Cardiac Gene Therapy: Methods and Protocols (Methods in Molecular Biology, 2573) 1071627066, 9781071627068

Table of contents :
Preface
Contents
Contributors
Part I: Overview
Chapter 1: Updates on Cardiac Gene Therapy Research and Methods: Overview of Cardiac Gene Therapy
1 Introduction
1.1 Cardiac Gene Therapy Approaches
1.2 Vectors and Promotors
1.3 Cardiac Gene Delivery Methods
1.4 Towards Clinical Translation
2 Conclusion
References
Part II: Gene Suppression, Editing, and Reprogramming
Chapter 2: Tough Decoy-Mediated Cardiac Gene Suppression
1 Introduction
2 Materials
2.1 Construction of Recombinant Adenovirus Expression Vector
2.2 Production of Recombinant Adenovirus
2.3 Luciferase Reporter Assay
2.4 Quantitative Real-Time PCR
2.5 Western Blotting
3 Methods
3.1 Obtain Shuttle Vector Carrying miRNA Tough Decoy
3.2 Linearization of a Shuttle Vector
3.3 Recombination in Bacteria
3.4 Purification of Recombinant Adenoviral Genome
3.5 Generation of Infectious Adenovirus
3.6 Preparation and Amplification of Primary Viral Stock
3.7 Purification of Virus
3.8 Luciferase Reporter Assay
3.9 Quantitative Real-Time PCR Analysis
3.10 Western Blotting
4 Notes
References
Chapter 3: Direct Reprogramming of Adult Human Cardiac Fibroblasts into Induced Cardiomyocytes Using miRcombo
1 Introduction
2 Materials
2.1 Reagents for Cell Culture
2.2 Reagents for microRNA Transfection
2.3 Reagents for RNA Purification and cDNA Synthesis
2.4 Reagents for Droplet Digital PCR
2.5 Reagents for Flow Cytometry
2.6 Reagents for Calcium Transient Imaging
3 Methods
3.1 Culture of Adult Human Cardiac Fibroblasts
3.2 AHCF Transfection with microRNAs
3.3 RNA Extraction
3.4 Evaluation of Transfection Efficiency: Day 2
3.5 Evaluation of Cell Reprogramming: Day 7
3.6 Evaluation of Cell Reprogramming: Day 15
3.7 Evaluation of Cell Reprogramming: Day 30
4 Notes
References
Chapter 4: CRISPR/Cas9 Gene Editing of RYR2 in Human iPSC-Derived Cardiomyocytes to Probe Ca2+ Signaling Aberrancies of CPVT A...
1 Introduction
2 Materials
2.1 Construction of Plasmid for RNA-Guided Cas9 Exonuclease and CRISPR/Cas9 Gene Edit
2.2 Cardiac Differentiation of hiPSCs and Dissociation of hiPSC-CMs
3 Methods
3.1 Design and Constructions of Materials for CRISPR/Cas9 Gene Editing
3.2 Introducing the Plasmid DNA and Repair Template into hiPSCs by Electroporation
3.3 Screening for Correctly Gene-Edited Cells
3.4 Cardiac Differentiation of the Mutant hiPSCs
3.5 Cardiomyocyte Dissociation for Single-Cell Experiments
3.6 Cardiac Differentiation of the Mutant hiPSCs in Maturation Media
4 Notes
References
Chapter 5: Enhancing Cardiomyocyte Transcription Using In Vivo CRISPR/Cas9 Systems
1 Introduction
2 Materials
2.1 Cloning
2.2 Validation
2.3 Downstream Application
3 Methods
3.1 Design gRNAs for Transcriptional Activation
3.2 Validate gRNAs for Transcriptional Activation In Vitro
3.3 Apply CRISPRa In Vivo
4 Notes
References
Chapter 6: AAV-Mediated Somatic Gene Editing for Cardiac and Skeletal Muscle in a Large Animal Model
1 Introduction
2 Materials
2.1 Vector Production
2.2 Vector Delivery in Pigs
3 Methods
3.1 Vector Design
3.2 Transfection of HEK 293 T Cells for rAAV Production
3.3 Iodixanol Purification
3.4 Gravity Flow Size Exclusion Purification
3.5 Virus Concentration
3.6 Anesthesia
3.7 Antegrade Coronary Injection
3.8 Selective Pressure Regulation of Retroinfusion
4 Notes
References
Part III: Cardiac Gene Therapy Vectors and Promotors
Chapter 7: Optimization of Synthesis of Modified mRNA
1 Introduction
2 Materials
2.1 Equipment
2.2 Solutions and Supplies
2.3 Synthesis of modRNA
2.3.1 Tailed DNA Template Preparation
2.3.2 PCR Product Purification Using QIAquick PCR Purification Kit
2.3.3 In Vitro Transcription (IVT) Reaction
2.3.4 RNA Phosphatase Treatment
2.3.5 modRNA Purification Using MEGAclear
2.3.6 modRNA Concentration for In Vivo Use Using Amicon Ultra-4 Centrifugal Filters
2.3.7 Quality Control of modRNA Using TapeStation
3 Methods
3.1 Tailed DNA Template Preparation
3.2 PCR Product Purification Using QIAquick PCR Purification Kit
3.3 In Vitro Transcription (IVT) Reaction
3.4 RNA Phosphatase Treatment
3.5 Purify modRNA Using MEGAclear
3.6 modRNA Concentration for In Vivo Use Using Amicon Ultra-4 Centrifugal Filters
3.7 Quality Control of modRNA Using TapeStation
4 Notes
References
Chapter 8: Design and Production of Heart Chamber-Specific AAV9 Vectors
1 Introduction
1.1 AAV9: Research Tool and Gene Therapy
1.2 AAV9: Structure
1.3 AAV9 vs. Transgenesis
1.4 AAV9: Tissue-Specific Enhancements
1.5 Atria vs. Ventricle
1.6 Objective: Generation of Chamber-Specific AAV9 Using Nppa and Myl2 Minimal Promoters
1.7 Identification of the Nppa and Myl2 Minimal Promoters
2 Materials
2.1 Cloning Reagents
2.2 AAV9 Production Reagents
2.3 AAV9 Purification Reagents
2.4 AAV9 Preparation for In Vivo Injection: Reagents and Delivery Equipment
3 Methods
3.1 Construction of Cardiac Chamber-Specific AAV Plasmids
3.2 Production of Cardiac Chamber-Specific AAV9
3.3 Purification of Cardiac Chamber-Specific AAV9
3.4 Preparation of Cardiac Chamber-Specific AAV9 for Usable and Safe Injection In Vivo
4 Notes
References
Chapter 9: Generation of Atrial-Specific Construct Using Sarcolipin Promoter-Associated CRM4 Enhancer
1 Introduction
2 Materials
2.1 Generation of SLN Promoter/CRM-Associated AAV Construct (See Note 1)
2.1.1 pTR.SLN.Luc./pTR.SLN.EGFP
2.1.2 pTR.CRM4.SLN.Luc
2.1.3 pTR.CRM4.SLN.EGFP
2.2 AAV Production
2.3 In Vitro Gene Transfer
2.4 In Vivo Gene Transfer
2.5 Luciferase Assay
2.6 Fluorescence Imaging
2.7 Western Blot Analysis
3 Methods
3.1 Generation of pTR.SLN.Luc Plasmid
3.2 Generation of pTR.SLN.EGFP Plasmid
3.3 Generation of pTR.CRM4.SLN.Luc Plasmid
3.4 Generation of pTR.CRM4.SLN.EGFP Plasmid
3.5 Generation of AAV Viral Vector
3.6 Preparation of PEI Stock Solution
3.7 Validation of In Vitro Gene Transfer (See Fig. 3a; See Notes 6 and 7)
3.8 In Vivo Sample Preparation for Luciferase Assay
3.9 BCA Assay for Protein Quantification
3.10 Luciferase Assay (See Fig. 3b)
3.11 Biodistribution by Western Blotting (See Fig. 4)
3.12 Fluorescent Imaging (See Fig. 5)
4 Notes
References
Part IV: Cardiac Gene Delivery Methods
Chapter 10: Cardiac Targeted Adeno-Associated Virus Injection in Rats
1 Introduction
2 Materials
2.1 Anesthesia
2.2 AAV Administration
3 Methods
3.1 Anesthesia, Intubation, and Ventilation of Rats (See Notes 1-3)
3.2 Local Intramyocardial Injection into the Ventricular Wall of the Heart
3.3 Intracoronary Injection (Fig. 1)
3.4 Postoperative Recovery
4 Notes
References
Chapter 11: Cardiac Gene Delivery in Large Animal Models: Antegrade Techniques
1 Introduction
1.1 Slow Intracoronary Perfusion
1.2 Intracoronary Perfusion + Coronary Artery Occlusion
1.3 Intracoronary Perfusion + Coronary Artery Occlusion + Coronary Sinus Occlusion
2 Materials
2.1 Slow Intracoronary Perfusion
2.2 Intracoronary Perfusion + Coronary Artery Occlusion
2.3 Intracoronary Perfusion + Coronary Artery Occlusion + Coronary Sinus Occlusion
3 Methods
3.1 Slow Intracoronary Perfusion
3.2 Intracoronary Perfusion + Coronary Artery Occlusion
3.3 Intracoronary Perfusion + Coronary Artery Occlusion + Coronary Sinus Occlusion
4 Notes
References
Chapter 12: Locked Nucleic Acid AntimiR Therapy for the Heart
1 Introduction
2 Materials
2.1 Arterial and Venous Peripheral Access
2.2 Angiography Equipment (Fig. 2)
2.3 Catheterization of the Coronary Artery (LAD or Left Circumflex Artery (RCx))
2.4 Infusion of LNA
2.5 Emergency Medication/Tools
2.6 AntimiR
3 Methods
3.1 Venous and Arterial Assess
3.2 Blocking of the Anterograde Flow in Coronary Arteries (LAD and RCx)
3.2.1 For the LAD Approach
3.2.2 For the RCx Approach
3.3 Anterograde Application (LAD and RCx)
4 Notes
References
Chapter 13: Cardiac Gene Delivery in Large Animal Models: Selective Retrograde Venous Injection
1 Introduction
2 Materials
3 Methods
4 Notes
References
Chapter 14: Endocardial Gene Delivery Using NOGA Catheter System
1 Introduction
2 Materials
2.1 Preparing the NOGA/MyoStar System
2.2 Preparing the Patient or the Experimental Animal
2.3 Electroanatomical Mapping
2.4 Gene Transfers
2.5 Removing the Femoral Sheath
3 Methods
3.1 Preparing the NOGA/MyoStar System
3.2 Preparing the Patient
3.3 Electroanatomical Mapping of the Left Ventricle
3.4 Excluding Points
3.5 Determining Injection Area
3.6 Gene Transfers
3.7 Removing Femoral Sheath
4 Notes
References
Chapter 15: Surgical Methods for Cardiac Gene Delivery in Large Animals
1 Introduction
2 Materials
2.1 Anesthesia and Pre- and Perioperative Medications
2.2 Direct Intramyocardial Gene Delivery
2.3 Surgical Gene Delivery with Heart-Lung Machine
3 Methods
3.1 Preoperative Care and Preparation
3.2 Direct Intramyocardial Gene Delivery (See Fig. 1)
3.3 Gene Delivery Using Heart-Lung Machine Including Method of Molecular Cardiac Surgery with Recirculating Delivery (See Fig....
3.4 Postoperative
4 Notes
References
Chapter 16: Atrial Gene Painting in Large Animal Model of Atrial Fibrillation
1 Introduction
2 Materials
2.1 Preparation
2.2 Surgery
3 Methods
3.1 Poloxamer Saline Solution Preparation
3.2 Animal Surgery
4 Notes
References
Chapter 17: Stent-Based Gene Delivery for Coronary Disease
1 Introduction
2 Materials
2.1 Priming Ad Vectors for Reversible Surface Immobilization on Thiolated Surfaces
2.2 Priming AAV Vectors for Reversible Surface Immobilization on Thiolated Surfaces
2.3 Formulation of Thiolated Metal Surfaces for Immobilization of Thiol-Reactive Viral Vectors
2.4 Assessment of Ad and AAV Vector Immobilization on the Metal Surfaces by Fluorescence Microscopy
2.5 In Vitro Reporter Gene Transduction Experiments with Stent- or Mesh Disk-Immobilized Ad and AAV Vectors
2.6 In Vitro Gene Transduction with Mesh-Immobilized Ad-SOD3: Immunocytochemistry
2.7 In Vitro Gene Transduction with Mesh-Immobilized Ad-SOD3: ROS Production
2.8 Deployment of Gene-Eluting Stent in the Carotid Artery (a Rat Model)
2.9 Optical Imaging of Vascular Gene Expression After Deployment of Gene-Eluting Stents
3 Methods
3.1 Priming Ad Vectors for Reversible Surface Immobilization on Thiolated Surfaces
3.2 Priming AAV Vectors for Reversible Surface Immobilization on Thiolated Surfaces
3.3 Formulation of Thiolated Metal Surfaces for Immobilization of Thiol-Reactive Viral Vectors
3.4 Assessment of Ad and AAV Vector Immobilization on the Metal Surfaces by Fluorescence Microscopy (See Fig. 2)
3.5 In Vitro Reporter Gene Transduction Experiments with Stent- or Mesh Disk-Immobilized Ad and AAV Vectors (See Figs. 3, 4, a...
3.6 In Vitro Gene Transduction with Mesh-Immobilized Ad-SOD3: Immunocytochemistry (See Fig. 6)
3.7 In Vitro Gene Transduction with Mesh-Immobilized Ad-SOD3: ROS Production (See Fig. 7)
3.8 Deployment of Gene-Eluting Stent in the Carotid Artery (a Rat Model)
3.9 Optical Imaging of Vascular Gene Expression After Deployment of Gene-Eluting Stents (See Fig. 8)
4 Notes
References
Chapter 18: Selective Anti-AAV Antibody Depletion by Hemapheresis and Immunoadsorption
1 Introduction
2 Materials
2.1 AAV-Immunoadsorbent Matrix Preparation (AAV Beads)
2.2 In Vitro Immunoadsorption
2.3 Catheter Implantation
2.4 In Vivo Hemapheresis and Immunoadsorption
2.5 Neutralization Luciferase Assay
2.6 ELISA
2.7 In Vivo Imaging
3 Methods
3.1 AAV-Immunoadsorbent Matrix Preparation (AAV Beads)
3.2 In Vitro Immunoadsorption
3.3 Elution of Anti-AAV Immunoglobulins from the Immunoadsorbent
3.4 Catheter Implantation
3.5 In Vivo Hemapheresis and Immunoadsorption
3.6 Neutralization and Luciferase Assay
3.7 ELISA
3.8 In Vivo Imaging
4 Notes
References
Chapter 19: Ex Vivo Delivery of Viral Vectors by Organ Perfusion for Cardiac Transplantation Gene Therapy
1 Introduction
2 Materials
2.1 Donor Blood Washing with Cell Saver Device
2.2 Priming the Ex Vivo Perfusion Device/Preparation of the Perfusate (See Note 1)
2.3 Ex Vivo Perfusion of the Cardiac Graft (See Note 1)
2.4 Administration of the Viral Vector to the Circuit
2.5 Removal of Cardiac Graft from Ex Vivo Perfusion Device
3 Methods
3.1 Donor Blood Washing of Neutralizing Components (See Note 2)
3.2 Preparing the Perfusate and Priming the Ex Vivo Perfusion Device
3.3 Ex Vivo Normothermic Perfusion of Cardiac Graft
3.4 Administration of the Viral Vector to the Circuit (See Note 7)
3.5 Removal of Cardiac Graft from Ex Vivo Perfusion Device
4 Notes
References
Part V: Pulmonary Hypertension
Chapter 20: Intra-Airway Gene Delivery for Pulmonary Hypertension in Rodent Models
1 Introduction
2 Materials
2.1 Rat Models of Pulmonary Artery Hypertension
2.1.1 Animals
2.1.2 Anesthesia
2.1.3 Surgery, PH Induction, Gene Delivery
2.2 Mouse Models of Pulmonary Artery Hypertension
2.2.1 Animals
2.2.2 PH Induction and Gene Delivery
2.3 Functional PH Characterization
2.4 Histological PAH Characterization
3 Methods
3.1 Animal Preparation and PH Model Creation in Rats
3.1.1 Monocrotaline-Induced PAH Model
3.1.2 Sugen-Chronic Hypoxia-Induced PAH Model
3.1.3 Pneumonectomy-Induced PAH Model
3.2 Animal Preparation and PH Model Creation in Mice
3.2.1 Sugen-Chronic Hypoxia-Induced PAH Model
3.3 Airway Gene Delivery
3.4 PAH Assessment and Gene Transduction Efficiency
4 Notes
References
Chapter 21: Endobronchial Gene Delivery for Pulmonary Hypertension in a Large Animal Model
1 Introduction
2 Materials
2.1 Animal Preparation
2.2 Bronchoscopy and Gene Delivery
3 Methods
3.1 Experiment Preparation
3.2 Animal Preparation
3.3 Bronchoscopy and Gene Delivery
4 Notes
References
Part VI: Patient Screening and Measuring the Efficacy of Cardiac Gene Therapy
Chapter 22: Cell-Based Determination of Neutralizing Antibodies Against Adeno-Associated Virus in Cardiac Gene Therapy
1 Introduction
2 Materials
2.1 Cell Culture and Maintenance
2.2 Neutralizing Antibody Assay
2.3 Luciferase Assay
3 Methods
3.1 Cell Culture and Cell Maintenance
3.2 ``Cell Plate´´ Preparation
3.3 Neutralizing Antibody Assay
3.4 Luciferase Assay
4 Notes
References
Chapter 23: Left Ventricular Pressure Volume Assessment Using Carotid Artery Access in the Rat
1 Introduction
2 Materials
3 Methods
4 Notes
References
Chapter 24: Assessing the Effect of Cardiac Gene Therapy Using Catheter-Based Pressure-Volume Measurement in Large Animals
1 Introduction
2 Materials
3 Methods
4 Notes
References
Chapter 25: ELISpot Assay for Gene Therapy in Large Animal Studies
1 Introduction
2 Materials
2.1 PBMC Isolation
2.2 ELISpot Assay
3 Methods
3.1 PBMC Isolation
3.2 ELISpot Assay
4 Notes
References
Chapter 26: Assessing Recombinant AAV Shedding After Cardiac Gene Therapy
1 Introduction
2 Materials
2.1 Storage Medium Preparation
2.2 Culture Medium Preparation
2.3 Specimen Sampling
2.3.1 Blood Sampling
2.3.2 Feces Sampling
2.3.3 Urine Sampling
2.3.4 Saliva Sampling
2.3.5 Nasal Mucus Sampling
2.3.6 qPCR Analysis
2.4 Infectious Replication Assay
3 Methods
3.1 Preparation of Storage Medium
3.2 Preparation of Culture Medium
3.3 Specimen Sampling
3.3.1 Blood Sampling (See Note 1)
3.3.2 Feces Sampling
3.3.3 Urine Sampling
3.3.4 Saliva Sampling
3.3.5 Nasal Mucus Sampling (Only Under Sedation)
3.4 qPCR Analysis to Determine Vector Concentration in Specimen Samples
3.4.1 Preparation of the rAAV DNA for the Standard Curve
3.4.2 Preparation of the Positive and Negative Control Samples
3.4.3 qPCR Protocol
3.5 Infectious Replication Assay
3.5.1 Day 1 (Morning): Seed Culture Plate with HeLa RC32 Cells
3.5.2 Day 2 (Morning): Prepare Specimen Samples and Adenovirus Dilutions to Add to Cells
3.5.3 Day 4 (Morning): Collect Cell Pellets
3.5.4 Total DNA Extraction from Infectious Replication Assay Cell Pellet
3.5.5 qPCR Analysis to Determine the Vector Concentration in Samples
4 Notes
References
Correction to: Direct Reprogramming of Adult Human Cardiac Fibroblasts into Induced Cardiomyocytes Using miRcombo
Index

Citation preview

Methods in Molecular Biology 2573

Kiyotake Ishikawa Editor

Cardiac Gene Therapy Methods and Protocols Second Edition

METHODS

IN

MOLECULAR BIOLOGY

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

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

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

Cardiac Gene Therapy Methods and Protocols Second Edition

Edited by

Kiyotake Ishikawa Icahn School of Medicine at Mount Sinai, New York, NY, USA

Editor Kiyotake Ishikawa Icahn School of Medicine at Mount Sinai New York, NY, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-2706-8 ISBN 978-1-0716-2707-5 (eBook) https://doi.org/10.1007/978-1-0716-2707-5 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022, Corrected Publication 2023 Chapter 3 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. The images or other third party material in this book are included in the book’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the book’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. This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface Since the publication of Cardiac Gene Therapy in 2016, there has been considerable progress in clinical translation of gene therapy, highlighted by clinical approvals for multiple gene therapy products. Unfortunately, no gene therapy specifically targeting the heart has translated into clinical practice yet, and to date, only a few early-phase clinical trials are underway. However, activity in preclinical areas of cardiac gene therapy has gained momentum and is rapidly growing toward the realization of next-generation therapy for cardiovascular diseases. In the second edition of Cardiac Gene Therapy, I recruited authors with long-term expertise in cardiac gene therapy experiments. Many of the collected protocol chapters are from expert investigators who have published novel work in the past few years since the publication of the first edition. With a focus on promoting further clinical translation, covered topics include gene suppression, editing, and reprogramming; cardiac gene therapy vectors and promotors; cardiac gene delivery methods; pulmonary hypertension; and patient screening and measuring the efficacy of cardiac gene therapy. Together with the protocols included in the first edition, these detailed and practical protocols will be valuable tools for researchers in cardiology to conduct cardiac gene therapy research. I thank all the expert authors for their dedication in describing step-by-step methodologies that will undoubtedly lead to further advancements and successful clinical translation of cardiac gene therapy. I am incredibly grateful to John M. Walker, the series editor, who provided me with this opportunity and guided the volume preparation process. I hope that readers will find Cardiac Gene Therapy: Methods and Protocols, Second Edition a useful reference for conducting and improving their projects. New York, NY, USA

Kiyotake Ishikawa

v

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

PART I

OVERVIEW

1 Updates on Cardiac Gene Therapy Research and Methods: Overview of Cardiac Gene Therapy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Francisco J. Romeo, Spyros A. Marvopoulos, and Kiyotake Ishikawa

PART II

3

GENE SUPPRESSION, EDITING, AND REPROGRAMMING

2 Tough Decoy-Mediated Cardiac Gene Suppression . . . . . . . . . . . . . . . . . . . . . . . . . Changwon Kho 3 Direct Reprogramming of Adult Human Cardiac Fibroblasts into Induced Cardiomyocytes Using miRcombo . . . . . . . . . . . . . . . . . . . . . . . . . . . . Camilla Paoletti, Carla Divieto, and Valeria Chiono 4 CRISPR/Cas9 Gene Editing of RYR2 in Human iPSC-Derived Cardiomyocytes to Probe Ca2+ Signaling Aberrancies of CPVT Arrhythmogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Naohiro Yamaguchi, Xiao-Hua Zhang, and Martin Morad 5 Enhancing Cardiomyocyte Transcription Using In Vivo CRISPR/Cas9 Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Eric Schoger and Laura C. Zelaraya´n 6 AAV-Mediated Somatic Gene Editing for Cardiac and Skeletal Muscle in a Large Animal Model. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tilman Ziegler, Tarik Bozoglu, and Christian Kupatt

PART III

v xi

13

31

41

53

63

CARDIAC GENE THERAPY VECTORS AND PROMOTORS

7 Optimization of Synthesis of Modified mRNA. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77 Jimeen Yoo and Lior Zangi 8 Design and Production of Heart Chamber-Specific AAV9 Vectors . . . . . . . . . . . . 89 Alina S. Bilal, Donna J. Thuerauf, Erik A. Blackwood, and Christopher C. Glembotski 9 Generation of Atrial-Specific Construct Using Sarcolipin Promoter-Associated CRM4 Enhancer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115 Dongtak Jeong

vii

viii

Contents

PART IV 10

11

12 13

14

15

16 17

18

19

Cardiac Targeted Adeno-Associated Virus Injection in Rats . . . . . . . . . . . . . . . . . . Michael G. Katz, Yoav Hadas, Adam S. Vincek, Nataly Shtraizent, Eric Schadt, and Efrat Eliyahu Cardiac Gene Delivery in Large Animal Models: Antegrade Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Spyros A. Mavropoulos, Kelly P. Yamada, Tomoki Sakata, and Kiyotake Ishikawa Locked Nucleic Acid AntimiR Therapy for the Heart . . . . . . . . . . . . . . . . . . . . . . . Sabine Samolovac and Rabea Hinkel Cardiac Gene Delivery in Large Animal Models: Selective Retrograde Venous Injection. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Philipp Schlegel and Philip W. J. Raake Endocardial Gene Delivery Using NOGA Catheter System . . . . . . . . . . . . . . . . . . Satu Siimes, Niko J€ a rvel€ a inen, Henna Korpela, and Seppo Yl€ a -Herttuala Surgical Methods for Cardiac Gene Delivery in Large Animals . . . . . . . . . . . . . . . Michael G. Katz, Yoav Hadas, Adam S. Vincek, Nataly Shtraizent, Hylton P. Gordon, Peter Pastuszko, Eric Schadt, and Efrat Eliyahu Atrial Gene Painting in Large Animal Model of Atrial Fibrillation. . . . . . . . . . . . . Weilan Mo and J. Kevin Donahue Stent-Based Gene Delivery for Coronary Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . Ivan S. Alferiev, Michael Chorny, Robert L. Wilensky, Robert J. Levy, and Ilia Fishbein Selective Anti-AAV Antibody Depletion by Hemapheresis and Immunoadsorption . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alejandro Orlowski and Thomas Weber Ex Vivo Delivery of Viral Vectors by Organ Perfusion for Cardiac Transplantation Gene Therapy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Michelle Mendiola Pla, Amy Evans, Paul Lezberg, and Dawn E. Bowles

PART V 20

21

CARDIAC GENE DELIVERY METHODS 135

147

159

171 179

189

205 217

235

249

PULMONARY HYPERTENSION

Intra-Airway Gene Delivery for Pulmonary Hypertension in Rodent Models. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 263 Malik Bisserier, Olivier Boucherat, Sebastien Bonnet, and Lahouaria Hadri Endobronchial Gene Delivery for Pulmonary Hypertension in a Large Animal Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 279 Olympia Bikou and Kiyotake Ishikawa

Contents

PART VI

ix

PATIENT SCREENING AND MEASURING THE EFFICACY OF CARDIAC GENE THERAPY

22

Cell-Based Determination of Neutralizing Antibodies Against Adeno-Associated Virus in Cardiac Gene Therapy . . . . . . . . . . . . . . . . . . . . . . . . . . Anjali J. Ravichandran, Renata Mazurek, and Kiyotake Ishikawa 23 Left Ventricular Pressure Volume Assessment Using Carotid Artery Access in the Rat . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Spyros A. Mavropoulos and Kiyotake Ishikawa 24 Assessing the Effect of Cardiac Gene Therapy Using Catheter-Based Pressure–Volume Measurement in Large Animals . . . . . . . . . . . . . . . . . . . . . . . . . . . Tomoki Sakata, Renata Mazurek, Spyros A. Mavropoulos, Francisco J. Romeo, Anjali J. Ravichandran, and Kiyotake Ishikawa 25 ELISpot Assay for Gene Therapy in Large Animal Studies . . . . . . . . . . . . . . . . . . . Renata Mazurek and Kiyotake Ishikawa 26 Assessing Recombinant AAV Shedding After Cardiac Gene Therapy . . . . . . . . . . Melad Farraha and Eddy Kizana Correction to: Direct Reprogramming of Adult Human Cardiac Fibroblasts into Induced Cardiomyocytes Using miRcombo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors IVAN S. ALFERIEV • The Children’s Hospital of Philadelphia, Philadelphia, PA, USA; The University of Pennsylvania Perelman School of Medicine, Philadelphia, PA, USA OLYMPIA BIKOU • Cardiovascular Research Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA ALINA S. BILAL • Translational Cardiovascular Research Center and Department of Internal Medicine, University of Arizona College of Medicine – Phoenix, Phoenix, AZ, USA MALIK BISSERIER • Cardiovascular Research Institute, Icahn School of Medicine at Mount Sina, New York, NY, USA ERIK A. BLACKWOOD • Translational Cardiovascular Research Center and Department of Internal Medicine, University of Arizona College of Medicine – Phoenix, Phoenix, AZ, USA SEBASTIEN BONNET • Pulmonary Hypertension Research Group, Que´bec Heart and Lung Institute Research Centre, QC, Canada; Department of Medicine, Laval University, QC, Canada OLIVIER BOUCHERAT • Pulmonary Hypertension Research Group, Que´bec Heart and Lung Institute Research Centre, QC, Canada; Department of Medicine, Laval University, QC, Canada DAWN E. BOWLES • Duke University, Durham, NC, USA TARIK BOZOGLU • Klinik und Poliklinik fu¨r Innere Medizin I, Klinikum rechts der Isar der Technical University Munich, Munich, Germany; German Center for Cardiovascular Research (DZHK), Munich Heart Alliance, Munich, Germany VALERIA CHIONO • Department of Mechanical and Aerospace Engineering, Politecnico di Torino, Turin, Italy; Department for Materials and Devices of the National Research Council, Institute for the Chemical and Physical Processes (CNR-IPCF UOS), Pisa, Italy MICHAEL CHORNY • The Children’s Hospital of Philadelphia, Philadelphia, PA, USA; The University of Pennsylvania Perelman School of Medicine, Philadelphia, PA, USA CARLA DIVIETO • Division of Advanced Materials and Life Sciences, Istituto Nazionale di Ricerca Metrologica, Turin, Italy J. KEVIN DONAHUE • Cardiovascular Medicine, UMass Chan Medical School, Worcester, MA, USA EFRAT ELIYAHU • Department of Genetics and Genomic Sciences, Icahn Institute for Genomics and Multiscale Biology, Icahn School of Medicine at Mount Sinai, New York, NY, USA; Icahn School for Data Science and Genomic Technology, Icahn School of Medicine at Mount Sinai, New York, NY, USA AMY EVANS • Duke University, Durham, NC, USA MELAD FARRAHA • Centre for Heart Research, The Westmead Institute for Medical Research, Westmead, NSW, Australia ILIA FISHBEIN • The Children’s Hospital of Philadelphia, Philadelphia, PA, USA; The University of Pennsylvania Perelman School of Medicine, Philadelphia, PA, USA CHRISTOPHER C. GLEMBOTSKI • Translational Cardiovascular Research Center and Department of Internal Medicine, University of Arizona College of Medicine – Phoenix, Phoenix, AZ, USA

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Contributors

HYLTON P. GORDON • Comparative Medicine and Surgery, Icahn School of Medicine at Mount Sinai, New York, NY, USA YOAV HADAS • Department of Genetics and Genomic Sciences, Icahn Institute for Genomics and Multiscale Biology, Icahn School of Medicine at Mount Sinai, New York, NY, USA LAHOUARIA HADRI • Cardiovascular Research Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA RABEA HINKEL • Deutsches Primatenzentrum GmbH, Go¨ttingen, Germany KIYOTAKE ISHIKAWA • Cardiovascular Research Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA NIKO JA€ RVELA€ INEN • A.I. Virtanen Institute for Molecular Sciences, University of Eastern Finland, Kuopio, Finland DONGTAK JEONG • Department of Molecular & Life Science, College of Science and Convergence Technology, Hanyang University ERICA, Ansan, South Korea MICHAEL G. KATZ • Department of Genetics and Genomic Sciences, Icahn Institute for Genomics and Multiscale Biology, Icahn School of Medicine at Mount Sinai, New York, NY, USA; Department of Cardiovascular Surgery and Pediatric Cardiac Surgery, Icahn School of Medicine at Mount Sinai, New York, NY, USA CHANGWON KHO • Division of Applied Medicine, School of Korean Medicine, Pusan National University, Yangsan, South Korea EDDY KIZANA • Centre for Heart Research, The Westmead Institute for Medical Research, Westmead, NSW, Australia; Faculty of Medicine and Health, The University of Sydney, Sydney, NSW, Australia; Department of Cardiology, Westmead Hospital, Westmead, NSW, Australia HENNA KORPELA • A.I. Virtanen Institute for Molecular Sciences, University of Eastern Finland, Kuopio, Finland CHRISTIAN KUPATT • Klinik und Poliklinik fu¨r Innere Medizin I, Klinikum rechts der Isar der Technical University Munich, Munich, Germany; German Center for Cardiovascular Research (DZHK), Munich Heart Alliance, Munich, Germany ROBERT J. LEVY • The Children’s Hospital of Philadelphia, Philadelphia, PA, USA; The University of Pennsylvania Perelman School of Medicine, Philadelphia, PA, USA PAUL LEZBERG • TransMedics, Inc, Andover, MA, USA SPYROS A. MARVOPOULOS • Cardiovascular Research Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA RENATA MAZUREK • Cardiovascular Research Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA MARTIN MORAD • Cardiac Signaling Center of University of South Carolina, Medical University of South Carolina, and Clemson University, Charleston, SC, USA; Department of Cell Biology and Anatomy, University of South Carolina, Charleston, SC, USA WEILAN MO • Cardiovascular Medicine, UMass Chan Medical School, Worcester, MA, USA ALEJANDRO ORLOWSKI • Cardiovascular Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA; Centro de Investigaciones Cardiovasculares “Dr. Horacio Cingolani,” Universidad Nacional de La Plata-CONICET, La Plata, Argentina CAMILLA PAOLETTI • Department of Mechanical and Aerospace Engineering, Politecnico di Torino, Turin, Italy PETER PASTUSZKO • Department of Cardiovascular Surgery and Pediatric Cardiac Surgery, Icahn School of Medicine at Mount Sinai, New York, NY, USA MICHELLE MENDIOLA PLA • Duke University, Durham, NC, USA

Contributors

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PHILIP W. J. RAAKE • Department of Internal Medicine III, Cardiology, University Hospital Heidelberg, University of Heidelberg, Heidelberg, Germany; DZHK [German Centre for Cardiovascular Research], Partner Site, Heidelberg, Germany; Department of Internal Medicine I, Cardiology, University Hospital Augsburg, University of Augsburg, Augsburg, Germany ANJALI J. RAVICHANDRAN • Cardiovascular Research Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA FRANCISCO J. ROMEO • Cardiovascular Research Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA TOMOKI SAKATA • Cardiovascular Research Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA SABINE SAMOLOVAC • Deutsches Primatenzentrum GmbH, Go¨ttingen, Germany ERIC SCHADT • Department of Genetics and Genomic Sciences, Icahn Institute for Genomics and Multiscale Biology, Icahn School of Medicine at Mount Sinai, New York, NY, USA; Mount Sinai Center for Transformative Disease Modeling, Icahn School of Medicine at Mount Sinai, New York, NY, USA PHILIPP SCHLEGEL • Department of Internal Medicine III, Cardiology, University Hospital Heidelberg, University of Heidelberg, Heidelberg, Germany; DZHK [German Centre for Cardiovascular Research], Partner Site, Heidelberg, Germany ERIC SCHOGER • Institute of Pharmacology & Toxicology, University Medical Center Go¨ttingen, Georg-August-University Go¨ttingen, Go¨ttingen, Germany; German Center for Cardiovascular Research (DZHK e.V.), Go¨ttingen, Germany; Georg-August-University Go¨ttingen, Go¨ttingen, Germany NATALY SHTRAIZENT • Grit Bio, Inc., New York, NY, USA SATU SIIMES • A.I. Virtanen Institute for Molecular Sciences, University of Eastern Finland, Kuopio, Finland DONNA J. THUERAUF • Department of Cellular and Molecular Biology, San Diego State University, San Diego, CA, USA ADAM S. VINCEK • Department of Genetics and Genomic Sciences, Icahn Institute for Genomics and Multiscale Biology, Icahn School of Medicine at Mount Sinai, New York, NY, USA THOMAS WEBER • Cardiovascular Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA; Spark Therapeutics Inc., Philadelphia, PA, USA ROBERT L. WILENSKY • The University of Pennsylvania Perelman School of Medicine, Philadelphia, PA, USA KELLY P. YAMADA • Cardiovascular Research Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA NAOHIRO YAMAGUCHI • Cardiac Signaling Center of University of South Carolina, Medical University of South Carolina, and Clemson University, Charleston, SC, USA; Department of Cell Biology and Anatomy, University of South Carolina, Charleston, SC, USA SEPPO YLA€ -HERTTUALA • A.I. Virtanen Institute for Molecular Sciences, University of Eastern Finland, Kuopio, Finland; Heart Center and Gene Therapy Unit, Kuopio University Hospital, Kuopio, Finland JIMEEN YOO • Cardiovascular Research Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA

xiv

Contributors

LIOR ZANGI • Cardiovascular Research Institute, Icahn School of Medicine at Mount Sinai, New York, NY, USA ´ LAURA C. ZELARAYAN • Institute of Pharmacology & Toxicology, University Medical Center Go¨ttingen, Georg-August-University Go¨ttingen, Go¨ttingen, Germany; German Center for Cardiovascular Research (DZHK e.V.), Go¨ttingen, Germany; Georg-August-University Go¨ttingen, Go¨ttingen, Germany XIAO-HUA ZHANG • Cardiac Signaling Center of University of South Carolina, Medical University of South Carolina, and Clemson University, Charleston, SC, USA; Department of Cell Biology and Anatomy, University of South Carolina, Charleston, SC, USA TILMAN ZIEGLER • Klinik und Poliklinik fu¨r Innere Medizin I, Klinikum rechts der Isar der Technical University Munich, Munich, Germany; German Center for Cardiovascular Research (DZHK), Munich Heart Alliance, Munich, Germany

Part I Overview

Chapter 1 Updates on Cardiac Gene Therapy Research and Methods: Overview of Cardiac Gene Therapy Francisco J. Romeo, Spyros A. Marvopoulos, and Kiyotake Ishikawa Abstract Gene therapy has made a significant progress in clinical translation over the past few years with several gene therapy products currently approved or anticipating approval for clinical use. Cardiac gene therapy lags behind that of other areas of diseases, with no application of cardiac gene therapy yet approved for clinical use. However, several clinical trials for gene therapy targeting the heart are underway, and innovative research studies are being conducted to close the gap. The second edition of Cardiac Gene Therapy in Methods in Molecular Biology provides protocols for cutting-edge methodologies used in these studies. In this chapter, we discuss recent updates on cardiac gene therapy studies and provide an overview of the chapters in the book. Key words Vector, Adeno-associated virus, Promoter, Delivery, Animal model, Assay, Experiments, Targeting, Specificity

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Introduction Gene therapy has recently graduated from the benchtops of researchers and has entered the realm of clinical usage for several applications, such as the treatment of retinal, spinal, and blood disorders. Several chimeric antigen receptor (CAR) T-cell therapies, in which genetic modification of T-cells is carried out ex vivo before reintroducing the cells to patients, have been clinically approved for blood cancer treatment [1]. Short interfering RNA (siRNA) therapy for hereditary ATTR amyloidosis was approved in 2018. A few AAV gene therapies are also approved by the US FDA, such as voretigene neparvovec for retinal dystrophy and onasemnogene abeparvovec for spinal muscular atrophy, and others are pending approval. Meanwhile, gene therapy that specifically targets the heart has been left behind, not having reached this stage of clinical translation yet. The major challenge for cardiac gene therapy is to achieve

Kiyotake Ishikawa (ed.), Cardiac Gene Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2573, https://doi.org/10.1007/978-1-0716-2707-5_1, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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efficient gene expression in the cardiac cells which is hindered by several barriers that prevent gene delivery to these cells. Nevertheless, novel approaches and tools have been and are continuing to be developed to overcome these barriers. The second edition of Cardiac Gene Therapy in Methods in Molecular Biology covers practical methods used in recent cardiac gene therapy studies. This chapter discusses the recent progresses in cardiac gene therapy studies while providing an overview of the chapters covered in the book. 1.1 Cardiac Gene Therapy Approaches

In addition to classic gene therapy approaches that focus on introducing new genes or adding copies of existing genes for supplementation, several novel approaches are now available including gene suppression, editing, and reprogramming. Gene suppression using siRNA, short hairpin RNA (shRNA), and microRNA (miRNA) offers inhibition or degradation of target mRNA and treats diseases that are associated with excessive gene signaling or abnormal gene expression. MicroRNA can target several genes and might provide opportunities to treat not only the single target gene but also large signaling cascades. There are also approaches to reduce endogenous microRNA in order to interfere with native RNA suppressive pathways using anti-miRNA [2] (Chap. 12), tough decoy [3, 4] (Chap. 2), and miRNA sponge [5]. Gene editing using zinc finger nucleases, transcription activator-like effector nucleases (TALENs), and, more recently, clustered regulatory interspaced short palindromic repeats (CRISPR)/Cas9 systems provides unique opportunities to edit the genes themselves, which can treat the fundamental pathology in variety of monogenic diseases. Specifically, CRISPR/Cas9 offers highly efficient and feasible gene editing and is now used in variety of cardiac applications including studying the function of specific genes [6] (Chap. 4), enhancing transcription of specific genomic loci [7] (Chap. 5), and treating diseases [8] (Chap. 6). Gene reprogramming is another unique approach that can potentially be used to change the cellular phenotype from fibroblast to cardiomyocytes in vivo [9] (Chap. 3) and treat patients with scars in the heart.

1.2 Vectors and Promotors

Gene delivery vectors are commonly divided into non-viral and recombinant viral vectors, each of them with advantages and disadvantages. Non-viral vectors are usually flexible in deliverable gene size and less immunogenic, whereas viral vectors offer higher efficiency. For non-viral vectors, naked nucleic acids are delivered either directly into the cardiac tissue or with scaffolds to protect them from degradation (proteins, lipids, or any other molecule). Plasmid DNA remains appealing as a carrier in human cardiac gene therapy due to the capacity of mass production, unlimited gene size, and relatively low immunogenicity [10]. However, its transfection efficiency remains low, and novel tools or approaches to enhance efficacy and cardiac specificity are likely needed. Recently,

Updates on Cardiac Gene Therapy Research and Methods. . .

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modified mRNA has proved its potential as a promising vector in coronavirus disease 2019 vaccines. Its cardiac application (Chap. 7) seems also promising, with positive results in several innovative applications including myocardial regeneration [11], myocardial remodeling inhibition [12], and myocardial cell reprogramming [13]. In addition, modified mRNA vascular endothelial growth factor A (VEGFA) is being tested in a clinical trial during elective surgical coronary revascularization with promising results [14]. Viral vectors used for cardiac gene therapy include adenovirus, adeno-associated virus (AAV), lentivirus, and Sendai virus. Among them, adenovirus and AAV have been the major vectors used for in vivo gene delivery. Adenovirus carries a double-stranded DNA genome, produces short-term gene expression (4 weeks), and infects a wide range of cells (nonselective tropism). It induces a strong immunological response against both viral particles and infected cells even after extracting viral genes (E1E2a//E3/E4 gene regions) [15] raising significant safety concerns. However, some clinical studies reported good safety profiles for new generation adenoviral vectors [16, 17]. Its greater transduction efficacy and larger gene capacity compared to AAV continue to render adenovirus a useful research tool. AAV is another option that is increasing in popularity, especially for therapeutic purposes. Several features make AAV vectors wellsuited for cardiac gene therapy including tropism for cardiac cells (serotypes AAV-1, AAV-6, AAV-8, AAV-9), persistent transgene expression, minimal risk of genomic integration, and low immunological reactivity [18]. The remaining challenges for clinical application of AAV in cardiac gene therapy include low cardiac uptake in vivo through intravenous delivery, reduced efficacy in patients with preexisting antibodies to AAV, and immune response that accompanies severe side effects at extremely high vector doses [19]. Improvements in cardiac delivery methods (discussed below), vector modification [20], vector packaging [21], patient screening for antibodies (Chap. 22), and antibody removal [22] (Chap. 18) are among the approaches taken to address these limitations. Lentivirus is a complex retroviral vector containing singlestranded RNA. Some of its main features include the ability to produce long-term transgene expression by means of host genome integration, a larger packaging capacity than AAV (9 kb), fast transgene expression (peak 6 days), and relatively low immunogenicity. However, genome integration carries the risk of inducing oncogenesis [23]. Together with lack of cardiospecificity, in vivo application of lentivirus remains problematic, and current applications are mainly focused on ex vivo gene transfer. One of the caveats of human gene transduction is the undesired off-target effects due to nonselective tissue targeting and subsequent adverse reactions including tissue inflammation. To

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improve specificity, cardiac-specific DNA regulatory sequences called promoters can be used to regulate gene expression. Initial research targeting myocytes used strong, constitutive promoter sequences like CMV (cytomegalovirus), RSV (respiratory syncytial virus), and elongation factor 1A (EF1A). Specificity was improved by identifying regulatory sequences unique to myocytes such as α-actin, muscle creatine kinase, myosin heavy chain promoter, myosin light chain promoter, and troponin T/I [24]. Along this line, chamber-specific promoters were also developed to specifically target the atrium [25, 26] or the ventricle [26] (Chaps. 8 and 9). 1.3 Cardiac Gene Delivery Methods

The heart is protected by various barriers which hinder effective cardiac uptake of therapeutic materials [27]. Unfortunately, many of gene delivery vectors lack cardiotropism and require selective delivery to the heart. Even some of the AAV serotypes that are considered cardiotropic have insufficient cardiac specificity when given systemically in species more advanced than rodents. In rodents, cardiac-targeted delivery requires direct access to the heart. Vectors can be injected directly into the myocardium using small needle or into the left ventricle during aortic clamp to direct the vectors into the coronary circulation (Chap. 10). In larger animals and humans, catheter-based approaches are also available in addition to open-chest surgical approaches. Catheter-based intracoronary antegrade delivery offers feasible delivery using established techniques used clinically for percutaneous coronary interventions [28, 29] (Chap. 11). Retrograde delivery also targets the coronary system from the venous side while avoiding the necessity of arterial access [30] (Chap. 13). It is important to note that despite targeting the same system, gene uptake patterns seem to be different between the two methods, with more basal and epicardial distribution seen after retrograde delivery as compared to antegrade delivery. Direct intramyocardial injection is also possible using endomyocardial injection catheters with image guidance. The NOGAMyostar system is one of the most frequently used tool both in large animal models and humans, which provides information of myocardial geometry and variability using electrical mapping [31] (Chap. 14). Although significantly more invasive, gene delivery during cardiopulmonary bypass has been shown to significantly improve cardiac gene uptake and minimize off-target gene expression [32] (Chap. 15). More selective deliveries were achieved by the atrial gene painting method for the left atrium [33] (Chap. 16) and gene eluting stents for the coronary arteries [34] (Chap. 17). More recently, a unique gene delivery method to the donor heart in cardiac transplantation was demonstrated [35] (Chap. 19) and holds promise in improving outcomes in cardiac transplantation. Finally, intra-airway delivery was shown effective in targeting the

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pulmonary vasculature to treat pulmonary hypertension in preclinical models (Chaps. 20 and 21) and awaits clinical testing. 1.4 Towards Clinical Translation

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Using the methods and tools described above, it is certain that we will see more clinical testing of cardiac gene therapy in the coming years. To achieve clinical success, it is of paramount importance to identify optimal endpoints. For gene therapies that aim to improve cardiac function, a detailed study of how a particular gene therapy affects cardiac physiology will provide key insights into the mechanisms underlying that therapy and help define surrogate endpoints. Pressure-volume relationship is the gold standard for evaluating cardiac contractility and compliance [36] (Chaps. 23 and 24). Establishing noninvasive means of functional assessment and efficacy of gene transduction are necessary steps in establishing a way to monitor the success of cardiac gene therapy in patients. Safety monitoring methods for off-target side effects such as immune responses [37] (Chap. 25) and arrhythmias are another important aspect of post-gene therapy monitoring. Although often disregarded in preclinical studies, vector shedding after gene delivery [38] can be an important component of clinical monitoring for vectors that are stable (Chap. 26).

Conclusion Cardiac gene therapy holds significant promise, but further refinement is necessary to overcome the remaining challenges discussed above. We expect that the method protocols covered in this book will help recruit more researchers to the field and promote innovative research that can aid in the clinical translation of cardiac gene therapy.

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5. Li Z, Liu L, Hou N, Song Y, An X, Zhang Y, Yang X, Wang J (2016) miR-199-sponge transgenic mice develop physiological cardiac hypertrophy. Cardiovasc Res 110(2):258–267. https://doi.org/10.1093/cvr/cvw052 6. Wei H, Zhang XH, Clift C, Yamaguchi N, Morad M (2018) CRISPR/Cas9 gene editing of RyR2 in human stem cell-derived cardiomyocytes provides a novel approach in investigating dysfunctional ca(2+) signaling. Cell Calcium 73:104–111. https://doi.org/10. 1016/j.ceca.2018.04.009 7. Schoger E, Carroll KJ, Iyer LM, McAnally JR, Tan W, Liu N, Noack C, Shomroni O, Salinas G, Gross J, Herzog N, Doroudgar S, Bassel-Duby R, Zimmermann WH, Zelarayan LC (2020) CRISPR-mediated activation of endogenous gene expression in the postnatal heart. Circ Res 126(1):6–24. https://doi.org/ 10.1161/CIRCRESAHA.118.314522 8. Moretti A, Fonteyne L, Giesert F, Hoppmann P, Meier AB, Bozoglu T, Baehr A, Schneider CM, Sinnecker D, Klett K, Frohlich T, Rahman FA, Haufe T, Sun S, Jurisch V, Kessler B, Hinkel R, Dirschinger R, Martens E, Jilek C, Graf A, Krebs S, Santamaria G, Kurome M, Zakhartchenko V, Campbell B, Voelse K, Wolf A, Ziegler T, Reichert S, Lee S, Flenkenthaler F, Dorn T, Jeremias I, Blum H, Dendorfer A, Schnieke A, Krause S, Walter MC, Klymiuk N, Laugwitz KL, Wolf E, Wurst W, Kupatt C (2020) Somatic gene editing ameliorates skeletal and cardiac muscle failure in pig and human models of Duchenne muscular dystrophy. Nat Med 26(2):207–214. https://doi.org/10. 1038/s41591-019-0738-2 9. Paoletti C, Marcello E, Melis ML, Divieto C, Nurzynska D, Chiono V (2022) Cardiac tissuelike 3D microenvironment enhances route towards human fibroblast direct reprogramming into induced cardiomyocytes by microRNAs. Cell 11(5). https://doi.org/10.3390/ cells11050800 10. Schmeer M, Schleef M (2014) Pharmaceutical grade large-scale plasmid DNA manufacturing process. Methods Mol Biol 1143:219–240. https://doi.org/10.1007/978-1-4939-04105_14 11. Magadum A, Singh N, Kurian AA, Munir I, Mehmood T, Brown K, Sharkar MTK, Chepurko E, Sassi Y, Oh JG, Lee P, Santos CXC, Gaziel-Sovran A, Zhang G, Cai CL, Kho C, Mayr M, Shah AM, Hajjar RJ, Zangi L (2020) Pkm2 regulates cardiomyocyte cell cycle and promotes cardiac regeneration. Circulation 141(15):1249–1265. https://doi.

org/10.1161/CIRCULATIONAHA.119. 043067 12. Magadum A, Singh N, Kurian AA, Sharkar MTK, Sultana N, Chepurko E, Kaur K, Zak MM, Hadas Y, Lebeche D, Sahoo S, Hajjar R, Zangi L (2021) Therapeutic delivery of Pip4k2c-modified mRNA attenuates cardiac hypertrophy and fibrosis in the failing heart. Adv Sci (Weinh) 8(10):2004661. https://doi. org/10.1002/advs.202004661 13. Kaur K, Hadas Y, Kurian AA, Zak MM, Yoo J, Mahmood A, Girard H, Komargodski R, Io T, Santini MP, Sultana N, Sharkar MTK, Magadum A, Fargnoli A, Yoon S, Chepurko E, Chepurko V, Eliyahu E, Pinto D, Lebeche D, Kovacic JC, Hajjar RJ, Rafii S, Zangi L (2021) Direct reprogramming induces vascular regeneration post muscle ischemic injury. Mol Ther 29(10): 3042–3058. https://doi.org/10.1016/j. ymthe.2021.07.014 14. Collen A, Bergenhem N, Carlsson L, Chien KR, Hoge S, Gan LM, Fritsche-Danielson R (2022) VEGFA mRNA for regenerative treatment of heart failure. Nat Rev Drug Discov 21(1):79–80. https://doi.org/10.1038/ s41573-021-00355-6 15. Andrews JL, Kadan MJ, Gorziglia MI, Kaleko M, Connelly S (2001) Generation and characterization of E1/E2a/E3/E4-deficient adenoviral vectors encoding human factor VIII. Mol Ther 3(3):329–336. https://doi. org/10.1006/mthe.2001.0264 16. Leikas AJ, Hassinen I, Hedman A, Kivela A, Yla-Herttuala S, Hartikainen JEK (2021) Long-term safety and efficacy of intramyocardial adenovirus-mediated VEGF-D(DeltaNDeltaC) gene therapy eight-year follow-up of phase I KAT301 study. Gene Ther. https://doi. org/10.1038/s41434-021-00295-1 17. Hammond HK, Penny WF, Traverse JH, Henry TD, Watkins MW, Yancy CW, Sweis RN, Adler ED, Patel AN, Murray DR, Ross RS, Bhargava V, Maisel A, Barnard DD, Lai NC, Dalton ND, Lee ML, Narayan SM, Blanchard DG, Gao MH (2016) Intracoronary gene transfer of adenylyl cyclase 6 in patients with heart failure: a randomized clinical trial. JAMA Cardiol 1(2):163–171. https://doi. org/10.1001/jamacardio.2016.0008 18. Chamberlain K, Riyad JM, Weber T (2017) Cardiac gene therapy with adeno-associated virus-based vectors. Curr Opin Cardiol 32(3): 275–282. https://doi.org/10.1097/HCO. 0000000000000386 19. Wilson JM, Flotte TR (2020) Moving forward after two deaths in a gene therapy trial of Myotubular myopathy. Hum Gene Ther

Updates on Cardiac Gene Therapy Research and Methods. . . 31(13–14):695–696. https://doi.org/10. 1089/hum.2020.182 20. Weinmann J, Weis S, Sippel J, Tulalamba W, Remes A, El Andari J, Herrmann AK, Pham QH, Borowski C, Hille S, Schonberger T, Frey N, Lenter M, VandenDriessche T, Muller OJ, Chuah MK, Lamla T, Grimm D (2020) Identification of a myotropic AAV by massively parallel in vivo evaluation of barcoded capsid variants. Nat Commun 11(1):ARTN5432. https://doi.org/10.1038/s41467-02019230-w 21. Adamiak M, Liang YX, Sherman C, Lodha S, Kohlbrenner E, Jeong D, Ceholski DK, Dogra N, Dubois N, Hajjar RJ, Sahoo S (2020) Exosome-mediated Encapsulation Alters AAV Antigenicity and Infectivity: Implications for Gene Delivery in the Heart. Circulat Res:127. https://doi.org/10.1161/ res.127.suppl_1.MP165 22. Orlowski A, Katz MG, Gubara SM, Fargnoli AS, Fish KM, Weber T (2020) Successful transduction with AAV vectors after selective depletion of anti-AAV antibodies by Immunoadsorption. Mol Ther-Meth Clin D 16:192–203. https://doi.org/10.1016/j. omtm.2020.01.004 23. David RM, Doherty AT (2017) Viral vectors: the road to reducing genotoxicity. Toxicol Sci 155(2):315–325. https://doi.org/10.1093/ toxsci/kfw220 24. Piekarowicz K, Bertrand AT, Azibani F, Beuvin M, Julien L, Machowska M, Bonne G, Rzepecki R (2019) A muscle hybrid promoter as a novel tool for gene therapy. Mol TherMeth Clin D 15:157–169. https://doi.org/ 10.1016/j.omtm.2019.09.001 25. Yoo J, Kohlbrenner E, Kim O, Hajjar RJ, Jeong D (2018) Enhancing atrial-specific gene expression using a calsequestrin cis-regulatory module 4 with a sarcolipin promoter. J Gene Med 20(12):ARTNe3060. https://doi.org/ 10.1002/jgm.3060 26. Bilal AS, Blackwood EA, Thuerauf DJ, Glembotski CC (2021) Optimizing adenoassociated virus serotype 9 for studies of cardiac chamber-specific gene regulation. Circulation 143(20):2025–2027. https://doi.org/10. 1161/Circulationaha.120.052437 27. Sahoo S, Kariya T, Ishikawa K (2021) Targeted delivery of therapeutic agents to the heart. Nat Rev Cardiol 18(6):389–399. https://doi.org/ 10.1038/s41569-020-00499-9 28. Watanabe S, Ishikawa K, Fish K, Oh JG, Motloch LJ, Kohlbrenner E, Lee P, Xie C, Lee A, Liang L, Kho C, Leonardson L, McIntyre M, Wilson S, Samulski RJ, Kranias EG, Weber T, Akar FG, Hajjar RJ (2017)

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Protein phosphatase Inhibitor-1 gene therapy in a swine model of nonischemic heart failure. J Am Coll Cardiol 70(14):1744–1756. https:// doi.org/10.1016/j.jacc.2017.08.013 29. Watanabe S, Leonardson L, Hajjar RJ, Ishikawa K (2017) Cardiac Gene Delivery in Large Animal Models: Antegrade Techniques. Methods Mol Biol 1521:227–235. https://doi.org/10. 1007/978-1-4939-6588-5_16 30. Raake PW, Schlegel P, Ksienzyk J, Reinkober J, Barthelmes J, Schinkel S, Pleger S, Mier W, Haberkorn U, Koch WJ, Katus HA, Most P, Muller OJ (2013) AAV6.betaARKct cardiac gene therapy ameliorates cardiac function and normalizes the catecholaminergic axis in a clinically relevant large animal heart failure model. Eur Heart J 34(19):1437–1447. https://doi. org/10.1093/eurheartj/ehr447 31. Jarvelainen N, Halonen P, Nurro J, Hatinen OP, Korpela H, Makinen P, Gan LM, Fritsche-Danielson R, Yla-Herttuala S (2021) Citrate-saline-formulated mRNA delivery into the heart muscle with an electromechanical mapping and injection catheter does not Lead to therapeutic effects in a porcine chronic myocardial ischemia model. Hum Gene Ther 32(19–20):1295–1307. https://doi.org/10. 1089/hum.2021.149 32. White JD, Thesier DM, Swain JB, Katz MG, Tomasulo C, Henderson A, Wang L, Yarnall C, Fargnoli A, Sumaroka M, Isidro A, Petrov M, Holt D, Nolen-Walston R, Koch WJ, Stedman HH, Rabinowitz J, Bridges CR (2011) Myocardial gene delivery using molecular cardiac surgery with recombinant adeno-associated virus vectors in vivo. Gene Ther 18(6): 546–552. https://doi.org/10.1038/gt. 2010.168 33. Liu Z, Hutt JA, Rajeshkumar B, Azuma Y, Duan KL, Donahue JK (2017) Preclinical efficacy and safety of KCNH2-G628S gene therapy for postoperative atrial fibrillation. J Thorac Cardiovasc Surg 154(5):1644–1651. e1648. https://doi.org/10.1016/j.jtcvs.2017. 05.052 34. Fishbein I, Guerrero DT, Alferiev IS, Foster JB, Minutolo NG, Chorny M, Monteys AM, Driesbaugh KH, Nagaswami C, Levy RJ (2017) Stent-based delivery of adenoassociated viral vectors with sustained vascular transduction and iNOS-mediated inhibition of in-stent restenosis. Gene Ther 24(11): 717–726. https://doi.org/10.1038/gt. 2017.82 35. Bishawi M, Roan JN, Milano CA, Daneshmand MA, Schroder JN, Chiang Y, Lee FH, Brown ZD, Nevo A, Watson MJ, Rowell T, Paul S, Lezberg P, Walczak R, Bowles DE (2019) A

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normothermic ex vivo organ perfusion delivery method for cardiac transplantation gene therapy. Sci Rep 9(1):8029. https://doi.org/10. 1038/s41598-019-43737-y 36. Bastos MB, Burkhoff D, Maly J, Daemen J, den Uil CA, Ameloot K, Lenzen M, Mahfoud F, Zijlstra F, Schreuder JJ, Van Mieghem NM (2020) Invasive left ventricle pressure-volume analysis: overview and practical clinical implications. Eur Heart J 41(12):1286–1297. https://doi.org/10.1093/eurheartj/ehz552 37. Lyon AR, Babalis D, Morley-Smith AC, Hedger M, Barrientos AS, Foldes G, Couch LS, Chowdhury RA, Tzortzis KN, Peters NS, Rog-Zielinska EA, Yang HY, Welch S, Bowles CT, Haley SR, Bell AR, Rice A, Sasikaran T, Johnson NA, Falaschetti E, Parameshwar J,

Lewis C, Tsui S, Simon A, Pepper J, Rudy JJ, Zsebo KM, Macleod KT, Terracciano CM, Hajjar RJ, Banner N, Harding SE (2020) Investigation of the safety and feasibility of AAV1/SERCA2a gene transfer in patients with chronic heart failure supported with a left ventricular assist device - the SERCALVAD TRIAL. Gene Ther 27(12):579–590. https://doi.org/10.1038/s41434-0200171-7 38. Farraha M, Barry MA, Lu J, Pouliopoulos J, Le TYL, Igoor S, Rao R, Kok C, Chong J, Kizana E (2019) Analysis of recombinant adenoassociated viral vector shedding in sheep following intracoronary delivery. Gene Ther 26(9):399–406. https://doi.org/10.1038/ s41434-019-0097-0

Part II Gene Suppression, Editing, and Reprogramming

Chapter 2 Tough Decoy-Mediated Cardiac Gene Suppression Changwon Kho Abstract MicroRNA (miRNA) is a small, non-coding RNA molecule (~22 nucleotides) that acts as a posttranscriptional gene regulator, primarily by inhibiting the translation of target mRNA transcripts or affecting cell mRNA stability. Since miRNAs are comprehensively involved in gene regulation, their abnormalities are associated with various human diseases, including cardiovascular disease. Additionally, targeted inhibition of disease-related miRNAs and their targets should have therapeutic potential. Therefore, this chapter describes the experimental steps for targeted inhibition of specific miRNAs using adenoviral vectorized tough decoys that efficiently silence miRNA function in cardiac cells. Key words MicroRNA, Tough decoy inhibitors, Adenoviral vectors, Functional assays of miRNA inhibition

1

Introduction MicroRNAs (miRNAs) are powerful molecules of gene regulation in eukaryotes that play an important role in various biological processes. It has been estimated that the human genome comprises more than 2500 miRNA species and that more than 60% of protein-coding genes can be regulated by miRNAs [1, 2]. Also, our understanding of the factors/mechanisms of the miRNAmediated regulatory circuit increases, with new evidence indicating that abnormalities in miRNA biology are implicated in many human diseases, including cardiovascular disease [3, 4]. Since miRNAs can be used as powerful diagnostic and prognostic biomarkers or drug targets are highlighted, effective molecular tools for manipulating miRNA activity are essential. Three approaches are used in miRNA inhibition studies: genetic knockouts, antisense oligonucleotides (ASOs), and sponges or decoys (Table 1) [5]. Gene knockout is becoming more adopted as genome editing techniques (i.e., transcription activator-like

Kiyotake Ishikawa (ed.), Cardiac Gene Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2573, https://doi.org/10.1007/978-1-0716-2707-5_2, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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Table 1 Strategies for miRNA loss-of-function studies Applications Technology

Characteristics

In vitro

In vivo

Genetic knockout/ knockdown

Conditional or conventional suppression

Primary cells Coupled with genome editing tools

Entire body or specific tissues (organs)

Anti-miRNA

Transient to persistent Transfection or inhibition gymnosis

oligonucleotides MiRNA sponges or decoys

Intravenous, intramuscular, intraperitoneal, subcutaneous injection

Transient to long-term Transfection or viral Lentivirus, Adenovirus or AAV-mediated delivery silencing infection

effector nucleases and clustered regularly interspaced short palindromic repeats/CRISPR-associated nuclease 9 (CRISPR/Cas9)) are used to knock out miRNA genes. Over the past decade, different miRNA inhibitors have been developed. ASOs (e.g., antagomirs) are the first miRNA inhibitors to suppress specific miRNAs by hybridizing to the complementary sequence of the target miRNA. This ASO technology has been used to downregulate gene expression for over 40 years. Several ASO-based RNA therapeutics, including cardiovascular drugs, received US Food and Drug Administration and European Medicines Agency approval [6, 7], demonstrating clinical utility. Overall, ASO-based drugs have many advantages, including design flexibility due to Watson-Crick-based recognition, optimized synthesis protocols, and extensive quality control during development. The miRNA sponge or decoy strategy is an alternative to ASO that interferes with the activity of specific miRNAs through miRNA sequestration. These tools can provide versatility to inexpensive chemical manufacturing and delivery agents in miRNA research. A competitive inhibitor, the miRNA sponge, is a highly expressed transgene that carries 4–12 repeated microRNA-binding sites (MBS) to scavenge microRNAs [8]. Thus, miRNA sponge technology can repress the entire family of related miRNAs . Another class of competitive inhibitors is “tough decoy” (TuD) RNAs. This hairpin-like RNA is typically 122 nucleotides (nts) long, comprising an intervening unpaired region that contains two bulged MBS exposed in the central part of the stem that supports miR repression (Fig. 1). The MBS regions complement target miRNA and prevent the association of endogenous miRs with target mRNAs. TuD RNA uses RNA polymerase III (H1 or U6 promoters) to express RNA-containing multiple MBSs between stem regions, preventing nuclease degradation of

Adenoviral Vector-Mediated Expression of Tough Decoys

15

Stem-loop Sequence-dependent interaction

Linker

5’

Endogenous miR

miR binding sites

3’

Linker

• Avoid binding of endogenous miRs to target genes • Lead to increased expression of target genes

Stem 3’ overhang

5’ 3’

Fig. 1 Scheme of a tough decoy. TuD vectors containing two multiple miRNA-binding site (MBS) regions increase the expression of a target gene by scavenging endogenous miRNAs that act as competitive inhibitors and avoid interactions between endogenous miRNA and its target mRNAs

RNA-attracted targets. For TuD RNA, the binding site is perfectly complementary to the target miRNA, containing 3 or 4 nts inserted into the Ago2 cleavage site to avoid endonucleolytic cleavage of TuD. As a result, TuDs are a more potent tool than miRNA sponges, as shown in a comparative study [9, 10]. Additionally, the viral vector-based TuD expression system is stable and suitable for long-term use [9, 11]. TuDs are being developed for more stable suppression and targeted delivery of selected miRs for clinical applications. TuDs can be expressed in cells through vector-based delivery or transfected with a synthetically generated inhibitor comprising 20 -O-methylated RNA oligonucleotides. However, reagent-based transfection of DNA in non-dividing primary cells, including cardiomyocytes, is highly inefficient due to inefficient nuclear entry and is nonviable. There are several advantages of the adenoviral vector system. First, it is easy to obtain a high titer. Once an initial stock is generated, the adenovirus can be easily amplified in HEK-293 cells to achieve a high titer. Each mammalian cell can produce on average 10,000 adenoviruses. Second, the adenovirus is stable and useful for production and storage. Third, unlike lentivirus, adenoviruses do not insert into the host genome, activating exogenous genes without activating other genes. Therefore, the recombinant adenovirus genome remains epichromosomal in host cells, making them ideal for in vivo studies. This chapter describes a protocol for generating adenoviral vectors expressing TuD and validating their effects on cardiac cells. Recombinant adenoviruses expressing TuDs are produced

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within 7–10 days after obtaining vectors. Purification of the recombinant adenovirus is performed by CsCl gradient purification, which takes 1.5 days. Functional validation, such as luciferase reporter assays, real-time PCR assays, and Western blotting, take 1–1.5 days. All procedures for virus production and handling should follow all published guidelines for Biosafety Level 2.

2

Materials

2.1 Construction of Recombinant Adenovirus Expression Vector

1. Plasmids containing miRNA target decoys (Addgene, http:// www.addgene.org) and negative control miRNAs (e.g., cel-mir-67). 2. pShuttle-IRES-hrGFP vector (Agilent Technologies). 3. BJ5183-AD electrocompetent cells (Agilent Technologies). 4. XL10-gold ultracompetent cells (Agilent Technologies). 5. PCR Purification Kit. 6. Gel Extraction Kit. 7. Plasmid DNA Miniprep Kit. 8. DNA Maxiprep kit for a Large-Construct. 9. Restriction enzymes: phosphatase (AP).

PacI

and

PmeI,

alkaline

10. DNA ladders. 11. DNA Loading Dye (6). 12. Non-toxic fluorescent dye for post-DNA-staining DNA stain (e.g., SYBR Safe). 13. Agarose gel. 14. Kanamycin. 15. TAE (Tris-acetate-EDTA) running buffer. 16. LB (Luria-Bertani) broth and LB agar plates. 17. SOB (Super Optimal Broth) medium. 18. MQ (Milli-Q ultrapure) water. 19. β-Mercaptoethanol. 20. Electroporation cuvettes (gap width 0.2 cm). 21. Round-bottom culture tube (17 mm  100 mm, 14 mL). 22. Microcentrifuge tube (1.7 mL, DNase-free, RNase-free). 23. Water bath. 24. Centrifuges. 25. Incubator for bacteria cell culture. 26. DNA gel electrophoresis system.

Adenoviral Vector-Mediated Expression of Tough Decoys

17

27. MicroPulser electroporation system. 28. UV transilluminator. 29. Spectrophotometer (NanoDrop spectrophotometer). 2.2 Production of Recombinant Adenovirus

1. PacI digested recombinant adenoviral vectors (transfection grade). 2. AD-293 cells (Agilent Technologies). 3. Cell culture medium: DMEM (Dulbecco’s Modified Eagle Medium), 10% FBS (fetal bovine serum), antibiotics (1% Penicillin/Streptomycin). 4. Serum-free DMEM or Opti-MEM (Gibco Invitrogen) for transfection reaction. 5. PBS (phosphate-buffered saline). 6. Transfection reagents (e.g., polyethylenimine (PEI)). 7. Ultracentrifuge tube. 8. PD-10 desalting column. 9. Cell lysis buffer: 10 mM HEPES, pH 8.0. 10. Syringes (3 mL) with needles (18G). 11. Light cesium chloride (CsCl) solution: CsCl/10 mM HEPES (pH 8.0) 1.25 g/mL . 12. Heavy cesium chloride (CsCl) solution: CsCl/10 mL HEPES (pH 8.0) 1.40 g/mL. The solutions should be autoclaved and pH adjusted to 7.4 for in vivo use. 13. Biosafety Level 2 Tissue Culture system. 14. Ultracentrifuge. 15. Spectrophotometer.

2.3 Luciferase Reporter Assay

1. pMirTarget reporter vector containing 30 -UTR of target gene (OriGene). 2. H9c2 cardiac myoblast cell line (ATCC). 3. Transfection reagents. 4. Luciferase assay kit. 5. 96-Well microplates (white bottom). 6. Luminometer.

2.4 Quantitative Real-Time PCR

1. 0.2 mL PCR tube. 2. miRNA isolation kit. 3. miRNA cDNA synthesis kit. 4. Specific primer pairs for detecting decoy target cardiac miRNAs, target cardiac genes of the miRNAs, and 18S ribosomal RNA (internal control).

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5. Real-time qPCR master mixtures (SYBR Green and ROX reference dye). 6. Real-time PCR thermal cycler. 2.5

Western Blotting

1. RIPA lysis buffer. 2. Protease inhibitor cocktail. 3. Phosphatase inhibitor cocktail. 4. BSA (bovine serum albumin). 5. BCA (bicinchoninic acid) Protein Assay Kit. 6. Laemmli sample buffer (4). 7. Reducing reagents (i.e., β-mercaptoethanol or dithiothreitol (DTT)). 8. SDS-PAGE gel. 9. SDS-PAGE Running system. 10. Wet Electroblotting systems. 11. SDS-PAGE running buffer (Tris-glycine-SDS buffer). 12. Electroblotting transfer buffer (Tris-glycine buffer). 13. TBS (Tris-buffered saline buffer) buffer. 14. TBST wash buffer (TBS with or without 0.1% Tween 20). 15. Blocking buffer (non-fat milk (5% w/v) and/or BSA (1–2% w/v) in TBS). 16. Pre-stained protein standards. 17. Blotting membrane (nitrocellulose membrane, 0.45 μm pore size). 18. Primary antibodies for target cardiac proteins. 19. Secondary antibodies conjugated).

(horseradish

peroxidase

(HRP)-

20. Western blot detection solution (ECL Western blotting substrate). 21. Western blot Rocker-shaker. 22. Developing system.

3

Methods

3.1 Obtain Shuttle Vector Carrying miRNA Tough Decoy

The most well-known decoy is a hairpin-sharped structure (see Note 1). The adenoviral vectors containing the TuD sequences are generated using the pAdEasy adenoviral vector system (see Note 2).

Adenoviral Vector-Mediated Expression of Tough Decoys

3.2 Linearization of a Shuttle Vector

19

1. Incubate 1 μg of pShuttle-IRES-hrGFP vector plus TuD segments (termed pShuttle-TuD) in a 50 μL reaction with 10 units (typically 1 μL) PmeI restriction enzyme at 37  C for 1 h. 2. Prepare sample to confirm complete digestion as follows: an aliquot of the reaction mixture (3 μL) + 1 μL DNA loading dye (6) + 1 μL MQ water. 3. Run 0.7% agarose gel with fluorescent dye in TAE running buffer at 100 V for 1 h (until the dye line is about 80% below the gel). 4. Visualize the DNA fragments using UV light. A full-size single band of the pShuttle-TuD plasmid should be detected. 5. Purify the PmeI-digested pShuttle-TuD vectors by spin column-based purification method (e.g., PCR purification kit). 6. Dephosphorylate the PmeI-digested pShuttle-TuD with AP at 37  C for 30 min. Approximately 4 μg of PmeI-digested pShuttle-TuD (~10 kb) is sufficient for 1 unit of AP treatment. 7. Inactivate AP enzymes for 20 min at 80  C. 8. Purify the dephosphorylated PmeI-digested pShuttle-TuD vectors by gel purification. 9. Check DNA concentration and purity by spectrophotometer. Expected range is approximately 3–4 μg of PmeI-digested pShuttle-TuD vectors.

3.3 Recombination in Bacteria

1. Prepare round-bottom temperature (RT).

culture

tubes

at

room

2. Pre-warm SOC medium (1 mL for each tube) in a 37  C water bath and LB agar plate containing 50 μg/mL of kanamycin at 37  C incubator for 1 h. 3. Pre-cool the microcentrifuge tube and electroporation cuvette on ice for at least 15 min. 4. Thaw BJ5183-AD cells on ice (approximately 10 min) and gently mix cells. The BJ5183-AD is a recombinant proficient E. coli strain carrying the pAdEasy plasmid that provides the essential machinery for obtaining the recombinant adenovirus genome. 5. Transfer 40 μL of BJ5183-AD cells to each of the chilled microcentrifuge tube. Add 100 ng of the linear form of pShuttle-TuD. Do not add more than 6 μL of DNA to 40 μL of cells. 6. Mix by gently tapping the tube. Do not pipet the cell/DNA mixture up and down. 7. Carefully transfer the BJ5183-AD cell/DNA mixture into a cooled cuvette without introducing air bubbles.

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8. Setup of the electroporator as follows: 25 μF, 2.5 kV, 200 Ω. 9. Dry the cuvette with a tissue to avoid bypass, and place the cuvette into the electroporator chamber. 10. Immediately after the pulse, add 1 mL of 37  C SOC medium (no antibiotics) to the cuvette, gently mix up and down twice, and then transfer to a round-bottom culture tube. 11. Shake the transformed cells vigorously at 37  C for 1 h. 12. Spread 50, 100, and 850 μL of the transformed cell suspension onto each of pre-warmed LB-kanamycin agar plates. 13. Incubate the plates 12–16 h at 37  C. 14. Pick at least 10 of the smallest and best isolated colonies from the recombinant plate, and inoculate a 5-mL culture of LB broth containing kanamycin. 15. Incubate the cultures 12–16 h at 37  C with vigorous shaking. 16. Harvest 2 mL of the BJ5183-AD cells at 6000  g for 15 min. 17. Isolate recombinant plasmids from BJ5183-AD cells using the plasmid miniprep kit. 18. Confirm recombination between pShuttle-TuD and pAdEasy plasmids by PacI digestion profiling analysis. For this, digest 0.5 μg of recombinant plasmids with 5 units of PacI restriction enzyme at 37  C for 1 h. 19. Run electrophoresis at 200 V for 1 h. If recombination is successful, PacI digestion produces DNA fragments of about 30 kb and 3.0 kb (if recombination occurs between the left arms) or 4.5 kb (if recombination occurs at origins of replication) size. A standard of over 10 kb of large size of DNA (e.g., HindIII digest of lambda DNA) is required. Before the next step, the presence of TuD segments should be double-checked using positive PCR and/or restriction enzyme reactions. 3.4 Purification of Recombinant Adenoviral Genome

1. Amplify the purified recombinant adenoviral vector containing TuD (termed recTuD) into XL10-gold ultracompetent cells. These cells are optimized for large-sized plasmids, and minimize the background when analyzing PacI digestion. For this, prepare a 42  C water bath, pre-heated SOC medium (42  C), and pre-warmed LB-kanamycin agar plates. 2. Thaw 50 μL of XL10 cells on ice (approximately 10 min). 3. Add 2 μL of the β-mercaptoethanol (β-ME) supplied with the kit to the XL-10 cells, and gently swirl the contents of the tube. 4. Incubate the cells on ice for 10 min, swirling gently every 2 min. 5. Add 50 ng of recTuD DNA (in 5 μL) to the cells and swirl gently.

Adenoviral Vector-Mediated Expression of Tough Decoys

21

6. Incubate the cell/recTuD DNA mix on ice for 30 min. 7. Heat shock the XL10 cells by placing tubes in a 42  C water bath for 30 sec. 8. Return the tubes to ice for 2 min. 9. Add 0.9 mL of pre-heated SOC medium to each transformation reaction. 10. Transfer 1 mL of the transformed cell suspension to a 17 mm  100 mm round-bottom culture tube, and incubate the tube for 1 h at 37  C while shaking vigorously. 11. Spread up to 200 μL of the transformed cells on LB-kanamycin agar plates. 12. Incubate the plates for 12–16 h at 37  C. 13. To scale up the recTuD, pick single colony from a XL10 recombinant plate, and inoculate 3–5 mL starter culture of LB-kanamycin broth for 8 h at 37  C. 14. Dilute the starter culture 1/500 into 500 mL LB-kanamycin broth, and grow at 37  C for 12–16 h with vigorous shaking. 15. Harvest the transformed XL-10 cells at 6000  g for 15 min. 16. Purify recombinant adenoviral vectors using a transfectiongrade DNA preparation kit for large-size DNA and digest the recTuD with PacI to expose its inverted terminal repeats. Adding an endotoxin removal step in the DNA preparation step is recommended to improve transfection efficiency (see Note 3). 3.5 Generation of Infectious Adenovirus

1. Prepare the low passage and healthy AD-293 cells, a HEK293derived cell line with improved cell adhesion and plaqueforming properties, 1 d before transfection (7–8  105 cells per 60-mm tissue culture dish). 2. Transfect 5 μg of purifying PacI-digested recTuD using a low-toxic reagent, PEI. The volume of PEI used is based on a 1:3 ratio of DNA (μg) to PEI (μg) [12]. Non-liposomal reagents can also be used. 3. Incubate the culture plates at 37  C for 5–10 d (see Note 4).

3.6 Preparation and Amplification of Primary Viral Stock

1. To prepare the primary viral vector stock, collect culture medium (approximately 2 mL per plate) from the transfected AD-293 cells 5–10 d after transfection. 2. Wash the cell monolayer with 0.5 mL of PBS. 3. Detach the AD-293 transfected cells using a cell scraper. 4. Collect the cells by centrifugation at 2000  g for 15 min. If most transfected cells are floating, be sure to resuspend the cells in the cell culture medium, and pellet the cells by centrifugation at 2000  g for 15 min.

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5. Discard the most supernatant, and resuspend the cell pellet in 0.5 mL of PBS. 6. Perform six freeze-thaw cycles between a 80  C freezer and a 37  C water bath. When the virus suspension is completely dissolved, remove it from the water bath immediately, and shake the thawed suspension vigorously for 30 sec. 7. Centrifuge at 20,000  g at RT for 15 min to remove cell debris. Store the supernatant, the primary virus stock (P1), at 80  C for later use. 8. To amplify the titer of the P1 virus, seed fresh AD-293 cells 24 h before infection to form an approximately 70% cell monolayer the following day. 9. Discard the medium upon infection, and infect the AD-293 monolayer with 0.5 mL of P1 for 1 h at 37  C. 10. After infection, cells are washed twice with PBS and maintained in cell culture medium. 11. Repeat steps 9–11 three times until P4 virus is obtained, and scale up virus production on a large scale through P4 virus infection. Infection of AD-293 cells and amplification of vector virus can be monitored by GFP expression using an inverted fluorescence microscope. Infected AD-293 cells with P4 virus are harvested when a maximal local cytopathic effect is achieved (Fig. 2). 3.7 Purification of Virus

1. Place 1.5 mL of heavy CsCl solution on the bottom of an ultracentrifuge tube (5 mL seal tube). 2. Add 2.5 mL of Light CsCl solution slowly on top of the first layer. 3. Add 1 mL of virus supernatant to this, and centrifuge at 100,000  g for 1 h at 18  C .

Fig. 2 Cell images infected with recombinant adenovirus showing cytopathic effect. On day 7 of transfection, cells begin to show plaque-like cytopathy. Bright field (a) and fluorescence field (b), magnification, 100

Adenoviral Vector-Mediated Expression of Tough Decoys

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4. Collect the virus band between light CsCl and heavy CsCl layers with a syringe. Insert needle about 0.5 cm below viral band, and approach the tube vertically, press it, and twist the needle. Aspirate the virus band from inside the tube with the needle pointing upwards. 5. Two-cycle ultracentrifugation can be applied to additional enrichment of adenovirus. Set up second CsCl gradients (2 mL of Light CsCl solution +2 mL of Heavy CsCl solution). 6. Perform a second ultracentrifugation at 100,000  g for 18 h at 18  C . 7. Desalt the virus using a PD-10 Sephadex column. 8. Perform a titration of the adenoviral vectors (see Note 5). In general, mammalian cells can produce 10,000 copies of adenovirus per cell. For example, the average yield of 293 cells in a T150 flask is ~107; thus the theoretical final titer is up to 109 ~ 1010 (see Note 6). 3.8 Luciferase Reporter Assay

1. Insert the sequence of the 30 -UTR region of the mRNA target containing the predicted binding site of a given miRNA (termed WT-UTR) into the downstream of the luciferase reporter reading frame of pMirTarget reporter vector. A mutation-predicted binding site (termed MUT-UTR) is also inserted into the pMirTarget luciferase reporter vector. 2. Seed 5  105 H9c2 embryonic cardiomyocytes in antibioticfree medium using a 6-well plate and grow to about 80% confluence. 3. Wash cells with 1.0 mL of PBS. 4. Change cell culture media to 1.0 mL of serum-free media without antibiotics per well. 5. Prepare transfection complexes by mixing 2–5 μL of transfection reagents and 0.5–2 μg of DNA to a total volume of 250 μL serum-free media. DNAs: WT-UTR (1 μg) + pShuttle-TuD (0 or 0.5 or 1 μg), WT-UTR (1 μg) + negative control, MUT-UTR (1 μg) + pShuttle-TuD, MUT-UTR (0 or 0.5 or 1 μg) + negative control. The MUT-UTR is used to evaluate the effect of a given miRNA on the expression of a reporter gene (see Note 7). 6. Incubate for 30 min at RT to allow DNA/transfection reagent complexes to form. 7. Add 250 μL of transfection complex to the cells, in a dropwise fashion all over the well. Gently rock the plate back and forth to ensure uniform transfection. Perform transfections in duplicate/triplicate.

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8. Incubate the cells at 37  C in a humidified CO2 incubator for 48 h. At 4–6 h post transfection, add 2 mL of cell culture medium, or replace the medium containing the transfection complex with 3 mL of fresh cell culture medium. 9. Remove the medium, and wash the cells carefully with PBS to avoid dissociation of the cells to be analyzed. 10. Collect the H9c2 cells by centrifugation at 2000  g for 15 min. 11. Add 200 μL of passive lysis buffer, and scrape the H9c2 cells from the plate. 12. Transfer cell lysates to a 1.7 mL microcentrifuge tube. 13. Vortex the tube for 15 sec, and centrifuge at 16,000  g for 5 min at 4  C to remove cell debris. Transfer the supernatant to a new microcentrifuge tube. 14. For luciferase activity, add 20 μL of cell lysate per well into the luminometer plate. 15. Read on a luminometer using luciferase assay substrate thawed and mixed at RT. The value is RLU (relative light unit). 16. Measure the protein concentration in 20 μL of the cell lysate. Use this value to normalize the luciferase value (calculate RLU/μg of protein). 3.9 Quantitative Real-Time PCR Analysis

1. Prepare freshly isolated and cultured adult cardiomyocytes (ACMs) [13]. Viral infection is possible after 1 h of incubation to achieve cardiomyocyte adhesion. 2. Aspirate the culture medium with unattached cardiomyocytes, and add a half-volume of FBS-free medium (e.g., 2.5 mL for 60-mm culture dishes) containing appropriate titers of adenovirus expressing control miRNA or TuD to the culture dish (see Note 8). 3. After an additional 1–2 h incubation, add another half-volume of culture medium without FBS. 4. After 24 h, remove the cell culture medium from infected cells, and wash the cells twice with PBS. Place the culture dishes on ice. 5. Isolate total cardiac RNAs, including miRNAs, using a miRNA isolation kit. 6. Determine the concentration and purity of purified RNA by measuring UV absorbance in a spectrophotometer. 7. Synthesize cDNA from isolated cardiac RNAs using a miRNA cDNA synthesis kit. For this, prepare poly (A) tailing reaction: mix 1 μg of miRNA (or total RNA), 5 units (typically 1 μL) of poly(A) polymerase, 2 μL of tailing reaction buffer (5), and nuclease-free water to a final volume of 10 μL.

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8. Incubate the reaction mix at 37  C for 1 h. 9. Inactivate the reaction by heating at 70  C for 5 min (e.g., PCR thermocycler). 10. Prepare first-strand cDNA synthesis reaction: mix 10 μL of poly (A) tailing reaction from step 8, 1 μL of reverse transcriptase (typically, 200 unit of M-MuLV enzyme), 9 μL of miRNA cDNA reaction mix consisting of Oligo(dT)/random hexamers, Rnase inhibitor, dNTPs, DTT, and reaction buffer to a final volume of 20 μL. 11. Incubate the reaction at 42  C for 20 min and then at 85  C for 5 min (e.g., PCR thermocycler). 12. Prepare qRT-PCR reaction mix as follows: 10 μL of SYBR Green Master Mix (2), 0.4 μL of each target cardiac miRNA primer (final conc. 200 nM) and 1–50 ng of cDNA from step 11 (up to 9.2 μL) to a final volume of 20 μL. 0.4 μL of ROX reference dye is optional. Use 18 s rRNA as a housekeeping gene. 13. Run the qRT-PCR reaction in a thermocycler. Perform a threestep cycling protocol as follows: Pre-incubation at 95  C for 2 min. 40 cycles of denaturation at 95  C for 5 sec, annealing at 60  C for 15 sec, extension at 72  C for 15 sec. Run all reactions at least in duplicate. Cq values >40 are considered negative, and check the melting point curves for all assays to confirm primer specificity. 14. Calculate the delta Ct values (relative expression) after normalizing the target to a reference gene, such as 18S rRNA, to determine the expression level of specific microRNAs, including the TuD target and its downstream genes. 3.10 Western Blotting

1. Prepare ACMs with or without TuD overexpression as described above. 2. Aspirate the medium and wash the cells once with ice-cold PBS. From this point on, keep ACMs and lysate on ice. 3. Add ice-cold RIPA buffer (e.g., 0.5 mL for 60-mm culture dishes) freshly supplemented with protease/phosphatase inhibitor cocktails (typically, a final concentration of 1% (v/v)) (see Note 9). 4. Scrape ACMs off the dish using a cell scraper, and transfer the ACM suspension into a pre-cooled microcentrifuge tube. 5. Incubate the tube on ice-cold for 30 min. Physical cell lysis steps, including homogenization and sonication, are optional. 6. Centrifuge the ACM suspension at 16,000  g for 15 min at 4  C.

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7. Collect the ACM supernatant (ACM lysate) in fresh microcentrifuge tube. Discard the pellet (cell debris). 8. Measure the protein concentration in the ACM lysate using the bicinchoninic acid (BCA) assay. For this, prepare 1 mg/mL bovine serum albumin (BSA) in RIPA buffer (BSA stock). 9. Make 6 BSA dilution as follows: Blank (100 μL RIPA), 1 μg/ 10 μL (10 μL BSA stock +90 μL RIPA), 2.5 μg/10 μL (25 μL BSA stock +75 μL RIPA), 5 μg/10 μL (50 μL BSA stock +50 μL RIPA), 7.5 μg/10 μL (75 μL BSA stock +25 μL RIPA, 10 μg/10 μL (100 μL BSA stock). This generates a standard curve. 10. For each ACM lysate, make 30 μL dilutions with RIPA buffer (1:10 dilution: 3 μL ACM lysate +27 μL RIPA). 11. Put 10 μL of each BSA standard and duplicates for each diluted ACM lysate (2 wells per sample) in a clear bottom 96-well plate. 12. Add 200 μL of working reagents provided by the kit to each of the wells. 13. Incubate plate in dark at 37  C for 30 min (e.g., covered with foil). 14. Cool the plate to RT, and read absorption at 562 nm with a plate reader. 15. Calculate protein concentration in the ACM lysate using the BSA standard curve and dilution factor. 16. Prepare sample for denaturing gel. Dilute ACM lysate with Laemmli buffer (4:1) supplied with a reducing reagent (e.g., mix the following components in a final volume 20 μL: 14 μL ACM lysat, 5 μL Laemmli buffer and 1 μL β-ME). 1.0-mmthick, 10-well combs will hold up to ~35 μL. 17. Heat the protein sample at 95–100  C for 5 min. 18. Place the tube on ice for 2 min. 19. Centrifuge the sample at 16,000  g for 1 min at 4  C. 20. Prepare pre-made gels with the appropriate polyacrylamide ratio based on molecular weight. A 4–20% gradient gel can separate a wide range of molecular weight sizes and commercially available. 21. Separate 10–50 μg of protein lysates on an SDS-PAGE gel. Include a pre-stained protein standard in a lane of the gel. Under typical condition, run gels in SDS-PAGE running buffer at 200 V (constant voltage) for 60–90 min or until the dye front reaches the bottom of the gel.

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22. Transfer the gel to blotting membranes (termed blot). Set up transfer apparatus, and transfer in transfer buffer at 300 mA/ 100 V for 1 h or 30 V for 12–16 h. 23. Block the blot with blocking buffer for 30 min at RT on a rocker-shaker. 24. Place the blot into 10 mL primary antibody (target cardiac proteins) diluted in a blocking buffer in a smallest tray on a rocker-shaker for 12–16 h at 4  C. 25. Wash the blot 3 times for 10 min each with 10 mL TBST on a rocker-shaker. 26. Place the blot with 10 mL HRP-conjugated secondary antibody diluted in blocking buffer for 60 min at RT on a rockershaker. 27. Wash the blot five times for 5 min each with 10 mL TBST on a rocker-shaker. 28. Wash the blot once for 5 min with 10 mL TBS on a rockershaker. 29. Remove excess TBS from the blot, and add the detection solution (ECL, enhanced chemiluminescent). For a minisized blot, 8–10 mL of detection solution is sufficient (typically, 0.1 mL per cm2 of membrane). 30. Incubate for 1–5 min at RT. 31. Remove the blot from the detection solution, and place the blot on a sample stage of the imaging system. The chemiluminescent signal is captured, recorded, and analyzed by digital imaging system (see Note 10). 32. Quantify the protein bands by suitable image analysis tools. For analysis of small-sized proteins (e.g., less than 10 kDa) or largesized proteins (>100 kDa), optimal transfer conditions should be tested. The sensitivity and specificity of Western blotting depend on several factors (see Note 11).

4

Notes 1. Generation of Tough Decoy Sequences Many TuD designs and prototypes have been published, and the most well-known decoy is a hairpin-sharped structure. Takeshi et al. published an optimal structure of TuD that contains two MBS domains connected by three-nucleotide linker to two flanked stem structures [11]. The MBS can be a perfect antisense MBS or a bulge on the MBS. Several web-based tools help design TuDs. Bioinformatics prediction algorithms (e.g., StarMir, TargetScan, miRmap, RNA hybrid, and miRNAsong) can be used to identify/verify targets and

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optimize TuD sequences. For example, StarMir, miRNAsong, and miRmap predict MBS potency designed within TuDs by computational analysis for free energy changes caused by miRNA-MBS interaction. Also, the miRmap algorithm provides all miRNAs that bind to the designed TuD sequence. These sequences can be modified to minimize off-target miRNA binding based on this information. Nucleotide substitution, sequence, and length of the linker can be determined by in silico analysis. RNAfold web server provides the secondary structure of TuD RNA. Moreover, several vectorized miRNA TuD RNAs are commercially available (e.g., Addgene, SigmaAldrich, OriGene, etc.) and directly synthesized on request. 2. Construction of a Shuttle Vector The designed TuD segment of the target miRNA containing U6 promoter is cloned into the pShuttle-IRES-hrGFP vector using the appropriate restriction enzyme site(s) in the multiple cloning site. However, due to the manipulation of the vector system, the PmeI and PacI target sites should not be present in the insert. If necessary, site-directed mutagenesis can be performed. 3. Preparation of Endotoxin-Free Plasmid Endotoxins are the cell membrane components of Gramnegative bacteria, such as E. coli. Endotoxins are released during the lysis of plasmid purification, reducing the transfection efficiency. Kits for endotoxin removal are commercially available (Qiagen, Invitrogen, Promega, etc.). Additionally, negative endonuclease strains, such as NEB stable competent E. Coli, can be used to obtain high-quality endotoxin-free plasmid. 4. Generation of Adenovirus It is necessary to verify adenovirus production through the reporter system after transfecting the PacI-digested recombinant adenovirus plasmid DNA into AD293 cells. Post 5–7 d transfection, transfected cells must show reporter expression and detach. If those observations are not seen, it is better to transfect more PacI-digested recombinant adenovirus plasmid DNA again. 5. Titration of Adenovirus Adenoviral titration can be divided into two types: physical titration and biological titration. Physical viral particles can be determined by measuring optical density (OD) at a wavelength of A260 nm. A conversion factor is then used that counts the number of virus particles per milliliter (VP/mL). Plaqueforming unit analysis, 50% tissue culture infectious dose assay, and infectious units analysis are classical biological methods. For example, infectious units can be determined by limiting the

Adenoviral Vector-Mediated Expression of Tough Decoys

29

dilution colony-counting method. The infected cultures are stored at 37  C in a CO2 incubator for 3 d. Vector titer is determined by counting the number of GFP-positive cells at the endpoint dilution as follows: vector titer (IU/mL) ¼ numbers of GFP positive cells  5 (IF)  DF, where IU, infectious unit, IF, inoculum factor, and DF, dilution factor. Biological efficacy can also be assigned based on infective genome titration using quantitative PCR [14]. 6. Replication-Competent Adenovirus Screening It is recommended to perform routine screening for wildtype contamination of recombinant adenovirus by detecting adenovirus E1 DNA [15]. 7. Assessment of TuD Specificity TuD specificity is assessed using control miRNAs by introducing mismatches into the TuD sequence or using scrambled sequences. Ideally, both control reporters with mutated MBS and mismatched or scrambled oligonucleotides should be included as specificity controls when performing luciferase assays. However, validated miRNA-negative controls, random sequences that do not produce identifiable effects on known miRNA functions, can be used as an alternative; they are commercially available. 8. Assessment of TuD’s Off-Target Effect qRT-PCR is used to assess the potential for off-target activity of TuDs by determining the relative abundance of various miRNAs and other miRNA-related families. Also, genome-wide transcriptional or proteomic approaches are used for more comprehensive analysis. 9. Preparation of Protein Lysates Cell lysis buffer contains various detergents that help release soluble proteins (e.g., Triton-X, Tween, NP-40, SDS, CHAPS). However, depending on the cellular location of the target protein, different lysis buffers are required. The most commonly used are RIPA and NP-40. RIPA buffers are useful for whole-cell extracts and membrane-bound proteins and may be preferable to NP-40-only buffers for nuclear protein extraction. In addition, subcellular fractionation is often used to increase the target protein concentration. 10. Reprobing Blots Signals from the blot can be removed with primary and secondary antibodies through stripping buffer (e.g. stripping by low pH) and re-probed with actin or GAPDH for loading control. However, multiple stripping/reprobing is not recommended as it causes significant sample loss and reduces detection sensitivity.

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11. Sensitivity and Specificity of Western Blotting Several factors affect the sensitivity and specificity of Western blotting. Since Western blotting is antibody-based detection, the quality of the antibody and optimal antibody-antigen reaction conditions are the most important factors. However, to obtain accurate results, the detection method, type of transfer membrane (e.g., nitrocellulose, PVDF), blotting reagents, and quantitative analysis method are also important.

Acknowledgments This work was supported by a 2-Year Research Grant of Pusan National University. References 1. Lewis BP, Burge CB, Bartel DP (2005) Conserved seed pairing, often flanked by adenosines, indicates that thousands of human genes are microRNA targets. Cell 120:15–20. https://doi.org/10.1016/j.cell.2004.12.035 2. Friedman RC, Farh KK, Burge CB et al (2009) Most mammalian mRNAs are conserved targets of microRNAs. Genome Res 19:92–105. https://doi.org/10.1101/gr.082701.108 3. Small EM, Olson EN (2011) Pervasive roles of microRNAs in cardiovascular biology. Nature 469:336–342. https://doi.org/10.1038/ nature09783 4. Nouraee N, Mowla SJ (2015) miRNA therapeutics in cardiovascular diseases: promises and problems. Front Genet 6:232. https://doi. org/10.3389/fgene.2015.00232 5. Stenvang J, Petri A, Lindow M, Obad S (2012) Inhibition of microRNA function by antimiR oligonucleotides. Silence 3:1. https://doi.org/ 10.1186/1758-907X-3-1 6. Crooke ST, Liang XH, Baker BF, Crooke RM (2021) Antisense technology: a review. J Biol Chem 296:100416. https://doi.org/10. 1016/j.jbc.2021.100416 7. Bennett CF (2019) Therapeutic antisense oligonucleotides are coming of age. Annu Rev Med 70:307–321. https://doi.org/10.1146/ annurev-med-041217-010829 8. Ebert MS, Neilson JR, Sharp PA (2007) MicroRNA sponges: competitive inhibitors of small RNAs in mammalian cells. Nat Methods 4:721–726. https://doi.org/10.1038/ nmeth1079 9. Xie J, Ameres SL, Friedline R et al (2012) Long-term, efficient inhibition of microRNA

function in mice using rAAV vectors. Nat Methods 9:403–409. https://doi.org/10. 1038/nmeth.1903 10. Hollensen AK, Bak RO, Haslund D, Mikkelsen JG (2013) Suppression of microRNAs by dualtargeting and clustered tough decoy inhibitors. RNA Biol 10:406–414. https://doi.org/10. 4161/rna.23543 11. Haraguchi T, Ozaki Y, Iba H (2009) Vectors expressing efficient RNA decoys achieve the long-term suppression of specific microRNA activity in mammalian cells. Nucleic Acids Res 37:e43. https://doi.org/10.1093/nar/ gkp040 12. Longo PA, Kavran JM, Kim MS, Leahy DJ (2013) Transient mammalian cell transfection with polyethylenimine (PEI). Methods Enzymol 529:227–240. https://doi.org/10.1016/ B978-0-12-418687-3.00018-5 13. Gorski PA, Kho C, Oh JG (2018) Measuring cardiomyocyte contractility and calcium handling in vitro. Methods Mol Biol 1816: 93–104. https://doi.org/10.1007/978-14939-8597-5_7 14. Gallaher SD, Berk AJ (2013) A rapid Q-PCR titration protocol for adenovirus and helperdependent adenovirus vectors that produces biologically relevant results. J Virol Methods 192:28–38. https://doi.org/10.1016/j. jviromet.2013.04.013 15. Zhang WW, Koch PE, Roth JA (1995) Detection of wild-type contamination in a recombinant adenoviral preparation by PCR. BioTechniques 18:444–447

Chapter 3 Direct Reprogramming of Adult Human Cardiac Fibroblasts into Induced Cardiomyocytes Using miRcombo Camilla Paoletti, Carla Divieto, and Valeria Chiono Abstract Direct reprogramming of fibroblasts into induced cardiomyocytes (iCMs) through microRNAs (miRNAs) is a new emerging strategy for myocardial regeneration after ischemic heart disease. Previous studies have reported that murine fibroblasts can be directly reprogrammed into iCMs by transient transfection with four miRNAs (miRs-1, 133, 208 and 499 – termed “miRcombo”). While advancement in the knowledge of direct cell reprogramming molecular mechanism is in progress, it is important to investigate if this strategy may be translated to humans. Recently, we demonstrated that miRcombo transfection is able to induce direct reprogramming of adult human cardiac fibroblasts (AHCFs) into iCMs. Although additional studies are needed to achieve iCM maturation, our early findings pave the way toward new therapeutic strategies for cardiac regeneration in humans. This chapter describes methods for inducing direct reprogramming of AHCFs into iCMs through miRcombo transient transfection, showing experiments to perform for assessing iCM generation. Key words microRNAs, Transient transfection, Direct cell reprogramming, Human cardiac fibroblasts, Induced cardiomyocytes

1

Introduction Myocardial infarction (MI) is the leading cause of death worldwide, causing massive loss of cardiomyocytes (CMs), inflammation and the formation of a non-functional fibrotic scar [1]. The poor regenerative capacity of adult cardiac tissue is the major barrier for heart regeneration. After MI, fibroblasts are the dominant cell type populating the cardiac scar [2]. Therefore, direct reprogramming of fibroblasts into iCMs provides a good therapeutic strategy to restore cardiac functions by repopulating cardiac scar with functional CMs [3]. In 2012, Jayawardena et al. have reported a

The original version of this chapter was previously published non-open access. A Correction to this chapter is available at https://doi.org/10.1007/978-1-0716-2707-5_27 Kiyotake Ishikawa (ed.), Cardiac Gene Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2573, https://doi.org/10.1007/978-1-0716-2707-5_3, © The Author(s) 2022, Corrected Publication 2023

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Fig. 1 Experimental outline for microRNA-mediated reprogramming of AHCFs into iCMs experiments including cell seeding and miRNA transient transfection using DharmaFECT1. We also indicated the expected timeline for induction of cardiomyocyte TFs (day 7), cardiomyocyte and fibroblast markers assessed by ddPCR expression or flow cytometry (day 15) and calcium transients analysis (day 30). (Figure created with Biorender.com under license)

combination of four miRNAs (miRcombo), able to induce direct reprogramming of mouse neonatal cardiac fibroblasts into iCMs [4]. Delivery of miRcombo was performed through cell transient transfection using the commercial transfection agent DharmaFECT1, and demonstrating that a single miRcombo dose may establish and maintain CM phenotype in cardiac fibroblasts [4]. Later, we have demonstrated that miRcombo is also able to directly reprogram AHCFs into iCMs, showing cardiomyocyte gene expression after 7 and 15 days of culture, 11% of reprogramming efficiency after 15 days of culture and spontaneous calcium transients after 30 days [5]. Here, we describe our protocol for inducing AHCF reprogramming into iCMs through single miRcombo transient transfection (Fig. 1).

2

Materials

2.1 Reagents for Cell Culture

1. Adult human cardiac fibroblasts (AHCFs) cryopreserved (Lonza). 2. Fibroblast expansion media: Fibroblast basal growth medium (FGM-3, Lonza), 10% fetal bovine serum (FBS), 0.1% insulin, 0.1% human fibroblast growth factor (hFGF), 0.1% gentamicin amphotericin (GA-1000).

Human Fibroblast Reprogramming into iCMs by MicroRNAs

33

3. Reprogramming media: Dulbecco’s Modified Eagle Medium (DMEM) with 4.5 g/L glucose (without L-glutamine), 10% fetal bovine serum, 1% L-glutamine, 1% penicillin/streptomycin (when specified in the text). 4. 0.05% Trypsin/EDTA. 2.2 Reagents for microRNA Transfection

1. DharmaFECT1 Transfection Reagent (Dharmacon).

2.3 Reagents for RNA Purification and cDNA Synthesis

1. QIAzol Lysis Reagent (QIAgen).

2. Synthetic mature miRNA mimics: hsa-miR-1a-3p, hsa-miR133a-3p, hsa-miR-208a-3p, hsa-miR-499a-5p, mirVana™ miRNA Mimic Negative Control #1 (mirVana).

2. Chloroform. 3. 70% ethanol in H2O stored at 4. 100% 2-propanol stored at

20  C.

20  C.

5. DNase- and RNase-free H2O. 6. High-Capacity cDNA Reverse Transcription Kit (Applied Biosystem). 7. miRCURY LNA RT Kit (Bio-Rad). 2.4 Reagents for Droplet Digital PCR

All reagents and instruments are from Bio-Rad: 1. DG8 Cartridges QX200. 2. DG8 Gaskets QX200. 3. QX200 Droplet Generator. 4. QX200 Droplet Reader. 5. QX200 droplet generation oil for EvaGreen assays. 6. Droplet generation oil for probes. 7. QX200 ddPCR EvaGreen Supermix. 8. ddPCR supermix for probes without dUTP. 9. ddPCR Gene Expression Assay (human) primers (Table 1).

2.5 Reagents for Flow Cytometry

1. 0.05% Trypsin/EDTA. 2. Phosphate buffer saline (PBS), 0.5% Tween-20 (v/v). 3. PBS, 10% FBS, 1% sodium azide. 4. Primary antibody for cardiac troponin T (Invitrogen). 5. Fluorescent-labelled secondary antibody.

2.6 Reagents for Calcium Transient Imaging

1. Modified Tyrode’s solution: 140 mM NaCl, 5 mM KCl, 1.8 mM CaCl2, 1 mM MgCl2, 10 mM glucose, 10 mM Hepes, in 2 mL of H2O, 0.1% bovine serum albumin (BSA).

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Table 1 List of markers analysed at different time points: cardiomyocyte TFs (GATA4, MEF2C, TBX5, HAND2 and NKX2.5), cardiomyocyte mature markers (TNNT2 and TNNI3), fibroblast markers (VIM, DDR2 and FSP-1), housekeeping gene (GAPDH) and miRNA-1 (miR-1) Official gene symbol

Official full name

GATA4

GATA binding protein 4

MEF2C

Myocyte enhancer factor 2C

TBX5

T-box transcription factor 5

HAND2

Heart and neural crest derivatives expressed 2

NKX2.5

NK2 Homeobox 5

TWF-1

Twinfilin actin binding protein 1

TNNT2

Cardiac troponin subunit T2

TNNI3

Cardiac troponin subunit I3

VIM

Vimentin

DDR2

Discoidin domain receptor 2

FSP-1

Fibroblast-specific-protein 1

GAPDH

Glyceraldehyde-3-phosphate dehydrogenase

miR-1

Hsa-miR-1-3p

2. Fluo-4, cell permeant (Invitrogen) used at final concentration of 5 μM in Dimethyl Sulfoxide (DMSO). 3. 35 mm cell treated-bottom μ-dishes (Ibidi).

3

Methods

3.1 Culture of Adult Human Cardiac Fibroblasts

1. Under laminar flow hood, for each cryovial containing the frozen cells, prepare 6 mL of Fibroblast expansion media in a sterile 15-mL conical tube and place it at 37  C. 2. Remove the cryovial from liquid nitrogen and place it in a 37  C water bath. Transfer the cells into the tube containing the pre-warmed complete fibroblast expansion media from previous step. 3. Centrifuge at 1000 g for 5 minutes at room temperature, then discard the supernatant and resuspend the cell pellet with fresh complete fibroblast expansion media Plate the cells in a 10-cm Petri dish and place in 37  C incubator with humidified atmosphere of 5% CO2. 4. When cells are 80% confluent, split them from passage 1 (P1) to P2 at a ratio of 1:3 by using 1 mL of 0.05% Trypsin/EDTA.

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Table 2 Table showing volume of microRNAs, DharmaFECT1 and serum- free medium for preparing transfection mix, according to well formats (see Note 2). Table shows also AHCF seeding densities required for setting up miRcombo-mediated reprogramming experiments in 6-well, 12-well and 35 mm dish Well format

Seeding density

6

110.000 cells/ well

55.000 12 or cells/ 35 mm well dish

Volume of Volume of antibiotic-free transfection mix medium

Final volume of culture medium

Tube 1

Tube 2

10 μL of microRNAs +190 μL of DMEM

400 μL 6 μL of DharmaFECT1 + 194 μL of DMEM

1,6 mL

2 mL

5 μL of microRNAs + 95 μL of DMEM

200 μL 2.5 μL of DharmaFECT1 + 97.5 μl of DMEM

800 μL

1 mL

Subsequently, split cells until P5. Change media every two days. Do not conduct microRNA transfections experiments in cells beyond P5. 3.2 AHCF Transfection with microRNAs

1. Plate AHCFs in antibiotic-free reprogramming media. Transfections can be performed in 6-well (RNA), 12-well (flow cytometry) or 35-mm dish (calcium transient) format, depending on the nature of the experiment. 2. Set up the transfection reaction under completely antibioticfree medium conditions. Start by preparing two tubes: (tube 1) dilution of microRNAs 5 μM in serum-free DMEM and (tube 2) dilution of DharmaFECT1 in serum-free DMEM. Use the proportions outlined in the table below (Table 2). Incubate at room temperate for 5 minutes. Then, add tube 1 content into tube 2, mix twice and incubate at room temperature for 20 min (transfection mix). In order to compare and validate the relative induction of cardiac reprogramming, transfect cells in controls versus microRNA-treated samples. 3. During the incubation time, change medium on cells to the appropriate volume of antibiotic-free reprogramming media as indicated in the table below (Table 2). 4. Add transfection mix directly to cells. 5. After 24 h, remove medium, rinse cells with PBS and add fresh complete reprogramming media. Continue cell culture up to desired time point (2, 7, 15 or 30 days post-transfection).

3.3

RNA Extraction

1. Remove medium from cells without washing the cells. 2. Under chemical hood, add 1 mL of QIAzol to each well and homogenize the cells. Collect cell lysate in 2-mL tube and either freeze lysate at 80  C or proceed immediately to RNA purification. During all steps, take samples in ice bath.

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3. Add 200 μL of chloroform, vortex and centrifuge RNA at 12400 g at 4  C for 30 min. 4. Ensure efficient phase separation of samples and transfer the upper aqueous phase into a fresh 2-mL tube. 5. Add 500 μL of ice-cold 2-propanol, invert tube twice and store at 20  C overnight. 6. Centrifuge at 12400 g for 30 minutes at 4  C. The RNA precipitate will form a pellet on the side and bottom of the tube. 7. Remove the supernatant and wash pellets twice with 1 mL of ice-cold 70% ethanol at 7400 g for 20 min at 4  C. 8. Remove supernatant and briefly dry the RNA pellet for 10 min by air-drying or under a vacuum. 9. Add an appropriate volume of DNase and RNAse free H2O and store at 20  C. 10. Assess RNA concentration and quality by absorbance ratio at 260 nm/280 nm (A260/A280) and 260 nm/230 nm (A260/ A230) wavelength. RNA concentration should be around 400 ng/μl with typical requirements of A260/A280 ratio >1.8–2.2 and A260/A230 ratio >1.7. 3.4 Evaluation of Transfection Efficiency: Day 2

1. To study effective miRNA transfection, plate AHCFs in 6-multiwell in antibiotic-free reprogramming media. 2. The day after, transfect cells with miR-1 and negmiR using DharmaFECT1 following Table 2 (see Note 1). Use untransfected AHCFs as controls. 3. After 24 h from transfection, remove medium, wash cells with PBS and incubate cells with fresh complete reprogramming media. 4. Perform RNA extraction as described in Subheading 3.3. 5. Perform miRNA reverse-transcription using miRCURY LNA RT Kit, diluting RNA samples to 5 ng/μl in DNase and RNAse free H2O and preparing the reaction following the manufacturer’s instructions. 6. Perform ddPCR analysis to assess the expression of miR-1 using EvaGreen supermix and primer for miR-1, following manufacturer’s instructions. 7. Set thermal-cycling conditions: 95  C for 5 minutes (1 cycle), 95  C for 30 s and 55  C for 1 minute (40 cycles), 90  C for 5 minutes (1 cycle), and a 4  C infinite hold. Load PCR plate on Bio-Rad QX200 droplet reader for quantification of cDNA copies/μl and analyze data using QuantaSoft analysis software. 8. In parallel, prepare cDNA synthesis to assess TWF-1 (miR-1 target) knockdown using High capacity cDNA kit. Dilute RNA

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to 200 ng in DNase and RNAse free H2O and prepare the reaction following the manufacturer’s instructions. 9. Perform ddPCR analysis using ddPCR supermix for probes (without dUTP) and primers for TWF-1. Use GAPDH primer as housekeeping gene. 10. Set thermal-cycling conditions: 95  C for 10 min (1 cycle), 94  C for 30 s and 55  C for 30 s (40 cycles), 98  C for 10 minutes (1 cycle), and a 4  C infinite hold. Load PCR plate on Bio-Rad QX200 droplet reader for quantification of cDNA copies/μL and analyze data using QuantaSoft analysis software. 11. Expected miR-1 expression in miR-1 transfected cells should be around 500-fold compared to controls. Expected TWF-1 mRNA expression in miR-1 transfected AHCFs should be between 10% and 20% compared to negmiR controls and untransfected cells. 3.5 Evaluation of Cell Reprogramming: Day 7

1. To study cardiomyocyte transcription factor (TF) expression in miRcombo-transfected cells, plate AHCFs in 6-multiwell in antibiotic-free reprogramming media. The day after, transfect cells with miRcombo and negmiR using DharmaFECT1 following Table 2. Use untransfected AHCFs as controls (see Note 3). After 24 h from transfection, remove medium, wash cells with PBS and incubate cells with fresh complete reprogramming media. 2. After 7 days of culture, harvest RNA and synthesize total cDNA using High capacity cDNA. Dilute RNA to 200 ng in DNase and RNAse free H2O and preparing the reaction following the manufacturer’s instructions. 3. For the first evidence of fibroblast reprogramming into iCMs in miRcombo-transfected cells, assess the expression of GATA4, MEF2C, TBX5, HAND2 and NKX2.5 cardiomyocyte TF expression performing ddPCR using ddPCR supermix for probes (without dUTP). Set thermal-cycling conditions as reported in Subheading 3.4, step 10. With the exception of NKX2.5 TF, whose upregulation is two-fold higher compared to controls, the expression of cardiomyocyte TFs in miRcombo-transfected cells is expected to be three- to sixfold higher compared to negmiR and untransfected control cells.

3.6 Evaluation of Cell Reprogramming: Day 15

1. To study cardiomyocyte and fibroblast marker expression and reprogramming efficiency after 15 days of culture, plate cells for RNA extraction (6-multiwell) and flow cytometry (12-multiwell) in antibiotic-free reprogramming media. The day after, transfect cells with miRcombo and negmiR using

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DharmaFECT1 following Table 2. Use untransfected AHCFs as controls. After 24 h from transfection, remove medium, wash cells with PBS and incubate cells with fresh complete reprogramming media. 2. After 15 days of culture, harvest RNA and synthesize total cDNA using High capacity cDNA. Dilute RNA to 200 ng in DNase and RNAse free H2O and preparing the reaction following the manufacturer’s instructions. 3. For mature cardiomyocyte markers, assess the expression of TNNT2 and TNNI3 genes through ddPCR using ddPCR supermix for probes (without dUTP). Set thermal-cycling conditions as reported in Subheading 3.4, step 10. Cardiomyocyte gene expression in miRcombo transfected cells should reach two- to three-fold compared to negmiR and untransfected controls. For fibroblast-associated markers, assess the expression of VIM, DDR2 and FSP-1 genes. Expected fibroblast gene expression in miRcombo transfected cells should be between 30% and 60% compared to negmiR and untransfected control cells. 4. In parallel, perform flow cytometry analysis to assess reprogramming efficiency in miRcombo and negmiR transfected cells. 5. Remove medium and wash cells with PBS. Trypsinize cells with 500 μL of 0.05% Trypsin/EDTA and permeabilize cells with 0.5% v/v Tween 20 in PBS for 5 minutes. 6. Wash cells with ice cold PBS with 10% FBS and 1% sodium azide. 7. Incubate cells with Cardiac Troponin T primary antibody for 1 h at 4  C. Then, wash samples twice in PBS with 10% FBS and 1% sodium azide and centrifuge at 1100 g for 5 min. 8. Incubate samples with fluorescently-labelled secondary antibody (choose proper fluorophore according to laser excitation wavelength) for 1 h at 4  C in the dark. Then, wash samples twice in PBS with 10% FBS and 1% sodium azide and centrifuge. 9. Evaluate cell reprogramming using flow cytometry. Percentage of cTnT+ miRcombo transfected cells is expected between 9% and 12% compared to negmiR control cells, which do not show cTnT+ cells. Use control cells stained only with secondary fluorescent antibody to set background fluorescence. 3.7 Evaluation of Cell Reprogramming: Day 30

1. To analyse calcium transient in miRcombo-transfected AHCFs, plate cells in 35 mm cell treated-bottom μ-dishes in antibioticfree reprogramming media. The day after, transfect cells with miRcombo and negmiR using DharmaFECT1 and following

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Table 2. Use untransfected AHCFs as controls. After 24 h from transfection, remove medium, wash cells with PBS and incubate cells with fresh complete reprogramming media. 2. After 30 days of culture, remove medium and wash cells twice with PBS. Load cells with 5 μM of Fluo-4 AM in modified Tyrode’s solution at 37  C for 30 min while shielded from light. 3. Wash cells twice in modified Tyrode’s solution and incubate them at 37  C for 30 min to allow complete de-esterification of intracellular compounds. 4. Record calcium transient using fluorescence microscope and performing high-speed time lapse lasting at least 2 min. 5. Analyse calcium transients using ImageJ software (NIH) and report calcium level as F/F0 ratio, where F is the intensity of fluorescence emission recorded for each cell, while F0 is the background fluorescence.

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Notes 1. To rapidly assess cell transfection efficiency with DharmaFECT1, fluorescent-labelled microRNA mimic can be used. Cell transfection can be assessed 24 h post transfection using flow cytometry. Use cells incubated with fluorescent-labelled microRNA (without DharmaFECT1) and untransfected cells as negative controls. 2. It is recommended to prepare 50 μM miRNA mimic stock solution in nuclease free H2O. This will reduce miRNA degradation during freeze-thaw cycles. 3. In the case of miRcombo transfection (miR-1, miR-133a, miR-208a, miR-499a-5p), add ¼ of the final volume indicated in the Table 2 for each miRNAs.

Funding The project has received funding from the European Research Council (ERC) under the European Union’s Horizon 2020 Research and Innovation Programe (Grant Agreement No. 772168).

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References 1. Talman V, Ruskoaho H (2016) Cardiac fibrosis in myocardial infarction-from repair and remodeling to regeneration. Cell Tissue Res 365: 563–581 2. Hinderer S, Schenke-Layland K (2019) Cardiac fibrosis – a short review of causes and therapeutic strategies. Adv Drug Deliv Rev 146:77–82 3. Sadahiro T, Yamanaka S, Ieda M (2015) Direct cardiac reprogramming. Circ Res 116: 1378–1391

4. Jayawardena TM, Egemnazarov B, Finch EA et al (2012) MicroRNA-mediated in vitro and in vivo direct reprogramming of cardiac fibroblasts to cardiomyocytes. Circ Res 110: 1465–1473 5. Paoletti C, Divieto C, Tarricone G et al (2020) MicroRNA-mediated direct reprogramming of human adult fibroblasts toward cardiac phenotype. Front Bioeng Biotechnol 8(529):1–14

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 4 CRISPR/Cas9 Gene Editing of RYR2 in Human iPSC-Derived Cardiomyocytes to Probe Ca2+ Signaling Aberrancies of CPVT Arrhythmogenesis Naohiro Yamaguchi, Xiao-Hua Zhang, and Martin Morad Abstract Human-induced pluripotent stem cells (hiPSCs) provide a powerful platform to study biophysical and molecular mechanisms underlying the pathophysiology of genetic mutations associated with cardiac arrhythmia. Human iPSCs can be generated by reprograming of dermal fibroblasts of normal or diseased individuals and be differentiated into cardiac myocytes. Obtaining biopsies from patients afflicted with point mutations causing arrhythmia is often a cumbersome process even when patients are available. Recent development of CRISPR/Cas9 gene editing system makes it, however, possible to introduce arrhythmiaassociated point mutations at the desired loci of the wild-type hiPSCs in relatively short times. This platform was used by us to compare the Ca2+ signaling phenotypes of cardiomyocytes harboring point mutations in cardiac Ca2+ release channel, type-2 ryanodine receptor (RyR2), since over 200 missense mutations in RYR2 gene appear to be associated with catecholaminergic polymorphic ventricular tachycardia (CPVT1). We have created cardiac myocytes harboring mutations in different domains of RyR2, to study not only their Ca2+ signaling consequences but also their drug and domain specificity as related to CPVT1 pathology. In this chapter, we describe our procedures to establish CRISPR/Cas9 gene-edited hiPSC-derived cardiomyocytes. Key words Genetic modification, Genetic mutation, Restriction fragment length polymorphism, Cardiac differentiation, Cardiac maturation medium, Catecholaminergic polymorphic ventricular tachycardia

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Introduction Heart disease remains as the leading cause of death worldwide in humans. Numerous factors including post-translation altered cardiac protein functions and/or inherited genetic mutations result often in lethal cardiac pathology. Most prevalent cardiac pathologies result from dysfunctional ion channel activities that alter cardiac action potential configurations, the speed of its conduction, the triggering calcium release, and the resultant contraction [1]. Large numbers of inherited mutations in cardiac ion channel

Kiyotake Ishikawa (ed.), Cardiac Gene Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2573, https://doi.org/10.1007/978-1-0716-2707-5_4, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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genes have been already reported that include missense mutations in surface membrane Na+, K+, and Ca2+ channels underlying the cardiac action potentials and the sarcoplasmic reticulum calcium release channel and the type-2 ryanodine receptors (RyR2). As of now, over 200 missense mutations in human RYR2 gene have been reported to associate with catecholaminergic polymorphic ventricular tachycardia (CPVT1), an arrhythmia induced by severe exercise or emotional stresses [2]. Inherited missense mutations of ion channels and their functional consequences have been examined either by construction and expression of mutant ion channel cDNA in the heterologous cells or by generating knock-in mouse models carrying the mutation. These approaches have advanced our understanding of cellular signaling mechanisms that underly cardiac ion channel pathologies but suffer from the drawbacks resulting from using non-muscle model platforms or non-human mouse cardiomyocytes. The development of human-induced pluripotent stem cells (hiPSCs) [3] may eventually overcome disadvantages of the traditional approaches, as it enables scientists to create human patient-specific stem cells from dermal fibroblast, adipose tissues, or blood cells and differentiate them into cardiomyocytes carrying patients’ specific genetic mutations. This approach when combined with CRISPR/Cas9 gene editing that introduces point mutations in wild-type cells [4] and bypasses harvesting patient biopsies may bring us closer to a model system where disease-associated mutations can be created and their pathophysiological phenotypes and drug sensitivities are compared. We have succeeded in combining these two techniques and have generated multiple hiPSC lines harboring RYR2 missense mutations [5–7]. Here we shall summarize CRISPR/Cas9 gene edit of hiPSCs and their differentiation into cardiomyocytes (hiPSC-CMs) and provide the reader with the modified procedures and technical tips that optimize mutant cardiomyocyte generations. More detailed general protocols and biological mechanisms of these methods can be found in other reports [4, 8, 9].

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Materials

2.1 Construction of Plasmid for RNAGuided Cas9 Exonuclease and CRISPR/Cas9 Gene Edit

1. The plasmid DNA for expressing Cas9 and cloning guide sequence (e.g., pX459). 2. Two single-strand oligonucleotides (20-mer) of guide sequence. 3. Plasmid construction enzymes. 4. Repair templates: (a) single-strand oligonucleotide with homologous DNA sequence of the target gene locus (~150bp) or (b) plasmid DNA carrying ~400-bp homologous sequence of the targeted gene.

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5. Wild-type human iPS cells. 6. StemPro™ Accutase™ Cell Dissociation Reagent (Thermo Scientific). 7. Electroporation medium (e.g., Neon electroporation system, Thermo Scientific). 8. StemFlex™ medium (Thermo Scientific). 9. Antibiotics for selection (e.g., 1 μg/ml puromycin). 10. Y27632 (ROCK inhibitor helping cell survival). 11. DNA extraction solution. 12. Taq polymerase and restriction enzymes (depend on the design of gene editing). 13. Agarose gel and TAE or TBE-based gel electrophoresis system. 14. PCR cloning kit. 2.2 Cardiac Differentiation of hiPSCs and Dissociation of hiPSCCMs

1. RPMI 1640 medium. 2. Insulin-free B27 supplement. 3. B27 supplement. 4. CHIR99021 (Wnt signaling activator). 5. IWR-1 (Wnt signaling inhibitor). 6. Fatty acid maturation medium: RPMI/B27(+) medium, 22.5 μM linoleic acid, 40.5 μM oleic acid, 52.5 μM palmitate, 23.8 μM BSA, 120 μM L-carnitine. 7. TrypLE™ Select Enzyme (10) (Thermo Scientific). 8. RevitaCell™ Supplement (100) (Thermo Scientific). 9. Matrigel diluted 75-fold with DMEM/F12 medium. 10. Vitronectin (diluted to 5 μg/ml concentration by DPBS).

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Methods CRISPR/Cas9 gene edit of hiPSCs is comprised from multiple Subheading (3.1, 3.2, and 3.3 below). In brief, RNA-guided Cas9 exonuclease recognizes Pam sequence (NGG) adjacent to the guide sequence and digests the double-stranded DNA (genome) at 3–4 bp upstream of the Pam sequence. The digested genome can be repaired by homology-dependent repair in the presence of DNA piece (called repair template) of the homologous sequence with the digested genome site. When the repair template carries a couple of mismatches of the nucleotide, such mismatches (mutations) can be incorporated in the host genome (see Fig. 1, Scheme1–3).

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Fig. 1 Schematic of CRISPR/Cas9 gene edit and screening of gene-edited clones. (Scheme 1) A desired gene locus is digested by Cas9 exonuclease. Cas9 is guided by RNA which sequence correspond to the 20-mer guide sequence (green) adjacent to Pam (NGG) sequence. Cas9 digests double-stranded DNA at 3–4 bp upstream of the Pam sequence. (Scheme 2) Repair of the digested genome in the presence of a piece of DNA with homologous sequence with the digested genome locus (repair template) can cause homology directed repair. A few nucleotide mismatches in the repair template (“M” in red) can be incorporated in the genome after the repair; thus, desired point mutations can be introduced during this step. (Scheme 3) To check whether the repaired genome has mutations, the repaired region of the genome is amplified by PCR. PCR primers (purple) should be outside of the repair template sequence. (Scheme 4) PCR products are digested by restriction enzyme. Incorporated mutations are designed to create a specific restriction enzyme site. Thus, PCR products with wild-type sequence are not digested by the enzyme (lower 4 PCR product in the cartoon), while PCR products from the mutated genome (upper 4 PCR products) are digested into two small DNA fragments (Scheme 5). PCR products after restriction digest are separated by gel electrophoresis. WT, wild type, HET, heterozygote; HOM, homozygote 3.1 Design and Constructions of Materials for CRISPR/ Cas9 Gene Editing

1. Determine the gene locus (the genome sequence (exon) encoding the amino acid residue) to be gene edited. Use Ensembl Genome Browser (www.ensembl.org) or similar genome browser. 2. Copy and paste genome sequence (~200 bp) around the target locus on the CRISPR design tool web sites to identify Pam sequences (NGG) near the desired gene-edited site. A number of CRSIPR design tools are freely available that can identify the Pam sequences and its specificity. 3. Select a Pam sequence and the adjacent 20 bp guide sequence on the 5-prime side of the Pam (Fig. 1, Scheme1). The desired point mutation should be on the 5-prime side of the Pam sequence. The choice of Pam sequence is preferred to be close to the mutation site (within ~20 bp) for better geneediting efficiency. Most of CRISPR design tools provide off-targeting scores, indicating specificity of each guide sequences. Off-targeting effect is one of the major concerns of CRISPR/Cas9 gene-editing approach, making it paramount to use multiple CRISPR design tools to identify specific Pam and adjacent guide sequences.

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4. Design two oligonucleotide primers that represent sense and antisense sequences of the guide sequence as determined in step 3. Additional nucleotides should be added to the primers to create sticky ends for the restriction enzyme to help the double-stranded oligonucleotides be incorporated into the plasmid vector. This restriction enzyme is a cloning site in the plasmid vector (item 1 in Subheading 2.1). The details of cloning into the pX459 plasmid are shown in Fig. 4 of Ref. [4]. 5. Design of the repair template. Repair templates should have the homologous DNA sequence to the target gene locus with a few mismatches (mutations). We have used two different repair templates; (1) single-stranded oligonucleotide (~150 bp) of the mutated gene sequence and (2) plasmid DNA carrying ~400 bp insert of the mutated gene sequence (see Note 1). The repair templates carry three mutations: (a) the desired mutation (in our study, a mutation that changes a single RyR2 amino acid residue), (b) a silent mutation (no amino acid change) creating a specific restriction enzyme site, and (c) a silent mutation eliminating the targeted Pam sequence to reduce a chance for recurring gene edit (see Note 2). 6. Construct plasmid DNA for expression of RNA-guided Cas9 exonuclease. Double-stranded oligonucleotides of the guide sequence (item 2 in Subheading 2.1) are cloned into the plasmid vector to express Cas9 exonuclease (item 1 in Subheading 2.1; see Ref. [4] for the detailed molecular biology strategies). Sequence the constructed plasmid DNA carrying the guide sequence. 3.2 Introducing the Plasmid DNA and Repair Template into hiPSCs by Electroporation

We have mostly used a modified protocol based on that described by Ran et al. [4]. Although there are number of electroporation systems, we have been successful using the Neon Transfection System. 1. Grow hiPSCs in three vitronectin-coated 60 mm dishes to nearly confluent levels, and then treat them with 5 μM Y27632 for 2 hours. 2. The cells are then dissociated using 1–2 mL Accutase reagent followed by addition of 1 mL StemFlex medium. Dissociated cells are collected by centrifugation (400  g for 2 min) and resuspended in 310 μL electroporation medium in the presence of 20 μg plasmid DNA with the guide sequence (step 6 in Subheading 3.1) and 0.4 nmol single-stranded oligonucleotide as a repair template (item 4 in Subheading 2.1). Alternatively, use 15 μg plasmid DNA with the guide sequence and 5 μg double-stranded repair template DNA (see Note 3).

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3. Transfect plasmid DNA and repair template into the cells by electroporation. We apply two pulses of 1050 V for 30 msec. The 100 μL of the transfected cells are then placed in each vitronectin coated 60 mm dish with StemFlex medium containing 5 μM Y27632. 4. Grow the cells overnight, and then change the medium to StemFlex on the following morning for several hours of incubation. 5. Add 1 μg/ml (final concentration) puromycin (for use of pX459 plasmid as Cas9 expression) for 48 hours incubation. At 24 hours after addition of puromycin, we exchange the medium with one containing fresh puromycin. 6. After 48 hours of puromycin treatment, the culture medium is replaced by StemFlex without antibiotics. Grow the cells until they form colonies, generally taking 1 week. 3.3 Screening for Correctly Gene-Edited Cells

1. After addition of 5 μM Y27632 for 2 hours to the 60 mm dishes, single-cell colony is scratched by 25-gauge needle syringe and then collected by 100 μl pipet tips (see Note 4). 2. Plate half of the collected cells from each colony in vitronectincoated 24-well plate containing StemFlex with Y27632. Collect the other half of the cells by centrifugation (2000  g for 10 min), and incubate them with DNA extraction solution to obtain genomic DNA. 3. Perform PCR using genomic DNA from each colony as a template. PCR primers are designed to amplify the targeted gene locus including whole sequence used for the repair template (Fig. 1, Scheme 3). 4. Digest PCR products with a restriction enzyme (designed in step 5 of Subheading 3.1), and determine whether PCR products are digested by gel electrophoresis (see Note 5). Intact size of the PCR product should be observed in wild-type cells. When both gene alleles are gene edited (homozygote), PCR products are completely digested, and there are two small sizes of PCR products. When only one gene allele is gene edited (heterozygote), the PCR products are only partly digested, and there will be three bands on the gel (one intact size and two smaller size) (Fig. 1, Scheme 4–5). 5. Sequence PCR products showing restriction digestion. Since PCR products are amplified from two gene alleles, the sequencing results will have different patterns (see Note 6). 6. Clone PCR products showing correct gene-edited sequence into plasmid vector using PCR cloning kit. Verify two gene allele sequence at the targeted area by sequencing PCR cloned plasmids (we usually sequence eight different plasmid DNAs

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per single-cell clone). The hiPSC clones with correct geneedited sequence are designated as “positive” clones. After subcloning (see Note 4), the cells are used for cardiac differentiation. 7. Sequence RT-PCR products of the targeted locus in the differentiated cardiomyocytes and confirm mutations in mRNA. This step would also reveal whether the mutations can induce splicing variants. 3.4 Cardiac Differentiation of the Mutant hiPSCs

We differentiate hiPSCs into cardiomyocyte by activating and inhibiting Wnt signaling pathway as described by Lian et al. [8] with some modifications. RPMI/B27 media is made by mixing RPMI 1640 medium and B27 supplement at 49 to 1 ratio. RPMI/B27( ) is made from insulin-free B27 supplement, while RPMI/B27(+) is from B27 supplement. A flowchart is shown in Fig. 2. 1. Passage hiPSCs as described in step 1 of Subheading 3.2. Cells collected by centrifugation (400  g for 2 min) are then resuspended in StemFlex and plated (~6  105 cells in 2 mL culture volume) on Matrigel (dilute, item 9 in Subheading 2.2)-coated 12-well plate. 2. Incubate cell culture in StemFlex containing 5 μM Y27632 overnight, and replace the medium with StemFlex only. 3. Once cells are spread all over the culture well, replace the culture medium with RPMI/B27( ) containing 6–12 μM CHIR99021.

Fig. 2 Flowcharts of cardiac differentiation protocols. Comparison of two methods of cardiac differentiation of hiPSCs is shown. We typically observe that cells start beating at day-8, and then we change the RPMI/B27( ) medium to RPMI/B27(+)

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4. Culture cells in the medium containing CHIR99021 for 24 hours (from day-0 to day-1) and then change medium to RPMI/B27( ) only. 5. Maintain cells with RPMI/B27( ) for 48 hours (from day-1 to day-3) and replace the medium with RPMI/B27( ) plus 5 μM IWR-1. 6. Culture cells with media containing IWR-1 for 48 hours (from day-3 to day-5), and change media with RPMI/B27( ) only. 7. Maintain cell culture with RPMI/B27( ). Change media with RPMI/B27(+) once majority of the cells start beating (typically at day-8). 8. Maintain the beating cardiomyocytes in RPMI/B27(+) medium for 1 month or longer before experiments. Change media twice a week. 3.5 Cardiomyocyte Dissociation for Single-Cell Experiments

We use single isolated cardiomyocytes for Ca2+ imagining and electrophysiological experiments (Fig. 3). We usually plate cells on 25 mm coverslips and maintain them in 35 mm dishes. The coverslip is coated with 1ml vitronectin (item 10 in Subheading 2.2) for 2 hours prior to cell plating. 1. Micro-dissect the beating area mechanically. Use a needle syringe for dissection. 2. Collect the cell clusters in a 15-mL centrifuge tube, and rinse them twice with PBS. 3. Add ~600 μl TrypLE™ reagent (10) (see Note 7), and digest the cells for 6–8 min at 37  C. 4. Add ~1 ml RPMI/B27(+) medium to stop digestion, and collect the cells by centrifugation (400  g for 2 min). 5. Wash the cells once with RPMI/B27(+) medium, and resuspend them with RPMI/B27(+) medium including 1 RevitaCell™ Supplement. When micro-dissected area is ~1/3 of one well (12-well plate), we resuspend the cells in 600–800 μl medium. 6. Plate the suspended cells (~200 μL/coverslip) on the 5 μg/ml vitronectin-coated 25 mm coverslip. Perform this step under the microscope to adjust the cell density. 7. Culture the cells in 2 mL RPMI/B27(+) medium including 1 RevitaCell™ Supplement. 8. Change the medium to RPMI/B27(+) medium only (without RevitaCell™ Supplement) on the following day.

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Fig. 3 Arrhythmogenic calcium sparks and action potentials in hiPSC-CMs harboring a RYR2 mutation. (a) Representative elementary Ca2+ release events (Ca2+ sparks) in wild-type (WT) and Q4201R mutant [6] hiPSCCMs. Ca2+ sparks in three different regions (black, red, and blue traces) of interest were measured by Ca2+ indicating dye, Fluo4, under total internal reflection fluorescence microscopy. While WT hiPSC-CMs exhibited transient and brief Ca2+ sparks, the mutant cells harboring CPVT-linked RyR2 mutations showed long-lasting Ca2+ sparks. Panels of 2D images show the development of single Ca2+ spark indicated by a and b in the Ca2+ spark traces of WT (region 1) and mutant (region 2) cells, respectively. (b) Action potentials recorded from patch-clamped hiPSC-CMs (current clamp mode). CPVT1-linked F2483I mutations [5] often cause irregular membrane depolarizations, early- and delayed-after depolarizations (red asterisks)

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Fig. 4 Cardiomyocytes differentiated from hiPSCs in the presence of fatty acids cocktail. (a) Relatively higher amounts of the elongated shapes of cardiomyocytes were observed by cardiac differentiation of wild-type hiPSCs in the presence of fatty acids. Cells were infected with adenovirus carrying FKBP12.6-GCaMP6 probe [10]. Since FKBP12.6 has a high affinity to RyR2, the striation pattern of the green fluorescence indicates the position of RyR2. (b) Double stain of the fatty acid cocktail-incubated hiPSC-CMs by FKBP-GCaMP6 probes (green) and TRITC-labeled WGA, wheat germ agglutinin (red). We observed some plasma membrane signals at the same position of FKBP-GCaMP probe (arrowheads), suggesting T-tubule formation. Blue is DAPI-stained nucleus 3.6 Cardiac Differentiation of the Mutant hiPSCs in Maturation Media

We use fatty acid cocktails [9] to drive hiPSC-CMs into more mature state. A concise flowchart of the process is shown in Fig. 2. 1. Differentiate hiPSCs into cardiomyocytes according to the same protocol as Subheading 3.4 for 10 days following the onset of cell beating. 2. Dissociate the differentiated cells using TrypLE™ reagent (10) at 10 days following cell beating (typically at day-18). 3. Collect the cells by centrifugation (400  g for 2 min), and resuspend them in RPMI/B27(+) medium that includes 1 RevitaCell™ Supplement. 4. Plate the resuspended cells in the Matrigel-coated 12-well plate. Change the medium to RPMI/B27(+) only on the next day (day-19). 5. Change the medium to the fatty acid maturation medium on the following day (day-20), and culture the cells at least for 2 weeks to obtain the cardiomyocytes with relatively matured morphology (Fig. 4). Change the medium twice a week.

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Notes 1. There are multiple designs for the repair template that in part depend on length of homologous sequence, position of the mutation, and whether the repair template is single or double stranded. We use asymmetric single-stranded oligonucleotides, which have about 25-bp homologous sequence on the 5-prime

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side of the mutation, and about 90-bp sequence on the 3-prime side from the Cas9 digested position. The single-stranded DNA has the same strand sequence as the targeted Pam, and the mutations are on the 5-prime side of the Cas9 digest site. The large double-stranded repair template (~400-bp) has mutations in the middle and 200-bp homologous DNA sequence on both sides of the mutations. 2. In some cases, the desired mutation creates a specific restriction enzyme site. In this case no additional mutation is required. A mutation in the Pam sequence is not always a “silent” mutation, i.e., mutations may cause amino acid change. In such cases we do not introduce the Pam site mutation but may choose a different Pam site. 3. The double-stranded repair template is linearized by a restriction enzyme, which digests the plasmid vector but not the targeted locus-specific sequence. The linearized DNA should be cleaned by ethanol precipitation before mixing with cells. 4. Scratch off a single isolated cell colony from the dish using a needle syringe, and collect the detached cells using a pipet. Since following colony isolation and collection, some (uncollected) cells float in the culture medium; we usually exchange the medium for every six-colony isolation to avoid contamination. To assure purity of the colony, we subclone the correctly gene-edited cell lines. 5. Since all the restriction enzymes are not active in the presence of PCR buffer, the selection of restriction enzymes and PCR buffer is important. New England Biolabs shows restriction enzyme activities for several different PCR buffers. 6. Sequencing may show PCR products that have (a) wild-type sequence; (b) double peaks (wild type and mutation) at the targeted nucleotide, indicating presence of heterozygote; (c) single peak of mutation, indicating homozygote; and (d) single or double peak of the mutation, but also continuous double peaks starting around targeted Pam sites. The last possibility most likely indicates that the genome carries not only the desired mutation but also has nucleotides insertion or deletion. If this insertion/deletion is only on one gene allele (and causes frameshift), and the other gene allele is correctly gene-edited, the cell may be useful for further experiments. The complete sequence of two gene allele must be then determined by cloning PCR products and sequencing. 7. Even though cells may be dissociated using 1 mg/ml collagenase (Type IV) or 0.05% Trypsin-EDTA, TrypLE reagent yields the best results in our hands.

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Acknowledgments This work was supported by the National Institutes of Health Grants HL147054 and HL153504. References 1. Napolitano C, Bloise R, Monteforte N, Priori SG (2012) Sudden cardiac death and genetic ion channelopathies: long QT, Brugada, short QT, catecholaminergic polymorphic ventricular tachycardia, and idiopathic ventricular fibrillation. Circulation 125(16):2027–2034. h t t p s : // d o i . o r g / 1 0 . 1 1 6 1 / CIRCULATIONAHA.111.055947 2. Kushnir A, Wajsberg B, Marks AR (2018) Ryanodine receptor dysfunction in human disorders. Biochim Biophys Acta Mol Cell Res 1865(11 Pt B):1687–1697. https://doi.org/ 10.1016/j.bbamcr.2018.07.011 3. Takahashi K, Tanabe K, Ohnuki M, Narita M, Ichisaka T, Tomoda K et al (2007) Induction of pluripotent stem cells from adult human fibroblasts by defined factors. Cell 131(5): 861–872. https://doi.org/10.1016/j.cell. 2007.11.019 4. Ran FA, Hsu PD, Wright J, Agarwala V, Scott DA, Zhang F (2013) Genome engineering using the CRISPR-Cas9 system. Nat Protoc 8(11):2281–2308. https://doi.org/10.1038/ nprot.2013.143 5. Wei H, Zhang XH, Clift C, Yamaguchi N, Morad M (2018) CRISPR/Cas9 Gene editing of RyR2 in human stem cell-derived cardiomyocytes provides a novel approach in investigating dysfunctional Ca(2+) signaling. Cell Calcium 73:104–111. https://doi.org/10. 1016/j.ceca.2018.04.009 6. Zhang XH, Wei H, Xia Y, Morad M (2021) Calcium signaling consequences of RyR2

mutations associated with CPVT1 introduced via CRISPR/Cas9 gene editing in humaninduced pluripotent stem cell-derived cardiomyocytes: comparison of RyR2-R420Q, F2483I, and Q4201R. Heart Rhythm 18(2): 250–260. https://doi.org/10.1016/j.hrthm. 2020.09.007 7. Fernandez-Morales JC, Xia Y, Renzo TJ, Zhang XH, Morad M (2022) Mutation in RyR2-FKBP Binding site alters Ca(2+) signaling modestly but increases “arrhythmogenesis” in human stem cells derived cardiomyocytes. Cell Calcium 101:102500. https://doi.org/ 10.1016/j.ceca.2021.102500 8. Lian X, Zhang J, Azarin SM, Zhu K, Hazeltine LB, Bao X et al (2013) Directed cardiomyocyte differentiation from human pluripotent stem cells by modulating Wnt/beta-catenin signaling under fully defined conditions. Nat Protoc 8(1):162–175. https://doi.org/10.1038/ nprot.2012.150 9. Yang X, Rodriguez ML, Leonard A, Sun L, Fischer KA, Wang Y et al (2019) Fatty acids enhance the maturation of cardiomyocytes derived from human pluripotent stem cells. Stem Cell Reports 13(4):657–668. https:// doi.org/10.1016/j.stemcr.2019.08.013 10. Pahlavan S, Morad M (2017) Total internal reflectance fluorescence imaging of genetically engineered ryanodine receptor-targeted Ca (2+) probes in rat ventricular myocytes. Cell Calcium 66:98–110. https://doi.org/10. 1016/j.ceca.2017.07.003

Chapter 5 Enhancing Cardiomyocyte Transcription Using In Vivo CRISPR/Cas9 Systems Eric Schoger and Laura C. Zelaraya´n Abstract Endogenous gene activation by programmable transcription factors offers gene-dose-dependent phenotyping of target cells embedded in their in vivo natural tissue environment. Modified CRISPR/Cas9 systems were developed to be used as guide (g) RNA programmable transcriptional activation platforms (CRISPRa) in vitro and in vivo allowing targeted or multiplexed gene activation studies. We specifically developed these tools to be applied in cardiomyocytes providing dCas9VPR expressing mice under the control of the Myosin heavy chain 6 (Myh6) promoter. Here, we describe a protocol for the efficient design and validation of newly identified gRNA for enhancing transcriptional activity of a selected gene of interest. Additionally, we are providing insights into a downstream application in a dCas9VPR expressing mouse model specifically for cardiomyocyte biology. Key words CRISPRa, Transgenic mouse models, Transcriptional control, Synthetic transcription factors, Gene-dose titration

1

Introduction Cardiomyocyte biology is ruled by transcriptional changes. These changes allow adaptation of the heart from development to postnatal cardiac homeostasis as well as from physiological to altered hemodynamic conditions driving maladaptive tissue remodeling [1, 2]. Our current knowledge on the contribution of specific genes to cellular functions has been traditionally investigated by employing transgenic small animal and in vitro models reflecting the effect of (over-)expression or lack of individual genes [3]. The adaptation of clustered regularly interspaced short palindromic repeats (CRISPR/Cas9) endonucleases as synthetic transcription factors made it possible to fine-tune the control of endogenous gene activity in a titratable and multiplexing-amenable manner [4] expanding the tool box for more precise functional studies. CRISPR-based endogenous gene activation (CRISPRa) employs

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enzymatically inactive Cas9 fused to transcriptional activation domains, which can be recruited by gRNAs to genomic regulatory elements of gene bodies to enhance target gene transcription [5]. Based on this system, we have generated a mouse model that enabled enhancement of cardiomyocyte-specific endogenous gene activation using an enzymatically inactive, dead (d)Cas9 fused to VP64, p65 transactivation domain, and RTA (VPR) [6]. Here, we summarize our protocol for efficient design and validation of newly designed gRNAs for targeted endogenous gene activation. We furthermore offer a protocol for CRISPRa in postnatal dCas9VPR transgenic cardiomyocytes and adeno-associated virus serotype 9 (AAV9)-mediated gRNA delivery in vivo (see Fig. 1).

2 2.1

Materials Cloning

1. LB agar plates (with appropriate selection antibiotic). 2. LB broth (with appropriate selection antibiotic). 3. Bacteria shaker and incubator (37
C). 4. Sterilized bacteria culture vessels. 5. Tabletop centrifuges. 6. Thermocycler. 7. dCas9VPR expression plasmid (i.e., addgene # 63798). 8. Empty gRNA expression plasmid (i.e., addgene #53187, addgene #60955, or modified pX330 and pX333 plasmids (removed Cas9) derived from addgene #110403 or addgene #64073). 9. Sense strand gRNA oligonucleotide. 10. Antisense strand gRNA oligonucleotide. 11. Polynucleotide kinase (PNK). 12. Alkaline phosphatase (FastAP). 13. FastDigest BpiI. 14. Gel and PCR Clean-Up Kit. 15. Ligation Kit. 16. NEBStable Competent E.coli. 17. SOC outgrow medium. 18. Plasmid Mini/Midi Preparation Kit. 19. Nanodrop. 20. Access to Sanger sequencing facility.

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Exon1 gRNAs

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caccNNNNNNNNNNNNNNNNNNNN NNNNNNNNNNNNNNNNNNNNcaaa 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20

B Validation

C Viral delivery preparation

D Downstream application

dCas9VPRMyh6 Fig. 1 Schematic overview of working steps for gRNA design and validation as well as downstream application. (a) In silico design of gRNAs and oligonucleotides for molecular cloning of CRISPRa components. Candidate gRNAs for endogenous gene activation should be ideally targeted to the 100 bp region upstream of the TSS. (b) Validation of gRNAs in easy-to-transfect cell lines (i.e. Neuro2a). (c) Preparation of viral constructs for cardiomyocyte transduction in vivo. (d) Possible downstream applications in CRISPRa mouse lines. This illustration contains Servier Medical Art (smart.servier.com) elements

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Validation

1. 24-well plates. 2. Mammalian cell culture incubator. 3. Laminar flow hood. 4. Transfection reagent (i.e., TurboFect). 5. Neuro2a (ATCC). 6. Neuro2A cell growth medium: Dulbecco’s Modified Eagle Medium, 10% v/v fetal calf serum, 100 U/mL Penicillin / 100 mg/mL Streptomycin. 7. Sterile PBS (without Mg2+ and Ca2+). 8. 0.25% Trypsin. 9. Minimal Essential Medium (MEM). 10. (Epi-)fluorescence microscope. 11. RNA cleanup columns. 12. cDNA synthesis Kit. 13. qPCR reaction mix. 14. qPCR cycler.

2.3 Downstream Application

1. Mouse handling facility. 2. 31G insulin syringe. 3. Sterile saline. 4. AAV gRNA expression plasmid (i.e., modified addgene #60958). 5. AAV (For gRNA delivery into cardiomyocytes in vivo, we recommend AAV serotype 9 (AAV9). It may be produced in the host laboratory or acquired commercially.) 6. Transgenic mouse model expressing Streptococcus pyogenes dCas9-based transcription factors. For endogenous gene activation in vivo, we established a mouse model for constitutive dCas9VPR expression in Myh6-expressing cells [6]. However, any other transgenic mouse model expressing Streptococcus pyogenes dCas9-based transcription factors may be used under consideration of the desired target cell population [7–11]. 7. Fluorescence stereomicroscope. 8. Trizol.

3

Methods

3.1 Design gRNAs for Transcriptional Activation

1. Extract the base pair sequence of the gene of interest from a genome browser database (e.g., Ensemble: https://www. ensembl.org/). 2. Identify the 50 region upstream of the transcriptional start site (TSS) (see Note 1).

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3. Select potential target sequences taking Streptococcus pyogenes Cas9 protospacer adjacent motifs into account (50 NGG 30 ). This can be facilitated by using gRNA design tools (e.g., ecrisp [12]: http://www.e-crisp.org/). We also recommend checking gRNA candidates for off-target hybridization with mismatchbased off-target search algorithms (e.g., Off-Spotter [13]: https://cm.jefferson.edu/Off-Spotter/). 4. Design oligonucleotides for cloning into suitable gRNA expression vectors. Most gRNA expression vectors generate overhangs by a class IIS restriction enzymes, which determines the to-be-inserted oligonucleotide. For matters of simplicity, we will exemplify the procedure with a pX333 (addgene #64073)-derivate [6] for which the oligonucleotide needs to have the following structure: 50 caccNNNNNNNNNNNNNNNNNNNN 30 . 30 NNNNNNNNNNNNNNNNNNNNcaaa 50 5. Cut gRNA expression construct with Type IIS restriction endonucleases (e.g. pX333-derivate [6] with BpiI): Mix 5 μg plasmid DNA with 1 μL FastDigest BpiI, 5 μL 10 FD Buffer, and nuclease-free H2O to a final volume of 50 μL. 6. Incubate reaction at 37
C for 3 h. 7. Add 1 μL FastAP to the reaction mix to dephosphorylate reaction products, and incubate at 37
C for 30 min. 8. Isolate restriction digest reaction product with a PCR CleanUp Kit, and determine DNA concentration. 9. Phosphorylate and anneal oligonucleotides with 1 μL PNK, 1 μL 10 Ligation Buffer, 1 μL 100 μM sense oligonucleotide, 1 μL 100 μM antisense oligonucleotide, and 6 μL nuclease-free H2O at 37
C for 30 min. 10. Heat reaction to 95
C for 5 min and decrease temperature in 5
C/min steps. 11. Dilute reaction 1:50 before proceeding with ligation reaction. 12. Ligate plasmid DNA and annealed oligonucleotide using: 50 ng digested plasmid DNA, 1 μL diluted annealed oligonucleotides, 2 μL 10 Ligation Buffer, 1 μL Ligase, and nucleasefree H2O to a final volume of 20 μL. 13. Incubate at room temperature for 30 min (see Note 2). 14. Transform 50 μL NEBStable Competent E.coli with 2.5 μL of ligation mix, and incubate on ice for 30 min. 15. Perform heat shock at 42
C for 30 sec. 16. Chill reaction on ice for 2 min, and add 200 μL SOC outgrow medium. 17. Incubate reaction at 37
C for 1 h shaking.

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18. Distribute reaction on LB agar plates with appropriate antibiotic, and let colonies grow at 37
C for 16 h. 19. Pick colonies and proceed with plasmid DNA cleanup (i.e., NucleoSpin Mini/Midi). 20. Confirm integration of gRNA oligonucleotide by Sanger sequencing. 3.2 Validate gRNAs for Transcriptional Activation In Vitro

1. Seed 1  105 Neuro2a cells/well in a 24-well plate format the day before transfection (see Note 3). 2. Mix 0.5 μg dCas9VPR-expressing plasmid and 0.5 μg gRNA expression plasmid together with 2 μL Turbofect in MEM, and incubate for 15 min. 3. Add transfection mix dropwise to cells. 4. Incubate cells at 37
C, 5% CO2, humidity controlled for 48 h. 5. Check transfection efficiency by fluorescence microscopy (see Note 4 and Fig. 2). 6. Isolate RNA (e.g., RNA (Macherey Nagel)), and perform qPCR to check for endogenous transcriptional activation (see Note 5). 7. Subclone selected gRNAs into appropriate viral vectors (AAV vectors) for downstream applications, and confirm construct identity by Sanger sequencing (see Note 6).

Fig. 2 Representative images of CRISPRa components transfected cells. (a) Expected transfection efficiencies in Neuro2A cells with gRNA (GFP) and dCas9VPR (tdTomato) expression plasmids. Scale bar ¼ 500 μm. (b) Endogenous gene activation of Nos2 as evaluated 48 h post-transfection by qPCR (unpublished data, Nos2 qPCR primer sequences: forward, 50 CAGCTGGGCTGTACAAACCTT 30 ; reverse, 50 CATTGGAAGTGAAGCGTTTCG 30 ; gRNA A, 50 GGAGGGGTATAAATACCTGA 30 ; B, 50 GCTAACTTGCACACCCAACT 30 ; C, 50 GACTCACTAGGGAGG GAGGC 30 ; D, 50 GCCAATATTCCAACACGCCC 30 ). (NT non-targeted gRNA: 50 GTCCAGCGGATAGAATGGCG 30 . Statistics: One-way ANOVA and Bonferroni correction, ***p  0.001)

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1. For postnatal day four (P4) intraperitoneal (i.p.) injections, genotype litter before the day of injection (P3; see Note 6), and prepare AAV9 (3.3  1011 vg/mouse dissolved in saline to a total volume of 80 μL) in a 31G insulin syringe. 2. Take up pups by the neck, and apply a gentle stretch in the abdominal area by adjusting your grip. 3. Aim for the lower abdominal quadrant and inject the entire volume. 4. Loosen your grip to reduce intraabdominal pressure preventing leakage of applied volume. We observed endogenous gene activation in whole heart tissue as early as four days postinjection (see Note 7). 5. For intravenous injections (i.v.) in adult mice, inject AAV9 (1  1012 vg/mouse dissolved in saline to a total volume of 80 μL) via the tail vein. We observed endogenous gene activation in whole heart tissue as early as 2 weeks post-injection. 6. After the desired time, harvest hearts and check reporter fluorescence by stereomicroscopy (see Fig. 3). 7. Isolate RNA (see Note 8), and perform qPCR to confirm endogenous gene activation of the target gene.

Fig. 3 Expected AAV9 transduction efficiencies for endogenous gene activation in mouse cardiomyocytes. (a) AAV9 delivered gRNAs (GFP) in Myh6 promoter-driven dCas9VPR (tdTomato) hearts next to non-transgenic, non-transduced littermate hearts. AAV9 (3.3  1011 vg/mouse) were injected at postnatal day four, and hearts were harvested 2 and 8 weeks post-injection. Scale bar ¼ 2 mm. (b) Isolated cardiomyocytes from 4 weeks post-injected mouse hearts representing high AAV-mediated gRNA transduction (GFP). Scale bar ¼ 200 μm

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Notes 1. CRISPRa gRNAs generally work best in a target window between 200 bp and 1 bp relative to the TSS with a fold gene activation maximum at 100 bp in our hands and in agreement with previous reports [14]. We test four to eight gRNAs per target gene to select the most effective candidate (s) for downstream applications. 2. We recommend considering a ligation control reaction (no gRNA oligonucleotide added to the ligation mix) to check for possible undigested/re-ligated plasmid DNA and to infer overall ligation efficiency. 3. Testing of newly designed gRNAs for endogenous gene activation should be performed in easy-to-transfect immortalized cell lines. We recommend Neuro2a for targeting the mouse genome. 4. Transfection efficiencies may vary depending on initial cell density and transfection reagent/method used. Our dCas9VPR expressing constructs as well as gRNA expression constructs co-express fluorescent proteins (tdTomato or GFP, respectively [6]), and we recommend checking for transfection efficiencies by appropriate means (e.g., fluorescence microscopy/flow cytometry) to reduce batch-to-batch differences. 5. qPCR/primer validation for the reliable detection and (semi-) quantification of target gene expression mRNA levels is crucial for accurate mRNA fold change compared to control groups. 6. It is recommendable to perform analytical restriction enzyme reactions to check for AAV’s inverted terminal repeat region before AAV production. 7. For neonatal systemic AAV injections, we recommend retrieving tail tip biopsies and tattooing of pups according to international and local animal experimental guidelines to genotype and later identify animals for injections. 8. While RNA isolation of Neuro2a cells may be performed with column-based isolation kits, we recommend RNA isolation from heart tissue with chloroform extraction/isopropanol precipitation (i.e., Trizol) to maximize overall RNA yield.

Acknowledgments This work was supported by the Deutsches Zentrum fu¨r HerzKreislauf-Forschung e.V. (DZHK) to ES and LCZ and the Deutsche Forschungsgemeinschaft e.V. (DFG), SFB1002 project C07 to LCZ.

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References 1. Pawlak M, Kedzierska KZ, Migdal M et al (2019) Dynamics of cardiomyocyte transcriptome and chromatin landscape demarcates key events of heart development. Genome Res 29: 506–519. https://doi.org/10.1101/gr. 244491.118 2. Nomura S, Satoh M, Fujita T et al (2018) Cardiomyocyte gene programs encoding morphological and functional signatures in cardiac hypertrophy and failure. Nat Commun 9:1–17. https://doi.org/10.1038/s41467-01806639-7 3. Dirkx E, da Costa Martins PA, de Windt LJ (2013) Regulation of fetal gene expression in heart failure. Biochim Biophys Acta Mol basis Dis 1832:2414–2424. https://doi.org/10. 1016/j.bbadis.2013.07.023 4. Chavez A, Tuttle M, Pruitt BW et al (2016) Comparison of Cas9 activators in multiple species. Nat Methods 13:563–567. https://doi. org/10.1038/nmeth.3871 5. Perez-Pinera P, Kocak DD, Vockley CM et al (2013) RNA-guided gene activation by CRISPR-Cas9-based transcription factors. Nat Methods 10:973–976. https://doi.org/ 10.1038/nmeth.2600 6. Schoger E, Carroll KJ, Iyer LM et al (2020) CRISPR-mediated activation of endogenous gene expression in the postnatal heart. Circ Res 126:6–24. https://doi.org/10.1161/ CIRCRESAHA.118.314522 7. Liao H-K, Hatanaka F, Araoka T et al (2017) In vivo target gene activation via CRISPR/ Cas9-mediated trans-epigenetic modulation. Cell 171:1495–1507.e15. https://doi.org/ 10.1016/j.cell.2017.10.025

8. Wangensteen KJ, Wang YJ, Dou Z et al (2018) Combinatorial genetics in liver repopulation and carcinogenesis with a in vivo CRISPR activation platform. Hepatology 68:663–676. https://doi.org/10.1002/hep.29626 9. Zhou H, Liu J, Zhou C et al (2018) In vivo simultaneous transcriptional activation of multiple genes in the brain using CRISPR–dCas9activator transgenic mice. Nat Neurosci 21: 440–446. https://doi.org/10.1038/s41593017-0060-6 10. Matharu N, Rattanasopha S, Tamura S et al (2019) CRISPR-mediated activation of a promoter or enhancer rescues obesity caused by haploinsufficiency. Science 363:eaau0629. https://doi.org/10.1126/science.aau0629 11. Colasante G, Qiu Y, Massimino L et al (2020) In vivo CRISPRa decreases seizures and rescues cognitive deficits in a rodent model of epilepsy. Brain 143:891–905. https://doi.org/10. 1093/brain/awaa045 12. Heigwer F, Kerr G, Boutros M (2014) E-CRISP: fast CRISPR target site identification. Nat Methods 11:122–123. https://doi. org/10.1038/nmeth.2812 13. Pliatsika V, Rigoutsos I (2015) “Off-spotter”: very fast and exhaustive enumeration of genomic lookalikes for designing CRISPR/Cas guide RNAs. Biol Direct 10:1–10. https:// doi.org/10.1186/s13062-015-0035-z 14. Gilbert LA, Horlbeck MA, Adamson B et al (2014) Genome-scale CRISPR-mediated control of gene repression and activation. Cell 159: 647–661. https://doi.org/10.1016/j.cell. 2014.09.029

Chapter 6 AAV-Mediated Somatic Gene Editing for Cardiac and Skeletal Muscle in a Large Animal Model Tilman Ziegler, Tarik Bozoglu, and Christian Kupatt Abstract Here we describe a protocol to produce a recombinant adeno-associated viral vector (rAAV)-based system to deliver the CRISPR-Cas9 complex into porcine skeletal muscle and myocardial cells. We initially describe the genomic composition of the rAAV-CRISPR vectors used in our lab. Furthermore, we give a step-bystep instruction into the production of recombinant viral vectors with high yields and purity. Lastly we describe the minimally invasive injection regimes to target the myocardium in a pig. Key words rAAV, Gene editing, CRISPR, Cas9, Duchenne muscular dystrophy, Gene vector

1

Introduction Since the advent of CRISPR-Cas9 technology, monogenic diseases such as Duchenne muscular dystrophy, cystic fibrosis, Huntington’s disease and others have been identified as disorders potentially treatable via in vivo gene editing. The CRISPR-Cas9 system has already found widespread use in basic and translation research with its use ranging from the generation of knockout cell lines to the treatment of diseases in mouse models. Considering the advance of the CRISPR-Cas9 system into a clinical setting however, additional experiments in large animal models, which more closely resemble the clinical situation, are needed. Furthermore, to introduce the CRIPSR-Cas9 system to facilitate gene editing, a suitable vector system is required. Here recombinant adeno-associated viral vectors (rAAVs) are of great utility, since they display low immunogenicity, show low rates of genomic integration, and allow for the prolonged expression of transgenes in the targeted cell, particularly in post-mitotic cells such as muscle fibers [1]. For the transduction of skeletal muscle and cardiomyocytes, a recombinant adenoassociated viral vector containing the genomic backbone of wildtype AAV2 together with the capsid from AAV9 has demonstrated

Kiyotake Ishikawa (ed.), Cardiac Gene Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2573, https://doi.org/10.1007/978-1-0716-2707-5_6, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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high efficacy in targeting muscle cells [2, 3]. One drawback of recombinant adeno-associated viral vectors lies in their limited packaging capacity of 4.5–5 kilobases, which is substantially exceeded by the ORF of Cas9 and gRNAs together with promoter regions and regulatory elements required. To overcome the packaging limitations of rAAV-based vectors, a two-vector system each encoding either the N-terminal or C-terminal portion of the Cas9 protein can be used. To facilitate the recombination of both Cas9 parts upon target cell transduction and protein expression, intein sequences are tagged to the C-terminal and N-terminal Cas9 parts, creating a split Cas9 version that spontaneously recombines intracellularly [4]. The extein/intein system used here is derived from the cyanobacterium Nostoc punctiforme and facilitates the recombination of both Cas9 segments together with the splicing and fusion of the C-intein and N-intein tags, leading to a fully functional Cas9 protein [5]. Recently this approach has successfully been utilized in our group to edit a diseased Dystrophin allele in a porcine model of Duchenne muscular dystrophy [6]. In larger animals more accurately representing a clinical setting, such as pigs, complicating factors come to bear. Crucial considerations regarding the use of rAAV systems in pigs range from the necessity to inject high numbers of viral particles requiring a largescale rAAV production, the potentially different toxicity of rAAV vectors, as well as alternative administration routes. Here we describe the considerations required to design rAAVbased vectors encoding the CRISPR-Cas9 system in an intein-split version, the method of generating said rAAVs, as well as potential administration routes to facilitate cardiac and skeletal muscle editing in the pig.

2

Materials

2.1 Vector Production

1. 293 T cells. 2. 72
15 cm cell culture dish. 3. DMEM. 4. PBS. 5. Pen/Strep. 6. Fetal calf serum (FCS). 7. PEI solution: 500 mL ddH2O, 500 mg PEI 500 (Polyethylenimine Max 40,000 Polysciences), adjust pH with 10 M NaOH (6.0–7.2). 8. Transfection medium: DMEM, 1% Pen/Strep. 9. Incubation medium: DMEM +16% FCS + 1% Pen/Strep.

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10. Transfection mix: 12 mL transfection medium, cis-plasmid, trans-plasmid, delta F6 in a ratio of 1:1:2 for a total of 4 μg. 11. Cell scrapers. 12. 100% Ethanol (denaturated). 13. Dry ice. 14. Benzonase. 15. Iodixanol (OptiPrep Density Gradient Medium, Sigma Aldrich). 16. Phenol Red: 5 mg/mL in PBS. 17. 10
Gradient buffer (GB): 50 mL 1 M Tris–HCl, 150 mL 5 M NaCl, 50 mL 1 M MgCl2, 250 mL ddH2O, filter sterilize using 0.22 μm vacuum filter, store at 4  C. 18. 5 M NaCl. 19. OptiSeal tubes 32,4 mL (Beckman Coulter). 20. Cannula 14G Sterican. 21. Cannula 18G BD Microlance. 22. Lint-free wipes. 23. Sephadex G-100 Superfine (Sigma Aldrich). 24. Econo-Pac Disposable Chromatography Columns 10 mL (BioRad). 25. Biorad Econo-Pac column. 26. Amicon Ultra-15 Centrifugal Filter Unit with Ultracel-10membrane (Millipore). 27. Millex-HV Filter 0.45 μm (Millipore). 2.2 Vector Delivery in Pigs

1. Azaperone. 2. Atropine. 3. Ketamine. 4. Midazolam. 5. Fentanyl. 6. Propofol. 7. Heparin. 8. Hi-Lo Oral/nasal tracheal tube. 9. Animal ventilator. 10. Pulse oximeter. 11. ECG. 12. Catheter sheath. 13. Retroinfusion catheter (ProMED – PTC Myoprotect SSR).

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Fig. 1 Schematic depiction of the two AAV plasmids encoding the components of the CRISPR-Cas9 machinery necessary for efficient target cell transduction and editing

3 3.1

Methods Vector Design

3.2 Transfection of HEK 293 T Cells for rAAV Production

Due to the size restrictions of adeno-associated viral vectors, we propose the use of a two-vector system to introduce the CRISPRCas9 components into the target cell. Firstly, during the production process, the transgene plasmids will be constructed with the ITR sequences flanking the packaged genome. As a pol III promoter to drive the guide RNA expression, the U6 promoter has been shown to be suitable. A truncated CBA hybrid promoter is used to drive the expression of both the N-Cas9 and the C-Cas9, both of which are tagged with the N-intein and C-intein. To later detect the protein, an HA and a FLAG-tag are added with a total of three nuclear localization signal sequences. The Cas9 ORF is completed by a bovine growth hormone polyA tail sequence (Fig. 1, see also Notes 1 and 2). 1. Culture 293 T cells to a confluence of 70%. In total 72 15 cm dishes are required. 2. Change media to transfection media (18 mL/dish). 3. Put dishes for about 1 hour back in the incubator. 4. Divide the transfection mix (Fig. 2a) into three 50 mL tubes (3
4 mL). 5. Add 44 mL transfection media to each tube. 6. Mix by inverting the tube three times. 7. Add 2.5 mL PEI solution dropwise to each tube. 8. Mix by inverting immediately three times.

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Fig. 2 (a) Transfection of HEK 293 T cells with the cis-plasmid, trans-plasmid, and delta F6 plasmid leads to the production of rAAV vectors. (b) For the iodixanol gradient centrifugation, a gradient of 15%, 25%, 40%, and 58% iodixanol is layered with the final addition of the virus-containing supernatant. After centrifugation for 130 minutes at 350,000 g, the purified virus can be found at the bottom of the 40% iodixanol layer and carefully removed

9. Incubate at RT for 15 min. 10. After incubation add 2 mL of the transfection mix dropwise to each dish. 11. Move the plates up and down and left and right a few times to make sure the transfection mix is evenly distributed over the plate. 12. Put the cells back in the incubator for 4 hours. 13. After 4 hours add 5 mL of incubation medium to each dish. 14. Incubate for 72 hours. 15. Suck off the medium until there are 2–3 mL left in the dish.

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16. Scrape the cells from the dish with a cell scraper, and collect them in a 50 mL tube. 17. Spin the cells for 20 min at 1100 g at 4  C. 18. Suck off the supernatant, and store the pellets at 80  C. 3.3 Iodixanol Purification

1. Put approximately 120 mL of 100% Ethanol in a 250 mL beaker, and carefully add dry ice until it’s saturated. 2. Thaw cells for 10 min at 37  C. 3. Resuspend the pellets in 30 mL (total) 1
GB, and redivide it into three tubes (10 mL each). 4. Put the tubes in ethanol/dry ice for 5 min. 5. Put the tubes at 37  C for approximately 13 min (make sure the suspension is completely thawed). 6. Put the tubes in ethanol/dry ice for 5 min. 7. Put the tubes at 37  C for 13 min. 8. Add 2 μL Benzonase to each tube, and mix by shaking the tubes gently. 9. Put the tubes at 37  C for 45–60 min; mix every 15 min. 10. Spin the tubes down for 15 min at 3220 g and 4  C. 11. Collect the supernatant in fresh tubes. 12. Freeze at 20  C until use. 13. Build an iodixanol gradient in a Beckman OptiSeal tube. Use three tubes per virus (for each 50 mL tube from step one, you need one OptiSeal tube). Start by adding a 15% solution to the bottom of the Beckman tube using a 10 mL syringe with a long 14 gauge metal cannula. Add subsequent layers below the previous layer, 25% ! 40% ! 58%, by extending the syringe needle to the bottom of the Beckman tube and slowly injecting. Care should be taken to ensure that no air is inside the syringe, which could disrupt the layering by generating bubbles. 14. Add the virus-containing supernatant from the processing step evenly to the three gradients by slowly dripping the solution onto the top layer of the gradient. Use a 14 gauge needle and drip very slowly, avoiding disruption of the gradient layers. 15. Fill the remaining empty volume of the Beckman tube to the top using 1x Gradient Buffer. Make sure there are no air bubbles left in the tube. 16. Remove 450 μL, and wipe the top part of the tube dry with a lint-free wipe. Close the tubes with the black lids. Weigh the tubes with the spacers for tara (accuracy 0.001). 17. Put the tubes into the rotor + spacers. The acceleration should be set to 3 and deceleration to 9.

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18. Centrifuge the tube at 350,000 g for 2 h 30 min, at 18  C. If you are using a different rotor, keep the RCF at 350,000 g, and vary the running time to keep the K-factor constant. 19. Carefully move the rotor under the hood. Remove the spacers and black lids, and mount the tube on a ring stand with a utility clamp. 20. Prepare a 5 mL syringe with an 18 gauge needle for extraction of the virus-containing layer. Accurately insert the needle approximately 1–2 mm below the interface between the 40% and 58% gradient buffer layers with the bevel of the needle facing up. The virus is visible as a subtle presence of color at the interface. Slowly extract 3 mL of solution, first with the beveled needle opening facing upwards during the first half of extraction and then facing downward for the rest (Fig. 2b). 21. Store the extracted virus at 4  C until needed for the next step. 3.4 Gravity Flow Size Exclusion Purification

1. Take one 25 mL Bio-Rad Econo-Pac column with upper bed support, lid, and stopper per AAV to be purified. Spray all down with 70% ethanol, and air-dry in the tissue culture hood under UV light to sterilize. 2. Rinse out the column with sterile filtered 1
GB to remove any excess ethanol. 3. Carefully load 25 mL of the hydrated Sepharose G100SF resin into each column, avoiding the formation of air bubbles in the column. 4. Add enough 1
GB to the top of the resin to fill up the column. Be careful not to disturb the top of the resin. 5. Allow all of the buffer to flow out of the column – It will stop dripping when it is done. 6. Attach the yellow stopper to the column tip. 7. Push the upper column bed support into the column and down onto the top of the resin. 8. Remove the yellow stopper. 9. Wash the resin with additional 1
GB – enough to fill up the column (about 10 mL). 10. Allow the wash buffer to flow through the resin until it stops dripping. 11. Attach the yellow stopper to the column tip, and add 1 mL of buffer to the upper reservoir. The column can now be capped, sealed with Parafilm and stored at 4 degrees until needed. 12. Set up 14 sterile 2.0 mL tubes per sample in a rack. Label with the vector name, and label from 1 to 14 from left to right. These will be used to collect fractions containing your virus and the flow-through from the column.

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13. Put an unlabeled tube under the column, and remove the stopper so that the excess buffer can drain. 14. Put the first labeled tube under the column. 15. Add the 3 mL of your OptiPrep purified AAV vector to the top of the column. 16. Collect 1 mL fractions from the column until the drips stop. 17. Add 11 mL of 1
GB to the top of the column, and continue collecting 1 mL fractions. 18. The early fractions should contain your AAV vector, and later fractions will contain the OptiPrep. 19. Take 20 μL of each fraction collected (normally it’s sufficient to measure from fraction 8–13), and measure the absorbance at 260 nm and 280 nm using a spectrophotometer. Use 1
GB as the BLANK. 20. Take all of the fractions from the start of collection up to the point where the absorbance begins to reach so high that the spec cannot measure it – these fractions contain your AAV vector without OptiPrep. Fractions above this point are where the OptiPrep is being eluted – it has a very high absorbance at 260 nm. 21. Pool all of the virus-containing fractions, and store at 4  C until needed for the next step. 3.5 Virus Concentration

1. Divide the collected fractions from step two onto two Amicon filter units, and if necessary fill the tubes up to 15 mL with PBS. 2. Spin down the tubes at 3220 g and 15  C until about 3 mL are left. 3. Fill the tubes up to 15 mL with PBS, and pipet it up and down to mix. 4. Repeat Subheading 3.5, step 2. 5. Combine the two tubes into one, and rinse the filter with 200 μL PBS; fill the tube up to 15 mL with PBS, and pipet it up and down to mix. 6. Repeat Subheading 3.5, step 2. 7. Fill the tube up to 15 mL with PBS and pipet it up and down to mix. 8. Repeat Subheading 3.5, step 2. 9. Pipet the virus into a 50 mL tube, and rinse the Amicon filter with 200 μL PBS. 10. Filter the virus through a 0.45 μm filter. 11. Take an aliquot for qPCR.

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12. Store the virus at 4  C for short term and at 80  C for long term. 13. Titrate the vector by linearizing via NcoI and PacI digestion (determine concentration with PicoGreen Assay). 14. Dilute to 1:8. 15. Highest standard is 1 ng/μL ! 1 ng ¼ 1,6912 gc. 16. Run PCR with primers directed at the ITR-sites. 3.6

Anesthesia

1. The anesthesia of the pig is initiated by an intramuscular injection of 280 mg azaperone, 500 mg of Ketamine, and 1 mg of atropine. 2. After sedation of the pig, an i.v. line into the ear vein of the animal is established. 3. For deeper anesthesia, 5 mg midazolam and 0.5 mg of fentanyl are injected into the ear vein. 4. After endotracheal intubation the pig is connected to a ventilator, and the anesthesia is continued with propofol (10 mg/kg/ h). 5. Oxygenation is monitored using a pulse 7oximeter (see also Note 3).

3.7 Antegrade Coronary Injection

1. For antegrade injection of viral vectors, the right carotid artery is surgically exposed. 2. A 7F catheter sheath is introduced into the right carotid artery and into the right external jugular vein via the Seldinger technique. 3. After the introduction of the sheath, 10,000 IE of heparin are administered. 4. A 7F guiding catheter is placed into the left main artery via the arterial sheath under fluoroscopic control using an X-ray C-arm. 5. An additional guiding catheter is placed into the coronary sinus via the venous sheath. 6. The artery of interest (either left anterior descending artery [LAD] or left circumflex artery [LCx]) is wired using a coronary wire as is the great cardiac vein (for injection into the LCx) or the anterior interventricular vein (for injection into the LAD). 7. Then an over-the-wire balloon is advanced into the coronary artery with an additional balloon placed into the coronary vein (see Note 4).

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8. After inflation of both balloons and occlusion of the artery and vein, the vector solution (in case of a two-vector system diluted to 1 mL in 0.9% saline, for vector amount see Notes 5–7) is injected via the over-the-wire balloon into the region of interest. The whole volume in injected during the vessel occlusion, which is upheld for a period of 2
5 minutes. 9. After deflation of the balloons, they are withdrawn from the vessels followed by a withdrawal of the coronary wires and the guiding catheters. 10. The sheaths are pulled followed by subcutaneous and cutaneous sutures. 11. To avoid infection a single dose of cefuroxime should be administered after removal of the catheter components and skin suture (250 mg/animal). 3.8 Selective Pressure Regulation of Retroinfusion

1. For selective pressure-regulated retroinfusion, follow steps 1–5 from Subheading 3.7. 2. After placement of the guiding catheters, wiring of the desired artery/vein pair, and introduction of an angioplasty balloon into the artery of interest, a modified pulmonary artery catheter (PTC Myoprotect SSR) is introduced into the vein of interest (again into the great cardiac vein for the LCx supplied area or the anterior interventricular vein for the LAD supplied area). 3. Upon inflation of both catheters and occlusion of the vessels, the solution carrying the viral particles is infused via the vein over a period of 2
5 minutes. 4. In between, both balloons are deflated to avoid excessive myocardial damage. 5. After infusion of the viral particles, the instrumentation is removed, and the wound is sutured as described in Subheading 3.7.

4

Notes 1. Prior to the use of a CRISPR-Cas9 system in large animal models, the proper guide RNAs need to be chosen and tested in a suitable in vitro setting with a porcine cell line. 2. While here we describe a two-vector system, which is necessary due to the packaging limitations of the rAAV vector, Cas9 proteins of smaller size might be suitable for a single-vector system. 3. Care should be taken during anesthesia of the pigs and manipulations of the heart during catheter-based delivery of the vectors due to a tendency of pigs to develop arrhythmias such

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as ventricular tachycardia and ventricular fibrillation. The operator should closely monitor heart rate and be aware of any malignant arrhythmias arising. 4. In general, catheter-based injections into the coronary vein and artery should be performed by experienced interventional cardiologist, since improper use of the guiding catheters and coronary wires can result in the triggering of arrhythmias (as mentioned above) as well as dissections of the coronary arteries. 5. The amount of viral vectors needed varies depending on the gene of interest, its availability dependent on the chromatin status, and the needed editing efficiency. As starting concentrations, we would suggest 5
1012 viral genomes per animal for each viral vector. 6. While here we describe a local application of viral particles, in some cases a systemic application might be desired. However, the amount of viral particles needed is significantly higher with an increased likelihood of unwanted transduction (in case of the rAAV 2/9 in the liver and kidney in particular) and thus higher risk of rAAV toxicity. For intravenous injections, the amount of viral genomes varies between 2
1013 to 2
1014 per kg bodyweight for each viral vector. 7. If an injection into the muscle is desired, 2.5
1013 viral genomes per kg bodyweight in a total volume of 200 μL should be performed (in a two-vector system 1.25
1013 per vector). If a functional effect is to be determined, we suggest multiple injections per animal. To help estimate the amount of injections needed, we perform nine injections into the porcine thigh and six injections into the upper arm of the pig. References 1. Nowrouzi A, Penaud-Budloo M, Kaeppel C, Appelt U, Le Guiner C, Moullier P, von Kalle C, Snyder RO, Schmidt M (2012) Integration frequency and intermolecular recombination of rAAV vectors in non-human primate skeletal muscle and liver. Mol Ther 20(6): 1177–1186. https://doi.org/10.1038/mt. 2012.47 2. Horstkotte J, Perisic T, Schneider M, Lange P, Schroeder M, Kiermayer C, Hinkel R, Ziegler T, Mandal PK, David R, Schulz S, Schmitt S, Widder J, Sinowatz F, Becker BF, Bauersachs J, Naebauer M, Franz WM, Jeremias I, Brielmeier M, Zischka H, Conrad M, Kupatt C (2011) Mitochondrial thioredoxin reductase is essential for early postischemic myocardial protection. Circulation 124(25):2892–2902.

h t t p s : // d o i . o r g / 1 0 . 1 1 6 1 / CIRCULATIONAHA.111.059253 3. Zincarelli C, Soltys S, Rengo G, Rabinowitz JE (2008) Analysis of AAV serotypes 1-9 mediated gene expression and tropism in mice after systemic injection. Mol Ther 16(6):1073–1080. https://doi.org/10.1038/mt.2008.76 4. Truong DJ, Kuhner K, Kuhn R, Werfel S, Engelhardt S, Wurst W, Ortiz O (2015) Development of an intein-mediated split-Cas9 system for gene therapy. Nucleic Acids Res 43(13): 6450–6458. https://doi.org/10.1093/nar/ gkv601 5. Cheriyan M, Pedamallu CS, Tori K, Perler F (2013) Faster protein splicing with the Nostoc punctiforme DnaE intein using non-native extein residues. J Biol Chem 288(9):

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6202–6211. https://doi.org/10.1074/jbc. M112.433094 6. Moretti A, Fonteyne L, Giesert F, Hoppmann P, Meier AB, Bozoglu T, Baehr A, Schneider CM, Sinnecker D, Klett K, Frohlich T, Rahman FA, Haufe T, Sun S, Jurisch V, Kessler B, Hinkel R, Dirschinger R, Martens E, Jilek C, Graf A, Krebs S, Santamaria G, Kurome M, Zakhartchenko V, Campbell B, Voelse K,

Wolf A, Ziegler T, Reichert S, Lee S, Flenkenthaler F, Dorn T, Jeremias I, Blum H, Dendorfer A, Schnieke A, Krause S, Walter MC, Klymiuk N, Laugwitz KL, Wolf E, Wurst W, Kupatt C (2020) Somatic gene editing ameliorates skeletal and cardiac muscle failure in pig and human models of Duchenne muscular dystrophy. Nat Med 26(2):207–214. https://doi. org/10.1038/s41591-019-0738-2

Part III Cardiac Gene Therapy Vectors and Promotors

Chapter 7 Optimization of Synthesis of Modified mRNA Jimeen Yoo and Lior Zangi Abstract Modified mRNA (modRNA) is a safe and effective vector for gene-based therapies. Notably, the safety of modRNA has been validated through COVID-19 vaccines which incorporate modRNA technology to translate spike proteins. Alternative gene delivery methods using plasmids, lentiviruses, adenoviruses, and adeno-associated viruses have suffered from key challenges such as genome integration, delayed and uncontrolled expression, and immunogenic responses. However, modRNA poses no risk of genome integration, has transient and rapid expression, and lacks an immunogenic response. Our lab utilizes modRNA-based therapies to promote cardiac regeneration following myocardial infarction and heart failure. We have also developed and refined an optimized and economical method for synthesis of modRNA. Here, we provide an updated methodology with improved translational efficiency for in vitro and in vivo application. Key words Modified mRNA, Synthetic mRNA, In vitro transcription, mRNA translation, Gene therapy, Gene-based therapy, Genetically based therapy, Cardiac gene therapy

1

Introduction Cardiovascular diseases (CVD) continue to maintain a high morbidity and mortality rate in the United States and globally. In the United States alone, the prevalence of CVD in adults over 20 years old is 49.2%, only increasing with age [1]. Currently, the majority of medical intervention for CVD is pharmacological interventions, addressing the symptoms but not the root of the development of disease. In severe cases, invasive, high-risk surgeries are the only option. Thus, CVD patients continue to suffer from chronic symptoms. In order to address CVD, our group has been focusing on developing and testing modified mRNA (modRNA) therapeutics for cardiac diseases. We have shown that by using modRNA, cardiac regeneration and reprogramming can be attained through the upregulation of therapeutic genes [2–4]. modRNA offers several advantages over other gene delivery methods such as plasmids,

Kiyotake Ishikawa (ed.), Cardiac Gene Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2573, https://doi.org/10.1007/978-1-0716-2707-5_7, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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lentiviruses, adenoviruses, and adeno-associated viruses. It has transient and rapid expression [5, 6], does not need to cross the nuclear membrane, does not have a risk of genome integration [7–10], does not have gene size constraints, and can be administered repeatedly due to the lack of immunogenicity [11, 12]. These characteristics have allowed it to be used in preclinical investigations for vaccinations [13, 14], cancer [15–17], and cardiac diseases [18, 19], among others. It has also been used in a variety of clinical trials [20–23]. Furthermore, global validation of modRNA-based therapeutics through effective COVID-19 vaccines demonstrates the potency and safety of modRNA [20, 22]. Previously, our group wrote several protocols for modRNA synthesis and delivery to cardiac tissue [24–29], adjusting our protocol to incorporate updates in literature to increase stability, translational efficiency, and lower immunogenicity. One essential adjustment is the alteration of the polyA tail length. The polyA tail length is critical for modRNA’s stability and translation capacity because it governs degradation speed [30]. We have shown that the elongation of the polyA tail length from 120 to 173 base pairs increased translational efficiency [29]. In addition to changing the polyA tail length, optimization of the nucleoside through synthetic modifications has been documented in making a more robust modRNA variation. Two major modifications that yielded non-immunogenetic mRNA with increased translational capacity and stability were the replacement of cytidine (C) with 5-methylcytidine (m5C) and uridine (U) with pseudouridine (Ψ) [11]. Our initial protocol incorporated these results [26]. New findings showed that the modification of U using 1-methyl-pseudouridine (1-M-Ψ) showed superior translation capacity [31]. We incorporated these new findings into our updated protocols [24, 28]. We compared luciferase (luc) containing modRNA with a Ψ or 1-M-Ψ modification of U to determine which modification had enhanced translation efficiency in vitro and in vivo (Fig. 1). In HeLa and 3 T3 cells lines transfected with Ψ or 1-M-Ψ containing modRNA (Fig. 1a, b), we observed a significantly higher luc signal when using 1-M-Ψ (Fig. 1 c, d; P < 0.0001, P < 0.01, respectively). We also observed a significantly higher luciferase signal when using 1-M-Ψ containing modRNA in vivo (Fig. 1 e, f; P < 0.05). Thus, we incorporated the 173 polyA tail with the 1-M-Ψ modification in this updated protocol to generate modRNA for therapeutic use.

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Fig. 1 Luc modRNA containing 1-M-Ψ had higher translation over Luc modRNA containing Ψ in vitro and in vivo. (a) Luc modRNA containing Ψ or 1-M-Ψ were transfected into the HeLa cell line. 24 hours postdelivery, Luc signal was evaluated using IVIS. (b) Quantification of a. (c) Luc modRNA containing Ψ or 1-M-Ψ were transfected into the 3 T3 cell line. 24 hours post-delivery, Luc signal was evaluated using IVIS. (d) Quantification of d. (e) Luc modRNA containing Ψ or 1-M-Ψ were delivered into the myocardium of CFW mice. Luc signal was evaluated 24, 48, and 72 hours post-delivery using IVIS. (f) Quantification of e. Unpaired two-tailed t-test was used for b&d. One-way ANOVA and Tukey’s multiple comparison test were used for f. ****, P < 0.0001, P < 0.01, *P < 0.05

2

Materials All solutions should be made in nuclease-free water unless specified. All materials used should be RNase
/nuclease-free.

2.1

Equipment

1. Thermocycler. 2. Microcentrifuge. 3. Vortex mixer. 4. ThermoMixer. 5. NanoDrop. 6. Gel electrophoresis system. 7. Agilent 4200 TapeStation system. 8. Centrifuge.

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2.2 Solutions and Supplies

1. Nuclease-free water. 2. TAE buffer. 3. 15 mL nuclease-free tubes. 4. Eppendorf tubes. 5. Nuclease-free strip PCR tubes. 6. Ethanol (100%). 7. 2 mL elution tubes.

2.3 Synthesis of modRNA 2.3.1 Tailed DNA Template Preparation

2.3.2 PCR Product Purification Using QIAquick® PCR Purification Kit 2.3.3 In Vitro Transcription (IVT) Reaction

1. DNA template with a final concentration of 50 ng/μL (see Note 1). 2. KAPA HiFi HotStart ReadyMix PCR Kit. 3. Primer solution: 1 μM of forward and reverse primer respectively (see Note 2). Forward Primer: 50 -TTG GAC CCT CGT ACA GAA GCT AAT ACG-30 Reverse Primer: 50 -TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTT TTC CTA CTC AGG CTT TAT TCA AAG ACC A-30 1. QIAquick® PCR Purification Kit. 2. Agarose gel (1%). 3. 1 KB Ladder. Please see reference on Table 2. 1. 5 MEGAscript® T7 Kit (Invitrogen). 2. GTP 75 mM solution (provided in MEGAscript® kit). 3. ATP 75 mM solution (provided in MEGAscript® kit). 4. CTP 75 mM solution (provided in MEGAscript® kit). 5. N1-methyl-pseudouridine (N1mΨ) 100 mM solution (TriLink BioTechnologies). 6. Anti-reverse cap analog, 30 -O-Me-m7G(50 ) ppp(50 )G 10 μmol (TriLink BioTechnologies). 7. T7 TURBO DNase enzyme (provided in MEGAscript® kit).

2.3.4 RNA Phosphatase Treatment

1. Antarctic phosphatase (New England BioLabs, M0289L). 2. Antarctic phosphatase buffer 10 (New England BioLabs, B0289S).

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Table 1 PCR cycle program for creating tailed DNA template Program Initialization

Temperature

Time

Cycles



5 minutes

1



95 C

Denaturation Annealing Extension

98 C 65  C 72  C

20 seconds 15 seconds 1 minute/kilobase

25

Final extension

72  C

5 minutes

1

2.3.5 modRNA Purification Using MEGAclear

1. 5 M Ammonium acetate salt solution (provided in Ambion MEGAclearTM Kit).

2.3.6 modRNA Concentration for In Vivo Use Using Amicon Ultra-4 Centrifugal Filters

1. Ultra-4 centrifugal filters 10 k (Amicon).

2.3.7 Quality Control of modRNA Using TapeStation

1. High-sensitivity RNA ScreenTape (Agilent).

2. Elution buffer (provided in Ambion MEGAclearTM Kit).

2. High-sensitivity RNA ScreenTape Sample Buffer (Agilent). 3. Loading tips (Agilent). 4. Tube strip (Agilent). 5. Caps (Agilent).

3

Methods All animal procedures should be performed under the protocols approved by an Institutional Care and Use Committee.

3.1 Tailed DNA Template Preparation

1. Create a 400 μL PCR master mix solution according to the reaction: 96 μL nuclease-free water, 4 μL plasmid, 100 μL forward + reverse primer mixture (1 μM of each primer), 200 μL KAPA 2 buffer. 2. Aliquot 50 μL of master mix per PCR tube. 3. Run PCR using a thermocycler. Follow the PCR protocol according to Table 1. Elongation will vary depending on open reading frame length, 1 minute per kilobase.

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Table 2 Preparation of NTPS reagents for a 400 μL in vitro transcription reaction

ARCA

Stock concentration (mM)

Amount (μL)

100

20

GTP

75

7.2

ATP

75

36.6

CTP

75

36.6

100

27.64

1-M-Ψ Nuclease-free water

3.2 PCR Product Purification Using QIAquick® PCR Purification Kit

–

32

Conduct all experiments in room temperature. 1. Collect the PCR product in a 15 mL nuclease-free tube. 2. Add 2 mL of PB buffer to the tube and mix with pipetting. Keep ratio of 1:5 of PCR product to PB buffer, respectively. 3. Add 600 μL of the mixture to the supplied filters. 4. Centrifuge at 12,879  g in a microcentrifuge for 1 minute and discard flow-through. 5. Add 750 μL of PE wash buffer to each tube. Centrifuge at 12,879  g for 1 minute. Discard flow-through. 6. Centrifuge at 12,879  g for 3 minutes to dry column. 7. Discard the collection Eppendorf tube.

tube

and

place

filter

in

an

8. Add 50 μL of nuclease-free water. Let stand for 1 minute. 9. Centrifuge at 12,879  g for 1 minute. Combine the purified product, and measure the concentration on a nanodrop. Make sure the ratio of 260/280 > 1.8 and 260/230 > 2.1. 10. To check the quality of the tailed DNA, run on a 1% agarose gel, and check for a single band at its respective size. 3.3 In Vitro Transcription (IVT) Reaction

1. Prepare custom nucleoside triphosphates (NTPS) according to Table 2. 2. Mix reagents in an Eppendorf tube at room temperature in the following order for a total volume of 400 μL: 160 μL custom NTPS (Table 2), 160 μL tailed template (50–100 ng/μL), 40 μL buffer 10 (MEGAscript kit from Ambion), and 40 μL T7 Enzyme (MEGAscript kit from Ambion). 3. Incubate for 4 hours at 37  C. 4. Add 12 μL of Turbo DNase (MEGAscript kit from Ambion) per 400 μL IVT reaction. 5. Mix gently and incubate for 15 minutes at 37  C.

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1. Mix the IVT reaction mixture with 1.1 mL of nuclear-free water to obtain a 1.5 mL solution. 2. Add 150 μL of Antarctic phosphatase and 150 μL of the Antarctic phosphatase buffer. Mix gently by pipetting. 3. Incubate for 1 hour at 37  C. 4. Measure concentration using nanodrop.

3.5 Purify modRNA Using MEGAclear

1. Preheat nuclease-free water 95  C and pre-cool centrifuge to 4  C. 2. Each filter cartridge can bind up to 100 ug of RNA. Prepare filters accordingly. 3. Add 6.3 mL binding solution concentrate to the modRNA obtained after phosphatase treatment. Mix gently by pipetting. 4. Add 4.5 mL of 100% ethanol to the above modRNA mixture. Mix gently by pipetting (see Note 3). 5. Immediately pipet ~700 μL of modRNA mixture to each filter. 6. Centrifuge for 1 minute at 10,000  g at 4  C. 7. Discard flow-through and repeat steps 5 and 6 until each filter binds up to 100 ug of RNA. 8. Add 500 μL of wash solution to each filter. 9. Centrifuge for 1 minute at 10,000  g at 4  C. 10. Discard flow-through, and add 500 μL of wash solution to each filter. 11. Centrifuge for 1 minute at 10,000  g at 4  C. 12. Discard flow-through, and centrifuge for 2 minutes at 10,000  g at 4  C to dry the filter. 13. Transfer filters into clean 1.5 mL collection tubes. 14. Elute RNA from the filter with 50 μL of 95  C nuclease-free water. Wait for 1 minute. 15. Centrifuge for 1 minute at 10,000  g at 4  C. 16. Repeat elution two more times for a total of 150 μL eluted solution per tube. 17. Collect all of the clean modRNA solution into one 15 mL tube (1.5 mL of clean modRNA solution). 18. Measure concentration of purified modRNA.

3.6 modRNA Concentration for In Vivo Use Using Amicon Ultra-4 Centrifugal Filters

1. Pre-cool the centrifuge to 4  C. 2. Wash the filter with 3 mL of nuclease-free water three times by pipetting to remove glycerin. The capacity for each filter is 1 mg of modRNA.

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3. Add 750 μL of nuclease-free water to the filter. 4. Add up to 1 mg of the purified modRNA and mix well by pipetting. 5. Add up to 4 mL of nuclease-free water to the filter, and centrifuge at 1431  g for 45 minutes at 4  C until the volume reduces to ~70 μL (see Note 4). 6. Collect the eluent within the filter into an Eppendorf tube. 7. Measure concentration of the concentrated modRNA in nuclease-free water using a 1:50 dilution (see Notes 5 and 6). 3.7 Quality Control of modRNA Using TapeStation

1. Equilibrate high-sensitivity RNA sample buffer at room temperature for 30 minutes. 2. Launch the Agilent 4200 TapeStation Controller software. Under settings, select RNA assay mode. 3. Load high-sensitivity RNA ScreenTape device into the 4200 TapeStation. 4. Place loading tips into the Agilent 4200 TapeStation instrument. 5. Vortex and spin down reagents before use. 6. For each sample, pipette 1 μL of high-sensitivity RNA sample buffer and 2 μL of RNA sample into a tube strip and apply caps. 7. Vortex sample vials at 358  g for 1 minute and spin down. 8. To denature samples, heat samples at 72  C for 3 minutes. Rest samples on ice for 2 minutes and then spin down. 9. For sample analysis, load samples into the Agilent 4200 TapeStation without strip caps. Use the electronic ladder for comparison. Select the required sample positions on the 4200 TapeStation Controller Software and click start. The Agilent TapeStation Analysis Software opens after the run and displays results.

4

Notes 1. Obtain DNA plasmid containing a recognition site for the forward primer, 50 UTR, ORF, 30 UTR, and reverse primer. This can be done through several companies such as GenScript, Life Technology, or IDT. 2. Depending on the desired translational efficiency and half-life of modRNA in vitro or in vivo, polyA tail lengths can be altered using different reverse primers. 3. In the purifying step using MEGAclear, when making several modRNA samples at once, add ethanol and mix each sample

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before proceeding to the next. This will prevent aggregation of the modRNA. 4. When concentrating, check every 15 minutes when centrifuging to achieve desired concentration. 5. modRNA can be stored for future use at
80  C. If cloudy check the quality using the TapeStation prior to use. 6. For use in vivo, modRNA concentration should not exceed 10 μg/μL to prevent precipitation. References 1. Virani SS, Alonso A, Aparicio HJ, Benjamin EJ, Bittencourt MS, Callaway CW, Carson AP, Chamberlain AM, Cheng S, Delling FN, Elkind MSV, Evenson KR, Ferguson JF, Gupta DK, Khan SS, Kissela BM, Knutson KL, Lee CD, Lewis TT, Liu J, Loop MS, Lutsey PL, Ma J, Mackey J, Martin SS, Matchar DB, Mussolino ME, Navaneethan SD, Perak AM, Roth GA, Samad Z, Satou GM, Schroeder EB, Shah SH, Shay CM, Stokes A, VanWagner LB, Wang NY, Tsao CW, American Heart Association Council on E, Prevention Statistics C, Stroke Statistics S (2021) Heart disease and stroke Statistics-2021 update: a report from the American Heart Association. Circulation 143(8):e254–e743. https://doi.org/10. 1161/CIR.0000000000000950 2. Kaur K, Hadas Y, Kurian AA, Zak MM, Yoo J, Mahmood A, Girard H, Komargodski R, Io T, Santini MP, Sultana N, Sharkar MTK, Magadum A, Fargnoli A, Yoon S, Chepurko E, Chepurko V, Eliyahu E, Pinto D, Lebeche D, Kovacic JC, Hajjar RJ, Rafii S, Zangi L (2021) Direct reprogramming induces vascular regeneration post muscle ischemic injury. Mol Ther 29(10): 3042–3058. https://doi.org/10.1016/j. ymthe.2021.07.014 3. Magadum A, Kurian AA, Chepurko E, Sassi Y, Hajjar RJ, Zangi L (2020) Specific modified mRNA translation system. Circulation 142(25):2485–2488. https://doi.org/10. 1161/CIRCULATIONAHA.120.047211 4. Magadum A, Singh N, Kurian AA, Sharkar MTK, Sultana N, Chepurko E, Kaur K, Zak MM, Hadas Y, Lebeche D, Sahoo S, Hajjar R, Zangi L (2021) Therapeutic delivery of Pip4k2c-modified mRNA attenuates cardiac hypertrophy and fibrosis in the failing heart. Adv Sci (Weinh) 8(10):2004661. https://doi. org/10.1002/advs.202004661 5. Kapeli K, Yeo GW (2012) Genome-wide approaches to dissect the roles of RNA binding proteins in translational control: implications

for neurological diseases. Front Neurosci 6: 144. https://doi.org/10.3389/fnins.2012. 00144 6. Lodish HF (2012) Translational control of protein synthesis: the early years. J Biol Chem 287(43):36528–36535. https://doi.org/10. 1074/jbc.X112.420356 7. Bernal JA (2013) RNA-based tools for nuclear reprogramming and lineage-conversion: towards clinical applications. J Cardiovasc Transl Res 6(6):956–968. https://doi.org/ 10.1007/s12265-013-9494-8 8. Cannon G, Weissman D (2002) RNA based vaccines. DNA Cell Biol 21(12):953–961. h t t p s : // d o i . o r g / 1 0 . 1 0 8 9 / 104454902762053882 9. Hacein-Bey-Abina S, von Kalle C, Schmidt M, Le Deist F, Wulffraat N, McIntyre E, Radford I, Villeval JL, Fraser CC, CavazzanaCalvo M, Fischer A (2003) A serious adverse event after successful gene therapy for X-linked severe combined immunodeficiency. N Engl J Med 348(3):255–256. https://doi.org/10. 1056/NEJM200301163480314 10. Kariko K, Buckstein M, Ni H, Weissman D (2005) Suppression of RNA recognition by toll-like receptors: the impact of nucleoside modification and the evolutionary origin of RNA. Immunity 23(2):165–175. https://doi. org/10.1016/j.immuni.2005.06.008 11. Kariko K, Muramatsu H, Welsh FA, Ludwig J, Kato H, Akira S, Weissman D (2008) Incorporation of pseudouridine into mRNA yields superior nonimmunogenic vector with increased translational capacity and biological stability. Mol Ther 16(11):1833–1840. https://doi.org/10.1038/mt.2008.200 12. Kormann MS, Hasenpusch G, Aneja MK, Nica G, Flemmer AW, Herber-Jonat S, Huppmann M, Mays LE, Illenyi M, Schams A, Griese M, Bittmann I, Handgretinger R, Hartl D, Rosenecker J, Rudolph C (2011) Expression of therapeutic

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proteins after delivery of chemically modified mRNA in mice. Nat Biotechnol 29(2): 154–157. https://doi.org/10.1038/nbt. 1733 13. Bahl K, Senn JJ, Yuzhakov O, Bulychev A, Brito LA, Hassett KJ, Laska ME, Smith M, Almarsson O, Thompson J, Ribeiro AM, Watson M, Zaks T, Ciaramella G (2017) Preclinical and clinical demonstration of immunogenicity by mRNA vaccines against H10N8 and H7N9 influenza viruses. Mol Ther 25(6): 1316–1327. https://doi.org/10.1016/j. ymthe.2017.03.035 14. Richner JM, Himansu S, Dowd KA, Butler SL, Salazar V, Fox JM, Julander JG, Tang WW, Shresta S, Pierson TC, Ciaramella G, Diamond MS (2017) Modified mRNA vaccines protect against Zika virus infection. Cell 169(1):176. https://doi.org/10.1016/j.cell.2017.03.016 15. Heiser A, Coleman D, Dannull J, Yancey D, Maurice MA, Lallas CD, Dahm P, Niedzwiecki D, Gilboa E, Vieweg J (2002) Autologous dendritic cells transfected with prostate-specific antigen RNA stimulate CTL responses against metastatic prostate tumors. J Clin Invest 109(3):409–417. https://doi.org/ 10.1172/JCI14364 16. Morse MA, Nair SK, Boczkowski D, Tyler D, Hurwitz HI, Proia A, Clay TM, Schlom J, Gilboa E, Lyerly HK (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. https://doi.org/10.1385/ IJGC:32:1:1 17. Morse MA, Nair SK, Mosca PJ, Hobeika AC, Clay TM, Deng Y, Boczkowski D, Proia A, Neidzwiecki D, Clavien PA, Hurwitz HI, Schlom J, Gilboa E, Lyerly HK (2003) Immunotherapy with autologous, human dendritic cells transfected with carcinoembryonic antigen mRNA. Cancer Investig 21(3):341–349. https://doi.org/10.1081/cnv-120018224 18. Lui KO, Zangi L, Silva EA, Bu L, Sahara M, Li RA, Mooney DJ, Chien KR (2013) Driving vascular endothelial cell fate of human multipotent Isl1+ heart progenitors with VEGF modified mRNA. Cell Res 23(10): 1172–1186. https://doi.org/10.1038/cr. 2013.112 19. Zangi L, Lui KO, von Gise A, Ma Q, Ebina W, Ptaszek LM, Spater D, Xu H, Tabebordbar M, Gorbatov R, Sena B, Nahrendorf M, Briscoe DM, Li RA, Wagers AJ, Rossi DJ, Pu WT, Chien KR (2013) Modified mRNA directs the fate of heart progenitor cells and induces vascular regeneration after myocardial infarction.

Nat Biotechnol 31(10):898–907. https://doi. org/10.1038/nbt.2682 20. Baden LR, El Sahly HM, Essink B, Kotloff K, Frey S, Novak R, Diemert D, Spector SA, Rouphael N, Creech CB, McGettigan J, Khetan S, Segall N, Solis J, Brosz A, Fierro C, Schwartz H, Neuzil K, Corey L, Gilbert P, Janes H, Follmann D, Marovich M, Mascola J, Polakowski L, Ledgerwood J, Graham BS, Bennett H, Pajon R, Knightly C, Leav B, Deng W, Zhou H, Han S, Ivarsson M, Miller J, Zaks T, Group CS (2021) Efficacy and safety of the mRNA-1273 SARS-CoV-2 vaccine. N Engl J Med 384(5): 4 0 3 – 4 1 6 . h t t p s : // d o i . o r g / 1 0 . 1 0 5 6 / NEJMoa2035389 21. Mullard A (2016) mRNA-based drug approaches phase I milestone. Nat Rev Drug Discov 15(9):595. https://doi.org/10.1038/ nrd.2016.182 22. Polack FP, Thomas SJ, Kitchin N, Absalon J, Gurtman A, Lockhart S, Perez JL, Perez Marc G, Moreira ED, Zerbini C, Bailey R, Swanson KA, Roychoudhury S, Koury K, Li P, Kalina WV, Cooper D, Frenck RW Jr, Hammitt LL, Tureci O, Nell H, Schaefer A, Unal S, Tresnan DB, Mather S, Dormitzer PR, Sahin U, Jansen KU, Gruber WC, Group CCT (2020) Safety and efficacy of the BNT162b2 mRNA Covid-19 vaccine. N Engl J Med 383(27):2603–2615. https://doi.org/10. 1056/NEJMoa2034577 23. Wilgenhof S, Van Nuffel AMT, Benteyn D, Corthals J, Aerts C, Heirman C, Van Riet I, Bonehill A, Thielemans K, Neyns B (2013) A phase IB study on intravenous synthetic mRNA electroporated dendritic cell immunotherapy in pretreated advanced melanoma patients. Ann Oncol 24(10):2686–2693. https://doi.org/ 10.1093/annonc/mdt245 24. Hadas Y, Sultana N, Youssef E, Sharkar MTK, Kaur K, Chepurko E, Zangi L (2019) Optimizing modified mRNA in vitro synthesis protocol for heart gene therapy. Mol Ther Methods Clin Dev 14:300–305. https://doi.org/10.1016/j. omtm.2019.07.006 25. Kaur K, Sultana N, Hadas Y, Magadum A, Sharkar MTK, Chepurko E, Zangi L (2020) Delivery of modified mRNA in a myocardial infarction mouse model. J Vis Exp 160: doi:10.3791/60832 26. Kondrat J, Sultana N, Zangi L (2017) Synthesis of modified mRNA for myocardial delivery. Methods Mol Biol 1521:127–138. https:// doi.org/10.1007/978-1-4939-6588-5_8 27. Sultana N, Hadas Y, Sharkar MTK, Kaur K, Magadum A, Kurian AA, Hossain N, Alburquerque B, Ahmed S, Chepurko E,

Modified mRNA Synthesis Zangi L (2020) Optimization of 5’ untranslated region of modified mRNA for use in cardiac or hepatic ischemic injury. Mol Ther Methods Clin Dev 17:622–633. https://doi. org/10.1016/j.omtm.2020.03.019 28. Sultana N, Magadum A, Hadas Y, Kondrat J, Singh N, Youssef E, Calderon D, Chepurko E, Dubois N, Hajjar RJ, Zangi L (2017) Optimizing cardiac delivery of modified mRNA. Mol Ther 25(6):1306–1315. https://doi.org/10. 1016/j.ymthe.2017.03.016 29. Sultana N, Sharkar MTK, Hadas Y, Chepurko E, Zangi L (2021) In vitro synthesis of modified RNA for cardiac gene therapy. Methods Mol Biol 2158:281–294. https:// doi.org/10.1007/978-1-0716-0668-1_21

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Chapter 8 Design and Production of Heart Chamber-Specific AAV9 Vectors Alina S. Bilal, Donna J. Thuerauf, Erik A. Blackwood, and Christopher C. Glembotski Abstract Adeno-associated virus serotype 9 (AAV9) is often used in heart research involving gene delivery due to its cardiotropism, high transduction efficiency, and little to no pathogenicity, making it highly applicable for gene manipulation, in vivo. However, current AAV9 technology is limited by the lack of strains that can selectively express and elucidate gene function in an atrial- and ventricular-specific manner. In fact, study of gene function in cardiac atria has been limited due to the lack of an appropriate tool to study atrial gene expression in vivo, hindering progress in the study of atrial-specific diseases such as atrial fibrillation, the most common cardiac arrhythmia in the USA. This chapter describes the method for the design and production of such chamber-specific AAV9 vectors, with the use of Nppa and Myl2 promoters to enhance atrial- and ventricular-specific expression. While several gene promoter candidates were considered and tested, Nppa and Myl2 were selected for use here because of their clearly defined regulatory elements that confer cardiac chamber-specific expression. Accordingly, Nppa (
425/+25) and Myl2 (
226/+36) promoter fragments are inserted into AAV9 vectors. The atrial- and ventricular-specific expression conferred by these new recombinant AAV9 was confirmed in a double-fluorescent Cre-dependent reporter mouse model. At only 450 and 262 base pairs of Nppa and Myl2 promoters, respectively, these AAV9 that drive chamber-specific AAV9 transgene expression address two major limitations of AAV9 technology, i.e., achieving chamber-specificity while maximizing space in the AAV genome for insertion of larger transgenes. Key words AAV9, Atria, Ventricle, Chamber-specific, Nppa, Myl2

1

Introduction

1.1 AAV9: Research Tool and Gene Therapy

As the leading cause of death in the USA [1], heart disease has been the focus of medical and academic research, with the aim to understand mechanisms of heart disease for the development of improved therapeutics that can effectively reduce mortality and improve quality of life. Therefore, it is essential to develop new technologies that can address cardiac pathologies in ways that could improve future treatments for heart disease. One recently

Kiyotake Ishikawa (ed.), Cardiac Gene Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2573, https://doi.org/10.1007/978-1-0716-2707-5_8, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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developed technology involves in vivo gene transfer, sometimes called gene therapy, a maneuver whereby the levels of proteins that are found to either be defective or absent in a diseased patient can be increased via ectopic expression of a functioning protein [2]. Additionally, as a research tool, being able to ectopically express transgenes in animal models can facilitate the elucidation of molecular mechanisms of diseases. One mode of ectopically expressing transgenes involves the use of viral vectors, such as adeno-associated virus serotype 9 (AAV9). Due to its cardiotropic nature, high transduction efficiency, low pathogenicity, and low cost of production, AAV9 is a promising therapeutic, and it is a commonly used research tool for heart disease [3–6]. 1.2

AAV9: Structure

1.3 AAV9 vs. Transgenesis

AAV9 has one of the smallest genomes of known viruses, comprising 4.7 kilobase pairs (kb) of single-stranded DNA encapsulated by a non-enveloped icosahedral capsid of 22 nm in diameter (Fig. 1). On both the 50 and 30 ends of the AAV9 genome are 145 base pair stretches of inverted terminal repeats (ITRs) that allow for the expression, replication, and integration of the AAV9 genome into its host. The genome itself is remarkably simple, containing only two genes, rep and cap, which encode four non-structural proteins necessary for viral replication, and three structural proteins that make up the AAV9 capsid (VP1, VP2, VP3) [7–9]. By removing the rep and cap genes and inserting tissue and celltype specific promoters with the objective of achieving tissuespecific transgene expression, recombinant strains of AAV9 can be generated that mediate robust and highly specific, targeted transgene expression (Fig. 1) [4, 10, 11]. However, even though the small size of the AAV9 genome facilitates construct design and development, there are limitations to the size of the regulatory elements and transgene that can be inserted. For example, studies have shown that attempting to package more than 4.7 kb of recombinant DNA into AAV9 results in an unstable viral structure and reduced transfection efficiency [12, 13]. Thus, in order to generate efficient recombinant strains of AAV9, it is essential to limit insert size in order not to exceed 4.7 kb. While AAV9 is one of many technological advancements that facilitate ectopic expression of transgenes in animal models, for the past three decades, the most common method of genetic manipulation in mice has been transgenesis [14]. While a longer and more expensive route, transgenesis results in the generation of genetically modified mice that can be propagated for many generations. Additionally, the genomic environment in which the ectopic DNA is inserted affords more flexibility, in terms of size, as well as greater and more reliable expression, compared to isolation of ectopic DNA into a small 5 kb circular vector, the approximate limit of AAV9. As such, challenges involved in the construction of

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Fig. 1 AAV9 structure and summary of recombinant AAV9 construction. (a) Diagram of the AAV9 virion, showing the icosahedral capsid enclosing its single-stranded genome. (b) Detailed diagram of the wild-type AAV9 genome (wtAAV9) and the sections to be replaced with mammalian promoter and transgene DNA to create the desired recombinant AAV9 (rAAV9); ITR, inverted terminal repeat; black boxes indicate promoters, where arrows direct for transcription; Rep: gene encoding replication proteins (White boxes); Cap: gene encoding capsid proteins (white crisscrossed boxes); green indicates a transgene of interest

recombinant strains of AAV9 include not only limited insert size but also the precise knowledge of regulatory elements that are necessary to achieve transgene expression in specific cells and tissues. 1.4 AAV9: TissueSpecific Enhancements

Studies using AAV9 to examine animal models of heart disease have focused primarily on targeted transgene expression to ventricular myocytes, due to their central role in cardiac function and due to the desire to enhance their function in many cardiac pathologies, such as heart failure [10, 11, 15]. In addition to the heart, AAV9 can infect hepatocytes, albeit, at a lower transduction efficiency than cardiac myocytes [16]. Accordingly, to enhance cardiac myocyte-specific expression of AAV9-encoded genes, two main approaches have evolved: the use of cardiac myocyte-specific promoters, such as the cardiac troponin T (Tnnt2) promoter [10], and use of miRNA targeting sequences (miR-TS), such as the miR-122 TS. miR-122 is a highly expressed miRNA in the liver specifically [17]. Therefore, implementation of a miR-122-TS in an AAV vector results in recognition and repression by endogenous miR-122 in the liver only, resulting in cardiac-specific expression [17].

1.5 Atria vs. Ventricle

Mice are often used for research of disease mechanisms [4, 14]. While mechanisms of heart disease in mouse models have been investigated using recombinant AAV9 developed with cardiac myocyte-specific promoters driving expression in both atrial and ventricular myocytes of the heart [10, 11], cardiac pathologies are often chamber-selective and involve myocytes in the atria or ventricles [18, 19]. Accordingly, reagents capable of conferring cardiac

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chamber-specific gene expression would be highly valuable for studying such cardiac diseases in mice [20]. While myocytes in both the atria and ventricles contribute to cardiac contractility, there are significant functional differences between atrial and ventricular myocytes. As the larger chambers of the heart, the ventricles primarily serve to generate the driving force that sends blood into circulation, while the atrial myocytes have functions in blood movement into the ventricles, as well as serving as endocrine cells that secrete peptide hormones and electrically excitable cells that ensure conduction of cardiac depolarization from the sinoatrial node, across the atria, down into the ventricles, thus ensuring rhythmogenesis and setting the pace for cardiac contractility [18–20]. Thus, both anatomically and physiologically, atria and ventricles have different, specialized roles; as such, defects in atrial or ventricular myocytes could produce distinct disease phenotypes—for example, atrial fibrillation and left ventricular hypertrophy, respectively. Atrial fibrillation and left ventricular hypertrophy are characterized by an irregular heartbeat and reduced cardiac output, respectively, exemplifying the disease phenotypes specific to compromised function of atrial and ventricular myocytes [19, 20]. 1.6 Objective: Generation of Chamber-Specific AAV9 Using Nppa and Myl2 Minimal Promoters

Since there are potential therapeutic and scientific benefits of expressing transgenes selectively in atrial or ventricular myocytes, it is desirable to generate recombinant forms of AAV9 for the expression of transgenes of interest in an atrial- or ventricular-, chamber-specific manner. To design cardiac chamber-specific AAV9, we first determined the minimal promoters of Nppa and Myl2, which encode for atrial natriuretic peptide (ANP) and myosin regulatory light chain 2, ventricular isoform (MLC2v), and inserted them into AAV9 vectors to achieve atrial- and ventricular-myocyte specific expression, respectively, while maximizing space for a transgene insert. Accordingly, the objective of this chapter is to describe the method for the construction and generation of such chamber-specific AAV9 vectors (Fig. 2).

1.7 Identification of the Nppa and Myl2 Minimal Promoters

Several chamber-specific gene promoters were analyzed in mouse hearts, and their chamber-specific expression tested in primary neonatal rat atrial and ventricular myocytes (data not shown; see Notes 1 and 2). Of the gene promoter candidates analyzed, Nppa and Myl2 demonstrated the most robust level of atrial and ventricular-specific expression, respectively, which was maintained with cardiac disease models (data not shown; see Notes 1 and 3). Furthermore, understanding the smallest regions of a promoter that confers cardiac chamber-specific gene expression is instrumental in identifying the minimal promoter necessary to achieve the desired expression in an AAV9 vector. Regulatory elements responsible for the atrial- and ventricular-specific expression of mouse

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Fig. 2 Approach to the design of chamber-specific AAV9. Detailed illustration utilizing an atrial-specific promoter, i.e., Nppa promoter (left) or ventricularspecific promoter, i.e., Myl2 promoter (right) for the design of chamber-specific AAV9; vent ¼ ventricular. Right-hand arrows indicate start of transcription for either Nppa (red; left), Myl2 (blue; right), or GFP (green); red and blue colors indicate atrial- and ventricular myocyte-specific expression, respectively

Nppa and Myl2, respectively, have been previously identified and illustrated in Fig. 3 [21–31]. Both mouse Nppa and Myl2 promoter fragments encompassing those regulatory elements are relatively small and, therefore, appropriate for constructing recombinant AAVs that retain maximal space in the genome to accommodate larger genes-of-choice (Fig. 3). For the mouse Nppa promoter, these regulatory elements are within 425 and 25 bp upstream and downstream of the Nppa 50 UTR, and for the mouse Myl2 promoter, they are 226 and 36 bp upstream and downstream of the Myl2 50 UTR, respectively.

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Fig. 3 Nppa and Myl2 gene and promoter map and sequence. (a) Map and (b) Sequence of Nppa gene and promoter. Map shows promoter elements (indicated by the shapes) and the regions responsible for chamberspecific expression, GATA: GATA-4/6 transcription factor binding element; SRE: serum response element recognized by serum response factor, NKE: Nkx2.5 transcription factor binding element; NRSE: neuronrestrictive silencer element recognized by the neuron-restrictive silencer factor. Blue text and box (spanning nucleotides
627 to
425) indicate promoter region responsible for ventricular expression during embryonic development; red text and box (spanning nucleotides
425 to +1) indicate promoter region used in this study to promote atrial-restricted expression; purple underlined text and purple boxes indicate NRSE [45]; Nppa sequence ID: ENSMUSG00000041616.9. (c) Map and (d) Sequence of Myl2 gene and promoter. Blue text and box (spanning nucleotides
226 to +1) indicate promoter region used in this study for ventricular-restricted expression, containing the hypoxia-inducible factor binding elements (HF3, 2, 1a, 1b) responsible for ventricular-specific expression, presented as bold, underlined blue text or rectangles on map; Myl2 sequence ID: ENSMUSG00000013936.12. (a–d) Plain black text and white boxes indicate UTRs; TSS: transcript start site; bold underlined black text and black boxes indicate exons; lower-case plain black text and lines indicate introns; note, gene sequence presented is partial for both Nppa and Myl2. Nppa (
425/+25) and Myl2 (
226/+36) were the promoter fragments used for construction of chamber-specific AAV9

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Materials Cloning Reagents

1. Restriction endonuclease MluI and restriction digest buffer. 2. Restriction endonuclease HindIII and restriction digest buffer. 3. Restriction endonuclease SmaI and restriction digest buffer. 4. PCR and gel purification kit. 5. DNA miniprep kit. 6. Primers (Table 1), oligos resuspended in molecular biology grade water. 7. T4 DNA Ligase and ligation buffer. 8. DNA maxi-prep kit. 9. TAE buffer: 4.84 g Tris base, 1.142 mL Glacial acetic acid, 2 mL 0.5 M EDTA, pH 8.0. 10. 1% agarose gel: 1% agarose (1 g/100 mL), TAE buffer, 0.1ug/ mL ethidium bromide. 11. Stellar competent bacteria (Takara). 12. SURE2 competent bacteria (Stratagene). 13. 14 mL round bottom polypropylene tubes. 14. SOC media: 3.603 g/L dextrose, 0.186 g/L KCl, 4.8 g/L MgSO4, 20 g/L tryptone, 5 g/L yeast extract. 15. Incubator shaker (Amerex Gyromax 737). 16. Ampicillin-resistant agar plates: 25 g LB broth/L, 15 g agaragar/L, 50ug/mL ampicillin. 17. 95 mm petri dishes. 18. Ampicillin-resistant lysogeny broth (LB) media: 25 g LB broth/L, 50ug/mL ampicillin. 19. Glass test tubes. 20. Alkaline phosphatase and alkaline phosphatase buffer.

2.2 AAV9 Production Reagents

1. AAVPro 293T cell line (Takara). 2. Water-jacketed incubator (37  C and 5% CO2). 3. DMEM/F12: L-glutamine, phenol red, high glucose, sodium pyruvate. 4. Fetal bovine serum. 5. Penicillin-streptomycin-glutamine (PSG). 6. Fungizone. 7. Culturing media: DMEM/F12, 10% fetal bovine serum, 1 PSG and fungizone.

ggaacgacgcgtAGAGAGCACACCCCATCATC

Myl2 (
226/+36)

Underlined: Clamp Bolded: Indicated restriction sites

ggaacgacgcgtATTGCCTCCTCTCCCGC

Nppa (
425/+25)

Forward with MluI

Table 1 Primers for PCR amplification of Nppa and Myl2 promoter fragments

ggaacgaagcttCCTCTGGAGAGTTCGAGGAG

ggaacgaagcttGTCTCCTAGCTGCCAGCATC

Reverse with HindIII

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8. Trypsin. 9. Transfection media: DMEM/F12, 2% fetal bovine serum, 1 PSG and fungizone. 10. Polyethylenimine (PEI) 0.517 mg/mL. 11. T-175 culture flasks. 12. 150 mm culture dishes. 13. pAAV2/9n: rep/cap gene expression construct (Addgene #112865). 14. pAdDeltaF6: AAV helper plasmid (Addgene #112867). 15. pds-hTnnt2-Cre (gift from Dr. Oliver Mu¨ller) [10]. 2.3 AAV9 Purification Reagents

1. Lysis buffer: 150 mM NaCl, 50 mM Tris–HCl, pH 8.5. 2. Benzonase 250 U/μL. 3. 1 M MgCl2. 4. OptiSeal polyallomer tubes. 5. 15% iodixanol: 45 mL Iodixanol (OptiPrep), 60 mL 3 M NaCl, 36 mL 5 TD buffer, 39 mL Molecular biology grade water for a total of 180 mL ¼ enough for 24 OptiSeal polyallomer tubes. 6. 25% iodixanol: 50 mL iodixanol (OptiPrep), 24 mL 5 TD buffer, 300 μL Phenol Red (0.5%), 46 mL molecular biology grade water for a total of 120 mL ¼ enough for 24 OptiSeal polyallomer tubes. 7. 40% iodixanol: 68 mL iodixanol (OptiPrep), 20 mL 5 TD buffer, 12 mL molecular biology grade water for a total of 100 mL ¼ enough for 24 OptiSeal polyallomer tubes. 8. 60% iodixanol: 100 mL iodixanol (OptiPrep), 250 μL phenol red ¼ enough for 24 OptiSeal polyallomer tubes. 9. 5 TD buffer: 5 PBS, 5 mM MgCl2, 12.5 mM KCl. 10. Spinal needle, 18G  3.5 inches. 11. 18G needles. 12. 10 mL syringes. 13. Ultracentrifuge. 14. 70Ti Rotor. 15. 12% SDS PAGE gels. 16. Coomassie blue stain (InstantBlue). 17. β-Mercaptoethanol (BME). 18. 4 Laemmli Sample Buffer.

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2.4 AAV9 Preparation for In Vivo Injection: Reagents and Delivery Equipment

1. Vivaspin 20: 100 kDa MWCO. 2. Lactated Ringer’s Solution (LR): 600 mg NaCl, 310 mg C3H5NaO3, 30 mg KCl, 20 mg CaCl2 dihydrate. 3. AAV9 standard (a virus of known titer – we used a virus gifted to us by the lab of Dr. Roger Hajjar). 4. 2 Laemmli Sample Buffer. 5. 1 phosphate-buffered saline (PBS). 6. 27 ½ G needles for tail-vein injections. 7. 1 mL syringes for tail-vein injections.

3

Methods

3.1 Construction of Cardiac ChamberSpecific AAV Plasmids

Here, we describe the method for cloning promoter fragments of Nppa and Myl2 into AAV plasmids to generate chamber-specific AAV9 vectors (Fig. 4). The cloning procedure detailed below applies to cloning promoter fragments into AAV plasmids but could be applied for any plasmid with minor changes in bacterial strain and growing temperature (see Note 4). 1. Choose the parent plasmid. We used the AAV plasmid pds-hTnnt2-Cre (a gift from Dr. Oliver Mu¨ller) [10], where hTnnt2 is a human Tnnt2 promoter fragment driving the gene Cre recombinase (Cre), making it an applicable vector to be used in transgenic mice using the Cre-lox system. Several restriction sites flank the hTnnt2 promoter in this plasmid, one set being MluI (upstream of the promoter) and HindIII (downstream of the promoter and upstream of the cmv enhancer). MluI and HindIII were the restriction sites chosen for the insertion of Nppa and Myl2 promoter fragments so as to completely replace hTnnt2 while maintaining the cmv enhancer (see Note 5). 2. Design primers with restriction sites MluI (at the 50 end) and HindIII (at the 30 end) to extract promoter fragments of Nppa (
425/+25) and Myl2 (
226/+36) from genomic DNA and subsequently clone into the parent plasmid (Table 1). When adding restriction sites, include a “clamp” sequence to facilitate restriction enzyme endonuclease activity, as underlined in Table 1. 3. Perform PCR to amplify the relevant promoter fragment. 4. Analyze 10 μL of the PCR product on a 1% agarose gel. The size corresponding to the promoter fragment should be the brightest and most distinct band. The percentage of agarose should be increased to 2% for smaller inserts under 200 bp (see Note 5).

AAV plasmid

Cre

Cre

Tnnt2 promoter MluI HindIII Nppa gene

Nppa

Rev

For

Myl2 gene

Myl2

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ligase

ligase

Transform in Stellar Pick colonies Myl2 prom

Nppa prom

Sequence plasmids to verify successful cloning Cre Nppa promoter

Cre Myl2 promoter

Transform in SURE2 Streak to isolate Myl2 prom

Nppa prom Inoculate for maxi-prep

Nppa pr Myl2 prom

Plasmid ready for transfection

Fig. 4 Illustration of AAV cloning workflow. Cloning of Nppa (
425/+25) and Myl2 (
226/+36) promoter fragments into an AAV plasmid is illustrated in Steps 1–7, starting with (1) digestion of parent AAV plasmid, resulting in a promoter-less parent plasmid with MluI (50 ) and HindIII (30 ) sticky ends, followed by (2) PCR product of Nppa (
425/+25) and Myl2 (
226/+36) generated from mouse genomic DNA using the appropriate forward (For) and reverse (Rev) primers, then (3) ligation of promoter fragments into parent plasmid, (4) transformation into Stellar competent bacteria, (5) verification of successful cloning by sequencing, (6) transformation of sequencing-verified clones into SURE2 for (7) large-scale production of AAV plasmid, and transfection to produce chamber-specific AAV9. Yellow, red and blue indicate Tnnt2, Nppa and Myl2, respectively

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5. Purify the PCR-amplified reaction product. 6. Digest 1 ug of the purified PCR product and 500–700 ng of the parent plasmid with MluI and HindIII (see Note 5). 7. Analyze on a 1% agarose gel. 8. Excise the digested PCR product and linearized parent plasmid (without the hTnnt2 insert) and gel purify. Elute in a small volume of 20–30 μL. 9. Perform a 10 μL ligation reaction with 1 μL of the linearized parent plasmid and 3 μL of the Nppa or Myl2 digested PCR product from Subheading 3.1, step 8. Ligate for at least 15 minutes at room temperature (RT). Include a “self” control which constitutes the parent plasmid alone to determine ligation efficiency, which will indicate how many colonies to harvest. 10. Perform transformations. Add 3 μL of the ligation products into 30 μL of Stellar Competent bacteria in a 14 mL round bottom polypropylene tube (see Note 4). Heat shock at 42  C for 45 seconds exactly; let cool on ice for 1 minute and then add 300 μL SOC media. Shake at 220–240 rpm for at least 45 minutes in a 37  C incubator shaker. 11. Spread 50 μL and 250 μL of the transformations each on two 95 mm ampicillin-containing agar plates. Incubate at 30  C for 24 hours. Harvest resulting colonies. The colonies will be very small, but do not allow colonies to grow any longer than 24 hours. If there are no colonies on the “self” control, harvest two colonies from the plates containing the promoter fragment insert, i.e., insert reaction plates. If the ratio of colonies on “self” control plate to insert is 1:10, then harvest 4 colonies from the insert reaction; if 5:10, then harvest 6–10 colonies. Inoculate selected colonies in 5–6 mL of lysogeny broth (LB) media with ampicillin in glass test tubes. Shake at 220–240 rpm at 30  C overnight (temperature is critical. See Note 4) in an incubator shaker. Do not let the bacterial cultures grow longer than 20 hours. 12. Isolate plasmid from bacterial culture. The yield will be low due to the low copy nature of AAV plasmids. Elute in a low volume when extracting plasmid. If the commercial kit calls for elution in 50–60 μL, aim for 30–40 μL to maintain a more highly concentrated DNA sample. 13. Verify successful construct production. To verify that the product plasmid contains the desired promoter insertion, restriction digest the original pds-hTnnt2-Cre and product plasmids (pds-Nppa-Cre and pds-Myl2-Cre) with MluI and HindIII to differentiate it from the original parent plasmid and to ensure the proper insert size (see Note 5).

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In addition, restriction digest with SmaI to verify integrity of inverted terminal repeats (ITRs). After digestion with SmaI, the original parent plasmid (pds-hTnnt2-Cre) used for this study will generate four DNA fragments on an agarose gel. Replacing the hTnnt2 promoter with the mouse Nppa and Myl2 promoter fragment will still result in four DNA fragments in total, but of slightly different sizes, owing to the different promoter sizes. Send plasmids for sequencing (e.g., Retrogen Sequencing, Inc.) or sequence in-house to verify that no mutations were introduced during the cloning process. Proceed to the next step with the confirmed correct clone. 14. Transform a single correct clone each for pds-Nppa-Cre and pds-Myl2-Cre into SURE2 competent bacteria. Add 5 ng of plasmid to 10 μL of SURE2 bacteria; heat shock at 42  C for 30 seconds exactly and then ice for 1 minute. Add 100 μL of SOC media and shake at 220–240 rpm for at least 45 minutes in a 37  C incubator shaker before plating. It is best to maintain and isolate large quantities of plasmid from SURE2 bacteria due to their assurance of low recombination frequency (see Note 4). 15. Plate transformed SURE2 bacteria by streaking to isolate on 95 mm ampicillin-containing agar plates. Incubate at 30  C for no more than 24 hours. 16. Harvest one isolated colony each. Inoculate in 5–6 mL LB media with ampicillin in a glass test tube. Incubate with shaking at 220–240 rpm at 30  C overnight in an incubator shaker. Do not grow for more than 20 hours. 17. Introduce 1 mL of culture into 1 mL of sterile glycerol to generate a glycerol stock for long-term storage and propagation in the future. Invert to mix until homogenous. Store at
80  C. You may use another 1 mL of the culture to inoculate a larger volume of LB media, or take directly from the glycerol stock in the future. A large volume, e.g., 1 L, will be required to produce enough plasmid for transfection in AAV9 production. Whenever growing bacterial cultures of AAV, grow in an incubator shaker at 30  C shaking at 220–240 rpm overnight. Never grow more than 20 hours. 18. Perform a restriction digest with SmaI to verify integrity of ITRs. Checking the integrity of the ITRs is necessary with every new plasmid preparation. Only transfect plasmids with intact ITRs for production of AAV9 (see Note 4). 3.2 Production of Cardiac ChamberSpecific AAV9

This section describes the procedure used to produce AAV9 (Fig. 5) [4], which was followed for the production of the new chamber-specific AAV9.

Plate HEK293 cells

Perform PEI transfection PEI 2/9

Ad Ad

Ad

Harvest cells

Ad

Purify and collect AAV9 Centrifugation

AAV9-containing cells

15% 25% 40% 60% Iodixanol gradient

Analyze collected fractions of AAV9 kDa

VP3 VP2 VP1

75 60

AAV9

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4x wash with LR Vivaspin Quantify virus Standard kDa

aAAV vAAV

AAV9 ready for injection

75 60 aAAV vAAV

Fig. 5 Illustration of AAV9 production and purification work-flow. Production of atrial- and ventricular-specific AAV9 (aAAV9 and vAAV9, respectively) is illustrated in steps 1–8, starting with (1) plating of HEK293 cells on T-175 flasks, (2) PEI transfection of HEK293 cells, with the entry of PEI-coated plasmids into one HEK293 cell shown, followed by (3) harvest of AAV9-containing HEK293 cells after 3–4-day incubation, (4) purification of AAV9 by iodixanol gradient centrifugation followed by collection of purified AAV9, (5) analysis of collected AAV9 for purity, where only the fractions containing robust AAV9 with minimal contaminants is saved; check marks and x’s indicate fractions to be saved (checks) and discarded (x’s). Then, (6) concentration of AAV9 in lactated ringer’s (LR) solution via centrifugation in Vivaspin 20 columns. After concentrating AAV9 in LR, (7) Quantification of aAAV9 and vAAV9 (technical replicate of 2) against a dilution series of a standard virus of known titer (technical replicate of 2) is performed for (8) Generation of aAAV9 and vAAV9 that is safe and ready for injection. N ¼ nucleus; PEI ¼ polyethylenimine (positively charged squiggles); Ad ¼ helper plasmid encoding adenoviral helper genes; 2/9 ¼ helper plasmid encoding rep and cap genes; red and blue circles indicate atrial- and ventricular-specific plasmids, respectively; note that the atrial- and ventricular-specific AAV plasmids would be transfected separately

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1. Culture HEK293 cells specifically engineered for AAV production on 150 mm dishes in culturing media. HEK293 cells specifically engineered for AAV production must be used to ensure production of AAV. Passage HEK293 cells 3–4 times until 24–32 plates are 80% confluent. At 80% confluency, each 150 mm dish contains 15–20 million cells. Trypsinize cells from 24 to 32 150 mm culture dishes, and seed onto 48 T175 flasks at 12 million cells per flask, in 18 mL of culturing media. Incubate the T-175 flasks at least 12–18 h before transfection at 37  C and 5% CO2. 2. Transfect HEK293 cells (see Note 6) to produce AAV9. Subheading 3.2, steps 3–8 describe the amount of transfection reagents for one T-175 flask at a density of 12 million cells/ flask. Scale volumes proportionally for a full AAV9 preparation of 48 T-175 flasks. 3. Prepare transfection reagent and plasmids by diluting separately in 1:1 DMEM:F12 (no antibiotic). First, dilute the transfection reagent, polyethylenimine (PEI), by adding 160 μL PEI per 0.5 mL of 1:1 DMEM:F12 (no antibiotic). Then, dilute plasmids by adding 5 μg of either the atrial- or ventricular-specific AAV plasmid cloned in Subheading 3.1, 5 μg of pAAV2/9n, and 10 μg of pAdDeltaF6 per 0.5 mL of 1:1 DMEM:F12 (no antibiotic). The pAAV2/9n plasmid is what will make the AAV serotype AAV9. Allow to sit at RT for 5 minutes. 4. Combine diluted PEI with the diluted plasmids. Vortex vigorously for 1 minute. 5. Allow the mixture to sit at RT for 30 minutes. 6. After 30 minutes, combine the mixture with 17 mL of transfection media. 7. Aspirate culturing media on one T-175 flask and replace with transfection media. 8. Rock for 1–2 minutes before placing in the 37  C/5% CO2 incubator to incubate for 3–4 days. Incubation at 3 or 4 days has given us the same titer. 9. After a 3–4 day incubation, collect HEK293 cells from all 48 T175 flasks by scraping in the transfection media it has incubated in. 10. Combine cells and the transfection media from 2 T-175 flasks into one 50 mL conical tube, resulting in 24 total 50 mL conical tubes. 11. Pellet the cells gently by centrifugation at 500  g for 10 minutes at RT. 12. Discard the supernatant of all 24 50 mL conical tubes.

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13. Add 20 mL of lysis buffer to one conical tube and resuspend and then transfer the resuspension into five additional conical tubes until six conical tubes have been combined into one. 14. There should now be four 50 mL conical tubes of resuspended AAV9-containing cell lysate. 15. The AAV9-containing cell lysate should then be subjected to three rounds of freeze-thaw. 16. Immediately after the third thaw, treat the lysate with 11 μL of Benzonase nuclease and 20 μL of 1 M MgCl2 at 37  C for 30 min. 17. Pellet what is now cellular debris by centrifugation at 3200  g for 20 min at RT. 18. Collect the supernatant and discard the pellet. The supernatant contains the AAV9 particles that can now be purified in the next step. 3.3 Purification of Cardiac ChamberSpecific AAV9

1. Purify AAV9 by use of iodixanol gradient centrifugation. Set up an iodixanol gradient in OptiSeal polyallomer tubes with four layers of iodixanol solutions: 15%, 25%, 40%, and 60% layers. Using a 5 mL serological pipet, add 7.3 mL of the 15% layer to the bottom of each tube. Then using a syringe needle, set up each subsequent layer slowly (4.9 mL of the 25% layer and 4.0 mL each of the 40% and 60% layers), placing the needle all the way at the bottom of the tube, so that each layer is below the other (Fig. 5). Then, layer the AAV9-containing supernatant from 3.2.17 (~10 mL) carefully on top of the 15% layer by use of an 18G needle and 10 mL syringe. Bend the 18G needle so that it touches the inner plastic tube and falls gently on top of the 15% layer. Careful not to disturb the layers. Each 50 mL conical tube of AAV9-containing supernatant will be layered onto two gradient tubes, i.e., four 50 mL conicals ¼ eight gradient tubes. 2. Seal and balance the iodixanol gradient tubes. 3. Ultracentrifuge in a 70Ti rotor at 69,000 rpm (or 490,000  g) at 16  C for 1 hour with maximum acceleration and low deceleration (i.e., coasting to a stop with no break). 4. Collect virus by breaking the seal on top of the tube and inserting a needle 2 mm below the 40–60% interface and collecting 3–5 fractions (~2–3 mL total) of this interface and as much of the 40% layer as possible. Try to avoid collecting any of the 25% layer, which contains empty AAV9 particles (no genome) and other cellular proteins. 5. Before proceeding to concentration of AAV9 for injection in vivo, ensure collected fractions of AAV9 contain only AAV9 and are free from contamination of other cellular

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proteins. To do this, prepare 21 μL of each collected fraction to 7 μL 4 Laemmli sample buffer supplemented with β-mercaptoethanol (5 μL per 100 μL of 4 Laemmli). Boil samples at 100  C for 5 minutes. 6. Run samples on a 12% SDS-PAGE gel, followed by staining with InstantBlue to visualize the viral capsid proteins, VP1, VP2, and VP3, whose molecular weights are 87, 73, and 62 kDa, respectively [9] (Fig. 5). The capsids will be readily apparent after 15 minutes. After this time, contamination of other cellular proteins will be readily apparent as a “smear” across the lane, or as distinct bands of molecular weights not corresponding to the viral capsids. Discard all fractions with contamination that is equal to or greater than the viral capsid band intensity, which is determined qualitatively with the naked eye. 3.4 Preparation of Cardiac ChamberSpecific AAV9 for Usable and Safe Injection In Vivo

This step is necessary not only to concentrate AAV9 into a workable titer but also to exchange the iodixanol solution with Lactated ringer’s solution (LR) for safe injection in vivo. 1. Combine clean fractions of purified AAV9 obtained from Subheading 3.3 (Fig. 5). 2. Dilute at least 3 with LR. 3. Add to four ultrafiltration devices, i.e., Vivaspin 20, and centrifuge at 3000  g for 10–20 minutes at RT. Discard the effluent. 4. Repeat Subheading 3.4, step 3 if more AAV9-containing diluted iodixanol (from Subheading 3.4, step 2) is remaining. 5. Once the AAV9-containing diluted iodixanol is packed below the 1.0 mL mark on the column of the Vivaspin, fill the Vivaspin up to 20 mL with LR and proceed to centrifuge at 3000  g for 10–20 minutes at RT. 6. Repeat centrifugation and discarding of effluent three more times. This repeated centrifugation is necessary for thorough exchange of iodixanol with LR, resulting in pure AAV9 in LR. 7. At the last centrifugation, ensure the level of AAV9-containing LR reaches the 0.2 mL mark for each Vivaspin column. 8. Combine the AAV9-containing LR from all Vivaspin columns. Since each Vivaspin should have 0.2 mL of AAV9-containing LR, there should be about ~800 μL total. 9. Proceed to the next step or store in aliquots at
80  C. Storing in aliquots is preferred to prevent continued freeze-thaws. 10. Quantify AAV9 by preparing a dilution series of an AAV9 of known titer, i.e., the standard. Add 16 μL of the standard to 4 μL of phosphate-buffered saline (PBS) in a tube. From this

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tube, take 10 μL and add to 10 μL of PBS in a new tube and so on until you have four tubes in total. Prepare a technical replicate of two. 11. Prepare the chamber-specific AAV9 for quantification by adding 8 μL to 2 μL of PBS. Prepare a technical replicate of two for each. 12. Supplement 100 μL of 2 Laemmli sample buffer with 5 μL BME. Then, add 10 μL of 2 Laemmli sample buffer supplemented with BME to all samples. Boil at 100  C for 5 minutes. 13. Run samples on a 12% SDS-PAGE gel, following by staining with InstantBlue to visualize the viral capsids (Fig. 5). 14. Perform densitometry analysis of the standard virus to yield a standard curve, using software like ImageJ and Microsoft Excel, which is used to determine the concentration of the new AAV9 preparation (see Note 6). 15. Inject the chamber-specific AAV9 via tail-vein injection into adult transgenic mice utilizing the Cre-lox system, i.e., containing a gene of interest flanked with LoxP sites for Cre recombinase to excise [32]. A dose of 1  1011 viral particles and 3x1011 viral particles has been tested with these vectors, where the 1  1011 dose is sufficient to drive Cre expression [33], with a dose-dependent increase at the 3  1011 dose (Fig. 6; see Notes 7 and 8).

4

Notes 1. Before cloning into AAV plasmids, the mouse Nppa and Myl2 promoters were first tested in cell culture in primary neonatal rat atrial and ventricular myocytes (NRAMs and NRVMs, respectively; data not shown). Several truncations of the Nppa and Myl2 promoter were cloned into GFP reporter constructs. While the Nppa promoter in an AAV9 vector has been recently published as an atrial-specific AAV9 [34], this promoter fragment contains a region of regulatory elements responsible for ventricular expression during early development [26, 27] and in our cell culture experiments promoted GFP expression in ventricular myocytes during hypertrophic signaling induced by phenylephrine. For this reason, the region in question (between 627 and 425 bp upstream of the 50 UTR) (Fig. 3) was removed, to maintain atrial specificity and minimize promoter size. The regulatory elements for Myl2 responsible for ventricular and cardiac specificity were well defined as four 10–13 bp elements (Fig. 3), residing within 226 bp upstream of the 50 UTR [28, 30, 31]. A range of promoter fragments from

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5’ ITR Negative control 3’ ITR Tnnt2 promoter Nppa promoter Cre Myl2 promoter recombinase

N P A

-425

+25

V

ria Ve n L i tric v e le G r as Br troc ai n Ki dn Ad e y ip os e

-226 +36

At

B

kDa

N

GAPDH GFP

37

P

GAPDH GFP

37

A

GAPDH GFP

37

V

GAPDH GFP

37

A

C

Ventricle

Atria

1X

V

3X

1X

3X

kDa

GAPDH GFP

37

GAPDH GFP

37

Fig. 6 Testing of chamber-specific AAV9 in vivo. (a) Diagram of the AAV9negative control (N) vector [46] driven by a distinct Myl2 promoter, AAV9positive (P) control vector driven by the human Tnnt2 promoter, atrial-specific AAV9 (A) vector driven by the mouse Nppa promoter, and ventricular-specific (V) AAV9 vector driven by the mouse Myl2 promoter; Cre recombinase is expressed by AAV9-P, AAV9-A, and AAV9-V, whereas AAV9-N has no transgene; ITR, inverted terminal repeat (b) Immunoblot (n ¼ 1 animal for each group) of GAPDH and GFP in extracts from atria, ventricle, liver, gastrocnemius (gastroc), brain, kidney, and adipose from double-fluorescent Cre-dependent reporter mice [32] injected with 1  1011 viral particles of AAV9N, AAV9-P, AAV9-A, and AAV9-V. This mouse model will express GFP only in the presence of Cre (see Notes 7 and 8). (c) Immunoblot (n ¼ 2 animals for each group) of GAPDH and GFP in atrial and ventricular extracts from the same double-fluorescent Cre-dependent reporter mice injected with 1  1011 (1) and 3  1011 (3) viral particles of AAV9-A and AAV9-V

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1000 bp and down to 226 bp upstream of the 50 UTR were tested in NRAMs and NRVMs. All truncations, including the smallest truncation (
226/+36), promoted robust ventricular-specific expression that did not change in ventricular specificity with hypertrophic signaling induced by phenylephrine. Based on the cell culture experiments, Nppa (
425/ +25) and Myl2 (
226/+36) promoter fragments were selected for further development of the chamber-specific AAV9 reported here. 2. In addition to Nppa and Myl2, Sln and Pln were candidate gene promoters considered for the generation of atrial- and ventricular-specific AAV9-mediated transgene expression, respectively. Sln and Pln encode sarcolipin and phospholamban, respectively, both of which regulate SERCA2a (the cardiac isoform of sarcoplasmic reticulum Ca2+ ATPase) activity in cardiac myocytes, with studies demonstrating that SLN and PLN act as the predominant form of SERCA2a regulation in atrial and ventricular myocytes, respectively [35, 36]. However, full characterization of the regulatory elements of the Sln and Pln promoters responsible for chamber-specific expression has not been performed. In an attempt to determine the minimal promoters that would confer chamber-specific expression, fragments of the mouse Sln and Pln promoters from 2000 bp to 500 bp upstream of the 50 UTR were used to generate luciferase reporter constructs, which were transfected into HeLa cells, NRAMs, and NRVMs (data not shown). Luciferase assays demonstrated no differences in the activities of large and small promoter fragments for either Sln or Pln. Between cell types, both Sln and Pln promoters expressed most robustly in NRVMs, with moderate expression in NRAMs, and significantly less expression in HeLa cells. While this trend was somewhat expected for the Pln promoter fragments, it was not for the Sln promoter fragments, where expression in atrial myocytes was expected to be greater than in ventricular myocytes. Therefore, we reasoned the regulatory elements responsible for Sln’s atrial-specific expression must not reside within 2000 bp upstream of the 50 UTR. Recently, a study published an atrial-specific AAV9 using the human Sln promoter and CRM4 enhancer [37]. In our hands, in atrial extracts, we were unable to detect Cre or GFP driven by the published Sln and CRM4 atrial-specific vector by qPCR or immunoblot at 1  1011 viral particles, delivered via tail-vein injection (Data not shown). However, it is possible that this dose of AAV9 was insufficient. Still, because of the large size of the promoters used in the generation of this AAV9, we sought to design an atrial-specific AAV9 vector with

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minimal promoter sequences that would still support atrialspecific transgene expression. 3. Because of our goal of developing chamber-specific vectors that maintain chamber-specific expression in normal mice, as well as in mouse models of cardiac pathology, we measured gene expression of several gene candidate promoters in the hearts of healthy mice, as well as those subjected to transverse aortic constriction, a surgical maneuver to induce pressure-overload induced cardiac hypertrophy [38] (Data not shown). We also measured gene transcripts in mouse atria and ventricles at various stages of development, to ensure that chamber specificity of the promoter selected does not change as a function of age. The nuances of the dynamic changes that can occur in gene expression during development [39] and pathology [26, 31, 40, 41] underscore the necessity to have a wellcharacterized promoter with defined regulatory elements responsible for temporal, pathological, and spatial expression. 4. One of our greatest cloning challenges has been the successful transformation of our ligations in bacterial strains that are designed to prevent recombination and deletion of secondary structures, such as the inverted terminal repeats (ITRs) that exist in AAV plasmids. One such bacterial strain used for this purpose is SURE2 (Stop Unwanted Rearrangement Events) [42]. The efficiency of transformation of ligated products in SURE2 is extremely low in our hands. For this reason, we use a more conventional strain, such as Stellar, and maintain growth of bacterial colonies and cultures at 30  C. Maintenance at this temperature has proven to be key for the prevention of recombination of the AAV plasmid and loss of ITRs. However, after successful transformation and generation of intact plasmid, if we continue to maintain AAV plasmids in Stellar bacteria, we sometimes observed mixed DNA, some with intact ITRs and some with one missing, despite maintenance at 30  C. Therefore, our protocol has been modified to transform ligations of AAV plasmids in Stellar competent bacteria but maintain and grow large volumes of intact AAV plasmids in SURE2 bacteria, which will always keep ITRs intact if cultures are grown at 30  C. With that said, we have seen that those AAV plasmids that are missing one ITR are successfully packaged into AAV9 capsids and have comparable expression to those with intact ITRs, in vivo. Therefore, the necessity for two intact ITRs may not be a requisite for expression of AAV9 in mouse models. 5. After generating a PCR product of the Nppa promoter with the primers listed in Table 1, a restriction digest of said PCR product with MluI and HindIII will result in two DNA fragments, due to an internal HindIII in the endogenous mouse

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Nppa promoter. The larger fragment (294 bp) will contain a 50 MluI and 30 HindIII overhang, and the smaller fragment (160 bp) will contain both a 50 and 30 HindIII overhang. We cloned the Nppa promoter by a two-step process: first, following the protocol described, we constructed a clone with the 294 bp Nppa fragment; then, using the aforementioned clone as the “parent” plasmid, we digested the new parent plasmid with HindIII and subjected only the new parent plasmid to alkaline phosphatase treatment, and ligated in the 160 bp Nppa fragment following the protocol described. It is important to save several clones and send for sequencing to check for correct orientation of the 160 bp fragment, as it will ligate in both directions. 6. When AAV9 production is low, it almost always is the result of poor transfection. Accordingly, we strive to ensure that helper plasmids are of high quality, i.e., are primarily supercoiled with no nicking or RNA contamination. Another source of reduced AAV9 production concerns the transfection method, i.e., a double vs triple transfection. In double transfection, the adenoviral “helper” genes and rep/cap genes specific for AAV9 are encoded in the same plasmid, while in triple transfection, these genes are on separate plasmids [43]. In our hands, both modes of transfection work, but only with specific AAV plasmids. For instance, we have two distinct types of AAV plasmids: pTRUFAAV, a gift from Dr. Roger Hajjar, and pds-AAV, a gift from Dr. Oliver Mu¨ller (the parent AAV plasmid used in this study). For pTRUF-AAV, double transfection results in high production of AAV9, but with pds-AAV, the production of AAV9 is significantly lower. To troubleshoot this, we performed a triple transfection with pds-AAV plasmids, and this significantly increased the yield of AAV9 by about threefold (Data not shown). In the current study, only the triple transfection method and pds-AAV plasmid were used to generate AAV9. 7. We recently published the experiments characterizing these chamber-specific AAV9 in vivo [33]. To ensure myocyte, as well as chamber-specific expression, we developed a protocol to isolate atrial and ventricular myocytes and non-myocytes from the same heart [44]. Using this method, we showed that the chamber-specific vectors demonstrated robust cardiac myocyte-specific Cre and GFP expression, with negligible expression in the non-myocyte fractions [33]. Examining expression in myocyte and non-myocyte cells has also proven useful for determining knockout/knockdown efficiency for cell-specific vectors. In data not shown, analyzing a myocytespecific knockdown/knockout in whole atrial or ventricular tissue can prove to be challenging if the gene being manipulated is robustly expressed in non-myocytes. The expression of

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said gene in non-myocytes will often obstruct the ability to discern knockdown in myocytes. This is primarily true when measured at the genetic or transcript level. To avoid this effect, we suggest measuring the gene of interest in isolated myocytes. 8. The mouse model used in Fig. 6 is a double-fluorescent Cre-dependent reporter, wherein membrane-targeted tandem dimer Tomato (tdTomato) and membrane-targeted enhanced GFP (eGFP) cassettes are driven by a chicken β-actin promoter coupled to a cytomegalovirus (CMV) enhancer [32]. Lox P sites flanking the tdTomato cassette are excised upon introduction of Cre, allowing eGFP to express. This model acts as a reliable indicator of the efficiency of chamber-specific AAV9s in the Cre-lox system, a common system exploited for generation of tissue-specific knockout strains of mice. The organ profile in Fig. 6b indicated a potential complication with the mouse model [32], in which there appeared to be gastrocnemius-specific expression of GFP in all conditions, implying GFP expression in the gastrocnemius is inherent to the mouse and not Cre dependent. Another possibility is the existence of a gastrocnemius-specific protein that is crossreactive with the GFP antibody used. For more thorough testing of organ-specificity between cardiac and gastrocnemius tissue in this mouse model, performing qPCR for GFP transcript would be more informative. In the remaining organs, the hTnnt2 and Nppa promoter showed cardiac-specific and atrialspecific expression of GFP, respectively (Fig. 6). Using the ventricular-specific AAV9, however, modest GFP expression in the liver was found. However, use of miR-122-TS [17] in addition to the Myl2 promoter fragment we designed will mitigate the liver expression and enhance ventricular specificity. References 1. Murphy SL, Kochanek KD, Xu J, Arias E (2021) Mortality in the United States. In: NCHS Data Briefs, No. 427. CDC: CDC 2. Pleger ST, Brinks H, Ritterhoff J et al (2013) Heart failure gene therapy: the path to clinical practice. Circ Res 113(6):792–809 3. Inagaki K, Fuess S, Storm TA et al (2006) Robust systemic transduction with AAV9 vectors in mice: efficient global cardiac gene transfer superior to that of AAV8. Mol Ther 14(1): 45–53. https://doi.org/10.1016/j.ymthe. 2006.03.014 4. Blackwood EA, Hofmann C, Santo Domingo M et al (2019) ATF6 regulates cardiac hypertrophy by transcriptional induction of the mTORC1 activator, Rheb. Circ Res 124(1): 79–93

5. Jessup M, Greenberg B, Mancini D et al (2011) Calcium upregulation by percutaneous Administration of Gene Therapy in cardiac disease (CUPID): a phase 2 trial of intracoronary gene therapy of sarcoplasmic reticulum Ca2+ATPase in patients with advanced heart failure. Circulation 124(3):304–313 6. Pleger ST, Shan C, Ksienzyk J et al (2011) Cardiac AAV9-S100A1 gene therapy rescues post-ischemic heart failure in a preclinical large animal model. Sci Transl Med 3(92): 92ra64 7. Kohlbrenner E, Weber T (2017) Production and characterization of vectors based on the cardiotropic AAV serotype 9. Methods Mol Biol 1521:91–107

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21. Small EM, Krieg PA (2004) Molecular regulation of cardiac chamber-specific gene expression. Trends Cardiovasc Med 14(1):13–18 22. Knowlton KU, Baracchinit E, Ross RS (1991) Co-regulation of the atrial natriuretic factor and cardiac myosin light chain-2 genes during a-adrenergic stimulation of neonatal rat ventricular cells: identification of cis sequences within an embryonic and a constitutive contractile protein gene which mediate inducible expression. J Biol Chem 266(12):7759–7768 23. LaPointe MS, Wug J, Greenberg B et al (1988) Upstream sequences confer atrial-specific expression on the human atrial natriuretic factor gene. J Biol Chem 263(19):9075–9078 24. Morissette MR, Sah VP, Glembotski CC et al (2000) The Rho effector, PKN, regulates ANF gene transcription in cardiomyocytes through a serum response element. Am J Physiol Heart Circ Physiol 278:H1769–H1774 25. Sprenkle AB, Murray SF, Glembotski CC (1995) Involvement of multiple cis elements in basal- and alpha-adrenergic agonist-inducible atrial natriuretic factor transcription. Roles for serum response elements and an SP-1-like element. Circ Res 77(6):1060–1069 26. Houweling AC, van Borren MM, Moorman AF et al (2005) Expression and regulation of the atrial natriuretic factor encoding gene nppa during development and disease. Cardiovasc Res 67:583–593 27. Small EM, Krieg PA (2003) Transgenic analysis of the atrial natriuretic factor (ANF) promoter: Nkx2-5 and GATA-4 binding sites are required for atrial specific expression of ANF. Dev Biol 261(1):116–131 28. Lee KJ, Hickey R, Zhu H et al (1994) Positive regulatory elements (HF-la and HF-lb) and a novel negative regulatory element (HF-3) mediate ventricular muscle-specific expression of myosin light-chain 2-luciferase fusion genes in transgenic mice. Mol Cell Biol 14(2): 1220–1229 29. Lee KJ, Ross RS, Rockman HA et al (1992) Myosin light chain-2 luciferase transgenic mice reveal distinct regulatory programs for cardiac and skeletal muscle-specific expression of a single contractile protein gene. J Biol Chem 267(22):15875–15885 30. Ross RS, Navankasattusas S, Harvey RP et al (1996) An HF-1a/HF-1b/MEF-2 combinatorial element confers cardiac ventricular specificity and establishes an anterior-posterior gradient of expression. Development 122: 1799–1809

AAV9 Cardiac Chamber-Specific Vectors 31. Zhu H, Garcia AV, Ross RS et al (1991) A conserved 28-base-pair element (HF-1) in the rat cardiac myosin light-chain-2 gene confers cardiac-specific and a-adrenergic-inducible expression in cultured neonatal rat myocardial cells. Mol Cell Biol 11(4):2273–2281 32. Muzumdar MD, Tasic B, Miyamichi K et al (2007) A global double-fluorescent Cre reporter mouse. Genesis 45(9):593–605 33. Bilal AS, Blackwood EA, Thuerauf DJ et al (2021) Optimizing adeno-associated virus serotype 9 for studies of cardiac chamberspecific gene regulation. Circulation 143(20): 2025–2027. https://doi.org/10.1161/ CIRCULATIONAHA.120.052437 34. Ni L, Scott L Jr, Campbell HM et al (2019) Atrial-specific gene delivery using an adenoassociated viral vector. Circ Res 124(2): 256–262 35. Bhupathy P, Babu GJ, Periasamy M (2007) Sarcolipin and phospholamban as regulators of cardiac sarcoplasmic reticulum Ca2+ ATPase. J Mol Cell Cardiol 42(5):903–911 36. Anderson DM, Makarewich CA, Anderson KM et al (2016) Widespread control of calcium signaling by a family of SERCA-inhibiting micropeptides. Sci Signal 9:457 37. Yoo J, Kohlbrenner E, Kim O et al (2018) Enhancing atrial-specific gene expression using a calsequestrin cis-regulatory module 4 with a sarcolipin promoter. J Gene Med 20(12):e3060 38. Rockman HA, Ross RS, Harris AN et al (1991) Segregation of atrial-specific and inducible expression of an atrial natriuretic factor transgene in an in vivo murine model of cardiac hypertrophy. Proc Natl Acad Sci 88:8277– 8281

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mice with cardiac hypertrophy and fibrosis. Am J Physiol Heart Circ Physiol 283: H707–H714 41. Minamisawa S, Wang Y, Chen J et al (2003) Atrial chamber-specific expression of sarcolipin is regulated during development and hypertrophic remodeling. J Biol Chem 278(11): 9570–9575 42. Gray SJ, Choi VW, Asokan A et al (2011) Production of recombinant adeno-associated viral vectors and use in in vitro and in vivo administration. Curr Protoc Neurosci. https://doi. org/10.1002/0471142301.ns0417s57 43. Nguyen TNT, Sha S, Hong MS et al (2021) Mechanistic model for production of recombinant adeno-associated virus via triple transfection of HEK293 cells. Mol Ther Methods Clin Dev 21:642–655. https://doi.org/10.1016/j. omtm.2021.04.006 44. Blackwood EA, Bilal AS, Azizi K et al (2020) Simultaneous isolation and culture of atrial myocytes, ventricular myocytes, and non-myocytes from an adult mouse heart. J Vis Exp. https://doi.org/10.3791/61224 45. Kuwahara K (2013) Role of NRSF/REST in the regulation of cardiac gene expression and function. Circ J 77(11):2682–2686 46. Jin JK, Blackwood EA, Azizi K et al (2017) ATF6 decreases myocardial ischemia/reperfusion damage and links ER stress and oxidative stress signaling pathways in the heart. Circ Res 120(5):862–875

Chapter 9 Generation of Atrial-Specific Construct Using Sarcolipin Promoter-Associated CRM4 Enhancer Dongtak Jeong Abstract Cardiac gene therapy has been hampered by off-target expression of gene of interest irrespective of variety of delivery methods. To overcome this issue, cardiac-specific promoters provide target tissue specificity, although expression is often debilitated compared to that of ubiquitous promoters. We have previously shown that sarcolipin promoter with an enhancer calsequestrin cis-regulatory module 4 (CRM4) combination has an improved atrial specificity. Moreover, it showed a minimal extra-atrial expression, which is a significant advantage for AAV9-mediated cardiac gene therapy. Therefore, it can be a useful tool to study and treat atrial-specific diseases such as atrial fibrillation. In this chapter, we introduce practical and simple methodology for atrial-specific gene therapy using sarcolipin promoter with an enhancer CRM4. Key words AAV9, Atrium, Cis-acting regulatory module, CRM4, Gene therapy, Sarcolipin

1

Introduction Atrial diseases are serious menaces to human health. For example, atrial fibrillation (AF) is one of the major cardiac arrhythmias with high mortality rate [1]. However, there are unmet needs for the current approaches to treat AF although cardiac gene therapies have emerged as a promising method [2, 3]. Currently, adenoassociated virus serotype 9 (AAV9) is the most promising delivery vehicle for transgene expression and the most widely used in cardiac preclinical trials due to its tropism for the heart [4–9]. Although the AAV9 vector in conjunction with the ubiquitous cytomegalovirus (CMV) promoter has shown cardiac expression, a major limitation is the expression of transgenes in other organs such as the liver, lung, thymus, brain, and kidney [5, 10]. Concerns for limited tissue specificity have been previously handled through transcriptional targeting [11, 12]. Since the virus regulates transgene production through its promoter, promoter alterations have been used to minimize off-target expressions. To overcome off-target

Kiyotake Ishikawa (ed.), Cardiac Gene Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2573, https://doi.org/10.1007/978-1-0716-2707-5_9, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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distribution, cardiac-specific promoters have been applied in previous literatures including α-myosin heavy chain, myosin light chain, enhanced myosin light chain, and cardiac troponin T promoters [13–16]. More recently, several promoters have been observed to regulate cardiac chamber specific gene expression. Among these are the atrial myosin light chain-2a, slow myosin heavy chain-3, atrial natriuretic factor, and sarcolipin (SLN) for the atrium and the myosin light chain-2 V for the ventricle [11, 17–20]. The uniqueness of these promoters has been exploited in other methods besides improving specificity such as allowing for cardiomyocytes generated from human-induced pluripotent stem cells to be separated by subtype-specific promoter-driven action potentials [19, 20]. However, the application of these promoters to drive chamber-specific transgene expression has been limited due to their compromised efficiency. To apply these promoters for use in gene therapy, it is essential to improve transduction efficiency of these promoters. To overcome this, we hypothesized that an improved atrial specificity can be achieved when two enhancer and promoter sequences, which are previously reported to show an atrial specificity, are combined with cardiotropic AAV9. Based on this hypothesis, we have taken advantage of SLN’s atrial specificity. The SLN promoter was chosen because of previous observation that the SLN protein is characteristically expressed in the atrial chamber of the heart. Furthermore, it has been observed to change its expression dependent on disease states. For example, sarcolipin has been found to be upregulated in rodent models of congenital heart disease and patients with preserved left ventricular ejection fraction and chronic isolated mitral regurgitation [21–23]. In contrast, sarcolipin was found to be downregulated in patients with chronic atrial fibrillation [24, 25]. In Duchenne muscular dystrophy mouse model, reducing sarcolipin expression was found to mitigate associated cardiomyopathy [26]. Alone, the SLN promoter activity was unsubstantial compared to that of ubiquitous promoters. Thus the addition of calsequestrin 2 cis-regulatory module 4 (CRM4), a cardiomyocyte-specific enhancer, which showed superior activity in the heart, was used to enhance transgene expression of SLN promoter within the heart [11, 27]. In addition to cardiac tissue, high SLN levels are found in the skeletal muscle and diaphragm [26]. The addition of CRM4 was thus hypothesized to improve cardiac specificity. When used in conjunction with the cardiotropic AAV9, we hypothesized even higher selectivity in the heart. As a result, we previously showed that the CRM4. SLN construct indeed induced atrial specific expression with minimized off-target effects in in vivo application.

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Therefore, the use of this construct can offer a noninvasive and technically simple approach for atrial-specific gene therapy. The CRM4 and SLN combination caused robust and highly specific atrial activity and is a promising method of targeting transcriptional mechanisms to improve atrial transgene transduction. Here we share practical and simple methodology for atrialspecific gene expression using SLN promoter and an enhancer CRM4 in this chapter.

2

Materials

2.1 Generation of SLN Promoter/CRMAssociated AAV Construct (See Note 1) 2.1.1 pTR.SLN.Luc./pTR. SLN.EGFP

1. The primary core promoter sequence by removal of polyadenylation sequence from a 1253 base pair human SLN promoter purchased from GeneCopoeia (HPRM12771, MD). 2. pTR-CMV-Luciferase and pTR-CMV-EGFP vectors (see Fig. 1a, b), 3. Primer pairs (forward and reverse) containing the shortened SLN promoter (1029 bps, see Fig. 2a) with overhangs compatible with the restriction enzymes. SLN promoters: Fwd: 50 CCTAGATCTGAATTCGGTACCTGAGGA ATGGGA-30 Rev: 50 -CGGTGTGCCTCTCATACCGGTTCTGCCTTTCT CATT-30 4. Restriction enzymes: KpnI and AgeI. 5. T4 DNA ligase. 6. Competent cells. 7. Super Optimal Broth (SOB) medium. 8. Water bath. 9. Antibiotic agar plate. 10. Shaking incubator.

2.1.2 Luc

pTR.CRM4.SLN.

1. First primer pairs for the 1029 bp SLN promoter from the pTR.SLN.Luciferase. Fwd: 50 -GAGCAAACACAATTGCTAGGG-30 Rev: 50 -AAGAGGATCAAAGACACACC-30 2. Second primer pairs with overhangs. Fwd: 50 -GATACAGTCTGTCCGAACGCGTGGAGCAAACA CAATTGCTAGGG-30 Rev: 50 - ACAGTACCGGAATGCCAAAGAGGATCAAAGA CACACC-30

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Fig. 1 Generation of AAV constructs with various promoters. (a) Schematic depiction of CMV and SLN promoters with luciferase or EGFP as a reporter gene. (b) Cloning scheme for CRM4-SLN-luciferase construct. (c) Cloning scheme for CRM4-SLN-EGFP construct

3. The pTR.CRM4.Luciferase plasmid from Dr. Thierry VandenDriessche (University of Brussels, Belgium) (see Fig. 2b). 4. Other materials are necessary as Subheading 2.1.1, item 5–10. 5. Seamless Assembly kit. 2.1.3 pTR.CRM4.SLN. EGFP

1. The pTR.CRM4.SLN.Luc. construct in Subheading 2.1.2. 2. CMV.EGFP shuttle vector obtained from Dr. Jude Samulski (University of North Carolina, Chapel Hill, USA). 3. Restriction enzymes: NcoI and Eco521 to remove the luciferase gene. 4. Other materials are necessary as Subheading 2.1.1, item 5–10. 5. Seamless Assembly kit.

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Fig. 2 The full sequence of sarcolipin promoter and CRM4 enhancer. (a) The full 1289 bp sarcolipin promoter sequence. (b) The full 207 bp CRM4 sequence 2.2

AAV Production

1. HEK293T cells (ATCC, Manassas, VA). 2. Growth medium: DMEM, 10% fetal bovine serum (FBS), 1% penicillin–streptomycin. 3. DMEM without antibiotics for the transfection reaction. 4. Transfection reagent: Polyethylenimine (PEI) linear MW 25,000, PBS. 5. DNase digestion buffer: 10 mM Tris–HCl, pH 7.5, 10 mM MgCl2, 50 U/mL DNase I. 6. 15% iodixanol solution: 4.5 mL of 60% iodixanol, 13.5 mL of 1  Tagment DNA Buffer (TD) buffer. 7. 25% iodixanol solution: 5 mL of 60% iodixanol, 7 mL of 1 TD buffer, 30 μL of phenol red. 8. 40% iodixanol solution: 6.7 mL of 60% iodixanol, 3.3 mL of 1 TD buffer. 9. 60% iodixanol solution: 10 mL of 60% iodixanol, 45 μL of phenol red. 10. Quick-Seal ultracentrifuge tubes and Plug (Beckman, USA). 11. Sterilized TD buffer: 0.38 g/L KCL, 0.1 g/L Na2HPO4, 8.0 g/L NaCl, 3.0 g/L Tris, pH 7.4. 12. Ultracentrifuge and fixed angle rotor.

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Table 1 qRT-PCR primers for AAV titration Primers

Sequences

CMV

F R

TCAATTACGGGGTCATTAGTTC ACTAATACGTAGATGTACTGCC

SV-40

F R

AGCAATAGCATCACAAATTTCACAA CCAGACATGATAAGATACATTGA

EGFP

F R

AAGTTCATCTGCACCACCG TCCTTGAAGAAGATGGTGCG

ITR

F

GGAACCCCTAGTGATGGAGTT

R

CGGCCTCAGTGAGCGA

13. qRT-PCR reagents including buffers, enzymes, primers, and detection methods (i.e., SYBR green or probe-based detection). 14. qRT-PCR primer pairs for AAV titration in Table 1 (SV40, CMV, ITR primer pairs, etc.) 15. Standard AAV vector. 16. qRT-PCR machine. 17. Vivaspin 20 Centrifugal concentrator 50 K MWCO. 2.3 In Vitro Gene Transfer

1. HL-1: Cardiac muscle cell line. 2. HL-1 cell culture medium: Claycomb medium 500 mL, 10% FBS, penicillin (100 U/mL), streptomycin (100 U/mL), glutamine (2 mM), and noradrenaline (0.1 mM). 3. PEI reagent. 4. Milli-Q H2O. 5. Hydrogen chloride (HCl).

2.4 In Vivo Gene Transfer

1. A single mouse restrainer.

2.5

1. Pierce Firefly Luciferase Glow Assay Kit (Thermo Fisher, USA; Kit contents: 1  Firefly Glow Assay Buffer; 100  D-Luciferin, 2  Cell Lysis Buffer; Luciferase assay working reagent: Mixture of Firefly Glow Assay Buffer and D-Luciferin with 100 to 1 ratio).

Luciferase Assay

2. 100 uL viral solution (1  1011 vg/mouse) of the pertaining vector.

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2. BCA assay reagents for protein quantification (Thermo Fisher, USA; Kit contents: 1  BCA Reagent A, 20  BCA Reagent B, BCA assay working reagent: Mixture of reagent A and B with 20 to 1 ratio; see 3.9 for detail protocol). 3. Luminometer. 2.6 Fluorescence Imaging

1. 4% paraformaldehyde. 2. Ethanol. 3. Xylene. 4. OCT compound. 5. Mounting medium with DAPI. 6. Confocal microscope. 7. 20% sucrose.

2.7 Western Blot Analysis

1. 10% SDS-PAGE gel (1.5 mm thick). 2. Nitrocellulose membrane (pore size 0.22 um). 3. Western running and transfer kits. 4. RIPA buffer for cell lysis. 5. Antibodies against EGFP and tubulin. 6. Secondary antibodies. 7. Western blot imager.

3

Methods

3.1 Generation of pTR.SLN.Luc Plasmid

1. Perform the PCR reaction using specific primers in Subheading 2.1.1, item 3. Detail condition is shown in Table 2. 2. Execute restriction enzyme digestion on SLN PCR product obtained from the above as shown in Table 3 (see Note 2). 3. Incubate in 37  C water bath for at least 4 hrs (O/N reaction is recommended). 4. Purify the enzyme-digested PCR product with PCR Purification Kit after 1% agarose gel running (see Note 3). 5. pTR.CMV.Luciferase plasmids are also digested by Kpn I and Age I restriction enzymes, and confirm the removal of the CMV promoter with agarose gel running. 6. Purify the digested pTR.Luciferase plasmid with the gel purification kit. 7. Perform the DNA ligation process shown in Table 4 using the SLN promoter and backbone CMV promoter-removed pTR. Luciferase plasmid (see Note 4).

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Table 2 PCR conditions for the SLN promoter Reagent

Amount (mL)

SLN promoter containing plasmid DNA from GeneCopoeia (100 ng/mL)

1

Taq polymerase

1

PCR buffer with Mgcl2 (10)

5

dNTP mix (10)

5

Forward primer (100 nM/mL)

1

Reverse primer (100 nM/mL)

1

D.W

36

Total

50

Table 3 Restriction enzyme digestion on the SLN promoter Reagent

Amount (mL)

PCR product (SLN promoter; 1253 bps)

16

KpnI (20 units/mL)

1

AgeI (20 units/mL)

1

Reaction buffer (10)

2

Total

20

Table 4 DNA ligation condition Reagent

Amount

T4 ligase buffer (10)

2 mL

Vector DNA (pTR.Luciferase)

50 ng

Insert DNA (SLN promoter)

37.5 ng

T4 ligase

2 mL

Nuclease free water

Up to 20 mL

Total

20 mL 

Incubate O/N at 16 C or 10 min at RT Sequencing the final ligated plasmid is highly necessary

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3.2 Generation of pTR.SLN.EGFP Plasmid

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1. Digest pTR-CMV-EGFP vector with Kpn I and Age I to remove the CMV promoter sequence. 2. The PCR-amplified and enzyme-digested SLN promoter used to construct pTR.SLN.Luc vector is used here again as Subheading 3.1, step 1. 3. Ligate the promoter and vector to create the pTR.SLN.EGFP vector. 4. Sequence and confirm the final ligated plasmid using a sequencing primer.

3.3 Generation of pTR.CRM4.SLN.Luc Plasmid

1. SLN promoter is amplified by PCR from the pTR.SLN.Luciferase with primers shown above in Materials Subheading 2.1.2, item 1. 2. A second PCR amplification using specific primers (Subheading 2.1.2, item 2) is done to add Gibson assembly overhangs (see Note 5). 3. Digest the CRM4.Luciferase (Fig. 1b) with MluI and HindIII to insert the SLN promoter between the CRM4 and luciferase. 4. Execute the PCR for SLN promoter with primer pairs in Subheading 2.1.2, and digest the PCR product by MluI and HindIII as shown in Subheading 3.1, step 1. 5. Perform the seamless cloning as shown in Table 5.

3.4 Generation of pTR.CRM4.SLN.EGFP Plasmid

1. Digest pTR.CRM4.SLN.Luciferase plasmid. with NcoI and Eco521 to remove the luciferase gene. 2. Digest pTR.CMV.EGFP plasmid with NcoI and Eco521 to remove EGFP as shown in Subheading 3.1, step 1. 3. Replace the luciferase gene with the EGFP gene obtained from plasmid pTR.CMV.EGFP.

3.5 Generation of AAV Viral Vector

1. Produce the recombinant AAVs from HEK293-T cells by the transfection of the cloned pTR plasmids produced by the Methods in Subheading 3.1, step 1–4. 2. Purify the AAV particles using the iodixanol gradient method followed by ultracentrifugation as follows. 3. Overlay each solution into a Quick-Seal tube in the order below using a 10 mL syringe and an 18 g needle, taking care to avoid bubbles. Detailed procedure and example picture are in Table 6. 4. Centrifuge at 350,000 g for 90 min in a T70i rotor at 10  C. 5. Carefully take the Quick-Seal tubes out of the rotor, and place them in a stable rack. (Make sure not to disturb the gradient). 6. Puncture the Quick-Seal tube slightly below the 60–40% interface with an 18 g needle attached to a 10 mL syringe.

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Table 5 Seamless cloning conditions for the pTR.CRM4.SLN constructs Reagent

Amount (mL)

Reaction buffer (5)

2

Linearized vector DNA (pTR.CRM4.Luciferase; 100 ng/mL)

2

Insert DNA (SLN promoter; 100 ng/mL)

4

10 enzyme mix

2

Nuclease-free water

8

Total

20 ml

Mix the reaction components by gently tapping the sides of the centrifuge tube, and incubate at room temperature for 30 minutes After 30 minutes of incubation, place the reaction mix on ice, and immediately proceed to the transformation step

Table 6 Iodixanol gradient for the AAV purification Reagent

Amount (mL)

60% iodixanol

1.5

40% iodixanol

2.5

25% iodixanol

3.6

15% iodixanol

4.8

7. The bevel of the needle should be up, facing the 40% step. 8. Collect up to 5 mL per tube, taking care to avoid collecting the proteinaceous material at the 40–25% interface. 9. The particles are concentrated by exchanging iodixanol for Lactate Ringer’s solution by multiple dilution and concentration steps using a Vivaspin 20 centrifugal concentrator 50 K MWCO. 10. The AAV titer is determined by quantitative real-time PCR and SDS-PAGE. 3.6 Preparation of PEI Stock Solution

1. Pour ~200 mL of Milli-Q H2O into a 500 mL glass beaker. 2. Add 250 mg of PEI to the beaker with stirring. 3. Add concentrated HCl dropwise to the solution to pH ¼ 1.9. 4. Stir at least 24 h until the PEI is completely dissolved. Maintain the pH < 2.0 throughout (dissolution appears to occur in 30–40 min but is not; maintain stirring for 24 h). Approximately 400 uL of 12 M HCl will be required for full PEI dissolution. There may still be some small fiber particles that will not dissolve.

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5. Neutralize by adding concentrated NaOH dropwise to the solution pH 7.0. 6. Pour the solution into a 500 mL glass cylinder. Adjust the final volume to 250 mL with Milli-Q H2O. 7. Filter-sterilize the solution through one 0.22-μm membrane. 8. Store aliquots of the desired volume at 80  C. 3.7 Validation of In Vitro Gene Transfer (See Fig. 3a; See Notes 6 and 7)

1. Prepare the HL-1 cells 1 day before transfection (2  105 cells per 60 mm dish). 2. Transfect cells with CMV.VLP, CMV.Luc or EGFP, SLN.Luc or EGFP, and CRM4.SLN.Luc or EGFP vectors using PEI reagent. 3. Forty-eight hours later, harvest the cells, and obtain protein lysates by RIPA buffer containing protein/phosphatase inhibitor cocktail or Pierce Firefly Luciferase Glow Assay Kit for western blot analysis and luciferase assay, respectively.

3.8 In Vivo Sample Preparation for Luciferase Assay

1. Inject AAV9 CMV.Luc, SLN.Luc, and CRM4.SLN.Luc into C57/B6 mice intravenously via tail vein (1  1011 vg/mouse for each viral vector). 2. Three weeks post injection, harvest the atrium, ventricle, skeletal muscle, diaphragm, brain, lung, liver, and kidney from each mouse. 3. Finely grind tissue samples and lyse 100 ug of samples from each tissue with 100 uL of 1 Cell Lysis Buffer in general luciferase assay kit. 4. Quantify protein amount in each sample with BCA assay.

3.9 BCA Assay for Protein Quantification

1. Pipette 10 μL of each standard and unknown sample replicate into an appropriately labeled test tube. 2. Add 1.0 mL of the BCA assay working reagent to each tube and mix well. 3. Cover and incubate tubes at selected temperature and time: Standard incubation time, 37  C for 30 minutes (working range ¼ 20–2000 μg/mL). 4. Cool all tubes to RT. 5. With the spectrophotometer set to 562 nm, zero the instrument on a cuvette filled only with water and then measure the absorbance of all the samples within 10 minutes. 6. Subtract the average 562 nm absorbance measurement of the Blank standard replicates from the 562 nm absorbance measurement of all other individual standard and unknown sample replicates.

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Fig. 3 Luciferase assay confirmed CRM4 enhanced atrial specificity and minimized off-target expression. (a) Luciferase assay was conducted 24 hours post incubation in HL1 cells. Gene expression of control and luciferase containing CMV, SLN, and CRM4.SLN vectors were measured. (b) In vivo gene expression was observed 3 weeks post tail vein delivery of control and luciferase containing CMV, SLN, and CRM4.SLN vectors. Luciferase assay was conducted to determine bio-distribution. Luciferase activity was normalized to background values. Statistical significance in (a) and (b) was measured with two-tailed Students T-test, and significant differences are demonstrated by a single asterisk (*), which indicates p < 0.05, a double asterisk (**), which indicates p < 0.01, or a triple asterisk (***), which indicates p < 0.005. Bar graphs represent mean  SD (n ¼ 4)

7. Prepare a standard curve by protein the average Blankcorrected 562 nm measurement for each BSA standard vs. its concentration in μg/mL. 8. Use the standard curve to determine the protein concentration of each unknown sample.

Generation of Atrial-Specific Construct

3.10 Luciferase Assay (See Fig. 3b)

127

1. Program the luminometer. 2. Add 10 μg/well of cell lysate to a white or black, opaque 96-well plate. 3. Add 50 μL of Luciferase assay working reagent to each well. 4. Wait 10 minutes for signal stabilization and detect the light output. 5. Measure the value of luminescence.

3.11 Biodistribution by Western Blotting (See Fig. 4)

1. Inject AAV9 CMV.EGFP, SLN. EGFP, and CRM4.SLN. EGFP into C57/B6 mice intravenously via tail vein (5  1010 and 1  1011 vg/mouse for each viral vector). 2. Harvest the atrium, ventricle, liver, and kidney from each mouse at 3 weeks post injection. 3. Lyse tissue homogenates using RIPA buffer with protease inhibitor cocktail. 4. Quantify protein amount using general BCA Protein Assay Kit as shown in Subheading 3.9. 5. Load and run proteins using 10% SDS-PAGE gels followed by transfer to nitrocellulose membrane. 6. Treat antibodies raised against EGFP and tubulin. 7. Image the protein bands. 8. Quantify the result of western blots with Image J software.

3.12 Fluorescent Imaging (See Fig. 5)

1. Inject AAV9 CMV.EGFP, SLN. EGFP, and CRM4.SLN. EGFP into C57/B6 mice intravenously via tail vein (1  1011 vg/mouse for each viral vector). 2. Three weeks post injection, harvest mouse heart tissues in a sagittal plane and fix in 4% paraformaldehyde for 24 hours, and move to 20% sucrose for cryoprotection for 24 hours before cryopreserving in OCT compound at 80  C. 3. Section tissues into 6-μm-thick slices. 4. Dehydrate the slides in 100% ethanol for 10 minutes, and dip in two changes of xylene. 5. Analyze the sectioned slides using a confocal microscope and image analyzer.

4

Notes 1. Tips for the AAV Vector Cloning It has been frequently observed that inverted terminal repeat (ITR) can be easily lost during the cloning procedure due to the instability of this region [28, 29]. Therefore, ITR

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Fig. 4 Atrial specificity and expression by CRM4 were validated by an alternative reporter, EGFP. In vivo gene expression was observed 3 weeks post tail vein delivery of control and EGFP containing CMV, SLN, and CRM4. SLN vectors. Western blot analysis was conducted to determine bio-distribution with antibody EGFP and normalized with tubulin

region should be checked by XmaI or SmaI restriction enzyme in every single step during the entire cloning process. 2. Definition of Unit One unit is defined as the amount of enzyme required to digest 1 μg of pXba DNA in 1 hour at 37  C in a total reaction volume of 50 μL. 3. Confirmation of PCR Product It can be acceptable even without agarose gel running procedure only if the PCR product was confirmed with a single band and the predicted size when it was previously applied. Otherwise, gel running procedure is essential to confirm

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Fig. 5 Visualization of biodistribution. In vivo gene expression in atrium and ventricle of control and EGFP containing CMV, SLN, and CRM4.SLN vectors were observed by fluorescent imaging using confocal microscope at 200 magnification. Bright field, DAPI, EGFP, and merged images were taken under same exposure and normalization settings. (The Figure was adopted from Yoo et al. [11])

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whether the PCR product has a correct identification or unexpected mutation, which possibly generates unusual restriction enzyme sites, during PCR procedure. 4. Tips for DNA Ligation T4 ligase is one of the most unstable enzymes; thus an aliquot of newly purchased T4 ligase and ligase buffer is highly recommended and stored at 20  C for long-term storage. 5. Tips for Seamless Cloning It is essential to design proper primer pairs for vector and insert (PCR product) in Seamless Cloning kit. Therefore, it is highly recommended to use GeneArt® Primer and Construct Design Tool in Thermo Fisher website (https://www. thermofisher.com/order/oligoDesigner). For maximum cloning efficiency, use 2:1 insert:vector molar ratio. Determine the concentration of your DNA insert solutions by OD260 or fluorescence, and use the concentrations to calculate the volume required to achieve the 2:1 molar ratio of insert to vector. In the case of very small inserts (200 bp), we recommend that you use at least 20 ng of insert DNA in the seamless cloning and assembly reaction. 6. Tips for the HL-1 Cell Culture It has been very difficult to generate cell lines from cardiac muscle; therefore, more studies are done in primary cardiomyocytes. However, there are two cell lines, HL-1 and H9c2, that are used occasionally. Particularly, HL-1 is a cardiac muscle cell line, derived from AT-1 mouse artial cardiomyocyte tumor lineage. These cells have the morphology of differentiated cardiomyocytes, maintaining the biochemical and electrophysiological properties and the ability to contract, while they can be passaged several times [30, 31]. However, HL-1 cells are very sensitive and notorious for the maintenance in differentiated state [31]. Therefore, they should be cultured in very specific medium and a couple of supplements such as 4 mM lglutamine and 10 μM norepinephrine. In addition, they also need to be plated on extracellular matrix such as fibronectin (0.5%) and gelatin (0.02%) [32, 33]. 7. Tips for the Transfection Into HL-1/H9c2 H9c2 and HL-1 cells are notorious for their transfection efficiency. For this reason, PEI solution is a superior alternative as a transfection reagent. Protocol to produce PEI transfection reagent is described in Subheading 3.6. However, PEI is also toxic for the cell if overused (recommended dose: 30ul for 100 mm culture dish (1E7 cells).

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Acknowledgments This work is supported by grants from the National Research Foundation of Korea (2021R1A2C1008058 and 2021R1A4A5032463) funded by the Korean Government (MSIP) and Hanyang University (HY-202000000002749). References 1. Carlsson L, Duker G, Jacobson I (2010) New pharmacological targets and treatments for atrial fibrillation. Trends Pharmacol Sci 31(8): 364–371 2. Bongianino R, Priori SG (2015) Gene therapy to treat cardiac arrhythmias. Nat Rev Cardiol 12(9):531–546 3. Bikou O, Thomas D, Trappe K, Lugenbiel P, Kelemen K, Koch M, Soucek R, Voss F, Becker R, Katus HA (2011) Connexin 43 gene therapy prevents persistent atrial fibrillation in a porcine model. Cardiovasc Res 92(2):218–225 4. Inagaki K, Fuess S, Storm TA, Gibson GA, Mctiernan CF, Kay MA, Nakai H (2006) Robust systemic transduction with AAV9 vectors in mice: efficient global cardiac gene transfer superior to that of AAV8. Mol Ther 14(1): 45–53 5. Zincarelli C, Soltys S, Rengo G, Rabinowitz JE (2008) Analysis of AAV serotypes 1–9 mediated gene expression and tropism in mice after systemic injection. Mol Ther 16(6): 1073–1080 6. Del Monte F, Williams E, Lebeche D, Schmidt U, Rosenzweig A, Gwathmey JK, Lewandowski ED, Hajjar RJ (2001) Improvement in survival and cardiac metabolism after gene transfer of sarcoplasmic reticulum Ca2+ATPase in a rat model of heart failure. Circulation 104(12):1424–1429 7. Jeong D, Lee M-A, Li Y, Yang DK, Kho C, Oh JG, Hong G, Lee A, Song MH, LaRocca TJ (2016) Matricellular protein CCN5 reverses established cardiac fibrosis. J Am Coll Cardiol 67(13):1556–1568 8. Miyamoto MI, Del Monte F, Schmidt U, DiSalvo TS, Kang ZB, Matsui T, Guerrero JL, Gwathmey JK, Rosenzweig A, Hajjar RJ (2000) Adenoviral gene transfer of SERCA2a improves left-ventricular function in aorticbanded rats in transition to heart failure. Proc Natl Acad Sci 97(2):793–798 ˜ oz A, 9. Wahlquist C, Jeong D, Rojas-Mun Kho C, Lee A, Mitsuyama S, van Mil A, Park WJ, Sluijter JP, Doevendans PA (2014)

Inhibition of miR-25 improves cardiac contractility in the failing heart. Nature 508(7497): 531–535 10. Mu¨ller O, Schinkel S, Kleinschmidt J, Katus H, Bekeredjian R (2008) Augmentation of AAV-mediated cardiac gene transfer after systemic administration in adult rats. Gene Ther 15(23):1558–1565 11. Yoo J, Kohlbrenner E, Kim O, Hajjar RJ, Jeong D (2018) Enhancing atrial-specific gene expression using a calsequestrin cis-regulatory module 4 with a sarcolipin promoter. J Gene Med 20(12):e3060 12. Nettelbeck DM, Je´roˆme V, Mu¨ller R (1998) A strategy for enhancing the transcriptional activity of weak cell type-specific promoters. Gene Ther 5(12):1656–1664 13. Prasad KR, Xu Y, Yang Z, Acton ST, French BA (2011) Robust cardiomyocyte-specific gene expression following systemic injection of AAV: in vivo gene delivery follows a Poisson distribution. Gene Ther 18(1):43–52 14. Aikawa R, Huggins GS, Snyder RO (2002) Cardiomyocyte-specific gene expression following recombinant adeno-associated viral vector transduction. J Biol Chem 277(21): 18979–18985 15. Werfel S, Jungmann A, Lehmann L, Ksienzyk J, Bekeredjian R, Kaya Z, Leuchs B, Nordheim A, Backs J, Engelhardt S (2014) Rapid and highly efficient inducible cardiac gene knockout in adult mice using AAV-mediated expression of Cre recombinase. Cardiovasc Res 104(1):15–23 16. Franz W-M, Rothmann T, Frey N, Katus HA (1997) Analysis of tissue-specific gene delivery by recombinant adenoviruses containing cardiac-specific promoters. Cardiovasc Res 35(3):560–566 17. Ni L, Scott L Jr, Campbell HM, Pan X, Alsina KM, Reynolds J, Philippen LE, Hulsurkar M, Lagor WR, Li N (2019) Atrial-specific gene delivery using an adeno-associated viral vector. Circ Res 124(2):256–262

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18. Small EM, Krieg PA (2004) Molecular regulation of cardiac chamber-specific gene expression. Trends Cardiovasc Med 14(1):13–18 19. Chen Z, Xian W, Bellin M, Dorn T, Tian Q, Goedel A, Dreizehnter L, Schneider CM, Ward-van Oostwaard D, Ng JKM (2017) Subtype-specific promoter-driven action potential imaging for precise disease modelling and drug testing in hiPSC-derived cardiomyocytes. Eur Heart J 38(4):292–301 20. Biendarra-Tiegs SM, Secreto FJ, Nelson TJ (2020) Addressing variability and heterogeneity of induced pluripotent stem cell-derived cardiomyocytes. Adv Exp Med Biol 1212(6): 1–29 21. Babu GJ, Bhupathy P, Petrashevskaya NN, Wang H, Raman S, Wheeler D, Jagatheesan G, Wieczorek D, Schwartz A, Janssen PM (2006) Targeted overexpression of sarcolipin in the mouse heart decreases sarcoplasmic reticulum calcium transport and cardiac contractility. J Biol Chem 281(7): 3972–3979 22. Zheng J, Yancey DM, Ahmed MI, Wei C-C, Powell PC, Shanmugam M, Gupta H, Lloyd SG, McGiffin DC, Schiros CG (2014) Increased sarcolipin expression and adrenergic drive in humans with preserved left ventricular ejection fraction and chronic isolated mitral regurgitation. Circ Heart Fail 7(1):194–202 23. Pashmforoush M, Lu JT, Chen H, St Amand T, Kondo R, Pradervand S, Evans SM, Clark B, Feramisco JR, Giles W (2004) Nkx2-5 pathways and congenital heart disease: loss of ventricular myocyte lineage specification leads to progressive cardiomyopathy and complete heart block. Cell 117(3):373–386 24. Uemura N, Ohkusa T, Hamano K, Nakagome M, Hori H, Shimizu M, Matsuzaki M, Mochizuki S, Minamisawa S, Ishikawa Y (2004) Down-regulation of sarcolipin mRNA expression in chronic atrial fibrillation. Eur J Clin Investig 34(11):723–730 25. Bhupathy P, Babu GJ, Periasamy M (2007) Sarcolipin and phospholamban as regulators of cardiac sarcoplasmic reticulum Ca2+ ATPase. J Mol Cell Cardiol 42(5):903–911

26. Voit A, Patel V, Pachon R, Shah V, Bakhutma M, Kohlbrenner E, McArdle JJ, Dell’Italia LJ, Mendell JR, Xie L-H (2017) Reducing sarcolipin expression mitigates Duchenne muscular dystrophy and associated cardiomyopathy in mice. Nat Commun 8(1): 1–14 27. Rincon MY, Sarcar S, Danso-Abeam D, Keyaerts M, Matrai J, Samara-Kuko E, Acosta-Sanchez A, Athanasopoulos T, Dickson G, Lahoutte T (2015) Genome-wide computational analysis reveals cardiomyocytespecific transcriptional cis-regulatory motifs that enable efficient cardiac gene therapy. Mol Ther 23(1):43–52 28. Wilmott P, Lisowski L, Alexander IE, Logan GJ (2019) A user’s guide to the inverted terminal repeats of adeno-associated virus. Hum Gene Ther Methods 30(6):206–213 29. Xie J, Mao Q, Tai PW, He R, Ai J, Su Q, Zhu Y, Ma H, Li J, Gong S (2017) Short DNA hairpins compromise recombinant adenoassociated virus genome homogeneity. Mol Ther 25(6):1363–1374 30. White SM, Constantin PE, Claycomb WC (2004) Cardiac physiology at the cellular level: use of cultured HL-1 cardiomyocytes for studies of cardiac muscle cell structure and function. Am J Phys Heart Circ Phys 286(3): H823–H829 31. Claycomb WC, Lanson NA, Stallworth BS, Egeland DB, Delcarpio JB, Bahinski A, Izzo NJ (1998) HL-1 cells: a cardiac muscle cell line that contracts and retains phenotypic characteristics of the adult cardiomyocyte. Proc Natl Acad Sci 95(6):2979–2984 32. Bajaj G, Sharma RK (2006) TNF-α-mediated cardiomyocyte apoptosis involves caspase-12 and calpain. Biochem Biophys Res Commun 345(4):1558–1564 33. Burt R, Graves BM, Gao M, Li C, Williams DL, Fregoso SP, Hoover DB, Li Y, Wright GL, Wondergem R (2013) 9-Phenanthrol and flufenamic acid inhibit calcium oscillations in HL-1 mouse cardiomyocytes. Cell Calcium 54(3):193–201

Part IV Cardiac Gene Delivery Methods

Chapter 10 Cardiac Targeted Adeno-Associated Virus Injection in Rats Michael G. Katz , Yoav Hadas, Adam S. Vincek, Nataly Shtraizent, Eric Schadt, and Efrat Eliyahu Abstract Gene therapy is a promising approach in the treatment of cardiovascular diseases. The vectors available for cardiovascular gene therapy have significantly improved over time. Cardiac tropism is a primary characteristic of an ideal vector along with a long-term expression profile and a minimal risk of cellular immune response. Preclinical and clinical studies have demonstrated that adeno-associated viral (AAV) vectors are one of the most attractive vehicles for gene transfer. AAV has gained great popularity in the last years because of its biological properties and advantages over other viral vector systems. In this chapter we will describe methods for intracardiac delivery of AAV vector in rats. Key words Cardiac gene therapy, Adeno-associated viral vector, Intramyocardial administration, Intracoronary administration, Cardiovascular diseases

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Introduction Over the past decade, gene therapy technology has significantly advanced, and viral cardiac gene delivery tools have progressed from application in animal research to FDA-approved clinical investigation in humans [1, 2]. Gene therapy is now considered an attractive strategy for combating cellular and molecular abnormalities underlying the pathogenesis of myocardial ischemia [3, 4]. Several recombinant viral vectors and nonviral gene delivery systems were tested for various cardiac applications [5]. Successful use of cardiac gene therapy requires that the DNA encoding for the gene of interest is efficiently delivered into cardiomyocytes, and in most cases, long-lasting gene expression is desired. Today, adeno-associated viruses (AAV)-based vectors are undoubtedly among the most promising DNA delivery vehicles. Advantages of AAV as a cardiac gene delivery vector include (a) efficient transduction into

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cardiomyocytes, (b) eliciting modest cellular immune response, and (c) triggering persistent expression of therapeutic gene of interest, even in the absence of genome integration. AAV is a small non-enveloped virus that can only replicate in the presence of a helper virus such as adenovirus or herpesvirus. The AAV genome consists of a single-stranded DNA that encodes only two genes, the Rep gene, which encodes proteins essential for DNA replication and packaging, and the Cap gene, which encodes the capsid proteins and, in an alternative reading frame, the assembly activating protein. The approximately 4.7-kb-long AAV genome is flanked by two inverted terminal repeats that are the only cis-elements required for the production of recombinant AAVs (rAAVs). An AAV naturally infects humans, usually as infants, and is not associated with any known disease or illness. AAV vectors gained significant attention for gene therapy due to important beneficial properties for therapeutic application in humans, showing advantages over other vectors. AAV-mediated gene delivery allows high transduction efficiency and prolonged transgene expression in a broad range of tissue and organs, including the heart while remaining much less pathogenic [6–8]. AAV vectors are single-stranded DNA vectors capable of achieving stable transgene expression with a good safety profile [9]. The infection and transduction of cells by AAV vectors has been reported to occur by a series of sequential events. After forming an interaction with cell surface receptors on the target cell, the viral capsid undergoes endocytosis. Then, intracellular trafficking through the endocytic/proteasomal compartment and endosomal escape provide the viral capsid for nuclear import. In the nucleus, virion uncoating and viral DNA doublestrand conversion lead to the transcription and expression of the transgene. Successful clinical application of the AAV vector therapeutic platform has been achieved for a variety of diseases, including Leber’s congenital amaurosis, spinal muscular atrophy, hemophilia A and B, and lipoprotein lipase deficiency among others [10, 11]. There may be some limitations to AAV tissue specificity in vivo or at least some significant differences in efficiency of transduction of different tissues and organs. Putatively, AAV9 vector is the most efficient serotype for cardiac gene delivery based on previous success in rodents [12]. However, AAV1, 6, 9-mediated gene expression does not plateau until 4–6 weeks after gene transfer to the myocardium. This slow gene expression profile significantly compromises the therapeutic impact, especially in regard to acute coronary events, which occur quickly and require immediate treatments in order to avoid irreversible myocardial injury. Known AAV vehicles also possess lower DNA packaging capacity, limiting the size of the transgene expression cassette, and may fail to transfect tissue because of preexisting immunity [13]. However, a new

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synthetic AAV serotype 2 lineage clone, Anc80L65, created from a reverse genetics approach and ancestral AAV sequence reconstruction, can help overcome the challenges of existing AAV vectors. Recently, it was demonstrated that Anc80L65 is an efficient gene transfer tool [14–16] with great therapeutic potential. Effective delivery solutions will be essential to advancing cardiac gene therapy efforts. The development of safe and efficient delivery systems is a prerequisite for successful clinical gene therapy. Critical elements of a delivery system consist of selectively targeting tissues of therapeutic relevance and minimizing systemic effects. Existing methods of cardiac gene delivery can be classified by the site and method of injection and interventional approach. Herein, a protocol is described for two most effective methods of cardiac gene transfer: intramyocardial and intracoronary administration. Intramyocardial injection offers the advantage of site selection, since multiple administrations can be carried out through the myocardium. Some important observations were delineated concerning intramyocardial injection: (a) the amount of recombinant protein produced increases with the amount of virus used, (b) reporter gene expression is rarely detected further than 5 mm from the injection site, and (c) the expression profile is similar in both ventricles, and the procedure itself causes minimal side effects in the heart. Intracoronary administration of genes is necessary for the many potential therapeutic targets that are inaccessible by intramyocardial injection. Effective therapy in systemic diseases like heart failure or cardiomyopathy will likely require an intracoronary delivery method capable of globally transducing the heart. This method provides the ability to obtain homogenous transduction. However, it is associated with rapid dilution of viral vectors in circulating blood and significant extracardiac expression.

2 2.1

Materials Anesthesia

1. Small animal ventilator equipped with veterinary anesthesia vaporizer, a nose cone device, and an inhalational isoflurane anesthetic chamber for small animals. 2. Isoflurane, 100 mL. 3. Ketamine HCl, 100 mg/mL. 4. Xylazine, 20 mg/mL. 5. Animal scale.

2.2 AAV Administration

1. Isothermal pad. 2. High-intensity fiber optic gooseneck light.

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3. Soft catheters. 16-gauge (G) for rats with weight of >200 g or 18-gauge for rats with weight of 95%) throughout the surgery. 11. Place rectal temperature probe in the animal, and maintain the body temperature with a heating lamp while the animal lies dorsally. 3.2 Local Intramyocardial Injection into the Ventricular Wall of the Heart

1. Perform a left anterolateral thoracotomy in the fourth intercostal space of the midclavicular line, length ~1 cm. 2. Insert a small self-retaining rib spreader. 3. Move the lung with gauze towards the posterior to allow access to the heart, and incise the pericardium. 4. Prepare the viral solution. After removing from 80  C, viral vector solution should be kept at room temperature around 30 min [14–16] (see Note 4). 5. Once the myocardium is exposed, inject the virus directly into the left ventricle with a 0.5 mL syringe fitted with a 30-gauge needle at an angle of 30–60 degrees in three places: in anterior, anterolateral, and lateral parts of the left ventricle. Each volume of injection is 0.15 mL/site. The overall injection volume should not be more than 0.5 mL. The puncture should go only a few millimeters deep since the ventricular wall is very thin. The heart injectable places should be chosen in the apex and posterolateral wall of the left ventricle (see Note 5).

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6. Once hemostasis is achieved with gentle gauze ball compression, begin closure of the ribs and intercostal muscles with 4-0 Prolene. Before tying the sutures, insert an 18-gauge catheter into the thoracic cavity as thoracoscopy tube, and then finish tying the sutures. Next, close the skin and subcutaneous space with running 5-0 Vicryl (absorbable) suture. After tying the skin sutures, use a 3 mL syringe to evacuate the air/fluid from the pleural cavity through the catheter that was inserted into the thoracic cavity, in order to regain the normal negative physiologic pressure. 7. Administer 2 mL sterile lactate Ringer solution subcutaneously to the animal to restore blood volume. 8. When the animal starts making spontaneous breathing motions and neck movements, disconnect the ventilator from the endotracheal tube, and remove the endotracheal tube from the animal. 9. Return the animal to the cage for recovery. 10. Inject subcutaneously 0.1 mg/kg of buprenorphine for pain control immediately after surgery and then every 12 hours for 3 days. 3.3 Intracoronary Injection (Fig. 1)

1. Follow Subheading 3.2, steps 1–3. 2. Once the myocardium is exposed, put small gauze under the left ventricle, and place a 6-0-purse string suture at the apex of the left ventricle. 3. Identify the aorta and pulmonary artery, and gently isolate these vessels. 4. Through the additional anterior thoracotomy in the second intercostal space, clamp the aorta and pulmonary artery distal to the coronary ostia with small Cooley aortic clamp. 5. Advance a 22 G angiocatheter with introducer from the left ventricular apex through the purse-string suture to the left ventricular outflow. 6. Remove introducer from the catheter, and observe the pulsatile blood flow from the left ventricle. Close the catheter with clamp, and connect to the syringe with viral construct. Slowly inject the 0.4 mL of viral solution during 2–3 sec. This allows the solution that contains the vectors to circulate down the coronary arteries and efficiently transfect the heart (see Note 6). 7. After 10–15 seconds, release the Cooley aortic clamp. 8. Follow Subheading 3.2, steps 6–10.

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Fig. 1 Indirect intracoronary gene delivery with clamping the aorta and pulmonary artery and advancing a catheter from the left ventricle to the aortic root 3.4 Postoperative Recovery

1. Turn off the isoflurane but continue giving the oxygen via endotracheal tube. Place the animal in the sternal position. Do not leave the animal unattended at any point until it has regained sufficient consciousness to ambulate and the animal is safely in its cage. 2. Monitor the heart rate (300–500 beats/min), oxygen saturation (95%), and animal’s limbs color to confirm the animal is breathing well. 3. When the animal starts making spontaneous breathing motions and neck movements and responds to physical stimuli (such as eyes widening, nose moving, and ears responding to sound), extubate the animal by pulling out the catheter and disconnect the ventilator. 4. Return the animal to an empty cage, away from the company of other animals. The animal should be alone for at least 3 days.

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Notes 1. Approval of the study by the local Institutional Animal Care and Use Committee is required before the initiation of the study. 2. Housing and environmental considerations for rats. Animals should be free of all endoparasites, ectoparasites, mycoplasma species, and common rodent’s viruses. Rats’ should be maintained at temperature 22  2  C and humidity 50  10% in a room with ventilated pre-filtered outside air and with controlled light cycles feeding a commercial laboratory diet and water ad libitum. 3. Males and females, as well as males from different litters, should be housed separately. 4. The temperature of the viral solution before administration should be approximately equal to rat’s body temperature according to manufacturer’s instructions. 5. Before intramyocardial injection the researcher should know the coronary anatomy to prevent damage of coronary arteries. For example, the left anterior descending (LAD) artery in rats branches off the ascending aorta and transverses down over the anterior lateral wall of the left ventricle parallel to the interventricular groove. The septal artery is the first large arterial branch of the LAD. The other branches include several diagonals that supply the LV lateral and posterior-free wall and pulmonary trunk. The most important surgical detail is that the LAD artery, through its course length to the apex, transverses completely intramurally and sometimes external inspection is difficult [17]. 6. Synthetic (e.g., AAV9) and non-synthetic AAV vectors (e.g., Anc80L65) are efficient tools for cardiac gene delivery with an early-onset and stable gene expression (Fig. 2). Also, GFP imaging revealed that both viral vectors effectively transduced muscles in the left ventricle. Should note that intramyocardial injection using synthetic AAV vector rendered the more efficient and persistent systemic transduction in cardiac muscles compared with intracoronary route (Fig. 3).

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Fig. 2 Fluorescence imaging of GFP and immunostaining with cardiomyocytes-specific markers a-actinin and DAPI which demonstrated high expression in CMs per field after Anc80L65 administration in rat heart sections after intramyocardial delivery AAV9.GFP and ANC80.GFP. (a) Average number of GFP-positive cells per field: (A) AAV9.GFP 1-day expression. (B) AAV9.GFP 3-day expression. (C) AAV9.GFP 6-week expression. (D) Anc80L65.GFP 1-day expression. (E) Anc80L65.GFP 3-day expression. (F) Anc80L65.GFP 6-week expression. (b) B, Fluorescence imaging of cardiomarkers in rat heart sections at 6 weeks: (A) AAV9.GFP coronal cutting. (B) AAV9.GFP sagittal cutting. (C) Anc80L65.GFP coronal cutting. (D) Anc80L65.GFP sagittal cutting. For nuclear labeling, cells were immunostained with red; actinin, blue; DAPI, and green-GFP

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Fig. 3 Mean area of total GFP fluorescence per cross section after intramyocardial and intracoronary administration of Anc80L65 and AAV9 at 6 weeks. GFP transgene expression was compared on heart cross sections after intramyocardial and intracoronary injections. Strong GFP expression was exhibited after injection of Anc80L65 vector. (a) Percentage of area of GFP fluorescence per section. (b) Fluorescence imaging of GFP: (A) AAV9.GFP intramyocardial delivery. (B) Anc80L65.GFP intramyocardial delivery. (C) AAV9. GFP intracoronary delivery. (D) Anc80L65.GFP intracoronary delivery References 1. Yla¨-Herttuala S, Baker AH (2017) Cardiovascular gene therapy: past, present, and future. Mol Ther 25:1095–1106 2. Ishikawa K, Weber T, Hajjar RJ (2018) Human cardiac gene therapy. Circ Res 123:601–613 3. Katz MG, Fargnoli AS, Kendle AP et al (2016) Gene therapy in cardiac surgery: clinical trials, challenges, and perspectives. Ann Thorac Surg 101:2407–2416 4. Scimia MC, Gumpert AM, Koch WJ (2014) Cardiovascular gene therapy for myocardial infarction. Expert Opin Biol Ther 14:183–195 5. Katz MG, Fargnoli AS, Williams RD et al (2013) Gene therapy delivery systems for enhancing viral and nonviral vectors for cardiac diseases: current concepts and future applications. Hum Gene Ther 24:914–927 6. Hammoudi N, Ishikawa K, Hajjar RJ (2015) Adeno-associated virus-mediated gene therapy in cardiovascular disease. Curr Opin Cardiol 30:228–234 7. Katz MG, Gubara SM, Hadas Y et al (2020) Effects of genetic transfection on calcium cycling pathways mediated by double-stranded adeno-associated virus in postinfarction remodeling. J Thorac Cardiovasc Surg 159:1809– 1819

8. Katz MG, Fargnoli AS, Weber T et al (2017) Use of adeno-associated virus vector for cardiac gene delivery in large-animal surgical models of heart failure. Hum Gene Ther Clin Dev 28: 157–164 9. Gao G, Vandenberghe LH, Alvira MR et al (2004) Clades of adeno-associated viruses are widely disseminated in human tissues. J Virol 78:6381–6388 10. Rangarajan S, Walsh L, Lester W, Perry D, Madan B, Laffan M et al (2017) AAV5-factor VIII gene transfer in severe hemophilia a. N Engl J Med 377:2519–2530 11. Keeler AM, Flotte TR (2019) Recombinant adeno-associated virus gene therapy in light of luxturna (and zolgensma and glybera): where are we, and how did we get here? Annu Rev Virol 6:601–621 12. Inagaki K, Fuess S, Storm TA et al (2006) Robust systemic transduction with AAV9 vectors in mice: efficient global cardiac gene transfer superior to that of AAV8. Mol Ther 14:45– 53 13. Asokan A, Samulski RJ (2013) An emerging adeno-associated viral vector pipeline for cardiac gene therapy. Hum Gene Ther 24:906– 913

Cardiac AAV Delivery 14. Hudry E, Andres-Mateos E, Lerner EP, Volak A, Cohen O, Hyman BT et al (2018) Efficient gene transfer to the central nervous system by single-stranded Anc80L65. Mol Ther Methods Clin Dev 10:197–209 15. Katz MG, Hadas Y, Bailey RA et al (2021) Efficient cardiac gene transfer and early-onset expression of a synthetic adeno-associated viral vector, Anc80L65, after intramyocardial administration. J Thorac Cardiovasc Surg.

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S0022-5223(21)00915-6. https://doi.org/ 10.1016/j.jtcvs.2021.05.050 16. Carvalho LS, Xiao R, Wassmer SJ et al (2018) Synthetic adeno-associated viral vector efficiently targets mouse and nonhuman primate retina in vivo. Hum Gene Ther 29:771–784 17. Katz MG, Fargnoli AS, Gubara SM et al (2019) Surgical and physiological challenges in the development of left and right heart failure in rat models. Heart Fail Rev 24:759–777

Chapter 11 Cardiac Gene Delivery in Large Animal Models: Antegrade Techniques Spyros A. Mavropoulos, Kelly P. Yamada, Tomoki Sakata, and Kiyotake Ishikawa Abstract Percutaneous antegrade coronary injection is among the least invasive cardiac selective gene delivery methods. However, the transduction efficiency of a simple bolus antegrade injection is quite low. In order to improve transduction efficiency in antegrade intracoronary delivery, several additional approaches have been proposed. In this chapter, we will describe the important elements associated with intracoronary delivery methods and present protocols for three different catheter-based antegrade gene delivery techniques in a preclinical large animal model. This is the second edition of this chapter, and it includes modifications we have made over the past several years that further enhance transduction efficacy. Key words Cardiovascular diseases, Gene therapy, Minimally invasive, Homogenous distribution, Intracoronary, Vectors, Gene delivery, Balloon occlusion, Impella

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Introduction Percutaneous antegrade coronary injection is the least invasive cardiac-selective delivery method with a relatively good safety profile. However, a simple antegrade bolus injection results in poor transduction efficiency [1]. To increase transduction efficiency, additional approaches such as slow intracoronary perfusion, intracoronary perfusion + coronary artery occlusion, and intracoronary perfusion + coronary artery occlusion + coronary sinus occlusion have been proposed and tested in preclinical animal models [2]. In this chapter, we will briefly describe features of each method and present practical protocols to perform preclinical gene transfer experiments in large animals. In ex vivo gene transfer experiments using adenovirus, it has been shown that higher coronary flow, increased vector dwell time, larger vector dose, and use of permeability enhancing agents are the

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Fig. 1 The important factors to increase gene transfer efficiency. Coronary flow and the perfusion pressure are somewhat related to each other. Vector dwell time, vector dose, and use of permeability enhancing agents affect gene transfer efficacy independently

most important factors that lead to increased transduction efficiency [3]. The importance of these factors was validated in vivo, with perfusion pressure becoming an additional factor that influences transduction efficiency [1, 4, 5]. Figure 1 shows the important factors for increasing gene transduction efficiency through vascular administration. 1.1 Slow Intracoronary Perfusion

This method is the simplest of the antegrade delivery approaches. The vectors are administered slowly into the coronary arteries without interrupting coronary flow [6–11]. This procedure is a well-established method that is very similar to coronary angiograms that are performed routinely in patients. It is an especially attractive option for gene delivery to patients with end-stage heart failure, as the procedure takes less time and is less invasive compared to other cardiac-targeting delivery methods. A number of clinical trials have already used this method for cardiac gene therapy in heart failure patients and have reported that there were no significant adverse effects associated with this gene delivery method [10, 12– 14]. However, this method suffers from the disadvantage of a lower transduction efficiency when compared to other antegrade delivery approaches. A visual guide on how to perform this procedure can be found in the Journal of Visualized Experiments [15].

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1.2 Intracoronary Perfusion + Coronary Artery Occlusion

This method delivers vectors through the lumen of an inflated balloon catheter during temporary occlusion of the coronary artery. It is slightly more technically complex than the slow intracoronary perfusion method and runs the risk of inducing myocardial ischemia, which can result in post-ischemic myocardial stunning and/or arrhythmias. In our laboratory, we use the Impella (Abiomed, Inc.) to support hemodynamics and alleviate ischemia during coronary artery occlusion. By temporarily blocking coronary artery flow with a balloon catheter and injecting the vectors distal to the occlusion, a higher concentration of delivered vector and longer dwell time are achieved compared to the slow intracoronary perfusion method [4, 16]. Depending on the injection rate, higher coronary flow and perfusion pressure can also be achieved. However, the efficacy of transduction in coronary arterial blockade remains controversial. Using adenovirus, Boekstegers et al. showed that ischemia during coronary artery infusion did not significantly increase myocardial transduction [17]. In contrast, Shah et al. reported that adenoviral infusion with temporary coronary occlusion using a considerably higher flow rate, and consequently elevated coronary pressures, resulted in higher gene expression, but this was also associated with more myocardial injury [18]. Our data suggest that coronary occlusion delivery increases transgene expression for AAV.

1.3 Intracoronary Perfusion + Coronary Artery Occlusion + Coronary Sinus Occlusion

Since the cardiac circulation is predominately supplied by the coronary arteries and drained through the coronary sinus, both vessels can be the target of gene delivery. A combination of coronary artery and coronary sinus blockade increases the vector dwell time in the closed circuit that is created between the balloons inflated in the coronary artery and the coronary sinus [1, 16]. Our data suggest that this method improves AAV gene transduction further compared to the coronary artery occlusion alone.

2

Materials

2.1 Slow Intracoronary Perfusion

1. Heating pad (a heat therapy pump). 2. Mechanical ventilator for large animals. 3. Catheter pack: syringes, drape, bowls, towels, needles, gauze, and scalpel. 4. Vital sign monitors including pressure sensors. 5. Y-connector with hemostatic valve. 6. Contrast agent for angiogram. 7. 5Fr or larger introducer sheath for vascular access. 8. Standard angiography introducer needle 18G
2.7500 and J wire.

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9. Guiding catheter for coronary artery access (see Note 1). 10. 0.035-inch guide wire. 11. Two 0.014-inch coronary guide wires with soft tip for coronary artery access. 12. Disinfectants: 70% isopropyl alcohol and povidone-iodine. 13. Telazol (tiletamine/zolazepam). 14. Buprenorphine. 15. Propofol. 16. Heparin sodium. 17. Vector solution. 18. Vector dilution solution: phosphate-buffered saline (PBS), 0.001% pluronic [19]. 19. Two 20-mL syringes for vector infusion. 20. Infusion pump tubing. 21. Infusion pump. 22. Nitroglycerin. 23. Phenylephrine. 24. ACT point-of-care device. 2.2 Intracoronary Perfusion + Coronary Artery Occlusion

1. Subheading 2.1, items 1–24. 2. Inflation device for a balloon catheter. 3. Over-the-wire (OTW) balloon dilation catheter for the occlusion of the coronary artery and injection of the vectors. The balloon size should be selected depending on the size of the coronary artery as determined during coronary angiography (see Note 2). 4. Impella (Abiomed, Inc.). 5. 14Fr introducer sheath. 6. 0.038-inch stiff wire. 7. 0.018-inch wire for Impella insertion. 8. Manual external defibrillator.

2.3 Intracoronary Perfusion + Coronary Artery Occlusion + Coronary Sinus Occlusion

1. Subheadings 2.1, items 1–24 and 2.2, items 1–8. 2. 7Fr long flexible sheath for jugular vein approach. 3. Guiding catheter for coronary sinus access for femoral vein approach (see Note 3). 4. 0.014-inch hydrophilic wire for the coronary venous access. 5. Balloon wedge catheter for coronary sinus occlusion. We use a 5Fr Swan-Ganz catheter. 6. 6Fr deflectable catheter for jugular access.

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1. Premedicate the animal (a pig or other large animal) using Telazol (8.0 mg/kg) for sedation and buprenorphine (0.01–0.03 mg/kg) for analgesia (see Note 4). 2. Intubate the animal and ventilate with 100% oxygen. Obtain venous access on the ear vein for infusion of fluids, propofol, and nitroglycerine (see Note 4). 3. Place the animal in dorsal position on a heating mat during the procedure, with its legs extended by ropes (see Note 5). 4. Connect the monitors on the animal. 5. Prepare the puncture site with 70% isopropyl alcohol followed by povidone-iodine. 6. Puncture the peripheral artery (femoral or carotid artery) to obtain an arterial access using the Seldinger method (see Notes 6 and 7). 7. Administer heparin sodium at the dose of 200–300 U/kg IV to achieve an activated coagulation time of 250–300 seconds. 8. Start intravenous nitroglycerin (1 μg/kg/min) infusion through the ear vein so that it can be infused for 15 minutes prior to vector injection [20]. Infusion will continue for the duration of the vector injection and will end after 10 minutes have elapsed from the completion of the injection procedure. 9. Advance the 0.035-inch guide wire with the guiding catheter to the ascending aorta. 10. Set up the injection lines as presented in Fig. 2, making sure that there is no air in the system. 11. Engage the left coronary artery (LCA), and after the angiogram, advance two 0.014-inch guide wires (see Note 8), one into the left anterior descending artery (LAD) and one into the left circumflex coronary artery (LCX) to stabilize the catheter position (see Notes 9 and 10). 12. Prepare two syringes. In the first syringe, dilute the vector solution to 15 mL with vector dilution solution. Prepare another syringe filled with more than 10 mL of vector dilution solution. The second solution (flush) will be used to deliver the residual virus within the catheter lumen. 13. Confirm that the catheter tip is stable at just proximal to the bifurcation of LAD and LCX. The pressure through the catheter is monitored to ensure that the catheter is not wedging. 14. Slowly inject contrast to ensure even flow to the LAD and LCX (see Note 11).

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Fig. 2 Slow intracoronary perfusion system. During the infusion, the Y-connector valve should be tightly locked to prevent the vector solution from loss. Pressure monitor line can be connected to femoral arterial sheath during injection. (a) Infusion pump. (b) Three-way stopcock for exchanging the infusion line and pressure monitor line (also to the manifold for angiography). (c) Y-connector with hemostatic valve and coronary wires. (d) Guiding catheter. (Reproduced from Watanabe et al. (first edition of this chapter) [22])

15. The vector solution (10 mL) is injected through the catheter over 10 minutes (1 mL/min) using an infusion pump into the LCA. After 10 minutes, the syringe is exchanged to the flush, and infusion is continued for 5 minutes (1 mL/min) (see Note 12 and Fig. 2). 16. Remove the catheter from the LCA, and engage the catheter to the right coronary artery (RCA) (see Note 13). 17. A 0.014-inch wire is inserted deep into the RCA to fix the catheter position. 18. The remaining 5 mL of vector solution is injected into the RCA over 5 minutes (1 mL/min) followed by 5 minutes (1 mL/ min) of flush. 19. Withdraw the guiding catheter and the wire. 20. Withdraw the sheath from the artery, and achieve hemostasis by applying direct pressure to the site for several minutes (see Note 14). 3.2 Intracoronary Perfusion + Coronary Artery Occlusion

1. Follow Subheading 3.1, steps 1–8. 2. Prepare a manual external defibrillator for reverting ventricular fibrillation that can be induced by the coronary artery occlusion (see Note 15). 3. Establish a 14Fr arterial access in the femoral artery for Impella delivery (see Note 16).

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4. Check ACT and administer additional heparin if ACT is below 250. 5. Insert 0.018-inch wire through the 14Fr sheath, and deliver Impella into the left ventricle. Run Impella at the maximum flow. 6. For the coronary artery occlusion, advance an angioplasty balloon along with the coronary guide wire into the target artery from the opposite side of the arterial access used for the Impella (see Note 17). 7. Perform an angiogram and place the balloon in a proximal part of the LAD. 8. Remove the wire inside the OTW balloon, and inflate the coronary balloon for 15 seconds three times for preconditioning. 9. After preconditioning, inflate the balloon and inject the vector solution through the wire lumen of the angioplasty balloon (see Notes 18–20). We inject 0.8–1 mL, and wait for 1 min before balloon deflation. This is repeated three times for each artery. 10. Upon finishing the injection, remove the balloon and the wire from the target coronary artery. 11. Advance a balloon along with the coronary wire into another target artery. Similarly occlude and inject the vectors. 12. Withdraw the guiding catheter, the balloon, and the wire. 13. Slowly wean the Impella over 15 minutes. Turn off and remove Impella (see Note 21). 14. Withdraw the sheaths from the arteries, and achieve hemostasis by applying direct pressure to the site for several minutes (see Note 14). 3.3 Intracoronary Perfusion + Coronary Artery Occlusion + Coronary Sinus Occlusion

1. Follow Subheading 3.2, steps 1–5. 2. For femoral access, advance the guide wire with the guiding catheter to venous system (see Note 22). For jugular access, use a 6Fr deflectable catheter to cannulate coronary sinus, and advance the flexible sheath into the coronary sinus along the catheter. A 6 or 7Fr sheath is recommended for this purpose. 3. Select the great cardiac vein with a 0.014-inch wire, and advance an occlusion balloon to the great cardiac vein (see Note 23). 4. Inflate the balloon and confirm rise in catheter tip pressure, which indicates complete occlusion. Systolic pressure will become similar to systolic aortic pressure if complete occlusion is achieved and there are no collateral branches. 5. Follow Subheading 3.2, steps 6–8.

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6. Occlude the coronary sinus first, and then occlude the coronary artery. 7. Follow Subheading 3.2, steps 9–13.

4

Notes 1. A Hockey-stick catheter is suitable for the femoral approach, and an Amplatz right catheter is suitable for the carotid approach in pigs. Usually, a 5Fr hockey stick catheter can cannulate both the LCA and RCA. 2. The shorter balloon is preferred to minimize the damage of the endothelium from the expansion of the balloon. 1–2 atm is sufficient to completely occlude the coronary artery using an appropriately sized balloon. 3. An Amplatz left catheter is suitable for the femoral approach to engage the coronary sinus in pigs. 4. Analgesia, anesthesia, and antibacterial drugs and all procedures are approved by the animal committee at the facility for all procedures. Inhalational anesthetics can be used instead of propofol. 5. The ropes should be tight during the vascular puncture (see Note 6), but loosened after vascular access is established. 6. Tightly pulling the leg of the puncture side will facilitate vessel puncture. In young healthy animals, vessels are mobile, and avoid needles when the legs are loosely pulled. 7. Echo guidance will increase the accuracy of the puncture. The femoral and the carotid arteries can also be accessed by a cutdown method. 8. Draw blood and flush the catheter every time a new wire/ balloon is removed/introduced to prevent air/thrombus injection into the body. 9. The administration of a vector solution into the coronary artery seems simple, but it can be tricky, especially in some species. An appropriate injection catheter should be chosen for each species. This can vary depending on the size of the animal. Inserting the coronary wires into the branches will ensure catheter stability at the coronary ostium. The catheter tip should remain inside the left main tract; however deep engagement of the catheter into the branches can cause an obstruction of the coronary artery and the formation of a thrombus. A large amount of heparin, continuous monitoring of the catheter position under fluoroscopy, and checking the catheter pressure and ECG for signs of ischemic changes are important.

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10. Insert the coronary wire into the LAD first, and then insert the second one into the LCX. Having the wire in the LCX first will make it very difficult to insert one into the LAD. 11. Alternatively, microcatheters can be delivered into each branch to inject them separately. However, this selective infusion can result in very proximal branches being bypassed during the delivery. 12. Make sure the Y connector valve is tightly locked. A leak from a loosely closed Y connecter results in loss of vectors. 13. An appropriate catheter for the RCA should be determined before the starting of LCA vector delivery. Catheters that fit both the LCA and RCA are preferred; however stable RCA cannulation and easy manipulation should be prioritized. We use Hockey-stick, but JR catheters also work well. 14. Protamine sulfate can be administered slowly over 5 min to accelerate hemostasis if the vector does not interact with the drug. We recommend pulling out the sheath prior to protamine injection to prevent clot formation around the sheath, which can cause thromboembolic complications. Although the frequency is low, protamine can cause hypotension. Continuous monitoring with ECG and pulse oximeter is necessary. If the signal of the pulse oximeter is lost, check the noninvasive blood pressure. If there is hypotension, inject 1–2 mL of atropine sulfate IV, or, in a severe case, inject low dose (0.5–1.0 mg) of phenylephrine IV. With timely administration of these drugs, the animal will recover without any adverse effects. 15. In case of ventricular fibrillation, apply 200 J shock as soon as possible. If the rhythm does not recover, deflate the balloon and apply another shock. Chest compression should be maintained during the arrhythmia. Monitor aortic pressure to evaluate the hemodynamics. 16. Pig arteries can be very spastic. We place the 5Fr sheath first and wait for at least 10 minutes so that the spasm is relieved. If the spasm is too severe during sheath insertion, do not attempt to overcome it by force; wait for the spasm to be relieved instead. 17. While 5Fr guiding catheter allows delivery of coronary balloons, it will compromise pressure monitoring through the catheter. We recommend using 6Fr or larger guiding catheters. 18. Continuously monitor the blood pressure, and deflate the balloon if the systolic aortic pressure falls below 60 mmHg. Consider early balloon deflation when there are frequent premature ventricular beats, since they often precede ventricular fibrillation.

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19. Intracoronary adenosine (25 μg) can be injected to increase cellular permeability before injection of the vector solution [21]. 20. It has been shown that high injection rate is associated with myocardial injury [4]. 21. Sudden removal can result in acute decompensation in heart failure animals. Gradual weaning is recommended. 22. The coronary sinus ostium is located in the anteroinferior atrial septum. An injection of contrast from the coronary artery can facilitate identifying the location of the coronary sinus. 23. In pigs, the azygos vein merges with the distal end of the coronary sinus (near the right atrium). The balloon should be located proximal to the merge. Coronary sinus pressure will not increase when the balloon is placed distal to the merge; thus pressure monitoring during coronary sinus occlusion will help determine optimal positioning.

Acknowledgments This work was supported by NIH R01 HL139963 (K.I). We acknowledge the Gene Therapy Resource Program (GTRP) of the National Heart, Lung, and Blood Institute, National Institutes of Health for providing the gene vectors used in this study. S.A.M. was supported by National Institutes of Health T32 HL007824-23. References 1. Logeart D, Hatem SN, Heimburger M, Le Roux A, Michel JB, Mercadier JJ (2001) How to optimize in vivo gene transfer to cardiac myocytes: mechanical or pharmacological procedures? Hum Gene Ther 12(13):1601–1610. h t t p s : // d o i . o r g / 1 0 . 1 0 8 9 / 10430340152528101 2. Ishikawa K, Tilemann L, Ladage D, Aguero J, Leonardson L, Fish K, Kawase Y (2012) Cardiac gene therapy in large animals: bridge from bench to bedside. Gene Ther 19(6):670–677. https://doi.org/10.1038/gt.2012.3 3. Donahue JK, Kikkawa K, Johns DC, Marban E, Lawrence JH (1997) Ultrarapid, highly efficient viral gene transfer to the heart. Proc Natl Acad Sci U S A 94(9):4664–4668 4. Emani SM, Shah AS, Bowman MK, Emani S, Wilson K, Glower DD, Koch WJ (2003) Catheter-based intracoronary myocardial adenoviral gene delivery: importance of

intraluminal seal and infusion flow rate. Mol Ther 8(2):306–313 5. Hajjar RJ, Schmidt U, Matsui T, Guerrero JL, Lee KH, Gwathmey JK, Dec GW, Semigran MJ, Rosenzweig A (1998) Modulation of ventricular function through gene transfer in vivo. Proc Natl Acad Sci U S A 95(9):5251–5256 6. Tilemann L, Lee A, Ishikawa K, Aguero J, Rapti K, Santos-Gallego C, Kohlbrenner E, Fish KM, Kho C, Hajjar RJ (2013) SUMO-1 gene transfer improves cardiac function in a large-animal model of heart failure. Sci Transl Med 5(211):211ra159. https://doi.org/10. 1126/scitranslmed.3006487 7. Kawase Y, Ly HQ, Prunier F, Lebeche D, Shi Y, Jin H, Hadri L, Yoneyama R, Hoshino K, Takewa Y, Sakata S, Peluso R, Zsebo K, Gwathmey JK, Tardif JC, Tanguay JF, Hajjar RJ (2008) Reversal of cardiac dysfunction after long-term expression of SERCA2a by gene transfer in a pre-clinical model

Intracoronary Gene Delivery of heart failure. J Am Coll Cardiol 51(11): 1112–1119. https://doi.org/10.1016/j.jacc. 2007.12.014 8. Ishikawa K, Fish KM, Tilemann L, Rapti K, Aguero J, Santos-Gallego CG, Lee A, Karakikes I, Xie C, Akar FG, Shimada YJ, Gwathmey JK, Asokan A, McPhee S, Samulski J, Samulski RJ, Sigg DC, Weber T, Kranias EG, Hajjar RJ (2014) Cardiac I-1c overexpression with reengineered AAV improves cardiac function in swine ischemic heart failure. Mol Ther 22(12):2038–2045. https://doi.org/10.1038/mt.2014.127 9. Fish KM, Ladage D, Kawase Y, Karakikes I, Jeong D, Ly H, Ishikawa K, Hadri L, Tilemann L, Muller-Ehmsen J, Samulski RJ, Kranias EG, Hajjar RJ (2013) AAV9.I-1c delivered via direct coronary infusion in a porcine model of heart failure improves contractility and mitigates adverse remodeling. Circ Heart Fail 6(2):310–317. https://doi.org/10.1161/ CIRCHEARTFAILURE.112.971325 10. Greenberg B, Yaroshinsky A, Zsebo KM, Butler J, Felker GM, Voors AA, Rudy JJ, Wagner K, Hajjar RJ (2014) Design of a phase 2b trial of intracoronary administration of AAV1/SERCA2a in patients with advanced heart failure: the CUPID 2 trial (calcium up-regulation by percutaneous administration of gene therapy in cardiac disease phase 2b). JACC Heart Fail 2(1):84–92. https://doi.org/ 10.1016/j.jchf.2013.09.008 11. Ishikawa K, Aguero J, Naim C, Fish K, Hajjar RJ (2013) Percutaneous approaches for efficient cardiac gene delivery. J Cardiovasc Transl Res 6(4):649–659. https://doi.org/10.1007/ s12265-013-9479-7 12. Hammond HK, Penny WF, Traverse JH, Henry TD, Watkins MW, Yancy CW, Sweis RN, Adler ED, Patel AN, Murray DR, Ross RS, Bhargava V, Maisel A, Barnard DD, Lai NC, Dalton ND, Lee ML, Narayan SM, Blanchard DG, Gao MH (2016) Intracoronary gene transfer of adenylyl cyclase 6 in patients with heart failure: a randomized clinical trial. JAMA Cardiol 1(2):163–171. https://doi. org/10.1001/jamacardio.2016.0008 13. Hulot JS, Salem JE, Redheuil A, Collet JP, Varnous S, Jourdain P, Logeart D, Gandjbakhch E, Bernard C, Hatem SN, Isnard R, Cluzel P, Le Feuvre C, Leprince P, Hammoudi N, Lemoine FM, Klatzmann D, Vicaut E, Komajda M, Montalescot G, Lompre AM, Hajjar RJ, Investigators A-H (2017) Effect of intracoronary administration of AAV1/SERCA2a on ventricular remodelling in patients with advanced systolic heart failure: results from the AGENT-HF randomized

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phase 2 trial. Eur J Heart Fail 19(11): 1534–1541. https://doi.org/10.1002/ ejhf.826 14. Lyon AR, Babalis D, Morley-Smith AC, Hedger M, Suarez Barrientos A, Foldes G, Couch LS, Chowdhury RA, Tzortzis KN, Peters NS, Rog-Zielinska EA, Yang HY, Welch S, Bowles CT, Rahman Haley S, Bell AR, Rice A, Sasikaran T, Johnson NA, Falaschetti E, Parameshwar J, Lewis C, Tsui S, Simon A, Pepper J, Rudy JJ, Zsebo KM, Macleod KT, Terracciano CM, Hajjar RJ, Banner N, Harding SE (2020) Investigation of the safety and feasibility of AAV1/SERCA2a gene transfer in patients with chronic heart failure supported with a left ventricular assist device – the SERCA-LVAD TRIAL. Gene Ther 27(12):579–590. https://doi.org/10.1038/ s41434-020-0171-7 15. Ishikawa K, Ladage D, Tilemann L, Fish K, Kawase Y, Hajjar RJ (2011) Gene transfer for ischemic heart failure in a preclinical model. J Vis Exp 51. https://doi.org/10.3791/2778 16. Hayase M, Del Monte F, Kawase Y, Macneill BD, McGregor J, Yoneyama R, Hoshino K, Tsuji T, De Grand AM, Gwathmey JK, Frangioni JV, Hajjar RJ (2005) Catheter-based antegrade intracoronary viral gene delivery with coronary venous blockade. Am J Physiol Heart Circ Physiol 288(6):H2995–H3000. https://doi.org/10.1152/ajpheart.00703. 2004 17. Boekstegers P, von Degenfeld G, Giehrl W, Heinrich D, Hullin R, Kupatt C, Steinbeck G, Baretton G, Middeler G, Katus H, Franz WM (2000) Myocardial gene transfer by selective pressure-regulated retroinfusion of coronary veins. Gene Ther 7(3):232–240. https://doi. org/10.1038/sj.gt.3301079 18. Shah AS, White DC, Emani S, Kypson AP, Lilly RE, Wilson K, Glower DD, Lefkowitz RJ, Koch WJ (2001) In vivo ventricular gene delivery of a beta-adrenergic receptor kinase inhibitor to the failing heart reverses cardiac dysfunction. Circulation 103(9):1311–1316 19. Bennicelli J, Wright JF, Komaromy A, Jacobs JB, Hauck B, Zelenaia O, Mingozzi F, Hui D, Chung D, Rex TS, Wei Z, Qu G, Zhou S, Zeiss C, Arruda VR, Acland GM, Dell’Osso LF, High KA, Maguire AM, Bennett J (2008) Reversal of blindness in animal models of leber congenital amaurosis using optimized AAV2mediated gene transfer. Mol Ther 16(3): 458–465. https://doi.org/10.1038/sj.mt. 6300389 20. Karakikes I, Hadri L, Rapti K, Ladage D, Ishikawa K, Tilemann L, Yi GH, Morel C, Gwathmey JK, Zsebo K, Weber T, Kawase Y,

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Hajjar RJ (2012) Concomitant intravenous nitroglycerin with intracoronary delivery of AAV1.SERCA2a enhances gene transfer in porcine hearts. Mol Ther 20(3):565–571. https://doi.org/10.1038/mt.2011.268 21. Wright MJ, Wightman LM, Latchman DS, Marber MS (2001) In vivo myocardial gene transfer: optimization and evaluation of

intracoronary gene delivery in vivo. Gene Ther 8(24):1833–1839. https://doi.org/10. 1038/sj.gt.3301614 22. Watanabe S, Leonardson L, Hajjar RJ, Ishikawa K (2017) Cardiac Gene Delivery in Large Animal Models: Antegrade Techniques. Methods Mol Biol 1521:227–235. https://doi.org/10. 1007/978-1-4939-6588-5_16

Chapter 12 Locked Nucleic Acid AntimiR Therapy for the Heart Sabine Samolovac and Rabea Hinkel Abstract Coronary heart disease is one of the leading causes of death in industrialized nations. Even though revascularization strategies improved the outcome of patients after acute myocardial infarction, about 30% of patients develop chronic heart failure. Ischemic heart disease and heart failure are characterized by an adverse remodeling of the heart, featuring cardiomyocyte hypertrophy, increased fibrosis, and capillary rarefaction. Therefore, novel therapeutic approaches for the treatment of heart failure, such as reducing ischemia/reperfusion injury, fibrosis, or hypertrophy, are needed. Here microRNAs (miRNAs) come into play. For heart failure, cardiac stress and several cardiovascular diseases, individual miRNAs, and whole miRNA clusters have been implicated as disease relevant and dysregulated. miRNAs are short non-coding RNA molecules of about 22 nucleotides, and their inhibitors are 8–15 nucleotides long plus a sugar-ring (LNA, locked nucleid acid) or cholesterol ending (AntagomiR). Modulation of miRNAs might serve as therapeutic targets through miRNA knockdown or overexpression via miRNA mimics. Due to their pleiotropic mode of action and the presence of individual miRNAs in a variety of tissues and cells, a local, target region-oriented application is important to achieve therapeutic effects and at the same time reducing adverse effects in other off-target organs and tissues. Due to their small size, the miRNA inhibitors are able to pass endothelial barrier at both arterial and venous sides of the bloodstream vessels. For these reasons, the gold standard administration route of miRNA modulators for therapeutic approaches of the left ventricle is the anterograde application into one or both coronary arteries via an over-the-wire (OTW) balloon. In this chapter we provide a comprehensive description of the anterograde application procedure in a large animal model such as pig. Of a particular note, this methodology is a standard procedure in catheter laboratories, a key characteristic that allows therapeutic translation from large animals to patients. Key words miRNA therapy, Inhibition, Oligonucleotides, Anterograde delivery, Ischemic heart disease, Hypertrophy, LNA, AntagomiR

1

Introduction Since the discovery of microRNAs (miRNAs) in 1993 [1] and their important role in the complex network of gene regulation, especially at the post-transcriptional level, they came to the attention to a variety of research areas [2]. The field of cardiovascular diseases is not an exception, being highly represented by numerous publications over the past two decades. An increase of cardiac-specific

Kiyotake Ishikawa (ed.), Cardiac Gene Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2573, https://doi.org/10.1007/978-1-0716-2707-5_12, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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microRNAs could be observed after myocardial infarction [3, 4], ischemia-reperfusion injury [5, 6], and aortic stenosis causing myocardial hypertrophy [7, 8]. The approach of an anti-miRNA therapy in the field of cardiovascular disease offers a new pathway on a molecular level to reduce the increased miRNA levels and allowing a regular protein expression in order to reduce the damage on the myocardium caused by acute and chronic heart failure [9, 10]. The structural design of miRNAs, which are short non-coding RNA molecules about 22 nucleotides long, allows different ways of inhibiting the mature miRNA leading to a loss of function [11]. The most common way to downregulate miRNA levels in vivo is the use of anti-microRNA (antimiRs or antimiRNA) [12], consisting of modified antisense oligonucleotides that are partially or fully complementary to the miRNA [13]. The antimiRs, which effectively inhibit miRNAs in the heart, are defined in two categories: (a) the 20 -O-methyl group (AntagomiRs) [14] and (b) the LNA-modified oligonucleotids (LNA-antimiRs) [10, 15]. While the AntagomiRs are conjugated to cholesterol, which facilitates their cellular uptake, the LNA-antimiRs have a phosphorothioate backbone, providing more stability, highbinding affinity, and good pharmacokinetic properties [13]. Compared to the AntagomiRs, LNAs are not complementary to the full mature miRNA sequence but exhibit 15 nucleotides including some mismatch to the sequence. Additionally, they can even be directed against the seed region of a miRNA (8 nucleotides), targeting multiple miRNA family members at once. In vivo treatments with LNA-antimiRs require lower doses (10–25 mg/kg) compared to AntagomiRs (80 mg/kg) and show a knockdown up to 60 days [13], depending on the species and the specific miRNA targeted. Considering these findings, the application of LNA-antimiR as therapeutic approach in small and large animal models is largely established in multiple fields such as wound healing [2, 16], blood cancer [17, 18] and hepatitis C [19, 20]. The large success of these studies allowed the development of treatments up to the stage of phase II clinical trials [2]. Regarding the deployment of LNA-antimiR as a potential therapy in cardiovascular diseases such as acute and chronic myocardial infarction [9, 10, 21, 22] and cardiac hypertrophy [23, 24], the main research focus was set on small animal models like mice and rats. One should consider that working with small animal models decreases the variety of possible translational application routes down to intravenous or subcutaneous. Large animal models like rabbits, dogs, pigs, and non-human primates are comparable to the body size of infants up to adults and allow clinically relevant application modalities. Since the application site determines the distribution pattern, the intracoronary direct administration of heart-specific LNA-antimiR shows a higher effect compared to the less invasive intravenous route [10]. Other approaches, such as direct injection into the myocardium, require either an open-chest model [25] or a special

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Therapeuc agent LAD

AIV Connector inflaon device Target region

miRNA inhibitor

Fig. 1 Scheme of anterograde miRNA inhibitor application into the LAD region via an over-the-wire balloon

catheter/technique [26] that are associated with a great medical effort, which drastically aggravates the feasibility and immensely increases the burden on the affected person/model. Regarding our own experiences on heart-specific LNA-antimiRs and different application routes, we recommend the minimal invasive percutaneous transluminal antegrade coronary approach [10, 27, 28]. This technique can be used for single coronary artery application, such as therapeutic approach for acute myocardial infarction [10], as well as for general left ventricular application, e.g., in chronic ischemic cardiomyopathy [27], or as antihypertrophic therapy [28]. In case of (a) therapeutic deliveries which are larger in size than LNA-antimiRs, e.g., peptides, cDNA, proteins, or (b) a diseaserelated obstruction of the left anterior descending artery (LAD), the retrograde application route via the cardiac vein (AIV, anterior intraventricular vein) is a safe but technical more advanced alternative [29]. The procedure of the percutaneous transluminal coronary antegrade application approach in a pig model is described in detail in this chapter (Fig. 1).

2

Materials

2.1 Arterial and Venous Peripheral Access

1. 8 French (F) sheath (including dilatator and wire) for the common carotid artery. 2. 11 F sheath (including dilatator and wire) for the external jugular vein. 3. Saline solution.

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Judkin right

Y-adapter

Torque handle

0.0014´´ guide wire wire /application line

guide wire insertion tool

balloon inflating devise

3-port angiography manifold Set

OTW balloon

Fig. 2 Materials needed for anterograde left coronary arterial miRNA-inhibitor application

4. Heparin. 5. Coagulation meter (activated clotting time; ACT). 6. Suture for vessel occlusion and skin closure. 2.2 Angiography Equipment (Fig. 2)

1. X-ray C-arm with angiography tools. 2. Contrast agent. 3. Saline solution. 4. Three-port angiography manifold set. 5. Pressure line for injection. 6. Y-Adapter. 7. Torque handle. 8. Guidewire insertion tool.

2.3 Catheterization of the Coronary Artery (LAD or Left Circumflex Artery (RCx))

1. Judkin right (JR) catheter 7F with side holes.

2.4

1. 5 ml syringe.

Infusion of LNA

2. 0.014 in. guide wire 300 cm length. 3. Over-the-wire balloon (2.5–3.5  8/10 mm). 4. Balloon inflation device (Fig. 2).

2. Saline solution or autologous blood as substrate.

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1. Nitroglycerin solution: 1 mg glycerol trinitrate, 10 mL saline. 2. Saline solution or autologous blood. 3. Defibrillator with external paddles. 4. Emergency medication (Suprarenin solution; either 1 mg epinephrine, 1 mL saline or 1 mg epinephrine, 10 mL saline) (Cordarex solution; 150 mg amiodarone).

2.6

3

AntimiR

LNA antimiRs are desalted and dried down in different nmol quantities. They should be stored at 15 to 30  C to be stable for at least 6 months. After re-suspension with nuclease-free TE buffer (10 mM Tris, 0.1 mM EDTA, pH 7.5 or 8.0), the stock solution should be aliquoted to avoid repeated freeze-thaw cycles, which will degrade the oligonucleotides. A storage at 15 to 30  C or below in a constant temperature freezer is recommended. Oligonucleotids are degraded by freeze-thaw cycles especially.

Methods

3.1 Venous and Arterial Assess

1. For the venous and arterial assess, place the animal in supine position, and tie the legs down. Shave the neck region, scrub, and drape in preparation for surgery (see Notes 1 and 2). 2. Perform a 3 cm neck skin incision; here the center of the incision is the middle of the triangle defined by the mandibular angle, the shoulder joint, and the tip of the sternum. Use the sternocloidomastoideus muscle as guidance for skin incision, blunt preparation of subcutaneous fat and muscle layer. 3. The carotid artery is on the medial part of the muscle, next to internal jugular vein and nerves from the cervical plexus. Prepare the carotid artery by freeing it from the surrounding tissue layers down to the adventitia while protecting the surrounding veins and nerves. Introduce an 8F sheath into the cranially ligated common carotid artery via an introducer. After successful introduction connect the sheath to a monitor system for internal blood pressure measurement. 4. Proceed in the same manner regarding the preparation as it is described above for the artery with the external jugular vein (vena jugularis externa) on the lateral side of the muscle. After ligating the vein as cranial as possible, introduce a 11F sheath. 5. Apply 10,000 IE heparin systemically (activated clotting time (ACT) > 250 s) (see Note 1).

3.2 Blocking of the Anterograde Flow in Coronary Arteries (LAD and RCx)

During the infusion time, the anterograde flow needs to be blocked, in order to achieve an optimal uptake efficacy up to the distal parts of the vein including the side branches.

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1. Use a Judkin right 7F catheter with side holes for blocking either the LAD or the RCx (see Subheading 3.2, step 5); connect the Y-adapter with the three-port manifold set/contrast syringe to the catheter (Fig. 2). Flush the whole system carefully with saline, and introduce the catheter into the 8F sheath on the carotid artery. 2. Adjust the X-ray to the area of the heart (valves), and proceed the catheter forward to the height of the aortic arch. Flush the catheter again with saline, turn the tip of the catheter to the right side, and move further towards the left ventricle. 3. Turn the catheter tip to the left side after passing the aortic arch branching, and carefully advance into the left main coronary artery (Fig. 3). 4. For visualizing the branching of the left coronary arteries, take an angiography via X-ray (Fig. 3). 5. Insert the 0.01400 guide wire into the JR catheter via the insertion tool, and push forward to the tip of the catheter. 3.2.1 For the LAD Approach

After reaching of the catheter tip with the guide wire, rotate the catheter tip into the direction of the coronary artery (LAD), and forward the wire into the LAD (Fig. 3). If needed, attach the torque handle to the distal end of the 0.01400 guide wire for a better rotation to place it in the LAD.

A

B

JR tip JR tip RCx

LAD

RCx

L AD

Guide wire tip

Occlusion OTW balloon

Fig. 3 Example for LAD miRNA-inhibitor application via an OTW balloon. (a) Coronary angiography of the left ventricular descending artery (LAD) as indicated by the red arrows. The tip of the Judkins right catheter is positioned in the left main before the branching of the LAD and RCx. Patency of the left coronary arteries should be assured, as displayed here with the contrast agent. (b) OTW balloon (inflated) at position of target area in the LAD via guide wire as indicated with the red arrows. miRNA-inhibitor is applied via the lumina of the OWT after withdrawal of the guide wire. Treatment follows the contrast agent distribution as displayed in a

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After reaching of the catheter tip with the guide wire, slowly advance the guide wire with a slight rotation into the direction of the RCx. Normally, the blood flow will take it following the anatomical conditions into the RCx (Fig. 3). 1. Connect the over-the-wire balloon to the balloon inflation device (filled with 50% saline: 50% contrast agent, Fig. 2). 2. Insert the balloon via the 0.01400  300 cm guide wire into the JR catheter, and proceed to the tip of the catheter. To position the balloon in the LAD (distal of the first diagonal branch), the balloon is pushed forward while holding the wire in the same position, in order to avoid wire-based vessel perforation as well as ventricular fibrillation. To position the balloon in the RCx (between the first and second left marginal artery), push the balloon forward while holding the wire at the same position. 3. Assure the correct positioning of the balloon by X-ray (Fig. 3).

3.3 Anterograde Application (LAD and RCx)

1. Dilute the LNA-antimiR in 5 ml saline solution or autologous blood in a syringe (see Note 3). 2. Occlude the LAD or the RCx (4–6 atm) by inflating the OTW balloon with the connected balloon inflation device. 3. Inject the diluted LNA-antimiR via the wire line of the overthe-wire (OTW) balloon continuously and slowly over the period of 3–5 min. 4. Flush the lumen of the OTW balloon with additional 2 mL of saline solution. 5. Deflate the balloon after a max. of 5–10 min (see Note 4). 6. Retract OTW balloon out of the JR catheter. 7. Control angiography for patency of the coronary artery (see Note 5). 8. Finally remove the JR catheter (see Note 6). 9. Remove the sheath and ligate the carotid artery (see Note 7). 10. Skin closure in two layers, muscle layer (platysma, resorbable suture) and the skin (non-resorbable suture, see Note 7).

4

Notes 1. Before starting the catheterization (before inserting the venous and arterial sheath), 10,000 IE of heparin are applied for a targeted ACT of >250 s. In the course of the intervention, especially if it is prolonged, the ACT must be checked regularly (every 30 min). If the value is design-line, and draw the injection area. 2. In the LLS map, select an area where LLS is less than 12%, as it is considered as a dyskinetic myocardium (see Fig. 1). 3. In the unipolar map, select an area where unipolar voltage is more than 5 mV, appearing as a violet color [19] (see Note 13).

3.6

Gene Transfers

1. The operator guides the catheter in the left ventricle and controls the curvature properties of the catheter, and an assistant controls needle movement and injections. 2. Connect the 7-Fr MyoStarTM injection catheter to the cable connection board. 3. Adjust the needle length to 3 mm by rotating the Luer-lock fitting connecting to the injectant syringe (see Notes 2, 14, and 15).

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Fig. 1 Electroanatomical map of the left ventricle. The colors of the map describe dyskinesia of the myocardium. The brown tags indicate locations of gene transfer injections

4. Place the injectant solution syringe into the catheter (see Note 16). With the needle revealed, push the syringe gently to clear any air out of the catheter. 5. Introduce the catheter to the left ventricle under fluoroscopic guidance (see Note 6). 6. After reaching the left ventricle, the start menu indicators should show green, and the catheter tip movement is seen on the reconstructed electroanatomical map. 7. Move to the desired location by catheter movement, rotation, and catheter tip curving. 8. Press down the injector thumb knob to introduce the needle into the myocardium (see Note 17). After reaching the previously determined injection site, introduce the needle, and confirm the valid injection placement by acquiring a point. Like previously during mapping, the LAT, CL, Loc, and LS parameters should indicate green. 9. Assess the quality of the placement by using the criteria described in Subheading 3.4. In addition to the current criteria, use a low LLS value, preferably below 6%.

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10. Once the placement is correct and the needle is attached to the myocardium, the therapeutic solution can be injected into the myocardium (see Note 18). 11. Before retracting the needle, wait an additional 5 seconds after the injection to prevent any backflow of the injectant (see Note 19). 12. Perform all the injections as described previously, and retract the catheter from the left ventricle. 3.7 Removing Femoral Sheath

1. Remove the femoral sheath, and apply pressure on the puncture site to prevent bleeding and hematomas. 2. Monitor the patient’s vital signs as necessary.

4

Notes 1. NogaStar and MyoStar catheters have four different curve options annotated B, C, D, and F in increasing size. We use D-curve for pigs. 2. Albumin coating of MyoStar catheter may be needed for some therapeutic agents, such as adeno-associated viruses (AAVs), as it to our experience prevents undesired binding to the catheter. 3. We use a small x-ray-positive paper clip in the reference location patch, making it easier to confirm that the location is in the plane with the heart. 4. After this step, the patient must remain in the same place during the procedure. 5. Operated animals must be under general anesthesia and ventilation. 6. It is easier to get through the aortic valve when the catheter tip is slightly curved. 7. Colors help to guide the catheter in the left ventricle as the catheters flex toward the red/yellow color. 8. Mapping can cause arrhythmias, such as ventricular tachycardia, possibly leading to ventricular fibrillation. For that reason, a defibrillator must be available. Gentle catheter movements reduce the risk of these events. 9. Our experience suggests that clicking points manually will lead to a better outcome than taking points automatically because invalid points can be rejected immediately. 10. Reject the point if LLS is 0%. 11. To ensure good quality and a reliable map, exclude all invalid points. 12. Remember to check also the other half of the left ventricle.

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13. The red area indicates non-viable scar tissue. 14. Our experience suggests that the needle length varies along the curvature used. 15. It is recommended to use the curved plastic model from the catheter package around the catheter, as it models the curvature of the aorta and thus helps your needle adjustment. 16. Select a smaller syringe for easier injections. 17. Needle length and placement should be verified with fluoroscopy. To our experience, the needle length is best seen in the angle RAO 90. 18. It is recommended to perform injections slowly, for example, 0.1 mL/15 s. We administer 10 injections, 0.2 mL each on the previously determined injection site. 19. Keep a light pressure on the syringe.

Acknowledgments This study was supported by Finnish Academy Flagship grant. References 1. James SL, Abate D, Abate KH et al (2018) Global, regional, and national incidence, prevalence, and years lived with disability for 354 diseases and injuries for 195 countries and territories, 1990-2017: a systematic analysis for the Global Burden of Disease Study 2017. Lancet 392:1789–1858. https://doi. org/10.1016/S0140-6736(18)32279-7 2. Neumann FJ, Sechtem U, Banning AP et al (2020) 2019 ESC guidelines for the diagnosis and management of chronic coronary syndromes. Eur Heart J 41:407–477 3. Kieserman JM, Myers VD, Dubey P et al (2019) Current landscape of heart failure gene therapy. J Am Heart Assoc 8. https:// doi.org/10.1161/JAHA.119.012239 4. Psaltis PJ, Worthley SG (2009) Endoventricular electromechanical mapping-the diagnostic and therapeutic utility of the NOGA XP Cardiac Navigation System. J Cardiovasc Transl Res 2:48–62. https://doi.org/10.1007/ S12265-008-9080-7 5. Duran JM, Taghavi S, Berretta RM et al (2012) A characterization and targeting of the infarct border zone in a swine model of myocardial infarction. Clin Transl Sci 5:416. https://doi. org/10.1111/J.1752-8062.2012.00432.X 6. Yee K, Malliaras K, Kanazawa H et al (2014) Allogeneic Cardiospheres delivered via

percutaneous Transendocardial injection increase viable myocardium, decrease scar size, and attenuate cardiac dilatation in porcine ischemic cardiomyopathy. PLoS One 9. https://doi.org/10.1371/JOURNAL. PONE.0113805 7. Garbayo E, Gavira JJ, de Yebenes MG et al (2016) Catheter-based Intramyocardial injection of FGF1 or NRG1-loaded MPs improves cardiac function in a preclinical model of ischemia-reperfusion. Sci Rep 6. https://doi. org/10.1038/SREP25932 8. J€arvel€ainen N, Halonen P, Nurro J et al (2021) Citrate-saline formulated mRNA delivery into the heart muscle with an electromechanical mapping and injection catheter does not lead to therapeutic effects in a porcine chronic myocardial ischemia model. Hum Gene Ther. https://doi.org/10.1089/HUM.2021.149 9. L€ahteenvuo JE, L€ahteenvuo MT, Kivel€a A et al (2009) Vascular endothelial growth factor-B induces myocardium-specific angiogenesis and arteriogenesis via vascular endothelial growth factor receptor-1- and neuropilin receptor-1dependent mechanisms. Circulation 119: 8 4 5 – 8 5 6 . h t t p s : // d o i . o r g / 1 0 . 1 1 6 1 / CIRCULATIONAHA.108.816454 10. Rutanen J, Rissanen TT, Markkanen JE et al (2004) Adenoviral catheter-mediated

Endocardial Gene Delivery Using NOGA Catheter System Intramyocardial gene transfer using the mature form of vascular endothelial growth factor-D induces transmural angiogenesis in porcine heart. Circulation 109:1029–1035. https:// doi.org/10.1161/01.CIR.0000115519. 03688.A2 11. Vale PR, Losordo DW, Milliken CE et al (2001) Randomized, single-blind, placebocontrolled pilot study of catheter-based myocardial gene transfer for therapeutic angiogenesis using left ventricular electromechanical mapping in patients with chronic myocardial ischemia. Circulation 103:2138–2143. https://doi.org/10.1161/01.CIR.103.17. 2138 12. Kastrup J, Jørgensen E, Ru¨ck A et al (2005) Direct intramyocardial plasmid vascular endothelial growth factor-a 165 gene therapy in patients with stable severe angina pectoris: a randomized double-blind placebo-controlled study: the Euroinject one trial. J Am Coll Cardiol 45:982–988. https://doi.org/10.1016/j. jacc.2004.12.068 13. Ripa RS, Wang Y, Jørgensen E et al (2006) Intramyocardial injection of vascular endothelial growth factor-a 165 plasmid followed by granulocyte-colony stimulating factor to induce angiogenesis in patients with severe chronic ischaemic heart disease. Eur Heart J 27:1785–1792. https://doi.org/10.1093/ eurheartj/ehl117 14. Fuchs S, Dib N, Cohen BM et al (2006) A randomized, double-blind, placebo-controlled, multicenter, pilot study of the safety and feasibility of catheter-based intramyocardial injection of AdVEGF121 in patients with refractory advanced coronary artery disease. Catheter Cardiovasc Interv 68:372–378. https://doi.org/10.1002/CCD.20859

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15. Stewart DJ, Kutryk MJB, Fitchett D et al (2009) VEGF gene therapy fails to improve perfusion of ischemic myocardium in patients with advanced coronary disease: results of the NORTHERN trial. Mol Ther 17:1109–1115. https://doi.org/10.1038/mt.2009.70 16. Kukuła K, Chojnowska L, Dabrowski M et al (2011) Intramyocardial plasmid-encoding human vascular endothelial growth factor A165/basic fibroblast growth factor therapy using percutaneous transcatheter approach in patients with refractory coronary artery disease (VIF-CAD). Am Heart J 161:581–589. https://doi.org/10.1016/j.ahj.2010.11.023 17. Hedman M, Hartikainen J, Syv€anne M et al (2003) Safety and feasibility of catheter-based local intracoronary vascular endothelial growth factor gene transfer in the prevention of postangioplasty and in-stent restenosis and in the treatment of chronic myocardial ischemia: phase II results of the Kuopio angiogenesis trial (KAT). Circulation 107:2677–2683. h t t p s : // d o i . o r g / 1 0 . 1 1 6 1 / 0 1 . c i r . 0000070540.80780.92 18. Hartikainen J, Hassinen I, Hedman A et al (2017) Adenoviral intramyocardial VEGFDDNDC gene transfer increases myocardial perfusion reserve in refractory angina patients: a phase I/IIa study with 1-year follow-up. Eur Heart J 38:2547–2555. https://doi.org/10. 1093/eurheartj/ehx352 19. Gyo¨ngyo¨si M, Dib N (2011) Diagnostic and prognostic value of 3D NOGA mapping in ischemic heart disease. Nat Rev Cardiol 8(7): 393–404. https://doi.org/10.1038/nrcardio. 2011.64

Chapter 15 Surgical Methods for Cardiac Gene Delivery in Large Animals Michael G. Katz , Yoav Hadas, Adam S. Vincek, Nataly Shtraizent, Hylton P. Gordon, Peter Pastuszko, Eric Schadt, and Efrat Eliyahu Abstract This chapter describes main strategies of surgical gene delivery in large animals. Existing methods of cardiac gene transfer can be classified by the site of injection, interventional approach, and type of cardiac circulation at the time of transfer. Randomized clinical trials have suggested that the therapeutic benefits of gene therapy are not as substantial as expected from animal studies. This discordance in results is largely due to gene delivery methods that may be effective in small animals but are not scalable to larger species and, therefore, cannot transduce a sufficient fraction of myocytes to establish long-term clinical efficacy. Ideally, an optimized gene transfer should incorporate the following: a closed-loop recirculation for extended transgene residence time; vector washout form the vascular system after transfer to prevent collateral expression; use of methods to increase myocardial transcapillary gradient for viral particles for a better transduction, probably retrograde route of gene delivery through the coronary venous system; and myocardial ischemic preconditioning. Key words Cardiac gene therapy, Gene delivery, Cardiopulmonary bypass, Intramyocardial administration, Cardiovascular diseases

1

Introduction Cardiac gene delivery received a strong inspiration to its development after the surgical direct intramyocardial injection of β-galactosidase/plasmid DNA construct into the left ventricle of beating rat hearts. Gene activity and myocyte expression were demonstrated 4 weeks after transfer [1]. These results showed for the first time that cardiac muscle has a unique ability to take up and express injected recombinant DNA without recipient cell division. The next historical step was the proof that viral-mediated gene transfer after intramyocardial injection is thousands of times more efficient than plasmid DNA [2]. Subsequent development involved

Kiyotake Ishikawa (ed.), Cardiac Gene Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2573, https://doi.org/10.1007/978-1-0716-2707-5_15, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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intracoronary infusion of adenoviral vector. Viral DNA was detected in the animal’s myocardium for 2 weeks after delivery [3]. Effective delivery solutions will be essential to advancing cardiovascular gene therapy efforts. Critical elements of a delivery system consist of the following: (i) selectively targeting tissues of therapeutic relevance and (ii) minimizing systemic effects [4, 5]. The development of safe and efficient delivery systems is a prerequisite for the translation from the experimental phase into clinical trials of a target that has been validated in pre-clinical models [6, 7]. It should be noted that a wide variety of techniques have already been designed and applied for cardiovascular applications [8, 9]. Unfortunately, most of these approaches have led to limited transfection and limited transduction efficiency, with a moderate-to-high incidence of systemic exposure [10–12]. Existing methods of gene delivery can be classified by the site and method of injection, interventional approach, and the variations of heart perfusion during transfer [13]. These and other data showed that the route of gene delivery are no less important than the choice of vector system, and the basic methods of cardiac surgical gene transfer were primarily intramyocardial injection and intracoronary administration with cardiopulmonary bypass (CPB) [10]. Direct intramyocardial techniques are very attractive and currently being used due to the fact that they are simple and reproducible, allow for targeting of specific areas of interest, and achieve high local vector concentration. Application of this method has resulted in successful therapeutic angiogenesis and focal arrhythmia therapy. However, a key limitation of this method includes the fact that transgene expression is limited to the injection sites so the method is therefore not sufficient to achieve a global distribution. The benefits of intracoronary delivery include the possibility of repeated vector deliveries to the whole myocardium with homogenous gene expression and minimal invasiveness. However, the limited cardiac transduction and varied results with systemic leakage leading to significant collateral organ uptake led researchers to identify parameters influencing the efficiency of intracoronary transfer. These parameters include the contact time of vector in coronary circulation, intravascular flow rate and perfusion pressure, composition of perfusate, and endothelial permeability. Limitations of intramyocardial and intracoronary delivery encouraged researchers to develop closed-loop recirculatory systems, which allowed separation of the coronary bed from systemic circulation. The principal strength of this technology includes a significant increase in transduction efficiency, the extension of vector residence time, the ability to manipulate endothelial permeability, the avoidance of an immune response to the vector, and the ability to wash out the vector post gene delivery limiting collateral organ exposure. Despite the attractiveness of cardiopulmonary bypass–mediated gene delivery approaches, it should be noted that one cannot

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exclude possible attendant morbidity, which may be related to technique-associated complications and the additional crossclamp time required.

2

Materials

2.1 Anesthesia and Pre- and Perioperative Medications

1. Intravenous (IV) ketamine 10 mg/kg. 2. IV midazolam 0.2 mg/kg. 3. Intramuscular (IM) buprenorphine 0.01 mg/kg. 4. Chlorhexidine scrub and solution. 5. Isoflurane. 6. Fentanyl patch, 75 mcg patch. 7. Eye lubricant. 8. Lidocaine gel. 9. Enrofloxacin (Baytril) 5 mg/kg. 10. IM buprenorphine 0.01 mg/kg. 11. IM morphine 5 mg. 12. IM banamine 1 mg/kg. 13. IM glycopyrrolate 0.02 mg/kg. 14. Anesthesia mask. 15. Laryngoscope. 16. Endotracheal tubes #7–8 Fr. 17. Pulse oximeter. 18. Nasal cannula for large animals. 19. Balloon with oxygen. 20. Triple lumen catheter for central venous pressure (CVP) and IV administration. 21. Thermometer. 22. Rumen tube. 23. Heating pad. 24. ECG pads.

2.2 Direct Intramyocardial Gene Delivery

1. Items in Subheading 2.1, items 1–13. 2. Surgical set: rib spreaders, blades, knife handle, army navy retractor, DeBakey forceps, Mayo and Metzenbaum scissors, regular hemostats, mosquito hemostats, needle holders, Kelly clamps, Schnidt hemostats. 3. Sutures: #2-0 silk pop-off on taper needle, #4-0 Prolene on taper needle, #0 silk ties, #1 vicryl on taper needle, #2 vicryl on taper needle.

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4. U-100 syringe, needle size 28G1/2. 5. 12-lead electrocardiogram machine. 6. Arterial line monitor. 7. Connection apparatus and pressure tubing. 8. 24–28 Fr angle chest tubes. 9. Pleur-evac. 10. Electric cautery. 11. Ringer lactate solution. 12. Ultrasound disposable sterile bags. 2.3 Surgical Gene Delivery with Heart– Lung Machine

1. Personnel: surgeon, assistant of surgeon, anesthesiologist, and cardiovascular perfusionist (operates a heart–lung machine). 2. 1–2 units of blood (see Note 1). 3. Items in Subheadings 2.1, items 1–13 and 2.2, item 2. 4. Sutures: #2-0 silk pop-off on taper needle, #4-0 Prolene on taper needle, #4-0 nonpledgeted Prolene on taper needle, #0 silk ties, #1 vicryl on taper needle, #2 vicryl on taper needle, #20 nylon on cutting needle, and #5 steel sternal wires for sternotomy. 5. Del Nido cardioplegia package. 6. Methylene blue solution. 7. Nitroglycerin solution. 8. Hespan 6%. 9. Solu-Medrol for injection. 10. Benadryl for injection. 11. Epinephrine for injection. 12. Lidocaine 1% for injection. 13. Protamine sulfate 1% for injection. 14. Penicillin G for injection. 15. Gentamicin for injection. 16. Normosol-R solution. 17. Perfusion equipment: 13 Fr retrograde catheter. 18. 26 Fr right angle cannula. 19. 26 Fr straight cannula. 20. 12 and 14 Fr arterial catheters. 21. 9 Fr DLP aortic root cannula. 22. Centrifugal pump. 23. ¼0 siliconized tubing. 24. Bentley bubble oxygenator.

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25. Siliconized polycarbonate connectors. 26. Echocardiogram machine. 27. Subheading 2.2, items 4–8. 28. Heating system. 29. Monitors: intraoperative monitor with ECG, arterial blood pressure, central venous pressure, cardiac output, and rectal temperature. 30. Blood gas analyzer. 31. Cell saver machine. 32. Large animal ICU with postoperative recovery: continuous oxygen supply, vacuum system for chest drainage, ECG, and infusion pump with 2–3 lines.

3

Methods

3.1 Preoperative Care and Preparation

1. Anesthetize the sheep with 10-mg/kg ketamine and 0.2-mg/ kg midazolam, delivered IV (see Note 2). 2. Transport the sheep from the colony to the preparation room and weigh. 3. Maintain anesthesia by isoflurane gas administration via mask. 4. Position the sheep in sternal recumbency on the preparation table. 5. Intubation: Using a long straight-edge laryngoscope and endotracheal suction (for clearance of secretions where necessary), visualize the vocal cords and insert the deflated, regularcuffed 8 Fr endotracheal tube (or alternate size depending upon sheep’s laryngeal space and vocal cord aperture) lubricated with viscous lidocaine, into endotracheal passage between vocal cords. Inflate the balloon cuff with 20 mL of air. 6. Confirm tube placement via auscultation, and connect the endotracheal tube to the mechanical ventilation apparatus with 2.5–3.5% isoflurane. The depth of anesthesia is monitored by vital signs, pupillary response to light, and response to physical manipulation. 7. Insert the rumen tube, lubricated with lidocaine gel, by feeding the tube gently, but firmly, over the base of the tongue into the esophagus and down into the rumen tract. 8. Administer 0.01-mg/kg buprenorphine IM, 0.02 mg/kg of glycopyrrolate IM, 1-mg/kg banamine IM, and 2.2-mg/kg naxcel IM. 9. Place fentanyl 75-mcg patch onto the groin area, and lubricate eyes with lacri-lube.

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Fig. 1 Direct intramyocardial gene delivery into the left ventricle

10. Clip the sheep, and cleanse the surgical areas with chlorhexidine scrub followed by chlorhexidine solution. 11. Place ECG pads on clipped forelimbs, and secure in place with silk tape. 3.2 Direct Intramyocardial Gene Delivery (See Fig. 1)

1. Connect the animal to the anesthesia machine with 1.5–2.5% isoflurane. 2. Connect the ECG leads to the ECG machine, and connect supportive IV lines. 3. Using the Seldinger technique, place a 7 Fr triple lumen catheter in the left jugular vein, which can be used intraoperatively for medication delivery and central venous pressure monitoring. 4. Place the animal in the supine position. Using sterile technique with a 7 Fr catheter, place a carotid or femoral arterial line for invasive blood pressure monitoring and Millar catheter measurements. 5. Position the animal in a right lateral decubitus position. After cleaning and disinfection using chlorhexidine scrub and iodine solution, drape the left chest in sterile fashion. 6. Enter the chest via a 5–7-cm left mini-thoracotomy incision at the IV–V intercostal space, reflect the left lung posteriorly, and incise and reflect the pericardium superiorly and laterally.

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7. Identify the area of gene delivery (see Note 3). 8. Perform multiple injections of gene construct into the wall of the left ventricle with U-100 syringe, needle size 28G1/2, volume 0.1 mL for each injection. 9. Inspect injected places for residual hemorrhage. 10. Flash the pericardial space with Ringer lactate three–five times to prevent future adhesions. Loosely approximate the pericardium with 3-0 silk. 11. Place a 28 Fr thoracostomy angle tube, and connect to low, continuous wall suction via a pleur-evac for 30–60 min. The thoracotomy incision is repaired by staged chest closure. Discontinue the thoracostomy tube, and extubate the animal in sequential fashion. Finally, transport the animal to a recovery room for postoperative care. 3.3 Gene Delivery Using Heart–Lung Machine Including Method of Molecular Cardiac Surgery with Recirculating Delivery (See Fig. 2)

1. Follow Subheading 3.1. 2. Position the sheep in supine position. After cleaning and disinfection, drape the sheep in sterile fashion from the neck to the knees. Place sterile large drapes, and cut holes over the chest, right groin, and right carotid artery. 3. Surgically expose the right femoral artery via cut-down procedure. Insert a 16G angiocatheter into the artery, and connect to the arterial line apparatus and monitor via pressure tubing. 4. Percutaneously cannulate the right brachial artery or femoral artery for blood pressure monitoring using arterial line kit. Once the arterial line is placed and appropriate waveform pressure is confirmed, secure the line using a 2-0 silk pop-off suture. 5. Expose the right carotid artery via cut-down procedure as it was done in Subheading 3.3, step 4 for introduction of the carotid catheterization to facilitate cardiopulmonary bypass. 6. Incise the chest skin till sternum. Advance the bone saw onto the field, and proceed with creating the median sternotomy, starting superiorly at the level of the sternal notch and proceeding distally to at least two fingerbreadths above the inferior aspect of the sternal plate, taking care not to divide the xiphoid. Open the chest, and control sternal bleeding with bone wax. 7. Reflect the lung parenchyma, and enter the pericardium, exposing the midline-positioned cardiac window ~5 cm. Place three pericardial stay sutures. 8. Perform a transepicardial echocardiographic examination to assess initial function of the heart (pre-cardiopulmonary bypass period) using sterile echo probe bag.

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Fig. 2 Cardiopulmonary bypass–mediated gene delivery. (a) Gene delivery via conventional cardiopulmonary bypass. (b) Gene transfer via cardiopulmonary bypass with complete cardiac isolation and recirculation (MCARD)

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9. Place a pledgeted #4-0 Prolene purse string around the right atrial appendage for superior vena cava (SVC) cannulation. Doubly place two #0 silk heavy sutures around the superior vena cava, and connect to tourniquets. Ensnare the ascending aorta using umbilical tapes. Place a purse string on the right atrium adjacent to the atrioventricular groove. This will become the cannulation site for the coronary sinus catheter (see Note 4). 10. Place one other purse string, on the inferior vena cava (IVC). 11. Give heparin (130 U/kg), and check active coagulation time (ACT). Ensure ACT is greater than 400 before proceeding. 12. Cannulate the right carotid artery using a 14 Fr cannula. 13. Using pericardial pledgets and #4-0 Prolene suture, a horizontal mattress, place a pledgeted suture on the ascending aorta, approximately 1 cm distal to the aortic root. Place the aortic cannula. 14. Cannulate the superior vena cava using a 26 French right-angle cannula. 15. Place the retrograde catheter into the coronary sinus via the right atrium at atrioventricular junction (see Note 5). 16. Connect the SVC cannula to a Y connector (with one limb clamped), and connect to the venous limb of the pump circuit. Bring the inflow lines from the coronary reservoir circuit and outflow lines up to the table. Recirculate the coronary reservoir solution through a stopcock with care to avoid entry of the solution into the systemic circulation or onto the field. 17. Ligate the right azygos (hemiazygos) veins with a 2-0 silk suture making sure there are no other veins draining into the SVC (see Note 6). 18. Initiate partial cardiopulmonary bypass (~50%) with one cannula in superior vena cava. Cannulate the IVC using a 26 Fr right-angle cannula. 19. Full cardiopulmonary bypass is initiated. Cool the heart to 30 degrees with bypass heat exchanger (see Note 7). 20. Doubly snare the IVC with a double loop of #0 silk suture. 21. The heart begins to fibrillate. Reduce aortic flow. 22. Cross clamp the aorta. Give 15-mL/kg del Nido cardioplegia via the aortic root cannula in an antegrade fashion. 23. Place a purse string #4-0 Prolene suture in the apex of the left ventricle. 24. Make a stab in the middle of the purse string, and place a cannula into the left ventricular cavity and clamp.

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25. Place a pledgeted purse string #4-0 Prolene suture in the right ventricular outflow tract. Then, place a cannula into the right ventricle, and clamp and snare the purse string. 26. The isolated cardiac circuit for the gene recirculation is constructed. 27. Give a second dose of cardioplegia 15 mL/kg via the aortic root cannula in an antegrade fashion. 28. Isolate the coronary circuit by tightening the vena caval snares. Infuse the coronary sinus catheter with only 3 mL to avoid rupture of the coronary sinus. 29. Connect the cardiac inflow limb to the retrograde catheter. 30. Inject methylene blue solution into the coronary sinus to confirm good cardiac isolation. Sinus pressure usually must be kept less than 80 mmHg with driving pressure of 100–110 mmHg. 31. Inject the nitroglycerin diluted in 10-mL normal saline into the coronary sinus. 32. Wait for 2 minutes. Then, inject 5 mL of 1014 gc gene construct solution to the coronary sinus. 33. Restore flow over one minute until coronary sinus pressure equals 90–100 mmHg, and recirculate the remaining reservoir solution for another 15 minutes. 34. Once the 15-minute recirculation interval is done, decompress the heart. 35. Remove the coronary sinus catheter. 36. The coronary circuit is flushed antegrade with approximately 1000 mL of hespan infused with 100 mg of Solu-Medrol, 50 mg of Benadryl. Decompress the heart again (into the yellow vent), and discard excess volume. 37. After flow has been reduced, remove the aortic and pulmonary cross clamps, and restore flow. 38. Initiate rewarming during 20 minutes. 39. Defibrillate the heart to restore normal sinus rhythm. 40. Remove the IVC and SVC cannulae. 41. Administer epinephrine at 1–2 mcg/min. Administer a 50-mg lidocaine bolus and an infusion at 1 mg/min. Once the heart is contracting well, remove the IV cannula. 42. Wean from bypass after 20 minutes of reperfusion. Once off bypass, give protamine and check ACT. 43. After the sheep has been weaned from bypass, obtain another trans-epicardial echocardiogram to grossly determine heart function after bypass.

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44. Place bilateral chest tubes to get rid of pneumothorax, and remove these 24–48 hours postoperatively. The thoracostomy tubes should subsequently be connected to wall suction through a pleur-evac. 45. Start using cell saver machine by connecting to the heart–lung machine to collect the blood loss during surgery. Place topical hemostatic agents to stop any bleeding. Close the sternal incision with seven #2-0 steel wires. Two wire sutures are placed around the manubrium, and five are placed around the edges of the body of the sternum. 46. Repair subcutaneous and skin layers with #1 vicryl and #20 nylon suture, respectively. Remove the femoral arterial line, and repair the artery, primarily using #7-0 Prolene. 47. When extubating the animal, care is taken to observe jaw motion, eye movement, and axial motion. Position the animal in sternal recumbency to prevent aspiration and to encourage recovery. Transfer the sheep to the postoperative care area once the animal has been taken off the ventilator (see Note 8). 3.4

Postoperative

1. Survival in large animal surgery with CPB depends highly on the quality of immediate critical care. Appropriate monitoring provides valuable information on the animal’s viability (see Note 9). We have created a postoperative protocol (see Table 1). A key aspect of this protocol is the constant supervision of the animal for the first 24 h after surgery. 2. After the animal recovers from anesthesia, extubate. 3. In the recovery room, place a nasal cannula to the animal’s rostrum, and connect the cannula to humidified oxygen. Initiate oxygen supplementation at 5 L/min until the animal is active and able to maintain a pulse oximetry value of 94% or greater. 4. Monitor the animal’s physiologic parameters, beginning with measurement of its temperature, pulse, respirations, heart rate, oxygen saturation, blood gases, etc.—obtained every 2–4 h— and document in the animal record. 5. Monitor central venous pressure (CVP) via triple lumen catheter in the jugular vein with a CVP manometer, and IV administration daily, which may remain in place and functional for up to 3 days. CVP monitoring (target, 4–10 mm H2O) provides important information regarding blood volume, which may allow for life-saving interventions with Lasix or fluids. 6. Obtain an arterial blood gas sample via the auricular artery of either ear, if possible, every 2 to 4 h for the first 12 h to evaluate acid–base level and electrolytes––especially potassium, calcium, and magnesium. Correct metabolic acidosis or alkalosis,

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Table 1 Postoperative care protocol: first 24–48 hours Monitoring

Medication

Fluids and feed

Vital signs every 4 hours SpO2 monitoring 24 hours CVP every 4 hours ECG every 8 hours Blood gases every 4 hours during the first day Chest tube drainage every 1 hour during the first day Urination monitoring Liver enzymes Kidney function CBC chemistry after 24 hours

Nasal O2 5 L/min, until stable Antibiotics (penicillin, gentamicin) Furosemide if CVP > 10-cm H 2O K+, ca+, and mg+ correction HCO3 if metabolic acidosis Inotropes if low cardiac output Normosol-R 50–80 ml/hr. if CVP < 4-cm H2O

Hetastarch IV first 24 hours Normosol-R IV first 48 hours Water when alert, feed after 6 hours post surgery

Postoperative care protocol in large animals after gene delivery with cardiopulmonary bypass Abbreviations: SpO2 blood saturation, CVP central venous pressure, ECG electrocardiography, K+, Ca+, Mg+ electrolytes, CBC complete blood count

hypokalemia, hypocalcemia, or hypomagnesemia with IV supplementation. In the absence of arterial access, use venous samples from the jugular CVP catheter instead. 7. Rewarm the animal with warming blanket to prevent hypothermia (>37  C). 8. Our antibiotic regime includes procaine penicillin G 22,000 IU/kg IV and gentamicin 6.6 mg/kg IV, given 1 h prior to incision and postoperatively daily for 5 days. 9. In the early postoperative period, we administer a crystalloid Normosol-R solution at a rate of up to 100 mL/h depending on CVP. 10. Perform laboratory analyses of blood count, electrolytes, and liver and kidney function daily for 3 days after surgery and whenever animals appear weak. 11. Assessment of pain in the postoperative period is very important. For pain care, buprenorphine and a fentanyl patch will be used for the first 3 days with the dose adjusted based on the animal’s activity and vitals. Give the first dose of each after induction of anesthesia. Place the fentanyl patch of 75 mcg following clipping. At this time, 2–5 mcg/kg buprenorphine will be given IV. The following signs will be used for pain distress: poor oral intake, increased heart rate/tachypnea, reluctance to move, pawing at the surgical site and inability to stand in the recumbent position, and hyperesthesia. 12. Observe and record urine and excrement production and chest thoracostomy tube output (see Note 10).

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13. Remove the thoracostomy tubes 24–36 h postoperatively. The proper use of thoracostomy tubes helps reduce the incidence of pleural effusions. However, delayed pleural effusions may still occur, requiring insertion of new thoracostomy tubes for 24 h. 14. Observe each sheep daily for evidence of surgical site infection, pain, and cardiovascular or respiratory compromise. This observation includes a physical exam, observation of the surgical site; assessment of attitude, activity, appetite, and fecal production; and measurement of temperature, pulse rate, respiratory rate, thoracic auscultation, and oxygen saturation via pulse oximetry. When necessary, perform blood pressure measurements, electrocardiography, or echocardiography (see Note 11). 15. Begin nourishment with water when the sheep is alert, and hay may be given when the animal is standing but neither is offered prior to 6 hr. postoperatively (see Note 12).

4

Notes 1. Since bleeding in cardiopulmonary bypass cases can occasionally be rapid, unpredicted, and life threatening, 1–2 units of blood from donor sheep should be used routinely. At the end of bypass, if the hematocrit is less than 15%, one should transfuse the blood. 2. All animal studies must follow the National Institute of Health guidelines (Guide for the Care and Use of Laboratory Animals, NIH publication No. 85–23, revised 1996) and be approved by the appropriate Institutional Animal Care and Use Committee. This protocol included sheep: Dorset male sheep, 0.8–1.4 years old, and weighing 35–55 kg. 3. The ideal sites for intramyocardial injection should be safe and effective. The sites included the left ventricle territory, anterior and posterolateral wall, and lateral to the left anterior descending artery. The number of injections is ~8–10. 4. During the administration of heart–lung machine, attention should be devoted to the cardioplegia purse string in the aortic root because large animals have a very short ascending aorta. 5. For retrograde vector infusion to the coronary sinus, care must be taken to confirm the proper catheter position. Placement is confirmed with injection of methylene blue and its distribution throughout the cardiac veins. 6. For complete cardiac isolation to prevent systemic loss of viral particles, azygos and hemiazygos veins should be ligated.

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7. Continuous monitoring of heart functions is important because of the possibility of hemodynamic instability. 8. When extubating the animal, care is taken to observe jaw motion, eye movement, and axil motion. The animal is positioned in sternal recumbency to prevent aspiration and to encourage recovery. 9. During pre- and post-bypass period and intramyocardial injection of viral particles, anesthesiologist should monitor the animal’s hemodynamic stability to avoid systemic vasodilatation with afterload reduction, tachycardia, and attempt to preserve sinus rhythm. 10. If postoperative blood loss from the chest thoracostomy tubes is greater than 100 mL, ACT and hematocrit should be checked. One must then consider a surgical source of bleeding. 11. After euthanasia as quickly as possible to avoid DNA, RNA, and protein degradation, cut a cube of tissue around 7–8-mm edge length in half so that it is no more than 5 mm thick. Fill cryotubes with the tissue, and store in the dry ice container until the completion of the tissue procurement. Store the samples in 80  C freezer until needed for future analysis. 12. Cardiopulmonary bypass featuring complete heart isolation and continuous cardiac perfusion is a very promising approach for solving the problem of efficient gene delivery. This system permits continuous isolated arrested heart perfusion through optimizing a number of delivery parameters including temperature, flow rate, driving pressure, ionic composition, and exposure time to the cardiac vessels. During complete cardiac isolation, the blood vector concentration was stable (see Fig. 3). After the completion of the recirculation interval and subsequent washing procedure, theinitially non-detectable systemic blood genome copy concentration significantly increased.

Fig. 3 Blood vector concentration during and after recirculation with cardiopulmonary bypass–mediated gene delivery gene delivery

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Acknowledgments We would like to thank Drs. Bridges CR, Fargnoli AS, and Gubara SM, all veterinarians and veterinary technicians for the help. References 1. Lin H, Parmacek MS, Morle G et al (1990) Expression of recombinant genes in myocardium in vivo after injection of DNA. Circulation 82:2217–2221 2. Guzman RJ, Lemarchand P, Crystal RG et al (1993) Efficient gene transfer into myocardium by direct injection of adenovirus vectors. Circ Res 73:1202–1207 3. Barr E, Carroll J, Kalynych AM et al (1994) Efficient catheter-mediated gene transfer into the heart using replication-defective adenovirus. Gene Ther 1:51–58 4. Ishikawa K, Tilemann L, Ladage D et al (2012) Cardiac gene therapy in large animals: bridge from bench to bedside. Gene Ther 19: 670–677 5. Katz MG, Fargnoli AS, Williams RD et al (2015) Cell and gene therapies for cardiovascular disease. In: Templeton NS (ed) Gene and cell therapy. CRC Press, New York, pp 861–901 6. Ishikawa K, Weber T, Hajjar RJ (2018) Human cardiac gene therapy. Circ Res 123:601–613 7. Katz MG, Fargnoli AS, Weber T et al (2017) Use of adeno-associated virus vector for cardiac gene delivery in large-animal surgical models of

heart failure. Hum Gene Ther Clin Dev 28: 157–164 8. Ishikawa K, Tilemann L, Fish K et al (2011) Gene delivery methods in cardiac gene therapy. J Gene Med 13:566–572 9. Katz MG, Fargnoli AS, Williams RD et al (2013) Gene therapy delivery systems for enhancing viral and nonviral vectors for cardiac diseases: current concepts and future applications. Hum Gene Ther 24:914–927 10. Hajjar RJ, Ishikawa K (2017) Introducing genes to the heart: all about delivery. Circ Res 120:33–35 11. Hulot JS, Ishikawa K, Hajjar RJ (2016) Gene therapy for the treatment of heart failure: promise postponed. Eur Heart J 37: 1651–1658 12. Katz MG, Fargnoli AS, Kendle AP et al (2016) Gene therapy in cardiac surgery: clinical trials, challenges, and perspectives. Ann Thorac Surg 101:2407–2416 13. Katz MG, Swain JD, White JD et al (2010) Cardiac gene therapy: optimization of gene delivery techniques in vivo. Hum Gene Ther 21:371–380

Chapter 16 Atrial Gene Painting in Large Animal Model of Atrial Fibrillation Weilan Mo and J. Kevin Donahue Abstract Gene therapy appears promising as a targeted treatment of cardiac diseases. Atrial fibrillation (AF) is the most common sustained cardiac arrhythmia and also a major contributor to stroke, heart failure, and death. Mechanisms that initiate and sustain AF are associated with structural and electrophysiological remodeling in the whole atria. Selection of the appropriate gene delivery method is critical for transduction efficacy. The ideal gene delivery method to manage AF should provide widespread and sufficient exposure to the transgene in atria only that safely maintains the homeostasis of the heart without off-target expression. All these requirements can be achieved using atrial gene painting that is directly applied to the atrial epicardial surface. In this chapter, we present the advantages of atrial gene painting and the experimental method, as applied to a large animal model of AF. Key words Gene painting, Atrial fibrillation, Large animal model, Cardiac arrhythmias, Gene therapy, Gene delivery method, Epicardial approach, Viral vectors

1

Introduction Atrial fibrillation (AF) is the most common sustained cardiac arrhythmia in adults and is associated with an increased incidence of stroke, heart failure, and death [1]. The occurrence of AF facilitates the continuation and recurrence of AF (“AF begets AF”) [2]. When AF has continued for a long time, AF management and restoration of sinus rhythm become more difficult [2]. Approximately 30% of AF cases are paroxysmal AF that stops within 7 days, whereas 70% cases are persistent (more than 7 days) or longstanding persistent AF (greater than 12 months) with higher AF burden that aggravate risks of stroke and mortality [3, 4]. Molecular studies point that both initiation and maintenance of AF are strongly related to structural and electrophysiological remodeling in the atria that creates arrhythmic substrates [1, 2, 5, 6].

Kiyotake Ishikawa (ed.), Cardiac Gene Therapy: Methods and Protocols, Methods in Molecular Biology, vol. 2573, https://doi.org/10.1007/978-1-0716-2707-5_16, © The Author(s), under exclusive license to Springer Science+Business Media, LLC, part of Springer Nature 2022

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The development of gene transfer vectors and delivery systems has given researchers the tools to target specific genes and pathways to regulate structural and electrophysiological abnormalities. Based on molecular mechanisms, transgenes can be delivered to increase or decrease the function of dysregulated targets in the heart. Viral and nonviral methods of gene transfer have been reported [7]. Viral vectors, in general, provide higher efficiency of gene expression and allows for easier delivery modalities [8]. Adenoviruses (Ads) and adeno-associated viruses (AAVs) are the two most commonly used viral vectors for cardiovascular applications [7]. In the heart, Ad vectors can mediate transient but robust transgene expression for 1–2 weeks [9]. It is a useful system for acute, time-limited risk of AF such as postoperative AF or other inflammation or metabolic triggered AF [10]. Although Ad vectors induce an immune response, all of the many clinical trials using Ad vectors on myocardium show no detectable toxicity [11, 12, 25– 34]. AAV has a small insert size for transgene (5 kb) but a favorable safety profile when delivered in reasonable amounts [8]. Another desirable property of AAV vectors is the potentially permanent expression that enables a prolonged treatment for diseases with permanent risk of recurrence, such as AF. Moreover, cardiotropic AAV serotypes (AAV1, 6, 8, and 9) hold promise as the unique platform to treat cardiovascular disease [13]. There is a misperception that the cardiac delivery problem has been solved by AAV, but the significant murine myocardial uptake of these cardiotropic AAVs, e.g., AAV9, is not nearly as effective in the large animal heart [14], pointing to the need for continued attention to physical, pharmacological, or other means of improved gene delivery. Other than transgene and delivery vectors, the method of delivery is a complementary component in determining the level of cardiac gene transfer. This is particularly important when considering the maximal potential of local expression while limiting systemic expression and possibly limiting the vector dose for safety and costs. A number of gene transfer methods have been reported, including intramyocardial injection [15], intracoronary infusion [16, 17], and gene painting [5, 14]. For the management of AF, the ideal delivery method should provide widespread and sufficient exposure to the transgene in the whole atria to achieve the desired on-target effects while minimizing off-target expression. All these requirements can be achieved using atrial gene painting method that directly applied to the atrial epicardial surface. Gene painting is an attractive option for a more efficient and atrial-specific gene delivery at high concentration. Viral vectors are diluted in poloxamer gel before painting. The unique property of poloxamer, being liquid at colder temperature and turning to

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gelatinous at body temperature, allows viral vectors to be mixed before warmed and solidified into the gel during delivery [23]. In cultures of vascular smooth muscle cells, complexing Ad vectors with poloxamer showed a tenfold increase in gene transfer [24]. In our previous studies, poloxamer has been used for Ad vectors to achieve high-density gene transfer [18–20]. To increase virus penetration, trypsin is used for complete transmural gene transfer with the epicardial painting method on the pig heart [18]. We have also seen preclinical efficacy for prevention of transient AF using adenoviral painting without trypsin, which gives epicardial-only gene transfer [10]. This painting method enables whole atrial gene transfer, including access to the avascular areas. Surgical delivery offers direct visual confirmation, allowing precise control of the treatment. Unlike all other reported delivery methods, no evidence of off-target gene expression is found in the ventricles, lungs, liver, spleen, kidneys, gonad, skeletal muscle, or brain [18]. Training requirements are not too demanding and imaging guidance systems such as fluoroscopy, echocardiography, and magnetic resonance imaging are not required. Compared to atrial gene painting, intramyocardial injection can only provide focused gene transfer with limited tissue volume [15]. Although the viral concentration is high at the injection site, it is insufficient to target the sprawling abnormalities of structure and electrophysiology in AF. Multiple injections would be necessary to solve this problem, which raises more problems related to the invasive nature of the injection procedure itself. Perforation at the thinned atrial tissue could be harmful. Tissue inflammation caused by injection may also create substrates for AF [14, 15]. The leakage of the vectors from the injection site can decrease the vector retention and also cause significant off-target exposure on the ventricles or lungs. Unlike the intracoronary infusion method, gene painting bypasses the blood and anatomical barriers. Although it is a wellestablished technique, intracoronary infusion faces a major obstacle that limits it in atrial gene transfer, the absence of an isolated atria vasculature [14]. As balloon blockade of the artery is required in this technique, it also increases the risk of acute ischemia [15]. Moreover, the delivery efficiency of intracoronary infusion has been questioned from the results of the CUPID2b clinical trial. Although this delivery method showed encouraging results in preclinical sheep modeling, it only resulted in 2% of all cardiomyocytes that contained the vector genome that would have been able to express the vector-delivered transgene [21]. In its current iteration, the atrial gene painting method is invasive; it currently needs an open chest surgery for delivery. In clinical settings, this delivery method could potentially be performed during open cardiac surgery or cardiac allografting [22] or through a thoracoscope after modifications [18]. Development

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of “painting” catheters or dedicated thoracoscopic tools would minimize the invasive nature of the procedure. The safety concern of trypsin has been studied carefully in our laboratory. A 0.5% trypsin concentration is an amount sufficient to achieve transmural gene transfer without affecting atrial integrity or function [18]. This method can be used in both small animal models such as rats, rabbits, and large animal models including sheep, pigs, and dogs, and this chapter describes the use on the pig heart for administration of gene painting method.

2 2.1

Materials Preparation

1. TKX mixture solution: 2.2-mg/kg Telazol, 2.2-mg/kg ketamine, 2.2-mg/kg xylazine. 2. Fentanyl patch (100 mcg/hr). 3. Dexamethasone (20 mg). 4. Isoflurane. 5. Antibiotic and antibacterial drugs. 6. A razor and a vacuum. 7. A heating pad and a heart therapy pump. 8. Monitors (vitals and 12-lead ECG). 9. A mechanical ventilator for the pig. 10. Disinfectants: a chlorhexidine scrub, a betadine scrub, and 5% betadine spray.

2.2

Surgery

1. A standard surgical pack for a thoracotomy (see Fig. 1). 2. An electrical cautery. 3. A dispersive electrode. 4. An oscillating saw. 5. A face shield. 6. Vector of interest. 7. Poloxamer F127. 8. Saline. 9. A filter with 0.45-um polyvinylidene difluoride (PVDF) membrane. 10. A pipette and tips. 11. A paintbrush with a round, flat, and soft synthetic tip (tip size: L, 2.5 cm; W, 0.5 cm). 12. A 10-mL glass beaker. 13. A water bath.

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Fig. 1 A standard surgical pack

14. A magnet stirrer. 15. 6 plastic ties. 16. A mediastinal drain tube. 17. Syringes. 18. Needles. 19. A stopcock. 20. 0 silk suture, 2-0 PDSII suture and 3-0 Vicryl suture. 21. Waterproof dressings.

3

Methods

3.1 Poloxamer Saline Solution Preparation

1. Two to three days before surgery, weigh out 1-g poloxamer F127 per 5-mL final volume. 2. In a beaker, first add a stir bar and saline in an amount that is approximately two-thirds of the required final volume. 3. Add the poloxamer F127 to the saline. 4. Place on a stir plate in a 4  C cold room, refrigerator, or similar cold space. 5. Cover the beaker with parafilm, and stir at a moderate rate. 6. Place pipettes, sterile filtration unit, and any necessary tools or equipment in the cold room so that they are cold when needed for use.

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Animal Surgery

1. One day before surgery, the pig receives a fentanyl patch and undergoes overnight fast. 2. On the surgery day, sedate the pig using TKX mixture solution (see Note 1). Shave the surgery site from the neck to the mid-abdomen and across to each armpit. 3. The pig is intubated, ventilated, and placed in dorsal position on a heating mat (see Note 2). Two front arms are tied down (see Note 3). General anesthesia is maintained with isoflurane throughout the procedure (see Note 4). 4. Connect the vital signs monitor (see Note 5) and 12-lead ECG. 5. At this point, everyone in the room should be in surgical attire including hat and mask. All materials and instruments should be kept in sterile fashion throughout the procedure. 6. The surgery site is scrubbed for 5 minutes with chlorhexidine scrub and then 5 minutes with betadine scrub and then sprayed with betadine in preparation for surgery. 7. Create an incision from the manubrium to the xiphoid process of the sternum (see Fig. 2a), and cut the body of the sternum using an oscillating saw (see Note 6). Use the electronic cautery to achieve hemostasis. 8. Spread the chest open (see Note 7). 9. Incise the pericardium carefully to expose the heart (see Fig. 2b and Note 8). 10. Add 0.5% trypsin in 20% poloxamer saline solution, and filter to sterilize at 4  C (see Notes 9, 10, and 11). 11. In the beaker, uniformly mix (see Note 12) the viral vectors (see Note 13) with the solution above to make the painting solution using a magnetic stirrer (see Fig. 3a). 12. Warm the beaker in the 37  C water bath until the painting solution becomes solid (see Fig. 3b and Note 14). 13. Dip the paintbrush (see Note 15) into the solid painting mixture (see Fig. 3c, d), and firmly paint on the epicardial surface of each atrium (see Fig. 3e and Note 16). Manipulate the heart to paint all exposed epicardial surfaces of the atria. Paint two rounds in total. In each round, paint each atrium for 1 minute. A 1-minute break is given to allow absorption before the next round (see Note 17). 14. Expose the heart to air for 10 minutes after gene painting, allowing virus penetration. 15. For closing the chest, administrate dexamethasone into the pericardial space before the chest is closed.

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Fig. 2 (a) Incision sites: a, manubrium; b, xiphoid process. Arrow indicating the moving direction of the saw. (b) Access to the heart after open pericardium

Fig. 3 (a) The liquid painting solution containing viral vectors, 20% poloxamer saline, and 0.5% trypsin. (b) Heating the liquid painting solution in the 37  C water bath. (c) The paintbrush. (d) The gel-like solid painting mixture after warmed. (e) Painting with the gel-like painting mixture on the atrial epicardial surface

16. The sternum is closed with sterile plastic ties placed between and around the opposing ribs so that the sternum is tightly held together (see Fig. 4a). 17. The overlying fascia and skin are closed in three layers: an initial 0 silk layer, a 2-0 PDSII intermediate layer, and then a 3-0 absorbable suture skin layer.

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Fig. 4 (a) Sternum closed with plastic ties. (b) Chest closed and sutured. The thoracic drain tube for air evacuation tunneled into the chest cavity through the incision on the skin. (c) The incision closed after air evacuation. The chest scar covered by clean gauze and dressings

18. Make a ~1-cm incision on the skin of the abdomen. Place the thoracic drain tube, and tunnel it diagonally through the incision to the chest cavity (see Note 18). 19. Evacuate all air from the chest cavity through the thoracic drain tube when chest is closed (see Fig. 4b). 20. Pull the tube out, and the resulted wound is sutured (see Note 19). 21. Clean the surgical site, and apply triple antibiotic ointment. Put a clean gauze pad on the scar before covering it with dressings (see Fig. 4c).

4

Notes 1. For heart failure in animal undergoing gene painting procedure, sedation should be performed using ketamine (20 mg/ Kg) only to avoid respiratory suppression. 2. Typical ventilator settings are intermittent mandatory ventilation at a rate of 8–25 breaths per minute, tidal volume of 10-mL/kg body weight adjusted to keep peak airway pressure