛ-Aminocaproic acid is used to prevent the fibrin patch from degrading during the culture. 9. Assessments of conduction velocity, calcium transients, tissue microresistivity, and force generation suggest that the 7-day, dynamic (rocking) culture period improves hCMP maturity and electrical integration [12]. 10. If ventricular fibrillation occurs, the animal can usually be rescued via immediate electrical defibrillation. 11. The S1 sample is taken from the anterior wall of the left ventricle, just to the left of the septal wall.

Acknowledgments This work was supported by the following funding sources: NIH RO1 HL 95077, HL114120, HL 131017, HL 138023, and UO1 HL134764. References 1. 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 67:2161–2176 2. Patsch C, Challet-Meylan L, Thoma EC, Urich E, Heckel T, O’Sullivan JF, Grainger SJ, Kapp FG, Sun L, Christensen K, Xia Y, Florido MH, He W, Pan W, Prummer M, Warren CR, Jakob-Roetne R, Certa U, Jagasia R, Freskgard PO, Adatto I, Kling D, Huang P, Zon LI, Chaikof EL, Gerszten RE, Graf M, Iacone R, Cowan CA (2015) Generation of vascular endothelial and smooth muscle cells from human pluripotent stem cells. Nat Cell Biol 17:994–1003 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/beta-catenin signaling under fully defined conditions. Nat Protoc 8:162–175 4. Bao X, Lian X, Dunn KK, Shi M, Han T, Qian T, Bhute VJ, Canfield SG, Palecek SP

(2015) Chemically-defined albumin-free differentiation of human pluripotent stem cells to endothelial progenitor cells. Stem Cell Res 15:122–129 5. Breckwoldt K, Letuffe-Breniere D, Mannhardt I, Schulze T, Ulmer B, Werner T, Benzin A, Klampe B, Reinsch MC, Laufer S, Shibamiya A, Prondzynski M, Mearini G, Schade D, Fuchs S, Neuber C, Kramer E, Saleem U, Schulze ML, Rodriguez ML, Eschenhagen T, Hansen A (2017) Differentiation of cardiomyocytes and generation of human engineered heart tissue. Nat Protoc 12:1177–1197 6. Ogle BM, Bursac N, Domian I, Huang NF, Menasche P, Murry CE, Pruitt B, Radisic M, Wu JC, Wu SM, Zhang J, Zimmermann WH, Vunjak-Novakovic G (2016) Distilling complexity to advance cardiac tissue engineering. Sci Transl Med 8:342ps13 7. Tzatzalos E, Abilez OJ, Shukla P, Wu JC (2016) Engineered heart tissues and induced pluripotent stem cells: macroand

Patch Fabrication and Transplantation microstructures for disease modeling, drug screening, and translational studies. Adv Drug Deliv Rev 96:234–244 8. Bolli R, Ghafghazi S (2015) Cell therapy needs rigorous translational studies in large animal models. J Am Coll Cardiol 66:2000–2004 9. Zhang JJ (2013) Mechanisms of cell therapy for clinical investigations: an urgent need for large-animal models. Circulation 128:92–94 10. Gao L, Gregorich ZR, Zhu W, Mattapally S, Oduk Y, Lou X, Kannappan R, Borovjagin AV, Walcott GP, Pollard AE, Fast VG, Hu X, Lloyd SG, Ge Y, Zhang J (2018) Large cardiac muscle patches engineered from human induced-

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pluripotent stem cell-derived cardiac cells improve recovery from myocardial infarction in swine. Circulation 137:1712–1730 11. Zhang L, Guo J, Zhang P, Xiong Q, Wu SC, Xia L, Roy SS, Tolar J, O’Connell TD, Kyba M, Liao K, Zhang J (2015) Derivation and high engraftment of patient-specific cardiomyocyte sheet using induced pluripotent stem cells generated from adult cardiac fibroblast. Circ Heart Fail 8:156–166 12. Jackman CP, Carlson AL, Bursac N (2016) Dynamic culture yields engineered myocardium with near-adult functional output. Biomaterials 111:66–79

Chapter 15 Isolation and Characterization of Intact Cardiomyocytes from Frozen and Fresh Human Myocardium and Mouse Hearts Honghai Liu, Kevin Bersell, and Bernhard Ku¨hn Abstract Procurement and characterization of intact human cells are essential for studies in regenerative medicine and translational medical research. The selection of the currently available approaches to isolate intact cells depends on the age of the hearts. To isolate cardiomyocytes from the fetal or neonatal myocardium, the myocardium can be minced into small tissue blocks followed by enzyme incubation. However, the fetal and neonatal cardiomyocytes are very soft and the morphology changes from long rod or spindle shape to spheres after isolation. Because of the dense packing of the cardiomyocytes and the strong cell-cell connection in adult myocardium, it is difficult to isolate the cardiomyocytes from adult myocardium by enzyme incubation only. A perfusion method is necessary to deliver the enzyme solution to the deep layers of the myocardium. However, intact hearts, which are very rare, are required for the perfusion method. Therefore, lacking methods to efficiently isolate cardiomyocytes from myocardium of various ages builds a barrier between basic research and clinical studies. Here, we describe a method for the isolation of intact cardiomyocytes from fresh or frozen human myocardium or fresh mouse hearts and the quantification of multinucleation, cardiomyocyte size, cell cycle activity, and total cardiomyocyte count per heart. We generalize this fixation-digestion method by isolating cells from a variety of mouse organs, including the liver, lung, and thymus. Key words Frozen and fresh myocardium, Intact cardiomyocytes, Fixation-digestion, Multinucleation, Cell volume, Total number of cardiomyocytes, Immunofluorescence

1

Introduction Regenerative medicine strategies have the potential to revolutionize therapies for a wide range of diseases [1]. These approaches require rigorous characterization of single cells and make the collection of intact cells essential for studies in regenerative medicine. The central experiments in the study of regenerative medicine require the determination of cell cycle activity, proliferation, cell size, and total number of cells. At present, the methods for characterizing intact cells are based on immunofluorescence

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_15, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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microscopy, which is currently the major approach for characterizing intact cardiomyocytes [2–4]. However, the nature of the cellcell connection and the limited thickness of the tissue sections make rigorously measuring single-cell characteristics (e.g., nucleation, cell volume) and the total number of cardiomyocytes in tissue sections challenging. Some researchers have isolated cardiomyocytes from heart samples using enzyme digestion for quantification of cardiomyocyte binucleation [3, 5, 6] and evaluating the total number of cardiomyocytes [5]. However, cell death and insufficient digestion decrease the quality and reproducibility of the results. Moreover, this kind of enzyme digestion requires re-optimization for context-specific myocardium, for instance, isolating cardiomyocytes from infant, aged, or diseased human myocardium. In addition, enzymatic digestion techniques cannot isolate cardiomyocytes from frozen human myocardium samples. We have developed the fixation-digestion method to isolate intact cardiomyocytes from fresh and frozen human myocardium through enzyme dissociation of formaldehyde-fixed samples [7]. The fixation-digestion method eliminates the viability issue of the cardiomyocytes, as it preserves the cardiomyocyte morphology and minimizes the tissue loss during cell dissociation. Here, we describe the details of the fixation-digestion method and demonstrate its use for characterization of intact cardiomyocytes from fresh and frozen human and mouse myocardium and on isolated cells from the mouse liver, lung, and thymus.

2

Materials

2.1 Tools and Instruments

1. Curved fine forceps. 2. Fine scissors. 3. Disposable scalpels. 4. Eppendorf tubes (2 mL). 5. Lab shaker. 6. Hybridization oven (37
C). 7. Multi-key differential counter. 8. Epifluorescence microscope connected to a CCD/CMOS camera. 9. Confocal microscope that acquires z-stack 3D images. 10. Fiji software (https://fiji.sc/).

2.2

Reagents

1. Fixation reagent: 3.7% Formaldehyde solution in distilled water. 2. Digestion buffer: Collagenase B (3.6 mg/mL), collagenase D (4.8 mg/mL). Dissolve the enzyme in PBS.

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3. Blocking and permeabilization solution: Donkey or goat serum (2.5%), Triton X-100 (0.05%). Dilute the chemicals in PBS. 4. Cardioplegia solution: KCl (25 μM). Dissolve the chemical in PBS.

3

Methods

3.1 Sample Fixation and Enzyme Digestion

1. For freshly isolated mouse hearts, remove the atria and other non-ventricle tissue (i.e., only ventricular cardiomyocytes will be isolated). Wash off the blood with ice-cold cardioplegia. For fresh-collected human myocardium, cut sufficient tissue but not larger than 5 mm  5 mm  2 mm for the experiment and immerge it in cardioplegia solution. For frozen myocardium, thaw the tissue on ice (~5 min) and immerge the tissue in ice-cold cardioplegia. 2. Cut a small piece of the myocardium (e.g., 5 mm  5 mm  2 mm), put it on a dry surface of a Petri dish, and drop 100 μL of cardioplegia to cover the myocardium piece. Alternative: Sample preparations will be conducted using a series of mechanical manipulations: cut a small piece of myocardium and place it onto a dry surface of a petri dish and drop 100 μL of cardioplegia to cover the myocardium piece. Return the remaining tissue to the ice. 3. Use a scalpel to mince the small piece of myocardium into ~0.5–1 mm tissue blocks, and then transfer all the minced tissue into a 2 mL Eppendorf tube containing 3.7% formaldehyde. 4. Repeat steps 2 and 3 until all of the myocardium for the experiment is minced and transferred to the same Eppendorf tube (see Note 1). 5. Place the Eppendorf tube on a lab shaker at room temperature, and fix the tissue blocks for 1 h and 40 min (see Note 2). 6. Wash the fixed tissue blocks with PBS three times. 7. The procedure is presented in Fig. 1. 8. Add 1 mL of the digestion buffer to the 2 mL Eppendorf tube containing the fixed heart tissue. 9. Put the Eppendorf tube in the hybridization oven (37
C) and shake the tube ten times per minute for 24 h. 10. Transfer the digested cells from the supernatant to a clean 2 mL Eppendorf tube and store upright at 4
C. 11. Resuspend the undigested heart tissue with 1 mL of fresh digestion buffer and shake the tube in the hybridization oven (37
C) at 10 rpm for 24 h.

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Digestion buffer

37°C, 24 hours, shaking

Dissociated cardiomyocytes in suspension

4°C, 24 hours Cardiomyocyte pellet

Remove the supernatant Transfer supernatant

Tissue blocks Add fresh digestion buffer

Remove the supernatant

37°C, 24 hours shaking

4°C, 24 hours

Re-suspension

Fig. 1 Isolation and collection of cardiomyocytes from tissue blocks

12. The dissociated cardiomyocytes in the first digestion will have formed a pellet at the bottom of the Eppendorf tube (step 10). Carefully remove the supernatant from the tube immediately prior to step 13. 13. Transfer the newly digested cells to the same Eppendorf tube, and store them at 4
C for 24 h to allow the cells in suspension to form a pellet at the bottom of the tube (see Note 3). 14. Repeat the above digestion procedure until all of the tissue is digested (see Note 4). Store the tube containing all the dissociated cardiomyocytes at 4
C for 24 h and remove the supernatant. 15. Resuspend the cell pellet at the bottom containing all of the cells dissociated from the tissue with PBS. Store the tube at 4
C for 24 h (see Note 5). 16. Remove the PBS from the supernatant and resuspend the cell pellet with clean PBS solution. Store the tube at 4
C for 24 h. 17. Remove the PBS from the tube as much as possible. Do not disturb the cell pellet at the bottom. 3.2 Count the Total Number of Cardiomyocytes in the Mouse Heart

1. Resuspend the pellet in PBS to make the volume of the cell solution 2 mL. 2. Transfer 10 μL of the cell solution to the hemocytometer to measure the concentration of the rod-shape cardiomyocytes, Ccardiomyocytes (cells/mL), under an inverted bright-field microscope (Fig. 2) (see Note 6).

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Fig. 2 Count the total number of cardiomyocytes in a mouse heart. The cardiomyocytes were isolated from adult mice (P30) using fixation-digestion method. No further treatment for immunofluorescence. The cells with blunt ends are considered half cells (white arrows). Scale bar: 100 μm

3. Calculate the total number of cardiomyocytes in the tube (2 mL, i.e., the total number of the cardiomyocytes of the heart), Ntotal ¼ Ccardiomyocytes2 mL. 4. Store the Eppendorf tube containing the cell solution at 4
C for 24 h to allow the cells to settle at the bottom. 3.3 Immunofluorescence Microscopy

1. Remove the supernatant carefully. Do not disturb the cell pellet at the bottom. 2. Use 100 μL of blocking and permeabilization solution to resuspend the cells. 3. Incubate at room temperature for 30 min (see Note 7). 4. Add 100 μL of 2X primary antibody working solution to the tube (e.g., for α-actinin and H3P antibodies, the working solution is 1:200 of stock solution; dilute the stock solution with PBS by 1:100 to make the 2X concentration of the working solution). Invert the solution several times to mix. 5. Lay the tube down at room temperature for 1 h (Fig. 3) (see Note 8). 6. Add 1.5 mL of PBS to the tube. 7. Store the tube upright at 4
C for 24 h. Let the cells settle at the bottom. 8. Remove the supernatant carefully. Do not disturb the cell pellet at the bottom of the tube (see Note 9).

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Cells and antibody solution

Fig. 3 Lay the tube down at room temperature and incubate the cardiomyocytes with antibody solution

9. Prepare the 2X concentration of the secondary antibody working solution in PBS (e.g., we usually dilute the secondary antibody by 1:200 for immunostaining; the dilution for 1:100 is 2X concentration) (see Note 10). 10. Add 100 μL of the secondary antibody working solution (2X) to the tube. Resuspend by gently flicking the tube. 11. Lay the tube down at room temperature for 1 h. 12. Add 100 μL of Hoechst solution (PBS, 1:500) to the tube. 13. Lay the tube down at room temperature for 5 min. 14. Add 1 mL of PBS to the tube. 15. Store the tube upright at 4
C for 24 h. 16. Remove the supernatant carefully. Do not disturb the cell pellet at the bottom. 17. Resuspend the pellet with 2 mL of PBS. 18. Store the tube upright at 4
C for 24 h. 19. Remove the supernatant carefully. Do not disturb the cell pellet at the bottom. 20. Resuspend the pellet with 1 mL of PBS. 21. Measure the concentration of the cardiomyocytes (CCM, unit: cardiomyocytes/mL) using a hemocytometer (see Note 11). 22. Transfer 100 μL of the cell solution to the wells of an 8-wellchambered cover glass. 23. Check the wells containing cells under an inverted bright-field microscope. Transfer more solution to the wells if 100 μL of the solution contains very few cells. Dilute the cell solution and redo the transfer if too many cells are transferred to the wells. 24. Calculate the total number of the cardiomyocytes (NCM ¼ CCM  100 μL) transferred to the wells. 25. Seal the wells with clear tape and cover the wells with a chamber lid (see Note 12). 26. The cells are ready for image acquisition.

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α-actinin, Hoechst, pan-Cadherin

P10

E19.5

B α-actinin, Hoechst, pan-Cadherin

P60

A

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Mouse

Fig. 4 Determine the nucleation of the intact cardiomyocytes of different nucleation dissociated from mice and humans of various ages. (a) Cardiomyocytes were isolated from mouse hearts. (b) Cardiomyocytes were isolated from diseased human hearts. Scale bar: 20 μm 3.4 Image Acquisition and Cardiomyocyte Characterization

1. Quantify the proportion of bi/multinucleated cardiomyocytes using an epifluorescence microscope. 2. Some rod-shaped cardiomyocytes can be broken during tissue block preparation, digestion, and the following procedures. Check the pan-cadherin to identify the intact cardiomyocytes (Fig. 4) (see Note 13). 3. Identify the cardiomyocytes and number of their nuclei through the eye port of the microscope to determine their nucleation. 4. Swipe through the well to count the number of mononucleated, binucleated, and multinucleated cardiomyocytes using a multi-key cell counter until the pre-designed total number of cardiomyocytes (e.g., 500 cardiomyocytes) has been reached. 5. Calculate the cardiomyocytes.

percentage

of

the

bi/multinucleated

6. Quantify the proportion of H3P+ cardiomyocytes using an epifluorescence microscope (Fig. 5). 7. Count the total number of the H3P+ cardiomyocytes (NH3PCM) in the well. 8. Calculate the percentage of the H3P+ cardiomyocytes in the myocardium or heart (PH3P-CM ¼ NH3P-CM/NCM  100%). 9. Measure the size of an intact cardiomyocyte by measuring the projection area (2D, unit: μm2) of the cardiomyocyte on the CCD/CMOS camera attached to an epifluorescence microscope. 10. Acquire multichannel images of the cardiomyocytes (bright field, α-actinin, pan-cadherin, Hoechst) (Fig. 6).

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P4

α-actinin, Hoechst, H3P

E19.5

Fig. 5 The intact mouse cardiomyocytes at M-phase. Cardiomyocytes were isolated from mouse hearts at various ages (E19.5: 19.5-day embryo; P4: 4 days after birth). Scale bar: 20 μm

Fig. 6 Measure the projection area of cardiomyocytes. The target cell is indicated by white arrows. (a) The snapshot of the isolated cardiomyocytes by a CMOS camera attached to an epifluorescence microscope. (b) The target cardiomyocyte is outlined for area measurement. (c) The measured area is reserved by cutting off in the bright-field channel. Scale bar: 40 μm

11. Open the image file with Fiji software in color 16-bit (8-bit) type, not RGB type. Use the (Image!Color!Channels Tool!Composite) tool to make all of the channels visible. 12. Use the (Analyze!Set scale) function to set the scale of the image if the scale is not set automatically. 13. Set the bright-field channel as the working channel. 14. Identify the cardiomyocytes (α-actinin+) and the nucleation (the number of nuclei, Hoechst+). 15. Use the “Freehand selections” tool to outline the bright-field boundary of the target cardiomyocytes. 16. Use the (Analyze!Measure) function to measure the “Area” value of the selected area. The “Area” value will appear in the popup “Results” window.

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Fig. 7 Measure the volume of cardiomyocytes from the z-stack. (a) The z-stack of the isolated cardiomyocytes is demonstrated as volume view. (b) Individual slices of the cardiomyocyte are indicated by a white arrow in (a) for the measurement of cardiomyocyte volume

17. Use the (Edit!Cut) function to permanently mark the selected area. It is important that the bright-field channel is the current working channel; it is only in this channel if the selected area is cut off but the immunostained pattern is preserved. 18. Save the modified image in tiff format to revisit in the future if needed. 19. Measure the volume (3D, unit: μm3) of a cardiomyocyte. 20. Using a confocal microscope, acquire a multichannel z-stack of the target cardiomyocytes (α-actinin, pan-cadherin, DAPI/ Hoechst). The smaller the step distance (dslice) of the z-stack, the more accurate the measurement. 21. Open the z-stack with Fiji software and use the (Analyze!Set scale) function to set the scale if the scale is not set automatically. 22. Measure the “Area” value (S) of the α-actinin+ area on each slice of the z-stack using the method described above (Fig. 7). 23. Calculate the volume VCM ¼ ∑S∙dslice. 3.5 Other Species and Organs

of

the

target

cardiomyocyte

We have applied the fixation-digestion method to isolate intact cells from a variety of organs of different species at various ages (Fig. 8).

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Fig. 8 The cells isolated from a variety of species and organs. (a) Cardiomyocytes from different species. The zoomed-in region of the human cardiomyocyte (lower panel) shows the characteristic striated structures. (b) The cells isolated from different mouse organs. Scale bar: 50 μm

4

Notes 1. To count the total number of cardiomyocytes in a mouse heart, it is important that all of the minced tissue pieces are collected and fixed with the fixation reagent. 2. It is important that the fixation procedure does not exceed 2 h; otherwise the cardiomyocytes will not be dissociated from the tissue. 3. Cardiomyocytes of different sizes need different lengths of time to settle at the bottom of the tube. Generally, larger cardiomyocytes settle at the bottom faster than smaller cardiomyocytes. Incubating the cardiomyocytes at 4
C for 24 h is sufficient for the smallest cardiomyocytes to settle at the bottom. For larger cardiomyocytes, the incubation time can be shorter. 4. When counting the total number of cardiomyocytes in mouse heart, it is important that all of the tissue is digested, i.e., no visible tissue blocks in the tube. Considering that the dissociated cardiomyocytes can also be digested by the digestion

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buffer, the digested cardiomyocytes in the suspension of the tube should be collected every day and stored at 4
C to inhibit enzyme activity. 5. This step washes off the digestion buffer from the isolated cardiomyocytes. 6. A scalpel can cut a few long-rod-shaped cardiomyocytes into halves (Fig. 2, the cells with blunt ends, white arrows). The number of cardiomyocyte halves should be divided by 2 to obtain the number of full cardiomyocytes. Because the permeabilization of the following procedures can destroy a few cardiomyocytes, the total number of cardiomyocytes in a heart should be counted before permeabilization. 7. After cell permeabilization, do not shake the tubes while incubating the cells with antibody solution; this could damage the cardiomyocytes in suspension. 8. Incubating the cell solution with the tube upright (see the position of the tube in Fig. 1) allows the cardiomyocytes to settle at the bottom of the tube and form a small cell pellet, which reduces the contact of the cells to the antibodies and causes nonuniform staining. Laying the tube down allows the cardiomyocytes spread onto a larger area on the wall of the tube in order to stain the cardiomyocytes in a more uniform manner. 9. After this step, use aluminum foil to wrap the tube. 10. The working concentration of the antibody solution should be determined by the try-and-test method. 11. Do not use the cardiomyocyte concentration obtained before permeabilization (see Note 6). 12. Sealing the well with clear tape can prevent the water from evaporating and keep the sample at 4
C for a longer period of time. 13. Embryonic and newborn cardiomyocytes exhibit spindle shapes with low cadherin expression. Therefore, the tapered ends can be regarded as the sign to identify intact cardiomyocytes for nucleation quantification.

Acknowledgments This research was supported by the Richard King Mellon Foundation Institute for Pediatric Research (UPMC Children’s Hospital of Pittsburgh), by a Transatlantic Network of Excellence grant by Foundation Leducq (15CVD03), Children’s Cardiomyopathy Foundation, and NIH grant R01HL106302 (to B.K.). This project was supported, in part, by UPMC Children’s Hospital of Pittsburgh (to H.L.), Genomics Discovery Award (UPMC Children’s

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Hospital), UPP Physicians, Vascular Medicine Institute, Aging Institute, and NIH grant UL1TR001857 from the Clinical and Translational Sciences Institute (University of Pittsburgh, to B.K.). References 1. Laflamme MA, Murry CE (2011) Heart regeneration. Nature 473(7347):326–335. https:// doi.org/10.1038/nature10147 2. Bensley JG, De Matteo R, Harding R, Black MJ (2016) Three-dimensional direct measurement of cardiomyocyte volume, nuclearity, and ploidy in thick histological sections. Sci Rep 6:23756. https://doi.org/10.1038/srep23756 3. Patterson M, Barske L, Van Handel B, Rau CD, Gan P, Sharma A, Parikh S, Denholtz M, Huang Y, Yamaguchi Y, Shen H, Allayee H, Crump JG, Force TI, Lien CL, Makita T, Lusis AJ, Kumar SR, Sucov HM (2017) Frequency of mononuclear diploid cardiomyocytes underlies natural variation in heart regeneration. Nat Genet 49(9):1346–1353. https://doi.org/10. 1038/ng.3929 4. Alkass K, Panula J, Westman M, Wu TD, Guerquin-Kern JL, Bergmann O (2015) No evidence for cardiomyocyte number expansion in preadolescent mice. Cell 163(4):1026–1036. https://doi.org/10.1016/j.cell.2015.10.035

5. Naqvi N, Li M, Calvert JW, Tejada T, Lambert JP, Wu J, Kesteven SH, Holman SR, Matsuda T, Lovelock JD, Howard WW, Iismaa SE, Chan AY, Crawford BH, Wagner MB, Martin DI, Lefer DJ, Graham RM, Husain A (2014) A proliferative burst during preadolescence establishes the final cardiomyocyte number. Cell 157 (4):795–807. https://doi.org/10.1016/j.cell. 2014.03.035 6. Soonpaa MH, Kim KK, Pajak L, Franklin M, Field LJ (1996) Cardiomyocyte DNA synthesis and binucleation during murine development. Am J Phys 271(5 Pt 2):H2183–H2189. https://doi.org/10.1152/ajpheart.1996.271. 5.H2183 7. Mollova M, Bersell K, Walsh S, Savla J, Das LT, Park SY, Silberstein LE, Dos Remedios CG, Graham D, Colan S, Kuhn B (2013) Cardiomyocyte proliferation contributes to heart growth in young humans. Proc Natl Acad Sci U S A 110(4):1446–1451. https://doi.org/10. 1073/pnas.1214608110

Chapter 16 Ex Vivo Techniques to Study Heart Regeneration in Zebrafish Sierra Duca and Jingli Cao Abstract Due to its pronounced regenerative capacity, the zebrafish heart represents an advantageous model system for exploring the cellular and molecular mechanisms of cardiac regeneration. Upon injury, the epicardium, the outermost mesothelial tissue layer of vertebrate hearts, serves dual purposes in the regenerating heart as both a signaling center and a source for crucial cell types. Traditional in vivo genetic approaches to study heart regeneration can be time consuming and are not applicable to large-scale approaches and live surveillance of cellular behaviors. Here, we demonstrate ex vivo methods to culture, maintain, and study the regenerative responses of epicardial tissue in excised zebrafish hearts. Epicardial cell proliferation and migration are monitored in real time after uninjured or injured hearts are excised, washed, and cultured for up to 30 days. In addition to these techniques, we describe ex vivo genetic ablation of the epicardium, cell proliferation assays, partial ventricular explant culturing, and chemical screening. Key words Heart regeneration, Zebrafish, Epicardium, Heart explant culture, Chemical screening, Live imaging

1

Introduction The minimal regenerative capacity of adult mammalian cardiomyocytes (CMs) is insufficient to repair damaged tissue after myocardial infarction [1]. By contrast, upon injury, the adult zebrafish heart can regenerate with minimal tissue scarring [2, 3]. The epicardium, a mesothelial tissue layer that covers vertebrate hearts, has been identified as necessary for myocardial regeneration in zebrafish [4]. In both fish and mammals, the epicardium has shared functions during heart regeneration and repair. Upon heart injury, the epicardium reactivates developmental programs to secrete paracrine signals for CM survival and proliferation, provide perivascular components (such as smooth muscle cells and pericytes) and other cell types (such as fibroblasts), and mediate inflammatory responses [5– 13]. Studies of epicardial regeneration responses will further elucidate the processes of heart regeneration and inform potential strategies for cardiac repair. Preexisting methods for studying heart regeneration have limitations, including a long generation time

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_16, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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for adult zebrafish (approximately 3 months) and restricted spatiotemporal capacity for large-scale screens and live imaging. To ameliorate some of these research obstacles, heart explants provide the opportunity for heightened live imaging in addition to enabling various experimental techniques to be performed, such as genetic ablation, cell proliferation assays, large-scale chemical screens, and tissue and bead grafts [4, 14, 15]. Here we describe standard explant culture, culture with epicardial ablation, and partial ventricular explant culture in coated dishes. This technique can be used to study both uninjured and injured hearts for up to 30 days in culture. Overall, heart explant cultures represent a novel ex vivo approach to spatiotemporally visualize epicardial cell behaviors.

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Materials Zebrafish

Zebrafish of the Ekkwill and Ekkwill/AB strains were maintained between 26 and 28.5
C with a light cycle of 14:10 h (light:dark) [2]. Adults between 4 and 12 months of both sexes were used. In these experiments, transgenic line Tg(tcf21:nucEGFP) pd41 was used to label epicardial cell nuclei (Fig. 2c) [3]. To ablate epicardial cells, a transgenic nitroreductase (NTR) line Tg(tcf21:mCherryNTR) pd108 was used [4, 6]. Other transgenic zebrafish can also be used with this protocol. Animal procedures were approved by the Animal Care and Use Committee at Weill Cornell Medical College.

2.2 Reagents and Key Equipment

1. Tricaine (MS-222): 15 mM Ethyl 3-aminobenzoate methanesulfonate (tricaine), 20 mM Tris–HCI at pH 7.4. The stock solution can be stored at 4
C for approximately 1 month, and at 20
C for longer term. To anesthetize adult fish, dilute stock solution in fish water to 0.042% (see Note 1).

2.1

2. Culture medium: DMEM plus 10% (vol/vol) FBS, 1% (vol/vol) MEM-NEAA, 100 U/ml penicillin, 100 μg/mL streptomycin, and 50 μM 2-mercaptoethanol (see Note 2). To reduce the incident of bacterial contamination, 0.1–0.2% (vol/vol) Primocin (antibiotic cocktail) was added to the culture medium. Filter sterilize with 0.22 μm filter and store the medium at 4
C for up to a month. 3. Ablation medium: 1 mM Metronidazole (Mtz). In a 50 mL Falcon tube, prepare 10–20 mL of 10 mM Mtz solution in culture medium just prior to use and vortex well. Dilute this solution 1:10 to the desired volume and filter sterilize with 0.22 μm filter (see Note 3). 4. Fish water: Use aquarium water from the zebrafish facility. 5. 70% (vol/vol) Ethanol.

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6. PBS: Use sterile PBS to wash the hearts. 7. 5-Ethynyl-20 -deoxyuridine (EdU): For proliferation assays, prepare a solution of 25 μM EdU in PBS (see Note 4). 8. 4% Paraformaldehyde (wt/vol) in PBS. 9. Fluoromount G. 10. Fibronectin: For partial ventricular explant culture, 12-well plates can be coated with 10 μg/mL fibronectin overnight. 11. Finger bowls (90  50 mm). 12. Plastic spoon. 13. Sponges with a single-center groove cut using scissors (0.5  2.5 cm). 14. 1.7-mL Eppendorf (EP) tubes. 15. 12-Well plates. 16. 60 mm Petri dish. 17. Syringe filter units (0.22 μm). 18. Sterile disposable plastic Pasteur pipettes. 19. Orbital shaker. 20. 1 mL Pipette and filter pipette tips. 21. Dissecting microscopes. 22. Cell culture incubator.

3 3.1

Methods Preparation

1. Prepare the necessary reagents as described in Subheading 2. If performing epicardial ablation experiments, prepare the ablation medium as described above. 2. Obtain a small sponge and create a 0.5  2.5 cm groove. It should be large enough to hold the fish in place during the heart dissection. 3. Place the orbital shaker in the cell culture incubator (28
C, 5% CO2, humidified). 4. Prewarm the DMEM and PBS to 28
C. 5. Sterilize the forceps, scissors, dissecting scope stage, benchtop surrounding dissecting scope, and sponge with 70% ethanol solution. Wash the sponge several times and moisten with fish water prior to heart isolation (see Note 5) (Fig. 1a). 6. Prepare a clean 60 mm petri dish containing 5 mL of culture medium and an EP tube with 1 mL of culture medium. Hearts will be placed in the EP tube (Fig. 1a).

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Fig. 1 Microscope and tool setup. (a) Equipment and tool setup for heart extractions including 1 mL pipette, 1 mL filter pipette tips, plastic Pasteur pipette, dissecting microscope, 60 mm petri dish with medium, sponge with groove, paper towels, forceps and scissors, PBS, EP tube with medium, plastic spoon, and finger bowl containing fish water with tricaine. (b) Heart explant with intact atrium (A), outflow tract (OFT), and ventricle (V). (c) Place the 12-well plate with the extracted hearts in an incubator (28
C, 5% CO2) shaking at 120–150 rpm. A polystyrene foam is placed in between to prevent extra heating from the shaker

7. If studying injured hearts, partial ventricular resection (removal of 10–20% of ventricle) can be performed prior to heart extraction as described previously [2]. 3.2 Heart Extraction and Cleaning

1. In a glass finger bowl, euthanize fish with tricaine solution (see Note 6).

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2. Transfer the fish carefully using a plastic spoon. Dry the fish briefly on a paper towel and transfer to the groove of the moistened sponge with the ventral side of the fish facing upwards. 3. Under the dissecting scope, open the ventral wall of the fish using sterile forceps or scissors to make a cut in between the gill region. Upon opening the pericardial sac, locate the outflow tract, which is anterior to the ventricle. Grasp the anterior end of the outflow tract with the forceps and carefully pull the heart out of the chest cavity. Sever the ventral aorta and sinus venosus with the other forceps (see Note 7). A video demonstrating a similar procedure can be found in a previous publication [16]. 4. Place the heart in the 60 mm petri dish with culture medium (Fig. 1a). Store the additional hearts in this petri dish until all heart extractions are finished. Keep the culture dish lid closed whenever possible. 5. Using forceps to hold the anterior end of the outflow tract, remove any excess tissue and blood clots from the extracted hearts using the other forceps (see Note 7) (Fig. 1b). Remove the atrium when required. 6. Transfer hearts from the petri dish to the EP tube (with medium) using a sterile Pasteur pipette (Fig. 1a). 7. Under a biological safety hood, rinse the hearts in the EP tube three times using sterile PBS and sterile pipette tips. 3.3 Standard Explant Culture

1. Add 1 mL of prewarmed medium (28
C) to each well of a 12-well plate. 2. Transfer each heart from the EP tube to a single well using a sterile Pasteur pipette (see Note 8). 3. Place the 12-well plate on the shaker in the 28
C incubator, shaking at 120–150 rpm for the entirety of the culture duration (see Note 9). Place a polystyrene foam between the plate and the shaker top to prevent extra heating from the shaker (Fig. 1c). 4. Change the culture medium every other day (see Note 10). Avoid touching the hearts while changing the medium (see Note 11). 5. Live imaging can be performed at desired timepoints for up to 30 days. Images can be taken using a stereo fluorescence microscope with a camera. Keep the lid of the culture plate closed while imaging to avoid bacterial contaminations (see Note 12). Typical images of heart explants carrying a tcf21: nucEGFP reporter are shown in Fig. 2a. 6. Proliferation assays and chemical treatments can be performed at required timepoints as described below (Fig. 2c).

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Fig. 2 Epicardial cell behaviors during explant cultures. (a) Explant hearts from tcf21:nucEGFP adults without outflow tract at 0 and 14 days of culture. Without blood pressure, the heart shrank. As a result of cell proliferation, the epicardial cell density increased. (b) In situ hybridization on a section of ventricular explant after 2 days of culture. Expression of raldh2 was detected primarily in the ventricular wall, suggesting epicardial activation. (c) Epicardial proliferation assay at 2 days of culture using adult tcf21:nucEGFP hearts incubated with 25 μM EdU for 1 h. The framed region is enlarged to show details below. Arrowheads indicate dividing epicardial cells and a single-channel image of EdU (white) is shown at the bottom right

7. Hearts can be fixed at selected timepoints for whole-mount imaging on a confocal microscope or sectioning followed by in situ hybridization or immunostaining (Fig. 2b). To fix hearts, use 4% paraformaldehyde (wt/vol) for 2 h at room temperature or overnight at 4
C. 8. For whole-mount imaging of fixed hearts, place each heart on a coverslip, add a drop of Fluoromount G, and then place another coverslip on top of the heart, compressing the heart. Dry the mounted hearts at room temperature for 2 h before imaging. Images can be taken for both sides of the heart using a confocal microscope (Fig. 2c).

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1. To ablate epicardial cells in explant cultures, use 1 mL of prewarmed (28
C) ablation medium (containing 1 mM Mtz) instead of the normal medium to each well of the 12-well plate. 2. Transfer each heart from the EP tube to a single well using a sterile Pasteur pipette. 3. Place the 12-well plate on the shaker in the 28
C incubator while shaking at 120–150 rpm for 24 h. 4. After the 24-h culture, carefully remove the ablation medium with Mtz, rinse the hearts twice with 1 mL of prewarmed PBS each time, and add 1 mL of normal culture medium (without Mtz) to each well. Avoid touching the hearts while removing the medium and PBS (see Note 11). Return the plate to the incubator and continue culturing while shaking. Dispose the Mtz solution according to institutional guidelines. 5. Change the culture medium every other day. Avoid touching the hearts while changing the medium (see Note 11). 6. Live imaging can be performed at desired timepoints as described above (Fig. 3). Keep the lid of the culture plate closed while imaging to avoid bacterial contaminations (see Note 12). 7. Proliferation assays and chemical treatments can be performed at required timepoints as described below. 8. Hearts can be fixed at selected timepoints for subsequent analyses, such as whole-mount imaging, sectioning, in situ hybridization, and immunostaining.

Fig. 3 Epicardial cell ablation and regeneration ex vivo. Hearts from tcf21:NTR; tcf21:nucEGFP adults were incubated with 1 mM Mtz for 24 h. After ablation, the remaining epicardial cells (labeled by a tcf21:nucEGFP reporter in green) regenerated from ventricular apex to base (arrows). Images depict explant hearts at 2, 5, 9, and 17 dpi (days post-injury)

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3.5 Proliferation Assay

1. Perform the heart culture procedure as described in Subheadings 3.3 and 3.4. 2. Prewarm the culture medium to 28
C and add 5-ethynyl20 -deoxyuridine (EdU) at a concentration of 25 μM. 3. Replace the medium with EdU medium at the desired timepoint. Avoid touching the hearts while changing the medium (see Note 11). Incubate on the shaker for 1 h at 28
C. 4. After 1 h, wash the hearts twice with prewarmed PBS. Hearts can be fixed at this point or you can continue culturing. 5. Fix the hearts with 4% paraformaldehyde (wt/vol) for 2 h at room temperature or overnight at 4
C. EdU staining procedures can be performed as previously reported [4]. 6. After staining, mount the hearts with coverslips and Fluoromount G as described in step 8 of Subheading 3.3. Images can be taken for both sides of the heart using a confocal microscope (Fig. 2c).

3.6 Partial Ventricular Explant Culture

1. The day prior to the experiments, coat 12-well plates with 10 μg/mL fibronectin and store overnight at 28
C. 2. Before culturing, prewarm the culture medium and PBS to 28
C. 3. Extract and clean the hearts as described in Subheading 3.2. Remove the atria and outflow tracts with forceps and only keep the ventricles. Cut each heart ventricle into three pieces using sterile forceps. 4. Rinse the ventricular pieces with warm PBS three times under a biological safety hood. 5. Remove the fibronectin solution right before plating explants. Add 1 mL of culture medium in each well. Plate the partial ventricular explants in the culture medium and culture for 72 h without agitation. 6. Change the culture medium every other day from 72 h (see Note 13). If performing chemical treatments or screens, add the appropriate compounds to the medium as described in Subheading 3.7. 7. Image the migrating epicardial cells after 72 h for up to 12 days using a stereo fluorescence microscope with a camera (Fig. 4). Keep the lid of the culture plate closed while imaging to avoid bacterial contaminations (see Note 12).

3.7 Chemical Treatments and Screens

1. Prepare the culture medium with an appropriate concentration of select small-molecule compounds to probe epicardial effects. Solvents such as DMSO, PBS, ethanol, or water can be used as a negative control. Filter-sterilize the medium using a syringe filter (0.22 μm) to avoid bacterial contamination.

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Fig. 4 Partial ventricular explant culture and drug treatment. Epicardial cell migration from partial ventricular explants carrying a tcf21:nucEGFP reporter. Cell nuclei are labeled with nuclear GFP (white). Cyclopamine (10 μM)-treated explants displayed decreased epicardial migration and proliferation as compared to the vehicle (DMSO). The framed regions are enlarged to show details at the bottom

2. Remove the culture medium from the explant cultures and add the medium with compounds or vehicles. If examining regeneration after epicardial ablation, add the compounds 2 days post-Mtz treatment. 3. Change the medium with the compounds or vehicles every other day. Do not touch the explants while changing the medium (see Note 11). 4. Live imaging can be performed before adding the compounds and any time after the treatment (Fig. 4). Keep the lid of the culture plate closed while imaging to avoid bacterial contamination (see Note 12). Epicardial cell survival, numbers (proliferation), and migration can be assessed using the tcf21: nucEGFP reporter (see Note 14).

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Notes 1. When using tricaine, avoid inhalation and direct contact with skin and eyes. 2. Nonessential amino acids (NEAA) have been used to increase the longevity of heart slice cultures. Additionally, 2-mercaptoethanol can act as a reducing agent to reduce oxygen radical levels [17]. 3. Metronidazole (Mtz) is a carcinogen. Wear gloves and eye protection. Avoid inhalation and contact with skin and eyes. 4. When handling EdU, avoid inhalation and contact with skin and eyes. Wear gloves and eye protection. 5. Sterile surfaces and tools are vital to reducing bacterial contamination. Use only sterile pipette tips. Washing the sponge with ethanol prior to each series of heart extractions reduces the risk of bacterial contamination of the culture. 6. When euthanizing the fish with tricaine, wait until gill movements cease and the fish no longer responds to a tail pinch. All animal procedures must be performed in accordance with standardized institutional guidelines. 7. When isolating and cleaning hearts, avoid damages to the epicardium, which may affect subsequent analyses. 8. More than one heart can be cultured in a well of a 12-well plate. However, hearts may adhere to each other, which may affect live imaging and other analyses. 9. The shaking speed of the orbital shaker can be adjusted according to its model and age. Generally, a speed of 120–150 rpm works well. However, the shaking speed should be held constant throughout the whole culture. 10. To reduce the risk of bacterial contamination, use 0.2% (vol/vol) Primocin in the culture medium immediately after heart extraction. When changing the medium every other day, reduce the percentage to 0.1% (vol/vol) and maintain it for the remainder of the culture duration. 11. Pipette tips may injure the epicardium, which could cause a regeneration response that may affect subsequent analyses. 12. While imaging, the plate cover should remain closed to avoid contamination. The dissecting scope surface and gloves can be cleaned with 70% ethanol prior to imaging. 13. Do not touch or blow on the explants while changing medium. Handle the culture plates gently. Do not shake the plates. 14. Other transgenic reporter lines can be utilized with partial ventricular explant cultures to probe the dynamics of

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regenerating epicardial cells: tcf21:H2A-mCherry labels epicardial cell chromatin [15] and tcf21:LifeAct-EGFP labels filamentous actin structures [15, 18]. In addition, the fluorescent ubiquitination-based cell cycle indicator (FUCCI) system [19] can be used to monitor epicardial cell proliferation and endoreplication events. Transgenic FUCCI lines tcf21:mAGzGeminin and tcf21:mKO2-zCdt1 can be used for this purpose [15].

Acknowledgments This work was supported by the American Heart Association Career Development Award (18CDA34110108) and Weill Cornell Start-up fund to J.C. References 1. Senyo SE, Steinhauser ML, Pizzimenti CL, Yang VK, Cai L, Wang M, Wu TD, GuerquinKern JL, Lechene CP, Lee RT (2013) Mammalian heart renewal by pre-existing cardiomyocytes. Nature 493(7432):433–436. https:// doi.org/10.1038/nature11682 2. Poss KD, Wilson LG, Keating MT (2002) Heart regeneration in zebrafish. Science 298 (5601):2188–2190. https://doi.org/10. 1126/science.1077857 3. Wang J, Panakova D, Kikuchi K, Holdway JE, Gemberling M, Burris JS, Singh SP, Dickson AL, Lin YF, Sabeh MK, Werdich AA, Yelon D, Macrae CA, Poss KD (2011) The regenerative capacity of zebrafish reverses cardiac failure caused by genetic cardiomyocyte depletion. Development 138(16):3421–3430. https:// doi.org/10.1242/dev.068601 4. Wang J, Cao J, Dickson AL, Poss KD (2015) Epicardial regeneration is guided by cardiac outflow tract and hedgehog signalling. Nature 522(7555):226–230. https://doi.org/10. 1038/nature14325 5. Huang GN, Thatcher JE, McAnally J, Kong Y, Qi X, Tan W, DiMaio JM, Amatruda JF, Gerard RD, Hill JA, Bassel-Duby R, Olson EN (2012) C/EBP transcription factors mediate epicardial activation during heart development and injury. Science 338(6114):1599–1603. https://doi.org/10.1126/science.1229765 6. Kikuchi K, Gupta V, Wang J, Holdway JE, Wills AA, Fang Y, Poss KD (2011) tcf21+ epicardial cells adopt non-myocardial fates during zebrafish heart development and regeneration. Development 138(14):2895–2902. https://doi.org/10.1242/dev.067041

7. Smart N, Bollini S, Dube KN, Vieira JM, Zhou B, Davidson S, Yellon D, Riegler J, Price AN, Lythgoe MF, Pu WT, Riley PR (2011) De novo cardiomyocytes from within the activated adult heart after injury. Nature 474(7353):640–644. https://doi.org/10. 1038/nature10188 8. Zhou B, Honor LB, He H, Ma Q, Oh JH, Butterfield C, Lin RZ, Melero-Martin JM, Dolmatova E, Duffy HS, Gise A, Zhou P, Hu YW, Wang G, Zhang B, Wang L, Hall JL, Moses MA, McGowan FX, Pu WT (2011) Adult mouse epicardium modulates myocardial injury by secreting paracrine factors. J Clin Invest 121(5):1894–1904. https://doi.org/ 10.1172/JCI45529 9. Smart N, Risebro CA, Melville AA, Moses K, Schwartz RJ, Chien KR, Riley PR (2007) Thymosin beta4 induces adult epicardial progenitor mobilization and neovascularization. Nature 445(7124):177–182. https://doi. org/10.1038/nature05383 10. Gemberling M, Karra R, Dickson AL, Poss KD (2015) Nrg1 is an injury-induced cardiomyocyte mitogen for the endogenous heart regeneration program in zebrafish. eLife 4. https:// doi.org/10.7554/eLife.05871 11. Lepilina A, Coon AN, Kikuchi K, Holdway JE, Roberts RW, Burns CG, Poss KD (2006) A dynamic epicardial injury response supports progenitor cell activity during zebrafish heart regeneration. Cell 127(3):607–619. doi: S0092-8674(06)01280-3 12. Wang J, Karra R, Dickson AL, Poss KD (2013) Fibronectin is deposited by injury-activated epicardial cells and is necessary for zebrafish

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heart regeneration. Dev Biol 382(2):427–435. https://doi.org/10.1016/j.ydbio.2013.08. 012 13. Wei K, Serpooshan V, Hurtado C, DiezCunado M, Zhao M, Maruyama S, Zhu W, Fajardo G, Noseda M, Nakamura K, Tian X, Liu Q, Wang A, Matsuura Y, Bushway P, Cai W, Savchenko A, Mahmoudi M, Schneider MD, van den Hoff MJ, Butte MJ, Yang PC, Walsh K, Zhou B, Bernstein D, Mercola M, RuizLozano P (2015) Epicardial FSTL1 reconstitution regenerates the adult mammalian heart. Nature 525(7570):479–485. https://doi. org/10.1038/nature15372 14. Cao J, Poss KD (2016) Explant culture of adult zebrafish hearts for epicardial regeneration studies. Nat Protoc 11(5):872–881. https:// doi.org/10.1038/nprot.2016.049 15. Cao J, Wang J, Jackman CP, Cox AH, Trembley MA, Balowski JJ, Cox BD, De Simone A, Dickson AL, Di Talia S, Small EM, Kiehart DP, Bursac N, Poss KD (2017) Tension creates an endoreplication wavefront that leads regeneration of epicardial tissue. Dev Cell 42 (6):600–615. e604. https://doi.org/10. 1016/j.devcel.2017.08.024

16. Kim J, Rubin N, Huang Y, Tuan TL, Lien CL (2012) In vitro culture of epicardial cells from adult zebrafish heart on a fibrin matrix. Nat Protoc 7(2):247–255. https://doi.org/10. 1038/nprot.2011.440 17. Habeler W, Pouillot S, Plancheron A, Puceat M, Peschanski M, Monville C (2009) An in vitro beating heart model for long-term assessment of experimental therapeutics. Cardiovasc Res 81(2):253–259. https://doi.org/ 10.1093/cvr/cvn299 18. Riedl J, Crevenna AH, Kessenbrock K, Yu JH, Neukirchen D, Bista M, Bradke F, Jenne D, Holak TA, Werb Z, Sixt M, Wedlich-Soldner R (2008) Lifeact: a versatile marker to visualize F-actin. Nat Methods 5(7):605–607. https:// doi.org/10.1038/nmeth.1220 19. Sugiyama M, Sakaue-Sawano A, Iimura T, Fukami K, Kitaguchi T, Kawakami K, Okamoto H, Higashijima S, Miyawaki A (2009) Illuminating cell-cycle progression in the developing zebrafish embryo. Proc Natl Acad Sci U S A 106(49):20812–20817. https://doi.org/10.1073/pnas.0906464106

Chapter 17 Purification of Pluripotent Stem Cell-Derived Cardiomyocytes Using CRISPR/Cas9-Mediated Integration of Fluorescent Reporters Francisco X. Galdos, Adrija K. Darsha, Sharon L. Paige, and Sean M. Wu Abstract Human induced pluripotent stem cell (hiPSC)-derived cardiomyocytes have become critically important for the detailed study of cardiac development, disease modeling, and drug screening. However, directed differentiation of hiPSCs into cardiomyocytes often results in mixed populations of cardiomyocytes and other cell types, which may confound experiments that require pure populations of cardiomyocytes. Here, we detail the use of a CRISPR/Cas9 genome editing strategy to develop cardiomyocyte-specific reporters that allow for the isolation of hiPSC-derived cardiomyocytes and chamber-specific myocytes. Moreover, we describe a cardiac differentiation protocol to derive cardiomyocytes from hiPSCs, as well as a strategy to use fluorescence-activated cell sorting to isolate pure populations of fluorescently labeled cardiomyocytes for downstream applications. Key words Cardiac differentiation, Fluorescent reporters, CRISPR/Cas9, Genome editing, Flow cytometry, hiPSC

1

Introduction With the advent of human induced pluripotent stem cells (hiPSCs), significant advances have been made toward the generation of patient-specific cell types that can be used for in vitro disease modeling and regenerative therapies [1]. Like human embryonic stem cells (hESCs), hiPSCs have the capacity to differentiate into all three embryonic germ layers and recapitulate key developmental events that give rise to all somatic cells in the body. Importantly, as iPSCs are directly derived from the somatic cells of donor patients with particular diseases of interest, they serve as powerful tools for modeling human disease without necessitating a priori knowledge of disease-causing mutations. Over the past decade, the hiPSC system has been applied for the study of a variety of human diseases

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_17, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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ranging from modeling schizophrenia using hiPSC-derived neurons to modeling cardiac arrhythmias and cardiomyopathies in hiPSC-derived cardiomyocytes [2–5]. While promising, a major limitation of the iPSC system is that current differentiation protocols generate mixed populations of cell types. In the cardiac field, extensive protocols have been generated for differentiating iPSCs into cardiomyocytes often with >80% purity [6, 7]. For example, a widely used protocol for differentiating iPSCs into cardiomyocytes by Lian et al. can result in a mixed population of fibroblasts, non-ventricular cardiomyocytes, and ventricular cardiomyocytes of varying maturity, which can confound downstream studies that rely on specific cardiomyocyte subtypes or degrees of maturation [6, 8]. Here, we detail the use of a CRISPR/Cas9 genome editing strategy as a way to develop cardiomyocyte-specific reporters that allow for the isolation of hiPSC-derived cardiomyocytes and chamber-specific myocyte subtypes for downstream studies. Fluorescent reporter systems have been extensively used to label specific cell types in animal models and have also been applied to optimize human ESC cardiac differentiations, thus providing a powerful tool for studying cardiomyocyte development and physiology in vitro [9–12]. With the development of single-cell RNA sequencing, murine and human studies have identified a growing number of chamber-specific cell surfaces, as well as intracellular markers, providing a wealth of potential candidate markers that can be used to isolate cardiomyocyte subtypes [13, 14]. These bodies of work highlight the importance of establishing a strategy that allows for rapid and specific labeling and purification of cardiomyocyte subtypes from hiPSC differentiation. In this chapter, we cover materials and methods that can be used for the design of cardiomyocyte-specific fluorescent reporters that can be integrated into the genome of hiPSCs for the isolation of pure populations of cardiomyocytes. The following methods allow for the insertion of a fluorescent reporter at the stop codon of the gene of interest separated by a P2A self-cleaving peptide such that the reporter’s expression is directly linked to the cardiac-specific marker’s gene expression. Lastly, we provide a detailed method for isolating fluorescently labeled cardiomyocyte populations by fluorescenceactivated cell sorting (FACS).

2

Materials

2.1 iPSC Culture Maintenance, Passaging, and Freezing

1. E8 media. 2. E8Y media: E8 media + 1:1000 dilution of 10 μM Y-27632 ROCK. 3. Accutase.

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4. Bambanker freezing media. 5. Matrigel media for coating culture plates: 1:350 dilution of Matrigel in DMEM/F12. For coating Matrigel plates, add 2 mL of media to each well of a 6-well plate with the Matrigel media. 2.2 SgRNA Cloning into PX458 Plasmid

1. Bacterial agar plates: Luria Broth-agar + sterile water + ampicillin (1:1000 dilution of 200 mg/mL stock). Mix all the components and microwave the mixture until completely dissolved. Carefully pour the mixture into bacterial plates and allow to cool and solidify. 2. Luria Broth: Luria Broth + sterile water + ampicillin (1:1000 dilution of 200 mg/mL stock). Microwave components until completely dissolved. We recommend an additional autoclave step to ensure long-term sterility. 3. PX458 (Addgene Plasmid #: 48138). 4. T4 PNK. 5. 10X T4 ligation buffer. 6. Nuclease-free water. 7. 10X FastDigest buffer. 8. Dithiothreitol (DTT) 10 mM stock. 9. Adenosine triphosphate (ATP) 10 mM stock. 10. FastDigest BbsI. 11. T4 DNA ligase. 12. Plasmid-Safe ATP-dependent DNase Kit. 13. Lipofectamine 3000 Kit.

2.3 Integration of Reporter Constructs into iPSCs

1. E8 and E8Y media (formulation above). 2. Blebbistatin stock solution: 10 millimolar of blebbistatin. 3. Accutase-blebbistatin solution: Prepare a 0.020% solution of blebbistatin in Accutase. For each well of a 6-well plate of iPSCs, add 0.33 uL of blebbistatin to 1.33 mL of accutase. 4. E8-blebbistatin solution: Prepare a 0.020% solution of blebbistatin in E8. For each well of a 6-well plate of iPSCs, add 1 uL of blebbistatin to 4 mL of E8. 5. E8-thiazovivin solution: 1:1000 dilution solution of 10 μM thiazovivin in E8 media. 6. P3 Primary Cell 4D-Nucleofector X Kit L (Lonza). 7. Transfection MasterMix (100 μL) that comes with P3 Primary Cell 4D Nucleofector Kit: P3 Solution, 82 μL, and Supplement 1, 18 μL. 8. PX458 with sgRNA cloned into plasmid.

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9. Donor plasmid containing the template for homology-directed repair. 10. Lonza 4D Nucleofector Machine. 11. DNA extraction kit. 12. PCR kit. 2.4 iPSC Cardiac Differentiation

1. RPMI media (with glucose). 2. RPMI media (without glucose). 3. B27 supplement. 4. B27 supplement minus insulin. 5. CHIR-99021 stock. 6. C59 stock.

2.5 FluorescenceActivated Cell Sorting of Fluorescently Labeled Cardiomyocytes

1. Thiazovivin. 2. Knockout serum or equivalent (i.e., fetal bovine serum). 3. DNase stock. 4. CM sorting media: 40% Knockout serum (or equivalent) in B27 plus insulin in RPMI media + 1:1000 10 μM of thiazovivin. 5. Stop solution: 50% Knockout serum (or equivalent) + 50% IMDM media + 1:1000 10 μM of thiazovivin + DNase (final concentration is 200 KU/mL). 6. 10X TrypLE Select. 7. 100-micron cell strainer. 8. Flow cytometer (FACSAria recommended).

3

Methods

3.1 Maintenance of Undifferentiated iPSCs

1. On the day after thawing or passaging iPSCs, aspirate E8Y, rinse the cells at least once with PBS to remove additional dead cells, and then feed with E8. Add 2 mL per well of a 6-well plate, 1 mL per well of a 12-well plate, and 10 mL for a 10 cm dish. 2. Replace E8 daily until the cells are ready to passage.

3.2 Passaging Undifferentiated iPSCs

1. Passage cells every 3–5 days, when hiPSC colonies appear to have started joining together and the plates are about 50–75% confluent. 2. Add E8Y to a conical tube (5 mL for each well of a 6-well plate).

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3. Aspirate the culture medium, rinse with PBS, and add Accutase (1 mL per well of a 6-well plate). Let the plate incubate at room temperature or 37
C for 2–5 min. 4. Monitor the plate under a microscope. When the cells start to detach, use a pipette to rinse the cells off of the dish and transfer them to the conical tube with E8Y (see Note 1). 5. Spin down the cells at 200  g for 3 min. 6. Aspirate the media and resuspend the cell pellet in 1 mL of E8Y, breaking up the pellet into a single-cell suspension. 7. Add additional E8Y to the cells depending on the dilution of the cells you want to plate. 8. Take a fresh 6-well Matrigel-coated plate (see Subheading 2.1 for Matrigel coating instructions), aspirate the excess Matrigel from the wells that will be used, and add 2 mL of E8Y to each well. 9. Pipette the cell suspension into the wells. Tilt the plate to ensure that the cells are evenly distributed within the wells. Return the plate to the 37
C incubator. 3.3 Freezing Undifferentiated iPSCs

1. Add E8Y to a conical tube (5 mL for each well of a 6-well plate). 2. Aspirate the culture medium, rinse the cells with PBS, and add Accutase. Let the cells incubate in Accutase at room temperature for 2–5 min. 3. When the cells start to detach, use a pipette to rinse the cells off of the dish and transfer the Accutase-cell solution to the conical tube with E8Y. 4. Spin the cells down at 1000 rpm for 3 min. 5. Aspirate the media and resuspend the cells in 1 mL of Bambanker freezing media. 6. Add enough Bambanker freezing media for the appropriate number of vials being frozen down. Each well of a 6-well plate can usually be frozen into two cryovials. 7. Transfer 1 mL of cells into each cryovial. Place the cryovials in a CoolCell or equivalent freezing cannister and store at 80
C to ensure control rate freezing at roughly 1
C/min. 8. After 2 h, transfer the cells to liquid nitrogen for long-term storage (see Note 2).

3.4

SgRNA Design

While several tools are available for the design of sgRNAs for CRISPR/Cas9 genome editing, we recommend free tools made available by www.Benchling.com. These allow for genomic sequences of interest to be easily downloaded and for sgRNA sequences to be designed for downstream applications based on a

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user-specified region of interest. Below we describe a step-by-step protocol for sgRNA design using Benchling.com. 1. Under the inventory tab in Benchling.com, click the plus sign button and click “Import DNA Sequences.” Several options will appear; click “Search External Databases.” 2. Search a desired gene by gene symbol (i.e., “MYL2”) or by NCBI accession number. 3. After clicking search, several options will appear to specify the genome database to be used to download the sequence of interest. For human sequences, we recommend the “GRCh38 (hg38, Homo Sapiens)” genome. Make sure to click the box indicating “Import in sense” orientation. This step will download the annotated gene of interest. 4. For sgRNA design to replace the stop codon region of a gene, navigate to the region of the gene that contains the stop codon (see Note 3). 5. Highlight a region that is 50 bp upstream and downstream on the stop codon of the gene of interest and click the “CRISPR” button on the rightmost panel of the screen (Fig. 1a).

Fig. 1 (a) Example of sgRNA design for ventricular marker gene MYL2. “TAG” represents the stop codon of the last exon of the gene. Red-highlighted region indicates the sgRNA sequence with the green PAM site labeled. (b) Sample design of the fluorescent reporter construct for integration using homology-directed repair. Left and right homology arms should flank the stop codon. After sgRNA/Cas9 cutting at the stop codon site, the design shown above will replace the stop codon with a P2A-fluorescent protein that is in frame with the rest of the gene of interest. This design tethers the expression of the fluorescent protein to that of the gene of interest. (c) To confirm successful genome editing and integration of fluorescent construct, PCR amplify both the 50 and 30 ends of the construct by designing one primer outside the homology region and its respective pair inside the homology region. This will ensure that the template being amplified is from the construct that is integrated into the desired locus rather than from residual donor plasmid that is present in the DNA sample

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6. Click “Design and Analyze Guides” and ensure that the following options are selected: “Single Guide,” “Guide Length ¼ 20,” “Genome ¼ GRCh38 (HG38, Homo Sapiens),” and “PAM ¼ NGG.” 7. Several sgRNA options will become available; we recommend selecting at least three sgRNA sequences that contain the stop codon within the sgRNA sequence (see Note 4). 3.5 Cloning and Validation of sgRNAs

The following protocol was adapted from Ran et al. (2013) and uses the PX458 (see www.addgene.org/crispr/zhang for more information) expression plasmid for cloning of sgRNAs [15, 16]. Using BbsI cloning sites, the sgRNAs designed above can be cloned into these expression vectors for downstream CRISPR genome editing applications of hiPSCs. We recommend testing three sgRNAs designed with Benchling.com to determine cutting efficiency. 1. Design two oligos where the bases indicated by the “N” region represent the sgRNA sequence obtained from Benchling.com (see Note 5): 50 -CACCGNNNNNNNNNNNNNNNNNNN-30 50 -AAACNNNNNNNNNNNNNNNNNNNC-30 2. Anneal and phosphorylate oligos using T4 phosphonucleotide kinase in T4 ligation buffer (Table 1). 3. Anneal in thermocycler using the following parameters: 37
C for 60 min and 95
C for 5 min. Ramp down to 25
C at 5
C per minute. 4. Dilute the annealed oligos 200-fold. 5. Set up digestion ligation reaction (Table 2). 6. Incubate the digestion ligation reaction using the following protocol on a thermocycler: 37
C for 5 min and 23
C for 5 min. Repeat first two steps for six cycles. Table 1 Oligo annealing and phosphorylation reaction Volume (μL)

Item

1

Oligo 1

1

Oligo 2

1

10X T4 ligation buffer

6

Nuclease-free water

1

T4 PNK

10

Total

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Table 2 SgRNA cloning into PX458 Volume (μL)

Item

X

PX458 (100 ng)

2

Phosphorylated and annealed oligos

2

10X FastDigest buffer

1

DTT (10 mM)

1

ATP (10 mM)

1

FastDigest BbsI

0.5

T4 DNA ligase

Y

Nuclease-free water

20

Total

Table 3 PlasmidSafe reaction Volume (μL)

Item

11

Ligation reaction

1.5

10X PlasmidSafe buffer

1.5

10 mM ATP

1

PlasmidSafe exonuclease

15

Total

7. Treat the ligation with PlasmidSafe exonuclease by setting up a reaction and incubating at 37
C for 1 h (Table 3). 8. Transform 1–2 uL of PlasmidSafe reaction into competent bacteria; we recommend using TOP10 competent cells. 9. Streak transformed bacteria onto agar plates with an ampicillin selection and incubate the plate overnight at 37
C. 10. The next morning, pick bacterial colonies for expansion in Luria Broth containing ampicillin. 11. Extract the plasmids using a Miniprep kit and sequence the plasmids using a U6 forward primer to confirm successful cloning of sgRNAs into PX458 plasmid. 12. To test sgRNA DNA cutting efficiency, use a Lipofectamine 3000 Kit (follow the manufacturer’s instructions) to transfect 1 ug of PX458 plasmids containing each of the cloned sgRNAs into HEK293 cells that are approximately 50% confluent.

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13. 48 h after transfection, harvest HEK293 cells for DNA extraction using a Qiagen DNeasy Kit. 14. To evaluate cutting efficiency, design PCR primers such that an ~700 bp amplicon is generated. Ensure that the forward primer is 200 bp upstream of the projected sgRNA cut site. 15. PCR amplify DNA extracted from HEK293 control cells (cell transfected without sgRNA plasmids) and the sgRNA PX458 transfected cells. 16. Run PCR products on a 1% agarose gel to ensure successful PCR amplification, and purify amplicons using a PCR purification kit. 17. Submit PCR amplicons for Sanger sequencing using the forward primer designed in step 13. 18. To assess sgRNA cutting efficiency, upload both the controland sgRNA-treated HEK293 sequencing chromatograms to https://tide.deskgen.com/. This will provide the cutting efficiency metric of each sgRNA. 19. Select the sgRNA that displays the highest cutting efficiency for downstream steps. 3.6 Design of Reporter Constructs for Isolation of PSC-Derived Cardiomyocytes

In order to isolate pluripotent stem cell-derived cardiomyocytes using fluorescent reporter constructs, we provide a walkthrough below for designing a donor plasmid that allows for the stop codon of a particular cardiac gene of interest to be replaced by a P2A sequence and fluorescent reporter sequence using CRISPR/Cas9 genome editing (Fig. 1b). This couples the expression of the fluorescent protein to the expression of the endogenous cardiac gene to be used for cardiomyocyte isolation. The P2A self-cleaving peptide separates the cardiac protein amino acid chain from the fluorescent reporter protein, thus giving rise to two independent peptides [17]. In Table 4, we list published markers for cardiomyocytes

Table 4 Selected markers for cardiomyocyte-specific fluorescent reporters Gene

Marker

TNNT2 All cardiomyocytes MLC2V Ventricular cardiomyocytes MLC2A Early stage—All cardiomyocytes; late stage—atrial cardiomyocytes SLN

Atrial cardiomyocytes

MYH6

All cardiomyocytes

NKX2.5 All cardiomyocytes and cardiac progenitor cells

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that can be used for the myocyte purification using a fluorescent reporter system [9, 13, 18–23]. 1. Identify the stop codon of the cardiac gene of interest and design left and right homology arms that are approximately 400 bp in length. Ensure that the homology arms flank the stop codon. 2. Place sequences encoding a P2A self-cleaving peptide and the coding region of a fluorescent protein of interest immediately adjacent to the left homology arm. Ensure that the P2A sequence is in frame with the cardiac gene to be targeted. 3. Add a poly-adenylation sequence 30 to the fluorescent protein of interest. 4. After the poly-A tail sequence, add an antibiotic resistance cassette that will be used for selecting pluripotent stem cell clones that have integrated the reporter construct of interest. This cassette must include a ubiquitously expressed mammalian promoter such as a Ubc, PGK, or CMV promoter, followed by an antibiotic resistance gene, such as puromycin, hygromycin, G418, or blasticidin resistance cassettes. 5. 30 to the antibiotic resistance cassette, place the right homology arm with a minimum length of 400 bp. 6. We recommend submitting plasmid designs to companies that are able to synthesize large plasmid inserts and clone into desired plasmid backbone. These companies can provide transformed bacterial stock with full sequencing information of the plasmid which cuts down on costs and time over the long run. 3.7 Integration of Reporter Constructs into iPSCs

For the integration of reporter constructs into hiPSCs, we recommend setting up a series of transfections to optimize the ratio of sgRNA/Cas9-expressing plasmid to donor plasmid. For example, we have found that 1 μg of sgRNA/Cas9 (PX458 plasmid) and 4 μg of donor plasmid give optimal viability and efficiency of genome editing. This must be determined empirically by testing different ratios of the CRISPR/sgRNA plasmid and donor plasmid. Include a donor-only control in order to use this as an antibiotic treatment control. Cells that have successfully integrated the construct will be permanently resistant to antibiotic selection, while the donor-only controls should all die after several rounds of treatment. 1. Culture the iPSCs in E8 media. Cells should be approximately ready to passage for optimal CRISPR/Cas9 genome editing (Fig. 2) (see Note 6). 2. Remove the media from the wells and add 1 mL of the Accutase-blebbistatin solution per well. 3. Incubate the plate at 37
C for 7 min.

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Fig. 2 Reporter integration. After the iPSCs have grown to 70–80% confluency, they can be combined in a cuvette with the plasmid as well as the CRISPR-Cas9 genome editing system (1) and electroporated using a nucleofector machine (2). Over the next 24 h, genome editing will take place (3). Cells can then be grown and treated with antibiotics to select for colonies that have integrated the plasmid

4. Take the plate out of the incubator and add 1 mL of the E8-blebbistatin solution on top of the Accutase-blebbistatin solution. 5. Dissociate to single cells, taking care to avoid bubbles. 6. Collect the cells in one 15 mL falcon tube. 7. Count the cells. 8. Put the cell solution in the centrifuge and spin down for 5 min at 1000 rpm. 9. Remove the supernatant and resuspend the cells in the E8-blebbistatin solution to get one million cells per mL. 10. Split the cells into individual 15 mL falcon tubes for each reaction. Each transfection reaction should have roughly one million cells. 11. Centrifuge the cells again for 3 min at 1000 rpm. 12. Aspirate the supernatant and add the transfection master mix (see Subheading 2.3). Pipette up and down gently ~5 times to get the solution into a single-cell suspension (see Note 7). 13. Pipette gently up to 110 uL per reaction into each cuvette. Tap the cuvette to prevent the formation of bubbles. 14. Turn on the nucleofector machine. Select the number of cuvettes you have. Select as many probes as you have (see Note 8). 15. Shock the cells and then immediately put 1 mL of E8-thiazovivin solution into each cuvette. 16. Let the cells sit for 10 min at room temperature prior to plating cell into 6-well plate. Failure to leave cells in the cuvette after transfection for 10 min will result in near-0% viability. 17. Pipette the cells from the cuvette into Matrigel-coated wells with 1 mL of E8-thiazovivin. Plate about 500,000 cells per well, which will roughly be half of the total volume of cells in

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the cuvette if one million cells were used for the transfection. Tilt plate to ensure an even distribution of cells (see Note 9). 18. 24 h after electroporation, rinse the cells with PBS and change the media in the plate to fresh E8. 19. 48 h after electroporation, rinse the cells with PBS and passage each well 1:12 onto a 6-well plate. Cells will be sparse when plating. Make sure to have single-cell suspensions. This will ensure that single cells plate and will allow for antibiotic selection of single cell-derived colonies that can subsequently be picked below. 20. The next day, rinse cells with PBS and begin treating cells with E8 + antibiotic (see Table 5) (see Note 10). 21. Treat with antibiotic + E8 for 4–10 days, until the donor-only transfection control has completely died. The sgRNA + donor group should now have formed individual colonies that can be picked. 3.8 Colony Picking and Clone Validation

Colony picking separates different colonies that have grown from a single cell that has incorporated the reporter construct. Thus, each colony is an individual clone and should be treated as an individual cell line (Fig. 3). 1. Fill a 12-well Matrigel-coated plate with 1 mL of E8Y in each well.

Table 5 Antibiotic selection table Antibiotic

Recommended concentration (μg/mL)

Puromycin

0.1–0.5

Hygromycin

200–400

G418

200–600

Fig. 3 Reporter validation and cell sorting. After antibiotic-treated colonies are picked and grown, each clone should be validated for the correct integration of the reporter construct into the genome by PCR and subsequent gel electrophoresis (1–2). hiPSC clones that show correct integration of the construct can be differentiated into cardiomyocytes (3) and sorted using the reporter system with a flow cytometer to obtain a pure population of fluorescently labeled cells

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2. Using a P200, gently scrape an individual colony until it is detached from the plate. Then, pipette up the colony and transfer it into one of the E8Y-filled wells on the previously prepared 12-well plate. 3. Repeat previous step 28 until 10–12 clones have been collected. Each colony is an individual clone. 4. Grow the colonies for 2–7 days in E8 in the 12-well plate. 5. When the colonies are getting big, passage them with EDTA so that each colony is dissociated and replated into a single well of a 6-well plate. 6. To passage each colony, aspirate the media in the well. 7. Rinse the colony with 1 mL of PBS. 8. Add 1 mL of EDTA and let it incubate the cells for 5–7 min either at room temperature or at 37
C. 9. While the cells are incubating in EDTA, add 1 mL of E8Y to each well of a Matrigel-coated 6-well plate. 10. When the cells begin to round up, aspirate the EDTA. 11. Add 1 mL of E8Y to the cells and pipette up and down to break up the colony into a single-cell suspension. 12. Transfer the single-cell suspension into one well of the previously prepared 6-well plate and expand the cells. 13. Grow the cells until 70% confluency is reached or until hiPSC colonies begin to merge. Add 1 mL of Accutase to each well for each clone and allow the cells to dissociate for 5 min. 14. Neutralize the Accutase dissociation with equal volume of E8Y and split; half of the well will be for freezing each clone and the other half will be for DNA collection. 15. Extract the DNA from each clone using a DNA extraction kit. 16. Design PCR primers that allow for inside-out PCR amplification that confirms integration of the desired genetic construct. One primer should bind to the region outside of the homology arm and the other primer should bind to a region inside of the homology arm of the reporter construct to be integrated. 17. PCR amplify the clone DNA for 50 and 30 construct integration using an available PCR kit (Fig. 1c). 18. Sanger sequence the 50 and 3’ PCR amplicons to confirm that no unintended mutations have been introduced during the genome editing process. 19. Confirm the heterozygous or homozygous integration of genetic constructs by conducting “outside-outside” PCR by taking the forward 50 outside primer and the reverse 30 outside primer.

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20. If selecting heterozygous integrated clones, sequence the outside-outside amplicon to confirm that no mutations were introduced by sgRNA/Cas9 cutting at the second allele of gene to be targeted. 21. Correctly edited hiPSC clones can then be selected for subsequent steps. 3.9 hiPSC Cardiac-Directed Differentiation

The following protocol is adapted from Lian et al. and uses a smallmolecule activation of the Wnt signaling pathway followed by Wnt signaling inhibition [6]. 1. Day 0 of differentiation: Add 2 mL of 4–8 μM CHIR-99021 in B27 minus insulin + RPMI media to each well of a 6-well plate containing hiPSCs or hESCs that are 85–95% confluent (see Note 11). 2. Day 2 of differentiation: Add 2 mL of B27 minus insulin in RPMI on top of the media containing CHIR. 3. Day 3 of differentiation: Change the media to 2 mL of 2 uM C59 in B27 minus insulin + RPMI media to each differentiating well. 4. Day 5 of differentiation: Change the media to 2 mL of B27 minus insulin + RPMI media to each well. 5. Day 7 of differentiation: Change the media to 2 mL of B27 plus insulin + RPMI media to each well. 6. Day 9 of differentiation: Change the media to 2 mL of B27 plus insulin + RPMI minus glucose media. Cardiomyocytes will survive on non-glucose substrates, thus allowing for enrichment of myocytes. 7. Day 10 of differentiation: Change the media to 2 mL of B27 plus insulin + RPMI media every 2 days, and glucose-starve as necessary to increase myocyte purity.

3.10 FluorescenceActivated Cell Sorting of Fluorescently Labeled Cardiomyocytes

Upon generation of a specific reporter line, the next step is to differentiate hiPSCs into cardiomyocytes and sort the CMs using FACS. Cardiomyocytes are large cells and, therefore, require special considerations when sorting. The following protocol is optimized to ensure single-cell dissociation of CMs and high viability. 1. Aspirate the culture media of CMs and rinse with PBS. 2. Add 1 mL of 10X TrypLE Select to each well of a 6-well plate and incubate at 37
C for a minimum of 8 min. Every minute, check under the microscope to ensure that CMs are dissociating. 3. When the cells start to detach from the plate, use a pipette to rinse the cells off the dish and triturate 7–8 times to ensure a single-cell suspension.

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4. Stop the TrypLE reaction with an equal volume of stop solution and transfer the cells to conical tubes. 5. Pellet the cells by centrifuging down at 300  g for 5 min. 6. Resuspend the cells in the appropriate CM sorting media to obtain a cell concentration of about 250,000 cells/mL (see Note 12). 7. Filter the cells through a 100-micron cell strainer after resuspending to ensure that the cells are in single-cell suspension. 8. Immediately after straining the cells, pipette ~2–4 mL of cell suspension into a FACS tube and place it on ice. 9. Ensure that proper negative controls are included for proper gating during the sorting process. This should consist of a CM population that is time-matched and negative for the fluorophore that was chosen for the reporter construct. 10. Using a FACSAria flow cytometer, make sure to use the largest nozzle size (130 micron in most cases) for sorting CMs. 11. When sorting CMs, sort one population at a time. For example, sort positive populations and negative populations sequentially, but not at the same time (see Note 13). 12. After sorting CMs, centrifuge down the cells and resuspend them in CM sorting media to the desired cell density. A minimum density of 200,000 cells per well of a 6-well plate will ensure high viability after sorting. 13. Change the media the day after sorting to B27 plus insulin. Cells can subsequently be used for downstream applications.

4

Notes 1. When working with hiPSCs, ensure that they are gently detached from the plate and they are in a true single suspension. This will ensure even plating of cells, which will subsequently provide the optimal conditions for successful cardiomyocyte differentiations. 2. Timely transfer of frozen cells from a 80
C freezer to liquid nitrogen will maintain good-quality hiPSCs. Leaving cells for extended periods of time (i.e., > 4 weeks) at 80
C can decrease viability and impair differentiation capacity. This is especially important to keep in mind in downstream steps for genome editing. 3. The stop codon can usually be found at the last exon of the gene of interest. Benchling.com automatically annotates an imported gene with the translation sequence that indicates

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each amino acid in the final protein. The amino acid annotation will indicate the stop codon. 4. When designing the sgRNA, ensure that it does NOT contain a BbsI cutting sequence (5’-GAAGAC-30 or 5’-GTCTTC-30 ). 5. We recommend designing the sgRNA such that the stop codon can lie at the center of the sgRNA sequence. The reason for this is to ensure that the sgRNA does not bind and cause cleavage of the donor plasmid used for homology-directed integration of reporter constructs. 6. We have found that hiPSCs with a confluency Open). 2. Zoom into a clone of interest for counting. 3. Initiate the Cell Counter (in MacOS, Plugins > Analyze > Cell Counter). 4. Remove the excess counters by clicking “Remove” in the Cell Counter window. 5. To begin counting, click “Initialize.” 6. Click on a specific Counter Type to record and append counts to that counter. 7. Record counts of individual cells by clicking on the cell to be recorded. 8. Record final counts to an Excel sheet (Fig. 4, see Note 7). Final counts are displayed on the right of the Counter. 9. Repeat steps 2–8 for all clones present within a tile scan.

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Fig. 4 Use of ImageJ Cell Counter plug-in to count the number of cells within a clone. Final cell counts are displayed in the marked red box 3.4 3-Dimensional Quantification of Clone Volumes

This section outlines the steps needed to perform quantification of clone volumes using CLARITY, light-sheet imaging, and Imaris processing. SeeNote 8 for safety precautions prior to starting.

3.4.1 Clearing of Postnatal Day 2 (P2) Mouse Hearts Using CLARITY

1. Rinse the P2 mouse hearts in PBS (3  10 min) (see Note 9). 2. Transfer the hearts to a 4% PFA solution and incubate at 4  C overnight. Repeat step 1. 3. Transfer the hearts to a 4% acrylamide solution along with 0.5% w/v of the photoinitiator 2,20 -azobis[2-(2-imidazolin-2-yl) propane]dihydrochloride and incubate overnight at 4  C. 4. To initiate polymerization, transfer hearts in the same solution to 37  C for 2–3 h. Repeat step 1. 5. Transfer the hearts to clearing solution at 37  C until cleared. This can take several hours depending on the thickness of the hearts. 6. Transfer the hearts to PBS for 1 day to remove residual SDS and store in RIMS until imaging.

3.4.2 Light-Sheet Calibration

1. Prepare the fluorescent bead sample by diluting the bead solution in RIMS with 1% low-melt-pointing agarose (1:150,000). Cut a piece of borosilicate glass tubing with an inner diameter of 6 mm and an outer diameter of 8 mm to a length of 30 mm. 2. Mix the diluted bead sample with 1% agarose and pipette the bead/agarose solution into the borosilicate tubing. Allow the agarose to solidify at room temperature (23  C).

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3. Fill the imaging chamber with a 99.5% glycerol solution. Place the borosilicate glass tubing containing the beads inside the chamber (see Note 10). 4. Attach a 3-D motorized translational stage to the borosilicate glass tubing to control the movement and orientation of the sample within the chamber. 5. Scan the beads to ensure that the thinnest light sheet covers the region of interest along the axial direction. If it does not, horizontally tune the illumination objective lens back and forth until the sharpest image appears. 6. Remove the beads from the imaging chamber. 3.4.3 Light-Sheet Imaging

1. Place the cleared heart in RIMS with a refractive index of 1.46–1.48 and 1% agarose. Insert the sample into borosilicate glass tubing and allow the agarose to solidify at room temperature (23  C) (see Note 11). 2. Attach a 3D motorized translational stage to the borosilicate glass tubing to control the movement and orientation of the heart within the chamber. 3. Position the sample so that it is in the center of the illumination beam created by the light-sheet imaging system. Acquire images using the sCMOS camera at a rate of 100 frames per second. 4. Using the motor controller, move the sample 1 μm in the axial direction and acquire images at each 1 μm increment with the sCMOS camera. Continue until the entire sample has been imaged. 5. Save the image at each increment as a TIFF file and stack all images along the axial direction in a single folder for each fluorescence channel. 6. Repeat steps 4 and 5 as needed for each fluorescence channel, making sure to change to proper filter sets and laser lines. 7. Set a pixel threshold intensity value to observe the contours of the heart and add pseudo-color to the images based on this grayscale intensity.

3.4.4 Image Reconstruction and 3D Clonal Volume Measurement Using Imaris

Familiarity with the basic functions of Imaris software is useful for this section (see Note 12). 1. If needed, remove the shadow or striping artifacts, which will interfere with accurate rendering of contiguous clone volumes, from the light-sheet images. This can be done with the ImageJ software (see Note 13). 2. To convert the light-sheet images to Imaris format, run the Imaris software and while remaining in the Arena view, run File > Batch Convert.

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3. In the Output section of the Imaris File Converter window, select the desired location for saving the Imaris files, and then click the “Add Files” button. 4. Navigate to a folder that contains the images of one fluorescence channel, select all files, and click “Open.” 5. Repeat above steps for each fluorescence channel of the lightsheet data. 6. Click “Start All” in the Imaris File Converter window to generate one Imaris file (.ims extension) for each channel. 7. When the conversion is complete, go to the Surpass view/3D view mode in Imaris to visualize and analyze the light-sheet images in 3D. From this point on, all steps to identify clones and quantify volume will be performed in Surpass/3D view. First, click the “Open” button, navigate to the location where the Imaris files were saved, and select one channel to open first. 8. To change the display color of the channel, run Edit > Show Display Adjustment, click on the name of the channel (i.e., “Channel 1”) in the Display Adjustment window, then choose a display color, and, if desired, type in a new name for this channel. 9. Run Edit > Add Channels and repeat step 8 for each channel to add the rest of the channels to the volume. 10. Volume quantification of clones is achieved by identifying distinct contiguous regions of positive signal and encapsulating each in a Surfaces object. Clones in each fluorescence color are created separately. Create a Surfaces object for the first channel by running 3D View > Surfaces or clicking the “Add New Surfaces” button in the Surpass Tree. When prompted, select the following parameters for each step in the Surfaces Creation Wizard to automatically create the Surfaces: 1/4: Use default selections (e.g., do not Skip automatic creation and do not Segment only a Region of Interest). 2/4: select a source channel; activate the Smooth option with either the automatically calculated value or a custom value for Surfaces Detail; and choose the Thresholding algorithm (see Note 14). 3/4: Set the Thresholding value and do not enable Split Touching Objects. 4/4: If needed, filter out unwanted objects that interfere with the visualization of true clones. For example, use a Number of Voxels filter to exclude small staining artifacts. Finish the Surfaces creation.

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11. Match the color of the Surfaces that were just created to the color of the fluorescence channel by selecting the Surfaces object in the Surpass Tree and clicking the “Color” tab. 12. Repeat steps 10 and 11 to create Surfaces for the rest of the channels. 13. The volume of each Surface is automatically computed when it is created. To view the volume data, select a Surfaces object in the Surpass Tree, activate the Statistics tab with the Detailed option, and choose “Specific Values” and “Volume” from the drop-down menus. Click the “Export” button to export the data to a spreadsheet for further analysis. Often it is necessary to exclude from the analysis artifacts or clones that were not delineated well. To select the subset of Surfaces that represent clones with high fidelity, activate the mouse pointer mode “Select” and click on those Surfaces while holding down the “Ctrl” or “Command” key. Rotate and zoom the scene to bring other objects into view by toggling back to the pointer mode “Navigate,” and then toggle back to “Select” to continue selecting additional Surfaces (press the “Esc” key to toggle between Navigate and Select pointer modes). Selected Surfaces should change color (e.g., yellow). Once all desired Surfaces are selected, activate the Statistics tab with the Selection option to view and export the volume data. The subset of selected Surfaces can be copied as a group to a new Surfaces object by clicking the “Duplicate Selection/Surfaces” button in the “Statistics” tab. This is useful for presentation purposes to show only the Surfaces that were analyzed and hide all others.

4

Notes 1. If you are having trouble dissolving 4-OHT in the concentrated 100 or 10 mg/mL solution, use a water bath sonicator at a low-speed setting and room-temperature water for about 30 min. 2. An alternative method for genotyping ɑMHCCreER is to use primers for the common Cre: Cre-F (50 -TGCAGGTTTTGAG CCCTAAC-30 ) and Cre-R (50 -CGAGAATGACTTCCCTG TCC-30 ). Additionally, primers for mOrange, mOrangeF (50 -CCCCGTAATGCAGAAGAAGA-30 ) and mOrangeR (50 -TCTTGGCCTTGTAGGTGGTC), or mCerulean, mCeruleanF (50 -ACGTAAACGGCCACAAGTTC-30 ) and mCeruleanR (50 -AAGTCGTGCTGCTTCATGTG-30 ), can be used to detect the mutant allele of Rainbow mice.

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3. To inject very small amounts of 4-OHT (i.e., for postnatal ɑMHCCreER;Rainbow), dilute the 10 mg/mL stock to 1 mg/ mL with corn oil in order to inject a volume > 50 μL, as there may be some leakage of 4-OHT at the needle injection site. To examine clonal expansion during embryonic development, 4-OHT should be administered at an embryonic period such as E9.5 or E12.5 and hearts harvested at a later postnatal time point, such as P1 or P2 for analysis. Labeling cells at embryonic stages requires timed mating for accuracy of developmental stage. For further information regarding timed mating of mice, please see reference [15]. 4. As complete clonal analysis of the mouse heart requires an extensive number of sections and glass slides, one option is to place two tissue sections on a slide and section through an additional 18 slices without collection. Collect the next two sections and repeat the process until the entire heart is sectioned. This enables the sampling of the entire heart at around 200 μm intervals while minimizing time and cost of supplies. 5. As the emission wavelengths of the fluorescent proteins have regions of partial overlap, the use of a confocal microscope may enable more precise control of emission detection. Using the Zeiss LSM880 Confocal, lasers 458, 488, and 594 nm can be utilized to excite mCerulean, mOrange, and mCherry, respectively. The respective excitation and emission spectrums of each of the fluorophores can also be uploaded into the Zeiss program and fine-tuned for better image quality. Note that in the Rainbow model, cells that are not recombined will express GFP; thus, clones of cells may also be initially screened as lacking GFP expression. 6. For more details on how to download and use the ImageJ Cell Counter plug-in, see reference [16]. 7. We recommend grouping the number of clusters and the number of cells within each cluster in the following manner: total number of clusters, clusters with single-labeled cells, clusters with two cells, and clusters with more than two cells (this group would contain the exact number of cells from the Cell Counter). 8. The following steps should be performed in a fume hood, as the chemicals being used are toxic. Subheading 3.4.1, step 2: Transfer the hearts to a 4% PFA solution and incubate at 4  C overnight. Subheading 3.4.1, step 3: Transfer the hearts to a 4% acrylamide solution along with 0.5% w/v of the photoinitiator 2,20 -azobis[2-(2-imidazolin-2-yl)propane]dihydrochloride and incubate overnight at 4  C.

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Subheading 3.4.1, step 5: Transfer the hearts to clearing solution at 37  C until cleared. This can take several hours depending on the thickness of the tissue. 9. Thoroughly rinse the heart in PBS to remove hemoglobin residue; otherwise strong autofluorescence will interfere with the genuine fluorescence signal. 10. Remove all of the bubbles around the heart in the tubing and chamber; otherwise stripe artifacts will degrade the image quality. 11. The borosilicate glass tubing is used to match the refractive index (1.47) for the CLARITY method. 12. To get started using Imaris, several “quick-start” tutorials are available. For a PDF document, go to reference [17]. For videos, visit the Oxford Instruments Learning Center or the YouTube channel, Imaris software. 13. One option to remove stripe artifacts from images is to process them in Fourier space with ImageJ using these steps (Fig. 5): Open an image and calculate a fast Fourier transform (Process > FFT > FFT). Draw two regions of interest (ROIs) around the clouds of intensity extending from the Y-axis in the power spectrum, without including the origin, and fill them in with black. Create a new image using the masked power spectrum (Process > FFT > Inverse FFT). To automatically remove the stripes from all of the images in a folder in this manner, use the Batch function of ImageJ (Process > Batch > Macro) with the following code: run (“FFT”); roiManager(“Select”, 0); run(“Clear”, “slice”); run (“Inverse FFT”); To use this code, first set the background color to black and the ROI Manager to contain only one element, a combined ROI containing the two ROIs used to mask the power spectrum. Use the ROI Manager button “More > OR (Combine)” to combine the two ROIs and the “Add” button to add

Fig. 5 ImageJ processing for removal of stripe artifacts. (a) Light-sheet image with stripe artifacts. (b) Power spectrum of the image in (a), calculated by fast Fourier transform with ImageJ software. (c) Power spectrum masked to remove horizontal stripe patterns. (d) Reverse FFT image calculated from the masked power spectrum in (c). Scale: 100 μm

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the new combined ROI to the list, and then delete the two original individual ROIs. 14. Thresholding is a critical step. Parameters must be set carefully to ensure that the entire clone volume is selected while keeping adjacent clones separate from each other. For a voxel size of 3.25  3.25  5 μm, we smoothed surface details to 6.50 μm. To deal with a varying intensity of background signals throughout the heart, we chose background subtraction (local contrast) with a sphere diameter of 70.0 μm. If the signal is strong and/or background is low or uniform, absolute intensity thresholding may be appropriate. The “Split Touching Objects” option should not be enabled to ensure that entire contiguous clones remain intact. It may be inevitable that some poorly formed surfaces are created due to low signal, unmanageably high background, and/or other factors. However, accurate quantification can be carried out by excluding undesired surfaces from the final analysis in step 9.

Acknowledgments This work was supported by the F31 HL144057 (N.B.N.), NIH DP2 HL127728 and UCLA Broad Stem Cell Research CenterRose Hills Foundation Research Award (R.A.). References 1. Feil R, Brocard J, Mascrez B, LeMeur M, Metzger D, Chambon P (1996) Ligandactivated site-specific recombination in mice. Proc Natl Acad Sci U S A 93 (20):10887–10890 2. Feil R, Wagner J, Metzger D, Chambon P (1997) Regulation of Cre Recombinase activity by mutated Estrogen receptor ligand-binding domains. Biochem Biophys Res Commun 237 (3):752–757 3. Feil R (2017) Conditional somatic mutagenesis in the mouse using site-specific recombinases. Handb Exp Pharmacol (178):3–28 4. Sereti K-I, Nguyen NB, Kamran P, Zhao P, Ranjbarvaziri S, Park S et al (2018) Analysis of cardiomyocyte clonal expansion during mouse heart development and injury. Nat Commun 9(1):754 5. Chung K, Wallace J, Kim S-Y, Kalyanasundaram S, Andalman AS, Davidson TJ et al (2013) Structural and molecular interrogation of intact biological systems. Nature 497(7449):332–337

6. Richardson DS, Lichtman JW (2015) Clarifying tissue clearing. Cell 162(2):246–257 7. Fei P, Lee J, Packard RRS, Sereti K-I, Xu H, Ma J et al (2016) Cardiac light-sheet fluorescent microscopy for multi-scale and rapid Imaging of architecture and function. Sci Rep 6:22489 8. Ding Y, Lee J, Ma J, Sung K, Yokota T, Singh N et al (2017) Light-sheet fluorescence imaging to localize cardiac lineage and protein distribution. Sci Rep 7:42209 9. Ding Y, Bailey Z, Messerschmidt V, Nie J, Bryant R, Rugonyi S et al (2018) Light-sheet fluorescence microscopy for the study of the murine heart. J Vis Exp (139):e57769 10. Fei P, Nie J, Lee J, Ding Y, Li S, Zhang H et al (2019) Subvoxel light-sheet microscopy for high-resolution high-throughput volumetric imaging of large biomedical specimens. Adv Photonics 1(1):1–13 11. Ding Y, Lee J, Hsu JJ, Chang C-C, Baek KI, Ranjbarvaziri S et al (2018) Light-sheet Imaging to elucidate cardiovascular injury and repair. Curr Cardiol Rep 20(5):35

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12. Ding Y, Abiri A, Abiri P, Li S, Chang C-C, Baek KI et al (2017) Integrating light-sheet imaging with virtual reality to recapitulate developmental cardiac mechanics. JCI Insight 2(22). https://doi.org/10.1172/jci.insight.97180 13. Lee J, Fei P, Packard RRS, Kang H, Xu H, Baek KI et al (2016) 4-dimensional light-sheet microscopy to elucidate shear stress modulation of cardiac trabeculation. J Clin Invest 126(5):1679–1690 14. Rasband WS (1997–2018) ImageJ. National Institutes of Health, Bethesda, MD. https:// imagej.nih.gov/ij/

15. Yeadon J. (2019) 6 steps for setting up timed pregnant mice The Jackson Laboratory. https://www.jax.org/news-and-insights/jaxblog/2014/september/six-steps-for-settingup-timed-pregnant-mice 16. Vos KD (2001–2010) Cell counter. University of Sheffield. https://imagej.nih.gov/ij/ plugins/cell-counter.html 17. BitPlane (2019) Imaris 9.3: Quick Start Tutorials United Kingdom. http://www.bitplane.com/ download/manuals/QuickStartTutorials9_3_0. pdf

Chapter 19 High-Fidelity Quantification of Cell Cycle Activity with Multi-Isotope Imaging Mass Spectrometry Frank Gyngard, Louise Trakimas, and Matthew L. Steinhauser Abstract The quantification of cell cycle activity is a prerequisite to defining the dynamics and scope of organ development or regeneration. Multi-isotope imaging mass spectrometry (MIMS) merges stable isotope tracers with an imaging mass spectrometry platform called NanoSIMS, which can quantitatively measure the incorporation of stable isotope tracers with high precision in suborganelle domains. MIMS has been applied to quantify the dynamics of postnatal cardiogenesis and mammalian cardiomyocyte regeneration during aging or in response to injury. Here, we present an approach to the conduct of MIMS experiments, with an emphasis on the application to the field of cardiac regeneration; however, the approach is also applicable, with, at most, minor modifications to broader biological questions. Key words Cell cycle activity, Imaging mass spectrometry

1

Introduction Successful heart regeneration requires the generation of new cardiac myocytes. Outside of narrow developmental windows in mammals or the hearts of lower and more regenerative model organisms, fully developed hearts display a limited capacity to regenerate cardiac myocytes via division of preexisting cardiac myocytes [1–3]. As such, methods used to track the generation of new cardiac myocytes should be able to detect rare cell cycle events with high fidelity. There are important limitations associated with existing methods of measuring cell turnover that are of particular relevance to studies of slow turnover tissues where modest absolute measurement errors captured during narrow time windows may be magnified when projected over longer and more physiologically relevant time periods. For label-free detection of cell cycle markers (e.g., Ki67, Aurora B kinase), it is challenging to convert an instantaneous staining frequency into a rate per unit of time. Inaccurate measurements may also arise when the cell cycle signal is obscured

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_19, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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by high background, as may occur with immunofluorescent detection of cell cycle markers in heart tissues that display autofluorescence. Multi-isotope imaging mass spectrometry (MIMS) is a method that can be applied to measure cell cycle activity in the animal and human tissues with high fidelity, including those that turn over slowly [3–6]. MIMS couples tracer methodology with a quantitative imaging mass spectrometry platform (NanoSIMS) that measures tracer incorporation in suborganelle domains with high accuracy and precision [7–9]. Stable isotope-enriched compounds are the most commonly used tracers for MIMS experiments. Stable isotopes are elemental variants that differ in atomic mass due to an alternative number of neutrons [9, 10]. Organisms ranging from plants to mammals can tolerate organism-wide enrichment of rare stable isotopic variants to two or more orders of magnitude above the natural concentration without evident growth inhibition of toxicity. As such, there is extensive precedent of stable isotope tracer utilization in even the most vulnerable human subjects, underscoring that they can generally be viewed as innocuous for studies in humans or model organisms [8]. In this chapter, we provide an approach to the conduct of MIMS. A complete guide to the complex operation of the NanoSIMS instrument is beyond the scope of this chapter; however, we describe critical details related to in vivo labeling, sample processing, instrumental configurations, image visualization, and data analysis. We also acknowledge common decision points that arise and describe relevant considerations to selecting the ideal methodological path to reflect the experimental goals.

2 2.1

Materials In Vivo Labeling

1.

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N-thymidine can be purchased in powder form from Cambridge Isotope Laboratories. Alternative labels for multiplexed experiments include 13C-thymidine, 2H-thymidine, or bromodeoxyuridine (Sigma or Invitrogen).

2. Make a stock solution at a concentration of 20 mg/mL in either sterile water or saline solution (e.g., 0.9% NaCl). Dissolve in water, except when delivering intravenously. Thymidine in solution is stable at room temperature; however, for prolonged storage of stock solution, store at
20  C. 3. Optional osmotic minipumps (Alzet): Because of the limited solubility of thymidine, select a pump model that delivers a sufficiently high volume per hour to achieve the target dose. 2.2

Tissue Fixation

1. Phosphate-buffered saline (PBS). 2. Dilute paraformaldehyde (Electron Microscopy Sciences) in phosphate-buffered saline (PBS) to 4%.

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1. Phosphate-buffered saline (PBS). 2. Ethanol at 100% (200 proof) or diluted in water to 50%, 70%, and 95% solutions. 3. LR white (Electron Microscopy Sciences). 4. Ethanol. 5. Small polyethylene embedding capsules (BEEM, Electron Microscopy Sciences). 6. Oven adjustable to 60  C.

2.4 Preparing Silicon Wafers

1. At 400 (~9.8 cm), cut the diameter of Si wafer (e.g., Purewafer) into square shapes that are 4.950 mm2. Si wafers of this nature can be acquired commercially from multiple retailers. Using gloves, separate these diced pieces by hand into ~75 square Si chips to place into the NanoSIMS sample holder. Non-square chips are unusable and can be discarded. 2. 600 mL Beaker or similar size. 3. Magnetic stir bar and stir plate. 4. Acetone. 5. Methanol. 6. Isopropanol. 7. Ethanol. 8. Sonicator. 9. Teflon-tipped forceps. 10. 0.2 μm Filter. 11. Petri dish. 12. Nitrogen or argon gas.

2.5 Sectioning and Preparing the Mounted Sample

1. Ultramicrotome equipped with diamond blade capable of cutting plastic resin-embedded samples to 500 nm.

2.6 NanoSIMS Analysis

1. NanoSIMS 50 or 50 L (Cameca): There are approximately 40 such instruments in the world, many of them available for collaborative analysis and/or as fee-for-service core facilities.

2. Hot plate.

2. Gold sputter coater. 2.7 Image Visualization and Data Analysis

1. For chain analysis, a script is needed to stitch adjacent images together. One such script can be accessed via https://github. com/BWHCNI/workflow/tree/master/mosaic_nrrd. 2. ImageJ software. 3. OpenMIMS plug-in to BWHCNI/OpenMIMS.

ImageJ:

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Methods In Vivo Labeling

1. Administer a nucleotide label, in vivo. Studies pertaining to cardiovascular regeneration have generally utilized 15N-thymidine; however, other stable isotope-tagged nucleotides (2H, 13 C, 18O) or nucleotide analogs (e.g., bromodeoxyuridine, iododeoxyuridine) can be used when multiplexing is desired. Due to potential toxicity with long-term administration, we advise restricting the role of nucleotide analogs to short labeling periods (e.g., 2.1. 3.1.6 Checking Quality of Tailed DNA

1. Measure the concentration of tailed DNA using a NanoDrop 2000C and adjust the concentration using nuclease-free water to 100–200 ng/μL. 2. For quality control, check the purity of the tailed DNA template product on a 1% agarose gel together with the original plasmid DNA (Fig. 3a).

3.1.7 In Vitro Transcription (IVT) Reaction

1. Prepare custom NTPS according to Table 1. The calculation is based on percentage nucleotides as below. 2. Mix reagents, at RT, in the following order to a total volume of 400 μL: 160 μL custom NTPS from Tables 1, 160 μL tailed template (50–100 ng/μL), 40 μL buffer X10 (MEGAscript kit from Ambion), and 40 μL T7 Enzyme (MEGAscript kit from Ambion). 3. Incubate for 4 h at 37  C. Table 1 Preparation of custom NTP solution for in vitro transcription reaction Name

Stock concentration (mM)

Amount (μL)

ARCA

100

100

GTP

75

36

ATP

75

183

CTP

75

183

1-M-pseudo-UTP

100

138

H2O

N/A

160

Total volume

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4. Add 12 μL of Turbo DNase (MEGAscript kit from Ambion) per 400 μL IVT reaction. Mix gently and incubate for 15 min at 37  C. 3.1.8 Purify modRNA Using MEGAclear

1. Insert a filter cartridge into the collection tube (supplied) and label it. Prepare one tube with a filter for ~0.1 mg of modRNA. 2. Preheat the nuclease-free water to 95  C. 3. Add 3.5 of binding solution concentrate to the modRNA obtained after phosphatase treatment. Mix gently by pipetting. 4. Add 2.5 of 100% ethanol to the above modRNA mixture. Mix gently by pipetting. 5. Immediately pipet ~700 μL of modRNA mixture to each filter. 6. Centrifuge for 1 min at 8944  g at 4  C. 7. Discard the flow-through and pipet 500 μL of wash solution to each filter. 8. Centrifuge for 1 min at 8944  g at 4  C. 9. Discard the flow-through and again pipet 500 μL of wash solution to each filter. 10. Centrifuge for 1 min at 8944  g at 4  C. 11. Discard the flow-through, centrifuge for 2 min at 8944  g at 4  C to dry the filter, and remove any residual wash solution. 12. Transfer filters into clean 1.5 mL collecting tubes and label them. 13. Elute RNA from the filter in clean tubes with 50 μL of pre-warmed elution solution. Apply 50 μL of the preheated elution solution to the center of the filter cartridge, close the cap of the new tube, and wait for 1 min. 14. Centrifuge for 1 min at 8944  g at 4  C. 15. Repeat elution procedure twice (total 150 μL per tube). 16. Collect all of the clean modRNA solution in each tube into one 15 mL tube (1.5 mL of clean modRNA solution). 17. Measure the concentration before proceeding to phosphatase treatment.

3.1.9 RNA Phosphatase Treatment

1. Add Antarctic phosphatase (5000 U/mL), 1/5 of the modRNA amount in the tube, and 10 Antarctic phosphatase buffer accordingly. Mix gently and incubate for 1 h at 37  C.

3.1.10 Purify modRNA Using MEGAclear

1. Insert a filter cartridge into the collection tube (supplied) and label it. Prepare one tube with a filter for ~0.1 mg of modRNA. 2. Preheat nuclease-free water to 95  C.

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3. Add 3.5 binding solution concentrate to the modRNA obtained after phosphatase treatment. Mix gently by pipetting. 4. Add 2.5 of 100% ethanol to the above modRNA mixture. Mix gently by pipetting. 5. Immediately pipet ~700 μL of modRNA mixture to each filter. 6. Centrifuge for 1 min at 8944  g at 4  C. 7. Discard the flow-through and pipet 500 μL of wash solution to each filter. 8. Centrifuge for 1 min at 8944  g at 4  C. 9. Discard the flow-through and again pipet 500 μL of wash solution to each filter. 10. Centrifuge for 1 min at 8944  g at 4  C. 11. Discard the flow-through, centrifuge for 2 min at 8944  g at 4  C to dry the filter, and remove any residual wash solution. 12. Transfer filters into clean 1.5 mL collecting tubes and label them. 13. Elute RNA from the filter into clean tubes with 50 μL of pre-warmed elution solution. Apply 50 μL of the preheated elution solution to the center of the filter cartridge, close the cap of the new tube, and wait for 1 min. 14. Centrifuge for 1 min at 8944  g at 4  C. 15. Repeat elution procedure twice (total 150 μL per tube). 16. Collect all of the clean modRNA solution in each tube and combine it into one 15 mL tube (1.5 mL of clean modRNA solution). 3.1.11 Concentrate modRNA for In Vivo Use by Amicon Ultra-4 Centrifugal Filters

1. Pre-cool the centrifuge machine to 4  C. 2. Label the tube and filter. Wash the filter with 3 mL of nucleasefree water three times to remove glycerin. One filter is good for cleaning 1 mg of modRNA. 3. Add 750 μL of water to the filter, then apply the volume containing 1 mg of modRNA from the IVT reaction to the filter, and mix well. Add nuclease-free water up to 4 mL and centrifuge at 1431  g for 45 min at 4  C until the volume reduces to ~70 μL. Remove the elute from the bottom. 4. Mix gently and collect the RNA in a different tube. 5. Measure the concentration by 1:50 dilution before checking the RNA quality.

3.1.12 Measure modRNA Quality by Agilent High-Sensitivity RNA Assay

1. Allow the high-sensitivity RNA sample buffer to equilibrate at room temperature for 30 min. 2. Thaw the RNA samples on ice.

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3. Launch the Agilent 4200 TapeStation Controller Software and, under settings, select RNA assay mode. 4. Flick the high-sensitivity RNA ScreenTape device and load it into the 4200 TapeStation instrument. 5. Place the loading tips into the Agilent 4200 TapeStation instrument. 6. Vortex the reagents and spin down before use. 7. For each sample, pipette 1 μL of high-sensitivity RNA sample buffer and 2 μL of RNA sample into a tube strip. Apply caps to tube strips with samples. 8. Mix liquids in sample vials using the vortex at 358  g for 1 min. 9. Spin down to position the sample at the bottom of the tube strip. 10. For sample denaturation, heat the samples to 72  C for 3 min. Place the samples on ice for 2 min. Spin down to position the samples at the bottom of the tube strip. 11. For sample analysis, load samples into the Agilent 4200 TapeStation instrument. Carefully remove the tube strip caps. Use the electronic ladder for comparison. Select the required sample positions on the 4200 TapeStation Controller Software. Click Start. The Agilent TapeStation Analysis Software opens after the run and displays results. 3.2 modRNA Preparation for In Vitro or In Vivo Delivery, Detection, and Analysis of modRNA Translation 3.2.1 In Vitro Transfection

1. Prepare the transfection mixture according to the manufacturer’s protocol. 2. Culture the cells in the DMEM medium containing 10% fetal bovine serum (FBS) and Anti-anti. 3. Using a 24-well plate with 2.5 μg of Luc modRNA (120 polyA or 173 polyA) in each well of a 6-well plate with 10 μg of nGFP modRNA (120 polyA or 173 polyA) in each well, transfect the modRNA neonatal rat CMs using the transfection reagent JetPEI (Polyplus). 4. 24 h Post-transfection, image the cells and measure the expression levels in IVIS. Collect the cell lysates and analyze by Western blot.

3.2.2 In Vivo Delivery

1. Deliver Luc modRNA (25 μg) in a total volume of 60 μL in TB buffer via direct injection to the myocardium in an open-chest surgery. 2. The sucrose-citrate buffer contains 20 μL of sucrose in nuclease-free water (0.3 g/mL) and 20 μL citrate (0.1 M, pH 7; Sigma) mixed with 20 μL of modRNA.

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3. The transfection mixture is directly injected (2–3 individual injections, 20 μL each) into the quadriceps femoris muscle or heart muscle. 3.2.3 Detecting Luciferase Expression Using the IVIS System

1. Either the transfected cells (24 h) or injected mice are bioluminescently imaged at different time points (24–144 h) in IVIS system. 2. To visualize cells expressing Luc in vitro, luciferin is added to cell culture plate, which was imaged in IVIS system. 3. To visualize tissues expressing Luc in vivo, mice are anesthetized with isoflurane (Abbott Laboratories), and luciferin (150 μg/g body weight; Sigma) is injected intraperitoneally. 4. Mice are imaged using an IVIS imaging system every 2 min until the Luc signal reaches a plateau. 5. Imaging data are analyzed and quantified with Living Image software.

4

Notes 1. To generate modRNA with different poly A tail lengths, we use two different reverse primers. 2. modRNA mixed to TB buffer can be stored at 80  C to be injected at a later date. 3. Transfection mixture can be injected via/using/in 2–3 individual injections of 20 μL each into the heart muscle. 4. modRNA should be aliquoted into smaller volumes. 5. For use in vivo, modRNA concentration should not be more than 10 μg/μL.

Acknowledgments We thank Nadia Hossain for her assistance. This work was funded in part by a seed package from the Icahn School of Medicine at Mount Sinai. Nishat Sultana and Mohammad Tofael Kabir Sharkar contributed equally to this work. References 1. Go AS et al (2014) Heart disease and stroke statistics—2014 update: a report from the American Heart Association. Circulation 129 (3):e28–e292 2. Dargie H (2005) Heart failure post-myocardial infarction: a review of the issues. Heart 91 (Suppl 2):ii3–ii6. discussion ii31, ii43-8

3. Magadum A, Kaur K, Zangi L (2019) mRNAbased protein replacement therapy for the heart. Mol Ther 27(4):785–793 4. Sahin U, Kariko K, Tureci O (2014) mRNAbased therapeutics—developing a new class of drugs. Nat Rev Drug Discov 13(10):759–780

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5. McIvor RS (2011) Therapeutic delivery of mRNA: the medium is the message. Mol Ther 19(5):822–823 6. Kariko K et al (2008) Incorporation of pseudouridine into mRNA yields superior nonimmunogenic vector with increased translational capacity and biological stability. Mol Ther 16 (11):1833–1840 7. Zangi L et al (2013) Modified mRNA directs the fate of heart progenitor cells and induces vascular regeneration after myocardial infarction. Nat Biotechnol 31(10):898–907 8. Magadum A et al (2018) Ablation of a single N-glycosylation site in human FSTL 1 induces cardiomyocyte proliferation and cardiac regeneration. Mol Ther Nucleic Acids 13:133–143 9. Sultana N et al (2017) Optimizing cardiac delivery of modified mRNA. Mol Ther 25 (6):1306–1315 10. Heiser A et al (2002) Autologous dendritic cells transfected with prostate-specific antigen RNA stimulate CTL responses against metastatic prostate tumors. J Clin Invest 109 (3):409–417 11. Morse MA et al (2002) The feasibility and safety of immunotherapy with dendritic cells loaded with CEA mRNA following neoadjuvant chemoradiotherapy and resection of pancreatic cancer. Int J Gastrointest Cancer 32 (1):1–6 12. Morse MA et al (2003) Immunotherapy with autologous, human dendritic cells transfected with carcinoembryonic antigen mRNA. Cancer Investig 21(3):341–349 13. Rittig SM et al (2011) Intradermal vaccinations with RNA coding for TAA generate CD8+ and CD4+ immune responses and induce clinical benefit in vaccinated patients. Mol Ther 19 (5):990–999 14. Su Z et al (2005) Telomerase mRNAtransfected dendritic cells stimulate antigenspecific CD8+ and CD4+ T cell responses in patients with metastatic prostate cancer. J Immunol 174(6):3798–3807

15. Weide B et al (2009) Direct injection of protamine-protected mRNA: results of a phase 1/2 vaccination trial in metastatic melanoma patients. J Immunother 32(5):498–507 16. Wilgenhof S et al (2013) A phase IB study on intravenous synthetic mRNA electroporated dendritic cell immunotherapy in pretreated advanced melanoma patients. Ann Oncol 24 (10):2686–2693 17. Creusot RJ et al (2010) A short pulse of IL-4 delivered by DCs electroporated with modified mRNA can both prevent and treat autoimmune diabetes in NOD mice. Mol Ther 18 (12):2112–2120 18. Mitchell DA et al (2008) Selective modification of antigen-specific T cells by RNA electroporation. Hum Gene Ther 19(5):511–521 19. Okumura K et al (2008) Bax mRNA therapy using cationic liposomes for human malignant melanoma. J Gene Med 10(8):910–917 20. Wang Y et al (2013) Systemic delivery of modified mRNA encoding herpes simplex virus 1 thymidine kinase for targeted cancer gene therapy. Mol Ther 21(2):358–367 21. Kariko K et al (2012) Increased erythropoiesis in mice injected with submicrogram quantities of pseudouridine-containing mRNA encoding erythropoietin. Mol Ther 20(5):948–953 22. Kormann MS et al (2011) Expression of therapeutic proteins after delivery of chemically modified mRNA in mice. Nat Biotechnol 29 (2):154–157 23. Mays LE et al (2013) Modified Foxp3 mRNA protects against asthma through an IL-10dependent mechanism. J Clin Invest 123 (3):1216–1228 24. Zimmermann O et al (2012) Successful use of mRNA-nucleofection for overexpression of interleukin-10 in murine monocytes/macrophages for anti-inflammatory therapy in a murine model of autoimmune myocarditis. J Am Heart Assoc 1(6):e003293 25. Kondrat J, Sultana N, Zangi L (2017) Synthesis of modified mRNA for myocardial delivery. Methods Mol Biol 1521:127–138

Chapter 22 Generation and Manipulation of Exosomes Shiqi Hu, Li Qiao, and Ke Cheng Abstract Exosomes are membrane-bound nano-vehicles shed by most eukaryotic cells. Exosomes contain specific proteins and RNAs from parent cells, and they play key signaling roles in cellular development, modulation, and tissue regeneration. Attempts to isolate and modify exosomes to increase their targeting efficiency to specific tissue are still in their infancy. Here, we describe generation of exosomes from biopsy, isolation of exosomes by centrifugal ultrafiltration method, and approaches for manipulation of cardiac homing exosomes by chemical engineering for the treatment of myocardial infarction. Key words Exosome generation, Exosome manipulation, Myocardial infarction, Cardiac homing peptide, Platelet membrane, Intravenous injection

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Introduction There are several principal methods for isolating exosomes: (1) differential centrifugation [1], (2) size exclusion (e.g., ultrafiltration [2] or chromatography [3]), (3) immunological separation (e.g., antibody-bead capture [4]), and (4) polymer-based precipitation [5]. The merits of using different methods also vary with the different origin of exosomes, from cell-conditioned media, urine, or plasma. Based on the purpose of studies, purity (i.e., protein-toexosome ratio, with the goal of minimizing non-exosome proteins that may be present in the biofluids), yield, or quality (preservation of exosome integrity and function), we should take advantage of the merits with different methods. In this chapter, we demonstrate the ultrafiltration centrifugation method due to its satisfactory yield and purity for exosomes from conditioned media. Exosomes from healthy cardiac spheroid-derived cells have the capacity to reduce myocardial infarction (MI)-related scar formation, increase cardiomyocyte proliferation, reduce fibrosis, support cardioprotection, and modulate inflammation [6]. However, the biodistribution of exosomes was not desired due to the low heart retention and accumulation through simple intravenous

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_22, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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(IV) injection. Targeting exosomes can be generated both by surface display (genetic modification of the patent cells) and by cloaking (chemical modification). We focus on chemical manipulation and describe two membrane engineering methodologies to directly embed exosome surfaces ex vivo with an anchor conjugated with cardiac homing peptide (CHP) or with injurytargeting platelet membranes [2]. CHP has been screened out through phage display techniques according to reports [7, 8]. Platelets can target the lesion region of the ischemic heart after IV administration [9–14], which provides a modular platform where any desired antibodies, peptides, and even partial foreign membranes can be introduced to facilitate engineered exosome uptake in tissues of interest.

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Materials Prepare all solutions using distilled water (prepared by purifying deionized water) and analytical grade reagents.

2.1 Exosome Generation

1. Cell culture medium (seeNote 1). 2. Fetal bovine serum (FBS) (seeNote 2). 3. Phosphate-buffered solution (PBS, 1
, pH 7.4) (seeNote 3). 4. T-75 flasks (ultralow attachment) (seeNote 4). 5. T-175 flask (seeNote 5). 6. Fibronectin (seeNote 6). 7. Collagenase solution (seeNote 7). 8. TrypLE Select (seeNote 8). 9. 0.22 μm Steriflip filters (seeNote 9). 10. 100 kDa Centrifugal filter tubes (seeNote 10).

2.2 Exosome Manipulation: Cloaking Ischemic Myocardium Homing Peptide

1. Cardiac homing peptide (CHP; CSRSMLKAC) (seeNote 11). 2. Dioleoylphosphatidylethanolamine N-poly (ethylene glycol)5000-hydroxysuccinimide (DOPE-PEG-NHS). 3. 3000 Da Centrifugal filter tubes. 4. 100 kDa Centrifugal filter tubes. 5. PBS, 1
, pH 7.4.

2.3 Exosome Manipulation: Cloaking Platelet Membrane

1. Platelet-rich plasma (seeNote 12). 2. HEP buffer (HEP refers to HEPES in the buffer): 140 mM of NaCl, 2.7 mM KCl, 3.8 mM of HEPES, 5 mM of EGTA, pH 7.4. Before use, warm up to room temperature and adjust pH if necessary.

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3. Platelet wash buffer (10 mM of sodium citrate, 150 mM of NaCl, 1 mM of EDTA, 1% (w/v) dextrose, pH 7.4). Before use, warm up to room temperature and adjust pH if necessary. 4. 100 kDa Centrifugal filter tubes. 5. Lipophilic green fluorescent dye (DiO) (DiOC18(3) (3,30 -dioctadecyloxacarbocyanine perchlorate) (seeNote 13). 6. CD41a FITC-conjugated antibodies (seeNote 14).

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Methods Exosomes should always be placed on ice during the chemical engineering process.

3.1 Exosome Generation

1. Cut the cardiac sample, place a 6 mm piece of tissue in a sterile 100
20 mm plate, add 5 mL of PBS, and then cut sample into 2 mm pieces. 2. Wash with 5 mL of PBS 3 times, and then incubate samples in 2 mL of collagenase solution at 37  C for 5 min. 3. Add 2 mL of 20% FBS media, and then cut the samples with a blade using a rolling method until you have 50 samples roughly 0.5 mm in size. 4. Coat a 150
20 mm plate with fibronectin in 37  C for 30 min, remove the fibronectin solution, and wash the fibronectin-coated dish with 10 mL of PBS. Remove PBS, and add 5 mL of 20% FBS media to the plate. 5. Transfer the samples to the fibronectin-coated plate. Pieces should be 1.5 cm apart. 6. Add 2 mL of 20% FBS media to the fibronectin plate, and then slowly transfer to a 37  C incubator for 12 h. 7. Add 10 mL of 20% FBS media to the plate. Slowly add to the edge of the plate to reduce tissue disturbance. Place the plate back into the 37  C incubator and change the media every 2 days until cells are confluent. 8. Remove the media from the plate and wash with PBS. Add 10 mL of TrypLE Select in the plate and incubate at 37  C for 5 min. Once cells are detached, add 10 mL of 20% media to the plate. Collect the cells into a 50 mL conical tube. Add 5 mL of 20% FBS media to wash the plate and collect it to the same tube. Centrifuge at 410
g for 5 min. Remove the supernatant and keep the pellet. 9. Resuspend the pellet in 1 mL of 10% FBS media, and add the resuspended solution to the ultralow attachment T-75 flask at a concentration of 10,000 cells per 1 cm2. Place the flask into the 37  C incubator for 1 week to make cardiac sphere cells.

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Fig. 1 (a) Illustration of the cell culture process. (b) Derivation and culture of cardiosphere-derived cells (CDCs) (reprinted with permission from [15]. Copyright © Creative Commons Attribution License 4.0)

10. Transfer the cells from the ultralow flask to a new fibronectin coated flask. Bring the flask up to a level with 20% FBS media. Place the flask in the 37  C incubator. The cardiac spheres will attach to the flask and grow out to form cardiosphere-derived cells (CDCs, Fig. 1) [15]. 11. Exosomes are isolated from the conditioned medium of these cardiac cells. Passage 1–3 cardiac sphere-derived cells were used. 12. Once the cells are 80% confluent, switch the medium to a serum-free medium and condition for another 3 days. 13. Isolate the exosomes from the conditioned medium by ultrafiltration. Filter the conditioned medium through 0.22 μm Steriflip filters to remove cellular debris and large vesicles. Add the filtrate to Amicon Ultra-15, 100 kDa filters and centrifuge at 3500
g for 20 min. Discard the flow-through. 14. Collect and wash the concentrated exosomes with PBS three times and store them at 80  C. 15. Nanoparticle tracking analysis (NanoSight) and transmission electron microscopy (TEM) (Fig. 2) showed that exosomes have a typical size (mode diameter ~ 100 nm) and concentration (109 particles/mL) found in the literature [6]. 3.2 Exosome Manipulation: Cloaking Ischemic Myocardium Homing Peptide

1. Combine DOPE-PEG5000-NHS (dioleoylphosphatidylethanolamine poly (ethylene glycol)5000 N-hydroxysuccinimide; Mn  5000, 10 mg) and CHP peptide (Mn ¼ 943.2, 20 mg in 1 mL DMSO) with a tenfold molar excess of peptide. Dissolve DOPE-NHS and CHP in 1 mL of PBS and allow them to react for 1 h to form the DOPE-CHP (seeNote 15).

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Fig. 2 (a) NanoSight analysis and (b) TEM images of exosomes (adapted with permission from [6]. Copyright © 2019 American Society for Clinical Investigation)

2. Purify the DOPE-CHP using 3000 Da MWCO centrifugal filter tubes with PBS. Wash twice to remove free CHP and discard the flow-through. 3. Incubate the DOPE-peptide with the exosomes with a lipid: exosome ratio of 6000:1 (i.e., 6000 molecules of DOPEpeptide for each exosome) (seeNote 16) for 10 min at 37  C. For example, 1013 exosomes need 0.6 mg DOPE-CHP (seeNote 17), and tenfold more DOPE-CHP (6 mg) should be used to enhance the reaction speed and efficiency of this step (seeNote 18). 4. Place the solution from step 1 in a 100 kDa ultracentrifugation tube and wash with PBS twice (10 mL
2) at 8  C. See Note 19 to get rid of extra DOPE-CHP . 5. Remove excess reagents through a 100 kDa centrifugal filter and adjust to the final concentration using PBS. 3.3 Exosome Manipulation: Cloaking Platelet Membrane

1. Add HEP buffer to the platelet-rich plasma (PRP) at a 1:1 ratio (v/v). Include prostaglandin E1 (PGE1, 1 μM final concentration) to prevent platelet activation. Mix very gently by inverting the tube slowly (seeNote 20). 2. Spin at 100
g for 15–20 min at room temperature (with no brake applied) to the pellet contaminating red and white blood cells. Transfer the supernatant into new plastic tube using a transfer pipette. 3. Pellet the platelets by centrifugation at 800
g for 15–20 min at room temperature (with no brake applied). Discard the supernatant. Rinse the platelet pellet with the platelet wash buffer (without resuspension in order to avoid unnecessary platelet activation) by gently adding the wash buffer and removing it slowly with a pipette. Repeat once.

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4. Obtain the platelet membrane by three freeze-thaw cycles. Sonicate for 5 min. Evenly distribute the platelet membranes. The solution may be a little white. 5. Determine the protein content in the purified platelet membrane by the BCA protein assay for further cloaking. 6. Cloak the platelet membrane onto the surface of exosomes. Incubate 0.5 mL of exosomes (1 mg protein) with 0.5 mL of platelet membrane containing 0.5 mg membrane protein. Extrude 15 times using an extruder (Avanti Polar Lipids, Inc) with a 200 nm membrane. 7. Characterize the engineered exosomes and compare them with the original exosomes by using a NanoSight NS300 to determine the size and total number. Use TEM to see the morphology, use Western blotting to detect the protein profiles, and use miRNA array to check the miRNA cargos (seeNote 21). 8. Label the exosomes and platelet membrane to calculate the cloak efficiency. To label with lipophilic green fluorescent dye (DiO), incubate purified exosomes (109 particles) with Fast DiO green fluorescent membrane dye at a final concentration of 2 μg/mL for 1 h at room temperature. Dilute the labeled exosomes with PBS to 15 mL, and then purify and concentrate them by centrifugation through a 10 kDa ultracentrifugation tube at 3000
g for 10 min to remove unbound dye and extra PBS. Repeat the purification process of washing and ultracentrifugation two times before resuspending the labeled exosome pellet in PBS. 9. Incubate platelet membranes with CD41a FITC-conjugated antibodies according to the manufacturer’s directions. The purification and condensation process is similar to what is described in step 7. 10. Fuse labeled exosomes and platelets for flow cytometry analysis to calculate the cloak efficiency (seeNote 22).

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Notes 1. Cell culture media for different types of cells were selected and prepared according to the literature. In this study, Iscove’s modified Dulbecco’s media (IMDM) was used as the basal media. 2. For cell isolation from biopsy, we used 20% FBS media since cells were vulnerable during this process. For general cell culture, we used 10% FBS media if no further description.

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3. 1
PBS was prepared by dissolving a phosphate-buffered saline tablet in distilled water according to the manufacturer’s instructions. 4. Ultralow attachment flasks were used to form spheroids. 5. General flasks were used for the growth and expansion of monolayer cells. 6. Fibronectin, as a widely distributed glycoprotein, was used as a substrate to promote the attachment of cells. Here, plates were incubated with 0.025 mg/mL of fibronectin in distilled water at 37  C for at least 30 min before using. 7. Add 10 mL of PBS to the bottle of collagenase (from Clostridium histolyticum, sterile-filtered, release of physiologically active rat hepatocytes tested, Type IV-S, 0.5–5.0 FALGPA units/mg solid, 125 CDU/mg solid, 50 mg) and transfer it to a 50 mL tube. Adjust the volume to 50 mL to make a 1 mg/ mL collagenase solution. Then, keep the collagenase solution on ice in the culture hood. 8. TrypLE Select is room-temperature stable and ready to use. 9. 0.22 μm Steriflip filters were used to remove all cell debris and microvesicles. We have found that there is no need to centrifugate cell debris. One can collect the media with exosomes and proteins in one step. 10. 100 kDa Centrifugal filters were used to remove proteins and other small molecular components in the media. One can collect pure exosomes easily after washing twice with PBS. 11. Cardiac homing peptides are synthesized by a company by providing the sequences. 12. Platelet-rich plasma can be collected from mice and rats, and commercial platelet-rich plasma from humans was available and affordable. One can process up to 200 mL of plasma at one time and obtain abundant platelet membranes with consistency. 13. Lipophilic fluorescent dyes are widely applied for exosome staining. 14. CD41a is a typical marker of platelet membranes. For this application, one must choose a different fluorescent color from the lipophilic dye in Note 13. 15. In our previous study, DOPE-NHS, without PEG, was used. PEG serves as a spacer and reduces the steric hindrance, which can enhance the flexibility of conjugated CHP and the targeting efficiency in vivo.

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16. The ration of lipid to exosomes is 6000:1 (i.e., 6000 molecules of DOPE-peptide for each exosome) according to our previous study. Peptide-to-exosome labeling ratio: Assuming an exosome radium of 90 nm the surface area of a single exosome can be calculated: r exosome ¼ 90 nm Asurface ¼ 4πr 2 A exosome ¼ 101788 nm2 In addition, the length of a peptide bond can be assumed as follows to give a predicted length of the CHP peptide as l peptide bond ¼ 3:5 Å or 0:35 nm l CHP ¼ 9  l peptide bond Assuming that the peptide occupies a circular shaped area, the maximum amount of space that can be occupied is calculated using the maximum hexagonal packing efficiency: 1 pffiffiffi ηhexagonal ¼ 3π 6 A paked ¼ A exosome  ηhexagonal Finally, it can be assumed that the peptide will only rarely be tangential to the surface of the exosome, but likely in a nearvertical position. To account for this, the occupied radius is l CHP ffiffiffi r CHP ¼ p 2 A single peptide ¼ r CHP2  π npeptides per exosome ¼

A packed ¼ 5922:61  6000 A single peptide

17. The weight of DOPE-CHP needed is shown by an example. For example, 1013 exosomes are going to be cloaked by DOPE-CHP. There are 6.022
1023 particles of exosomes per mole. The mole of DOPE-CHP needed is M DOPEXHP ¼ 6000


1013  107 mol 6:022
1023

The molecular weight of DOPE-CHP is Mw DOPECHP ¼ 5000 þ 943:2 ¼ 5943:2 g=mol The weight of DOPE-CHP needed is

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Fig. 3 Schematic of cloaking process to generate myocardium-targeting exosomes. First, DOPE-NHS was reacted with cardiac homing peptide (CHP) to generate the phospholipid membrane anchor. The lipophilic tails of the DOPE-CHP then spontaneously insert into the exosomal membrane

W DOPECHP ¼ M ∗Mw ¼ 107
5943:2  0:6 mg 18. More DOPE-CHP can enhance the insertion efficiency and reduce the incubation time. The insertion rate depends on the membrane fluidity, which mainly depends on incubation temperature. Exosomes should always be on ice during the whole process if not otherwise specified. Thus, to reduce the exposure time of exosomes at 37  C, tenfold DOPE-CHP was added to enhance efficiency. The process is demonstrated in Fig. 3. 19. We found that when the temperature of centrifuge is set at 4  C, the membrane of ultracentrifuge tube was inclined to freeze and exosomes block the membrane. We assume that low temperature would induce more mechanical injury to the morphology and membrane of exosomes after centrifugation. 8  C is preferred for this step. 20. Platelet responses to thrombin are mediated by proteaseactivated receptors (PAR). Thrombin binds to the extracellular domain of PAR-1 and PAR-4. Both PAR-1 and -4 activation must be inhibited to prevent platelet activation. 21. Extrusion over polycarbonate membranes alters the morphology of vesicles. The size of engineered exosomes is more evenly distributed after extrusion. On the other hand, surface charge and protein profiles are similar before and after, indicating no alteration in exosome stability. In addition, miRNA cargos of exosomes before and after show no significant difference. The size of platelet membranes is uneven, and visible turbidity can be seen. The membrane we used is 200 nm, which alters the size of platelet membranes and cloaks the platelet membranes onto exosomes during extrusion. The size of the exosomes is around 100 nm before the manipulation, which does not change after the process. The process is demonstrated in Fig. 4. 22. The double-positive area indicates successfully cloaked particles, while the single-positive areas indicate unfused platelet

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Fig. 4 Schematic illustration of cloaking process to generate platelet membrane-fused exosomes. Nanoscale pieces of platelet membranes were generated by ultrasonication, and then fused with exosomes by extrusion

membranes and exosomes, respectively. The coating efficiency is calculated accordingly: E¼

Double positive%
100% Single positive ðexosomes þ platelet membranesÞ% þ double positive%

Acknowledgments This work was supported by grants from the National Institutes of Health to Dr. Ke Cheng (R01 HL123920, HL137093, HL144002, and HL146153) and the American Heart Association (18TPA34230092 and 19EIA34660286). References 1. Momen-Heravi F, Balaj L, Alian S, Mantel PY, Halleck AE, Trachtenberg AJ, Soria CE, Oquin S, Bonebreak CM, Saracoglu E, Skog J, Kuo WP (2013) Current methods for the isolation of extracellular vesicles. Biol Chem 394(10):1253–1262. https://doi.org/ 10.1515/hsz-2013-0141 2. Vandergriff A, Huang K, Shen DL, Hu SQ, Hensley MT, Caranasos TG, Qian L, Cheng K (2018) Targeting regenerative exosomes to myocardial infarction using cardiac homing peptide. Theranostics 8(7):1869–1878. https://doi.org/10.7150/thno.20524 3. Boing AN, van der Pol E, Grootemaat AE, Coumans FA, Sturk A, Nieuwland R (2014) Single-step isolation of extracellular vesicles by size-exclusion chromatography. J Extracell Ves 3. https://doi.org/10.3402/jev.v3.23430 4. Koliha N, Wiencek Y, Heider U, Jungst C, Kladt N, Krauthauser S, Johnston IC, Bosio A, Schauss A, Wild S (2016) A novel multiplex bead-based platform highlights the diversity of extracellular vesicles. J Extracell

Ves 5:29975. https://doi.org/10.3402/jev. v5.29975 5. Martins TS, Catita J, Rosa IM, Silva OABDE, Henriques AG (2018) Exosome isolation from distinct biofluids using precipitation and column-based approaches. PLos One 13(6): e0198820. https://doi.org/10.1371/journal. pone.0198820 6. Qiao L, Hu SQ, Liu SY, Zhang H, Ma H, Huang K, Li ZH, Su T, Vandergrif A, Tang JN, Allen T, Dinh PU, Cores J, Yin Q, Li YJ, Cheng K (2019) microRNA-21-5p dysregulation in exosomes derived from heart failure patients impairs regenerative potential. J Clin Invest 129(6):2237–2250. https://doi.org/ 10.1172/Jci123135 7. Kanki S, Jaalouk DE, Lee S, Yu AYC, Gannon J, Lee RT (2011) Identification of targeting peptides for ischemic myocardium by in vivo phage display. J Mol Cell Cardiol 50(5):841–848. https://doi.org/10.1016/j. yjmcc.2011.02.003 8. Won YW, McGinn AN, Lee M, Bull DA, Kim SW (2013) Targeted gene delivery to ischemic

Generation and Manipulation of Exosomes myocardium by homing peptide-guided polymeric carrier. Mol Pharm 10(1):378–385. https://doi.org/10.1021/mp300500y 9. Su T, Huang K, Ma H, Liang H, Dinh P-U, Chen J, Shen D, Allen TA, Qiao L, Li Z, Hu S, Cores J, Frame BN, Young AT, Yin Q, Liu J, Qian L, Caranasos TG, Brudno Y, Ligler FS, Cheng K (2019) Platelet-inspired nanocells for targeted heart repair after ischemia/reperfusion injury. Adv Funct Mater 29(4):1803567. https://doi.org/10.1002/adfm.201803567 10. Li ZH, Hu SQ, Cheng K (2018) Platelets and their biomimetics for regenerative medicine and cancer therapies. J Mater Chem B 6 (45):7354–7365. https://doi.org/10.1039/ c8tb02301h 11. Junnan T, Teng S, Ke H, Phuong-Uyen D, Zegen W, Adam V, Michael TH, Jhon C, Tyler A, Taosheng L, Erin S, Emily M, Leonard JL, Laura R, Alex L, Ashley B, Thomas GC, Deliang S, George AS, Zhen G, Jinying Z, Ke C (2018) Targeted repair of heart injury by

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stem cells fused with platelet nanovesicles. Nature Biomedical Engineering 2(1):17–26 12. Zhenhua L, Deliang S, Shiqi H, Teng S, Ke H, Feiran L, Lei H, Ke C (2018) Pretargeting and Bioorthogonal Click Chemistry-Mediated Endogenous Stem Cell Homing for Heart Repair. ACS Nano 12(12):12193–12200 13. Zhenhua L, Shiqi H, Ke H, Teng S, Jhon C, Ke C (2020) Targeted anti–IL-1β platelet microparticles for cardiac detoxing and repair. Science Advances 6(6):eaay0589 14. Deliang S, Zhenhua L, Shiqi H, Ke H, Teng S, Hongxia L, Feiran L, Ke C (2019) AntibodyArmed Platelets for the Regenerative Targeting of Endogenous Stem Cells. Nano Letters 19 (3):1883–1891 15. Michael TH, James de A, Bruce K, Kathryn M, Junnan T, Zegen W, Thomas GC, Jorge P, Tao-Sheng L, Ke C (2015) Cardiac regenerative potential of cardiosphere-derived cells from adult dog hearts. Journal of Cellular and Molecular Medicine 19(8):1805–1813

Chapter 23 Epigenetic Assays in Purified Cardiomyocyte Nuclei Matthew C. Hill and James F. Martin Abstract The adult mammalian heart’s potential for regeneration is very inefficient. Importantly, adult mammalian cardiomyocytes (CMs) are characterized as a cell population with very limited mitotic potential. Conversely, the neonatal mouse heart possesses a brief, yet robust, regenerative capacity within the first week of life. Cell type-specific enrichment procedures are essential for characterizing the full spectrum of epigenomic landscapes and gene regulatory networks deployed by mammalian CMs. In this chapter, we describe a protocol useful for purifying CM nuclei from mammalian cardiac tissue. Furthermore, we detail a low-input procedure suitable for the parallel genome-wide profiling of chromatin accessibility, histone modifications, and transcription factor-binding sites. The CM nuclei purified using this process are suitable for multi-omic profiling approaches. Key words Nuclei isolation, Density gradient centrifugation, Immunopurification, Epigenomics, Histones, Transcription factors, Chromatin, Cardiomyocytes, Regeneration

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Introduction The heart is comprised of several diverse cell types that together work to maintain proper organ function. However, this multicellularity makes bulk tissue-based epigenomic datasets difficult to untangle and interpret. Among the distinct cardiac cell populations, mammalian cardiomyocytes (CMs) are particularly difficult to isolate given their massive size and intractability. Indeed, a single healthy CM can measure over 100 μm long. Further, in response to stress, organ dysfunction, and disease, CMs often increase in size through hypertrophy [1]. Fluorescence-activated cell sorting (FACS) of such large cells requires nontraditional large-particle FACS instruments [2]. Laser capture microdissection and differential plating are useful purification methodologies; however, they are not well suited for quickly isolating CMs and may cause unwanted epigenomic perturbations. Overall, it is technically challenging to obtain pure populations of CMs for performing in vivo epigenetic studies without expensive and elaborate instruments.

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_23, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Tools for genetically labeling a cell type of interest have proven useful for investigating tissue heterogeneity in model organisms [3]. Among these technologies, several elegant approaches have emerged that rely on nuclei isolation techniques, including INTACT (isolation of nuclei tagged in specific cell types), BiTS-ChIP (batch isolation of tissue-specific chromatin for immunoprecipitation), and FANS (fluorescence-activated nuclear sorting) [4–7]. Importantly, working with isolated nuclei is an ideal approach for CM-specific epigenomics, as it eliminates the need to handle whole CMs. Technologies like INTACT rely on the cell type-specific expression of a nuclear envelope-localized genetic tag, which is easily affinity purified [4]. Such procedures require extensive breeding schemes and cannot be applied to human tissue samples. However, studies have found that CM nuclei are coated with PCM1 (pericentriolar material 1) and that this expression pattern can be exploited for CM enrichment [8– 10]. Therefore, to carry out CM-specific epigenomic profiling for the purposes of answering questions pertaining to heart regeneration, heart failure, and arrhythmogenesis, we have adopted a PCM1-based magnetic assisted nuclei sorting approach. Affinity-purified CM nuclei are ideal for use with a multitude of epigenomic and transcriptomic assays, including low-input epigenomic profiling technologies, like the assay for transposase-accessible chromatin using sequencing (ATAC-seq), ChIPmentation, and CUT&RUN [11–13]. CM nuclei can also be used as input for chromosome conformation capture (3C) techniques, including 3C, 4C, HiC, and capture HiC [14]. The approach outlined in this chapter allows for the isolation of both human and mouse CM nuclei from fresh, frozen, and fixed tissue samples with a single commercially available antibody. Additionally, the chapter details how to profile chromatin accessibility, histone modifications, and transcription factor binding in a single experiment (Fig. 1). The workflow detailed here is ideal for use with patient-derived cardiac tissue samples.

Fig. 1 Overview of the workflow for isolating cardiomyocyte nuclei as input for epigenomic profiling. First, cardiac tissue is processed and then nuclei are purified via density gradient centrifugation. Next, a PCM1based magnetic affinity purification step is performed to enrich for cardiomyocyte nuclei. Finally, enriched cardiomyocyte nuclei are used as input for epigenomic profiling (e.g., ATAC-seq, ChIP-Seq, Methyl-seq, HiChIP)

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Materials Unless explicitly stated otherwise, make all buffers with ultrapure DNase- and RNase-free water.

2.1 Cardiomyocyte Nuclei Isolation

1. 7 mL Dounce homogenizer. 2. Magnetic Eppendorf stand. 3. 32 mL thick-wall polycarbonate ultracentrifuge tubes. 4. Swinging bucket ultracentrifuge rotor. 5. Refrigerated ultracentrifuge. 6. End-to-end rotator, kept at 4
C. 7. Barrier pipette tips, RNase/DNase-free. 8. PCM1 antibody (Sigma HPA023370). 9. Cell strainer, 40 μm. 10. Cell strainer, 20 μm. 11. Tissue homogenizer (electric). 12. Dithiothreitol stock solution (DTT; 1000): 1 M DTT in water. Store aliquots at 20
C. 13. Spermine tetrahydrochloride stock solution (spermine; 1000): 0.15 M Spermine in water. Store aliquots at 20
C. 14. Spermidine trihydrochloride (spermidine; 1000x): 0.5 M Spermidine. Store aliquots at 20
C. 15. Tricine stock solution (tricine-KOH): 1 M Tricine in water, add concentrated KOH until a pH of 7.8 is achieved. 16. Buffer HB: 0.25 M Sucrose, 25 mM KCl, 5 mM MgCl2, 20 mM tricine-KOH, pH 7.8. Filter, sterilize, and store at 4
C. 17. Wash buffer (WB): 0.4% IGEPAL CA-630 in buffer HB. Filter, sterilize, and store at 4
C. 18. 5% IGEPAL CA-630: 5% IGEPAL CA-630 in buffer HB. Filter, sterilize, and store at 4
C. 19. Diluent buffer: 150 mM KCl, 30 mM MgCl2, 120 mM tricineKOH, pH 7.8. Filter, sterilize, and store at 4
C. 20. Working solution: Mix 5 volumes of OptiPrep Density Gradient Medium (Sigma, D1556) with 1 volume of diluent buffer. Make fresh and keep on ice. 21. DAPI. 22. Hemocytometer. 23. Epifluorescence microscope.

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ATAC-Seq

1. Wash buffer (ATAC wash buffer): 10 mM Tris–HCl pH 7.4, 10 mM NaCl, 3 mM MgCl2, 0.1% Tween-20. Store at 4
C. 2. Thermomixer. 3. TD buffer (TD, 2): 20 mM Tris–HCl pH 7.6, 10 mM MgCl2, 20% dimethyl formamide (DMF). Adjust pH to 7.6 with glacial acetic acid prior to addition of DMF. 4. Transposition buffer (Tn5, 1): 25 μL of 2 TD buffer, 2.5 μL Tn5 transposase (Illumina, 15027865), 16.5 μL PBS, 0.5 μL 1% digitonin, 0.5 μL 10% Tween-20, 5 μL H2O. Make fresh 50 μL reaction per individual experiment/sample. 5. DNA purification kit. 6. Custom primers: Ad1 AATGATACGGCGACCACCGAGATCTACACTC GTCGGCAGCGTCAGATGTG; Ad2.1 TAAGGCGACAAGCAGAAGACGGCATACGAG ATTCGCCTTAGTCTCGTGGGCTCGGAGATGT; Ad2.2 CGTACTAGCAAGCAGAAGA CGGCATACGAG ATCTAGTACG GTCTCGTGGGCTCGGAGATGT; Ad2.3 AGGCAGAACAAGCAGAAGA CGGCATACGAG ATTTCTGCCTGT CTCGTGGGCTCGGAGATGT; Ad2.4 TCCTGAGCCAAGCAGAAGACGG CATACGAG ATGCTCAGGAGTCTCG TGGGCTCGGAGATGT; Ad2.5 GGACTCCTCAAGCAGAAGA CGGCATACGAG ATAGGAGT CCGTCTCGTGGGCTCGGAGATGT; Ad2.6 TAGGCATGCAAGCAGAAGAC GGCATACGAG ATCATGCCTA GTCTCGTGGGCTCGGAGATGT; Ad2.7 CTCTCTACCAAGCAGAAGA CGGCATACGAG ATGTAGAGAGGTCTCGTGGGCTCGGAGATGT; Ad2.8 CAGAGAGGCAAGCAGAAGACGGCATACGA GATCCTCTCTGGTCTCGTGGGCTCGGAGATGT; Ad2.9 GCTACGCTCAAGCAGAAGAC GGCATACGAG ATAGCGTAGCG TCTCGTGGGCTCGGAGATGT; Ad2.10 CGAGGCTGCAAGCAGAAGACGGCATACG AGATCAGCCTCGGTCTCGTGGGCTCGGAGATGT; Ad2.11 AAGAGGCACAAGCAGAAGACGGCATACGA GATTGCCTCTTGTCTCGTGGGCTCGGAGATGT; Ad2.12 GTAGAGGACAAGCAGAAGACGGCATACGA GATTCCTCTACGTCTCGTGGGCTCGGAGATGT; Ad2.13 GTCGTGATCAAGCAGAAGACGGCATACGA GATATCACGACGTCTCGTGGGCTCGGAGATGT; Ad2.14 ACCACTGTCAAGCAGAAGACGGCATACG AGATACAGTGGTGTCTCGTGGGCTCGGAGATGT; Ad2.15 TGGATCTGCAAGCAGAAGACGGCATACGA GATCAGATCCAGTCTCGTGGGCTCGGAGATGT; Ad2.16 CCGTTTGTCAAGCAGAAGACGGCATACGA GATACAAACGGGTCTCGTGGGCTCGGAGATGT;

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Ad2.17 TGCTGGGTCAAGCAGAAGACGGCATACGA GATACCCAGCAGTCTCGTGGGCTCGGAGATGT; Ad2.18 GAGGGGTTCAAGCAGAAGACGGCATACGA GATAACCCCTCGTCTCGTGGGCTCGGAGATGT; Ad2.19 AGGTTGGGCAAGCAGAAGACGGCATACGA GATCCCAACCTGTCTCGTGGGCTCGGAGATGT; Ad2.20 GTGTGGTGCAAGCAGAAGACGGCATACGA GATCACCACACGTCTCGTGGGCTCGGAGATGT; Ad2.21 TGGGTTTCCAAGCAGAAGACGGCATACGA GATGAAACCCAGTCTCGTGGGCTCGGAGATGT; Ad2.22 TGGTCACACAAGCAGAAGACGGCATACGA GATTGTGACCAGTCTCGTGGGCTCGGAGATGT; Ad2.23 TTGACCCTCAAGCAGAAGACGGCATACGA GATAGGGTCAAGTCTCGTGGGCTCGGAGATGT; Ad2.24 CCACTCCTCAAGCAGAAGACGGCATACGA GATAGGAGTGGGTCTCGTGGGCTCGGAGATGT. 7. Amplification master mix: 25 μL of 2 Q5® Hot Start HighFidelity Master Mix, 2.5 μL of 25 μM primer Ad1 (universal), 2.5 μL of 25 μM primer Ad2 (index). 8. Thermocycler. 9. Real-time PCR instrument (qPCR) and consumables. 10. qPCR master mix: 4.5 μL Nuclease-free H2O, 0.25 μL 25 μM primer Ad1, 0.25 μL 25 μM primer Ad2, and 10 μL KAPA SYBR FAST qPCR 2 master mix. 11. Qubit fluorometer and dsDNA HS Assay Kit. 12. Fragment analyzer instrument (AATI) and NGS fragment kit (1–6000 bp). 13. EB elution buffer (Qiagen). 14. Agencourt AMPure XP beads. 2.3 CUT&RUN and High-Throughput Sequencing

1. CUT&RUN wash buffer: Combined 1 mL of 1 M HEPES pH 7.5, 1.5 mL NaCl, 50 μL of 0.5 M spermidine, and bring the total volume to 50 mL with ultrapure H2O. Add a single EDTA-free protease inhibitor tablet and vortex thoroughly. Make fresh. 2. Digitonin wash buffer (Dig-wash): Combine 200 μL of a 5% digitonin solution with 40 mL of CUT&RUN wash buffer. Make fresh. 3. Antibody buffer: Add 8 μL of 0.5 M EDTA with 2 mL of the freshly prepared Dig-wash buffer. Make a separate aliquot for each antibody to be used and add antibody at a 1:100 concentration unless a manufacturer or experimenter recommends differently. Vortex to mix. Store on ice.

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4. Stop solution (STOP, 2): Add 340 μL of 5 M NaCl, 200 μL 0.5 M EDTA, 100 μL 0.2 M EGTA, 20 μL 5% digitonin, 25 μL RNase A (Thermo Fisher, EN0531), 125 μL glycogen (2 mg/ mL), and 2 pg/mL of heterologous spike-in DNA (e.g., yeast DNA provided by Dr. Steven Henikoff). Make fresh. 5. Antibodies for targeting transcription factors, and/or histone modifications of interest. 6. Protein A-MNase fusion protein (e.g., aliquot provided by Dr. Steven Henikoff). 7. Calcium stock solution: 100 mM CaCl2. 8. DNA extraction reagents: Phenol:chloroform:isoamyl alcohol (25:24:1, v/v), 10% SDS stock solution, proteinase K (20 mg/ mL), chloroform, 2 mg/mL glycogen, 100% ethanol, and phase-lock microcentrifuge tubes (Qiagen, 129046). 9. EB elution buffer. 10. Metal block for 1.5 mL tubes. 11. Refrigerated centrifuge. 12. Thermomixer. 13. Thermocycler. 14. Qubit fluorometer and dsDNA HS Assay Kit. 15. Fragment analyzer instrument (AATI) and NGS fragment kit (1–6000 bp). 16. KAPA Hyper prep kit and KAPA single-indexed adapters. 17. Agencourt AMPure XP beads. 18. Illumina sequencing instrument (e.g., NextSeq 500).

3

Methods The entire procedure from cardiac tissue processing to sequenceready ATAC-seq, and CUT&RUN libraries take approximately 3 days. The major steps in the protocol are as follows: 1. Cardiac tissue collection or thawing. 2. Cardiac tissue homogenization. 3. Density gradient centrifugation to enrich for cardiac nuclei. 4. PCM1-based magnetic affinity purification of CM nuclei. 5. Quantification of CM nuclei purity and yield. 6. ATAC-seq. 7. CUT&RUN. 8. Library preparation. 9. High-throughput sequencing and analysis.

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The following protocol for nuclei purification is primarily derived from Mo et al. [14]. It is important that samples be kept on ice and that all vessels and instruments used to store CM nuclei be pre-chilled. The following volumes are for processing a single sample: 1. Prepare all buffers and solutions for tissue processing and density gradient centrifugation (see Note 1). First, make fresh working solution, and then prepare the density gradient solutions. For the 30% OptiPrep solution, mix 4.5 mL of working solution with 3 mL of buffer HB. For the 40% OptiPrep solution, mix 3.2 mL of the working solution with 0.8 mL of buffer HB. Aliquot 50 mL of WB, 10 mL of buffer HB, and 1 mL of 5% IGEPAL CA-630. Thaw aliquots of DTT, spermine, and spermidine stock solutions. Add a protease inhibitor tablet to the 10 mL aliquot of buffer HB. Finally, add appropriate amounts of DTT, spermine, and spermidine to all buffers to achieve 1x concentration (seeNote 2). All buffers with DTT, spermine, and spermidine added are designated with “++” (e.g., HB++). 2. Collect cardiac tissue and place it in a petri dish containing 8 mL of buffer HB (no additives) and mince the tissue into small ~1 mm  ~1 mm sized chunks using sterile curved scissors. 3. Using a wide-bore P1000 tip, collect the minced cardiac tissue and place it into an Eppendorf tube. Let the tissue settle and remove as much liquid as possible. Next, add 1 mL of buffer HB++. 4. Homogenize the tissue on ice with an electric tissue homogenizer (Biogen PRO200) until chunks are no longer visible (see Note 3). 5. Transfer the homogenized sample to a pre-chilled 7 mL Dounce homogenizer and place on ice. Add 4 mL of buffer HB++ and homogenize the tissue with the loose pestle for 30 strokes. Next, carefully homogenize with the tight pestle for 30 strokes. Add 320 μL of 5% IGEPAL CA-630++ and continue homogenizing with the tight pestle (see Note 4). 6. Run the sample through a pre-wetted 40 μm cell strainer placed in a 50 mL tube. Add 5 mL of working solution to the sample (total volume is now ~10 mL). 7. Pour the sample into pre-chilled polycarbonate ultracentrifuge tubes. 8. Underlay the homogenized cardiac tissue-containing solution with the 30% OptiPrep solution and then the 40% OptiPrep solution (from step 1).

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9. Weigh the sample and make a balance with water. To adjust the weight of a sample, make a 1:1 mixture of leftover working solution and buffer HB. 10. Place balanced samples in the pre-chilled rotor and spin at 10,000  g for 18 min at 4
C. Use a slow break speed (see Note 5). 3.2 Magnetic Immunoprecipitation of Cardiomyocyte Nuclei

1. While the samples are spinning, aliquot 80 μL of Protein G Dynabeads (per sample) into a 1.5 mL Eppendorf tube and wash two times with 800 μL of ice-cold WB++. After the second wash, resuspend the beads in 80 μL of WB++. Transfer 20 μL of these beads into a new 1.5 mL Eppendorf tube, labeled “Preclear.” 2. When the spin is finished, vacuum aspirate the top 25% OptiPrep layer and about 2/3 of the 30% OptiPrep layer. Next, using a P1000, transfer 1.5–1.6 mL of the sample band at the 30–40% interface into the “Preclear” tube and mix well. Endto-end rotate the preclear sample at 4
C for 15 min (see Note 6). 3. While the sample is preclearing, prepare two 1.5 mL Eppendorf tubes (tubes “Ab#1” and “Ab#2”) with 2 μL of anti-PCM1 antibody (see Note 7) and 400 μL of wash buffer++. 4. Place the magnetic rack in ice. 5. Place precleared sample on the magnet for 5 min. Transfer the supernatant (this is “supernatant #1”) from each sample into a clean 1.5 mL tube and place on the magnet. Resuspend the dry precleared beads in 400 μL of WB++ and place back on the magnet. Remove supernatant and place in a new tube (this is “supernatant #2”) on the magnet. 6. Transfer ~800 μL from “supernatant #1” and 200 μL from “supernatant #2” into each of the 400 μL antibody/WB++ tubes (tubes “Ab#1” and “Ab#2”). End-to-end rotate at 4
C for 45 min. 7. Mix the remaining 60 μL of washed protein-G beads and add an equal amount of these beads (~30 μL) to the two tubes containing lysate and antibody (tubes “Ab#1” and “Ab#2”) from the previous step. End-to-end rotate at 4
C for 25 min. 8. Place the samples on the magnet for no longer than 1 min. Take samples off of the magnet. Rock tubes back and forth to resuspend beads. Repeat seven times (see Note 8). 9. Place the Eppendorf tubes with the sample on the magnet for 4 min. 10. Remove the supernatant, and then add 500 μL mL of WB++. Remove from the magnet and let it sit on ice for ~30 s.

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11. Carefully pipet up and down to mix the bead-bound nuclei. 12. Pre-wet a 20 μm CellTrics filter with ~300 μL of WB++, shake off any liquid, and transfer the filter to a labeled 1.5 mL Eppendorf tube per sample. Run 500 μL of “Ab#1” and “Ab#2” through the single filter to combine the tubes. 13. Remove the filters after ensuring that they have drained into the tubes completely, shake the edges off, close the tubes, and transfer them to the magnet. 14. After the solution has cleared, remove the supernatant. Next, keeping the beads on the magnet, add 1 mL of WB++. Repeat this wash procedure six times (see Note 9). 15. Resuspend the washed bead-bound CM nuclei in 100–1000 μL WB++ (see Note 10), remove it from the magnet, and place on ice. 16. Stain a small aliquot of nuclei with DAPI and quantify with a hemocytometer (see Note 11). 3.3 CM-Specific ATAC-Seq

The following ATAC-seq protocol is derived from methodologies described previously [11, 15, 16]. 1. Aliquot ~50,000 bead-bound CM nuclei into a fresh 1.5 mL Eppendorf tube. Place on the magnet until the solution clears, and then remove the supernatant. 2. Add 1 mL of ATAC wash buffer and let it sit on the magnet for ~30 s. 3. Remove the supernatant and place the tube on ice. Next, add 50 μL of the Tn5 transposition buffer to bead, and pipet mix carefully. 4. Place resuspended the sample in a thermomixer set at 37
C with 1000 rpm mixing for 30 min (see Note 12). 5. Purify the tagmented DNA with the Zymo DNA Clean and Concentrator-5 kit (see Note 13). Elute DNA in 21 μL of EB. This is a good stopping point. The purified DNA can be stored at 20
C for several weeks.

3.4 ATAC-Seq Library Preparation and Quality Control

1. Begin the pre-amplification by adding the 20 μL of transposed DNA to the amplification master mix. Mix thoroughly and spin down briefly. Next, place the sample in a thermocycler and carry out the following PCR program: 72
C for 5 min, 98
C for 30 s, then 5 cycles of 98
C for 10 s, 63
C for 30 s, and 72
C for 1 min; then hold at 4
C. 2. Remove the sample from the thermocycler and place it on ice. 3. To determine the number of necessary additional PCR cycles, set up the qPCR reaction (see Note 14). Start by adding 5 μL of the pre-amplified sample to 15 μL of the qPCR master mix in a qPCR tube strip.

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4. Perform the qPCR amplification with the following program: 95
C for 3 min, then 20 cycles of 95
C for 10 s, 63
C for 30 s, and 72
C for 1 min. 5. To calculate the additional number of cycles needed, plot linear Rn versus cycle and determine the cycle number that corresponds to 1/3 of the maximum fluorescent intensity. Typically, the number of additional cycles required is 2–4. 6. Amplify the remaining 45 μL of pre-amplified product on a thermocycler for the number of cycles determined via qPCR. 7. Purify the amplified ATAC-seq library by adding 81 μL of AMPure XP beads to the sample (1.8 bead to sample ratio). Wash the sample as directed by the manufacturer (using fresh 80% ethanol). Elute in 20 μL of EB buffer. 8. Determine the concentration of the ATAC-seq library with the Qubit quantification system. This concentration will allow you to determine the correct Fragment Analyzer NGS kit to select. 9. Run a Fragment Analyzer lane with 2 μL of the library according to the manufacturer’s instructions (Fig. 2a). 10. The library is ready for Illumina sequencing (Fig. 2b). For standard interpretation of chromatin accessibility, ~25–50 million paired-end reads are sufficient. For footprinting, sequence at a much higher depth (>150 million reads). 3.5

CUT&RUN

The following CUT&RUN protocol is derived from Skene et al. (2017) [13]. 1. Aliquot 100–250,000 nuclei into a fresh 1.5 mL Eppendorf tube (see Note 15). 2. Place the tube on the magnetic stand and remove supernatant.

Fig. 2 Quality control and expected results from CM-specific ATAC-seq. (a) An example of a fragment analyzer results from a good-quality cardiomyocyte-enriched ATAC-seq library. (b) Example genome browser tracks showing the CM-enriched ATAC-seq results from three individual cardiac tissue samples

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3. Add 1 mL of CUT&RUN wash buffer. Let it sit for 30 s. 4. Remove the supernatant. 5. Add 50 μL of antibody buffer to the assay tube and carefully resuspend the bead-bound CM nuclei. 6. Incubate overnight at 4 (see Note 16).




C with end-over-end rotation

7. If the primary antibody is rabbit or guinea pig, then skip to step 9. If not, then place the sample on the magnet stand. Remove the supernatant. Add 50 μL of antibody buffer containing the appropriate secondary antibody (raised in rabbit or guinea pig), and carefully mix the sample. 8. End-to-end rotate the sample for 15 min at room temperature. 9. Place the sample on the magnetic stand, and then remove the supernatant. 10. Wash with 1 mL of Dig-wash buffer (mix by inversion or careful pipetting). 11. Place the sample on the magnetic stand and remove the clear supernatant. Next, remove from the magnet. 12. Add 50 μL of Dig-wash buffer, and then add 2.5 μL of a 1:10 dilution of the protein A-MNase fusion protein stock. Mix carefully to resuspend the CM nuclei. 13. End-to-end rotate the sample for 10 min at room temperature. 14. Place on the magnetic stand and remove the supernatant. 15. Add 1 mL of Dig-wash buffer and mix carefully. 16. Place the sample on the magnetic stand and remove the supernatant. 17. Add 1 mL of Dig-wash buffer and mix carefully. 18. Place on the magnetic stand and remove the clear supernatant. 19. Add 100 μL of Dig-wash buffer to the sample and mix carefully to resuspend the nuclei. 20. Place the tube in a 1.5 mL heat block resting in wet ice, and incubate until the sample reaches 0
C. 21. Remove the cooled sample from the block and add 2 μL of the 100 mM CaCl2 stock solution. Mix carefully and immediately place back into the 0
C block. 22. Incubate for 30–45 min. 23. To terminate the reaction, add 100 μL of STOP solution and mix carefully. 24. Incubate for 10 min at 37
C in a thermomixer. 25. Centrifuge the sample at 16,000  g for 5 min at 4
C.

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26. Decant the supernatant cleanly from the pellet and transfer to a fresh 1.5 mL microcentrifuge tube. 3.6 CUT&RUN DNA Extraction

1. Add 2 μL of 10% SDS and 2.5 μL of proteinase K to each sample tube. Mix by inversion and incubate for 10 min at 70
C. 2. Add 300 μL of phenol:chloroform:isoamyl alcohol (PCI) and mix by vortexing. 3. Transfer the sample to a prepared phase-lock tube, and, with a centrifuge set, spin down the sample for 5 min at 16,000  g at room temperature. 4. Add 300 μL of chloroform, invert ten times to mix, and spin down again at room temperature for 5 min at 16,000  g. 5. Remove the aqueous liquid (top layer) and carefully transfer it to a fresh 1.5 mL tube containing 2 μL of 2 mg/mL glycogen. 6. Add 750 μL of 100% ethanol and mix by vortexing. 7. Chill at 20
C for 20 min, and then centrifuge for 10 min at 4
C at 16,000  g. 8. Carefully pour off the liquid and drain on a paper towel. 9. Rinse the pellet in 1 mL of 100% ethanol, and then centrifuge for 5 min at 4
C at 16,000  g. 10. Carefully pour off the liquid and drain on a paper towel. Air-dry the pellet until all residual ethanol has evaporated (see Note 17). 11. Resuspend the DNA pellet in 32 μL of buffer EB. 12. Determine the concentration of the sample with the Qubit system. This information is important for determining the concentration of adapters to use during library preparation.

3.7 CUT&RUN Library Preparation

1. Use the remaining volume (~30 μL) of DNA as input for library preparation with the KAPA Hyper Prep Kit. 2. Perform end repair and A-tailing, adapter ligation, and all cleanup steps according to the manufacturer’s instructions. 3. For library amplification, use the following two-step PCR program: 98
C for 45 s, 14 cycles of 98
C for 15 s, 60
C for 10 s, then a single hold at 72
C for 1 min, and finally hold at 4
C. 4. After AMPure XP purification, elute the library in 20 μL of buffer EB. 5. Determine the concentration of the CUT&RUN library with the Qubit quantification system. This concentration will allow you to determine the correct Fragment Analyzer NGS kit to select.

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Fig. 3 Quality control and expected results from CM-specific CUT&RUN. (a) An example of a fragment analyzer result from a good-quality cardiomyocyte-enriched H3K27ac CUT&RUN library. (b) Example genome browser tracks showing the CM-enriched CUT&RUN and ATAC-seq results derived from a single cardiac tissue sample

6. Run a Fragment Analyzer lane with 2 μL of the library according to the manufacturer’s instructions (Fig. 3a). 7. The library is ready for paired-end Illumina sequencing (see Note 18) (Fig. 3b).

4

Notes 1. It is best to start the protocol in the morning. Having several containers full of ice makes organizing all of the buffers, homogenizers, and tubes much easier. 2. If collecting RNA, then add RNase inhibitors to buffers. For native ChIP-seq, add Na butyrate. 3. Take care not to overhomogenize the sample with the mechanical homogenizer. A low-medium speed is adequate. Keep the probe clean and cold. Resting the probe inside a 15 mL Falcon tube full of HB buffer in ice is preferable. Alternatively, preform this step in a cold room. 4. To monitor the progress of the homogenization, take 10–20 μL of the sample from the Dounce homogenizer and place on a microscope slide. Add 1 μL of DAPI, and then visualize with an epifluorescence microscope. Ideally, the overwhelming majority of nuclei should be free in solution. If clumps of vessels and CMs are still evident, then add more strokes with the tight pestle. 5. For a SW 32 Ti, this will be 9024 RPM. For a SW 28, this will be 8693 RPM. 6. You can take a larger volume of nuclei from the interface as long as you scale up the other reagents appropriately (e.g., the number of tubes containing antibody and WB++ and washed protein-G beads).

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7. A PCM1 antibody concentration of ~1.5–4 μg/mL works well. This concentration may need to be optimized to adjust for differences in lot numbers. 8. This procedure increases the number of beads bound to each nucleus. 9. If you possess a large magnetic stand, then the wash steps can be done in a large-volume vessel (e.g., 10–15 mL) with a greater volume of WB++. 10. Resuspend bead-bound nuclei in a volume commensurate with the amount of input tissue. Typically, for a 50 mg piece of heart tissue, the final pellet of bead-bound CM nuclei is resuspended in 500 μL. 11. It is often best to dilute the nuclei (~1:50) prior to quantification with a standard hemocytometer. 12. This is a good time, during the 30-min transposition reaction, to process your remaining nuclei for CUT&RUN, and/or RNA isolation. 13. 250 μL of the provided DNA-binding buffer works well. The Qiagen MinElute PCR Purification kit (28004) can be substituted for the Zymo kit here. 14. For the qPCR reaction, alternative master mix recipes may be used if the KAPA qPCR master mix is not available (e.g., adding SYBR Green stock to reaction containing the pre-amplification master mix components) [16, 17]. 15. Approximately 50,000 nuclei is a good starting point. 16. After all wash steps and incubations, it is best to spin down the sample(s) with a quick low-speed pulse (~100 g) on a microcentrifuge. 17. After you have poured off the ethanol, you can spin down the sample at 16,000  g for 1 min and then carefully remove any excess ethanol with a p10 pipet. Be very careful not to touch the pellet. 18. The Illumina NextSeq 500 75 cycle kit comes with enough reagents to complete 92 cycles. A 43  43 bp sequencing run with a 6 bp index performs well for up to 24 libraries (~16 million reads per library).

Acknowledgment We would like to thank Drs. Alisa Mo and Jeremy Nathans for providing the original nuclear isolation protocol. We thank Dr. Steven Henikoff for providing CUT&RUN reagents. This work was supported by grants from the National Institutes of

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Health (DE023177, HL127717, HL130804, HL118761 (J.F. M.)), F31HL136065 (M.C.H.), Vivian L. Smith Foundation (J.F.M.), State of Texas funding (J.F.M.), and Fondation LeDucq Transatlantic Networks of Excellence in Cardiovascular Research (14CVD01) “Defining the genomic topology of atrial fibrillation” (J.F.M.). References 1. Go¨ktepe S, Abilez OJ, Parker KK et al (2010) A multiscale model for eccentric and concentric cardiac growth through sarcomerogenesis. J Theor Biol 265:433–442 2. Lo´pez J, Sharma J, Avila J et al (2017) Novel large-particle FACS purification of adult ventricular myocytes reveals accumulation of myosin and actin disproportionate to cell size and proteome in normal post-weaning development. J Mol Cell Cardiol 111:114–122 3. Handley A, Schauer T, Ladurner AG et al (2015) Designing cell-type-specific genomewide experiments. Mol Cell 58:621–631 4. Deal RB, Henikoff S (2010) A simple method for gene expression and chromatin profiling of individual cell types within a tissue. Dev Cell 18:1030–1040 5. Deal RB, Henikoff S (2011) The INTACT method for cell type-specific gene expression and chromatin profiling in Arabidopsis thaliana. Nat Protoc 6:56–68 6. Bonn S, Zinzen RP, Perez-Gonzalez A et al (2012) Cell type-specific chromatin immunoprecipitation from multicellular complex samples using BiTS-ChIP. Nat Protoc 7:978–994 7. Haenni S, Ji Z, Hoque M et al (2012) Analysis of C. elegans intestinal gene expression and polyadenylation by fluorescence-activated nuclei sorting and 30 -end-seq. Nucleic Acids Res 40:6304–6318 8. Bergmann O, Jovinge S (2012) Isolation of cardiomyocyte nuclei from post-mortem tissue. J Vis Exp 10(65):e4205

9. Preissl S, Schwaderer M, Raulf A et al (2015) Deciphering the epigenetic code of cardiac myocyte transcription. Circ Res 117:413–423 10. Gilsbach R, Preissl S, Gru¨ning BAA et al (2014) Dynamic DNA methylation orchestrates cardiomyocyte development, maturation and disease. Nat Commun 5:5288 11. Buenrostro J, Giresi P, Zaba L et al (2013) Transposition of native chromatin for fast and sensitive epigenomic profiling of open chromatin, DNA-binding proteins and nucleosome position. Nat Methods 10:1213–1218 12. Schmidl C, Rendeiro AFF, Sheffield NC et al (2015) ChIPmentation: fast, robust, low-input ChIP-seq for histones and transcription factors. Nat Methods 12:963–965 13. Skene PJ, Henikoff S (2017) An efficient targeted nuclease strategy for high-resolution mapping of DNA binding sites. Elife 6:e21856 14. Monroe TO, Hill MC, Morikawa Y et al (2019) YAP partially reprograms chromatin accessibility to directly induce adult cardiogenesis in vivo. Dev Cell 48:765–779.e7 15. Mo A, Mukamel E, Davis F et al (2015) Epigenomic signatures of neuronal diversity in the mammalian brain. Neuron 86:1369–1384 16. Buenrostro J, Wu B, Chang H et al (2015) ATAC-seq: a method for assaying chromatin accessibility genome-wide. Curr Protoc Mol Biol 109:21–29 17. Corces MR, Trevino AE, Hamilton EG et al (2017) An improved ATAC-seq protocol reduces background and enables interrogation of frozen tissues. Nat Methods 14:959–962

Chapter 24 Genetic Lineage Tracing of Non-cardiomyocytes in Mice Zhongming Chen and Jop H. van Berlo Abstract Genetic lineage tracing is accomplished using bi-transgenic mice, where one allele is altered to express Cre recombinase, and another allele encodes a Cre-dependent genetic reporter protein. Once Cre is activated (constitutive or in response to tamoxifen), the marker gene-expressing cells become indelibly labeled by the reporter protein. Therefore, daughter cells derived from labeled cells are permanently labeled even if the marker gene that drove Cre recombinase expression is no longer expressed in these cells. This system is commonly used to label putative progenitor cells and determine the fate of their progeny. Here, we describe the use of c-kit-based genetic lineage-tracing mouse line as an example and discuss caveats for performing these types of experiments. Key words Progenitor cell, Genetic lineage-tracing, Transgenic mice, Tamoxifen, Differentiation

1

Introduction Genetic lineage tracing is a powerful tool in biomedical research. The concept of genetic lineage tracing is that a marker gene is specifically expressed in one type of cell, typically a progenitor or stem cell [1, 2]. The method relies on the presence of two components in the cell that is to be labeled; the first is a site-specific recombinase, and the second is the sites that are specifically recognized by the recombinase [3]. When both are present and active, the recombinase will induce genetic recombination with the excision of a cassette, whereby a reporter gene can become expressed. The consequence of genetic recombination is definite, and even when the recombinase is no longer expressed, the reporter gene stays expressed due to it being controlled by a generic promoter. The result is that this system can be used to identify the cellular progeny of stem and progenitor cells in a multitude of tissues. The first component of genetic lineage tracing relies on the expression of a site-specific recombinase, where three things are important: the recombinase, whether constitutively active or ligand-activated recombination, and a reliable promoter. First,

Kenneth D. Poss and Bernhard Ku¨hn (eds.), Cardiac Regeneration: Methods and Protocols, Methods in Molecular Biology, vol. 2158, https://doi.org/10.1007/978-1-0716-0668-1_24, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Zhongming Chen and Jop H. van Berlo

one must choose a recombinase. In mice, the bacteriophagederived Cre recombinase is the most widely used [4]. Others include the baker’s yeast-derived Flipase (Flp) [5]. However, due to issues with thermostability and efficiency, this enzyme was initially much less widely used, although recent versions, such as Flp-e and codon-optimized Flp-o, resulted in similar efficiencies as Cre recombinase [6]. Lastly, another bacteriophage-derived recombinase termed Dre recombinase with similar efficiencies as Cre recombinase was recently identified [7, 8]. These three recombinase enzymes display constitutive nuclear localization, where they can bind to the specific sites that are recognized by these recombinases. Second, one must choose a constitutively active version of the recombinase, or a mutated version that is only activated upon binding to a ligand [9]. The main difference between these two versions is that the constitutively active recombinase will label all cells that express the chosen promoter from embryonic development onward, while the ligand-inducible version will only become activated when sufficient levels of ligand are present in the target cell to induce nuclear localization of the recombinase. Typically, a mutated estrogen receptor that can no longer bind endogenous estrogens is used to cause cytoplasmic localization of the recombinase in the absence of a ligand (and, hence, is inactive, since the sites that are recognized by the recombinase are localized in the nucleus) [10]. Upon administration of the estrogen analog tamoxifen or raloxifene, the recombinase translocates to the nucleus and thus can bind to specific sites to induce recombination [1]. Third, one must choose the promoter to drive the expression of the recombinase. After a specific promoter is chosen to label a certain cell type, the most important decision when generating genetic lineagetracing mouse models is that of a transgenic DNA construct that is incorporated randomly, as opposed to a knock-in approach where the endogenous promoter is used to drive the expression of the recombinase [11]. Important considerations include the fidelity of expression and level of expression, with the most important trade-off being low-level expression, and potentially inefficient recombination when using the endogenous promoter (i.e.,